Cellular Migration and Formation of Axons and Dendrites: Comprehensive Developmental Neuroscience [2 ed.] 0128144076, 9780128144077

Table of contents :
Cellular Migration and Formation of Axons and Dendrites
Copyright
Contributors
1. Development of neuronal polarity in vivo
1.1 Introduction
1.2 Axon initiation in vitro versus in vivo
1.2.1 Axon initiation in vitro
1.2.2 Axon initiation in vivo
1.3 Distinction between cues regulating axon specification versus axon growth
1.4 Extracellular cues regulating neuronal polarization and axon initiation
1.4.1 Netrin-1 and Wnt control axon initiation in Caenorhabditis elegans
1.4.2 Polarized emergence of the axon in retinal ganglion cells of Xenopus
1.4.3 Extracellular cues underlying the emergence of axon and dendrites in mammalian neurons
1.5 Intracellular pathways underlying neuronal polarization
1.5.1 Role of protein degradation and local translation in axon specification and axon growth
1.5.2 Role of cytoskeletal dynamics in axon initiation and growth
1.5.3 Major signaling pathways involved in axon initiation and growth
1.5.3.1 LKB1 and its downstream kinases SAD-A/B and MARK1-4
1.5.3.2 PAR3-PAR6-APKC
1.5.3.3 Ras- and Rho-family of small GTPases
1.5.3.4 PI3K and PTEN signaling during axon specification
1.5.3.5 AKT/protein kinase B
1.5.3.6 GSK3 and axon specification
1.6 Conclusion and future directions
References
2. Role of the cytoskeleton and membrane trafficking in axon-dendrite morphogenesis
2.1 Introduction
2.2 Developmental stages
2.3 Role of cytoskeleton in establishment of neuronal polarity
2.3.1 Actin
2.3.2 Actin dynamics during axon formation
2.3.3 Microtubules
2.3.4 Microtubules dynamics during axon formation
2.3.5 Cytoskeletal dynamics during dendritic growth and arborization
2.3.6 Subcellular cytoskeletal specializations
2.4 The role of (membrane) trafficking during neuronal polarization
2.4.1 Trafficking during early neuronal development
2.4.2 Motor protein-based transport in axons and dendrites
2.4.3 The secretory and endosomal pathway
2.4.4 RNA transport and local translation
2.4.5 Barriers for the segregation of functional domains
2.4.6 Protein stabilization and degradation
2.5 Maintaining neuronal polarity
2.6 Future work on neuronal morphogenesis
References
3. Axon growth and branching
3.1 Introduction
3.2 Cell biological mechanisms
3.2.1 Growth cones: structure and function
3.2.2 Regulation of cytoskeleton assembly
3.2.2.1 Actin
3.2.2.2 Microtubules
3.2.3 Interaction between F-actin and microtubules
3.2.4 Membrane trafficking and axonal transport
3.2.5 Protein translation and stability
3.3 Extracellular regulation of axon growth and branching during neural development
3.3.1 Nerve growth factor and neurotrophic factors
3.3.2 Guidance molecules: netrin, slit, semaphorin, ephrin, and wnt
3.3.3 Cell adhesion molecules: permissive or instructive
3.3.4 Glial cells and myelination
3.3.5 Neural activity
3.3.6 Additional axon branching molecules
3.4 Intracellular signaling mechanisms that mediate axon growth and branching
3.4.1 Rho family small GTPases: linking receptors to the cytoskeleton
3.4.2 Calcium
3.4.3 Cyclic nucleotides as second messengers and modulators
3.5 Concluding remarks
References
4. Axon guidance: Netrins
4.1 Introduction
4.2 Netrins and their receptors
4.2.1 Netrin discovery and structure
4.2.2 Netrin receptors
4.2.3 Interactions with other signaling systems
4.2.4 Netrin functional domains and interactions with receptors
4.3 Netrin function in axon guidance and cell migration
4.3.1 Mammalian spinal cord
4.3.1.1 Guidance by midline-derived Netrin-1 in the spinal cord
4.3.1.2 Guidance by ventricular zone-derived Netrin-1 in the spinal cord
4.3.1.3 Synergy between Netrin-1 from floor plate and from ventricular zone in the spinal cord
4.3.1.4 Interpreting the guidance defects caused by loss of Netrin-1 in the spinal cord
4.3.2 Mammalian hindbrain
4.3.2.1 In hindbrain, Netrin-1 from ventricular zone is more important than from floor plate
4.3.2.2 Control of neuronal cell migration by Netrin-1 in the hindbrain
4.3.3 Guidance of other classes of mammalian axons and cells: attraction, repulsion, and modulation
4.3.4 Invertebrate systems
4.3.4.1 Attraction and repulsion by UNC-6 in Caenorhabditis Elegans
4.3.4.2 Attraction and repulsion by Netrins in Drosophila
4.4 Beyond axon and cell guidance: additional roles for Netrins in the nervous system
4.5 Involvement of Netrin signaling in disorders of the nervous system
4.6 Netrins: players outside the nervous system
4.7 Conclusion
References
5. Axon guidance: semaphorin/neuropilin/plexin signaling
5.1 Introduction
5.2 Structural features
5.3 Mechanisms of intracellular signaling
5.4 Function in neural circuit development
5.5 Semaphorins, plexins, and neuropilins in neurological disorders
5.5.1 Autism spectrum disorder
5.5.2 Kallmann's syndrome
5.5.3 Amyotrophic lateral sclerosis
5.5.4 Late-onset neurodegenerative diseases
5.6 Conclusions and perspectives
References
6. Ephrin/Eph signaling in axon guidance
6.1 The setting of the play
6.1.1 Ephs and ephrins
6.1.2 Rules of interaction
6.1.3 Fundamental action modes
6.1.4 Phylogeny
6.2 Mechanisms of ephrin/Eph signaling in axon guidance
6.2.1 Biophysical aspects
6.2.1.1 Membrane distribution
6.2.1.2 Cis interactions
6.2.1.3 Trafficking
6.2.2 Biochemical aspects
6.2.2.1 Signal transduction of forward signaling
6.2.2.2 Signal transduction of reverse signaling
6.3 Ephrins and Ephs in invertebrate axon guidance
6.3.1 Caenorhabditis elegans
6.3.2 Insects
6.4 Binary ephrin/Eph signaling-pathfinding
6.4.1 Peripheral pathfinding-limb bud innervation
6.4.2 Pathfinding in the spinal cord
6.4.3 Pathfinding in the brain stem-auditory system
6.4.4 Central pathfinding
6.4.4.1 Optic chiasm
6.4.4.2 Corpus callosum and anterior commissure
6.5 Proportional ephrin/Eph signaling-mapping
6.5.1 Olfactory wiring
6.5.2 Retinotectal/retinocollicular projection
6.5.2.1 Mechanisms of mapping along the anterior-posterior axis
6.5.2.2 Mechanisms of mapping along the dorsoventral axis
6.5.2.3 Computational modeling
6.5.3 Retinogeniculate projections
6.5.4 Thalamocortical projections
6.5.5 Corticocollicular projections
6.6 Ephrins and Ephs in regeneration
6.7 Perspectives and open questions-``curtain down and nothing settled''
Acknowledgments
References
7. Axon guidance: Slit-Robo signaling
7.1 Introduction
7.2 Slits and their receptors
7.2.1 Slit discovery and structure
7.2.2 Identification of the slit receptor robo
7.2.3 Slit and Robo interactions
7.2.3.1 Regulation of Slit-Robo interactions
7.3 Slit-Robo function in midline crossing
7.3.1 Spatial expression patterns of Slit and Robo
7.3.2 Posttranscriptional Robo regulation
7.3.3 Regulation of Robo protein expression at the midline
7.3.3.1 Drosophila and vertebrate midlines
7.3.3.2 Caenorhabditis elegans midline
7.3.4 Regulation of Robo signaling at the midline in vertebrates
7.3.5 Slit-Robo signaling for exiting the midline
7.4 Modulation of Slit-Robo signaling
7.4.1 Transcriptional control
7.4.2 Regulation of Slit-Robo signaling by metalloprotease cleavage
7.4.3 Regulation of Slit-Robo signaling by ubiquitination
7.5 Signaling downstream of Robo
7.5.1 Rho family of small GTPases
7.5.2 Abelson tyrosine kinase
7.5.3 Actin-interacting proteins
7.6 Beyond the midline: additional roles for Slit-Robo in the nervous system
7.6.1 Lateral positioning
7.6.2 Cell migration and cell polarity
7.6.3 Dendritic and axonal outgrowth and branching
7.7 Slit-Robo contribution to axon targeting in a complex target field
7.8 Involvement of Slit-Robo in disorders of the nervous system
7.9 Conclusion
References
8. Nonconventional axon guidance cues: Hedgehog, TGF-β/BMP, and Wnts in axon guidance
8.1 Introduction
8.1.1 Morphogens as axon guidance cues
8.2 Sonic hedgehog in axon guidance
8.2.1 Canonical Shh signaling
8.2.2 Shh is a chemoattractant for spinal cord commissural axons
8.2.3 Shh binding to Boc attracts commissural axons through a noncanonical signaling pathway to modulate the growth cone cytoskeleton
8.2.4 Shh guides axons along the longitudinal axis of the spinal cord
8.2.5 14-3-3 proteins regulate a cell-intrinsic switch from Shh-mediated attraction to repulsion of commissural axons after midli ...
8.2.6 Shh guides contralateral and ipsilateral retinal ganglion cell axons
8.2.7 Shh is a chemoattractant for midbrain dopaminergic axons
8.2.8 Shh binding to Gas1 repels enteric axons
8.3 TGF-β superfamily members in axon guidance
8.3.1 Canonical bone morphogenetic protein signaling
8.3.2 BMP7:GDF7 repels spinal cord commissural axons
8.4 Wnts in axon guidance
8.4.1 Canonical and noncanonical Wnt signaling
8.4.2 Wnt5 repels commissural axons from the Drosophila posterior commissure via derailed, a Ryk tyrosine kinase family member
8.4.3 Wnt5, complexed with derailed, repels Drosophila mushroom body axons
8.4.4 Wnt binding to Ryk repels axons of the corticospinal tract and corpus callosum through a Ca2+-dependent mechanism
8.4.5 Wnt binding to Fz attracts postcrossing commissural axons via protein kinase C ζ and planar cell polarity signaling
8.4.6 Wnt binding to Fz regulates dopaminergic axon guidance through planar cell polarity signaling
8.4.7 Wnt3 mediates mediolateral retinotectal topographic mapping
8.4.8 Wnts guide axons of Caenorhabditis elegans mechanosensory neurons and D-type motoneurons via Fz-type receptors
8.4.9 The Wnt ligand CWN2 regulates Caenorhabditis elegans motor neuron axon guidance through a Ror-type receptor CAM-1
8.5 Cross-talk between axon guidance cues
8.5.1 Shh induces the response of commissural axons to semaphorin repulsion during midline crossing
8.5.2 Shh regulates Wnt signaling in postcrossing commissural axons
8.5.3 The TGF-β family member unc-129 regulates Unc6/Netrin signaling in Caenorhabditis elegans
8.6 Conclusions and perspectives
List of Acronyms and Abbreviations
Glossary
Acknowledgments
References
9. Axon regeneration
9.1 Introduction
9.2 Anatomy of the spinal cord
9.3 Spinal cord injury repair: a complex problem
9.4 Axon regeneration in the injured central nervous system versus peripheral nervous system
9.4.1 Intrinsic mechanisms of dorsal root ganglion neuron axon regeneration
9.5 Extrinsic mechanisms: inhibitors of central nervous system axon regeneration
9.6 Extrinsic mechanisms: growth factors
9.6.1 The anatomical substrate of neurorepair
9.7 Axon regeneration in the retinofugal system
9.8 Lessons learned from an evolutionary perspective
9.8.1 Immune-mediated neurorepair mechanisms
9.9 Conclusions
Acknowledgments
References
10. Axon maintenance and degeneration
10.1 Introduction
10.2 Essentials of axonal transport in axon maintenance
10.2.1 Cellular components that are transported along the axons
10.2.2 Regulations of microtubule stability and organization during axon maintenance
10.2.3 Defects in motor proteins cause axon degeneration
10.2.4 Role of mitochondria transport in axon maintenance
10.2.5 Membrane transport and insertion are essential for axon maintenance
10.3 Proteasome and autophagy pathways in axonal homeostasis
10.3.1 Ubiquitin-proteasome system in axon maintenance
10.3.2 Role of autophagy/lysosome pathway in maintaining axonal homeostasis
10.4 Role of glial cells in axon maintenance
10.5 Maintaining axon track positions and other structural features
10.6 Axon pruning and axon degeneration
10.6.1 Developmental axon pruning
10.6.2 Pathological axon degeneration
10.6.3 Molecular mechanisms of pathological axon degeneration
References
11. Dendrite development: invertebrates
11.1 Structure and anatomy of invertebrate dendrites
11.2 Methods for studying dendrite morphology in Drosophila
11.3 Anatomical background for key model systems in which dendritic morphogenesis is studied in invertebrates
11.3.1 Drosophila dendritic arborization sensory neurons
11.3.2 Drosophila motoneurons
11.3.3 Drosophila olfactory projection neurons
11.3.4 Caenorhabditis elegans PVD neurons
11.4 Cell biology of dendritic growth
11.4.1 Microtubule polarity differs between dendrites and axons
11.4.2 Dynein-dependent trafficking controls dendritic branching
11.4.3 Role of the secretory pathway and Golgi outposts in dendritic elaboration
11.5 Transcriptional control of dendritic morphology
11.5.1 Control of dendrite morphological identity of Drosophila PNS neurons
11.5.2 Transcriptional control of dendritic targeting of olfactory PNs
11.5.3 Chromatin remodeling factors and dendritic development
11.6 Posttranscriptional control of dendritic development
11.6.1 Control of mRNA translation in dendritic development
11.6.2 miRNAs in dendritic development
11.7 Control of dendritic field formation I: guidance and targeting
11.7.1 Slit and netrin signaling during midline dendritic guidance
11.7.2 A combinatorial ligand-receptor complex guides dendritic branches
11.7.3 Coarse and specific control of PN dendritic targeting
11.7.4 Glial control of dendritic targeting
11.8 Control of dendritic field formation II: dendritic self-avoidance and tiling
11.8.1 Interactions between dendrites generate evenly covered territories
11.8.1.1 Dendritic self-avoidance
11.8.1.2 Dendritic tiling
11.8.2 Scaling growth of arbors and maintenance of evenly covered territories
11.9 Dendritic remodeling
11.9.1 Transforming growth factor-β signaling and ecdysone receptor expression during dendritic remodeling
11.9.2 Sox14 and mical function downstream of ecdysone receptor
11.9.3 Signaling mechanisms for dendritic pruning
11.9.3.1 Ubiquitin-proteasome system
11.9.3.2 Caspases
11.9.4 The cell biology of dendritic pruning
11.9.4.1 Microtubule disassembly
11.9.4.2 Local endocytosis and compartmentalized calcium transients
11.9.5 Similarities between dendrite pruning and injury-induced axon degeneration
11.9.6 Similarities and differences in dendrite development, dendrite regrowth after pruning, and dendrite regeneration after injury
11.10 Concluding remarks
See also
References
12. Dendrite development: vertebrates
12.1 The structure and function of vertebrate dendrites
12.1.1 Methods for manipulating and studying dendrite morphology in vertebrates
12.2 The cell biology of dendritic growth
12.2.1 Regulators of the microtubule network in dendrite formation
12.2.2 Regulators of the actin cytoskeleton
12.2.3 Dendrite elaboration requires a satellite secretory pathway
12.2.4 RNA translation in dendrites
12.2.5 Powering dendrite growth
12.2.6 Intracellular cascades that translate extrinsic signals into changes in dendrite structure
12.3 Control of dendritic field formation I: size
12.3.1 Afferent-derived neurotrophins limit size
12.3.2 Control of arbor size by neurotransmission
12.3.3 Activity-dependent mechanisms that influence dendrite growth and stabilization
12.4 Control of dendritic field formation II: shape
12.4.1 Apical dendrite initiation and outgrowth of cortical pyramidal neurons
12.4.2 Activity-dependent orientation of dendrite growth in the somatosensory cortex
12.4.3 Positional cues shape asymmetric dendritic arbors in the mouse retina
12.5 Control of dendritic field formation III: targeting and synapse selectivity
12.5.1 Formation of a Proto-IPL by retinal amacrine cells
12.5.2 Laminar targeting of retinal dendrites is coordinated by adhesive and repellent cues
12.5.3 Transcriptional control of laminar-specific targeting of dendrites in retina
12.5.4 Local recognition mechanisms to control synapse selectivity
12.5.5 An integrated, multistep model for synaptic wiring in the retina IPL
12.6 Space-filling mechanisms to optimize dendritic field distribution
12.6.1 Tiling and mosaics
12.6.2 Dendrite self-avoidance
12.7 Emergence of dendrite compartmentalization
12.7.1 Subcellular patterning of synaptic inputs along dendritic domains
12.7.2 Patterning the membrane excitability of dendritic compartments
12.8 Neurodevelopmental disorders: the price of poor dendritic development?
12.9 Conclusion
Abbreviations
Acknowledgments
References
13. Cell polarity and initiation of migration
13.1 Introduction
13.2 Migratory behaviors during radial migration in the developing cerebral cortex
13.2.1 Bipolar migrating neurons along the radial glial fibers: locomotion
13.2.2 Radial glial fiber-independent mode of migration: somal translocation and terminal translocation
13.2.3 Multipolar migration
13.2.4 Transformation from multipolar migrating neurons to bipolar locomoting neurons
13.2.5 Departure from the ventricular zone: differences in migratory behavior between direct progeny of the apical progenitors in ...
13.2.6 Behaviors of the progenitor cells in the subventricular zone
13.3 Molecular mechanisms that regulate the initiation of migration and cell polarity during migration
13.3.1 Coupling between neuronal differentiation and migration
13.3.2 Controlling the initiation of radial migration
13.3.3 Regulation of multipolar migration
13.3.4 Extracellular molecules that affect migrating cells
13.4 Conclusion
See also
List of abbreviations
Glossary
Supplementary data
References
14. Nucleokinesis
14.1 Nucleokinesis: introduction
14.2 The nucleus
14.2.1 The nuclear membrane and nuclear pores
14.3 Chromatin
14.4 Membraneless organelles in the nucleus
14.5 Higher order structure of the nucleus
14.6 Diseases
14.6.1 Cohesinopathies
14.6.2 Affecting the nuclear envelope
14.7 Interactions between the nucleus and the cytoskeleton
14.7.1 The LINC complex, structure
14.8 The LINC complex, function
14.9 The LINC complex in nuclear positioning
14.10 The link between the nucleus and the centrosome
14.11 The LINC complex in nucleokinesis
14.12 Nucleokinesis during interkinetic nuclear movement
14.13 Microtubule binding motors
14.13.1 Dynein
14.13.2 Kinesin Kif1a
14.14 Cytoskeleton dynamics as nuclear drivers
14.15 Collective mechanisms for nuclear migration
14.15.1 Intercellular signaling
14.15.2 Mechanical interactions
14.16 The role of INM during neurodevelopment
14.17 INM summary
14.18 Conclusions and future directions
Acknowledgments
References
15. Radial migration in the developing cerebral cortex
15.1 Introduction
15.2 Production of cortical projection neurons
15.3 Organization of the neocortex
15.4 Trajectory of migrating neurons in the developing brain
15.5 Modes of migration
15.6 Radial migration in the developing human neocortex
15.7 Factors that regulate the radial migration of cortical neurons
15.7.1 Secreted molecules
15.7.1.1 Reelin
15.7.1.2 Semaphorins
15.7.2 Neurotransmitters
15.7.2.1 GABA
15.7.2.2 Glutamate
15.7.2.3 ATP
15.7.3 Adhesion molecules
15.7.3.1 Integrins
15.7.3.2 Gap junctions
15.7.4 Cytoskeletal regulators
15.7.4.1 Lis1
15.7.4.2 Doublecortin
15.7.4.3 Filamin A (FLNA/FLN1)
15.7.4.4 Cdk5
15.7.5 Transcription factors
15.7.5.1 Pax6
15.7.5.2 Tbr2
15.7.5.3 Neurogenins
15.8 Summary
References
16. Mechanisms of tangential migration of interneurons in the developing forebrain
16.1 Birth of distinct interneuron subtypes and onset of their migration from the subpallium
16.2 Molecular cues drawing the path of cortical interneuron migration
16.3 Molecular cues controlling the integration of interneurons into the cortical migratory streams
16.4 Molecular cues controlling the intracortical dispersion of interneurons
16.5 Signals dictating the arrest of interneuron migration within the cortical wall
16.6 Role of subpallial transcription factors in the tangential migration of interneurons into the cortex
16.7 Cell-intrinsic regulation of cortical interneuron migration
16.8 Dynamic remodeling of the cytoskeleton during interneuron migration
16.9 Regulation of the tangential migration of interneurons in the rostral migratory stream to the olfactory bulb
16.10 Molecular regulation of the migration of striatal interneurons
16.11 Evolutionary perspective of the tangential migration
16.12 Conclusions and perspectives
List of acronyms and abbreviations
References
17. Migration in the hippocampus
17.1 Overview of hippocampal structure and lamination
17.1.1 Terminology important for studying hippocampal structure
17.2 Developmental specification of hippocampal fields
17.2.1 The basic developmental scheme of the hippocampus
17.2.2 The cortical hem
17.2.3 The cortical hem organizes the hippocampal fields
17.2.4 The role of canonical Wnt signaling in hippocampal development
17.3 Migration of Cajal-Retzius cells in the hippocampus
17.3.1 What are Cajal-Retzius cells?
17.3.2 What are the functions of Cajal-Retzius cells?
17.3.3 What are the origins of Cajal-Retzius cells?
17.3.4 The cortical hem is the major source of Cajal-Retzius cells for the dorsal telencephalon
17.3.5 The extent of the cortex covered by hem-derived Cajal-Retzius cells
17.3.6 Recruitment of hem-derived Cajal-Retzius cells to the meninges
17.3.7 Tangential dispersion of Cajal-Retzius cells in the marginal zone
17.4 Migration of hippocampal pyramidal neurons
17.5 Migration of hippocampal interneurons
17.5.1 Cellular and distributional diversity of interneurons in the hippocampus
17.5.2 Origins and migration of hippocampal interneurons
17.6 Migration of neural progenitors and granule cells in the dentate gyrus during development
17.6.1 The basic developmental scheme of the dentate gyrus
17.6.2 Migration of granule neurons to form the granule cell layer
17.6.3 Emergence and migration of long-lived neural stem cells and establishment of subgranular zone
17.7 Conclusions
References
18. Hindbrain tangential migration
18.1 Introduction
18.2 Tangential migration: a historical overview
18.3 Molecular mechanisms controlling the tangential migration of precerebellar neurons
18.3.1 Influence of the midline on tangentially migrating precerebellar neurons
18.3.2 Why do PCN neurons migrate near the pial surface?
18.4 Molecular mechanisms controlling the tangential migration of facial motor neurons
18.4.1 Origin and migration of facial motor neurons
18.4.2 The caudal migration of FBM neurons
18.4.2.1 The planar cell polarity pathway
18.4.2.2 Other molecules controlling FBM caudal migration
18.4.3 Role of chemoattraction and chemorepulsion
18.4.4 Role of the meninges in the tangential migration of FBM neurons
18.5 Ending tangential migration
18.6 Conclusion
Acknowledgments
References
19. Neuronal migration in the developing cerebellar system
19.1 Introduction
19.1.1 Part I. Diverse migration pathways and guidance cues during cerebellar system development
19.1.1.1 Distinct cerebellar germinal zones: the ventricular zone and rhombic lip
19.1.1.1.1 Early patterning
19.1.1.1.2 The rhombic lip and Atoh1 domain define the glutamatergic lineage
19.1.1.1.3 The ventricular zone and Ptf1a domain define the GABAergic lineage
19.1.1.1.4 Other Rh1 derivatives
19.1.1.2 Migration of purkinje cells
19.1.1.3 Migration of minor ventricular zone derivatives: Pax2-positive interneurons, basket cells, golgi cells, and stellate cells
19.1.1.4 Migration of precerebellar nuclei
19.1.1.5 Migration of upper rhombic lip derivatives
19.1.1.5.1 Deep cerebellar nuclei
19.1.1.5.2 Granule neuron progenitors and cerebellar granule neurons
19.1.1.5.3 Unipolar brush cells
19.1.2 Part II. The cytoskeletal organization of cerebellar granule neurons
19.1.2.1 Cerebellar granule neuron migration diversity after the establishment of the secondary germinal zone
19.1.2.2 The road to the two-stroke motility paradigm
19.1.2.3 The roles of the microtubule cytoskeleton and associated motors
19.1.2.4 The role of the actin cytoskeleton
19.1.2.5 The role of microtubule-actin cross talk
19.1.3 Part III. The facets of cerebellar granule neuron polarity: timing cell recognition, differentiation, germinal zone exit, a ...
19.1.3.1 Cerebellar granule neuron recognition/adhesion: the contribution of astrotactins and the siah2-Pard3-JamC pathway
19.1.3.2 The Zeb1-Pard6/3A transcriptional pathway
19.1.3.3 The foxo polarization pathway
19.1.4 Part IV. Migration deficits in cerebellar medulloblastomas: the effects of perturbed migration pathways are no longer limit ...
Acknowledgments
References
20. Neuronal migration of guidepost cells
Chapters cited
20.1 An introduction to guidepost cells
20.1.1 Neuronal migration in the context of axonal tracts formation
20.1.2 Defining the notion of guidepost cells
20.2 Role of neuronal migration in the formation of the lateral olfactory tract
20.2.1 Anatomy and development of the lateral olfactory tract
20.2.2 Diffusible guidance cues in the pathfinding of lateral olfactory tract axons
20.2.3 Roles of guidepost ``lot cells''
20.2.4 Tangential migration of lot cells: specification, routes, and molecular mechanisms
20.2.5 Fate of lot cells
20.3 Hippocampal Cajal-retzius cells in the formation of axonal connections
20.3.1 Anatomy and development of the hippocampus and entorhinohippocampal projections
20.3.2 Cajal-Retzius cells as putative guidepost neurons for the formation of entorhinal projections
20.3.3 Toward a more generic role of Cajal-Retzius cells as guideposts?
20.4 Migration of neuronal guidepost cells in the formation of thalamocortical connections
20.4.1 Anatomy and development of thalamocortical and corticofugal axons
20.4.2 Pioneer cortical subplate axons in the pathfinding of thalamocortical projections
20.4.3 Origin and migration of subplate neurons
20.4.4 The subpallium is a major intermediate target for thalamocortical axons
20.4.5 Guidepost cells in the diencephalic and subpallial pathfinding of thalamocortical projections
20.4.6 Migration of guidepost corridor cells: routes and guidance cues
20.4.7 Fate of guidepost cells for thalamocortical projections
20.5 Neuronal migration of guidepost cells in the formation of the corpus callosum
20.5.1 Anatomy and development of the corpus callosum
20.5.2 Roles of glial cells in the development of the corpus callosum
20.5.3 Tangentially migrating neurons in the development of the corpus callosum
20.6 Neuronal migration of guidepost cells and evolution of brain wiring
20.6.1 Tangential migration of guidepost neurons: a hallmark of the telencephalon?
20.6.2 Neuronal migration of guidepost cells in the evolution of the internal capsule
20.7 Towards an integration of migrating guidepost neurons in normal and pathological brain development
20.7.1 Guidepost neurons in the shaping of axonal tract organization and topography
20.7.2 Integrating tangential neuronal migration of guideposts in normal and pathological brain development
20.8 Conclusions
References
21. Neuronal migration in the postnatal brain
21.1 Introduction
21.2 Regulation of neuronal migration in the normal brain
21.2.1 Migratory scaffolds
21.2.1.1 Neighboring cells in the neuronal chain
21.2.1.2 Astrocytes
21.2.1.3 Blood vessels
21.2.2 Directional control from the V-SVZ toward the OB
21.2.3 Migration termination in the OB
21.3 Regulation of neuronal migration in the injured brain
21.3.1 Migratory scaffolds in the injured brain
21.3.1.1 Neighboring cells in the neuronal chain
21.3.1.2 Astrocytes
21.3.1.3 Blood vessels
21.3.1.4 Radial glial cells
21.3.2 Directional control toward a lesion
21.3.3 Enhancement of neuronal migration as a strategy for endogenous neuronal regeneration
21.4 Postnatal neuronal migration in primates
21.5 Summary
References
22. Transcriptional and posttranscriptional mechanisms of neuronal migration
22.1 Introduction to neuronal migration
22.1.1 Different ways to migrate: ``I did it my way''
22.2 Transcriptional and posttranscriptional control of neuronal migration
22.2.1 Radial migration
22.2.1.1 Radial migration: locomotion
22.2.1.2 Radial migration: translocation
22.2.1.3 Subtypes of neocortical radial glia; outer radial glia and the somal translocation mode of migration
22.2.1.3.1 Interplay of transcription factors and radial migration guidance cues
22.2.1.3.2 Posttranscriptional events in radial migration: the role of RNA-binding proteins, microRNA, and long noncoding RNA
22.2.1.4 RNA-binding proteins
22.2.1.5 lncRNAs
22.2.1.6 MicroRNAs
22.2.2 Tangential migration: transcriptional and posttranscriptional control
22.2.2.1 Interplay of transcription factors and tangential migration guidance cues
22.2.2.1.1 Posttranscriptional events in tangential migration: the role of RNA-binding proteins and microRNA
22.3 Conclusion and future directions
List of acronyms and abbreviations
References
23. Migration of myelin-forming cells in the CNS
23.1 Introduction
23.1.1 Genesis of myelin-producing cells during development
23.1.2 Oligodendrocyte precursor cells: born to migrate
23.2 Migratory paths followed by oligodendrocyte progenitor and precursor cells
23.3 Chemokinetic factors: the motility of oligodendrocyte precursors
23.4 Adhesion and chemotactic mechanisms: how the movement of oligodendrocyte precursors is guided?
23.4.1 Adhesion and surface molecules
23.4.2 Secreted factors
23.5 Concluding remarks
Acknowledgments
References
24. Coordination of different modes of neuronal migration and functional organization of the cerebral cortex
24.1 Introduction
24.1.1 Arealization of the cortex
24.1.2 Cortical columns constitute cortical areas
24.1.3 Minicolumns constitute columns
24.2 Migration of related projection neurons into the same minicolumn
24.2.1 Early lack of evidence that sister projection neurons migrate into the same minicolumn
24.2.2 Sister projection neurons migrate into the same minicolumn and intersynapse
24.3 Integration of projection neurons into cortical minicolumns
24.3.1 Migratory scaffolds restrict tangential movement of projection neurons
24.3.2 Molecular signaling limits tangential movement of projection neurons during multipolar stage
24.4 Integration of interneurons into cortical columns
24.4.1 Interneuron subtypes areally distribute via tangential migration
24.4.2 Do sister interneurons migrate into the same minicolumn?
24.4.3 Sister interneurons preferentially intersynapse
24.4.4 Regulating the timing of the shift from tangential to radial migration
24.4.5 Projection neurons attract migrating interneurons into cortical plate
24.4.6 Radial glial cells trigger a shift in migration mode
24.5 Genetic and cellular mechanisms controlling shifts in migratory modes
24.6 Conclusion
List of abbreviations
References
25. The impact of different modes of neuronal migration on brain evolution
25.1 Types of neuronal migration in vertebrate brain development-radial and tangential migration shaping vertebrate brains
25.2 The impact of radial migration on brain evolution
25.2.1 Evolution of radial migration
25.2.2 Radial migration on laminar brains
25.2.3 Radial migration on elaborated brains
25.2.4 The influence of radial migration on pallial internal circuitry
25.2.4.1 Somal translocation
25.2.4.2 Glial-guided locomotion
25.2.4.3 Evolutionary origin of glial-aided locomotion
25.3 The impact of tangential migration on brain evolution
25.3.1 Pallial interneurons and the modulation of brain circuits
25.3.1.1 Conserved features of tangential migration of pallial interneurons in vertebrates
25.3.1.2 Divergence in tangential migratory routes of pallial interneurons
25.3.1.3 Diversifying complexity of GABAergic subtypes
25.3.2 Glutamatergic tangential contributions as developmental scaffolds
25.3.3 Tangential migration shaping brain connections-guidepost neurons in evolution
25.3.4 Tangential migrations along the central nervous system
25.4 Conclusions
Glossary
References
26. Neuronal migration disorders
26.1 Introduction
26.2 Types of malformations
26.2.1 Pachygyria
26.2.2 Lissencephaly
26.2.3 Cobblestone lissencephaly
26.2.4 Subcortical band heterotopia
26.2.5 Periventricular heterotopia
26.2.6 Polymicrogyria
26.2.7 Mammalian target of rapamycin complex pathway-related malformations
26.2.8 Microcephaly
26.3 Identified mutations and mechanisms in neuronal migration disorder
26.3.1 Mutations in microtubule-associated proteins (LIS1, DCX, KIF5C, KIF2A, DYNC1H1, and EML1)
26.3.2 Tubulin mutations (TUBA1A, TUBB2B, TUBB3, TUBG1, TUBA8, and TUBB5)
26.3.3 Periventricular heterotopia and mutations in FLNA, ARFGEF2, C6orf70, FAT4, DCHS1, and MOB2
26.3.4 Variant lissencephalies and mutations in ARX and RELN
26.3.5 Cobblestone malformations and mutations in dystroglycan genes
26.3.6 Focal cortical dysplasias and dysplastic megalencephaly and mutations in mTOR, PIK3CA, DEPDC5, AKT3, NPRL3, and PIK3R2
26.4 Summary and concluding remarks
References
Index
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B
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D
E
F
G
H
I
J
K
L
M
N
O
P
R
S
T
U
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Citation preview

Cellular Migration and Formation of Axons and Dendrites Comprehensive Developmental Neuroscience Second Edition Senior Editors-in-Chief John Rubenstein

Department of Psychiatry & Weill Institute for Neurosciences University of California, San Francisco, San Francisco, CA, United States

Pasko Rakic

Department of Neuroscience & Kavli Institute for Neuroscience Yale School of Medicine, New Haven, CT, United States

Editors-in-Chief Bin Chen

Department of Molecular, Cell & Developmental Biology University of California, Santa Cruz, Santa Cruz, CA, United States

Kenneth Y. Kwan

Michigan Neuroscience Institute & Department of Human Genetics University of Michigan, Ann Arbor, MI, United States

Section Editors Alex Kolodkin Solomon H. Snyder Department of Neuroscience Johns Hopkins University School of Medicine, Baltimore, MD, United States

Eva Anton

UNC Neuroscience Center & Department of Cell and Molecular Physiology University of North Carolina School of Medicine, Chapel Hill, NC, United States

Academic Press is an imprint of Elsevier 125 London Wall, London EC2Y 5AS, United Kingdom 525 B Street, Suite 1650, San Diego, CA 92101, United States 50 Hampshire Street, 5th Floor, Cambridge, MA 02139, United States The Boulevard, Langford Lane, Kidlington, Oxford OX5 1GB, United Kingdom Copyright © 2020 Elsevier Inc. All rights reserved. No part of this publication may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopying, recording, or any information storage and retrieval system, without permission in writing from the publisher. Details on how to seek permission, further information about the Publisher’s permissions policies and our arrangements with organizations such as the Copyright Clearance Center and the Copyright Licensing Agency, can be found at our website: www.elsevier.com/permissions. This book and the individual contributions contained in it are protected under copyright by the Publisher (other than as may be noted herein). Notices Knowledge and best practice in this field are constantly changing. As new research and experience broaden our understanding, changes in research methods, professional practices, or medical treatment may become necessary. Practitioners and researchers must always rely on their own experience and knowledge in evaluating and using any information, methods, compounds, or experiments described herein. In using such information or methods they should be mindful of their own safety and the safety of others, including parties for whom they have a professional responsibility. To the fullest extent of the law, neither the Publisher nor the authors, contributors, or editors, assume any liability for any injury and/or damage to persons or property as a matter of products liability, negligence or otherwise, or from any use or operation of any methods, products, instructions, or ideas contained in the material herein. Library of Congress Cataloging-in-Publication Data A catalog record for this book is available from the Library of Congress British Library Cataloguing-in-Publication Data A catalogue record for this book is available from the British Library ISBN: 978-0-12-814407-7 For information on all Academic Press publications visit our website at https://www.elsevier.com/books-and-journals

Publisher: Nikki Levy Acquisitions Editor: Natalie Farra Editorial Project Manager: Andrae Akeh Production Project Manager: Surya Narayanan Jayachandran Cover Designer: Mark Rogers Cover Image: Jason Keil Typeset by TNQ Technologies

Contributors François Beaubien, Montreal Neurological Institute, McGill University, Department of Neurology and Neurosurgery, Montréal, QC, Canada Franck Bielle, Institut du Cerveau et de la Moelle Epinière, Paris, France Frank Bradke, Laboratory for Axon Growth and Regeneration, German Center for Neurodegenerative Diseases (DZNE), Bonn, Germany Frédéric Charron, Institut de Recherches Cliniques de Montréal (IRCM), Montreal, QC, Canada; Integrated Program in Neuroscience, McGill University, Montreal, QC, Canada; Department of Anatomy and Cell Biology, Department of Biology, McGill University, Montreal, QC, Canada; Department of Medicine, University of Montreal, Montreal, QC, Canada; Division of Experimental Medicine, McGill University, Montreal, QC, Canada Alain Chédotal, Institut de la Vision, Sorbonne Université, INSERM, CNRS, Paris, France Jean-François Cloutier, Montreal Neurological Institute, McGill University, Department of Neurology and Neurosurgery, Montréal, QC, Canada Christopher L. Cunningham, Solomon H. Snyder Department of Neuroscience, Johns Hopkins University School of Medicine, Baltimore, MD, United States Fernando de Castro, Instituto Cajal-CSIC, Spanish Research Council/Consejo Superior de Investigaciones Científicas-CSIC, Madrid, Spain Kevin C. Flynn, Stem Cell and Gene Therapy, BioTechne, Minneapolis, MN, United States Fernando García-Moreno, Achucarro Basque Center for Neuroscience, Parque Científico UPV/EHU Edif. Sede, Leioa, Spain; Ikerbasque Foundation, Bilbao, Spain Sonia Garel, Institut de Biologie de l’École Normale Supérieure (IBENS), PSL Université, Paris, France; Institut National de la Santé et de la Recherche Médicale (INSERM) U1024, Paris, France; Centre National de la Recherche Scientifique (CNRS) UMR 8197, Paris, France

R.J. Giger, The University of Michigan, Medical School, Ann Arbor, MI, United States Wesley B. Grueber, Columbia University, New York, NY, United States K. Hayashi, Keio University School of Medicine, Tokyo, Japan Zhigang He, Kirby Center of Neuroscience, Children’s Hospital Boston, Harvard Medical School, Boston, MA, United States Holden Higginbotham, Department of Biology, Brigham Young University, Rexburg, ID, United States Katrine Iversen, Montreal Neurological Institute, McGill University, Department of Neurology and Neurosurgery, Montréal, QC, Canada Artur Kania, Neural Circuit Development Laboratory, Institut de Recherches Cliniques de Montréal (IRCM), Montreal, QC, Canada; Integrated Program in Neuroscience, McGill University, Montreal, QC, Canada; Department of Anatomy and Cell Biology, Division of Experimental Medicine, McGill University, Montreal, QC, Canada Eyal Karzbrun, Kavli Institute of Theoretical Physics and Department of Physics, University of California, Santa Barbara, CA, United States Arnold R. Kriegstein, Department of Neurology, University of California, San Francisco, CA, United States; The Eli and Edythe Broad Center of Regeneration Medicine and Stem Cell Research, University of California, San Francisco, CA, United States Zeljka Krsnik, Croatian Institute for Brain Research, School of Medicine, University of Zagreb, Zagreb, Croatia Christophe Laumonnerie, Department of Developmental Neurobiology, St. Jude Children’s Research Hospital, Memphis, TN, United States Julie L. Lefebvre, The Hospital for Sick Children, Toronto, ON, Canada; Department of Molecular Genetics, University of Toronto, Toronto, Canada

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Contributors

Fanny Lepiemme, GIGA-Stem Cells / GIGA-Neurosciences, University of Liège, Liège, Belgium Guangnan Li, Department of Neurology, University of California, San Francisco, CA, United States Joseph J. LoTurco, University of Connecticut, Mansfield, CT, United States Le Ma, Thomas Jefferson University, Philadelphia, PA, United States Jean-Bernard Manent, Inmed Inserm, Marseille, France Julie Marocha, The Hospital for Sick Children, Toronto, ON, Canada; Department of Molecular Genetics, University of Toronto, Toronto, Canada Zoltán Molnár, Department of Physiology, Anatomy and Genetics, University of Oxford, Oxford, United Kingdom K. Nakajima, Keio University School of Medicine, Tokyo, Japan Laurent Nguyen, GIGA-Stem Cells / GIGA-Neurosciences, University of Liège, Liège, Belgium Stephen C. Noctor, Department of Psychiatry and Behavioral Sciences, UC Davis MIND Institute, Sacramento, CA, United States

Masato Sawada, Department of Developmental and Regenerative Neurobiology, Institute of Brain Science, Nagoya City University Graduate School of Medical Sciences, Nagoya, Japan Kazunobu Sawamoto, Department of Developmental and Regenerative Neurobiology, Institute of Brain Science, Nagoya City University Graduate School of Medical Sciences, Nagoya, Japan; Division of Neural Development and Regeneration, National Institute for Physiological Sciences, Okazaki, Japan K. Sekine, Keio University School of Medicine, Tokyo, Japan Carla Silva G., GIGA-Stem Cells / GIGA-Neurosciences, University of Liège, Liège, Belgium David J. Solecki, Department of Developmental Neurobiology, St. Jude Children’s Research Hospital, Memphis, TN, United States Constantino Sotelo, Institut de la Vision, Sorbonne Université, INSERM, CNRS, Paris, France H. Tabata, Keio University School of Medicine, Tokyo, Japan Marc Tessier-Lavigne, Stanford University, Stanford, CA, United States

Hirofumi Noguchi, Department of Neurology, University of California, San Francisco, CA, United States

Stephen R. Tymanskyj, Thomas Jefferson University, Philadelphia, PA, United States

R. Jeroen Pasterkamp, Department of Translational Neuroscience, UMC Utrecht Brain Center, University Medical Center Utrecht, Utrecht University, Utrecht, The Netherlands

Marieke G. Verhagen, Department of Translational Neuroscience, UMC Utrecht Brain Center, University Medical Center Utrecht, Utrecht University, Utrecht, The Netherlands

Samuel J. Pleasure, Department of Neurology, University of California, San Francisco, CA, United States

Fan Wang, Department of Neurobiology, Duke University, Durham, NC, United States

F. Polleux, Columbia University, New York, NY, United States

Franco Weth, Karlsruhe Institute of Technology, Zoological Institute, Department of Cell and Neurobiology, Karlsruhe, Germany

Tatiana Popovitchenko, Department of Neuroscience and Cell Biology, Rutgers University, Robert Wood Johnson Medical School, New Brunswick, NJ, United States Janet E.A. Prince, Montreal Neurological Institute, McGill University, Department of Neurology and Neurosurgery, Montréal, QC, Canada Mladen-Roko Rasin, Department of Neuroscience and Cell Biology, Rutgers University, Robert Wood Johnson Medical School, New Brunswick, NJ, United States Orly Reiner, Department of Molecular Genetics, The Weizmann Institute of Science, Rehovot, Israel

Patricia T. Yam, Institut de Recherches Cliniques de Montréal (IRCM), Montreal, QC, Canada Jing Yang, School of Life Sciences, Peking University, Beijing, China Bing Ye, University of Michigan, Ann Arbor, MI, United States Bernard Zalc, Université Pierre & Marie Curie-Paris 6, Centre de Recherche de l’Institut du Cerveau et de la Moelle épinière, UMR_S 975, Inserm U 975, CNRS UMR 7225, Hôpital de la Salpêtrière, Paris, France

Chapter 1

Development of neuronal polarity in vivo F. Polleux Columbia University, New York, NY, United States

Chapter outline 1.1. Introduction 1.2. Axon initiation in vitro versus in vivo 1.2.1. Axon initiation in vitro 1.2.2. Axon initiation in vivo 1.3. Distinction between cues regulating axon specification versus axon growth 1.4. Extracellular cues regulating neuronal polarization and axon initiation 1.4.1. Netrin-1 and Wnt control axon initiation in Caenorhabditis elegans 1.4.2. Polarized emergence of the axon in retinal ganglion cells of Xenopus 1.4.3. Extracellular cues underlying the emergence of axon and dendrites in mammalian neurons 1.5. Intracellular pathways underlying neuronal polarization

3 4 4 5 5 6 6 6 7 8

1.5.1. Role of protein degradation and local translation in axon specification and axon growth 1.5.2. Role of cytoskeletal dynamics in axon initiation and growth 1.5.3. Major signaling pathways involved in axon initiation and growth 1.5.3.1. LKB1 and its downstream kinases SAD-A/B and MARK1-4 1.5.3.2. PAR3ePAR6eAPKC 1.5.3.3. Ras- and Rho-family of small GTPases 1.5.3.4. PI3K and PTEN signaling during axon specification 1.5.3.5. AKT/protein kinase B 1.5.3.6. GSK3 and axon specification 1.6. Conclusion and future directions References

8 9 10 10 11 12 13 14 14 15 16

1.1 Introduction The ability of neurons to form a single axon and multiple dendrites underlies the directional flow of information transfer in the central nervous system (CNS). Dendrites and axons are fundamentally distinct domains of neurons at the molecular, ultrastructural, and functional levels. Dendrites integrate synaptic inputs, triggering the generation of action potentials at the level of the soma and the axon initiation segment (AIS). Action potentials then propagate along the axon and trigger neurotransmitter release at presynaptic contacts onto their postsynaptic partners. This chapter reviews what is currently known about the cellular and molecular mechanisms underlying the ability of neurons to polarize and form a single axon and multiple dendrites during development. This question has received much attention over the past three decades using mainly in vitro approaches following some of the pioneering work of Gary Banker and colleagues on dissociated hippocampal neuron cultures (Craig and Banker, 1994; Goslin and Banker, 1989). In these assays, immature hippocampal or cortical pyramidal neurons (PNs) are dissociated and maintained in culture for variable periods of time (typically 1e 3 weeks). Remarkably, in these conditions, PNs can repolarize to form a single axon and multiple dendrites and later establish functional synaptic contacts in these reductionist conditions. This approach became, and remains, the dominant model to study multiple aspects of neuronal polarization, including axon specification as well as axon and dendrite growth, and it has yielded the identification of many molecules that regulate axon formation in vitro. However, the central question for the field has been to determine if the cellular and molecular mechanisms identified in these in vitro conditions are conserved in vivo. At present, only a few of the genes identified using in vitro approaches have been shown to be required for axon initiation and outgrowth in vivo. In vitro, axon initiation and elongation is thought to reflect the intrinsic ability of neurons to polarize in the absence of relevant extracellular cues. However, extracellular cues have been shown to play an

Cellular Migration and Formation of Axons and Dendrites. https://doi.org/10.1016/B978-0-12-814407-7.00001-8 Copyright © 2020 Elsevier Inc. All rights reserved.

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4 PART | I Formation of axons and dendrites

important role during neuronal polarization in vivo. Here, we focus on our current understanding of the complex interplay between extracellular cues and intracellular signaling pathways underlying the emergence of axons and dendrites during neuronal polarization in vivo.

1.2 Axon initiation in vitro versus in vivo 1.2.1 Axon initiation in vitro Historically, the advent of in vitroedissociated neuronal cultures provided an experimental template for improving our understanding of the cell biology of neuronal polarity, including the specification of axons and dendrites. Pioneering work using these cultures established a paradigm in which isolated neurons in culture can adopt spatially and functionally distinct dendritic and axonal domains (Craig and Banker, 1994; Goslin and Banker, 1989). Careful analysis of these cultures led to the observation that cultured hippocampal neurons transition through several stages: from freshly plated stage 1 cells bearing immature neurites to stage 5 cells that exhibit mature axons, dendrites, dendritic spines, and functional synapses (Fig. 1.1A) (Craig and Banker, 1994; Dotti et al., 1988). It should be noted that, in the classical E18 rat hippocampal cultures, most plated

FIGURE 1.1 Parallel between neuronal polarization in vitro and in vivo. Comparison of the sequence of events leading to the polarization of cortical pyramidal neurons in vivo and in vitro. (A) In dissociated cultures, postmitotic cortical neurons display specific transitions as classically described for hippocampal neurons by Dotti et al. (1988). At stage 1, immature postmitotic neurons display intense lamellipodial and filopodial protrusive activity, which leads to the emergence of multiple immature neurites, stage 2. Stage 3 represents a critical step when neuronal symmetry breaks and a single neurite grows rapidly to become the axon (purple) while other neurites acquire dendritic identity. Stage 4 is characterized by rapid axon and dendritic outgrowth. Finally, stage 5 neurons are terminally differentiated pyramidal neurons harboring dendritic spines and the axon initiation segment (AIS). (B) The axonedendrite polarity of pyramidal neurons is derived from the polarized emergence of the trailing (TP) and leading processes (LP), respectively. In vivo, pyramidal neurons acquire other key features of their terminal polarity such as the AIS (yellow cartridge) and dendritic spines (gray protrusions) during the first postnatal weeks of development.

Development of neuronal polarity in vivo Chapter | 1

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cells were polarized postmitotic neurons before dissociation; therefore, neuronal polarization using this in vitro model likely corresponds to the “repolarization” of previously polarized neurons in vivo. It is therefore important to keep in mind that molecular manipulations in this in vitro model act on previously polarized neurons that may retain some aspects of polarization, and this can be critical for interpreting results from such manipulations. Recent advances in techniques that allow manipulation of gene expression more specifically in neural progenitors, such as in utero or ex utero cortical electroporation (Hand et al., 2005; Hatanaka and Murakami, 2002; Saito and Nakatsuji, 2001; Tabata and Nakajima, 2001), provide a paradigm to (1) manipulate gene expression in progenitors, before neuronal polarization occurs upon cell cycle exit; and (2) visualize the earliest stages of neuronal polarization in a contextual cellular environment, including in organotypic slices or intact embryonic brain (Barnes et al., 2007; Calderon de Anda et al., 2008; Hand et al., 2005).

1.2.2 Axon initiation in vivo Neuronal polarization can be divided into several specific steps in vivo. Upon cell cycle exit, mammalian neurons usually migrate over long distances before reaching their final destinations. In vivo, most neurons undergo axonedendrite polarization during migration. While initiating radial migration, neocortical PNs form a leading process and a trailing process, each becoming the axon or the dendrite, respectively (Fig. 1.1B). Careful examination of the morphological transition between neural progenitor and postmitotic neuron reveals that neurons can inherit their axon and dendrite polarity directly from the apicobasal polarity of their progenitors. This is the case for retinal ganglion cells (RGCs) and bipolar cells in the developing vertebrate retina (Hinds and Hinds, 1978; Morgan et al., 2006; Zolessi et al., 2006, reviewed in Barnes and Polleux, 2009). In other neuronal subpopulations undergoing long-range migration, neuronal morphogenesis involves extensive stereotypical changes, leading to polarized outgrowth of their axons and dendrites. This is the case for cerebellar granule neurons (CGNs) as well as cortical and hippocampal PNs, two of the best-studied models of neuronal polarization (Gao and Hatten, 1993; Hatanaka and Murakami, 2002; Komuro et al., 2001; Noctor et al., 2004; Rakic, 1971, 1972; Shoukimas and Hinds, 1978). Both CGNs and PNs acquire their axonedendrite polarity from the polarized emergence of their trailing and leading processes, respectively, during migration (reviewed in Barnes and Polleux, 2009). Precise examination of the process dynamics occurring shortly after cell cycle exit in dorsal telencephalic progenitors suggests that there is often a slight delay between formation of trailing (axon initiation) and leading (dendrite elaboration) processes, with trailing process formation frequently preceding leading process formation (Calderon de Anda et al., 2008; Hand and Polleux, 2011; Namba et al., 2014). However, one thing is clear in both PNs and CGNs: Axon formation starts before or during radial migration. Interestingly, different neuronal populations display distinct modes of axon formation, reflecting their mode of migration, lineage, and type of axon projection. For example, cortical interneurons, which will form axons projecting only locally within the cortex, originate from the ventral telencephalon and must migrate over very long distances before initiating their axon upon reaching their final destination in the cortex (Bortone and Polleux, 2009; Cobos et al., 2007; Yamasaki et al., 2010). The precise mechanisms underlying axon emergence in cortical interneurons are largely unknown. However, they are strikingly different from those employed by radially migrating pyramidal cortical neurons, which initiate axon formation during migration. This leads to the hypothesis that the ability of interneurons to form an axon is inhibited during tangential migration and that axon initiation in cortical interneurons may depend upon factor(s) present only in their target environment, the cortex. As discussed later in the chapter, an emerging concept from recent work done primarily in Caenorhabditis elegans suggests that, in vivo, the “symmetry-breaking” events that lead to the emergence of the dendrite and the axon require the ability of postmitotic neurons to sense gradients of extracellular cues, leading to the asymmetric activation of signaling pathways underlying the emergence of the axon. These data are supported by recent evidence in mammals showing that extracellular cues such as diffusible TGFb or membrane-bound cell adhesion molecules such as TAG-1 play roles in axon specification in vivo by triggering specific signaling pathways in the neurite’s trailing process that becomes the axon (Yi et al., 2010; Namba et al., 2014).

1.3 Distinction between cues regulating axon specification versus axon growth Most studies published over the past two decades in this field have been performed using in vitro approaches. The classic paradigm for confirming the regulatory role of a gene in neuronal polarity is to show that downregulation of its expression using shRNA technology or gene knockout technology is required for axon formation. These experiments are typically done using staining with axon-specific makers and measurement of neurite length since the axon usually grows 5e10 times faster than neuritis, which will eventually become dendrites. However, this type of evidence may not be sufficient to

6 PART | I Formation of axons and dendrites

distinguish unambiguously an effector of axon specification from a molecule simply required for axon growth (Jiang et al., 2005). Conversely, showing that overexpression or overactivation of a candidate molecule leads to the emergence of multiple neurites that display the molecular identity of an axon is generally used to suggest that this molecule is sufficient to confer axon identity. However, this approach is limited by the fact that it relies on overexpression, which can be confounded by abnormal activation of a pathway normally not involved in axon specification or neuronal polarity. Recent technical advances allow for the manipulation of gene expression in vivo and include in utero cortical electroporation in rodent cortex or cerebellum (Famulski et al., 2010; Saito and Nakatsuji, 2001) and also transgenic approaches in Xenopus (Zolessi et al., 2006). Therefore, more biologically relevant validation of candidate gene function during neuronal polarization often includes testing its requirement using gene knockout or shRNA-mediated knockdown technologies or the analysis of conventional or conditional knockout in combination with in utero electroporation allowing single cell resolution analysis of axon formation (Barnes et al., 2007; Shelly et al., 2007; Yi et al., 2010; Namba et al., 2014). Finally, one important caveat of the in vivo approaches mentioned in this chapter to study mammalian axon specification is that several of the molecules identified by loss-of-function analysis not only affect axon/dendrite (trailing/leading process) generation but also often when compromised lead to abnormal initiation of radial migration. This effect on neuronal migration can complicate the interpretation of experimental results since defects in axon specification could be the direct result of the molecule’s function in axon specification, or these defects could result from secondary effects that are a result of preventing the postmitotic neuron from responding to extracellular cues that are required for neuronal polarization.

1.4 Extracellular cues regulating neuronal polarization and axon initiation 1.4.1 Netrin-1 and Wnt control axon initiation in Caenorhabditis elegans Is there any in vivo evidence for the role of extracellular cues in the specification of neuronal polarity? Significant progress in our understanding of the molecular and cellular mechanisms specifying axon initiation during neuronal polarization has been made using C. elegans as a model. This pioneering work has markedly enhanced our understanding of how extracellular cues instruct axon initiation in vivo (reviewed by Yogev and Shen, 2017). The neurons of the nematode have a stereotyped morphology, as can be seen with respect to specific projections along the dorsoventral and anterioreposterior body axes. Elegant experiments involving genetic screens have identified an extracellular cue, UNC-6 (Netrin), along with its receptor UNC-40 (DCC), as critical genes for orchestrating axon initiation in vivo (Adler et al., 2006). This work also identified downstream proteins in this pathway, including (mammalian orthologs are shown in parenthesis, when known) AGE-1 (phosphoinositide-3 kinase [PI3K]), DAF-18 (PTEN), UNC-34 (Enabled), CED-10 (Rac), UNC-115/AbLIM, and MIG-10/lamellipodin. The current model for the relationship among these genes and UNC-6/netrin signaling involves DAF-18’s limitation of AGE-1 activity following UNC-40/DCC stimulation and the asymmetric recruitment of MIG-10/ lamellipodin to the plasma membrane. This recruitment requires activated CED-10/Rac binding directly to MIG-10/ lamellipodin and the involvement of the PAK-like kinase, Pak-1 (Adler et al., 2006). The function of a kinase in the regulation of cytoskeletal rearrangement is consistent with a similar role played by MIG-10/lamellipodin and the likely mechanism by which it operates once recruited to the plasma membrane to stimulate directed neurite outgrowth. Another regulator thought to act in concert with MIG-10/lamellipodin to drive filopodial formation is the Enabled homolog, UNC34 (Chang et al., 2006). Finally, SLT-1 (Slit) is another extracellular cue that likely acts through MIG-10 recruitment to control neuronal polarization (Chang et al., 2006). Two other studies have identified the diffusible signal Wnt and its receptor as critical regulators of axon specification and neuronal polarity (Hilliard and Bargmann, 2006; Prasad and Clark, 2006). In addition to characterizing loss of function mutants in the genes encoding Lin-44 (Wnt) and its receptor Lin-17 (Frizzled [Fzl]), a genetic screen has identified VPS35, a component of retromer complex that regulates vesicular traffic and is required for proper Wnt secretion, as an important regulator of neuronal polarity (Pan et al., 2008; Prasad and Clark, 2006).

1.4.2 Polarized emergence of the axon in retinal ganglion cells of Xenopus Detailed live imaging experiments of Xenopus RGC polarization revealed that polarized axon outgrowth requires unidentified extracellular cues present in the basal lamina (Randlett et al., 2011; Zolessi et al., 2006). The axon of developing RGCs normally grows on the basal side of the neuron. In a mutant called Nok, characterized by the absence of retinal pigmented epithelium, some postmitotic RGC neurons show defective polarized axon outgrowth on the apical side along the now-exposed basal lamina. In this context, the polarized emergence of the axon on the basal side of the RGC is correlated with the position of the centrosome, Par3, and the apical complex (containing the atypical protein kinase C

Development of neuronal polarity in vivo Chapter | 1

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[aPKC], b-catenin, and F-actin) on the apical side of the cell where the dendrite will emerge. Taken together, this work strongly suggests that (1) the basal lamina contains important extracellular cues that play a role in organizing the polarized emergence of the RGC axon and (2) RGC neurons inherit the intrinsic apicobasal polarity of their progenitor at least with respect to the Par3/aPKC components of the polarity complex. Recently, a signaling cascade has been linked to potential extracellular cues regulating axon initiation in vivo (Barnes and Polleux, 2009; Barnes et al., 2008). Conditional deletion of LKB1 in pyramidal cortical neurons (also called Par4 or STK11) demonstrated that LKB1 is required for axon initiation in cortical neurons but does not impact their radial migration (Barnes et al., 2007). Structure/function analysis indicates that phosphorylation of LKB1 at Serine 431 is required for its function in axon specification (Barnes et al., 2007). This phosphorylation event has been linked to the ability of extracellular cues such as brain-derived neurotrophic factor (BDNF) to stimulate cAMP production and protein kinase A (PKA)edependent phosphorylation of LKB1 on S431 in the nascent axon (see below for details; Shelly et al., 2007, 2010, 2011).

1.4.3 Extracellular cues underlying the emergence of axon and dendrites in mammalian neurons Several lines of evidence suggest that extracellular cues can direct the polarized emergence of the axon and dendrites both in vitro and in vivo. One paradigm involves dissociated cortical or hippocampal PNs plated on striped substrates coated with two different cell adhesion molecules (e.g., laminin and NgCAM, reviewed in Barnes and Polleux, 2009). The first immature neurite of E18 hippocampal neurons contacting the boundary between two stripes systematically becomes the axon. This occurs despite initial outgrowth of immature neurites occurs on laminin or NgCAM, suggesting that immature neurites can detect changes in the nature of the extracellular substrate rather than the absolute nature of the novel substrate they are encountering (Esch et al., 1999). Using a similar approach, Shelly and colleagues showed that neurites of immature hippocampal neurons growing on a patterned substrate can detect the presence of BDNF, which plays an instructive role in axon specification because the first neurite contacting a BDNF stripe systematically becomes the axon (Shelly et al., 2007). The effect of BDNF on axon specification requires cAMP-dependent PKA activation and phosphorylation of LKB1 at position 431 by PKA (Shelly et al., 2007, 2010), suggesting that LKB1 phosphorylation on S431 acts as a detector that breaks neuronal symmetry following encounters with extracellular cues such as BDNF in this in vitro context. To detect the existence of extracellular cues that play a role in cortical axon guidance and neuron polarization, an overlay in vitro assay was developed. This rather simple assay involves plating fluorescently labeled dissociated cortical neurons onto cortical slices to test whether polarized axon emergence in vivo is mainly the result of asymmetric activation of intracellular effectors (perhaps inherited by progenitors) or extracellular cues that direct axon specification. Polleux et al. demonstrated that the second scenario is most likely because only a couple of hours after plating, the vast majority of cortical neurons displayed a single, short axon directed ventrally toward the ventricle (Polleux et al., 1998), as observed for radially migrating neurons in vivo. These authors went on to demonstrate that the class 3 secreted semaphorin, Sema3A, which is enriched in the most superficial part of the cortical wall (the top of the cortical plate), plays a role in repulsing axon initiation ventrally toward the ventricle (Polleux et al., 1998). More recently, Sema3A was shown to also regulate the polarized emergence of the leading process/apical dendrite both in the overlay assay, that is, independently of radial migration where it requires cGMP production and PKG activation (Polleux et al., 2000), and also in vivo during radial migration (Chen et al., 2008). Interestingly, Sema3A can play a role in the specification of dendritic identity by activating a cGMP-dependent pathway involving activation of cGMP-gated calcium channels (Nishiyama et al., 2011) and also by repressing axonal identity in a LKB1-dependent manner (Shelly et al., 2007, 2010, 2011; see Fig. 1.2). Recently, TGFb signaling was shown to be required for the polarized emergence of the axon of radially migrating PN in vivo (Yi et al., 2010, Fig. 1.3). TGFb ligands are expressed in the germinal zone of the cortex, where they could act as an instructive “ventral” cue for the polarized emergence of the axon in multipolar neurons before engaging radial migration. In vitro experiments demonstrated that local application of TGFb on a single neurite in immature stage 1 cortical neurons is sufficient to trigger fast axonal extension (Yi et al., 2010). Importantly, conditional genetic deletion of TGFb receptor 2 expression leads to the production of neurons without trailing process/axon in vivo. One noticeable difference compared with the conditional ablation of LKB1 (Barnes et al., 2007), which also leads to the absence of trailing process/axon formation but not radial migration defects, is that TGFb receptor 2 conditional deletion leads to retardation of radial migration in a subset of cortical neurons (Yi et al., 2010) (Fig. 1.2). TGFb receptor function during axon specification also requires the phosphorylation of Par6 on S435, which was previously shown to mediate the epithelial-to-mesenchymal transition (EMT; Ozdamar et al., 2005). As discussed later in this chapter, this “noncanonical” TGFb receptoredependent signaling represents an attractive in vivo signaling pathway

8 PART | I Formation of axons and dendrites

Sema3A

‘prodendrite’ signal(s) Polleux et al. (2000) Shelly et al. (2010) Shelly et al. (2011) Nishiyama et al. (2011)

MZ 6 5 CP LP 4 TP

IZ

Barnes et al. (2007) Shelly et al. (2007)

LKB1

3 2

SVZ

? VZ

Others?

SAD-A/B Kishi et al. (2005)

1

?

?

MAPs TGFE

‘proaxon’ signal(s) Yi et al. (2010)

Axon specification

FIGURE 1.2 Extracellular signals regulate axon and dendrite specification in neocortical pyramidal neurons (PNs). Time-lapse analysis has revealed that polarization of axodendritic polarity in PN occurs in vivo in a stepwise manner on cell cycle exit: Following asymmetric division of radial glial cells (step 1) or intermediate progenitors in the subventricular zone (SVZ) (not shown), recently generated postmitotic neurons first form transient, dynamic neurites (multipolar stage; step 2). Axon specification occurs when a trailing process (TP) is stabilized (red) either before (step 3) or after the formation of a leading process (LP) in neurons engaging radial translocation (step 4). On reaching their final destination at the top of the cortical plate (CP) (step 5), neurons detach from the radial glial scaffold and start an extensive program of axon growth (see Fig. 1.1) and dendritic branching concomitant with synaptogenesis (step 6).

for axon specification since it is known to involve recruitment of the ubiquitin ligase Smurf1, which induces local degradation of RhoA during EMT and has previously been implicated in axon specification (Schwamborn et al., 2007a, 2007b). Therefore, these results suggest that neuronal polarization might have coopted signaling pathways that regulate EMT for the purpose of axon specification. Overall, this work suggests that the polarized emergence of a single axon/trailing process and the apical dendrite/ leading process are controlled at least in part by extracellular cues such as TGFb and Sema3A, respectively, expressed in a graded manner along the neuron’s migratory path (Fig. 1.2). More recently, another type of extracellular cue that is membrane-bound (the cell adhesion molecule TAG-1) and expressed along the axon of preexisting neurons has been shown to regulate the emergence of the trailing axonal process during the multipolar-to-bipolar transition (Namba et al., 2014). This work suggests that, at different times during cortical neurogenesis (when neurons destined to populate different layers are generated), different types of cues such as diffusible graded TGFb or membrane-bound axonal CAMs such as TAG-1 play roles in axogenesis and may coordinate axogenesis and the directionality of axon outgrowth (for example, projecting medially for callosal projections vs. laterally for corticofugal projections). More work is needed to clarify whether or not other extracellular cues are required for the polarized emergence of axons and dendrites in diverse neuronal cell types and how these extracellular cues mediate their effects on the specification of the unique molecular axonal and dendritic identity (Fig. 1.2).

1.5 Intracellular pathways underlying neuronal polarization 1.5.1 Role of protein degradation and local translation in axon specification and axon growth Spatial regulation of protein expression by selective degradation has been demonstrated in several contexts during neuronal development, including axonal pruning (Watts et al., 2004), various aspects of axon guidance decisions (Bloom et al., 2007; Campbell and Holt, 2001; DiAntonio et al., 2001; Lewcock et al., 2007), synapse formation (DiAntonio et al., 2001; Nakata et al., 2005), synapse maintenance (Aravamudan and Broadie, 2003; DiAntonio et al., 2001; Ehlers, 2003; Speese et al., 2003), and synapse elimination (Ding et al., 2007, reviewed in DiAntonio and Hicke, 2004). Acute treatment with the proteosome inhibitor lactacystin blocks axogenesis in dorsal root ganglion cells (Klimaschewski et al., 2006). Furthermore, more prolonged inhibition of protein degradation with lactacystin leads to the formation of multiple axons

9

Sema3A concentration ?

TGFβ concentration ?

Development of neuronal polarity in vivo Chapter | 1

TGFβ TGFβ-R1/2 (Par3?)-aPKC Par6S435

?

(PKA, p90RSK)? LKB1S431

Smurf1 RhoA

SAD-A/B MAP phosphorylation–microtubule dynamics? Presynaptic vesicle trafficking?

Axon spcification FIGURE 1.3 Intracellular pathways underlying axon specification in response to extracellular cues in vivo. As shown in Fig. 1.2, recent data (Yi et al., 2010) provided evidence that TGFb receptor activation is required for axon specification/trailing process stabilization in vivo. This work also suggested that TGFb receptor signaling in the context of axon specification required a “noncanonical” branch of TGFb involved in EMT, which triggers Par6 phosphorylation by TGFb receptor 2 and recruits the ubiquitin ligase Smurf1 and locally degrades RhoA, three steps previously shown to be required for axon specification. Another kinase previously shown to be required in vivo for axon specification is LKB1 (Par4), which is phosphorylated on S431 specifically in the axon in response to several signaling pathways (PKA, p90RSK, or aPKC). LKB1 phosphorylates and activates several other kinases, including SAD-A/B kinases, which have been shown to be required for axon specification in vivo. The actual link between TGFb receptor/Par6/Smurf1/ RhoA and the LKB1/SAD kinase pathway is currently unknown but might involve direct interaction between Par6 and LKB1 or phosphorylation of LKB1 by aPKC, which is known to form a complex with Par6. The downstream effectors of SAD kinases and other LKB1-dependent kinases are currently unknown but might involve local phosphorylation of microtubule-associated proteins such as Tau or MAP1b but probably involve many other effectors, all involved in axon specification. Proteins indicated in red have been shown to be required in vivo for axon specification, whereas proteins indicated in orange have only been studied in dissociated neuronal cultures in vitro.

(Yan et al., 2006). The protein kinase AKT, which we described earlier as being critical for neuronal polarity, appears to undergo selective degradation (Yan et al., 2006). In fact, this degradation selectively targets the inactive pool of AKT in neurites, resulting in a net enrichment of AKT in a single process that contains active AKT, the nascent axon. This phenomenon is consistent with the negative feedback signal model proposed by Arimura and Kaibuchi (2007) to explain axon specification of a single axon during neuronal polarization. Schwamborn et al. showed that the small GTPase Rap1b is regulated by a similar scheme because the active form of Rap1b is spared from degradation and is ultimately enriched in the axon (Schwamborn et al., 2007b). In this case, the ubiquitin ligase acting on Rap1b is Smurf2, whereas the related Smurf1 appears to affect only neurite outgrowth. Additional work demonstrates that an interaction between Smurf2 and the polarity scaffold PAR3 must exist for proper neuron polarization (Schwamborn et al., 2007a). This finding appears to be related to PAR3 targeting of Smurf2 to the axon because perturbation of the interaction of either PAR3eSmurf2 or PAR3eKIF3A results in Rap1b increase in all neurites. The converse situation exists for LIM kinase (LIMK) since levels of this protein must be reduced for axon initiation in vitro (Tursun et al., 2005). Future experiments should clarify the molecular mechanisms regulating the function of Smurf1/2 ubiquitin ligases in axon specification in vivo especially in the context of TGFb signaling (Yi et al., 2010) (Fig. 1.2).

1.5.2 Role of cytoskeletal dynamics in axon initiation and growth Appropriate regulation of the actin and microtubule cytoskeleton is critical for neuronal polarization and has been the focus of numerous studies. Experiments using the actin-destabilizing agents latrunculin B and cytochalasin D indicate that remodeling of the actin-based cytoskeleton is an important regulatory step in axon formation (Bradke and Dotti, 1997).

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PART | I Formation of axons and dendrites

Specifically, actin depolymerization localized to a single neurite in unpolarized stage 2 hippocampal neurons is sufficient to confer axonal identity. A proposed mechanism is that loose actin filaments allow the egress of microtubules and lead to rapid elongation of a given neurite, perhaps outpacing the transport of negative regulators of axonal identity. The idea of cellular asymmetries being reinforced by localized microtubule stabilization and invasion proposed by Kirschner and Mitchison (1986) was elegantly demonstrated using a photoactivatable form of the tubulin-stabilizing compound taxol, which can direct axonal specification to a single immature neurite (Witte et al., 2008). Collectively, these results suggest that a dynamic equilibrium between actin depolymerization and microtubule stabilization plays a role in specifying axonal initiation and axonal identity. Future investigations will need to identify the effectors regulating this balance locally and also characterize how specific upstream signaling pathways might regulate the spatial and temporal control of these cytoskeletal dynamics.

1.5.3 Major signaling pathways involved in axon initiation and growth 1.5.3.1 LKB1 and its downstream kinases SAD-A/B and MARK1-4 A pioneering genetic screen performed by Kemphues and colleagues in the late 1980s identified six Par genes encoding distinct protein families. Many studies have since demonstrated that invertebrate and vertebrate Par genes play critical roles in epithelial cell polarity during development as well as in the context of cell transformation and metastasis (Goldstein and Macara, 2007; Kemphues et al., 1988). Although this pathway is critical to polarity in many species, the signal linking this pathway to extracellular cues has remained elusive. The furthest upstream component known in this cascade is an evolutionarily conserved kinase named LKB1 (also called PAR4). LKB1 translocates from the nucleus and is activated by heterodimerization with one of two related pseudokinases known as Strad-a and Strad-b (Dorfman and Macara, 2008). In addition to binding Strad, LKB1 function in neuronal polarity requires its phosphorylation at S431, a target of both PKA and p90RSK kinases (Collins et al., 2000; Sapkota et al., 2001). This phosphorylation can be triggered by extracellular cues including BDNF (Shelly et al., 2007, Fig. 1.2). This event might be mediated in part by cues that provide chemotactic attraction of radially migrating neurons toward the cortical plate, such as Sema3A (Chen et al., 2008; Polleux et al., 2000), or by other extracellular cues including the neurotrophins (NTs) BDNF/ NT4/NT3 (Shelly et al., 2007), Netrin (Adler et al., 2006), FGFs, or other cues that can activate cAMP-dependent PKA or p90 RSK (RSK1-3). At this point, we can speculate that any other presently uncharacterized serine/threonine PKA, of which there are several, able to phosphorylate S431 on LKB1 might mediate the polarizing function of extracellular cues found in different developing brain regions (Barnes and Polleux, 2009). Future investigation will identify the relevant extracellular cues and the corresponding signaling pathways that trigger phosphorylation of LKB1 at position S431, thereby specifying the axon in developing cortical PNs in vivo (Fig. 1.3). Once LKB1 is activated by binding to its obligate coactivator Strad-a and S431 phosphorylation occurs (only in the neurite that becomes the axon), LKB1 phosphorylates SAD-A/B kinases (and probably the microtubule affinityeregulated kinases MARKs1e4; Matenia and Mandelkow, 2009). These signaling events are required for axon specification in part by the resulting phosphorylation of microtubule-associated proteins (MAPs) enriched in the axon, such as Tau. On the basis of known functions of SAD kinases in presynaptic vesicular clustering in C. elegans (Crump et al., 2001), we hypothesize that SAD-A/B kinases also specify axon identity by directing presynaptic vesicular trafficking in the neurite that is in the process of becoming the axon. Most importantly, genetic deletion of LKB1 in cortical PNs prevents axon formation, whereas overexpression of LKB1 and its coactivator Strad in neural progenitors, or LKB1 alone in postmitotic cells, is sufficient to lead to the formation of multiple axons (Barnes et al., 2007; Shelly et al., 2007). Experiments in Xenopus laevis suggested that LKB1 may regulate aPKC inactivation of glycogen synthase kinase 3 (GSK3)a/b (Ossipova et al., 2003), two proteins involved in neuronal polarity (see later). However, at this point, the exact contribution of LKB1 in adenomatous polyposis coli (APC)/GSK3 function in neuronal polarity is poorly understood. LKB1 also phosphorylates and activates a family of 13 PKAs related to the C. elegans PAR1 protein (Lizcano et al., 2004). To date, three of these have been implicated in regulating axon formation: SAD-A and SAD-B kinases as well as MARK-2 (microtubule affinity regulating kinase-2; also called Par1b). RNAi knockdown of SAD kinases partially abrogates the ability of LKB1 overactivation to induce the formation of multiple axons in cortical neurons, indicating that LKB1’s function in promoting axogenesis largely is derived from its activation of SAD-A/B kinases (Barnes et al., 2007). Double knockout mice for SAD-A and SAD-B results in neurons that cannot form axons in vivo (Kishi et al., 2005), and overexpression of SAD-A/B induces a modest but significant increases in the formation of multiple axons (Choi et al., 2008). SAD and MARK kinases target several MAPs, including MAP2, MAP4, and Tau by phosphorylating three K-X-G-S motifs within each protein, reducing their microtubule binding affinity and thus destabilizing microtubules (Drewes et al., 1997; Illenberger et al., 1996). Little is known about SAD kinase regulation; however, a recent study suggested that the protein phosphatase PP2 might downregulate SAD catalytic activity by reversing LKB1-mediated phosphorylation

Development of neuronal polarity in vivo Chapter | 1

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(Bright et al., 2008). Another study has recently implicated the tuberous sclerosis complex (TSC) genes TSC1/2 in regulating SAD protein abundance (Choi et al., 2008). The microtubule regulatory scheme is the same for the four members of MARK kinase family, but to date only MARK2 has been implicated in neuronal polarity (Biernat et al., 2002; Chen et al., 2006). Because RNAi-mediated knockdown of MARK2 induces supernumerary axons and overexpression of MARK2 inhibits axon formation, it is tempting to hypothesize that MARK2 is a negative regulator of axogenesis (Chen et al., 2006). Intriguingly, MARK2 can interact with the serine/threonine kinase PAK5, and this interaction is thought to inhibit MARK2 kinase activity while simultaneously destabilizing actin cytoskeleton (Matenia et al., 2005). Thus, the MARK2/PAK5 dyad might coordinate actin and microtubule cytoskeletal dynamics during the establishment and/or maintenance of neuronal polarity (discussed below). Several lines of evidence reveal that other potential regulators of neuronal polarity act by regulating MARK2. GSK3a/b can inactivate MARK2 catalytic activity through phosphorylation, and similarly, aPKC can inhibit MARK2 activity through T595 phosphorylation (Timm et al., 2008). The planar cell polarity signaling molecules Dishevelled1 (Dvl1) and Wnt5a also appear to be involved in the MARK2/aPKC pathway of neuronal polarization (Zhang et al., 2007). In this scenario, Wnt5a activation of its receptor Fzl leads to the stabilization of aPKC through its direct interaction with Dvl1. This increase in aPKC activity then leads to an increase in the inhibitory phosphorylation of MARK2. Consistent with this model, increased Dvl1 expression leads to multiple axons, and RNAi knockdown inhibits axon formation (Zhang et al., 2007). Furthermore, the combination of RNAi inhibition of both MARK2 and Dvl1 results in normal axon formation. cJun N-terminal kinase (JNK) is another potential target for Dvl1 signaling (Ciani and Salinas, 2007) and plays a role in neuronal polarization, and inhibition of JNK blocks neuronal polarization in a reversible manner (Oliva et al., 2006).

1.5.3.2 PAR3ePAR6eAPKC The other core components of the polarity complex identified in C. elegans are the scaffolding proteins PAR-3 and PAR-6. Many binding partners for this complex have been implicated in regulating the polarity of epithelial cells. Neuroepithelial radial glia distribute the PAR3/6 complex to their apical domain along the ventricular wall (Bultje et al., 2009; Costa et al., 2008; Manabe et al., 2002). Proteins reported to exist in a complex with PAR3/6 include aPKC and the small GTPase Cdc42 (Joberty et al., 2000; Lin et al., 2000; Qiu et al., 2000), the kinesin motor protein KIF3A (Nishimura et al., 2004), the guanine exchange factor Tiam1/STEF (Chen and Macara, 2005; Nishimura et al., 2005), the lipid and protein phosphatase PTEN (Feng et al., 2008; von Stein et al., 2005), the GTPase-activating protein (GAP) p190RhoGAP (Zhang and Macara, 2008), the tumor suppressor lethal giant larvae (Lgl) (Plant et al., 2003), the scaffold protein inscuteable (Schober et al., 1999), the ubiquitin ligases Smurf1 (Ozdamar et al., 2005) and Smurf2 (Schwamborn et al., 2007b), and the TGFb receptors 1/2 (TGFR1/2; Ozdamar et al., 2005). Each of these proteins has also been implicated in controlling polarity in nonneuronal cells as part of the PAR3/6 complex. PAR3/6 are enriched in the nascent axon in stage 3 hippocampal neurons, and overexpression of wild-type and also truncated forms of either PAR3 or PAR6 perturbs the formation of a single axonal process in hippocampal neurons (Shi et al., 2003). However, in Drosophila, orthologs of PAR3 (bazooka), PAR6, or aPKC do not appear to be required for proper axonedendrite specification (Rolls and Doe, 2004). This could mean that PAR3/6 have acquired a function in neuronal polarity late during evolution in the vertebrate radiation. Alternatively, there is so far no genetic loss-of-function evidence in vertebrates (especially in mammals) demonstrating that Par3 and Par6 are required for axon specification. This evidence will be clearly more challenging to obtain in mammals than in Drosophila because of potential genetic redundancy: There are four Par6 genes (PAR6A-D) and two Par3-like genes in mammalian genomes (Barnes et al., 2008; Goldstein and Macara, 2007). So far, in vivo assessment of Par3 function in the developing cortex using shRNA-mediated knockdown has revealed a clear function with respect to the ability of radial glial progenitors to divide asymmetrically to produce neurons (Bultje et al., 2009), but not as yet in neuronal polarity. Future investigation of in vivo Par3/Par6 functions in axon specification and neuronal polarization will be important. The role of aPKC in neuronal polarity is tightly linked to its ability to associate with the PAR3ePAR6 complex. The activity of aPKC is greatly reduced when associated with PAR6 (Yamanaka et al., 2001). This partnership provides a regulatory scheme that requires additional signaling events to produce a spatially limited pool of activated aPKC, since guanosine triphosphate (GTP)ebound cdc42 binding relieves this inhibition. Phosphorylation targets of aPKC include the PAR3/6-binding partner Lgl, and this posttranslational modification is thought to play a crucial role in regulating the subcellular localization of Lgl during polarization in several cellular contexts (Betschinger et al., 2003; Plant et al., 2003; Yamanaka et al., 2003). As mentioned previously, aPKC can also phosphorylate the PAR1 ortholog MARK2 on threonine 595 (Hurov et al., 2004; Suzuki et al., 2004), in this case inhibiting its kinase activity. Global inhibition of aPKC activity in polarizing neurons clearly demonstrates a role for this kinase in the establishment of neuronal polarity, at least in vitro (Shi

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PART | I Formation of axons and dendrites

et al., 2003), consistent with observed enrichment of activated aPKC in the nascent axon (Schwamborn and Puschel, 2004; Zhang et al., 2007). PAR3/6 axonal localization likely occurs via PAR3 interaction with the microtubule plus-end-directed kinesin KIF3A because interference with KIF3A function leads to the delocalization of PAR3 and aPKC from the nascent axon tip (Nishimura et al., 2004). As discussed previously, inhibition of GSK3 or perturbation of APC localization also eliminates PAR3 targeting from the tip of the nascent axon (Shi et al., 2004), suggesting a hierarchical scheme that enriches the Par3/6 complex in the developing axon.

1.5.3.3 Ras- and Rho-family of small GTPases Small GTPases are critical regulators of cytoskeletal and membrane dynamics underlying cell motility, cell polarity, and cell growth. Small GTPase proteins are molecular switches that generally act on downstream effectors when bound to GTP and are inactive when this GTP is hydrolyzed to guanosine diphosphate (GDP). Rho-GTPases possess relatively slow intrinsic GTP hydrolysis activity, and their catalytic activity is regulated by GTPase-activating proteins (GAPs: 53 predicted in the human genome). GAPs therefore act as negative regulators of GTPase activity by promoting the GDP-bound (inactive) state. Activation of small GTPases by exchanging GDP for GTP is controlled by guanine nucleotide exchange factors (GEFs: 69 predicted in the human genome). Not surprisingly, both Ras- and Rho-family small GTPases have been shown to be involved in axon specification and axon growth. Several members of the Ras-family of small GTPases have been shown to regulate neuronal polarity including H-Ras, R-Ras, K-Ras, and N-Ras. Overexpressing either wild-type or a constitutively active mutant (V12 or Q61L) of H-Ras, or the related protein R-RasQ87L, leads to the production of multiple axons (Fivaz et al., 2008; Oinuma et al., 2007; Yoshimura et al., 2006). Ras proteins regulate both the MAP kinase and PI3K pathways, and pharmacological inhibition of either pathway was sufficient to inhibit the production of additional axons; surprisingly, this did not impact axon formation, in general (Yoshimura et al., 2006). Ras activation is coupled to many cell surface receptors including growth factor receptors, and an EGFR tyrosine kinase inhibitor, AG1478, can inhibit axon formation (Shi et al., 2003). Elegant work using a fluorescent reporter of Ras activation demonstrates the restricted nature of Ras signaling and its recruitment during axon determination to contribute to a positive feedback loop with PI3K (Fivaz et al., 2008). Additional work remains to identify which upstream activators may regulate Ras during the disruption of neuronal symmetry to determine the nascent axon; we discuss in the following some potential candidates in this chapter. The best studies of all mammalian Rho-family small GTPases (there are 22 in total) are Cdc42, RhoA, and Rac1. Expression of dominant-negative (locked in GDP-bound state) or constitutively active (locked in GTP-bound state) mutants of each of these small GTPases in polarizing neurons, or treatment with the Rho-GTPase inhibitor toxin B (Bradke and Dotti, 1999), demonstrates a critical role for both Cdc42 and Rac1 in vitro in rodent neurons (Nishimura et al., 2005; Schwamborn and Puschel, 2004) and in vivo in the Drosophila CNS (Luo et al., 1994). Specifically, the expression of Cdc42L28, a mutant Cdc42 that constitutively cycles between a GDP- and GTP-bound state, leads to the formation of multiple axons in rodent neurons. The loss of Cdc42 expression, through either siRNA knockdown (Schwamborn and Puschel, 2004) or genetic ablation (Garvalov et al., 2007), leads to a strong axon specification defect. In the case of Cdc42 conditional knockout mice, the axon phenotype may be due to increased levels of phosphorylated, inactive, cofilin, which is a regulator of actin dynamics enriched in developing axons (Garvalov et al., 2007). This phosphorylation is achieved by LIMK, an activity stimulated by the Cdc42 effector kinase Pak1. Paradoxically, Pak1 activity is greatly reduced in Cdc42null mice, suggesting that the deregulation of another pathway regulating cofilin occurs in the absence of Cdc42, most likely the RhoA-regulated kinase ROCK (Maekawa et al., 1999). The loss of Pak1 itself also inhibits neuronal polarization, and conversely, constitutively active Pak1 induces multiple tau1-positive processes (Jacobs et al., 2007). The latter effect can be partially reduced by coexpression of either an unphosphorylatable form of cofilin or a GDP-locked Rac1, suggesting that Rac1 may act downstream of Pak1 activation. Taken together, these results demonstrate a role for activated Cdc42 in neuronal polarization beyond its association with the PAR3/6 complex, which is described later in this chapter. RhoA is another small GTPase, and it is typically associated with destabilization of the actin cytoskeleton and also with myosin-based contractility. Experiments using a constitutively active form of RhoA show that it inhibits neuritogenesis, whereas a dominant-negative form of RhoA enhances neurite outgrowth (Schwamborn and Puschel, 2004). This finding is consistent with the regulatory role proposed for p190RhoGAP and the effect of inhibiting the RhoA-activated kinase, ROCK, on axogenesis (Bito et al., 2000). Future experiments will test if RhoA activation is regulated by local degradation through Smurf1/2 activity specifically in the axon downstream of local TGFb receptor activation and recruitment of Par6 (Yi et al., 2010), as previously shown for EMT (Ozdamar et al., 2005). The examination of Rac1’s role in neuronal polarization has led to some confounding results. In Drosophila, the expression of either dominant-negative (GDP-locked) Rac (Luo et al., 1994) or loss of Rac expression (Hakeda-Suzuki

Development of neuronal polarity in vivo Chapter | 1

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et al., 2002; Ng and Luo, 2004; Ng et al., 2002) affects outgrowth but not polarity. Similarly, siRNA knockdown of mammalian Rac1 typically does not affect axon identity (Gualdoni et al., 2007), although some reports detected unpolarized neurons following expression of the dominant-negative form of Rac1 (Nishimura et al., 2005). In cultured neurons, a constitutively active version of Rac1 does not affect axon specification (Schwamborn and Puschel, 2004). These results, while mixed, do hint at a more complex regulation of Rac1 in neuronal polarization. This point becomes clearer, as we discuss later, since the only GEF proteins shown to be crucial for axon formation appear to control Rac1. Small GTPases have a plethora of effectors within cells, and proper activation of these effectors, both spatially and temporally, requires exquisite control of both activation and inactivation by GEFs and GAPs, respectively. Apart from p190RhoGAP, most studies have so far focused on the function of GEFs in neuronal polarity. This includes the GEFs Tiam1 and STEF, described in more detail later, and the DOCK7 GEF, which has recently been reported to be a regulator of axon specification by activating Rac1 and thereby triggering phosphorylation of Stathmin/Op18, a microtubuledestabilizing factor critical for axogenesis (Watabe-Uchida et al., 2006). Another axonally enriched, unconventional Rac1 regulatory protein is the cytoplasmic dynein light chain TcTEX-1 (Chuang et al., 2005). Increased levels of TcTEX-1 result in increases in GTP-loaded Rac1, and a drop in GTP-Rac1 levels is observed following TcTEX-1 knockdown. Multiple axons result from overexpression of TcTEX-1, and this effect is preserved using a mutant form (T94E) that cannot bind dynein heavy chain. Consistent with a role in controlling Rac1 in the neurite specified to become the axon, this supranumerary axon phenotype is suppressed by constitutively active RhoA or dominant-negative Rac1. Therefore, several GEFs seem to cooperate in the local activation of Rac1 during axon specification. Rap1b, a member of the Ras superfamily of GTPases, is also required for proper neuronal polarization in vitro (Schwamborn and Puschel, 2004; Schwamborn et al., 2007b) and in vivo (Jossin and Cooper, 2011). It is found at the tip of the nascent axon, and its overexpression leads to hippocampal neurons bearing multiple axons. The loss of Rap1b following siRNA knockdown abrogates axon formation, and expression of autocycling Cdc42 can rescue the phenotype. Expression of a constitutively active Rap1b fails to reverse the loss of axons observed following loss of Cdc42, indicating that Rap1b lies upstream of Cdc42 in this neuron polarization pathway. Similarly, suppressing axogenesis via pharmacological inhibition of PI3K can be reversed by autocycling either Cdc42 or constitutively active Rap1b, placing both of these small GTPases downstream of PI3K signaling during axon specification. In addition to its role in one of the canonical polarity pathways, studies on Rap1b have explored a novel mechanism for protein localization during neuronal polarity, namely, selective protein degradation (Schwamborn et al., 2007a, 2007b).

1.5.3.4 PI3K and PTEN signaling during axon specification The phosphatidylinositol-3 kinase (PI3K) family regulates diverse biological functions, including cell polarity, cell motility, chemotaxis, and also neuronal migration and polarization; however, these results are based almost exclusively on the use of pharmacological inhibitors such as Wortmanin or LY294002 (Jossin and Goffinet, 2007; Polleux et al., 2002; Shi et al., 2003; Zhou et al., 2007). The best characterized class Ia PI3-kinase (PI3KcIa) is involved in the formation of phosphatidylinositol (3,4,5)-triphosphate (PIP3) and lies downstream of Ras and upstream of protein kinase B ((PKB) or AKT) during signal transduction. Work from several groups has implicated PI3K in axon specification based on the observation that pharmacological inhibition of PI3K activity using LY294002 or Wortmannin prevents axon formation (Jiang et al., 2005; Menager et al., 2004; Shi et al., 2003; Yoshimura et al., 2006). However, these data have to be interpreted carefully since these inhibitors are not class specific. They inhibit the three main classes of PI3K (Stack and Emr, 1994), including phosphatidylinositol-3 kinase class III (PI3Kc3 also called Vps34) that produces exclusively phosphatidylinositol-3 monophosphate (PI(3)P) and regulates endocytosis, vesicular trafficking, trimeric G-protein signaling, and the mTOR (mammalian target of rapamycin) pathway (Backer, 2008). However, overexpression of the constitutively active catalytic subunit of PI3K (p110) leads to the formation of multiple axons (Yoshimura et al., 2006), suggesting that PI3K activation is both required and sufficient for axon specification. Using the pleckstrin homology (PH) domain of AKT fused to GFP (PHAKTeGFP) as a biosensor for PIP3 formation, Menager et al. (2004) have shown that PIP3 accumulates selectively within a single neurite following local application of laminin to stage 2 hippocampal neurons. Future investigations will need to address when and where PI3K activation occurs in vivo during neuronal polarization and also which class of PI3K is required (class 1, 2, or 3), using genetic loss-of-function approaches. Two PI3K-interacting proteins, Shootin1 (Toriyama et al., 2006) and Singar1/2 (Mori et al., 2007), were recently identified as potential regulators of axon formation using a mass spectrometry approach. Overexpression of Shootin1 leads to the generation of supernumerary axons, and RNAi knockdown inhibits axon formation. Shootin1 is transported via a myosin-dependent mechanism to axonal growth cones, and its overexpression leads to aberrant accumulation of PI3K in minor neurites, likely leading to the observed alteration of axon specification. Shootin1 colocalizes with active pools of

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PI3K, and inhibition of PI3K activity significantly reduces the ability of Shootin1 to induce multiple axons. These data suggest a role for Shootin1 in regulating PI3K activity, and its selective transport to the nascent axon is likely critical for establishing axonal identity. Singar exists as at least two protein isoforms generated by alternative splicing, Singar1 and Singar2, and both are expressed in developing neurons. RNAi against both forms causes cultured neurons to form multiple axons, and this effect is prevented when PI3K activity is inhibited. Unlike Shootin1, overexpression of Singar is not sufficient to affect axon formation. When coexpressed with Shootin1, Singar1, but not Singar2, can reduce the multiple axon phenotype caused by Shootin1 overexpression alone. This result suggests an antagonistic relationship between Shootin1 and Singar 1 and also that Singar protein may inhibit PI3K activity. PTEN (phosphatase and tensin homolog deleted on chromosome 10) is a lipid and protein phosphatase that acts in direct opposition to PI3K activity since PTEN dephosphorylates PIP3 into PIP2 and thus limits PIP3 signaling both spatially and temporally. Increasing levels of PTEN expression lead to a loss of axon formation (Jiang et al., 2005; Shi et al., 2003), whereas reduction of PTEN expression via RNAi-mediated knockdown leads to a multiple axon phenotype (Jiang et al., 2005). This effect is consistent with the gain-of-function mutation of PI3K described earlier and highlights the critical need to maintain the delicate balance of phospholipid composition at the membrane to ensure proper neuronal polarization and axon formation. Remarkably, PTEN serves a major role in suppressing injury-induced CNS axon growth via regulation of the mTOR pathway (Park et al., 2008; see later).

1.5.3.5 AKT/protein kinase B Several proteins are recruited via their PIP3-specific PH domains to sites of membrane created by PI3KcIa activity. The protein kinase AKT, also called PKB, undergoes such a translocation to the membrane via its PH domain, a step required for its dual phosphorylation on T308 and S473 and subsequent activation by the membrane-targeted protein kinases PKD1 and PKD2, respectively. This activated form of AKT is enriched in growth cones of polarized neurons (Shi et al., 2003). When a myristoylation site is added to recombinant AKT (myr-AKT), it is constitutively targeted to the membrane, independent of PI3K, and therefore is constitutively active. When overexpressed in neurons, this form of AKT is sufficient to generate multiple axon formation (Yoshimura et al., 2006), consistent with a unified pathway in which AKT acts downstream of PI3K to regulate axon formation. In addition to PKD, another PI3K-regulated kinase, ILK (integrin-linked kinase), increases AKT activity via S473 phosphorylation (Delcommenne et al., 1998). Similar to other regulators of AKT, hyperactive ILK (S343D) increases multiple axon formation in cultured neurons, and RNAi reduction or pharmacologic inhibition of ILK leads to the failure of axon formation without affecting the dendritic formation (Guo et al., 2007). These experiments point to an important, but not exclusive, role for ILK in regulating AKT. A common target protein of both ILK and AKT is GSK3b (Delcommenne et al., 1998), and phosphorylation by either protein is thought to inactivate GSK3 kinase activity (see later). Future experiments will need to address which class of PI3K is required for axon specification and also if PTEN and AKT are required for neuronal polarization in vivo. Finally, we also need to know whether these important effectors are specifically required for neuronal polarization and axon specification, or rather, if they are more generally required for neural progenitor polarity. It remains possible that the defects in polarization of postmitotic neurons caused by alteration of PTEN or AKT function are a consequence of an earlier requirement in specifying neuroepithelial polarity (Barnes et al., 2008).

1.5.3.6 GSK3 and axon specification GSK3 proteins are well-studied serine/threonine protein kinases that function in the regulation of multiple intracellular processes, including pathways downstream of receptor tyrosine kinases and Wnt/Fzl signaling. The two genes encoding GSK3 kinases (a and b) in mammals produce proteins that perform essentially redundant functions. GSK3s have the unusual property of being constitutively active, a state that is reversed following phosphorylation at Ser9 in GSK3b or Ser21 in GSK3a by multiple kinases, including AKT, ILK, and aPKC (Etienne-Manneville and Hall, 2003). Recent in vitro work implicates GSK3b as a critical regulator of neuronal polarity. Experiments using several types of GSK3 inhibitors indicate that GSK3b acts as a negative regulator of axon formation because this treatment leads to the formation of multiple axons (Jiang et al., 2005; Yoshimura et al., 2005). Gartner et al. (2006) suggested that the situation is clearly more complex in vivo. Using double knock-in mice bearing single point mutations GSK3ßS9A and GSK3aS21A, Gartner et al. reported no obvious deficits in neuronal morphogenesis in vivo and in vitro (Gartner et al., 2006). In fact, these mice are viable and do not show any obvious CNS developmental phenotypes. However, using inhibitors of GSK3a/b such as lithium chloride or, more specifically, SB-415286, SB216763, or AR-A014418, Gartner et al. were able to replicate the multiple axon phenotype obtained by others (Garrido et al., 2007; Jiang et al., 2005). These results indicate that although the exact role of Ser9/Ser21 phosphorylation in GSK3

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inactivation remains to be understood and may involve an alternate site (Thornton et al., 2008), it is clear that the catalytic activity GSK3s are a critical regulator of neuronal polarity in standard in vitro paradigms. Several downstream targets of GSK3s are potential effectors of neuronal polarity, and many involve regulation of the cytoskeleton. Collapsin response mediator protein-2 (CRMP-2) is one such microtubule-binding protein that is enriched in tips of the nascent axon and is regulated by GSK3b such that phosphorylated CRMP-2 displays a decreased binding affinity for tubulin heterodimers (Inagaki et al., 2001; Yoshimura et al., 2005; reviewed in Arimura et al., 2004). As observed for other polarity regulators, overexpression of CRMP-2 is sufficient to induce the formation of multiple axons, and truncated forms of CRMP-2 can impair axon formation (Inagaki et al., 2001). Although the ability of CRMP-2 to facilitate microtubule assembly is important in regulating axon formation, CRMP-2 is also known to associate with several other proteins, including the actin polymerization-regulating Sra-1/WAVE1 complex, which might contribute to its function in axogenesis. Kawano et al. showed that CRMP-2 links the Sra-1/WAVE1 complex with the microtubule-based motor protein Kinesin 1, and Sra1/WAVE expression is likely required for CRMP-2’s induction of multiple axons (Kawano et al., 2005). APC is another well-established effector of GSK3a/b that are enriched in the neurite that will become the axon early during neuronal polarization (Shi et al., 2004). Most cell biological evidence suggests that APC enhances microtubule stability, and it is well established that APC can bind microtubule plus ends via its EB1-binding domain. Phosphorylation of APC by GSK3b blocks its ability to bind the plus ends of microtubules, and inhibition of GSK3b leads to an accumulation of APC in multiple neurites (Shi et al., 2004). Expression of truncated forms of APC is sufficient to inhibit axon formation and elongation (Shi et al., 2004; Zhou et al., 2004). Reduction of APC using shRNA has also been demonstrated to interfere with efficient axon elongation (Purro et al., 2008). Additional work shows that the APC/GSK dyad regulates targeting of another polarity protein, PAR3, since overexpression of full-length or truncated APC disrupts neuronal polarization, and inhibition of GSKb changes the distribution of the pool of APC in the nascent axon (Shi et al., 2004; Zhou et al., 2004). Furthermore, growth factoretriggered inactivation of GSK3b by PI3K signaling acts through the APC to control axon elongation (Zhou et al., 2004). Investigators have observed similar results for two other GSK targets, the MAPs, MAP1b (Gonzalez-Billault et al., 2004) and Tau (Sperber et al., 1995), that, when phosphorylated by GSK3b, alter microtubule dynamics. These results emphasize a key principle that underlies much of what is known about neuronal polarizationdthe microtubule cytoskeleton is a major endpoint for polarity regulators. Interestingly, PTEN was also recently identified as a GSK3 substrate (Maccario et al., 2007), which may represent a negative feedback loop for AKT signaling following activation via stabilization of PTEN. As noted above, the two GSK-3 family members, GSK-3a and GSK-3b, have largely redundant functions. Elimination of either family member alone has little effect on axon growth either in vitro or in vivo (Kim et al., 2006). However, complete inhibition of GSK-3 activity via inhibitors or shRNA directed at both family members appears to block axon growth altogether in vitro (Garrido et al., 2007; Kim et al., 2006; Shi et al., 2004).

1.6 Conclusion and future directions The mechanisms that underlie neuronal polarization, the specification of axon and dendrite identity, their morphology, and directed outgrowth in vivo have parallels with the mechanisms determined using reductionist in vitro models. However, there are several important distinctions. They include the complex interactions between extracellular cues, such as graded diffusible cues such as TGFb and/or cell adhesion molecules presented by preexisting axons such as TAG-1 (Yi et al., 2010; Namba et al., 2014), and the intracellular pathways they activate specifically in the neurite (trailing process) that becomes the axon can only be studied in vivo. Among the many outstanding issues for the field to address relating to the link between these extracellular/intracellular signaling pathways and the cellular events they control are (1) process-specific neurite outgrowth (significantly faster for the neurite becoming the axon than neurites becoming dendrites); (2) the selection of the cargoes that will be specifically targeted and transported to the axon versus the dendrite (see, for example, reviews by Gumy and Hoogenraad, 2018; Lewis et al., 2013); and (3) the striking, compartment-specific, distribution and morphology of organelles such as the endoplasmic reticulum or mitochondria (see, for example, Lewis et al., 2018). As an excellent example of this last point, in cortical PNs, mitochondria display compartment-specific morphologies: axonal mitochondria are short and of a standard length (w1 mm), whereas dendritic mitochondria are long and fused and fill almost the entire dendritic tree (Lewis et al., 2018). This observation suggested that mitochondrial fusion must dominate over fission in dendrites and vice versa in the axon of these same neurons. In a recent study, Lewis et al. identified the Drp1 receptor Mff as a critical regulator of the high levels of mitochondrial fission observed in axons, but the mechanisms underlying this compartment-specific level of mitochondrial fission are still poorly understood. This study constitutes a new paradigm for exploring some of these

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mechanisms of neuronal polarization for organelles as well as other cargoes. These include neurotransmitter vesicles specifically in the axon, or postsynaptic cargoes such as neurotransmitter receptors specifically in dendrites (Gumy and Hoogenraad, 2018). The basic mechanisms underlying neuronal polarization are beginning to be understood at the cellular and molecular levels using in vitro models; however, our understanding of how these mechanisms operate in vivo is still fragmentary but will undoubtedly be the focus of many investigations in the near future.

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Chapter 2

Role of the cytoskeleton and membrane trafficking in axonedendrite morphogenesis Kevin C. Flynn1 and Frank Bradke2 1

Stem Cell and Gene Therapy, Bio-Techne, Minneapolis, MN, United States; 2Laboratory for Axon Growth and Regeneration, German Center for

Neurodegenerative Diseases (DZNE), Bonn, Germany

Chapter outline 2.1. Introduction 2.2. Developmental stages 2.3. Role of cytoskeleton in establishment of neuronal polarity 2.3.1. Actin 2.3.2. Actin dynamics during axon formation 2.3.3. Microtubules 2.3.4. Microtubules dynamics during axon formation 2.3.5. Cytoskeletal dynamics during dendritic growth and arborization 2.3.6. Subcellular cytoskeletal specializations 2.4. The role of (membrane) trafficking during neuronal polarization

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2.4.1. Trafficking during early neuronal development 2.4.2. Motor proteinebased transport in axons and dendrites 2.4.3. The secretory and endosomal pathway 2.4.4. RNA transport and local translation 2.4.5. Barriers for the segregation of functional domains 2.4.6. Protein stabilization and degradation 2.5. Maintaining neuronal polarity 2.6. Future work on neuronal morphogenesis References

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2.1 Introduction Neurons can be among the most morphologically complex cells with multiple extensions traversing far-reaching distances and convoluted pathways to connect with other cells (e.g., upper motor neurons of the cerebral cortex), but they can also be relatively simple with short processes making local contacts (e.g., GnRH neurons). All neurons, however, share a common specialization that is crucial to their function: They are polarized. This specialization entails an intricate morphogenesis in which initially spherical cells extend processes in a precise manner to create circuits that, although plastic and subject to modification, can function throughout an organism’s lifetime. As the signaling units of the nervous system, neurons require discrete functional domains. To receive, store, and transmit electricechemical signals, a neuron must develop subcellular compartmentalization, with each compartment assuming a specific morphology and molecular constitution. The development of a neuron with such specialization requires a sophisticated developmental program that is collectively called neuronal polarization (Craig and Banker, 1994). Although Ramon y Cajal introduced the theory of neuronal polarity with dendrites receiving information and axons transmitting signals to downstream cells in the early 1900s (Shepard, 1991), debate over the nature and development of neuronal polarity continued well into the 20th century. It was not until the pioneering work by Gary Banker, Carlos Dotti, and colleagues optimized mammalian hippocampal cultures that the concept of neuronal polarity become mainstream and ability to study its development feasible (Banker, 2018). At the broadest level, the differentiation of the somatodendritic compartment and the axon is common to nearly all neurons (Fig. 2.1A). Additionally, many neurons have even more

Cellular Migration and Formation of Axons and Dendrites. https://doi.org/10.1016/B978-0-12-814407-7.00002-X Copyright © 2020 Elsevier Inc. All rights reserved.

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(A)

Somatodendritic compartment

Axonal compartment

Direction of signal propagation Synapse

AIS

Synapse (NMJ)

Nodes of Ranvier

UMN

LMN

Muscle

(B)

Stage 1 0–0.25div

Stage 2 0.25–2div

Stage 3 2–4div

Stage 4 4–14div

Stage 5 14–21div

FIGURE 2.1 Morphology of polarized neurons and the stages of neuronal development. (A) A lower motor neuron (LMN) exemplifies the functional and morphological polarity of a neuron. On the anatomical and functional level, neurons are subdivided into the somatodendritic compartment, which receives signals (in this case from an upper motor neuron [UMN]) and the axon, which transmits signals to the next cell (in this case a muscle). The axon initial segment (AIS) separates these two domains acting as a selective barrier preventing the mixing of the distinct molecular components of the two domains. Neurons exhibit further subcellular polarity into distinct domains based on function and underlying molecular composition. For example, dendritic spines are sites of synaptic input and enriched in neurotransmitter-gated ion channels responsible for producing synaptic potentials. Myelinated axons have nodes of ranvier, which, like the AIS, are enriched in voltage-gated channels necessary for the propagation of action potentials. (B) Neuronal development can be divided into a series of five consecutive stages of development based on observations from hippocampal neurons in culture (Dotti et al., 1988). This developmental program is stereotypical and highly reproducible. Shortly after plating, the spherical cell bodies extend filopodia and lamellipodia and assume a “fried egg” morphology (stage 1). Within hours, neurons begin extending immature neurites, which are indistinguishable (stage 2). During the stage 2e3 transition, one of the immature neurites elongates and begins acquiring the morphological and molecular features of an axon. This occurs at 2e3 days in vitro (div) and marks the first sign of polarity (stage 3). After 4 days to a couple weeks in culture, the remaining neurites begin to arborize and acquire dendritic characteristics (stage 4). Between 2 and 3 weeks in culture, there is the continued maturation of the axon and dendritic arbors with the formation of dendritic spines and mature neuronal circuits (stage 5).

complex patterns of intracellular specialization necessary for their function. Thus, it is not surprising that even when subtle problems arise during the establishment or maintenance of neuronal polarity, serious neurological diseases can develop including lissencephaly, autism, and amyotrophic lateral sclerosis (Maussion et al., 2008; Rasband, 2010; Kanning et al., 2010). The establishment of neuronal polarity is, thus, an essential developmental process underlying the function of individual neurons and the nervous system as a whole. Cell polarity is not unique to neurons, as many diverse cell types require some form of polarization. From budding yeast to dividing stem cells, from epithelial cells to dendritic cells of the immune system, a subcellular division of labor is a commonality necessary not only for the formation and function of individual cells but also for complex multicellular tissues. Many of the basic principles of cell polarization are common to all these diverse cell types, including neurons. However, distinct mechanisms are also employed to establish the unique functions of each cell type. Neuronal function demands a unique morphology and division of labor to ensure the fidelity of the unidirectional flow of communication in the nervous system (Fig. 2.1A). To this end, neurons typically have multiple dendrites, which receive synaptic inputs, and a single axon, which transmits signals via action potentials to the next cell. To

Role of the cytoskeleton and membrane trafficking in axonedendrite morphogenesis Chapter | 2

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perform these functions, the axon and dendrites are strikingly different in their morphology, cytoskeletal organization, molecular constitution, and membrane composition. Thus, the morphogenesis and the segregation of specific molecular components of the axon and dendrites underlie the core process of neuronal polarization and, thus, neuronal function. The morphogenesis occurring during neuronal polarization requires the spatially and temporally coordinated regulation of the actin and microtubule cytoskeletons. The foremost event in neuronal polarization is the formation of the axon. Thus, the remodeling of the cytoskeleton first occurs to induce the growth and branching of the developing axon, while concomitantly repressing the growth of the other processes, the future dendrites. Only later, specialized cytoskeletal dynamics transpire to facilitate dendritic growth, arborization, and the formation of discrete postsynaptic specializations (e.g., dendritic spines). During the early events of neuronal polarization, the elongation of the axon requires material to support increase in surface area and volume. Trafficking of membranous and cytosolic components is regulated to generally support this overall increase of axonal mass. As development proceeds, the formation of functional domains during neuronal development depends on trafficking mechanisms that result in a polarized delivery system. This molecular “sorting” is achieved by diverse-layered and interdependent mechanisms including discriminatory secretory and endosomal transport, physical barricades barring the mixing of intercompartmental material, and selective degradation of unwanted material. The fact that the maintenance of neuronal polarity also requires the continued regulation of these intracellular sorting mechanisms indicates that these events are not only important during development but also throughout the life span of an organism. This chapter focuses on the essential contributions of the cytoskeleton and intracellular trafficking to neuronal polarization. General features and signaling pathways implicated in the process of neuronal polarization are covered in the previous chapter (Chapter 123). Therefore, here, we will only generally discuss principles of these signaling pathways, as they pertain to the direct regulation of the cytoskeleton and cellular trafficking. Since axon formation is the foremost event during neuronal polarization, we will largely focus on the cytoskeletal and trafficking mechanisms underlying the initial specification of the axon. However, as a neuron matures, trafficking mechanisms are established that continue to differentiate the axonal compartment from the somatodendritic compartment. Therefore, our discussion of neuronal polarization will also cover various sorting mechanisms that occur as neurons become functionally mature. We will briefly discuss dendritic development in the context of the cytoskeleton, but specific details of dendritic development are reviewed elsewhere in this book (Chapter 11 and 12). The basic principles of cellular polarization, some of which are applicable for neuronal polarization, are also reviewed in other chapters of this book (Chapter 24e25).

2.2 Developmental stages Dissociated neuronal culture systems, in particular cortical and hippocampal neurons and, to a lesser degree, cerebellar granule neurons (CGNs), have provided the majority of our knowledge regarding the mechanisms of neuronal polarization. Hippocampal and cortical neurons undergo a stereotypical developmental progression in a manner similar to their in vivo development, divided into five consecutive stages (Dotti et al., 1988; Craig and Banker, 1994, Flynn, 2013; Fig. 2.1B). Within hours of plating, the neurons assume a fried egg morphology extending broad circumferential lamellipodia and filopodia protrusions (Stage 1). During neuritogenesis, lamellipodia and stable filopodia form growth cones and begin extending forming multiple nascent neurites (stage 2). At this point in development, the neuron is still symmetric, as any of these neurites has the potential to form the axon. During the stage 2e3 transition, the initial symmetry breaking event occurs as one neurite begins growing more rapidly transforming into the morphologically and molecularly distinct axon (stage 3). The remaining processes later grow and arborize, developing into dendrites (stage 4). After around 2 weeks in culture, neurons mature and develop dendritic spines, and mature neuronal networks are established (stage 5). The process of axon formation (i.e., axonogenesis) is the decisive step underlying neuronal polarization and therefore will be the focus of this chapter. However, other later events in polarization are equally important for neuron function. Following axonogenesis, neurons continue to acquire subcellular specializations, which are crucial for development and maintenance of neuronal polarity. The orchestration of a dynamic cytoskeleton and intracellular trafficking are the principle mechanisms for the continued process of neuronal polarization. Our knowledge regarding the development of neuronal polarity largely relies on studies performed in cell culture. Although recent advances in imaging and tissue culture platforms have confirmed many aspects of the basic concepts of neuronal polarization, there are notable differences in more physiological contexts. This is dependent on the brain region and developmental stage with the surrounding playing a key role in the determination of spacial and temporal features of polarization. In mammalian embryonic development, the majority of cortical and hippocampal neurons are born in the ventricle zone (VZ) and subventricular zone (SVZ) and migrate toward the intermediate zone of the brain. During this migration, they extend multiple minor neurites, and these multipolar stage (MP) neurons in vivo resemble stage 2 neurons

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PART | I Formation of axons and dendrites

in culture (Noctor et al., 2004; Takano et al., 2015). Neuronal polarization largely occurs coincidentally with migration; as the neuron moves into the outer layers of the cortex, a trailing process extends rapidly and develops into the axon. Meanwhile, other processes retract with the exception of a leading process (a future dendrite) as the neuron becomes bipolar and continues its migration to the cortical plate. Although this progression occurs in the majority of neurons, pyramidal neuronal development can occur in different ways; pioneer callosal axons extend from one process of a multipolar neuron before migration early in mammalian brain development (Hand and Polleux, 2011). Indeed, there are many examples of perturbations of key molecular players that have similar effects on neuronal development in vivo and in cell culture (Garvalov et al., 2007; Tahirovic et al., 2010; Flynn et al., 2012). Furthermore, in vivo neuronal polarization of pyramidal neurons proceeds from a progenitor void of inherit polarity, which proceeds to a multipolar stage whereby axon formation is the first symmetry breaking event (Funahashi et al., 2014). While axon initiation is random in cell culture, environmental cues such as TGF-b impinge upon this naïve neuron in vivo to direct axon formation (Yi et al., 2010). Thus, at least in pyramidal neurons, axon growth initiates neuronal polarity in culture in a manner analogous to the in vivo setting. This is not the case in other systems; in retinal ganglion neurons, apicalebasal polarity is inherited from neuroepithelial progenitors, and the direction of axon growth is preordained (Randlett et al., 2011). Overall, with caution to not over interpret findings, the key cytoskeletal features of polarization found in neuronal culture are likely applicable to the in vivo situation.

2.3 Role of cytoskeleton in establishment of neuronal polarity Like all eukaryotic cells, neurons contain networks of fibrous elements including actin, microtubules, and intermediate filaments, known collectively as the cytoskeleton. It not only provides structural support of the neuron itself but also serves as tracks on which the intracellular movement of organelles, chromosomes, proteins, and RNAs depends. Moreover, the cytoskeleton powers the movement of the neuron and the extension of neurites. As the cytoskeleton undergoes dynamic changes, a neuron moves, extends neurites, and undergoes the morphological transformation of polarization. Therefore, the intricate regulation and coordination of the components of the cellular cytoskeleton is crucial to neuronal morphogenesis. The initial symmetry breaking event in neuronal polarization is the elongation of the axon. This is when the morphological and molecular distinctions that later define neuronal function are established. Strong evidence has accumulated over the past few decades that a network of feedback loops specifies the future axon as one neurite extends longer than the other neurites of an immature, multipolar neuron (stage 2). These feedback loops are positive in the emerging axon, reinforcing axon growth, and fate commitment while concomitantly generating global negative feedback loops that repress neurite growth in the other processes (Schelski and Bradke, 2017). Invariably, these feedback loops involve the transport and regulation of signaling molecules that converge on the actin and microtubule cytoskeletons. Beside kinases, phosphatases, cyclic nucleotides, and calcium ions, the Rho family of small GTPases controls diverse aspects of the cellular cytoskeleton to elicit changes in the shape and motility of diverse cell types. For neurons, the Rho GTPases are regarded as pivotal signaling switches that manage neuronal morphogenesis through a diverse set of effectors controlling specific changes in actin and microtubules (Govek et al., 2005; Hall and Lalli, 2010). Indeed, several proposed feedback signals that promote axon polarization regulate PI3 kinase activity (PIP3), which steers morphological changes via the modulation of the cytoskeleton via Rho GTPases (Schelski and Bradke, 2017). The three canonical Rho GTPases, Rho, Rac, and cdc42, are particularly important in neuronal polarization. These pivotal signaling proteins have divergent effects on the cytoskeleton, effecting actin structural changes via actin nucleators, actomyosin contractility, and actin-severing proteins (e.g., cofilin) while concomitantly altering microtubules via end-binding proteins (e.g., CLASPs, EBs), CRMP2, and microtubule-associated proteins (MAPs) (e.g., Tau) (Govek et al., 2005). Whereas Rac1- and cdc42-mediated signaling tends to promote neuronal polarization, RhoA signaling pathways attenuate axon growth (Hall and Lalli, 2010). For example, the genetic deletion of cdc42 results in a loss of filopodia and prevalence of looping microtubules, leading to a marked attenuation of axon development (Garvalov et al., 2007). Rac1 deletion results in a reduction of neuronal polarization in cerebellar neurons with a loss of lamellipodia (Tahirovic et al., 2010). Interestingly, while both of these Rho GTPases affect neuronal polarization, they do so by affecting different cytoskeletal effectors. Thus, it may be that neuronal polarization can be regulated by different cytoskeletal regulators as long as the cytoskeletal dynamics are permissive for neuronal polarization.

2.3.1 Actin Actin structural organization and dynamics are vitally important for neuronal polarization. During polarization, various signaling pathways and actin-binding proteins (ABPs) modulate dynamics and/or the structural organization of the actin cytoskeleton.

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Of the six actin isoforms found in mammals, there are two actin isoforms predominantly expressed in neurons at near equal levels: b-actin and g-actin. Three a-actin isoforms are generally expressed at lower levels in neurons. Despite over 90% amino acid sequence homology and overlapping biochemical properties, b-actin and g-actin have distinct localizations and functions in vivo. Whereas g-actin localizes to the cell body (Otey et al., 1986; Kashina, 2006), b-actin is more specifically enriched to dynamic actin networks of the cell, such as growth cones (Bassell et al., 1998). Functional differences between the different isoforms are normally attributed to differences in their N-termini, with elasticity inversely related to polymerizationedepolymerization kinetics. b-Actin-rich actin polymers are highly dynamic and show high depolymerization rates compared with filaments consisting of higher levels of a- and/or g-actin (Bergeron et al., 2010). These differences may also reflect the increased propensity for actin dynamizing proteins such as cofilin to bind b-actinrich actin filaments (De La Cruz, 2005). More recently, knockout mice have provided evidence for the specific and overlapping functions of actin isoforms (Cheever and Ervasti, 2013). Interestingly, a smooth muscle a-actin isoform, which is normally nominally expressed in the brain, is massively upregulated in b-actin knockout mice, which together with g-actin can compensate for the loss of b-actin (Cheever et al., 2011). A recent study using CRISPR/Cas9 elegantly showed that swapping of the g-actin into b-actin gene position, thereby completely ablating all b-actin, had no developmental defects (Vedula et al., 2017) in comparison with the complete b-actin knockout mice, which is embryonic lethal (Bunnell et al., 2011). Therefore, b-actin is only important for neuronal development in that it is simply the most highly expressed isoform, which is dependent on the gene’s favorable position in the genome rather than the gene itself. Actin exists in two forms: a monomeric, globular (G-actin) and a polymerized, filamentous form (F-actin). In individual actin molecules, there is a deep nucleotide binding cleft that is occupied by one of three adenine nucleotides: ATP, ADP-Pi, or ADP (Pollard et al., 1992, Box 2.1). In neurons, the majority of G-actin is bound with ATP, whereas ADP-Pi and ADP bound actin predominate in actin filaments. Actin filaments are long chains of G-actin formed into two parallel polymers twisted around each other into a helical orientation with a diameter between 6 and 8 nm. There are two rotational states of F-actin in regard to this twist: a more twisted form and a less twisted form. This rotational status of F-actin has important consequence for actin structures and dynamics; more twisted actin has less thermodynamic stability and can be disassembled more easily. Furthermore, the changes the topology of the actin filament and can influence the binding of proteins that modulate actin organization and turnover. For example, actin-depolymerizing factor (ADF)/cofilin proteins, which are crucial for neuronal polarization, bind to the more twisted form of actin. In neurons, as in other cells, actin filaments are assembled into higher-order networks with distinct spatial organizations determined by specific repertoires of ABPs. These include filament meshworks in lamellipodia, linear bundles in microspikes and filopodia, actin arcs, and cortical actin juxtaposed to the cell membrane. In growth cones, the motile tip of neurites, some combination of these actin structures, is always present (Fig. 2.2). Growth cones, although pleiomorphic,

BOX 2.1 Actin dynamics Actin monomers can spontaneously self-assemble into isolated actin filaments. Following nucleation and an elongation phase, actin filaments can reach a steady-state polymer mass in which they are still dynamic. An actin filament is structurally and functionally polarized such that two ends of the filament have different equilibrium constants with actin monomers for assembly, called critical concentrations. When the concentration of G-actin lies in between the different critical concentrations, net assembly occurs at one end (the barbed or plus end), and net disassembly occurs at the other end (the pointed or minus end) in a process called treadmilling (Bugyi and Carlier, 2010 This polarity of the filament is also reflected by the adenine nucleotide binding status of the actin subunits. ATPeactin, which dominates the actin monomer pool, is added preferentially to the plus end of actin filaments. The ATP rapidly hydrolyzes, although the inorganic phosphate does not disassociate, resulting in ADPePieactin. Eventually, the Pi dissociates resulting in ADPeactin accumulation toward the minus end of the filament. The kinetics of ATP hydrolysis and Pi release results in a molecular aging or stratification, such that ATPeactin is scarce and specifically enriched toward the,

“young,” plus ends of filaments, whereas ADPePi makes up the middle-aged portion of the filament and ADPeactin makes up the older portion of the filament toward the minus end. This inherent polarity and molecular stratification have important implications for cellular dynamics, as different actin-binding proteins have different affinities for specific nucleotide-bound actin subunits, and this could influence the formation and dynamics of actin structures (Pak et al., 2008). Actin treadmilling Cc– > G-actin concentration > Cc+ Pointed end (−)

Barbed end (+)

ADP-actin

ADP-Pi-actin

ATP

ADP

ATP-actin

26

PART | I Formation of axons and dendrites

(A)

Stage 2–3 neuron

(B)

Peripheral domain

Transitional domain

Central domain Microtubules/F-actin

(C)

(D)

Actin nucleation and filament growth at the plus ends of filaments pushes leading edge forward

F-actin retrograde flow is result of F-actin polymerizationdepolymerization and actomyosin contraction

Actin arcs and bundles formed in the transitional domain by actomyosin contractility constrain microtubules

(E)

F-actin retrograde flow

Actin severing and depolymerization

Microtubule growth is enhanced by various complementary factors Microtubules grow along F-actin bundles

Legend ATP-actin

Increased F-actin polymerization pushes filopodia forward

Protrusion

ADP-Pi-actin ADP-actin Arp2/3 Formin Drebrin

GTP-E tub GTP-D tub GDP-E tub GTP-D tub Stathmin CRMP-2 KIF2A EB-1

ADF/cofilin

F-actin retrograde flow Protrusion

Ena/Vasp Myosin II

EB-3 Clip170/115

FIGURE 2.2 The neuronal cytoskeleton and the growth cone. (A) Immunostaining of a stage 2e3 hippocampal neuron shows the two of the major components of the neuronal cytoskeleton: microtubules (green) and actin (red). Microtubules comprise the major structural element of the cell body and the neurite processes, whereas F-actin localizes mainly to the growth cones. (B) Expanded schematic of the large growth cone from the neuron in (A) Growth cones are classically subdivided into three distinct regions: the peripheral domain, the central domain, and the transition zone. The peripheral domain contains linear actin bundles underlying filopodia and a meshlike actin network in lamellipodial veils. The central domain is largely devoid of actin filaments but contains radially oriented microtubules that are bundled proximally and splay out distally occasionally extending into the peripheral growth cone. The transition zone demarcates the central domain and the peripheral domain and contains transverse bundles of actin filaments called actin arcs. (C and D) Expanded regions indicated in panel B show details of cytoskeletal organization and dynamics in different regions of the growth cone. (C) Actin arcs are generated in the transition domain, as actin filaments are reorganized and compressed via the actions of myosin II contractility and the pressure of actin polymerization at the leading edge. Microtubules are constrained into the central domain or reoriented by the actin arcs. Microtubules can undergo rapid disassembly or catastrophies, which can be aided by KIF2A. The occasional microtubule extends beyond the transition zone into the peripheral domain. (D) The meshwork actin organization and dynamics in lamellipodial veils is thought to be the result of various actin-binding proteins. Arp2/3 complex network nucleates actin branches off existing actin filaments and other actin-binding proteins (e.g., filamin, not shown) maintain actin filaments in a branching pattern. The barbed (þ) ends are oriented toward the membrane, so actin polymerization occurs mainly at the leading edge of the

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typically have three distinct regions: a peripheral domain, a transition zone, and a central domain (Lowery and Van Vactor, 2009). The peripheral domain has multiple radially oriented filopodia with lamellipodia veils in between. In the transition zone, transversely oriented actin arcs are located. The central domain is largely devoid of F-actin but rich in microtubules and organelles. Like the underlying individual filaments, these actin structures in the peripheral growth cone are extremely dynamic, being constantly remodeled to direct the movement of the growth cone. A vital aspect of this dynamics is actin retrograde flow, which is the rearward displacement of actin filaments away from the leading edge toward the center of the growth cone. Actin retrograde flow is the combined result of actin polymerization and depolymerization kinetics (treadmilling) in coordination with actomyosin contractility (Mederios et al., 2006, Fig. 2.2). The polymerization of actin filaments occurs preferentially at the leading edge of growth, which is likely the result of the enrichment of proteins that promote de novo F-actin assembly, such as formins or WASP, which adjoin actin assembly to the membrane (Pak et al., 2008). In growth cones, this results in an actin network organizational matrix with the plus ends of actin filaments oriented distally and the pointed ends oriented proximally. The important consequence of this is that the direction of filament growth is always perpendicular to the leading edge membrane and parallel to the direction of neurite advance. The combined effect of the polymerization of networks of actin filaments can provide a pushing force to drive membrane expansion, but only under certain conditions. The so-called “clutch hypothesis” ventures that the physical coupling of actin filaments to components of the extracellular matrix (ECM) via transmembrane focal contacts transmits a traction force (the clutch is engaged), allowing actin polymerization to push the membrane forward while diminishing retrograde flow rates (Mitchison and Kirschener, 1988; Suter and Forscher, 1998). Increasing the concentrations of ECM, such as N-cadherin, does indeed increase axon outgrowth and growth coneesubstrate adherence (Bard et al., 2008). Conversely, disrupting the coupling of actin to transmembrane focal contacts (e.g., L1CAM, integrins, or N-cadherin) increases actin retrograde flow, decreases traction force, and limits growth cone advance (Gomez and Letourneu, 2014). However, it is possible that, in the absence of adhesion molecules, growth cones, such as leukocytes, can have an adaptive modulation of actin turnover kinetics and increase retrograde flow rates to drive forward protrusions (Renkawitz et al., 2009). Thus, actin retrograde flow has been likened to the engine that powers growth cone motility and axon extension. However, in the absence of actin filaments, the axon drives forward at even faster rates, propelled solely by the action of microtubules (Marsh and Letourneau, 1984; Bradke and Dotti, 1999). With depolymerized actin, neurites grow abnormally without direction ignoring environmental cues that normally influence navigation. The F-actin network in growth cones therefore may actually moderate neurite extension by limiting microtubule advance. As a barrier to uncontrolled microtubule growth, the modulation of F-actin structure and turnover are required for the changes in velocity and direction of neurite growth. The growth cone actin network is thus more like a combination of the brakes, steering, drive train as well as parts of the engine of the neurite (Lowery and Van Vactor, 2009). Therefore, the modulation of how actin filaments are assembled, organized, and disassembled has bearing on growth cone dynamics, axon formation, and the development of neuronal polarity.

2.3.2 Actin dynamics during axon formation Actin superstructures largely determine growth cone shape and dynamics, which in turn determines axon growth rates. During neuronal polarization, changes in growth cone shape and dynamics precede axonogenesis. In stage 2 neurons, one growth cone enlarges and displays increased dynamics before elongating into an axon (Bradke and Dotti, 1999; Kunda et al., 2001, Fig. 2.3). The F-actin in this growth cone also has increased sensitivity to cytochalasin D, analogous to the axonal growth cone of stage 3 neurons, suggesting that increased actin turnover occurs prior to axon growth (Bradke and Dotti, 1999). The remaining growth cones of stage 2 neurons remain small and quiescent with a more rigid actin

=

growth cone, and the combined effect of multiple sites of polymerization pushes the leading edge forward. Actin filaments continuously flow in a retrograde manner due to the combined effects of actin treadmilling (polymerization distal and depolymerization proximal) and actomyosin contractility toward the transition zone. F-actin retrograde flow can cause microtubules to buckle and break as they grow along F-actin bundles. (E) Uniformly polar actin filaments are bundled in filopodia via specific actin-binding proteins (e.g., Fascin, not shown). Actin is polymerized exclusively at the tips of filopodia pushing the membrane forward. In filopodia, anticapping proteins such as Ena/Vasp aid in actin polymerization. Away from the membrane, actin-depolymerizing factor/cofilin proteins bind to older portions of the actin filaments and sever and disassemble actin, which can then be recycled for further polymerization. Microtubule growth along filopodia is mediated by various proteins and/or protein complexes such as EB3edrebrin. Microtubule polymerization is regulated in part by end-binding proteins such as EB1 and by tubulin heterodimer binding proteins such as CRMP-2 and stathmin, which aid or impede microtubule polymerization, respectively.

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PART | I Formation of axons and dendrites

Stage 2

Stage 3

Stage 2–3

Dynamic microtubules

Stable microtubules

Actin filaments

FIGURE 2.3 Changes in microtubules and actin facilitate axon formation. Axon formation is the foremost event in neuronal polarization, and specific changes in microtubules and actin are necessary for this to occur. Generally, increased microtubule stability and actin instability cooperate to promote axon initiation. In stage 2 neurons, all of the immature neurites have the potential to become the axon. All of the neurites have quiescent growth cones and similar microtubule organization with microtubules largely constrained in the central domain of the growth cones. During the stage 2e3 transition, one growth cone enlarges and displays increased dynamics, which is reflected in increased F-actin turnover. The increased dynamics of the actin relieves inhibitory constraints on the microtubules, which can extend into the periphery of the growth cone. The remaining neurites have less dynamic growth cones and more rigid F-actin. The microtubules of the presumptive axon also exhibit increased stability compared with the other minor neurites. Following axon formation, in stage 3 neurons, the growth cone continues to display increased dynamics and actin turnover. In addition, the increased microtubule stability persists in the axonal neurite shaft, whereas the microtubules of nascent dendrites remain, in comparison, largely unstable.

cytoskeleton. From these observations, a hypothesis emerged that the regulation of actin in growth cone was crucial for the development of neuronal polarity. The first evidence that the regulation of actin dynamics could be involved in axon growth comes from seminal work by Marsh and Letourneau who showed that destabilization of the neuronal cytoskeleton with cytochalasin D induces neurite growth in dorsal root ganglion (DRG) neurons (Marsh and Letourneau, 1984). Later work extended these finding specifically to neuronal polarization: Global treatment of cultured hippocampal neurons of cytochalasin D induced the formation of multiple axons (Bradke and Dotti, 1999). Furthermore, the local application of pulses of cytochalasin D treatment on one undifferentiated neurite of a stage 2 neuron could induce the treated neurite to become an axon. Even after axonogenesis, the minor processes of stage 3 neurons can be transformed into axons with the pharmacological actin depolymerization (Bradke and Dotti, 2000), demonstrating that axonedendrite fate is plastic and that this plasticity is governed, in part, by the regulation of actin dynamics. Although these studies showed that increased actin destabilization or turnover was sufficient to induce axon initiation, they did not address the endogenous regulators of actin that could be involved in axonogenesis. A plethora of ABPs are expressed in developing neurons (Dent and Gertler, 2003; Dent et al., 2011), with a growing list implicated in the regulation of neuronal polarization. These ABPs affect different features of the actin network including actin nucleation, severing, branching, contraction, and bundling (Fig. 2.2). Arp2/3 proteins are actin nucleators that initiate actin polymerization on the sides of existing actin filaments, resulting in a branched actin structures underlying lamellipodial veils (Ishikawa and Kohama, 2007). Formins are actin nucleators that are thought to mediate actin polymerization in filopodia (Faix and Grosse, 2006; Kovar, 2006), although recent studies also implicate Arp2/3 in filopodia formation (Korobova and Svitkina, 2008). Another class of WH2 domain actin nucleators Spire and Cordon Bleu promotes the formation of unbranched and unbundled actin filaments in neurons (Kessels et al., 2011). All of these actin nucleators have been implicated in neuronal morphogenesis. Silencing of Arp2/3 decreases axon growth via abnormal actin remodeling and

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decreased actin dynamics (Korobova and Svitkina, 2008). Likewise, the expression of dominant-negative mDia or siRNA knockdown of mDia attenuates axon growth in cerebellar granule neurons (Arakawa et al., 2003). However, it was shown by a different group that inhibition of Arp2/3 by scavenging the protein complex from its place of action enhances axon growth (Strasser et al., 2004). Along the shafts of spinal cord axons, Arp2/3 localizes to actin patches where it is essential for inducing filopodia and collateral axonal branches (Spillane et al., 2012). While inhibition of Arp2/3 in DRG neurons generally attenuates F-actin accumulation and filopodia formation, actin retrograde flow, axon growth, and guidance are differentially regulated by Arp2/3 on the different substrates L1 and laminin (San Miguel-Ruiz and Letourneau, 2014). As all these conflicting data are from neuronal cell culture experiments and using siRNAs or pharmacological inhibitors, the physiological role for Arp2/3 in neuronal polarization remains to be elucidated. While formins and Arp2/3 have been implicated in axon formation and branching, Cordon bleu has been shown to drive filopodia formation and dendritic branching (Kessels et al., 2011). This suggests that different actin nucleators are employed to direct the development of distinct morphological changes during the complex process of neuronal polarization in a context-dependent manner. Regulators of actin nucleators have also been implicated in neuronal polarization. Wave (WiskotteAldrich syndrome protein [WASP] family verprolin-homologous protein) forms a complex with Sra1 and Nap1, which localizes to growth cone lamellipodia where it regulates actin polymerization via Arp2/3 and profilin (Takenawa and Miki, 2001; Takenawa and Suetsugu, 2007; Pilo Boyl et al., 2007). The Wave complex promotes axon growth and the knockout of Nap1; an essential component of the Wave complex results in reduced axon extension (Kawano et al., 2005; Yokota et al., 2007). Furthermore, Wave complex has been shown to mediate axon extension downstream of Rac1 in cerebellar granule neurons, presumably through regulation of Arp2/3 (Tahirovic et al., 2010). Proteins of the Ena/Vasp family also enhance actin polymerization kinetics via their anticapping activity and in growth cones localize to the leading edge of extending filopodia and lamellipodia (Drees and Gertler, 2008; Lebrand et al., 2004). Genetic ablation of all three Ena/Vasp proteins in cortical neurons results in severely reduced axonal tract formation in vivo and a failure of neuritogenesis in vitro (Kwiatkowski et al., 2007; Dent et al., 2007). These mutant neurons have aberrant actin bundling, filopodia formation, and a reduction of actin retrograde flow (Dent et al., 2007). Ena/Vasp proteins can also cooperate with profilin to deliver actin monomers to the plus end of growing actin filaments (Barzik et al., 2005). Profilins are actin monomer-binding proteins that aid in the exchange of ADP for ATP, priming G-actin for incorporation into the barbed ends of actin filaments (Witke, 2004). The silencing of profilin IIa decreases actin filament density and leads to increased neurite outgrowth (DaSilva et al., 2003). High profilin activity in the absence of increased actin depolymerization, thus, may attenuate axon development by increasing actin filament polymerization and density. The aforementioned pharmacological experiments on actin depolymerization alluded to the presence of endogenous proteins that would accelerate actin turnover to facilitate axon formation. Members of the ADF/cofilin family are such proteins. ADF/cofilin proteins bind preferentially to the ADPeactin subunits toward the minus ends of actin filaments and increase actin turnover by severing actin filaments and increase depolymerization at the minus end (Bamburg, 1999). Cofilin and, to a lesser degree, ADF are expressed in brain and specifically localized to growth cones (Garvalov et al., 2007). During early neuronal polarization, ADF/cofilin activity increases in the largest growth cone of a stage 2 neuron and in the axonal growth cone of a stage 3 neuron, and this increased activity is mediated by the Rho GTPase, cdc42. Reduction of cofilin levels with siRNA attenuates axon development, whereas the overexpression of cofilin facilitates axon growth (Garvalov et al., 2007; Meberg and Bamburg, 2000). Although the genetic ablation of cofilin has drastic consequences on neuronal cell proliferation and migration, there is only a subtle effect on axon growth (Bellenchi et al., 2007). However, ADF expression is elevated and could partially compensate for reduced cofilin expression. In support of this, the genetic ablation of both ADF and cofilin results in a drastic reduction of neurite formation and axon outgrowth, which is the consequence of a near-complete loss of actin retrograde flow and turnover (Flynn et al., 2012). In vivo, the conditional ablation of ADF/cofilin in the brain results in neurons that fail to form neurites and have lost migratory behavior, leading to a severely hypoplastic cortex lacking axonal tracts. Since pharmacological actin depolymerization rescues neurite growth, actin turnover is thus required for neurite growth and axon development. Myosin II-mediated actin contractility also contributes to retrograde flow and regulates actin dynamics in growth cones (Mederios et al., 2006; Lowery and Van Vactor, 2009, Fig. 2.2). Myosin II is a motor protein that generates contractile forces in actin filament networks, which leads to compression of actin gels and the sliding of bundled actin filaments (Brown and Bridgman, 2003; Ishikawa et al., 2003), and can even lead to actin disassembly (Haviv et al., 2008). Notably, myosin II is highly enriched in the transition domain of growth cones, where it compresses actin filament bundles into actin arcs (Mederios et al., 2006, Fig. 2.2). Inhibition of myosin II activity with pharmacological inhibitors or genetic knockout of myosin II isoforms leads to increased neurite growth, accelerated axonal development, and increased development of ectopic axons (Kollins et al., 2009; Flynn et al., 2009). Increased myosin II activity may oppose axon development by increasing the rigidity of the actin network and decreasing actin-based growth cone dynamics and by indirectly limiting

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PART | I Formation of axons and dendrites

microtubule dynamics in growth cones. Indeed, inhibition of myosin II with pharmacological inhibitors reduces F-actin levels in growth cones, decreases actin retrograde flow, and allows microtubules to protrude farther into the peripheral domain of growth cone (Mederios et al., 2006; Ketschek et al., 2007). The effect of myosin II activity on growth cone dynamics and axon elongation is also, in part, due to the coupling of actin to the substratum via adhesion complexes in the growth cone. These focal contacts consist of linkages of ABPs to different transmembrane proteins, for example, shootin and ezrin to L1, or talin, paxillin, a-actinin, and vinculin to integrins (Gomez and Letourneu, 2014). The general concept is that retrograde flow, driven in part by actin polymerization and in part by myosin II contractility, exerts a force that is transduced to the substratum via focal contacts (the “clutch”) and growth cone advances. However, how the clutch is engaged is context dependent with the substratum dictating how myosin II activity translates into changes in axon growth; for example on laminin, while the clutch is likely fully engaged, the myosin inhibition slows axon advances clutch, but on polylysine, where the focal contacts are minimal, myosin inhibition facilitates axon growth due to released constraints on microtubules (Ketchek et al., 2007). In other contexts, specific ABPs have been shown to play a role in clutch engagement during axon growth. A shootinecortactin complex engages L1CAM in growth cones of pyramidal neurons to increase traction forces and axon elongation (Kubo et al., 2015). This has important implications in the development of L1-dependent axon tracts such as the corpus callosum, which display severe dysgenesis reduced in the absence of shootin (Baba et al., 2017). In sensory neurons, ezrin/radixin/moesin (ERM) proteins tether actin to L1 adhesions to promote growth cone advance, as the inhibition of this interaction results in increased retrograde flow and decreased growth rates (Marsick et al., 2012). Actin organization and dynamics are regulated to facilitate growth cone motility and axon growthdthe first step of neuronal polarization. Although nuances in the exact mechanisms depend on timing and environmental cues in vivo, changes in actin organization and/or dynamics that increase actin turnover and growth cone dynamics are conducive for axon elongation, whereas changes that lead to actin stabilization are refractory to axon development. To achieve this, there is a feedback signal (e.g., PI3K activity) that regulates actin polymerization and depolymerization (severing) as well as actomyosin contractility in the growth cone during axon formation: Disruptions in any of these processes disrupt neuronal polarization. These actin rearrangements not only lead to more actin turnover and a more dynamic growth cone but also directly influence microtubule dynamics and advance, which is the driving force for axon growth. The remodeling of the actin cytoskeleton also plays a role in later events of neuronal polarization, including the development of diffusion barriers between the axon and dendrites (the axon initial segment [AIS]), the organization of actin rings, the outgrowth and arborization of dendrites, and synaptogenesis. These processes are briefly discussed later in this chapter and in greater detail in other chapters of this book Chapters 103e114, 127, 140).

2.3.3 Microtubules Microtubules play important roles in many cellular functions, including neuronal morphogenesis. During neuronal development, microtubules must form stable bundles, which grow and reorganize to provide the main structural framework for the shafts of axons and dendrites. In fact, microtubules are the driving force underlying neurite extension. As with actin, many of the signaling pathways involved in neuronal polarization impinge upon proteins that modulate microtubule stability and dynamics. Furthermore, microtubules serve as the tracks for intracellular trafficking. Recent work has indicated that microtubules are actively regulated during neuronal polarization, changing in their dynamics, stability, and organization during axon formation and the subsequent neuronal morphogenesis (Hoogenraad and Bradke, 2009). Microtubules are assembled from soluble tubulin dimers, which, like actin, can self-assemble into polymers (Desai and Mitchison, 1997; Box 2.2). Soluble tubulin exists as a heterodimer, consisting of a- and b-tubulin, which are the separate products from different genes and share about 50% amino acid homology. Furthermore, there are multiple isoforms of aand b-tubulin, which can be differentially modulated by posttranslational modifications such as tyrosination, detyrosination, acetylation, polyglutamylation, and phosphorylation (Janke and Kneussel, 2010). The tubulin isoform composition of microtubules and the modifications they are subject to can influence the binding of microtubule-binding proteins, microtubule motors, and the dynamic properties of microtubules. An additional tubulin isoform, g-tubulin shares around 30% homology with a- and b-tubulin (Moritz and Agard, 2001). g-Tubulin is organized in large complexes that form an open ring structure, called the g-tubulin ring complex (gTuRC), which plays an important role in microtubule nucleation (Raynaud-Messina and Merdes, 2007). Microtubules nucleate and polymerize spontaneously in vitro when a/b-tubulin concentrations are high. However, in cells, the intracellular monomer concentration seems too low for spontaneous nucleation, although this possibility has not been excluded (Job et al., 2003). Therefore, microtubule formation is assisted by specific structures called microtubuleorganizing centers (MTOCs) (Luders and Stearns, 2007). MTOCs allow the cell to control where and when to

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BOX 2.2 Microtubule dynamics Howard and Hyman, 2003). To maintain the dynamic instability, microtubules consume energy by the hydrolysis of GTP. b-Tubulin has a GTP-hydrolyzing activity that is strongly activated when the dimer is incorporated into the polymer. This hydrolyzing activity leads only to a small layer of tubulin dimers at the plus end that are bound to GTP, the so-called GTP cap. It stabilizes the plus end, because GDP-bound microtubules are intrinsically more unstable. If new polymerization is slower than the GTP hydrolysis, the plus end becomes unstable and results in catastrophe (Mitchison and Kirschner, 1984; Howard and Hyman, 2003). The dynamic behavior of the minus ends is not of interest in vivo, because they are generally capped and thus stabilized (Dammermann et al., 2003). Because GTP hydrolysis is not necessary for microtubule polymerization, the GTP hydrolysis is only important for the dynamic properties of microtubules. GTP ‘cap’ J tubulin Capping (−) protein

E-tubulin D-tubulin GDP

Shrinking microtubule

D-tubulin GDP

(+)

Tubulin heterodimer GTP

Rescue

Growing microtubule Catastrophe

During polymerization, the a/b-tubulin heterodimers arrange into linear protofilaments that associate laterally to form the hollow microtubule cylinders. In most mammalian cells, microtubules form a tube of 13 protofilaments. Within a protofilament, the tubulin heterodimers associate in a head-to-tail fashion. This makes microtubules intrinsically polar, resulting in two structurally and kinetically different ends: the highly dynamic plus end and the less dynamic minus end. The a-tubulin within the dimer is oriented toward the plus end, and the b-tubulin subunit toward the minus end (Desai and Mitchison, 1997; Howard and Hyman, 2003). Microtubules are intrinsically dynamic, a feature termed dynamic instability. They undergo periods of growth and shrinkage at the microtubule plus end. Dynamic instability allows microtubules to switch abruptly from growth to shrinkage (catastrophe) and from shrinkage to growth (rescue) (Mitchison and Kirschner, 1984; Howard and Hyman, 2003). The assembly and disassembly of microtubules are important for not only their generation but also their dynamic properties. The assembly of microtubules can be characterized by three steps: The first phase is defined by a thermodynamically unfavorable and therefore rate-limiting nucleation step. It is followed by rapid elongation of the polymer and finally by a steady-state phase. In the nucleation step, small oligomers of a/b-tubulin heterodimers form a nucleus. Once a stable oligomer of a certain size is reached, rapid polymerization of the microtubule occurs. During the steady state, microtubules display the dynamic instability, when microtubules switch randomly at their plus ends between “catastrophe” and “rescue,” leading to their highly dynamic behavior (Mitchison and Kirschner, 1984;

GDP

Tubulin GTP heterodimer GTP

assemble microtubules. The conventional MTOC in animal cells is the centrosome, an organelle next to the nucleus (see Chapter 2.5.5). Recently, also centrosome-independent and decentralized microtubule formation has been identified in many organisms and cell types (Bartolini and Gundersen, 2006; Luders and Stearns, 2007). After assembly, individual microtubules assume a polarized tubule structure, which are arranged together into linear arrays in the axon. They are dynamic, yet rigid cylindrical, polymers of a/b-tubulin heterodimers with a diameter of about 25 nm (Box 2.2). Microtubules have a unique organization in neurons. In contrast to many somatic cells, neuronal microtubules are not anchored at the centrosome but are abundant in the cytoplasm throughout the whole cell body and funnel into the processes (Baas, 1999). The microtubules reach lengths up to 100 mm within the neurite shafts and are organized in regularly spaced, parallel arrays. During polymerization, the a/b-tubulin heterodimers arrange into linear protofilaments that associate laterally to form the hollow microtubule cylinders. After nucleation, microtubule minus ends are capped in cells, so that most of the interesting dynamics relevant for neuronal polarity occur at the plus ends. Microtubule plus ends oscillate between periods of slow growth and rapid shortening events called “catastrophies,” which can be “rescued” and growth reinitiated. This polymerization and depolymerization behavior is called dynamic instability (Box 2.2). The polymerization and depolymerization dynamics of microtubules are critical for their cellular functions, including their role in neuronal morphogenesis (Conde and Caceres, 2009). In neurons, the number of microtubules growing distally into the peripheral domain of the growth cone is increased during rapid axon growth (Grabham et al., 2007, Fig. 2.3). In the growth cone, microtubules can grow methodically or rapidly disassemble and reorient their direction of growth. These features facilitate growth cone turning and axon guidance. Furthermore, microtubules can generate pushing forces during growth phases and pulling forces during shrinking phases that influence neurite elongation and retraction, respectively. Microtubule assembly and disassembly are regulated by various microtubule-binding proteins that can promote assembly, stabilize microtubules, or destabilize microtubules. Many of these proteins play an active role in neuronal polarization.

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In addition to the modulation of microtubule dynamic instability, the structural regulation of microtubules into bundles is also essential for neuronal morphogenesis. Dense bundles of microtubules make up the main structural framework of both dendrites and axons. However, in mature neurons, axons and dendrites differ in their microtubule organization. In proximal dendrites, microtubules have a mixed polarity, with a population of plus ends facing the cell body and a population facing the distal dendrite. In axons, the microtubules predominantly have a plus-ends distal orientation. This difference is not the case before axonogenesis, where all neurites have uniform polarity microtubules with the plus ends distal. The conversion of the uniform polarity to mixed polarity microtubules in dendrites is a key process underlying axonedendrite differentiation. It partially underlies differences in the growth behavior and the morphological differences of axons and dendrites. Furthermore, these differences in microtubule organization contribute to selective intracellular trafficking as discussed in the following.

2.3.4 Microtubules dynamics during axon formation Microtubules are essential for neurite growth. Early studies demonstrated that the expression of neuronal-specific microtubule-binding proteins, such as Map2c and Tau, which promote microtubule bundling and stabilization, induces neurite-like protrusions in nonneuronal cells (Edson et al., 1993). The importance of microtubule dynamics in axon growth was first demonstrated with drugs that specifically stabilize or destabilize microtubules. Treatment of cultured neurons with either taxol, which stabilizes microtubules, or nocodazole, which depolymerizes microtubules, leads to an inhibition of axon outgrowth (Letourneau and Ressler, 1984; Tanaka et al., 1995; Rochlin et al., 1996), demonstrating that microtubule dynamics were essential for axon extension. More recent studies have specifically shown that modest microtubule stabilization is sufficient to induce axon formation (Witte et al., 2008). When a low concentration of taxol was applied to stage 2 hippocampal neurons, multiple axons formed. Even selective microtubule stabilization in one neurite with photorelease of caged taxol resulted in site-directed axon formation. The supernumerary axons in taxol-treated neurons also had increased levels of acetylated tubulin, a posttranslational modification associated with microtubule stabilization. The stable microtubules could serve as the basis for further microtubule assembly, as local taxol application also promotes the extension of newly polymerized microtubules into the growth cone (Witte et al., 2008). Additionally, stable microtubules may serve as preferred tracks for microtubule-based motors transporting membrane, organelles, and proteins supporting axon elongation. Various posttranslational modifications influence neuronal polarization via microtubule stabilization and via the modulation of microtubule-binding proteins and motors. In axons, it was recognized early on that there were two domains of microtubules based on a-tubulin modifications. The first domain, consisting of the proximal neurite shaft, is enriched in detyrosinated tubulin or Glu-tubulin and is resistant to the microtubule-depolymerizing drug nocodazole. The distal region of the axon is enriched in tyrosinated tubulin and rapidly depolymerizes with nocodazole (Condi and Caceres, 2009). Acetylation is another modification that occurs on stable long-lived microtubules, although it itself does not contribute to microtubule stability (Janke and Kneussel, 2010). Studies on developing hippocampal neurons have shown that in stage 2 neurons there is a polarized distribution of the ratio of acetylated to tyrosinated tubulin in one neurite (Witte et al., 2008). This polarized accumulation of acetylated microtubules is also observed in the axon of stage 3 neurons, suggesting that microtubule stabilization proceeds axon specification (Fig. 2.3). Furthermore, increased posttranslational modifications including acetylation and detyrosination specifically promote the binding and transport of kinesin-1 motors (Reed et al., 2006; Hammond et al., 2010), which can help molecular sorting of axonal cargos. In axons, further stabilization to microtubules occurs by the irreversible removal of the penultimate glutamine residue of detyrosinated tubulin resulting in D2 tubulin (Paturle-Lanfanechere et al., 1991, 1994). The detyrosination of tubulin can be reversed by tubulin tyrosine ligase (TTL), which could lead to more dynamic microtubules (Barra et al., 1988). Genetic ablation of TTL abolishes the presence of tyrosine tubulin in neurons and increases the resistance of microtubules to nocodazole-mediated depolymerization and the abnormal binding of microtubule plus endebinding proteins, such as Clip-170 (Erck et al., 2005). Interestingly, cultured TTL knockout neurons display accelerated axonogenesis and increased formation of supernumerary axons, indicating that TTL-mediated microtubule modifications (resulting in increased microtubule stability) mediate axon specification. Since TTL only retyrosinates free tubulin, the result is that the majority of newly polymerized tubulin is composed of tyrosinated tubulin (Janke and Kneussel, 2010). Tyrosinated tubulin favors the binding of certain plus endebinding proteins that regulate microtubule dynamics and interactions with actin in the growth cone, which in turn facilitates the rapid remodeling of the cytoskeleton necessary for regulating outgrowth rate and steering the growth cone. In the shafts of developing axons, detyrosinated tubulin (and acetylated tubulin) dominates. This favors the binding of KIF5 (kinesin-1) into the axonal compartment (Konichi and Setou, 2009). Thus, it appears that the tyrosination/detyrosination

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state of microtubules contributes to a selectivity filter for the binding of specific sets of microtubule-binding proteins and molecular motors, which influences neuronal polarization. The tyrosination/detyrosination state of microtubules does not seem to affect the binding of the structural MAPs, MAP2 or MAP4 (Janke and Kneussel, 2010). However, other MAPs are affected by these and other posttranslational modifications. The canonical, structural MAPs are a group of proteins that copurify with tubulin after repeated rounds of depolymerization and reassembly. These proteins have been demonstrated to promote the assembly of microtubules and stabilize microtubules presumably via the formation of cross-bridging between adjacent microtubules (Hirokawa, 1994). The most abundant neuronal Maps, Map1B, Map2, and Tau are differentially distributed in axons and dendrites and thought to contribute to neuronal polarization via stabilizing effects on microtubules and by influencing the interactions of other microtubule-binding proteins. Although Map1B is found throughout neurons, a phosphorylated form that has increased microtubule stabilizing activity is specifically localized in developing axons (Boyne et al., 1995). Specific Map2 isoforms (A and B) are localized to dendrites later in neuronal development, whereas tau is enriched in axons (Conde and Caceres, 2009). In spite of specific subcellular localizations, there are overlapping functions of these Maps. The ablation of Map1B results in stunted axon development, decreased growth cone size, and a decreased microtubule polymerization dynamics (Gonzalez-Billault et al., 2001). Interestingly, the deletion of the Tau-1 or Map2 does not affect axon formation, gross microtubule organization, or growth cone size in younger neurons (Takei et al., 2000; Teng et al., 2001). However, the combinatorial deletion of Map1B with either Map2 or Tau results in similar abnormalities: reduced axon and minor neurite growth, disorganized microtubules, and growth cone defects (Takei et al., 2000; Teng et al., 2001). At first glance, these data suggest that these Maps are functionally redundant. This is certainly true, to some degree. However, there are distinctions that are likely important to neuronal polarization. Map1B has synergistic effects with both Map2 and Tau and therefore may represent a more comprehensive Map that can compensate for deficiencies in either of the other Maps. Map2 and Tau are more finely tuned. In the absence of Map1B, the ablation of Map2 has greater consequences on dendritic growth, whereas the ablation of Tau causes more severe axon growth defects, correlating with their localization. In later stages of development, the importance of Map2 in dendritic development becomes more apparent as Map2-deficient neurons have impairments in dendritic arborization (Harada et al., 2002). The binding of MAPs also affects the microtubule growth dynamics in neurons. For example, the neuronal polarization deficits of Map1B-mutant neurons are partially rescued by the expression of end-binding protein 1 (EB-1) (Jimenez-Mateos et al., 2005), which can enhance microtubule polymerization. The modulation of microtubule polymerization and depolymerization dynamics also influences axon growth. In simple terms, a positive potential for microtubule growth exists in the developing axon during neuronal polarization. In addition to the availability of free tubulin, other microtubule-binding proteins influence microtubule growth and shrinkage. Proteins and protein complexes that bind the ends of microtubules can aid the polymerization of microtubules or, alternatively, facilitate the rapid dismantling of microtubules. Furthermore, free tubulin-binding proteins that aid or prohibit free tubulins’ ability to incorporate into microtubule plus ends also impact neuronal development (Fig. 2.2). Microtubule plus endetracking proteins (þTips) are a diverse group of proteins that accumulate at the plus ends of microtubules. By influencing microtubule growth dynamics and directionality, cellular signaling, and interactions with actin filaments and the cell cortex, the þTips influence various aspects of neuromorphogenesis, including the development of neuronal polarity (Akhmanova and Steinmetz, 2008; Conde and Caceres, 2009, Fig. 2.2). During axonogenesis, the þTip, adenomatous polyposis coli (APC) was shown to accumulate specifically in the developing axon and not in the minor processes. Inhibiting APC with function-blocking mutants or siRNA inhibits axon growth either by interfering with microtubule interaction with the cell cortex or by aberrant targeting of Par3, a key protein in polarization of diverse cell types (Shi et al., 2004; Zhou et al., 2004; Purro et al., 2008). The aforementioned feedback signal, PI3K activity, influences microtubule plus-end dynamics via GSK3 beta pathway (Schelski and Bradke, 2017). Tight regulation of GSK3 beta activity can promote axon outgrowth via the APC and another þTip, CLASP, promoting microtubule polymerization while limiting interactions with retrograde actin flow, allowing microtubules to penetrate the peripheral domain of growth cones (Cammarata et al., 2016). The end-binding proteins, EB-1 and EB-3, are the essential, core scaffolding elements of plus-tip-binding protein complexes, which are highly and differentially expressed during neuronal development. EB-1 expression is elevated during axonogenesis, and depletion of EB1 in neuronal cell lines decreases neurite outgrowth (Stepanova et al., 2010). Significantly, these decreases in neurite outgrowth coincided with decreased microtubule growth rates and the lengths, suggesting that EB1 regulates neurite growth directly by influencing microtubule growth dynamics. EB-3 also plays a role in neuronal development, albeit via a different mechanism. EB-3 interacts with the ABP drebrin to coordinate microtubules and actin during neurite initiation and growth (Geraldo et al., 2008). Members of the cytoplasmic linker protein family, Clip115/

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PART | I Formation of axons and dendrites

Clip170, also play an important role in axonogenesis. The expression of dominant-negative Clip170 inhibits axon formation, whereas the expression of microtubule-binding domain of Clip170 promotes the formation of multiple axons (Neukirchen and Bradke, 2011). Clip170 may exert its effects on neuronal polarization by promoting growth cone consolidation into the neurite shaft at the wrist of the advancing growth cone. Although most þTips bind to growing microtubules, some bind to the plus ends of shrinking microtubules. The motor protein KIF2A, a member of the kinesin 13 family that binds to plus ends, is enriched in growth cones and uses ATP hydrolysis to fuel its microtubule depolymerization activity (Hirokowa and Noda, 2008). The ablation of KIF2A results in abnormal axonal outgrowth patterns with increased collateral branch growth due to decreased microtubule depolymerization in growth cones (Homma et al., 2003). Doublecortin (DCX) is another MAP that is found preferentially near the plus ends of microtubules that plays a role in microtubule stability, bundling, and protofilament number. In regard to microtubule dynamics, DCX binding between microtubule protofilaments reduces microtubule catastrophies by limiting their tendency to splay outward (Moores et al., 2006). DCX also may facilitate crosstalk between microtubules and actin via neurabin II. Mutations in this X-linked gene lead to neurological disorders in humans: In females, some cortical neurons migrate abnormally and result in a “double cortex” (hence doublecortin), whereas in males, it results in type 1 lissencephaly (des Portes et al., 1998). Recent work has indicated that in addition to neuronal migration defects, axon outgrowth is also impaired in doublecortin knockout mice and even in humans with the disease-causing doublecortin mutations (Bielas et al., 2007). In mice, the DCX deletion causes excessively branched axons in culture and severely reduced axonal tracts in vivo. These phenotypes are due to the increased splaying of microtubules in DCX knockout neurons. A protein phosphatase 1espinophilin pathway mediates the dephosphorylation and activation of DCX in the zone between the axon shaft and the growth cone, where microtubules are being bundled to facilitate the consolidation phase of axon growth (Bielas et al., 2007). Thus, DCX has a role in axonogenesis via its bundling activity and by mediating interactions with actin to promote the consolidation of the growth cone into the elongating axonal shaft. LIS1, another gene implicated in lissencephaly, has also been implicated in axon formation by acting in concert with dynein to allow dynamic microtubules to resist F-actin retrograde flow and penetrate the peripheral domain in growth cones (Grabham et al., 2007). The collapsing response mediator protein 2 (CRMP2) was first identified for its role in semaphorin-mediated growth cone collapse (Goshima et al., 1995). It is highly and specifically expressed in the developing nervous system and was identified as a crucial microtubule-regulating protein during axonogenesis (Inagaki et al., 2001). CRMP-2 binds to free tubulin heterodimers and enhances microtubule assembly (Fukata et al., 2002). While the expression of wild-type CRMP-2 promotes axonogenesis, mutant forms of CRMP-2 that lack the domain responsible for binding free tubulin inhibit axon growth and branching (Fukata et al., 2002). CRMP-2 is regulated by glycogen synthase 3 beta (GSK3b), which phosphorylates CRMP2, inhibiting its ability to bind tubulin heterodimers and mediate axon induction (Yosimura et al., 2005). Once again, we see GSK3b as one of the pivotal signals at the center of signaling pathways engaged during neuronal polarization (Jiang and Rao, 2005). Thus, the regulation of CRMP-2 downstream of established polarization signaling pathways promotes its role as carrier of tubulin dimers, enhancing their delivery to the plus ends of microtubules, thereby aiding axon elongation. Additionally, CRMP-2 transports also the Wave complex into the axon and thereby also influences actin dynamics (Kawano et al., 2005). CRMP-2 could work as a more general cargo adaptor for kinesin-1, which could also influence axon formation. Lastly, CRMP-2 regulates Numb-mediated endocytosis of adhesion molecules, yet another possible mechanism whereby CRMP-2 affects axon growth (Nishimura et al., 2003). While these cell culture experiments suggest that CRMP-2 activity aids axon formation, its ablation in the CNS apparently can form axons that normally only results in deficits in dendrite branching and spine formation, which may lead to a schizophrenia-like phenotype (Zhang et al., 2016). Although this may be due to developmental redundancy by related proteins (e.g., CRMP-1), this underscores the need to carefully interpret findings from cell culture and perform physiologically relevant studies in vivo to verify and further investigate mechanisms underlying neuronal polarization. Other free tubulin-binding proteins act to destabilize microtubules by preventing polymerization. Proteins of the SCG10, stathmin/Op-18 family bind to free tubulin heterodimers and sequester them in the neuronal cytoplasm. This allows the disintegration of the GTP-cap, which can lead to rapid disassembly of microtubules. Stathmin/Op-18 activity is regulated by phosphorylation, which inhibits its ability to bind to free tubulin and destabilize microtubules (Wittmann et al., 2004; Manna et al., 2006). Laminin, an extracellular matrix molecule that induces axon formation (Esch et al., 1999), induces stathmin dephosphorylation and inactivation downstream of the Rac GTPase activator, Dock 17 (Watabe-Uchida et al., 2006). There is a polarized distribution of the inactive, phosphorylated version of stathmin/Op-18 in the developing axon compared with the minor processes, suggesting that the microtubule-destabilizing effects of stathmin/Op-18 are refractory to axon growth. Likewise, the expression of an unphosphorylatable, constitutively active stathmin/Op-18 reduces neuronal polarization (Watabe-Uchida et al., 2006).

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Analogous to the actin cytoskeleton, not only microtubule destabilization, but also microtubule severing, is important for neuronal development. At various locations throughout a neuron, including the centrosome, branch points, and the growth cone, microtubule severing can lead to the reorganization of microtubules and contribute to neuronal morphogenesis, especially in collateral branching. Furthermore, locally severed microtubules could also serve as new seeds for microtubule growth and thereby increase microtubule number and mass, as described for the formation of the meiotic spindle in Caenorhabditis elegans (Roll-Mecak and Vale, 2006). There are two main microtubule-severing proteins expressed in neurons, katanin and spastin, which have not only some overlapping functions but also important distinctions. Katanin is distributed throughout the neuron and has its highest level of expression during axon growth, which reduces after target selection. The tight modulation of moderate katanin activity is important for axon growth: both blocking katanin function and enhancing it cause defects in axon growth (Karabay et al., 2004). Thus, both too little and too much microtubule severing are deleterious for axon growth. Interestingly, in the absence of tau binding, katanin activity induces collateral branching in axons, indicating that tau protects axonal microtubules from katanin-mediated severing and branching (Quiang et al., 2006). Spastin has more of a role in microtubule rearrangements, leading to collateral branching (Yu et al., 2008). Axonal branching is increased with increased spastin levels and decreased with spastin depletion. Unlike katanin, spastin activity does not affect neuronal polarization. Although axon growth depends on microtubule polymerization (Tanaka et al., 1995), it is rather the formation of new microtubules than only the extension of existing microtubules that delivers the polymers necessary for axon extension (Yu and Baas, 1994). During the past decades, it has been intensely debated how and where the microtubule arrays in axons are formed. First mentioned by Lasek (Lasek, 1986), the “polymer transport model” proposed that microtubules are nucleated at the centrosome, released from the centrosome through the microtubule-severing protein katanin (Ahmad et al., 1994; Ahmad et al., 1998; Baas et al., 2005), and then transported along the axon by the motor protein dynein (Ahmad et al., 1998; Wang and Brown, 2002). In contrast, other groups reported that, if nothing else, growing microtubules are not transported in axons, and most microtubules are stationary (Ma et al., 2004; Kim and Chang, 2006). Moreover, tubulin is transported into the axon and to the growth cone in its nonpolymerized form (Terada et al., 2000; Kimura et al., 2005). The alternative hypothesis was therefore termed the “subunit transport model.” It postulated that tubulin is transported into the processes in single subunits or oligomers that are then locally incorporated into the microtubules (Hirokawa et al., 1997). Indeed, local microtubule polymerization occurs in comparable rates throughout all neuronal compartments (Stepanova et al., 2003). Consistently, local inhibition of microtubule assembly at the axon tip inhibits axonal growth, whereas inhibition at the cell body does not have an effect on axon growth (Bamburg et al., 1986). To date, both models are under discussion, especially the aspect of microtubule transport into the axon, with different groups still presenting data for both models by the use of live-cell imaging approaches (Wang and Brown, 2002; Ma et al., 2004; Kim and Chang, 2006). As the same experiments can be interpreted in different ways (Terada, 2003; Myers et al., 2006), finding the definitive answer is a complicated task. Recently, the model of centrosomal-microtubule nucleation has been challenged, at least in the context of axon growth following initial polarization: The ablation of the centrosome in stage 3 neurons did not affect axon elongation, and at later stages, no new polymers were nucleated at the centrosome (Stiess et al., 2010). Therefore, there is also no new supply of new microtubule polymers from the cell body. How new microtubules are then generated remains still unclear. In addition to microtubule severing as described earlier, new microtubule could be also generated by decentralized, noncentrosomal microtubule nucleation as it is observed, for example, in fission yeast, plants, or the mitotic spindle (Janson et al., 2005; Murata et al., 2005; Uehara et al., 2009). This might be a general mechanism as the centrosome becomes deactivated also in different CNS neurons (Leask et al., 1997). However, it is still unclear if the centrosome position plays an active role in the determination of the axon. Polarized microtubule assembly at the centrosome could support the outgrowth of the future axon. Furthermore, the Golgi apparatus localizes next to centrosome (Sütterlin and Colanzi, 2010) and thus could support axon growth by polarized membrane trafficking to the developing axon (Bradke and Dotti, 1997). However, the data are contradicting and differ from neuronal cell type (de Anda et al., 2005; Dotti and Banker, 1991; Zolessi et al., 2006; Sharp et al., 1995; Zmuda and Rivas, 1998; de Anda et al., 2010). Furthermore, it is debated if the observed correlation of centrosome position is cause or just an epiphenomenon of the polarization itself (Higginbothham and Gleeson, 2007; Arimura and Kaibuchi, 2007; Witte and Bradke, 2008).

2.3.5 Cytoskeletal dynamics during dendritic growth and arborization Thus far, we have predominantly focused on cytoskeletal dynamics in the context of axon initiation and growth. This is for good reason, as it is the foremost event in neuronal polarization. However, to achieve a fully functional neuron, the dendrites must differentiate into the signal receivers of the neuron. An essential aspect of this is the elaboration of their

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PART | I Formation of axons and dendrites

arbors, which increases their receptive field and the sites of potential contacts with presynaptic neurons. Dendritic growth and differentiation are considered in depth in other chapters of this work (Chapters 127 and 140). Thus, we will only briefly provide an overview of the cytoskeletal changes that occur during dendritic arborization. Like axons, the major structural components of dendrites are microtubules and actin. Many of the molecules that play a role in axon development play a similar role in dendrite growth (Luo, 2002), but at a delayed time, as dendritic growth and arborization typically occurs later, in stage 4 neurons (Craig and Banker, 1994). In the developing cortex, dendritic growth begins after neurons are situated in their final cortical layer and undergoes phases of arbor growth coinciding with the arrival of afferent input (Jan and Jan, 2010). As in axons, the canonical Rho GTPases affect dendritic morphogenesis in a manner analogous to their influence on axon growth and mainly via their influence on the cytoskeleton. Rac1 and Cdc42 activities have a positive effect on dendritic development, whereas RhoA acts as a negative regulator of dendritic growth and branching (Urbanska et al., 2008). In spite of their similarity, important differences exist between axons and dendrites, including growth rates, microtubule organization, neurite caliber, and branching patterns (Craig and Banker, 1994). In particular, the complex and increased branching pattern distinguishes dendrites from axons. Discrepancies in the organization and regulation of the cytoskeleton during dendritic development likely account for these differences. A couple notable differences between axons and dendrites are their microtubule organization and growth cone actin dynamics. Whereas axons have unipolar, axial microtubule orientation, dendrites have biaxial microtubules with mixed polarity. The acquisition of the biaxial orientation of the microtubules during stage 4 contributes to dendritic growth patterns and corresponds temporally to the acquisition of the proximodistal taper unique to dendrites (Baas et al., 1989). The acquisition of the mixed polarity microtubule array in dendrites involves the motor CHO-1 (KIF23) (Sharp et al., 1997), whereas dynein seems to maintain uniform microtubules in the axon (Zheng et al., 2008). Interestingly, at the time when dendrites grow, the centrosome does not nucleate microtubules anymore (Stiess et al., 2010). Therefore, it is unknown how the microtubule arrays are generated in neurons during dendritogenesis. Furthermore, dendrites have a less densely packed microtubule organization, which also influences dendritic morphogenesis (Conde and Caceres, 2009). The spacing of microtubules in dendrites is mediated by Map2, as neurons deficient in Map2 display a decrease in the spacing of dendritic microtubules (Teng et al., 2001; Harada et al., 2002). Furthermore, the outgrowth of dendrites and the elaboration of dendritic arbors are impaired in the absence of Map2 (Teng et al., 2001; Harada et al., 2002). Another Map, Map1A, becomes upregulated and specifically enriched in dendritic branch points concomitantly with dendritic differentiation in vivo and in cultured neurons (Szebenyi et al., 2005). The downregulation of Map1A reduces total dendritic length, but mainly via the inhibition of higher-order arbor initiation and growth. In addition to the binding of specific Maps, the stability of microtubules is different in dendrites compared with axons. The ratio of microtubule acetylation/tyrosination, an indication of the levels of stable microtubules, is lower in dendrites of stage 4 neurons in integrated neuronal networks (Gomis-Ruth et al., 2008). Consistent with this observation, microtubules in dendrites of mature neurons are more susceptible to nocodazole-mediated microtubule depolymerization, resulting in the degeneration of dendrites. Furthermore, the induction of microtubule stability with Taxol in established neuronal networks can transform dendrites into axons (Gomis-Ruth et al., 2008). It is likely that some of these differences in the cytoskeleton are established early on in neuronal development, as microtubules in young, developing dendrites are also more susceptible to nocodazole-mediated microtubule depolymerization, resulting in the loss of minor neurites (Witte et al., 2008). The decreased stability and density of the microtubules undoubtedly influence the slow growth behavior of dendrites. This also may relate to why dendrites are more reliant on cellular adhesion for their growth than axons (Chamak and Prochiantz, 1989). Increasing cellular adhesion to very high levels negatively affects axonal growth disproportionally compared with dendritic growth (Lafont et al., 1993). Dendritic microtubules may be less able to resist the tensile forces of F-actin retrograde flow in growth cones, limiting their extension and causing them to be more reliant on cellular adhesion for growth. Inhibition of myosin II, which partly mediates F-actin retrograde tensile forces, increases the extension rates of immature dendritic processes (Kollins et al., 2009). The reduction of F-actin-mediated tension may allow the dendritic microtubules to grow unimpeded and facilitate dendrite growth. However, under normal growth conditions, the sparsity of microtubules and decreased stability of the microtubules are insufficient to drive extension to a level on par with axonal growth. Growing microtubules also have to overcome a generally less dynamic and more rigid actin cytoskeleton in dendritic growth cones compared with axonal growth cones. Brief application of actin destabilizing drugs can completely depolymerize F-actin in the axonal growth cone, whereas actin filaments remain intact in the developing dendrites (Bradke and Dotti, 1999). In addition to the increased myosin IIemediated actin contractility, decreased cofilin activity also likely keeps the actin of immature dendrites in a quiescent state (Garvalov et al., 2007). In support of this, BMP7 signaling, which induces dendritogenesis (Guo et al., 2001), can activate Lim kinase downstream of BMP receptor 2, leading to decreased cofilin activity (Wen et al., 2007). Even in stage 4 neurons, more rigid F-actin is responsible for modulating dendritic growth as treatment with cytochalasin D converts dendrites into axons (Bradke and Dotti, 2000). Since both microtubule

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stabilization and actin instability transform dendrites into axons, dendrite fate is likely specified by microtubule-actin interactions that limit microtubule growth and maintain a quiescent actin cytoskeletondthis limits the extension rates of the dendrites. Even when dendritic growth occurs, it never reaches the velocities of axon growth (Dotti et al., 1988). Rather, the increased emergence of secondary and higher-order branches accounts for the overall increase in dendritic surface area. The extensive branching of dendrites occurs primarily via interstitial branching (Luo, 2002). Along the shafts of dendrites, actin-rich filopodia are extremely dynamic, constantly extending and retracting. When dendritic filopodia become stable, they engorge and begin elongating. The trigger for elongation and branch formation involves a synchronization of actin dynamics and microtubule invasion into the filopodia, though a direct comprehensive study into this is lacking. A couple possible candidates specifically mediating these interactions during dendritic branching are Map2C and Map1A, both of which bind microtubules in dendrites and can induce F-actin rearrangements (Rogar et al., 2004; Szebenyi et al., 2005). The importance of actin nucleation in shaping dendritic arbors was elucidated in studies that showed when NWasp-Arp2/3-mediated actin nucleation activity was disrupted, dendritic arbors were irregular (Rocca et al., 2008). The expression of another novel ABP, cordon bleu, which promotes the growth of unbranched actin filaments and filopodia, greatly increases dendritic arborization (Ahuja et al., 2007). Direct regulation of microtubules is also important for dendritic branching as cypin regulates dendrite patterning by affecting microtubule assembly (Akum et al., 2004; Chen and Firestein, 2007). In conclusion, axon and dendrite development are both based on the cytoskeleton and share common mechanisms of growth. However, these mechanisms are differentially regulated in axons and dendrites. How these differences occur and how also the different timing during development is regulated still remain unclear.

2.3.6 Subcellular cytoskeletal specializations In the later stages of neuronal development, subcellular specializations are formed that carry out certain functions necessary for the development and maintenance of neuronal polarization. These include the AIS, nodes of Ranvier, axonal actin rings, actin trails, presynaptic terminals, and postsynaptic specializations such as dendritic spines. These structures represent the pinnacle of neuronal subcellular functional specialization. The AIS first starts forming in stage 3e4 hippocampal neurons, following initial axon formation. AIS formation, like axonogenesis itself, is an intrinsic property of the neuron; no external cues or cellular interactions are needed. As indicated by the accumulation of voltage-gated sodium (Naþ) channels, the AIS forms from DIV (day in vitro) 2 until DIV 7, at which time nearly all neurons contain voltage-gated Naþ channels and other AIS markers such as ankyrinG and bIV spectrin (Yang et al., 2007). The accumulation of ankyrinG, an organizer of membrane domains in many cell types, at the axon hillock is the master switch, leading to the formation of the AIS, and triggers the recruitment of other AIS-resident proteins (Rasband, 2010). For example, ankyrin recruits Naþ channels to the AIS via targeting motifs in the loop between domains II and III. The AIS also contains additional ion channels, other transmembrane proteins, and a specialized cortical cytoskeleton organization, which are all tethered directly or indirectly to ankyrinG (Rasband, 2010). It is currently unknown exactly how ankyrinG itself localizes to the axon hillock to initiate AIS formation. Importantly, ankyrin directly recruits bIV spectrin, another important scaffolding protein that links the AIS to the cortical actin cytoskeleton (Yang et al., 2007). The interaction with the underlying actin filament network is crucial to the integrity of the AIS. The disruption of actin filaments with the actin-depolymerizing drug cytochalasin D uncouples Naþ channels from the AIS, resulting in reduced Na currents (Kole et al., 2008, Rasband, 2010). The high Naþ density is particularly important for the AIS since it facilitates the low threshold for generating an action potential, a defining feature of the axon. In addition to its role as the start site for axonal electrical properties, the AIS serves another crucial function in neuronal polarization. AIS development marks a second phase of axonedendritic segregation and subcompartmentalization by acting as a selective barrier for the lateral diffusion of membrane components and the intracellular diffusion of cytosolic proteins. AIS development underlies the polarized segregation of the membrane components that determine the function of the axon as distinct from the dendrites. The AIS forms (about 30e60 mm) at the base of the axon distal to the axon hillock where it begins to function to prevent the mixture of somatodendritic and axonal compartments. In stage 3e4 neurons by DIV 5, it begins to act as a selective cytoplasmic diffusion barrier (Song et al., 2009) and later (DIV 11) assumes its role to limit the lateral diffusion of lipids and proteins of the plasma membrane (Nakada et al., 2003). As discussed in the following, the function of ankyrinG and the AIS as a molecular fence is also essential for the maintenance of neuronal polarity. Like the AIS, nodes of Ranvier are formed by the same cytoskeletal network with ankyrinG as the master regulator. Functionally, nodes of Ranvier are very similar to the AIS, with high densities of voltage-gated ion channels underlying comparable electrical properties, such as the low threshold for generating action potentials. However, in

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contrast to the AIS, the development of the nodes of Ranvier depends on glia-derived signals, namely myelination (Girault and Peles, 2002). The ability for a neuron to chemically transmit information to an effector cell, be it another neuron, an endothelial cell, or striated muscle, depends on a functional synaptic terminal. Different neurons may differ in the organization of their individualized synapses, but they all are variations of a common theme. Synaptogenesisdthe formation of synaptic connectionsdoccurs as a growth cone reaches its target and transforms into the presynaptic terminal, a process that requires the reorganization of the actin cytoskeleton. The growth cone loses filopodia, forms tight contacts with the postsynaptic terminal, and begins accumulating synaptic vesicles. The formation and development of new synapses is reliant on F-actin rearrangements. In newly formed synapses of DIV 5 hippocampal neurons, actin depolymerization with latrunculin A abolishes presynaptic terminal integrity: synaptophysin clusters disappear, vesicle recycling is reduced, and ultrastructurally evident presynaptic terminals are no longer observed in axons (Zhang and Benson, 2001). Increasingly stable actin structures drive synaptic stabilization, whereas synapses with unstable actin structures are eliminated. In mature synapses, presynaptic scaffolding proteins such as Bassoon accumulate and maintain the integrity of synapses independently from F-actin, and actin structures play more of a background modulatory role in functional properties of synaptic transmission (Halpain, 2003). Not much is known about the role of microtubules in the presynaptic entity. However, it was shown in synaptic boutons of the drosophila neuromuscular junction that microtubules form a hairpin loop in the developing synapse and that this is necessary for synaptogenesis (Roos et al., 2000; Hummel et al., 2000; Conde and Caceres, 2009). With the advent of more advanced live-cell imaging techniques and superresolution imaging methods, extraordinary detail in cytoskeletal structures in polarized neurons have been elucidated. While it has long been recognized that microfilaments line the cortical regions of the axon and extend deep into the lumen of the axon, specific regular organization was not observed until recently (Leterrier et al.2017). Axons were found to contain circumferential F-actin rings lining the interior of the plasma membrane spaced at regular intervals of 190 nm done the length of the axon (Xu et al., 2013, Fig. 2.4). The rings were connected and presumably organized by bII spectrin tetramers and contain adducin, which caps and stabilizes short-actin filaments. While actin rings are present along the entire axon shaft, their composition is altered proximally as the AIS is established in maturing neurons whereby bII spectrin is replaced by bIVeankyrinB scaffolds (Leterrier et al., 2017). Although drug studies and live-cell imaging experiments suggest actin rings are very stable, how they are formed is unknown. They are not in immature stage 2 neurons but first appear after initial axon growth and neuronal polarization has commenced and become more distinct as neurons become mature. The function of these actinspectrin rings in the axon remains elusive, but they may serve to maintain the cylindrical shape of axons, support microtubules in the neurite shafts, and endow the axon with crucial mechanical properties such as flexibility while still resisting stretching and compressive forces. In support of this, b-spectrin ablation in C. elegans results in axons that break Stage 2

Stage 2–3

Mitochondria Vesicle

Peroxisome Axonal cargo

Stage 3

Ribosome Dendritic cargo

FIGURE 2.4 Trafficking in polarized neurons. (A) A mature, stage 5 neuron has an axon and dendrites that are morphologically, molecularly, and functionally distinct. The distinct molecular repertoires of axonal and dendritic compartments are established during development and maintained via a sophisticated trafficking program. (B) Expanded region of a dendrite from panel (A) The general organization of microtubules and F-actin is depicted. Golgi outposts are common at branch points in dendrites. In the proximal dendrites, microtubules have a mixed orientation with plus ends and minus ends oriented distally. The orientation of microtubules gradually scales with dendritic length such that the distal dendrite is dominated by a plus-end distal orientation. (C) Conversely, the axon contains a uniform microtubule array with the majority of the plus ends oriented distally throughout the length of the axon. The axon initial segment is a subcellular F-actin-rich specialization of the axon that acts as a selective barrier separating the axon from the somatodendritic compartment. In mature neurons, periodic cortical actin rings (arrows) corral microtubules and maintain the cylindrical structure of axons. (D) Expanded region from panel B is shown. Trafficking mechanisms along microtubules in the proximal dendrites contributes to the specific sorting of dendritic cargos. The mixed orientations of microtubules are particularly important in dendritic sorting. The minus endedirected motor dynein navigates.

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down due to mechanical strains of normal body movement (Hammarlund et al., 2007). The regular spacing of sodium channels mirrors that of the underlying actinespectrin periodicity, suggesting an additional role organizing ion channels crucial for action potential propagation (Xu et al., 2013). While the F-actin in the cortical actinespectrin rings is stable, other deeper actin filaments in the axon are remarkable dynamic. Actin hot spots generate rapidly growing longitudinally oriented actin “trails” in a formin-dependent manner (Ganguly et al., 2015). These actin trails move both in the anterograde and in the retrograde directions and may help deliver cargo (actin and ABPs) to En-Passant boutons along the actin shaft. In addition, actin trails may represent homeostatic pool of local actin for other actin-dependent processes (actin-dependent trafficking, bouton formation, filopodia extensions, etc.) as part of a larger axonal actin network (Leterrier et al., 2017). On the postsynaptic side of excitatory synapses, dendritic spinesdspecialized actin-based sites of postsynaptic signal transmissiondare required for function of dendrites as signal receivers. Dendritic spines typically begin developing after 14 days in stage 5 cultured hippocampal neurons (Craig and Banker, 1994). In vivo, there is a similar timeframe for dendritic spine development, beginning in the cortex shortly after dendrites have extended from the neuronal soma and concomitant with the arrival of afferents into their prospective receptive fields. The latter suggests that an intimate association of synaptic membranes triggers spine morphogenesis. Indeed, live-cell imaging studies show that dynamic filopodia extend from the shafts of dendrites, explore their surroundings, and occasionally make contact with axons, initiating synapse formation and transforming into spines (Ziv and Smith, 1996; Bhatt et al., 2009) These movements of the filopodia and their transformation into spines occur via actin-mediated rearrangements (Hotulainen and Hoogenraad, 2010). Once formed, spines continue to display activity-dependent dynamics. Spines occur in different morphologies and can transition between these morphologies in an actin-dependent manner, with the acquisition of a mushroom-type morphology being the strongest type of postsynaptic structure. Actin is organized into long- and short-branching filaments in the spine neck and shorter-branching actin filaments in the spine head (Fig. 2.4). Numerous ABPs have been identified that modulate spine shape and dynamics including cofilin, Arp2/3, and myosin (Hotulainen and Hoogenraad, 2010). In addition to organized actin filaments, spines contain membranous organelles, such as endoplasmic reticulum (ER) and the postsynaptic density, which organizes receptors, adhesion proteins, channels, and signaling proteins in the postsynaptic membrane. Microtubules have also been shown to invade subsets of dendritic spines and regulate their dynamics (Hu et al., 2008; Jaworksi et al., 2009). These invasions seem to be also dependent on neuronal activity and involved in synaptic plasticity. Periodic membrane-associated actin rings, which are more prominent in axons, were also intermittently observed in distal dendrites and in the neck region of dendritic spines (He et al., 2016). In these regions, spectrineactin rings may support dendritic shape, corral microtubules, aid in the distribution of proteins, and regulate dendritic spine elasticity (Konietzny et al., 2017).

2.4 The role of (membrane) trafficking during neuronal polarization To support the extension of neurites, biomaterial must be delivered and incorporated into the growing processes. This biomaterial includes organelles, cytosolic proteins, components of the cytoskeleton, and vesicles delivering membrane proteins and lipids destined for different organelles or the neuronal plasma membrane. During neuronal morphogenesis, as the simple sphere of a stage 1 neuron develops into the highly complex, reticular stage 5 neuron with extremely long axons and elaborates dendritic trees, there is about a 200-fold increase in surface area (Pfenniger, 2009). The expansion of area and extensive outgrowth of a neuron in and of itself is impressive, but even more so is the discriminatory trafficking of distinct molecules to specific locations in the neuron. Accumulating evidence has pointed to a complicated hierarchical system regulating the specificity of trafficking (Fig. 2.4). This entails layered and interdependent mechanisms that operate at different levels, interacting to contribute to molecular “sorting” program.

2.4.1 Trafficking during early neuronal development Early in neuronal development, however, the sorting program is not so sophisticated, and the unspecific delivery of materials predominates to support the initiation and growth of nascent neurites. Simple, diffusion-based movements are an effective way for short-range transport of cytosolic material to supply materials supporting the extension of newly formed neurites (Popov and Poo, 1992). But even at these early stages, not all trafficking is completely diffusion based. The delivery and the exocytosis fusion of vesicles to the membrane does not rely on microtubule-based transport but does seem to rely, at least partially, on active, actin-based transport mechanisms during neuritogenesis (Gupton and Gertler, 2010). Diffusion likely continues to play a role in the movement of materials at all stages of neuronal development, but since diffusion is a promiscuous process, it acts in opposition to polarization. Therefore, neurons acquire strategies later in

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development to reign in the ambiguous influence of diffusion. Nevertheless, at the early phases of polarization, in the absence of external cues, small stochastic differences in transport of growth-promoting molecules lead to a random selection of the axon, which is amplified into an all-or-nothing response. While local activation of positive feedback loops (promoting axonal cytoskeletal dynamics) occurs in one neurite promoting its outgrowth, these signals are globally reduced in other neurites (less favorable cytoskeletal dynamics) (Scelski and Bradke, 2017). As the morphology of neurons becomes more complex, additional active mechanisms are harnessed to facilitate the directed delivery of membranous and cytosolic material to support the growth and differentiation of growing axons and dendrites. During axon growth, the vectoral flow of material into the axon supports its elongation (Fig. 2.5). This not only results in a semiselective transport with the accumulation of agents that support axon growth but also results in the buildup of cargos considered dendrite-specific in the nascent axon (Bradke and Dotti, 1997; Craig and Banker, 1994). The bulk flow of material supports the surface area expansion of the membrane and the increased cytosolic volume of the growing axon. Although there is some indiscriminate transport at these early stages of neuronal development, directly proceeding axon growth, there is specific accumulation of motors and proteins that support axon elongation (Jacobson et al., 2006; Arimura and Kaibuchi, 2007), suggesting that, in addition to diffusion and bulk flow, there are selective transport mechanisms at work early on. The kinesin motor Kif5C (kinesin-1) transiently explores multiple immature neurites of a stage 2 neurons but then selectively remains in one before that neurite begins elongating into the axon (Jacobson et al., 2006). This observation suggests that the cargos transported by kinesin-1 may provide some impetus for axon initiation. In hippocampal neurons, the occurrence of anterograde “waves” is correlated to axon growth and transport materials that support axon growth such as actin, the ABP cofilin, and shootin, a regulator of PI3K activity (Ruthel and Banker, 1999; Flynn et al., 2009; Toriyama et al., 2010). These waves are dynamic actin-based structures, moving at rates of slow axonal transport, and may represent an alternative way to transport some intracellular proteins independent of microtubule motors during axon formation. During axonogenesis, retrograde diffusion of cytosolic proteins counteracts active transport and contributes to the tugof-war type of periodic growth and retraction observed among neurites of stage 2 neurons. Retrograde diffusion is indiscriminate, occurring similarly in all neurites, depriving them of potentially positive cues for axon growth. Therefore, it seems that neurite length is a major determinant for feedback of axon formation. Since the rate of retrograde diffusion is lower in the longest neurite, the concentration of a positive cue would be higher in that neurite. Indeed, when a stage 3 neuron’s axon is cut to above 10 mm longer than the other neurites, it regenerates as the axon (Goslin and Banker, 1989). However, when the axon is cut to a length less than 10 mm longer than the other neurites, the determination of a new axon is random. Thus, there is a neurite length-dependent feedback. This same phenomenon is observed in mature neurons integrated in functional circuits (Gomis-Ruth et al., 2008). These observations have led to the hypothesis that a growthpromoting factor (or factors) is actively transported to growth cone but diffuses back to the soma. One such protein exhibiting this behavior is shootin. The active transport of shootin via waves to one growth cone of stage 2 neurons is correlated with a burst of growth. In stage 2 neurons, this accumulation is transient, due to the periodic and stochastic transport of shootin to all the processes and because diffusion back to the soma depletes the growth cone of shootin. However, as the axon forms, shootin accumulation is stabilized in the axonal growth cone via more preferential active transport (i.e., increased wave frequency) and because retrograde diffusion has less of an effect over long distances (Toriyama et al., 2010). Since the genetic ablation of shootin does not inhibit neuronal polarization per se (Baba et al., 2017), the transport of other cues is likely important to axon initiation. Indeed, the enrichment of other regulators of neuronal polarity such as Cdc42, Rac1, and cofilin to the growing axon follows a similar mechanism (Winans et al., 2016; Flynn et al., 2009). Diffusion will continue to influence the distribution of cytosolic elements in developing neurons; however, the development of the AIS diffusion barrier limits the contribution of diffusion during later neuronal polarization (see the following text).

2.4.2 Motor proteinebased transport in axons and dendrites Other studies have shown polarized accumulation of specific proteins such as Par3/Par6/aPKC, Akt, Rho GTPases, and cytoskeleton-regulating proteins into the neurite destined to become axon during stage 2, before it starts elongating. All of these proteins mediate axon initiation, so it seems that active transport mechanismsdperhaps kinesin-1-mediated transport or wavesdare harnessed even at these early stages to achieve the specific accumulation of axon growth-promoting agents. But how are these transports directed to one compartment only? The analogy of a ground transport company like the United Parcel Service (UPS) or FedEx can help understand the complexity of intracellular trafficking during neuronal polarization. In the neuron, the base of operations would be the in the soma, including the sites of biosynthesis such as the ER and ribosomes and the secretory pathway, including the ER

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FIGURE 2.5 Trafficking during axonogenesis. In stage 2 neurons, there is uniform delivery of material to all the neurites. At this stage, all of the neurites have the potential to become the axon and have equal distributions of proteins, vesicles, and organelles. During the stage 2e3 transition, there is increased bulk trafficking of organelles, axonal-specific proteins, and ribosomes into the presumptive axon. This supports the enlargement and dynamics

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=

of the growth cone. Continued polarized delivery of materials supports the elongation of the axon in stage 3 neurons. Although there is some specific delivery of key axonal proteins, trafficking of many proteins is indiscriminate, as many dendritic proteins can also be found in the developing axon. Only later, are molecular sorting mechanisms more finely tuned to lead to segregation of dendritic cargo and the maturation of functional subdomains. Microtubules specifically in dendrites in a bidirectional manner to deliver important cargo such as AMPA receptors. Other studies implicate the plus endedirected kinesin, KIF17 in dendritic targeting. Microtubule modifications also can affect the targeting of cargos in a mature neuron. Typically, microtubules in dendrites are tyrosinated, and axon-specific motors such as kinesin 1 (KIF5) avoid tyrosinated tubulin and therefore do not enter dendrites. The binding of the microtubule-associated protein, MAP2, may also influence motor-driven transport in dendrites. E. Expanded region from panel B shows distal dendrite with a dendritic spine. The distal regions of dendrites have microtubule arrays where plus-end distal orientations dominate. Kinesins, such as KIF17, likely transport dendritic cargo in the shaft of distal dendrites. Importantly, local transport occurs along actin filaments in cortical regions such as dendritic spines. The actin motor, myosin V, is involved in the local transport and delivery of AMPA receptors in neuronal dendrites by navigating toward the plus ends of actin filaments. F. Expanded region from panel C displays microtubule transport and the role of the axon initial segment (AIS) in the axon. Specific kinesins, such as kinesin 1 (KIF5), navigate toward the plus ends of microtubules in axons. Microtubule posttranslational modifications in the axon such as detyrosination and acetylation help direct the attachment and movement of kinesin 1 along axonal microtubules. The binding of MAPs such as tau also can influence motor-based transport. The AIS located at the proximal axon is crucial for molecular sorting of axonal versus somatodendritic cargos. The AIS is composed of various scaffolding proteins including ankyrinG and bIV spectrin, which bind to and/or organize various components of the AIS. Actin filaments are tethered to the AIS and are essential for integrity of the AIS and maintaining its role as a diffusion barrier. The AIS acts to limits the lateral mobility of membrane components and the diffusion of cytosolic components, thereby segregating the axon from the somatodendritic compartment. At the AIS, microtubule-based axonal cargos, those carried by KIF5, for example, are allowed through, whereas dendritic cargos are prevented from entering the axon.

and Golgi apparatus. The highways in the neurons are the microtubule networks radially extending away from the somatic home base. The 18 wheelers that transport cargo long distances are the microtubule motors, kinesin and dynein. ATP is the gas. Local transport stations are sometimes utilized for more local shipments, and in neurons, dendritic Golgi outposts serve this purpose. In some cases, there are also local roads near the destination, which in neurons are actin filament networks, and myosin motors are local transporters. The final destination of the cargo can be the cell membrane, a specific organelle or the cytoplasm. As with ground transport, sometimes, the shipments go awry and are delivered to the wrong address and need to be sent back to the right location. The cell also has a mechanism for sending back or getting rid of unwanted material. At all levels along the transport pathway, there is the possibility for regulation. This seems true in the neuron; complex layers of regulation ensure that the right materials are located in the right positions at the right times necessary for neuronal polarization. In the following, we will discuss the single components of this trafficking system. As fast and reliable ground transport depends on good highways and fast trucks with a certain destiny, polarized trafficking depends a lot on the motor-based transport of cargos along microtubules (Goldstein and Yang, 2000; Hirokawa and Takemura, 2005, Fig. 2.4). Microtubules are optimal highways for long-distance transport in neurons because they are long, rigid, and unbranched. Although individual microtubules do not extend the entire length of the neurites, the ends of adjacent microtubules overlap, providing a more-or-less continuous and easily accessible track for microtubule motors to drive along. These motors equate to the delivery trucks that carry cargo along the microtubule highways of the neuron. These motors actually “walk” (rather than drive, see Gennerich and Vale, 2009 for details regarding the molecular mechanism of motors) in a directional manner, along the microtubules either toward the minus or the plus end of the microtubule. Therefore, one could imagine the selective transport of cargos into dendrites with minus endedirected motors, since dendrites are the only compartment with minus-end distal microtubules (Black and Baas, 1989). As a matterof-fact, selective transport of dendritic proteins is found in hippocampal neurons (Burack et al., 2000), and also minus endedirected motors localize to dendrites such as dynein and KIF2C (Hanlon et al., 1997; Saito et al., 1997; Kapitein et al., 2010). It was already shown that dynein is required for polarized transport into dendrites of Drosophila neurons (Zheng et al., 2008), which have in dendrites microtubules with a uniform minus-end distal orientation opposite to the axon (Stone et al., 2008). A recent study extended this observation to hippocampal neurons and showed that dynein drives cargo selectively into dendrites (Kapitein et al., 2010). Furthermore, the authors showed by quantitative modeling that bidirectional dynein-driven transport on bipolar microtubules provides a potential mechanism for selective transport into dendrites. Plus endedirected kinesins such as kinesin-2/Kif17 that also bind specifically dendritic proteins and localize to dendrites (Setou et al., 2000) might be necessary for the cargo transport within the distal dendrite, which comprises a uniform plus-end distal microtubule polarity. In conclusion, it seems that the bipolar organization of microtubules is key to selective transport into dendrites and thus for neuronal polarization (Fig. 2.4). Is the targeted transport into axons also based on the orientation of microtubules? An important consequence of the uniform plus-end distal polarity of microtubules is that dynein-based transport is prevented into the axon. This already provides some targeting specificity. Another plus endedirected microtubule motor, kinesin-1/Kif5, which is important for

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neuronal polarity, binds preferentially to axonal microtubules (Nakata and Hirokawa, 2003; Konishi and Setou, 2009). This preference actually precedes axon formation: the overexpressed motor domain enriches in the tip of the future axon shortly before axon formation starts (Jacobson et al., 2006). Furthermore, inhibition of the polarized transport by kinesin-1/Kif5 caused polarity defects in the neurons (Konishi and Setou, 2009). However, orientation of microtubules cannot explain this selective targeting during axon specification, because microtubules are uniformly oriented in all processes early at this stage (Baas et al., 1988; Stepanova et al., 2003). In a recent study, Konishi and Setou showed that it is not microtubule orientation, but rather microtubule modification that steers kinesin-1/Kif5 into the axon (Konishi and Setou, 2009). They showed that tyrosination, a posttranslational modification of tubulin that occurs more frequently in dendrites, prevents kinesin-1/Kif5 to bind to these modified microtubules. Therefore, kinesin-1/Kif5 binds preferentially to detyrosinated microtubules, which are abundant in the axon, leading to axonal targeting of the motor. Consistently, inhibition of tyrosination or stabilization of microtubules abolished the polarized localization of the motor and caused the formation of multiple axons (Konishi and Setou, 2009; Hammond et al., 2010). Additional to the dendritic exclusion by tyrosinated microtubules (Konishi and Setou, 2009), posttranslational modifications of stable microtubules such as acetylation, detyrosination, and polyglutamylation seem therefore to promote kinesin-1/Kif5 binding and transport into the axon (Reed et al., 2006; Hammond et al., 2010). Intriguingly, one of the cargo proteins of kinesin-1/Kif5 is CRMP-2, which brings necessary cytoskeleton regulators such as WAVE-1 or tubulin into the axon (Kawano et al., 2005; Kimura et al., 2005). This could explain the polarity defects by inhibiting kinesin-1/Kif5 (Konishi and Setou, 2009). However, also other kinesins bring polarity regulators into the developing axon, for example, KIF3A transports APC (Shi et al., 2004) or the kinesin-like protein GAKIN transports PIP3 into the axon, whereby it regulates polarization (Horiguchi et al., 2006). Microtubule-based transport is not the only cytoskeleton-based trafficking mechanism required for the establishment of neuronal polarity. If microtubules are the superhighways used for long-distance transport in the neuron, actin filaments are the local street system used for local delivery of some cargos to their final destination. In neurons, actin filaments are concentrated just underneath the membrane toward which they are oriented, in bundles or in a branching network. These actin “streets” are also under a constant state of construction, extending some tracks while disassembling others over the course of seconds. These dynamic tracks are navigated by myosin motors, the local delivery trucks (Bridgman, 2009, Fig. 2.4). The actin-based motor myosin Va is both necessary and sufficient to target transmembrane proteins, such as GluR1, to dendrites (Lewis et al., 2009). Other myosin motors have been shown to play a role in the local delivery of membrane-associated cargos into filopodia and dendritic spines (Bridgman, 2009). The collective influence of the cytoskeleton is that a high degree of selective transport is based on cytoskeletal differences in axons and dendrites (Arnold, 2009; Kapitein and Hoogenraad, 2011). The polarization, orientation, and posttranslational modifications of microtubules impart integral differences in these highways that help direct traffic to different compartments of the neuron. By recognizing these differences in the microtubule highways, motors navigate them in a selective manner. The orientation and regulation of the local streets of the actin filament network also contribute to the delivery of cargos to specific locations. However, the regulation of the neuronal highways and local street systems (roadwork) is only the beginning of the sorting mechanisms at work in a polarized neuron.

2.4.3 The secretory and endosomal pathway In line with our transport analogy, the correct delivery of parcels includes the sorting of parcels according to their destiny and then their correct addressing. Similarly, the selective targeting of membrane proteins involves the recognition, sorting, and targeting of molecules to specific locales in the cell. Membrane proteins are typically sorted to the cell membrane via secretory and/or endocytic pathways. In developing neurons, the bulk of protein and lipid synthesis occurs in the soma (Pfenninger, 2009), highlighting the importance of a sorting program in which newly made materials are targeted to their appropriate destinations. This is also true in mature neurons; however, there is some peripheral biosynthesis occurring in distal neurites that can influence axonal growth and dendritic differentiation. After biosynthesis at the ER, proteins transit the Golgi apparatus and the secretory pathway via lipid-enclosed vesicles. Lipids themselves, synthesized in the smooth ER, also enter the secretory pathway via the Golgi apparatus. There are three main mechanisms that are responsible for the selective targeting of membrane proteins that have been also adapted by neurons. First, membrane proteins can be selectively targeted to either axons or dendrites, via a sorting process at the trans-Golgi apparatus whereby they are packed into specifically targeted vesicles. This mechanism is reported for the insulin-like growth factor-1 receptor (IGFR) in the axonal growth cone (Pfenninger et al., 2003). The targeted transport of these vesicles will be then performed by axon selective motor proteins such as kinesin-1. Second, proteins are targeted to one membrane domain and are then endocytosed and redirected to their target membrane domain. This process, called transcytosis, was shown for the axonal cell adhesion molecule L1 (Yap et al., 2008). Third, membrane proteins are not specifically targeted but are only retained at the

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specific location by scaffolding proteins or binding to other cells and everywhere else endocytosed and degraded. This process works for voltage-gated Naþ channels and Vamp2 (Garrido et al., 2001; Sampo et al., 2003). All of these processes rely on molecular “postal codes” that assign the cargo to a particular localization. For direct targeting of membrane proteins, specific amino acid motifs act as these postal codes, resulting in an intrinsic sorting mechanism targeting them to the plasma membrane or to intracellular organelles. In epithelial cells, these targeting signals help localize proteins via direct binding to coat protein components at the cytoplasmic face of membranes (Bonifacino and Traub, 2003). In neurons, transferrin receptors are specifically targeted to dendrites via a cytoplasmic tyrosine-rich motif (Burack et al., 2000). Interestingly, the transcytosis of the axonal cell adhesion molecule L1 seems to be also regulated by amino acidebased motifs. L1 contains a tyrosine-based signal, YRSLE, which mediates its targeting from the trans-Golgi network to the somadendritic domain (Yap et al., 2008). In addition to the somadendritic signal, L1 contains a 15-amino acid axonal signal in the cytoplasmic tail, which are both necessary for the transcytotic pathway (Yap et al., 2008). A mutation of this tyrosin-based motif inhibits transcytosis, and L1 is directly transported to the axon. As the YRSLE motif can be phosphorylated (Schaefer et al., 2002), cells could use that as a possibility to turn on and off the transcytotic pathway and therefore modulate L1 localization. In addition to the 15-amino acid signal stretch involved in transcytosis, L1 contains a second axonal targeting signal, consisting of extracellular fibronectin-like repeat motifs (Sampo et al., 2003). The reason why L1 comprises two sufficient axonal targeting signals, although they show additive effects, and what role the transcytosis pathway plays remains unclear. Other types of postal codes are also employed to label the proteins for specific targeting including GPI anchors, palmitoylation, and glycolization. For instance, palmitoylation of proteins, the addition of fatty acids to cysteine residues, plays an important role in the targeting of many proteins (Fukata and Fukata, 2010). The palmitoylation of PSD-95, for example, is essential for its postsynaptic targeting (El-Husseini et al., 2000) and is involved in synaptic plasticity (Noritake et al., 2009). In contrast, the palmitoylation of NCAMs is necessary for axonal growth cone targeting (Ponimaskin et al., 2008). GPI anchors and palmitoylation have been also proposed to be important for the selective retention of the membrane proteins by interacting with lipid rafts (Allen and Chilton, 2009). The addition of new membrane proteins to the plasma membrane requires also the fusion of vesicle with the plasma membrane. This process adds another possibility to specifically target proteins, as the fusion mechanism is highly specialized and precise. The vesicular SNAREs (SNAP receptors) bind only to the complementary target SNARE at the plasma membrane and thereby restrict fusion to specific sites (Jahn and Scheller, 2006). The expression, functional role, and specific localization of more than 35 mammalian SNAREs are not clear yet. The vesicle docking before membrane fusion is mediated by Rab proteins, a family of more than 60 proteins that also bind only to specific molecules at the target membrane, adding yet another targeting mechanism (Stenmark, 2009). These small GTPases can recruit specific motors as it was shown that Rab3-containing vesicles recruit Kif1Bb and Kif1A motors via the linker protein DENN/MADD for axonal transport (Niwa et al., 2008). Furthermore, Rab27 regulates the axonal transport of TrkB via Slp1/CRMP-2 and kinesin 1 (Arimura et al., 2009). All the vesicles of the secretory pathway emerge out of the Golgi apparatus. In young neurons, the Golgi apparatus is localized adjacent to the emerging axon and is necessary for axon growth as its disruption inhibits axon growth (Jareb and Banker, 1997; de Anda et al., 2005). In maturing hippocampal neurons, the Golgi changes position following axonogenesis: in stage 4 neurons, it rotates around the cytoplasm to reside at the site of apical dendrites, where it will remain for the duration of a neuron’s life (Horton et al., 2005). Furthermore, as dendrites increase in complexity, they form Golgi outposts at dendritic branch points, where they are actively engaged in post-Golgi trafficking. In fact, dendrites contain ribosomes, ER, and functional ER exit sites (Gardiol et al., 1999; Horton and Ehlers, 2003), which are found together with Golgi outposts meaning that dendrites contain all the components for a local secretory pathway. The localization of mRNAs in dendrites further validates that all the ingredients are present for local protein biosynthesis (Bramham and Wells, 2007). Indeed, the plasma membrane contains both lipids and proteins generated locally in dendrites (Ye et al., 2006). This is likely to support the intensified development of the dendrites at these later stages of development. The disruption of Golgi integrity with Brefeldin-A in stage 4 neurons decreases dendritic growth, arborization, and the accumulation of dendrite-specific molecules (Horton et al., 2005). Even in more mature stage 5 neurons, inhibition of Golgi dynamics reduces dendritic length. Furthermore, in vivo laser ablation of Golgi outposts retard dendrite growth (Ye et al., 2006). These studies all indicate that a local protein synthesis and a local secretory pathway are crucial to dendritic differentiation.

2.4.4 RNA transport and local translation Local protein synthesis in dendrites requires the presence of mRNA. Thus, the transport of mRNA is crucial for both dendritic growth and remodeling. Local translation is not unique to dendrites, however, as growing axons also contain

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mRNAs and exhibit local translation, which is of importance for guided axon growth (Lin and Holt, 2008). The closest ground transport analogy for RNA transport would be receiving something (e.g., furniture) that requires assembly. You receive this unfunctional item in nice protective packaging and have to unwrap it and assemble it: Luckily, you have all the tools you need. The neuron also sends incompletely assembled material in the form of mRNAs, and it also has the tools it needs in axons and dendrites to locally translate that mRNA into functional proteins. mRNA transport is complex and involves different types of RNA-binding protein complexes, or granules, such as ribonucleoprotein particles (RNPs) serving as the protective packaging for transport (Bramham and Wells, 2007). These RNPs bind mRNAs and sequester them from translation machinery in the soma and are necessary for their transport into axons and dendrites. In axons, local mRNA translation has been demonstrated to modulate chemotropic responses during growth cone guidance (Lin and Holt, 2008). For example, the binding of b-actin mRNA translation to zipcode-binding protein 1 (ZBP1) regulates growth cone motility (Zhang et al., 2001) and attractive growth cone turning (Yao et al., 2006). In lower motor neurons, the binding and transport of b-actin mRNA with heterogeneous RNP-R and survival motor neuron 1 (SMN1), the gene deleted in the developmental disorder spinal muscular atrophy, facilitates axon growth (Rossol et al., 2003). In dendrites, the transport and local translation of mRNAs encoding cytoskeletal proteins, neurotransmitter receptors, scaffolding proteins, and signaling enzymes play a role in activity-dependent synaptic plasticity, such as longterm potentiation (LTP) (Bramham and Wells, 2007). The movement kinetics and sensitivity of RNPemRNA complexes to nocodazole indicated that microtubule motors transport RNPs in dendrites. Thus far, only the conventional kinesin, KIF5, has been specifically implicated in the transport of RNPs with mRNAs targeted to dendrites, such as the a-subunit of calcium/calmodulin-dependent kinase II (aCaMKII) (Kanai et al., 2004; Ohashi et al., 2002). Whether in axons or dendrites, the directed transport and local translation of mRNA has important and specific roles at different locales during neuronal development. The local transport and translation of proteins may also coordinate with local protein degradation to control the localization and activity of proteins in specific compartments of neurons (Segreffe and Hoppe, 2009).

2.4.5 Barriers for the segregation of functional domains As discussed earlier, the development of the AIS is a key developmental event enabling the distinct compositions of axonal and dendritic compartments. The AIS represents a molecular wall in the neuron that separates the functionally distinct axon from the somatodendritic compartment. However, the wall is more like a molecular sieve or fence, acting more as a selective barrier than an impassible barricade, allowing some molecular crosstalk among the two sides, but still maintaining the functional integrity of the distinct compartments. The AIS serves as a barrier for both membrane constituents and intracellular components in the cytosol. In the analogy to ground transportation, the AIS likens to the customs officials, at the borders of two sovereign nations, checking if the appropriate documents are in place to warrant passage between the two domains. Materials with the proper credentials are allowed entry into the axon, whereas those without the proper credentials remain in the somatodendritic domains. Likewise, once inside the axonal compartment, exit is prohibited except for the occasional rogue breakout. What are the credentials? It seems there are a few contributing factors that the AIS recognizes to filter material into the axon, including molecular size, cargo type, and motorecargo interactions. The ability of the AIS to act as both a membrane diffusion barrier and cytoplasmic barrier depends on ankyrinG and intact actin filaments. Indeed, the accumulation of F-actin at the AIS correlates to its ability to act as a diffusion barrier. Moreover, the destabilization of actin results in the redistribution of cytoplasmic molecules and increases the motility of membrane components through the AIS (Winckler et al., 1999; Song et al., 2009). Concurrent with the appearance of the AIS, there is a selective redistribution of the NMDA receptor into the somatodendritic compartment. The vesicles that carry NMDA receptor subunits are stalled at the AIS, whereas other vesicles carrying the axonal synaptic vesicle protein VAMP2 are not impeded. This selectivity is determined by the interactions of different kinesins and, with their specific cargos, dictates the ability to pass through the AIS. For example, KIF17 and KIF5B transport NR2B and VAMP2, respectively. The movement of KIF17 is impeded through the AIS, whereas the movement of KIF5B is not. This depends on the cargo or cargo-binding regions of the kinesins since the motor domains of KIF17 or KIF5B are distributed throughout the neuron. Interestingly, swapping the cargo-binding tail domains of KIF5B and KIF17 can redistribute NMDA receptors to the axon in addition to the somatodendritic compartment (Song et al., 2009). Likewise, swapping the tail domain of KIF5B for KIF17 redistributes VAMP2 to the somatodendritic compartment in addition to the axon. These observations, as well as the fact that KIF5 can carry both axonal and dendritic cargos, suggest that cargoemotor interactions play a role in axonedendritic targeting as mediated by the selectivity filter at the AIS. However, how the cargo is recognized by the AIS is still unclear. Axonal exclusion mechanisms based only on the cargo size seem not be involved as experimentally manipulated cargo transport by KIF17 predominantly targets the axon (Kapitein et al., 2010).

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PART | I Formation of axons and dendrites

2.4.6 Protein stabilization and degradation As with postal delivery, the trafficking system in neurons occasionally sends packages to the wrong address. When this package is, for example, an axonal-specific protein that could adversely affect dendritic function, the neuron needs to get rid of it. One option is to forward the misrouted material to its appropriate destination. Indeed, this can occur via transcytosis, which helps establish polarized membrane domains. Alternatively, this unwanted material can be marked for destruction. To this end, the neuron harnesses the temporal and spacial control of protein turnover by the ubiquitin proteosome system (UPSdnot to be confused with the United Parcel Service). This not only is a mechanism to get rid of missorted proteins but also seems to serve as a rather specific way to localize proteins at their site of action. The development and maintenance of neuronal polarity critically depends on this local regulation of protein degradation. The UPS pathway is the main protein degradation pathway in eukaryotes and influences a diverse cellular process including cell cycle progression, cell growth, polarity, cellular signaling, and apoptosis (Segref and Hoppe, 2009). In response to a variety of signals, the UPS engages the sequential activation of Ub-activating enzymes (E1), Ub-conjugating enzymes (E2), and Ub ligases (E3), resulting in the conjugation of the small protein, Ubiquitin to lysine residues in targeted proteins. The subsequent tagging of the initial ubiquitin with at least four ubiquitins (polyubiquitylation) targets the protein for destruction via proteolysis mediated by the proteosome (Welchman et al., 2005). In addition to the degradation pathway, ubiquitination can also mediate other cellular behaviors, such as endocytosis and signaling. During neuronal development, the UPS pathway is crucial to the establishment of neuronal polarity. Treatment of developing neurons with specific inhibitors of the UPS or via the expression of a dominant-negative ubiquitin leads to a loss of neuronal polarity and the formation of multiple axons (Yan et al., 2006). In these studies, the protein kinase Akt was identified as a particularly important target for selective degradation in developing dendrites during neuronal polarization. Akt is known to be involved in signaling pathways, leading to the induction of axon formation (Arimura and Kaibuchi, 2007). The selective degradation of inactive Akt in the neurites leads to the polarized distribution of active Akt in one neurite, which becomes the newly formed axon. The Rho GTPase Rap1b, which induces axonogenesis via cdc42 activation (Schwamborn et al., 2006), shows a similar UPS-dependent regulation. The enrichment of Rap1b to the nascent axon depends on the activity of the ubiquitin ligase Smurf-2, which leads to its degradation in the minor processes (Schwamborn et al., 2007). In the cases of Akt and Rap1B, ubiquitylation and degradation leads to the enrichment of these proteins in the developing axon. Conversely, proteins refractory to axon growth can be marked for degradation specifically in the axon and retained in the developing dendrites. Indeed, this is the case for Lim kinase, which phosphorylates and inactivates cofilin, whose activity is necessary for actin turnover during axonogenesis. The ubiquitin ligase, Rnf6 targets Lim kinase for destruction specifically in the axon (Tursun et al., 2005), and this could contribute to increased cofilin activation in the developing axon. More recently, the E3 ubiquitin ligase, TRIM2, was also shown to regulate neuronal polarization (Khazaei et al., 2011). Ubiquitin and protein degradation are also important in later events of neuronal development, such as dendritic remodeling (Kuo et al., 2005), axon guidance (Campbell and Holt, 2001), and synapse formation (DiAntonio and Hicke, 2004). How exactly the UPS is synchronized at specific locals in neurons is unclear. It could be regulated directly at the level of the UPS system, by the (in)activity of the inappropriately localized protein, or may also be coupled with mRNA transport and local translation (Segref and Hoppe, 2009). Regardless of how its function is coordinated, local protein degradation is crucial for the development of neuronal polarity.

2.5 Maintaining neuronal polarity A particularly wondrous feature of the nervous system is its resilience. Generally, neurons are born during embryonic and early postnatal development, develop into functional units of the nervous system, and remain stable and active for the lifetime of an organismdwhich in humans can exceed 100 years. Neurons must retain their polarization to ensure the function of the nervous system. This entails a complete generation and reincorporation of all biological components of a neuron every few weeks. With this constant biosynthesis of proteins, lipids, carbohydrates, and nucleic acids mechanisms are in place to organize the appropriate localization to ensure the uninterrupted function of a polarized neuron. Clearly, this continued maintenance of neuronal polarity is important to an aging organism. Indeed, in humans, the breakdown of subcellular compartmentalization may underlie neurodegenerative diseases such as amyotrophic lateral sclerosis (ALS), leading to severe neurological impairment and even death (Maussion et al., 2008; Rasband, 2010; Kanning et al., 2010). Some of these maintenance mechanisms are simply the continued usage of developmental programs that remain active throughout the life of a neuron. Other mechanisms are constituted as neurons mature, further ensuring the fidelity of subcompartmentalization in functional neurons.

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The maintenance of polarization is achieved, substantially, by the cytoskeleton. As discussed earlier, cytoskeletal specializations such as the AIS are crucial for the segregation of functional domains of neurons. The integrity of the cortical actin structure, ankyrinG levels, and the density of Naþ and Kþ channels of the AIS are essential to the continued function of mature neurons. When ankyrinG is downregulated in mature neurons, the axon becomes dedifferentiated, with a proximal loss of Naþ channels and bIV spectrin, and reprogrammed with dendrite-like features, including increased levels of the dendritic proteins MAP2, and the Kþ/Cl transporter (Hedstrom et al., 2008). The disruption of the AIS via ischemia causes proteolysis of ankyrinG and bIV spectrin, resulting in a loss of Naþ channels and a disruption of neuronal polarity (Rashband, 2010). Purkinje neurons deficient in ankyrinG exhibit functional dendritic spines on their axons, demonstrating the requirement for the AIS in axonedendrite identity (Sobotzik et al., 2009). Reduced ankyrinG and AIS function has also been linked to cognitive disorders such as schizophrenia (Cruz et al., 2009, Rashband, 2010). Interestingly, mature neurons display an amazing degree of plasticity in regard to neuronal polarization, and this plasticity may depend on the integrity of the AIS. As discussed earlier, axonal injury to functionally mature neurons can reprogram an existing dendrite to transform into an axon, depending on the site of the injury (Gomis-Ruth et al., 2008). Whether the original axon regenerated or a dendrite converted into an axon depended on the lesion site: Proximal lesions closer than 35 mm from the cell body resulted in dendrite reprogramming, whereas more distal lesions resulting in axon regrowth. The localization of this switch is correlated to microtubule stability, as it demarcates where microtubules distal to the lesion are more stable and where microtubules proximal to the lesion are similar to dendritic microtubules (Gomis-Ruth et al., 2008). Since the transport of axonal cargo by kinesin-1 relies on differences in microtubule stability, the loss of stable microtubules may reinitiate an “axon lottery,” allowing a preexisting dendrite to transform into an axon. Alternatively, the fate switch may be related to the AIS. The 35 mm proximity of the fate-switching lesion is coincidentally close to the AIS. Thus, in these axotomy experiments, axonedendritic fate could relate to a loss of the AIS and therefore a loss of polarity maintenance (Schafer et al., 2009). In at least one example, the plasticity of neuronal polarity may be beneficial during traumatic injury to the nervous system. Following spinal cord injury of adult felines, commissural spinal neurons have been observed to transform dendrites into axons that can then grow through the injury site (Fenrich et al., 2007, Fendrich and Rose, 2009). This plasticity and reorganization of neuronal polarity could lead to the compensatory formation of adaptive neural circuits following such injuries. In addition to the AIS, the selective targeting of proteins is necessary to maintain neuronal polarity. This is done either by selective retention of membrane proteins or by selective transport maybe mediated by the different axonal and dendritic microtubule arrays (Kapitein et al., 2010). However, the mechanisms of selective trafficking are tailored during development, reaching their peak in specificity in mature neurons (Ledesma and Dotti, 2003).

2.6 Future work on neuronal morphogenesis The majority of our current understanding about the mechanisms underlying neuronal polarization is from work in neuronal cultures (Arimura and Kaibuchi, 2007). Over the past decade, work has boldly begun to characterize neuronal development in vivo and ex vivo (Noctor et al., 2004; Barnes et al., 2008; Calederon de Anda, 2010; Funahashi et al., 2014) and explored the mechanisms in vivo with in utero and ex utero electroporation studies and using genetically modified organisms (Hand et al., 2005; Garvalov et al., 2007; Tahirovic et al., 2010; Flynn et al., 2012). Interestingly, in some cases, in vivo studies have validated earlier cell culture experiments regarding the importance of cytoskeletal regulators in early neuronal development such as ADF/cofilin (Flynn et al., 2012), but other in vivo studies validate some conclusions drawn from cell culture work while challenging others (Baba et al., 2017; Zhang et al., 2016). Progress has certainly been made, but future work needs to continue to identify the physiologically relevant players in neuronal polarization. For the actin cytoskeleton, there are various ABPs that are highly expressed during neuronal morphogenesis but remain uncharacterized. The same is true of microtubule-regulating proteins. Another particularly contentious issue is how cytoskeletal subunits are generated and transported during neuronal polarization. For example, microtubules were traditionally believed to be nucleated exclusively at the centrosome and then released and transported to support axon growth (Baas et al., 2005). However, recent work has contested this view by showing that axon growth (which is dependent on microtubule nucleation) can occur independently from a centrosomal-based mechanism (Stiess et al., 2010). In addition, the advances of the past years have tremendously improved our understanding of selective transport based on cytoskeletal differences in axons and dendrites (Arnold, 2009; Kapitein and Hoogenraad, 2011). However, we are still at the beginning to fully understand this process. For example, in the early phase of polarization, we have only knowledge about kinesin-1/ kif5, and with respect to the extensive amount of different motors involved in axonal transport (Hirokawa and Noda, 2008), we are just starting to piece the puzzle of polarized transport together.

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Furthermore, axon development occurs in a highly directed manner in vivo as compared with in culture, which proceeds in a stochastic manner, indicating the influence of extracellular cues in the determination of neuronal polarization (Yi et al., 2010). For example, growth factors such as TGF-b and extracellular matrix molecules such as laminin seem to provide spatial directives for axon formation in vivo (Yi et al., 2010; Lei et al., 2012). Future work will need to elucidate the role of additional extracellular factors in neuronal polarization and how they affect the regulation of the cytoskeleton and intracellular trafficking. Yet another challenge for the future is to determine if differences in the mechanism underlying neuronal polarization are different in the diverse neuronal subtypes in the body. The majority of our knowledge comes from rodent hippocampal, cortical, and cerebellar neurons (Barnes and Polleux, 2009; Tahirovic and Bradke, 2009), but less is known regarding the polarization of peripheral neurons, interneurons, and spinal cord neurons. Are similar mechanisms at work in diverse neurons? We already see some similarities and differences in cerebellar and hippocampal neurons, which may reflect their unique morphologies and functions in vivo. It will be interesting to see how the mechanisms of overlap differ in different neuronal systems. Beyond the examination of different types of neurons from animal models is the more clinically relevant necessity to characterize neuronal development in human neurons. With the advancement of neuronal differentiation protocols from pluripotent stem cells, studying the cytoskeletal mechanisms of neuronal development in different types of human neurons is an impending reality (Steinbeck and Studer, 2015; Kelava and Lancaster, 2016; Tao and Zhang, 2016). Indeed, some initial work has already begun in this area (Nakashima et al., 2018). Moreover, the refinement of brain organoids that display remarkable similarities to human brain structures will make it possible to probe the development of neuronal polarization in an in vivoelike setting (Lancaster et al., 2013; Kelava and Lancaster, 2016). The generation of neurons and brain organoids from induced PSCs isolated from patients suffering a host of neurodevelopmental disorders will not only allow for a deeper understanding for disease mechanisms in humans but also provide more grounded leverage to develop treatments for these disorders. As cell therapy progresses, with precise gene editing techniques, replacement of affected neuron populations may even eventually hold promise for clinical therapy (Steinbeck and Studer, 2015). Finally, axon regrowth following pathological conditions or traumatic injury may use the same intrinsic factors that govern axon formation and growth during development. In the case of spinal cord injury, the manipulation of intrinsic regulators of axon growth, including some regulators of the cytoskeleton, can indeed promote some axonal regeneration (Kwon et al., 2002). Conditioning lesions to peripheral nerves are correlated to increased axonal transport, and this is one of the requirements necessary to support axon regeneration (Hoffman, 2009) Therefore, the elucidation of cytoskeletal and trafficking mechanisms underlying axon growth (and neuronal polarization) during development could reveal alternative targets for promoting axon regeneration following CNS injury or disease.

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Chapter 3

Axon growth and branching Le Ma and Stephen R. Tymanskyj Thomas Jefferson University, Philadelphia, PA, United States

Chapter outline 3.1. Introduction 3.2. Cell biological mechanisms 3.2.1. Growth cones: structure and function 3.2.2. Regulation of cytoskeleton assembly 3.2.2.1. Actin 3.2.2.2. Microtubules 3.2.3. Interaction between F-actin and microtubules 3.2.4. Membrane trafficking and axonal transport 3.2.5. Protein translation and stability 3.3. Extracellular regulation of axon growth and branching during neural development 3.3.1. Nerve growth factor and neurotrophic factors 3.3.2. Guidance molecules: netrin, slit, semaphorin, ephrin, and wnt

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3.3.3. Cell adhesion molecules: permissive or instructive 3.3.4. Glial cells and myelination 3.3.5. Neural activity 3.3.6. Additional axon branching molecules 3.4. Intracellular signaling mechanisms that mediate axon growth and branching 3.4.1. Rho family small GTPases: linking receptors to the cytoskeleton 3.4.2. Calcium 3.4.3. Cyclic nucleotides as second messengers and modulators 3.5. Concluding remarks References

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3.1 Introduction One striking feature of neurons is their long processes extending out from their cell bodies. These processes, called axons, are responsible for transmitting electrical signals that are collected at the dendrites and integrated at the axonal hillock. Each neuron has only one axon, which can reach synaptic targets as far as 1 m away in humans. To connect with multiple targets, axonal branches sprout at different locations, either along the axonal shaft as interstitial collaterals or at the nerve endings as terminal arbors. The growth of axons and the formation of their branches are intimately associated with axonogenesis and axon guidance, two processes that are discussed in detail elsewhere in this book section. Together, they help establish appropriate synaptic connections, generate topographic maps, define receptive fields, and confer structural plasticity during critical periods. To ensure the proper assembly of functional neural circuits, axon growth and branching are tightly regulated during development; any misregulation could cause synaptic dysfunction and lead to impaired sensory, motor, cognitive, and mental abilities, deficits found in many neural developmental and psychiatric disorders. In the mature nervous system, axon growth and branching are less frequent (Portera-Cailliau et al., 2005), but their regulation is no less important; how to regrow damaged axons that have lost their growth ability and how to sprout new branches from both injured and spared axons have profound implications in regenerative medicine (Mahar and Cavalli, 2018; Tedeschi and Bradke, 2017; Tuszynski and Steward, 2012). How do axons grow and branch? This is a century-old question that has intrigued many developmental neurobiologists since the days of Ramon y Cajal (1904). With the advent of technological advances in molecular genetics, high-resolution imaging, and neuronal culture, tremendous progress has been made over the past three decades in elucidating the molecular and cellular mechanisms governing axon growth and branching. In this chapter, we attempt to provide a comprehensive overview of our current knowledge of how these processes are regulated. This knowledge is synthesized from a large body

Cellular Migration and Formation of Axons and Dendrites. https://doi.org/10.1016/B978-0-12-814407-7.00003-1 Copyright © 2020 Elsevier Inc. All rights reserved.

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of work in many neuronal cell types of both the peripheral nervous system (PNS) and the central nervous system (CNS) in invertebrate (Aplysia, Drosophila, Caenorhabditis elegans) and vertebrate (frog, zebrafish, and rodent) models using primary neuronal culture and in vivo manipulation strategies. Owing to space limitations, we only include representative studies to illustrate the general principles of axon growth and branching. For more details on each specific topic, the reader is advised to consult the original articles and also several recent reviews (Armijo-Weingart and Gallo, 2017; Bilimoria and Bonni, 2013; Dent et al., 2010; Gallo, 2011, 2016; Gibson and Ma, 2011; Hall and Lalli, 2010; Kalil and Dent, 2014; Kapitein and Hoogenraad, 2015; Omotade et al., 2017; Pacheco and Gallo, 2016; Quiroga et al., 2018; Winkle et al., 2016).

3.2 Cell biological mechanisms 3.2.1 Growth cones: structure and function The growth cone is a distinctive and highly motile structure located at the tip of a growing axon or branch. It was originally described by Ramon y Cajal as “a club or battering ram endowed with exquisite chemical sensitivity, rapid ameboid movements, and a certain motive force allowing it to circumvent obstacles in its path” (Ramon y Cajal, 1904). Direct observation of live neurons over the past century has provided ample evidence to support its leading role in controlling axon growth and branching. Structurally, the growth cone is characterized by the expansion of the cytoplasm at the end of the axon shaft (Lowery and Van Vactor, 2009). On a two-dimensional substrate, it spreads out like a fan with the center (C) domain that is sometimes engorged with cytoplasm. The growth cone is dependent on the coordination of two cytoskeleton components, actin and microtubules. In the axon shaft, microtubules are organized into parallel bundles, but they become splayed out in the C-domain, which is enriched with membrane vesicles. Outside the C-domain is the leading edge, or the peripheral (P) domain, that has veil-like lamellipodia populated by needle-like filopodia. Both structures are rich in actin filaments (F-actin), which are organized as bundles in filopodia and branched networks in lamellipodia. Growth cones cultured in three-dimensional (3D) matrices, or observed within tissues, have similar domain structures, albeit smaller and less flat (Harris et al., 1985; PorteraCailliau et al., 2005; Sabry et al., 1991). For some large growth cones, such as those of Aplysia, growing on a highly adhesive substrate results in an observable transitional (T) region between the C and P domain that contains a contractile arc formed by actinemyosin bundles, which provide contractile forces on microtubules (see later) (Medeiros et al., 2006). Although growth cones may come in different sizes and shapes, their movement generally can be broken down into three steps: (1) F-actin-mediated membrane protrusion in the P-domain allowing the growth cone to expand in size; (2) advancement of the microtubule-rich C domain to the P-domain to form a new front; and (3) consolidation to generate a new segment of the axon through retraction of the peripheral membrane at the rear end of the growth cone. The coordination of these steps determines the rate of axonal growth and also the growth direction of extension (Lowery and Van Vactor, 2009; Omotade et al., 2017). In culture, growth cones can split in the middle to generate two new growth cones that extend in different directions, forming two daughter branches (Gallo, 2011). Alternatively, a new growth cone may form from an existing axonal shaft either along the trailing axon following growth cone collapse (Davenport et al., 1999) or from a region demarcated by the growth cone that pauses during its advancement (Szebenyi et al., 1998). However, in vivo imaging of axons from Xenopus retinal ganglion cells (RGC) or mouse cortical neurons suggest that branches are rarely formed by growth cone splitting (Harris et al., 1987; Portera-Cailliau et al., 2005), indicating the latter two mechanisms might be used predominantly to generate different types of branches in vivo. The growth of new collateral branches is often preceded by membrane protrusion initiated in the middle of axonal shafts, as shown in sensory and cortical neurons (Hand et al., 2015; Ketschek and Gallo, 2010). While the grow cone is vital for axon growth and branching during development, injured axons in the mature nervous system often lack this motile structure. Instead, an oval structure called retraction bulb is usually formed at the tip of the proximal stump of a lesioned axon. It contains disorganized microtubules and lacks the ability to grow, thereby contributing to the intrinsic failure of CNS nerve regeneration (Bradke et al., 2012; Erturk et al., 2007).

3.2.2 Regulation of cytoskeleton assembly F-actin and microtubules are two polymers that are dynamically assembled from protein subunits (Kueh and Mitchison, 2009), and which are important for neuronal migration and morphogenesis (see Section 1.1). Pioneering studies using inhibitors that block their assembly have demonstrated that axon growth and branching is dependent on these two cytoskeletal components (Yamada et al., 1970), and extensive imaging analyses have uncovered general principles regarding their assembly during axon growth and branching (Omotade et al., 2017; Pacheco and Gallo, 2016).

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Axon growth is primarily driven by the dynamic remodeling of the actin cytoskeleton in the growth cone, which is led along its trajectory by lamellipodia and filopodia that constantly probe its surroundings. Once a positive cue is received, these filopodia are invaded by dynamic microtubules that subsequently become stabilized. Generating collateral branches along the axon requires a similar sequence of events. Actin-driven filopodia first arise from actin-rich regions called actin patches in axons of dissociated neurons (Ketschek and Gallo, 2010) or from F-actin pools in axons growing in tissue slices (Hand et al., 2015). Microtubules, normally bundled within the axon, are then destabilized and enter the filopodia either by the transport of small microtubule fragments or by dynamic polymerization (Gallo, 2011; Kalil and Dent, 2014; Pacheco and Gallo, 2016). The coordinated assembly of these two cytoskeleton systems in axon growth and branching depends on a vast number of regulatory proteins that we discuss in the following sections.

3.2.2.1 Actin F-actin consists of polarized actin polymers that usually assemble to elongate at one end called barbed ends and disassemble to shorten at the opposite end called pointed ends. If elongation is equal to shortening, this leads to a treadmilling phenomenon that allows the F-actin to move in space (Bugyi and Carlier, 2010). In cells, the barbed ends of F-actin face the plasma membrane, whereas the pointed ends are anchored at the F-actin networks deep inside (see later). There, actin treadmilling happens on a large scale with polymerization right underneath the membrane and depolymerization in the back of F-actin network, thus providing the driving force for membrane protrusion (Gomez and Letourneau, 2014; Insall and Machesky, 2009; Rottner and Schaks, 2018). In the growth cone, such force drives the dynamic extension of filopodia that contain actin bundles or lamellipodia that are populated with crisscrossed actin networks (Lowery and Van Vactor, 2009). In addition, the entire F-actin network is pulled back due to membrane resistance as well as by the myosin II motor anchoring to the T domain, resulting in a commonly seen phenomenon called retrograde flow (Gomez and Letourneau, 2014; Lowery and Van Vactor, 2009; Omotade et al., 2017). Since F-actin assembly powers growth cone motility, regulation of actin dynamics is important for axon growth and branching (Fig. 3.1).

FIGURE 3.1 Subcellular structures in axon growth and guidance. Microtubules (blue lines with arrows) are filled in the axon, the branch, and the growth cone. They may also extend into filopodia in the leading edge. F-Actin (shown in red lines) is mainly enriched under the membrane and its assembly provide the protrusive force. Membrane vesicles (yellow circles) are transported in the axon along microtubule tracks and fuse with cell membrane at different sites. Cell adhesions (shown in green patches) provide anchor for the growth cone. These structures are the targets for regulation shown in Figure 3.2.

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As is observed in nonneuronal cells, F-actin dynamics are regulated by several factors that control different events in the F-actin assembly/disassembly cycle (Luo, 2002; Pollard and Borisy, 2003; Rottner et al., 2017). One event is actin nucleation, a rate-limiting step that can be activated by several factors to generate an actin nucleus for F-actin assembly. The first identified activator was the Arp2/3 complex (Insall and Machesky, 2009), which initiates actin polymerization and stays at the plus end to generate branched actin networks from existing filaments. It is allosterically regulated by factors like the WASP protein and the WAVE complex (Insall and Machesky, 2009), which link F-actin to extracellular signaling. The Arp2/3 complex is critical for growth cone motility as well as axon growth and branch formation (Korobova and Svitkina, 2008; Spillane et al., 2011; Strasser et al., 2004), and its function in the growth cone is substrate dependent (San Miguel-Ruiz and Letourneau, 2014). Another actin nucleator is formin, which promotes actin nucleation but remains bound to the barbed end afterward (Courtemanche, 2018; Goode and Eck, 2007; Grikscheit and Grosse, 2016). Formins define a large family of proteins (Higgs, 2005) that regulate axon growth by coordinating F-actin and microtubules (Matusek et al., 2008; Szikora et al., 2017). Finally, a novel actin nucleator called Cordon-bleu was found to promote axon branching by hippocampal neurons in culture (Ahuja et al., 2007). Thus, there is no doubt that actin nucleation is a key step for regulating actin reorganization during axon growth and branching. Another step subject to regulation in the actin assembly cycle is filament capping, which is best illustrated by the studies of Ena/VASP proteins. These proteins promote actin assembly by binding to the barbed ends of F-actin and recruiting actin monomers (Drees and Gertler, 2008). Genetically deleting all three homologs (Mena, EVL, and VASP) in mice led to a failure of axon initiation in cortical neurons (Kwiatkowski et al., 2007). This defect is attributed to the loss of actin bundles and, hence, filopodia (Dent et al., 2007). Interestingly, however, other neurons, such as RGCs or dorsal root ganglion (DRG) sensory neurons, still extend axons. One explanation is that these neurons normally grow on the extracellular matrix (ECM) protein laminin, but cortical neurons do not, pointing to differential regulation of filament capping by distinctive extracellular environments. Dynamic assembly of F-actin also requires the disassembly of old F-actin in the back of actin networks to recycle the protein building blocks. This step is controlled mainly by cofilin, a cytoplasmic protein that severs and depolymerizes Factin (Pollard and Borisy, 2003). Cofilin is regulated by phosphorylation via LIM kinase and the phosphatase Slingshot. Studies of these two upstream regulators in neuronal culture and in intact animals suggest a critical role for cofilin in axon growth and branching (Dent et al., 2010; Endo et al., 2003, 2007; Hocking et al., 2009). Although most studies of F-actin regulation in neurons focus on actin assembly in the growth cone, F-actin filaments are present in axons but are difficult to resolve by conventional methods, such as electron microscopy (Schnapp and Reese, 1982). Recent advances in superresolution imaging have led to the identification of periodic actin rings in the axonal shaft in a variety of neuronal cell types from several different organisms (D’Este et al., 2016; Leterrier et al., 2015; Xu et al., 2013). These actin rings are separated from each other by bII-spectrin, with a space of w180e190 nm. The actin binding protein adducin, which caps F-actin barbed tips, is thought to stabilize the actinespectrin lattice, rendering it resistant to actindepolymerizing drugs (Xu et al., 2013; Zhang et al., 2014). Though actin rings are stable, other axonal F-actin structures are surprisingly dynamic and are able to polymerize and depolymerize. These structures appear either as actin waves that undergo treadmilling and move along axons (Ruthel and Banker, 1999), or as actin trails that polymerize from “hot spots” via formin (Ganguly et al., 2015; Roy, 2016). Although their exact functions remain to be determined, it has been proposed that actin rings support membrane proteins such as integrins and ion channels, whereas actin waves and trails mediate actin transport (Papandreou and Leterrier, 2018; Roy, 2016). Furthermore, actin patches rich in Arp2/3 complexes have been shown to serve as precursors of collateral branches (Ketschek and Gallo, 2010; Spillane et al., 2011, 2012).

3.2.2.2 Microtubules The assembly and transport of long microtubule polymers has been shown to contribute to axon growth and branching (Conde and Caceres, 2009; Dent and Gertler, 2003; Kapitein and Hoogenraad, 2015; Leterrier et al., 2017). In growth cones, individual microtubule polymers extend out from the neck regions into the C-domain, growing in different directions and sometimes entering the P-domain, where dynamic microtubule assembly helps determine the direction of growth cone navigation (Fig. 3.1). At the neck region, bundled microtubules contribute to the consolidation step during growth cone advancement. In the axon, microtubules are more stable and provide tracks for fast axonal transport of proteins and membranes needed for axon and branch growth. In addition, microtubules can form loops in the growth cone or regions of axons that will eventually develop into branches (Dent et al., 1999; Dent and Kalil, 2001). Like F-actin, microtubules are polarized polymers that are assembled from tubulin heterodimers. In cells, the assembly usually happens at one end called plus ends, cycling between growth, shrinkage, and occasional pauses. This intrinsic microtubule assembly property, termed dynamic instability (Mitchison and Kirschner, 1984), can support axonal

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development (Dent et al., 1999; Dent and Gertler, 2003; Mitchison and Kirschner, 1988; Sabry et al., 1991). In axons, individual microtubule polymers can be further subdivided into three regions based on their stability: (1) a dynamic region at the plus end, also known as a labile domain, which undergoes dynamic assembly and disassembly; (2) a stable region in the middle, which is less susceptible to catastrophe; and (3) a cold stable region at the minus end, which is resistant to depolymerization induced by low temperature (4
C) and/or millimolar calcium ions (Baas et al., 2016; Song and Brady, 2015). The stability of microtubules changes throughout axonal development, usually increasing as the axon ceases to grow. The presence of dynamic and labile microtubules allows microtubules to explore the intracellular volume of the growth cone (Dent et al., 1999; Sabry et al., 1991) and, subsequently, the transition from dynamic to stable microtubules on one side of the growth cone consolidates growth cone motility in a specific direction (Dent and Gertler, 2003; Lowery and Van Vactor, 2009; Mitchison and Kirschner, 1988). This function is supported by studies that locally perturb microtubule stability in the growth cone (Buck and Zheng, 2002). In addition, treatment of Xenopus neurons with the microtubuledepolymerizing drug vinblastine at doses that block only dynamic microtubules increased growth cone wandering and reduced persistent forward movement, suggesting the involvement of dynamic microtubules in controlling the processivity of normal axon growth (Tanaka et al., 1995). Finally, inhibiting microtubule dynamics prevents branch formation but not axon elongation in cultured cortical neurons (Dent and Kalil, 2001). Thus, microtubule dynamics serve a key role in regulating axon growth and branching. The dynamic assembly of microtubules at plus ends is influenced by two groups of plus endetracking proteins (þTIPs). One group, including end binding proteins (EBs), recognizes and binds microtubule plus ends directly and thus forms the core of the þTIP complex. The other group binds microtubule plus ends by association with the first group via a CAP-Gly domain or an SxIP motif (Akhmanova and Steinmetz, 2008, 2015). Many of these proteins have been found to contribute to axon growth (van de Willige et al., 2016). For example, two plus endebinding proteins, APC and CLASP2, are important for NGF-dependent axon growth of sensory neurons and neocortical development (Dillon et al., 2017; Zhou et al., 2004), and the microtubule plus endetracking proteins SLAIN1/2, ch-TOG, and TACC3 can promote axonal development (Erdogan et al., 2017; van der Vaart et al., 2012). While microtubule assembly happens at plus ends, the opposite ends, called minus ends, are bound and stabilized by a family of calmodulin-regulated spectrin-associated proteins (CAMSAP1-3) (Akhmanova and Hoogenraad, 2015) both in vitro and in vivo (Jiang et al., 2014). Since they prevent depolymerization of microtubules, CAMSAPs are important for neuronal polarization and dendrite development (Yau et al., 2014, 2016), but their function in axon growth and branching remains to be determined. Interestingly, ninein, another protein that binds to the minus end of microtubules, can influence growth and branching when targeted by its upstream effector Sip1 (Srivatsa et al., 2015). Other cellular factors that affect microtubule dynamics can regulate axon growth and branching. For example, CRMP2, a protein that promotes microtubule assembly in vitro, regulates neuronal polarity and branch formation (Fukata et al., 2002; Yoshimura et al., 2005) and axon growth (Higurashi et al., 2012) in cultured neurons. Furthermore, SCG10 and SCG10-like proteins that bind to tubulin heterodimers are highly expressed in the brain and are implicated in regulating axon growth and branching in culture (Grenningloh et al., 2004; Poulain and Sobel, 2007). Where are new microtubules generated in neurons in the first place? New microtubules are usually assembled from nucleation, a rate-limiting step, with the aid of g-tubulin at the minus end (Oakley et al., 2015). Like in nonneuronal cells, it has been long thought that axonal microtubules are nucleated from the centrosomes in the cell body and then released and transported to axons (Baas et al., 2005), since antibody blocking of g-tubulin abolished or compromised axon growth of cultured sympathetic neurons (Ahmad et al., 1994). However, Drosophila neurons lacking centrosomes develop nearly normal axons (Basto et al., 2006), and ablation of centrosomes in hippocampal neurons does not affect axon growth (Stiess et al., 2010). Although manipulating g-tubulin alters in microtubule polarity, neurons are still able to develop normally (Nguyen et al., 2014). These studies suggest that there are multiple ways of nucleating new microtubules in axons. Indeed, microtubules have been found to be nucleated at noncentrosomal nucleation sites in the cell body (Stiess et al., 2010), and acentrosomal microtubules are nucleated with the aid of a RAN-dependent protein TPX2, which is required for maintaining axon length (Chen et al., 2017). The stability of axonal microtubules is important for axon growth and branching. It is achieved by many lattice-bound microtubule-associated proteins (MAPs), which often have multiple microtubule-binding sites to stabilize microtubules (Dehmelt and Halpain, 2005; Kapitein and Hoogenraad, 2015). A large number of MAPs have been shown to impact axon morphology, with regulation often governed by their phosphorylation state (Ramkumar et al., 2018). For example, the first identified MAP, tau, is localized to the axon, and its dephosphorylation leads to its dissociation from microtubules and the access of katanin to microtubules (Qiang et al., 2006; Yu et al., 2008b). Another axonal MAP, MAP1B, is also important for axon growth and branching since knockdown results in enhanced axonal branching, suggesting it represses branching (Barnat et al., 2016; Tymanskyj et al., 2012). Phosphorylation of doublecortin, a MAP found in inherited mental

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retardation and epilepsy (Allen and Walsh, 1999), can also restrict axon branching (Bilimoria et al., 2010). Recently, a less well-studied MAP, MAP7, was found to regulate and promote collateral branch formation in DRG neurons (Tymanskyj et al., 2017). Despite the presence of many MAPs, understanding how they are spatially and temporally controlled to regulate microtubule stability and function remains an exciting area of further investigation. Though axonal microtubules are relatively stable, they need to be broken (destabilized) to adapt to morphological changes of axons, often associated with branch formation. This is achieved by a family of microtubule-severing proteins: katanin, spastin, and fidgetin. These cytoplasmic proteins contain an “AAA domain,” which provides the ATPase activity needed to break the microtubule lattice. The breakage produces free microtubule plus ends that can polymerize and also minus ends that can depolymerize (Baas et al., 2005; Salinas et al., 2007). The level of katanin in neurons correlates with the ability of axons to grow, with the highest expression during rapid axon growth and decreased expression when axons cease to grow (Karabay et al., 2004). Spastin was found to be mutated in patients with spastic paraplegia, a neurodegenerative disease that is characterized by a progressive spasticity and lower limb weakness (Errico et al., 2002). When expressed in cultured neurons, spastin proteins harboring the human mutation can aggregate, altering microtubule dynamics, and inhibit axon outgrowth (Solowska et al., 2014, 2017). Loss of spastin also reduces axon outgrowth in zebrafish (Wood et al., 2006). Both katanin and spastin can promote branching by generating short microtubules necessary for branch formation, but only katanin depends on the phosphorylation and subsequent dissociation of tau, a MAP that protects against katanin-dependent severing (Chen et al., 2014; Qiang et al., 2010; Yu et al., 2008b). Spastin has also been shown to disassemble microtubules during branch remodeling and synaptic elimination (Brill et al., 2016). Recently, a third severing protein called fidgetin has been identified and shown to regulate axon growth, guidance, and regeneration (Austin et al., 2017; Fassier et al., 2018; Leo et al., 2015). Microtubule severing generates short microtubule fragments that can be transported along long and stationary microtubule polymers (Baas et al., 2005) and thus may facilitate axonal branch formation. This model is supported by electron microscopic studies of axonal branches (Yu et al., 1994) and real-time imaging of microtubules in live cultured neurons (Dent and Gertler, 2003; Gallo and Letourneau, 1999). In addition, transport of microtubules has been suggested to provide tubulin to axons and growth cones. However, the mechanism of transport had been open to debate (Baas and Buster, 2004). Early pulse-chase studies suggested that microtubules move along axons at a rate that is significantly slower than motor-based fast axonal transport; however, later studies indicate that this slow transport is due to frequent pauses intermingled with fast transport. In fact, in the squid giant axon, tubulin oligomers are transported as large complexes by a kinesin-based motor (Terada et al., 2000). In cultured mammalian neurons, motor-based fast transport has been observed, and the movement is asynchronous and infrequent, contributing to the overall slow rate as previously observed (He et al., 2005; Wang and Brown, 2002). Such transport has been suggested to be mediated by the kinesin heavy chain (Jolly et al., 2010) and provides a mechanism for regulating axon growth (Barlan and Gelfand, 2017; Lu and Gelfand, 2017; Lu et al., 2016; Winding et al., 2016). Interestingly, recent work also suggests that cytoplasmic dynein, a minus-end motor, is responsible for microtubule transport (He et al., 2005; Rao et al., 2017). Regardless of the mechanism, microtubule transport could provide another site for regulating axon growth and branching. Finally, microtubules are subject to several modes of posttranslational modification (PTM), including tyrosination, acetylation, and glutamination (Janke and Bulinski, 2011). PTMs are responsible for regulating microtubule stability as well as the interaction between MAPs and motor proteins with microtubules. For example, microtubule tyrosination and glutamination increase the activity of the severing protein spastin (Roll-Mecak and Vale, 2008; Valenstein and RollMecak, 2016), and microtubule acetylation promotes the binding of kinesin and its cargos (Balabanian et al., 2017; Reed et al., 2006). These modifications are enriched in specific parts of the neuron (Hammond et al., 2010) and developmentally regulated (Song and Brady, 2015), and so elucidating their functional roles in axon growth and branching is under active investigation (Hammond et al., 2010).

3.2.3 Interaction between F-actin and microtubules Since axon growth and branching are dependent on actin and microtubule assembly, proteins that interact with both filaments provide a mechanism to coordinate their behaviors. Drebrin is an F-actin-binding protein that also binds to microtubules via EB3 (Geraldo et al., 2008; Merriam et al., 2013). In the growth cone, it is localized to the T-domain, modulating the interaction between F-actin and microtubules and regulating axon outgrowth. During collateral branch formation, drebrin localizes to filopodia and promotes entry of microtubules into filopodia (Ketschek et al., 2016). Microtubules can overcome the actin retrograde flow when coupled to actin bundles within the growth cone. This is achieved through drebrin and Spectraplakins (ACF7-EB1), which when cross-linked to actin in filopodia can regulate axon outgrowth (Alves-Silva et al., 2012). There is also evidence that, when the minus endedirected motor dynein is anchored to the membrane, via the actin cortex, it can promote forward sliding microtubules and axon extension (del Castillo et al., 2015; Roossien et al., 2014).

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Another family of proteins able to link actin and microtubules are septins (Mostowy and Cossart, 2012), small GTP-binding proteins that form heterooligomeric complexes and assemble into filaments and rings. There are 13 septin genes (SEPT1-12, 14) in humans, and many undergo alternative splicing (Russell and Hall, 2011). In axons, SEPT6 localizes with actin patches and promotes actin polymerization, leading to the formation of filopodia. SEPT7 then promotes entry of microtubules into filopodia, facilitating axonal branch formation (Hu et al., 2012). SEPT9 modulates the transport of kinesin-1 and kinesin-3, regulating trafficking into axons and dendrites (Karasmanis et al., 2018). Thus, septins may represent a new class of polymers with great potential to control axon growth and branching.

3.2.4 Membrane trafficking and axonal transport The soma of an average mammalian neuron is about 20 mm in diameter, but its axon can extend up to 1 m in humans. If the axon diameter is w1 mm, the surface area of the axon is about 3000 times that of the soma (3  106 vs. 1  103 mm2). Considering a normal growth rate of w20 mm/h, each neuron must add at least 1.5 times its surface area every day. Including the formation of axonal branches, this growth requires a considerable amount of plasma membrane to be synthesized and incorporated into the extending axon (Quiroga et al., 2018). There has been major debate in the field as to where new membranes are added along axons. Conceivably, membranes could be added to the growth cone via anterograde transport, discussed later, and then moved back toward the cell body by retrograde flow. However, labeling studies suggest that new membrane can be added along the axon as well (Futerman and Banker, 1996). Regardless, lipids are synthesized in the smooth endoplasmic reticulum (ER), which is located in the cell body and also present in axons, whereas membrane proteins are made in the rough ER and modified posttranslationally in the Golgi apparatus. Unlike dendrites (Sann et al., 2009; Ye et al., 2007), no Golgi apparatus has been observed in mammalian axons. Thus, membrane trafficking via vesicular transport is likely the main route by which new membranes are delivered to the surface of axons and growth cones (Fig. 3.1). As a result, new branching sites can be determined by sites of membrane vesicle fusion (Winkle and Gupton, 2016). Two major events related to this process are important for axon growth and branching. The first is membrane trafficking between different post-Golgi compartments, including early and late endosomes (Sann et al., 2009). Early endosomes are bound by the small GTPase Rab5 and are associated with Unc51, a conserved protein kinase. Loss of Unc51 function in worm and mammalian neurons leads to shortened axons and increased axonal branch formation (Ogura et al., 1994; Zhou et al., 2007). Late endosomes deliver secretory vesicles to the plasma membrane for exocytosis. Disrupting their membrane targeting interferes with axon growth in cultured hippocampal neurons. Moreover, protrudin, a protein that binds to the GDP-form of Rab11 associated with recycling endosomes, was shown to regulate axon extension in culture (Shirane and Nakayama, 2006). Recently, it has been observed that local SNARE-mediated exocytosis is critical for axon branching induced by the guidance cue Netrin-1 (Winkle et al., 2014). Additional regulation of membrane trafficking is mediated by the BAR domain containing proteins srGAP and syndapin (Dharmalingam et al., 2009; Guerrier et al., 2009). BAR domains are conserved structural motifs that dimerize via their coiled-coil domains to form modules with positively charged residues aligned on one surface. This surface allows BAR-containing proteins to interact with the plasma membrane and change membrane curvature in a manner that is important for exocytosis and for filopodia and axonal branch formation (Frost et al., 2009). The second event relates to fast axonal transport, which is mediated by molecular motors that travel along microtubules and deliver both membrane and protein cargos to different sites (Hirokawa and Takemura, 2004). This is an important process considering the large distances that any protein or membrane organelle synthesized in the cell body needs to travel to reach the nerve terminal. Early labeling studies showed that transport is achieved in two modes: fast, w400 mm/day (1 mm/s), often seen with organelles, and slow, w8 mm/day (0.1 mm/s), mainly associated with the transport of cytoskeletal proteins (Maday et al., 2014). Fast axonal transport is achieved by motor proteins called kinesins and dyneins that consume ATP and move along microtubules (Hirokawa and Takemura, 2004). Because microtubules in the axon have their plus ends projecting toward the growth cone, the kinesin superfamily proteins (also known as KIFs), which move toward the plus end of microtubules, generally support anterograde transport and move proteins or organelles away from the cell body (Hirokawa and Takemura, 2005), whereas cytoplasmic dynein moves toward the minus end and supports the retrograde transport toward the cell body. Because of the importance of axonal transport for axon growth and branching, motor proteins are undoubtedly critical. Depletion of kinesin-5 increases short microtubule transport, reduces axonal retraction, increase axon length and branching frequency in cultured neurons (Myers and Baas, 2007), whereas deletion of kinesin-1 increases RGC axon branching in zebrafish (Auer et al., 2015). JIP3, a kinesin activator, also promotes axon elongation (Watt et al., 2015). Interestingly, one

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type of kinesin molecule, KIF2A, has a microtubule-depolymerizing activity and suppresses collateral branch formation, since it has been observed that the KIF2A mutant mouse has multiple brain abnormalities that include increased axon length and branching in cortical and hippocampal neurons (Homma et al., 2003). Retrograde transport by dynein is also critical for delivering “signaling endosomes” formed at the nerve terminals to the nucleus in the soma (Cosker et al., 2008; Ibanez, 2007) (see later for its role in neurotrophin function). Mutations in dynein and its associated proteins have been found to cause neural degeneration in mice and humans (Cosker et al., 2008). Dynein has also been shown to sort microtubule polarity (Rao et al., 2017) and generate the force that drives the cytoskeleton forward during axon elongation (Roossien et al., 2014). Overall, the large family of motor proteins and their regulators are important in regulating axon development and nerve regeneration. One organelle that is particularly interesting in transport regulation is the mitochondrion, which undergoes bidirectional movement in axons. Since the generation and growth of new axonal branches relies on many ATP-dependent processes, local ATP production provided by mitochondria at sites of growth is important. Mitochondria can be transported up and down the axon, but many remain stationary (Miller and Sheetz, 2006; Sheng, 2014). The mitochondrial binding protein syntaphilin, and its upstream regulators LKNB1 and NUAK1, are responsible for anchoring mitochondria to the cytoskeleton. Deletion of these molecules increases the number of motile mitochondria and results in a decrease in axonal branch formation (Courchet et al., 2013; Spillane et al., 2013).

3.2.5 Protein translation and stability During axon growth and branching, axon volume can approach w200-fold that of a typical cell body (8  105 vs. 4  103 mm3), and this is achieved by adding w10% of total soma mass per day. Several intracellular signaling pathways involved in cell size control have been implicated in axonal development. One pathway involves the mammalian target of rapamycin (mTOR), a protein kinase that is part of a key regulatory complex that controls protein synthesis during general cell growth (Bhaskar and Hay, 2007). Cell culture studies using DRG neurons show a requirement of this pathway and protein synthesis for regenerating axons (Verma et al., 2005). This was also demonstrated in an injury study in the optic nerve, where blocking the upstream mTOR inhibitor, phosphatase and tensin homolog (PTEN), led to increased optic nerve regeneration following a crush lesion in mice (Park et al., 2008). Furthermore, deletion of PTEN in the mouse brain results in increased dendritic arborization (Kwon et al., 2006) and axon branching (Drinjakovic et al., 2010). The suppressing effects of PTEN on axon outgrowth have been attributed to its role in regulating microtubule stability (Kath et al., 2018). Moreover, molecular perturbation of another upstream mTOR regulator, Tsc1/2, affects axon growth (Choi et al., 2008). These studies suggest that pathways critical to cell growth are important for generating neuron morphology during development and axon regeneration after injury. Cytoplasmic proteins, just like membrane proteins, are mainly synthesized in neuronal cell bodies. However, free ribosomes have been observed in axons by electron microscopy (Tennyson, 1970), and mRNA is found in axons and growth cones (Bassell et al., 1998; Piper and Holt, 2004). Like proteins, mRNA can be transported by cellular factors that recognize specific sequences in their untranslated region (Zhang et al., 1999). Local protein synthesis in axons and growth cones contributes to axon guidance in response to extracellular cues (Brittis et al., 2002; Campbell and Holt, 2001; Ming et al., 2002). In addition, culture studies show that local protein synthesis is needed for signal-induced growth instead just for basal axon growth (Gracias et al., 2014; Hengst et al., 2009). Similar observations have also been made in regenerating DRG neuron axons (Verma et al., 2005; Vogelaar et al., 2009). Interestingly, local protein translation is also important during axonal branch development since mitochondrial recruitment to branch sites is a prerequisite for local translation and branch stabilization (Spillane et al., 2012, 2013). Consistent with these data, local actin synthesis at the base and interiors of new branches contributes to their stabilization (Wong et al., 2017). Thus, local protein synthesis in neuronal subcompartments is important for axon growth and branching (Rangaraju et al., 2017). Another pathway that regulates cytoplasmic protein levels involves the ubiquitin-proteasome system (UPS), which controls protein stability through the action of many substrate-specific E3 ligases (Hershko and Ciechanover, 1998). The UPS modulates growth cone responses to guidance cues in Xenopus neurons (Campbell and Holt, 2001); it promotes local degeneration of axon terminals during Drosophila development (Watts et al., 2003); and it mediates the intrinsic program of Wallerian peripheral neuronal degeneration in mammalian neurons following injury (Zhai et al., 2003). In addition, the cell cycle regulator Cdh1-APC, a ubiquitin ligase, controls axon growth and morphology in cerebellar granule cells via regulation of its target, SnoN, a transcriptional corepressor (Konishi et al., 2004; Menon et al., 2015; Stegmuller and Bonni, 2005). The E3 ubiquitin ligase Phr1 was shown in the mouse to be important for axon growth of sensory and motor axons by interacting with microtubules (Lewcock et al., 2007). Phr1, also found in Drosophila (Highwire) and C. elegans (RPM-1), is suggested to be a signaling hub to regulate neuronal development and axon degeneration (Grill et al., 2016).

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Furthermore, another E3 ligase, Nedd4, controls terminal axon arborization in Xenopus RGCs by directly regulating PTEN (Drinjakovic et al., 2010). Recently, the E3 ligase TRIM9 was shown to interact with VASP to modulate filopodia stability in axon outgrowth (Menon et al., 2015). The function of E3-ubiquitin ligases appears conserved through both vertebrates and invertebrates, suggesting an important role for protein regulation during axon growth and branching. Finally, microRNAs define a mechanism for the regulation of mRNA stability, which is important for a variety of cellular processes (Bushati and Cohen, 2007; Fiore et al., 2008; Wang and Bao, 2017). Two microRNAs, miR-9 and miR132, control the mRNAs encoding cytoskeleton regulators, including proteins such as MAP1B and Rasal1, a Ras GTPase, and in turn they affect the extent of axon outgrowth (Dajas-Bailador et al., 2012; Hancock et al., 2014). miR-124 has been shown to control axon growth in cultured neurons (Yu et al., 2008a), whereas miR-16 binds to and decreases levels of the protein translation initiation factors eIF2b2 and eIF4G2, leading to decreased axon growth (Kar et al., 2013). Furthermore, two enzymes important for miRNA function have been identified through a genetic screen of olfactory axon development in Drosophila (Berdnik et al., 2008), and the miR-17-92 cluster was found to enhance axon outgrowth (Zhang et al., 2013). Thus, a very large number of distinct microRNAs provide another regulatory mechanism in axon growth and branching, and identification of their targets will be key to understanding their functions.

3.3 Extracellular regulation of axon growth and branching during neural development 3.3.1 Nerve growth factor and neurotrophic factors Nerve growth factor (NGF) was originally isolated from the mouse sarcoma tissue for its activity to promote neuronal growth and survival of sympathetic ganglion (SG) explants in culture (Levi-Montalcini, 1987). It belongs to a family of related molecules called neurotrophins, which include neurotrophin-3 (NT-3) and brain-derived neurotrophic factor (BDNF) (Hallbook, 1999). Each class of neurotrophin binds selectively and with high affinity to a specific Trk receptor (TrkA, TrkB, or TrkC), which dimerizes upon ligand binding to its Ig domains and signals through its intracellular tyrosine kinase domain (Chao, 2003). In addition, all neurotrophins bind to the low-affinity receptor p75. As implied by their name, neurotrophins regulate neuronal survival (Snider, 1994), but in addition to this trophic function, they are important for axon growth and branch morphology (Huang and Reichardt, 2001). Extensive evidence defining neurotrophin signaling pathways and function has been provided by studies focused on peripheral sensory neurons in the DRG, trigeminal ganglion (TG), and SG (Glebova and Ginty, 2004; Kuruvilla et al., 2004; Patel et al., 2000). The growth-promoting activity of neurotrophic factors was evident in early days of NGF analysis, when it was observed that NGF stimulates radial outgrowth of axons from SG explants (Levi-Montalcini, 1987). Most neurotrophins are able to promote sensory axon elongation in dissociated or slice cultures. In vivo, it has been shown that sensory axons in the developing limb bud preferentially grow toward ectopically placed beads saturated with different neurotrophic factors (Tucker et al., 2001). However, a big debate ensued as to whether or not the growthepromoting activity of neurotrophins is simply a result of promoting cell survival. To separate these functions, several studies took advantage of deleting BAX, a proapoptotic protein required for programmed cell death. In a double knockout that removes both TrkA and BAX in the mouse, in which cell death normally associated with the loss of NGF is prevented, DRG sensory neurons fail to innervate the superficial layers of neonatal skin (Patel et al., 2000). Similar observations were made of sympathetic innervation in NGF/; BAX/ mice, in which reduced innervation of sympathetic target organs was observed to varying degrees (Glebova and Ginty, 2004). Such a growth requirement for neurotrophin signaling is consistent with the role of p75, a lowaffinity neurotrophin receptor that is required for the innervation of cutaneous and sympathetic targets (Lee et al., 1992). In addition to axon growth, neurotrophins regulate axonal branch morphology. Culture studies using dissociated embryonic sensory neurons show that NGF and NT-3 are required to control different aspects of axon growth and terminal arborization (Lentz et al., 1999). This morphological requirement was demonstrated in TG explants, in which NGF promoted axon elongation and NT-3 enhanced axonal collaterals (Ulupinar et al., 2000). In addition, the p75 knockout mouse has short and poorly branched peripheral axons during embryonic development (Bentley and Lee, 2000). Glial cell lineederived neurotrophic factor (GDNF) belongs to a unique family of neurotrophic factors that include three other members (Airaksinen and Saarma, 2002). Their receptors consist of the receptor tyrosine kinase Ret, which functions as a signaling subunit, and the GDNF receptor, a GPI-linked ligand-binding subunit (Airaksinen et al., 1999). Early studies showed that GDNF produced robust axon outgrowth of ciliary ganglia in culture and that GDNF antibodies blocked ciliary axon outgrowth in chick embryos (Hashino et al., 2001). The dependence on GDNF signaling in axon development is best revealed by animals lacking the Ret receptor, which fail to project axons from all of SGs (Enomoto et al., 2001). Furthermore, studies of Ret signaling in DRG neurons suggest that it is important for collateral axon formation in mechanosensory neurons (Luo et al., 2009).

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Neurotrophins are usually secreted by target tissues far away from the neuronal cell body or soma, suggesting that the receptors in the growth cone are important for relaying signals responsible for axon growth. Early studies using compartmentalized cultures showed that NGF applied exclusively to distal growth cones is sufficient to promote axon extension, whereas NGF applied to the cell body is not (Campenot, 1977). In vivo studies based on antibody perturbation or genetic deletion support this idea (Glebova and Ginty, 2004; Ruit et al., 1992), suggesting the presence of local and long-range signaling. In fact, activation of TrkA receptors in the axons leads to not only local cytoskeleton reorganization in the axons (Gallo, 2011; Spillane et al., 2012) but also retrograde endosome signaling within the cell body (Barford et al., 2017; Yamashita and Kuruvilla, 2016). Recently, it was shown that TrkA-containing endosomes are retrogradely transported in heterogenous vesicles called multivesicular bodies (Ye et al., 2018). Transport of signaling endosomes to the soma leads to transcriptional regulation mediated by transcription factors (Scott-Solomon and Kuruvilla, 2018). Such regulation is not limited to NGF signaling, since in vivo studies show that target-derived NT-3 is required for proprioceptive DRG neurons to make connections via NT-3 regulation of the transcription factor ER81, whereas target-derived GDNF is important for motor axons to correctly innervate muscle targets owing to the regulation of the transcription factor PEA3 (Ladle et al., 2007). Interestingly, NGF and NT3 are differentially regulated by a hierarchical neurotrophin signaling cascade that coordinates sequential stages of axon growth, innervation of targets, and neuronal survival in sympathetic neurons (Kuruvilla et al., 2004), suggesting that more complex developmental regulation by neurotrophins can be at play. Because of their activities that affect axon growth and branching, neurotrophins have been proposed to stimulate axon regeneration and sprouting after injury. Indeed, injection of NT-3 into the lesion sites in adult rat spinal cords causes enhanced sprouting of regenerating axons in the corticospinal tract (Schnell et al., 1994). Furthermore, NGF, NT-3, and GDNF promote varying degrees of functional regeneration of DRG sensory neurons into the adult spinal cord following injury (Ramer et al., 2000). Although the therapeutic use of NGF is hampered by increased neuropathic pain associated with extensive NGF-induced sprouting of pain sensing axons, the development of treatments with other neurotrophins, or more targeted activation of neurotrophin intracellular signaling pathways, still holds the promise for treating nerve injury and degeneration (Keefe et al., 2017).

3.3.2 Guidance molecules: netrin, slit, semaphorin, ephrin, and wnt Since axon guidance is intimately associated with axon growth and branching, it is not surprising that molecules controlling axon guidance (see Sections 1.4e1.8) can influence both processes (Dickson, 2002; Gibson and Ma, 2011; Kolodkin and Tessier-Lavigne, 2011). In the early attempts to isolate axon guidance molecules such as Netrin, the growthpromoting property was often used to quickly assay their activity during purification (Serafini et al., 1994). Both in vitro and in vivo studies demonstrated that, like axon guidance, these molecules can both positively and negatively regulate axon growth and branching of neurons in the CNS and the PNS (Fig. 3.2). Netrin is a target-derived factor that is secreted by the floor plate to attract commissural axons to cross the midline (Serafini et al., 1994) (see Section 1.4). It also promotes the growth of commissural axons, since mice lacking functional netrin have much shorter commissural axons (Serafini et al., 1996). Knockout analysis showed that Netrin-1 promotes the growth of thalamic axons through an intermediate structure on their path to the cortex and demonstrated that Netrin-1 is capable of influencing both axon growth and branching during CNS development (Braisted et al., 2000). In cultured cortical neurons, addition of Netrin-1 results in an increase in axon branching without affecting axon length (Dent et al., 2004). The positive influence on axon branching has been demonstrated for RGC axons in the Xenopus tectum, where Netrin-1 increases axon terminal remodeling dynamics, including branch addition and retraction, and results in an overall increase in axon arborization (Manitt et al., 2009). One of the first guidance molecules identified as a positive regulator of axon branching was the N-terminal fragment of the repulsive guidance molecule Slit2 (see Section 1.7). This molecule was found to promote both axon elongation and branching of dissociated DRG sensory neurons in 3D collagen gels (Wang et al., 1999). This branching activity was supported by mouse mutants with deletion of the three Slit genes, or their cognate Robo1 and Robo2 receptors, in which the TG ophthalmic branch showed significant reduction of axon arborization (Ma and Tessier-Lavigne, 2007). This branching activity was also demonstrated in the peripheral projection of zebrafish and Drosophila sensory neurons (Yeo et al., 2004; Zlatic et al., 2009). In addition, Slit regulates CNS axon growth and branching, since application of Slit2 induces arborization of the central trigeminal axon within the brainstem target region in culture, and this activity correlates well with the expression of Slit2 and Robo receptors during development (Ozdinler and Erzurumlu, 2002). However, studies on RGC axons in the zebrafish and Xenopus optic tectum suggest that Slits and Robos exhibit no positive or even inhibitory effect on axon arborization (Campbell et al., 2007; Hocking et al., 2010), raising the possibility of differential

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Negative regulators: Semaphorins, ephrins, Nogo, etc. Neural activity

Ca2+

GTPases cAMP/cGMP

Positive regulators: Neurotrophins Netrin, etc. FIGURE 3.2 Regulation of axon growth and branching by extracellular and intracellular signaling. Molecule cues in the embryonic environment provide positive or negative signals that promote or inhibit axon growth and branching during development. Neural activity can also influence these two developmental processes. These signals lead to the remodeling of the subcellular structures described in Figure 3.2 via the intracellular signaling molecules, such as Ca2þ, Rho family small GTPases, and cAMP/cGMP.

regulation of PNS versus CNS neurons by Slit. A recent study in Drosophila shows that Slit also binds to Down syndrome cell adhesion molecule (Dscam), a cell surface receptor belonging to the immunoglobulin superfamily (Ig-SF), and thereby regulates localized axonal branching (see detailed discussion later) (Dascenco et al., 2015). The semaphorin family of guidance molecules represents a large group of extracellular cues that can negatively regulate axon growth and branching based on their repulsive activities (see Section 1.5). The best studied semaphorin is the secreted cue Semaphorin 3A (Sema3A), which controls the growth and branching patterns of peripheral DRG and TG axon projections during embryonic development in the mouse (Kitsukawa et al., 1997; Taniguchi et al., 1997). This activity is mediated by its ligand-binding coreceptor Neuropilin1 (Nrp1) and the signaling coreceptor PlexinA3 and A4 (Jongbloets and Pasterkamp, 2014). A recent study shows that the Sema3-dependent Nrp1 function is required for controlling motor nerve growth in the diaphragm (Saller et al., 2016). In the CNS, Sema3A inhibits axon branching of cortical neurons in culture (Dent et al., 2004), and this inhibitory activity has been linked to pruning of hippocampal mossy fibers and pyramidal axonal branches (Bagri et al., 2003). Interestingly, studies of the semaphorin coreceptor Neuropilin2 and PlexinA3/ A4 in knockout mice have demonstrated the requirement of Sema3A function in stereotypic pruning of corticospinal collaterals from the visual cortex, but not the collicular collaterals of the motor cortex (Low et al., 2008), indicating regionspecific regulation by this extracellular cue. Such local specificity is supported by the role of Sema3A in controlling layerspecific interneuron branching in the cerebellum (Cioni et al., 2013). Interestingly, the repulsive activity of Sema3A induces axonal branch formation in Xenopus RGC axons after growth cone collapse in culture (Campbell et al., 2001), supporting the intriguing potential connection between repulsion and axon branching mentioned earlier (Davenport et al., 1999), but the underlying mechanism remains to be uncovered. Members of the ephrin family are surface-associated proteins that mediate short-range axon guidance (see Section 1.6). Their involvement in axon growth and branching is best illustrated in the development of RGC axon projections, which form terminal arbors in spatially restricted terminal zones (TZs) of the optic tectum (or superior colliculus in mammals) (McLaughlin et al., 2003). Studies of RGCs grown on alternating stripes containing different TZ membranes demonstrated that this is partially mediated by ephrin-A, a GPI-anchored ephrin that prevents ectopic branch formation (Yates et al., 2001). This conclusion is supported by inactivating ephrin-A function in developing chick embryos (Sakurai et al., 2002). Interestingly, this function is mediated not only by forward signaling via the EphA receptor (McLaughlin and O’Leary, 2005) but also by reverse signaling mediated via the interaction between ephrin-A with two neurotrophin receptor, TrkB and p75 (Lim et al., 2008; Marler et al., 2008). Since similar functions were also found for thalamic axons (Mann et al., 2002), ephrins thus regulate target-specific terminal axon arborization by restricting branching in the CNS. A recent study extended such function to the C. elegans Ephrin (EFN-4) protein, which coordinates with heparan sulfate proteoglycans (HSPGs) in the worm body wall muscle to control axon outgrowth and branching of interneurons and motoneurons (Schwieterman et al., 2016).

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Lastly, morphogens have been shown to regulate axon guidance (Charron and Tessier-Lavigne, 2005; Yam and Charron, 2013) as well as axon growth and branching. The best example is the Wnt family of ligands, which has been implicated in this process in both the CNS and the PNS. Initial evidence came from the study of WNT7a, which regulates cerebellar granule cells axon remodeling (Hall et al., 2000). In the PNS, WNT3 appears to be a factor secreted from the ventral spinal cord that inhibits axon elongation of NT-3 responsive DRG neurons, while simultaneously promoting axon arborization (Krylova et al., 2002). This activity is likely mediated by its Ryk receptor (Lu et al., 2004). Another family member, WNT5a, works with NGF to regulate sympathetic axon growth and branching (Bodmer et al., 2009). Genetic deletion of Wnt5 or its receptor, Ror, demonstrates the role of this pathway in controlling arborization of sympathetic axons within the targets in vivo (Ho et al., 2012; Ryu et al., 2013). The role of Wnts in axon branching has also been assessed in Drosophila, where impaired Wnt signaling involved in planar cell polarity affects axonal branch extension of mushroom body neurons (Ng, 2012). Thus, it is possible that distinct classes of morphogens are capable of differentially regulating axon elongation and branching in various neuronal cell types.

3.3.3 Cell adhesion molecules: permissive or instructive Cell adhesion is important for axon growth since it provides growth cones with an anchor to extend axons and branches. A molecular clutch model was proposed to account for growth cone movement driven by three coordinated events: actin polymerization at the membrane, depolymerization at the rear end of the growth cone, and retrograde flow of F-actin (see Section 3.2.2.1) (Mitchison and Kirschner, 1988). This model is supported by studies of growth cones from Aplysia neurons (Medeiros et al., 2006), Xenopus spinal neurons (Nichol et al., 2016), and rodent hippocampal neurons (Bard et al., 2008). A strong correlation was found between the rate of growth cone movement and the mechanical coupling between the ligand-bound adhesion receptors and the retrograde actin flow (Bard et al., 2008), thus linking cell adhesion to actin dynamics in axon growth and branching. Interestingly, an early study in cell culture showed that growing neurites exhibit little selectivity between different molecular substrates and no difference in growth rates on these substrates (Lemmon et al., 1992), raising the possibility that adhesion could simply provide a permissive environment for axon growth. Nonetheless, extensive molecular studies in the past two decades have demonstrated diverse functions of cell adhesion in axon and branch development (Missaire and Hindges, 2015; Short et al., 2016; Zipursky and Sanes, 2010). Cell adhesion is achieved in at least two ways: by interaction with the ECM or by binding to cell surface molecules present in neighboring cells. The ECM is a complex macromolecular network consisting of glycoproteins and proteoglycans that are often assembled into large repeating structures (Kiryushko et al., 2004). These molecules, including laminin, fibronectin, collagens, HSPG, and chondroitin sulfate proteoglycans, are often found in the extracellular space between neurons and other cell types such as glial cells, and also in surrounding structures such as basement membranes. Early in vitro studies showed that these molecules are preferred substrates for neurons to grow axons and interact with integrin receptors, a type of cell adhesion molecule (CAM) located on the neuronal cell surface (Bixby and Harris, 1991). In addition to serving as a “glue,” the ECM provides an anchorage site for extracellular factors to influence axon growth and branching (Wiese and Faissner, 2015). One such factor is anosmin, a secreted protein encoded by the KAL-1 gene that was found to be mutated in a form of Kallman syndrome in humans (de Castro et al., 2014). This syndrome is associated with anosmia and the absence of the olfactory tract in the brain. Interestingly, the homolog of this gene was first shown in C. elegans to regulate axon branching, since its loss during development results in the formation of an extra branch in a single neuron (Rugarli et al., 2002). The role in branching was also observed in rodent olfactory neurons (SoussiYanicostas et al., 2002) and cerebellum Purkinje cells (Gianola et al., 2009). Anosmin associates with the ECM and binds integrin on the cell surface to regulate adhesion. A study of branch regulation in C. elegans further showed that anosmin serves as an autocrine cofactor for FGF and signals through a receptor complex (Diaz-Balzac et al., 2015). It is worth pointing out that glycoproteins and proteoglycans can be associated with both the ECM and the neighboring cell surface. They often provide an anchorage site or act as cofactors for other extracellular molecules to control axon growth and branching (Beller and Snow, 2014; Poulain and Yost, 2015). Deletion of these proteins or modifying enzymes that control the constellation of posttranslational glycosylation events for these proteins can lead to impaired axon growth and guidance defects that are similar to those seen in the absence of extracellular cues. One example is the discovery of dystroglycan in a mouse forward genetic screen based on sensory axon growth phenotypes and the observation of a direct interaction between dystroglycan and Slits in regulating axon guidance owing to guidance cue sequestration (Wright et al., 2012). The interaction between axons and neighboring cells is mediated by several families of CAMs. One is the cadherin superfamily of transmembrane proteins, which contain multiple cadherin repeats in their extracellular domains and require calcium for homophilic interactions (Takeichi, 2007). This superfamily can be divided into three subfamilies: classic, flamingo/CELSRs, and protocadherins (Pcdhs). Members of these families have been shown to regulate axon growth and

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branching in different species. N-cadherin is a classic cadherin that regulates terminal arborizations of olfactory axons in Drosophila (Zhu and Luo, 2004), axonal extension of RGCs in Xenopus (Riehl et al., 1996), and motor axon growth and branching in the zebrafish (Bruses, 2011). Flamingo is a nonclassical cadherin that was found in Drosophila to regulate dendritic branching (Gao et al., 2000), sensory axon growth (Steinel and Whitington, 2009), and axon arborization in the visual system (Chen and Clandinin, 2008; Lee et al., 2003; Senti et al., 2003). The mammalian Flamingo homologs CELSR1-3 regulate neurite growth and guidance both in vitro and in vivo (Feng et al., 2016; Shima et al., 2007; Tissir et al., 2005; Zhou et al., 2008). Pcdhs are newly characterized members of the cadherin superfamily. They can be divided into two groups: nonclustered Pcdhs that are scattered throughout the genome and clustered Pcdhs that are organized into three gene clusters containing large variable exons in tandem (Morishita and Yagi, 2007). Clustered Pcdhs use alternative promoters to yield a large number of protein isoforms with various combinations of extracellular domain sequences (Morishita and Yagi, 2007). Early genetic studies of a- and d-Pcdh clusters demonstrated their involvement in the elongation of serotonergic projections and striatal axons in the mouse brain (Katori et al., 2009; Uemura et al., 2007). However, their main functions appear to be patterning axon and dendritic branches to allow self/noneself-recognition and contact-dependent repulsion based on expression of selective isoforms (Lefebvre, 2017) (see later). Interestingly, deletion of all three Pcdh clusters in mouse olfactory neurons leads to loss of axonal branches and collapse of axons (Mountoufaris et al., 2017). These functions are also seen with Pcdh-18b, a nonclustered Pcdh that controls motor axon growth and arborization in the zebrafish (Biswas et al., 2014). In addition to the cadherin superfamily, CAMs of the Ig-SF have been implicated in regulating axon growth and branching (Kamiguchi and Lemmon, 2000). These molecules are characterized by their large extracellular sequence containing multiple Ig domains that can mediate homophilic or heterophilic interactions (Maness and Schachner, 2007). Interestingly, these CAMs are capable of binding in cis to other membrane proteins such as integrins, neuropilins, and even other Ig CAMs, thus serving as coreceptors in axon growth and guidance regulation (Schmid and Maness, 2008). Some of the best studied Ig-CAMs are L1-CAM and neural CAM (NCAM), which have multiple isoforms/homologs and play important roles in neural development in different species (Brennaman and Maness, 2008; Kamiguchi et al., 1998; Siegenthaler et al., 2015). The importance of Ig-CAMs in controlling axon development is illustrated by a unique role of Dscam, a protein found in the study of Down syndrome (Hattori et al., 2008). Studies of Drosophila Dscam1 reveal tens of thousands of protein isoforms with variable Ig domains that are generated through alternative splicing (Schmucker et al., 2000). Like clustered Pcdhs, this molecular diversity, in conjunction with exquisite isoform specificity for homophilic binding, provides a robust molecular mechanism for self/noneself-recognition that is critical for axon and dendrite patterning (Hattori et al., 2008; Zipursky and Grueber, 2013). This mechanism allows the same class of axons or dendrites to avoid each other but at the same time achieve maximum coverage of target or receptor fields (Chen et al., 2006; Matthews et al., 2007; Millard et al., 2007). Recently, it was found that the diversity of Dscam1 homophilic interactions also contributes to the morphology of axon collateral branches (He et al., 2014). Interestingly, the mammalian Dscam genes do not generate diverse alternative isoforms. Although the self/noneselfrecognition function is likely mediated by other molecules, such as Pcdhs, homophilic interactions of the mammalian Dscam are important for many related processes, including arborization and laminar termination of RGC dendrites, and the spacing of amacrine cell bodies in the retina (Fuerst et al., 2008, 2009; Yamagata and Sanes, 2008). Dscam has also been shown to bind guidance molecules (Alavi et al., 2016; Andrews et al., 2008; Liu et al., 2009; Ly et al., 2008), although the in vivo role of this association is not fully understood (Palmesino et al., 2012). Furthermore, recent studies showed that Dscam can promote fasciculation and growth of mouse RGC axons (Bruce et al., 2017) and growth of retinal bipolar cell processes (Simmons et al., 2017), demonstrating diverse functions of the mammalian Dscam.

3.3.4 Glial cells and myelination The extracellular environment through which neuronal processes navigate changes dramatically later in development. Synapses are formed between specific axons and dendrites when glial cells start to differentiate and participate in sculpturing synaptic connections. While glial cells have many functions (Allen and Lyons, 2018), their roles in axon growth and branching during development have just begun to be appreciated (Rapti et al., 2017). However, one notable role that has been studied extensively in the mature CNS is myelination, which occurs during postnatal periods and increases the conduction velocity of the adult nervous system. CNS myelin has long been known to be inhibitory to axon growth (Schwab and Thoenen, 1985). Extensive studies in the context of axon injury and regeneration have identified a number of myelin-associated inhibitors, including myelin-associated glycoprotein (MAG), Nogo,

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and oligodendrocyte-myelin glycoprotein (OMgp) (Geoffroy and Zheng, 2014; Yiu and He, 2006) (see Section 1.9). Interestingly, these three molecules bind to a common receptor, the Nogo receptor (NgR), which also interacts with p75 (Wang et al., 2002), as well as the paired immunoglobulin-like receptor B (PirB) protein (Atwal et al., 2008). However, targeting these factors or their receptors in mice failed to significantly promote regeneration of injured axons, suggesting that additional molecules are present to mediate the inhibitory effects of myelin (Geoffroy and Zheng, 2014; Kim et al., 2004; Zheng et al., 2005). The function of these molecules in normal development has begun to be elucidated (Baldwin and Giger, 2015). MAG is required for the initiation of myelin formation and subsequent maintenance of myelination in uninjured animals (Schachner and Bartsch, 2000). More relevant to this section, these molecules have been shown to regulate axon growth and branching. For example, deletion of MAG in mice has subtle myelination defects but increased axon atrophy in aging mice (Baldwin and Giger, 2015); Nogo is involved not only in glial cell development and myelination (Chong et al., 2012) but also in axonal branch development of olfactory and sensory neurons (Eckharter et al., 2015; Iketani et al., 2016); and OMgp is present at the nodes of Ranvier and may function to prevent axon collateral formation (Huang et al., 2005). Furthermore, studies of mice lacking the NgR and PirB receptors have revealed interesting roles for these molecules in the formation of ocular dominance columns (Atwal et al., 2008), a process involving the remodeling of axonal branches (see later) (Antonini and Stryker, 1993). Therefore, additional functions of these myelin-associated factors, especially in dynamic regulation of axon growth and branching, await future investigation (Chen and Zheng, 2014; Kondiles and Horner, 2018).

3.3.5 Neural activity Neural activitydboth spontaneous and evokeddare critical for axon growth and branching in the formation and maturation of neural circuits at different developmental stages (Holtmaat and Svoboda, 2009; Spitzer, 2006). An excellent example is provided by axonal projections in the vertebrate visual system, where both types of neural activity play a crucial role during the critical period of retinal circuit development (Fig. 3.2). First, topographic map formation (see Section 1.6) requires complex control of axon growth and branching, which is in part regulated by spontaneous neural activity. Mice in which spontaneous activity has been genetically disrupted exhibit a disturbance of the normal pattern of axonal projections from the lateral geniculate nucleus (LGN; an RGC axonal target) to the cortex and also precise retinotopic mapping (Cang et al., 2005). In the superior colliculus (another RGC axonal target), which contains two visual maps, the corticocollicular projection is refined over time so that it aligns with each of the duplicated maps via bifurcation of its projections. However, when the spontaneous retinal activity is genetically disrupted in mice, the bifurcation of the corticocollicular projection is lost, resulting in a loss of map alignment (Triplett et al., 2009). Second, evoked neural activity plays a critical role in axonal competition. An early study of retinal projections in cats experiencing a loss of visual input by monocular deprivation demonstrated that terminal arbors from the deprived eye were withdrawn from the target field, suggesting that visual activity stabilizes or maintains axonal branches (Antonini and Stryker, 1993). Later, it was demonstrated that correlated RGC activity plays a critical role in regulating both the elimination of inappropriate terminal arbors and the selective stabilization of axonal branches (Ruthazer et al., 2003). This was further supported by studies demonstrating that new axonal branches are preferentially initiated at nascent synapses, whereas mature synapses stabilize branches: two effects enhanced by visual stimulation (Meyer and Smith, 2006; Ruthazer et al., 2006). Studies of retinal axons have shown that reducing neural activity leads to decreased axon terminal growth and reduced formation of new terminal arbors, a deficiency that is relieved when the activity of neighboring RGCs is also suppressed (Hua et al., 2005). This intriguing finding suggests a model in which activity-based competition for limited target space between neighboring axon terminals results in the mature terminal morphology found in the adult animal. It is clear from these studies in the visual systems that neural activity of various kinds is required for normal development of axon branching patterns. A recent study of callosal axons that link the two hemispheres in the sensory cortex further demonstrates that evoked activity from sensory inputs and spontaneous cortical activity are both important for appropriate axon growth and the formation of correct synaptic connections (Suarez et al., 2014). The direct role of electrical activity on axon growth regulation has also been demonstrated in cultured neurons. First, normal RGC axon growth requires BDNF, but RGCs do not extend axons very far (Goldberg et al., 2002). However, this basal axon growth can be significantly enhanced if the RGCs are subjected to levels of electrical stimulation that evoke levels of spontaneous activity equivalent to those experienced during normal development. Similarly, in cultured sympathetic neurons, electrically stimulated axon collaterals exhibit a growth advantage over unstimulated axons of the same neuron, or other neurons (Singh and Miller, 2005). Finally, studies using organotypic cocultures suggest that electrical activity regulates branching of thalamocortical projections (Uesaka et al., 2005, 2007).

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What mediates this unique regulation of axon growth and branching by neural activity? The immediate effect of neural activity is increased calcium signaling, which can alter axon growth and branching (discussed later; see Section 3.4.2). In addition, mechanisms mediated by the extracellular cues discussed above could be in play. For example, neurotrophins have been shown to mediate synaptic plasticity (McAllister et al., 1999; Poo, 2001). Studies of cultured sympathetic neurons suggest an intriguing employment of such a mechanism, since electrical activity results in increased expression and secretion of BDNF from “winning” axons, which then bind to p75 receptors on “losing” axons, resulting in their being pruned (Singh and Miller, 2005). Consistent with these observations, mice bearing a mutant version of the BDNF gene that is insensitive to electrical activityestimulated expression, or mice lacking the p75 receptor, do not exhibit developmental pruning, resulting in overlapping innervation of adjacent target compartments (Singh et al., 2008). More recently, studies of thalamocortical neurons have shown that Netrin-4, a membrane-associated Netrin family member (Hayano et al., 2014), and HDAC9, a protein involved in transcription regulation (Alchini et al., 2017), control axon branching in an activitydependent fashion. Thus, transcriptional regulation of the extracellular cues could mediate activity-dependent regulation of axon growth and branching as well.

3.3.6 Additional axon branching molecules In addition to the extracellular cues discussed earlier, evidence has emerged that implicates factors previously known for their functions in vascular physiology in regulating axon growth and branching. One example is endothelin, which is secreted by muscle tissues and regulates the growth and guidance of sympathetic axons during early development (Makita et al., 2008). The other is a member of the natriuretic peptide hormone family, which controls axon DRG sensory neurons branching. Normally, the central DRG neuron projection enters the dorsal spinal cord and bifurcates to form two axonal branches, each of which then forms collateral branches. Loss of the C-type natriuretic peptide (CNP), or its receptor, results in bifurcation failure, although the subsequent formation of collateral branches from the remaining axon occurs normally (Schmidt et al., 2009; Zhao and Ma, 2009). Loss of the CNP receptor Npr2 disrupts axon bifurcation in DRG neurons (Schmidt et al., 2007; Zhao et al., 2009) and also in TG neurons, which produces behavioral deficits (Ter-Avetisyan et al., 2014, 2018). Additionally, CNP is capable of promoting axon branching in dissociated DRG neurons and stimulates axon outgrowth from DRG explants (Zhao and Ma, 2009). These findings suggest that a variety of factors can serve as extracellular cues to regulate axonal development and likely more remain to be discovered.

3.4 Intracellular signaling mechanisms that mediate axon growth and branching 3.4.1 Rho family small GTPases: linking receptors to the cytoskeleton Members of the Rho family of small GTPases, consisting of Rac, Rho, and Cdc42, act as molecular switches to link cell surface receptors to downstream signaling pathways involved in cytoskeletal regulation (Hall and Lalli, 2010). They are activated by guanine nucleotide exchange factors (GEFs) that catalyze the exchange of guanosine diphosphate (GDP) for guanosine triphosphate (GTP) and link cell surface signals to specific cellular responses. The active GTP form binds to specific downstream effector proteins that are involved in cytoskeleton reorganization. This activity is terminated by GTPases-activating proteins (GAPs), which increase the intrinsic GTPase activity of these proteins. The functional role of Rho family GTPases in axonal development has been established through the studies of GTPases as well as their specific GEFs and GAPs in cultured neurons and in intact animals (Hall and Lalli, 2010; Luo, 2000; Spillane and Gallo, 2014). For example, in hippocampal cultured neurons, the Rac/Cdc42 effectors N-WASP and IQGAP3 can both promote axon extension (Banzai et al., 2000; Wang et al., 2007), whereas Rho and its effector ROCK negatively regulate axon outgrowth (Govek et al., 2005). In vivo evidence comes from early studies in Drosophila, in which constitutively active or dominant-negative mutants of Rac caused axon outgrowth defects (Luo et al., 1994), and in mice, in which a constitutively active Rac reduced axonogenesis as well as axon growth of cerebellar Purkinje cells (Luo et al., 1996). Further analysis of three Rac isoforms in Drosophila mushroom body neurons shows a sequential requirement for axon branching, then guidance, and finally axon growth (Ng et al., 2002). Interestingly, genetic interaction studies show that Rac appears to have opposite activities on axon growth: inhibition via the effector kinase PAK and promotion via a PAK-independent manner (Ng and Luo, 2004). These studies have thus revealed complex functions of Rho GTPases mediated by their effectors to link different extracellular cues to regulation of axon growth and branching (Fig. 3.2).

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Regardless of the signaling complexity, Rho family GTPases have conserved subcellular functions involving the assembly and disassembly of the actin and microtubule cytoskeletons (Hall and Lalli, 2010). Early studies in fibroblasts showed that Rac, Rho, and Cdc42 influence actin-mediated motile structures, including lamellipodia, stress fibers, and filopodia, respectively (Ridley and Hall, 1992; Ridley et al., 1992). These functions also contribute to different stages of axonal development. For example, Cdc42 is involved in filopodium formation in growth cones, whereas Rac is found in the actin patches along axons to induce Arp2/3-dependent actin assembly via the WAVE complex for branch formation (Spillane and Gallo, 2014). Rho usually activates myosin-based contraction through the ROCK kinase and thus contributes to retraction of axons and branches (Hall and Lalli, 2010). Rho family GTPases are known to regulate microtubule assembly (Zheng, 2004), but their role in microtubule stabilization during axon growth was only appreciated in several recent studies (Borgen et al., 2017; Gujar et al., 2018; Tivodar et al., 2015).

3.4.2 Calcium Calcium ions (Caþþ) are a common second messenger, mediating a wide variety of cellular functions that include cell motility (Zheng and Poo, 2007). Calcium also mediates spontaneous and evoked neural activities during development (Rosenberg and Spitzer, 2011; Yamamoto and Lopez-Bendito, 2012). Cells maintain a constant intracellular calcium concentration of about 100 nM, several orders of magnitude lower than found in the extracellular space. Upon stimulation, Caþþ concentration can increase dramatically by entering the cytoplasm from the extracellular space via ion channels in the plasma membrane or from internal stores via IP3-sensitive channels. Owing to the high levels of immobile calciumbinding proteins present in the cytoplasm, calcium ions do not diffuse very far and so their action is limited to local areas near the site of entry. The main influence of calcium on axon growth and branching is growth cone motility (Fig. 3.2). While an optimal level of calcium is critical, spontaneous fluctuations of calcium in the form of waves and spikes appear to be more important in regulating the rate of axonal elongation, as observed in Xenopus axons (Gomez and Spitzer, 2000). When present globally in the growth cone, calcium regulates overall growth motility and axonal extension, but asymmetric changes in calcium concentration could lead to changes in growth direction (Gomez and Zheng, 2006). Such changes can result in asymmetric clathrin-mediated endocytosis as well as an imbalance of endocytosis and exocytosis, as shown in two studies on this topic (Tojima et al., 2010, 2014). In addition, both spontaneous calcium transients and calcium waves elicited by extracellular stimuli correlate with branch formation in cultured cortical neurons (Hutchins and Kalil, 2008; Tang and Kalil, 2005). Intracellular calcium mediates Wnt5a- or NCAM2-induced axon branching (Hutchins et al., 2012; Sheng et al., 2015). Moreover, a number of downstream targets that bind to calcium have been implicated in mediating these activities by regulating cytoskeletal dynamics and cell adhesion locally in the growth cone (Zheng and Poo, 2007). Finally, the long-term changes elicited by calcium may be mediated by calcineurin and N-FAT nuclear complexes, which have been found to regulate axonal development (Graef et al., 2003). Thus, calcium plays important but complex roles in axon growth and branching by regulating cytoskeleton reorganization (Gasperini et al., 2017) along with other critical cellular functions described earlier.

3.4.3 Cyclic nucleotides as second messengers and modulators Cyclic nucleotides, including cyclic adenosine-30 ,50 -monophosphate (cAMP) and cyclic guanosine-30 ,50 -monophosphate (cGMP), are another group of second messengers important for cell signaling. Both are produced by specific cyclases that are regulated by upstream activators. In vitro, they modulate guidance responses to specific extracellular cues in the growth cones, with high levels favoring attraction and low levels for repulsion (Song and Poo, 1999). For axonal development, it has been shown that cAMP can promote axon regeneration of adult PNS neurons both in culture and in a rodent injury model (Neumann et al., 2002; Qiu et al., 2002). However, in embryonic PNS neurons, cAMP suppresses branching since loss of adenylate cyclase 1, an enzyme that synthesizes cAMP, increases peripheral branches (Haupt et al., 2010). Additional studies of other adenylate cyclases demonstrate this role in axon pruning (Nicol and Gaspar, 2014). Interestingly, cGMP promotes branch formation (Zhao et al., 2009) and regulates sensory afferent bifurcation in the mouse spinal cord (Schmidt et al., 2007; Zhao et al., 2009). In sensory axons, cGMP is produced by membrane-bound guanylate cyclase (Schmidt et al., 2007, 2009; Zhao and Ma, 2009). However, nitric oxide that activates the soluble guanylate cyclase can also modulate axonal arbors of RGC in Xenopus (Cogen and Cohen-Cory, 2000), suggesting the presence of multiple ways of activating cGMP signaling. Studies of axon/dendrite formation in cultured hippocampal neurons suggest a reciprocal interaction that results in local amplification and long-range suppression of the two cyclic nucleotide signals (Shelly et al., 2010). Thus, these two small

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molecules may interact with each other and provide a unique ying-yang signaling mechanism to modulate axon growth and branching during development (Fig. 3.2). Such interaction is likely mediated by the downstream cAMP- and cGMPdependent protein kinases, which can locally control cytoskeleton dynamics, as shown by recent studies in growth cone guidance (Akiyama et al., 2016). Although several cytoskeleton regulators, including GSK3 and CRMP2, have been implicated in the regulation of axon growth and branching (Shelly et al., 2011; Xia et al., 2013), additional molecular mechanisms involving these two classes of signaling molecules remain to be identified.

3.5 Concluding remarks Extensive work over the past three decades has provided a wealth of knowledge regarding molecular and cellular regulation of axon growth and branching. However, many questions remain to be addressed. First, the relationship between axon growth and branching at the cell biological level needs to be firmly established. What are the key sites for cytoskeletal regulation? How are cytoskeletal components, in particular microtubules, regulated to switch from axon growth to branch growth? How different is axonal branch elaboration from axon growth? How are different forms of axonal branches generated through coordinated regulation of axon growth, guidance, and branching? In addition to new imaging technology, quantitative analysis of axon growth and branching (Chalmers et al., 2016) should help us understand these key cell biological mechanisms. Second, although many extracellular factors that regulate axon growth and branching have been identified in different regions of the nervous system at different developmental stages, a common theme driving growth and branching regulation has yet to emerge. What confers cell-type specificity? What are the major differences between axons in the PNS and CNS? Similarly, what are the differences between embryonic and adult neurons that render them differentially regulated by common factors? What are the shared and unique mechanisms that influence axon and dendrite branching? Importantly, how do these factors that function during development affect axon and branch regeneration (Chisholm et al., 2016; Hollis, 2016)? Finally, a developing axon encounters a myriad of extracellular signals en route to its target. How does the axon integrate all of these signals and convey discrete responses to the cellular machinery that defines its morphology, controls rates of growth and branching, and determines correct branch locations? As each neuron undergoes considerable intrinsic changes during development (Goldberg, 2004), how do these changes, via transcriptional regulation, influence responses to the extracellular environment? And how do glial cells influence the development and regeneration of axons and their branches? Answering these questions will further advance our understanding of these important processes in circuit development. This will, in turn, help us to better understand the molecular and cellular basis of many developmental disorders and to find new ways to promote nerve regeneration following injury or degeneration.

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Chapter 4

Axon guidance: Netrins Marc Tessier-Lavigne Stanford University, Stanford, CA, United States

Chapter outline 4.1. Introduction 4.2. Netrins and their receptors 4.2.1. Netrin discovery and structure 4.2.2. Netrin receptors 4.2.3. Interactions with other signaling systems 4.2.4. Netrin functional domains and interactions with receptors 4.3. Netrin function in axon guidance and cell migration 4.3.1. Mammalian spinal cord 4.3.1.1. Guidance by midline-derived Netrin-1 in the spinal cord 4.3.1.2. Guidance by ventricular zoneederived Netrin-1 in the spinal cord 4.3.1.3. Synergy between Netrin-1 from floor plate and from ventricular zone in the spinal cord 4.3.1.4. Interpreting the guidance defects caused by loss of Netrin-1 in the spinal cord 4.3.2. Mammalian hindbrain

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4.3.2.1. In hindbrain, Netrin-1 from ventricular zone is more important than from floor plate 96 4.3.2.2. Control of neuronal cell migration by Netrin-1 in the hindbrain 96 4.3.3. Guidance of other classes of mammalian axons and cells: attraction, repulsion, and modulation 96 4.3.4. Invertebrate systems 97 4.3.4.1. Attraction and repulsion by UNC-6 in Caenorhabditis Elegans 97 4.3.4.2. Attraction and repulsion by Netrins in Drosophila 98 4.4. Beyond axon and cell guidance: additional roles for Netrins in the nervous system 99 4.5. Involvement of Netrin signaling in disorders of the nervous system 99 4.6. Netrins: players outside the nervous system 100 4.7. Conclusion 101 References 102

4.1 Introduction A fundamental feature of the organization of nervous systems of all bilateral animals, from roundworms to fruit flies to vertebrates, including humans, is the presence of numerous axonal projections across the midline of the animal that serve to connect and coordinate the two sides of the body. Sets of axons crossing the midline are termed “commissures,” and the very diverse types of neurons that send their axons to and across the midline through these commissures are, as a group, termed “commissural neurons.” Defects in guidance of commissural axons across the midline result in impaired coordination of the two body sides, and, as we shall see, in humans they underlie several inherited neurological disorders. Prominent among the guidance cues that steer axons to the midline are Netrins, a small family of cues that are conserved across species not just in structure but also in their function of attracting axons to the midline. They are also multifunctional, guiding axons through not just attraction but also repulsion, and regulating other aspects of development and function not just in the nervous system but also in nonneural systems. This chapter focuses on the role of Netrins in axon guidance, particularly at the midline, but we also discuss briefly their role in guidance beyond the midline and in other developmental, physiological, and pathological processes.

Cellular Migration and Formation of Axons and Dendrites. https://doi.org/10.1016/B978-0-12-814407-7.00004-3 Copyright © 2020 Elsevier Inc. All rights reserved.

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4.2 Netrins and their receptors 4.2.1 Netrin discovery and structure The Netrins provide a wonderful example of convergence in scientific discovery, as they were identified independently in the roundworm Caenorhabditis elegans and in vertebrates as cues that steer axons to the midline in these very divergent organisms. Embryological studies had demonstrated that commissural axons in the spinal cord of rat embryos can be attracted in vitro by a factor(s) secreted by midline cells (Tessier-Lavigne et al., 1988), whereas genetic screens in C. elegans identified a group of genes required for axons to project toward or away from the midline (Hedgecock et al., 1990). Cloning of the affected genes in C. elegans identified one that codes for a secreted protein termed UNC-6 that is required for midline guidance (Ishii et al., 1992), whereas biochemical purification from chick brain extracts led to the identification of two related proteins, Netrin-1 and Netrin-2, which can mimic the in vitro guidance activity of rat midline cells (Serafini et al., 1994; Kennedy et al., 1994). Remarkably, UNC-6 and the vertebrate Netrins were found to be species homologs, which implied that mechanisms for midline guidance are conserved across the 600 million years of evolution that separates nematodes from vertebrates. At the time of this discovery, this came as a surprise, as it was widely believed that the mechanisms involved in wiring the nervous system would be more complex and sophisticated for higher organisms (Easter et al., 1985; Goodman, 1994). Instead, these studies underscored that more complex nervous systems use the same building blocks as simpler nervous systems, just deploying them in more intricate ways. Netrins were identified soon thereafter in Drosophila (Mitchell et al., 1996; Harris et al., 1996), and indeed in all other animal species where they have been sought, from planaria (Cebria et al., 2005) to humans (Meyerhardt et al., 1999). There are several subfamilies of Netrins in vertebrates. In this chapter, we focus exclusively on the subfamily implicated in axon guidance, which includes UNC-6 in C. elegans; Netrin-1 and Netrin-2 in chick; Netrin-1 and Netrin-3 in mammals (Netrin-2 is specific to birds and fish, whereas Netrin-3 is specific to mammals (Serafini et al., 1996; Wang et al., 1999)); and Netrin-A and Netrin-B in Drosophila. These Netrins are all secreted glycoproteins of w75e80 kDa and possess three structural regions (Fig. 4.1A): a globular amino-terminal region, termed domain VI; three epidermal growth factor (EGF)elike repeats in a rigid rodlike middle region, termed domain V; and a C-terminal region, the NTR or C domain, that is highly basic and contains six conserved cysteine residues that are likely to be paired in internal disulfide

(A)

Mammals:

(B)

Netrin-1, 3

(C) DCC/neogenin

C. elegans:

UNC-6

UNC-40

Drosophila:

Netrin-A, B

Frazzled UNC-5 family

Globular

VI

EGF-like

V

NTR

C

Ca++

Signaling moiety

FIGURE 4.1 Structure of Netrins and Netrin receptors. (A) Netrin-1 and Netrin-3 in mammals, UNC-6 in Caenorhabditis elegans, and Netrin-A and Netrin-B in Drosophila have similar structural domains (green diagram): a globular N terminal domain (domain VI); a middle domain (V) that possesses three EGF-like repeats (LEs1-3); and a C terminal domain (C or NTR). Domains VI and V are the major signaling domains. Right: crystal structure of domains VI and V of chick Netrin-1 at 2.8Å, showing the globular nature of domain VI and the rigid rodlike structure of domain V (Xu et al., 2014). (B) Structure of Netrin-1, Netrin-2, and Netrin-3 (Netrin-2 is found in birds), showing the homology of Netrin domains VI and V to similar domains in the g chain of Laminins. Also shown are Netrin-4, Netrin-G1, and Netrin-G2, which show greater homology to the b chain of Laminins. (C) Netrin receptors. Left, members of the DCC family (DCC and neogenin in mammals, UNC-40 in C. elegans and Frazzled in Drosophila), which in their extracellular domain possess four immunoglobulin (Ig) domains and six Fibronectin type III (FNIII) domains, and in their intracellular domain three evolutionarily conserved domains (P1-3). Right, members of the UNC-5 family (UNC-5 in C. elegans and Drosophila, and UNC5A-D in mammals) have two Ig and two Thrombospondin type I (TSP) domains extracellularly, and three conserved domains (ZU5, DB, and DD [Death Domain]) intracellularly. Adapted from Sun, K.L.W., Correia, J.P., Kennedy, T.E., 2011. Netrins: versatile extracellular cues with diverse functions. Development 138 (11), 2153e2169. https:// doi.org/10.1242/dev.044529.

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bonds. Domains VI and V derive their names from domains in the g-chain of laminins to which the Netrin domains are highly homologous (Fig. 4.1B). The NTR domain of Netrins is homologous to NTR domains in a variety of other proteins, including complement proteins C3, C4, and C5, secreted frizzled-related proteins, and tissue inhibitors of metalloproteinases. Mammals also possess other more divergent Netrins: Netrin-4, Netrin-G1, and Netrin-G2, which are more homologous to the b-chain of laminins than to the g-chain (Nakashiba et al., 2000, 2001; Yin et al., 2000) (Fig. 4.1B). In addition, whereas Netrin-4 is secreted, the G Netrins are anchored to cell membranes via glycosylphosphatidylinositol anchors. Mammals also possess Netrin-5, which lacks domain VI and is more related to Netrin-1 in its other domains (Yamagishi et al., 2015). These proteins have not so far been implicated in axon guidance and are not discussed further here.

4.2.2 Netrin receptors The screen in C. elegans for genes affecting midline guidance that led to the identification of the Netrin unc-6 also identified two additional genes, unc-40 and unc-5, which affect those migrations (Hedgecock et al., 1990), and which turned out to code for receptors that mediate attractive and repulsive actions of Netrins, respectively (Fig. 4.1C). Attractive Netrin receptors, including UNC-40, are members of the conserved Deleted in Colorectal Cancer (DCC) family of transmembrane receptors, which has two members in mammals (DCC and Neogenin), one in C. elegans (UNC40) and one in Drosophila (Frazzled). These proteins possess four amino-terminal immunoglobulin (Ig) domains, followed by six fibronectin type III (FNIII) repeats, a transmembrane domain, and a long cytoplasmic domain with several motifs that are conserved across evolution (Fig. 4.1C). These receptors all behave functionally as attractive Netrin receptors (Chan et al., 1996; Keino-Masu et al., 1996; Kolodziej et al., 1996; Fazeli et al., 1997; Xu et al., 2014), and mammalian DCC and Neogenin have been shown directly to bind Netrin-1 with high affinity (Keino-Masu et al., 1996). Mammalian DCC and Neogenin each have two isoforms, short and long, arising from alternative splicing that affects the length of the linker between FNIII repeats 4 and 5, which in turn constrains their interaction with Netrin-1 (Xu et al., 2014); for DCC at least, the longer isoform appears to be required for its function as an attractive receptor (Leggere et al., 2016). Repulsive Netrin receptors are members of a conserved family of transmembrane proteins defined by C. elegans UNC5 (Leung-Hagesteijn et al., 1992), which includes four members in mammals (UNC5A-D) (previously known as UNC5H1eH4; Ackerman et al., 1997; Leonardo et al., 1997; Englekamp, 2002) and one in Drosophila, UNC-5 (Keleman and Dickson, 2001). These proteins possess two amino-terminal immunoglobulin (Ig) domains, two thrombospondin type I (TSP1) repeats, a transmembrane domain, and a cytoplasmic domain with several conserved motifs (Fig. 4.1C). Again, these proteins behave functionally as repulsive Netrin receptors (Leung-Hagesteijn et al., 1992; Hamelin et al., 1993; Hong et al., 1999; Keleman and Dickson, 2001), and those members that have been tested, i.e., mammalian UNC5A and UNC5B, bind Netrin-1 with high affinity (Leonardo et al., 1997). What specifies attraction versus repulsion? Studies using chimeric receptors, in which cytoplasmic domains of DCC or UNC5 family receptors are combined with heterologous ectodomains, show that the cytoplasmic domains carry the information needed to mediate attraction or repulsion (Bashaw and Goodman, 1999; Hong et al., 1999; Stein et al., 2001; Brankatschk and Dickson, 2006).

4.2.3 Interactions with other signaling systems Although DCC family and UNC-5 family proteins are the central receptors for attraction and repulsion by Netrins, their function is modulated in important ways by a dizzying array of interactions with other signaling systems. First, although UNC-5 family proteins are sufficient to mediate Netrin repulsion, their activity is potentiated by interactions with DCC/UNC-40 family receptors, an effect that may be important for mediating repulsion of axons exposed to low Netrin concentrations at a distance from a Netrin source (Hedgecock et al., 1990; Hong et al., 1999; Keleman and Dickson, 2001; Brankatschk and Dickson, 2006). In C. elegans, this UNC-5þUNC-40 signaling is promoted at the expense of UNC-5-only signaling by the binding to UNC-5 of the TGFb family member UNC-129, produced dorsally, which thereby potentiates the repulsion that guides dorsal migrations away from the UNC-6/Netrin source (Colavita et al., 1998; MacNeil et al., 2009). In addition, in mammals, UNC5 family members bind and can mediate repulsive responses not only to Netrins but also to another important family of repellents that are members of the FLRT (fibronectin and leucine-rich transmembrane protein) family (Karaulanov et al., 2009; Sollner and Wright, 2009; Yamagishi et al., 2011; Visser et al., 2015). And in blood vessels, UNC5B can bind and be activated by transmembrane Robo4, a divergent member of the Roundabout (Robo) family (discussed in Chapter 7), functioning as a ligand expressed on other endothelial cells (Koch et al., 2011).

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DCC family members similarly show interactions with other receptors and other ligands. First, although DCC family proteins are sufficient to mediate Netrin attraction, several interactions modulate this activity. In mammals, transmembrane Robo3, another divergent member of the Robo family, is expressed by commissural axons (Sabatier et al., 2004; Marillat et al., 2004) (see Fig. 4.2B), and, although it does not bind Netrin-1, functions as a coreceptor with DCC to potentiate Netrin/DCC attraction of these axons in vitro (Zelina et al., 2014). It is interesting to speculate that this potentiation facilitates long-range attraction by Netrin-1 in vivo, much as DCC proteins potentiate UNC5 proteins for long-range repulsion. This possibility is supported by the fact that, in mammals, Robo3 expression is largely confined to commissural axons that cross the midline at the floor plate (Sabatier et al, 2004; Marillat et al., 2004; Friocourt et al., 2019), which can be attracted at long range by Netrin1 (Kennedy et al., 1994; Shirasaki et al., 1996; Moreno-Bravo et al., 2019; Wu et al., 2019), whereas retinal ganglion cell axons, which are guided only at short range by Netrin/DCC attraction (Deiner et al., 1997), do not express Robo3 (Friocourt et al., 2019). Commissural axon guidance defects in Robo3 knockout mice are consistent with a role in potentiating Netrin/DCC attraction (Sabatier et al., 2004; Jaworski et al., 2015). However, Robo3 in mammals has additional activities, including silencing Slit/Robo1/2 repulsion (Sabatier et al., 2004) and mediating NELL2 repulsion (Jaworski et al., 2015); further analysis is required to dissect the relative contributions of these activities to guidance. Mechanisms also exist to counteract Netrin/DCC attraction. In Xenopus, attractive Netrin/DCC signaling can be inhibited by Slit/Robo signaling (Stein and Tessier-Lavigne, 2001), an effect that might contribute to a known reduction in responsiveness of axons to Netrin-1 when they reach the midline (Shirasaki et al., 1998; Zou et al., 2000). Another mechanism that can attenuate this signaling in mammals is cleavage of DCC by surface metalloproteases, leading to DCC ectodomain shedding (Galko and Tessier-Lavigne, 2000). In addition, the secreted protein Draxin can bind DCC and affect its activity (Ahmed et al., 2011; Liu et al., 2018), and the C1q-tumor necrosis factor family member Cerebellin-4 can compete with Netrin-1 for binding to DCC (Haddick et al., 2014). In C. elegans, attractive UNC-6/UNC-40 signaling is dampened by the receptor protein tyrosine phosphatase CLR-1 (Chang et al., 2004). And in Xenopus, not only can Netrin/ DCC attraction be attenuated, but in some growth cones it can even be converted to actual repulsion by modulation of second messenger pathways (Ming et al., 1997; Nishiyama et al., 2003) and by an extracellular cue (Laminin-1)dan effect that may contribute to axons leaving a region of coexpression of Laminin and Netrin to enter a Netrin-only region (Höpker et al., 1999). Second, DCC family proteins can also mediate guidance effects by non-Netrin ligands. For example, in addition to functioning as a Netrin receptor, in mammals the DCC family receptor Neogenin also binds and mediates repulsion by the ligand repulsive guidance molecule (Rajagopalan et al., 2004; Matsunaga et al., 2006). In C. elegans, UNC-40/DCC functions to mediate attractive signaling by the ligand MADD-4 (a homolog of vertebrate ADAMTS-like proteins) (Seetharaman et al., 2011); this does not appear to reflect direct binding and instead involves binding of MADD-4 to EVA1, a conserved transmembrane protein with predicted galactose-binding lectin domains, which recruits UNC-40 into a ternary complex (Chan et al., 2014). And, perhaps most intriguingly, in Drosophila, when axons reach the midline, Frazzled/DCC functions in a Netrin-independent fashion to trigger transcription of the key midline regulator Commissureless (Yang et al., 2009), an effect that involves release of the Frazzled cytoplasmic domain and its action as a transcription factor (Neuhaus-Follini and Bashaw, 2015); how this unusual Frazzled function is activated on midline contact is unknown but could involve a non-Netrin ligand. A few major themes emerge from these findings. First, UNC5 family receptors appear specialized for repulsion, and at least two distinct ligand families in addition to Netrins can tap into this repulsive system. Second, while DCC family receptors are major mediators of attraction, they appear more promiscuous, being capable of potentiating repulsive UNC5 signaling, of being coopted to mediate attraction by other ligands, and of directly mediating repulsive signaling as well. Third, multiple mechanisms exist to modulate Netrin/DCC and Netrin/UNC5 signaling, providing opportunities to fine-tune the function of these ligand/receptor signaling systems for particular guidance tasks, including determining whether UNC5 proteinsdand possibly DCC toodmediate only short-range or also long-range guidance effects. Are there Netrin receptors beyond the DCC and UNC5 families? Netrins have been found to interact with a number of other transmembrane proteins, including the adenosine receptor 2B (A2B), the immunoglobulin gene superfamily member down syndrome cell adhesion molecule (DSCAM), the amyloid precursor protein (APP), and integrins (the latter via the Netrin NTR domain), though potential in vivo guidance roles mediated by these interactions remain incompletely characterized (Corset et al., 2000; Stein et al., 2001; Yebra et al., 2003; Andrews et al., 2008; Ly et al., 2008; Palmesino et al., 2012; Rama et al., 2012).

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FIGURE 4.2 Effects of the floor plate and Netrin-1 on commissural axons in the spinal cord. (A) Diagram of the embryonic rodent spinal cord at midgestation. Commissural neurons (green), with cell bodies in the dorsal spinal cord, send axons (green dotted line) ventrally along the pial edge and then redirect their growth to the midline, skirting the developing motor column (MC). The axons cross the midline when they reach the floor plate (FP) and then leave the midline to project to their eventual targets (not shown). DRG, dorsal root ganglia flanking the spinal cord. (B) Micrographs of a transverse section of the E10.5 mouse spinal cord, showing commissural axons in green (immunolabeling for Robo3) and radial ventricular zone neural progenitor cells in red (immunolabeling for Nestin). (C) Expression of Netrin-1 mRNA in a section from a comparably staged mouse embryo, visualized by in situ hybridization. Netrin-1 transcripts are highly expressed not only in floor plate (FP-Netrin-1) but also at a lower level in the ventral two-thirds of the spinal cord (VZ-Netrin1). Arrow at top shows the dorsal-most third of the spinal cord, where Netrin-1 is not expressed. (D) Floor plate and Netrin-1 promote commissural (C) axon outgrowth from explants. Coculture of explants of embryonic rat dorsal spinal cord (D) with microdissected floor plate (FP, left panel), an aggregate of COS cells secreting Netrin-1 (middle), or an aggregate of control COS cells (right), shows that floor plate and cells secreting Netrin-1, but not control cells, stimulate outgrowth of commissural axons. Cultures performed for 40 h in three-dimensional collagen gels. (E) Floor plate and Netrin-1 cause turning of commissural axons within explants. Explants of dorsal spinal cord were cultured in collagen gels for 40 h, and then commissural axons within the explants were labeled (red) with an antibody to a specific marker (TAG-1). Axons are viewed from the side. After 40 h in control cultures (bottom), axons have grown from a dorsal position (D) along a vertical (ventral) trajectory and are not affected by control COS cells placed to their left. However, when cultured with either microdissected floor plate (FP, top) or COS cells secreting Netrin-1 (middle), the axons reorient their growth toward the source of Netrin-1, showing that Netrin-1 has turning activity. (F) Purified Netrin-1 can attract individual growth cones. An isolated embryonic Xenopus spinal axon grown on a two-dimensional substrate was exposed to a point source of Netrin-1 (pipette in upper right corner). Top, start of experiment (0 h); bottom, 1 h later, the growth cone has turned toward the source. (A) Reproduced from Wu, Z., Makihara, S., Yam, P.T., Teo, S., Renier, N., Balekoglu, N., Moreno-Bravo, J.A., et al., 2019. Long-range guidance of spinal commissural axons by Netrin1 and sonic Hedgehog from midline floor plate cells. Neuron 101 (4), 635e647.e4. https://doi.org/10.1016/j.neuron.2018.12.025; (B) Micrograph courtesy of Dr. Z. Wu; (C) Reproduced from Serafini, T., Colamarino, S.A., Leonardo, E.D., Wang, H., Beddington, R., Skarnes, W.C., Tessier-Lavigne, M., 1996. Netrin-1 is required for commissural axon guidance in the developing vertebrate nervous system. Cell 87 (6), 1001e1014. https://doi.org/10.1016/s0092-8674(00)81795-x;

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4.2.4 Netrin functional domains and interactions with receptors Domains VI and V together mediate the major signaling functions of Netrins in attraction and repulsion (Keino-Masu et al., 1996; Lim et al., 1999, 2002; Moore et al., 2012) (Fig. 4.1A). The highly basic NTR domain mediates tight binding of Netrins to negatively charged residues on cell surfaces (such as heparan sulfate proteoglycans) and in the extracellular matrix (ECM); domains VI and V also bind surfaces and the ECM, but to a lesser extent (Serafini et al., 1994; Kennedy et al., 1994; Moore et al., 2012). A consequence of this binding is that, although Netrins are diffusible proteins, their range of diffusion in any particular setting is dependent on the extent to which they are bound by extracellular factors in their environment (Deiner et al., 1997; Moore et al., 2012). Interaction with surfaces and the ECM also serves to aggregate Netrins and hence Netrin receptors, which appears to be important for receptor activation (Keino-Masu et al., 1996; Stein et al., 2001). In fact, Netrin/DCC function in commissural neurons appears to require heparan sulfate expression by these neurons, presumably to mediate such aggregation (Matsumoto et al., 2007). In addition to this cell-binding role, the NTR domain may have a direct signaling role in suppressing axonal branching, at least in C. elegans (Lim et al., 1999). How Netrins interact with DCC and UNC5 family receptors is an area of intense study but remains incompletely understood. Netrin-1 interacts with the FNIII repeats of DCC and with the Ig domains of UNC5B (Geisbrecht et al., 2003). Structures of domains VI and V of Netrin-1 bound either to FNIII domains 4 and 5 of DCC or Neogenin or to FNIII domains 5 and 6 of DCC have been reported, which reveal three regions of Netrin-1 that can interact with these FNIII domains: domain VI, EGF repeat 2, and EGF repeat 3 (Finci et al., 2014; Xu et al., 2014). Biochemical analysis indicates that the Netrin-1 EGF repeat 2 is also the region of Netrin-1 that interacts with the two Ig domains of UNC5B (Grandin et al., 2016). Models for the interaction of Netrin-1 with these receptors, and to explain how DCC attraction can be converted to repulsion by UNC5 receptors, have been proposed (Finci et al., 2014, 2015; Xu et al., 2014; Liu et al., 2018), but further work is required to fully elucidate these interactions.

4.3 Netrin function in axon guidance and cell migration Netrins play an evolutionarily conserved role in guiding axons and cells to the nervous system midline, which has been studied most extensively in mammals, in C. elegans, and in Drosophila. Here, we discuss axon guidance functions of Netrins in these three systems, with a major but not exclusive focus on midline guidance.

4.3.1 Mammalian spinal cord A major model for the analysis of axon guidance, including Netrin function, is the developing spinal cord of mice and rats. Spinal commissural neurons are born in the dorsal spinal cord and extend axons to a specialized set of cells at the ventral midline termed floor plate cells, following a stereotyped trajectory: the axons initially grow near and parallel to the pial edge and then break away from the edge when they reach the developing motor column and reorient toward the midline floor plate largely skirting the motor column (Fig. 4.2A and B). Analysis of this system has helped provide a detailed understanding of Netrin-1 functions and shows that there are two sources of Netrin-1 that contribute to guiding the axons (Fig. 4.2C): the floor plate cells, which function to attract the axons from a distance (i.e., over a few hundred micrometers) and help them reorient toward the midline, and ventricular zone (radial progenitor) cells, which function to guide them more locally.

4.3.1.1 Guidance by midline-derived Netrin-1 in the spinal cord Midline floor plate cells secrete two major diffusible chemoattractants, Netrin-1 and Sonic Hedgehog (Shh, which is discussed in greater detail in Chapter 8), which attract the axons at a distance. In vitro, floor plate cells both stimulate outgrowth of commissural axons from explants, a permissive effect (Tessier-Lavigne et al., 1988) (Fig. 4.2D), and cause

=

(D) Left panel from Tessier-Lavigne, M., Placzek, M., Lumsden, A.G.S., Dodd, J., Jessell, T.M., 1988. Chemotropic guidance of developing axons in the mammalian central nervous system. Nature 336 (6201), 775e778. https://doi.org/10.1038/336775a0, Right two panels from Kennedy, T.E., Serafini, T., de la Torre, J.R., Tessier-Lavigne, M., 1994. Netrins are diffusible chemotropic factors for commissural axons in the embryonic spinal cord. Cell 78 (3), 425e435. https://doi.org/10.1016/0092-8674(94)90421-9; (E) Reproduced from Kennedy, T.E., Serafini, T., de la Torre, J.R., Tessier-Lavigne, M., 1994. Netrins are diffusible chemotropic factors for commissural axons in the embryonic spinal cord. Cell 78 (3), 425e435. https://doi.org/10.1016/00928674(94)90421-9. (F) Reproduced from Hong, K., Hinck, L., Nishiyama, M., Poo ,M.-ming, Tessier-Lavigne, M., Stein E., 1999. A ligand-gated association between cytoplasmic domains of UNC5 and DCC family receptors converts netrin-induced growth cone attraction to repulsion, Cell 97 (7), 927e941. https://doi.org/10.1016/s0092-8674(00)80804-1.

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them to reorient within explants over w150e250 mm, an attractive effect (Placzek et al., 1990) (Fig. 4.2E). These two activities, outgrowth promotion and attraction, are dissociable, as Netrin-1 possesses both the outgrowth-promoting and the turning activities of floor plate cells in explant cultures (Serafini et al., 1994; Kennedy et al., 1994), whereas Shh only has turning activity (Charron et al., 2003). Similarly, Netrin-1 can both support axon extension on a substrate and cause turning in dissociated cell culture (Fig. 4.2F), whereas Shh only causes turning (de la Torre et al., 1997; Charron et al., 2003; Yam et al., 2009; Wu et al., 2019). Genetic analysis in vivo has defined the contribution of Netrin-1 and Shh from the floor plate. Regional deletion of Netrin-1 from just the floor plate results in several phenotypes, including increased invasion of the motor column and a thinned commissure (Fig. 4.3AeC). The increased invasion appears to reflect loss of the long-range attractive effect of Netrin-1, whereas the thinned commissure appears to result from stalling of the axons as they approach the midline and may reflect loss of the permissive, outgrowth-promoting function of Netrin-1 (Wu et al., 2019; Moreno-Bravo et al., 2019). Blocking Shh signaling in commissural axons similarly resulted in more axons inappropriately entering the motor column but not a thinned commissure (Charron et al., 2003; Okada et al., 2006; Wu et al., 2019), perhaps reflecting the fact that Shh has only attractive, not permissive activity. Genetic deletion of both Netrin-1 from floor plate and Shh signaling results in additive defects (Wu et al., 2019). Collectively, these results show that Netrin-1 and Shh secreted from floor plate collaborate as long-range attractants over w150e200 mm to help reorient the axons away from the edge and toward the midline and that Netrin-1 functions more locally at the floor plate to guide the axons too. Having two rather than one chemoattractant increases the fidelity of the guidance. A third candidate chemoattractant from floor plate, VEGF, may increase the fidelity even further (Ruiz de Almodovar et al., 2011).

4.3.1.2 Guidance by ventricular zoneederived Netrin-1 in the spinal cord Neural progenitor cells in the developing spinal cord have a radial morphology that extends from the ventricular zone, where their nuclei reside, to the pial surface (Fig. 4.2B). In the ventral two-thirds of the spinal cord, these ventricular zone cells produce Netrin-1 mRNA (Kennedy et al., 1994; Serafini et al., 1996; Varadarajan et al., 2017) (Fig. 4.2C), and Netrin1 protein is found to decorate the entire span of these cells in that region but is significantly enriched near the pial edge of the spinal cord, marking a corridor in the dorsal spinal cord that the axons initially follow (MacLellan et al., 1997; Kennedy et al., 2006; Varadarajan et al., 2017; Varadarajan and Butler, 2017) (Fig. 4.4). Studies using mice lacking a floor plate showed that this ventricular zoneederived Netrin-1 is also important for guidance (Charron et al., 2003; Varadarajan et al., 2017), and this was demonstrated directly using regional deletion of Netrin-1 from part (Varadarajan et al., 2017) or all of the ventricular zone (leaving just the floor plate as a source) (Moreno-Bravo et al., 2019). Loss of this source of Netrin-1 causes several defects, including some axons initiating growth in a dorsal rather than a ventral direction, impaired growth along the edge, some axons leaving the spinal cord via dorsal or ventral roots, and more axons inappropriately invading the motor column rather than making a bee-line to the midline floor plate (Varadarajan et al., 2017; Moreno-Bravo et al., 2019).

4.3.1.3 Synergy between Netrin-1 from floor plate and from ventricular zone in the spinal cord Removing both sources of Netrin-1 leads to both additive and supraadditive effects. This was done either using germline Netrin-1 mutant mice (Serafini et al., 1996; Laumonnerie et al., 2014; Bin et al., 2015; Yung et al., 2015; Varadarajan et al., 2017; Varadarajan and Butler, 2017; Wu et al., 2019) or by removing the two sources separately in the same mice (Moreno-Bravo et al., 2019). In mice lacking Netrin-1 from both sources, the defects in dorsal spinal cord are similar to those in mice lacking just ventricular zone Netrin-1: some axons initiate growth dorsally rather than ventrally, there is impaired growth along the edge, and some axons leave the spinal cord via dorsal roots (Fig. 4.3D and E). In the ventral spinal cord, however, the extent of axon misguidance is more severe than would be predicted from the combined effects of deleting either source of Netrin-1 alone: the axons become very defasciculated and exhibit little directionality, and almost no axons reach or cross the midline (Fig. 4.3D and E). Therefore, in the ventral spinal cord, Netrin-1 from the two sources is partially redundant, as each source can partially compensate for the absence of the other.

4.3.1.4 Interpreting the guidance defects caused by loss of Netrin-1 in the spinal cord Collectively, these studies have suggested a model for the function of Netrin-1 in the spinal cord that involves both local and long-range actions and both permissive (growth-promoting) and guidance (attractive) effects (Fig. 4.4).

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FIGURE 4.3 Netrin-1 from floor plate and from ventricular zone both contribute to commissural axon guidance. (A)e(C) Regional deletion of Netrin-1 selectively from the floor plate results in both long-range and short-range guidance defects. (A) Micrograph of a section from a control embryo (left) or an embryo in which Netrin-1 has been genetically deleted from floor plate cells but not ventricular zone cells (DFP-Netrin1; right), labeled for Robo3 to visualize commissural axons. The growth of the axons is normal in the dorsal spinal cord, and many axons grow ventrally toward the midline, but a thinning of the ventral commissure is evident at higher magnification (bottom panels). (B) In such embryos, deletion of Netrin-1 from floor plate also results in greater invasion of the motor column (delineated in red) by commissural axons. (C) Diagram of changes in the DFP-Netrin1 mutants: thinning of the commissure, lateral displacement of the commissural bundle, and increased invasion of the motor column. (D) and (E) Loss of Netrin-1 from both floor plate and ventricular zone results in much more extensive defects in commissural axon guidance. (D) Sections through a control embryo (left) and a Netrin-1 null mutant (right), labeled with Robo3 to visualize commissural axons. Multiple defects in guidance are evident, which are summarized in the diagram in (E), including (1) misprojecting axons (in the dorsal spinal cord, some project dorsally, and, in the ventral spinal cord, there is dramatic invasion of the motor column) (blue arrows in (D) and text in (E)); (2) inappropriate entry of peripheral axons (not shown) and exit of commissural axons from the dorsal roots (orange arrows in (D) and text in (E)); and (3) an almost completely absent commissure (magenta arrow in (D) and text in (E)). Adapted from Wu, Z., Makihara, S., Yam, P.T., Teo, S., Renier, N., Balekoglu, N., Moreno-Bravo, J.A., et al., 2019. Long-range guidance of spinal commissural axons by Netrin1 and Sonic Hedgehog from midline floor plate cells. Neuron 101 (4), 635e647.e4. https://doi.org/10.1016/j.neuron.2018. 12.025).

First, in terms of protein distribution, both immunochemical (Kennedy et al., 2006) and functional (Moreno-Bravo et al., 2019; Wu et al., 2019) data indicate that Netrin-1 made by floor plate cells diffuses over w150e250 mm and can affect axons over such distances (Fig. 4.4). In contrast, immunochemical data (Kennedy et al., 2006; Varadarajan et al., 2017; Varadarajan and Butler, 2017) suggest that Netrin-1 made by radial ventricular zone cells does not travel far from the cells that make it and becomes enriched in a corridor near the pial edge, where the endfeet of these cells is located (Fig. 4.4). Why does Netrin-1 from the two sources behave differently in terms of its apparent diffusion? One possibility is that there are distinct secretory mechanisms, such that the radial cells deposit the protein locally and the floor plate cells do

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FIGURE 4.4 Summary of the roles in spinal commissural axon guidance of Netrin-1 from ventricular zone progenitors and from floor plate, and of Shh from floor plate. Netrin-1 protein is enriched along the pial edge in the dorsal spinal cord and found throughout the ventral spinal cord. Netrin-1 is produced by cells in the ventral two-thirds of the ventricular zone (VZ-Netrin-1) and by the floor plate (FP-Netrin-1). Netrin-1 at the pial edge, which appears to derive primarily from VZ-Netrin1, acts locally, and serves to confine commissural axons to the pial edge in the dorsal spinal cord (left). When axons enter the ventral spinal cord, they leave the pial edge and navigate around the motor column (MC) to the midline (middle and right). This guidance is directed by FP-Netrin1 acting at a distance (middle), as well as by VZ-Netrin1, which synergizes with FP-netrin1 in guidance in this region (not shown). In addition, Shh (also secreted by the floor plate) acts at a distance to guide commissural axons toward the floor plate (right). From Wu, Z., Makihara, S., Yam, P.T., Teo, S., Renier, N., Balekoglu, N., Moreno-Bravo, J.A., et al., 2019. Long-range guidance of spinal commissural axons by Netrin1 and sonic Hedgehog from midline floor plate cells. Neuron 101 (4), 635e647.e4. https://doi.org/10.1016/j.neuron.2018.12.025.

not (Varadarajan et al., 2017). Another is that the protein could be modified differentially by the two types of cells, such that the protein made by floor plate is more diffusible (Wu et al., 2019). Whatever the mechanism, the net effect is that Netrin-1 made by one source appears to act more locally than Netrin-1 made by the other source. In dorsal spinal cord, early-born commissural axons enter and grow ventrally down an enriched corridor of Netrin-1 protein along the pial edge (Fig. 4.4), and as discussed, in its absence, axon growth in this corridor is impaired, as some axons grow dorsally and some exit via the dorsal roots (Fig. 4.3D and E). This could reflect simply a permissive action of Netrin-1, such that in its absence, the axons are less effectively confined to this corridor. Although a simple permissive function could potentially explain the guidance defects that are seen, the observation of an increasing dorsal-toventral gradient of Netrin protein in this corridor (Kennedy et al., 2006) raises the possibility that the axons are guided by a gradient of protein, something that remains to be evaluated. As also discussed, once the axons reach the ventral spinal cord, breaking away from the edge and redirection toward the midline requires Netrin-1 from both sources, and in its absence, the axons grow ventrally but without directionality (Fig. 4.3D and E). In this case, it seems less likely that Netrin-1 simply has a permissive effect on the axons and instead appears to be guiding them. Again, the presence of an increasing dorsal-to-ventral gradient of Netrin protein in this region (Kennedy et al., 2006) suggests a mechanism for that guidance. This guidance is assisted by Shh (and potentially also VEGF) made by floor plate; other factors appear to contribute as well, such as the repellent NELL2, made by cells in the motor column, which appears to contribute to keeping the axons out of that region (Jaworski et al., 2015). Finally, the observation that, in the absence of Netrin-1, commissural axons leave the spinal cord via the dorsal and ventral roots (Laumonnerie et al., 2014; Wu et al., 2019) (Fig. 4.3D and E) could again reflect either permissive or guidance effects, though studies in the hindbrain (see later) suggest that it might just reflect a permissive effect, i.e., that Netrin-1 simply makes the spinal cord a more favorable environment for the axons, thereby deterring their exit.

4.3.2 Mammalian hindbrain As in the spinal cord, in the hindbrain, commissural axons born dorsally extend to the ventral midline, and Netrin-1 is again critical for their guidance. Interestingly, though, in this region, ventricular zone cells appear to be the more important source of Netrin-1.

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4.3.2.1 In hindbrain, Netrin-1 from ventricular zone is more important than from floor plate As in the spinal cord, in the hindbrain, commissural axons are also attracted in vitro by floor plate cells and by Netrin-1 (Shirasaki et al., 1996), and in vivo complete loss of Netrin-1 results in profound defects in growth of these axons to and across the midline (Dominici et al., 2017; Yamauchi et al., 2017). However, a similar approach of regional deletion showed that removal of Netrin-1 from just the floor plate had little to no effect on these axons, whereas its removal from the ventricular zone resulted in defects that were almost (though not quite) as extensive as in the null mutant (Dominici et al., 2017; Yamauchi et al., 2017). While the effect of removing Netrin-1 from floor plate together with other chemoattractants such as Shh has not yet been examined, these results already show that in hindbrain the major source of Netrin-1 is the ventricular zone, with the floor plate making a lesser contribution. What accounts for the difference between spinal cord and hindbrain? It has been pointed out that the challenges facing commissural axons in these two regions are different (Moreno-Bravo et al., 2019; Wu et al., 2019). In the spinal cord, after growing near the pial edge, the axons break away and steer to the midline, largely avoiding the motor column. This may be important to prevent them from leaving the spinal cord through the ventral roots (as occurs in Netrin-1 mutants), and the main purpose of the chemoattractants appears to be to orchestrate this redirection toward the midline. The situation in the hindbrain is different, as the axons grow near the pial edge over very long distances (millimeters) to the midline and never leave the edge since there is no motor column to avoid, so a chemoattractant to redirect the axons seems less necessary. Chemoattractants such as Netrin-1 and Shh that act over w150e250 mm may therefore be most useful when reorientation of axons is required within that range, as is the case in spinal cord but not hindbrain.

4.3.2.2 Control of neuronal cell migration by Netrin-1 in the hindbrain Not only the axons of commissural neurons but also their cell bodies can migrate along a dorsoventral, circumferential trajectory in the subpial region in the spinal cord and hindbrain. Impaired migration of these neuronal cell bodies was observed in the hindbrain of Netrin-1 and DCC mutants (Serafini et al., 1996; Fazeli et al., 1997; Bloch-Gallego et al., 1999; Zelina et al., 2014) and also in the spinal cord (Junge et al., 2016; Leggere et al., 2016). In vitro experiments on one of these classes of neurons, pontine neurons in the hindbrain, show that they are attracted in vitro by floor plate cells and by Netrin-1 in a DCC-dependent manner (Yee et al., 1999; Zelina et al., 2014). Robo3 is also a major regulator of these migrations in vivo (Marillat et al., 2004), functioning at least partly as a co-receptor with DCC to potentiate responses to Netrin-1 (Zelina et al., 2014)eas discussed above for axons. Interestingly, just as axons abnormally exit the spinal cord in Netrin-1 mutants, in the hindbrain of these mutants, the migrating neurons were found to leave the central nervous system via various cranial nerve roots (Moreno-Bravo et al., 2018; Yung et al., 2018). Experiments in which Netrin-1 gene expression was reintroduced in the central nervous system in a Netrin-1 mutant showed that the mere presence of Netrin-1 protein is sufficient to prevent the cells from leaving the hindbrain, even if its distribution is not completely normal (Yung et al., 2018). This suggested that the ability of Netrin-1 to retain the migrating cells in the CNS reflects a permissive action, rather than a guidance action that would be dependent on the precise, graded distribution of the protein.

4.3.3 Guidance of other classes of mammalian axons and cells: attraction, repulsion, and modulation Netrin-1, acting via DCC, has been implicated in the attractive guidance of a variety of other classes of axons, including retinal (Deiner et al., 1997), thalamocortical (Braisted et al., 2000; Powell et al., 2008; Bielle et al., 2011), cortical efferent (Métin et al., 1997; Richards et al., 1997; Fothergill et al., 2014), hippocampal (Barallobre et al., 2000), dopaminergic (Lin et al., 2005; Xu et al., 2010; Li et al., 2014), and sympathetic (Brunet et al., 2014) axons. Interesting similarities and differences in the guidance of these axons compared to those in the spinal cord and hindbrain are worth noting. For instance, as mentioned, retinal ganglion cell axons require Netrin-1 to exit the retina at the optic disc and enter the optic nerve, and this appears to reflect a local, not a long-range, action of Netrin-1 made by cells at the optic disc (Deiner et al., 1997); these axons express DCC but not Robo3 (Deiner et al., 1997; Friocourt et al., 2019). In the case of axons that form the corpus callosum, the evidence indicates that Netrin-1 attracts the early wave of callosal pioneer axons, which derive from the cingulate cortex. However, Netrin-1 is not attractive for the majority of callosal axons, which derive from the neocortex, and instead regulates their guidance through a different mechanism, by blocking repulsion by the chemorepellent Slit2 (discussed in Chapter 7) (Fothergill et al., 2014). Netrin-1 may similarly help guide thalamocortical axons by modulating responses to Slit2, in this case actually converting Slit2 repulsion to attraction (Bielle et al., 2011). And in the case of dopaminergic axons, the two well-known classes of these axonsdfrom the substantia nigra

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and from the ventral tegmental area, which project to different regions of the striatumdshow preferences for different concentrations of Netrin-1 that contributes to the topographic organization of their projections (Li et al., 2014). Thus, Netrin-1 shows a great deal of versatility in how it can contribute to attractive axon guidance. While in mammals many of the best characterized responses to Netrin-1 have been of the attractive variety, repulsion by Netrin-1 has also been well documented. For example, trochlear motor axons in the hindbrain, which migrate away from a Netrin-1 source, are repelled by Netrin-1 in vitro (Colamarino and Tessier-Lavigne, 1995), although their guidance in vivo is not impaired by loss of Netrin-1, indicating the operation of redundant cues (Serafini et al., 1996). Sensory axons that project from the dorsal root ganglia into the spinal cord initially pause before entering the spinal cord, and this appears to reflect a repellent action of Netrin-1 in the subpial area in the spinal cord, since the axons are repelled by Netrin-1 in vitro via the UNC5C receptor, and in Netrin-1 and Unc5C knockouts, they project into the spinal cord prematurely (Masuda et al., 2008; Varadarajan et al., 2017; Watanabe et al., 2006; Wu et al., 2019). Netrin-1 also contributes to guiding the axons of Unc5C-expressing motoneurons through repulsion into the developing ventral limb bud, since Netrin-1 is expressed in the dorsal limb, and in Netrin-1 or Unc5C knockouts there is significant misprojection of the axons into that region (Poliack et al., 2015). Netrin/UNC5 signaling also regulates various noncircumferential cell migrations in the embryo, including to repel migrating cerebellar cells and thereby establish certain cellular boundaries (Przyborski et al., 1998), and to stimulate the dispersal of developing oligodendrocyte precursor cells in the embryonic spinal cord (Jarjour et al., 2003; Tsai et al., 2003).

4.3.4 Invertebrate systems Netrin function in axon guidance has also been extensively studied in invertebrates, particularly in C. elegans and Drosophila. Here, we summarize special insights obtained in those systems.

4.3.4.1 Attraction and repulsion by UNC-6 in Caenorhabditis Elegans C. elegans has proven to be an especially powerful system for genetic analysis of Netrin function. Initial genetic analysis showed that mutations in the Netrin gene unc-6 and in the Netrin receptor genes unc-40 and unc-5 affect the guidance of multiple populations of circumferentially migrating axons and cells without affecting their longitudinal movements; unc-5 regulates dorsal migrations, unc-40 primarily regulates ventral migrations, and unc-6 regulates both dorsal and ventral migrations (Hedgecock et al., 1990) (Fig. 4.5). Molecular, genetic, and expression analysis led to the model that UNC-6 is a ventrally-derived Netrin ligand that mediates attraction via the DCC family receptor UNC-40 and repulsion via the UNC5 receptor, with UNC-40 also potentiating UNC-5 activity for long-range repulsion (Ishii et al., 1992; Leung-Hagesteijn et al., 1992; Hamelin et al., 1993; Chan et al., 1996; Wadsworth et al., 1996). Importantly, circumferentially migrating axons and cells throughout the animal are guided by these mechanisms, indicating that UNC-6 provides a global guidance wild type nematode

unc-6 null loss-of-function

unc-5 loss-of-function

unc-40 loss-of-function

unc-6 dorsal defective

unc-6 ventral defective

FIGURE 4.5 Mutations in the Netrin unc-6 and its receptors differentially affect dorsal and ventral migrations. Schematic of defects in dorsally migrating and ventrally migrating axons in C. elegans in different mutant backgrounds. Left: diagram of a cross-section through a control embryo illustrating a neuron with its cell body (left black circle) near the ventral pole (V) that sends an axon dorsally (arrow pointing up), and a neuron with its cell body (right black circle) located closer to the dorsal pole (D) that sends its axon ventrally (arrow pointing down). Next panel: null mutations in the Netrin unc-6 result in defects in both dorsal and ventral migrations. Next panel: null mutations in unc-5, which codes for the repulsive Netrin receptor UNC-5, impair dorsal migrations, a defect also seen in specific alleles of unc-6 (the “dorsal-only” alleles). Far right panel: null mutations in unc-40, which codes for the attractive Netrin receptor UNC-40, primarily impair dorsal migrations, a defect seen in yet other alleles of unc-6 (the “ventral-only” alleles). Some dorsal migration defects are also seen in unc-40 mutants (not shown). Reproduced from Goodman, C.S., 1994. The likeness of being: phylogenetically conserved molecular mechanisms of growth cone guidance, Cell 78 (3), 353e356. https://doi.org/10.1016/0092-8674(94)90413-8.

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system, although the extent to which different classes of axons and cells are affected by loss of unc-6 varies, with some being very affected and others only modestly (Hedgecock et al., 1990; Wadsworth et al., 1996), reflecting the operation of other redundant cues, such as dorsally derived SLT-1/Slit (Hao et al., 2001). These cell and axon migrations have in common that they occur on a specific basal lamina, with cells migrating on one side and axons on the other, suggesting that UNC-6 itself is associated with the basal lamina (Ishii et al., 1992). How does UNC-6 guide these migrations? It has been proposed that UNC-6 secreted by ventrally-located cells could establish a decreasing ventral-to-dorsal gradient that directs guidance (Wadsworth et al., 1996). Support for such a mechanism was provided by time-lapse analysis of initial axon formation of a specific circumferentially projecting neuron (HSN), since its leading edge is normally polarized from the get-go toward the distant ventral UNC-6 source and this polarization is abolished in unc-6 and unc-40 mutants (Adler et al., 2006). Importantly, the distinct functions of UNC-6 in guiding cells versus axons, and dorsal versus ventral migrations, are genetically separable, as some partial loss-of-function alleles of unc-6 selectively affect only cell but not axon migrations, others selectively affect just dorsal migrations, and yet others just ventral migrations (Hedgecock et al., 1990; Wadsworth et al., 1996; Lim et al., 2002) (Fig. 4.5). Of note, unc-6 mutations that selectively impair dorsal migrations result in the selective deletion of the second EGF-like repeat of UNC-6 (Wadsworth et al., 1996), anticipating the observation that, in mammals, the homologous domain of Netrin-1 mediates the Netrin-1/UNC5B interaction (Grandin et al., 2016).

4.3.4.2 Attraction and repulsion by Netrins in Drosophila Netrins also contribute to directing the guidance of commissural axons in Drosophila, but with an interesting twist. The two fly Netrin genes, Netrin-A and Netrin-B, are both expressed by midline cells; consistent with a role in midline attraction, removing both causes severe defects in commissure formation, which can be rescued by reexpression of either one alone in midline cells (Mitchell et al., 1996; Harris et al., 1996). These Netrin actions are mediated by the DCC-family receptor Frazzled (Kolodziej et al., 1996). These findings, which parallel those in vertebrates and C. elegans, might suggest that fly Netrins also function to attract the axons to the midline at a distance. Contradicting this, however, a subsequent study (Brankatschk and Dickson, 2006) showed that, in the absence of the two Netrins, the axons still grow normally to the midline, and only once they get there do they show defective midline crossing that appears to be the ultimate cause of defective commissure formation. Moreover, a Netrin construct that remains tethered to cell membranes was fully capable of rescuing the commissural defect in Netrin mutant flies (Brankatschk and Dickson, 2006). Together, these results imply that, at the Drosophila midline, Netrins only function locally to enable midline crossing and not at a distance to attract axons to the midlinedunlike in the mammalian spinal cord where Netrin-1 does both (Fig. 4.3)dand leave open the nature of the guidance system(s) that directs the axons to the midline. A similar theme of local action is evident in the Drosophila visual system, where the growth cones of a particular class of photoreceptors require Netrins to form normal projections to a specific target layer expressing these ligands, but this function can again be mediated by a membrane-tethered Netrin construct (Timofeev et al., 2012). Furthermore, time-lapse analysis shows that, in the absence of Netrins, guidance of photoreceptor axons to this layer is normal, but, on arriving, the axons fail to remain attached to the target layer (Akin and Zipursky, 2016)devidence of a contact-mediated attractive and/ or adhesive action of Netrins. Whether Netrins mediate attractive effects at a distance for any axons in Drosophila remains to be determined. This is not to say, however, that Netrins only have short-range actions in Drosophila. In fact, both short- and long-range actions of Drosophila Netrins have been documented for repulsion. This was shown through ectopic expression experiments in which the single UNC5 family receptor was misexpressed in different classes of neurons (Keleman and Dickson, 2001; Brankatschk and Dickson, 2006). When commissural neurons are forced to express UNC5, they fail to grow to the midline, reflecting a repulsive action of Netrin that does not require Frazzled/DCC and that occurs at short range, since the defect is suppressed by removal of Netrin genes but restored by reexpression of tethered Netrin. In contrast, misexpression of UNC5 in some interneurons that normally do not project to the midline drives them out of the central nervous system, a long-range effect that requires Frazzled/DCC function, is suppressed by removal of Netrin genes, but is not restored by reexpression of tethered Netrin. Collectively, these results imply that in Drosophila, as in other species, Netrins can function at both short- and long-range in repulsion, with UNC5 mediating both actions, and with long- but not short-range action requiring Frazzled/DCC (Keleman and Dickson, 2001; Brankatschk and Dickson, 2006). While these misexpression experiments establish that Drosophila UNC5 is a repellent Netrin receptor, its normal function has been more difficult to pinpoint, perhaps because of redundancy. For example, SNa motoneurons avoid a muscle normally expressing Netrin-B and can be caused to avoid other muscles by ectopic expression of Netrin-Bdan effect that requires UNC5; however, removal of Netrins or UNC5 does not cause SNa to enter the muscle it normally avoids, presumably reflecting the operation of redundant guidance cues that are sufficient to keep it out (Winberg et al., 1998; Keleman and Dickson, 2001).

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4.4 Beyond axon and cell guidance: additional roles for Netrins in the nervous system Although Netrins were initially identified based on their roles in axon guidance, they have proven to be remarkable versatile, playing key roles in other aspects of neural development and physiology. First, Netrins can stimulate the branching of axons (Dent et al., 2004), and in the case of cortical neurons, Netrin-1 shifts from stimulating the outgrowth of their axons to stimulating their branching as the neurons age (Matsumoto and Nagashima, 2017). Netrin signaling also regulates aspects of dendrite development. In both zebrafish and Drosophila, Netrin/DCC signaling guides certain dendrites (Furrer et al., 2003; Suli et al., 2006; Brierley et al., 2009; Mauss et al., 2009). In C. elegans, UNC-6/UNC-40 signaling stimulates the growth but not the guidance of a particular dendrite (Teichmann and Shen, 2011). And again, in C. elegans, dendritic self-avoidance and dendritic tiling, which involve dendrite-dendrite repulsion, can be directed by UNC-6/UNC-5/UNC-40 signaling (Smith et al., 2012; Yip et al., 2016). Naturally occurring neuronal cell death is widespread in the developing vertebrate nervous system. Knockout of Unc5A in mice reduces apoptotic neuronal death in the developing spinal cord, implying that UNC5A triggers cell death, perhaps via its C-terminal Death Domain (Fig. 4.1C); the presumptive ligand that activates UNC5A is not known but does not appear to be Netrin-1 (Williams et al., 2006). Netrins also regulate synapse formation. In C. elegans, UNC-6/UNC-40 signaling is required for presynaptic assembly of a particular neuron (Colon-Ramos et al., 2007), whereas UNC-6/UNC-5 signaling inhibits presynaptic assembly of another (Poon et al., 2008). Evidence for synaptogenic actions of Netrins exists in vertebrates as well, specifically in the Xenopus tectum (Manitt et al., 2009), in the mammalian mesolimbic dopaminergic system (Grant et al., 2007; Xu et al., 2010; Manitt et al., 2011) and in the mammalian cortex (Goldman et al., 2013). Beyond regulating synapse formation, Netrin/DCC signaling has been implicated in synaptic function and plasticity. Loss of DCC impairs long-term potentiation (LTP) at cortical synapses (Horn et al., 2013), and evidence has been provided that activity-dependent release of Netrin-1 is an effector of LTP in the hippocampus (Glasgow et al., 2018). Netrins also contribute to aspects of myelination. Newly postmitotic oligodendrocytes express Netrin-1, which contributes via DCC to the branching of their processes and the formation of myelin sheets (Rajasekharan et al., 2009). In mature oligodendrocytes, DCC localized to oligodendroglial membrane loops appears to bind Netrin-1 on axons, and in the absence of DCC or Netrin-1, the nodes of Ranvier are disrupted (Jarjour et al., 2008).

4.5 Involvement of Netrin signaling in disorders of the nervous system An exciting development in the past decade has been the discovery that mutations in the Netrin-1 or DCC genes in humans underlie several inherited neurological deficits that result from defective midline axonal crossing. Heterozygous mutations in DCC or Netrin-1 in humans can give rise to so-called congenital mirror movements (CMMs), which are characterized by involuntary movements of one limb that mirrors the intended movement of the opposite limb (Srour et al., 2010; Depienne et al., 2011; Méneret et al., 2017). More detailed analysis of CMM patients with a DCC mutation revealed that the corticospinal tract (CST), which is normally largely crossed, has an enhanced ipsilateral projection in these patients (Fig. 4.6), and each cortical hemisphere activates not just the correct (contralateral) side but also the ipsilateral side, helping explain the mirror movements in these patients (Welniarz et al., 2017). In addition to CMM, mutations in DCC have been shown to underlie some familial cases of agenesis of the corpus callosum (Marsh et al., 2017) and split-brain syndrome (Jamuar et al., 2017). Why the mutations cause one phenotype in some families and different ones in other families is uncertain but may result either from the specific mutations in the families or from genetic modifiers. In addition, while it is natural to think that the defects in CST midline crossing in individuals with DCC or Netrin-1 mutations reflect reduced attraction of these axons to the midline, evidence in certain DCC mutant mice, in which similar CST projection defects are observed, suggests that the phenotype may not result directly from malfunction of DCC in CST neurons and may instead reflect a cascading effect on CST axons of some other midline crossing defect(s) that results from impaired Netrin/DCC attraction (Welniarz et al., 2017). Of note, defects in midline crossing are also seen in individuals with inherited mutations in the human ROBO3 gene, which cause an autosomal recessive disorder termed “horizontal gaze palsy with progressive scoliosis” (HGPPS), which is

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FIGURE 4.6 Some mutations in the human DCC gene result in congenital mirror movements, which involve miswiring of the corticospinal tract. (A) Diagram showing the projection of the corticospinal tract (CST) from the motor cortex (M1) to the spinal cord. In control subjects, most fibers project contralaterally (crossed CST, blue), and many fewer remain ipsilateral (red). (B) Tractography enables visualization of the crossed (c-cst, blue) and uncrossed (u-cst, red) CST at the level of the pyramidal decussation in a control subject (top) and two subjects with DCC mutations (middle and bottom). In the latter, the uncrossed CST is greatly exaggerated. (C) A quantitative measure of “laterality” (i.e., the size of the uncrossed vs. the crossed projections) shows that the CST is more crossed in two controls, but more uncrossed in two patients with DCC mutations. Adapted from Welniarz, Q., Morel, M.-P., Pourchet, O., Gallea, C., Lamy, J.-C., Cincotta, M., Doulazmi, M., et al., 2017. Non cell-autonomous role of DCC in the guidance of the corticospinal tract at the midline. Sci. Rep. 7 (1), 410. https://doi.org/10.1038/s41598-017-00514-z.

marked behaviorally by impairment of coordinated eye movements on the horizontal axis and anatomically by defects in medullary decussation of the corticospinal tract and somatosensory axon tracts (Jen et al., 2004; Sicotte et al., 2006; Chan et al., 2006). These anatomical and behavioral defects can be recapitulated in mice using region- and site-specific genetic deletion of the Robo3 gene and appear to be cell autonomous (Renier et al., 2010). As mentioned earlier, Robo3 can function both to potentiate Netrin-1/DCC signaling and to silence Slit/Robo1/2 repulsion; whether defects in Robo3 cause these midline crossing defects by impairing one or otherdor bothdof these functions is not known. Mutations in UNC5 family receptors in humans have not, to date, been found to cause inherited brain wiring defects. However, human genetic analysis identified a rare inherited mutation in UNC5C that predisposes to Alzheimer’s disease (Wetzel-Smith et al., 2014). How it confers this risk is uncertain, but it may be through a direct effect on neurons since UNC5C mRNA is enriched in the hippocampus, and since the mutation increases death of cultured human cells and rodent neurons expressing UNC5C (Wetzel-Smith et al., 2014). As in the prior discussion of UNC5A’s involvement in developmental cell death, the death-promoting action of UNC5C may involve its conserved C-terminal cytoplasmic “Death Domain” (Fig. 4.1C).

4.6 Netrins: players outside the nervous system Outside the nervous system, Netrins have been implicated in a variety of adhesive, migratory, recognition, and survival events. We highlight a few of these here. Netrins have been implicated in regulating the morphogenesis of various branched epithelial and endothelial structures. During mammary gland ductal development, Netrin-1 secreted by luminal epithelial cells binds to the DCC family receptor Neogenin made by neighboring ductal cap cells, mediating adhesion between the two layers (Srinivasan et al., 2003). In the embryonic lung, Netrin-1 made by epithelial stalk cells surrounds the developing endoderm bud and regulates bud morphogenesis via UNC5B (Liu et al., 2004). In the pancreas, ductal epithelial cells produce Netrin-1, which promotes their adhesion to the ECM, in this case via some integrin receptorsdat least in vitro (Yebra et al., 2003). And UNC5B plays a major role in vascular patterning. Unc5B is expressed in arterial endothelial cells, capillaries, and tip cells, and

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Unc5B mutant mice show aberrant extension of tip cell filopodia and excessive vessel branching, implying that UNC5B negatively regulates these processes (Lu et al., 2004). Although exogenously added Netrin-1 can trigger tip cell filopodial retraction that is dependent on UNC5B, and Netrin-1 is therefore a candidate ligand for regulating the vessels, the expression of Netrin-1 does not fit with such a role in development (Lu et al., 2004). Instead, the divergent Robo family member Robo4 appears to be at least one ligand that regulates vascular UNC5B function (Koch et al., 2011). Netrins are also implicated in regulating immune cell recruitment in acute and chronic inflammation. Leukocytes express UNC5B, and their migration is potently inhibited by Netrin-1, which in the adult is expressed by vascular endothelium and downregulated by immune modulators. This leads to a model in which Netrin-1 functions to inhibit migration into tissues under normal conditions, and its downregulation helps enable an acute inflammatory response (Ly et al., 2005; Rosenberger et al., 2009). In the pathological setting of chronic atherosclerosis, Netrin-1, acting via UNC5B, appears to promote pathology by causing the retention of cholesterol-laden macrophage “foam cells” in the artery wall (van Gils et al., 2012). In addition, Netrins and their receptors have also been implicated in cancer. In certain cellular settings, unbound DCC and UNC5 family receptors can trigger cellular apoptosis that is blocked when the receptors bind Netrin-1, leading to the proposal that these receptors might function as tumor suppressors (Mehlen et al., 1998; Llambi et al., 2001). In colorectal cancer, DCC is lost at high frequency, consistent with a possible tumor suppressor role (Fearon et al., 1990). Evidence in support of this possibility was obtained using a mouse in which the proapoptotic function of DCC was blocked by genetic mutation, which caused increased intestinal neoplasia and increased susceptibility to tumorigenesis triggered by another cancer-causing gene (Castets et al., 2011). Furthermore, an antibody to Netrin-1 that blocks its interaction with UNC5B triggers death in vitro of tumor cells that express both factors (Grandin et al., 2016), which has motivated a clinical trial to evaluate the utility of this antibody as a treatment for various cancers. These provide just some of the most prominent examples of a growing list of functions that are being discovered for Netrins outside the nervous system.

4.7 Conclusion Circumferential axon and cell migrations toward and away from the midline are a prominent feature of the development of all bilateral animals. Netrins are an ancient set of molecules that were coopted early in evolution to provide a global coordinate system that directs many of these migrations. Studies of Netrins have also revealed many important insights into the logic and mechanisms of axon guidance, including the findings that: l

l

l l

l

l

l

l

Netrins are bifunctionaldattractive to some axons and repulsive to others, depending of the receptors expressed by the axon (DCC family for attraction, UNC5 family for repulsion); the function of these receptors can be potentiated by interaction with other coreceptors (DCC receptors potentiate UNC5 repulsion in multiple species; Robo3 potentiates DCC attraction in mammals); the function of DCC family receptors can also be attenuated or inhibited by interactions with yet other receptors; Netrin receptors are also promiscuous, capable of mediating signals from other attractive, repulsive, or modulatory ligands, either by direct binding of the ligands or through other coreceptors; Netrins can function to guide axons at long range (i.e., over a few hundred micrometers) in some biological settings but only at short range (i.e., in a contact-mediated fashion) in others, apparently reflecting the extent to which the Netrin protein travels from its source and/or the sensitivity of the responding axon, which in turn may depend on expression of a potentiating coreceptor; Netrins possess multiple activities for axons that can be distinguished biologically, including being able to guide (i.e., attract or repel) when presented from a point source, to stimulate outgrowth when presented uniformly, and to promote adhesion Netrins, acting via the same receptors, can influence many other aspects of neural development and function, including axon branching, pruning, synaptogenesis, and plasticity Netrins have also been coopted to control many other aspects of development and physiology in nonneural systems and are implicated in a variety of pathological processes.

Despite this progress, however, many questions remain, including how Netrins interact with their receptors at a structural level, how attractive and repulsive signals are transduced, whether there are specialized cellular and/or molecular mechanisms to regulate the range of action of Netrins, and, most foundationally, whether long-range guidance effects of Netrins in vivo are mediated by gradients of Netrins, as has been hypothesized in vertebrates and in C. elegans. The availability of powerful genetic, biochemical, and imaging tools to study Netrin function and mechanism should help

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answer these questions in coming years. Importantly, the discovery that mutations in the Netrin-1, DCC, and Robo3 genes cause inherited neurological defects in humans provides powerful tools for understanding how normal and abnormal patterns of brain wiring control key aspects of human behavior.

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104 PART | I Formation of axons and dendrites

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106 PART | I Formation of axons and dendrites

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Axon guidance: Netrins Chapter | 4

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108 PART | I Formation of axons and dendrites

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Chapter 5

Axon guidance: semaphorin/neuropilin/ plexin signaling Marieke G. Verhagen and R. Jeroen Pasterkamp Department of Translational Neuroscience, UMC Utrecht Brain Center, University Medical Center Utrecht, Utrecht University, Utrecht, The Netherlands

Chapter outline 5.1. 5.2. 5.3. 5.4. 5.5.

Introduction Structural features Mechanisms of intracellular signaling Function in neural circuit development Semaphorins, plexins, and neuropilins in neurological disorders

109 111 113 115 116

5.5.1. Autism spectrum disorder 5.5.2. Kallmann’s syndrome 5.5.3. Amyotrophic lateral sclerosis 5.5.4. Late-onset neurodegenerative diseases 5.6. Conclusions and perspectives References

116 116 117 117 117 118

5.1 Introduction To establish functional neuronal connections during development, axons follow specific paths that are marked by instructive proteins known as axon guidance proteins. These proteins act to attract or repel axons and function over short or long distances. Although initially identified for their role in axon guidance, axon guidance proteins are now known to have much broader functions, both within and outside the nervous system. In the nervous system, axon guidance proteins are, for example, involved in the regulation of cell migration, dendrite morphology, and synaptogenesis (Pasterkamp and Giger, 2009; Tran et al., 2007; Yoshida, 2012; Koropouli and Kolodkin, 2014). Our nervous system consists of millions of neurons and glial cells, axonal connections, and synaptic contacts, but only a limited set of axon guidance cues (w100) has been identified that can regulate the formation of this complex organ (Pasterkamp and Kolodkin, 2003; Pasterkamp, 2012). Accumulating evidence indicates that distinct molecular mechanisms act to diversify the effects of axon guidance cues, allowing them to control a disproportionally large number of different cellular events. Several different families of axon guidance cues have been identified (Pasterkamp and Kolodkin, 2003; Pasterkamp, 2012), but the focus of this chapter is on the largest family of canonical axon guidance proteins, the semaphorins. The first semaphorins were reported in the early 1990s, i.e., collapsin-1 (now called Sema3A (Luo et al., 1993)) and fasciclin IV (now called Sema1a (Kolodkin et al., 1992)). Since then, 28 additional semaphorins have been identified and categorized into eight subclasses on the basis of sequence similarity and structural features (Bamberg et al., 1999; Alto and Terman, 2017). Classes 1 and 2 contain invertebrate semaphorins; classes 3e7 contain vertebrate semaphorins; and class V contains viral semaphorins. Sema5c is an exception, as it is also found in invertebrates (Fig. 5.1). Semaphorins play important roles in various biological processes ranging from the regulation of immune responses to angiogenesis and tumorigenesis (Casazza et al., 2007; Nishide and Kumanogoh, 2018; Neufeld et al., 2016; Neufeld et al., 2012). In addition to their role in axon guidance, semaphorins control the migration of various neuronal cell types such as neural crest cells in the peripheral nervous system (Osborne et al., 2005) and GABA-ergic interneurons (Marín et al., 2001), CajaleRetzius cells (Bribián et al., 2014), cortical neurons (Chen et al., 2008), and cerebellar granule neurons (Kerjan et al., 2005; Maier et al., 2011; Renaud et al., 2008) in the central nervous system. Other neural processes regulated by semaphorins include neuronal proliferation

Cellular Migration and Formation of Axons and Dendrites. https://doi.org/10.1016/B978-0-12-814407-7.00005-5 Copyright © 2020 Elsevier Inc. All rights reserved.

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110 PART | I Formation of axons and dendrites

FIGURE 5.1 Semaphorins and semaphorin receptor complexes. Thirty semaphorins have been identified and can be categorized into eight classes based on structural similarities. In neural cells, semaphorins signal mainly through plexins. Eleven plexins have been identified (in both invertebrates and vertebrates), and these are subdivided into four classes (AeD). Sema3s signal through PlexinAs but require neuropilins (only detected in vertebrates) for activating plexins, with the exception of Sema3E. Sema3E binds PlexinD1 directly in the absence of neuropilins. Sema2a, Sema2b, and Sema3s are secreted proteins. Transmembrane semaphorins are Sema1a, Sema1b, and Sema5c in invertebrates and Sema4s, Sema5s, and Sema6s in vertebrates. Sema7A is the only glycosylphosphatidylinositol (GPI)-linked semaphorin. SemaVA and SemaVB are found in viral genomes. Sema6s, and most likely other membrane-anchored semaphorins, can influence signaling through inhibitory cis interactions with PlexinAs. In addition, several transmembrane semaphorins function as receptors (reverse signaling). Neuronal semaphorin receptor complexes often contain coreceptors that can affect signaling outcome, for example: RTKs, receptor tyrosine kinases; OTK, off track; Gyc76c, guanylyl cyclase, proteoglycans, integrins; CAMs, immunoglobulin superfamily cell adhesion molecules; Tim-2, T-cell Ig and mucin domainecontaining protein 2; CD-72, B cell differentiation antigen CD72; CLCP1, CUB, LCCL-homology, coagulation factor V/VIII homology domains protein 1; TREM2, triggering receptor expressed on myeloid cells 2.

and polarity, neurite growth and pruning, synapse formation and function, and dendrite morphology (Pasterkamp and Giger, 2009; Tran et al., 2007; Yoshida, 2012). Semaphorins exert their effects by binding to multisubunit cell surface receptors. In neurons, plexins constitute the major semaphorin receptors. The first plexin receptor identified, virus-encoded semaphorin protein receptor (VESPR) (now known as PlexinC1), binds viral semaphorins and mediates cell adhesion (Ohta et al., 1995; Comeau et al., 1998; Winberg et al., 1998). Plexins are large transmembrane signaling molecules that are subdivided into four classes (PlexinAeD). In addition to their interaction with plexins, semaphorins can bind other (co)receptors such as CD72 (Ishida et al., 2003), Tim2 (Kumanogoh et al., 2002), integrins (Jongbloets et al., 2017), proteoglycans (Cho et al., 2012), and neuropilins (Fig. 5.1).

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Intriguingly, semaphorins not only function as ligands, but some transmembrane semaphorins can also act as receptors, a process called reverse signaling, depending on the molecular and cellular context (Battistini and Tamagnone, 2016). In this chapter, we further discuss the semaphorin, plexin, and neuropilin protein families and provide an overview of their signature functions in the nervous system. First, we discuss general features and novel aspects of semaphorin, plexin, and neuropilin molecules and of the signaling pathways activated downstream of these molecules, stressing general principles and recent insights. The focus here is on cell migration and the formation of neuronal connections. Second, we discuss examples of recent work showcasing how dysregulation of axon guidance proteins might cause pathological changes and neurological diseases, and we explore the potential these insights have for developing therapeutic strategies and improving functional recovery.

5.2 Structural features Semaphorins are defined by a conserved characteristic feature called the semaphorin (sema) domain (Bamberg et al., 1999). This extracellular N-terminal w400 amino acid region facilitates homodimerization and interactions with semaphorin receptors. In addition, semaphorins, except class V (viral) semaphorins, contain a cysteine-rich plexinesemaphorine integrin (PSI) domain located at the C-terminus (Love et al., 2003; Antipenko et al., 2003). Other class-specific domains are immunoglobulin (Ig)-like domains found in most semaphorins, a C-terminal basic domain specific for class 3 semaphorins (Luo et al., 1993) and thrombospondin domains that are specific for class 5 semaphorins (Adams et al., 1996). Semaphorin classes 1, 4, 5, and 6 are composed of transmembrane proteins. Members of classes 2, 3, and V are secreted, and the only semaphorin in class 7 (Sema7A) is a glycosylphosphatidylinositol (GPI)-linked semaphorin (Pasterkamp, 2012; Alto and Terman, 2017; Jongbloets and Pasterkamp, 2014; Kolodkin and Tessier-Lavigne, 2011). Structural analysis of the sema domain revealed a propeller-like appearance composed of secondary structure b-sheet twists (Love et al., 2003; Antipenko et al., 2003). The protein topology of the sema domain broadly conforms to the typical architecture of a seven-blade b-propeller fold, which is a commonly found structural scaffold in various proteins (Fülöp and Jones, 1999). Structural analysis revealed distinct insertions in different sema domains that determine binding specificity (Janssen et al., 2010). The sema and PSI domains of different semaphorins can differ slightly, and the various classspecific domains that follow the sema and PSI domains are likely to influence the stability of proteineprotein complexes. These differences among semaphorins influence specific proteineprotein interactions such as dimerization, semaphorineplexin binding, and semaphorineneuropilin interactions (Gherardi et al., 2004). Like semaphorins, plexins contain a sema domain followed by PSI domains that together facilitate proteineprotein interactions. These shared domains and the generic architecture of semaphorins and plexins suggest common modes of interaction and signaling. Additional domains found in plexins are immunoglobulineplexinetranscription (IPT) domains and a highly conserved intracellular region with a GTPase-activating protein (GAP) homology domain (containing a Rho GTPase binding domain [RBD] insertion). The GAP homology domain mediates the repulsive effects of semaphorins through regulation of cytoskeletal components. Class A plexins (PlexinAs) are the best-characterized semaphorin receptors and cluster, along with PlexinBs, in the two largest subfamilies of plexins. PlexinC1 and D1 are phylogenetically the most distant classes of plexins (Tamagnone et al., 1999). Over the past decades, crystal structures of distinct semaphorins and plexins, individually and in complexes, have been reported and highlight key residues and structural features required for proteineprotein interactions and signaling (Love et al., 2003; Antipenko et al., 2003; Janssen et al., 2010; Liu et al., 2010; Nogi et al., 2010; Tong et al., 2009; He et al., 2009; Appleton et al., 2007; Lee et al., 2003; Shang et al., 2017; Kong et al., 2016; Pascoe et al., 2015; Wang et al., 2013; Janssen et al., 2012; Wang et al., 2012; Bell et al., 2011) (see Box 5.1). At the structural level, it has been observed that semaphorins and plexins interact in a 2:2 complex. The sema domain facilitates homodimerization of two semaphorin proteins (Siebold and Jones, 2013). Each protein binds a monomeric plexin protein through the sema domain (Janssen et al., 2010; Liu et al., 2010; Nogi et al., 2010; Janssen et al., 2012). PlexinA1, PlexinA2, and PlexinA4 exist in a ring-like conformation or in a less-frequent, twisted-open chairlike, conformation that may modulate PlexinA signaling (Kong et al., 2016). In the absence of ligands, plexins are positioned on the membrane in a head to stalk conformation in cis, preventing cytoplasmic dimerization (Kong et al., 2016). This mechanism of autoinhibition provides spatiotemporal control over signaling and prevents premature activation of receptors. Upon ligand binding, the PlexinA cytoplasmic domains dimerize, presumably because of intramolecular changes in protein conformation, and intracellular signaling is initiated (Fig. 5.2). Multiple classes of semaphorins can directly interact with plexins; however, class 3 semaphorins, with the exception of Sema3E, require either neuropilin 1 (Nrp-1) or neuropilin 2 (Nrp-2) for assembling a holoreceptor Nrp/plexin signaling complex (Pasterkamp, 2012; Tamagnone et al., 1999; Janssen et al., 2012; Takahashi et al., 1999; Gu et al., 2005) (Fig. 5.1). Neuropilins play an important role during neuronal development. Although a detailed discussion of Nrp function

112 PART | I Formation of axons and dendrites

BOX 5.1 Structural studies of semaphorin, plexin, and neuropilin proteins and complexes

Molecule(s) Semaphorins Sema3AS

A˚ 2.8

Sema3AS-P-I

3.3

Sema3AS-P-PlexinA21-4-Nrp11-4 SEMA4Decto

7.0

SEMA4DectoPLXNB11-2

3.0

Sema6Aecto

2.3

Sema6AS-P

2.5

Sema6Aecto-PlexinA21-4 Sema6AS-P-PlexinA2S-P SEMA7AS-P-IPLXNC1ecto

2.2

2.0

3.6 2.4

Ref Antipenko et al. (2003) Janssen et al. (2012) Janssen et al. (2012) Love et al. (2003) Janssen et al. (2010)

Plexins A˚ PlexinA1cyto-Rac1 3.6

Janssen et al. (2010) Nogi et al. (2010) Janssen et al. (2010) Nogi et al. (2010) Liu et al. (2010)

Bell et al. (2011) PLXNB1cyto 2.4 Tong et al. (2009) PlexinB2cyto-PDZ- 3.2 Pascoe et al. RhoGEF e5.0 (2015) PlexinC1cyto 3.3 Wang et al. (2013) PlexinC1cyto3.3 Wang et al. Rap1B (2013) 2.7 Shang et al. PlexinD1cyto (2017) 3.2 Shang et al. PlexinD1cytoGIPC1 (2017)

PlexinA21-4

2.3

PlexinA2S-P

2.1

PlexinA3cyto

2.0

PlexinAecto PlexinA11-10 PlexinA17-10 PlexinA24-5 PlexinA41-10 PLXNB1cyto-Rac1

4.0 2.2 1.36 7.5 4.4

Ref Wang et al. (2012) Janssen et al. (2010) Nogi et al. (2010) He et al. (2009) Kong et al. (2016)

Neuropilins A˚ NRP13 1.9

Ref Lee et al. (2003)

Nrp11-4

2.7

NRP13-4

1.8

Janssen et al. (2012) Appleton et al. (2007) Appleton et al. (2007) Appleton et al. (2007)

NRP12-42.0 FAB NRP13-FAB 2.2

NRP13-4 NRP13-4Tuftsin NRP15 NRP23-4 NRP23-4 NRP21-4FAB*

2.4

Vander Kooi et al. (2007) 2.15 Vander Kooi et al. (2007) 2.24 Yelland Djordjevic (2016) 1.95 Appleton et al. (2007) 2.3 Appleton et al. (2007) 2.75/ Appleton et al. 3.1 (2007)

A˚, resolution in A˚ngstro¨m; GIPC1, GAIP-interacting protein, C-terminus 1; Neuropilin1-5, a1-a2 (CUB)1-2, b1-b2 (coagulation factor V/VIII homology)3-4, membrane proximal MAM (c)5 domains; PDZ-RhoGEF, PDZ domain guanine nucleotide exchange factor (GEF); Plexin1-10/ecto, 1e10 include sema, PSI, IPT domains/ectodomain; Plexincyto, juxtamembrane (JM) segment, Rho GTPases-binding RBD and GTPase-activating protein (GAP) domains/cytoplasmic domain; Rac1, Rho family small GTPases; Rap1B, Ras family small GTPases; Ref, reference; Semaphorinecto, ectodomain; SemaphorinS-P-I, sema, PSI, Ig domains; Tuftsin, VEGF HBD analog. *FAB ¼ Fab fragment of anti-NRP antibody blocking either vascular endothelial growth factor (VEGF) “anti-Nrp1B” or Sema3 binding “anti-panNrp1A”.

is beyond the scope of this chapter, Nrps are multifunctional molecules well known for their role during cardiovascular development and other physiological and disease processes (Pellet-Many et al., 2008). Neuropilins contain a CUB (a1/a2) domain, a two-factor V/VIII homology domain (b1/b2), and a membrane proximal c (MAM) domain. These domains are required for sema domain binding of requisite interacting semaphorins and influence protein stability. Neuropilins contain a single transmembrane domain and a small cytoplasmic region (Kawakami et al., 1996). In the nervous system, neuropilins bind class 3 semaphorins and interact with PlexinA1e4 to form a holoreceptor complex that can mediate growth cone collapse and axon guidance. The neuropilin coreceptor modulates and stabilizes the 2:2 semaphorineplexin complex, creating a 2:2:2 complex shown for Sema3A, PlexinA1, and Nrp1 (Janssen et al., 2012). The semaphorineplexin and semaphorineplexineneuropilin complexes often contain additional proteins, such as receptor protein kinases and cell adhesion molecules, which modulate downstream intracellular signaling pathways critical for cellular function, often in a cell typeespecific and tissue-specific manner (Pasterkamp, 2012). Semaphorins and plexins can be expressed on separate cells and interact in trans or on the same cell interacting in cis (Fig. 5.2). For example, axonal Sema6B interacts with PlexinA2 expressed at the chick floorplate in trans to induce commissural axon turning postcrossing. In the same system, cis interactions between Sema6B and PlexinA2 on precrossing commissural axons are thought to regulate Sema6B (Andermatt et al., 2014). Interestingly, examination of Sema6Ae PlexinA2 trans versus cis interactions in vitro reveals that cis interactions have a specific modulatory role and require different protein interfaces compared with those needed for trans binding (Perez-Branguli et al., 2016). Another example of

Axon guidance: semaphorin/neuropilin/plexin signaling Chapter | 5

monomer

reverse signaling trans

Plexins

Semaphorins

dimer

113

dimer autoinhibition

monomer

forward signaling trans

no signaling cis inhibition

FIGURE 5.2 Structural basis of different modes of semaphorineplexin signaling. Semaphorins and plexins exist as monomers and homodimers. During binding, semaphorin homodimers bind two plexin monomers to form a symmetrical 2:2 complex. Plexins engage in intramolecular head-to-stalk interactions in the absence of ligand. Semaphorin ligand binding in trans disrupts this state of plexin autoinhibition. Consequently, the cytoplasmic domains of plexins dimerize to trigger downstream signaling. Plexins can also trigger signaling through transmembrane semaphorins, a process termed reverse signaling. Finally, transmembrane semaphorins can bind plexins in cis and inhibit plexin signaling.

an inhibitory semaphorineplexin cis interaction is the binding of Sema6A and PlexinA4 on dorsal root ganglion neurons that prevents activation of PlexinA4 by Sema6A in trans (Haklai-topper et al., 2010). Furthermore, in a subset of starburst amacrine cells (SACs) in the mouse retina, Sema6A binds PlexinA2 in cis to control Sema6A repulsive responses in vitro and SAC stratification and morphology in vivo (Sun et al., 2013). Interestingly, in Caenorhabditis elegans, binding of the transmembrane semaphorin SMP-1 to Plex1 in cis triggers receptor signaling, leading to the inhibition of synapse formation (Mizumoto and Shen, 2013). Thus, cis interactions constitute an excellent regulatory mechanism for providing tight spatiotemporal control of semaphorineplexin signaling (for more details on semaphorin trans vs. cis regulatory mechanisms, please see Haklai-topper et al. (2010)).

5.3 Mechanisms of intracellular signaling Interactions between semaphorins, plexins, and neuropilins activate intracellular signaling pathways that trigger cell typeespecific and tissue-specific responses. Intracellular events that initiate these functional outcomes of semaphorine plexineneuropilin complexes are only partially understood. The following section discusses mechanisms and modes of intracellular signaling involving semaphorins and their receptors during neural circuit development. Emphasis is on the role of small GTPases, which constitute the best studied components of intracellular semaphorin signaling pathways.

114 PART | I Formation of axons and dendrites

Plexins trigger cytoplasmic signaling pathways through their intracellular GAP homology domain (Hota and Buck, 2012). Different signaling routes have been described downstream of plexins that control different aspects of cellular morphology and function. Small GTPases play a crucial function downstream of semaphorins and plexins. GTPases occur in active or inactive states corresponding to their interaction with GTP or GDP, respectively (Vetter and Wittinghofer, 2001; Hall and Lalli, 2010; Wennerberg, 2004). Small GTPases are regulated by GTP/GDP exchange factors. GEFs (guanine nucleotide exchange factors) activate GTPases by increasing binding to GTP. GAPs (GTPase-activating proteins) terminate signal transduction by activating intrinsic GTPase activity to hydrolyze GTP to GDP (Vetter and Wittinghofer, 2001; Bos et al., 2007). In the developing nervous system, small GTPases regulate cell dynamics affecting actin dynamics, cell shape, movement, motility, and position (Etienne-Manneville and Hall, 2002; Gonzalez-Billault et al., 2012; Luo, 2000). Small GTPases are one of the few intracellular signaling proteins that have been implicated in plexin signaling in vivo. For example, the distribution of dentate gyrus granule cells is controlled by Sema5A signaling through interactions of the PlexinA2 GAP homology domain and Rap1, a member of the Rap subfamily of Ras small GTPases. Loss of PlexinA2 GAP activity or the ablation of Rap1 in neurons results in malformation of the dentate gyrus (Zhao et al., 2018). In addition, the Rac-GAP b2chimaerin has been shown to mediate Sema3F-dependent mossy fiber pruning in the hippocampus in vivo. During this process, NRP-2 selectively binds b2-chimaerin, and Sema3F binding activates this GAP to restrain Rac1-dependent effects on the cytoskeleton. Intriguingly, this pathway is required for Sema3F-mediated axon pruning but is dispensable for the effects of Sema3F on axon repulsion and spine remodeling (Riccomagno et al., 2012). Small GTPases are regulated by several other proteins such as receptor tyrosine kinases and serine/threonine kinases. For example, PlexinB1 associates with ErbB2, which is a transmembrane receptor tyrosine kinase. Sema4D ligand binding to PlexinB1 activates ErbB2 tyrosine kinase activity, resulting in autophosphorylation and phosphorylation of PlexinB1. This triggers RhoGEF proteins to activate RhoA and RhoC GTPases, enabling signal transduction (Swiercz et al., 2004). Another example of GTPase regulation is seen in glioma cells. Sema5A binding to PlexinB3 promotes Rac1 recruitment to RhoGDI (RhoGDP dissociation inhibitor), which is a negative regulator of Rho GTPases that directly interacts with PlexinB3. RhoGDI inactivates Rac1 and reduces its membrane localization, resulting in actin remodeling (Li and Lee, 2010). In addition to providing a link to the cytoskeleton, small GTPases also connect plexins to integrin signaling. For example, plexins negatively regulate integrin-mediated cell adhesion through FERM domain-containing guanine nucleotide exchange factor (FARP2) (Kuo et al., 2018). Sema3Aeneuropilin interactions trigger FARP2 to dissociate from PlexinA1 and activation of FARP2 Rac GEF activity. Dissociation of FARP2 is essential for Sema3A-mediated axon repulsion as it causes inhibition of integrin function (Toyofuku et al., 2005). Other downstream processes required for Sema3A-mediated repulsion include recruitment of Rnd1 to PlexinA1 that causes downregulation R-Ras, a small GTPase that normally promotes cell adhesion (Zhang et al., 1996). Sema5AePlexinA2 signaling also affects integrin receptors through small GTPases (Zhao et al., 2018; Duan et al., 2014). PlexinA2 inhibits Rap1 GTPase, thereby negatively regulating integrin-mediated cell adhesion. Loss of plexin GAP activity or ablation of Rap1 results in aberrant granule cell distribution, indicating that PlexinA2 forward signaling through Rap1 regulates granule cell migration and distribution during DG development (Zhao et al., 2018). For more information on other proteins associated with integrin signaling in cellecell and cellesubstrate adhesion events in the nervous system, see Toyofuku et al. (2005), Zhang et al. (1996), Duan et al. (2014), Burridge and Wennerberg (2004). Different small GTPases interact directly with the plexin GAP homology domain to initiate downstream signaling events (Burridge and Wennerberg, 2004; Jaffe and Hall, 2005). Centrally located in the GAP homology domain is the RBD (Rho GTPase-binding domain), which is important for plexin dimerization and receptor activation (Tong et al., 2007). Upon ligand binding, a weak plexineplexin interaction of cytoplasmic RBD regions is triggered, forming an intracellular dimer. These regions contain binding sites for distinct small GTPases. The Rho GTPases Rnd1 and Rac1 disrupt the plexin dimer, leading to conformational changes and receptor activation (Tong et al., 2009). Plexin family members have different binding affinities for Rho GTPases and consequently differ in the mechanism of GAP homology domain regulation. PlexinC1 and PlexinD1 seem to have different requirements of GTPases to regulate GAP activity (Uesugi et al., 2009) compared with members of the more extensively studied PlexinA and PlexinB classes (Wylie et al., 2017). Intracellular signaling through plexins can also occur through direct interaction with cytosolic proteins that function as binding partners of the intracellular part of the plexin receptor. In addition to small GTPases, several other proteins can bind the plexin cytoplasmic domain or act further downstream. An intriguing example of a plexin-interacting protein is Molecule Interacting with CasL (MICAL) (Terman et al., 2002). MICALs are large cytosolic proteins that negatively regulate and enzymatically modify actin through posttranslational residue oxidization, resulting in remodeling and disassembly. SeIR/MsrB antagonizes the effect of MICALs by restoring actin polymerization (Hung et al., 2010; Hung et al., 2011; Hung et al., 2013). Another prominent group of plexin effectors are protein kinases. For example, the tyrosine kinases Pyk2, Syk, FAK, Fer/Fes, Fyn, and Scr play a role downstream of semaphorins and plexins (Hota and Buck, 2012).

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115

A full description of these interactions is beyond the scope of this review, and for more details, see Pasterkamp (2012), Jongbloets and Pasterkamp (2014), Zhou et al. (2008). Semaphorins can mediate both forward (ligand function) and reverse (receptor function) signaling (Fig. 5.2). Class 4e6 semaphorins (Fig. 5.1) can trigger “reverse signaling” pathways via their intracellular domains. Our knowledge of reverse signaling is mostly based on in vitro data, and in vivo evidence is strongest for Sema-1a in Drosophila (Godenschwege et al., 2002; Jeong et al., 2012; Jeong et al., 2017; Yu et al., 2010). However, several studies suggest that reverse signaling could also play an important role during vertebrate neuronal circuit development (Andermatt et al., 2014; Mauti et al., 2007; Bernard et al., 2012; Burkhardt et al., 2005; Leighton et al., 2001; Sun et al., 2015). For example, at the chick spinal cord midline, Sema6B is expressed in commissural axons and is suggested to function cell-autonomously as a receptor. PlexinA2 expressed in floorplate cells acts as an instructive turning signal for commissural axons, and it is detected by axonal Sema6B (Andermatt et al., 2014). Although it remains unclear how Sema6B signaling is mediated intracellularly in this particular example, some downstream effectors of semaphorin reverse signaling have been characterized. For example, during cardiac development, Sema6D reverse signaling through Abl mediates actin dynamics and regulates cell migration (Toyofuku et al., 2004). Sema6A can also act as a receptor through Abl in cultured neurons (Perez-Branguli et al., 2016; Hou et al., 2006). Chemically inducing multimerization of Sema6A receptors causes activation of Abl and other signaling proteins such as GSK3alpha/beta, p130CAS, Ezrin, Radixin, Moesin, and MARCKS (Perez-Branguli et al., 2016). In silico predictions and biochemical studies reveal Ena/VASP-like protein, involved in actin dynamics (Drees and Gertler, 2008), as a potential interactor for Sema6A through its cytoplasmic zyxin domain (Klostermann et al., 2000) and Src as a potential interactor for Sema6B through its Src-homology 3 (SH3) domain (Eckhardt et al., 1997). Class 4 semaphorins have been suggested to function as receptors through their cytoplasmic PDZ domain (Burkhardt et al., 2005). For more details on semaphorin reverse signaling, please see Battistini and Tamagnone (2016).

5.4 Function in neural circuit development In the nervous system, semaphorins control a wide variety of developmental processes such as axon guidance, cell migration, dendrite morphology, and synaptogenesis (Pasterkamp and Giger, 2009; Tran et al., 2007; Yoshida, 2012; Koropouli and Kolodkin, 2014). The best characterized function of semaphorins is their ability to attract or repel axons, directing these structures toward or away from specific regions. A small selection of cellular functions of semaphorins and receptors in aspects of neural circuit development is discussed here. The establishment of neural connections between distant brain regions is essential for normal nervous system function. A well-characterized effect of semaphorins is their ability to restrict neurites to specific lamina (Huberman et al., 2010). Functionally similar neuronal subtypes and projections are often anatomically segregated and patterned to establish functional neural circuits. The retina and hippocampus are two examples of systems that are organized into well-defined lamina. Many different semaphorins and receptors have been described to be involved in the development of retinal and hippocampal mossy fiber projections (Sun et al., 2013; Sun et al., 2015; Tawarayama et al., 2010; Suto et al., 2007; Matsuoka et al., 2011; Matsuoka et al., 2012; Belle et al., 2016). For example, Sema3E-mediated chemorepulsion through PlexinD1 was found to be important for the lamina-specific targeting of entorhinal cortex (EC) and mossy fiber axons in the hippocampus. The EC is the main source of excitatory input to the hippocampus, and the establishment of connections between these brain regions is important for proper brain function (Mata et al., 2018). In mice lacking Sema3EePlexinD1 signaling, the ECehippocampal pathway is disrupted, and mossy fibers are misrouted. Sema3EePlexinD1 signaling has also been implicated in CajaleRetzius cell migration in the cortex (Bribián et al., 2014), development of other neural connections in hippocampus (Chauvet et al., 2007), and synaptogenesis in the striatum (Ding et al., 2012). To establish neuronal connections, axons grow over long distances and rely on intermediate targets and guidance cues along their route. For example, Sema6A was found to be involved in subpallial pathfinding of thalamocortical axons (TCAs) (Little et al., 2009). Thalamic axons are topographically sorted while extending through the subpallium en route to appropriate cortical areas. Sema6A knockout mice show subcortical misrouting of TCAs early during development. TCAs extend aberrantly in the ventral subpallium and initially fail to innervate the visual cortex. Interestingly, axons follow an alternative route, and thalamocortical connectivity is reestablished postnatally. PlexinA2 and PlexinA4 are not expressed by the misrouted axons but along the path of these axons, and therefore, Sema6A is suggested to serve as a receptor on TCAs (Mitsogiannis et al., 2017). At later stages of development, after neuronal connections have been established, further refinement and modification of neuronal networks occurs. For immature neural circuits to transform into functionally mature networks, the elimination of redundant synapses is crucial (Watanabe and Kano, 2011). Sema3E and Sema7A function as retrograde signaling cues for climbing fiber (CF) to Purkinje cell (PC) synapses in the developing mouse cerebellum. Only strengthened CFs extend

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to dendrites of PCs. Weaker CFs are left on the PC soma and eliminated. Normally, PC-derived Sema3A interacts with PlexinA4 and strengthens and maintains CFs. PC-derived Sema7A interacts with PlexinC1 and b1-integrin. The downstream effectors of Sema7A cofilin and focal adhesion kinase (FAK) promote elimination of presynaptic CFs during postnatal cerebellar development (Pasterkamp, 2012; Uesaka et al., 2014). Knockdown of Sema3A in postsynaptic cells accelerates elimination, whereas knockdown of Sema7A impairs synapse elimination during early postnatal development in vivo. Thus, the balance between different semaphorin proteins seems to be important for triggering synapse strengthening or elimination in the cerebellum. Also, the subcellular localization and specific distribution of these semaphorins in the cerebellum is important (Uesaka and Kano, 2018).

5.5 Semaphorins, plexins, and neuropilins in neurological disorders Defects in the expression or function of axon guidance cues have been linked to various neurological diseases. Many canonical axon guidance proteins have been implicated in neurological disease, and for an overview of this work, we direct the reader to other reviews (Pasterkamp and Giger, 2009; Van Battum et al., 2015; Lin et al., 2009; Schmidt et al., 2009; Nugent et al., 2012; Geschwind and Levitt, 2007; Giger et al., 2010; Giacobini and Prevot, 2013; Yaron and Zheng, 2007). Here, we highlight a few recent studies that examine how semaphorins, plexins, and/or neuropilins contribute to neurological diseases that are characterized by pathological changes in neuronal connectivity. Semaphorins, plexins, and neuropilins have been linked to neurological disorders such as autism spectrum disorder (ASD), Kallmann’s syndrome (KS), spinal muscular atrophy (SMA), epilepsy, congenital disorders, schizophrenia, and late-onset brain neurodegenerative diseases such as amyotrophic lateral sclerosis (ALS), Parkinson’s disease (PD), and Alzheimer’s disease (AD). Our understanding of how semaphorin signaling is linked to these diseases derives from human studies but also from experiments using in vitro and animal models. A selection of neurological diseases in which semaphorins and their receptors may play a role is discussed here.

5.5.1 Autism spectrum disorder Abnormal wiring of the brain is a hallmark of ASD. ASD is believed to be caused by neural connectivity disturbances resulting in hypo- or hyperconnectivity between and within brain regions (Geschwind and Levitt, 2007; Amaral et al., 2008; McFadden and Minshew, 2013). Axon guidance proteins may mediate these disturbances, and human genetic studies implicate defective axon guidance protein function. For example, a de novo microdeletion of SEMA5A was found in a patient with ASD and intellectual disability (Mosca-Boidron et al., 2016). Additional evidence for the involvement of SEMA5A gene deletions was found in a subset of patients with Cri du Chat syndrome, showing autistic traits such as repetitive movements, obsessive attachment to objects, and social isolation (Mosca-Boidron et al., 2016). In addition to Sema5A, multiple other axon guidance genes have been associated with ASD (Van Battum et al., 2015). Genetic alterations in the integrin downstream signaling interactor SH3 and multiple ankyrin repeat domains 3 (SHANK3) were linked to ASD and other neuropsychiatric disorders such as schizophrenia, intellectual disability, and manic-like behavior (Durand et al., 2007; Jiang and Ehlers, 2013; Gauthier et al., 2010; Lilja et al., 2017). In knockout mice, silencing of SHANK3 triggered increased Rap1 activity that activates integrin-mediated signaling, leading to altered cell spreading and invasion, whereas restoration of SHANK3 improved autistic-like symptoms in mice (Mei et al., 2016). Interestingly, similar to SHANK3, PlexinA2 GAP activity regulates Rap1 GTPases, and PlexinA2 knockout mice show schizophrenialike behavior. It is therefore tempting to speculate that SHANK3, PlexinA3, and Rap1 are part of a common disease pathway (Zhao et al., 2018).

5.5.2 Kallmann’s syndrome KS is a pathological condition characterized by aberrant GnRH neuron migration, reduced fertility, and olfactory axon guidance defects. Mutations in SEMA3A, SEMA3E, SEMA7A, PLXNA1, NRP1, and NRP2 have been associated with this pathological condition (Cariboni et al., 2015; Känsäkoski et al., 2014; Marcos et al., 2017; Hanchate et al., 2012). Knockout mouse models showed that Sema3A is needed for guidance of GnRH neurons and that removal of Sema3A induces KS-like phenotypes (Cariboni et al., 2015; Cariboni et al., 2011). Sema3A knockout mice, or Sema3A receptor knockout mice, display reduced fertility and failure of olfactory axons to grow to the hypothalamus. As a result, GnRH neurons stall or migrate aberrantly. Another essential cue for GnRH neurons to migrate toward the hypothalamus is Sema7A. Sema7A is expressed on olfactory axons, which are used as a scaffold by GnRH neurons (Messina et al., 2011). In the absence of Sema7A, olfactory axons grow normally, but GnRH neurons stall and fail to migrate into the brain. How

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specific semaphorin (receptor) mutations observed in patients cause KS remains incompletely understood. In addition to Sema3A and Sema7A, Sema4D may also be a potential candidate protein for KSs (Giacobini et al., 2008); however, no mutations have been reported in SEMA4D in KS patients to date (Cariboni et al., 2018).

5.5.3 Amyotrophic lateral sclerosis Defects in semaphorins and semaphorin receptors may also underlie the pathogenesis of ALS (Schmidt et al., 2009). ALS is characterized by loss of motor neurons and their axonal connections. In a mouse model for ALS, SOD1G93A mice, Sema3A expression is increased in terminal Schwann cells that surround neuromuscular junctions (NMJs) (Winter et al., 2006), whereas the Sema3A coreceptor Nrp-1 is located on axonal terminals at NMJs (Venkova et al., 2014). Human postmortem tissue analysis also reveals that Sema3A expression is increased in ALS patients specifically in the motor cortex and to a lesser extent in the spinal cord (Körner et al., 2016). These changes in Sema3A signaling may trigger loss of adhesion or repulsion of motor axons that cause muscle denervation and, consequently, motor neuron degeneration. Interestingly, in vivo antibody administration prolongs life span in SOD1G93A mice (Venkova et al., 2014). However, disruption of Sema3AeNeuropilin1 signaling complex in knockout mouse models did not alter muscle reinnervation (Shadrach and Pierchala, 2018). Therefore, the precise therapeutic potential of targeting Sema3A following neuromuscular injury or disease remains unclear. Interestingly, miR-126-5p was reported to be reduced at presymptomatic stages in ALS mouse models. miR-126-5p targets Sema3A and Nrp-1, and anti-Nrp-1 antibodies can rescue axonal degeneration and NMJ dysfunction in vitro. Similarly, overexpression of miR-126-5p was able to rescue axonal degeneration and NMJ disruption in vitro and in vivo. Together, these data identify miR-126-5p as a therapeutic target in ALS, which may function by acting on aberrant Sema3A signaling (Maimon et al., 2018). Changes in the actin cytoskeleton are a hallmark of ALS and other progressive motor neuron diseases such as SMA (Hensel and Claus, 2018). SMA is characterized by motor neuron degeneration due to deletions of survival of motoneuron 1 (SMN1). PlexinD1 is cleaved by metalloproteases, and in SMA mouse models, cytoplasmic PlexinD1 fragments bind to actin aggregates that perturb cytoskeletal regulation in the growth cone (Rademacher et al., 2017). If and how mislocalization of the PlexinD1 cytoplasmic domain leads to SMA remains to be investigated. Overall, further functional studies are needed to understand how the dysregulation of semaphorins and their receptors contributes to neurodegenerative diseases such as ALS and SMA.

5.5.4 Late-onset neurodegenerative diseases Expression profiling studies reveal changes in the expression of several axon guidance proteins in patients with AD and PD (Van Battum et al., 2015). Polymorphisms located in the coding regions of axon guidance genes have been established as risk factors using genome-wide association studies (GWAS) and metaanalyses. Recent studies provide evidence for new high-risk factors for PD (Yu et al., 2014) linked to semaphorins and also elucidate the roles of certain previously linked semaphorin signaling pathways in the pathogenesis of PD in mouse models (Qi et al., 2016). Although axon guidance proteins play a crucial role in the development and maintenance of dopaminergic circuits (Van den Heuvel and Pasterkamp, 2008), how their dysregulation contributes to the pathogenesis of PD remains to be established. In AD, the expression of several semaphorins and receptors is changed as shown by profiling studies in patients (Van Battum et al., 2015). The Sema3A signaling molecule collapsin response mediator protein-2 (CRMP2) also has been firmly linked to AD. Neurofibrillary tangles that characterize the disease contain hyperphosphorylated CRMP2 that might potentiate Sema3Amediated repulsion. Amyloid-b protein increases the phosphorylation of CRMPs through RhoA small GTPases in vitro, and levels of RhoA and CRMP2 surrounding amyloid plaques in the cortex are increased in a mouse model for AD (Tg2575) (Good et al., 2004; Cole et al., 2004; Soutar et al., 2009; Petratos et al., 2008). Interestingly, GWAS approaches identified a novel PLXNA4 variant linked to AD pathogenesis. PlexinA4 affects tau phosphorylation, leading to neurofibrillary tangle formation in neurons (Jun et al., 2014). These data suggest that PLXNA4 variants may be linked to tau pathology in AD. Overall, these data implicate semaphorins and their receptors in late-onset neurodegenerative disease. It is unlikely that single-gene mutations in semaphorins or their receptors cause neurodegenerative diseases, but they may contribute to disease risk, onset, or progression. Functional studies are needed to assess the precise contribution of semaphorin and semaphorin receptor mutations to AD and PD.

5.6 Conclusions and perspectives The semaphorin family of axon guidance molecules has been implicated in various aspects of neural circuit development. Novel insights into signaling pathways activated by semaphorins and their receptors, plexins and neuropilins, explain their

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ability to exert highly complex and tightly regulated functions. Transmembrane semaphorins function as both ligand and receptors and can form, together with (co)receptors, signaling complexes that are regulated through cis and trans interactions. In addition, presignaling autoinhibition of plexins contributes to the spatiotemporal control of semaphorin signaling. Important future challenges are to determine how different binding modes are initialized to trigger forward or reverse signaling pathways and also cis or trans interactions. Furthermore, we also need to know how appropriate downstream interactors are recruited in specific contexts in vivo. Structural work has significantly advanced our understanding of how semaphorins and receptors interact. The determination of intracellular binding interfaces also has aided the identification of novel downstream interactors involved in several semaphorin signaling pathways. Altered expression and function of semaphorins plays a role in neurological disease. Understanding the fine-tuned regulation and mechanistic details of semaphorin signaling and function is therefore an imperative step toward designing strategies to modulate neural injury and disease. Functional studies are needed to establish causality of patientassociated mutations and to understand underlying disease mechanisms.

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Chapter 6

Ephrin/Eph signaling in axon guidance Franco Weth1 and Artur Kania2, 3, 4 1

Karlsruhe Institute of Technology, Zoological Institute, Department of Cell and Neurobiology, Karlsruhe, Germany; 2Neural Circuit Development

Laboratory, Institut de Recherches Cliniques de Montréal (IRCM), Montreal, QC, Canada; 3Integrated Program in Neuroscience, McGill University, Montreal, QC, Canada; 4Department of Anatomy and Cell Biology, Division of Experimental Medicine, McGill University, Montreal, QC, Canada

Chapter outline 6.1. The setting of the play 6.1.1. Ephs and ephrins 6.1.2. Rules of interaction 6.1.3. Fundamental action modes 6.1.4. Phylogeny 6.2. Mechanisms of ephrin/Eph signaling in axon guidance 6.2.1. Biophysical aspects 6.2.1.1. Membrane distribution 6.2.1.2. Cis interactions 6.2.1.3. Trafficking 6.2.2. Biochemical aspects 6.2.2.1. Signal transduction of forward signaling 6.2.2.2. Signal transduction of reverse signaling 6.3. Ephrins and Ephs in invertebrate axon guidance 6.3.1. Caenorhabditis elegans 6.3.2. Insects 6.4. Binary ephrin/Eph signalingdpathfinding 6.4.1. Peripheral pathfindingdlimb bud innervation 6.4.2. Pathfinding in the spinal cord 6.4.3. Pathfinding in the brain stemdauditory system

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6.4.4. Central pathfinding 6.4.4.1. Optic chiasm 6.4.4.2. Corpus callosum and anterior commissure 6.5. Proportional ephrin/Eph signalingdmapping 6.5.1. Olfactory wiring 6.5.2. Retinotectal/retinocollicular projection 6.5.2.1. Mechanisms of mapping along the anterioreposterior axis 6.5.2.2. Mechanisms of mapping along the dorsoventral axis 6.5.2.3. Computational modeling 6.5.3. Retinogeniculate projections 6.5.4. Thalamocortical projections 6.5.5. Corticocollicular projections 6.6. Ephrins and Ephs in regeneration 6.7. Perspectives and open questionsd“curtain down and nothing settled” Acknowledgments References

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Abbreviations BDNF brain-derived neurotrophic factor c-Cbl cellular casitas B-lineage lymphoma c-Ret cellular “rearranged during transfection” cAMP cyclic adenosine monophosphate CST corticospinal tract Dock1 dedicator of cytokinesis 1 dpy18 dumpy 18 EGF epidermal growth factor En engrailed Erk1/2 extracellular signal regulated kinase 1/2 ESCRT endosomal sorting complex required for transport Fezf2 FEZ family zinc finger 2 FF fiber/fiber FT fiber/target

Cellular Migration and Formation of Axons and Dendrites. https://doi.org/10.1016/B978-0-12-814407-7.00006-7 Copyright © 2020 Elsevier Inc. All rights reserved.

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fwd forward GAP GTPase-activating protein GDNF glia cell lineederived neurotrophic factor GEF guanosine nucleotide exchange factor GFRa1 GDNF family receptor alpha 1 GoF gain of function HD-PTP His domainecontaining protein tyrosine phosphatase Isl2 insulin-regulated protein 2 LAD2 leukocyte adhesion deficiency 2 LIMK1 lim domain kinase 1 LMC lateral motor column LoF loss of function mTOR mammalian target of rapamycin p75NTR p75 neurotrophin receptor PDZ postsynaptic density protein 95/disc large 1/zonula occludens 1 Pou3f4 Pou class 3 homeobox 4 PTP1B protein tyrosine phosphatase 1B rev reverse RGC retinal ganglion cell RTK receptor tyrosine kinase S1 primary somatosensory cortex SAM sterile alpha motif Satb2 special AT-rich sequence binding protein 2 SC superior colliculus SFK Src family kinase SH2/3 src homology 2/3 Syk spleen tyrosine kinase Ten-m teneurin-m Tiam1 T-cell lymphoma invasion and metastasis 1 TrkB tropomyosin receptor kinase B Tsc2 tuberous sclerosis 2 Unc5c uncoordinated 5c V1 primary visual cortex VAB-1 variable abnormal morphology 1 Zic2 zinc finger protein of the cerebellum 2

6.1 The setting of the play 6.1.1 Ephs and ephrins EphA1, the first discovered Eph transmembrane receptor tyrosine kinase (RTK), was named after an overexpressing erythropoietin-producing hepatoma cell line (Hirai et al., 1987), and it was deorphanized by the discovery of ephrin-A1 (Bartley et al., 1994). The mouse and human genomes contain 14 paralogs of Ephs and 8 of ephrins, which, based on their sequences, can be subdivided into A- and B-clades. Their domain composition is shown in Fig. 6.1A. In vertebrates, ephrin-Bs are transmembrane and ephrin-As are glycosylphosphatidylinositol (GPI)-anchored. Thus, both receptors and ligands are membrane bound, making the ephrin/Eph system a cellecell contact signaling system. Complexes of receptor/ ligand ectodomains have been resolved in atomic detail (Nikolov et al., 2013; Seiradake et al., 2016). The minimal signaling complex consists of two ephrin and two Eph monomers (heterotetramer, Fig. 6.1B), and it juxtaposes two receptor molecules for autophosphorylation in trans, which is needed for RTK activation (Lemmon and Schlessinger, 2010). The catalytic center of the monomeric kinase is autoinhibited by the interaction of the juxtamembrane and kinase domains. Phosphorylation of two tyrosines within the juxtamembrane domain relieves the autoinhibition and results in activation (Wybenga-Groot et al., 2001). EphA10 and EphB6 are kinase dead due to substitutions of catalytically relevant amino acids.

6.1.2 Rules of interaction Surprisingly, ephrins and Ephs do not form specific receptor/ligand pairs but, rather, interact promiscuously within their clades: EphAs bind ephrin-As and EphBs bind ephrin-Bs (Gale et al., 1996). However, this oversimplification has

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FIGURE 6.1 Schematic of ephrin/Eph structures and binding affinities. (A) Domain structure of ephrin-As, ephrin-Bs, and the EphA/B monomer. PBM, PDZ-binding motif; ICD, intracellular domain; RBD, receptor-binding domain; LBD, ligand-binding domain; EGF-like, EGF-like domain; FN III, fibronectin-type III domain; TK, tyrosine kinase domain; SAM, sterile alpha motif. The approximate positions of the tetramerization and clustering interfaces in LBD and Sushi domains (dark lines) are shown. The inset shows simplified symbols and color scheme used in this review to indicate ephrin and Eph dimers: EphAs: blue, ephrin-As: pink, EphBs: turquoise, ephrin-Bs: peach. (B) Top views of exemplary ephrin-A/EphA aggregate according to X-ray crystallography: heterotetramer (top), ephrin-A5/EphA4 heterohexamer (middle), detail of an ephrin-A5/EphA2 linear cluster (bottom). (C) Binary ephrin/Eph interaction matrix based on Noberini et al. (2012). Green: binding, red: no binding. Ephrins and Ephs are sorted according to sequence similarity indicated by the neighbor joining trees.

exceptions that are depicted in the interaction matrix shown in Fig. 6.1C (Noberini et al., 2012). Although all indicated interactions are of very high affinity, with dissociation constants (Kds) in the low nanomolar range, qualitative specificity attributions might be of limited value, particularly in a system that can operate in a quantitative mode. In addition, the measured values have to be viewed critically. First, data obtained by different methods are not fully concordant. Second, binding studies using surface-tethered molecules and soluble ectodomain-Fc fusion proteins poorly reconstitute native membraneeprotein interactions. Third, omitting transmembrane and intracellular domains eliminates their potential influences on binding parameters. Thus, in vivo affinities might differ from what in vitro Kds suggest.

6.1.3 Fundamental action modes Whenever ephrins and Ephs are involved in a guidance system, multiple potential modes of action (Fig. 6.2) have to be considered, which will be exemplified in further detail in the following: l l

l

l

l

Signals can be transduced in forward (fwd, ephrin:Eph) as well as in reverse (rev, Eph:ephrin) directions. Interactions do occur not only in trans but also in cis: for example, with ephrin and Eph residing on the same growth cone membrane. Cis interactions are thought to result in signal attenuation. Ephrin/Eph signaling can be attractive or repulsive to the growth cone, with “attraction” describing reinforcement of growth in the direction of the contacted cue. Cues sensed in trans could be located on adjacent cells, serving as guides (fiberepath or fiberetarget [FT] interactions), but also on neighboring fibers including their terminals (fiberefiber [FF] interactions). Most notably, the ephrin/Eph system can operate in a binary mode, instructing choices among discrete guidance alternatives (whether or not to turn at a choice point, to follow a fiber tract, or to invade a selected territory) or in a proportional mode, allowing for the discrimination of subtle differences in surface label concentrations on otherwise indistinguishable cells as occurs in projection axon mapping.

6.1.4 Phylogeny The evolutionary diversification of Ephs and ephrins has been reconstructed from the sequences of orthologous genes in phyla ranging from sponges to vertebrates (Mellott and Burke, 2008), indicating that the B-system is more ancient than the A-system. In urochordates, which still possess only one type of Eph (B-like), two clades of ephrins (A/B) appear for the first time, whereas the A/B distinction of Ephs is a vertebrate innovation. The GPI-anchor is not indicative of the ephrin-A/ B distinction, as it was gained and lost repeatedly during evolution.

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FIGURE 6.2 Fundamental modes of ephrin/Eph action occurring alternatively or in combinations. Ephrin/Eph cues can be displayed in trans on adjacent cells (fiber/path or fiber/target [FT] interactions) or on other growth cones/fiber terminals (fiber/fiber (FF) interactions). In the same cell, interactions can also happen in cis. Signal transduction can occur in forward (FWD) or reverse (REV) direction, leading to either attraction (ATTR) or repulsion (REP) of the fiber terminal. Cis interactions often lead to signal attenuation (ATTN). The integration of ephrin/Eph signaling can evoke binary or proportional responses as exemplified in path finding choices or in topographic mapping, respectively.

Cnidarians are the most ancient animals containing differentiated tissues. Hydra, a model cnidarian, has 4 B-type Ephs and 3 B-type ephrins (Tischer et al., 2013). Both are coexpressed in the endoderm in unequal patterns, indicating ephrin enrichment at sites of reduced cell adhesion. Thus, Hyephrin-B1 expression forms a sharp line at the location where a new bud will be pinched off. This suggests an ancient function of Ephs in epithelial cell adhesion, which was possibly attenuated by coexpressed ephrins. The eventual diversified deployment of the ephrin/Eph system is closely related to the evolution of the mammalian brain. About one-fifth of cortex-specific enhancers are mammalian innovations. These novel enhancers are enriched near genes related to cell migration and neurite wiring, in particular near genes encoding semaphorin and ephrin/Eph signaling components (Emera et al., 2016). This suggests a correlation between the evolutionary unfolding of connectomic complexity and the use of these molecules in axon guidance.

6.2 Mechanisms of ephrin/Eph signaling in axon guidance 6.2.1 Biophysical aspects 6.2.1.1 Membrane distribution GPI-anchored ephrin-As are enriched in membrane lipid rafts (Davy et al., 1999). EphA fwd signaling occasionally needs cAMP, with the requirement being spatially restricted to lipid raft regions (Averaimo et al., 2016). This indicates that Ephs

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might also reside in membrane microdomains. On spinal motor neuron growth cones, ephrins and Ephs are mostly laterally segregated in the membrane (Marquardt et al., 2005). The partitioning among membrane domains, however, can be modulated through variation of ephrin/Eph surface concentrations (Kao and Kania, 2011) or by the membrane lipid composition (Fiederling et al., 2017), which might contribute importantly to signal modulation (see next section). Ligands, subsequent to initial receptor heterodimerization, nucleate the formation of larger ephrin/Eph clusters (Fig. 6.1B). Ephrin-A1 anchored in supported lipid bilayers gets dramatically aggregated by contact with EphA2expressing cells, and inhibition of EphA2 clustering causes abnormal cell responses (Salaita et al., 2010). Pharmacological control of receptor oligomerization indicated that EphB2 needs at least receptor trimers to trigger a biological response (Schaupp et al., 2014). X-ray crystallography, apart from the heterotetramerization interface, revealed additional clustering interfaces (Fig. 6.1A) on Eph ectodomains (Seiradake et al., 2013; Xu et al., 2013). Clustering of activated receptors will cause spatial focusing of the signal, which could help to sharpen guidance decisions. Crystallography also suggests coclustering of unliganded receptors, poising them for binding. An unresolved critical issue, however, relates to the activation of such unliganded receptors, which might be incompatible with quantitative signaling. By direct observation of ephrin-B1/EphB2 cluster dynamics and computational modeling, a mechanism has been suggested for keeping track of ligand concentrations despite the activation of unliganded receptors (Ojosnegros et al., 2017): Clustering might proceed through an activating polymerization phase (addition of monomers to receptor/ligand seed complexes) and a subsequent inactivating phase of polymer condensation. The rates of both phases, which shape the signal profile, are determined by the initial number of seed complexes, i.e., the ligand concentration, which would thus be encoded in the time course of signaling.

6.2.1.2 Cis interactions EphAs and ephrin-As are coexpressed on retinofugal axons, suggesting possible cis interactions. The evidence for these leading to attenuation of EphA signaling is that a loss of function (LoF) of axonal ephrin-As sensitizes EphA fwd signaling and a gain of function (GoF) desensitizes it (Hornberger et al., 1999). Two distinct spinal motor neuron classes differentially innervate vertebrate dorsal and ventral limb muscles, in part, through EphA and EphB signaling. This innervation pattern is directed by ephrin-As in the ventral limb and ephrin-Bs in the dorsal limb, respectively (see Section 4.1 and Fig. 6.5). However, EphB-expressing motor neurons also express some EphAs, and their activity is thought to be attenuated by coexpressed ephrin-A since ephrin-A LoF sensitizes these motor axons to ephrin-As (Kao and Kania, 2011). In contrast, the motor axon population that expresses high levels of EphAs also coexpresses ephrin-As but in separate membrane microdomains. This decreases the incidence of cis interactions and allows axonal ephrin-As to respond to limb mesenchyme EphAs (Marquardt et al., 2005). Thus, in limb bud innervation, cis interactions can be regulated by membrane microdomain localization and switch off a minor, potentially confounding, guidance system, sharpening the binary pathfinding choice. In topographically projecting retinal axons (see Section 5.2.1 and Fig. 6.6A), which use the ephrin/Eph proportional mode, cis attenuation seems to have a more finely tuned role in adaptative modulation, with the appropriate extent of this interaction presumably being adjusted by altering membrane lipid composition and subsequent vesicular trafficking (Fiederling et al., 2017). Mechanistically, cis interactions are generally thought to abrogate trans forward and reverse signal transduction. Cis attenuation could, however, equally well be explained by counterbalanced fwd and rev signaling (Gebhardt et al., 2012). Signaling receptor/ligand interactions in cis require an antiparallel molecular arrangement. This might be enabled by membrane curvature (Schmick and Bastiaens, 2014), perhaps in membrane ruffles, between filopodia or through vesicular localization. Supposedly, a basal degree of cis signaling will in fact be inevitable in every dynamic growth cone and might represent a significant source of noise in this ultrasensitive sensor. Concurrent fwd and rev signaling could be a fitting adaptation to this problem, as it would cause the perturbance (fwd signaling induced by cis ligands) to self-report its extent (via the concurrent rev signaling), enabling appropriate subtraction of the noise.

6.2.1.3 Trafficking Vesicular trafficking might have important roles in ephrin/Eph signaling beyond homeostatic regulation of membrane protein turnover. Thus, cis signaling has been suggested to harness the membrane curvature of endosomes (see previous section). Furthermore, as has been observed for other RTKs, the role of signaling endosomes is beginning to be recognized in the context of ephrin/Eph signaling (Boissier et al., 2013). Theoretically, cell surface signaling delivers an instantaneous picture of the extrinsic conditions, whereas signaling endosomes could provide a short-term memory of recent signaling history, which might help growth cones extract ultraweak signals from a noisy background through temporal averaging.

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Using a novel Förster resonance energy transferebased conformation sensor (Sabet et al., 2015), it was shown in Cos-7 cells that unliganded EphA2 occasionally gets phosphorylated by random collisions with other EphA2 molecules. It is then endocytosed to a Rab11þ compartment, bringing phosphorylated EphA2 into apposition with PTP1B phosphatase for recycling. Ligand binding, in contrast, induces tight clustering and additional phosphorylation events that in turn recruit the E3-ubiquitin ligase c-Cbl. These activated and ubiquitinated clusters are trafficked to Rab5þ early endosomes and eventually to Rab7þ late endosomes. The endocytosis that occurs in the adaptive modulation of proportional ephrin/Eph signaling (“coadaptation”; see Section 5.2.1) appears to use yet another pathway, since this mechanism relies neither on random collisions nor on ligand binding (Fiederling et al., 2017). In addition, there are some peculiarities of ephrin/Eph trafficking that impact signaling. At ephrin/Eph-mediated cellecell contact sites, complete receptor/ligand complexes can be bidirectionally engulfed in a signaling-dependent manner (Marston et al., 2003; Zimmer et al., 2003). Functionally, this trans-endocytosis is assumed to contribute to the conversion of high-affinity ephrin/Eph adhesive binding into repulsion. Another mechanism achieving the same effect is the specific proteolytic cleavage of trans ephrin/Eph complexes by the metalloprotease ADAM10 (Hattori et al., 2000; Janes et al., 2005). More recently, it was found that upon depolarization, EphB-expressing neurons can release exosomes containing Ephs and ephrins in their membranes (Gong et al., 2016). Potentially, the existence of these exosomes extends ephrin/Eph signaling to contact-independent processes. Furthermore, His domainecontaining protein tyrosine phosphatase (HD-PTP), a protein associated with the endosomal sorting complex required for transport (ESCRT) complex that is also needed for exosome formation, is required for ephrin-B2:EphB2 signaling-induced collapse of cultured cells and axonal growth cones, resulting in aberrant guidance of chick spinal motor neuron axons in vivo. HD-PTP depletion abrogates ligand-induced EphB2 clustering and also EphB2 and Src family kinase activation. The exact mechanism underlying this effect is unknown, but this signaling mechanism once again highlights the importance of Eph receptor trafficking at the very earliest signaling steps (Lahaie et al., 2019).

6.2.2 Biochemical aspects 6.2.2.1 Signal transduction of forward signaling There are multiple studies linking specific Eph fwd signal transduction mechanisms to growth cone collapse or repulsion. One of the first ephrin:Eph signaling events is the phosphorylation of specific intracellular tyrosine residues on Eph receptors (Section 1.1), which serve as docking sites for some downstream effectors. Hippocampal growth cones expressing EphA3 in which these different tyrosines have been mutated show diminished collapse responses to ephrin-A; however, at least two of these residues have to be mutated simultaneously to achieve a complete attenuation of collapse (Shi et al., 2010). Although direct evidence for Src family kinases (SFKs) phosphorylation of these residues remains elusive, the involvement of SFKs in Eph-dependent axon guidance is evident from both in vivo and in vitro studies (Kao et al., 2009). Eph receptor kinase activity is required for specific axon guidance decisions (e.g., Kullander et al., 2001), but its requirement for early developmental events and the partial redundancy of Eph class members have precluded a more systematic analysis of this question. A chemogenetic approach using a small molecule blocker of Eph kinase activity has allowed for the efficient inactivation of all three B class receptors in mice, leading to the conclusion that EphB tyrosine kinase activity is required for retinal growth cone collapse and in vivo RGC axon guidance at the optic chiasm, as well as the formation of the corpus callosum (Soskis et al., 2012). An early consequence of Eph receptor tyrosine phosphorylation is that it promotes the binding of SH2 and SH3 adaptor proteins to Eph receptors (Fig. 6.3A). One such adaptor is Nck2 (Holland et al., 1997), and its function is essential for the normal axon guidance of spinal motor neurons in vivo and also responses of cultured motor neurons to ephrin-As and ephrin-Bs. Interestingly, Nck2 also modulates motor axon ephrin-A:EphA responses and phenotypes induced by a-chimaerin GoF (Chang et al., 2018b and see later). Following the phosphorylation of Eph receptor juxtamembrane tyrosines, activation of the Eph tyrosine kinase, and adaptor binding, the most common cellular response to fwd Eph signaling is the destabilization of the growth cone cytoskeleton. This depends on the balance of Rho family GTPase (Rho, Rac, Cdc42) activity levels: High levels of Rho activation and low levels of Rac and Cdc42 activation result in cytoskeleton destabilization (Hall, 2012), which may occur locally in a growth cone such that only the side of the growth cone with a stable cytoskeleton advances, leading to growth cone turning; in more extreme situations, when destabilization occurs globally, the growth cone collapses. This cellular outcome, common to almost all axon guidance signaling pathways, raises the question of the identity of proteins that control the function of Rho family GTPases in response to ephrin/Eph signaling. There are two classes of such proteins:

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FIGURE 6.3 Major biochemical aspects of forward and reverse ephrin/Eph signaling. (A) Known and recently identified transducers of EphA/B fwd signaling. Subsequent to activating phosphorylations of the Eph intracellular domain, the indicated intracellular transducers are recruited. Major downstream pathways converge on actin, which regulates growth cone dynamics and is involved in endocytosis. Key regulators of the actin turnover are Rho-GTPases, such as Rac and Rho, which are activated by GEFs and inhibited by GAPs. Rac induces the formation of lamellipodia or promotes endocytosis, whereas Rho induces actomyosin contraction. Cofilin is an actin-severing protein. The effects of SFKs on actin stability are more indirect. A second pathway influences translation via Tsc2, an indirect negative regulator of the mTOR translation activator. (B) Known transducers of ephrin reverse signaling. The phosphorylation of the ephrin-B intracellular domain recruits transducers, which also eventually target actin. GPI-anchored ephrin-As, however, use other families of transmembrane receptors as coreceptors for signal transduction. One pathway involves signaling via the GDNF/GFRa1/cRet complex and is branch promoting/attractive; the other pathway involves the proBDNF/p75NTR complex and is branch suppressive/repulsive. BDNF, Brain-derived neurotrophic factor; GEF, guanosine nucleotide exchange factor; GAP, GTPase-activating protein; GDNF, glia cell lineederived neurotrophic factor; SFK, Src family kinase; mTOR, mammalian target of rapamycin; GPI, glycosylphosphatidylinositol; GFRa1, GDNF family receptor alpha 1; p75NTR, p75 neurotrophin receptor.

GTPase-activating proteins (GAPs) and guanine nucleotide exchange factors (GEFs). a-Chimaerin is a Rac-GAP that binds EphA4 and relays its activation to the cytoskeleton by decreasing Rac activity. It is required for EphA4-mediated growth cone collapse in response to ephrin-B3 and ephrin-A1 and also for the guidance of corticospinal axons that relay motor signals from the cortex to the spinal cord (Beg et al., 2007; Iwasato et al., 2007). Biochemical data suggested that it may also act downstream of EphB receptors. However, in the context of spinal motor neuron axon guidance, which depends on both EphA and EphB signaling (see section 4.1), a partial a-chimaerin LoF disables EphA signaling while leaving EphB signaling relatively intact. This suggests that a-chimaerin may be an EphA-specific Rac-GAP (Kao et al., 2015). One of the EphA-regulated GEFs involved in axon guidance is ephexin1, whose phosphorylation by activated EphA4 enhances RhoA activity, shifting the GTPase balance toward cytoskeleton destabilization and growth cone collapse (Sahin et al., 2005). Both in vitro and in vivo, ephexin1 LoF affects EphB-mediated axon guidance of spinal motor neurons to a greater extent than EphA-mediated axon guidance, suggesting that although ephexin1 appears to be an effector of both classes of Eph signaling, this ability may be context dependent. Interestingly, SFK GoF axon guidance phenotypes are suppressed by a loss of ephexin1 function, in line with the suggestion that ephexin1 is a phosphorylation target of ephrin:Eph-activated SFKs (Chang et al., 2018a). Another GEF essential for endocytosis, possibly functioning in parallel with ephexin1 is Vav2. It is required for in vitro growth cone collapse by ephrin-A1 and in vivo axon guidance. These observations link ephrin/Eph complex removal from the surface of growth cones and repulsive Eph-mediated guidance (Cowan et al., 2005). One of the most notable exceptions to the general principle that ephrin/Eph signaling mediates growth cone collapse and retraction through GEFs and GAPs is the requirement of the Rac1 GEF Tiam1 for cortical neurite outgrowth induced

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by ephrin-A1. Tiam1 binds to the intracellular domain of EphA2, but it can also function in EphB:ephrinB signaling by binding to the cytoplasmic tail of ephrin-B1 and activating Rac1 (Tanaka et al., 2004). Eph receptors relay extrinsic signals to the growth cone cytoskeleton, and this may implicate the actin-destabilizing protein cofilin (Zhou et al., 2007). Mice mutant for the Lim domain kinase 1 (Limk1), a cofilin inactivator, are characterized by the absence of the peroneal nerve, a phenotype also seen in humans with congenital talipes equinovarus, known as clubfoot. This mutation causes the upregulation of Limk1 in mouse spinal motor neurons, resulting in reduced outgrowth of spinal motor axons and, consequently, the absence of the peroneal nerve. This is likely caused by a deactivation of cofilin, which increases actin stability to the point of inhibiting axon outgrowth. A genetic interaction between EphA4 and Limk1 mutations suggests that the Limk1ecofilineactin pathway may lie downstream of the EphA4 receptor, although the details of this association and similarity to dendritic signaling remain unclear (Collinson et al., 2018). An interesting variation on the classical Eph fwd signal transduction is the cooperation between the Engrailed transcription factors (Ens) and ephrin-A5 in growth cone collapse (Brunet et al., 2005). Surprisingly, Ens have been proposed to act as secreted guidance cues that can enter the growth cone cytoplasm and repel some retinal axons through their effects on protein translation. Ens may potentiate the retinal growth cone collapse caused by ephrin-A5 by increasing growth cone ATP synthesis, whose hydrolysis causes elevation of adenosine and the activation of its A1 receptor (Stettler et al., 2012). In addition to nearly instantaneous signal relay to the cytoskeleton, axon guidance cues also have longer-term effects on protein synthesis within axons and growth cones. EphrinA:EphA signaling is thought to relieve the inhibition of protein synthesis via the tuberous sclerosis complex 2 (Tsc2)ephosphatidylinositol 3ekinase (PI3K)eAktemTOR pathway. The evidence for this includes the impact of Tsc2 mutation on topographic mapping of RGC axons in the visual system and also in vitro growth cone collapse responses to ephrin-As (Nie et al., 2010). In contrast, EphA fwd signaling events that can be recapitulated in embryonic stem cellederived spinal motor neurons do not depend on local protein synthesis (Nedelec et al., 2012).

6.2.2.2 Signal transduction of reverse signaling Ephrin rev signaling was first described for transmembrane ephrin-Bs (Holland et al., 1996), but later on also for GPIanchored ephrin-As (Davy et al., 1999). In both cases, SFKs (Src and Fyn, respectively) are early elements of the signal transduction pathway. It has been suggested that Eph-induced ephrin clustering reorganizes membrane microdomains and thus concentrates SFKs, causing their activation through trans-autophosphorylation. SFKs directly phosphorylate tyrosines of ephrin-B intracellular domains (Fig. 6.3B; Palmer et al., 2002). Thus, docking sites for adaptor proteins such as Nck1/2 are formed that recruit the Rac-GEF Dock1 to activate Rac and trigger a repulsive response. In this context, Rac may have a role in endocytosis (Xu and Henkemeyer, 2009). Since they are GPI anchored, ephrin-As need transmembrane coreceptors to transduce signals, and neurotrophic factor receptors contribute to this function. Thus, free ephrin-A forms a physical complex with the brain-derived neurotrophic factor (BDNF)ebound TrkB RTK to enhance the branch-promoting activity of BDNF in murine retinal axons (Marler et al., 2008). This enhancement is abolished when EphA binds ephrin-A, and the EphA-bound ephrin-A now directly interacts with the proBDNF-bound p75NTR (Fig. 6.3B) and thereby enables the antibranching activity of proBDNF. In addition to regulation of axon collateral branches proximal to the growth cone, this quaternary complex also contributes to the repulsive guidance of retinal axons by ephrin-A rev signaling (Lim et al., 2008; Marler et al., 2010). Notably, a comparable complex, using glia cell lineederived neurotrophic factor (GDNF), GFRa1, and the RTK c-Ret as coreceptors, seems to mediate attractive ephrin-A rev signaling of spinal motor axons ((Bonanomi et al., 2012); see 4.1). The two sections above highlight an interesting relationship between in vivo studies of ephrin/Eph signaling in axon guidance and the biochemical experiments that implicate specific signaling events in these guidance processes. Generally, past studies of Eph function in axon guidance have been the driving force in identifying new principles of Eph fwd signaling. However, over the recent years, specific axon guidance paradigms emerged as testing grounds for signaling effectors suggested by biochemical experiments (Fig. 6.4). One interesting outlook for the future is the development of a concerted pipeline of biochemically proven effectors, to be tested in two to three specific paradigms in vivo. This work will be an important prerequisite for the next wave of structural experiments that examine how Ephs/ephrins interact with their intracellular effectors.

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FIGURE 6.4 Summary of evidence for function of various effectors and the tyrosine kinase domain in EphA/B forward and ephrin-B reverse signaling. Green checkmarks represent the presence of loss of function (LoF) in vivo phenotypes in motor axons in the limb, corticospinal axons, and retinal ganglion cell axons at the optic chiasm and in their targets (e.g., tectum or superior colliculus) and callosal (CC) or anterior commissure (AC) axons. The presence of evidence for in vitro isolated growth cone collapse and deduced axoneaxon interactions is also shown. Red minuses indicate a reported lack of a phenotype. In the case of CC/AC, the presence of EphB LoF phenotypes and the absence of EphA LoF function phenotypes are shown. Ephrin-A rev signaling uses coreceptors not belonging to the ephrin/Eph family. The evidence in this table is based on the articles referenced in Section 2.2.1 and others. SFK, Src family kinases; TK, Tyrosine kinase.

6.3 Ephrins and Ephs in invertebrate axon guidance 6.3.1 Caenorhabditis elegans The power of Caenorhabditis elegans genetics has been seminal in uncovering cardinal axon guidance mechanisms. One advantage of studying Eph signaling in this organism is its reduced complexity. C. elegans has only one Eph receptor (VAB-1) and four potential B-class ephrin ligands, and the most conspicuous function for ephrin/Eph signaling in worms is in axon crossing at the midline of the ventral nerve cord, a process related to commissural axon guidance in vertebrates. Genetic evidence suggests that vab-1 may function in a kinase-dependent and independent manner, although so far only evidence of fwd signaling has been found (George et al., 1998; Wang et al., 1999; Zallen et al., 1999). Vab-1 also functions to instruct the precise termination of axons projecting along the anteroposterior axis and also localization of the neuronal soma, as suggested by gain and LoF phenotypes (Mohamed and Chin-Sang, 2006). For some axon guidance events, efn-4 ephrin LoF in C. elegans has more severe phenotypes than vab-1, suggesting that some aspects of this ephrin’s function may be VAB-1 independent. Indeed, biochemical and genetic experiments argue that the L1 neuronal cell adhesion molecule (CAM) protein LAD-2 can act as a noncanonical receptor for EFN-4 (Dong et al., 2016). The function of the extracellular matrix (ECM) in the distribution of axon guidance cues has been suggested by in vitro and biochemical studies. Evidence of this comes from the loss of the prolyl 4-hydroxylase enzyme encoded by C. elegans dpy-18. Its action is most likely through secondary modification of ECM proteins, changing their affinity for various axon guidance ligands, which is indicated by widespread axon guidance defects in dyp-18 mutants. At least a part of this phenotype is caused by decreased expression levels of C. elegans ephrins (Torpe and Pocock, 2014). The observations described earlier exemplify the power of C. elegans genetics to identify novel Eph/ephrin signaling components and interactions with other signaling pathways. Given the clear phenotypes of vab-1 LoF that could form a basis of suppressor or enhancer screens, there is great potential for additional work in this area. Perhaps one reason this line of experimentation is somewhat underexplored stems from the lack of a demonstration that in C. elegans growth cones, ephrin/Eph signaling functions in a mechanistically similar way to how it does in vertebrates. Some simple studies based

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on Eph/ephrin signaling ideas gleaned from vertebrates would certainly clarify this point, since there are few homologs of vertebrate Eph-interacting proteins on the current WormBase list of vab-1 genetic interactors (https://www.wormbase.org).

6.3.2 Insects The Drosophila genome contains one single Eph receptoreencoding gene, and it is expressed in developing axons. Initially, its GoF did not lead to any overt axon guidance phenotypes (Scully et al., 1999). More insights into Eph signaling came from the analysis of the only Drosophila ephrin, which is predicted to have two transmembrane domains. Its RNAimediated LoF and GoF produces ventral nerve cord axon guidance phenotypes consistent with a function in axon repulsion (Bossing and Brand, 2002). However, Eph LoF argues against any function in the embryonic nervous system and instead suggests a requirement for Eph in the normal development of axons that form the mushroom body, a brain structure responsible for olfaction-related learning. These axons coexpress ephrin and Eph, suggesting their function in axoneaxon interaction (Boyle et al., 2006). Similarly, in the olfactory receptor neurons of the moth Manduca sexta, ephrin and Eph are coexpressed in complementary fashion, and exogenous Eph and ephrin inhibit the axon growth of explanted neurons (Kaneko and Nighorn, 2003). Recent experiments suggest that Drosophila Neurexin contributes to ephrin function in the developing visual system, but these findings have not been confirmed by ephrin genetic LoF experiments (Liu et al., 2017). As for C. elegans, Drosophila has many things to offer as a genetic model organism. Unfortunately, in the context of ephrin/Eph signaling, its classical genetics have been underused, and sometimes are in conflict with RNAi-based data. However, the specific Eph phenotypes are an excellent starting point for an enhancer screen designed to identify novel effectors and interactors.

6.4 Binary ephrin/Eph signalingdpathfinding 6.4.1 Peripheral pathfindingdlimb bud innervation One of the best understood axon guidance decisions is a binary choice made at the base of the vertebrate limb by spinal motor neuron axons extending from motor neurons residing in the lateral motor column (LMC, Fig. 6.5). Upon limb entry, lateral LMC axons select a dorsal limb trajectory, whereas medial LMC axons select a ventral one. Cellular-level experiments argued that this decision was made in response to short-range nondiffusible signals (Ferns and Hollyday, 1993). In line with these predictions, the molecules that mediate this choice are symmetrically deployed ephrins and Ephs. GoF and LoF experiments show that lateral LMC axons expressing EphA receptors are repelled into the dorsal limb from ephrin-As in the ventral limb mesenchyme. Similarly, medial LMC axons expressing EphB receptors are repelled from the ephrin-Bs in the ventral limb mesenchyme (Helmbacher et al., 2000; Kania and Jessell, 2003; Luria et al., 2008). This pleasing molecular symmetry is a platform for more complex Eph/ephrin interactions and signaling paradigms. LMC neurons also express ephrins, which attenuate some of the LMC neuron Eph fwd signaling (see Section 2.1.2). Ephrin-As expressed in the ventral limb, mesenchyme may also be cis-attenuated by EphA4, which is also expressed there. This idea is supported by analysis of a mutation, which blocks a protease cleavage site near the transmembrane domain of EphA4, resulting in overall higher EphA4 protein levels in the ventral limb and increased incidence of lateral LMC axons entering the ventral limb. This is, perhaps, a consequence of increased ventral limb mesenchyme ephrin-A attenuation by EphA4 and therefore less repulsion of lateral LMC axons attempting to aberrantly extend there (Gatto et al., 2014). In addition to fwd Eph signaling, the growth of lateral LMC axons into the dorsal limb is promoted by attractive rev signaling from EphA4 in the dorsal limb mesenchyme to ephrin-A5 on lateral LMC axons (Dudanova et al., 2012; Marquardt et al., 2005). Since ephrin-As lack an intracellular domain, how is the response to EphA4 in the limb transduced into lateral LMC axons? C-Ret, expressed by lateral LMC axons, appears to fulfill that function by associating with ephrinA (see also Fig. 6.3B). Additionally, it allows lateral LMC axons to respond to limb-derived GDNF, and the activation of the GDNF:c-Ret pathway in the presence of EphA4:ephrin-A binding results in enhanced accuracy of dorsal limb trajectory selection (Bonanomi et al., 2012; Dudanova et al., 2010). Genetic evidence suggests that this rev signaling pathway involves the seven-pass cadherin Celsr3, which associates with ephrin-A2, ephrin-A5, and c-Ret, and the knockout of which decreases EphA:ephrin-A signaling responses in LMC neurons, disrupting their in vivo guidance (Chai et al., 2014). In addition to motor axons, vertebrate spinal nerves also contain sensory axons, and these two classes of axons interact with each other via Eph and ephrin fwd and rev signaling. Sensory axon tracking along preexisting motor axons relies on rev signaling from EphA3 and EphA4 in motor axons to ephrin-As in sensory axons, which induces attraction. Loss of EphA3 and EphA4 in motor axons causes sensory axons to be repelled from motor axons, suggesting that axon tracking (also called axon bundling, or fasciculation) involves a balance of positive and negative signals (Wang et al., 2011).

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FIGURE 6.5 Model of Eph/ephrin signaling pathways guiding spinal motor axons in the limb. Schematic shows cross section through the spinal cord and limb of mouse or chick, with lateral motor column (LMC) neuron cell bodies in the ventral spinal cord. Lateral LMC neurons are colored in green and project to the dorsal limb (d), whereas medial LMC neurons are in red and project to the ventral (v) limb. The Eph/ephrin signaling pathways operating in the two axonal populations as they enter the limb mesenchyme (circle) are highlighted in the respective colors. Thus, coloring in this schematic differs from the other figures of this chapter. Lateral LMC axons express mainly EphA4 and are repelled from ephrin-A2 and ephrin-A5 expressed in the ventral limb. They also express ephrin-A5 that mediates attraction to the dorsal limb mesenchyme expressing EphA4. EphA4 and ephrin-A5 are present in different membrane domains so that they are able to signal in parallel. In contrast, the high level of ephrin-B2 expression is able to attenuate the potential signaling from dorsal limb ephrin-Bs (see later) to EphBs in these neurons. Lateral LMC axons also express Neogenin and other attractive netrin1 receptors. Exposure to ephrin-A increases the levels of Neogenin in the lateral LMC growth cones, increasing their preference for growth on netrin1. GDNF expression in the dorsal limb activates the Ret receptor, which in the presence of EphA4:ephrin-A5 signals increases the attraction of these axons for the dorsal limb mesenchyme. Medial LMC axons express mainly EphB-class receptors and are repelled from ephrin-B2 expressed in the dorsal limb. Expression of ephrinAs attenuates the potential signaling from the limb mesenchyme ephrin-As to trace expression of EphAs in this population. Unc5c, the repulsive netrin1 receptor, forms a molecular complex with EphB2, and coexposure to netrin1 and ephrin-B2 results in synergistic activation of Src family kinases (SFKs) and repulsion from the dorsal limb. Main references: Helmbacher, F., Schneider-Maunoury, S., Topilko, P., Tiret, L., and Charnay, P. (2000). Targeting of the EphA4 tyrosine kinase receptor affects dorsal/ventral pathfinding of limb motor axons. Development 127, 3313-3324; Kania, A., and Jessell, T.M. (2003). Topographic motor projections in the limb imposed by LIM homeodomain protein regulation of ephrin-A:EphA interactions. Neuron 38, 581-596; Kramer, E.R., Knott, L., Su, F., Dessaud, E., Krull, C.E., Helmbacher, F., and Klein, R. (2006). Cooperation between GDNF/Ret and ephrinA/EphA4 signals for motor-axon pathway selection in the limb. Neuron 50, 35-47. https://doi.org/10.1016/j.neuron.2006.02.020; Luria, V., Krawchuk, D., Jessell, T.M., Laufer, E., and Kania, A. (2008). Specification of motor axon trajectory by ephrin-B:EphB signaling: symmetrical control of axonal patterning in the developing limb. Neuron 60, 1039-1053. https://doi.org/10.1016/j.neuron.2008.11.011; Marquardt, T., Shirasaki, R., Ghosh, S., Andrews, S.E., Carter, N., Hunter, T., and Pfaff, S.L. (2005). Coexpressed EphA receptors and ephrin-A ligands mediate opposing actions on growth cone navigation from distinct membrane domains. Cell 121, 127-139. https://doi.org/10.1016/j.cell.2005.01.020; Poliak, S., Morales, D., Croteau, L.P., Krawchuk, D., Palmesino, E., Morton, S., Cloutier, J.F., Charron, F., Dalva, M.B., Ackerman, S.L., Kao, T.J., and Kania, A. (2015). Synergistic integration of Netrin and ephrin axon guidance signals by spinal motor neurons. eLife 4. https://doi.org/10.7554/eLife.10841; Croteau, L., Kao, T. and Kania, A. (2019) Ephrin-A5 potentiates netrin-1 axon guidance by enhancing Neogenin availability. Sci Rep 9, 12009 https://doi.org/10.1038/s41598-019-48519-0

Layered on top of the complex motor axon fwd and rev ephrin/Eph signaling, and its cis attenuation, are the interactions between ephrin:Eph signaling and Netrin signaling pathways. Netrin is expressed in the dorsal limb mesenchyme, and the lateral LMC motor axons that traverse it express receptors that enable attraction to Netrin, whereas the medial LMC motor axons that avoid the Netrin domain express the Unc5 class of repulsive Netrin receptors. These distinct Netrin receptors are

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required for LMC axon attraction to, and repulsion from, Netrin. This raises the question: What happens when LMC axons are exposed to Netrin and ephrins simultaneously? It turns out that ephrins synergize with Netrin so that even at very low concentrations of both ligands, where either one has no effect on LMC axon guidance, these cues together robustly guide LMC axons. This enhanced sensitivity correlates with the perdurance of SFK activation by ephrin and Netrin, which may be the result of EphB2-Unc5c receptor complex formation and likely increases the fidelity of the dorsal versus ventral trajectory LMC motor neuron axon guidance choice (Poliak et al., 2015). The study of LMC axons at the base of the limb has extended the biochemical studies of ephrin:Eph signaling (Section 2.2), but it has also revealed novel principles of Eph signaling, including cooperation and synergy in the collaboration between Eph and non-Eph receptors. These ideas have not yet been validated outside of motor neurons, but since Netrin and ephrin receptors are expressed by other developing neurons, it is possible that such interactions between these signaling pathways are generally applicable principles of axon guidance. Furthermore, unpublished data suggest that synergy between attractive Netrin receptors and ephrin-As could be a consequence of ephrin-A upregulation of Netrin receptor expression (Croteau et al., 2019).

6.4.2 Pathfinding in the spinal cord Axons that cross the midline of the vertebrate nervous system to link its two sides also rely on Eph/ephrin signaling. The development of descending axon trajectories from the cortex to the spinal cord, and those that link left and right spinal motor circuits that coordinate locomotion, depends on Eph/ephrin signaling. The corticospinal tract (CST) axons from the motor cortex decussate (cross the midline) in the medulla and innervate the contralateral spinal cord; however, mice with an EphA4 mutation have a hopping gait, which is most likely caused by some CST axons remaining in the ipsilateral medulla and projecting to the ipsilateral spinal cord (Dottori et al., 1998). One explanation for this phenotype is that ephrinB3 present at the midline of the spinal cord normally prevents EphA4-expressing CST axons from entering the midline through fwd signaling (Yokoyama et al., 2001). Since EphA4 and ephrin-B3 are also required in local spinal circuits that control walking (Kullander et al., 2003), the extent of the contribution of aberrant ipsilateral CST projections to the hopping phenotypes of EphA4 mutants was not clear. Forebrain-specific EphA4 knockouts have abnormal ipsilateral spinal CST projections, but normal spinal-level reflexive locomotion. On the other hand, voluntary locomotor behaviors that require the movement of a single limb resulted in bilateral movements in these mutant mice. These data suggest that the ipsilateral spinal innervation of EphA4-mutant CST axons can contribute to abnormal behaviors, but only in certain contexts (Serradj et al., 2014). Extending these data in the context of signaling mechanisms downstream of EphA4 and cellular interactions at the spinal cord midline, cortex-specific knockout of the gene encoding the EphA4 Ras GAP a-chimaerin showed abnormal ipsilateral spinal cord CST axons, similar to those seen in EphA4 mutants. Additionally, a spinal corde specific loss of a-chimaerin resulted in abnormal development of the spinal midline, reminiscent of tissue boundary defects seen in Eph LoFs (Katori et al., 2017), and highlighted the potential importance of spinal EphA4 in midline development. The early development of the CST pathway depends on EphB1, such that mice mutant for this gene have CST axons inappropriately crossing the midline of the ventral forebrain via the anterior commissure (AC) (Section 4.4.2) and a drastic reduction of CST axons that reach the more caudal CST waystations. The transcription factor Fezf2, which controls the identity of CST projection neurons, controls the expression of EphB1, and phenotypes in Fezf2 mutants are similar to those seen in EphB1 mutants, revealing the transcriptional logic that controls the development of the longest axons in vertebrates (Lodato et al., 2014). In parallel to axons descending from the brain to the spinal cord, at least some ipsilateral ascending axonal pathways are also guided by the Eph/ephrin system in mice. For some of these connections, the Zic2 transcriptional program that controls laterality in the visual system (Section 4.4.1) also functions to control the expression of the EphA4 receptor in commissural neurons. Many ipsilateral ascending projection neurons express Zic2, which activates EphA4 expression and facilitates their repulsion from ephrin-B3 present at the spinal midline. EphA4 deletion in the spinal cord results in these axons crossing the spinal midline and is phenocopied by a mutation in which only the intracellular domain of EphA4 is deleted, suggesting a fwd signaling mode. Underscoring the possibility of interactions between these ascending axons with descending CST axons, spinal cordespecific deletion of EphA4 also affects the guidance of CST axons (Escalante et al., 2013; Paixao et al., 2013). A different set of spinal commissural axons is thought to be repelled from ephrin-B3 at the ventral spinal midline through EphB fwd signaling (Kadison et al., 2006), a process that may require the Syk kinase such that a change in its phosphorylation state switches the response of ephrin-B-stimulated EphBs from repulsion to attraction (Noraz et al., 2016).

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6.4.3 Pathfinding in the brain stemdauditory system Axons of the mammalian ventral cochlear nucleus (VCN) located in the auditory brain stem also must cross the nervous system midline. These neurons project axons contralaterally to the medial nucleus of the trapezoid body (MNTB), and this arrangement is essential for localization of an auditory stimulus through the computation of an interaural differential. In this context, genetic analysis in mice suggests that EphB2:ephrin-B2 rev signaling prevents the innervation of the ipsilateral MNTB by VCN axons (Hsieh et al., 2010). A separate genetic study also implicates ephrin-A2 and ephrin-A5 in this event (Abdul-Latif et al., 2015). Upstream of the cochlear nuclei, spiral ganglion sensory neurons (SGNs) innervate cochlear hair cells, which detect auditory stimuli. En route to the cochlea, SGN axons traverse the otic mesenchyme, where the transcription factor Pou3f4 directly controls the expression of EphA4. Genetic LoF and GoF experiments suggest that EphA4 binding to ephrin-B2 on SGN axons promotes their fasciculation (Coate et al., 2012). In the cochlea, SGN axons make a binary target choice: type 1 SGNs target inner hair cells (IHCs), and type 2 SGNs target in outer hair cells (OHCs). OHCs, but not IHCs, express ephrin-A5, and its loss results to inappropriate innervation of OHCs by type 1 SGNs. The repulsion of SGN axons from OHC depends on EphA4 fwd signaling and possibly involves signal transduction through ephexin1 (Defourny et al., 2013). So far, Ephrin/Eph signaling appears to be a principal effector of binary and discrete axon guidance events in the auditory system. Although some of these guidance events are emerging as similar to those in the visual system (Sections 5.2 and 5.3), the mechanisms that assemble other auditory connections remain to be discovered. For example, the tonotopic arrangement of afferent auditory axons in the inferior colliculus has been proposed to also depend on ephrin gradients (Cramer and Gabriele, 2014).

6.4.4 Central pathfinding 6.4.4.1 Optic chiasm Arguably one of the best understood axon guidance decisions at the midline of the nervous system is that executed by retinal ganglion neuron (RGC) axons at the optic chiasm. Most RGC axons traverse the optic chiasm apparently unimpeded, but those that express a high level of EphB1 are repelled from ephrin-Bs expressed there and form ipsilateral projections. This decision controls the ipsilateral versus contralateral RGC innervation ratio that determines the extent of binocularity in different vertebrate species (Williams et al., 2003). Mouse RGCs express all three EphB receptors, but GoF and receptor chimera experiments confirm that the EphB1 receptor is most likely the principal EphB controlling RGC laterality. For example, despite similar ephrin-B binding constants (see Section 1.2), EphB2 cannot functionally replace EphB1 (Petros et al., 2009). The restriction of EphB1 expression to ipsilaterally projecting RGCs occurs via the expression of the Zic2 transcription factor (Garcia-Frigola et al., 2008). However, an additional mechanism controlling EphB1 expression in RGCs is suggested by the analysis of mice mutant for the gene encoding the transmembrane glycoprotein Ten-m/Odz/teneurin Ten-m3, which results in lowered EphB1 expression in RGCs and, consequently, an increased number of ipsilateral RGC axons (Young et al., 2013).

6.4.4.2 Corpus callosum and anterior commissure The corpus callosum (CC) is the principal connective structure linking the two cerebral hemispheres. It is organized topographically such that axons from the medial and lateral regions of the cortex traverse its dorsal and ventral aspects, respectively. This arrangement is in part the result of medial axons expressing ephrin-A5 and repelling EphA3-expressing lateral axons prior to crossing the midline (Nishikimi et al., 2011). The transcription factor Satb2 is one of the principal determinants of callosal projections, since its loss results in their absence. This phenotype can be in part rescued by overexpression of EphA4, despite the lack of callosal phenotypes in EphA4 mutant mice (Srinivasan et al., 2012). Following midline crossing by CC axons, their expression of ephrin-B1 silences responsiveness to Semaphorin 3C through its receptor Neuropilin1, as suggested by interactions between ephrin-B1 and Neuropilin1. Surprisingly, this may occur independently of Eph receptor activation (Mire et al., 2018). Compared with the CC, the AC is a relatively minor connection that also links the cerebral hemispheres. However, it was one of the first axon pathways to be shown to rely on Eph signaling and provided genetic evidence for ephrin-B rev signaling: EphB2 null mice display AC defects, whereas mice expressing an EphB2 lacking an intracellular domain do not (Henkemeyer et al., 1996). Similarly, although EphA4 is required for AC formation, it does so in a kinase and SAM domain-independent manner, further implicating Eph:ephrin signaling in CC formation (Kullander et al., 2001).

136 PART | I Formation of axons and dendrites

6.5 Proportional ephrin/Eph signalingdmapping In the binary signaling mode described earlier, ephrin/Eph signals are distinguished by step-like differences (present vs. absent, one ephrin vs. two different ephrins, etc.). As an example of a binary response, axons may avoid or prefer a region of the nervous system that contains an ephrin or an Eph. In contrast, in the proportional signaling mode, the same cues are distinguished incrementally, and a graded response is generated so that a growth cone can be guided with high precision inside the target region. Proportional ephrin/Eph signaling occurs in the mapping of the main olfactory system and in the formation of topographic projections.

6.5.1 Olfactory wiring The mammalian nose contains the main olfactory epithelium, recognizing general odorants and the vomeronasal organ recognizing pheromones. Apical and basal layers of the vomeronasal epithelium express different classes of vomeronasal receptors and project to the anterior and posterior accessory olfactory bulb, respectively. This segregation is accomplished by binary EphA6:ephrin-A5 attractive rev signaling (Knoll et al., 2001), and ephrin-A5 LoF results in a corresponding mistargeting of apical projections. Sensory discrimination in the main olfactory system is based on olfactory sensory neurons (OSNs) expressing only one of the more than a thousand different odorant receptor (OR) genes. OSNs expressing the same OR, though randomly scattered on the epithelial plane, converge their axons into one or few OR-specific stereotyped target points (glomeruli) of the olfactory bulb. The expression of ephrin-A5 and EphA5 is counterregulated in OSNs at incrementally varying levels correlated with the OR type (Cutforth et al., 2003; Serizawa et al., 2006). Thus, OSNs of each ephrin-A5/EphA5 ratio are randomly scattered like their corresponding ORs, and ephrin-A5 and EphA5 do not form spatial countergradients. But when ordered according to the ephrin/Eph expression levels, smooth countergradients become visible. The glomeruli on the bulb are also not systematically arrayed according to ephrin/Eph expression levels, arguing against a role of ephrins and Ephs in their stereotypic positioning. Mosaic deregulation of ephrin-A in a random half of OSNs leads to glomerular segregation of otherwise homotypic fibers. Apparently, the ephrin/Eph system, in this case, provides a wide range of ratiometric axon identity labels, which are used together with several other surface molecules for self/noneselfdiscrimination to support glomerular segregation, but not for mapping according to similarity.

6.5.2 Retinotectal/retinocollicular projection 6.5.2.1 Mechanisms of mapping along the anterioreposterior axis The retinotectal projection is the topographic, i.e., neighborhood-preserving projection of retinofugal axons onto the midbrain’s optic tectum (corresponding to the superior colliculus [SC] in mammals, Fig. 6.6A). This projection has played a paradigmatic role for axon guidance research in general, since it formed the basis of Sperry’s classical regeneration experiments that gave rise to the influential hypothesis that axon guidance was chemoaffinity based (Sperry, 1963). In seminal in vitro experiments, Bonhoeffer and Huf (1982) showed that tectal cell membranes indeed display a graded biochemical guidance cue for retinal axons. Neuronal activity, in contrast, is dispensable for primary mapping, although it seems to be required for the final compaction of termination zones (Benjumeda et al., 2013). Identifying the graded tectal cues as ephrin-As, and their retinal detectors as countergraded EphAs (Cheng et al., 1995; Drescher et al., 1995), first established the role of the ephrin/Eph system in axon guidance. In artificial striped substrates that allow for clear assessment of binary axon guidance choices (“stripe assays” (Walter et al., 1987)), ephrin-A5 repels retinofugal axons through fwd signaling irrespective of their topographical identity and is therefore insufficient to explain topographic mapping of these projections onto the tectum. Only at very low cue concentrations is the sensitivity of nasal axons completely lost due to cis attenuation (section 2.1.1). But, in addition to these expression patterns of EphAs and ephrin-As, there are also countergradients of ephrin-As in the retina and EphAs in the tectum, enabling rev signaling, which in stripe assays mediates topographically nondifferential repulsion (Hornberger et al., 1999; Rashid et al., 2005). If, in vitro, ephrin-As and EphAs are presented in alternating stripes, topographically differential choice behavior emerges, where nasal axons prefer ephrin-A, and temporal axons prefer EphA, indicating a need for concurrent fwd and rev signaling for mapping (Gebhardt et al., 2012). These conclusions are corroborated in vivo by studies of mouse mutants (Feldheim et al., 2004; Frisen et al., 1998; Pfeiffenberger et al., 2006; Rashid et al., 2005). Deleting fwd signaling components results in the formation of ectopic nasal terminations shifted anteriorly, and deleting components of rev signaling mostly leads to ectopic temporal termination zones shifted posteriorly (Fig. 6.6C).

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FIGURE 6.6 Proportional ephrin/Eph signaling in topographic mapping. (AeC) Topographic retinotectal mapping. (A) The retinal nasotemporal (n, t) and dorsoventral (d,v) axes are projected onto the posterioreanterior (p,a) and lateralemedial (l,m) axes of the tectum. Ephrin-As and EphAs are expressed as countergradients on both retina and tectum (color shading). The expression of the B-system (mouse) along the dorsoventral axis is indicated by the colored wedges. Inset: Molecular underpinnings of exemplary midtemporal and midnasal fiber terminals interacting with an anterior target cell (fiber/target [FT] interaction) and between each other (fiber/fiber [FF] interaction). Horizontal bars indicate cell membranes in appropriate color shades. Both fwd and rev interactions act repulsively () on the fiber terminal, cis interactions attenuate (x) signaling. (B) According to a current model (Gebhardt et al., 2012), a mapping fiber terminal (magenta growth cone) seeks the minimum of an FT potential originating from ephrin/Eph fwd and rev interactions with the target and of an FF potential, which originates from corresponding interactions with other terminals and which coemerges upon fiber sorting. The schematic shows the FF “potential well” at the end of the mapping process. The minimum is reached when total fwd and total rev signals are balanced, corresponding to a map position displaying an ephrin-A/EphA ratio (color shading) similar to the terminal’s site of origin. (C) Projection phenotypes of major mouse deletion mutants (gray oval: superior colliculus [SC]; blue and red circles: temporal and nasal termination zones, respectively). Murine ephrin-A/EphA expression patterns are shown below, with red X indicating a deletion (modified from Weth et al., 2014). (D) Topographic thalamocortical maps. Left: Schematic drawing of a horizontal section through the left embryonic forebrain. An ephrin-A gradient in the ventral telencephalon sorts thalamic axons expressing graded amounts of EphA4 for anterior versus posterior cortical destinations (interareal topography). Right: Schematic drawing of a coronal section through the embryonic forebrain at later stages. Intraareal topography, exemplified for the visual projection from the dorsal geniculate nucleus (dLGN) to the primary visual cortex (V1), relies on the indicated ephrin-A/EphA gradients for mapping.

More key elements of the topographic mapping mechanism were discovered when constant amounts of EphA3 were knocked into a random population of RGCs (Brown et al., 2000). This led to a map duplication, with mutant axons projecting anteriorly and displacing wild-type axons to a second posterior map, which recalls indications of pronounced mapping plasticity observed in older regeneration studies (Goodhill and Richards, 1999). Notably, these observations can be interpreted in terms of ephrin/Eph-mediated FF interactions, which eventually supersede the FT interactions (Gebhardt et al., 2012). Albeit less well noticed, Bonhoeffer originally had found similar discriminatory power of RGC growth cones for other retinal axons as for tectal membranes (Bonhoeffer and Huf, 1985). The role of FF interactions has also gained support from the effects of conditionally deleting retinal ephrin-A5 (Suetterlin and Drescher, 2014), which revealed repulsion of temporal axons by ephrin-A5 on nasals. In sum, topographic mapping seems to be based on six dimensions of concurrent ephrin/Eph signaling (cis, FF, FT, each with fwd and rev signaling Fig. 6.6A), which, according to computational modeling (Gebhardt et al., 2012), can be summarized by one straightforward concept: Growth cones seek a balance of total fwd to total rev signaling. Mechanistically, this involves the growth cone sensing the ephrin/Eph ratio in its vicinity, which includes target cells and other

138 PART | I Formation of axons and dendrites

terminals, and responding to it with graded avoidance in proportion to the overall dissimilarity. The crucial importance of ratiometric sensing in the proportional signaling mode (Gebhardt et al., 2012; Reber et al., 2004) has been highlighted by the discovery of a unique mechanism of ratio conservation upon signal modulation, termed coadaptation (Fiederling et al., 2017). EphAs and ephrin-As adapt to fwd and rev signaling, respectively, potentially confounding quantitative signaling. Ratiometric quantification is rescued in this situation by coadaptation, which implies strict coregulation of both sensors independent of whether both or only one has been activated. Mechanistically, coadaptation is based on clathrin-mediated endocytosis, presumably triggered by reorganization of membrane microdomains downstream of signaling, which, based on computational modeling, has been suggested to eventually increase cis attenuation. As a side note, it should be mentioned that due to adaptation, retinal axons can grow continuously on homogeneous ephrin-A substrates, although with efficiencies depending on the topographic origin and ephrin concentration (Hansen et al., 2004). Differential outgrowth, however, does not involve any nonlinear choice response of the growth cones and should, therefore, not be confused with topographically differential targeting.

6.5.2.2 Mechanisms of mapping along the dorsoventral axis Retinotopic mapping along the dorsoventral axis has received much less attention than the anterioreposterior projection. Ephrin-Bs and EphBs are countergraded along both the retinal dorsoventral and collicular mediolateral axes (Fig. 6.6A) of the mouse (Hindges et al., 2002). Axons expressing high levels of each guidance sensor project to targets expressing high levels of cognate signals, suggesting attractive FT interactions. Similar conclusions were drawn for Xenopus (Mann et al., 2002). The retinofugal axons of EphB2/B3 LoF mutants show a preponderance of lateral over medial branches, which are equally distributed in the wild type. Similar phenotypes were shown in a mouse knock-in of kinase-dead EphB2, which was interpreted as an indication of fwd signaling (Hindges et al., 2002). However, there are several hints that important factors might still be missing in this model. Deleting ephrin-B1, the only collicular ephrin-B, has only minor effects (Thakar et al., 2011). Stripe assays with chick retinal axons reveal repulsive instead of attractive effects for ephrinB/EphB fwd and rev signaling (unpublished observations, F.W.). Therefore, repulsive FF interactions among axon branches should be considered as effectors of termination zone positioning along the dorsoventral axis. Finally, all of the effects described earlier observed in ephrin-B/EphB mutants relate to axon side branches. Substantial mediolateral fiber preordering is visible in the fan-shaped innervation via the brachium of the SC, which seems, however, not to be affected by the ephrin-B/EphB mutants. Notably, axonal ephrin-Bs are needed for tectal receptive field plasticity mediated by synaptic activity and terminal arbor dynamics in Xenopus (Lim et al., 2010), indicating that the observed effects on the side branches might relate more to terminal zone refinement than to primary axon guidance.

6.5.2.3 Computational modeling The vast body of experimental evidence on retinotopic mapping has spurred many computational models, but we chose to focus here only on the most comprehensive ones (Fraser and Perkel, 1990; Gebhardt et al., 2012; Weber et al., 1997; Willshaw, 2006; Yates et al., 2004). In the era of classical embryology, models mainly addressed the conflict between rigid chemoaffinity and the pronounced long-term plasticity of mapping observed upon regeneration from microsurgical manipulations. Thus, regrowing axons from a half-retina map smoothly onto a full tectum, a full retina forms a compressed map on a half-tectum, and a half-retina forms a normally oriented map on a mismatching half-tectum or even an inverted map, if the other half-tectum is left occupied with the original innervation (for review, see Goodhill and Richards, 1999). Growth cone targeting can be modeled as a potential energy minimization process, like the rolling of a ball to the bottom of a bowl. The potential is high when the growth cone is far from its target, and it is minimal at the target site (Fig. 6.6B). For rigid FT chemoaffinity, this potential energy results from the differential interactions of the fiber terminals with topographic cues on the target (FT potential). It increases with higher topographic mismatch between the terminal and its current position. The FT potential is needed to anchor the map and to coarsely presort the fibers. All comprehensive models agree, however, that, for understanding mapping plasticity on suboptimal, or even nonmatching, targets, a second, dynamically evolving potential is needed, which emerges from interactions among the terminals (FF potential). This potential has often been explained by a combination of Hebbian plasticity and nonspecific short-range repulsion/ competition among the fiber terminals, preventing their convergence into one common target point. During development, spontaneous activity waves sweep over the retina, resulting in synchronization of the activity of neighboring RGCs. Thus, the Hebbian principle of “fire together, wire together” could provide a neighborhood preserving drive for topographic

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mapping even without appropriate target cues. However, mapping plasticity is also observed in the absence of activity (Meyer and Wolcott, 1987). Therefore, the sorting drive of the FF potential must originate from activity-independent, chemical cues on the fiber terminals, which encode topographic information. Classical models, building on these principles, differed in their detailed assumptions, but all successfully reconciled rigidity and plasticity. Plasticity, in the molecular era, gained renewed interest when genetic manipulation of a chemoaffinity cue (EphA3) led to comparable plastic remapping ((Brown et al., 2000), see Section 5.2.1), prompting novel ephrin/Eph-based comprehensive models. A model exclusively relying on observed ephrin/Eph interactions for FT as well as for FF interactions can in fact explain rigid chemoaffinity in vitro and in vivo and also mapping plasticity (Gebhardt et al., 2012). The model can be summarized by the rule that the growth cones minimize their guidance potential by seeking a balance of total fwd and rev signaling. The latest implementation also includes the coadaptation mechanism (Fiederling et al., 2017; see Section 5.2.1).

6.5.3 Retinogeniculate projections In mammals, retinofugal axons send collaterals to the thalamic dorsal lateral geniculate nucleus (dLGN). Input from both eyes, which is needed for binocular vision, is segregated by eye in the dLGN, but the two eye-specific topographic maps are registered (¼aligned). Registration and segregation involve the ephrin/Eph system in ways that depend upon the extent of binocularity. In mice (low binocularity), ipsilaterally projecting RGCs residing in the ventrotemporal retinal crescent extend axons that terminate in a segregated patch in the dorsomedial dLGN, and their mapping and segregation occur prenatally. Ipsiand contralateral RGC axons use the same ephrin/Eph gradient systems for mapping, and both get misplaced in ephrinA2,3,5 triple knockouts (Pfeiffenberger et al., 2005). Notably, however, these axons read out the ephrin gradients differently, as they align in antiparallel orientation along the anterioreposterior target axis for functional registration (Haustead et al., 2008). This might be based on FT versus precisely timed FF interactions in contra- and ipsilateral RGC axons, respectively. Segregation, however, is retained in the ephrin-A2,3,5 triple mutants. It turns out to be activity dependent and relies on the correlation of spontaneous retinal activity, which differs among the two eyes. In carnivores and primates with frontal eyes and a high degree of binocularity, RGCs in the temporal retina project axons ipsilaterally, and the nasal RGCs project contralaterally, in each case using a modified retinal gradient system. In humans, EphAs are high in the central retina and fade out peripherally, whereas ephrin-As are countergraded, allowing for the parallel mapping of ipsi- (temporal) and contralateral (nasal) axons, which is needed for functional binocularity in animals with frontal eyes (Lambot et al., 2005). In the ferret, with a high degree of binocularity, mapping and segregation into eye-specific layers occur pre- and postnatally, respectively. Postnatal EphA overexpression retains topographic registration, but it destroys layering, indicating a second, later use of the ephrin/Eph system for spatial layer segregation. This mode of ephrin/Eph signaling, however, depends on spontaneous (not necessarily correlated) retinal activity (Huberman et al., 2005).

6.5.4 Thalamocortical projections Thalamic relay nuclei typically maintain reciprocal topographic connections with their respective cortical areas. In addition, the relationship between thalamus and cortex is globally topographic. Both intra- and interareal topographic mapping deploys the ephrin/Eph system. Matching ephrin-A/EphA countergradient systems have been found in the dLGN and the primary visual cortex (V1) (Fig. 6.6D) as well as in the thalamic ventrobasal (VB) complex and the associated somatosensory cortex (S1). In accordance with these gradient patterns, LoF of ephrin-A5 results in a deformation of the VB > S1 thalamocortical map (Vanderhaeghen et al., 2000), whereas the ephrin-A2,3,5 triple knockout mouse reveals a distortion in the dLGN > V1 map (Cang et al., 2005). Manipulation of EphA7 expression in deep layer neurons of S1 shifts the terminals of their corticothalamic projections in the VB complex (Torii and Levitt, 2005). This suggests that thalamocortical and corticothalamic intraareal topographic projections rely on similar ephrin/Eph-based mapping mechanisms as the paradigmatic retinotectal projection. Global thalamocortical topography is supported by an earlier anterioreposterior ephrin-A gradient in the horizontal plane of the ventral telencephalon (Fig. 6.6D), which is on the pathway of ascending thalamocortical axons and fans them out for anterior versus posterior cortical destinations through repulsion (Dufour et al., 2003). Interestingly, this system, used for axon sorting instead of termination mapping, does not appear to require EphA countergradients but instead the guidance cues Netrin-1 (Powell et al., 2008) and Slit2 (Bielle et al., 2011) and thus lacks major features of the topographic mapping systems.

140 PART | I Formation of axons and dendrites

6.5.5 Corticocollicular projections In addition to retinal inputs, the SC receives multimodal inputs form other brain structures such as the primary visual and somatosensory cortices, and these inputs are topographic and aligned with the retinocollicular map. Upon investigation of the S1 > SC and V1 > SC maps in Isl2:EphA3 knock-in mutants, in which the retinocollicular map is duplicated (see Section 5.2.1), it was found that the V1 projections split up and follow this duplication, whereas the S1 projections map normally (Triplett et al., 2012). This was interpreted as evidence for an activity-dependent (“fire together, wire together”) mechanism for directing the visual cortical projection and of a genetic ephrin/Eph-based mechanism for directing somatosensory cortical projections. The splitting of the V1 > SC projection is no longer observable when the correlated spontaneous retinal activity is disrupted. Alternatively, these results could also be explained by ephrin/Eph-based fwd and rev FF signaling mechanisms for V1 and S1 inputs, respectively. This is because in the Isl2:EphA3 knock-in mutants, the ephrin-A distribution on the fiber terminals should be transformed to a discontinuous, saw-toothed shape, with the potential to induce map duplication, whereas the EphA3 distribution should stay continuous and allow for a uniform map. The loss of splitting of the V1 projection upon experimental desynchronization of retinal activity would then be interpreted as a loss of resolution due to a lack of termination zone compaction. The genetic interpretation is more consistent with the results of Isl2-mediated overexpression of ephrin-A3 randomly in one-half of the RGC population (Savier et al., 2017), forcing the V1 projection to split up locally among normal and overexpressing retinal terminals.

6.6 Ephrins and Ephs in regeneration It is well established that, following spinal cord injury, one of the most potent signals preventing reinnervation and regeneration are the various protein components of myelin (Filbin, 2003). Given the role of ephrin/Eph signaling in growth cone collapse and repulsion in the developing nervous system, it is perhaps not surprising that ephrin-B3 can mediate at least some of the axon growth inhibitory functions of myelin in vitro (Benson et al., 2005). These observations are also corroborated in vivo by the observation that mice mutant for ephrin-B3 recover more readily from certain kinds of spinal cord and optic nerve injuries (Duffy et al., 2012). Similarly, blockade of ephrin-A5 in mouse stroke models results in enhanced motor and sensory cortex axonal sprouting, which is further potentiated by forced limb usage (Overman et al., 2012). The development of new strategies to interfere with Eph receptor function will be one of the ways to exploit these observations in the search for new axon injury therapies.

6.7 Perspectives and open questionsd“curtain down and nothing settled” Eph/ephrin signaling may appear to be a mature field; however, there remain many important questions about this pathway, particularly since it is employed in a wide variety of normal and pathological processes. Arguably, studying Eph/ephrin signaling in the context of axon guidance has begun to answer these questions, and this line of investigation has generated very useful tools and paradigms that pave the way toward a comprehensive general model of signaling and, more specifically, the logic of its deployment in the developing nervous system. One open question is the specificity of receptoreligand interactions. Some experiments suggest that Eph receptors of a particular class may be functionally equivalent, but this simplistic view is undermined by the observations that not all EphA or EphB receptors can functionally substitute for each other in axon guidance, no matter the level of their expression (e.g., Luria et al., 2008; Petros et al., 2009). Perhaps, part of the answer is in the different structural configurations of two related receptors complexed with the same ligand (Seiradake et al., 2013). These experiments suggest that different ephrineEph structures may engage different intracellular signaling effectors, but until all such proteins are discovered, the answer to this question will remain elusive. New signaling insights may also come from the use of next-generation imaging tools that allow for the visualization of Eph signaling in real time, for example, through the use of biosensors, or artificial (small molecule and optogenetic) activation of Eph receptors. One big advantage of biosensors is that they allow the visualization of Eph signaling dynamics with subcellular resolution, linking local cell biological- and molecular-level events. Intriguingly, these tools have yet to be extensively used in the context of neuronal growth cone guidance and axon targeting (Mao et al., 2018; Schaupp et al., 2014). Another avenue that merits further exploration is how the spatial organization of Eph receptors affects their downstream signaling transduction. This has been exploited in the context of cellecell adhesion and cancer invasiveness, raising the interesting possibility that changes in the adhesion of a growth cone to its substrate elicited by axon guidance cues may

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change the distribution of Eph receptors via sheer physical forces, which in turn may alter sensitivity to ephrins (Salaita et al., 2010). The question of cross-talk between axon guidance signaling pathways was raised many years ago, and although some of the simpler paradigms involving additive effects have been explored, more complex modes involving Eph receptors, such as synergistic interactions, remain poorly understood. Eph receptor signaling complexes include a wide variety of other unrelated receptors, raising the possibility that although Eph receptors can be activated by ephrins, the simple one ligandeone receptor class signaling mode may be an exception in an in vivo context (Marquardt et al., 2005; Poliak et al., 2015). Growth cones can perform guidance decisions in response to extremely small concentration differences, in the range of less than one molecule across the growth cone diameter on average (Rosoff et al., 2004). Responses based on such minute differences most probably necessitate Turing-like mechanisms of local autocatalysis and lateral inhibition in the signal transduction networks. The proportional mode, in addition to making such ultrasensitive guidance decisions, requires the growth cone to keep track of quantitative concentration ratios. How this is achieved in ephrin/Eph signaling is far from being understood. On the systemic level, the well-studied processes of limb bud innervation and retinotectal mapping might serve as paradigms for other ephrin/Eph-based guidance systems. Given the diversity of ephrin/Eph paralogs and signaling modes, a detailed analysis of protein expression patterns in vivo is of utmost importance and might in the future be improved by the use of genetic tags such as the SNAP-tag or the HaloTag. Improved in vitro assays, reconstituting the in vivo situation more realistically, such as using gradient substrates manufactured with microfluidic networks or systems based on supported lipid bilayers, will help to better single out the functions of all involved ephrin/Eph signaling channels, as will in vivo experiments with more specific conditional mutations. Eventually, computational modeling will be invaluable for an integrated understanding of how biological function emerges from complex ephrin/Eph signaling networks. In light of the ubiquitous use of the ephrin/Eph system in neural development, these approaches will help us to better understand the intriguing process of brain wiring in health and disease.

Acknowledgments Franco Weth is supported by the German Research Foundation (KSOP, DFG GSC 21). Artur Kania is supported by the Canadian Institutes of Health Research (MOP-97758 and PJT-153053) and Fondation IRCM. The authors would like to apologize to those colleagues whose work was omitted due to space limitations.

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Chapter 7

Axon guidance: SliteRobo signaling Katrine Iversen, Franc¸ois Beaubien, Janet E.A. Prince and Jean-Franc¸ois Cloutier Montreal Neurological Institute, McGill University, Department of Neurology and Neurosurgery, Montréal, QC, Canada

Chapter outline 7.1. Introduction 147 7.2. Slits and their receptors 147 7.2.1. Slit discovery and structure 147 7.2.2. Identification of the slit receptor robo 148 7.2.3. Slit and Robo interactions 149 7.2.3.1. Regulation of SliteRobo interactions 149 7.3. SliteRobo function in midline crossing 150 7.3.1. Spatial expression patterns of Slit and Robo 152 7.3.2. Posttranscriptional Robo regulation 152 7.3.3. Regulation of Robo protein expression at the midline152 7.3.3.1. Drosophila and vertebrate midlines 152 7.3.3.2. Caenorhabditis elegans midline 153 7.3.4. Regulation of Robo signaling at the midline in vertebrates 154 7.3.5. SliteRobo signaling for exiting the midline 155 7.4. Modulation of SliteRobo signaling 155 7.4.1. Transcriptional control 155

7.4.2. Regulation of SliteRobo signaling by metalloprotease cleavage 7.4.3. Regulation of SliteRobo signaling by ubiquitination 7.5. Signaling downstream of Robo 7.5.1. Rho family of small GTPases 7.5.2. Abelson tyrosine kinase 7.5.3. Actin-interacting proteins 7.6. Beyond the midline: additional roles for SliteRobo in the nervous system 7.6.1. Lateral positioning 7.6.2. Cell migration and cell polarity 7.6.3. Dendritic and axonal outgrowth and branching 7.7. SliteRobo contribution to axon targeting in a complex target field 7.8. Involvement of SliteRobo in disorders of the nervous system 7.9. Conclusion References

156 157 157 158 158 159 159 159 160 161 162 164 164 165

7.1 Introduction The regulation of axonal growth and guidance during nervous system development relies on a plethora of molecules expressed in the environment as well as on the surface of growing axons. Several of these molecules are secreted in the environment and can have an effect on axonal growth from a distance through the formation of gradients. The Slits belong to a family of potent secreted chemorepellents that can regulate the growth of axons. In addition, this family of molecules plays a critical role in regulating several other developmental processes, including cell migration, cell adhesion, and angiogenesis. In this section, we focus our discussion on the role of Slits in controlling axon guidance at the spinal cord midline, and we provide examples of the role of Slits in regulating additional biological processes in the developing nervous system.

7.2 Slits and their receptors 7.2.1 Slit discovery and structure Genetic screens in Drosophila, popularized in the past two decades of the 20th century, led to the discovery of a large number of molecules that play important roles in development. Included in this group of identified molecules are the canonical axon guidance molecule families. The Slit family of axon guidance molecules was first discovered in a genetic screen for embryonic patterning defects in Drosophila (Nussleinvolhard et al., 1984). Their role as axon guidance

Cellular Migration and Formation of Axons and Dendrites. https://doi.org/10.1016/B978-0-12-814407-7.00007-9 Copyright © 2020 Elsevier Inc. All rights reserved.

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molecules was further observed in genetic screens for commissural axon guidance pathfinding defects in Drosophila (Hummel et al., 1999; Seeger et al., 1993). Although the Slits were originally characterized in Drosophila (Nussleinvolhard et al., 1984), their homologs have also been characterized in Caenorhabditis elegans (Hao et al., 2001), Xenopus (Li et al., 1999; Chen et al., 2000), zebrafish (Yeo et al., 2001; Fricke et al., 2001; Hutson and Chien, 2002), chicken (Li et al., 1999; Holmes and Niswander, 2001), zebra finch (Warren et al., 2010), and mammals (Brose et al., 1999; Li et al., 1999; Holmes et al., 1998; Itoh et al., 1998; Marillat et al., 2002; Niclou et al., 2000; Wang et al., 1999). There are three homologous mammalian Slit family members (Slit1, Slit2, and Slit3), whereas invertebrates express a single Slit molecule (Brose et al., 1999; Holmes et al., 1998; Itoh et al., 1998; Li et al., 1999; Nakayama et al., 1998; Yuan et al., 1999b). Slits are large (w190 kilodaltons (kDa)) secreted glycoproteins containing four leucine-rich repeat (LRR) domains at their N-terminus followed by six epidermal growth factorsdsuch as motifs, laminin G domains (one in invertebrates and three in vertebrates), and a C-terminal cysteine-knot motif (Fig. 7.1A).

7.2.2 Identification of the slit receptor robo The Roundabout (Robo) gene was first identified in Drosophila in a genetic screen for genes regulating midline crossing in the CNS (Seeger et al., 1993) and later shown to encode an evolutionarily conserved transmembrane protein (Kidd et al., 1998a). Genetic (Kidd et al., 1999) and biochemical (Brose et al., 1999) evidence revealed that Slit can bind to Robo, which is expressed on commissural axons at the midline. Robo proteins are large (150e180 kDa) transmembrane type 1 receptors that belong to the immunoglobulin (Ig) superfamily (Fig. 7.1B). Three homologs of Robo (Robo1, 2, and 3 [also known as Rig-1]) are highly expressed in the nervous system and are found in many species including Drosophila (Simpson et al., 2000b; Kidd et al., 1998a), zebrafish (Lee et al., 2001; Challa et al., 2005), chick (Vargesson et al., 2001),

(A)

(B)

(C)

FIGURE 7.1 (A) Structure of the Slit proteins. Slits are large w190-kDa secreted glycoproteins containing four leucine-rich repeat domains at their Nterminus followed by six epidermal growth factors (EGF)elike motifs, one laminin G domain in invertebrates (three in vertebrates), and a C-terminal cysteine-knot motif. Slit is cleaved by a protease in a large N-terminal fragment and a shorter C-terminal fragment. (B) Structure of the Robo proteins. Contrary to Slit, the structure of Robo receptors varies more across different species. The Robos contain in their extracellular portion variable number of immunoglobulin (Ig)-like domains and fibronectin type III (FNIII) repeats and of up to four conserved intracellular motifs (catalytic activity [CC]). Two isoforms of Robo3 with distinct carboxy terminals are generated by alternative splicing. Robo3.1 transcript arises from the splicing out of intron 26 and intron 27 resulting in a 78 amino acids peptide, whereas retention of the intron 26 leads to the generation a 43 amino acid peptide in the case of Robo3.2. (C) Proposed mode of activation of the Robo receptor complex. Robo dimers are in a compact conformation close to the membrane. Slit binding induces a conformational change that relaxes the structure and leads to activation of signaling downstream of Robo.

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and mammals (Yuan et al., 1999a; Li et al., 1999; Brose et al., 1999). A unique Robo ortholog has also been identified in C. elegans (SAX-3) (Zallen et al., 1998). Structurally, the Robos are composed of five extracellular Ig-like domains, three fibronectin type III (FNIII) repeats, and of up to four conserved intracellular motifs without catalytic activity (CC) (Robo1/ 2 CC0-CC3, Robo3 CC0, CC2-3). A fourth Robo (also known as magic Roundabout) is also present in mammals and zebrafish, although its ability to directly bind to Slit remains a question of debate (Park et al., 2003; Suchting et al., 2005; Zhang et al., 2009; Adams and Eichmann, 2010; Rama et al., 2015; Koch et al., 2011). While Robo4 shares some structural similarity with the other Robos, it contains fewer Ig-like domains and FNIII repeats and lacks the CC1 and CC3 motifs in its intracellular region (Adams and Eichmann, 2010). Robo4 is highly expressed in the vasculature during embryonic development (Park et al., 2003) and plays a role in the regulation of angiogenesis (Bedell et al., 2005; Jones et al., 2008). Although Robo4 was initially believed to be absent from the nervous system, some evidence suggests that it is also expressed in the nervous system and that it contributes to the migration of pyramidal neurons in the cortex (Zheng et al., 2012).

7.2.3 Slit and Robo interactions Mammalian Slits bind Robo1e3 with different affinities. Whereas Slit2 binds Robo1/2 with high affinity, it interacts with mammalian Robo3 with lower affinity (Zelina et al., 2014). Slit2 can be cleaved into a 140-kDa N-terminal (Slit-N) fragment that binds Robo and a 55- to 60-kDa C-terminal fragment (Slit-C) that does not interact with Robo but can associate with the axon guidance receptor PlexinA1 and with the basement membrane protein dystroglycan (Wang et al., 1999; Brose et al., 1999). Although the protease involved in the cleavage of Slit in mammals remains to be identified, the Drosophila homolog of pheromone convertase 2 (PC2), Amontillado (Amon), can cleave Slit (Ordan and Volk, 2016). The binding of Slit to Robo was suggested to take place between the second LRR motif of Slit (D2) and the first two Ig domains of Robo (Liu et al., 2004). Deletion of Ig1 in Drosophila Robo1 abolishes Slit binding, and mutagenesis analyses revealed that residues Thr74, Phe114, and Arg117 (human Robo1 residues Thr86, Phe128, and Arg131, respectively) are critical for Slit binding (Reichert et al., 2016). Recent studies have unveiled potential effects of Slit binding on the conformation and activation of Robo receptor complexes. Robo1 can dimerize and form a tetrameric oligomer composed of “dimer of dimers” in a compact structure in proximity to the cell membrane (Zakrys et al., 2014; Yom-Tov et al., 2017; Aleksandrova et al., 2018). It is proposed that binding of Slit2-N to Robo1 leads to a conformational change in Robo1, which releases the compact nature of the receptor without affecting dimerization and results in activation of its signaling (Fig. 7.1C) (Aleksandrova et al., 2018). Several additional Slit-binding partners and receptors have been identified and implicated in nervous system development. EVA-1 was identified in C. elegans as a cell-autonomous receptor for Slit (Fujisawa et al., 2007). In eva-1 mutants, as in slt-1/slit and sax-3/robo mutants, the initial pioneer phase of the anterior ventral microtubule (AVM) sensory neuron axon extension toward the ventral nerve cord frequently fails, and axons grow along the lateral epidermis toward the head (Fujisawa et al., 2007). Interestingly, the mammalian homolog EVA-1C is expressed in the developing nervous system, specifically by axons contributing to commissures, tracts, and nerve pathways of the developing spinal cord and forebrain, whose growth and guidance are influenced by Slit (James et al., 2013). In Drosophila, Slit-N can bind to Dscam1 and promote the formation of Robo1-Dscam1 complexes, preventing Slitmediated repulsion and promoting longitudinal axon growth across the segment boundary (Alavi et al., 2016). Slit-N binding to Dscam1 can also regulate axon collateral extension from Drosophila mechanosensory neurons in a Roboindependent manner. In this case, the binding of Slit-N to Dscam1 enhances the formation of a complex between Dscam1 and the receptor tyrosine phosphatase RPTP69D, leading to Dscam1 dephosphorylation (Dascenco et al., 2015). While the Slit-C fragment does not appear to interact with Robo, it can bind directly to PlexinA1, a cell surface receptor implicated in semaphorin signaling. PlexinA1 and slits genetically interact in the crossing of spinal cord commissural axons in mice, and PlexinA1 is required for Slit-induced commissural axon repulsion (Delloye-Bourgeois et al., 2015). The cell surface amyloid precursor protein (APP) has also been shown to bind Slit. The binding of Slit to APP promotes its ectodomain shedding and activation of the small Rho GTPase Rac1. Analyses in C. elegans indicate that the APP ortholog, APL-1, genetically interacts with SLT-1 and SAX-3 in the guidance of pioneer axons (Wang et al., 2017). Although there are an increasing number of Slit-binding cell surface receptors identified, the majority of Slit functions in development described to date rely on Slit binding to Robo family receptors.

7.2.3.1 Regulation of SliteRobo interactions The regulation of SliteRobo interactions can serve to either promote or inhibit SliteRobo signaling. For example, Slit2 binding to the secreted BMP protein antagonist Gremlin antagonizes SliteRobo interactions (Tumelty et al., 2018).

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In contrast, considerable evidence suggests that, for SliteRobo binding to lead to functional signaling, the action of heparan sulfate proteoglycans (HSPGs) as a coreceptor is required (Van Vactor et al., 2006; Lee and Chien, 2004). The HSPGs are composed of a series of sulfated disaccharide units (heparan sulfate [HS]) attached to a core protein that is either secreted or membrane bound. The binding of Slit to HSPGs was initially identified when Slit was found to bind to an affinity matrix of the HSPG glypican-1 (Liang et al., 1999). It has been suggested that O-sulfation of heparan sulfate is the critical structural feature for glypican binding to Slit and that N-sulfation has a lesser role in this process (Ronca et al., 2001). Slit binds heparin through its second LRR domain, the same region that interacts with Robo (Hussain et al., 2006). In addition, Robo can also bind directly to heparin with high affinity at the interface between Robo domains Ig1 and Ig2 (Fukuhara et al., 2008), whereas a second low affinity binding site is found on Ig2 (Li et al., 2015). In Drosophila, mutations in a Syndecan homolog of a membrane-bound HSPG affect all aspects of Slit activity and cause robo null-like phenotypes (Steigemann et al., 2004). Syndecan (sdc) interacts genetically with robo and slit, and double mutations cause a synergistic strengthening of the single-mutant phenotypes, suggesting that Syndecan is a critical component of the Slit/ Robo signaling pathway (Steigemann et al., 2004). In contrast, genetic interactions between slit and other HSPGs, such as dally (Nakato et al., 1995) and perlecan (Voigt et al., 2002), were not observed, suggesting that Syndecan may be the major HSPG involved in SliteRobo function in Drosophila. In zebrafish, Slit interacts with the basement membrane component type IV collagen Col4a5 and with the HSPG agrin to regulate lamina-specific targeting of retinal ganglion cell (RGC) axons in the optic tectum (Xiao et al., 2011). HSPGs may also be required for SliteRobo signaling in mammals. The Slit-C fragment can bind to dystroglycan, which localizes Slit within the basement membrane and floor plate of the neural tube. Disruption of dystroglycan expression leads to commissural axon guidance defects that are similar to alterations observed in Robo1/Robo2 double-null mice (Wright et al., 2012). Genetic ablation of ext1, a member of the exostosin family of genes required for heparan sulfate biosynthesis, leads to axon guidance defects that are reminiscent of the phenotype observed in slit mutant mice. In ext1enull mice, RGC axons project ectopically into the contralateral optic nerve at the optic chiasm as observed in slit1/ slit2 double-null mice (Plump et al., 2002; Inatani et al., 2003). Interestingly, although few guidance defects in RGC projections are observed in slit2 mutant mice, removing one allele of ext1 in these mice causes profound retinal axon misguidance (Inatani et al., 2003). Moreover, different modifications to the sugar structures of HS by HS modification enzymes such as the C5-epimerase encoded by hse-5, the 6O-sulfotransferase encoded by hst-6, and the 2O-sulfotransferase encoded by hst-2 are instructive in Slit-dependent guidance of motor neurons in C. elegans acting through Eva-1 signaling (Bulow et al., 2008). In mice, knockout animals for two different HS modifiers, hs2st and hs6st1, exhibited phenotypes that closely match those of slit1
/
and slit2
/
embryos (Conway et al., 2011). These observations support a model whereby the effect that HS interactions have on Slit-dependent guidance depends on the specific sugar structures generated by HS modifying enzymes. Altogether, these results indicate that heparin sulfate plays a physiologically essential role in Slit-mediated signaling.

7.3 SliteRobo function in midline crossing SliteRobo functions have been extensively studied in commissural axon pathfinding, one of the best understood axon guidance systems. In the mouse spinal cord, commissural neurons are born in the dorsal spinal cord, and their axons navigate toward the floor plate in the ventral part of the spinal cord to form commissures that connect the two symmetrical halves of the nervous system. Upon crossing the midline to the contralateral side, these axons turn longitudinally and grow toward the lateral funiculus. Other neurons in the spinal cord project axons that grow toward the midline but innervate targets on the ipsilateral side without crossing the midline. In the classical model of commissural axon crossing at the floor plate, the floor plate secretes attractive cues, namely Netrin1 (Dickson, 2002; Tessier-Lavigne and Goodman, 1996) and Sonic Hedgehog (Charron et al., 2003), which draw commissural axons toward the midline (Fig. 7.2B). More recent evidence indicates that expression of Netrin1 by cells within the ventricular zone of the neural tube promotes the growth of commissural axons to the midline in the hindbrain and spinal cord (Dominici et al., 2018; Varadarajan et al., 2017). Upon reaching the midline, commissural axons are proposed to acquire sensitivity to the floor plate-derived repellent Slit, which promotes their exit from the floor plate and prevents contralateral axons from recrossing. Ubiquitous ablation of Slit expression in mice led to several midline crossing defects, including stalling of axons inside the floor plate (Long et al., 2004). Nonetheless, a specific requirement for floor plateederived Slit in midline axonal crossing remains to be addressed using cell-specific ablations of Slit expression. In Drosophila, commissural axon guidance in the ventral nerve cord is similar to midline crossing in vertebrates. While subsets of axons do not cross the midline and grow on the ipsilateral side along the midline, commissural axons grow

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FIGURE 7.2 SliteRobo in midline crossing. (A) Midline crossing in Drosophila. The Drosophila midline cells secrete Netrin and Slit. Ipsilateral axons (red), which express high levels of Robo protein, grow adjacent to the midline but never cross it due to their Slit sensitivity. Precrossing commissural axons express Comm that acts to prevent Robo proteins from reaching the axonal cell surface. This allows commissural axons to be attracted to and cross the midline. Postcrossing commissural axons downregulate Comm expression, allowing Robo proteins to reach the axon cell surface and prevent recrossing. Three distinct lateral pathways are formed following midline crossing. Medial axons express Robo1, intermediate tract axons express Robo1 and 3, whereas the most lateral tract axons express Robo1, 2, and 3. (B) Midline crossing in mice. In the mouse, spinal cord ipsilateral axons (red), which express Robo1, 2, and 3 grow toward the midline, turn and grow longitudinally to innervate targets on the same side of the spinal cord. Precrossing commissural axons (black) express low levels of Robo1, 2, and 3.2 but high Robo3.1 protein. These axons project to the floor plate by responding to the chemoattractants Netrin1 and Shh. Upon reaching the floor plate, commissural axons cross the midline to the contralateral side, turn longitudinally, and grow toward the lateral funiculus. Postcrossing commissural axons upregulate Robo1, 2, and 3.2 proteins allowing these axons to continue to grow adjacent to the midline and never recross it. The spatial expression of Robo3.2 is regulated by local transcription of its transcript and nonsense-mediated mRNA decay. (C) Axonal guidance in the Caenorhabditis elegans nerve cord: Cross section of anterior ventral microtubule (AVM) ventral guidance at the nerve cord in C. elegans. SLT-1 is secreted from dorsal (D) muscles (blue), whereas UNC-6 is secreted by ventral (V) muscles (purple). AVM axons express SAX-3 and UNC-40 receptors allowing for their simultaneous repulsion from slit and attraction to UNC-6. (D) Longitudinal view of AVM ventral guidance in C. elegans. AVM axons project ventrally until the midline where they turn to travel anteriorly (A).

toward the midline in response to Netrin and cross and exit the midline in response to Slit. Once on the contralateral side of the midline, these axons grow longitudinally in specific lateral tracts (Fig. 7.2A). Analysis of slit mutations in the fly provided the initial evidence that Slits act as midline repellents in axon guidance in the nerve cord. In Drosophila, slit-null mutations lead to collapse of axon tracts onto the midline (Kidd et al., 1998a; Rothberg et al., 1990). Similarly, in vertebrate slit mutants, commissural axons show defects such as stalling at the midline or aberrant crossing (Long et al., 2004). Robo mutations also lead to defects in commissural axon pathfinding that include random crossing of ipsilateral axons as well as recrossing of longitudinally projecting commissural axons (Kidd et al., 1998a; Seeger et al., 1993). Integrins also play a role in Slit responsiveness at the midline. Drosophila embryos lacking one copy of slit along with one copy of the integrin gene encoding aPS1, aPS2, aPS3, or bPS1 show pathfinding errors at the midline that are more severe than slit heterozygous embryos alone, indicating that the response of growing axons in the nerve cord to midline Slit can be potentiated by adhesive interactions from integrins (Stevens and Jacobs, 2002). The study of Slit and Robo function in commissural axon midline crossing has shed light on important questions that include the following: How do commissural axons exit the midline when exposed to powerful midline attractants? How do

152 PART | I Formation of axons and dendrites

these axons avoid recrossing the midline upon continued exposure to attractants? The answers to these questions have revealed intricate molecular mechanisms by which SliteRobo signaling is regulated that will be discussed in the following sections.

7.3.1 Spatial expression patterns of Slit and Robo To begin to discuss how Slits and Robos contribute to commissural axon guidance, their patterns of expression in the developing vertebrate spinal cord and Drosophila ventral nerve cord must be discussed. In both Drosophila and vertebrates, Slits are expressed and secreted by midline cells in Drosophila (Rothberg et al., 1988, 1990) and Slits1e3 in vertebrates (Brose et al., 1999; Holmes et al., 1998; Itoh et al., 1998; Li et al., 1999; Nakayama et al., 1998), whereas Robo receptors (Robo1e3) are expressed by commissural axons as well as by axons projecting ipsilaterally (Dickson and Gilestro, 2006; Kidd et al., 1998a; Brose et al., 1999; Long et al., 2004; Sabatier et al., 2004; Camurri et al., 2004). In Drosophila, the glycosylation pathwayeassociated enzyme Mummy is required for Slit glycosylation and secretion and also for maintaining the abundance of Robo on axons (Manavalan et al., 2017). Notably, in mice and humans, the Robo-1 locus has two alternative promoters that lead to the production of two transcript isoforms, Robo1 and Dutt1, both sharing considerable sequence similarity (Nural et al., 2007). Interestingly, the alternative promoters appear to have distinct spatial distributions and temporal transcriptional activities (Clark et al., 2002). A study examining the expression patterns of the Robo1 and Dutt1 isoforms by in situ hybridization revealed that they are differentially expressed in the embryonic mouse brain (Nural et al., 2007). Furthermore, and importantly for this discussion, Dutt1 is the main isoform expressed in embryonic commissural axons (Nural et al., 2007). Nonetheless, we will use the term Robo1 to refer to both Robo1 isoforms for the remainder of this chapter.

7.3.2 Posttranscriptional Robo regulation An interesting guidance decision to examine in vertebrates is the difference in pathways that commissural and ipsilateral axons take in the spinal cord. Both sets of axons encounter the chemoattractants Netrin1 and Shh during their growth in the spinal cord. Despite the potent attractive effects of these chemoattractants, only commissural axons will enter the floor plate and cross the midline. The different routes taken by these two classes of axons are not due to differential responsiveness to these chemoattractants but are instead due to the repulsion of ipsilateral axons away from the midline by Slits. In fact, commissural axons will eventually become sensitive to midline repellents as well once they cross the midline, but it is the tight regulation of Slit responsiveness that allows for these two distinct groups of axons to follow different paths despite being exposed to identical chemical environments. How is this regulation of sensitivity to midline repellents conveyed? Both invertebrates and vertebrates have distinct molecular mechanisms to control the posttranscriptional regulation of the Robo family receptors in commissural neurons.

7.3.3 Regulation of Robo protein expression at the midline 7.3.3.1 Drosophila and vertebrate midlines The difference in responsiveness to midline Slit by commissural and ipsilateral axons can be explained by the differential expression of Robo protein at the surface of these axons. In Drosophila, both ipsilateral and commissural neurons express similar amounts of robo transcript. In contrast, the Robo protein levels on axon tracts from these two populations of neurons are quite different (Kidd et al., 1998a). Precrossing commissural axons harbor low levels of Robo protein, whereas ipsilateral axons express much higher levels of Robo protein (Kidd et al., 1998a). This leads to diverging responses at the midline by these two populations of axons; ipsilateral axons are highly repelled by midline Slit, whereas commissural axons show overall attraction to the midline and negligible repulsion by Slit (Fig. 7.2A). Robo2 and Robo3 are also absent from the surface of precrossing commissural axons (Rajagopalan et al., 2000a, 2000b; Simpson et al., 2000a). While forced expression of any of the three Robos can prevent commissural axon crossing (Kidd et al., 1999; Simpson et al., 2000b; Rajagopalan et al., 2000a), loss-of-function studies reveal that only Robo is required to control midline crossing. Indeed, while robo mutants show prevalent midline crossing errors, robo2 and robo3 single or double mutants seldom contain defects (Simpson et al., 2000b; Rajagopalan et al., 2000a). Akin to Robo regulation in Drosophila, vertebrate midline axon guidance also has an elegantly tuned system involving the regulation of SliteRobo signaling in precrossing commissural axons. Robo1 and Robo2 are expressed at low levels on precrossing commissural axons, but their expression is upregulated following crossing (Long et al., 2004) (Fig. 7.2B). Robo3 expression, on the other hand, differs from what is observed in Drosophila; its expression is high on precrossing

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and low on postcrossing commissural axons (Sabatier et al., 2004). As observed in Drosophila, overexpression of Robo1 in mouse spinal cord commissural neurons can also prevent commissural axon crossing (Sabatier et al., 2004). In Drosophila, the regulation of Robo protein levels in commissural axons is controlled by the multipass transmembrane protein Comm (Keleman et al., 2002). This protein was identified in the same genetic screen that led to the discovery of Robo (Seeger et al., 1993). In comm mutants, commissural axons fail to cross the midline and instead share the path of ipsilateral axons (Seeger et al., 1993). This phenotype is in stark contrast to the defects observed in robo mutants and suggested that Comm may regulate Robo expression. Comm is expressed by both commissural neurons and midline cells, but it is not expressed in ipsilateral neurons (Bonkowsky et al., 1999) (Fig. 7.2A). A series of experiments whereby Comm expression was first ablated in the entire embryo and selectively reexpressed in either commissural neurons or midline cells revealed that expression of Comm in commissural axons is required for proper midline crossing of these axons (Keleman et al., 2002, 2005; Georgiou and Tear, 2002). In addition, forced expression of Comm in ipsilateral neurons results in midline crossing of their axons (Bonkowsky et al., 1999). Finally, comm;robo and comm;slit double mutants show defects in midline crossing that resemble the phenotype observed in single robo (Seeger et al., 1993) or slit mutants, suggesting that Comm functions to interfere with SliteRobo signaling (Dickson and Gilestro, 2006). How does Comm antagonize this signaling and control the crossing of axons at the midline? Studies examining the subcellular localization of Comm provide better understanding of how it interferes with the SliteRobo signaling pathway. In the Golgi, Comm can bind Robo and directly traffic it to endosomes, thus preventing it from ever reaching the cell surface (Keleman et al., 2002; Myat et al., 2002). It is still unclear how Comm couples Robo to the endosomal system (Myat et al., 2002; Keleman et al., 2005). These data support a model whereby Comm blocks SliteRobo signaling and allows midline crossing in commissural neurons by sequestering Robo from these axons, thus rendering them irresponsive to midline Slit (Kidd et al., 1998a; Keleman et al., 2002; Araujo and Tear, 2003; Dickson and Senti, 2002; Georgiou and Tear, 2003; Simpson et al., 2000b). Following midline crossing, Comm expression is downregulated, leading to the expression of Robo protein at the surface of commissural axons (Kidd et al., 1998b; Tear et al., 1996). As a result, commissural axons become responsive to Slit expressed by the midline, which prevents them from recrossing the midline (Kidd et al., 1998a; Long et al., 2004). There is also evidence that additional mechanisms exist to regulate Slit responsiveness in Drosophila commissural neurons. In early stages of commissural axon guidance, Robo2 is highly expressed in midline cells, and its overexpression can partly rescue the comm mutant phenotype. The ability of midline celleexpressed Robo2 to promote midline crossing is dependent on its Ig1/2 domains but does not require its cytoplasmic domain, showing that Robo2 binding to Robo1 on commissural axons can suppress Slit repulsion (Evans et al., 2015). These observations suggest a model in which Comm and Robo2 work together to suppress Slit repulsion in commissural axons while crossing the midline through cell autonomous and noncell autonomous mechanisms, respectively. The regulation of Robo expression may also play a critical role in controlling Slit responsiveness in other regions of the fly nervous system, such as for projections of small lateral neurons (sLNv) in the brain. Slit and robos interact genetically with the receptor protein tyrosine phosphatase rptp69d in the guidance of these axons. Furthermore, RPTP69D forms a complex with Robo3 and increases its surface expression, leading to enhanced SliteRobo signaling (Oliva et al., 2016).

7.3.3.2 Caenorhabditis elegans midline The study of axon guidance in C. elegans involves examination of the extension of axons into one major axon bundle of either the nerve ring in the head or the ventral nerve cord (White et al., 1976, 1986). Ventral nerve cord guidance at the midline involves UNC-6 (Netrin)-mediated attraction of peripheral axons to the ventral midline where the major nerve cords form (Ishii et al., 1992). In C. elegans, this attraction is mediated by UNC-40 (DCC/frazzled) (Chan et al., 1996; Hedgecock et al., 1990). In addition, SAX-3 (Robo) acts together with SLT-1 (Slit), expressed in the dorsal muscles of the nerve cord to repel SAX-3 expressing axons toward the ventral midline (Hao et al., 2001) (Fig. 7.2C and D). The expression of SAX-3 is dynamic; early in development, expression is found in all epidermal cells, whereas later on in development, during initial axonal outgrowth, it is highly expressed in the neurons of the nerve ring (Zallen et al., 1998). This transient expression of SAX-3 is reminiscent of the changes in Robo protein levels in commissural axons in other species. Mutations in the sax-3 gene lead to phenotypes analogous to those seen in Robo mutants in Drosophila and vertebrates (Zallen et al., 1998). Indeed, axons often take on more lateral positions, some stalled at the midline, whereas others cross the midline multiple times (Zallen et al., 1998). The AVM mechanosensory neurons, whose cell bodies are located in the lateral hypodermis, send axons that extend ventrally to the ventral midline. In mutants lacking the slt-1 gene, AVM axons do not cross the midline and instead grow anteriorly in positions lateral to the midline, a defect similar to the phenotype observed in sax-3 mutants (Hao et al., 2001).

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Interestingly, the defects observed in sax-3 mutants are not all present in slt-1 mutants, suggesting that SAX-3 can function independently of SLT-1 (Hao et al., 2001). This is highlighted in the case of the trajectory followed by the nerve ring axons en route to the ring head, a large neuropil in the head of the worm. Here, sax-3 mutants exhibit widespread pathfinding errors, including premature termination, anterior rerouting, and formation of an ectopic posterior process, whereas slt-1 mutants show none of these defects (Hao et al., 2001).

7.3.4 Regulation of Robo signaling at the midline in vertebrates As mentioned earlier, Robo protein regulation at the midline is also observed in vertebrates although Comm-like proteins have yet to be identified in vertebrates (Dickson and Gilestro, 2006). The WAGR syndromeeassociated protein PRRG4 has been proposed as a candidate to regulate Robo expression in commissural neurons, although functional analyses in vertebrates will be needed to establish its role in the guidance of these axons to the floor plate (Justice et al., 2017). A component of the vesicle fusion machinery, RabGDI, has been implicated in the regulation of Robo1 expression in vertebrates. RabGDI can promote Robo1 expression at the surface of commissural axons, and its downregulation in chick embryos prevents axons from exiting the floor plate, a phenotype also observed upon Robo1 downregulation (Philipp et al., 2012). While the spatiotemporal control of Robo expression may regulate Slit responsiveness, SliteRobo signaling in vertebrate commissural neurons appears to also be regulated by the expression of Robo3 in these neurons (Chen et al., 2008). Close examination of the expression levels of Robo3 on commissural axons revealed a spatially distinct pattern of expression that mirrored the expression of the other two Robo family members in these axons: high Robo3 expression in precrossing commissural axons together with low expression in postcrossing commissural axons (Sabatier et al., 2004; Chen et al., 2008) (Fig. 7.2B). While this pattern of expression suggested Robo3 may have a redundant function to Robo1 and Robo2 in midline crossing of commissural axons, examination of these projections in the spinal cord of robo3 mutant mouse embryos supports the idea that Robo3 may in fact inhibit Robo1- and Robo2-dependent Slit signaling. In robo1;robo2, double-mutant embryos commissural axons enter the floor plate but fail to exit on the contralateral side (Chen et al., 2008). In contrast, commissural axons assume the path of ipsilateral axons and never approach the midline in robo3 mutant embryos (Sabatier et al., 2004; Marillat et al., 2004). Furthermore, these defects can be partially rescued by ablating expression of Robo1, Slit1, or Slit2 in robo3 mutant embryos, indicating that Robo3 expression normally inhibits Slit signaling (Sabatier et al., 2004). Another potential mechanism of action for Robo3 in commissural axons is promoting Netrin-dependent attraction of these axons to the floor plate. Though mammalian Robo3 does not bind to Slit, it can interact with the Netrin receptor DCC and potentiate Netrin/DCC-mediated attraction of pontine neurons toward the midline (Zelina et al., 2014). In an explant culture assay, Netrin1 fails to attract pontine neurons from robo3 null explants, and this can be rescued by expression of mammalian Robo3. It is therefore possible that Robo3 also contributes to Netrinmediated attraction of commissural axons to the floor plate. In addition to promoting guidance of commissural axons to the floor plate by inhibiting Slit signaling, and possibly promoting Netrin attraction, Robo3 expression on these axons is important for preventing them from entering the ventral horn of the spinal cord, as they project to the floor plate. The secreted molecule NELL2 is highly expressed in the ventral horn region, can bind to Robo3, and can repel commissural axons in vitro in a Robo3-dependent manner (Jaworski et al., 2015). Hence, Robo3 plays multiple roles in the guidance of commissural axons toward the floor plate. The ability of Robo3 to interfere with Slit-induced repulsion appears to be mediated by spatially regulated expression of two Robo3 protein isoforms encoded by two alternatively spliced Robo3 mRNA isoforms (Chen et al., 2008). Both Robo3 isoforms are expressed in the spinal cord during commissural axon guidance but are located in distinct axonal compartments (Chen et al., 2008). Precrossing commissural axons express isoform Robo3.1, whereas the second Robo3 isoform, Robo3.2, is transiently expressed in postcrossing commissural axons (Fig. 7.2B). This seemingly rapid transition in expression profiles is thought to contribute to the different responses of pre- and postcrossing commissural axons to midline guidance cues. The expression of Robo3.2 is regulated through the local translation of its transcript in postcrossing axons in response to an unidentified signal from the floor plate (Colak et al., 2013). Following translation of the robo3.2 transcript, nonsense-mediated mRNA decay of this transcript ensures the transient expression of the Robo3.2 protein isoform. To understand the functional differences between these two Robo3 isoforms, they were ectopically expressed in the chick spinal cord embryo. Persistent expression of Robo3.1 in commissural axons favored midline crossing since all commissural axons expressing Robo3.1 crossed and inappropriately recrossed the midline, consistent with the notion that this isoform promotes midline crossing (Sabatier et al., 2004; Chen et al., 2008). In contrast, Robo3.2 overexpression led to a failure of most axons to cross the midline. Furthermore, the robo-3 mutant mouse phenotype could be largely rescued by in utero expression of Robo3.1, but not Robo3.2 (Chen et al., 2008). Inhibition of Robo3.1 expression using in vivo siRNA

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confirms that Robo3.1 expression accounts for the majority of the Robo3 activity required to inhibit precocious repulsion from midline by Slit. Collectively, these experiments suggest that Robo3.1 acts to prevent premature axon repulsion during midline crossing by limiting the response to Slit by precrossing axons, whereas Robo3.2 may function to turn on repulsion from the midline postcrossing (Chen et al., 2008; Sabatier et al., 2004). Robo3.1 may block Slit-mediated repulsion by interfering with Robos1 and 2 through direct physical interaction or by interfering with their downstream signaling pathway(s), although direct evidence for either of these possibilities remains to be obtained (Chen et al., 2008). In contrast to Robo3.1, Robo3.2 does not interfere with Slit signaling and is proposed to contribute to repulsion of postcrossing commissural axons (Chen et al., 2008). However, it remains unclear how Robo3.2 may mediate repulsion of these axons since it does not bind to Slits (Zelina et al., 2014).

7.3.5 SliteRobo signaling for exiting the midline Commissural axons that cross the midline must then exit the floor plate, an environment rich in chemoattractants. In addition to promoting the exit of axons through repulsion, SliteRobo signaling also plays a critical role in inhibiting Netrin-mediated attraction in the floor plate. One of the Netrin receptors, DCC, belongs to the Ig superfamily of transmembrane proteins and contains an ectodomain composed of four Ig domains, six fibronectin (FN) type III repeats, and a long intracellular region (Keino-Masu et al., 1996). Binding of Netrin to DCC induces intracellular signaling events that promote the growth and turning of axons toward a source of Netrin. When spinal neurons isolated from Xenopus are exposed to a source of Netrin in vitro, their growth cones are attracted to the Netrin source (Ming et al., 2002). In contrast, when these axons are simultaneously presented with Netrin1 and Slit2, the attractive effect of Netrin1 is completely abolished, suggesting that Slit2 inhibits NetrineDCC signaling (Stein and Tessier-Lavigne, 2001). Interestingly, Slit2 does not affect Netrin1-mediated extension of axons, indicating that SliteRobo signaling regulates only specific signaling events downstream of DCC (Stein and Tessier-Lavigne, 2001). SliteRobo-dependent silencing of Netrin1 attraction is also necessary to prevent the growth of motor neuron axons toward the floor plate. Inhibition of SliteRobo signaling, achieved by expressing a dominant-negative form of Robo in explant cultures or through ablation of Robo1 and Robo2 in vivo, induces growth of motor neuron axons toward a Netrin-1 source (Bai et al., 2011). The silencing effect of Slit on Netrin-mediated attraction could be a receptor-mediated event whereby Slit binds to Robo on the growth cone and antagonizes Netrin-mediated attraction through DCC. This model is supported by the observation that Robo and DCC form a complex through their cytoplasmic domains upon exposure to Slit. Blocking this interaction prevents the Slit-induced silencing of Netrin attraction in vitro (Stein and Tessier-Lavigne, 2001). Furthermore, the accumulation of a membrane-bound cleaved form of the intracellular domain of DCC in mice lacking the g-secretase Presenilin1 blocks RoboeDCC interactions, resulting in erroneous Netrin1 attraction of motor neuron axons toward the floor plate (Bai et al., 2011). Another example of cross-talk between the Slit and Netrin signaling pathways is observed in C. elegans. UNC-73, a homolog to the mammalian Rho-GEF Trio protein, regulates the cell surface localization of SAX-3 and also the subcellular localization of UNC-40 (Levy-Strumpf and Culotti, 2007; Watari-Goshima et al., 2007). In the case of SAX-3, there is a direct interaction between Vab-8L, a kinesin-related protein, UNC-73, and the intracellular domain of SAX-3 to increase its cell surface levels (Watari-Goshima et al., 2007). CRML-1, the C. elegans homolog of the mammalian cell migration regulator CARMIL, can also antagonize UNC-73, thus reducing cell surface SAX-3 levels (Vanderzalm et al., 2009). In parallel, UNC-40 trafficking is influenced by UNC-73, which activates the Rho-family GTPase MIG-2 in collaboration with Vab-8L (Levy-Strumpf and Culotti, 2007).

7.4 Modulation of SliteRobo signaling The control of SliteRobo signaling at the midline is achieved through the regulated transport of the Robo receptors to the membrane as well as through expression of alternatively spliced versions of a specific Robo receptor, as discussed earlier. Other modes of regulation exist including transcriptional control of Slit and Robo expression, cell surface cleavage of the Robo receptors, and endocytic trafficking of the Robo receptors.

7.4.1 Transcriptional control Transcription factors that regulate expression of the slit gene were originally identified in Drosophila. The regulation of slit expression at the midline involves interactions between three distinct types of transcription factors: single-minded (Sim), a basic helixeloopehelix (bHLH)ePAS transcription factor; Fish-hook (Fish), a Sox HMG domain protein; and Drifter

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(Dfr), a POU domain protein (Ma et al., 2000). Midline is another transcription factor that has been implicated in the direct control of both slit and robo transcription in flies (Liu et al., 2009). Loss-of-function of this T-box transcription factor severely reduced the expression of the receptor and the ligand. The role of Midline might be conserved throughout evolution since it has orthologs (such as Tbx20) that are also expressed in motor neurons in human, mouse, and zebrafish (Ahn et al., 2000; Takeuchi et al., 2005). Interestingly, both slit1;slit2 double mutants and tbx20 mutant mice show defects in the development of hindbrain motoneurons (Hammond et al., 2005; Song et al., 2006). The well-characterized Hox family of homeodomain-containing transcription factors has also been proposed to act as transcriptional regulators of the Slit and Robo genes. The neuronal migration defects observed in hoxa2 and hoxb2 mutant mice are phenocopied in compound robo1;robo2, slit1;slit2, and robo2;slit2 mutant mice (Geisen et al., 2008). Indeed, Hoxa2 binds directly to the robo2 locus in vivo, and maintenance of high Robo and Slit expression levels is impaired in Hoxa2 mutant mice (Geisen et al., 2008). In addition, the homeobox gene irx4 specifically inhibits Slit1 expression in the chicken retina (Jin et al., 2003), and ablation of Isl1 and Isl2 led to decreased expression of Slit2 in mice (Lee et al., 2015). The expression of Robo1 and Robo2 in thalamic neurons is regulated by the LIM homeodomain 2 (Lhx2) and gastrulation brain homeobox 2 (Gbx2) transcription factors (Chatterjee et al., 2012; Marcos-Mondejar et al., 2012). Ablation of Lhx2 expression in thalamic neurons resulted in increased expression of robo1 and robo2 transcripts in these neurons, suggesting that Lhx2 represses their expression (Marcos-Mondejar et al., 2012). Although Lhx2 is proposed to directly repress expression of robo1 and robo2 transcripts, Gbx2 promotes robo2 expression by inhibiting regulators of LIM transcription factors (Chatterjee et al., 2012). The changes in robo1 and robo2 expression observed in Lhx2 and Gbx2 mutant mice are thought to underlie defects observed in thalamocortical axonal projections in these mice (Chatterjee et al., 2012; Marcos-Mondejar et al., 2012). In Drosophila motor neurons, loss of the homeodomain transcription factor Hb9 resulted in decreased expression of Robo2 and axonal guidance defects, indicating that Hb9 is required for Robo2 expression in these neurons (Santiago et al., 2014). Nkx2.9 has also been proposed as a regulator of Robo2 expression but in motor neurons. Loss of Nkx2.9 leads to defects in projections of spinal accessory motor neurons that are associated with a decrease in Robo2 expression (Bravo-Ambrosio et al., 2012). As illustrated in our discussion of midline axon guidance, the production of Robo3 protein is tightly regulated through alternative splicing of the gene in commissural neurons. Several families of transcription factors have been implicated in the control of Robo3 expression. Robo3 was originally identified in a screen for genes with elevated expression in a retinoblastoma protein (Rb) mutant mouse. In rb mutant embryos mice, Robo3 expression is upregulated in the hindbrain and spinal cord, suggesting that Rb may inhibit transcription of the robo3 gene (Yuan et al., 1999a, 2002). The transcription factor Pax2 can interact with Rb and reverse its transcriptional suppression at the robo3 promoter (Yuan et al., 2002). The transcription of the robo3 gene is also regulated by members of the Sim family of transcription factors. In Sim1;Sim2 double mutant mice, Robo3 is expressed ectopically in a subset of hypothalamic neurons, whereas Robo1 and Robo2 expression remains unchanged (Marion et al., 2005). These double mutant mice are characterized by abnormal axon crossing of the midline with the presence of axons either directed toward the midline or crossing it (Marion et al., 2005). Furthermore, ablation of Sim1a and Artn2 in zebrafish leads to increased expression of the robo3a.1 isoform in the hypothalamus (Schweitzer et al., 2013). The two closely related LIM homeodomain proteins Lhx2 and Lhx9 also regulate robo3 expression in the dorsal spinal cord (Wilson et al., 2008). The control of robo3 expression in cerebellar neurons involves the RNA-binding protein Musashi1 (Msi1) (Kuwako et al., 2010). Msi1 null mice share similar defects in axonal midline crossing and neuronal migration of precerebellar neurons as those observed in robo3 null mice, supporting the notion that Robo3 expression is controlled by Msi1 (Kuwako et al., 2010).

7.4.2 Regulation of SliteRobo signaling by metalloprotease cleavage Common approaches used to regulate ligandereceptor signaling include the cleavage of the ligand into fragments that are biologically inactive or the shedding of the receptor ectodomain from the cell surface. These mechanisms have been implicated in the control of signaling for other axon guidance molecules such as Netrin and Ephrins. For example, cleavage of the Netrin receptor DCC attenuates Netrin signaling (Galko and Tessier-Lavigne, 2000). In contrast, cleavage of the EphrinA2 ligand at the cell surface is necessary for Ephrin-dependent contact repulsion (Hattori et al., 2000). In Drosophila, a dose-dependent genetic interaction between slit and the metalloprotease kuzbanian (Kuz) (ADAM10 in humans) has been described (Schimmelpfeng et al., 2001). Kuz is a single-pass transmembrane metalloprotease belonging to the Adam family and is expressed at the cell surface where it cleaves its substrates within their extracellular domain, resulting in ectodomain shedding (Fambrough et al., 1996). Thus, the dose-dependent genetic interaction between kuz and slit suggests that Kuz regulates the cleavage of Slit during midline axon repulsion. However, more recent evidence indicates that Robo is the actual target of Kuz and that its cleavage is important in the context of midline axon guidance

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(Coleman et al., 2010). This mechanism of action is supported by the observation that Slit proteolysis is not required for its repulsive effects at the midline (Coleman et al., 2010). Furthermore, the crystal structure of the juxtamembrane (JM) region of human Robo1 revealed that the JM region is folded in a conformation that precludes protease accessibility and cleavage of the extracellular region (Barak et al., 2014). Mutagenesis analyses showed that exposure of the JM region can facilitate Robo1 cleavage and shedding from transfected cells (Barak et al., 2014). Binding of ECM-attached Slits to Robo1 on the growth cone of axons may generate a tensile force that reveals the cleavage site and promotes Robo1 shedding (Barak et al., 2014). Based on these combined in vivo and in vitro observations, protease-dependent cleavage of the Robo receptors is likely the main regulatory mechanism for SliteRobo repulsion at the Drosophila midline.

7.4.3 Regulation of SliteRobo signaling by ubiquitination Using the yeast two-hybrid strategy to identify proteins interacting with the intracellular domain of Robo1, multiple clones encoding ubiquitin-specific protease 33 (USP33) were isolated (Wong et al., 2001). The deubiquitinating enzyme USP33 is widely expressed in the brain and in other tissues (Li et al., 2002). Robo1 interacts directly with USP33 in both heterologous cells and mouse brain lysates (Yuasa-Kawada et al., 2009). Robo1 protein has the potential to be ubiquitinated, and USP33 is involved in its deubiquitination (Yuasa-Kawada et al., 2009). Furthermore, using a loss-of-function approach, USP33 was found to be required for Slit-induced, but not Sema3F-induced, growth cone collapse in mouse commissural neurons (Yuasa-Kawada et al., 2009). Also, USP33 is required for commissural axons to cross the midline. A significant number of DiI-labelled axons stall in the midline when siRNAs directed against Usp33 are electroporated in ovo in chick embryos (Yuasa-Kawada et al., 2009). This phenotype is similar to those observed in slit1;slit2;slit3 triple knockout mice and also in robo1 mutant mice (Long et al., 2004).

7.5 Signaling downstream of Robo The binding of Slit to Robo induces intracellular signaling events that regulate cytoskeletal dynamics and, ultimately, axon guidance (Fig. 7.3). Though the cytoplasmic region of Robo does not contain any obvious catalytic domains, it contains

FIGURE 7.3 SliteRobo signaling. The binding of Slit to Robo takes place between the second leucine-rich repeat (LRR) motif of Slit and the first two Ig domains of Robo. Following this interaction, multiple downstream events occur to regulate cytoskeletal rearrangements and axonal guidance. Robo signaling is downregulated by phosphorylation of the CC1 domain by the tyrosine kinase Abelson (Abl), which binds directly to the CC3 domain. The actin-binding protein Enabled (Ena) that binds the Robo CC1 and CC2 domains is also phosphorylated by Abl. Ena, in collaboration with the monomeric actin-binding protein Profilin, positively regulates actin polymerization by associating with the barbed end of actin filaments. CrossGAP/Vilse binds directly to the CC2 intracellular domain of Robo and promotes the hydrolysis of RacGTP and, less efficiently, of Cdc42GTP. srGAPs may also promote GTP hydrolysis in RhoGTPases to promote Slit-mediated repulsion. Having an opposite effect on Rho family GTPases, the guanine nucleotide exchange factors (GEF) protein Son of Sevenless (Sos) activates Rac1. Sos forms a ternary complex with Robo and the adaptor protein Dreadlocks (Dock)/Nck, which binds the CC2 and CC3 domains, to regulate Rac-dependent cytoskeletal rearrangement and axonal repulsion in response to the Slit ligand. The Rac1 effector p21-activated kinase (Pak), which binds to Robo CC2-3 domains, is thought to contribute to stabilization of the new actin filaments.

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several conserved motifs, and it is necessary for Slit-mediated repulsion (Simpson et al., 2000b). Upon Slit binding, Robo receptors are endocytosed and trafficked in endosomes where they are proposed to recruit downstream effectors involved in regulating the activity of Rho family GTPases and cytoskeletal rearrangements (Chance and Bashaw, 2015).

7.5.1 Rho family of small GTPases The family of Rho-GTPases, including Rac, Cdc42, and Rho, has emerged as including key regulators of cytoskeletal dynamics (Hall, 1998). Rho-GTPases are present either in an active form (GTP-bound) conformation or in an inactive state (GDP-bound). This tightly regulated balance relies on the antagonistic function of guanine nucleotide exchange factors (GEFs) and GTPase-activating proteins (GAPs). GEFs promote the exchange of GDP to GTP, thereby inducing signaling downstream of Rho GTPases, and GAPs enhance GTP hydrolysis, which blocks signaling. Since Robo receptors do not possess autonomous CC in their intracellular domains, the family of Rho-GTPases and their regulators represent interesting candidates for signaling. A yeast two-hybrid screen using a portion of the intracellular domain of Robo1 as bait leads to the identification of the SliteRobo GTPase-activating proteins (srGAPs) (Wong et al., 2001). Four srGAPs (srGAPs1, 2, 3, and 4) have been identified (Lucas and Hardin, 2018). srGAP1e3 are highly conserved (60%e80% identical), whereas srGAP4 only shares 51% identity with srGAP-3. srGAP proteins are highly expressed in the brain, lung, and spleen (Wong et al., 2001). In the central nervous system, srGAPs have diverse patterns of expression that are often distinct from each other and are therefore likely to be important for many different aspects of development (Bacon et al., 2009). srGAPs interact with a proline-rich region of the Robo cytoplasmic region through their Src homology domain 3 (SH3). The SH3 domain of srGAP can also bind to the actin reorganization proteins WASP/WAVE (Soderling et al., 2007; Linkermann et al., 2009). It appears that different srGAPs selectively activate the GTPase activity of specific Rho family GTPases and block their signaling. The different srGAPs have different but overlapping effects on the GTPases. In a heterologous cell system, srGAP-1 promotes GTP hydrolysis by Cdc42 and RhoA (Wong et al., 2001); however, current reports differ with respect to the ability of srGAP-1 to inactivate Rac1 (Wong et al., 2001; Ma et al., 2013; Yamazaki et al., 2013). srGAP2 can stimulate Rac1 activity (Guerrier et al., 2009; Ma et al., 2013; Mason et al., 2011), whereas srGAP3 inactivates Cdc42 and Rac1 (Soderling et al., 2002; Waltereit et al., 2012). Despite different reports on the effects of srGAP-1 on Rac1, it has been shown that srGAP-1 can regulate Slit-mediated inhibition of cell migration in vitro. Interestingly, when tissue explants of precursor neurons from the anterior subventricular zone (SVZ) are infected with a virus that expresses an srGAP1 mutant lacking CC, these neurons are no longer repelled by a source of Slit and migrate normally out of the explants (Wong et al., 2001). A model has been proposed where, upon meeting Slit in the environment, the growth cone binds Robo locally, activating srGAPs that in turn results in inactivation of RhoGTPases. Since RhoGTPases stimulate actin polymerization, this leads to an asymmetric inactivation of actin polymerization, thereby making the growth cone move away from the source of Slit (Lucas and Hardin, 2018). Using Drosophila genetics, two independent groups have identified an additional Rho-GAP, CrossGAP/Vilse, that interacts both biochemically and genetically with Robo (Hu et al., 2005; Lundstrom et al., 2004). Moreover, this Rho-GAP is present at the right developmental time and place to interact with the Robo receptor (Hu et al., 2005). Strong midline guidance defects in wild-type embryos are generated only by CrGAP/Vilse gain-of-function, but not by the loss-offunction, suggesting that Robo might normally function to downregulate CrGAP/Vilse to control midline crossing (Hu et al., 2005). GEFs have also been implicated in the regulation of SliteRobo signaling. The Rho-GEF protein Son of Sevenless (Sos) is recruited to the plasma membrane, where it forms a ternary complex with Robo and the adaptor protein Dreadlocks (Dock) to regulate Rac-dependent cytoskeletal rearrangement in response to the Slit ligand (Yang and Bashaw, 2006; Fan et al., 2003). Rac1, in turn, activates p21-activated kinase (Pak), leading to stabilization of new actin filaments (Rex et al., 2009). In C. elegans, mutations in the Rac GEF Rin-1 resulted in altered dorsoventral axonal guidance due to improper localization of actin in hermaphrodite-specific neurons (HSNs). Genetic interaction analyses indicate that Rin-1 acts downstream of Robo to regulate dorsoventral guidance of HSN axons (Doi et al., 2013). It remains possible, however, that Rin-1 may also regulate atypical Rho GTPases, such as CHW-1, which has been implicated in SAX3/Robo-mediated axonal guidance (Alan et al., 2018).

7.5.2 Abelson tyrosine kinase The cytoplasmic CC2 sequence (LPPPP) of Robo is a consensus binding site for the Ena-VASP-homology (EVH1) domain of Drosophila Enabled (Ena). The actin-binding protein Ena is a member of a small family of evolutionarily conserved proteins that are implicated in the regulation of the actin cytoskeleton during cell motility and growth cone

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guidance (Wills et al., 1999). Ena binds to the abelson tyrosine kinase (Abl) (Gertler et al., 1995, 1996), and both Abl and Ena can directly bind to the cytoplasmic domain of Robo (Bashaw et al., 2000). Binding of Ena/VASP family proteins to Robo is necessary to enhance Slit-mediated repulsion, at least in part by stimulating filopodial extension in the growth prior to repulsion (Bashaw et al., 2000; McConnell et al., 2016). In contrast, Abl was proposed to antagonize Robo signaling (Bashaw et al., 2000). Moreover, Abl kinase activity is necessary for inhibition of repulsive Robo signaling (O’Donnell and Bashaw, 2013). However, the observation that Drosophila capulet (capt), a homolog of the adenylyl cyclasee associated protein that regulates actin polymerization, collaborates with Abl to mediate Slit function challenges the previous model whereby Abl acts purely to antagonize and/or downregulate Robo signaling (Wills et al., 2002). It is likely that the differential effect of Abl on SliteRobo repulsive signaling is context dependent (Bashaw et al., 2000; Hsouna et al., 2003; Lee et al., 2004; Wills et al., 2002). In C. elegans, the UNC-34/Enabled protein is also known to modulate SAX-3-mediated repulsion (Yu et al., 2002). Another Abl-binding partner has also been implicated downstream of Robo. The microtubule-associated protein (MAP) Orbit/MAST was identified in Drosophila for its ability to genetically interact with Abl. This MAP is necessary for accurate axon guidance at the midline choice point (Lee et al., 2004). Epistasis analyses support a role for Orbit/MAST in modulating the action of Slit and its receptors by acting downstream of Abl (Lee et al., 2004).

7.5.3 Actin-interacting proteins The Drosophila actin-binding protein Canoe (Cno) promotes SliteRobo repulsive signaling by stabilizing Robo expression at the membrane of the growth cone (Slovakova et al., 2012). Loss of Cno expression resulted in decreased Robo localization in growth cone filopodia and rescued the commissureless phenotype observed in comm mutants, indicating that it is necessary for SliteRobo signaling (Slovakova et al., 2012). Genetic interactions with cno were also observed with several components of the SliteRobo signaling pathway, including sos and Rac GTPases. Sax3/Robo activation in C. elegans is also linked to the regulation of subcellular localization of the WAVE/SCAR actin nucleation complex during development of the embryonic epidermis (Bernadskaya et al., 2012). The regulated subcellular localization of this complex is thought to be essential for the localization of polarized F-actin during cell movement and may play a similar role during axonal repulsion.

7.6 Beyond the midline: additional roles for SliteRobo in the nervous system 7.6.1 Lateral positioning Following commissural axon midline crossing in the Drosophila ventral nerve cord, axons turn longitudinally and position themselves in lateral pathways next to the midline. Insight into the mechanisms that regulate the lateral positioning of these axonal fascicles was first provided by the observation that Robo proteins are differentially expressed in these axons. Axons that occupy the most medial pathway, adjacent to the midline, express only Robo1 (Rajagopalan et al., 2000b). Intermediate tract axons express Robo1 and Robo3 while axons belonging to the lateral pathway express all three Robos (Rajagopalan et al., 2000b; Simpson et al., 2000b) (Fig. 7.2A). The differential expression of Robo family members on longitudinal axons could provide a “Robo code” that mediates responsiveness to a gradient of midline-derived Slit proteins. Alternatively, lateral positioning of axons could be determined by the total levels of all Robo proteins on axons, with axons in the most lateral pathway expressing the highest levels of Robos. Loss- and gain-of-function analyses demonstrated that Robos contribute to lateral positioning of axon fascicles. Loss of Robo2 or Robo3 function leads to a shift in axon position closer to the midline, whereas gain of Robo2 or Robo3 function shifts axons away from the midline (Rajagopalan et al., 2000b; Simpson et al., 2000b). In contrast, Robo1 does not seem to play an instructive role in lateral positioning of commissural axons (Simpson et al., 2000a; Rajagopalan et al., 2000b; Evans and Bashaw, 2010). Structural differences between Robo family members may play a role in determining lateral positioning of axons. While the intracellular domain of Robo receptors is dispensable for lateral positioning (Evans and Bashaw, 2010; Spitzweck et al., 2010), differences in the extracellular regions of Robo1 and Robo2 appear to contribute to their divergent roles in lateral positioning. In medial apterous-expressing axons, which project directly adjacent to the FasII-positive axonal pathways within the ventral nerve cord, ectopic expression of Robo2 directs these axons to more lateral pathways (Evans and Bashaw, 2010). Furthermore, inserting the Ig1 and Ig3 domains of Robo2 into Robo1 is sufficient to confer lateral positioning activity to Robo1 (Evans and Bashaw, 2010). The differences in lateral positioning activity of Robo1 and Robo2 do not appear to be due to higher affinity of Slit for Robo2 (Evans and Bashaw, 2010; Spitzweck et al., 2010) but rather due to differences in multimerization and receptoreligand stoichiometry established by the Ig domains (Evans and Bashaw, 2010).

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While the gain-of-function experiments described earlier support a model whereby structural differences in Robo receptors underlie their specific activity in lateral positioning, Robo allele swapping experiments suggest that it is the overall levels of Robo receptors expressed on the axons that determine their lateral positioning (Spitzweck et al., 2010). Swapping robo1 or robo2 into the robo3 allele is sufficient to rescue lateral positioning defects observed in robo3 mutants, indicating that the overall levels of expression of Robo receptors on axons, rather than which type of receptor is expressed, instructs their lateral position (Spitzweck et al., 2010). Under these conditions, axons that express all three Robo family members could respond more strongly to a gradient of midline-secreted Slit and select the most lateral pathway. Thus, there exists convincing, yet conflicting, evidence in support of the two models proposing that either structural differences in Robos or total levels of Robo expression on axons contribute to the lateralizing effect of Robos. Additional experiments combining loss-of-function and allele swapping approaches will hopefully clarify the mechanism of action of Robo proteins in axon positioning lateral to the midline. In addition to promoting the positioning of longitudinal tracts, SliteRobo signaling also regulates their fasciculation and maintenance. In Drosophila, Slit is expressed within longitudinal tracts, and reduced Slit levels lead to the defasciculation and mixing of these tracts (Bhat, 2017). A function for SliteRobo signaling in the regulation of axonal fasciculation has also been reported in the vertebrate motor system where Slit2 and Robos are expressed by motor neurons and their ablation results in premature defasciculation of motor nerves at muscle targets (Jaworski and Tessier-Lavigne, 2012).

7.6.2 Cell migration and cell polarity Slits are well-characterized regulators of cell migration in nonneuronal as well as neuronal systems. In the nervous system, Slits are implicated in the control of neuronal precursor cell migration, precerebellar neurons, cortical pyramidal neurons, and interneurons. In the developing and mature brain, neuronal precursors from the SVZ migrate along the rostral migratory stream (RMS) to the olfactory bulb (OB) (Hu, 1999; Nguyen-Ba-Charvet et al., 2004; Sawamoto et al., 2006; Wu et al., 1999). Neuroblasts in the SVZ and in the RMS express the Slit receptors Robo2 and Robo3 (Nguyen-Ba-Charvet et al., 2004). It is thought that these neuroblasts respond to Slit during their migration, orienting so as to migrate toward the OB (Hu, 1999; Nguyen-Ba-Charvet et al., 2004; Sawamoto et al., 2006; Wu et al., 1999; Guerrero-Cazares et al., 2017). Importantly, in cultured SVZ explants or neuroblasts isolated from slit1 mutant mice, migration is altered (Nguyen-BaCharvet et al., 2004). In vivo, chains of migrating cells are misdirected and migrate toward the corpus callosum in slit1 mutants (Nguyen-Ba-Charvet et al., 2004). The choroid plexus (CP) is thought to be a source of repellents for migrating OB precursor cells (Hu, 1999). slit2 mutant mice show an absence of repulsive activity from the CP when it is cocultured along with SVZ explants, suggesting that the repulsive CP activity is indeed Slit2 (Nguyen-Ba-Charvet et al., 2004). In addition, the septum is also a source of Slit1 and Slit2 repellents for migrating OB neuroblasts (Nguyen-Ba-Charvet et al., 2004). SliteRobo signaling has also been implicated in the migration of human neural precursor cells from the subventricular zone to the OB at fetal stages of development (Guerrero-Cazares et al., 2017). Neurons comprising the hindbrain precerebellar nuclei are derived from neuroepithelium in the fourth ventricle and migrate along dorsoventral and anteroposterior axes during development to reach their eventual position in the adult brain. The expression of Netrin1 and Slits in the floor plate has been proposed to attract these neurons ventrally and prevent their crossing of the midline, respectively (Bloch-Gallego et al., 1999; Marcos et al., 2009; Sotelo and Chedotal, 2005; Yee et al., 1999; Geisen et al., 2008; Gilthorpe et al., 2002; Hammond et al., 2005). Precerebellar neurons express Robo1-3, and robo1;robo2 double knockout mice show disturbed migration that includes crossing by precerebellar neurons across the floor plate (Geisen et al., 2008; Marillat et al., 2002, 2004; Di Meglio et al., 2008). A similar phenotype is also observed in compound slit1/slit2 knockout embryos, supporting the idea that Slit expression in the floor plate prevents Roboexpressing precerebellar neurons from crossing the midline (Di Meglio et al., 2008). However, conditional ablation of robo2 in precerebellar neurons in a robo1 knockout background does not lead to defects in migration of these neurons, demonstrating a surprising noncell autonomous role for Robos in this process (Dominici et al., 2018). Furthermore, the specific ablation of Slit2 expression in the floor plate of slit1;slit3 double knockout embryos did not recapitulate the migration alterations observed in germline knockouts suggesting that floor plateederived Slits are not the main chemorepellent preventing these neurons from crossing the midline (Dominici et al., 2018). These observations emphasize the need to further examine and dissect the function of Slits and other axon guidance cues using cell typeespecific genetic ablation of these molecules. Slit and Robo also influence the migration of pyramidal neurons and cortical interneurons that populate the cortex (Andrews et al., 2006, 2008). Robos1 and 2 are expressed by cortical interneurons during their period of migration from the ganglionic eminence (GE) in the ventral telencephalon to the cortex (Andrews et al., 2008). Slit, secreted from the ventricular zone of the GE, was first thought to repel Robo-expressing interneurons to their correct positions in the cortex

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(Hu, 1999; Wu et al., 1999; Zhu et al., 1999); however, the lack of migratory defects in slit1;slit2 double mutants did not support this hypothesis (Andrews et al., 2008; Marin et al., 2003; Marin and Rubenstein, 2003). Interestingly, in robo1 mutant embryos, some cortical interneurons failed to be repelled by the striatum on their way to the cortex and invaded this region. Further studies demonstrated that Robo1 can associate with Npn1, a receptor for secreted semaphorins, which are chemorepulsive cues expressed in the striatum that can repel cortical interneurons (Marín et al., 2001). Loss of Robo1 expression led to decreased expression of Npn1 and decreased responsiveness to secreted semaphorins, resulting in invasion of the striatum (Hernandez-Miranda et al., 2011). Robo family members have also been implicated in the radial migration and positioning of pyramidal neurons within the cortex. Knockdown of either Robo1 or Robo4 expression leads to migration delays and misorientation of cortical pyramidal neurons, respectively (Zheng et al., 2012; Gonda et al., 2013). Interestingly, SliteRobo signaling also plays a role in regulating the maintenance of neuronal progenitor pools during development of these neurons (Borrell et al., 2012). Neuronal migration involves many changes in cell shape via membrane deformation and remodeling (Ayala et al., 2007). The SliteRobo effector proteins srGAPs (described earlier) have been implicated as regulators of neuronal migration and neurite initiation by influencing such changes in the cell membrane (Guerrier et al., 2009). The F-BAR domain of srGAP2 was shown to attenuate the formation of filopodia-like membrane protrusions in cortical neurons, whereas the F-BAR region of srGAP1 and srGAP3 promotes it (Guerrier et al., 2009; Coutinho-Budd et al., 2012). Furthermore, shRNA knockdown of srGAP2 expression in radial glial progenitors accelerates their migration in cortical slices, suggesting that srGAP2 negatively regulates radial glia migration (Guerrier et al., 2009). Although srGAP2 is known as a regulator of SliteRobo signaling, its functions in the context of neuronal migration have yet to be directly linked to Slit or Robo. An interesting question in the study of migration relates to how external cues, such as Slit, are able to direct cells to specific targets. While the activation of Rho GTPases by Slit can have a direct impact on cytoskeletal rearrangements necessary for migration, there is growing evidence that Slit influences cell polarity, which is essential for cytoskeleton organization (Killeen and Sybingo, 2008; Raman et al., 2018). In migrating cerebellar granule cells, Slit induces an asymmetric calcium burst in the leading process and in the soma of a migratory cell, and this is thought to regulate migration (Guan et al., 2007). Cell polarity in RGC axons extending toward the optic disk is also manipulated by Slits (Thompson et al., 2006). In migrating neuroblasts isolated from the SVZ and grown in culture, Slit induces a repositioning of the microtubule-organizing center at one pole of the cell to influence its migration (Higginbotham et al., 2006). Hence, the influence of Slits on neuronal migration relies on changes in the cytoskeleton of the cell. Whether these cytoskeletal rearrangements are induced directly through activation of Rho GTPases or are influenced by Slit-induced modulation of cell polarity proteins remains to be established.

7.6.3 Dendritic and axonal outgrowth and branching One fundamental issue in modern neuroscience is understanding how synapses between neurons are established. The morphological characteristics of a neuron, including the degree of branching and of elongation of both dendrites and axons, are crucial for the establishment of appropriate connectivity (London and Hausser, 2005). In addition to its repulsive role in axonal guidance, SliteRobo signaling is also implicated in axonal branching. This process is characterized by extensive branching of the primary axon tips or by collateral branching from primary axon shaft when or after the axonal growth cone reaches the target zone. The mechanisms that regulate branch formation are thought to be independent from those regulating axon targeting, although these mechanisms may often work in cooperation with one another (Yamamoto et al., 2002). In this context, Slit2 was isolated in a screen that targeted molecular regulators of axon branch initiation or extension (Wang et al., 1999). When applied to dissociated dorsal root ganglion cells, purified recombinant human Slit2 stimulates axon branching (Wang et al., 1999). Slit can also promote axonal branching in trigeminal ganglion explants isolated from the CNS (Ozdinler and Erzurumlu, 2002). Furthermore, loss-of-function genetic experiments also support a role for Slit in inducing axonal branching. Indeed, slit2;slit3 double and slit1;slit2;slit3 triple mutants have both reduced trigeminal sensory branching above the eye, and robo-1;robo-2 double mutant mice have a similar peripheral branching phenotype (Ma and Tessier-Lavigne, 2007). Interestingly, in zebrafish, overexpression of Slit2 promotes trigeminal sensory axons branching in a Robo2-dependant manner (Yeo et al., 2004). In contrast, Slit1a, a zebrafish slit ortholog (Hutson et al., 2003), negatively regulates arborization in the zebrafish retinotectal system and seems to also be a negative regulator of synaptogenesis (Campbell et al., 2007). The nature of this dual role still needs to be resolved. Another mechanism through which Slits can influence axonal branching is by binding DSCAM. In Drosophila, the binding of Slit to DSCAM enhances DSCAM interaction with the receptor protein tyrosine phosphatase RPTP69D, promoting the formation of collateral axon branches (Dascenco et al., 2015).

162 PART | I Formation of axons and dendrites

Mechanisms for regulating dendritic outgrowth or arborization have received less attention than those controlling axons (Chisholm and Tessier-Lavigne, 1999; Song and Poo, 1999). However, the extensive growth and branching of primary dendrites, after cell polarity is established, must be tightly regulated for correct synaptic connections to form. Several molecules that were first characterized for their roles in guiding axons have also been implicated in regulating dendritic morphology (Polleux et al., 2000; Furrer et al., 2003, 2007; Suli et al., 2006). The control of dendritic growth and branching by SliteRobo signaling was first demonstrated during in vitro experiments where Slit1 proteins were added to cortical neuron cultures. Slit1-treated neurons show increased dendritic length and more complex dendritic morphologies (Whitford et al., 2002). The stimulation of cortical dendrite branching is dependent on Robo receptors and their signaling through the adaptor protein Nck2 (Round and Sun, 2011). Furthermore, the ectopic expression of Robo in Drosophila giant fiber neurons causes defects in dendrite extension and guidance, whereas expression of Robo2 or Robo3 has no effect (Godenschwege et al., 2002). The authors, however, note that these experiments do not seem to reflect an endogenous role for Robo in this system (Godenschwege et al., 2002). Using single-cell labeling in Drosophila, Robo was shown to at least partially control midline crossing for both axons and dendrites (Furrer et al., 2003). The role of SliteRobo signaling in dendritogenesis was attributed to the control of timing, positioning, and size of dendrites in the embryonic CNS using the aCC motoneuron, one of the first CNS neurons to generate dendrites in the Drosophila embryonic CNS (Furrer et al., 2007). These proposed roles for SliteRobo signaling in dendrites are supported by a similar analysis in class IV multidendritic dendriteearborization (md-da) neurons in the Drosophila embryonic peripheral nervous system (Dimitrova et al., 2008). In vertebrates, SliteRobo signaling is required for dendritic selfavoidance during elaboration of dendritic fields (Gibson et al., 2014). During dendritic development, dendrites avoid overlapping to maximize receptive field coverage and minimize redundant signaling. In the mouse cerebellum, Purkinje cells express high levels of both Slit2 and Robo2, and ablation of either molecule specifically in Purkinje cells results in excessive dendritic overlap, demonstrating a cell-autonomous role for SliteRobo repulsion in dendritic self-avoidance (Gibson et al., 2014). In summary, SliteRobo signaling is important for both dendritic and axonal maturation during development. In an attempt to consolidate the differential effects of Robo signaling in the two neurite types, it has been proposed that distinct receptor combinations function in axons and dendrites (Hocking et al., 2010). This model relies on two main observations made in Xenopus RGCs. First, antisense knockdown and dominant-negative forms of Robo2 and Robo3 help demonstrate that Slit/Robo signaling is necessary to stimulate dendrite branching, primarily via Robo2 but has no role in RGC dendrite guidance (Hocking et al., 2010). Second, the dominant-negative Robo molecules inhibited the extension of axons and caused misrouting of some axons (Hocking et al., 2010). Another explanation for the differential effects of SliteRobo signaling on dendritic and axonal development might be that while axonal development relies on both cell- and nonecellautonomous SliteRobo signaling (Aoki et al., 2013; Dominici et al., 2018), dendritic development relies mainly on SliteRobo signaling through cell-autonomous mechanisms (Gibson et al., 2014).

7.7 SliteRobo contribution to axon targeting in a complex target field The study of axon guidance involves understanding not only how axons move from point to point and make decisions at intermediate targets but also how they innervate specific regions within complex target fields. The role of Slits and Robos in regulating target recognition is best exemplified in the mammalian olfactory systems. In the main olfactory system of the mouse, proper connectivity between olfactory sensory neurons (OSNs) located in the olfactory epithelium (OE) and second-order neurons in the OB allows for the detection and identification of odors from the environment. During development of these connections, OSN axons must navigate an intricate three-dimensional field to target specific regions termed glomeruli in the OB. SliteRobo signaling is critical for these axons to reach their appropriate positions in the dorsoventral axis (Cho et al., 2007). Robo2 is expressed in a high dorsomedial to low ventrolateral gradient in OSNs within the OE throughout development (Fig. 7.4A). OSN axons expressing high levels of Robo2 project to the dorsal region of the OB. Additionally, slit1 is expressed in the ventral region of the OB. In robo2 and slit1 mutant mice, a subset of axons that normally innervate the dorsal region of the OB mistarget to the ventral aspect of the OB (Cho et al., 2007). Furthermore, specific ablation of Robo2 expression in OSNs results in a ventral shift in the targeting of specific populations of OSNs, demonstrating that a cell-autonomous role for Robo2 is OSN axonal guidance (Cho et al., 2011, 2012). The expression of Robo1 in olfactory ensheathing cells is also necessary for the targeting of OSN axons, suggesting that Robo signaling contributes to OSN axon targeting through both cell- and nonecell-autonomous mechanisms (Aoki et al., 2013). In addition, SliteRobo interactions appear to control the entry of OSN axons in the OB. Double-mutant robo1;robo2 as well as slit1;slit2 mutant mice show ectopic OSN projections past the OB inside the diencephalon, along with

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(A)

(B)

FIGURE 7.4 Control of olfactory sensory neuron and vomeronasal sensory neuron axonal targeting by Slit and Robo. (A) Axonal targeting within the dorsoventral axis of the olfactory bulb (OB) is controlled by the graded expression of axon guidance cues and their receptors. The graded expression of the Slit receptor Robo2 in a high dorsal to low ventral gradient within the olfactory epithelium (OE) (yellow) is required to segregate axons emanating from olfactory sensory neurons (OSNs) in the dorsal region of the OE to the dorsal aspect of the OB. Expression of Slit1 in the ventral region of the OB prevents Robo2-expressing axons from entering the ventral region of the OB. (B) Vomeronasal sensory neuron (VSN) axon targeting involves the projection of apical neuron axons (black) to the anterior accessory olfactory bulb (AOB) and the projection of basal VSNs (yellow) to the posterior AOB. The graded expression of Slits extending from anterior (high) to posterior (low) guides Robo2-positive basal VSN axons to target to glomeruli in the posterior AOB.

perturbed convergence to some glomeruli (Nguyen-Ba-Charvet et al., 2008). This indicates that SliteRobo signaling has a dual role in controlling both the entry of OSN axons inside the OB and the convergence of axons into specific regions within the OB. SliteRobo interactions also play a role in controlling the projections emanating from second-order neurons in the OB to form the lateral olfactory tract (LOT). While Robo1 and Robo2 are expressed by LOT axons, Slit1 and Slit2 are secreted by the septum and function to repel LOT axons (Fouquet et al., 2007; Nguyen Ba-Charvet et al., 1999; Nguyen-Ba-Charvet et al., 2002). In robo2 and robo1;robo2 mutant mice, the LOT is completely disorganized, suggesting that these Robo receptors mediate Slit-induced repulsion of LOT axons and proper formation of the tract (Fouquet et al., 2007). Slits and Robos have also been implicated in the regulation of axonal targeting in the accessory olfactory system. Here, vomeronasal sensory neurons (VSNs) located in the basal region of the vomeronasal organ (VNO) project their axons to the posterior part of the accessory olfactory bulb (AOB), whereas VSNs in the apical region of the VNO project their axons to the anterior part of the AOB (Fig. 7.4B). SliteRobo signaling plays a critical role in ensuring the correct segregation of these two populations of axons within the AOB. While Robo1 is expressed by all VSNs, Robo2 expression is restricted to basal VSNs (Prince et al., 2009; Cloutier et al., 2004; Knoll et al., 2003). Slits1, 2, and 3 are all expressed in the AOB; however, prominent Slit1 expression is restricted to cells located at the anterior tip of the AOB (Prince et al., 2009; Cloutier et al., 2004; Knoll et al., 2003). Ablation of Robo2 expression in VSNs leads to mistargeting of subsets of basal VSN axons to the anterior region of the AOB (Fig. 7.4C). Similar defects are observed in slit mutant mice (Prince et al., 2009). Taken together, studies in the olfactory systems indicate that SliteRobo signaling plays a critical in the segregation of axons into specific regions of complex targeting fields.

164 PART | I Formation of axons and dendrites

7.8 Involvement of SliteRobo in disorders of the nervous system Despite their critical role in nervous system development, surprisingly few studies have so far linked Slits and Robos to neurological disorders. SLIT3 was found to be the most significant locus in a genome-wide association analysis in major depressive disorder (Glessner et al., 2010). Also, mutations in the ROBO3 gene were found in patients with a rare congenital syndrome (Jen et al., 2004). Horizontal gaze palsy with progressive scoliosis (HGPPS) is characterized by the absence of conjugate horizontal eye movement and also scoliosis that often requires surgical intervention early in life. Despite defects in conjugate horizontal eye movements, patients are reasonably coordinated and do not present with other obvious neurological deficits (Haller et al., 2008). In HGPSS patients, corticospinal and somatosensory axons fail to cross the midline to reach their appropriate targets, a phenotype reminiscent of the midline crossing defects observed in robo3 mutant mice (Renier et al., 2010). Initially, 10 different homozygous mutations scattered throughout the robo3 gene were identified, including 9 in the region encoding the extracellular domain (Jen et al., 2004). Since the first report, 11 additional mutations have been described in the literature (Chan et al., 2006; Abu-Amero et al., 2009; Amouri et al., 2009) or reported in the human gene mutation database. The observation that HGPSS is only observed in patients with mutations in both alleles of ROBO3 suggests that these mutations may be loss-of-function mutations. To better understand the etiology of HGPPS syndrome, a mouse model containing a robo3 deletion has been a valuable tool since both precerebellar neuron cell bodies and their axons fail to cross the midline in these mice (Marillat et al., 2004). The development of a floxed robo3 allele has provided important information on the role of Robo3 in various populations of neurons. The selective deletion of robo3 in two hindbrain rhombomeres, including rhombomere 5 comprising the abducens nucleus, leads to selective horizontal eye movement defects in mice (Renier et al., 2010). Moreover, when robo3 expression is ablated in inferior olivary neurons, the mice exhibit profound locomotor deficits, including an ataxic gait that persists into adulthood (Renier et al., 2010). Interestingly, motor performance deficits in these mice are so severe that their ataxia appears worse than in mice with a complete lack of cerebellar output. This ataxic behavior can be primarily attributed to the ipsilateral rerouting of a large number of olivocerebellar axons. The Rho-GTPase-activating protein srGAP3, which signals downstream of Robo, has also been associated with mental disorders. The SRGAP3 gene was initially found to be disrupted by a de novo balanced translocation in a patient with facial dysmorphism, hypotonia, and severe mental retardation (Endris et al., 2002). However, a subsequent sequencing of the SRGAP3 gene in patients with idiopathic mental retardation (n ¼ 95) or autism spectrum disorders (ASDs) (n ¼ 142) failed to identify any association between SRGAP3 haploinsufficiency and mental retardation (Hamdan et al., 2009). Another study found mutations in SRGAP2 in a patient presenting with early infantile epileptic encephalopathy (Saitsu et al., 2012), whereas two studies separately found mutations in SRGAP2 in patients with ASD (Dennis et al., 2012; Nuttle et al., 2013). Considering the wide array of functions of Slits during development, combined with the numerous mutations identified in genes involved in their signaling, it is likely that disruption of SliteRobo signaling contributes to a wide variety of neurodevelopmental disorders. Emerging evidence also supports a role for SliteRobo signaling in controlling repair and regeneration following nervous system injury. Ischemic stroke induced an upregulation of Slit2, Robo1, and Robo4 in mouse astrocytes, but not in neurons or microglia (Park et al., 2016). Slit2 and Robos may contribute to the formation of the astroglial scar that occurs following ischemic injury, since their expression was increased and sustained long term in the astroglial scar following ischemic injury in mice (Jin et al., 2016). In the peripheral nervous system, Slit1 and Robo2 are also upregulated in the dorsal root ganglia following sciatic nerve injury (Yi et al., 2006; Chen et al., 2012). srGAP3 expression was also increased, suggesting that SliteRobo signaling in regeneration following nerve injury may involve signaling through srGAP3.

7.9 Conclusion Many families of molecules that were originally identified for their role as potent regulators of axonal growth and guidance have now been shown to be critical in regulating a wide range of physiological processes both inside and outside of the nervous system, including axonal guidance, cell migration, cell adhesion, and the establishment of cell polarity. As outlined in this chapter, great progress has been made in understanding the cell intrinsic and extrinsic mechanisms that control the effect of Slits in several of these processes. More investigation will be necessary to fully understand how Slits can have diverse effects on different populations of neurons, or even on different compartments within a specific neuron. The recent identification of several new receptors that can mediate Slit function has provided some insight into this question; however, further understanding of the signaling pathways activated by these receptors in response to Slit will be necessary. In addition, the continued development and availability of more sophisticated approaches for gene manipulation in mice will

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allow scientists in the field to define further the role of Slits in specific populations of cells and assess the effect of Slitdependent wiring on specific behaviors. Hence, a combination of genetic, biochemical, and physiological approaches will provide further insight into the regulation of Slit function.

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Chapter 8

Nonconventional axon guidance cues: Hedgehog, TGF-b/BMP, and Wnts in axon guidance Patricia T. Yam1 and Fre´de´ric Charron1, 2, 3, 4, 5, * 1

Institut de Recherches Cliniques de Montréal (IRCM), Montreal, QC, Canada; 2Integrated Program in Neuroscience, McGill University, Montreal,

QC, Canada; 3Department of Anatomy and Cell Biology, Department of Biology, McGill University, Montreal, QC, Canada; 4Department of Medicine, University of Montreal, Montreal, QC, Canada; 5Division of Experimental Medicine, McGill University, Montreal, QC, Canada

Chapter outline 8.1. Introduction 176 8.1.1. Morphogens as axon guidance cues 176 8.2. Sonic hedgehog in axon guidance 176 8.2.1. Canonical Shh signaling 176 8.2.2. Shh is a chemoattractant for spinal cord commissural axons 178 8.2.3. Shh binding to Boc attracts commissural axons through a noncanonical signaling pathway to modulate the growth cone cytoskeleton 179 8.2.4. Shh guides axons along the longitudinal axis of the spinal cord 180 8.2.5. 14-3-3 proteins regulate a cell-intrinsic switch from Shh-mediated attraction to repulsion of commissural axons after midline crossing 180 8.2.6. Shh guides contralateral and ipsilateral retinal ganglion cell axons 181 8.2.7. Shh is a chemoattractant for midbrain dopaminergic axons 182 8.2.8. Shh binding to Gas1 repels enteric axons 182 8.3. TGF-b superfamily members in axon guidance 183 8.3.1. Canonical bone morphogenetic protein signaling 183 8.3.2. BMP7:GDF7 repels spinal cord commissural axons 183 8.4. Wnts in axon guidance 184 8.4.1. Canonical and noncanonical Wnt signaling 184 8.4.2. Wnt5 repels commissural axons from the Drosophila posterior commissure via derailed, a Ryk tyrosine kinase family member 186 8.4.3. Wnt5, complexed with derailed, repels Drosophila mushroom body axons 187

8.4.4. Wnt binding to Ryk repels axons of the corticospinal tract and corpus callosum through a Ca2þ-dependent mechanism 187 8.4.5. Wnt binding to Fz attracts postcrossing commissural axons via protein kinase C z and planar cell polarity signaling 188 8.4.6. Wnt binding to Fz regulates dopaminergic axon guidance through planar cell polarity signaling 189 8.4.7. Wnt3 mediates mediolateral retinotectal topographic mapping 190 8.4.8. Wnts guide axons of Caenorhabditis elegans mechanosensory neurons and D-type motoneurons via Fz-type receptors 190 8.4.9. The Wnt ligand CWN2 regulates Caenorhabditis elegans motor neuron axon guidance through a Rortype receptor CAM-1 192 8.5. Cross-talk between axon guidance cues 192 8.5.1. Shh induces the response of commissural axons to semaphorin repulsion during midline crossing 192 8.5.2. Shh regulates Wnt signaling in postcrossing commissural axons 193 8.5.3. The TGF-b family member unc-129 regulates Unc6/ Netrin signaling in Caenorhabditis elegans 193 8.6. Conclusions and perspectives 193 List of Acronyms and Abbreviations 194 Glossary 195 Acknowledgments 195 References 196

* Senior author.

Cellular Migration and Formation of Axons and Dendrites. https://doi.org/10.1016/B978-0-12-814407-7.00008-0 Copyright © 2020 Elsevier Inc. All rights reserved.

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176 PART | I Formation of axons and dendrites

8.1 Introduction During nervous system development, neurons project axons over long distances to reach their appropriate targets and establish neural circuits that are crucial for proper motor, sensory, and cognitive functions. Axons are tipped by a growth cone, which responds to diverse axon guidance cues as they navigate through the extracellular milieu to arrive at their target location. The axon guidance cues present in the extracellular environment can be attractive or repulsive. Four families of guidance cues were originally identified that mediate axon pathfinding: netrins, semaphorins, Slits, and ephrins (Dickson, 2002; O’Donnell et al., 2009). While these classical guidance molecules were found to regulate a variety of axon pathfinding events, they did not account for all axon guidance activities in the developing embryo. About 15 years ago, it was discovered that the morphogens Sonic hedgehog (Shh), bone morphogenetic proteins (BMPs), and Wnts can also guide axons. Now, we are uncovering the diversity of signaling pathways involved in morphogenmediated axon guidance. Some morphogens, such as BMPs and Wnts, regulate axon growth as well as guidance (Li et al., 2009; Phan et al., 2010), but this chapter will focus primarily on the guidance mechanisms of these molecules. We will also discuss the crosstalk between axon guidance cues.

8.1.1 Morphogens as axon guidance cues Morphogens are defined as diffusible molecules produced in a restricted region of a tissue that can impart specific differentiation programs to target cells through the establishment of a long-range concentration gradient. They typically regulate cell fate specification and tissue patterning during embryonic development. To be considered a morphogen, a signaling molecule must meet two criteria: (1) it must have a concentration-dependent effect on its target cells, and (2) it must exert a direct action at a distance. Members of the Hedgehog (Hh), TGF-b and Wnt families, in addition to fitting these criteria, also function as axon guidance molecules. The canonical Shh, BMP, and Wnt signaling pathways employed in cell fate specification transduce a signal to the nucleus, yet axon guidance by morphogens involves local changes at the growth cone. We will discuss here how the signaling pathways used by morphogens to guide axons diverge from the canonical signaling pathways used by morphogens to specify cell fate.

8.2 Sonic hedgehog in axon guidance 8.2.1 Canonical Shh signaling Hh proteins are found in insects and vertebrates but have no clear orthologs in nematodes. While a single Hh gene is present in flies, three exist in mammals: Shh, Indian hedgehog (Ihh), and Desert hedgehog (Dhh). In the neural tube, Shh is secreted at the ventral midline by the notochord and floor plate cells, where it functions as a graded signal for the generation of distinct classes of ventral neurons along the dorsoventral (DeV) axis (Dessaud et al., 2008; Jessell, 2000; Martı and Bovolenta, 2002). Consistent with its role as a morphogen, Shh induces a range of ventral spinal cord cell fates in a concentration-dependent manner and exerts a direct action at a distance to specify neural tube cell fate. Cell fate specification and tissue patterning activities of Hh molecules are mediated by canonical Hh signaling. Canonical Shh signaling (Fig. 8.1A) induces changes in the transcription of target genes via members of the Gli/Ci transcription factor family, which are zinc finger transcription factors that are capable of functioning as both repressors and activators. When Shh is absent, Ptch1 inhibits the activity of the transmembrane protein Smoothened (Smo), an activator of the pathway (Dessaud et al., 2008). Full-length Gli2, which functions as a transcriptional activator, is degraded, and Gli3 is proteolytically cleaved to generate Gli3R, which functions as repressor following nuclear translocation. Upon the binding of Shh to its receptors Ptch1, together with either Boc, Cdon, or Gas1, Ptch-mediated Smo inhibition is relieved (Allen et al., 2011; Dessaud et al., 2008; Izzi et al., 2011). Smo activation inhibits the proteolytic cleavage of Gli3 and the targeting of Gli2 for proteosomal degradation. Gli2 then translocates to the nucleus where it activates transcription of target genes involved in cell fate specification. In addition to its activity as a morphogen, Shh is also an important axon guidance cue for commissural neurons (Bourikas et al., 2005; Charron et al., 2003; Yam et al., 2012), retinal ganglion cells (RGCs) (Fabre et al., 2010; Gordon et al., 2010; Sánchez-Camacho and Bovolenta, 2008; Trousse et al., 2001), midbrain dopaminergic neurons (Hammond et al., 2009), and enteric neurons (Jin et al., 2015). It guides axons through a transcription-independent noncanonical signaling pathway(s), distinct from the canonical Shh signaling pathway that specifies cell fate.

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(A) canonical Shh signaling Boc/Cdon/ Gas1

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FIGURE 8.1 Canonical and noncanonical Shh signaling pathways. (A) Canonical Shh signaling. In the absence of Shh, Ptch1 inhibits the activity of Smo. Full-length Gli2 is targeted for proteasomal degradation, whereas full-length Gli3 is proteolytically cleaved to generate the Gli3 repressor form (Gli3R), which blocks transcription. Shh induces signaling by binding to its receptors Ptch1 together with Boc, Cdon, or Gas1, which relieves Ptch1mediated inhibition of Smo. Smo activation elicits a downstream signaling cascade that ultimately results in proteolytic cleavage of Gli2 and its translocation to the nucleus where it induces transcription of target genes. (B) In precrossing commissural axon attraction, the Dock/ELMO complex is bound to Boc in the absence of ligand. Shh binding to Boc releases Dock/ELMO from Boc and relocalizes it to the growth cone periphery where it activates Rac. Shh binding to Boc also leads to Smo activation and activation of SFKs. SFKs phosphorylate ZBP1, releasing b-actin mRNA and leading to local translation of b-actin. (C) In contralateral RGCs, Shh acts via a Smo-dependent pathway to repel RGC axons through PKCa and ILK. Note that Ptch1 is assumed to also bind Shh in axon guidance, but it has not been directly proven. Shh, Sonic hedgehog; SFK, Src family kinase; RGC, retinal ganglion cell; PKCa, protein kinase C a; ILK, integrin-linked kinase.

178 PART | I Formation of axons and dendrites

8.2.2 Shh is a chemoattractant for spinal cord commissural axons During spinal cord development, commissural neurons, which are located in the dorsal neural tube, project axons that extend along the DeV axis toward the floor plate (Fig. 8.2). Upon reaching the floor plate, they cross the floor plate to form axon commissures and then turn anteriorly and migrate along the anteroposterior (AeP) axis to their targets in the brain (Colamarino and Tessier-Lavigne, 1995). One major chemoattractant that guides commissural axons toward the midline is netrin1 (Kennedy et al., 1994; Serafini et al., 1994, 1996; Varadarajan et al., 2017). A detailed analysis of netrin1 null mice demonstrated that even in the absence of netrin1, some commissural axons do reach the midline, suggesting that additional diffusible attractant(s) might be expressed by floor plate cells to help guide commissural axons toward the midline (Charron et al., 2003; Serafini et al., 1996). Shh, which is expressed by the floor plate at this stage of neural development and has long-range effects in the spinal cord, was shown to function as an attractive guidance cue for commissural axons, mimicking the netrin1-independent chemoattractant activity of the floor plate in in vitro spinal cord explant turning assays (Charron et al., 2003). Given that Shh is a potent morphogen, its ability to reorient the trajectory of commissural axons toward the source of Shh in spinal cord explant turning assays could be indirect. Shh could repattern the explant, thus altering the expression of other guidance cues within the explant, which then secondarily cause axon reorientation. However, analysis of a panel of DeV patterning markers showed that they were unaffected by Shh and revealed that the spinal cord explants used to assess chemoattractant activity were no longer competent to be repatterned by Shh. Thus, the chemoattractive effect of Shh on commissural axons was likely to be direct (Charron et al., 2003). Three different approaches were used to demonstrate that Shh functions as a direct chemoattractant cue for commissural axons. Firstly, crossing of Wnt1-Cre mice, which specifically express Cre recombinase in the dorsal neural tube, with mice bearing a floxed allele of Smo, gave rise to mutant embryos with commissural axon trajectories that were defective in the ventral spinal cord (Charron et al., 2003). Because Cre is not expressed in the ventral spinal cord, this genetic evidence strongly implies that the axonal misrouting is not due to repatterning or other changes in the ventral spinal cord but instead reflects a guidance defect arising from loss of Smo in commissural neurons. In a second approach, Shh induced the turning of growth cones of isolated Xenopus spinal neurons in dissociated cell culture, confirming that Shh can function as a chemoattractant for Xenopus spinal axons (Charron et al., 2003). Finally, in a third approach, dissociated rat commissural axons turned up a Shh gradient in an in vitro Dunn chamber turning assay, an assay where the angle turned of axons exposed to a Shh gradient can be measured over a short time period (Yam et al., 2009). Taken together, these results suggest that Shh functions to guide commissural axons both in vitro and in vivo by acting directly as a chemoattractant on these axons. Although both Shh and netrin1 are secreted by the floor plate and guide commissural axons, there are important differences between the two guidance cues. Shh is secreted solely from the floor plate, forms a gradient in the ventral spinal cord, and guides commissural axons over the last 200e300 mm toward the floor plate (Sloan et al., 2015). However, netrin1 is produced by both the ventricular zone and the floor plate (Kennedy et al., 1994; Serafini et al., 1996; Varadarajan

D

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FIGURE 8.2 Axon guidance cues in the neural tube. Left. Precrossing commissural axons are repelled from the roof plate by BMPs and attracted to the floor plate by Shh, netrin1, and VEGF. Right. Following midline crossing at the floor plate, commissural axons migrate anteriorly due to an attractive Wnt4 gradient and a repulsive Shh gradient. D, dorsal; V, ventral; A, anterior; P, posterior; BMP, bone morphogenetic protein; VEGF, vascular endothelial growth factor. Reprinted from Yam, P.T., Charron, F. Signaling mechanisms of non-conventional axon guidance cues: the Shh, BMP and Wnt morphogens. Current Opinion in Neurobiology 23, 965e973 Copyright (2013), with permission from Elsevier.

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et al., 2017; Wu et al., 2019). Ventricular zone netrin guides commissural axons locally at the pial edge in the dorsal and ventral spinal cord (Varadarajan et al., 2017), whereas floor plate netrin (FP-netrin1), like Shh, guides commissural axons at long range over the last 200e300 mm toward the floor plate (Wu et al., 2019). Analysis of mice with a deletion of FPnetrin1 and Boc (the receptor for Shh) shows that FP-netrin1 and Shh collaborate to guide commissural axons in the ventral spinal cord. However, the individual FP-netrin1 and Boc mutants do not have the same commissural axon guidance phenotype in the ventral spinal cord, most likely owing to the fact the netrin1 promotes axon outgrowth in addition to growth cone turning and thus may also act permissively, whereas Shh only promotes growth cone turning and has no effect on axon outgrowth (Wu et al., 2019).

8.2.3 Shh binding to Boc attracts commissural axons through a noncanonical signaling pathway to modulate the growth cone cytoskeleton Shh-mediated guidance of commissural axons requires Smo, since conditional removal of Smo in commissural neurons generated mice with defects in their commissural axon trajectories in the ventral spinal cord (Charron et al., 2003). This was further corroborated with in vitro experiments where treatment of commissural neurons with SANT-1, a highly specific Smo antagonist, prior to exposure to the Shh gradient, abrogated axon turning toward a Shh gradient in vitro (Yam et al., 2009). Thus, similar to canonical Shh signaling, Shh-mediated axon guidance requires the signaling activator Smo in a cell autonomous manner. However, Shh induces commissural axon growth cone turning within 10 min (Yam et al., 2009). This rapid growth cone response suggests that, for axon guidance, Shh does not signal to the nucleus via the canonical pathway but instead acts rapidly and locally at the growth cone (Charron and Tessier-Lavigne, 2005; Yam et al., 2009). Indeed, commissural neurons can still respond to Shh and reorient up a gradient of Shh even when global transcriptional activity in neurons is pharmacologically inhibited, or when Gli-mediated transcription is specifically inhibited by Gli3R expression (Yam et al., 2009). This provided the first direct evidence that Shh guides axons through a novel signaling pathway that is different to the transcription-dependent canonical Shh pathway (Fig. 8.1B). Shh-mediated attraction of commissural axons requires the receptor Boc (Okada et al., 2006) (Fig. 8.1B). Boc (together with its homolog Cdon) are type I transmembrane proteins that consist of 4e5 immunoglobulin (Ig) domains and two to three fibronectin type III (FNIII) repeats in its extracellular domain, making it closely related to the axon guidance receptors of the Robo and DCC families (Okada et al., 2006). Unlike canonical Shh signaling where Boc acts redundantly with Cdon and Gas1 to receive the Shh signal (Allen et al., 2011; Izzi et al., 2011), in commissural axon guidance, Boc is absolutely required to receive the Shh signal. Boc/ mice, but not Cdon/ mice, have abnormal commissural axon projections that are highly dispersed and invade the motor column (Okada et al., 2006), similar to the phenotype of conditional removal of Smo in commissural neurons. Downstream of Boc, Shh activates Src family kinases (SFKs) in a Smo-dependent manner to guide commissural axons (Yam et al., 2009) (Fig. 8.1B). Stimulation of commissural neurons with Shh increased the kinase activity of Src and Fyn. Active SFKs are characterized by phosphorylation at Y418, and Shh stimulation also increased phospho-SFK at Y418, as detected by immunofluorescence and western blotting. Furthermore, Shh stimulates local changes at the growth cone, with extrinsic Shh gradients inducing a polarized distribution of activated SFKs within the growth cone within 30 min. Interestingly, the highest levels of activated SFKs are found on the growth cone side facing the highest concentration of Shh. Inhibition of SFK activity with PP2, a pharmacological inhibitor, and C-terminal Src kinase, a negative regulator of SFKs, both blocked the ability of commissural neurons to respond to Shh in the in vitro Dunn chamber turning assays and also spinal cord explant turning assays. In contrast, SFK inhibition by PP2 had no effect on Shh induction of Gli-luciferase reporter activity, a standard assay for Shh-dependent canonical transcriptional activity (Yam et al., 2009). Thus SFKs are required for Shh-mediated guidance of commissural axons, but not for induction of Gli transcriptional activity, indicating a point of divergence between canonical and noncanonical Shh pathways. Axon guidance signaling ultimately leads to cytoskeletal remodeling that drives growth cone turning. We have begun to elucidate how Shh regulates the changes in the actin cytoskeleton that are required for growth cone turning. One target of SFKs is zipcode-binding protein 1 (ZBP1/Igf2bp1), an mRNA-binding protein that transports b-actin mRNA and releases it for local translation upon phosphorylation. Shh increases b-actin protein levels in the growth cone in an SFK-dependent manner (Lepelletier et al., 2017). The Shh-mediated increase in b-actin protein occurs even when the cell bodies have been removed, demonstrating that it is the result of local translation. Furthermore, a Shh gradient polarizes the b-actin distribution in the growth cone, illustrating that Shh gradients act locally at the growth cone to regulate b-actin. Shh increases ZBP1 phosphorylation in the growth cone, and disruption of ZBP1 phosphorylation in vitro abolished the turning of commissural axons toward a Shh gradient. Disruption of ZBP1 function in vivo resulted in commissural axon guidance

180 PART | I Formation of axons and dendrites

errors that phenocopy the defects seen in Boc/ and Smo conditional mice. Therefore, ZBP1 is required for Shh-mediated axon guidance by transporting and releasing b-actin mRNA for the local translation of b-actin (Fig. 8.1B). The newly synthesized pool of b-actin can then be used to promote polarized actin polymerization. Recently, Makihara et al. (2018) discovered that Shh-mediated axon guidance requires the activity of the unconventional guanine nucleotide exchange factors (GEFs), Dock3 and 4, and their binding partner ELMO1 and 2 (Makihara et al., 2018) (Fig. 8.1B). In vitro knockdown of Dock3 and 4 and ELMO1 and 2 inhibits the ability of commissural neurons to turn up a Shh gradient in a Dunn chamber assay. Likewise, in vivo knockdown of Dock3 and 4 and ELMO1 and 2 in mice leads to defects in the trajectory of commissural axons. GEFs are key regulators of Rho-GTPases, which are extremely important for the precise spatiotemporal regulation of actin cytoskeleton dynamics (Dickson, 2001; Hall and Lalli, 2010). GEFs promote Rho-GTPase activity by exchanging GDP with GTP to promote the active GTP-bound state. In commissural neurons, Shh induces activation of the RhoGTPase, Rac1, in a Dock3/4-dependent manner. The receptor for Shh, Boc, binds Dock through ELMO, and Shh stimulation reduces the interaction between Boc and the Dock/ELMO complex. The Dock/ELMO complex then relocates to the growth cone periphery, where it activates Rac1 to enable growth cone turning (Makihara et al., 2018) (Fig. 8.1B). Furthermore, modulating Dock activity in a polarized manner is sufficient to induce growth cone turning. This demonstrates that modulation of GEF activity is sufficient to elicit growth cone turning and highlights the instructive role of GEFs in axon guidance. Shh signaling through pSFK/ZBP1 and Dock/ELMO induces changes in the levels of b-actin and active Rac1, respectively, both of which contribute to the regulation of actin cytoskeleton assembly. This highlights how modulation of the growth cone cytoskeleton is a key step in Shh-mediated axon guidance. Whether or not the pSFK/ZBP1 and Dock/ ELMO pathways are independent or whether pSFK may also regulate Dock/ELMO activity is an exciting area for future research.

8.2.4 Shh guides axons along the longitudinal axis of the spinal cord Once commissural axons reach and cross the floor plate at the midline, they make a sharp turn and migrate anteriorly toward the brain. In addition to attracting commissural axons along the DeV axis toward the floor plate, a posterior-high/ anterior-low Shh gradient guides postcrossing commissural axons along the AeP axis of the spinal cord (Bourikas et al., 2005; Yam et al., 2012) (Fig. 8.2). Using in vitro Dunn chamber turning assays with commissural neurons, Shh was shown to have a direct effect on the guidance of postcrossing commissural axons by repelling them (Yam et al., 2012). Shh secreted from the ventral floor plate has also been suggested to guide descending serotonergic axons in the raphespinal tract via Ptch1 and Smo (Song et al., 2012). Current evidence suggests that the Shh signaling pathway in the guidance of postcrossing commissural axons differs between chick and rodents. In chick, the receptor for Shh repulsion of postcommissural axons is Hhip1, a Shh-binding membrane protein transiently expressed in the periventricular region and in commissural neurons at the time of floor plate midline crossing (Bourikas et al., 2005). Hhip expression itself is regulated transcriptionally by Shh, and this depends on the heparan sulfate proteoglycan, glypican1 (GPC1), with loss of GPC1 also causing commissural axon guidance defects in chick (Wilson and Stoeckli, 2013). However, in Hhip1/ mice, postcrossing commissural axons project correctly along the AeP axis (Yam et al., 2012), suggesting that Hhip1 is not required for postcrossing commissural axon guidance in mice. The Shh receptor for postcrossing commissural axon guidance in mice remains to be identified. In rodents, Shh-mediated postcrossing commissural axon guidance requires Smo. Genetic inactivation of Smo specifically in commissural neurons in mice (Yam et al., 2012) and downregulation of Smo in rat open-book cultures (Parra and Zou, 2010) leads to AeP guidance defects and demonstrate that Smo is required cell autonomously for AeP guidance of postcrossing commissural axons. However, in chick, postcrossing commissural axon guidance by Shh does not depend on Smo (Bourikas et al., 2005). Since Wnt-mediated axon guidance in chick can be regulated differently to that in rodents (see Section 8.5.2 “Shh regulates Wnt signaling in postcrossing commissural axons”), it is possible that Shh signaling mechanisms also differ between chick and rodents.

8.2.5 14-3-3 proteins regulate a cell-intrinsic switch from Shh-mediated attraction to repulsion of commissural axons after midline crossing Commissural axons are initially attracted by Shh along the D-V axis (precrossing). After crossing the floor plate, commissural axons switch their response to Shh from attraction to repulsion, so that they are repelled anteriorly by a decreasing posteroanterior Shh gradient along the longitudinal axis (Bourikas et al., 2005; Yam et al., 2012) (Fig. 8.2). This

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switch from Shh-mediated attraction to repulsion also occurs in vitro with dissociated commissural neurons as they age in culture (Yam et al., 2012). This recapitulates the change in response to Shh between pre- and postcrossing commissural axons in vivo, indicating that the switch in response is cell intrinsic and time dependent. The switch in polarity of the turning response depends on 14-3-3 proteins, which are enriched in postcrossing commissural axons. 14-3-3 protein inhibition converted Shh-mediated repulsion of aged dissociated neutrons to attraction and prevented the correct anterior turn of postcrossing commissural axons in vivo, an effect mediated by protein kinase A (PKA). Conversely, premature overexpression of 14-3-3 proteins in vitro and in vivo drives the switch in Shh responsiveness from attraction to repulsion. Thus, modulating 14-3-3 protein levels is sufficient to change the polarity of the turning response of commissural axons to Shh gradients. Notably, while 14-3-3 proteins regulate the turning response to Shh downstream of Shh reception, Shh signaling itself does not seem to regulate 14-3-3 levels or activity (Yam et al., 2012).

8.2.6 Shh guides contralateral and ipsilateral retinal ganglion cell axons In the visual system, RGCs extend axons from the retina through the optic nerve toward the diencephalic ventral midline (Fig. 8.3A). Their primary target is the superior colliculus in mammals or the optic tectum in chick. In chick, which has no binocular vision, all RGC axons project contralaterally. In vertebrates with frontally located eyes, subpopulations of RGC axons segregate at the optic chiasm to project to targets on the ipsilateral or contralateral sides of the brain and establish binocular vision. Contralateral axons cross the midline at the chiasm, whereas ipsilateral axons deviate from the chiasm and continue in the ipsilateral optic tract. Shh guides RGC axons at the optic chiasm (Fabre et al., 2010; Sánchez-Camacho and Bovolenta, 2008; Trousse et al., 2001) and the optic tract (Gordon et al., 2010). Shh expression at the chiasm border defines a constrained pathway within the ventral midline that guides contralateral RGC axons (Sánchez-Camacho and Bovolenta, 2008; Trousse et al., 2001) (Fig. 8.3A). The receptor(s) involved in Shh-mediated axon guidance of contralateral RGCs have not yet been determined.

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optic tract optic chiasm FIGURE 8.3 Shh guides contralateral and ipsilateral retinal ganglion cell axons. (A) Shh bordering the optic chiasm (red) guides contralateral RGCs through the optic chiasm. At the optic chiasm, VEGF-A is also a chemoattractant for contralateral RGCs. In mice, ipsilateral RGCs, which originate in the ventrotemporal retina, are repelled from the optic chiasm by remotely produced Shh and locally produced EphrinB2, through their receptors Boc and EphB1, respectively. (B) Shh (orange), produced by contralateral RGC cell bodies of the retina and transported anterogradely down the axon, is secreted at the optic chiasm. Ipsilateral RGCs, arriving after contralateral axons at the chiasm, are repelled from the midline by this axon-derived Shh, into the ipsilateral tract. (C) Knockdown of Shh in contralateral RGCs reduces the proportion of ipsilateral axons in a nonecell-autonomous manner. D, dorsal; V, ventral; A, anterior; P, posterior; N, nasal; T, temporal; Shh, Sonic hedgehog; RGC, retinal ganglion cell; VEGF, vascular endothelial growth factor.

182 PART | I Formation of axons and dendrites

However, some of the downstream signaling molecules have been elucidated. In rodents, inhibition of Smo by Ptch1Dloop2 in RGCs suggests that Shh controls the pathfinding of contralateral RGC axons cell autonomously through Smo (SánchezCamacho and Bovolenta, 2008). In chick, Shh rapidly activates protein kinase C a (PKCa) and integrin-linked kinase (ILK) in RGC growth cones. Expression of a dominant-negative PKCa or ILK inhibited Shh repulsion of RGC axons in vitro and resulted in aberrant RGC axon pathfinding at the optic chiasm in vivo, demonstrating that Shh guides contralateral RGC axons through PKCa and ILK (Guo et al., 2012) (Fig. 8.1C). Shh also induces macropinocytosis in RGCs, and this is important for Shh-mediated RGC growth cone repulsion and collapse in vitro (Kolpak et al., 2009), although the in vivo relevance of this observation has yet to be demonstrated. The guidance of ipsilateral RGCs is also dependent on Shh (Fig. 8.3A). Boc expressed in ipsilateral RGCs allows them to be repelled by Shh at the optic chiasm in a Smo-dependent manner. In vitro assays demonstrated that while RGC axons expressing high levels of Boc retracted in response to Shh, axons with low levels of Boc did not. Boc/ mice have almost 50% fewer ipsilateral projections than wild-type mice (Fabre et al., 2010). Conversely, gainof-function experiments demonstrated that expression of Boc in contralateral RGCs resulted in their projection to the ipsilateral side of the brain. Together, these experiments demonstrate that Boc mediates Shh-dependent repulsion of RGC axons and is essential for the correct pathfinding of ipsilateral RGCs at the optic chiasm (Fabre et al., 2010). Since Boc is not expressed in contralateral RGCs (Fabre et al., 2010), it is unlikely to be involved in contralateral RGC axon guidance. Shh repulsion of ipsilateral RGCs also depends on axoneaxon interactions (Fig. 8.3B). Shh mRNA is absent from the optic chiasm itself; thus, Shh is not synthesized locally at the chiasm. Instead, Shh at the chiasm comes from Shh produced by contralateral RGCs in the retina, which is then transported anterogradely along the axon and accumulates at the optic chiasm (Peng et al., 2018). Ipsilateral RGCs, which arrive at the chiasm after contralateral RGCs, are then repelled by this source of Shh (Fig. 8.3B). In vitro, contralateral RGC axons, which secrete Shh, repel ipsilateral RGCs in a Boc- and Smodependent manner. In vivo, knockdown of Shh in the contralateral retina causes a decrease in the proportion of ipsilateral RGCs in a nonecell-autonomous manner (Peng et al., 2018) (Fig. 8.3C), demonstrating that contralateral RGCs produce Shh remotely to guide ipsilateral RGCs at the optic chiasm.

8.2.7 Shh is a chemoattractant for midbrain dopaminergic axons In mammals, midbrain dopaminergic neurons (mDNs) migrate rostrally toward the forebrain. Shh is expressed at the ventral midline of the midbrain, adjacent to the location of mDNs. In vitro cultures of midbrain explants revealed that dopaminergic axons can grow toward the source of Shh in a Smo-dependent manner (Hammond et al., 2009). Conditional inactivation of Smo using Nestin-cre, which drives Cre expression throughout the CNS, leads to reduced and misprojected medial mDN axons (Hammond et al., 2009). In contrast, projections of lateral mDN were not affected. Marker analysis suggested that hypothalamic, forebrain, and ventral midline structures differentiated normally in Nestrin-cre;Smo conditional knockout mice, supporting the idea that the phenotype observed in medial mDN is due to a guidance defect related to Shh signaling and probably not due to an abnormal differentiation of the tissues (Hammond et al., 2009).

8.2.8 Shh binding to Gas1 repels enteric axons Enteric neurons, which control digestive functions such as gut movement, extend their axons between two peripheral smooth muscle layers to form a tubular meshwork arborizing the gut wall. Shh secreted from the gut epithelium prevents enteric axons from projecting centrally into the intestinal villi, thereby forcing their peripheral tubular distribution (Jin et al., 2015). Both mutants in Shh and its receptor Gas1 have enteric axon projection defects. Furthermore, Wnt1-Cremediated conditional inactivation of Gas1 and Smo in the dorsal neural tube and neural crest cells (the cells of origin of enteric neurons) also generate mice with more centrally projecting enteric axons, with no effect on enteric progenitor or neuron positioning. Enteric axons in vitro are also repelled from Shh in a Gas1-dependent manner. Together, these results suggest that Gas1 and Smo act cell autonomously to mediate Shh signaling in enteric neurons to prevent central projections (Jin et al., 2015). Given that Gas1 is a GPI-anchored protein, it is unlikely to directly recruit cytoplasmic signaling mediators, unlike the cytoplasmic tail of Boc, which binds to the ELMO/Dock complex in commissural neurons. Of the Gai proteins that can couple with Smo, G protein a Z (Gnaz) is found in enteric axons. Knockdown and dominant-negative inhibition of Gnaz reduced the turning response of enteric axons to Shh in vitro, and Gnaz mutant intestines contained more centrally projecting enteric axons in the intestinal villi. Therefore, Gnaz is an intracellular effector of Shh-mediated repulsion of enteric axons (Jin et al., 2015).

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8.3 TGF-b superfamily members in axon guidance 8.3.1 Canonical bone morphogenetic protein signaling The TGF-b superfamily is a large family of growth factors that comprises at least 50 distinct molecules identified in mammals, fish, worms, and flies. Members of this family include the prototypic TGF-bs, the BMPs, the activins, and the growth and differentiation factors (GDFs) (Attisano and Wrana, 2002; Massagué, 1998; Schmierer and Hill, 2007). Roof plate cells, which are located at the dorsal midline of the neural tube, produce several inductive signals that control the specification of dorsal neural tube cell types (Lee and Jessell, 1999). Many members of the TGF-b/BMP family are expressed during the time of dorsal neuron generation, and some, such as GDF7, are required for the normal specification of dorsal neurons (Lee et al., 1998). Members of the TGF-b superfamily induce signaling by bringing together a heteromeric complex of type I and type II serine/threonine kinase receptors (Attisano and Wrana, 2002; Massagué, 1998) (Fig. 8.4A). Upon ligand binding, type II receptors transphosphorylate the type I receptors. Once activated, the type I receptor kinase directly phosphorylates receptor-regulated Smads (R-Smads), which can then associate with the co-Smad, Smad4, and translocate to the nucleus where they interact with DNA-binding proteins to regulate transcriptional responses that control cell fate specification (Attisano and Wrana, 2002; Massagué, 1998). BMPs also guide commissural axons through type I and type II BMP receptors (BMPRs). However, there are differences in the role of individual receptor subunits, and current data suggest that the activation of particular BMPR complexes can lead to specific downstream signaling events in axon guidance that differ from that in cell fate specification.

8.3.2 BMP7:GDF7 repels spinal cord commissural axons Although commissural neurons from netrin1 and Dcc-null mice display severe guidance defects, their initial trajectory from the roof plate in the dorsal region of the spinal cord appears normal (Fazeli et al., 1997; Serafini et al., 1996). This suggests that an additional guidance cue might be controlling the dorsal migration of commissural axons. This was confirmed when roof plate tissue appended next to dorsal spinal cord explants deflected commissural axons in spinal cord

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FIGURE 8.4 Canonical and noncanonical BMP signaling pathways. (A) Canonical BMP signaling. Ligand binding initiates TGF-b/BMP signaling by inducing the heterodimerization of type I and type II serine/threonine kinase receptors. The type II receptor transphosphorylates and activates the type I receptor, which in turn phosphorylates and activates the R-Smads. Activated R-Smads associate with the co-Smad, Smad4 and translocate to the nucleus where they interact with transcription factors to regulate gene transcription. (B) Precrossing commissural axon repulsion is mediated by BMP7:GDF7 binding to BMPRIB:BMPRII, followed by PI3K activation. BMP, bone morphogenetic protein; TGF-b, Transforming growth factor b.

184 PART | I Formation of axons and dendrites

explant turning assays (Augsburger et al., 1999). BMP7, which is expressed by the roof plate, mimics the repellent activity of the roof plate in vitro, without affecting spinal cord cell fate specification, at the concentrations used for chemorepulsion. Inhibition of BMP7 activity with Follistatin, function-blocking antibodies, and genetic inactivation of Bmp7 showed that BMP7 contributed to the chemorepellent activity of the roof plate for commissural axons (Augsburger et al., 1999). Subsequent experiments demonstrated that genetic ablation of Gdf7, a more divergent member of the TGF-b superfamily, reduced the ability of the roof plate to deflect commissural axons to the same extent as elimination of Bmp7 (Butler and Dodd, 2003). Indeed, biochemical studies show that GDF7 and BMP7 form heterodimers and that coexpression of both molecules enhanced the chemorepellent activity of either factor alone on commissural neurons. Thus, BMP7:GDF7 heterodimers secreted by the roof plate repel commissural axons ventrally (Augsburger et al., 1999; Butler and Dodd, 2003) (Fig. 8.2). The minimum BMP7 concentration required to elicit growth cone collapse is over two orders of magnitude lower than that for inducing cell fate changes (Perron and Dodd, 2011). Together with evidence that the highly related BMP6 has no effect on growth cone collapse (Perron and Dodd, 2011) and only weakly repels axons (Augsburger et al., 1999), this suggests that there may be selective engagement of receptor complexes specific to axon guidance by BMP7 that can occur at low BMP7 concentrations. Indeed, whereas both type I BMPRs (BMPRIA and BMPRIB) contribute to the specification of dorsal cell fates in the spinal cord, axon guidance is principally mediated by BMPRIB (Yamauchi et al., 2008) (Fig. 8.4B). BMPRIB, but not BMPRIA, is expressed by postmitotic dorsal neurons, and it is necessary for the correct axon guidance response to roof plateesecreted cues. The commissural axon mispolarization phenotype in the dorsal spinal cord of BmprIb/- mutants is similar to that observed in Bmp7/ and Gdf7/ mice (Yamauchi et al., 2008). Using a pharmacological inhibitor of BMPRI kinase activity, it has also been shown that although BMPRIB is required for BMP-induced axon repulsion, BMPRI kinase activity, which is required for cell fate specification, is not required for axon guidance (Perron and Dodd, 2011). Consistent with BMPRI kinase activity not being required for axon guidance, downstream of receptor engagement, at BMP7 concentrations that can elicit growth cone collapse, R-Smads are not phosphorylated (Perron and Dodd, 2011), suggesting that BMP-mediated axon guidance is independent of R-Smad signaling. This is consistent with the rapid growth cone collapse response to BMP7 (Augsburger et al., 1999), which argues against BMP7 acting through a transcriptionbased mechanism. Instead, experiments with spinal cord explants and dissociated spinal neurons demonstrated that BMP7-mediated axon repulsion requires phosphatidylinositol-3-kinase (PI3K) activity (Fig. 8.4B), which is not implicated in classical BMP signaling for cell fate induction (Perron and Dodd, 2011). In vitro experiments with Xenopus spinal neurons suggest that BMP7 can also induce growth cone turning by controlling cofilin levels (Wen et al., 2007); however, further experiments are needed to see if this translates to an in vivo role for cofilin in BMP-mediated commissural axon guidance. In fact, experiments in mouse and chick show that LIM kinase phosphorylation of cofilin regulates commissural axon growth but not guidance downstream of BMP7 (Phan et al., 2010; Yamauchi et al., 2013).

8.4 Wnts in axon guidance 8.4.1 Canonical and noncanonical Wnt signaling Wnts are a large family of ligands that can activate at least three different signal transduction pathways (Niehrs, 2012; Nusse, 2012). The canonical b-catenin-dependent pathway is primarily involved in cell differentiation and proliferation and controls gene expression through stabilization of b-catenin and subsequent transcription (Fig. 8.5A). In the absence of Wnt ligand, b-catenin is constitutively targeted by the destruction complex, which is composed of axin, casein kinase Ia (CKIa), glycogen synthase kinase 3b (GSK3b), and adenomatous polyposis coli (APC), and mediates proteasomal degradation. The binding of Wnt to its receptor Frizzled (Fz), together with the coreceptor components LRP5 or LRP6, results in the disruption of the destruction complex by the cytoplasmic protein Dishevelled (Dvl). This leads to the suppression of GSK3b activity and results in the stabilization and nuclear accumulation of b-catenin. In the nucleus, b-catenin associates with the transcription factors TCF (T cell factor) and LEF (lymphoid enhancerebinding factor) to regulate the transcription of Wnt target genes. There are also noncanonical b-catenin-independent Wnt signaling pathways, and they include the Wnt/Ca2þ and Wnt/ PCP (planar cell polarity) pathways (Niehrs, 2012; Nusse, 2012). The Wnt/Ca2þ pathway is involved in various aspects of development, cancer, inflammation, and neurodegeneration (De, 2011). In the Wnt/Ca2þ pathway, Wnts trigger Fz-mediated activation of heterotrimeric G proteins, which activates phospholipase C (PLC). This leads to the generation of diacylglycerol and IP3, the release of Ca2þ from intracellular stores and activation of effectors such as Ca2þ/calmodulin-dependent protein kinase II (CaMKII), calcineurin, and PKC (Gao and Chen, 2010; Niehrs, 2012) (Fig. 8.5B).

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FIGURE 8.5 Canonical and noncanonical Wnt signaling pathways. (A) Canonical Wnt/b-catenin signaling. In the absence of Wnt, b-catenin is phosphorylated by GSK3b and CKIa, which targets it for polyubiquitination and proteosomal degradation. Wnt binding to its receptor Fz and coreceptors LRP5/6 activates Dvl and suppresses GSK3b activity. This leads to b-catenin stabilization and its accumulation in the nucleus where it associates with LEF/TCF to regulate gene transcription. (B) Activation of the Wnt/Ca2þ pathway induces a signaling cascade involving G-proteins, PLC, the mobilization of intracellular Ca2þ, calcineurin, CaMKII, and NFAT. In some contexts, PKC is implicated in this signaling cascade. (C) In the Wnt/PCP pathway, WnteFz interactions leads to Dvl-dependent activation of the small GTPases Rho and Rac, which activate ROCK and JNK to regulate cell polarity. Other core components of PCP signaling include Vangl2 and Celsr. (D) In Drosophila, Wnt5a promotes the repulsion of axons from the posterior commissure to the anterior commissure through interaction with the Ryk-like receptor Drl and the activation of the SFK Src64B. (E) Wnt5a repels callosal axons by binding to its receptor Ryk, and possibly Fz. This leads to extracellular influx of Ca2þ from TRP channels and CaMKII activation. CaMKII can phosphorylate the MT-associated protein, tau, which regulates MT dynamics. (F) Wnt4/5a binding to Fz3 attracts postcrossing commissural axons through a signaling cascade involving PI3K, aPKC/Par3/Par6, and GSK3b. Postcrossing commissural axon guidance is also regulated by the PCP proteins Fz3, Vangl2, Celsr3, and JNK. Dvl1 inhibits Fz internalization. When the pathway is activated, aPKC and Vangl2 repress the Dvl1-mediated inhibition of Fz internalization. In contrast, Dvl2 promotes aPKC activity and pathway activation. GSK3b, glycogen synthase kinase 3b; CKIa, casein kinase Ia; LEF, lymphoid enhancerebinding factor; TCF, T cell factor; PLC, phospholipase C; CaMKII, calmodulin-dependent protein kinase II, PKC, protein kinase C; PCP, planar cell polarity; ROCK, Rho kinase; JNK, Jun-N-terminal kinase; SFK, Src family kinase; TRP, transient receptor potential; MT, microtubule; PI3K, phosphatidylinositol-3-kinase; aPKC, atypical PKC; GSK3b, glycogen synthase kinase 3b.

186 PART | I Formation of axons and dendrites

PCP signaling regulates cell polarity and tissue morphogenesis such as the patterning and organization of epithelia, including skin hair follicles and the inner ear epithelium. PCP signaling occurs by Wnt signaling through Fz and Dvl independently of b-catenin (Fig. 8.5C). Dvl associates with the small GTPase RhoA through the Formin homology domain protein Daam1. This interaction leads to the activation of RhoA and its downstream effector Rho kinase (ROCK). Dvl can also stimulate Rac activity, which leads to activation of Jun-N-terminal kinase (JNK). Although their mode of action remains poorly understood, other core components of the Wnt/PCP pathway include the atypical cadherin Celsrs, the fourpass transmembrane protein Vangl2 (Van Gogh-like 2), and the LIM domainecontaining protein, Prickle (Pk) (Gao and Chen, 2010). Wnts guide many types of axons, including postcrossing commissural axons (Lyuksyutova et al., 2003), corticospinal tract (CST) axons (Liu et al., 2005), postcrossing callosal axons (Keeble et al., 2006), and axons of serotonergic and dopaminergic neurons of the brain stem (Fenstermaker et al., 2010) in vertebrates, as well as several axon types in Drosophila (Bonkowsky et al., 1999; Reynaud et al., 2015; Yoshikawa et al., 2003) and C. elegans (Hilliard and Bargmann, 2006; Maro et al., 2009; Pan et al., 2006; Prasad and Clark, 2006; Song et al., 2010). Several Wnt pathways and receptors appear to be involved in axon guidance, and there is emerging evidence that PCP proteins may be important in many Wnt-mediated axon guidance pathways.

8.4.2 Wnt5 repels commissural axons from the Drosophila posterior commissure via derailed, a Ryk tyrosine kinase family member The first direct demonstration that Wnt proteins function as axon guidance molecules was obtained from studies of commissural neurons in the Drosophila CNS. The embryonic Drosophila CNS is composed of an array of axons displayed as a ladder-like structure. Each body segment comprises an anterior and a posterior commissural tract that crosses the midline and joins one of the two lateral longitudinal tracts extending along the length of the embryo. Axons need to correctly project to either the anterior or posterior commissure (Fig. 8.6A). How axons choose to project to either the anterior or posterior commissure is determined by the receptor Derailed (Drl) and its ligand Wnt5 (Bonkowsky et al., 1999; Yoshikawa et al., 2003). Drl, a member of the Ryk tyrosine kinase family, is expressed in the axons and growth cones of neurons that project in the anterior commissure (Bonkowsky et al., 1999; Callahan et al., 1995). In Drl mutant embryos, many neurons that should normally project into the anterior commissure are instead rerouted into the posterior commissure. Moreover, ectopic expression of Drl in neurons that normally projected into the posterior commissure forced their projection across the anterior commissure (Fig. 8.6B).

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FIGURE 8.6 Wnt5a repels commissural axons from the Drosophila posterior commissure. In the developing Drosophila embryo, the axons composing the CNS form a ladder-like structure, with each segment of the embryo comprising a posterior (PC) and an anterior (AC) commissure which cross the midline and join the longitudinal tracts. (A) In wild-type animals, Wnt5a is expressed in the PC and repels Drlþ (green) axons from the PC toward the AC. (B) In mutants where Drl is ectopically expressed in PC neurons, Drlþ PC axons (indigo) are repelled from the PC and cross at the AC. (C) In Wnt5a mutants, Wnt5a expression is lost and Drlþ neurons are no longer repelled from the PC; thus, some Drlþ neurons cross at the PC instead of the AC. (D) In mutants ectopically expressing Wnt5a at the AC, Drlþ neurons that usually cross at the AC stall and fail to migrate across. This leads to a thinning of the AC where Wnt5a is overexpressed.

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To elucidate the mechanism underlying Drl function, a soluble version of the extracellular domain of Drl was used to identify potential Drl ligands expressed at the surface of the Drosophila ventral nerve cord (Bonkowsky et al., 1999; Yoshikawa et al., 2003). Drl binding was detected in the posterior commissure, implying that Drl guides axons into the anterior commissure by repelling them from the site of ligand expression in the posterior commissure. Immunohistochemistry and RNA in situ hybridization studies demonstrated that Wnt5 is expressed in the posterior commissure (Fradkin et al., 2004; Yoshikawa et al., 2003). Furthermore, endogenous Wnt5 from fly extracts binds to the extracellular domain of Drl. In agreement with this, specific binding of the Drl extracellular domain to the posterior commissure of fly ventral cord disappeared in wnt5 mutants (Yoshikawa et al., 2003). Loss of wnt5 function leads to misprojection of anterior commissure axons through the posterior commissure (Fig. 8.6C) and diminished the ability of misexpressed Drl to force axons into the anterior commissure. Moreover, while the ectopic expression of Wnt5 throughout the midline inhibited the formation of the anterior commissure (Fig. 8.6D), overexpression of Wnt5 in drl mutants did not. Thus, biochemical and genetic data indicate that Drl is a receptor for Wnt5 and that this ligandereceptor complex is important for the repulsion of axons away from the posterior commissure. Members of the Ryk family are thought to be catalytically inactive due to amino acid substitutions at highly conserved kinase sites that are normally required for the phosphorylation of substrates. Two members of the SFK family, Src64B and Src42A, were identified as effectors of the Wnt5/Drl signaling pathway in Drosophila (Wouda et al., 2008) (Fig. 8.5D). Expression analysis demonstrated that like Drl, Src64B and Src42A are expressed in anterior commissural axons. Analysis of the ventral nerve cord commissure in embryos mutant for both Src64B and Src42A revealed “fuzzy” commissures, longitudinal breaks, and axon stalling, phenotypes similar to those observed in Wnt5-null embryos (Wouda et al., 2008). Ectopic expression of both Src64B and Src42A forced posterior commissural axons into the anterior commissure in a Drldependent manner. Biochemical studies in Drosophila cells demonstrated that Drl physically interacts with Src64B. This interaction is both required and potentiated the kinase activity of Src64B, and it promoted Drl phosphorylation. Interestingly, the mammalian orthologs of Drl and Src, Ryk and c-Src, are also able to interact in mammalian cells, suggesting that this interaction is evolutionarily conserved (Wouda et al., 2008). However, a role for Src downstream of Ryk in vertebrate axon guidance has yet to be identified.

8.4.3 Wnt5, complexed with derailed, repels Drosophila mushroom body axons The mushroom body a lobe plays roles in long-term aversive memory in the Drosophila adult brain. WNT5, DRL, and DRL-2 are required for mushroom body a axon guidance, with mutants having normal a axon branching but extending inappropriately along the medial trajectory and displaying aberrant midline crossing (Reynaud et al., 2015). WNT5 is localized adjacent to mushroom body axons via binding to the DRL receptors, a member of the Ryk family. DRL captures and presents WNT5 to mushroom body axons. Consistent with this, the cytoplasmic domain of DRL is not required for its function in a axon guidance. DRL-2, another Ryk family receptor, is expressed by mushroom body neurons and is also required for a axon guidance, suggesting that it is a mushroom body axon-intrinsic WNT5 receptor. DRL’s ectodomain must be cleaved and shed to guide axons, and the ectodomain (but not the intracellular domain) is detectable at the tip of the mushroom body a lobe. Biochemical experiments demonstrate that the DRL extracellular domain forms a complex, via WNT5, with transmembrane DRL-2. Thus, WNT5, in complex with the shed DRL ectodomain, guides mushroom body axons via repulsion through their intrinsic DRL-2 receptor (Reynaud et al., 2015).

8.4.4 Wnt binding to Ryk repels axons of the corticospinal tract and corpus callosum through a Ca2D-dependent mechanism Located in the cortex, CST neurons extend axons that project through the mid- and hindbrain, cross the midline, and migrate down the spinal cord in the dorsal funiculus. An AeP decreasing gradient of Wnt1 and Wnt5a is produced in a region neighboring the dorsal midline and dorsal funiculus, and this repels CST axons down the spinal cord. In agreement with a chemorepellent role for Wnts on CST axons, Wnt1 and Wnt5a were found to deflect motor cortical axons away from the source of ligand in explant assays. Ryk is expressed by CST axons, and blocking Ryk function with anti-Ryk blocking antibodies inhibits Wnt1- and Wnt5a-mediated axon repulsion of explants in vitro and the posterior growth of CST axons in mice in vivo. Thus, Ryk mediates the repulsive effect of Wnt1 and Wnt5a in CST axon guidance (Liu et al., 2005). Wnt5a also repels cortical axons at the corpus callosum, the major forebrain commissure. Wnt5a is expressed in a region surrounding the corpus callosum in a high-to-low mediolateral gradient and repels postcrossing callosal axons through its receptor Ryk. Supporting this, axons project aberrantly across the corpus callosum in Ryk/ mice, which have a thicker corpus callosum and a failure of axons to project into the contralateral hemisphere (Keeble et al., 2006).

188 PART | I Formation of axons and dendrites

Electroporation of Ryk siRNA in cortical slices demonstrated that silencing of the Ryk receptor reduced postcrossing callosal axon growth and led to axon pathfinding errors, which included premature dorsal turning toward the cortex or abnormal ventral turning toward the septum (Hutchins et al., 2011). Downstream of receptor activation, pharmacological experiments with cortical slice cultures of the corpus callosum, and also with dissociated cortical neurons isolated from the sensorimotor cortex at a time when cortical axons begin to enter the corpus callosum, suggest that extracellular calcium entry through transient receptor potential (TRP) channels is required for Wnt5a-mediated repulsion of callosal axons (Hutchins et al., 2011; Li et al., 2009) (Fig. 8.5E). A similar mechanism appears to be involved in Wnt5a guidance of CST axons (Li et al., 2009). Furthermore, in callosal axons, Ca2þ may act through CaMKII to regulate Wnt5a-mediated callosal axon guidance (Hutchins et al., 2011). Dynamic microtubules (MTs) are also required for Wnt5a-mediated cortical axon repulsion in vitro (Li et al., 2014). Wnt5a gradients induced asymmetric redistribution of growing MTs toward the far side of the growth cone exposed to lower Wnt5a concentrations. Experiments in vitro suggest that Wnt5a regulates MT dynamics through the MT-associated protein tau, which stabilizes MTs. Wnt5a increases phospho-tau levels at Ser262, an MT-binding site (Li et al., 2014). Phosphorylation of tau by CaMKII at Ser 562 detaches it from MTs and leads to increased MT dynamics. Overexpression of a dominant-negative tau S262A blocks Wnt5a-mediated repulsion in vitro and Wnt5a-mediated MT redistribution (Li et al., 2014). This suggests that tau phosphorylation at Ser262 is necessary for Wnt5a-induced growth cone repulsion (Li et al., 2014) and provides a possible link between Ca2þ signaling downstream of Wnt5 and MT reorganization required for axon guidance (Fig. 8.5E). In addition to repelling growth cones, Wnt5a can also promote cortical axon outgrowth. In contrast to Wnt-mediated axon repulsion, which requires calcium influx through TRP channels, Wnt5a-mediated axon outgrowth involves calcium release from intracellular stores through IP3 receptors in addition to calcium influx through TRP channels (Li et al., 2009). Both axon guidance and axon outgrowth require the Ryk receptor. Addition of secreted Frizzled-related protein 2 (sFRP2), which blocks Wnt/Fz interactions, inhibits Wnt5a-mediated axon repulsion in vitro, but not Wnt5a-mediated axon outgrowth (Li et al., 2009), suggesting that Wnt5a-mediated axon repulsion may also require Fz(s), but this needs to be confirmed with more direct genetic or knockdown approaches. Thus, Wnt5a promotes cortical axon outgrowth and growth cone repulsion through distinct receptors and signaling pathways.

8.4.5 Wnt binding to Fz attracts postcrossing commissural axons via protein kinase C z and planar cell polarity signaling Wnt4, present in an increasing posteroanterior gradient along the neural tube, guides postcrossing commissural axons anteriorly (Lyuksyutova et al., 2003), together with Shh (Bourikas et al., 2005; Yam et al., 2012) (Fig. 8.2). Experiments with spinal cord explant cultures showed that an ectopic posterior source of Wnt4 rerouted postcrossing axons posteriorly, suggesting that Wnt4 is an instructive, rather than a permissive cue, and that it acts as an attractant. Addition of the Wnt inhibitors Sfrp1, Sfrp2, and Sfrp3 caused stalling and random rostrocaudal turning of postcrossing commissural axons in explant cultures. Fz3, a receptor for Wnt4, is expressed by commissural neurons. fz3/ mice have postcrossing commissural axon A-P guidance defects, but no spinal cord patterning defects, suggesting that Wnt4 acts through its receptor Fz3 to guide postcrossing commissural axons (Lyuksyutova et al., 2003). LRP6 is an Fz coreceptor important for canonical Wnt/b-catenin signaling (He et al., 2004). Commissural axon trajectory is normal in LRP6 mutant mouse embryos, suggesting that Wnt-mediated commissural axon guidance does not require canonical Wnt signaling (Lyuksyutova et al., 2003). This concept has been challenged by experiments showing that dsRNA and siRNA knockdown of the canonical Wnt/b-catenin signaling pathway genes Lrp5, Lrp6, and b-catenin in chick generates postcrossing commissural axon phenotypes, without appearing to grossly affect neural tube patterning (Avilés and Stoeckli, 2016). This suggests that the canonical Wnt/b-catenin signaling pathway is involved in Wntmediated postcrossing axon guidance. The difference in results between mouse and chick may be due to LRP5 expression in Lrp6/ mice that compensates for the lack of LRP6 function. Another confounding factor is that the phenotypes of Lrp5, Lrp6, and b-catenin knockdown in chick are different from the Wnt signaling postcrossing commissural axon phenotypes previously described in the fz3/ mouse (Lyuksyutova et al., 2003) and in open-book cultures with perturbations in AeP guidance signaling (Wolf et al., 2008), where the predominant phenotype is randomized AeP turning rather than floor plate stalling or no turns. In contrast to the mouse, in chick, the predominant phenotype observed is floor plate stalling and no turns at the floor plate boundary, which may reflect a floor plate crossing and/or exiting function for the canonical Wnt pathway genes and may point to a difference in function of the genes in chick versus rat/mouse. Many signaling components downstream of Wnt in postcrossing commissural axon guidance have been elucidated, and they include the atypical PKC (aPKC), PKCz, a key component of apicobasal polarity signaling, and the PCP signaling

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molecules Celsr3, Vangl2, and JNK (Fig. 8.5F). Thus, even if it is unclear whether canonical Wnt/b-catenin signaling is involved in Wnt-mediated commissural axon guidance, it is clear that noncanonical transcription-independent pathways are required. Wnt4 attracts postcrossing commissural neurons through PKCz (Wolf et al., 2008). The activated form of PKCz (phosphoPKCz) is expressed in postcrossing commissural axons, and aPKC activity is required for Wnt-mediated attraction of commissural axons and proper AeP pathfinding in neural tube explants. Inositol phospholipid signaling activates PKCz, and consistent with this, PI3Ks were also found to be required for AeP guidance. Pharmacological experiments suggest that heterotrimeric G-proteins may be required to transduce the signal from Fz to PI3K. Further experiments demonstrate that GSK3b, which is downstream of the PKCz/Par6/Par3 complex required for establishing cell polarity, is also expressed in postcrossing commissural neurons. Chemical inhibition of GSK3b also resulted in AeP guidance randomization and ablation of the response to Wnt4 in vitro, implicating GSK3b downstream of aPKC (Wolf et al., 2008) (Fig. 8.5F) in Wnt-mediated axon guidance. The core PCP components Celsr3 and Vangl2 and the PCP signaling mediator JNK are also present in postcrossing commissural axons. In dissociated commissural neurons, JNK is activated by Wnt5a, a Wnt present in the spinal cord. Both the Vangl2 mutant mice (Looptail, Lp/Lp) and Celsr3/ mice show AeP guidance defects of commissural neurons, and pharmacological inhibition of JNK in open-book cultures also generates AeP guidance defects in commissural neurons. Thus, Vangl2, Celsr3, and JNK are required for proper AeP guidance of postcrossing commissural axons (Shafer et al., 2011) (Fig. 8.5F). Classical PCP signaling requires Fz internalization. In commissural neuron growth cones, Wnt5a promotes the colocalization of Fz3 with AP-2 at the tips of filopodia, and this is assumed to reflect an increase in endocytosis (Onishi et al., 2013). Hyperphosphorylation of Fz3 causes it to be targeted to the plasma membrane, and it is postulated that this regulates the balance between plasma membrane and internalized Fz3 (Shafer et al., 2011). Using heterologous cell lines, expression of Dvl1 increased Fz3 hyperphosphorylation, thereby increasing cell surface Fz3 levels and presumably decreasing its internalization. This was associated with a decreased ability of Wnt5a to activate JNK, an intriguing result given that Dvl1 is normally considered an activator of PCP signaling. Vangl2 antagonizes this by reducing Fz3 phosphorylation and promoting its internalization (Shafer et al., 2011) (Fig. 8.5F). In contrast to Dvl1 that induces Fz3 hyperphosphorylation and inhibits signaling (Shafer et al., 2011), Dvl2 does not induce Fz3 hyperphosphorylation and cell surface accumulation. Instead, Dvl2 knockdown suppresses Wnt5a-induced JNK activation and Dvl1-induced Fz3 hyperphosphorylation, suggesting that Dvl2 is required as a positive mediator of WntePCP signaling (Onishi et al., 2013) (Fig. 8.5F). Together, these experiments suggest that PCP signaling is involved in Wnt-mediated guidance of postcrossing commissural axons in rodents (Shafer et al., 2011), an observation that has also been seen in chick (Avilés and Stoeckli, 2016). aPKC, a component of apicobasal polarity signaling, and PCP signaling cooperate to guide axons in response to Wnt. Wnt5a activates aPKC and JNK in commissural neurons. Constitutively active aPKC inhibits Dvl1-induced Fz3 hyperphosphorylation. Furthermore, PAR6, a protein that forms a complex with aPKC in apicobasal polarity signaling, cooperates with aPKC to inhibit Dvl1-induced Fz3 hyperphosphorylation (Onishi et al., 2013) (Fig. 8.5F). PKCz can be activated by phosphorylation at T410. Overexpression of Dvl2, but not Dvl1, increased T410 phosphorylation in dissociated spinal cord neurons. Wnt5a also increased T410 phosphorylation of PKCz (Onishi et al., 2013). These results suggest that Wnt5a-Dvl2 signaling activates aPKC (Fig. 8.5F). Given that Dvl2 is required for Wnt5a-mediated JNK activation, further experiments are required to determine if this is due to Dvl2 activation of aPKC and inhibition of Fz hyperphosphorylation, or if Dvl2 signals to JNK via other PCP components. How PCP signaling, which normally regulates the organization of cells in a sheet, is used to dictate growth cone asymmetry and steer growth cones is a fascinating area of ongoing research (Zou, 2012). Several PCP signaling components are also required for attractive EphrinA reverse signaling in motor neuron guidance (Chai et al., 2014), raising the intriguing possibility that PCP signaling may be a fundamental axon steering mechanism that extends beyond Wnt signaling.

8.4.6 Wnt binding to Fz regulates dopaminergic axon guidance through planar cell polarity signaling Serotonergic (5-HTþ) and dopaminergic (THþ) neurons in the brain stem are organized in discrete nuclei located in the midbrain and hindbrain and project along the rostrocaudal axis to reach their targets in both the brain and spinal cord. Wnt/ PCP signaling is also important for axon guidance of dopaminergic and serotonergic neurons of the brain stem (Fenstermaker et al., 2010). Open-book preparations of hindbrain explants demonstrated that Wnt7b attracts dopaminergic axons, whereas Wnt5a repels dopaminergic axons. In vivo, dopaminergic axons migrate anteriorly due to an attractive Wnt7b gradient and a repulsive Wnt5a gradient. Dopaminergic neurons have AeP guidance defects in fz3, Celsr3, and Vangl2 (Lp) mutant mice, including abnormal lateral trajectory and aberrant posterior projections. This implicates the PCP

190 PART | I Formation of axons and dendrites

signaling components Fz3, Celsr3, and Vangl2 in Wnt-mediated dopaminergic axon guidance. Dopaminergic neurons from fz3 mutant mice are unresponsive to Wnt5a and Wnt7b, implying that Fz3 is the receptor for these Wnts in axon guidance. In addition, Wnt5a mutant mice have transient AeP guidance defects for dopaminergic axons, the transient nature probably reflecting the role of other redundant Wnts such as Wnt7b in their guidance (Fenstermaker et al., 2010).

8.4.7 Wnt3 mediates mediolateral retinotectal topographic mapping After crossing the optic chiasm, RGC axons project toward their targets in the optic tectum in chick or the superior colliculus in mouse. The projections of RGC axons form a topographic map on the tectum such that the topography of the image projected on the retina is recapitulated in the tectum. AeP topographic mapping is specified by gradients of repulsive EphrinAs along the AeP axis of the tectum acting through EphA receptors on RGC axons (Cheng et al., 1995; Drescher et al., 1995). Topographic mapping along the medialelateral axis is determined by gradients of attractive EphrinBs acting through EphB receptors (Hindges et al., 2002; Mann et al., 2002). Experimental evidence and modeling studies suggested that, in addition to EphrinBs, other cues are necessary to account for RGC axon guidance along the medialelateral axis (Hindges et al., 2002). Interestingly, as observed for EphrinB molecules, Wnt3 is expressed in a mediolateral decreasing gradient in the chick tectum (Schmitt et al., 2006). Moreover, RGCs express both Fz5 and Ryk. While Fz5 appears to be expressed uniformly throughout RGCs, Ryk is expressed in a gradient that decreases ventrodorsally. In vitro explant assays demonstrated that ventral axons are repelled by Wnt3, but dorsal axons are attracted by low Wnt3 and repelled by high Wnt3. Wnt3 repulsion is mediated by Ryk, expressed in a ventral-to-dorsal decreasing gradient, whereas attraction of dorsal axons at lower Wnt3 concentrations is mediated by Fz. Providing evidence that Wnt3/Ryk-mediated repulsion plays a role in RGC axon targeting along the medialelateral axis in vivo, overexpression of Wnt3 in the tectum caused RGC axons to avoid the source of ectopic Wnt3. Furthermore, expression of dominant-negative Ryk in dorsal RGC axons resulted in a medial shift of the termination zone, a phenotype that is opposite to RGC axons mutant for EphB. Therefore, Wnt3 repels RGC axons laterally, and the Wnt3 gradient provides a repulsive force that counterbalances the medially directed attractive force of EphrinBs.

8.4.8 Wnts guide axons of Caenorhabditis elegans mechanosensory neurons and D-type motoneurons via Fz-type receptors The C. elegans genome contains five Wnt genes (cwn-1, cwn-2, egl-20, mom-2, and lin-44), four Fz receptors (lin-17, mig-1, cfz-2, and mom-5), one Ryk family member (lin-18), and one Ror-type receptor (cam-1) (Eisenmann, 2005). Expression and genetic analysis revealed that EGL-20, CWN-1, and LIN-44 are expressed in the tail of worm embryos and larvae and function as chemorepellents in the AeP guidance of ALM, PLM, AVM, and PVM mechanosensory neurons, which mediate the response to light touch (Herman et al., 1995; Hilliard and Bargmann, 2006; Pan et al., 2006) (Fig. 8.7A). EGL-20 and CWN-1 act redundantly to guide the anterior migration of PVM, AVM, and ALM axons, whereas LIN-44 regulates the rostral pathfinding of PLM axons (Hilliard and Bargmann, 2006; Pan et al., 2006; Prasad and Clark, 2006; Zheng et al., 2015). Analysis of compound mutants showed that LIN-44 acts through LIN-17 to mediate its guidance function, whereas EGL-20 and CWN-1 function through the MIG-1 and MOM-5 receptors (Hilliard and Bargmann, 2006; Pan et al., 2006; Zheng et al., 2015). Wnt-mediated guidance of PLM neurons may also involve PCP signaling. Genetic experiments suggest that the PCP components Van Gogh (VANG-1), Prickle (PRKL-1), and Flamingo (FMI-1) have a modulatory role in Wnt signaling in PLM neurons, since the phenotype of these mutants is weaker than the Wnt phenotype (Zheng et al., 2015). In addition to regulating the longitudinal axon pathfinding of mechanosensory neurons, LIN-44 and LIN-17 are also involved in the guidance of GABAergic D-type motor neurons, which innervate the body wall muscles and are responsible for reciprocal inhibition during locomotion (Maro et al., 2009) (Fig. 8.7B). D-type motor neurons are located along the ventral midline. They initially project axons anteriorly; however, along their path, these extensions take a sharp turn and migrate dorsally forming a commissure. Once they reach the dorsal nerve cord, they bifurcate and grow in both the rostral and caudal directions. Interestingly, a large number of lin-44 mutants show an overextension of the posterior projections of D-type motor neurons (Fig. 8.7B). This overextension was further enhanced in lin-44;egl-20 double mutants, which suggested that LIN-44 and EGL-20 cooperatively regulate the guidance of these axons (Maro et al., 2009). Ectopic expression of LIN-44 from dorsal body wall muscles resulted in the premature arrest in migration of axons, suggesting that LIN-44 acts as an instructive repellent cue for guidance. Analysis of genetic mutants of Fz receptors demonstrated that lin17 mutants had both over- and underextension defects. Double lin-44;lin-17 mutants did not exhibit enhanced overextension defects compared with lin-17 mutants; thus, LIN-44 appears to function through LIN-17 to mediate the posterior guidance of D-type motor neurons (Maro et al., 2009).

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FIGURE 8.7 Wnt signaling in axon guidance in Caenorhabditis elegans. (A) CWN-1, EGL-20, and LIN-44 are expressed in the tail region in an anterior to posterior gradient. While CWN-1 and EGL-20 regulate the guidance of ALM, AVM, and PVM motor neuron axon guidance, LIN-44 regulates PLM axon pathfinding. (B) LIN-44 regulates axon outgrowth from D-type motor neurons. The cell bodies of D-type motor neurons are located along the ventral midline. They first extend axons anteriorly, and along their path, their projections take a sharp turn and migrate dorsally forming a commissure. Once they reach the dorsal nerve cord end, they bifurcate and grow in the anterior and posterior directions. Lin-44 mutants display an overextension of the posterior axon of the DD6 motor neuron. Arrows indicate the normal length of the posterior DD6 axon, and arrowheads indicate overextensions. (C) CWN-2 promotes the extension of RMED/V motor axons along the longitudinal axis of the worm. CWN-2 is expressed in the posterior region of the developing pharynx and in the intestine of the developing worm.

Genetic analysis points to a b-catenin-dependent and a b-catenin-independent pathway downstream of LIN-17 in Dtype axon guidance. For example, gsk-3 and pry-1/axin mutants showed overextension defects that were reminiscent of lin-44 mutants. Consistent with this, single mutants of all four b-catenin orthologs in C. elegans (BAR-1, WRM-1, HMP-2, and SYS-1) and POP-1/TCF showed underextension defects. Unexpectedly, mutants for mig-5/Dsh, an ortholog of Dvl, also showed strong underextension phenotypes, which is in contrast to a positive role of Dvl in the canonical Wnt pathway (Maro et al., 2009), but nevertheless consistent with the negative role for Dvl1 identified in other Wnt-mediated axon repulsion pathways (Shafer et al., 2011; Zheng et al., 2015). However, given that lin-17 mutants display both over- and underextension defects and mutation of bar-1 in a lin-17 mutant background did not fully rescue the lin-17 phenotype, it was hypothesized that both b-catenin-dependent and b-catenin-independent pathways are involved in the regulation of Dtype motor axon guidance (Maro et al., 2009).

192 PART | I Formation of axons and dendrites

8.4.9 The Wnt ligand CWN2 regulates Caenorhabditis elegans motor neuron axon guidance through a Ror-type receptor CAM-1 CWN-2 attracts axons from RME motor neurons that innervate head muscles and regulate foraging movements in C. elegans. The cell bodies of RME motor neurons are located middorsally (RMED), midventrally (RMEV), left laterally (RMEL), and right laterally (RMER), and they each extend two processes in opposite directions running around the nerve ring (the main circumferential nerve tract in C. elegans) near the anterior surface. In addition to processes that migrate around the circumference of the nerve ring, RMED and RMEV axons send out two processes that migrate along the dorsal and ventral cords, respectively, before arresting at midbody (Fig. 8.7C). A genetic screen identified several components of the Wnt signaling pathway as regulators of RMED/V axon outgrowth (Song et al., 2010). cwn-2 mutants lacked both RMED and RMEV axons (Fig. 8.7C). Given that marker analysis demonstrated that RMED or RMEV cell fate specification was not affected in cwn-2 mutants, it was suggested that CWN-2 is involved in AeP neurite outgrowth. Ectopic expression of cwn-2 in locations other than its normal expression site redirected axon outgrowth toward the site of cwn-2 expression, indicating that CWN-2 functions as a chemoattractive cue on RMEV/D axons (Song et al., 2010). Further analysis of mutants suggested that CAM-1, MIG-1, and CFZ-2 function as receptors of CWN-2 to mediate RMEV/D axon guidance. Given that cam-1 null mutants exhibit a stronger phenotype than single mig-1 or cfz-2 mutants, CAM-1 is likely to function as the main CWN-2 receptor, whereas MIG-1 and CFZ-2 may function as coreceptors. Genetic analysis also supports a role for DSH-1 in RMEV/D axon outgrowth downstream of CWN-2 signaling. However, none of the other components of the canonical Wnt/b-catenin pathway, including BAR-1, PRY-1, WRM-1, and SYS-1, appear to have a role in RMEV/D neurite outgrowth downstream of CWN-2 (Song et al., 2010). cam-1 mutations that disrupt the intracellular domain of CAM-1 have a weaker RMED/V phenotype than the cam-1 null, suggesting that other receptor components may also facilitate transmission of the CWN-2 signal (Wang and Ding, 2018). Yeast two-hybrid screen and coimmunoprecipitation experiments showed that SAX-3, the sole Robo receptor in C. elegans, associates with the CAM-1 intracellular domain. SAX-3 binds directly to CWN-2, as is demonstrated by GST pull-down assays. SAX-3 also forms a complex with CAM-1 and the downstream effector DSH-1. Genetic analysis showed that SAX-3 functions with CAM-1 in RMED/V neurite outgrowth, but this function is independent of Slit, the classic Robo ligand (Wang and Ding, 2018). This interaction between Robo and Wnt signaling reveals new ways in which Wnt signaling can be modulated. CWN-2 also functions through CAM-1 and MIG-1 in motoneurons to regulate nerve ring placement (Kennerdell et al., 2009). This has been hypothesized to occur through regulation of axon guidance of SIA and SIB neurons, which in turn instruct the positioning of the nerve ring (Kennerdell et al., 2009), although this has not been directly demonstrated. Genetic experiments show that SAX-3 might also act with CAM-1 in CWN-2-mediated nerve ring placement (Kennerdell et al., 2009), suggesting that the molecular function of SAX-3 in Wnt-mediated REMD/V axon guidance may also be conserved in nerve ring formation. To date, there is no evidence that the Robo receptors or Ror receptors are involved in Wnt-mediated axon guidance in vertebrates. It will be interesting to determine whether the functions of these molecules are evolutionary conserved.

8.5 Cross-talk between axon guidance cues 8.5.1 Shh induces the response of commissural axons to semaphorin repulsion during midline crossing Shh not only directly guides axons but can also modulate the response of axons to other guidance cues. During midline crossing, spinal cord commissural axons acquire responsiveness to secreted members of the semaphorin family of guidance cues, which regulate their floor plate exit and restrict their postcrossing trajectory into a longitudinal pathway. Shh at the floor plate, together with NrCAM and gdnf, activates the repulsive response of commissural axons to certain secreted semaphorins, which allows for the correct exit of commissural axons from the floor plate (Charoy et al., 2012; Nawabi et al., 2010; Parra and Zou, 2010). In explant outgrowth assays in vitro, Shh activates the repulsive response of precrossing axons to Sema3B and Sema3F. In open-book spinal cord explants treated with a PKA inhibitor or forskolin, an activator of adenylate cyclase, axons stalled, knotted inside the floor plate, recrossed the floor plate, or misprojected posteriorly. These defects are similar to those observed when Shh function is blocked or in mutants for neuropilin-2, a receptor for the secreted semaphorins Sema3B and Sema3F, implicating PKA activity in the induction of semaphorin responsiveness. Furthermore, treatment of precrossing explants with forskolin blocked the induction of the repulsive response to Sema3F, and Shh treatment decreased the levels of activated PKA in commissural axons. Together, these results suggest that Shh may induce semaphorin responsiveness by reducing the activity of the cAMP/PKA pathway (Parra and Zou, 2010).

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8.5.2 Shh regulates Wnt signaling in postcrossing commissural axons In chick, Wnts are also axon guidance cues for postcrossing commissural axons. However, in contrast to rodents, the expression of Wnt does not vary along the AeP axis (Domanitskaya et al., 2010). Instead, the decreasing posteroanterior Shh gradient along the spinal cord induces a decreasing posteroanterior gradient of the Wnt antagonist Sfrp1 (secreted frizzled-related protein 1). This effectively creates an increasing posteroanterior gradient of Wnt activity that attracts postcrossing commissural axons (Domanitskaya et al., 2010). Both gain- and loss-of-function studies demonstrated that disruption of the graded expression of Sfrp1 resulted in aberrant turning or stalling of postcrossing axons at the floor plate. In addition, Sfrp1 blocks the attractive effect of Wnt5a and Wnt7a on postcrossing commissural axons. Thus, in chick, Shh also establishes an attractive functional gradient of Wnt, which guides axons, (Domanitskaya et al., 2010) in addition to directly repelling postcrossing commissural axons anteriorly (Bourikas et al., 2005). Whether this indirect guidance of postcrossing axons by Shh through shaping a Wnt activity gradient is conserved in rodents is not yet known. Wnt signaling in postcrossing axons in rodents is also regulated by Shisa2, a transmembrane protein known to inhibit Wnt signaling. Shisa2 inhibits Wnt signaling by blocking Fz3 glycosylation and cell surface presentation. Constitutive Shisa2 expression causes defects in AeP turning of postcrossing commissural axons. Loss of Shisa2 leads to precocious anterior turning of commissural axons before or during midline crossing, due to premature activation of Wnt signaling. It is postulated that the downregulation of Shisa2 switches on responsiveness to Wnts in postcrossing commissural axons via regulation of cell surface Fz. Shisa2 expression depends on Smo (Onishi and Zou, 2017), although a direct effect of Shh on Shisa2 expression has not yet been demonstrated.

8.5.3 The TGF-b family member unc-129 regulates Unc6/Netrin signaling in Caenorhabditis elegans The C. elegans gene unc-129, a divergent TGF-b family member, is required for proper guidance of pioneer motor axons along the DeV axis (Colavita et al., 1998; Colavita and Culotti, 1998). Mutants in daf-1 and daf-4, the known TGF-b serine-threonine kinase receptors in C. elegans, and genes encoding the Smads had no axon guidance defects, suggesting that UNC-129 may function independently of the canonical TGF-b signaling (Colavita et al., 1998; MacNeil et al., 2009). Instead, Unc-129 mutants have defects in the dorsal trajectories of motor axons resembling those found in unc-5, unc-6/Netrin, and unc-40/Dcc mutants, with no detectable patterning defects (Colavita et al., 1998; Colavita and Culotti, 1998; Hedgecock et al., 1990). Subsequent studies showed that UNC-129 interacts with the UNC-5 receptor (MacNeil et al., 2009). The UNC-5 receptor mediates axon repulsion from UNC-6/netrin through two distinct signaling pathways, one that is dependent on the UNC-40 coreceptor (UNC-5 þ UNC-40) and one that is independent of UNC-40 (UNC-5 alone) (Hedgecock et al., 1990). UNC-6 is expressed by cells in the ventral nerve cord to generate a ventral-high dorsal-low gradient, whereas UNC-129 is expressed by dorsal body wall muscle cells to generate an opposing ventral-low dorsalhigh gradient (Colavita et al., 1998). Thus, motor axons migrate down an UNC-6 gradient and up an UNC-129 gradient. Genetic studies showed that UNC-129 modulates both of these pathways (MacNeil et al., 2009). UNC-129 enhances UNC-5þUNC-40 signaling to promote the long-range repulsive activity of UNC-6 on dorsal motor axons guidance. At the same time, it inhibits the UNC-5 alone pathway. Thus, UNC-129 appears to increase the sensitivity of the growth cone to continuously lower concentrations of UNC-6/netrin encountered as the growth cone moves down the UNC6 gradient by interacting with UNC-5 to enhance UNC-5 þ UNC-40 signaling, allowing for long-range guidance by UNC6/netrin (MacNeil et al., 2009).

8.6 Conclusions and perspectives The initial discoveries that morphogens function as axon guidance cues generated considerable excitement. Subsequent research has highlighted the many instances where Shh, BMPs, and Wnts guide axons, underscoring their diverse role in axon guidance and nervous system development. Morphogens regulate cell fate specification by controlling gene expression through canonical signaling pathways. While the signaling pathways involved in axon guidance by Shh, BMPs, and Wnts are not fully understood, axon guidance by these morphogens occurs predominantly, if not entirely, through transcription-independent noncanonical signaling pathways. It has been demonstrated that at least some of these pathways result in polarization of signaling intermediates in the growth cone, e.g., pSFKs or b-actin (Lepelletier et al., 2017; Yam et al., 2009), or implicate known polarity pathways, e.g., PCP and apicobasal polarity proteins (Fenstermaker et al., 2010; Shafer et al., 2011; Wolf et al., 2008). Thus, sensing of the extracellular gradient and establishing growth cone polarity are important features of axon guidance, for both morphogens and classical axon guidance cues.

194 PART | I Formation of axons and dendrites

Shh signaling in commissural neurons has been linked to the regulation of actin dynamics, whereas Wnt signaling in cortical callosal axons has been linked to the regulation of MT dynamics. We anticipate that further links between morphogens and cytoskeletal regulators will be discovered. Modulation of either actin dynamics (Makihara et al., 2018; Zhou et al., 2002) or MT dynamics (Buck and Zheng, 2002) is sufficient to induce growth cone turning. Given that modulation of either actin or MTs may be instructive for growth cone turning and the interplay between actin and MTs, it is possible that signaling to induce growth cone turning needs only target actin or MTs, but not both. Some of the downstream signaling molecules implicated in axon guidance by morphogens, such as SFKs and Ca2þ/ CaMKII, have also been implicated in the signaling pathway of classical axon guidance cues (Li et al., 2004; Liu et al., 2004; Meriane et al., 2004; Wen et al., 2004). This is not surprising given that, regardless of the axon guidance molecule, they ultimately modulate cytoskeletal dynamics at the growth cone to enable growth cone turning. The use of common signaling mediators allows for information from multiple guidance cues to be integrated in one growth cone. For example, both Shh and netrin1 signal though SFKs (Liu et al., 2004; Meriane et al., 2004; Yam et al., 2009). Under conditions where individual Shh and netrin1 gradients are too shallow to induce a growth cone turning response or SFK polarization, combined gradients of Shh and netrin1 synergize to enable growth cones to sense these shallow gradients of Shh and netrin1 and turn toward the gradient. Notably, SFK activity is one point of integration of signaling between the two guidance cues, and SFK activity in the growth cone is polarized only in the presence of combined shallow gradients of Shh and netrin1, and not the individual gradients (Sloan et al., 2015). How information from multiple guidance cues is integrated by the growth cone is an exciting area of research. Studies on noncanonical Shh signaling in axon guidance were the first to define a transcription-independent noncanonical Hh pathway. Shh also directs cortical microcircuit formation (Harwell et al., 2012), and this may be mediated by a transcription-independent noncanonical Hh pathway. Noncanonical BMP signaling pathways have been implicated in dendritogenesis and synaptic stability (Eaton and Davis, 2005; Lee-Hoeflich et al., 2004), and noncanonical Wnt signaling is involved in dendritogenesis (Rosso et al., 2005). In many cases, these noncanonical effects of Shh, BMPs, and Wnts lead to activation of molecules involved in cytoskeletal rearrangement. Further research on how Shh, BMPs, and Wnts guide axons will not only add to our understanding of axon guidance and neural circuit formation but also add to our knowledge of noncanonical signaling by morphogens.

List of Acronyms and Abbreviations AeP anteroposterior AC anterior commissure APC adenomatous polyposis coli aPKC atypical PKC BMP bone morphogenetic protein CaMKII Ca2þ/calmodulin-dependent protein kinase II CKIa casein kinase Ia CNS central nervous system CST corticospinal tract DeV dorsoventral Dhh Desert hedgehog Drl Derailed Dvl Dishevelled FNIII fibronectin type III FP-netrin1 floor plate netrin1 Fz Frizzled GEF guanine nucleotide exchange factor GPC1 Glypican1 GSK3b glycogen synthase kinase 3b Hh Hedgehog Ig Immunoglobulin Ihh Indian hedgehog ILK integrin-linked kinase JNK Jun-N-terminal kinase LEF lymphoid enhancerebinding factor

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mDN midbrain dopaminergic neurons MT microtubule PC posterior commissure PCP planar cell polarity PI3K phosphatidylinositol-3-kinase Pk Prickle PKA protein kinase A PKC protein kinase C PLC phospholipase C R-Smad receptor-regulated Smad RGC retinal ganglion cell ROCK Rho kinase SFK Src family kinase Sfrp1 secreted frizzled-related protein 1 Shh Sonic hedgehog Smo Smoothened TCF T cell factor TGF-b transforming growth factor b TRP transient receptor potential Vangl2 Van Gogh-like 2 VEGF vascular endothelial growth factor ZBP1 zipcode-binding protein 1

Glossary Chemoattractant Inorganic or organic substance that induces a cell or organism to move toward it. Chemorepellent Inorganic or organic substance that induces a cell or organism to move away from it. Commissural neuron Neuron that passes between the two hemispheres of the brain or between the two sides of the brain stem or spinal cord. Commissure A band of nerve fibers crossing from one side to another of the brain, spinal cord, or body. Distal tip cell Large somatic cell located at the tip of each gonad arms in Caenorhabditis elegans. Dunn chamber assay An in vitro assay for axon guidance where dissociated neurons are exposed to a gradient of a chemoattractant in the Dunn chamber. The turning of axons in response to the gradient is imaged and directly measured over 1e2 h. Floor plate Region of the ventral midline of the developing spinal cord important for guidance and crossing of commissural axons. Growth cone Specialized actin-based structure at the end of a growing axon or dendrite that drives its elongation and can respond to directional cues. Morphogen A diffusible substance that forms a concentration gradient and influences the specification and differentiation of a cell or tissue during embryonic development. Nerve ring Major nerve tract found in Caenorhabditis elegans that encircles the isthmus of the pharynx to form a tightly packed ring-like structure on the outside of the pharynx. Notochord A transient, cylindrical structure of mesodermal cells underlying the neural plate (and later the neural tube) in developing vertebrate embryos. Open-book preparations Spinal cord tissues cut along the longitudinal axis at either the dorsal or ventral midline. Optic chiasm Region where the axons of contralateral retinal ganglion cells from each eye cross the midline. Optic nerve Nerve containing the axons of retinal ganglion cells. It extends from the eye to the optic chiasm. Optic tectum Roof of the midbrain constituting a major visual center in vertebrates. In mammals, it is known as the superior colliculus. Retinal ganglion cells Neurons located in the retina that transmit visual information to the optic tectum. Roof plate Region of the dorsal midline of the developing spinal cord, which secretes factors important for the guidance commissural axons. Spinal cord explant turning assay An in vitro turning assay where spinal cord explants are cultured adjacent to cells secreting a chemical cue. The reorientation of axon paths within the explant in response to the chemical cue is measured. Transient receptor potential channels Group of ion channels located mostly on the plasma membrane of animal cell types that mediate a variety of sensations.

Acknowledgments Work performed in the Charron lab is supported by grants from the Canadian Institutes of Health Research (CIHR), the Fonds de Recherche du Québec-Santé (FRQS), and the Canada Foundation for Innovation (CFI). FC holds the Canada Research Chair in Developmental Neurobiology. We thank L. Izzi for assistance with the figures and manuscript draft.

196 PART | I Formation of axons and dendrites

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Development 135, 2277e2287. https://doi.org/10.1242/dev.017319. Wu, Z., Makihara, S., Yam, P.T., Teo, S., Renier, N., Balekoglu, N., Moreno-Bravo, J.A., Olsen, O., Chédotal, A., Charron, F., Tessier-Lavigne, M., 2019. Long-range guidance of spinal commissural axons by Netrin1 and Sonic hedgehog from midline floor plate cells. Neuron 101, 635e647. e4. https://doi.org/10.1016/j.neuron.2018.12.025. Yam, P.T., Kent, C.B., Morin, S., Farmer, W.T., Alchini, R., Lepelletier, L., Colman, D.R., Tessier-Lavigne, M., Fournier, A.E., Charron, F., 2012. 14-3-3 proteins regulate a cell-intrinsic switch from Sonic hedgehog-mediated commissural axon attraction to repulsion after midline crossing. Neuron 76, 735e749. https://doi.org/10.1016/j.neuron.2012.09.017. Yam, P.T., Langlois, S.D., Morin, S., Charron, F., 2009. Sonic hedgehog guides axons through a noncanonical, Src-family-kinase-dependent signaling pathway. Neuron 62, 349e362. https://doi.org/10.1016/j.neuron.2009.03.022. Yamauchi, K., Phan, K.D., Butler, S.J., 2008. BMP type I receptor complexes have distinct activities mediating cell fate and axon guidance decisions. Development 135, 1119e1128. https://doi.org/10.1242/dev.012989. Yamauchi, K., Varadarajan, S.G., Li, J.E., Butler, S.J., 2013. Type Ib BMP receptors mediate the rate of commissural axon extension through inhibition of cofilin activity. Development 140, 333e342. https://doi.org/10.1242/dev.089524. Yoshikawa, S., McKinnon, R.D., Kokel, M., Thomas, J.B., 2003. Wnt-mediated axon guidance via the Drosophila derailed receptor. Nature 422, 583e588. https://doi.org/10.1038/nature01522. Zheng, C., Diaz-Cuadros, M., Chalfie, M., 2015. Dishevelled attenuates the repelling activity of Wnt signaling during neurite outgrowth in Caenorhabditis elegans. Proc. Natl. Acad. Sci. USA 112, 13243e13248. https://doi.org/10.1073/pnas.1518686112. Zhou, F.-Q., Waterman-Storer, C.M., Cohan, C.S., 2002. Focal loss of actin bundles causes microtubule redistribution and growth cone turning. J. Cell Biol. 157, 839e849. https://doi.org/10.1083/jcb.200112014. Zou, Y., 2012. Chapter six e does planar cell polarity signaling steer growth cones? In: Yang, Y. (Ed.), Current Topics in Developmental Biology. Academic Press, pp. 141e160.

Chapter 9

Axon regeneration R.J. Giger The University of Michigan, Medical School, Ann Arbor, MI, United States

Chapter outline 9.1. 9.2. 9.3. 9.4.

Introduction 201 Anatomy of the spinal cord 202 Spinal cord injury repair: a complex problem 202 Axon regeneration in the injured central nervous system versus peripheral nervous system 203 9.4.1. Intrinsic mechanisms of dorsal root ganglion neuron axon regeneration 203 9.5. Extrinsic mechanisms: inhibitors of central nervous system axon regeneration 204

9.6. Extrinsic mechanisms: growth factors 9.6.1. The anatomical substrate of neurorepair 9.7. Axon regeneration in the retinofugal system 9.8. Lessons learned from an evolutionary perspective 9.8.1. Immune-mediated neurorepair mechanisms 9.9. Conclusions Acknowledgments References

205 206 207 208 208 209 210 210

9.1 Introduction In vertebrates, including humans, the brain and spinal cord are well protected by hard bones of the scull and segmentally aligned vertebrae bodies. While this provides substantial protection to delicate central nervous system (CNS) tissue, the skull or vertebrae column can be fractured or dislocated during a car accident, sports injury, or in combat. In severe cases, this may result in brain or spinal cord tissue damage and lead to permanent neurological deficits. In a similar vein, damage to the retinofugal system, an elaborate fiber system that originates from the neural retina and conveys visual input to the brain, can lead to loss of vision. Finding treatments that improve or reverse the neurological deficits caused by CNS injury is a long-standing and ambitious goal of medicine and biomedical research. Accounts of spinal cord injury (SCI) and attempts to treat it date back to ancient times. The Greek physician Hippocrates (460e377 BCE) once wrote: “there are no treatment options for SCI that resulted in paralysis and unfortunately, those patients suffering from such injuries are destined to die.” While recent advances in emergency medicine and acute care greatly improved the survival rates following SCI, treatment options remain scarce, and patients are faced with a poor prognosis for complete recovery. To improve the quality of life of the estimated 3 million SCI patients worldwide, new treatment strategies are urgently needed. Despite the very limited treatment options, recent advances in our understanding of axon regeneration (Bradke et al., 2012), epidural spinal cord stimulation (Capogrosso et al., 2016; Bloch et al., 2017), biomedical engineering, and stem cell technology (DeBrot and Yao, 2018; Rosenzweig et al., 2018) are fueling new optimism that effective treatment strategies can be developed. Studies in animal models across different phyla have identified key mechanisms that positively or negatively influence regenerative growth of severed axons. This includes pioneering studies in invertebrates and lower vertebrates where a CNS injury typically leads to robust axonal regeneration, offering opportunities to explore the underlying mechanisms and compare them to the mammals where the regenerative capacity of CNS neurons is very limited (Turner et al., 1982; Becker and Becker, 2014). Important advances in our mechanistic understanding of axon regeneration in invertebrates (Chisholm et al., 2016; Hao and Collins, 2017), amphibians (Diaz Quiroz and Echeverri, 2013; Borodinsky, 2017), and reptiles (Jacyniak et al., 2017) have recently been reviewed. Here, we discuss key concepts and mechanisms of axon regeneration, focusing on the injured adult mammalian spinal cord and retinofugal system, neural tissues in which long-distance axon regeneration does not occur spontaneously. Findings in mammals, primarily in mouse

Cellular Migration and Formation of Axons and Dendrites. https://doi.org/10.1016/B978-0-12-814407-7.00009-2 Copyright © 2020 Elsevier Inc. All rights reserved.

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202 PART | I Formation of axons and dendrites

and rat models, will be compared with cellular strategies and molecular mechanisms that enable long-distance axon regeneration in the injured teleost CNS (zebrafish and goldfish), where spontaneous anatomical and functional regeneration typically does occur. Key principles and molecular strategies of axon regeneration and failure will be highlighted. Recent success with combinatorial treatments in mammals subjected to experimental SCI or axonal transection in the retinofugal system will also be discussed.

9.2 Anatomy of the spinal cord A typical human spinal cord is a w50-cm-long cylindrical structure, 1e1.5 cm in diameter, composed of white matter and gray matter. The spinal white matter contains large numbers of ascending and descending groups of fibers, called tracts, which carry electrical impulses from the brain down the spinal cord and sensory information from the periphery up the spinal cord toward the brain. Myelin sheaths, lipid-rich membrane structures tightly wrapped around most long axons, expedite propagation of electrical impulses and allow effective communication among remote groups of neurons. The spinal gray matter, a substance rich in synapses and neural cell bodies, harbors different types of interneurons, glia, and spinal motoneurons (Fig. 9.1). An injury to the spinal cord can damage a few, many, or all spinal fibers. In rare cases, complete recovery from a mild SCI can occur, whereas in most cases, SCI will result in permanent functional impairment, including sensory deficits and loss of voluntary movement. In addition, autonomic dysreflexia is a major complication of SCI. This occurs because autoregulatory organs, such as the heart, gastrointestinal tract, and many glands, are controlled by autonomic nerves. Depending on severity, and level at which the spinal injury occurs, individuals will suffer from partial or full para- or tetraplegia. Because neural elements located above and below the injury site remain largely intact, functional deficits in SCI patients are primarily the result of lost neuronal connectivity and silencing of vital communication lines.

9.3 Spinal cord injury repair: a complex problem Initial damage to the spinal cord, caused by the impact (primary damage), not only destroys tissue and axonal tracts near the injury site but also typically causes spinal bleeding and disruption of the bloodebrain barrier (BBB). Neural tissue damage triggers a number of cellular and biochemical cascades that cause cell death and degeneration over the next several days and weeksdprocesses generally referred to as secondary damage. Secondary damage includes glutamate excitotoxicity and hypoxia, complications that happen within minutes following SCI, and for logistical reasons, these are very difficult to block. Subsequent waves of secondary damage involve a proinflammatory immune response, oxidative damage, axonal dieback, tissue scarring, and protracted cell death. Because the molecular and cellular environment of the spinal cord is constantly changing, from the moment of trauma until several weeks to months later, therapies need to be designed and applied to target specific mechanisms at different postinjury time points. Strategies for the repair of acutely injured

DC

DRG WM

GM

PNS FIGURE 9.1 Cross section through the lumbar spinal cord with attached dorsal root ganglion (DRG). The spinal column consists of white matter (WM) rich in fiber tracts that run up and down the spinal cord. The butterfly-shaped spinal gray matter (GM) harbors synapses, different types of spinal interneurons, including propriospinal neurons, as well as spinal motoneurons. Motoneurons extend their axons into the peripheral nervous system (PNS) to innervate muscle fibers. Sensory neurons located within DRGs send out a short axon that splits into two branches. One branch projects peripherally and becomes part of the PNS. The other branch projects centrally into the spinal cord. Some axons of DRG neurons project rostrally toward the brain and form the dorsal columns (DCs). A conditioning injury to the PNS, typically a sciatic nerve crush injury, activates growth programs in DRG neurons that facilitate regeneration of severed central axons in the DC.

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neural tissue may target mechanisms that regulate neuronal survival, inflammation, tissue scarring, or axonal dieback. At later time points, or in subjects with chronic SCI, treatments will need to focus on promoting axonal growth, formation of new synapses, activity-dependent adaptive neuroplasticity, and axon myelination.

9.4 Axon regeneration in the injured central nervous system versus peripheral nervous system It is well established that the regenerative capacity of severed axons in the adult mammalian peripheral nervous system (PNS) is very robust. This stands in stark contrast to the injured CNS, where spontaneous axonal regeneration is very limited following spinal cord or optic nerve injury. In rodents, transection or crush injuries to the sciatic nerve lead to complete hindlimb paralysis and full functional recovery within 2e3 weeks. This dichotomy between injured PNS and CNS tissue was first described by Ramon y Cajal and prompted studies aimed at understanding what is “special” about the PNS. A series of elegant nerve grafting experiments in the rat, carried out by Aguayo and colleagues, established that some populations of adult CNS neurons possess the capacity to extend long axons following injury if provided with a favorable environment, such as a peripheral nerve graft. Conversely, optic nerve segments transplanted into the PNS failed to support growth of denervated sciatic nerve axons; only few sciatic nerve axons entered the optic nerve transplant, whereas the majority bypassed the transplant before reentering the distal stump of the sciatic nerve (Aguayo et al., 1978, 1981). Collectively, nerve transplantation experiments in the rat established two important principles of axonal regeneration: First, some populations of mammalian CNS neurons retain a capability for long-distance axon growth throughout adulthood, and second, the PNS milieu, but not the CNS milieu, supports long-distance axon regeneration in vivo.

9.4.1 Intrinsic mechanisms of dorsal root ganglion neuron axon regeneration Sciatic nerve injury leads to activation of neuron-intrinsic growth programs in lumbar dorsal root ganglion (DRG) sensory neurons and spinal motoneurons that support robust axonal regeneration (Chen et al., 2007). DRG neurons are pseudounipolar cells with a short axon that splits into two prominent branches: one projecting toward peripheral targets and the other centrally to innervate the spinal cord, including ascending axons within the dorsal columns (DCs) (Fig. 9.1). While injury to the peripheral branch results in robust axon regeneration and target innervation, the tip of the injured central branch forms a clublike structure, called a retraction bulb, and fails to undergo axon regeneration. Thus, depending on which of the two axonal branches of an individual DRG neuron is injured, a vastly different regenerative response is observed. Consistent with observations by Aguayo et al. (1981), this suggests that the CNS environment is a major reason for the failed regeneration of severed axons. Quite remarkably, regeneration of injured DC axons can be significantly enhanced by a prior injury to the sciatic nerve, an experimental manipulation called “conditioning injury” (CI) (Richardson and Issa, 1984; Neumann and Woolf, 1999). This seminal observation has been exploited extensively to uncover mechanisms that promote axon regeneration in the injured adult mammalian CNS (Abe and Cavalli, 2008). The CI response can be mimicked by elevated levels of cyclic adenosine monophosphate (cAMP) and activation of protein kinase A (PKA). Conversely, blocking of PKA abolishes CI-induced neurite outgrowth of cultured DRG neurons (Cai et al., 2002). PKA is thought to promote neuronal growth through inhibition of the small GTPase RhoA (Snider et al., 2002) and activation of the cAMP response element-binding protein (CREB), a nuclear transcription factor that influences injuryinduced gene expression (Gao et al., 2004). Recently, work identified dual-leucine zipper kinase as an effector of cAMP/PKA signaling important in neurons to initiate new axonal growth (Hao et al., 2016; Asghari Adib et al., 2018). While CI-induced regeneration of severed DC axons is quite robust, axons fail to regenerate over long distances, growth is typically twisted, and it appears to lack direction. This stands in contrast to the highly organized and directed growth of injured peripheral axons and indicates that regenerating CNS axons show a more complex growth response. Gene expression analyses of lumbar DRGs identified numerous gene products that are upregulated following sciatic nerve injury and led to the discovery of regeneration-associated genes (RAGs) and neuron-intrinsic pathways that underlie CIinduced central axon regeneration (Tanabe et al., 2003; Seijffers et al., 2006; Blesch et al., 2012; Omura et al., 2015). Epigenetic reprogramming of gene expression, through acetylation of histone 3, enhances RAG expression following a peripheral but not central axonal injury. Functional studies showed that epigenetic reprogramming in sensory neurons is required for CI-induced regeneration of the central branch (Puttagunta et al., 2014). In the absence of CI, injury to the central axon fails to induce expression of RAGs, indicating that sensory neurons do not enter a regenerative state following central injury. Differential gene expression analysis in DRGs, following injury to the peripheral or central branch, identified hub genes and coexpression modules associated with axon regeneration (Chandran et al., 2016). In primary DRG neurons,

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coexpression of one or two transcription factors that regulate hub gene expression is sufficient to enhance axon outgrowth, demonstrating that activation of networks identified in silico increases growth. Activation of core elements of the regeneration-associated transcriptional network can be mimicked by treatment of primary neurons with ambroxol, a small molecular compound that promotes axon outgrowth in vitro. To ask whether ambroxal treatment is sufficient to promote axonal regeneration of injured CNS neurons, mice were subjected to optic nerve crush injury and treated with or without amboxol. In ambroxal-treated mice, a modest but significant increase in retinal ganglion cell (RGC) axon regeneration was observed (Chandran et al., 2016). Collectively, these studies show that activation of neuron-intrinsic growth programs is sufficient to partially overcome the growth inhibitory constraints of injured CNS tissue. They also show that the core networks of transcriptional programs that enable axon regeneration may be similar among different types of adult neurons.

9.5 Extrinsic mechanisms: inhibitors of central nervous system axon regeneration The growth inhibitory nature of injured adult mammalian CNS tissue is well established and has been studied extensively, leading to the identification of numerous growth-inhibitory molecules (Schwab et al., 1993; Hu and Strittmatter, 2004). Prominent examples include the reticulon family member 4A (RTN4A/Nogo-A), myelin-associated glycoprotein (MAG/ Siglec-4), oligodendrocyte glycoprotein (Omgp) (Filbin, 2003; Giger et al., 2008; Xie and Zheng, 2008), sulfatide (Winzeler et al., 2011), and several canonical axon guidance molecules (Giger et al., 2010). Attempts to promote neurorepair following functional ablation of one or several growth inhibitory activities have met with variable success, suggesting that lowering the growth inhibitory environmental influences alone may not be sufficient to achieve robust axonal regeneration. For example, in spinal cordeinjured rats (Bregman et al., 1995) and in nonhuman primates subjected to a unilateral cervical injury (Freund et al., 2009), acute blockage of Nogo-A with a neutralizing antibody is associated with modest functional improvement. However, independent studies using a mouse genetic approach, including adult mice deficient for Nogo-a, either alone or in combination with Mag and Omgp, found either little evidence or no evidence for enhanced axon regeneration following SCI and no improvement in functional outcomes (Lee and Zheng, 2012). Proteoglycans are a large and diverse family of cell surface and extracellular matrix (ECM) proteins that consist of one or more glycosaminoglycan (GAG) chains covalently linked to a core protein. Certain types of chondroitin sulfate proteoglycans (CSPGs) strongly inhibit neurite outgrowth, a feature determined by specific modifications of their GAG moieties rather than the protein core (Smith et al., 2015; Pearson et al., 2018). In the CNS, CSPGs are found in association with glial scar tissue and perineural nets (Busch and Silver, 2007). Enzymatic digestion of GAGs with bacterial chondroitinases attenuates their growth inhibitory action, facilitating axon sprouting and neuronal plasticity in the injured (Bradbury et al., 2002) and naïve (uninjured) mammalian CNS (Pizzorusso et al., 2002). Neuronal cell surface receptors that participate in CNS myelin and CSPG-mediated growth inhibition have been identified and include members of the Nogo receptor family (NgRs), leukocyte common antigen-related protein (LAR), and its homolog RPTPs), leukocyte immunoglobulin-like receptor family (LILRB3/PirB), and sphingosine 1 phosphate receptor 2 (S1PR2) (Fournier et al., 2001; Venkatesh et al., 2005; Atwal et al., 2008; Shen et al., 2009; Dickendesher et al., 2012; Kempf et al., 2014). Studies in transgenic mice deficient for one or several receptors for CNS regeneration inhibitors showed either no regeneration or very limited regeneration following CNS injury (Kim et al., 2004; Zheng et al., 2005; Dickendesher et al., 2012). The limited regeneration observed may in part be due to the presence of inhibitors that signal through different receptors, lack of growth-promoting factors in the injured mammalian CNS, or failure of injured CNS neurons to activate and sustain intrinsic growth programs. ECM components are regulated by CNS injury and influence axonal growth and regeneration (Hoffman and O’Shea, 1999; Quraishe et al., 2018; Roumazeilles et al., 2018; Song and Dityatev, 2018). Integrin receptors play a major role in how a neuron interacts with ECM components. Integrin activation can occur through “inside-out” signaling and the binding of the proteins kindlin and talin to the integrin cytoplasmic tail. This enables binding of extracellular ligands to integrins and activation of intracellular cascades. Tenascin-C promotes growth of axons if they express a tenascin-binding integrin, particularly integrin a9b1. Additionally, integrins can be inactivated by CSGPs, and this inhibition can be overcome by the presence of a b1-binding integrin activator, kindlin-1. In the adult rat, viral vector-mediated overexpression of integrin a9 and kindlin in DRG neurons leads to long-distance regeneration of ascending axons and recovery of sensation after a dorsal root crush injury (Cheah et al., 2016). In the adult PNS, integrins are transported along the axon, but this many not be the case for injured CNS neurons (Andrews et al., 2016), indicating that axonal transportation of molecules associated with growth is a limiting factor in injured adult CNS neurons and may be targeted to promote axon regeneration (Koseki et al., 2017; Petrova and Eva, 2018). Scar tissue formed at the CNS injury is composed of different cellular components, including reactive astrocytes, infiltrating meningeal cells, macrophages, and microglia. In response to injury, astrocytes become activated, proliferate,

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and show increased expression of ECM proteins, including fibronectin, laminin, and CSPGs (Busch and Silver, 2007; Orr and Gensel, 2018). Activated as a self-protection mechanism, the formation of the glial scar is meant to facilitate wound healing by sealing off the injury site, helping to rebuild the BBB, and thereby serves to prevent secondary degeneration of surrounding healthy tissue from infection or exposure to a toxic inflammatory milieu triggered by spinal tissue damage (Rolls et al., 2009). Work from the Sofroniew laboratory showed that targeted elimination of reactive astrocytes after SCI causes failure of BBB repair, reduces axonal sprouting, and causes more severe functional deficits (Sofroniew, 2014; Anderson et al., 2016). Astrocyte progenitors exposed to bone morphogenic protein-4, but not ciliary neurotrophic factor (CNTF), prior to transplantation into a rat SCI form glial bridges that guide and promote axon regeneration (Davies et al., 2008). Thus, tissue scarring following CNS injury poses a very difficult problem, since scarring has both beneficial and detrimental effects on neural regeneration (Fawcett et al., 2012; Anderson et al., 2016). A more nuanced understanding of the cellular heterogeneity in the scar and molecular mechanisms associated with neuroprotective and detrimental effects is required to develop treatment strategies that promote neurorepair.

9.6 Extrinsic mechanisms: growth factors On the other end of the spectrum of extrinsic mechanisms are molecules that promote neuronal growth and axonal regeneration. These include growth-promoting ECM molecules, cell adhesion molecules, and growth factors. Growth factors contribute to growth, guidance, and survival of several neuronal populations during development. The functions of growth factors in the injured PNS and CNS have been extensively studied. Numerous reports show that administration of growth factors promotes growth of different populations of severed spinal axons (Hollis and Tuszynski, 2011; Keefe et al., 2017). In the adult mammalian CNS, growth factors are scarce and are not robustly upregulated following injury. However, a strong regenerative response is observed in spinal cordeinjured rodents subjected to a combinatorial treatment that included activation of neuron-intrinsic growth programs with cAMP, implantation of a permissive matrix at the injury site, and expression of growth factors along the path of regenerating axons (Lu et al., 2012a). In a similar vein, administration of the proregenerative factors IGF1 and osteopontin is sufficient to promote regenerative growth and sprouting of injured corticospinal tract axons and improves behavioral outcomes following thoracic SCI (Liu et al., 2017). In a recent study, robust regrowth of descending propriospinal axons, across a complete SCI, was accomplished in adult rodents. This required a combination treatment comprising (1) activation of neuron-intrinsic growth programs, (2) induction of growthsupportive substrates at the lesion site, and (3) chemoattraction of regenerating axons by glia-derived neurotrophic factor. Individually, none of these treatments promote regeneration of propriospinal axons across the lesion, demonstrating the synergistic effects of combinatorial treatments (Anderson et al., 2018). While regeneration of propriospinal axons was robust and associated with propagation of electrical impulses across the lesion, behavioral outcomes, as assessed by hindlimb motor function, were not improved (Anderson et al., 2018). Collectively, these studies show that combination treatments can achieve impressive axon regeneration in adult rodents; however, the morphology of regenerated axons is elaborate and does not resemble the directed axonal growth observed in the injured PNS. Future challenges concern guidance of regenerating axons and formation of new synapses on appropriate targets to build neural circuits that lead to improved behavioral outcomes. A recent and very promising strategy to reestablish lost connectivity following SCI is the transplantation of neural stem cells (NSCs) into the injury site. NSCs embedded into fibrin matrices containing growth factors were grafted into rats with a complete spinal cord transection. Within 2e3 months following transplantation, grafted cells differentiated into neurons and showed exuberant axonal outgrowth from the graft into the host spinal cord. Importantly, graft-derived neurons formed electrophysiological relays across the injury site, associated with recovery of hindlimb function (Lu et al., 2012b). A similar approach was successful in nonhuman primates. Human neural progenitor cells (NPCs) were grafted into a cervical SCI site in rhesus monkeys. Hundreds of thousands of human axons extended out from grafts through monkey white matter and synapsed in distal gray matter and improved function beginning several months after grafting. These preclinical studies demonstrate feasibility of NPCs graft therapy following SCI (Rosenzweig et al., 2018). To achieve rapid propagation of action potentials along regenerated axons, nearby oligodendrocyte progenitors (OPCs) need to differentiate and myelinate new axons. While OPCs are abundant in the adult mammalian CNS, spontaneous myelination of regenerated axons is very inefficient. The underlying molecular mechanisms that block axon myelination are poorly understood and an area of intense scientific inquiry (Franklin and Goldman, 2015; Mayoral and Chan, 2016; Meireles et al., 2018). Even if regenerated axons synapse with target neurons and become myelinated, newly formed circuits are likely not capable of spontaneously restoring lost function but need to be trained and refined in an activity-dependent manner. Extensive training of newly formed circuits may stabilize and strengthen meaningful connections and outcompete others (Jakeman et al., 2011; Houle and Cote, 2013).

206 PART | I Formation of axons and dendrites

9.6.1 The anatomical substrate of neurorepair It is well established that incomplete SCI in mammals is associated with spontaneous, yet limited, improvement of motor function over a prolonged time course following injury (Raineteau and Schwab, 2001). Numerous studies showed that behavioral outcomes in animals with incomplete SCI can be further improved if subjected to experimental treatments that promote neuronal growth. While hindlimb function is improved, morphological studies show limited regeneration of served axons and no evidence for target reinnervation (Thallmair et al., 1998; Lang et al., 2015). The functional recovery in some behavioral tests is quite impressive, but in no sense can these tests be considered a linear measure of axonal regeneration past the injury site. This begs the question of what is the anatomical substrate of the improved behavior. Many experimental treatments either block growth inhibitory cues or introduce growth-promoting factors, manipulations known to increase neuronal sprouting and growth of injured and uninjured CNS neurons. This leaves open the possibility that some, perhaps even most, of the recovery was due to compensatory plastic changes elsewhere rather than to the regeneration of damaged axons past the lesion site (Fig. 9.2A). In support of this idea, CNS regeneration inhibitors, including Nogo-A, CSPGs, and their canonical receptors, have been shown to negatively regulate neuronal sprouting, circuit remodeling, and synaptic strength (Mironova and Giger, 2013; Schwab and Strittmatter, 2014). Because most SCIs in humans are anatomically incomplete (Tuszynski et al., 1999), but lead to complete paralysis distal to the injury site, functional reorganization of spared neuronal elements may be harnessed to improve behavioral outcomes in the absence of long-distance axon regeneration. This was nicely demonstrated in experimental animals subjected to two temporally separated unilateral spinal hemisections, at thoracic level T12 and 10 weeks later at T7, a surgical paradigm that essentially results in a complete loss of all projections from the brain to lumbar locomotor circuits (Fig. 9.2B). Spontaneous recovery of hindlimb function was assessed 4 weeks after the second lesion. Remarkably, recovery of bilateral stepping was observed only when the lesions were placed at different times but not after simultaneous lesion. This strongly argues that intrinsic spinal circuits, presumably propriospinal interneurons that connect multiple spinal cord segments, are able to reorganize and relay information past the lesions (Bareyre et al., 2004; Courtine et al., 2008; Flynn et al., 2011). These studies have important implications and show that plasticity of intrinsic spinal circuits can restore partial hindlimb function in the absence of long-distance axon regeneration (Fig. 9.2B). In a more recent study, pharmacological manipulation of

(A)

(B)

FIGURE 9.2 Longitudinal view of spinal cord with supraspinal axonal projections (green), propriospinal neurons (blue), and spinal motoneurons (red). (A) Following unilateral spinal cord injury on the right side, connections between the brain and distal spinal motoneurons are lost. In the absence of longdistance axon regeneration, new circuits are formed via collateral sprouts (dotted lines) of supraspinal and propriospinal axons, thereby restoring connectivity with spinal motoneurons. (B) Staggered spinal cord injuries at different rostrocaudal levels disrupt all connections between the brain and caudal spinal cord. Axon collaterals from supraspinal and propriospinal neurons form relays to restore connectivity with motoneurons.

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dormant spinal circuits met with success. Mice with staggered bilateral spinal hemisection at thoracic levels T7 and T10, inflicted at the same time, show permanent hindlimb paralysis. Chronic treatment with a small molecule agonist of the neuron-specific potassium/chloride cotransporter KCC2 significantly improved hindlimb stepping 7 weeks post-SCI (Chen et al., 2018). Mechanistic studies showed that KCC2 activity in inhibitory neurons, located between the two injury sites, is required to improve hindlimb function (Chen et al., 2018). These studies are significant as they demonstrate that pharmacological activation of dormant spinal circuits enables persistent hindlimb stepping. Additional studies are needed to investigate whether pharmacological activation of KCC2 combined with rehabilitation training leads to further improvement of hindlimb function.

9.7 Axon regeneration in the retinofugal system The retina is a thin tissue layer at the back of the eye that detects and converts light into electrical impulses. The principal output neurons of the retina are the RGCs, a heterogeneous cell population composed of many different subtypes. The retinofugal pathway is formed by RGC axons and is a heavily myelinated and topographically organized fiber system that connects the retina with the visual brain. Damage to the optic nerve as a result of trauma or disease can disrupt connections between the retina and the brain. Damaged axons in the mammalian optic nerve show a transient sprouting response that does not support long-distance regeneration. Retroorbital optic nerve crush injury in rodents is an established preclinical model to study RGC degenerative mechanisms (Villegas-Perez et al., 1993; Niwa et al., 2016), neuroprotection (Thomas et al., 2017), and axon regeneration (Benowitz et al., 2017). Studies in the injured mammalian visual system have been very productive and identified numerous experimental manipulations that increase the regenerative capacity of adult RGCs. These include manipulations of key signaling pathways that regulate gene expression, protein synthesis, and axonal transport (He and Jin, 2016; Weng et al., 2016; Fawcett and Verhaagen, 2018; Petrova and Eva, 2018). The mechanistic target of rapamycin (mTOR) has emerged as a major player. Many growth factors signal through the PI3K/AKT pathway. This pathway leads to activation of the small GTPase Rheb, a strong activator of mTOR, and subsequent promotion of mechanisms that control protein synthesis, cell growth, and survival. The PI3KeAKTeRhebemTOR pathway is kept in check by the tumor suppressor phosphatase and tensin (PTEN) homolog. In adult mice, RGC-selective deletion of Pten leads to constitutive activation of mTOR and, following optic nerve injury, lengthy axon regeneration (Park et al., 2008). In mice, RGC axon regeneration is further enhanced if Pten deletion is combined with loss of suppressor of cytokine signaling 3 (SOCS3), a negative regulator of the Janus kinase/signal transducers and activators of transcription (JAK/STAT) pathway (Sun et al., 2011). In adult RGCs, activation of the mTOR pathway through a constitutively active mutant of Rheb, combined with visual stimulation or chemogenetic approaches to increase RGC activity, enables many axons to regenerate over long distances. Regenerated axons formed synapses with brain targets and partially restored visual function (Lim et al., 2016). Overexpression of the transcription factor c-myc in RGCs promotes axon regeneration and strongly augments regeneration induced by deletion of Pten and Socs3 when combined with administration of CNTF. The c-myc, Pten, Socs3, and Cntf combination treatment enables large numbers of RGC axons to cross the optic chiasm, and some axons enter the ipsi- and contralateral optic tracts (Belin et al., 2015); however, most regenerated axons wander astray, impeding significant innervation of more remote brain targets (Luo et al., 2013; Pernet and Schwab, 2014; Belin et al., 2015). The mechanism(s) by which mTOR activation promotes axon regeneration are not fully understood but are thought to involve elevated S6 kinasedependent protein synthesis and inhibition of GSK3b (Yang et al., 2014; Zhang et al., 2018). A recent study found that mTOR controls translation of axonal mRNAs in injured DRG neurons (Terenzio et al., 2018) and evidence suggests that mTOR functions in both retrograde injury signaling and axonal synthesis of regeneration-associated proteins. Upon the transition from embryonic stages to adult, the intrinsic neuronal growth activity is repressed (Goldberg et al., 2002; Koseki et al., 2017). This transition is regulated, at least in part, at the transcriptional level by members of the Krüppel-like factor (Klf) family, powerful regulators of neuronal growth. In vitro, overexpression of the Klf-4 and Klf-9 suppresses axonal growth, whereas Klf-6 and Klf-7 function as positive regulators of axon regeneration in vitro (Moore et al., 2009; Blackmore et al., 2012) and of optic nerve axon regeneration in zebrafish in vivo (Veldman et al., 2007). Moreover, loss of Klf-4 or Klf-9 in mouse RGCs promotes axon regeneration following optic nerve injury, at least in part through increased activation of the Jak-STAT pathway (Moore et al., 2009; Qin et al., 2013). Collectively, these studies show that activation of neuron-intrinsic growth programs is sufficient to achieve longdistance axon regeneration in the injured mammalian retinofugal system. These studies also show that proper navigation of regenerating axons does not occur spontaneously, similar to observations in the spinal cord, axons frequently wander astray and fail to robustly innervate preinjury targets.

208 PART | I Formation of axons and dendrites

9.8 Lessons learned from an evolutionary perspective Extrinsic mechanisms: The regenerative failure of injured RGC axon in adult mammals stands in stark contrast to zebrafish and goldfish, teleosts in which severed RGC axons regenerate extensively and fully restore vision (Becker and Becker, 2014; Bollaerts et al., 2018). Differences in the RGC regenerative response in the zebrafish and mouse, two vertebras with a very similar genetic makeup, have been investigated extensively and identified environmental and neuronintrinsic mechanisms that enable or block axon regeneration, pathfinding, and target innervation in the retinofugal system. In adult mammals, damage to the optic nerve leads to protracted RGC death (Watanabe et al., 1997; Fernandes et al., 2012; Maes et al., 2017). Only few RGC subtypes survive an optic nerve injury, including alpha-RGCs and intrinsically photosensitive RGCs, neither of which is capable of spontaneous long-distance axon regeneration (Duan et al., 2015; Li et al., 2016). In the mouse, RGC axon transection leads to activation of the proapoptotic molecule Bax and loss of w80% of RGCs within weeks (Libby et al., 2005; Maes et al., 2017). In the zebrafish, optic nerve injury does not cause RGC death, and axons regenerate to faithfully and topographically innervate their termination fields in the adult brain (Becker et al., 2000). The striking differences in CNS regeneration between mammals and fish promoted studies at the underlying cellular and molecular mechanisms. In mammals, myelin debris from dying oligodendrocytes is laden with growth inhibitory molecules and contributes to a growth hostile environment (Schwab et al., 1993; Hu and Strittmatter, 2004; Giger et al., 2008). The lesioned zebrafish CNS shows little expression of molecules that inhibit axon regeneration. CNS myelin prepared from fish does not inhibit neurite outgrowth in vitro, since fish and mammalian neurons readily extend axons on fish myelin, but neither fish nor mammalian neurons grow neurites in the presence of mammalian CNS myelin (Bastmeyer et al., 1991). This is quite remarkable as it shows that fish neurons already possess the surface receptor(s) to respond to inhibitory activities present in mammalian CNS myelin. This argues that mammalian inhibitors function through “evolutionarily old” surface receptors and signaling pathways already present in teleosts. Unlike mammals, the lesioned zebrafish CNS shows little scarring or upregulation of CSPGs (Becker and Becker, 2008). While myelin inhibitors do exist in the zebrafish CNS, they are less inhibitory. For example, mammalian Nogo-A is a potent inhibitor of neurite outgrowth with multiple inhibitory regions (Schweigreiter, 2008; Abdesselem et al., 2009). The zebrafish genome encodes two Nogo-A homologs, RTN4a and RTN4b. While RNT4b inhibits neurite outgrowth, RTN4a supports neurite outgrowth. Only low levels of RTN4b are present following optic nerve injury in adult zebrafish (Bodrikov et al., 2017). In contrast to mammalian oligodendrocytes, optic nerve injury in the fish leads to oligodendrocyte de-differentiation, and glial cells start to express growth-promoting molecules on their surface (Bodrikov et al., 2017). Astrocytes in the fish optic nerve align with the direction of axonal growth and are permissive for regenerating axons; no obvious alignment is observed in injured mammals, and reactive astrocytes do not provide guidance (Garcia and Koke, 2009; Yang et al., 2015). Collectively, these studies indicate that CNS (extrinsic) influences that impair axon regeneration are abundantly present in mammals but largely absent in fish. When coupled with limited tissue scarring in the fish, and the presence of glial cells that provide guidance, this may account at least in part for the regenerative differences in fish and mammals. Intrinsic mechanisms: There are differences in intrinsic growth capabilities within different populations of CNS neurons. For example, following SCI, mammalian corticospinal axons are more refractory to regeneration than serotonergic axons (Giger et al., 2010). In a similar vein, subsets of mammalian RGCs show different regenerative potentials (Duan et al., 2015; Li et al., 2016). This observation is not unique to mammals but extends to teleost CNS neurons. In zebrafish, a thoracic SCI leads to complete paralysis and injured fish regain normal swimming behavior within 4e6 weeks (Becker and Becker, 2014). While many axons show nearly complete regeneration, others, including axons of Mauthner neurons, exhibit poor axonal regrowth. Thus, the intrinsic regeneration capability in fish CNS neurons is variable; moreover, it seems that not all axons need to regenerate to fully restore function. Similar to mammalian neurons, Mauthner axon regeneration can be enhanced by elevation of intracellular levels of cAMP (Becker and Becker, 2014). Conversely, in zebrafish, where RGC axon regeneration occurs spontaneously, pharmacological inhibition of mTOR with rapamycin leads to significantly reduced axon regeneration (Diekmann et al., 2015). Similar to mammals, RGC axon regeneration in zebrafish is sensitive to manipulation of the Gp130 Jak/STAT pathway (Smith et al., 2009; Elsaeidi et al., 2014). Together, these findings demonstrate that evolutionarily conserved mechanisms regulate intrinsic regenerative programs in adult CNS neurons.

9.8.1 Immune-mediated neurorepair mechanisms CNS injury causes rapid activation of nearby microglia and recruitment of blood-derived macrophages; their combined action is thought to contribute to secondary damage, including axon dieback, neuronal death, and tissue scarring (Fitch and Silver, 2008; Sofroniew, 2009; Gadani et al., 2015; Cekanaviciute and Buckwalter, 2016). Similar to mammals, CNS

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injury in zebrafish triggers a strong immune response (Bollaerts et al., 2017). Strikingly, in zebrafish, inflammation is a key process required for regeneration. Following treatment with dexamethasone to block the innate branch of the immune system, immunosuppressed zebrafish are observed to have impaired neuroregeneration following SCI (Ohnmacht et al., 2016). Because innate immune cells, including monocytes, macrophages, and microglia, can acquire different activation stages associated with specific functions (Xia et al., 2015), it has been proposed that, depending on context, neuroinflammation leads to a micromilieu that is either detrimental or supportive for repair (Kigerl et al., 2009). Recent work established that proinflammatory molecules released by microglia activate astrocytes, and depending on the context of a CNS insult, reactive astrocytes may adopt a cytotoxic phenotype or a proregenerative phenotype (Zamanian et al., 2012; Liddelow et al., 2017; Shinozaki et al., 2017). In the mouse, induction of cytotoxic retinal astrocytes is observed following optic nerve crush injury and can be blocked by intraocular injection of TNFa, IL1a, and C1q neutralizing antibodies. Blockade of cytotoxic astrocytes has a strong protective effect toward injured RGCs (Liddelow et al., 2017). This suggests that, in the injured mammalian CNS, acute inflammation impedes neural tissue regeneration, and the astrocyte phenotype may ultimately determine survival and regeneration of injured CNS neurons (Brambilla et al., 2012; Calkins et al., 2017). Multiple lines of evidence show that, under certain circumstances, neuroinflammation can be beneficial. Intraocular inflammation, either triggered by a cataract-inducing lens injury or by intraocular injection of zymosan, causes an inflammatory milieu with strong RGC protective and proregenerative features (Fischer et al., 2000; Yin et al., 2003). The proregenerative effects of inflammation toward injured neurons are not restricted to the visual system, since injection of zymosan into DRGs or spinal cord parenchyma triggers a local immune response that promotes growth of sensory neurons (Steinmetz et al., 2005; Gensel et al., 2009). Despite these encouraging findings, inflammation elicited by zymosan causes bystander toxicity, undermining the beneficial effects (Gensel et al., 2009, 2015; Baldwin et al., 2015). Zymosan is a crude yeast cell wall preparation; the active ingredient enabling axon regeneration is a particulate form of b-glucan, a ligand for the innate immune receptors dectin-1 and Toll-like receptor 2 (TLR2). Intraocular injection of a purified form of particulate b-glucan (called curdlan), a ligand for dectin-1, is sufficient to promote RGC axon regeneration in wild-type but not in dectin-1-deficient mice (Baldwin et al., 2015). Of clinical interest, administration of b-glucan at the time of nerve injury, or even 2 days later, promotes equally robust RGC axon regeneration, suggesting a large therapeutic window for b-glucan/ dectin-1-elicited neurorepair (Baldwin et al., 2015). There are many instances where neuroinflammation is detrimental and does not lead to enhanced neuronal survival or axon growth (Gensel et al., 2009; Evans et al., 2014; Kroner et al., 2014; Gadani et al., 2015; DiSabato et al., 2016). In the injured visual system, intraocular injection of bacterial lipopolysaccharide, a ligand for Toll-like receptor 4 (TLR4), causes a strong inflammatory response but does not increase RGC survival or axon regeneration (Baldwin et al., 2015). A future challenge is to identify specific immune cell types and biochemical pathways associated with neurotoxic (“bad”) and proregenerative (“good”) inflammation. Many questions remain how activation of subsets of immune cells leads to enhanced RGC protection and axon regeneration or toxicity. While our understanding of immune-mediated neurorepair is still fragmented, there is a keen interest in the identification of the factors associated with inflammation-induced RGC protection and axon regeneration. These may include CNTF, leukemia inhibitory factor, and IL-6, factors that activate the glycoprotein 130/STAT3 pathway (Leibinger et al., 2009), the calcium-binding protein oncomodulin (Yin et al., 2006; Kurimoto et al., 2013), and ApoE (Lorber et al., 2009). Over the past years, the list of factors that promote RGC regeneration has steadily increased and now includes CXCL12, HGF, IGF1, osteopontin, and Wnt (Harvey et al., 2012; Heskamp et al., 2013; Ogai et al., 2014; Duan et al., 2015; Fischer, 2017; Patel et al., 2017). Despite this impressive list, little is known about the specificity of these factors toward RGC subpopulations, potential synergistic actions on RGC survival and protection, or effects on other retinal cell types (Duan et al., 2015). Of interest, immune-mediated repair mechanisms seem to be largely distinct from other growth-promoting manipulations, such as deletion of PTEN or intraocular injection of cAMP/oncomodulin. Combining these manipulations resulted in robust additive growth effects, greatly exceeding RGC axon regeneration achieved by individual treatments (de Lima et al., 2012).

9.9 Conclusions Following nervous system injury, axon regeneration is controlled by neuron-intrinsic and neuron-extrinsic mechanisms (Table 9.1). Neuron-intrinsic signaling pathways that regulate axonal growth and regeneration in the injured vertebrate CNS appear to be evolutionarily conserved. While lower vertebrates spontaneously activate and sustain neuron-intrinsic growth programs, injured mammalian CNS neurons fail to do so. Experimental activation of neuron-intrinsic growth programs in mammalian CNS neurons is sufficient to promote robust axon regeneration and can be further enhanced by simultaneous lowering of extrinsic growth constraints. However, the growth pattern of regenerated axons in the mammalian CNS is complex and appears to lack guidance. As a consequence, axons typically wander astray and fail to

210 PART | I Formation of axons and dendrites

TABLE 9.1 Overview of extrinsic and intrinsic mechanisms that positively and negatively influence axon regeneration in the injured adult mammalian central nervous system. Extrinsic mechanisms

Intrinsic mechanisms

Growth promoting -

Growth factors Morphogens Permissive ECM Cell adhesion molecules Integrin signaling Neuronal activity Alternatively activated immune cells

-

RAGs cAMP/PKA/CREB Rheb/mTOR gp130 Jak/STAT3 Klf6, Klf7 DLK c-myc Epigenetic mechanisms

-

RhoA/ROCK PTEN SOCS3 Klf4, Klf9 Impaired axonal transport Impaired axonal translation non-muscle myosin II

Growth inhibitory -

CNS myelin Certain types of CSPGs Nonpermissive ECM Guidance cues Cytotoxic astrocytes Tissue scarring Classically activated immune cells

ECM, extracellular matrix; CSPGs, chondroitin sulfate proteoglycans; PTEN, phosphatase and tensin; RAGs, regeneration-associated genes; cAMP, cyclic adenosine monophosphate; PKA, protein kinase A; CREB, cAMP response element-binding protein; mTOR, mechanistic target of rapamycin; DLC, dual-leucine zipper kinase; SOCS3, suppressor of cytokine signaling 3.

contact and synapse on preinjury targets. In injured teleosts, the environment lacks many of the growth inhibitory cues present in the mammalian CNS, and unlike mammals, fish glia form conduits that support regenerating axons and provide guidance. In both fish and mammals, CNS injury triggers a strong immune response. Neuroinflammation is important for regeneration in fish but largely detrimental in mammals. Certain experimental manipulations that skew injury-induced neuroinflammation toward a reparative phenotype lead to elevated production of growth factors and elicit robust axon regeneration. The extent of neuronal growth and plasticity required to restore function following CNS injury is an ongoing debate. In spinal cordeinjured mammals, sprouting, rather than long-distance regeneration of supraspinal axons, is sufficient for the formation of relays with intraspinal neurons that bridge the injury site and improve behavioral outcomes. Pharmacological activation of dormant spinal circuits can restore substantial hindlimb function, presumably in the absence of axon regeneration, suggesting that neuroplasticity of spinal circuits is a key mechanism to improve function. Extensive rehabilitation training will be needed for the reorganization and shaping of new spinal circuits to maximize behavioral outcomes. Combination treatments that make use and integrate the most promising strategies will likely have additive effects and lead to improved treatment protocols suitable for translation into human subjects.

Acknowledgments I thank my colleagues Daniel Goldman, Catherine Collins, and members of my laboratory for critical reading of the manuscript.

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Chapter 10

Axon maintenance and degeneration Fan Wang1, Jing Yang2 and Zhigang He3 1

Department of Neurobiology, Duke University, Durham, NC, United States; 2School of Life Sciences, Peking University, Beijing, China; 3Kirby

Center of Neuroscience, Children’s Hospital Boston, Harvard Medical School, Boston, MA, United States

Chapter outline 10.1. Introduction 217 10.2. Essentials of axonal transport in axon maintenance 218 10.2.1. Cellular components that are transported along the axons 218 10.2.2. Regulations of microtubule stability and organization during axon maintenance 220 10.2.3. Defects in motor proteins cause axon degeneration 221 10.2.4. Role of mitochondria transport in axon maintenance 221 10.2.5. Membrane transport and insertion are essential for axon maintenance 222 10.3. Proteasome and autophagy pathways in axonal homeostasis 223

10.3.1. Ubiquitineproteasome system in axon maintenance 10.3.2. Role of autophagy/lysosome pathway in maintaining axonal homeostasis 10.4. Role of glial cells in axon maintenance 10.5. Maintaining axon track positions and other structural features 10.6. Axon pruning and axon degeneration 10.6.1. Developmental axon pruning 10.6.2. Pathological axon degeneration 10.6.3. Molecular mechanisms of pathological axon degeneration References

223 223 224 224 225 225 227 227 229

10.1 Introduction After embryonic development establishes initial wiring of the nervous system, neuronal connections need to be maintained both during postnatal growth and throughout an animal’s entire life span. The drastic increase in body size as animals mature into the adulthood, together with mechanical and physical stresses caused by body movement, environmental insults, and aging, poses constant challenges to the stability of neuronal architecture. In this chapter, we will discuss several key mechanisms that are involved in axon maintenance, and we summarize the current cellular and molecular understanding of the axon degeneration processes. Axon maintenance requires delivery of new materials and removal of unwanted proteins to maintain the structure and function of axons. Consistent with this principle, human diseases with axon degeneration are often caused by mutations in genes involved in axonal transport and protein degradation. In addition to these neuronal-intrinsic mechanisms, interactions with glial cells and the extracellular matrix are also important for maintaining axonal integrity and organization. Axon pruning and axon degeneration occur both during development and in many types of neurodegenerative diseases caused by genetic mutations or injuries resulting from toxic, ischemic, or traumatic insults. In pathological conditions, axon degeneration can be either a part of a neuronal death program or brought about by death-independent pathways. Thus, identifying cellular and molecular mechanisms underlying axon pruning and degeneration is important for understanding neural circuit development, and this will also help in the identification of therapeutic strategies that interfere with the process of axon degeneration in neurodegenerative diseases and neural injuries.

Cellular Migration and Formation of Axons and Dendrites. https://doi.org/10.1016/B978-0-12-814407-7.00010-9 Copyright © 2020 Elsevier Inc. All rights reserved.

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218 PART | I Formation of axons and dendrites

10.2 Essentials of axonal transport in axon maintenance 10.2.1 Cellular components that are transported along the axons The axon is a specialized structure unique to neurons. The main function of the axon is to conduct action potentials from the cell body to synapses. It also delivers biological materials required for synaptic structure and neurotransmission. Since most neuronal proteins and membranes are synthesized in the cell body, while the distal portion of an axon can be as far as 1 m away from the soma (as is the case for some human sensory and motor neurons), efficient and coordinated transport of materials and organelles is especially important for maintaining axon integrity and function. In the mid-19th century, Augustus Waller showed that axons could not self-sustain when separated from their parental cell body (see Section 6). The first direct experimental demonstration of anterograde axonal transport was made by Paul Weiss in the 1940s. He found that if an axon was ligated, materials would accumulate on the side of the axon close to the soma. When the knot was untied, the accumulated materials continued to move down the axon at a rate of 1e10 mm per day (Weiss and Hiscoe, 1948). However, if all materials in the axons were transported at this rate, they would take an unfeasibly long time (at least half a year in the longest axons) to reach the axon terminus. In the late 1960s, experimental methods were developed to trace the movements of proteins labeled with radioactive amino acids. In this case, the radioactive amino acids were injected into the soma of neurons so that newly synthesized proteins would incorporate these radioactive amino acids. The movement of these labeled proteins along the axons was measured, and this allowed for an estimate of the rate of transport. Using this approach, Bernice Grafstein discovered the presence of fast axonal transport (distinct from the slow axonal transport described by Weiss) that could occur at a rate as fast as 1000 mm per day (Grafstein and Forman, 1980). A diverse array of cargoes including mitochondria and other membranous organelles (such as synaptic vesicle precursors and endosomes), membrane proteins such as sodium and potassium channels, lipids, soluble signaling molecules, and cytoskeleton components are actively transported along axons (Fig. 10.1). Generally, membranous organelles (mitochondria and various vesicles), neurotransmitter vesicles, channel proteins, and multivesicular bodies are transported at a rate of 50e400 mm/day by fast axonal transport. On the other hand, soluble proteins, including unassembled actin and tubulin monomers, clathrin, and glycolytic enzymes, are transported at a net rate of 3e8 mm/day, which is considered to be the slow component b (SCb) of axonal transport. Assembled microtubules (MTs) and neurofilaments are transported at a rate of 0.1e1 mm/day, also referred to as slow component a (SCa) axonal transport (Fig. 10.2). Recent studies showed that, in many cases, the “slow” transport is in fact caused by frequent pauses of the cargoes, and the speed during actual movement is comparable with that observed in the “fast” transport process. The mechanism is known as the “Stop and Go” model of the slow axonal transport (Brown, 2003; Roy et al., 2007).

Dendrites Actin filament Cargo Endosome Microtubule Mitochondrion Nucleus Neurofilament Ribosome Soma Smooth ER Synaptic vesicle precursor Synaptic vesicle

Initial segment

Axon hillock Presynaptic terminal

FIGURE 10.1 Organelles and structural components in axons. Despite recent demonstration of translation of mRNA in axons, the vast majority of axonal proteins are synthesized in the neuronal cell body (soma) and transported to the axons. While the protein synthesis machineries such as rough endoplasmic reticulum (ER) (polyribosome tethered ER, also called Nissl bodies) and Golgi apparatus are highly enriched in the neuronal perikarya, they are largely excluded from the axon at the axon hillock/initial segment. The molecular basis for this sorting is not well understood. Cytoplasm in the axon hillock does not appear to act as a physical “sizing” barrier because a variety of organelles such as mitochondria, smooth ER, and lysosome enter the axon readily. The unique shape of an axon is the result of interplays between membrane components (the lipid matrix and associated proteins) and cytoskeletal elements such as actin, microtubule, and neurofilaments. In many cases, the axons could be myelinated starting from the initial segment, which permits the saltatory conduction of the action potential.

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Anterograde axonal transport

Kinesins

Type of transport Fast (50a400 mm day–1): Organelles (mitochondria and vesicles) Neurotransmitters Channel proteins Multivesicular bodies Endosomes

Retrograde axonal transport

Slow component a (0.1a1 mm day–1): Assembled neurofilaments and microtubules

Dynein

Slow component b (3–8 mm day–1): Soluble proteins

FIGURE 10.2 Schematic representation of axonal transport. Axonal transport occurs bidirectionally along microtubules, which run along the length of the axon and provide the main cytoskeletal “rails” for transportation. The motor proteins, kinesins and cytoplasmic dynein, are mechanochemical enzymes that move cargoes in the anterograde (toward the axon tip) or retrograde (toward the cell body) directions, respectively. Motor proteins bind and transport multiple different cargoes. Vesicular cargoes move relatively fast, whereas transport of proteins takes much longer. Recent studies have revealed that the movement of individual “slow” cargoes is actually rapid but unlike fast cargoes, they pause frequently, making the overall transit rate much slower.

The primary mechanism that moves these cellular “cargoes” along axons is long-range MT-based transport. Here, we briefly summarize, using a railroad analogy, three key components, i.e., the “rails,” “engines,” and “fuel suppliers” of the transport machinery (Fig. 10.2). The “rails” are axonal MTs, with their “plus” ends pointing toward the distal termini of axons and their “minus” ends facing the neuronal soma. The “engines” are molecular motors that bind and move cargoes along the MT rails. Kinesin family proteins are the motors carrying cargoes in the anterograde direction (moving toward plus end of MTs). In mammals, the kinesin superfamily consists of about 45 members (KIFs) that are grouped into 14 subfamilies. Kinesins are composed of one to four motor polypeptides called heavy chains that contain a highly conserved motor domain with ATPase and MT-binding regions and also a divergent tail domain. Regulatory and accessory subunits, such as the kinesin light chains, are thought to interact with the tail domain of the kinesin heavy chains to confer cargobinding specificity and regulation of cargo destination (Hirokawa and Takemura, 2005). In addition to anterograde transport, axons often receive trophic signals from their synaptic targets and need to deliver these signals back to their cell bodies. Movement in this direction, from the axonal terminal to the soma (toward the minus end of MTs), is called retrograde transport. All membranous organelles also undergo retrograde transport. The molecular “engine” for this direction is provided by cytoplasmic dynein (Kardon and Vale, 2009). Dynein is a large and complex MT motor, composed of two dynein heavy chains motor subunits and various light chain subunits. Different from kinesins, which many family members that have evolved to carry a variety of different cargoes and execute different functions, a single cytoplasmic dynein carries out similarly diverse transport activities in the retrograde direction. It appears that dynein

220 PART | I Formation of axons and dendrites

employs a “subunit heterogeneity” strategy in which an array of light chains link dynein to different cargoes and mediate dynein’s interaction with several adaptor/regulatory proteins (Kardon and Vale, 2009). Moreover, although dynein advances most frequently in 8 nm steps (same as kinesins) toward the minus end of MTs, it has been observed that dynein can take steps of other sizes (4e32 nm) and can also move backward and sideways. Both kinesin and dynein motors generate force through hydrolyzing ATP, and local glycolysis and mitochondria are the ATP “fuel suppliers.” Emerging evidence indicates that the ATP energy deficit in axons results in halted transport and also the degeneration of axonal structures (see Section 6). The length and narrow caliber of axons, coupled with the high demand for materials that must be transported, make axons highly vulnerable to perturbations involving the transport machinery.

10.2.2 Regulations of microtubule stability and organization during axon maintenance As described earlier, MTs are the essential “rails” of the axonal transport machinery, and not surprisingly, perturbations of MT organization or dynamics, or disruption of interactions between MTs and motor proteins, can interfere with axonal transport of materials and thereby disrupt axon maintenance. MT dynamics and stability are modulated by a variety of proteins, including MT assembly-promoting factors, MT-stabilizing factors (such as MT-associated proteins [MAPs]), MT-destabilizing factors, MT-severing proteins, and also MT-based motors of the kinesin superfamily and dynein (Desai and Mitchson, 1997). Among these, structural MAPs that include MT assemblyepromoting proteins MAP1 (1a and 1b), MAP2, and Tau function to stabilize MTs. One important characteristic of a neuron is its unique MAP components. For example, Tau is concentrated in axons, whereas MAP2 is localized in dendrites. MAP1b and Tau proteins were among the first proteins to be implicated in regulating MT organization and dynamics. Both MAP1b and Tau are required for axon extension, but there is functional redundancy between them. Mice lacking Tau or MAP1b have severe axon elongation defects, underscoring the importance of these MAPs in axon formation and maintenance (Takei et al., 2000). Tau has been studied extensively for its involvement in neurofibrillary tangle formation in Alzheimer’s disease and in frontotemporal dementia with parkinsonism-17 (FTDP-17). A possible mechanism underlying these diseases is that the presence of too many Tau proteins, which can be caused by genetic mutations that affect Tau expression, becomes a roadblock that hinders the movement of motor proteins along MTs, impairing axonal transport. In addition, neurons often respond to a stress, such as accumulation of amyloid-b (Ab) peptides derived from the amyloid precursor protein (APP), by phosphorylating Tau, which causes it to dissociate from MTs. Subsequently, hyperphosphorylated Tau forms filamentous aggregates that further damage axons. In addition, Tau-depleted MTs become highly vulnerable to MT-severing proteins, and as a consequence, the axon structure breaks down (Baas and Qiang, 2005). Furthermore, it has been shown that genetically reducing Tau can prevent the Ab-induced defects in axonal transport (Vossel et al., 2010). The function of these MAPs can be regulated through phosphorylation by mitogen-activated protein kinases (MAPKs) (Sanchez et al., 2000). The c-Jun N-terminal protein kinase (JNK) subfamily of MAPKs is particularly relevant to axon maintenance. JNK1 can phosphorylate MAP1b and MAP2B to promote its MT-binding ability and facilitate MT assembly. In Jnk1
/
mutant mice, the initial development of the nervous system is normal. However, during the postnatal growth of Jnk1
/
mice, MTs are progressively lost in axons and dendrites, and the anterior commissural and spinal cord axons become swollen and gradually degenerate (Chang et al., 2003). The function of JNK in axon maintenance is apparently evolutionally conserved, since in the absence of JNK protein in Drosophila (called Basket in Drosophila), mushroom body (MB) axons in the central brain become unstable and degenerate (Rallis et al., 2010). Surprisingly, heterozygous loss of JNK (Basket) causes MB axons to overextend. This and other studies suggest that the JNK kinases are “double-edged” swords. It turns out that JNK also phosphorylates JNK-interacting protein 1 (JIP1). JIP1 is a linker protein that associates kinesin-1 with certain membrane proteins or vesicle cargoes. Elevated JNK activity disrupts the interaction between kinesin-1 and JIP1, leading to the dissociation of cargoes from kinesin motors and thereby negatively affecting axonal transport (Koushika, 2008). The JNK signaling pathway is generally activated in response to stress. It is thus conceivable since axons are subject to moderate, yet frequent, stresses caused by growth, body movement, or other physiological/ metabolic processes that MT destabilization is a common occurrence. The “stressed” MTs activate JNK signaling, which in turn phosphorylates MAPs, promotes MT assembly, and restabilizes axonal MTs. Moreover, JNK signaling also induces transcriptional changes that may lead to expression of the genes important for axon maintenance. In addition to MAPs, tubulins have been shown to undergo a variety of posttranslational modifications that regulate the stability of MTs. The common modifications include tyrosinationedetyrosination, acetylationedeacetylation, Snitrosylation, polyanimation, phosphorylation, and others (Yu et al., 2015). These modifications have been suggested to contribute to the formation of stable MT segments that are resistant to MT-destabilizing factors (Yu et al., 2015). These

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stable domains of MT may serve to regulate the axonal cytoskeleton by organizing and perhaps nucleating MTs. For example, mutations of mec-17 protein, the enzyme responsible for the acetylation of MTs in Caenorhabditis elegans, lead to the MT instability and impair the axonal transport (Neumann and Hilliard, 2014).

10.2.3 Defects in motor proteins cause axon degeneration Axonal transport is crucially dependent on molecular motors to carry cargoes to and from axons (De Vos et al., 2008). Mutations in several kinesin motors (anterograde) and in proteins associated with the cytoplasmic dynein motor (retrograde) have been identified as being associated with diseases of motor or sensory axon degeneration, including hereditary spastic paraplegia (HSP), axonal forms of CharcoteMarieeTooth disease (CMT2), and also amyotrophic lateral sclerosis (ALS). For example, mutations of the conserved residues in KIF5A, a kinesin heavy chain for the kinesin-1 motor, cause a rare form of HSP. A loss-of-function mutation in KIF1b disrupts the transport of certain synaptic vesicles and causes CMT2A. Dynactin is a protein complex copurified with dynein. The complex has a molecular mass of 1 MDa, nearly as large as that of cytoplasmic dynein itself, and it is composed of 11 different subunits that include the largest subunit, p150Glued. Dynactin functions to enhance the processivity of dynein motor, targets dynein to specific subcellular locations, and coordinates the bidirectional movement of dynein-attached cargoes. Missense mutations in p150Glued have been linked to both familial and sporadic ALS, and the mutant form of p150Glued is shown to directly impair vesicular transport in motor axons (De Vos et al., 2008). Two elegant studies revealed a role of p150Glued in initiating retrograde transport at synaptic termini, and mutations of the protein’s CAP-Gly domain lead to both Perry syndrome and the distal hereditary motor neuropathy 7B (Lloyd et al., 2012; Moughamian and Holzbaur, 2012). The interactions between motor proteins and MTs play important roles in axonal transport and thus axon maintenance. Studies found that kinesins and dynein show different sensitivities to the amount of Tau on MTs. Kinesins often stall on MTs at the sites where Tau binds, and they can even detach from MTs at these locations. By comparison, the binding of Tau to MTs has a milder effect on the dynein motor (Dixit et al., 2008). The ability of dynein to move bidirectionally and to take sideway steps to an adjacent MT, which kinesins rarely do, makes dynein less sensitive to Tau and enables it to bypass a Tau obstacle. In cases of abnormal Tau expression, such differential effects of Tau on kinesins versus dynein movement would result in the reduced anterograde transport and a net increase in retrograde transport. As a consequence, distal axons become gradually depleted of transported materials and thereby undergo axon degeneration.

10.2.4 Role of mitochondria transport in axon maintenance The essential role of mitochondria as the ATP fuel supplier for axonal functions such as transport and synaptic vesicle release implies that any damage to mitochondria and/or to their subcellular distribution will have deleterious effects on axon maintenance. Mitochondria are highly dynamic organelles that constantly undergo fusion and fission. Dynamin-related GTPases are the core components of the machineries that mediate mitochondrial fusion and fission. For example, mutations in OPA1, the gene encoding an intramitochondrial dynamin, result in dominant optic atrophy. Also, abnormal function of the mitochondrial fission protein dynamin-related protein 1 contributes to Huntington’s disease (Guo et al., 2013). Several studies support the role of KIF5b as the major kinesin motor responsible for the anterograde transport of mitochondria, and KIF1B may also participate in this process. Moreover, several adaptor proteins that specifically couple mitochondria to kinesin or dynein motors have been identified (De Vos et al., 2008; Salinas et al., 2008). For example, Syntabulin and Milton are two major adaptor proteins that link mitochondria to kinesins. Syntabulin loss-of-function impairs the anterograde but not retrograde transport of mitochondria, though how Syntabulin associates with mitochondria is unclear. Milton was first identified in Drosophila in the context of mutations in this gene that cause the depletion of mitochondria in axons and synapses. The association of the Miltonekinesin (KIF5) motor with mitochondria requires two mitochondrial outer membrane proteins: Miro, a Rho-like GTPase (Stowers et al., 2002), and Mitofusins (Mfn1 or Mfn2) (Fig. 10.3). Importantly, a mutation in Mfn2 gene was found to cause CMT2A disease with degeneration of peripheral sensory and motor axons. The disease-associated form of Mfn2 leads to mitochondria clustering in neuronal cell bodies and proximal axons. Loss of Mfn2 function significantly slows down axonal transport of mitochondria, which is independent of its role in the fusion of mitochondria (Misko et al., 2010). Notably, reduction of mitochondria numbers in axons leads to diminished ATP production and thereby impairs the transport of other cargoes. It has been proposed that the anterograde transport of mitochondria delivers these organelles to the sites of high ATP demand in axons, whereas retrograde mitochondria transport delivers damaged mitochondria back to cell bodies for degradation (Miller and Sheetz, 2004). This idea may be particularly relevant to understanding neurodegenerative diseases associated with mitochondria damage. For instance, the number of transported mitochondria in axons decreases with age,

222 PART | I Formation of axons and dendrites

Kinesin Mitochondrion KIF5b

Motor

Neck

Adaptor proteins

Stalk

Tail

Mitochondrial protein

MILTON

Miro

Syntabulin

Mitofusins

Anterograde Retrograde

−

+

Microtubules FIGURE 10.3 Schematic representation of anterograde transport of mitochondrion. Kinesins (KIF5 or KIF1) mediate anterograde transport of mitochondria. Milton and Syntabulin are adaptor proteins that link mitochondria to kinesins. Miro and Mitofusins are mitochondria outer membrane proteins that interact with Milton.

and such decrease precedes the degenerative process in a mouse model of glaucoma (Takihara et al., 2015). Also, mutations in the gene encoding LRRK2, a protein associated with Parkinson’s disease, delay the sequestration of damaged mitochondria and their consequent degradation via mitophagy (Hsieh et al., 2016). Similarly, lysosomal deficits impair the timely clearance of damaged mitochondria in ALS-inflicted motor axons (Xie et al., 2015). It should be pointed out that, in addition to ATP production, mitochondria also participate in Ca2þ signaling and Ca2þ storage in axons. Furthermore, mitochondria are in constant contact with the endoplasmic reticulum (ER). In fact, the aforementioned Mfn2 protein localizes to both ER and mitochondria. Mfn2 forms trans homo- or heterodimers with Mfn2 or Mfn1 located on mitochondria to help tether mitochondria to the ER. The function of such close apposition between the ER and mitochondria is only beginning to be explored (de Brito and Scorrano, 2010). These functions include cooperation in synthesizing and exchanging phospholipids, as well as collaboration in regulating Ca2þ release and uptake, both of which are important for axon maintenance.

10.2.5 Membrane transport and insertion are essential for axon maintenance Among all of the cellular contents that are required for axon maintenance, membranes deserve special consideration here. The surface area of a nonneuronal cell, roughly spherical shape with a diameter of 20 mm, is about 1256 mm2. By contrast, owing to elaborate dendritic and axonal processes, the surface area of a typical vertebrate neuron is over w250,000 mm2. This area can easily amount to millions of mm2 for neurons with long axons. Maintaining this vastly expanded plasma membrane (also called plasmalemma) over long distances is a complex yet crucial process for neurons (Pfenninger, 2009). In addition, after the axons reach their targets, initially through de novo outgrowth in early development, the continued and often dramatic growth of animals during postnatal and adolescent periods requires a concomitant expansion and addition of axonal plasmalemma. This phase of axon elongation, with its terminal tethered to its target, is called “networked axon growth” and posts yet another challenge for axon maintenance. The bulk of phospholipids and membrane proteins in neuronal cell bodies are synthesized in the ER. Most of our knowledge on membrane transport and insertion in neurons came from studies of de novo axon outgrowth of cultured embryonic neurons. Since an embryonic axon 1 mm in diameter typically extends 0.5 mm in length per day, this translates into a rate of surface area increase at 1 mm2 per minute. Thus, a significant amount of membrane must be synthesized and

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delivered to the growing axon. During such de novo outgrowth, new membranes are preferentially added to the growth cones, at the tips of extending axons, rather than to the existing axon shaft. Pleomorphic vesicles, called plasmalemmal precursor vesicles (PPVs) with a diameter of w150 nm and enriched for newly synthesized phospholipids, are the source of new membranes. These PPVs are thought to be transported by KIF2, KIF4, or kinesin1 to growth cones. Subsequent fusion of PPVs with the plasma membrane, a process aided and regulated by the exocytosis machinery, enables the growth and extension of axons (Pfenninger, 2009). Some studies suggest that fusion of PPVs with the plasma membrane is partly regulated by signaling through the insulin-like growth factor 1 receptor (IGFR1) in cultured neurons. IGFR1 stimulates PI3K-Akt signaling, which could in turn activate a Rab family member that then recruits and facilitates Exocyst-mediated docking of PPVs with the growth cone membrane (laurino et al., 2005). Surprisingly, little is known about membrane transport and insertion during networked axon growth in postnatal animal development. Once growth cones are transformed into presynaptic termini, there is no longer any apparent focal point for new membrane insertion, at least none that are currently known. From an economical point of view, adding new membranes to the proximal end of networked axons followed by lateral diffusion reduces the need for long-distance transport. On the other hand, membranes could be inserted at multiple sites along the entire length of an axon. It should be noted that the smooth ER (SER) is distributed throughout axons as a continuous tubular network, raising the possibility that phospholipids may be synthesized locally in the axonal SER. In this respect, it is worth pointing out that mutations affecting SER formation can cause HSP, a disease characterized by degeneration of distal axons from neurons that project the body’s longest axons such as the corticospinal neurons. Autosomal-dominant mutations in the genes encoding Spastin, Atlastin, or REEP1 are the most common causes of HSP. Recent studies indicate that these three proteins interact with each other, localize to the ER, and mediate the formation of SER as well as the interactions of ER network with the MT cytoskeleton. Furthermore, ER morphological defects appear to be one of the main pathogenic mechanisms in HSP. Thus, although these findings do not directly prove the role of axonal SER in providing membrane sources for plasmalemma, they do lend support for the importance of local ER network in maintaining the health and integrity of axons (Renvoisé and Blackstone, 2010).

10.3 Proteasome and autophagy pathways in axonal homeostasis 10.3.1 Ubiquitineproteasome system in axon maintenance Misfolded or unwanted proteins can be degraded by the ubiquitineproteasome system (UPS). The UPS has a general housekeeping role in cellular homeostasis. Here, we discuss findings supporting a crucial role for the UPS in maintaining axonal functions and dynamics. These findings are from studies of Gigaxonin gene, which is mutated in human giant axonal neuropathy (GAN) (Yang et al., 2007). Patients with GAN have a postnatal-to-childhood onset of axon degeneration. The most severely affected axons are those that form long-distance projections, including peripheral nerves and the corticospinal tract. Gigaxonin is a cytoplasmic protein containing a BTB domain and six Kelch repeats. It binds directly to the ubiquitin-activating enzyme E1 and controls the ubiquitin-mediated degradation of MAP1B light chain (MAP1B-LC), MAP8, and tubulin folding cofactor B (TBCB). Loss-of-function Gigaxonin mutations result in the accumulation of all three substrates in neurons. The MT-associated protein MAP1b is cleaved posttranslationally into a heavy chain and a light chain. MAP8 is another MAP with expression enriched in neurons. Neurons overexpressing either MAP1b-LC or MAP8 exhibit fragmented axons and progress to cell death. It is hypothesized that accumulation of MAP1B-LC or MAP8 either alters MT organization or obstructs motor proteins from binding to MTs, thereby impairing axonal transport. Thus, the level of these MAPs must be exquisitely regulated for maintaining axonal integrity. On the other hand, TBCB belongs to the family of tubulin-specific chaperons. TBCB is an MT-destabilizing factor, and its accumulation leads to loss of MTs. In fact, reduction of MTs in the enlarged and swollen axons is a common pathological feature of GAN. Therefore, Gigaxonin-mediated UPS degradation of MAP1B-LC, MAP8, and TBCB, together with other putative substrates, is essential for proper MT organization and dynamics to ensure axonal transport and integrity.

10.3.2 Role of autophagy/lysosome pathway in maintaining axonal homeostasis Autophagy is an evolutionarily conserved catabolic pathway in which cytoplasmic contents, including proteins and organelles, are wrapped by double-membraned vesicles, which are then fused with lysosomes for bulk degradation. Recent studies have revealed connections between autophagy and major neurodegenerative disorders such as Alzheimer’s, Parkinson’s, and Huntington’s diseases. In particular, autophagy plays essential roles in degrading disease-related, aggregate-prone mutant proteins such as tau, a-synuclein, and huntingtin. Here, we will focus on the role of autophagy in axonal homeostasis.

224 PART | I Formation of axons and dendrites

Studies of cultured neurons showed that autophagosomes are formed in axons and retrogradely transported to neuronal soma, to be fused with lysosomes. Deletion of autophagy-specific genes in Purkinje neurons resulted in the accumulation of abnormal membranous organelles, including structures resembling autophagosomes in the axonal termini (Nishiyama et al., 2007; Yue et al., 2008). These observations lead to the hypothesis that basal autophagy in axons is required for recycling of axonal membranes and/or vesicles at the axonal termini, perhaps to support synaptic activity and maintain axonal homeostasis. As discussed earlier, another major substrate of autophagy in axons is damaged mitochondria, which need to be cleared in a timely manner to prevent their destructive effects on axonal energy production.

10.4 Role of glial cells in axon maintenance The interaction between neurons with surrounding glia is a cardinal feature of all nervous systems, and this is discussed extensively in Chapters 16, 17, 23 and 25. Readers are encouraged to read these other chapters for details on glial cell’s involvement in neuronal migration and axon guidance. Here, we will briefly summarize two key points that are particularly relevant to the roles of glial cells in axon maintenance, which include both myelin-dependent and myelin-independent mechanisms. A major function of oligodendrocytes in the CNS and Schwann cells in the PNS is to generate large amounts of myelin, which wraps around axon segments many times. Myelin sheaths insulate axons and thus enable rapid salutatory electrical conduction. Loss of intact myelin underlies many neurological diseases, including multiple sclerosis, inherited leukodystrophies of the CNS, and several types of peripheral neuropathies. The causes of axon degeneration in these demyelinating diseases are complex. After all, neurons can grow and maintain axons in cultured conditions without glia and myelin. It is believed that in vivo myelin protects axons from the cytotoxic attack of peripheral immune cells or activated microglia (Neumann et al., 2002). In addition, loss of myelin will cause sodium channels to be redistributed along the axon, and thereby, more sodium channels are required per unit length to conduct electric impulses. Consequently, demyelinated axons consume considerably more energy and could overtake ATP production capacity by local glycolysis and mitochondria. Such an axonal energy deficit will lead to the failure of the Naþ/Ca2þ exchanger and Ca2þ-ATPases and thus result in an increase in cytosolic Ca2þ in axons. Notably, high Ca2þ levels trigger Ca2þ-dependent proteases called calpains that can degrade a variety of cytoskeleton proteins. Also, accumulation of axonal Ca2þ further impairs mitochondrial functions and eventually causes axon degeneration (Trapp and Stys, 2009). Importantly, myelinating glial cells also provide trophic support for axon functions in addition to generating myelin for physical protective barriers. This was discovered through genetic studies of mouse mutants that affect glial cells without impairing myelin synthesis (Griffiths, 1998; Lappe-Siefke et al., 2003). For example, in the absence of PLP and DM20, two myelin membrane proteolipid proteins, oligodendrocytes in the mutant mice still assemble compact myelin sheaths that are sufficient for rapid impulse propagations. However, axonal transport is significantly slowed in adult mutant mice, and subsequently, axons undergo degeneration in all regions of the CNS. Myelin-associated glycoprotein (MAG) is another myelin component important for axonal maintenance. Mutant mice lacking MAG have normal myelination and no symptoms in early adulthood; however, their sciatic nerves exhibit reduced diameter and progressive axon degeneration (Li et al., 1994; Montag et al., 1994). The molecular mechanisms underlying such myelin-independent functions of glial cells in axon maintenance have begun to be understood. In particular, it is now clear that myelinating glial cells provide essential metabolic support for axonal survival and function. For instance, pyruvate and lactate produced from glycolysis in oligodendrocytes serve as an important energy source for local axon metabolism (Fünfschilling et al., 2012). Also, mice with the mitochondrial dysfunction specific in Schwann cells exhibit the progressive peripheral neuropathy (Viader et al., 2011).

10.5 Maintaining axon track positions and other structural features In addition to the mechanisms responsible for maintaining axonal structure and functions, studies in the nematode nervous system reveal dedicated molecular mechanisms that maintain axon track positions (Benard and Hobert, 2009). As the worm undergoes developmental growth from larva to adult, its nervous system must keep pace with the growth of its body and also withstand the mechanical stress resulting from locomotion. Genetic screens identify a set of genes that are required for maintaining the position of axons within fascicles. Intriguingly, axons in mutant animals are indistinguishable from those in wild-type animals during early development, and their positioning defects only appear in later stages of life. These axon-positioning factors include several Ig-domain containing proteins such as SAX-7, ZIG family proteins, DIG-1, and EGL-15/FGFR. For example, worms carrying loss-of-function mutations in the sax-7 gene fail to maintain the proper axon position in the ventral nerve cord, and they also have altered position of the neuronal soma in various head ganglia. SAX-7 is an

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ortholog of the vertebrate L1 family members (L1, CHL1, Neurofascin, and NrCAM). Interestingly, mutations in human L1 are associated with several neurological diseases including X-linked complicated spastic paraplegia type 1 and X-linked agenesis of the corpus callosum. These observations highlight the importance of axon interactions with extracellular components for positioning and maintaining their position in a dynamic local environment. It will be interesting to examine whether failure in maintaining axon track position contributes to the etiology of certain neurological diseases. Also worth noting is the membrane-associated periodic actin skeleton, which consists of F-actin rings separated by w190 nm spacers of spectrin tetramer and other auxiliary structural proteins (Xu et al., 2013). This actin cytoskeleton is ubiquitously present in mature axons. The precise function of this periodic cytoskeleton structures in axon maintenance is not yet clear, and how such periodic structures are regulated in axons also awaits further investigation. Since axons need to endure stretch and movements, one speculated function of the periodic actinespectrin scaffold is to provide axonal shafts with rigidity and stability (Dubey et al., 2018).

10.6 Axon pruning and axon degeneration Selective elimination of axons and/or their presynaptic structures without death of the parental neurons occurs both under physiological conditions during normal development and also in pathological conditions in adulthood. Axon pruning in physiological contexts is critical for establishing precise neuronal connections by enabling the removal of exuberant or misguided axonal branches. Like other morphological changes during neural development, axon pruning is tightly regulated by a combination of extracellular and intracellular signals. On the other hand, axon degeneration in pathological conditions happens frequently in many types of neurodegenerative diseases and in injuries to axons caused by toxic, ischemic, or traumatic insults (Salvadores et al., 2017).

10.6.1 Developmental axon pruning A common feature of neuronal development is that initially axons often connect to more targets, and targets are often innervated by more axons. Thus, transforming these exuberant connections to precise circuits requires the removal of axonal branches, either at a small scale such as observed during the elimination of axon terminal arbors or at a large scale involving the removal of inappropriate axon collaterals of significant length (Fig. 10.4). Importantly, this sort of developmental axon pruning is highly selective and does not damage the neuronal cell body or the branches that remain preserved. Small-scale axon terminal arbor pruning is a local process that eliminates extra or ectopic synapses and, in many cases, involves a mechanism of neural activity-dependent competition. A classic example of such developmental synapse elimination is the refinement of polyneuronal innervation to mononeuronal innervation at the vertebrate neuromuscular

Hippocampus CA1 CA3

CA3 DG

Visual cortex Layer V visual cortical neuron

FIGURE 10.4 Examples of large-scale axon pruning during vertebrate development. Upper panel: In the hippocampal mossy fiber projections, the infrapyramidal branches of the mossy fibers originated from the dentate granule cells to the CA3 region are pruned. Lower panel: Long projections from layer V visual cortical neurons to multiple subcortical regions are pruned.

226 PART | I Formation of axons and dendrites

junction. This process occurs during postnatal period whereby through competition, the terminal presynaptic arbors from all but one “winning” axon are removed. In vivo imaging studies showed that the small branches of the “losing” axons initially form swellings, which the researchers termed “axosomes.” Subsequently, the connections among axosomes thin out and disappear. Finally, axosomes are phagocytosed by Schwann cells and degraded in lysosomes (Bishop et al., 2004). Similar axosome engulfment was observed during the elimination of climbing fiber inputs onto Purkinje cells in the cerebellum, suggesting that this may be a common cellular mechanism for small-scale axon terminal pruning (Neukomm and Freeman, 2014). One of the best-known examples of large-scale developmental axon collateral elimination is the removal of inappropriate subcortical axonal projections originating from visual and motor cortices. Initially, layer 5 neurons from both cortices all project to the spinal cord, multiple targets in the brain stem, and the superior colliculus. Within the first 2e 3 weeks of postnatal development, only the collateral branches that are functionally appropriate for each cortical region are retained. Specifically, neurons from the visual cortex eliminate the projections into the spinal cord as well as other brain stem nuclei related to motor functions (Fig. 10.4). An intrinsic factor, the homeodomain transcription factor Otx1, was found to regulate the pruning of axons from the visual cortex, but the exact mechanism by which it functions remains to be fully determined. Recent studies indicate that Sema3F, a member of the semaphorin family of axon guidance molecules, is strongly expressed in the dorsal spinal cord at the time of visual cortical axon pruning. Interestingly, the Sema3F holoreceptor components, which include Neuropilin-2, Plexin-A3, and Plexin-A4, are specifically expressed in the visual, but not motor, cortical neurons. Genetic analyses confirm that Sema3F repulsive signaling through its holoreceptor is required for the proper removal of visual cortical axons from the spinal cord (Vanderhaeghen and Cheng, 2010). In addition, in the developing hippocampus, dentate gyrus granule cells extend a transient bundle of mossy fiber collaterals that course adjacent to the basal dendrites of pyramidal neurons in the CA3 region, which is termed the infrapyramidal bundle (IPB). Axons in this transient IPB are later retracted, and this retraction is also triggered by Sema3F-mediated repulsive signaling (Riccomagno and Kolodkin, 2015). Repulsive axon guidance molecules also play important roles in stereotyped axon pruning during the development of topographic neural maps. For instance, in the retinotopic map formed by the retinal ganglion cells (RGCs) projecting to the superior colliculus, RGCs initially overextend their axons beyond their intended targets. This is followed by the sprouting of interstitial axon arbors in the appropriate terminal zone. The removal of overextended axons is regulated by graded expression of ephrins, a family of repulsive guidance molecules, in the superior colliculus. RGC axons express Eph family tyrosine kinases that serve as receptors for ephrins. Graded ephrin-Eph repulsive signals therefore enable pruning of the overextended axon branches beyond the terminal zone (Schuldiner and Yaron, 2015). Axons extending from neurons that have been deprived of their physiological trophic factors also undergo selfdestruction, and this may represent an additional mechanism of developmental axon pruning. For example, when the distal part of an axon of cultured sympathetic neuron is locally deprived of nerve growth factor (NGF), that part of the axon degenerates, whereas the rest of the cell body and axon survives (Raff et al., 2002). Recent studies have begun to reveal the molecular mechanisms underlying such trophic deprivation-induced axon pruning. It has become clear that the classical apoptosis pathway exerts an essential role in this process. Genetic deletion of caspase-3, caspase-9, or Bax, key components in executing apoptosis, all dramatically delay the degeneration of NGF-deprived sensory axons (Simon et al., 2012). Additionally, calpains, another family of intracellular proteases, function downstream of caspases to cause cytoskeleton degradation and promotion of axon degeneration (Yang et al., 2013). Several upstream signals can trigger the apoptosis pathway. For instance, DR6, a TNF receptor family member, is involved in the activation of caspases in NGFdeprived sensory axons. This DR6-dependent pathway also appears to participate in the developmental axon pruning of the retinotopic projections in the superior colliculus, as discussed earlier (Nikolaev et al., 2009). In invertebrates, it is known that the insect nervous system undergoes extensive remodeling of dendritic and axonal structures and connections during metamorphosis. The insect hormone ecdysone is the major extrinsic factor that triggers metamorphosis and the global pruning of many, but not all, types of axons and dendrites. This pruning process has been studied in great detail for one class of neurons called g neurons in the Drosophila MB, a brain structure implicated in learning and memory. MB g neurons lose the majority of their dendrites and dorsal and medial axonal collaterals at the onset of ecdysone signaling through the degenerative process. It begins with depolymerization of MTs, followed by neurofilament degradation, synapse disassembly, axon fragmentation, and engulfment of axonal debris by glial cells (Luo and O’Leary, 2005). Furthermore, the MB g neuron pruning process cell autonomously requires activities of nitric oxide synthase (NOS) that produce NO. Overexpressing NOS or elevating NO concentrations stimulates axon pruning even when ecdysone level is low, whereas loss of NOS resulted in pruning defects (Rabinovich et al., 2016). Notably, NOS activity is downregulated at the onset of axonal regrowth after metamorphosis; thus, NO functions as a switch between axon pruning and regrowth in Drosophila. In addition, another related model in Drosophila is the class IV dendrite

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arborization sensory neurons, which undergo extensive dendrite pruning during early metamorphosis. Curiously, this dendrite pruning process is regulated by caspase activity as well as calpains (Williams et al., 2006; Kanamori et al., 2013), reminiscent of the involvement of caspases and calpains in the trophic deprivation-induced axon pruning of mammalian sensory neurons as discussed earlier.

10.6.2 Pathological axon degeneration Acute axon degeneration: In certain traumatic brain or spinal cord injuries, CNS axons may be severed. Surgeries and wounding could also lead to acute transection of peripheral nerves. Within minutes to a few hours immediately after such axotomy, there is a sudden axonal destruction that extends for approximately a few hundred micrometers proximal and distal to the lesion site. This process is termed acute axon degeneration (AAD) (Kerschensteiner et al., 2005). It is thought that the rapid influx of extracellular Ca2þ upon injury triggers AAD. Electron microscopic studies of cultured neurons revealed that immediately following axotomy, double-membraned vacuoles resembling autophagosomes accumulate in the axon segments close to the lesion site. A study on optic nerve axons in vivo confirmed the appearance of autophagosomes within the first few hours following injury (Knöferle et al., 2010). It has been suggested that axotomy-induced calcium influx triggers axonal autophagy and underlies the cellular mechanism of AAD. In addition, the influx of Ca2þ activates the Ca2þ-dependent calpains, which can further disrupt the axonal integrity. Chronic axon degeneration/Wallerian degeneration: Our current understanding of mechanism underlying chronic pathological axon degeneration has benefitted from studies on Wallerian degeneration, which was first described by Augustus Waller in his nerve transection experiments in frogs (Waller, 1850). In rodents, within 24e48 h after traumatic axotomy, distal axons and their synaptic termini undergo stereotyped degeneration that includes loss of membrane potential, fragmentation of cytoskeleton, membrane swelling, and destruction of axonal integrity. The debris of degenerated axons and also surrounding myelin sheaths is then cleared by macrophages in the PNS and by microglia in the CNS. It is important that although direct axonal transection is not common in clinics, chronic axon degeneration prevails in many neurodegenerative diseases. In particular, the process of axon breakdown in pathological conditions shares many similarities with Wallerian degeneration. For example, axonal swelling, an early sign of axon degeneration, has been identified in mouse models of Alzheimer’s disease and also in the early stages of Alzheimer’s disease in patients. These swollen axons accumulate abnormal levels of MT-associated proteins, motor proteins, organelles, and vesicles. In addition, in many cases, the initial degeneration of distal regions of long nerve fibers is followed by subsequent distal-to-proximal progression over time: A pathological pattern termed “dying back” or distal axonopathy. Notably, strategies used to rescue neurons or their axons from apoptotic cell death have been largely unsuccessful in attenuating the progression of such axonal disease phenotypes (Duncan and Goldstein, 2006), suggesting the presence of nonapoptotic mechanisms in pathological axon degeneration. Efficient therapeutic intervention directed at neurodegenerative diseases requires axonal protection, and thus, understanding the mechanisms underlying pathological axon degeneration is crucial for developing such therapeutics.

10.6.3 Molecular mechanisms of pathological axon degeneration The serendipitous discovery in the 1980s of the Wallerian degeneration slow (Wlds) mutant mouse, in which axons distal to a traumatic injury site remain intact and functional for about 3e4 weeks, provided significant insights into the molecular mechanisms regulating pathological axon degeneration (Fig. 10.5). The Wlds phenotype is caused by overexpression of a chimeric Wlds mutant gene, which contains the coding sequence of the N-terminal 70-amino-acid fragment of the ubiquitination factor E4B Ube4b (homologous to yeast Ufd2) fused to the entire coding region of the NADþ biosynthetic enzyme nicotinamide nucleotide adenylyltransferase Nmnat1 Mack et al. (2001). The axon-protective effect of Wlds is genetically dominant and intrinsic to axons. Apoptotic cell death is not affected in Wlds mice, providing additional support for the idea that pathological axon degeneration and apoptotic cell death are regulated by distinct molecular mechanisms. It has subsequently been shown that the Wlds gene is a potent inhibitor of axon degeneration upon traumatic injury both in the PNS and in the CNS in a variety of species, ranging from rodents to fruitflies. Interestingly, the Wlds mutant protein exhibits axonal protective effects in pathological conditions beyond traumatic axotomy, including those caused by neurotoxic damage or neurodegenerative insults such as glaucoma, raising the possibility that pathological axon degeneration shares certain common regulatory mechanisms (Coleman and Freeman, 2010). Mechanistically, a large body of studies point to the role of Wlds protein in maintaining axonal NADþ levels. NADþ is a central coenzyme for normal metabolic processes, and its concentration plummets in axons immediately preceding to their fragmentation. Axotomy-induced degeneration can be blocked by exogenous NADþ or its precursors (Wang et al., 2005). The axon protective effect of the Wlds protein depends on the activity of Nmnat1 and results from its cytosolic localization in axons. In fact, direct delivery of recombinant Nmnat1 protein to axotomized axons is sufficient to suppress their degeneration (Sasaki and Milbrandt, 2010). The current model postulates that axon degeneration after injury is

228 PART | I Formation of axons and dendrites

Mouse sciatic nerve

Drosophila olfactory sensory axons

Rat DRGs axon (in vitro culture)

Injured WIds

Injured WT

Uninjured

Mouse phrenic nerve

FIGURE 10.5 Conserved protective effects of Wlds on delaying Wallerian degeneration. Schematic representations of experimental observations in four different axon injury models are shown here. In each model, top row illustrates uninjured nerve or axons, middle row shows degenerative phenotypes after axotomy in wild-type animals, and bottom row depicts results when these neurons overexpress Wlds. The most left column represents transverse section of the phrenic nerve, and each circle represents a myelinated axon. The lines in the right three columns represent axons. Wlds overexpression protects axons from degeneration after axotomy both in vivo and in vitro models, and the protective effect is conserved in both mammalian axons and olfactory receptor neurons in Drosophila.

activated upon depletion of Nmnat2, a short-lived Nmnat and the only NADþ-synthesizing enzyme normally present in axons. The Wlds/Nmnat1 protein, which is stable in axons, compensates for the rapid degradation of Nmnat2, thereby maintaining NADþ homeostasis in axotomized axons (Gilley et al., 2013). Beyond the serendipitous discovery of WldS, genetic screens in Drosophila played a critical role in identifying molecular mechanisms dedicated to pathological axon degeneration in response to traumatic axotomy. Loss-of-function mutations of the dSarm/Sarm1 gene (which encodes sterile a/Armadillo/Tolleinterleukin receptor homology domain protein) have been identified to confer strong protection of axotomized axons in a variety of neuronal types in mice and also in Drosophila (Osterloh et al., 2012). How might dSarm/Sarm1 execute axon death? The ortholog of Sarm1 gene, named tir-1, is originally reported in C. elegans as a key regulator of asymmetric neuronal development through its downstream MAPK signal (Chuang and Bargmann, 2005). A recent study showed that this evolutionarily conserved Sarm1-MAPK pathway drives pathological axon degeneration. Genetic depletion of MAPKKK proteins (DLK, MLK2, and MEKK4) or MAPK proteins (JNK1, JNK2, and JNK3) in the Sarm1-MAPK pathway results in the significantly prolonged survival of axotomized axons (Yang et al., 2015). In addition, similar to that observed with the Wlds mutant protein, the Sarm1-MAPK pathway can function in the scenarios besides traumatic axotomy, such as axon degeneration caused by neurotoxic or ischemic damages (Yang et al., 2015), lending further support for the shared molecular mechanism of pathological axon degeneration. Moreover, it is intriguing to note that inhibition of MAPKKK protein DLK, and its downstream JNK proteins, also suppresses degeneration of NGF-deprived axons (Ghosh et al., 2011), revealing a potential convergent regulation for physiological pruning and pathological degeneration. On the other hand, another study argues that MAPKs function upstream of Sarm1 signaling. MAPKs promote axon degeneration by accelerating the turnover of the Nmnat2 enzyme, thereby limiting the axonal level of NADþ (Walker et al., 2017). Decreased NADþ activates Sarm1 through an as yet unidentified mechanism. Interestingly, axonal NADþ loss after axotomy is blocked in Sarm1
/
mutant (Gerdts et al., 2015). The Sarm1 TIR domain appears to have endogenous NAD þ hydrolase activity (Essuman et al., 2017). This surprising finding has led to the idea that activated Sarm1 further depletes axonal NADþ through its NADþ hydrolase function, causing catastrophic energy failure and ultimately resulting in explosive axon fragmentation. However, a recent study suggests that this linear MAPK/Nmnat2/Sarm1-dependent NADþ-depletion model is oversimplified (Neukomm et al., 2017). A screen in Drosophila identifies a BTB/BACK domain protein called Axundead (Axed). The fly has one Nmnat gene. Only Axed mutant, but not dSarm mutant, can fully block the neurodegeneration induced by loss of Nmnat in fly. Axed is the convergent downstream target of both Nmnat elimination- and dSarm activation-mediated axon death pathway (Neukomm et al., 2017). It will be interesting to know whether loss of Axed also prevents axon degeneration in the mammalian system.

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Finally, Wlds expression or Sarm1 deletion delays axon degeneration in traumatic injuries and some, but not all, disease models, suggesting the existence of other molecular pathways. Indeed, ongoing work continues to reveal novel mechanisms involved in pathological axon degeneration. For example, a recent report suggests that the necroptosis pathway might contribute to the process of axon degeneration in familial ALS condition (Ito et al., 2016). To this end, it is also important to note that certain invertebrate axons intrinsically exhibit very slow Wallerian degeneration. For instance, evoked neurotransmitter releases can occur in the severed axonal segments in crustaceans for up to 1 year after the initial transection. These axons are often invaded by hypertrophic adaxonal glia, which are thought to directly transfer proteins to the severed axons to sustain their long-time survival (Parnas et al., 1991). It will be interesting to explore whether such naturally occurring mechanisms that counter pathological axon degeneration are instructive for potential therapeutic applications in humans. Thus, greater in-depth understanding of how axon degeneration pathways exhibit cross-talk with the various mechanisms of axon maintenance will be critical for future directions in the field of axon biology and the therapeutic extensions of this work in health and disease.

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Mack, T.G., Reiner, M., Beirowski, B., Mi, W., Emanuelli, M., Wagner, D., Thomson, D., Gillingwater, T., Court, F., Conforti, L., Fernando, F.S., Tarlton, A., Andressen, C., Addicks, K., Magni, G., Ribchester, R.R., Perry, V.H., Coleman, M.P., 2001. Wallerian degeneration of injured axons and synapses is delayed by a Ube4b/Nmnat chimeric gene. Nat. Neurosci. 4, 1199e1206. Miller, K.E., Sheetz, M.P., 2004. Axonal mitochondrial transport and potential are correlated. J. Cell Sci. 117, 2791e2804. Misko, A., et al., 2010. Mitofusin 2 is necessary for transport of axonal mitochondria and interacts with the Miro/Milton complex. J. Neurosci. 30, 4232e4240. Montag, D., et al., 1994. Mice deficient for the myelin-associated glycoprotein show subtle abnormalities in myelin. Neuron 13, 229e246. Moughamian, A.J., Holzbaur, E.L., 2012. Dynactin is required for transport initiation from the distal axon. Neuron 74, 331e343. Neukomm, L.J., Freeman, M.R., 2014. 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Neuronal autophagy: going the distance to the axon. Autophagy 4, 94e96.

Chapter 11

Dendrite development: invertebrates Wesley B. Grueber1, * and Bing Ye2 1

Columbia University, New York, NY, United States; 2University of Michigan, Ann Arbor, MI, United States

Chapter outline 11.1. Structure and anatomy of invertebrate dendrites 232 11.2. Methods for studying dendrite morphology in Drosophila 232 11.3. Anatomical background for key model systems in which dendritic morphogenesis is studied in invertebrates 233 11.3.1. Drosophila dendritic arborization sensory neurons 233 11.3.2. Drosophila motoneurons 233 11.3.3. Drosophila olfactory projection neurons 233 11.3.4. Caenorhabditis elegans PVD neurons 234 11.4. Cell biology of dendritic growth 235 11.4.1. Microtubule polarity differs between dendrites and axons 235 11.4.2. Dynein-dependent trafficking controls dendritic branching 235 11.4.3. Role of the secretory pathway and Golgi outposts in dendritic elaboration 236 11.5. Transcriptional control of dendritic morphology 237 11.5.1. Control of dendrite morphological identity of Drosophila PNS neurons 237 11.5.2. Transcriptional control of dendritic targeting of olfactory PNs 239 11.5.3. Chromatin remodeling factors and dendritic development 241 11.6. Posttranscriptional control of dendritic development 241 11.6.1. Control of mRNA translation in dendritic development 241 11.6.2. miRNAs in dendritic development 242 11.7. Control of dendritic field formation I: guidance and targeting 242 11.7.1. Slit and netrin signaling during midline dendritic guidance 242

11.7.2. A combinatorial ligandereceptor complex guides dendritic branches 244 11.7.3. Coarse and specific control of PN dendritic targeting 244 11.7.4. Glial control of dendritic targeting 244 11.8. Control of dendritic field formation II: dendritic selfavoidance and tiling 245 11.8.1. Interactions between dendrites generate evenly covered territories 245 11.8.1.1. Dendritic self-avoidance 245 11.8.1.2. Dendritic tiling 247 11.8.2. Scaling growth of arbors and maintenance of evenly covered territories 248 11.9. Dendritic remodeling 249 11.9.1. Transforming growth factor-b signaling and ecdysone receptor expression during dendritic remodeling 249 11.9.2. Sox14 and mical function downstream of ecdysone receptor 249 11.9.3. Signaling mechanisms for dendritic pruning 251 11.9.3.1. Ubiquitineproteasome system 251 11.9.3.2. Caspases 251 11.9.4. The cell biology of dendritic pruning 251 11.9.4.1. Microtubule disassembly 251 11.9.4.2. Local endocytosis and compartmentalized calcium transients 252 11.9.5. Similarities between dendrite pruning and injuryinduced axon degeneration 252 11.9.6. Similarities and differences in dendrite development, dendrite regrowth after pruning, and dendrite regeneration after injury 252 11.10. Concluding remarks 252 References 253

* Senior author.

Cellular Migration and Formation of Axons and Dendrites. https://doi.org/10.1016/B978-0-12-814407-7.00011-0 Copyright © 2020 Elsevier Inc. All rights reserved.

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11.1 Structure and anatomy of invertebrate dendrites Dendrites are the input regions of neurons, and their morphologies determine how information is received and processed by individual neurons. Understanding how complex dendritic arbors develop their shapes presents an important and challenging problem, and one that is under investigation in both vertebrate and invertebrate systems. Invertebrate studies offer numerous fundamental insights that are the focus of this chapter. In particular, studies in model genetic systems have allowed researchers to identify and study genes that control morphogenesis cell autonomously within individual neurons, as well as to identify extrinsic signals that impact dendritic branching and targeting. Advancements in our understanding of dendrite development have followed close behind the development of new approaches for manipulating and studying dendrites. Dendrites and axons can be defined in part by the distinct polarity of their microtubule cytoskeletons. Vertebrate dendrites and axons differ in the orientation of microtubule ends, with microtubules oriented plus-end distal in axons, and mixed plus-end distal, plus-end proximal polarity in dendrites (Baas et al., 1988). An implication of these findings, pointed out by Baas and colleagues, is that “organelle compartmentalization in the neuron is secondary to the generation of microtubule arrays of different polarity orientation in the axon and the dendrite. Thus, the establishment of these distinct microtubule arrays may provide a structural basis for many of the differences that distinguish the dendrite from the axon” (Baas et al., 1988). Indeed, it has been shown that these differences in microtubule polarity impact the trafficking machinery and the cargos carried to dendrites and axons, since kinesin and dynein trafficking complexes are primarily plusend or minus-end directed, respectively. In contrast to typical multipolar vertebrate neurons, invertebrate neurons are often unipolar, meaning that dendrites often, but not always, emerge from a proximal neurite that is continuous with the axon. As discussed below, there are several exceptions to this typical invertebrate neuron, so the differences between vertebrate and invertebrate dendrites are quantitative rather than qualitative (Grueber and Jan, 2004). The unipolar nature of insect neurons means that, by and large, dendritic and axonal compartments are not separated by a neuronal soma as is the case with most vertebrate neurons. Despite these differences, invertebrate dendrites and vertebrate dendrites appear to be homologous and likely derived from similar developmental programs (Sánchez-Soriano et al., 2005). Microtubule polarity likewise differs in the axons and dendrites of insects, but rather than the mixed dendritic microtubule arrangement observed in vertebrates, insect dendrites show almost exclusively minus-end directed microtubules, suggesting that a minus-end microtubule arrangement is the common evolutionarily conserved feature of all dendrites (Stone et al., 2008). These conserved aspects of dendrite biology indicate that knowledge gained in insect systems is likely to be generally applicable to processes that occur in vertebrates.

11.2 Methods for studying dendrite morphology in Drosophila Dendritic morphology is studied by analysis of numerous quantitative arbor features, for example, total dendrite length, branch number and distribution, and territory size. There is therefore good reason to achieve the highest resolution possible of dendrites during experiments, and the standard required for such studies is single-neuron resolution. This is even the case in animals with simpler nervous systems with relatively fewer neurons, such as Drosophila. Several major technical advancements, too numerous to outline in detail, have provided for specific genetic manipulations of neurons, allowing for high-resolution assessment of dendritic morphology in individual neurons. Of particular importance is the binary GAL4eUAS (upstream activation sequence) expression system (Brand and Perrimon, 1993), in which the production of the yeast transcriptional activator GAL4 by individual or small groups of cells can activate transgenes of choice that are linked to the GAL4-binding UAS DNA sequence. The identities of neurons that produce the transgene are determined by the specificity of the enhancer elements that drive the Gal4 cDNA. A large collection of Gal4 enhancer trap lines has been generated and can be used to misexpress genes selectively in different neurons. Building on the GAL4eUAS system, the mosaic analysis with a repressible cell marker (MARCM) technique allows labeling individual wild-type or mutant neurons positively with membrane-associated fluorescent proteins, such as GFP (Lee and Luo, 1999). By virtue of ubiquitous expression of the GAL80 repressor of GAL4eUAS interaction, UASeGFP expression is uniformly suppressed. However, by a clever genetic trick involving targeted recombination, GAL80 suppression is relieved in small groups of cells and so these alone glow brightly in a nonfluorescent background. The remarkable feature of this system is that those same fluorescent neurons can be made mutant for nearly any gene of interest, allowing single-cell resolution of mutant cells. There are several advantages to this system, including the ability to test cell autonomy since the GFP-labeled neurons are also mutant. Many of the insights into dendritic morphogenesis that are introduced in this chapter derive from experiments involving this powerful genetic approach. Variations of this technique, such as twin-spot MARCM (Yu et al., 2009), which independently labels paired sister clones from the same neural progenitor cell, offer additional advantages for dissecting the relationship between dendrite morphogenesis and neuronal lineages.

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A key technology in Drosophila and Caenorhabditis elegans is the forward genetic screen. Forward genetics involves searching for genetic pathways that are important for a particular biological property by examining genomic disruptions that cause a particular phenotype of interest. For example, interest in identifying genes that control dendrite morphology led to a screen for disruptions in dendrite branching complexity, targeting, or growth (Gao et al., 1999). Different strains were generated that carried random genetic lesions, and these were screened usually by microscopic analysis. This classical unbiased genetic screening approach is extremely powerful, and it is in use today. Once a lesion is identified as disruptive for the trait of interest, the lesion is mapped to the gene if possible, and the gene is then studied in great detail using more refined manipulations of individual neurons, some of which include the GAL4eUAS and MARCM approaches described above.

11.3 Anatomical background for key model systems in which dendritic morphogenesis is studied in invertebrates 11.3.1 Drosophila dendritic arborization sensory neurons Sensory neurons in the Drosophila embryo and larva cover the body wall and subserve different sensory functions. One neuronal subtype, called the dendritic arborization (da) neuron, spreads complex dendrites to cover the body wall (Fig. 11.1A and B). da neurons have polarized neurites, with axons and dendrites leaving from opposite sides of the soma. There are several distinct morphological, and likely functional, classes of da neurons, and they are referred to as class I, II, III, or IV neurons; each class has an increasing dendritic arbor complexity (Grueber et al., 2002). The dendrites of individual da neurons do not overlap since they project so as to maximally cover their entire receptive field; to do so they display self-avoidance (Grueber et al., 2002; Sweeney et al., 2002). In addition, two classes of body wall sensory neurons, the class III and class IV neurons, provide a complete and nonoverlapping tiling of the body wall, much like the way in which tiles cover a floor completely without any tiles overlapping with one another (Grueber et al., 2002). The da neurons are sandwiched between epidermal cells and muscles. Because larval epidermal cells and the cuticle above them are transparent, the cell bodies, dendrites, and proximal axons of da neurons can be imaged in living larvae using techniques that include confocal microscopy. This accessibility makes da neurons ideal for cell biological analysis in vivo (Stone et al., 2008; Ori-McKenney et al., 2012; Ye et al., 2007). The da neurons have been used to dissect several different molecular aspects of dendrite formation, including mechanisms of dendritic diversification, self-avoidance and tiling, pruning, and the cellular basis of dendritic growth.

11.3.2 Drosophila motoneurons Insect motor neurons have cell bodies that are situated in an outer cortex of the central nervous system (CNS), and they send dendrites into a central neuropil where they receive inputs (Fig. 11.1C). The basic organization of Drosophila motoneuron dendrites is that of a myotopic map in which the position of dendrites in the central neuropil reflects the position of muscle targets for axons in the periphery (Landgraf et al., 2003). By larval stages each motor neuron dendrite projects to a particular domain along the mediolateral axis of the CNS. Unlike sensory neuron dendrites, these domains can overlap regardless of cell type (Kim et al., 2009). It is possible that overlap rules are relaxed and that input specificity relies more on the selection of appropriate synaptic partners rather than delimitation of inputs by dendritic territorial boundaries. The motor system has been used to identify the roles of midline guidance cues in dendrite guidance (Brierley et al., 2009; Mauss et al., 2009). There is also evidence that motor neuron growth is subject to regulation by neuronal activity (Duch et al., 2008; Hartwig et al., 2008; Tripodi et al., 2008).

11.3.3 Drosophila olfactory projection neurons The Drosophila olfactory system is fundamental for detection of food, mates, and warning signals from the environment. It is a remarkable example of precise neuronal wiring between neurons that detect olfactory information and the neurons that relay this information to the brain for processing and signal integration (Fig. 11.1D). The olfactory system comprises w400 olfactory receptor neurons (ORNs) responsible for detecting a large variety of odorant cues. ORNs project axons to the primary olfactory center, called the antennal lobe (AL). At the AL, ORNs synapse with second-order neurons, the olfactory projection neurons (PNs). The cell bodies of PNs surround the AL, and each projects dendrites to one of approximately 60 glomeruli in the AL; this is where synapses are formed with ORNs. PNs, in turn, extend axons to higher-order brain centers for further olfactory processing. The projections of PN dendrites to individual glomeruli in the AL are highly stereotyped and present a powerful system to dissect the mechanisms of dendritic targeting by transcription factors and guidance cues (Jefferis et al., 2001).

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Olfactory receptor neurons (ORNs) FIGURE 11.1 Representative systems in which dendritic morphogenesis is studied in invertebrates. (A) Confocal image of a dendritic arborization sensory neuron in Drosophila. (B) Schematic of dendritic arbor (red) and axonal projection (blue) of da neuron used in this chapter. (C) Motoneuron dendrites (red) in the central nervous system, with axon (blue) shown projecting to the periphery. (D) Antennal lobe projection neurons (PNs). Three lineages of PNs surround the antennal lobe. See text for details of nomenclature. PNs project dendrites (red) to specific glomeruli within the antennal lobe where they receive input from olfactory receptor neurons. PN axons (blue) project to higher brain centers including the mushroom body and lateral horn (not shown). (E) Caenorhabditis elegans PVD neurons elaborate an extensive dendritic arbor along the body wall. Major branches resolve into higherorder assemblages of dendrites that are repeated along the length of the animal. Part (E) reproduced from Shrestha, Grueber, 2010. Neuronal morphogenesis: worms get an EFF in dendritic arborization. Curr. Biol. 20, R673eR675; with permission.

11.3.4 Caenorhabditis elegans PVD neurons The PVD neurons in C. elegans provide another important model for studying dendrite morphogenesis (Albeg et al., 2011; Dong et al., 2013; Oren-Suissa et al., 2010; Salzberg et al., 2013). These two bilateral polymodal sensory neurons have a remarkably complex dendritic arbor, yet they show a highly ordered arrangement of primary, secondary, and higher-order

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branches (Fig. 11.1E). Branches sprout from parent branches at predominantly right angles all along the length of the animal body wall, resulting in a complete tiling of the skin surface with sensory fibers. This regular arrangement, combined with the powerful genetic approaches available in worms, has led to insights into the mechanisms that underlie dendritic branching, targeting, self-avoidance, and tiling.

11.4 Cell biology of dendritic growth Dendrites have a vast surface area much greater than a typical animal cell. How this large investment in plasma membrane growth is supported is a fascinating and unresolved problem. In addition, axons and dendrites usually differ significantly, even in the same neuron, in targeting, growth, and elaboration, raising the question of how differential investments are made in dendritic versus axonal morphogenesis. The answers to several of these issues appear to reside in differential trafficking of cargos to dendrites versus axons, and this in turn relies on the organization of microtubules along dendrites.

11.4.1 Microtubule polarity differs between dendrites and axons The growth of a dendritic arbor is intimately tied to the polarity of dendritic microtubules, as well as the differences between microtubules in axons and dendritic arbors. As such it is useful to review the basic organization of the cell’s cytoskeleton, particularly, as it pertains to neuronal compartmentalization. In the soma of a developing neuron lies a microtubule-organizing center, termed the centrosome and consisting of two centrioles and surrounding pericentriolar material. Microtubules have an intrinsic polarity of plus-ends and minus-ends, with tubulin subunits being added to growing microtubules at the plus end. Microtubule minus-ends are anchored at the centrosome and extend out into neuronal compartments. Early studies made by Bass and colleagues showed that the first process to grow from cultured hippocampal neurons contains uniformly plus-end distal microtubules (Baas, 1999; Baas et al., 1989). This process rapidly grows into the axon, whereas later processes to emerge differentiate into dendrites and harbor both plus-end and minus-end distal microtubules and thus have a mixed microtubule arrangement. Although these studies were performed in vertebrate neurons, invertebrate neurons follow a similar (but not identical) set of rules. Insect axons, like their vertebrate counterparts, contain microtubules that are exclusively plus-end distal (Stone et al., 2008). Dendrites contain minus-end distal microtubules as well in Drosophila, but in contrast to vertebrates have only a very small representation of plus-end distal microtubules (Stone et al., 2008). Thus, a shared feature of vertebrate and invertebrate dendrites is the presence of minusend distal microtubules. Paradoxically, the establishment of microtubule polarity in neurons requires microtubule-associated molecular motors. In Drosophila da neurons, loss of dynein genes leads to the appearance of minus-end distal microtubules in axons (Zheng et al., 2008; Satoh et al., 2008). In C. elegans DA9 and PHC sensory neurons, kinesin-1 is required for the presence of minus-end distal microtubule in dendrites (Yan et al., 2013). In mutants with defective kinesin-1, microtubules in dendrites adopt an axon-like plus-end distal orientation. It has been proposed that the molecular motors transport plus-end microtubules out of dendrites and minus-end microtubules out of the axon, leading to the microtubule polarity observed in normal dendrites and axons.

11.4.2 Dynein-dependent trafficking controls dendritic branching Microtubule polarity influences the trafficking of proteins and organelles throughout the cell. Microtubule cargos are trafficked by microtubule motors, and different types of motors move with either minus-end or plus-end directed polarity. The kinesin family of proteins includes the major plus-end motors, whereas dyneins carry cargos in a minus-end direction. Dynein, a very large protein complex, is the major microtubule motor for dendrites. The dynein complex consists of core components: dynein heavy chain, dynein light chain, light intermediate chain (Dlic), and light chain. Both dynein and dynein cargos are important for the spatial distribution of branches along a dendritic arbor (Zheng et al., 2008; Satoh et al., 2008). Dlic was identified in mutagenesis screens for genes affecting dendritic morphogenesis. These studies of Dlic focused on a group of neurons, the da neurons that have branches concentrated near the margins of the arbor but are sparse in the center of the receptive field near the cell body (Fig. 11.2A). Neurons lacking Dlic show an aberrant shift of dendritic branches from distal regions of the arbor to proximal regions, suggesting that trafficking of branching machinery is normally required to place branches in specific locations along an arbor and is disrupted in neurons lacking Dlic (Fig. 11.2B). The dynein complex is essential for the distribution of dendritic Golgi outposts, which regulate dendritic branching (Ye et al., 2007; Zheng et al., 2008) (see details below). In addition, disruption of the Rab5 protein, a small GTPase of the Rab family and a regulator of the early endocytic pathway, in Dlic mutant neurons blocks the proximal

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FIGURE 11.2 Spatial control of branching by dynein trafficking. (A) Wild-type class IV arbor shows branching most concentrated near the margin of the arbor (arrow) and more sparse branching nearer to the cell body (arrowhead). (B) In mutants of Dynein light intermediate chain 2 (Dlic2), the arborization becomes concentrated near the cell body, with distal dendrites showing poor elaboration. (C) Expression of a dominant negative (DN) Rab5 construct in Dlic2 neurons suppresses the proximal branching phenotype. Data are summarized from Satoh, D., Sato, D., Tsuyama, T., Saito, M., Ohkura, H., Rolls, M. M., Ishikawa, F., Uemura, T., 2008. Spatial control of branching within dendritic arbors by dynein-dependent transport of Rab5-endosomes. Nat. Cell Biol. 10, 1164e1171; Zheng, Y., Wildonger, J., Ye, B., Zhang, Y., Kita, A., Younger, S.H., Zimmerman, S., Jan, L.Y., Jan, Y.N., 2008. Dynein is required for polarized dendritic transport and uniform microtubule orientation in axons. Nat. Cell Biol. 10, 1172e1180.

hyperbranching Dlic phenotype without restoring branching at the margins of the arbor (Fig. 11.2C). Thus, Golgi outposts and Rab5 are components of the trafficking machinery required for dendritic arbor elaboration (Zheng et al., 2008; Satoh et al., 2008).

11.4.3 Role of the secretory pathway and Golgi outposts in dendritic elaboration The secretory pathway that delivers membrane to the cell surface consists of the endoplasmic reticulum (ER), and Golgi apparatus, and, in neurons, Golgi outposts that become prominent during periods of rapid dendritic growth. Golgi outposts are a conserved component of the dendrite branching machinery in vertebrates and invertebrates. The discovery of isolated Golgi outposts in mammalian neurons (Gardiol et al., 1999; Horton and Ehlers, 2003; Horton et al., 2005) provides context for genetic studies carried out in Drosophila. Somatic Golgi were found concentrated at the base of the largest branch in vertebrate hippocampal and cortical pyramidal neurons, the apical dendrite, and that isolated Golgi resided out along the arbor, primarily at branch points. Pharmacological disruption of forward Golgi trafficking in neurons using brefeldin A in dissociated cell culture resulted in decreased dendrite growth. Dispersion of the Golgi into multiple dendrites caused elaboration of a nonpolarized arbor with all branches of fairly equal length and branching complexity. These experiments reveal how polarized trafficking of membrane components can lead to specific patterning features of dendritic arbors. Forward genetics identified several components of the secretory pathway as being important for normal dendritic growth in invertebrate neurons (Ye et al., 2007). Class IV da neurons were screened for molecules that regulate dendritic growth, and among those identified were several proteins that fit into a common forward secretory pathway: Sec23, Sar1, and Rab1 (Fig. 11.3). Interestingly, in these mutant lines, axon growth was disrupted to a lesser extent than dendrite growth, suggesting a differential reliance of dendrite versus axon growth on ER-to-Golgi transport (Fig. 11.3). Consistently, pathogenic mutations in the Golgi SNARE protein Membrin, which causes a severe neurological disorder in humans, partially reduce SNARE-mediated membrane fusion and preferentially impair dendrite growth in Drosophila da neurons (Praschberger et al., 2017). These studies demonstrate that dendritic growth is particularly sensitive to deficits in the secretory pathway. Golgi outposts are likewise observed in the da neurons, and da dendrite morphology depends on intact Sar1 (Ye et al., 2007). Laser-induced damage of Golgi outposts halts dendrite branch dynamics, supporting a role for these organelles in arbor dynamic extension. Golgi outposts nucleate microtubules in dendrites and contribute to dendrite branch growth and stability (Ori-McKenney et al., 2012; Zhou et al., 2014; Yalgin et al., 2015). Whether or not dendritic branches grow depends on the direction of outpost-nucleated microtubule growth. The protein centrosomin localizes to cis-Golgi and recruits microtubule nucleation activity to dendritic Golgi outposts for retrograde microtubule polymerization away from nascent dendrite branches, leading to the simple dendritic arbors in class I da neurons (Yalgin et al., 2015). Centrosomin is

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Axon FIGURE 11.3 A forward genetic screen identifies components of forward secretory trafficking in dendritic growth. To the left are confocal images of wild-type class IV dendrites (upper panel) and axons (lower panel) marked with green fluorescent protein (GFP) driven by an enhancer of the pickpocket gene, which is specifically expressed in class IV neurons. Other panels indicate dendrite and axon phenotypes in Sec23, Sar1, and Rab1 mutants. The phenotypes of these lines are very similar suggesting that they operate in a common genetic pathway for dendritic elaboration. Reproduced from Ye, B., Zhang, Y., Song, W., Younger, S.H., Jan, L,Y., Jan, Y.N., 2007. Growing dendrites and axons differ in their reliance on the secretory pathway. Cell 130, 717e729; with permission.

a major transcriptional target of the transcription factor Abrupt, which is only expressed in class I neurons among the four classes of da neurons. Thus, it seems that microtubule nucleation activity associated with Golgi outposts is used by cell typeespecific regulators to differentiate the morphologies of different neuronal types. Evidence from studies in Drosophila da neurons suggests that dendritic Golgi outposts differ from Golgi in the cell body in their compartmental organization. While the cell bodies of da neurons contain Golgi consisting of stacks of cis, medial, and trans compartments, these compartments are often disconnected in da neuron dendrites, likely due to the lack of the Golgi structural protein GM130 in dendrites (Zhou et al., 2014). Golgi compartments in dendrites exhibit dynamic movements, and this may determine the degree of stacked Golgi outposts and, consequently, has the potential to regulate acentrosomal microtubule growth. Studies of C. elegans PVD sensory neurons suggest that not only membrane deposition but also membrane shaping by the fusogen protein called EFF-1 (epithelial fusion failure-1) is critical for dendritic branching and morphogenesis (OrenSuissa et al., 2010). Fusogens are important for cell fusion events during development and control fusion by altering membrane curvature. The level of EFF-1 sets the proper branching complexity of PVD neurons, with higher levels suppressing branching and lower levels leading to more branching and disorganization of the normal near 90 degrees branching angles seen in PVD arbors (Fig. 11.1E).

11.5 Transcriptional control of dendritic morphology 11.5.1 Control of dendrite morphological identity of Drosophila PNS neurons Dendritic arbors have remarkably diverse morphologies, but within a cell type, dendritic morphology is consistent to the extent that characteristic dendrite shapes are often used to classify different functional groups of neurons. Neuronal typespecific morphogenesis is dictated by intrinsic mechanisms of transcriptional control (Corty et al., 2009; Grueber and Jan, 2004; Jan and Jan, 2010). The peripheral nervous system (PNS) of Drosophila embryos and larvae has been a useful model for identifying several of these mechanisms. PNS neurons are classified into several different morphological and functional types: among them are external sensory neurons and chordotonal organs, which have single unipolar ciliated dendrites, and the multidendritic (md) neurons with complex, multipolar dendrites. The md neurons are further distinguished based on their complexity and the substrate upon which they arborize. The md-da neurons extend dendrites along the epidermis, mdtracheal dendrite (td) neurons project along tracheal respiratory organs, and the md-bipolar dendrite (bd) neurons extend along connective strands typically along the anteroposterior axis of the animal (Bodmer and Jan, 1987). Most work that addresses mechanisms of dendrite development has been performed on the da subtype. This cell type is further classified into four different morphological classes that are distinguished by the complexity of their dendritic arbors (Grueber et al., 2002). Class I neurons have simple dendrites, class II neurons show additional branching, class III neurons have extensive short extensions, and class IV neurons have complex space-filling arbors (Fig. 11.4). This system provides a useful model for understanding genetic programs that specify diversity of dendritic architecture. The first fundamental choice that neurons make is whether to extend a single dendrite or multiple dendrites. Mono- and multipolar neurons are related by lineage in Drosophila, and the choice as to whether a cell will generate a single dendrite versus

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Gene expression key Abrupt Cut Spineless Knot FIGURE 11.4 da neuron morphology and differential transcription factor expression in fly peripheral nervous system. (A) Organizational chart of Drosophila sensory neurons for developmental studies, including the da neurons. (BeE) Characteristic dendritic morphologies of class IeIV da neurons. For each tracing, a pie chart showing known transcription factor expression patterns for the transcription factors Abrupt (red), Cut (shades of blue depending on the level at which it observed in the class), Spineless (yellow), and Knot (green). These four transcription factors are known to mediate classspecific dendritic development (see text for details). Part (A) and tracings reproduced from Grueber, W.B., Jan, L.Y., Jan, Y.N., 2003a. Different levels of the homeodomain protein cut regulate distinct dendrite branching patterns of Drosophila multidendritic neurons. Cell 112, 805e818; with permission.

many is controlled by the zinc-finger protein Hamlet (Ham) and the Krüppel-like transcription factor Dendritic arbor reduction 1 (Dar1) (Moore et al., 2002; Wang et al., 2015). Ham is expressed transiently in the precursor cells and neurons that generate a monopolar dendritic arbor. Forced expression of Ham in a postmitotic md neuron will reduce branching. Conversely, loss of Ham from the external sensory (es) neuron lineage will generate additional md neurons (the extra md neuron phenotype is actually how mutations in the ham gene were originally identified). While Ham appears to act in precursors to specify cell identity and transiently in postmitotic neurons to specify the bipolar morphology of neurons, Dar1 postmitotically defines the multipolar morphology of da neurons (Wang et al., 2015). Dar1 is expressed in postmitotic multipolar, but not bipolar or unipolar, neurons. Loss of dar1 gradually converts multipolar neurons into the bipolar or unipolar morphology without changing neuronal identity, while misexpression of Dar1 or its mammalian homolog in unipolar and bipolar neurons causes them to assume multipolar morphologies. These findings also raise the possibility that similar transcriptional control is involved in the formation of multipolar dendritic morphologies, which is yet to be determined. Once a cell is fated for a multipolar morphology, several other transcriptional programs take over responsibility that specify different class-specific morphologies. The transcription factor Cut, which also acts to specify sensory organ identity of bristle neurons (Bodmer and Jan, 1987), was found to be expressed in different levels in different morphological classes of da neurons (Grueber et al., 2003a). Cut is absent in class I neurons, expressed at low levels in class II neurons, intermediate levels in class IV neurons, and highest levels in class III neurons. Overexpression of Cut in neurons that normally express low levels leads to branching characteristic of high-level expressing neurons, whereas loss of Cut leads to dendritic simplification characteristic of nonexpressing neurons. The particular level of Cut expressed by different neurons appears important, since for those neurons that express low levels, up or down modulation of Cut levels leads to bidirectional shifts in dendritic complexity. Class I da neurons do not express Cut, but express Abrupt, a zinc-finger

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transcription factor of the Broad, Tramtrack, Bric-a-brac (BTB) family (Li et al., 2004; Sugimura et al., 2004). Loss of Abrupt from these neurons leads to extra dendritic branches, whereas overexpression of Abrupt leads to fewer dendrites being formed by neurons that are normally highly complex. It is interesting to note that the cell typeespecific roles of Cut and Abrupt in the dendrite development of Drosophila neurons are paralleled by a similar mechanism in mammalian cerebral cortex, at least at the conceptual level. The mammalian homologs of Cut, Cux-1, and Cux-2, which are specifically expressed in the upper cortical layers (II/III and IV) (Nieto et al., 2004), are required for dendritic branching in the pyramidal neurons in those layers (Cubelos et al., 2010). By contrast, the zinc-finger transcription factor Zfp312, which is specifically expressed in the deep cortical layers (V and VI), is required for dendritic development of deep-layer, but not that of upper-layer, pyramidal neurons (Chen et al., 2005). Another important transcription factor for dendritic diversity is the helix-loop-helix (HLH) transcription factor (TF) Knot/Collier. Among da neurons, Knot is expressed only in class IV neurons and expression in other classes is sufficient to convert them to a class IVelike morphology (Crozatier and Vincent, 2008; Hattori et al., 2007b; Jinushi-Nakao et al., 2007). Knot does not act by itself to generate class IV arbors but, rather, exhibits a common mode of regulation by transcription factors in general, that of combinatorial action. In the case of dendrite morphology, Knot acts in combination with Cut to specify class IV neuron morphology such that both Cut and Knot are required together to implement the transcriptional program that helps to generate class IV branching patterns. Other TFs likely act together with the above TFs, including the aryl hydrocarbon receptor family member TF Spineless (Kim et al., 2006). Spineless is expressed in all classes of da neurons, and loss of Spineless function in different neurons leads to convergence of the various neuronal classes of both simple and complex morphologies to an intermediate dendritic complexity. Thus, Spineless activity may be necessary for the activity of other transcriptional programs during dendritic diversification, without which neurons fail to elaborate into diverse dendritic arbor shapes. Transcription factors can presumably promote or inhibit growth and branching by promoting or repressing the expression of multiple possible target genes; however, these are largely unknown. One prediction is that transcriptional targets must exist that mediate the effect of TFs on the dendritic cytoskeleton. One known example is the spastin gene, which was identified as a target of Knot in the highly complex class IV neurons (Jinushi-Nakao et al., 2007). Spastin encodes a microtubule-severing protein implicated in the disease hereditary spastic paraplegia. Knot promotes the expression of Spastin in class IV neurons, which is proposed to generate microtubule breaks that can seed the construction of new dendritic branches critical for forming complex dendritic arbor morphologies.

11.5.2 Transcriptional control of dendritic targeting of olfactory PNs Studies of dendritic development of olfactory PNs illustrate the principle of combinatorial control of dendritic morphogenesis by suites of transcription factors. Rather than acting as single dedicated switches that control the fates of cells in a binary fashion, many transcription factors show context-dependent activity. For example, the output of 1 TF can depend on whether another TF is present or absent in the same cell, and this TF activity can in turn depend on the presence of yet another TF. Such combinatorial transcriptional codes vastly increase the possible developmental outputs that might be controlled by any single TF. For dendrite morphogenesis, codes have been described most extensively among olfactory PNs, where they are important for ensuring precise dendritic targeting to specific AL glomeruli (Komiyama and Luo, 2007; Komiyama et al., 2003, 2004). The major outputs of the code are both global positioning and specific targeting of dendritic arbors in the AL. Global positioning refers to the position of dendrites along the major axes of the AL: anteroposterior, dorsoventral, and mediolateral (Fig. 11.1D). Although different AL glomeruli reside in these broad regions, global position cues act without regard for the specific glomerulus, and they are responsible for getting dendrites to the proper region of the AL. By contrast, specific targeting refers to targeting of dendrites to specific glomeruli within these global positions. Some transcription factors specify only global dendritic position along the major AL axes, including Cut, which controls position along the mediolateral axis (Komiyama and Luo, 2007). Cut gain-of-function shifts dendrites medially, and loss of cut shifts dendrites laterally in the AL. Cut may regulate responsiveness to attractive or repulsive cues that are distributed in a spatial pattern in the AL. Numerous transcription factors have also been identified in the control of specific glomerular targeting, acting within the context of the TFs that specify global position (Corty et al., 2009; Komiyama and Luo, 2007). A good example of the interplay between TFs that act at these different levels of specificity is given by the innervation of the DL1 glomerulus by the DL1 PN (Fig. 11.5). To understand the code used by DL1, we have to consider in more detail the basic lineage relationships among AL PNs. There are about 150e200 PNs in adult Drosophila born from three major lineages, the anterodorsal (ad) lineage, the lateral (L) lineage, and the ventral (v) lineage. These lineages give rise to clusters of PNs called adPNs, lPNs, and vPNs, respectively, whose cell bodies surround the AL (Fig. 11.1D). The DL1 neuron is part of

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FIGURE 11.5 Combinatorial transcriptional control of projection neuron dendritic targeting. Shown are schematics of the antennal lobe with the DL1 projection neuron and glomerulus highlighted. (A) Wild-type DL1 projects exclusively to the DL1 glomerulus. (B) In acj6 mutant cells, DL1 is targeted, but dendrites spill out into areas outside of the glomerulus. (C) Drifter misexpression in an acj6-mutant cell shifts the dendritic arbor out of the DL1 gomerulus and causes innervation of glomeruli that are normally targeted by Drifterþ projection neurons. (D) If Cut is misexpressed in acj6-mutant DL1 PNs, the effect is very different: the dendrites are shifted medially, but still innervate glomeruli that are targeted by neurons in the adPN lineage. (E) When both Drifter and Cut are misexpressed in acj6-mutant cells, the dendrites are both shifted medially, and the specificity of targeting is shifted to glomeruli that are innervated by the lPN lineage. Data are summarized from Komiyama, T., Sweeney, L.B., Schuldiner, O., Garcia, K.C., Luo, L., 2007. Graded expression of semaphorin-1a cell-autonomously directs dendritic targeting of olfactory projection neurons. Cell 128, 399e410.

the adPN lineage, and all neurons of this lineage express the POU domain TF Abnormal chemosensory jump 6 (Acj6). By contrast, neurons in the lPN lineage, but not those of the adPN lineage, express another POU domain TF called Drifter (Komiyama et al., 2003). Thus, these two TFs are expressed in a complementary and mutually exclusive group of neurons. Both TFs control lineage-specific targeting such that loss of one and coincident misexpression of the other in specific lineages leads to a switch in characteristic dendritic targeting patterns. However, these phenotypes are modified by the expression status of the global positional signal provided by Cut (Fig. 11.5) (Komiyama and Luo, 2007). Loss of acj6 from

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DL1 causes dendrites to spill out of their normal target, but the specificity of dendrite projection to the DL1 glomerulus is not strongly affected (Fig. 11.5B). By contrast, if acj6 is lost and different instructional cues are given by misexpression of Drifter, the specificity of targeting is switched and the neurons innervate a completely different glomerulus (Fig. 11.5C). The role of Cut in dendritic targeting depends on the other transcription factors present in DL1. Cut misexpression in acj6neurons causes dendrites to miss DL1 and instead project to much more medial target glomeruli (Fig. 11.5D). However, all glomeruli targeted are still those innervated by other adPN lineage neurons (the same lineage as the DL1 PN). If both Cut and Drifter are misexpressed in acj6 DL1 neurons, dendrites spill out of the DL1 glomerulus to innervate the glomeruli that are normally innervated by neurons of the Drifter-expressing lPN lineage (Fig. 11.5E). This set of genetic manipulations provides a clear example of how the particular morphogenetic program of individual neurons can depend on the combinatorial expression and activity of specific transcription factors.

11.5.3 Chromatin remodeling factors and dendritic development Gene transcription is not only a consequence of the presence or absence of particular transcription factors. Rather, gene expression is subject to modification by epigenetic mechanisms that regulate the accessibility of DNA to these factors. One mechanism of epigenetic modification is through ATP-dependent chromatin remodeling by neuronal-specific Brahmarelated gene/Brahma-associated factor complexes (nBAFs). Multisubunit nBAF complexes modify DNAehistone interactions to regulate the accessibility of DNA. An RNAi-based screen in Drosophila identified several BAF subunits as being important regulators of dendritic morphogenesis (Parrish et al., 2006). In mammalian neurons, an elegant mechanism has been uncovered for control of dendritic development by nBAFs. Undifferentiated neuronal progenitors require the inclusion of the BAF53a subunit for proper progenitor proliferation. In postmitotic neurons, BAF53a is replaced by a homologous subunit, BAF53b, and this switch is required for proper neuronal differentiation and dendrite development of hippocampal, cerebellar, and cortical neurons (Wu et al., 2007). One possibility is that neuron-specific epigenetic mechanisms for controlling gene expression may be particularly important in neurons that must maintain a basic dendritic architecture over days, weeks, or years, and simultaneously maintain some degree of plasticity (Wu et al., 2007). Chromatin structure is also modified by acetylation and deacetylation of histone lysine residues. Deacetylation of histones by histone deacetylases (HDACs) correlates with chromatin compaction and gene repression. Olfactory PN targeting and axon targeting are regulated by a specific HDAC, Rpd3, a close homolog of the mammalian genes HDAC1 and HDAC2 (Tea et al., 2010). In addition, dendrite branching of da sensory neurons is increased upon Rpd3 RNAi knockdown, so the role of HDACs in dendrite arborization may be widespread (Parrish et al., 2006). It is conceivable, but not yet shown, that HDACs may act to modulate expression or activity of some of the transcription factors described earlier in this section. Histone modifications by Polycomb group (PcG) genes are another avenue for silencing gene expression. Two repressor complexes, Polycomb repressor complexes 1 and 2 (PRC1 and PRC2) act via distinct mechanisms to modify chromatin accessibility, and components of both complexes are important for dendritic morphogenesis. Mutations in PRC1 and PRC2 components lead to decreased da resulting from a failure of dendritic maintenance (Parrish et al., 2007).

11.6 Posttranscriptional control of dendritic development Posttranscriptional control of gene activity is widely used throughout development as a mechanism to finely control protein expression and localization. These mechanisms include control of mRNA translation and regulation by microRNAs (miRNAs). In some systems, such as in the olfactory system where the miRNA processing pathway is important for dendritic targeting, and in da neurons discussed below, posttranscriptional control of gene expression plays important roles in dendritic development.

11.6.1 Control of mRNA translation in dendritic development Translational control of gene expression plays an important role in dendritic development in invertebrates. A conserved translational repressor complex consists of the RNA-binding proteins Nanos (Nos) and Pumilio (Pum). Nos and Pum were first studied for their role in the localization of mRNAs in early fly embryos. In the early embryo, the posterior localization of Nos represses specific mRNAs, such as the anterior determinant hunchback. Nos and Pum are both expressed in da neurons, and overexpression of either one leads to reduced dendritic branching. As in the early embryo, Pum and Nos work together during the elaboration of more complex class III and IV dendrites in da sensory neurons. Knockout of either one leads to elongated F-actin-enriched dendritic filopodia unique to class III da neurons and simplification of class IV dendrites (Ye et al., 2004). These complex phenotypes coincide with those observed in hippocampal neurons in mammals

242 PART | I Formation of axons and dendrites

(Vessey et al., 2010). In immature neurons, loss of Pum2 (a mammalian homolog of Pum) increases dendrite growth, but in mature neurons it reduces dendritic spines yet promotes the extension of dendritic filopodia. Like in flies, overexpressing Pum2 reduces dendritic branching in rat hippocampal neurons. In the early embryo, the localization of nos mRNA is not perfectly restricted, and other mechanisms exist to repress incorrectly localized mRNA. Nos is subject to translational repression via a sequence in the 30 untranslated region (UTR) of nos mRNA called the translational control element (TCE). The TCE is important for proper localization of Nos within cells, and thus spatial regulation of mRNAs by PumeNos. Overexpression of nos without a native 30 UTR results in more severe defects in dendritic branching than overexpression of nos with the native 30 UTR, suggesting that nos translation in neurons is repressed by other factors (Ye et al., 2004). This repression depends on the RNA-binding proteins Glorund and Smaug. Glorund and Smaug mutants also show dendrite defects, so these proteins might either restrict mRNA translation during transport out to the dendrites or may restrict mislocalized mRNA from being translated (Brechbiel and Gavis, 2008). Either way, these results indicate that localized translation of nos is critical for dendritic morphogenesis. Since Nos is a translational repressor, the next questions to address are the identity and functions of the mRNAs whose translation is directly regulated by Nos.

11.6.2 miRNAs in dendritic development A critical pathway for posttranscriptional regulation of gene expression is via miRNA-mediated translational repression. Mature miRNAs are generated by successive enzymatic steps involving the generation of pre-miRNAs by Drosha, an RNase III, and Pasha, a double-stranded RNA (dsRNA)-binding protein. Dicer, another RNase III, then cleaves the premiRNAs to generate mature miRNAs. Roles for miRNAs in dendritic development are, at present, poorly understood. However, a few results suggest important roles for this type of gene regulation. Both pasha and dicer were identified in a forward genetic screen for mutations that affect olfactory PN dendritic targeting (Berdnik et al., 2008). Specific subsets of PNs showed cell autonomous defects in both dendritic targeting and axon morphology in higher brain centers upon knockdown of either of these two components. Dendrites of all PNs are not equally affected, but those that are show poor innervation of their normal glomerulus as well as ectopic innervation of additional glomeruli. These results implicate the miRNA processing pathway in dendritic targeting, but they leave open the identity of specific miRNA(s) that mediate PN targeting. Additionally, in Drosophila da neurons increased levels of a pre-miRNA, pre-miR124a, strongly suppress dendritic branching (Xu et al., 2008). The normal role for miR124 in dendritic development, and whether this could be one of the miRNAs responsible for the dicer and pasha mutant phenotypes in PNs, is not clear. Finally, as will be discussed below, the bantam miRNA plays a nonautonomous role in the control of dendritic elaboration in Drosophila sensory neurons (Parrish et al., 2009).

11.7 Control of dendritic field formation I: guidance and targeting Dendritic arbors are often represented as a collection of processes that extend more or less equally around a central soma. While this is a fair depiction of certain types of arbors with so-called radial territories, many dendrites actually show highly polarized outgrowth and branching. Some prominent examples from vertebrates include cerebellar Purkinje cells, which show a highly complex arbor that extends unidirectionally from the cell body; pyramidal cells in the cortex and hippocampus, which show a long apical dendrite and shorter more numerous basal dendrites; and certain classes of direction selective retinal ganglion cells in the vertebrate retina. As discussed above, polarized membrane transport, for example, in pyramidal neurons, is important for the establishment of initial polarized dendritic growth. In addition, it is now well demonstrated that dendrites, like axons, respond to specific guidance signals in their environment that elicit directional growth toward specific targets. These findings have important implications for how neural circuits are assembled because they show that both dendrites and axons are active players in wiring of the nervous system, and that though their growth and targeting progresses independently it must be coordinated in order for proper patterns of connectivity to emerge.

11.7.1 Slit and netrin signaling during midline dendritic guidance Proper dendritic targeting generally depends on instructive signals from the extracellular environment and reception of these signals by dedicated receptors (Corty et al., 2009; Furrer et al., 2007). The major molecules that control axon guidance in both vertebrate and invertebrate nervous systems, including Slit and Robo, Netrin and Frazzled, and Semaphorins and Plexins, have all been implicated in dendritic targeting in flies, but these molecules have complex and cellspecific roles. Motoneuron dendrites can choose to avoid crossing the midline and arborize in ipsilateral neuropil, or cross

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FIGURE 11.6 Control of dendrite targeting by midline signaling systems. (A) The RP3, aCC, and RP2 neurons show different patterns of midline crossing by dendrites and axons. Axons are indicated by an “a” for each of the neurons. Both RP3 and aCC normally have contralateral and ipsilateral dendritic arbors, whereas RP2 is restricted to ipsilateral neuropil. (B) In netrin or frazzled mutants, neither RP3 nor aCC project across the midline and dendrites elaborate only in ipsilateral neuropil. (C) In robo mutants, RP3 dendrites elaborate at the midline and RP2 dendrites project across the midline to contralateral neuropil. (DeF) Studies of the aCC neuron have additionally showed that both slit and robo are required for elaboration of dendrites. Dendrites grow in the zone of Slit expression (yellow) and are absent when animals are made mutant for either Slit (E) or the Robo receptor (F). Data redrawn from Furrer, M.P., Kim, S., Wolf, B., Chiba, A., 2003. Robo and Frazzled/DCC mediate dendritic guidance at the CNS midline. Nat. Neurosci. 6, 223e230; Furrer, M.P., Vasenkova, I., Kamiyama, D., Rosado, Y., Chiba, A., 2007. Slit and Robo control the development of dendrites in Drosophila CNS. Development 134, 3795e3804.

the midline and arborize in contralateral neuropil. In addition, dendrites select particular mediolateral positions within the neuropil in which to arborize, and do so in a cell typeespecific manner. These different decisions and their molecular control will be considered below to exemplify several emerging themes in dendritic guidance. Different neurons make cell-specific decisions with regard to dendritic navigation at the midline, and remarkably within the same cell axons and dendrites can navigate differently with respect to the midline. This diversity is exemplified by findings on the RP2, RP3, and anterior corner cell (aCC) motoneurons (Furrer et al., 2003, 2007) (Fig. 11.6A). The aCC neuron elaborates dendritic trees in ipsilateral and contralateral neuropil and sends an axon ipsilaterally to innervate a peripheral muscle (Fig. 11.6A). In the absence of Netrin and Frazzled, aCC extends only ipsilateral dendrites (the ipsilateral axon is unchanged) because those dendrites that normally cross the midline do not receive appropriate crossing cues (Fig. 11.6B) (Furrer et al., 2003). The RP3 neuron likewise elaborates contralateral and ipsilateral dendrites, but it extends an axon contralaterally to innervate its peripheral muscle. Without Netrin or Frazzled, the contralateral projection is likewise lacking (Fig. 11.6B) (Furrer et al., 2003). Thus, Netrin and Frazzled are typically required in dendrites, as they are in axons, for midline crossing. RP3 neurons lacking Robo arborize almost exclusively at the midline, suggesting that midline SliteRobo signaling normally prevents midline dendrite growth (Fig. 11.6C) (Furrer et al., 2003). Similar results have been seen for other types of neurons in addition to RP3 (Fig. 11.6C). Different behaviors by dendrites in response to the same cues could arise in part via differential localization of receptors, or differential timing of receptor expression; however, these possibilities have yet to be rigorously tested. As described earlier most, though not all, insect neurons generate at least some of their dendritic arbor from a proximal neurite that is continuous with the axon. In these neurons the targeting of dendrites depends partly on the positioning of dendrite sprouting along the neurite. The aCC motoneuron is one of the first central neurons to extend dendrites in Drosophila and the expression of Slit prefigures the location of aCC dendritic growth (Fig. 11.6D) (Furrer et al., 2007). In slit mutant embryos that generate no functional protein, aCC neurons do not grow dendrites (Fig. 11.6E). Similar to the case with slit mutants, robo mutants show very little aCC dendritic growth (Fig. 11.6F). Ectopic expression of Slit near other parts of the axon is not sufficient to induce dendritic outgrowth, thus it appears that other cues exist to prime parts of the axon for SliteRobo-induced dendritic growth. Slit and Robo are clearly partners in dendritic targeting of motoneurons,

244 PART | I Formation of axons and dendrites

but they do not act alone. Rather, Netrin- and Frazzled-mediated attraction acts in opposition to Slit- and Robo-mediated repulsion to specify mediolateral positioning of dendrites (Brierley et al., 2009; Mauss et al., 2009). The relative levels of the receptor activity in different neurons operates in a sort of tug-of-war, with high levels of Frazzled and low levels of Robo activity moving dendrites closer to the midline, and higher levels of Robo (and/or lower Frazzled) activity moving dendrites further away.

11.7.2 A combinatorial ligandereceptor complex guides dendritic branches Studies focused on the PVD neurons in C. elegans demonstrate that a multiprotein ligandereceptor complex of adhesion molecules plays an important role in dendriteesubstrate interaction and guides the patterning of dendritic branching (Dong et al., 2013; Salzberg et al., 2013; Zou et al., 2016). PVD dendrites extend between hypodermal cells and internal organs. Hypodermal cells express the ligands SAX7 (a homolog of vertebrate L1-CAM) and MNR-1 (Menorin), and muscles secrete a worm homolog of LECT2 (leukocyte cellederived chemoeaxin-2). PVD neurons express the receptor DMA-1 (dendrite-morphogenesis-abnormal). Biochemical and genetic studies show that these proteins form a multiprotein ligandereceptor complex. Loss of any of these molecules leads to misdirected 2 degrees branches and reduced number of 3 degrees branches. Strikingly, ectopic expression of the ligands in cells that do not normally express them directs PVD dendrites to these cells. These findings suggest that a high local concentration of the ligands SAX/MNR-1/LECT2 activates the DMA-1 receptor, which then regulates cell adhesion between neurites or could activate signaling events that ultimately promote dendritic branching and stabilization.

11.7.3 Coarse and specific control of PN dendritic targeting Members of the Semaphorin family are ligands for the Plexin and Neuropilin families of Semaphorin holoreceptors, but in Drosophila AL development the transmembrane Semaphorin Sema-1a acts in a unique manner. Sema-1a is expressed in a gradient across the AL with high levels in dorsolateral AL and low levels in ventromedial AL (Komiyama et al., 2007). Increasing levels of Sema-1a in neurons leads to shifts in PN dendritic targeting toward high-level AL regions, whereas loss of Sema-1a leads to shifting of dendrites toward lower Sema-1a level regions. These manipulations of Sema-1a levels, performed in neurons rather than in the extracellular environment, indicate that Sema-1a acts cell autonomously as a receptor. Further studies suggest that two secreted Semaphorins, Sema-2a and Sema-2b, are possibly the ligands for Sema1a (Sweeney et al., 2011). These two Semaphorins bind to Sema-1a-expressing cells and form gradients that oppose the Sema-1a gradient. Loss of both Sema-2a and Sema-2b shifts dorsolateral PN dendrites in the AL to ventromedial regions. The function for Sema-1a in dendritic targeting is to specify a coarse map across the AL, but not to direct the specific targeting of dendrites to particular glomeruli. This is accomplished in part by the leucine-rich repeat (LRR) proteins Tartan (Trn) and Capricious (Caps), which specify discrete glomerular targeting for PN dendrites that express these proteins during development (Hong et al., 2009). It is very likely that Trn and Caps make up part of a cell surface code for specific targeting, but do not specify targeting alone since misexpression of Caps in neurons that show no expression directs their dendrites to a reproducible subset of the available glomeruli that are normally innervated by Caps-positive PNs. One possibility is that targeting to these other glomeruli, for which Caps alone is not sufficient, also involves other components of the code that are not normally expressed by the cells in which the Caps misexpression experiments were performed (Hong et al., 2009).

11.7.4 Glial control of dendritic targeting The guidance factors that are presented to dendrites are often produced and secreted by specific glial cells in the nervous system. These glia, therefore, play an important role in the generation of dendritic morphology by instructing dendrites where to arborize (Procko and Shaham, 2010). In some cases, glia support dendrite growth and targeting by providing an adhesive substrate. This mode of development has been observed in a set of primary sensory neurons, the amphid sensilla in C. elegans, which project sensory processes to the nose of the worm (Heiman and Shaham, 2009; Procko and Shaham, 2010). The tips of these dendrites associate directly with an amphid sheath (AMsh) glial cell. Time-lapse studies show that dendrite elongation occurs as a result of cell body migration away from the nose, rather than as directed growth of the dendrite itself. Dendritic anchoring requires extracellular matrix (ECM) components provided by both the neuron and hypodermal cells at the dendrite tip. A protein called DYF-7, which is a secreted zona pellucida (ZP) domain protein, is provided by neurons, while DEX-1, a secreted zonadhesin domain protein, is provided by hypodermis. The partnership between ZP domains and zonadhesin proteins has an interesting counterpart in vertebrates (Procko and Shaham, 2010).

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The ECM of vertebrate oocytes is made up of ZP domains, and zonadhesin is a sperm protein important for fertilization. Similarly, in Drosophila chordotonal organs, a type of mechanosensory organ, dendrite orientation is determined by migration patterns of an organ accessory cell with which the dendrite maintains a close association (Mrkusich et al., 2010). Development of the accessory cell is dependent upon Netrin and the Netrin receptor Frazzled, which provide a context in which this pathway can nonautonomously influence dendrite orientation.

11.8 Control of dendritic field formation II: dendritic self-avoidance and tiling Dendritic territories define how and where neurons receive synaptic or sensory information from other neurons or from the environment. Invertebrate neurons have served as a model for understanding how receptive fields are specified during development to ensure appropriate connectivity and information processing. It is worth considering that dendrites are growing through a highly dynamic environment consisting of other cells, ECM, and glial cells, and thus any cue that they encounter on their way to forming a normal receptive field is changing and likely transient. In this environment, however, dendrites must assume a predictable, characteristic, shape to ensure proper information flow. In this section, we consider the mechanisms that generate the sizes and arrangements of dendritic fields in individual neurons, and also relative to other neurons and nonneuronal cells; an important step in achieving the correct dendritic arbor patterning.

11.8.1 Interactions between dendrites generate evenly covered territories 11.8.1.1 Dendritic self-avoidance Dendrites are typically depicted in textbook images as branched structures extending outward more or less evenly from the neuronal soma. Dendrites arising from the same neuron are called sister dendrites, and these indeed typically spread away from one another without overlap. This spreading property of dendrites (and axons) is termed self-avoidance and results in largely nonoverlapping arrangements of sister dendrites. By contrast, axons and dendrites from different types of cells must coexist in the nervous system for proper connectivity that includes overlapping receptive fields with different tuning properties (Fig. 11.7A). How do dendrites distinguish among sister dendrites, which they avoid, and dendrites from other neurons, with which they can share space and overlap? Early studies of this problem were carried out in highly branched leech sensory axons (Blackshaw et al., 1982; Kramer and Kuwada, 1983; Wang and Macagno, 1998). Lesioning of individual stereotyped axon branches led to ingrowth of remaining neighboring sister branches, indicating that selfavoidance arises through a repulsive mechanism. Kramer and Stent concluded that, given the large number of neurons in the nervous system and large number of nonsister arbors that would need to coexist, any molecular solution has to provide a great deal of recognition diversity (Kramer and Stent, 1985). A solution to this problem was uncovered decades later by experiments in Drosophila. Multiple studies indicate that self-avoidance depends critically on Down syndrome cell adhesion molecule 1 (Dscam1), which is a member of the immunoglobulin superfamily of transmembrane adhesion molecules. Drosophila Dscam1 is a homolog of a human gene with similar protein domain organization. The Dscam1 locus supports the extensive recognition diversity that is required for the discrimination of self versus nonself by encoding up to 38,016 isoforms; this is achieved via extensive alternative splicing that leads to isoform diversity encoded in the extracellular and transmembrane regions of the protein (Fig. 11.7A) (Schmucker et al., 2000). Splicing in the extracellular region occurs in three variable parts of different Ig domains, potentially giving rise to over 19,000 isoforms with strict homophilic binding activity (Fig. 11.7A) (Schmucker et al., 2000; Wojtowicz et al., 2004, 2007). Each neuron expresses a unique set of Dscam1 isoforms to “barcode” its dendrites (Neves et al., 2004; Zhan et al., 2004). In neurons lacking Dscam1 function, dendrites that normally spread across their territory without overlap now cross each other extensively or form tight bundles, indicating a loss of self-avoidance of dendritic processes from individual neurons (Fig. 11.7B) (Hughes et al., 2007; Matthews et al., 2007; Soba et al., 2007; Zhu et al., 2006). Single isoforms are sufficient for dendritic self-avoidance within a single neuron; however, coexistence of dendrites is eliminated if two neurons are forced to express the same Dscam1 isoform (Fig. 11.7C). Genetic engineering to reduce the number of isoforms capable of being produced from the Dscam1 locus has shown that thousands of isoforms are required to ensure that dendrites can distinguish among sister and nonsister dendrites (Hattori et al., 2007a, 2009). In the current model for Dscam1 function, the presentation of Dscam1 isoforms is the same for all dendrites that come from the same soma, so only other sister dendrites share the same molecular tag, defined by Dscam1 isoform expression, that signals recognition and repulsion (Hattori et al., 2008). This feature also explains how nonsister dendrites are able to coexist, since the chance of two neurons stochastically selecting the same set of Dscam1 isoforms from such a very large possible pool would be exceedingly small. How binding between cell adhesion molecules ultimately initiates repulsion

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FIGURE 11.7 Dscam controls self-avoidance of dendrites. (A) Genomic, mRNA, and protein organization of Dscam. The Dscam locus can generate thousands of isoforms via alternative splicing of the extracellular and transmembrane domains. Alternative splicing of the extracellular domain gives rise to transmembrane molecules with isoform-specific binding ability. The two transmembrane splice forms determine axon versus dendrite targeting. (B) da neurons normally show self-avoidance of dendrites, that is, sister dendrites do not overlap. Dscam mutant da neurons show extensive dendritic crossing indicating a defect in self-avoidance. Single arbitrary Dscam isoforms provided back to Dscam- neurons can fully rescue self-avoidance defects. (C) Neurons in different classes can cover overlapping fields, that is, their arbors coexist. When both cells are forced to express a single isoform, their dendrites now show ectopic repulsion. These results indicate that Dscam molecular diversity is not strictly required for self-avoidance, but is required for self versus nonself discrimination that is essential for coexistence of arbors. Data summarized from Hughes, M.E., Bortnick, R., Tsubouchi, A., Baumer, P., Kondo, M., Uemura, T., Schmucker, D., 2007. Homophilic Dscam interactions control complex dendrite morphogenesis. Neuron 54, 417e427; Soba, P., Zhu, S., Emoto, K., Younger, S., Yang, S.J., Yu, H.H., Lee, T., Jan, L.Y., Jan, Y.N., 2007. Drosophila sensory neurons require Dscam for dendritic self-avoidance and proper dendritic field organization. Neuron 54, 403e416; (A) Reproduced from Wojtowicz, W.M., Flanagan, J.J., Millard, S.S., Zipursky, S.L., Clemens, J.C., 2004. Alternative splicing of Drosophila Dscam generates axon guidance receptors that exhibit isoform-specific homophilic binding. Cell 118, 619e633, with permission.

likely depends on sequences within the intracellular tail since expression of a truncated Dscam1 molecule with an intact extracellular domain but no intracellular domain leads to adhesion but not repulsion. The signaling pathways downstream of Dscam1-mediated self-avoidance remain to be defined. The mammalian Dscam gene does not produce extensive splicing variants as the Drosophila Dscam1 gene does. In mammals the neurite self-avoidance is mediated by another set of genes, called clustered protocadherins (Pcdhs), which

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provides extensive molecular diversity through molecular mechanisms that are distinct from the alternative splicing of Dscam1 (Zipursky and Sanes, 2010). Thus, there appears to be a convergent evolution that leads to similar functions of mammalian Pcdhs and Drosophila Dscam1 in neurite self-avoidance. The role of Pcdhs in dendrite self-avoidance in mammals is discussed in detail in Chapter 12, “Dendritic Development: Vertebrates.” For neurons that elaborate their dendrites in a space that is almost two-dimensional (2D), such as the Drosophila da neurons, a key mechanism that limits dendritic crossingsdincluding both self-avoidance and tiling (see below)dis to confine dendrites on a 2D plane. This is achieved through interactions between dendrites and ECM or epidermal cells. Neuronal integrins and epidermis-derived laminins, but not Dscam1, play a critical role in such interactions (Han et al., 2012; Kim et al., 2012). Moreover, the epidermis secretes Sema-2b, which binds to PlexB on dendrites, to regulate dendriteeECM interaction (Meltzer et al., 2016). Therefore, dendritic self-avoidance in Drosophila is achieved through a multistep process that involves homotypic repulsion mediated by Dscam1 and the confinement of dendritic branches on 2D planes by epidermisedendrite interactions mediated by Sema-2b/PlexB and laminins/integrins.

11.8.1.2 Dendritic tiling Dendritic tiling refers to the contiguous and nonoverlapping arrangement of arbors often observed among different neurons of the same functional type, similar to nonoverlapping tiles covering a floor. This optimal coverage is an efficient way to ensure that all sensory input space is covered at least once by each functional type of neuron, and it is also seen in vertebrate neurons including certain retinal ganglion cells and amacrine cells in the visual system, and in sensory neurons of insects and worms (Gallegos and Bargmann, 2004; Grueber et al., 2001, 2002; Smith et al., 2010). In Drosophila, tiling is observed in two classes of da neurons, the class III and class IV neurons. Laser ablation of single class IV neurons early in development, before territory boundaries are formed, leads to invasion of the vacated region by dendrites from remaining neurons on all sides, indicating that neurons prevent other neurons from projecting dendrites into their territory (Fig. 11.8) (Grueber et al., 2003b; Sugimura et al., 2003). One possibility is that a factor is presented by the membranes of dendrites that is capable of being sensed by other like-type neurons and that this factor induces cessation of further dendritic growth, or maybe induces an intracellular response that leads to growth cone turning. Genetic approaches have, however, revealed signaling mechanisms that apparently impact tiling. Two genes, furry and tricornered, are each necessary for self-avoidance and tiling in class IV neuron dendrites (Emoto et al., 2004). Tricornered is a serine/threonine kinase and a member of the NDR kinase family, members of which are generally required for the outgrowth of branched structures such as bristles and dendrites in Drosophila. The function of Furry is unknown, but does interact physically and genetically with Tricornered. This pathway is important for turning behavior of dendrites, presumably in response to repulsive cues (Emoto et al., 2004). Tricornered is also important for the branching of dendrites but the roles in tiling and branching are separable in that only the latter requires the small GTPase Rac1. Additional components of the Tricornered tiling pathway include the target of rapamycin complex 2 (TORC2) members TOR, Rictor, and Sin1 (Koike-Kumagai et al., 2009). TORC2 is critical for Tricornered kinase activity. Links have not yet been made between Tricornered and cell surface receptors important for dendriteedendrite repulsion. Like dendritic self-avoidance, dendritic tiling also requires the confinement of dendrites on 2D planes. In fact, consistent with their roles in both tiling and self-avoidance, the dendritic crossing defects in tricornered and furry mutants are caused by impaired confinement of dendritic growth on a 2D plane, rather than a defect in repulsion (Han et al., 2012). The Sema-2b/PlexB signaling activates the Tricornered kinase to regulate dendriteeECM adhesion, confining dendritic branches in 2D planes (Meltzer et al., 2016). A role for Tricornered and Furry is conserved in C. elegans sensory neuron tiling (Gallegos and Bargmann, 2004). The worm gene sax-1 encodes the homolog of trc, and worm sax-2 encodes a homolog of furry. These genes were studied in the tiling of mechanosensory neurons called posterior lateral microtubule (PLM) and anterior lateral microtubule (ALM). Tiling by PLM and ALM is accomplished by a mechanism that is different from fly sensory neurons. Instead of repelling each other when they meet, the PLM neuron initially overshoots its territory and overlap ensues. However, as the animal grows, PLM extension ceases and the territories of the neurons come into register with their intended targets. After this matching, the sensory neuron and substrate grow together and tiling is maintained. Animals defective in sax1/2 are unable to cease their initial dendrite overgrowth and consequently, owing to the scaling phase of growth, end up with overlapping dendritic fields. In contrast to the fly system described above, ALM and PLM do not appear to signal mutual repulsion. These studies show that it is important to determine how phenotypes arise during development since it allows one to better determine the dynamic roles played by any signaling component involved in regulating neuronal morphology.

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FIGURE 11.8 Tiling of dendritic arbors. Upper panels show normal dendritic development of dendrites in embryonic and larval stages. By early larva neighboring, dendrites belonging to the same class fully and nonredundantly tile their territories. Lower panels show the consequences of single-cell ablation of one of the tiling neurons in embryos. By larval stages, the vacated territory has been invaded by neighboring same class neurons such that much of the field is still covered. These results imply the existence of repulsive signals that are passed between cells of the same class and normally lead to the setting of strict territory boundaries. Data are from Sugimura, K., Yamamoto, M., Niwa, R., Satoh, D., Goto, S., Taniguchi, M., Hayashi, S., Uemura, T., 2003. Distinct developmental modes and lesion-induced reactions of dendrites of two classes of Drosophila sensory neurons. J. Neurosci. 23, 3752e3760; Grueber, W.B., Jan, L.Y., Jan, Y.N., 2003a. Different levels of the homeodomain protein cut regulate distinct dendrite branching patterns of Drosophila multidendritic neurons. Cell 112, 805e818; Grueber, W.B., Ye, B., Moore, A.W., Jan, L.Y., Jan, Y.N., 2003b. Dendrites of distinct classes of Drosophila sensory neurons show different capacities for homotypic repulsion. Curr. Biol. 13, 618e626.

11.8.2 Scaling growth of arbors and maintenance of evenly covered territories Dendritic arbor growth in vertebrates and invertebrates typically proceeds with a period of rapid extension followed by relative stability of arbor shape and size. However, as animals and organs grow and are reshaped during development, coordinated growth of neuronal structures is required in order to maintain coverage of inputs and appropriate neuronal connectivity. This later scaling phase of neuronal growth has been investigated in Drosophila larval da sensory neurons, which scale their growth to the extensive growth of the body wall (Parrish et al., 2009). Scaling is robust, since neuronal growth matches various genetic mutant strains with larger or smaller body sizes. The molecular basis of scaling growth involves signals from the epidermis to the neuron. Expression of the miRNA bantam in epidermal cells is responsible for limiting the growth of overlying epidermal cells. The signal is very local, since loss of bantam from individual epidermal cells leads to overgrowth of only those dendrites that grow over that mutant cell. The basis of this signaling event is not currently known, but it seems that the signal from epidermis to neuron is capable of activating expression of the Akt kinase in the neuron. The robust scaling of class IV arbors provides an interesting contrast to ALM and PLM neurons in the worm, discussed above, in which a lack of scaling growth is actually essential for matching neuronal arbor size to the size of the animal body surface.

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11.9 Dendritic remodeling Metamorphosis is a dramatic developmental transition that is widespread among invertebrates. Extensive reorganization of the nervous system is necessary for the specification of new, adult-specific, behaviors such as flying, walking, and mating (Truman, 1990). The changes that occur in the nervous system during the complete metamorphosis from a larval to an adult form have provided a robust model for understanding dendritic pruning and regrowth during development. In the moth Manduca sexta, about 90% of adult motoneurons are derived from motoneurons that were present in the larval stage. During metamorphosis, larval neurons either die or remodel their dendrites and axons to be integrated into adult neural circuits. In Manduca, different motoneurons, named MN1, MN4, and MN6, show contrasting fates during metamorphosis, implying that different neurons have distinct genetic programs that specify whether they will die or persist and remodel. Whereas MN6 degenerates, MN1 persists through metamorphosis and remodels its dendritic arbor. Remodeling of MN1 involves loss of higher-order (third- and fourthorder) branches in the pupal stage, along with dendritic swellings along primary and secondary branches. When the adult stage is reached, higher-order branches have reemerged from persisting second-order branches, and an entirely new dendritic field is generated (Truman and Reiss, 1976). Two-photon microscopy allows for live imaging of sensory arbors undergoing regression, and these events are likely representative of what occurs in other neurons (Williams and Truman, 2005a). Reduction of dendritic arbors involves local degeneration and branch retraction. Pruning begins with disruption of the microtubule cytoskeleton, then thinning, breaking, and fragmentation of dendrites (Fig. 11.9) (Williams and Truman, 2005b). The process shares several features with cellular apoptosis, a theme that we discuss below.

11.9.1 Transforming growth factor-b signaling and ecdysone receptor expression during dendritic remodeling Knowledge about the genetic programs that control remodeling during metamorphosis has emerged primarily from studies in Drosophila. Several intrinsic factors have been identified as being critical for dendrite pruning, including ecdysone receptors, transforming growth factor-b (TGF-b) signaling pathway, the ubiquitineproteasome system, and caspase activation. 20Hydroxyecdysone (ecdysone) is a steroid hormone that coordinates the metamorphic transition in insects. Expression of different ecdysone receptor isoforms (EcR-A, EcR-B1, and EcR-B2) correlates with different cellular responses to circulating hormone. EcR-B1 expression in neurons is generally correlated with dendritic pruning (Truman et al., 1994), and knockout and rescue studies show that EcR-B isoforms, but not EcR-A, are required cells autonomously for pruning in several distinct types of neurons (Kirilly et al., 2009; Kuo et al., 2005; Lee et al., 2000; Schubiger et al., 1998; Williams and Truman, 2005a; Zheng et al., 2003). Genetic screens have also clarified pathways that are required for EcR-B1 expression and function. Disruption of TGF-b signaling in mushroom bodies prevents dendritic remodeling at metamorphosis (Fig. 11.10) (Zheng et al., 2003). These TGF-b mutant neurons also lack EcR-B1 expression, and provision of EcR-B1 in TGF-b mutant neurons rescues remodeling, placing TGF-b signaling upstream of patterned EcR-B1 expression.

11.9.2 Sox14 and mical function downstream of ecdysone receptor Patterned EcR expression is clearly critical for generating different responses of neurons to circulating ecdysone. Whether cells carry out pruning, maintenance, or death programs depends on the cascades of factors induced downstream of EcR transcriptional activity. An RNAi screens for factors that are regulated by ecdysone signaling identified the high-mobility group (HMG)

(A)

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FIGURE 11.9 Cytoskeletal changes in remodeling dendrites. (A) A larval ddaC neuron showing the approximate region of the arbor depicted in panels (B) and (C). (beb0 ) Branches from a late larval stage labeled for a membrane marker (mCD8::GFP) and the microtubule-binding protein Futsch (b0 ). Futsch labeling looks smooth and continuous. (cec0 ) A few hours after puparium formation, the branches are decorated with flipodia (f) and the Futsch staining is beaded (arrows along dendrite). Figure reprinted from Williams, D.W., Truman, J.W., 2005a. Cellular mechanisms of dendrite pruning in Drosophila: insights from in vivo time-lapse of remodeling dendritic arborizing sensory neurons. Development 132, 3631e3642; with permission.

250 PART | I Formation of axons and dendrites

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FIGURE 11.10 Molecular basis of dendritic remodeling in Drosophila. (A) Mushroom body g neurons in larval stages showing dendritic (red) and axonal (blue) arbors. Eighteen hours after puparium formation, the dendrites have regressed. Dendritic regression is under the control of the TGF-b pathway, which induces expression of the EcR-B1 isoform of the ecdysone receptor, allowing neurons to show appropriate responses to ecdysone hormone. (B) Molecular pathways of pruning have been worked out in the ddaC da neuron. A boxed region of ddaC is shown in (C) as an area that in wild type (w1118 genotype) undergoes complete pruning by 14 h after puparium formation. By contrast, the axon and cell body show no overt changes at this time, suggesting local signaling leading to dendrite-specific pruning. (D) A transgenic reporter for Caspase activity. mCD8 localizes the reporter to membranes, and the PARP sequence is a target of Caspases. The cleaved peptide is then recognized by immunohistochemistry. By around 7e12 h APF, the class IV neuron shows Caspase reporter activity restricted to dendrites. (E) A summary of the molecular mechanisms of dendrite severing in class IV dendrites. See text for details. Part (BeD) reproduced from Williams, D.W., Kondo, S., Krzyzanowska, A., Hiromi, Y., Truman, J.W., 2006. Local caspase activity directs engulfment of dendrites during pruning. Nat. Neurosci. 9, 1234e1236, with permission; Part (E) adapted from Kuo, C.T., Zhu, S., Younger, S., Jan, L.Y., Jan, Y.N., 2006. Identification of E2/E3 ubiquitinating enzymes and caspase activity regulating Drosophila sensory neuron dendrite pruning. Neuron 51, 283e290, with permission.

transcription factor Sox14 (Kirilly et al., 2009). Sox14 RNAi leads to severe dendrite pruning defects in the class IV da neuron, ddaC, such that by 18 h after puparium formation (APF) a large number of dendrites is still attached to Sox14 knockdown neurons (wild-type control neurons are completely pruned by this stage of development). Interestingly, sensory neurons that normally undergo apoptosis do not die upon Sox14 knockdown, indicating a common requirement for Sox14 in both pruning and apoptosis. Expression of Sox14 depends on EcR-B1 isoform expression in ddaC, and Sox14 levels are slightly higher in neurons destined to undergo apoptosis than those destined to prune their dendrites. Whether differential levels of Sox14

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determine pruning versus apoptotic fates is an interesting remaining question. Mical, a large multidomain cytosolic protein, is upregulated in ddaC coincident with, and dependent upon, Sox14 (Kirilly et al., 2009). Furthermore, mical mutant neurons show disrupted dendrite severing. Thus, a pathway composed of EcR, Sox14, and Mical governs severing of dendrites during pruning and might act similarly in different types of neurons. Notably, Mical is dispensable for normal apoptosis, suggesting that it is involved in a select subset of Sox14 functions during metamorphic remodeling of the nervous system. How Mical controls severing in dendrites is unknown. However, given the evidence that strongly suggests its role in F-actin disassembly (Alto and Terman, 2018), it may rearrange the F-actin cytoskeleton in dendrite severing.

11.9.3 Signaling mechanisms for dendritic pruning 11.9.3.1 Ubiquitineproteasome system Key signaling events initiated during pruning have been worked out for the ddaC neuron (Fig. 11.10). Pruning relies on EcR-B1, similar to other systems, as well as the ubiquitineproteasome system and local caspase activity in dendrites (Kuo et al., 2005, 2006; Williams and Truman, 2005a; Williams et al., 2006). The ubiquitineproteasome system is a conserved mechanism for protein degradation. Proteins to be degraded that are bound by E3 ubiquitin ligases are targets for ubiquitin transfer from protein carriers of activated ubiquitin. Disruption of this system leads to aberrant accumulation of proteins, which may be deleterious for cell function. Conversely, activation of the system can initiate axon degeneration after neuronal damage. Blocking ubiquitin activation by expressing into neurons, a yeast ubiquitin protease, UBP2, prevents the severing step of pruning (Kuo et al., 2005). In addition, mutations in the uba1 gene, a gene that is required for ubiquitin activation, also prevents dendrite severing. The fly genome encodes numerous E2/E3 ubiquitin ligases, but surprisingly a single E3 ubiquitinating enzyme activity encoded by the ubcD1 gene is specifically required for dendritic pruning of ddaC (Kuo et al., 2006). Moreover, the pruning of ddaC dendrites also requires the ATPase Valosin-containing protein (VCP) (Rumpf et al., 2011), which is a chaperone for ubiquitylated proteins and in human is involved in ubiquitin-positive frontotemporal lobar degeneration and amyotrophic lateral sclerosis. The ubiquitin system seems to regulate pruning through two separate pathways: by degrading the caspase inhibitor DIAP1 (see below) (Kuo et al., 2006; Rumpf et al., 2011) and by regulating the splicing and expression of the pruning factor Mical (Rumpf et al., 2014).

11.9.3.2 Caspases The exquisite timing of dendrite pruning at the onset of metamorphosis is a remarkable feature of the remodeling process, but also noteworthy is the selective destruction of dendrites and sparing of axons (Kuo et al., 2005; Williams and Truman, 2005a). The similarities between dendrite pruning and cellular apoptosis prompted investigation of apoptotic machinery in ddaC pruning, and this provided insight into the spatial regulation of pruning (Fig. 11.10). The activity of caspases, proteases critical for apoptosis, can be inhibited in Drosophila by disruption of DRONC, the sole initiator caspase. Removal of DRONC activity by either knockout or expression of a dominant negative protein largely inhibits pruning at the stage of branch detachment, indicating that apoptotic machinery is important for dendrite pruning (Kuo et al., 2005; Williams and Truman, 2005a). The activity of DRONC is, in turn, controlled by the negative regulator DIAP1 (Drosophila inhibitor of apoptosis), and overexpression of DIAP1 inhibits branch removal. Interestingly, though, expression of proapoptotic proteins (that function as negative regulators, or antagonists, of DIAP1) in ddaC promotes apoptosis. This suggests that if apoptotic machinery is indeed active (as the DRONC results suggest), then it must normally be localized in the dendrites and not throughout the rest of the cell. Indeed, results obtained with a transgenic reporter of caspase activity (Fig. 11.10D), and also the localization of antibodies that detect activated caspases, indicate that caspase activity is restricted solely to dendritic arbors and, furthermore, that caspase activity is activated only locally in pruning dendrites. Selectivity for dendrites could involve specific localization or activation of proteins that promote pruning to dendrites; sequestering these proteins from axons; or, alternatively, ubiquitous localization of factors that promote pruning but selective protection of axons from their action.

11.9.4 The cell biology of dendritic pruning 11.9.4.1 Microtubule disassembly Dendrite severing also involves local microtubule disassembly, which in sensory neurons involves two kinase, the Ik2 and Par-1 kinases (Lee et al., 2009; Herzmann et al., 2017). Ik2 is closely related to mammalian IKK proteins and promotes the degradation of DIAP1; Par-1 is involved in cell polarity and is known to phosphorylate microtubule-associated proteins. In

252 PART | I Formation of axons and dendrites

Ik2 or Par-1 mutants, the microtubule cytoskeleton remains intact during the early phase of dendrite pruning. Premature activation of Ik2 leads to premature severing of dendrites, suggesting that activation of Ik2 provides a temporal signal for the initiation of severing (Lee et al., 2009). Moreover, Par-1 seems to promote the disappearance of the microtubuleassociated protein Tau during pruning (Herzmann et al., 2017). Although how Ik2 and Par-1 promote microtubule disassembly during dendrite pruning is still unclear, this process might involve the microtubule-severing protein Katatin-p60-like1. In a screen for microtubule destabilizing or severing proteins that are required for efficient dendrite severing, Jan and colleagues found that ddaC neurons mutant for Kataninp60-like1 show delayed dendrite severing, and dendrites become separated from the soma several hours later than normal (Lee et al., 2009). Whether or not Ik2 and Par-1 regulate Katanin-p60-like1 remains to be determined.

11.9.4.2 Local endocytosis and compartmentalized calcium transients Before dendrite pruning, the presumptive locations of severing undergo thinning (Kanamori et al., 2015). This event coincides with enhanced local endocytosis at these locations and the appearance of Ca2þ transients in the branches distal to them (Kanamori et al., 2015). The small GTPases Rab5 and dynamin, which regulate endocytosis, are required for these local endocytosis and compartmentalized Ca2þ transients. These Ca2þ transients activate the protease calpain to facilitate severing of dendritic branches (Kanamori et al., 2013). Thus, it has been proposed that this local endocytosis-mediated dendrite thinning compartmentalizes dendrites for pruning, possibly by promoting compartmentalized Ca2þ transients (Kanamori et al., 2015).

11.9.5 Similarities between dendrite pruning and injury-induced axon degeneration Dendrite pruning also shares certain morphological properties and molecular mechanisms with injury-induced axonal degeneration, termed Wallerian degeneration. In remodeling ddaC neurons overexpressing Wallerian degeneration slow (or WldS), a mutant protein that dominantly delays the degeneration of severed axons (Coleman and Freeman, 2010) causes the severed dendritic branches to persist for an unusually long period. The C-terminal portion of WldS is derived from a 285 amino acid sequence of nicotinamide mononucleotide adenylyltransferase 1 (Nmnat1), which is important for NADþ salvage. During pruning, severed dendrites are lost as normal if an Nmnat1 enzyme dead version of WldS is overexpressed. Thus, NADþ-sensitive pathways are important both for injury-induced axonal severing and developmental dendrite pruning.

11.9.6 Similarities and differences in dendrite development, dendrite regrowth after pruning, and dendrite regeneration after injury The dendrites of Drosophila da neurons regenerate after injury-induced severing (Song et al., 2012). Although the number of branches can be completely regenerated to that of uninjured neurons(Stone et al., 2014), the regenerated dendrites differ from uninjured ones in their greatly reduced lengths and the lack of self-avoidance among branches of the same neuron (Thompson-Peer et al., 2016). The underlying molecular differences remain unclear. The regenerated dendrites of da neuron are precociously pruned (Thompson-Peer et al., 2016). Moreover, they display fragmentation of the distal dendrites before severing near the cell body, which is opposite to the pruning process in uninjured dendrites. However, dendritic injuries before pruning do not appear to affect the postpruning regrowth, as the dendrite morphologies of injured and uninjured neurons are indistinguishable in after the metamorphosis is completed. The molecular mechanisms that underlie these differences between injured and uninjured neurons are unclear.

11.10 Concluding remarks The cellular and molecular biology of dendrite development have high degree of similarity between invertebrates and vertebrates (see also in Chapter 12, “Dendritic Development: Vertebrates”). Drosophila and C. elegans offer effective genetic analyses for the identification of mechanisms that underlie dendrite development in vivo. Many important concepts about dendrite development were established based on studies in invertebrates. Nevertheless, a number of fundamental questions in dendrite development remain unanswered. For example, what are the cellular and molecular determinants of the spacing between dendritic branchesda key aspect of dendrite morphology? What are the molecular and cellular underpinnings that direct neurons to assume multipolar, unipolar, or bipolar morphologies? What is the signaling mechanism that underlies Dscam-mediated dendritic self-avoidance? To answer these questions, we likely need the combination of all the approaches that invertebrate and vertebrate systems offer.

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See also Formation of Axons and Dendrites: Axon growth and branching; Dendritic Development: Vertebrates; Development of Neuronal Polarity in vivo.

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Sugimura, K., Yamamoto, M., Niwa, R., Satoh, D., Goto, S., Taniguchi, M., Hayashi, S., Uemura, T., 2003. Distinct developmental modes and lesioninduced reactions of dendrites of two classes of Drosophila sensory neurons. J. Neurosci. 23, 3752e3760. Sweeney, L.B., Chou, Y.H., Wu, Z., Joo, W., Komiyama, T., Potter, C.J., Kolodkin, A.L., Garcia, K.C., Luo, L., 2011. Secreted semaphorins from degenerating larval ORN axons direct adult projection neuron dendrite targeting. Neuron 72, 734e747. Sweeney, N.T., Li, W., Gao, F.B., 2002. Genetic manipulation of single neurons in vivo reveals specific roles of flamingo in neuronal morphogenesis. Dev. Biol. 247, 76e88. Tea, J.S., Chihara, T., Luo, L., 2010. Histone deacetylase Rpd3 regulates olfactory projection neuron dendrite targeting via the transcription factor Prospero. J. Neurosci. 30, 9939e9946. Thompson-Peer, K.L., Devault, L., Li, T., Jan, L.Y., Jan, Y.N., 2016. 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Chapter 12

Dendrite development: vertebrates Julie L. Lefebvre1, 2, a and Julie Marocha1, 2 1

The Hospital for Sick Children, Toronto, ON, Canada; 2Department of Molecular Genetics, University of Toronto, Toronto, Canada

Chapter outline 12.1. The structure and function of vertebrate dendrites 257 12.1.1. Methods for manipulating and studying dendrite morphology in vertebrates 259 12.2. The cell biology of dendritic growth 259 12.2.1. Regulators of the microtubule network in dendrite formation 259 12.2.2. Regulators of the actin cytoskeleton 260 12.2.3. Dendrite elaboration requires a satellite secretory pathway 261 12.2.4. RNA translation in dendrites 262 12.2.5. Powering dendrite growth 262 12.2.6. Intracellular cascades that translate extrinsic signals into changes in dendrite structure 262 12.3. Control of dendritic field formation I: size 263 12.3.1. Afferent-derived neurotrophins limit size 263 12.3.2. Control of arbor size by neurotransmission 263 12.3.3. Activity-dependent mechanisms that influence dendrite growth and stabilization 264 12.4. Control of dendritic field formation II: shape 265 12.4.1. Apical dendrite initiation and outgrowth of cortical pyramidal neurons 265 12.4.2. Activity-dependent orientation of dendrite growth in the somatosensory cortex 266 12.4.3. Positional cues shape asymmetric dendritic arbors in the mouse retina 266

12.5. Control of dendritic field formation III: targeting and synapse selectivity 268 12.5.1. Formation of a Proto-IPL by retinal amacrine cells 269 12.5.2. Laminar targeting of retinal dendrites is coordinated by adhesive and repellent cues 269 12.5.3. Transcriptional control of laminar-specific targeting of dendrites in retina 271 12.5.4. Local recognition mechanisms to control synapse selectivity 273 12.5.5. An integrated, multistep model for synaptic wiring in the retina IPL 273 12.6. Space-filling mechanisms to optimize dendritic field distribution 273 12.6.1. Tiling and mosaics 273 12.6.2. Dendrite self-avoidance 275 12.7. Emergence of dendrite compartmentalization 277 12.7.1. Subcellular patterning of synaptic inputs along dendritic domains 277 12.7.2. Patterning the membrane excitability of dendritic compartments 278 12.8. Neurodevelopmental disorders: the price of poor dendritic development? 278 12.9. Conclusion 279 Abbreviations 279 Acknowledgments 280 References 280

12.1 The structure and function of vertebrate dendrites Neurons are diverse in their morphology and function. Each neuron develops a dendritic tree that is complex in structure and integral to the ways in which the cell receives and processes neural information. The dendritic morphologies of neurons bear cell typeespecific features in dendritic field size, shape, branch length and complexity, and synaptic patterns along dendritic branches, which have important implications for specifying neuronal connectivity (Fig. 12.1). The location and density of dendrites constrain the types and numbers of presynaptic inputs the neuron can receive. The electrical properties of dendrites, combined with the patterns of synapses and distances from the soma, influence the computations performed by dendrites and the contributions of individual postsynaptic potentials to the firing of the neuron. Thus, a Senior author.

Cellular Migration and Formation of Axons and Dendrites. https://doi.org/10.1016/B978-0-12-814407-7.00012-2 Copyright © 2020 Elsevier Inc. All rights reserved.

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FIGURE 12.1 Eight key neuronal cell types used for the study of dendritic morphogenesis in vertebrates. Neurons were labeled with fluorescent dyes or genetically encoded fluorescent proteins and captured in intact tissue by confocal light microscopy. These cell types are distinguishable by their dendritic arbor morphology and characteristic size, shape, targeting, and branch complexity. (A) A cortical pyramidal neuron with a large dendritic arbor that spans multiple cortical layers. Apical dendrites elaborate along the outer layer of the cortex, and basal dendrites radiate from the cell body. Image courtesy: M. Scofield, Medical University of South Carolina. (B) The dendritic arbor of a mouse Purkinje cell bears a dense and complex branching pattern. The nonoverlapping arrangement of Purkinje dendrites is a characteristic of dendrite self-avoidance. (C) A cerebellar granule cell has a diminutive arbor with four, short claw-like dendrites (black arrows) that form postsynapses around large mossy fiber terminals. (D) A tectal neuron from a Xenopus tadpole. This model system is particularly suited for studies on the influence of visual stimuli and neural activity on dendritogenesis and developing circuits using in vivo live imaging. Image courtesy: K. Haas, University of British Columbia. (E) En face view of a retinal starburst amacrine interneuron with a stereotyped, radial arrangement of dendrites, confined to a narrow layer. The uniform spacing of dendrites is dependent on a process called dendrite selfavoidance. (F) J-RGC is a retinal ganglion cell (RGC) subtype that bears asymmetric dendrites pointing ventrally along the retinal plane. (G) Dendritic morphology of an alpha-ON RGC with an arbor and soma that are characteristically larger than other RGC subtypes. (H) An ONeOFF direction-selective RGC (ooDSGC) has a bistratified arbor in which dendrites are confined to two narrow layers (see Figs. 12.4 and 12.5). In this en face view, both ON and OFF arbors project into the same field of view. En face views of retinal neurons show the dendrite morphology along the retina plane, but dendrites are confined to narrow layers when viewed in cross sections (see Fig. 12.4).

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specializations of neuronal function and circuit outputs are in large part determined by cell typeespecific features in dendritic patterns. How do neurons develop their distinctive dendritic morphology? In this chapter, we review mechanisms of dendrite development in vertebrate nervous systems with a focus on cell typeespecific patterning. We begin with an overview of the cell biological mechanisms that build the dendritic structure. We discuss developmental strategies and molecular players that pattern aspects of dendritic field formation: (1) size; (2) shape; (3) targeting; and (4) dendrite coverage. Each feature is controlled by multiple factors such as transcriptional regulators, cues from neighboring cells and synaptic partners, and activity-dependent remodeling. Developing dendrites undergo dynamic changes in shape to optimize their wiring and synapse formation. We draw examples from multiple vertebrate model systems including mice, chick, frog, and zebrafish.

12.1.1 Methods for manipulating and studying dendrite morphology in vertebrates We are just beginning to understand how cell typeespecific dendrite patterns emerge in vertebrate nervous systems. As with invertebrate models (Chapter 11), the standard for quantifiable studies of dendrite morphogenesis is single neuron resolution. Such studies were previously challenging in vertebrate models due to lack of reliable and high throughput access to cell types of interest and the difficulty of prospectively identifying the cells during development. With a growing repertoire of molecular markers for vertebrate cell types, we can now apply genetic approaches similar to those used in invertebrates. Genetic tools provide access to monitor and manipulate defined subpopulations of neurons in vivo. Cre recombinaseebased technologies permit a diversity of manipulations, such as confinement of gene deletions (conditional knockouts) or overexpression of reporters or gene of interest (conditional knock-in) to specific cell types or regions. Targeted gene expression and recombination strategies pioneered in Drosophila, such as the Gal4-UAS and Flp-FRT systems, have also been modified for vertebrate model systems. Sparse labeling of single neurons can be achieved by viral delivery or electroporation of vectors that encode fluorescent proteins driven by selective promoters or Cre-dependent recombination. The genetic toolkit is expanding in versatility and sophistication: diversity of genetic recombinases (e.g., Flp, Dre) for intersectional approaches; improved gene delivery methods such as viral vectors and transposasemediated insertions; options to monitor or manipulate neuronal activities; and improved spatiotemporal and single cell resolution of manipulations. Transcriptional profiling of purified populations or single cells also opens windows on cell typeespecific gene programs that regulate dendrite morphogenesis. Many of the developmental mechanisms discussed here were discovered in studies that combined gene expression profiling and manipulations of neuronal cell types.

12.2 The cell biology of dendritic growth Dendrite arbor development poses enormous demands on the cellular machines that support growth. Neurons grow long, highly branched dendrites through a dynamic process of branch addition and remodeling. Neurite elaboration results in expansion of the cytoskeleton and surface area that is far greater in neurons than in any other vertebrate cell type. The extreme dimensions require massive production of proteins and lipids, and a complex organization of organelles, cellular machines, and satellite structures, many of which are not present in axons. In this section, we review cell biological mechanisms that build the dendritic arbor, and we highlight their specializations for dendritic development and functions.

12.2.1 Regulators of the microtubule network in dendrite formation Dendrite development is a reiterative process of branch extension, retraction, and stabilization that relies on dynamic remodeling of the neuronal cytoskeleton, which is composed of microtubules, actin, and intermediate filaments. Dendritic branches emerge from slim actin-rich protrusions called filopodia. Filopodia are profuse and most retract (Niell et al., 2004). Whether a filopodium stabilizes and matures into a dendrite or synaptic spine depends on extrinsic effectorsdsuch as permissive substrates, guidance receptors, and neuronal activitydthat stimulate actin assembly and microtubule invasion. Similar to axons, dendritic growth cones steer and advance through coordinated microtubule and actin polymerization (Conde and Caceres, 2009). The size and shapes of dendrites are determined by spatial control of actin and microtubule remodeling by extrinsic cues. Microtubules and actin filaments (F-actin) have an intrinsic polarity with a molecularly and functionally distinct plusend that favors polymerization (or barbed end for F-actin). Microtubules are composed of alpha- and beta-tubulin polymers; the exposed beta-tubulin on the plus-end leads to a higher rate of growth and is a key site for regulation of

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microtubule dynamics (Kapitein and Hoogenraad, 2015). Microtubules remodel through alternate spurts of growth and disassembly, which are influenced by GTP-bound states of tubulin and by microtubule-associated proteins (MAPs) and plus-end tracking proteins (þTIPs). Diverse MAPs and þTIPs are required to stabilize or destabilize microtubules, convert to modes of growth or shrinkage, and some act downstream of cell surface signals. For example, MAP2 is required for dendrite elongation by forming cross-bridges to stabilize microtubules (Harada et al., 2002). Neuronal depolarization and phosphorylation by CAMKII enhance MAP2 binding to microtubules, which then facilitate dendritic remodeling and synaptic plasticity (Vaillant et al., 2002). Microtubule polymerization required for dendritic outgrowth occurs largely after the microtubule organization center loses its function (Stiess et al., 2010), suggesting that most dendritic microtubules are generated by local acentrosomal mechanisms. Recent studies support this idea. Acentrosomal microtubules are nucleated by the g-tubulin ring complex and the associated HAUS/augmin complex, and are subsequently stabilized by the minus-end-associated CAMSAP2 (Yau et al., 2014; Sanchez-Huertas et al., 2016; Cunha-Ferreira et al., 2018). Depletion of any of these components reduces dendritic length and branching. Dendritic microtubules can also be generated at Golgi outposts, as shown in Drosophila dendritic arborization neurons (Ori-McKenney et al., 2012; Yalgin et al., 2015) or along preexisting microtubules (Sanchez-Huertas et al., 2016), but the prevalence of these microtubule initiation mechanisms in vertebrate dendrites is unknown. Microtubules also support growth by transporting organelles, and protein and membrane cargos along growing branches. Cargo transport is determined by microtubule polarity and motor proteins, such as kinesins that transport cargo toward the plus-end and dyneins that transport toward the minus-end. In contrast to axons with uniformly plus-ends out microtubule arrays, microtubules within dendrites have a mixture of minus-end and plus-ends-out orientation (Baas et al., 1988). Mixed microtubule polarity is observed in neurites that emerge from nonpolarized neurons, suggesting that axons eventually develop a uniform plus-end-out orientation while dendrites maintain mixed orientations (Stepanova et al., 2003; Yau et al., 2016). This balance might be critical for dendritic outgrowth: increasing the ratio of minus-end-out microtubules in hippocampal dendrites by inhibiting kinesin-5 reduces branch length and thickness (Kahn et al., 2015). By contrast, Drosophila and Caenorhabditis elegans dendrites have a predominantly minus-end-out organization (Chapter 11). Dendritic microtubule orientation may differ among species, cell types, and branch locations. However, Drosophila dendrites also contain mixed polarity early in development (Hill et al., 2012), suggesting that the navigational challenges of dendritic cargo delivery may be best met with both plus-end and minus-end transport. Microtubules with similar orientations and modifications are further sorted into bundles to ensure the specificity and polarized transport of dendritic cargo (Nirschl et al., 2017; Tas et al., 2017). Motor proteins regulate the delivery of dendritic components and prevent them from entering axons and are critical for dendritic compartmentalization by transporting membrane proteins such as neurotransmitter receptors to specific dendritic domains (Kapitein et al., 2010; Petersen et al., 2014; Franker et al., 2016; Karasmanis et al., 2018).

12.2.2 Regulators of the actin cytoskeleton Actin rearrangements that drive dendrite morphogenesis require continual actin turnover and a multitude of actin assembly regulators. Actin nucleators promote either linear F-actin elongation (i.e., formins) or branched F-actin and expansion (i.e., Arp2/3 complex), and their combined actions determine dendrite morphology. Ena/VASP proteins, which act downstream of several guidance receptors, are F-actin elongation factors that promote profilin-mediated actin polymerization and antagonize capping proteins at barbed ends (Kwiatkowski et al., 2007). In Purkinje cell dendrites, the formin Daam1 promotes filament nucleation and elongation, but its activity is limited by MTSS1, an I-BAR family membrane-bending protein, that also upregulates Arp2/3-dependent branching (Kawabata Galbraith et al., 2018). Cordon-bleu (Cobl) is a nucleator that promotes dendritic elaboration in hippocampal and Purkinje neurons by recruiting multiple actin monomers for linear F-actin assembly (Ahuja et al., 2007; Haag et al., 2012). The Arp2/3 complex generates branched F-actin by nucleating new branches on the side of existing filaments. Branched actin networks are prominent in cortical actin and lamellipodia, where they produce outward forces against the plasma membrane to drive neurite enlargement and motility (Strasser et al., 2004; Konietzny et al., 2017). Filament severing proteins such as the actin depolymerization factor and cofilin promote dendrite branching and arbor complexity by increasing uncapped ends available for polymerization and enhancing F-actin turnover (Rosario et al., 2012; Konietzny et al., 2017). Actin-based motility is tightly coupled to microtubule growth by factors that link microtubule plus-ends with localized actin assembly. To advance, turn, or retract dendrites, microtubules interact with the lamellipodial actin mesh through contractile actomyosins and other cross-linking proteins (Coles and Bradke, 2015). MAPs bridge actin filaments with microtubules. Microtubule þ TIPs such as CLIP-170 associate with formins to accelerate actin filament elongation (Henty-

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Ridilla et al., 2016). Downregulation or overexpression of CLIP-170 greatly simplifies or enhances dendritic complexity, respectively, revealing a mechanism to coordinate rapid cytoskeleton reorganization required for dendrite morphogenesis (Swiech et al., 2011; Henty-Ridilla et al., 2016). Spatiotemporal control of cytoskeletal dynamics is coordinated by signals from cell surface receptors that activate Rhorelated GTPases and the WAVE regulatory complex. The Rho-GTPasesdwhich include RhoA, Rac1, and Cdc42dare molecular switches that control cytoskeletal effectors such as formins, Ena/VASP, and MAPs. Rho-GTPases can have antagonizing roles. In dendritic arbors of Xenopus tectal neurons and rodent hippocampal neurons, Rac1 and Cdc42 promote actin polymerization and branch addition while RhoA inhibits dendrite extension (Li et al., 2000; Nakayama et al., 2000; Hayashi et al., 2007). Rho-GTPases are regulated by guanineenucleotide exchange factors (GEFs) that catalyze active GTP-bound states, and by GTPase-activating proteins (GAPs) that render them inactive. GEFs and GAPs link extracellular signals to local actin cytoskeleton rearrangements. For instance, Rac1-specific GEF Tiam1 is activated by NMDA receptor to induce activity-dependent dendrite arborization of pyramidal neurons (Tolias et al., 2005). Purkinje cell dendrite complexity is dependent on the Rac1-specific GEF Dock1, which acts downstream of adhesion-G protein-coupled receptor BAI3 (Lanoue et al., 2013). Plexin1B receptors contain a GAP in their cytoplasmic region that, upon Sema4D binding, downregulates M-RAS and actin polymerization to reduce dendritic outgrowth (Oinuma et al., 2004; Tasaka et al., 2012). With more than 70 GEF and GAP members identified so far in mammals (Huang et al., 2017), these molecular switches might specify cytoskeletal responses to particular stimuli. Arp2/3 activity is stimulated by members of the WiskotteAldrich syndrome protein (WASP) family and the WAVE regulatory complex (WRC). In addition to activation by Rac1, the WRC might be activated by a large, diverse set of transmembrane receptors through direct interaction with a short consensus motif (WIRS) in their intracellular domains (Chen et al., 2014). Several neuronal receptors with WIRS motifs are implicated in neurite patterning, including members of the protocadherin, ROBO, DCC, neuroligin, and G-protein-coupled receptor families. By connecting cell surface receptors to the actin nucleation machinery, receptoreWRC interactions could instruct cytoskeletal changes that dictate dendrite morphology. While this idea has been demonstrated for axonal branching in vertebrates and neurite branching in C. elegans (Chia et al., 2014; Hayashi et al., 2014; Zou et al., 2018), control of actin remodeling by a cell surface receptor that signals through the WRC remains to be shown in vertebrate dendrite models.

12.2.3 Dendrite elaboration requires a satellite secretory pathway Dendrite morphogenesis is a period of immense plasma membrane growth. In addition to expansion, membrane composition is exquisitely patterned with guidance receptors to direct branch growth, and with neurotransmitter receptors and ion channels to mediate dendritic excitability and signal propagation. To meet these demands, the biosynthesis and trafficking of membrane components are supported by secretory organelles located in the soma and by satellite structures located in dendrites. The secretory system includes the endoplasmic reticulum (ER), the ER-to-Golgi intermediate compartment (ERGIC), the Golgi apparatus (GA), and the trans-Golgi network (TGN). From the earliest stages of neuronal polarization, secretory organelles polarize and first direct their delivery toward the axon and then reorganize toward the growing dendrites (Bradke and Dotti, 1997). In pyramidal neurons, polarization of the Golgi apparatus toward the long dendrite directs membrane trafficking and rapid, asymmetric outgrowth of apical dendrites (Horton et al., 2005). Long-range trafficking to dendrites distantly located from the soma may be insufficient. A solution for dendrites is a satellite system of secretory structuresdincluding ER, ER exit sites, and Golgidfor local processing and insertion of dendritic membrane proteins (Pierce et al., 2001; Horton and Ehlers, 2003). This satellite system is excluded from axons. The ER extends a continuous membrane structure throughout axons and dendrites, with ribosome-bound rough ER primarily in somatodendritic compartments and smooth ER in distal neurites. In dendrites, however, there are zones of ER enriched with ER exit sites and Golgi outposts (see below), suggesting that the entire secretory machinery is spatially compartmentalized and available to process cargo to local dendritic domains (Cui-Wang et al., 2012). This form of local trafficking enhances dendritic branching and is promoted by synaptic activity to increase surface expression of AMPA glutamate receptors, suggesting a mechanism for compartmentalized trafficking of receptors (Cui-Wang et al., 2012; Evans et al., 2017). Vertebrate dendrites also contain Golgi outposts, which have a mini-stack morphology and operate similarly to somatic Golgi: they receive membrane cargo from ERGICs and release them for membrane delivery, including transmembrane and secreted proteins such as brain-derived neurotrophic factor (BDNF) (Horton and Ehlers, 2003). In pyramidal neurons, Golgi outposts are most abundant near branch points in major dendrites during dendritogenesis but decline dramatically in mature neurons (Horton et al., 2005; Quassollo et al., 2015). Thus, the extent to which dendritic satellite EReGolgi structures are employed to modify transmembrane and locally translated proteins is not known. Recent innovations in live

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imaging and molecular reporters have revealed that dendritic post-ER cargo transits through a variety of routes. These include long-range retrograde transport to somatic Golgi, direct trafficking from the ERGIC and recycling endosomes to the plasma membrane in a Golgi-independent manner, and transit through Golgi microcompartments that are widely distributed in the dendritic arbor (Hanus et al., 2014; Mikhaylova et al., 2016; Bowen et al., 2017; Evans et al., 2017).

12.2.4 RNA translation in dendrites Satellite secretory systems and mRNA translation machinery endow dendrites with a degree of autonomy over the production of proteins required for growth and plasticity. mRNAs are abundant in dendrites and encode hundreds of proteins critical for dendrite development, including neuronal receptors and cellular effectors that support growth (Cajigas et al., 2012). A handful of dendritically targeted mRNAs have been studied, including those encoding a variety of proteins: MAP2, b-actin, BDNF, the a subunit of the calcium-/calmodulin-dependent protein kinase II (CaMKIIa), the NMDAR NR1 subunit, and in addition a range of miRNAs (Doyle and Kiebler, 2011). Transport and translation of dendritic mRNAs require RNA-binding proteins (RBPs), but only a few have been studied in dendrite development such as FMRP and ZBP1 (Dictenberg et al., 2008; Perycz et al., 2011). Although cue-induced local protein synthesis has mainly been studied in the context of synaptic plasticity and axon guidance (Holt and Schuman, 2013; Ainsley et al., 2014; Cioni et al., 2018), these mechanisms are likely relevant in dendritogenesis. With rapid progress in technologies for sequencing and imaging of RNA transcripts in intact neural tissue and for tracking de novo protein synthesis (Sambandan et al., 2017), new insights on the spatiotemporal control of dendritic translation will certainly emerge.

12.2.5 Powering dendrite growth Dendritic outgrowth poses significant cellular energy demands. Local production of ATP by dendritic mitochondria supports the modifications and trafficking of lipids and proteins. An abundant supply of ATP also supports many phosphorylation reactions such as those that drive cytoskeleton dynamics and set the membrane potential via ionic pumps. In developing dendrites, the motility and density of mitochondria increase significantly, particularly at sites of branching and synapse formation (Fukumitsu et al., 2015; Faits et al., 2016). As circuits mature, dendritic mitochondria become stationary and remain at branch points and synapses (Faits et al., 2016). Perturbations to mitochondrial transport or creatine kinase activity in dendrites lead to ATP deficiency and abnormal aggregation of actin filaments, which diminish branch outgrowth (Fukumitsu et al., 2015). Delivery of mitochondria into dendrites is coordinated by microtubule adaptor proteins such as TRAK2 and dynein-mediated minus-end transport (van Spronsen et al., 2013). The spatiotemporal cues that deliver mitochondria to sites of high-energy demands in growing dendrites are not known.

12.2.6 Intracellular cascades that translate extrinsic signals into changes in dendrite structure Growth and guidance cues instruct changes in dendritic shape through receptor activation and signaling cascades. Receptor tyrosine kinases (RTKs), such as tropomyosin-related kinases (Trk) and Eph receptors, phosphorylate effectors and interact with adaptor proteins. Other guidance receptors, such as Plexins and Robo, signal through Rho-GTPases or other actin regulators. Among the intracellular proteins commonly activated by neuronal receptors and shown to control dendrite growth are calmodulin-dependent protein kinases (CaMK) (Wu and Cline, 1998; Redmond et al., 2002; Wayman et al., 2006), extracellular signaleregulated/mitogen-activated protein (ERK/MAP) kinases (Kumar et al., 2005), phosphoinositide-30 kinase (PI3K)/AKT (Dijkhuizen and Ghosh, 2005), beta-catenin (Yu and Malenka, 2003), and protein kinase C (Garrett et al., 2012). The signaling cascades are complex, cross-interact with each other, and add specificity through recruitment of coreceptors or effectors expressed by the responding cell. We highlight a few examples here and throughout the chapter where we discuss how signals lead to cell typeespecific changes in morphology. In pyramidal neurons, the neurotrophin BDNF stimulates dendrite branching through the MAP kinase and PI3-kinase pathways and independently of protein synthesis (Dijkhuizen and Ghosh, 2005). However, as neurons elaborate larger arbors, they require protein synthesis stimulated by signals downstream of PI3K, including mammalian target of rapamycin complexes (mTORC1 and mTORC2) (Jaworski et al., 2005; Urbanska et al., 2012). Pyramidal dendrite branching is also dependent on cellecell interactions mediated by the cadherin-related clustered protocadherins (cPcdhs). cPcdhs negatively regulate the activities of focal adhesion kinases FAK/Pyk2 and downstream, protein kinase C (PKC) (Chen et al., 2009; Garrett et al., 2012; Suo et al., 2012). As a result, myristoylated alanine-rich C-kinase substrates (MARCKS) are maintained in an active, nonphosphorylated state to interact with the actin cytoskeleton and promote dendrite arborization.

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Neural activity is also a potent regulator of dendrite growth through activation of calcium (Ca2þ) dependent pathways. Upon depolarization, the influx of cytoplasmic Ca2þ activates CaMKIV and leads to phosphorylation of cAMP-response element binding protein (CREB) (Hu et al., 1999; Redmond et al., 2002; Wayman et al., 2006). Other Ca2þ effectors include CaMKII, which is essential for dendritic branch initiation in hippocampal neurons by promoting filopodia motility, extension, and branching (Fink et al., 2003). CREB/CBP induces activity-dependent transcriptional programs that regulate multiple aspects of dendrite maturation and synaptogenesis. CREB targets include genes encoding BDNF and Wnt-2, which further promote dendrite elaboration and synaptogenesis. BDNF itself also stimulates CREB phosphorylation (Finkbeiner et al., 1997), highlighting positive feedback effects of spontaneous or early synaptic activity on the subsequent integration of those neurons into circuits. Other Ca2þ-dependent transcriptional regulators include calcium-responsive transactivator (CREST), NeuroD, MECP2, and MEF2 (Aizawa et al., 2004; Ince-Dunn et al., 2006; Zhou et al., 2006; Flavell et al., 2008). These proteins regulate expression of intrinsic effectors that influence particular aspects of dendrite development, but their gene targets and how they shape morphology are largely unknown.

12.3 Control of dendritic field formation I: size The size and complexity of the dendritic tree define the information a neuron receives by delimiting the types and numbers of presynaptic inputs. As discussed in Section 12.2.6 above, in vitro studies of cortical and hippocampal neurons led to the identification of roles for neuronal activity, neurotrophins, and their downstream signals in dendrite elongation and branching. In vivo, dendrite size must also be adjusted according to the architectural and functional demands of the circuit. One requirement is that the dimensions of the postsynaptic arbor match the size and density of presynaptic terminals. Second, dendritic size should fulfill circuit properties such as convergence or divergence. Neurons that pool information from multiple pathways must be sufficiently large to accommodate numerous inputs. On the other hand, information from a common input can diverge into parallel pathways through multiple target cells bearing small dendritic arbors relative to the afferent territory. Third, topographical variations in dendrite size for a given cell type might be accompanied by differences in cell number or changes in afferent territories to preserve constant synaptic densities (Yu et al., 2018). Here we discuss examples in which neurotrophic information or neural activity are coordinated between afferent and target neurons to tune dendrite size.

12.3.1 Afferent-derived neurotrophins limit size Given the effects of neurotrophins on dendrite outgrowth in a variety of neuronal cell types (McAllister et al., 1995; Lom et al., 2002; Cohen-Cory et al., 2010), neurotrophins released by afferents could influence dendrite arbor size of target cells. This idea has been tested in a few ways. First, sparse labeling of BDNF-secreting neurons in the ferret cortex revealed the potent but exquisitely short-range effects of BDNF on dendritic growth of nearby neurons (Horch and Katz, 2002). The short-range action by BDNF (a mere 4.5 mm) could be relevant to synaptically connected cells. Evidence for afferentderived neurotrophic action comes from a genetic study in the mouse cerebellum, in which the growth of Purkinje cell dendrite arborization is limited by available neurotrophin-3 (NT-3) secreted from parallel fiber afferents (Joo et al., 2014). Remarkably, knockout of the NT-3 receptor TrkC in all Purkinje cells does not affect dendrite arborization, but elimination of TrkC in sparse numbers of Purkinje cells leads to arbor shrinkage in a cell-autonomous manner. This effect was rescued by removing parallel fiber-derived NT-3, suggesting that growing dendrites compete for NT-3 from their presynaptic partners to maximize their dendritic territories.

12.3.2 Control of arbor size by neurotransmission Neural transmission from afferents has a potent influence on dendrite growth and distribution, ensuring that postsynaptic cells fine-tune their connectivity with stronger inputs. Sensory neurons, in particular, are sensitive to deafferentation or inhibition of neural activity. Depending on the cell type and age, decreased activity causes global, cell-wide adaptations or local adjustments to dendritic growth. A classic example of local structural plasticity occurs in the nucleus laminaris (NL) in the chick auditory system. NL neurons bear two dendritic tufts that are similar in size and complexity but receive distinct streams of excitatory inputs from the ipsilateral or contralateral ear. If inputs to one set of NL dendrites are blocked due to axonal transection or inhibition of activity, the denervated NL tuft shrinks while the innervated one maintains its dendritic structure (Deitch and Rubel, 1984; Wang and Rubel, 2012). Loss of neurotransmission, rather than loss of afferent contact, is responsible for this effect because removal of the cochlea two synapses away causes similar shrinkage. Conversely, activation of NL inputs stimulates dendrite growth in the corresponding tuft (Sorensen and Rubel, 2006, 2011). Dendritic

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changes, both in terms of branch retractions and additions, occur within a few hours and are reversible. Thus, developing NL can rapidly and locally adjust the dendritic size to functional changes. This adaptive property may facilitate their function as coincidence detectors for sound localization. Global effects on dendritic arbor size are observed when inhibitory GABAergic transmission is blocked and excitatory drive is increased. In Xenopus tectal neurons and rodent hippocampal neurons, manipulations to GABA-A receptor function increase dendrite arbor growth and area (Wayman et al., 2006; Shen et al., 2009). Similarly, decreased glycinergic transmission in the gerbil auditory system expands dendritic territories, highlighting the importance of balanced excitatory and inhibitory inputs on dendritic maturation (Sanes et al., 1992; Sanes and Hafidi, 1996). In younger neurons, GABA transmission is initially depolarizing due to an immature chloride reversal potential, and these excitatory effects also promote dendrite outgrowth in the developing mouse neocortex in vivo (Cancedda et al., 2007). One source of GABA that stimulates pyramidal dendrite arborization comes from long-range GABAergic projection neurons that terminate in layer 1 and synapse with apical dendrites. During the first postnatal week, these GABAergic inputs exert excitatory responses in cortical dendrites, stimulating branching and synaptogenesis (Chen and Kriegstein, 2015). The effects are limited to apical dendrites receiving GABAergic input since blockade of GABA release attenuates outgrowth of apical dendrites but not those of distant basal dendrites. As these connections mature into inhibitory ones, perturbations during their development could alter inhibitory functions in the mature cortical circuitry.

12.3.3 Activity-dependent mechanisms that influence dendrite growth and stabilization What are the mechanisms that adjust dendritic arbor size during development according to the sensory stimuli from the outside world? In vivo imaging experiments of developing optic tectal neurons in Xenopus tadpoles have provided insights into the cellular dynamics and molecular players (Fig. 12.2). Young tectal cells have simple dendritic arbors that first undergo a rapid growth and branching phase. As they reach a characteristic size, tectal arbors stabilize and are less dynamic (Wu et al., 1999). Thus, dendrites switch from responding to “grow” to “stop-growing” signals. Moreover, dendritogenesis and synaptogenesis occur concurrently, and early synaptic activity promotes the elaboration and stabilization of the dendritic arbor to adjust the size and shape of the arbor accordingly (Cline and Haas, 2008). A potential starting point is the formation of activity-independent, transient contacts between dendritic filopodia and axonal processes through adhesion molecules such as Neurexin and Neuroligin-1 (Chen et al., 2010). Filopodia are then stabilized by NMDA-type receptordependent glutamatergic transmission. Driven by visual activity, NMDA-dependent transmission also stimulates branch

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FIGURE 12.2 Activity-dependent mechanisms that influence dendrite growth and stabilization in the Xenopus visual system. (A) The trajectory of dendritic arbor development of optic tectal neurons in the Xenopus tadpole. Since dendrite development and synaptogenesis occur concurrently, visual activity influences arbor size and complexity at different phases and through distinct mechanisms. Arbor development is highly dynamic, with branch addition and lengthening (green branches) and elimination (red branches). Arbor outgrowth is initially influenced by NMDA-type glutamatergic transmission and increased RhoGTPase activity. At later stages, AMPA receptoremediated synaptic transmission promotes branch stabilization and maintains the arbor structure. (B) Manipulations to neural activity reveal experience-dependent structural plasticity of dendritic arbors. Dark-rearing or blockade of NMDA receptor or AMPA receptoredependent synaptic maturation leads to reduced arbor size and complexity. Visual stimulation enhances synapse formation and transmission while increasing dendritic size and complexity.

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growth through activation of the cytoskeleton by Rho-GTPAses, Rac and Cdc42 (Li et al., 2002; Sin et al., 2002). As synapses recruit AMPA-type receptors, AMPAR-dependent transmission of visual activity contributes to arbor growth and complexity by promoting branch stabilization (Sin et al., 2002; Haas et al., 2006). RhoA and CAMKII limit process elaboration and might mediate stop-growing signals (Wu and Cline, 1998; Li et al., 2000). Early visual experience promotes the interplay between dendritogenesis and synaptogenesis to establish an arbor of appropriate size while enhancing synaptic connectivity, experience-dependent plasticity, and sensitivity to visual stimuli. Dendrite sampling and stabilization are regulated by local calcium signaling. Live calcium imaging studies in intact Xenopus tadpole and zebrafish larvae, as well as in chick and rodents ex vivo and in vivo have captured local calcium transients that emerge during dendrite filopodia dynamics and synaptogenesis. In embryonic chick retinal ganglion cells (RGCs), blockade or focal uncaging of local calcium fluxes affect dendritic branch structures (Lohmann et al., 2002). Moreover, calcium signals emerge upon contacts between a filopodium and prospective axonal partner, and they correlate with increased filopodia stabilization (Lohmann et al., 2005; Lohmann and Bonhoeffer, 2008). As filopodia sample potential axonal targets, calcium-regulated signaling appears to be crucial for the selective stabilization and maturation of appropriate contacts into synapses. Thus, the interplay between dendritogenesis and synaptogenesis is critical for shaping dendritic arborizations. As proposed by the synaptotropic hypothesis, dendritic branch growth and stabilization will be biased toward regions receiving more excitatory drive, as they are likely to encounter additional synaptic partners (Cline and Haas, 2008).

12.4 Control of dendritic field formation II: shape Dendrite orientation, distribution, and branching complexity are features that together give rise to dendritic field shape. As with size, the shape and location of dendrites determine the number and types of synaptic inputs received by a neuron and the integration of synaptic responses. Fine spatial structures of the arbor, such as branch density and distribution, define the density of receptive field information that can be sampled by a neuron. A core principle of dendrite shape development is the interplay between intrinsic and extrinsic regulation. The influence of environmental cues on growing dendrites is determined by gene expression programs that set up the basic pattern and deploy effectors and surface receptors specific to the cell type. In the next section, we discuss some developmental strategies and molecular mechanisms shown to pattern dendrite development of specific neuronal cell types.

12.4.1 Apical dendrite initiation and outgrowth of cortical pyramidal neurons For many neuron types, arbor shape is established by growing dendrites in a directionally biased manner. In cortical pyramidal neurons, the direction of dendrite growth is determined as soon as neurons terminate migration and polarize to extend axons and dendrites. After migration, the neurons are already oriented with the leading edge pointing toward the pia layer, defining a process that will differentiate into the primary apical dendrite. Transition from migration to dendrite initiation in these cells is regulated in part by the transcription factor Sox11 (Hoshiba et al., 2016). Sox11 expression is coupled to migration and suppresses dendrite initiation. Manipulations that downregulate Sox11 expression or disrupt migration prematurely induce branching. Genetic regulation of the transition from migration to dendrite initiation serves at least two roles: (1) ensures that neurons have reached their target location before elaborating dendrites and (2) turns off the expression of genes that respond to promigratory cues or suppress growth, and turns on genes for dendrite growth and guidance (Prigge and Kay, 2018). Once migratory pyramidal neurons reach their cortical layer, they extend an axon toward the subventricular zone and one major apical dendrite toward the pial surface. The pia is a source of diffusible chemoattractants, as shown by dissociated embryonic neurons cocultured on a slice of neonatal cortex that all point their dendrites toward the pia (Polleux et al., 2000). One cue is Sema3A, a secreted member of the semaphorin protein family. Disrupting the Sema3A gradient by overexpression or removal of Sema3A in cultured cortical slices results in outgrowth in random directions. Interestingly, axons from the same pyramidal population are repelled by Sema3A (Polleux et al., 1998). Dendrite-specific attraction to Sema3A might be due to asymmetric localization of soluble guanylate cyclase and production of cGMP in the nascent dendrite, which transduces Sema3A signals into outgrowth responses (Song et al., 1998; Polleux et al., 2000). In vivo, the chemoattractive activities of Sema3A are more complex as ablation of Sema3A signaling through its receptor Neuropilin-1 disrupts branching of basal dendrites but not those of apical dendrites (Gu et al., 2003; Tran et al., 2009). Sema3A may also act at the earliest stages of axon-dendrite polarization, as demonstrated for hippocampal pyramidal neurons (Shelly et al., 2011). Other cues that coordinate polarized properties of radial migration and apical dendrite formation include transcription factor Neurogenin2 and Reelin signaling (Hand et al., 2005; Olson et al., 2006; O’Dell et al., 2015). As discussed

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in Section 12.2.3 above, apical dendrite growth is supported by polarized secretory trafficking, and disruptions to Golgi polarity abolish asymmetric outgrowth (Horton et al., 2005). The polarization of Golgi may be regulated downstream of these cues, or it may regulate the polarized distribution of guanylate cyclase and receptors for Sema3A. The polarization of signal transduction components and cell surface receptors to nascent dendrites may be a general principle for setting up oriented growth. Subsequently, the apical and basal domains undergo substantial branching to form characteristic apical and basal dendritic trees. Sema3A has ongoing roles in apical and basal dendrite outgrowth by signaling with receptor protein tyrosine phosphatase PTP-sigma and tyrosine kinase Fyn (Nakamura et al., 2017). In basal dendrites, Sema3A signals through distinct downstream factors, TAOK2 (thousand-and-one-amino acid 2 kinase) and JNK (c-Jun N-terminal kinase), which are dispensable for apical dendrite morphogenesis (de Anda et al., 2012).

12.4.2 Activity-dependent orientation of dendrite growth in the somatosensory cortex Other neurons elaborate dendrites in a random direction but reorient their growth to areas receiving stronger or more numerous inputs. In developing sensory circuits, dendrite distributions change in response to spatial or activity differences in sensory inputs. This adaptive process leads to new growth and/or retraction of branches, and it eliminates weak connections and strengthens stronger ones. By restricting the connections of sensory afferents to a limited number of target neurons, a point-to-point sensory map is produced to better transmit the spatial information of stimuli from the outside world. A key model in which to study this problem is the rodent somatosensory system. In the cortex, “barrel” structures emerge from experience-dependent remodeling of dendritic structures and synaptic connectivity during the first postnatal week. Barrels comprise a wall of spiny stellate cells that extend an asymmetric arbor into the barrel hollow, which contains thalamocortical axon arborizations (Fig. 12.3). This circuit forms a point-to-point map such that each barrel is topographically arranged in a layout similar to the whisker pads and receives sensory information from one whisker relayed through multiple intermediates (Petersen, 2007). To confine the transfer of information from one whisker to one barrel at the cortical level, the stellate cells reorganize their dendritic arbors to maximize synaptic connectivity with thalamic axons, which enter in a topographically segregated manner. Stellate cells initially extend dendrites that are diffuse and overlap multiple proto-barrels. As they receive thalamic activity, dendrites asymmetrically reorient their growth toward the center of a single barrel with more active inputs. This process is experience-dependent: sensory deprivation, or global or stellate cell-autonomous blockade of NMDA-dependent transmission during the neonatal period results in profuse branching and diminished barrels (Iwasato et al., 1997, 2000; Datwani et al., 2002; Espinosa et al., 2009). In live imaging studies of the neonatal mouse barrel cortex, stellate dendrite tips are observed to be highly motile and to grow and branch increasingly toward the barrel hollow, where they stabilize in an NMDAR-dependent manner (Mizuno et al., 2014; Nakazawa et al., 2018). The transcription factors LIM homeobox 2 (Lhx2) and BTB domain containing 3 (BTBD3) regulate a gene network responsible for this activity-dependent remodeling. In the somatosensory cortex, BTBD3 is expressed exclusively by stellate neurons. BTBD3 expression is regulated by Lhx2, but its translocation from the cytoplasm to the nucleus is stimulated by NMDAR-dependent activity. Loss of Lhx2 or BTBD3 in stellate neurons disrupts dendritic asymmetry and barrel formation, similar to the loss of activity, while restoration of BTBD3 in Lhx2 mutant cortex rescues dendrite asymmetry (Matsui et al., 2013; Shetty et al., 2013; Wang et al., 2017). BTBD3 also controls dendritic remodeling and ocular dominance column formation in the ferret visual cortex. Thus, BTBD3 is a conserved regulator of experiencedependent structural plasticity in the sensory cortex. BTBD3 is a member of the BTB/POZ family along with the Drosophila Abrupt, which controls branch formation of class I dendritic arborization neurons through gene regulation of effectors such as microtubule organizer proteins (see Chapter 11). Mining BTBD3-dependent transcriptional programs could yield insights into effector pathways that regulate experience-dependent dendritic plasticity.

12.4.3 Positional cues shape asymmetric dendritic arbors in the mouse retina How do neuronal subtypes that are related in origin and develop in shared environments elaborate their unique dendritic patterns? Neuronal cell typeespecific arbor remodeling in response to positional cues is one mechanism to diversify shapes. The mouse retina is well suited to study type-specific dendritic patterning. It contains >40 types of RGCs, which differ in transcriptional signatures as well as in their dendritritic arbor patterns such as diameter, symmetry, and branch density (Rheaume et al., 2018; Sanes and Masland, 2015). While the majority of RGCs elaborate symmetric arbors, the JRGC and F-RGC types bear a wedge-shaped asymmetric arbor (Fig. 12.1). Most J-RGCs point their dendrites toward the ventral pole of the retina, a pattern that correlates with their tuning to ventrally directed (upward) motion (Kim et al., 2008).

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FIGURE 12.3 Activity-dependent orientation of dendrite growth in the somatosensory cortex. Postnatal reorganization of spiny stellate neurons in layer IV contributes to the cortical cytoarchitecture of “barrels.” (A) The barrel field in the mature rodent somatosensory cortex receives sensory information from whisker pads in a topographic manner. (B) At postnatal day 0, dendrites of stellate neurons (blue) grow radially. They receive active inputs from topographically organized bundles of thalamic-cortical axons (TCA, beige area). (C) By P7, dendrite outgrowth is oriented toward TCA bundles to maximize afferent contact. Stellate cell bodies align along barrel walls, and arbors become asymmetric with dendrites pointing toward barrel hollow. (D) Upon sensory deprivation, barrel fields fail to emerge or are weakly defined. (E) Inhibition of NMDA-dependent glutamatergic transmission disrupts stellate dendritic reorganization (orange). (F) Loss of transcription factors Lhx2 and activity-dependent BTBD3 disrupt the stellate cell alignment and oriented dendritic outgrowth.

F-RGCs comprise four subtypes with significantly smaller dendritic fields, but they, too, point their dendrites along the same dorsoventral axis (Rousso et al., 2016). How are these asymmetric arbors established? Among the J-RGCs, the degree of asymmetry varies depending on their position in the retinal plane. Early in dendritogenesis, dorsally located JRGCs extend an asymmetric arbor but ventrally located ones are radial (Liu and Sanes, 2017). By the following week, the remaining ventral J-RGCs remodel into asymmetric arbors through a combination of selective branch growth, pruning and reorientation of primary dendrites. J-RGC remodeling is not influenced by neural activity and visual experience (Elias et al., 2018). The identities of these positional cues and cognate receptors in J-RGCs are unknown; candidates include secreted guidance cues, morphogens, or cellecell interactionebased mechanisms akin to planar cell polarity (Liu and Sanes, 2017). Interestingly, the direction selectivity of J-RGCs to upward motion is weaker in those few J-RGCs that maintain a symmetrical arbor, suggesting a correspondence between asymmetric arbor shape and J-RGC feature selectivity (Kim et al., 2008).

268 PART | I Formation of axons and dendrites

12.5 Control of dendritic field formation III: targeting and synapse selectivity Precise connections among synaptic partners form from complex, multistep interactions between dendrites and their environment. The task of circuit assembly seems insurmountable given the dynamic states of neurite growth and sampling. One organization strategy is to target developing axons and dendrites to shared regions, which then organize into densely innervated layers (laminae) or columns. Restricting dendrite growth to layers offers advantages for: (1) facilitating synapse selectivity by limiting potential partners to those that project to the same location; (2) maximizing synapse formation between them due to proximity; (3) reducing the number of recognition molecules required to match synaptic partners; and (4) connecting neurons with related circuit functions, giving rise to anatomically and functionally distinct sublayers. As a result, sublaminae acquire distinct synaptic and circuit properties, and together they compute different features of neuronal information in parallel. Laminar structures in the retina are effective models for the discovery of cellular mechanisms and recognition molecules that control dendritic wiring and synapse specificity (Fig. 12.4). Dendrites and synaptic connections are contained in compact layers called the outer plexiform layer and inner plexiform layer (IPL). The IPL is stratified into a dozen sublaminae, each of which process different features of visual information received by the retina. Dendrites of w40 types of RGCs stratify to particular sublaminae, where they receive synapses from specific subtypes of bipolar and amacrine interneurons. These stereotyped laminar patterns of connectivity confer each RGC type selective responses to visual features, such as contrast, directional motion, and edges. IPL formation and dendrite lamination therein is genetically determined.

FIGURE 12.4 The anatomy of the mammalian retina. Laminar organization of retinal cells and synapses. In the outer retina, the rod and cone photoreceptors are located in the outer nuclear layer (ONL) and form synapses onto horizontal cells (one is illustrated in dark blue) and bipolar cell dendrites (one is illustrated in green) in the outer plexiform layer (OPL). Interneurons in the inner retinal layer (INL) include the horizontal cells (dark blue), bipolar cells (green), and amacrine interneurons (dark red). Excitatory bipolar cells (green) and inhibitory amacrine cells (dark red) project to sublayers in the inner plexiform layer (IPL), where they synapse with RGC subtypes (blue) and shape their responses. The IPL is functionally stratified into w a dozen sublaminae, each of which contains distinct circuits that process particular aspects of visual information received by the retina. The outer IPL layers are part of the OFF visual pathways that produce responses to decrements of light, while the lower IPL layers contain the ON pathways. For simplicity, the IPL is subdivided into five sublayers, S1eS5. The IPL on the left depicts three RGC subtypes (cyan) that target their dendrites to specific sublaminae: a JRGC with dendrites to S1; a local edge detector/Sidekick-2-positive RGC with dendrites confined to S3; and a bistratified ONeOFF direction-selective retinal ganglion cell (ooDSGC) (RGCs) with dendrites that target S2 in the OFF half, and S4 in the ON half. The IPL on the right shows the directionselective circuit to illustrate how inner retinal circuits are organized. The direction-selective circuit occupies OFF and ON sublayers, S2 and S4. ooDSGCs (blue) extend two arbors in the OFF and ON sublayers, where they receive glutamatergic excitatory inputs from OFF and ON bipolar cells (green) and GABAergic inhibitory inputs from OFF and ON starburst amacrine cells (dark red).

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Aside from modest structural changes, manipulations affecting neural activity do not affect IPL structure, at least in mice (Lefebvre et al., 2015). The IPL is a framework to present positional cues for guidance and to cluster presumptive synaptic partners in the same space, while also restricting the presence of connections that do not contribute to circuit function. Here we focus on IPL formation in the vertebrate retina to discuss mechanisms of dendrite targeting and synapse selectivity. The mouse retina model has yielded several recent discoveries, in large part due to a detailed classification of retinal cell types and a powerful toolkit for genetically accessing them during development.

12.5.1 Formation of a Proto-IPL by retinal amacrine cells The developing IPL is classically viewed as a scaffold of positional cues for incoming dendrites. If retinal neurons use cues from the IPL, how does the first layer form? Insights into the formation of a proto-IPL come from time-lapse imaging of fluorescently labeled retinal cells in zebrafish larvae. Amacrine cells are the first cell types to extend dendrites toward the presumptive IPL (Godinho et al., 2005). They rapidly form a narrow plexus between amacrine cells residing in the INL and those displaced in the ganglion cell layer. The plexus stratifies into sublaminae and serves as a scaffold for subsequent targeting of amacrine and RGC dendrites. Targeting behaviors of RGC dendrites are dynamic and diverse: early RGCs added dendritic layers sequentially and in an inside-out manner, while other RGCs biased dendrite growth within narrow sublamina (Mumm et al., 2006). Variations in RGC dendrite targeting reflect subtype-specific differences as demonstrated in the mouse retina (Kim et al., 2010). A proto-IPL assembly model was proposed with the following predictions: (1) amacrine cell dendrites use homotypic, cell surface cues to adhere to each other to form a plexus and to self-assemble into a scaffold; (2) postsynaptic partners, the RGCs, are not required for scaffold assembly; and (3) RGC dendrites use surface cues on the amacrine scaffold to position their dendrites to the correct sublaminae. Molecules that organize the proto-IPL and target RGC dendrites onto the scaffold were recently identified in the mouse retina (Fig. 12.5). A proto-IPL organizer is MEGF10, a transmembrane protein that mediates homotypic interactions between starburst amacrine cells (Kay et al., 2012). As predicted by the proto-IPL model, once migrating starburst amacrine reach the inner retina, they begin to express MEGF10 and grow dendrites that form homotypic contacts and fasciculate into two sublayers (Ray et al., 2018). In the absence of homotypic interactions or MEGF10, starburst cell dendrites grow exuberant processes outside the IPL. Their synaptic partners, the ONeOFF direction-selective RGCs (ooDSGCs) and bipolar cells, have IPL targeting defects because their neurites follow the starburst dendritic sublaminar errors. The latter confirms another aspect of the proto-IPL model: starburst dendrites use cell adhesion molecules to position ooDSGC dendrites and bipolar terminals to their IPL sublayer. Starburst cells are essential for ooDSGC dendrite stratification, and they guide ooDSGCs dendrites through homophilic adhesive interactions mediated by type II cadherin molecules 6, 9, and 10 (Duan et al., 2018). Moreover, other cadherin members mediate selective laminar targeting of bipolar inputs and other circuit components to assemble functional direction-selective circuits (Duan et al., 2014, 2018). Thus, combinatorial adhesive interactions mediated by cadherin members specify synaptic connectivity and neural circuit assembly in the retina and may play similar roles in synaptic matching in the brain. The starburst scaffold is not required for laminar targeting of at least some non-DS-interneurons (Ray et al., 2018), suggesting that other IPL-organizing amacrine scaffolds exist. One candidate is a Fat3-positive amacrine cell population. Similar to MEGF10 mutant retinas, amacrine cells lacking Fat3 fail to polarize dendrite growth toward the IPL and instead send dendrites to ectopic locations (Deans et al., 2011). Formation of a laminar scaffold from interneurons is thus one strategy to recruit pre- and postsynaptic partners belonging to the same circuit and facilitate synaptic connectivity. Scaffolds could also exclude processes from inappropriate neurons. Development of the IPL strata, which in the mouse contains >10 functionally and anatomically distinct circuits, will require several stratification strategies and molecular recognition events.

12.5.2 Laminar targeting of retinal dendrites is coordinated by adhesive and repellent cues Dendrite laminar targeting in retina can be accomplished through a variety of guidance strategies, including (1) selective adhesion onto existing layers similar to amacrine scaffolds; (2) repulsive cues localized to sublayers prevent invasion of dendrites from inappropriate cell types; (3) mutual expression of recognition or adhesion molecules in cell types that form connections, or target similar layers. Dendrite laminar patterning is coordinated by various molecular recognition events but the specifics will vary by cell type. For example, some RGC types begin with broad dendritic outgrowth and then prune their arbor to specific sublayers, while others bias dendritic outgrowth to sublayers within the IPL (Kim et al., 2010). Laminar restriction of dendrites and connections is mediated by a battery of attractive and repellent cues that successively ramify processes to narrow layers. They include members of the immunoglobulin superfamily of cell adhesion

270 PART | I Formation of axons and dendrites

FIGURE 12.5 Factors that assemble direction-selective circuits in the mouse retina. (A) Development and laminar targeting of direction-selective synaptic partners in the IPL. i. Starburst amacrine cell dendrites (red) extend into the IPL and form a proto-IPL scaffold that instructs subsequent targeting of ooDSGCs and bipolar cells. ii. ooDSGC target dendrites to the starburst scaffold. iii. ooDSGC dendrites are stabilized and costratify with the ON and OFF starburst sublayers. iv. ON and OFF bipolar cell axons terminate onto respective ON and OFF sublayers. (B) IPL organization of the mature direction-selective circuit. Dendrites of bistratified ooDSGCs (blue) connect to OFF starburst amacrine dendrites (red, upper layer) in S2, and to dendrites of ON starburst cells (red, lower layer) in S4. (C) The starburst dendritic scaffold is necessary for ooDSGC dendrite stratification, as demonstrated by

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molecules (IgSF), cadherins, and semaphorins. IgSF moleculesdSidekick, Dscam, and Contactindmediate homophilic adhesion with minimal binding to related members. They are proposed to bias the connectivity between specific pairs of RGC and amacrine subtypes that cotarget IPL sublaminae (Yamagata and Sanes, 2008, 2012). In support of this idea, misexpression of Sidekick-1, Sidekick-2, Dscam, DscamL, or Contactin-2 in the chick retina redirect dendrites to the sublayer corresponding to the misexpressed molecule. In mice, the Sidekicks and Dscams are dispensable for laminar targeting but mediate other aspects of wiring specificity (see Sections 12.5.4; 12.6.1). Just as dendrites require adhesive cues to target the correct location, they require repellents to exclude them from other locations. Semaphorins are key repellents for IPL stratification. Repulsive signaling between Sema6A localized in the inner ON half of the IPL prevents entry of dendrites of OFF-amacrine subtypes that express PlexinA2 (Matsuoka et al., 2011a; Sun et al., 2013). Sema5A and Sema5B establish a “no-go” zone within the inner cellular layers and prevent neurites expressing Sema receptors Plexin A1 or A3 from extending out of the IPL (Matsuoka et al., 2011b). Thus, an initial sorting of dendrites by no-go cues facilitates targeting to sublaminae. As shown for FLRT and Unc5 receptors in retinal neurons, and for many neuronal receptors in general, recognition molecules cross-bind to other family members for additional selectivity (Visser et al., 2015). The high degree of specificity required during the wiring process will require combinatorial and hierarchical interactions among many recognition molecules.

12.5.3 Transcriptional control of laminar-specific targeting of dendrites in retina Genetic control of dendritic morphology involves multiple layers of regulation. Some aspects of dendrite development, such as dendritic initiation and orientation, may be tied to neuronal birthdate. Other characteristics shared by subsets of cells might be tied to neuronal subtype diversification. On the other hand, dendritic targeting or synaptic partnering decisions likely result from exquisite typeespecific gene regulatory programs. Few transcription factors belonging to the third category have been identified. This is in contrast to progress in Drosophila, for which there is some knowledge of transcriptional factors that determine class-specific features of dendritic arborization neurons (see Chapter 11), or the combinatorial logic of “morphology transcription factors” that dictate distinct patterns of motorneurons (Enriquez et al., 2015). Similar gene regulatory logic will surely be relevant to vertebrate nervous systems. In mouse retina, a recipe for identifying transcriptional programs for type-specific dendrite targeting decisions combines categorization of cell types, tools for genetic access, and expression profiling of purified cell types. Type-specific neuronal morphology may be encoded by combinatorial transcription factor codes. Mouse RGCs can be segregated into >40 distinct subtypes based on transcriptomic signatures, including combinatorial expression of transcription factors (Sanes and Masland, 2015; Rheaume et al., 2018). Combinatorial expression of five transcription factors defines the set of four F-RGCs subtypes, which also differ in dendritic shape, size, and lamination patterns; the roles of this code are unknown (Rousso et al., 2016). Transcription factor T-box brain 1 (Tbr1) marks another set of four RGC subtypes, and in this case, Tbr1 was shown to act as a transcriptional determinant of laminar identity (Fig. 12.6) (Liu et al., 2018). The Tbr1-positive RGC subtypes commonly direct their dendrites to upper OFF layers of the IPL but differ in other molecular markers and dendritic arbor shape. In the absence of Tbr1, these RGCs mistarget their dendrites to lower ON layers, while ectopic expression of Tbr1 in other RGC types redirects their dendrites to upper layers. Other aspects of neuronal identity and dendrite morphology are unchanged. In Tbr1-positive J-RGCs, Tbr1 mediates laminar targeting of JRGC dendrites through the expression of cell surface molecules cadherin-8 and Sorcs3. Cadherin-8 and Sorcs8 are not expressed by the other Tbr1-positive RGC subtypes. Thus, the Tbr1 program controls dendrite targeting to the upper IPL but diverges in individual subtypes to regulate distinct effectors that generate four unique dendritic morphologies. To date, we know few genetic programs that control unique dendritic morphologies but are not essential for fate. Recently, Satb homeobox 1 (Satb1) was discovered to be a key regulator for the bistratified dendritic morphology of ooDSGCs (Peng et al., 2017). In the absence of Satb1, ooDSGCs bear only one dendritic arbor in the OFF layer because they

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genetic ablation of starburst amacrine cells (SACs). (D) In Cadherin 6-9-10 triple mutants, starburst dendrites stratify, but ooDSGC dendrites fail to follow. Homophilic cadherin 6-9-10 adhesion stabilizes ooDSGC dendrites onto starburst dendrites (Duan et al., 2018). (E) Megf10 stabilizes starburst dendrites into a laminar scaffold that guides ooDSGC dendrites. In the absence of Megf10, starburst dendrites form ectopic layers; ooDSGC dendrites costratify with them because adhesive interactions are intact (Ray et al., 2018). (F) In Sema6A mutants, OFF starburst dendrites misproject from S2 to the ON sublayer S4. Normally, Sema6A proteins are expressed by neurites located in the ON IPL, including ON starburst cells, and repel OFF starburst dendrites that express the Sema6A receptors PlexinA2 (Sun et al., 2013). (G) In Satb1 mutants, ooDSGCs are monostratified because they fail to stabilize an ON arbor in S4. Satb1 regulates S4-specific arborization by inducing Contactin 5 expression in ooDSGCs, which in turn mediates homophilic adhesion with Contactin-5-expressing ON starburst dendrites (Peng et al., 2017).

272 PART | I Formation of axons and dendrites

FIGURE 12.6 Dendrite targeting patterns of retinal ganglion cell types. Gallery of genetically defined RGC types with their characteristic IPL stratification profile. (AeD) Four types of RGCs that commonly express transcription factor Tbr1 and target dendrites to upper OFF layers. Each Tbr1þ subtype differs in their precise IPL stratification pattern and other dendritic features (Liu et al., 2018). (DeF) Three alpha (a)-type RGCs bear characteristically large monostratified arbors and cell bodies. The four aRGC subtypes differ in their IPL stratification, molecular signatures, and responses to light (Krieger et al., 2017). OFFa-transient RGC is not shown. (G) ONeOFF direction-selective ganglion cells (ooDSGC) bear bistratified arbors that target S2-OFF or S4-ON layers. (H and I) Two RGC subtypes target the S3 sublaminae, but form distinct intralaminar connections through the expression of Sidekick2 and Sidekick1 adhesion molecules, respectively (Krishnaswamy et al., 2015).

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fail to stabilize branches in the ON layer (Fig. 12.5). Consequently, ooDSGCs fail to receive ON direction-selective information. The ON dendrite targeting errors in Satb1-mutant ooDSGCs result from loss of Satb1-dependent expression of IgSF member Contactin-5 (Cntn5). Cntn5 is required in ooDSGCs and in their synaptic partners, the ON-SACs, to stabilize branches through homophilic adhesion of Cntn5/Casper4 complexes. Defects in Cntn5-deficient ooDSGCs were less severe than in Satb1-mutant cells, suggesting that other Satb1-dependent genes are required for ON arbor patterning. Satb1 has global and gene-specific regulatory functions, and so defining the Satb1-dependent gene network responsible for ON-arbor targeting will yield insights into how dendritic features are genetically encoded and deployed during development.

12.5.4 Local recognition mechanisms to control synapse selectivity Despite laminar patterning mechanisms, most neuronal cell types intermingle their dendrites in dense neuropils that contain diverse potential partners. Even within the mouse IPL, distinct RGC subtypes often arborize to identical sublayers but wire up into distinct circuits (Sanes and Masland, 2015). Therefore, local adhesive and repulsive interactions are required to restrict synaptic formation between appropriate partners. For example, two functionally distinct RGC subsets express either IgSF proteins Sidekick-1 (Sdk-1þ) or Sidekick-2 (Sdk-2þ), but they are morphologically similar and arborize dendrites within the same IPL sublayer (Fig. 12.6). Both Sdk-1þ and Sdk-2þ RGC dendrites intermingle with those of Sdk-2þ amacrine interneurons. Homophilic Sdk2þ interactions strongly bias synaptic connectivity between Sdk-2þ RGCs and Sdk-2þ amacrine cells, compared to mismatched Sdk1þ RGC and Sdk-2þ amacrine cells (Krishnaswamy et al., 2015). Sdk-2 proteins localize to synapses, and Sdk-2þ is required in both synaptic partners. This synaptic matching mechanism is also functionally critical for enhancing the sensitivity of a retinal circuit for detection of small objects and local changes in motion. Together, this study illustrates a molecular strategy: mutual expression of homophilic synaptic adhesion molecules biases synaptic selectivity to neurons pairs that coexist with many other neurons. Selective molecular interactions likely regulate the proportion of types of afferent inputs received by one given neuron, such as observed in convergent innervation. For example, ONa-RGCs are innervated by several bipolar cell types, but the majority of the synapses come from one type (Tien et al., 2017). This suggests a requirement for cues to bias synapse formation to preferred inputs or limit them with minor inputs. If the preferred bipolar inputs are reduced during development by genetic ablation, ON-alpha (ONa) RGC dendrites compensate by forming more connections with the available inputs with no obvious changes to their structure. However, ONa-RGC dendrites favor the recruitment of some bipolar inputs versus others, revealing biases in synapse selectivity. Importantly, this rewiring precisely preserved ONa-RGC function and responses to light. These findings suggest a form of homeostatic plasticity in which postsynaptic dendrites selectively recruit inputs to maintain proper outputs.

12.5.5 An integrated, multistep model for synaptic wiring in the retina IPL Based on the studies described in this section, one can propose a multistep model for the dendritic patterning and synaptic assembly of retinal circuits (Krishnaswamy et al., 2015). First, a proto-IPL generates a scaffold and uses a set of recognition molecules to direct arbors to appropriate sublayers. Repulsive molecules further confine the laminar restriction of dendrites. Within the sublayers, the proximity between inappropriate partners may yield a low rate of connectivity. However, recognition molecules mediate selective interactions to bias connectivity between specific pairings. The partnering decisions that occur during circuit assembly are astoundingly complex, but further studies in models such as the vertebrate IPL will reveal general principles and molecular regulators of wiring specificity.

12.6 Space-filling mechanisms to optimize dendritic field distribution Competitive dendritic interactions between neurons of the same type (homotypic) provide a space-filling mechanism to ensure that the entire field is covered by dendrites of a single neuron type and that sampling is complete but nonredundant. Tiling, mosaic spacing, and dendrite self-avoidance are three homotypic distributions that optimize the spacing of dendritic arbors for uniform sampling of the receptive field.

12.6.1 Tiling and mosaics Tiling refers to an arrangement in which dendritic arbors from the same neuronal type fill the entire receptive surface while minimizing overlap with neighboring homotypic arbors (Fig. 12.7). Other cell types cover their field with extensive dendritic overlap with homotypic neighbors. The degree of dendritic overlap is also described in terms of coverage factor,

274 PART | I Formation of axons and dendrites

FIGURE 12.7 Tiling, mosaics, and self-avoidance. Competitive and repulsive interactions between developing neurites of the same cell type (homotypic) ensure uniform coverage of the receptive field. Variations in the timing and degree of repulsion produce a range of coverage factors and cellular arrangements. (A) Strong contact-dependent repulsion between dendrite tips prevent overlap between arbors, resulting in tiling. (B) Developmentally transient repulsion regulates intercellular spacing (top) but then attenuates, leading to arbor overlap in mature cells (bottom). (C) Mosaic refers to the regular spacing of somas belonging to an individual cell type. Depending on the cell type, mosaic arrays can be generated by homotypic dendritic repulsion and be accompanied by minimal or extensive arbor overlap. (D) In tiling, dendritic arbors of homotypic neurons occupy separate territories with minimal overlap. (E) Dendrite self-avoidance: sibling dendrites emanating from a single neuron distribute evenly across the field. Mutual repulsion between sibling dendrites ensures minimal branch overlap and maximal territory coverage.

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which refers to the number of neurons and their dendritic fields that cover any point in the tissue (Reese and Keeley, 2015). Neurons that tile have a coverage factor of w1 while those that overlap span larger values. For instance, starburst amacrine cells have a dendritic coverage factor of w30 (Keeley et al., 2007), and their arbors fasciculate into a dense plexus. However, their cell bodies are distributed across the retinal plane in a mosaic, which refers to a nonrandom array in which each soma is surrounded by an exclusion zone devoid of cell bodies of the same type (Fig. 12.7) (Lefebvre et al., 2015). Neurons that tile also form mosaics, as tiling leads to a regular soma distribution. Nearly all retinal neuronal subtypes are organized into mosaics. This organizing feature of the vertebrate retina ensures that each point of the image is sampled by a similar complement of microcircuits and their constituents. The regular spacing of retinal neurons and their arbors is established by contact-dependent interactions between homotypic dendrites during development. In live imaging studies of ONa-RGCs in ferret retina, dendroedendritic contacts were detected between ONa-RGCs, but not with those of other neighboring RGCs (Lohmann and Wong, 2001). Manipulations of neuron density by laser or genetic ablation revealed that boundaries between homotypic RGC neighbors are established by contact-dependent repulsion or retraction. First, the ablation of RGCs led to a biased reorganization of dendrites from homotypic neurons into the vacant space (Perry and Linden, 1982; Eysel et al., 1985). In mice, however, the remaining dendritic arbors are unchanged when RCG density is genetically reduced, suggesting that a lack of positive growth cues may also be associated with loss of RGCs (Lin et al., 2004). Second, among perfectly tiled Type 7 cone bipolar retinal cells (Wassle et al., 2009), genetically induced reduction of cell density increases dendrite arbor size of homotypic neurons, and vice versa (Lee et al., 2011). These changes were not influenced by manipulations of afferent activity. The degree of overlap depends on the onset and attenuation of homotypic repulsion (Fig. 12.7). For instance, horizontal retinal cells initially tile via transient processes to position their soma into mosaics before reaching their final laminar position (Huckfeldt et al., 2009). Young starburst dendrites contact each other as they arrive in the IPL and repel to adjust their soma position tangentially (Ray et al., 2018). But mutual inhibition must subsequently be attenuated to allow dendritic arbors to grow over each other. Cell surface molecules MEGF10 and MEGF11 are required for patterning the horizontal and starburst cell mosaics (Kay et al., 2012). Loss of these molecules leads to a random distribution of soma. MEGF10 acts cell autonomously and is proposed to mediate contact-dependent repulsive interactions between transient neurites by mediating homophilic recognition and signaling in trans (Ray et al., 2018). In the mouse retina, Dscams are required for the formation of dopaminergic amacrine and bNOS-positive neuron mosaics (Fuerst et al., 2008). In their absence, mosaics fail to form due to excessive adhesion that causes rearrangement of somas. A transmembrane protein, Dscam1, is proposed to act as a mask to attenuate adhesion among dendrites mediated by cadherin members (Garrett et al., 2018).

12.6.2 Dendrite self-avoidance Dendrite self-avoidance produces a uniform distribution of branches within the neuron’s territory through competitive interactions among dendrites emanating from the same neuron, or “self” dendrites (Fig. 12.7). This process ensures efficient sampling of inputs by a given neuron. Otherwise, neurons receive fewer synapses and expend resources on dendrites devoid of connections. During development, self-dendrites contact and repel each other to prevent overlap but freely interact with dendrites belonging to homotypic but “non-self” neurons. Vertebrate neurons such as starburst amacrine cells and Purkinje cells exhibit dendrite spacing and behaviors (Fujishima et al., 2012) similar to self-avoiding mechanosensory dendrites in C. elegans and Drosophila (Fig. 12.8). How do dendrites recognize and repel self-dendrites? One strategy is the use of ligandereceptor pairs that signal in trans on apposing dendrites to induce contact-dependent repulsion. In starburst cells, the radial distribution and the nonoverlapping arrangement of distal dendrites require transmembrane ligand Sema6A and its receptor PlexA2 (Sun et al., 2013). In cerebellar Purkinje cells, the secreted Slit2 is proposed to anchor to the membrane and bind to its receptor Robo2 on apposing self-dendrites (Gibson et al., 2014). Neurons that overlap extensively with homotypic neighbors, such as starburst cells, require a molecular system for dendrite self-/non-self-discrimination. Starburst cell dendrites not only overlap but fasciculate and form GABAergic synaptic connections with neighboring starburst cells. To achieve this pattern, starburst dendrites discriminate and selectively repel self-dendrites while interacting with non-self-dendrites (Fig. 12.8). The clustered protocadherins (cPcdhs) are a large family of cadherin-like cell surface molecules that mediate self-avoidance and self-/non-self-discrimination (Lefebvre et al., 2012). In mice, the Pcdh locus is organized into three gene clusters alpha (Pcdha), beta (Pcdhb), and gamma (Pcdhg) and encodes 58 isoforms (Wu and Maniatis, 1999). Single Purkinje cells and olfactory sensory neurons express random combinations of Pcdha, Pcdhb, and Pcdhg isoforms (Kaneko et al., 2006; Mountoufaris et al., 2017).

276 PART | I Formation of axons and dendrites

FIGURE 12.8 Dendrite self-/non-self-avoidance is mediated by the clustered protocadherins. (A) Retinal starburst amacrine cells develop radial dendritic arbors through self-avoidance, but overlap and form reciprocal connections with neighbors. This pattern requires dendrite self-/non-selfdiscrimination, which is mediated by unique combinations of clustered protocadherin molecules (cPcdhs) on neighboring cells. (B) Dendrite self-/nonself-discrimination model: Self-dendrites (dark red) bear the same cPcdh molecule (orange) and therefore recognize and self-avoid each other. Self-dendrites overlap with non-self-dendrites (blue) expressing different cPcdhs (magenta). (C) The clustered Protocadherin gene loci include the 3 Pcdhalpha, -beta, and egamma genes, which encode 14, 22, and 22 isoforms. Pcdha and Pcdhg mRNA transcripts are produced from splicing an alternate variable exon to 3 constant exons. One variable exon is transcribed upon promoter choice. Pcdha6 mRNA is shown as an example. The variable exon encodes the extracellular region, composed of 6 cadherin-like extracellular domains (EC), the transmembrane (TM) and juxtamembrane domains. The 3 constant exons encode the intracellular region (ICR), common in all Pcdh-as or Pcdh-gs isoforms. (D) Subsets of cPcdh isoforms interact in cis to produce unique cPcdh combinations. Matching cPcdh combinations interact in trans to produce repulsion for self-avoidance, or adhesion in other contexts (Molumby et al., 2016). (E) Wild-type cerebellar Purkinje cell (PC on left, blue) exhibit dendrite self-avoidance (inset), whereas dendrites self-overlap in a Pcdh-gamma mutant PC (orange) (Lefebvre et al., 2012). (F) Similar to a wild-type starburst cell (black), self-avoidance is not affected in the absence of Pcdh-alpha (blue). Pcdh-gammas are essential for starburst dendrite self-avoidance, as demonstrated by branch crossings and clumps in Pcdhg mutant cells (orange). Dendritic morphology defects are more severe in double Pcdh-alpha/-gamma mutant starburst cells (green). Thus, cPcdhs from different clusters cooperate in complex, redundant ways to pattern starburst cells (Ing-Esteves et al., 2018).

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Thus, combinatorial expression of cPcdhs might endow each neuron a unique cPcdh identity, generating an extraordinary scale of recognition diversity sufficient for self-/non-self-discrimination. Pcdhgs are essential for dendrite self-avoidance in starburst cells and Purkinje cells (Lefebvre et al., 2012). In genetic manipulations that rendered all starburst cells to express an identical Pcdhg isoform, self-avoidance was intact but neighboring starburst cells recognized each other as self and failed to form synapses (Lefebvre et al., 2012; Kostadinov and Sanes, 2015). Since GABAergic connections between starburst cells are critical for direction selectivity, the loss of Pcdhg diversity perturbed the retinal responses to moving stimuli (Kostadinov and Sanes, 2015). Moreover, Pcdhg and Pcdha members interact with each other in complex and redundant ways, as illustrated by the severe effects of removing Pcdha and Pcdhg on dendrite self-avoidance (Fig. 12.8) (Ing-Esteves et al., 2018). In the cortex, Pcdhgs promote cortical dendritic branching and stabilization through homophilic and adhesive activities (Molumby et al., 2016). Thus, cPcdh isoform diversity is critical for organizing complex dendrite arrangements and circuit functions. Regulation of dendrite self-avoidance by the cPcdhs parallels Drosophila Dscam1-mediated self-avoidance (Chapter 11; (Lefebvre et al., 2015)). Although the vertebrate Dscam genes do not undergo splicing diversity they mediate avoidance of homotypic dendrites by masking cell surface molecules such as cPcdhs and cadherins (Fuerst et al., 2008; Garrett et al., 2018). Intracellular signals that mediate dendrite self-avoidance remain to be elucidated. They could regulate self-avoidance receptor delivery, such as has been observed for LKB1-SIK-dependent trafficking of Robo2 to Purkinje dendrites (Kuwako and Okano, 2018), or regulate actin to modulate branch protrusion and retraction (Kawabata Galbraith et al., 2018).

12.7 Emergence of dendrite compartmentalization To perform local operations and process different streams of information, dendrites develop exquisite patterns of synaptic and ion membrane channels within separate compartments. Dendrites are molecularly specialized at multiple scales, from the entire arbor to the single spine. For example, the apical arbors of hippocampal pyramidal neurons are subdivided into distal and proximal domains that receive distinct inputs and have independent signal processing capacities (Spruston, 2008). Synaptic and electrical properties are patterned along the length of single branches to perform local computations that influence input signals (Branco and Hausser, 2010). Development of dendritic compartmentalization is particularly understudied. Here we describe a few known examples of synaptic and ion channel patterning along dendritic branches of the hippocampal apical arbor.

12.7.1 Subcellular patterning of synaptic inputs along dendritic domains Hippocampal pyramidal neurons receive distinct streams of inputs that innervate nonoverlapping domains of distal and proximal apical dendrites. Targeting of inputs onto specific dendritic domains is accomplished by selective adhesion to connect correct inputs and by repulsive cues to exclude incorrect inputs (Lefebvre et al., 2015). Selective adhesion is mediated by the complementary actions of two Netrin-G/NGL receptoreligand pairs (Kim et al., 2006). Binding of GPIlinked Netrin-G1 to NGL-1 sorts axonal afferents from the entorhinal cortex onto the distal apical dendrites, while NetrinG2/NGL-2 interactions connect axonal afferents from hippocampal CA3 to the proximal dendritic compartment of CA1 pyramidal neurons. Netrin-G/NGL do not direct axon targeting: in the absence of netrin-G1 or -G2, afferents properly target their dendritic compartments but the cognate NGL proteins are ectopically distributed across the dendritic arbor (Nishimura-Akiyoshi et al., 2007). Moreover, as trans-synaptic adhesion molecules known to organize postsynaptic proteins (Kim et al., 2006), Netrin-G and NGL, are proposed to nucleate formation of compartment-specific excitatory synapses. Therefore, axonal afferents bearing netrin-G1 or netrin-G2 target their dendritic domain and there might mobilize their cognate NGL-1 or NGL-2 receptor to organize synapse formation in the distal and proximal domains, respectively. In support of this model, the absence of NGL-2 disrupts excitatory synaptic transmission and spine density in the proximal, but not distal, apical dendrites (DeNardo et al., 2012). Thus, input-derived adhesive cues provide one mechanism to pattern dendritic compartments to receive certain synapses. Repulsive actions by semaphorins regulate the subcellular patterning of synaptic inputs to dendritic domains. Hippocampal CA3 cells express tethered Sema6A throughout their dendritic arbors, but its activity is spatially attenuated in the proximal dendrites through cis-interactions with its receptor PlexinA2 (Suto et al., 2007). However, mossy fiber afferents express another Sema6A receptor, PlexinA4, which causes repulsion from other dendritic domains. Therefore, mossy fiber afferents are confined to a proximal dendritic domain due to local PlexinA2-dependent attenuation of Sema6A activity, and repulsion by flanking dendritic domains. In a second example, a secreted semaphorin establishes a gradient of spine density along dendritic branches. Secretion of Sema3F restricts the formation of excitatory synapses in proximal dendritic regions, resulting in a decreased spine density toward the soma (Tran et al., 2009). Interestingly, Sema3F signaling might also be

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spatially restricted by enriched localization of its Neuropilin-2 receptor to zones closest to cell bodies. Both examples illustrate that spatial control of subcellular trafficking of membrane proteins is critical for patterning dendritic domains.

12.7.2 Patterning the membrane excitability of dendritic compartments Establishing dendrite compartmentalization requires mechanisms that spatially control the trafficking and insertion of membrane proteins to dendritic domains. This problem has been explored in CA1 neurons, which develop a graded distribution of HCN1 channels toward the distal apical dendrites. Dissociation of hippocampal neurons abolished the distal dendritic enrichment of HCN1 channels, suggesting that the gradient is established by a local extrinsic cue (Kupferman et al., 2014). HCN1 targeting to distal dendrites requires Reelin signaling in dendrites, resulting from diffusible Reelin activating its cytosolic counterpart Disabled-1 (Dab1) and Src family tyrosine kinases. Conditional deletion of Dab1 from CA1 neurons abolishes the graded distribution of HCN1 and other distally targeted ion channel proteins such as GIRK1 (Kupferman et al., 2014). As a result, the electrical properties of distal dendrites are altered, which likely disrupts hippocampal circuit functions since the loss of HCN1 is known to inappropriately strengthen local inputs and impair synapse formation (Nolan et al., 2004). Estrogen is another extrinsic factor that promotes HCN1 enrichment (Meseke et al., 2018). How extrinsic cues accomplish HCN1 localization is not known, but it does not appear to regulate general trafficking or expression of HCN1 channels. One attractive model for establishing dendritic compartmentalization involves extrinsicdependent patterning of dendrites that locally organize the local molecular and cell biological specializations for compartment-specific protein regulation and trafficking.

12.8 Neurodevelopmental disorders: the price of poor dendritic development? Deficits that impede dendrite development can have long-lasting consequences on neuron and circuit function and can thereby contribute to the pathogenesis of neurodevelopmental disorders (NDDs). Dendritic anomalies are prominent clinical features across NDDs (Kulkarni and Firestein, 2012). For example, progressive dendrite atrophy is observed in postmortem brains of individuals with Down syndrome (Becker et al., 1986) and reduced dendrite branching in schizophrenia and autism spectrum disorders (ASDs) (Raymond et al., 1996; Broadbelt et al., 2002). Modeling NDD-associated gene candidates from human genetic studies using human-induced pluripotent stem cells (hiPCS) and animal models are powerful entry points for investigating relationships between dendritic defects and NDD disease pathogenesis. Modeling NDD-associated dendrite pathology is relatively simpler for monogenic diseases or those associated with defined syntenic chromosomal regions. For example, Timothy syndrome is caused by a point mutation in CACNA1C, encoding a calcium channel subunit. Cortical neurons derived from Timothy syndrome patient IPSCs show increased activity-dependent dendrite retraction (Krey et al., 2013). Ectopic activation of RhoA, independent of Ca2þ influx, is implicated in this process and can be suppressed by overexpressing the GTPase GEM. In a mouse model of Angelman syndrome, defined by a deficiency in the E3 ubiquitin ligase gene Ube3a, pyramidal neurons fail to develop a polarized morphology (Miao et al., 2013). In William’s syndrome, defined by hemizygosity for w26 genes on chromosome 7q11.23, hiPSC-derived cortical neurons exhibit higher total dendritic length and number (Chailangkarn et al., 2016). Similar dendritic anomalies are observed in cortical pyramidal neurons in postmortem William’s syndrome brains. Other disease models used to study dendritic anomalies include the MeCP2-deficient Rett syndrome model, the Fmr1-deficient Fragile X syndrome, and the chromosome 21 trisomy Down syndrome model (Kulkarni and Firestein, 2012). For polygenic diseases, another strategy is to focus on single, NDD-associated genes with known roles in dendrite development to uncover links between altered dendritic connectivity and pathogenesis. Dyrk1A is located on the human chromosome 21 interval with three copies in Down syndrome, but gene disruptions have been also identified in human ASD and intellectual disabilities (Park and Chung, 2013; van Bon et al., 2016). Dyrk1A encodes a kinase that regulates a myriad of processes, including microtubule and actin dynamics, and disease-associated mutations lead to dendritic branching and connectivity defects (Dowjat et al., 2007; Lepagnol-Bestel et al., 2009; Dang et al., 2018). TAOK2, a gene located in the 16p11.2 chromosomal susceptibility region for ASD, schizophrenia (SCZ), and other NDDs, encodes a kinase that regulates actin reorganization and basal dendrite complexity in mouse cortical neurons (Weiss et al., 2008; Richter et al., 2018). Cntnap2, a gene disrupted in ASD, SCZ, and ID stabilizes dendrites of inhibitory neurons via interaction with CASK, a cytoskeletal scaffold protein (Poot, 2015; Gao et al., 2018). The levels and activity of members of the collapsing response mediator protein family (CRMPs), which function in dendrite initiation and outgrowth, are also altered in patients and animals models from various NDDs (Quach et al., 2015). These studies demonstrate that alterations in disease-associated genes affect the structure of dendritic arbors and may provide insights into circuit alterations. The integration of experimental models will help translate human genetic findings into biological understandings of the connectivity defects that underlie cognitive dysfunction and developing brain disorders.

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12.9 Conclusion Dendritic arborizations form from the coordinated development of features such as dendrite size, shape, branch complexity, targeting, and synaptic and ion channel patterning. Exquisite patterning of these features requires the complex integration of extrinsic and intrinsic information that are received by cell type-specific repertoires of receptors and effectors. We are just beginning to uncover pathways that pattern individual features, but have much more to do to understand how they are coordinated during development. Advances in gene expression profiling and genetic access to cell types in intact nervous systems are greatly facilitating the study of vertebrate neurons at the single-cell resolution. Most examples discussed in this chapter come from studies in the mouse vertebrate retina, a model with a simpler architecture, well-characterized cell types and transgenic markers to probe them. The sophisticated toolkit available in the mouse retina has led to the discovery of developmental strategies and molecular pathways that regulate type-specific dendritic patterning, and their importance for circuit function. Similarly, new findings come from well-studied cells in the brain such as the cortical pyramidal, stellate and cerebellar Purkinje neurons. While studies in a handful of cell types will yield general principles and molecular pathways, we anticipate that a similar framework will be applied to other neuronal populations and circuits. Cellular taxonomy, single cell and spatial transcriptomics, microscopy, and genetically encoded reporters are rapidly advancing the state of the field. Together, this research will not only tell us how neurons develop their unique dendritic morphology, but also how they compute information and contribute to circuit function.

Abbreviations DTIPs Plus-end tracking proteins AMPA-R a-Amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid receptor, an ionotropic transmembrane receptor for glutamate BDNF Brain-derived neurotrophic factor BTBD3 BTB (Broad-complex, tramtrack and Bric a brac) domain 3 CaMK Calmodulin-dependent protein kinases Cdc42 Rho-GTPase cell division cycle 42 cPcdhs Clustered protocadherins Dscam Down syndrome cell adhesion molecule Ena/VASP proteins Enabled/Vasodilator-stimulated phosphoprotein F-actin Filamentous actin FAK/Pyk2 Focal adhesion kinase/protein tyrosine kinase 2 FLRT Fibronectin leucine rich transmembrane protein GABA Gamma-aminobutyric acid GAPs GTPase activating proteins GCL Granule cell layer GEFs Guanineenucleotide exchange factors GPI-linked Glucose-6-phosphate isomerase HCN1 Hyperpolarization-activation cyclic nucleotide-gated channel 1 hiPCS Human-induced pluripotent stem cells hippocampal CA1/3 Hippocampal cornu ammonis 1/3 IgSF Immunoglobulin superfamily of cell adhesion molecules INL Inner nuclear layer IPL Inner plexiform layer J-RGC Junction adhesion molecule B retinal ganglion cell Lhx2 LIM (Lin-11, Islet-1, Mec-3) homeobox 2 MAPs Microtubule-associated proteins MEGF10, 11 Multiple epidermal growth factor-like domains 10 or 11 mTORC1 and mTORC2 Mammalian target of rapamycin complexes 1 and 2 NMDAR N-methyl-D-aspartate receptor NR1 N-methyl-D-aspartate receptor subunit 1 NT-3 Neurotrophin-3 ONa-RGCs ON-alpha retinal ganglion cells ooDSGCs ONeOFF direction-selective RGCs Rac1 Ras-related C3 botulinum toxin substrate 1 RGCs Retinal ganglion cells Satb1 Special AT-rich binding protein 1 Sdk-1D and 2D Sidekick cell adhesion molecule 1 and 2

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Tbr1 T-box brain 1 Tiam1 T-lymphoma invasion and metastasis 1, a Rac1-specific GEF WASP WiskotteAldrich syndrome protein family WAVE WiskotteAldrich syndrome protein (WASP)/verprolin homologue WIRS A short consensus motif WRC WAVE regulatory complex

Acknowledgments We thank Bing Ye, and Lefebvre lab members Madison Gray and Samantha Esteves for comments, and Wendy Xueyi Wang for the figures and illustrations. J.L.L. is supported by a Canada Research Chair Tier 2.

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Signaling mechanisms underlying reversible, activity-dependent dendrite formation. Neuron 34, 985e998. van Bon, B.W., et al., 2016. Disruptive de novo mutations of DYRK1A lead to a syndromic form of autism and ID. Mol. Psychiatry 21, 126e132. van Spronsen, M., Mikhaylova, M., Lipka, J., Schlager, M.A., van den Heuvel, D.J., Kuijpers, M., Wulf, P.S., Keijzer, N., Demmers, J., Kapitein, L.C., Jaarsma, D., Gerritsen, H.C., Akhmanova, A., Hoogenraad, C.C., 2013. TRAK/Milton motor-adaptor proteins steer mitochondrial trafficking to axons and dendrites. Neuron 77, 485e502. Visser, J.J., Cheng, Y., Perry, S.C., Chastain, A.B., Parsa, B., Masri, S.S., Ray, T.A., Kay, J.N., Wojtowicz, W.M., 2015. An extracellular biochemical screen reveals that FLRTs and Unc5s mediate neuronal subtype recognition in the retina. eLife 4, e08149. Wang, C.F., Hsing, H.W., Zhuang, Z.H., Wen, M.H., Chang, W.J., Briz, C.G., Nieto, M., Shyu, B.C., Chou, S.J., 2017. Lhx2 expression in postmitotic cortical neurons initiates assembly of the thalamocortical somatosensory circuit. Cell Rep. 18, 849e856. Wang, Y., Rubel, E.W., 2012. In vivo reversible regulation of dendritic patterning by afferent input in bipolar auditory neurons. J. Neurosci. 32, 11495e11504. Wassle, H., Puller, C., Muller, F., Haverkamp, S., 2009. Cone contacts, mosaics, and territories of bipolar cells in the mouse retina. J. Neurosci. 29, 106e117. Wayman, G.A., Impey, S., Marks, D., Saneyoshi, T., Grant, W.F., Derkach, V., Soderling, T.R., 2006. Activity-dependent dendritic arborization mediated by CaM-kinase I activation and enhanced CREB-dependent transcription of Wnt-2. Neuron 50, 897e909. Weiss, L.A., et al., 2008. Association between microdeletion and microduplication at 16p11.2 and autism. N. Engl. J. Med. 358, 667e675. Wu, G.Y., Cline, H.T., 1998. Stabilization of dendritic arbor structure in vivo by CaMKII. Science 279, 222e226. Wu, G.Y., Zou, D.J., Rajan, I., Cline, H., 1999. Dendritic dynamics in vivo change during neuronal maturation. J. Neurosci. 19, 4472e4483. Wu, Q., Maniatis, T., 1999. A striking organization of a large family of human neural cadherin-like cell adhesion genes. Cell 97, 779e790. Yalgin, C., Ebrahimi, S., Delandre, C., Yoong, L.F., Akimoto, S., Tran, H., Amikura, R., Spokony, R., Torben-Nielsen, B., White, K.P., Moore, A.W., 2015. Centrosomin represses dendrite branching by orienting microtubule nucleation. Nat. Neurosci. 18, 1437e1445. Yamagata, M., Sanes, J.R., 2008. Dscam and Sidekick proteins direct lamina-specific synaptic connections in vertebrate retina. Nature 451, 465e469. Yamagata, M., Sanes, J.R., 2012. Expanding the Ig superfamily code for laminar specificity in retina: expression and role of contactins. J. Neurosci. 32, 14402e14414. Yau, K.W., Schatzle, P., Tortosa, E., Pages, S., Holtmaat, A., Kapitein, L.C., Hoogenraad, C.C., 2016. Dendrites in vitro and in vivo contain microtubules of opposite polarity and axon formation correlates with uniform plus-end-out microtubule orientation. J. Neurosci. 36, 1071e1085. Yau, K.W., van Beuningen, S.F., Cunha-Ferreira, I., Cloin, B.M., van Battum, E.Y., Will, L., Schatzle, P., Tas, R.P., van Krugten, J., Katrukha, E.A., Jiang, K., Wulf, P.S., Mikhaylova, M., Harterink, M., Pasterkamp, R.J., Akhmanova, A., Kapitein, L.C., Hoogenraad, C.C., 2014. Microtubule minus-end binding protein CAMSAP2 controls axon specification and dendrite development. Neuron 82, 1058e1073. Yu, W.Q., El-Danaf, R.N., Okawa, H., Pacholec, J.M., Matti, U., Schwarz, K., Odermatt, B., Dunn, F.A., Lagnado, L., Schmitz, F., Huberman, A.D., Wong, R.O.L., 2018. Synaptic convergence patterns onto retinal ganglion cells are preserved despite topographic variation in pre- and postsynaptic territories. Cell Rep. 25, 2017-2026 e2013. Yu, X., Malenka, R.C., 2003. 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Chapter 13

Cell polarity and initiation of migration K. Hayashi, K. Sekine, H. Tabata and K. Nakajima* Keio University School of Medicine, Tokyo, Japan *Corresponding author.

Chapter outline 13.1. Introduction 289 13.2. Migratory behaviors during radial migration in the developing cerebral cortex 290 13.2.1. Bipolar migrating neurons along the radial glial fibers: locomotion 290 13.2.2. Radial glial fibereindependent mode of migration: somal translocation and terminal translocation 290 13.2.3. Multipolar migration 292 13.2.4. Transformation from multipolar migrating neurons to bipolar locomoting neurons 293 13.2.5. Departure from the ventricular zone: differences in migratory behavior between direct progeny of the apical progenitors in the VZ and the basal progenitors in the subventricular zone /intermediate zone 294

13.2.6. Behaviors of the progenitor cells in the subventricular zone 13.3. Molecular mechanisms that regulate the initiation of migration and cell polarity during migration 13.3.1. Coupling between neuronal differentiation and migration 13.3.2. Controlling the initiation of radial migration 13.3.3. Regulation of multipolar migration 13.3.4. Extracellular molecules that affect migrating cells 13.4. Conclusion See also List of abbreviations Glossary Supplementary data References

296 297 298 299 299 301 301 301 301 302 302 302

13.1 Introduction One of the features of neurons in the developing central nervous system is that they migrate away from their birthplaces to their final destinations. One of the best-described mechanisms of neuronal migration is the radial migration of projection neurons in the developing cerebral cortex. Neurons are generated in the ventricular zone (VZ) or the subventricular zone (SVZ), migrate radially through the intermediate zone (IMZ) to reach the cortical plate (CP), and finish their migration beneath the marginal zone (MZ). The combination of classical studies, such as Golgi staining and electron microscopy (EM) analyses, and the more recent studies involving time-lapse analyses has revealed that, during the course of neuronal migration, the neurons dynamically change their migratory behaviors. In the current model, the majority of neurons that are born in the VZ during the early embryonic stage exit the VZ using a migratory mode called somal translocation (Section 13.2.2); these cells then transform into multipolar cells that exhibit a unique behavior called multipolar migration (Section 13.2.3). In the late embryonic stage when the CP develops, multipolar migrating neurons transform into bipolar cells before entering the CP (Section 13.2.4), where the neurons then move toward the MZ using a migratory mode called locomotion (Section 13.2.1). Finally, these neurons change their migration mode to terminal translocation and reach beneath the MZ (Section 13.2.2). In contrast, the pyramidal neurons in the Ammon’s horn of the hippocampus migrate in a climbing mode (Kitazawa et al., 2014; Hayashi et al., 2015) after undergoing multipolar-tobipolar transition (see Chapter 17. Migration in the hippocampus). Some progenitors move to the SVZ or IMZ, where they are called basal progenitors or intermediate progenitors, and produce neurons (Sections 13.2.5 and 13.2.6). Analyses of human diseases and the advancement of molecular biology techniques have also revealed the key molecules regulating these morphological changes in neuronal migration (Section 13.3). In this chapter, we introduce the historical

Cellular Migration and Formation of Axons and Dendrites. https://doi.org/10.1016/B978-0-12-814407-7.00013-4 Copyright © 2020 Elsevier Inc. All rights reserved.

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debates concerning neuronal migration, the current models of morphological changes during the initial phase of radial neuronal migration, and the molecular mechanisms that underlie the initiation and the polarity regulation of migration in the developing cerebral cortex.

13.2 Migratory behaviors during radial migration in the developing cerebral cortex 13.2.1 Bipolar migrating neurons along the radial glial fibers: locomotion An important feature of the developing cerebral cortex is the “inside-out” arrangement of projection neurons; that is, laterborn neurons settle more superficially than earlier-born neurons in the CP. Consequently, later-born neurons need to migrate a much longer distance than earlier-born neurons. Considering that some earlier-born neurons are already differentiated and have already started to develop dendrites when later-born neurons are passing through the region of these differentiated neurons, later-born neurons are thought to use a specialized mode of migration. With this in mind, Rakic (1972) proposed a highly sophisticated mode of neuronal migration. Using serial-section EM findings, he showed that, during the late stages of fetal monkey development, when the thickness of the cerebral wall was more than 3000 mm, the entire length of a bipolar migrating neuron was attached to the ascending fibers of specialized cells called radial glial cells (RGCs). Rakic proposed that neurons migrate along these radial glial fibers (RGFs). This mode of migration along RGFs was well accepted and was directly observed once time-lapse imaging was developed. For example, Nadarajah et al. (2001) observed neuronal migration in the early embryonic mouse cortex (E13e14) using a combination of slice cultures and time-lapse analyses. By labeling the sections with Oregon Green BAPTA AM, they showed that bipolar migrating neurons with a short (30e50 mm) leading process did not attach to the pial surface. The length of the leading process remained almost constant, with fluctuations of up to 12%. The authors reported that these cells were often flattened on one side, suggesting that they may be apposed to an RGF. Since these morphological features resembled the features of RGF-guided migrating neurons that had been reported on fixed sections, they named this mode of movement “locomotion.” The locomoting cells moved in a saltatory manner; that is, intervals of no movement occurred between periods of rapid forward movement. RGCs are defined primarily by their morphology; that is, they span the entire thickness of the developing cerebral wall, from the ventricular surface to the pial surface, and possess subcellular structures that resemble those of astrocytes. RGCs have also been shown to function as neurogenic progenitor cells (Miyata et al., 2001; Noctor et al., 2001; Tamamaki et al., 2001). Thus, locomoting neurons are born from RGCs and use RGFs as a scaffold for migration.

13.2.2 Radial glial fibereindependent mode of migration: somal translocation and terminal translocation While this important concept of locomotion along the RGFs has been widely observed and established, several reports have also suggested the existence of RGF-independent migration. For example, based on an analysis of serial EM sections, Shoukimas and Hinds (1978) argued that neurons did not associate with RGFs during the early stages of cortical development. Early Golgi preparations of rat embryonic cortex by Berry and Rogers (1965) showed the presence of “binucleate” cells in the VZ with connections to both the ventricular surface and the pial surface. The authors proposed that the cytokinesis of these ventricular cells might be preceded by the movement of one daughter nucleus to the CP through the long, radially oriented basal (or pia-directed) process. Morest (1970) studied neurogenesis in the forebrain of opossum pouch young using rapid Golgi preparations. The author found that, when the cerebral cortex was just starting to form, most of the primitive epithelial cells extended from the ventricular surface to the pial surface and suggested that the nuclei of these cells moved after the internal process (apical process or process toward the ventricle) had been withdrawn. Thus, the migration of these cells involves the movement of the nucleus through the external process (basal process or process toward the pial surface) in a manner that is likely independent of the RGFs. These cells retained their attachment to the pial surface through their external processes, whereas their internal processes lost their attachments to the ventricular surface and formed retraction bulbs. This type of migration was named “perikaryal translocation.” Perikaryal translocation seems to end just beneath the MZ. This type of migration was suggested to require a very short time for completion, perhaps a matter of minutes or at most hours. Additional evidence for RGF-independent migration was reported by Brittis et al. (1995). These authors used the monoclonal antibody 2G12, which recognizes not only mitotic cells but also the early neuronal (nonmitotic) population. In rats, 2G12-positive cells were observed just beneath the preplate at E15. These cells were nonmitotic and positive for the

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neuronal marker Tuj1. When the E15 cortex was labeled with DiI from the pial surface, labeled cells with a long, unbranched radial shape could be identified. Some cells had detached their ventricular end feet but still retained a pial process, with the cell bodies approaching the CP. This morphology suggests that many young neurons move radially via nuclear translocation. They also reported that, at E16, most of the labeled cells were located in the IMZ and exhibited a locomoting cell morphology, whereas some cells were located just above the VZ and exhibited a tangentially oriented axon-like process. These data suggest that neurons may change their migratory behavior during the developmental course, and nuclear translocation may be the main migratory mode during the early stages. However, both experiments lacked direct evidence of nuclear movement. Nadarajah et al. (2001) directly observed RGF-independent neuronal migration using time-lapse imaging of cultured mouse cortical slices (E13eE14). These authors reported that the cell soma moved toward the pial surface, whereas the long leading process (60e95 mm) remained attached to the pial surface and became thicker and progressively shorter. They referred to this mode of migration as somal translocation, rather than nuclear translocation or nucleokinesis, because the entire cell body is translocated (Fig. 13.1 and Video Clip 1). The translocating cells often had a short trailing process directed toward the VZ, but not attached to the ventricular surface. The leading process was often branched, and the soma of the translocating cells changed their shape as they moved toward the MZ, possibly because of the distortion of the cell body as it squeezed between other cellular structures, and the soma moved rapidly up to the branch point of the leading process. Some of the translocating cells were calbindin-positive neurons. The dynamics of somal translocation and locomotion were markedly different. The translocating cells moved continuously, and the soma moved at a relatively constant pace (1e3 mm min
1) without pause. On the other hand, the locomoting cells moved in a saltatory manner, as mentioned previously. Because of the pauses, the average speed of locomotion was significantly slower (35 mm h
1) than that of the translocating cells (60 mm h
1). These observations provided the first direct evidence that somal translocation was actually a frequently observed migratory mode, especially during the early stage of cortical development. Another type of radial glial fibereindependent migration is terminal translocation, which is observed in the late stage of cortical development (Sekine et al., 2011). When the somata of the migrating neurons in the locomotion mode arrived beneath the primitive cortical zone (PCZ), which is the outermost region of the CP composed of densely packed immature neurons (Sekine et al., 2011, 2012), and the tips of the leading processes of migrating neurons reached the MZ, the neurons left the RGFs and changed their migration mode from locomotion to terminal translocation. During the terminal translocation, only the somata moved rapidly over short distances through the PCZ toward the top of the CP, whereas the tips of the processes remained attached to the MZ. Miyata et al. (2001) also showed that postmitotic neurons left the VZ by somal translocation during the early stage of cortical development. They studied the fate of the RGFs using DiI labeling of the E14 mouse cerebral wall. A time-lapse analysis of the DiI-labeled dividing RGCs in the VZ showed that the RGF was asymmetrically inherited by one of the daughter cells without retraction or degeneration and maintained its attachment to the pial surface both before and after cell division. The daughter cells that inherited the ascending fibers of the RGCs became postmitotic neurons, whereas their

(A)

Pial surface

(B) CP

IMZ

100

VZ

FIGURE 13.1 (A) A schematic diagram of somal translocation. A neuronal soma moves toward the pial surface, whereas the leading process keeps attached to the pial surface. (B) Morphology of a somal translocation cell visualized using in utero electroporation of GFP. Note that the basal process (arrowheads) of a neuron exhibiting somal translocation (arrows) keeps in contact with the pial surface. CP, cortical plate; IMZ; intermediate zone; VZ, ventricular zone. Scale bar: 100 mm.

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siblings became the next generation of the RGCs. The daughter neurons left the VZ by somal translocation once their descending processes lost contact with the ventricular surface. What triggers the initiation of migration, and what kinds of mechanisms regulate the somal translocating neurons once they leave the VZ? Morest (1970) proposed the idea that the detachment of the descending process from the ventricular surface was the initial event of neuronal migration and differentiation. Shoukimas and Hinds (1978) also supported this idea using an EM serial-section analysis of the early histogenesis of the mouse cerebral cortex. They also found that the pair of centrioles was located within the most distal part of the ventricular process in the undifferentiated cells in the VZ, whereas these centrioles moved toward the soma of the neurons in the IMZ. During “locomotion,” centrosome-dependent “nuclear pulling” is thought to be an important regulator (see Chapter 14). In the case of somal translocation, however, nuclear pulling was unlikely because centrosomes were located in the trailing (descending or ventricular) process. Miyata and Ogawa (2007) noticed that the pial process of the somal translocating neuron appeared to be twisted and proposed a “twist-plus-stretch” model. By microsurgically transecting the ventricular attached process, they observed the initiation of somal translocation through the contraction of the pia-attached process (basal process), similar to a spring. They also found that intermediate filaments, but not microfilaments or microtubules, in the pia-attached process were the major factors for the basis of this spring-like force. These data suggest that the intermediate filament-mediated twisting of pial processes produces a spring-like force that functions to propel the cell soma away from the VZ.

13.2.3 Multipolar migration In contrast to the bipolar or monopolar morphology of cells that migrate using locomotion or somal translocation, histological analyses of fixed sections of developing cerebral cortex have demonstrated the presence of multipolar cells in the lower IMZ/SVZ. Stensaas (1967) analyzed the morphologies of the migrating neurons in the fetal rabbit cerebral cortex using Golgi staining. He reported that the IMZ (which he originally described as the intermediate lamina) contained two kinds of zones that differed in cell density and the orientation of the nuclei of the migrating neurons. The inner zone was dark and contained cells with obliquely oriented nuclei, whereas the outer zone was light and contained elements with radially oriented nuclei. He also reported a difference in the morphologies of the neurons in these two zones, with the migrating neurons in the inner IMZ exhibiting a stellate soma and a horizontal or vertical axon with many short processes, while the migrating neurons in the outer IMZ exhibited an oblique to vertical soma with an axon descending and turning near the VZ and with a single preapex (or leading process) arising from the opposite side of the axon. Shoukimas and Hinds (1978) studied the detailed morphologies of migrating neurons using a combination of EM serial-section analyses and autoradiography. These authors then reported the morphological changes of migrating neurons in the IMZ/SVZ. Just above the VZ, migrating neurons had a short descending process that extended only a short distance into the VZ, suggesting that these neurons had just left the VZ. Many neurons observed in the inner IMZ (and lower SVZ) had one tangentially oriented long process and many shorter processes. The long process was thought to be the primitive axon. Below the CP, the migrating neurons had one long tangentially oriented process and one pia-directed process extending into the CP. Using autoradiography, they suggested that these neurons became cortical neurons at a later stage. They also showed that the position of centrioles varied depending on the location of the cell within the IMZ. At the bottom of the IMZ (or lower SVZ), the pair of centrioles was located at the ventricular side of the nucleus in cells that had a relatively simple and radially oriented morphology. On the other hand, no consistent centriole location was observed in cells with multiple processes. In the migrating neurons just beneath the CP (or in the upper IMZ), the pair of centrioles was located at the base of the pia-directed radial processes. Multipolar cells with numerous short processes were also observed on fixed sections using virus vectors expressing green fluorescent protein (GFP) (Noctor et al., 2001; Tamamaki et al., 2001). The radially oriented bipolar or monopolar morphology of locomotion or somal translocation cells could not account for the presence of the large proportion of multipolar cells in the lower IMZ/SVZ, and the means by which these cells moved was not known. Tabata and Nakajima (2001, 2008) developed the technique of in utero electroporation and directly showed the dynamic behavior of multipolar cells in the lower IMZ/SVZ by combining this technique for mouse embryonic cortex with time-lapse analyses of slice cultures (Tabata and Nakajima, 2003). By introducing the GFP expression plasmid at E14.5, they were able to visualize the morphology of migrating neurons. At 36 h after electroporation, many of the GFP-positive neurons within the lower IMZ/SVZ exhibited a multipolar morphology (Fig. 13.2). Most of the thin processes extended independently from the RGFs, suggesting that the multipolar cells were not associated with RGFs. The authors analyzed the behavior of multipolar neurons using time-lapse analyses and showed that multipolar neurons did not have a fixed cell polarity but, rather, extended and retracted thin processes in various directions in a very dynamic manner (Video Clip 2).

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(B)

(A)

CP IMZ SVZ MAZ VZ FIGURE 13.2 Morphology of multipolar migrating cells. (A) A lower magnification view of multipolar migrating cells. Electroporation of GFP was performed at E14.5, and the brain was examined at E16. Note that multipolar cells are accumulated in the MAZ (just above the VZ). CP, cortical plate; GFP, green fluorescent protein; IMZ, intermediate zone; SVZ, subventricular zone; MAZ, multipolar cell accumulation zone; VZ, ventricular zone. Scale bar: 100 mm. (B) A higher magnification view of multipolar migrating cells.

Many of the multipolar cells remained within a narrow band above the VZ (multipolar cell accumulation zone, MAZ (Tabata et al., 2009), which overlaps with the lower part of the SVZ; see the following text) for a considerable time period (w1 day) and, in the lower IMZ, migrated slowly toward the pial surface and sometimes jumped tangentially. Since the multipolar neurons moved in various directions and changed direction frequently, the mean net change in their positions in the lower IMZ was 2.2 mm h
1, which is consistent with the in vivo migration speed reported by Takahashi et al. (1996). Because the movement of multipolar neurons resembled neither locomotion nor somal translocation, they termed this movement “multipolar migration” (Tabata and Nakajima, 2003). Although these cells are morphologically “multipolar,” it is likely that the “multipolar” migrating cells do not have a fixed polarity, since the position of the centrosome was not constant, as mentioned above. What is the biological meaning of multipolar migration? Multipolar migrating neurons in the MAZ begin to extend a tangentially oriented long process that is thought to be a primitive axon (Shoukimas and Hinds, 1978; Stensaas, 1967; Tabata et al., 2009). Indeed, Namba et al. (2014) demonstrated that the axons extending from multipolar cells established contact with the preexisting axons of early-born neurons through a cell surface molecule TAG-1 (transient axonal glycoprotein-1) and that this interaction was required for the cellular polarization of multipolar cells. Considering that multipolar cells extend and retract thin processes dynamically, these cells may be searching for environmental cues related to the direction of axon growth. Another feature of multipolar migrating cells is that they can move in tangential directions, as described previously (Tabata and Nakajima, 2003). Using triple ephrin-As knockout mice and in utero electroporation, Torii et al. (2009) showed that the tangential dispersion of migrating neurons was crucial for the integration of cortical neurons in the proper radial columns and that this dispersion depended on the ephrin-AeEphA system. The flexible movement of multipolar migrating neurons, including the “tangential jump,” may also partly contribute to enabling a passage through the lower IMZ without requiring a specific scaffold for migration. This possible mechanism could be important, since several obstacles to radial migration exist in the IMZ, such as bundles of tangential axon fibers. Further study is needed to elucidate how multipolar migration is regulated and what role this movement plays during cortical development.

13.2.4 Transformation from multipolar migrating neurons to bipolar locomoting neurons Now that three modes of neuronal migration in the developing cerebral cortex (locomotion, somal translocation, and multipolar migration) have been identified, the next question concerns the relationship among these migratory behaviors; that is, are these mechanisms independent of each other and used by separate neuronal populations, or do the transition from one mode to another occur, with the same neurons using multiple modes during the course of migration? Although Tabata and Nakajima (2003) observed that multipolar cells accounted for the major population of the GFPlabeled migrating cells in the lower IMZ and SVZ/MAZ, they did not find multipolar migrating cells in the CP, where most of the migrating cells exhibited a locomotion morphology. One possible explanation for this observation is that these

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locomoting cells in the CP might have migrated by locomotion all the way from the cortical VZ, which comprises a population independent of the multipolar cells in the lower IMZ and SVZ/MAZ. Another possibility is that the locomoting CP cells might have derived from multipolar IMZ/SVZ/MAZ cells. If the latter is true, the multipolar cells must have transformed into locomoting cells before entering the CP. When Tabata and Nakajima investigated the morphology of the migrating cells in the lower IMZ/SVZ/MAZ of a mouse cortex in detail, they found only a small population of cells exhibiting locomotion. In addition, the multipolar cells generally ultimately migrated toward the pial surface in time-lapse analyses, expressed neuronal markers, exhibited a locomoting-cell-like bipolar morphology beneath the CP, and mostly were not apoptotic, as revealed by TUNEL (terminal transferase dUTP nick end labeling) staining. Based on these results, the authors proposed that the multipolar cells entered the CP as bipolar locomoting cells (Tabata and Nakajima, 2003). To systematically analyze the relationship between the cellular identity and the mode of migration across the entire thickness of the cerebral wall, Hatanaka et al. (2004) compared the position and morphology between cortical neurons derived from E12.5 (early-born neurons) and from E15.5 (late-born neurons) mouse VZ cells, both of which had been labeled with GFP-expression plasmids using in utero electroporation. Since somal translocation was reportedly observed frequently during the early stages of cortical development (Nadarajah et al., 2001), the early-born neurons were expected to leave the VZ by somal translocation. Above the VZ (IMZ or preplate), the authors observed that many GFP-positive neurons had a multipolar morphology, whereas the labeled cells in the VZ assumed a bipolar shape at E13.5 (24 h after electroporation). When the morphology of early-born neurons was examined at E14 (40 h after electroporation), the GFP-positive neurons were found in the CP and were oriented radially, extending a thick process that reached the pial surface and had a thin axon-like process running through the IMZ. These observations suggest the possibility that the earlyborn neurons leave the VZ as somal translocation, assume a multipolar morphology, and transform into radially oriented bipolar cells as they migrate into the CP. On the other hand, the late-born neurons derived from E15.5 VZ exhibited a multipolar morphology above the VZ at 1 day after electroporation. Two days after electroporation, some labeled cells with a multipolar morphology were located in the lower IMZ, but other cells with a bipolar shape were seen in the upper IMZ. These neurons possessed a short and thick process extending toward the pial surface and an axon-like process running through the IMZ. These bipolar neurons were apposed to the RGFs. These observations suggest that late-born neurons may change their morphology from a multipolar morphology to a bipolar shape during the course of migration. However, these observations did not provide direct evidence that a single neuron really changes its mode of migration after leaving the VZ. Noctor et al. (2004) and Tabata et al. (2009) directly observed the transformation from multipolar migration to locomotion on cultured cortical slices after the injection of GFP-expressing retrovirus vectors into the rat embryonic cortex or by in utero electroporation in the mouse embryonic cortex, respectively. They analyzed the migratory behaviors using time-lapse analyses of cortical slices and showed that the majority of multipolar migrating cells transform into locomoting cells before entering the CP. The quantitative data using long-term slice cultures, however, may not reflect the in vivo situation exactly. Therefore, in vivo quantitative analyses of the migratory behaviors were next performed (Tabata et al., 2009), which are summarized and discussed in the next section.

13.2.5 Departure from the ventricular zone: differences in migratory behavior between direct progeny of the apical progenitors in the VZ and the basal progenitors in the subventricular zone /intermediate zone Although multipolar migrating cells were initially described as the neurons that become postmitotic in the VZ, neuronal progenitor cells exist not only in the VZ but also in the SVZ (and the lower IMZ). Smart (1973) confirmed the presence of mitotic figures away from the ventricular surface of the developing cerebral wall, an observation that was originally made in the 19th century. Several reports followed, describing these cells as a secondary proliferative population, nonesurfacedividing cells, basal progenitors, or intermediate progenitors (Englund et al., 2005; Haubensak et al., 2004; Miyata et al., 2004; Noctor et al., 2004; Takahashi et al., 1995a,b). Although this population was previously reported as being mainly comprised of glial progenitors, Noctor et al. (2004) directly showed that the majority of these cells (about 70%, at least) produced a pair of neurons in slice cultures of late-embryonic rat cerebral wall. A “basal progenitor” in the SVZ is thought to originate from an “apical progenitor” in the VZ and to produce a pair of neurons apart from the ventricular surface. Indeed, the SVZ was originally defined as the region containing proliferative cells that are not attached to the ventricular surface and do not show interkinetic nuclear movement during the mitotic cycle (Boulder Committee, 1970). Despite the aforementioned general view of the SVZ as a secondary proliferating zone, multipolar cells in the “SVZ” were shown to be basically postmitotic (Tabata and Nakajima, 2003). To elucidate the relationship between multipolar migrating cells and basal progenitors in the SVZ, the migratory behaviors of “SVZ” cells and their mitotic activity in vivo were examined in detail (Tabata et al., 2009).

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Tabata et al. (2009) labeled VZ cells with GFP using in utero electroporation at E15 and further applied thymidine analogs to distinguish the postmitotic cells in all VZ-derived GFP-positive cells in E15eE16 mouse cerebral cortex (late stage of cortical development). At 12 h after electroporation, the major population of GFP-positive cells was still located in the VZ, but some GFP-positive cells had rapidly exited the VZ and were found in the “SVZ” or IMZ and exhibited a somal translocation morphology. At 36 h after electroporation, most of the GFP-positive postmitotic cells had accumulated just above the VZ (76% in the dorsomedial cortex and 92% in the lateral cortex) and showed the typical morphology of multipolar migrating cells. Time-lapse analyses using slice cultures confirmed that, during the initial phase of migration, the direct progenies of VZ cells stayed in the VZ for more than 10 h, mainly assuming a pin-like morphology with an apical process attached to the ventricular surface, and then accumulated rather specifically just above the VZ as multipolar migrating cells. Ochiai et al. (2007) also observed in an E14 cortical slice that some VZ cells with a pin-like morphology directly became neurons without further mitosis and exhibited a multipolar morphology above the VZ. The application of thymidine analogs in vivo showed that many of the somal translocation cells observed at 12 h after electroporation on E15 in the upper part of the “SVZ” or IMZ had mitotic activities (58%). Immunostaining for mitotic markers at 36 h after electroporation also showed that a high proportion of GFP-positive cells within the upper part of the SVZ/IMZ were mitotically active (62%), whereas most of the multipolar migrating cells observed above the VZ or the lower part of the SVZ were postmitotic (76%) (Tabata et al., 2009). These multipolar cells remained just above the VZ without further mitosis for almost 24 h, transformed to locomoting neurons with a bipolar morphology before entering the CP, and finally differentiated into layer II/III pyramidal neurons in vivo. The zone where the multipolar cells accumulated above the VZ was dubbed the multipolar cell accumulation zone (MAZ) (Tabata et al., 2009), which overlaps with the lower part of the SVZ. These data also indicated that there are two distinct populations that migrate from the VZ during the late stage of cortical development (E15eE16 in mice). One population completes mitosis in the VZ and mainly exhibits a pin-like morphology. These cells then slowly migrate into the MAZ, where they accumulate as postmitotic multipolar cells. The other population migrates into the SVZ/IMZ quickly via somal translocation and undergoes further cell division. Because these populations differ with regard to the timing of their exit from the VZ, Tabata et al. described them as a “slowly exiting population (SEP)” and a “rapidly exiting population (REP),” respectively (Fig. 13.3). The REP includes the basal progenitors, which divide symmetrically to produce multipolar migrating neurons. It is also possible that the REP includes the glial progenitor cells, because some of them are positive for Olig2 (7.6%). Whereas the REP enters the SVZ/IMZ earlier than the SEP, the SEP transforms into locomoting cells and enters the CP earlier than the REP (Tabata et al., 2009). The major means of the basal progenitor production may change during the course of development. Miyata et al. (2004) examined the behavior of the basal progenitors using DiI labeling and time-lapse analyses of E13e14 mouse cortical slices (early stage of cortical development) (Fig. 13.4). The basal progenitors were mainly observed in the SVZ and also in the lower IMZ. When an RGC asymmetrically divided in the VZ at E13e14, one of the daughter cells inherited the RGF and the other daughter cell exhibited a pin-like morphology. Both daughter cells left the VZ through the retraction of the apical (ventricular) process. As described previously, the former cells exhibited a somal translocation morphology, and most of them (about 80%) were postmitotic neurons (Miyata et al., 2001), whereas the others produced pairs of neurons (Miyata et al., 2004). Ochiai et al. (2007) observed a direct transition from somal translocation to multipolar migration without mitosis in E14 cortical slices. On the other hand, most of the pin-like morphology cells (about 70%) underwent subsequent mitosis and

Somal Translocation oRG

SVZ

IMZ MAZ VZ

REP SEP

FIGURE 13.3 Differences of migratory profiles between direct progeny of the apical progenitors in the VZ and the basal progenitors in the IMZ/SVZ in the late stage of cortical development. After asymmetrical division of a radial glial cell (RGC) in the VZ, one daughter cell rapidly leaves the VZ with a somal translocation morphology (REP, magenta) and the other cell stays for more than 10 h in the VZ and slowly enters the MAZ (SEP, green). The main population of the REP becomes the basal progenitors in the IMZ/SVZ or the oRG cells (pink), but some cells differentiate into postmitotic neurons without further mitosis (orange dashed line). The REP may also include glial progenitor cells. On the other hand, the SEP mainly differentiates into postmitotic multipolar cells without further mitosis, whereas some of the SEP cells may still have mitotic activity (purple dashed line).

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SVZ

IMZ

MAZ VZ

FIGURE 13.4 Morphological changes around the initial phase of radial migration in the early stage of cortical development. When a radial glial cell (RGC, blue) asymmetrically divides, one of the daughter cells inherits the basal process (green) and assumes a somal translocation morphology, and the other becomes a pin-like morphology cell (magenta). The somal translocation cells mainly become postmitotic multipolar neurons without further mitosis, whereas some become basal progenitors (purple dashed line). The majority of pin-like morphology cells become basal progenitors, whereas some pin-like morphology cells differentiate into postmitotic multipolar neurons without further mitosis (orange dashed line).

produced pairs of neurons (Ochiai et al., 2007), whereas others directly transformed to multipolar migrating neurons without further mitosis. These data suggest that the pin-like morphology cells are the main source of the basal progenitors during this early stage. Thus, especially during the early stage of cortical development (E13e14), somal translocation cells could directly become postmitotic neurons, and the pin-like morphology cells could become the basal progenitors. However, as mentioned previously, in the late stage of cortical development (E15e16 in mice), the majority of somal translocation cells in the REP undergo further cell division (at least 60%), and the pin-like morphology cells in the SEP become multipolar migrating neurons without further mitosis (76%e92%). Studies on the regulation of these processes during development are needed in the future. Like somal translocation, the centrosomes of pin-like morphology cells were located at the tip of the retracting ventricular process, suggesting that the centrosome-dependent “nuclear pulling” model proposed for locomotion cannot be applied to the departure of pin-like morphology cells (Ochiai et al., 2007). However, the “twist-plus-stretch” model of the “spring-like” pia-attached process may also be not applicable, since the pin-like morphology cells do not have a pia-attached process. Thus, how these pin-like morphology cells leave the VZ remains unknown. One possible mechanism is an active force mediated by the actomyosin system on the ventricular side of the soma, similar to the migration of interneurons (Bellion et al., 2005). Another model is the passive pushing out model mediated by the interkinetic movement of adjacent RGCs. Okamoto et al. (2013) reported that retraction of the neocortical progenitors’ basal processes by knockdown of TAG-1 molecule resulted in the failure of interkinetic nuclear migration. This migratory deficit caused overcrowding of the progenitors at the apical surface, leading to mislocation of the progenitors out of the VZ and disruption of the histogenesis. This suggests that the apical progenitors may sense mechanical stress. Further study is needed to uncover the mechanisms responsible for the initiation of the migration of pin-like morphology cells.

13.2.6 Behaviors of the progenitor cells in the subventricular zone Whereas the REP cells are defined by the migratory profiles of the direct progeny of the apical progenitors in the VZ, recent histological studies and time-lapse analyses have revealed the presence of a novel type of neuronal progenitor in the outer subventricular zone (OSVZ) of the human and ferret neocortices (Hansen et al., 2010; Fietz et al., 2010). The developing human SVZ has a massively expanded OSVZ, and the increase in the OSVZ cells is thought to be one of the major evolutionary events for the neocortical expansion (Fish et al., 2008). Hansen et al. (2010) revealed that, in the developing human OSVZ, there are many radial gliaelike neuronal progenitors with a basal process attached to the pial surface. Interestingly, these progenitors lack an apical process that reaches the ventricular surface, unlike the RGCs. These progenitors are shown to be PAX6/SOX2/HES1-positive and have the ability of both proliferative and self-renewing asymmetric divisions to generate the daughter neuronal progenitors that can further proliferate. However, they are negative for TBR2, suggesting that these progenitors are different from the basal progenitors. The authors referred to this new progenitor type as the oRG (OSVZ radial glia-like) cells. Similar neuronal progenitors were also observed in the ferret neocortex (OSVZ progenitors of ferret) (Fietz et al., 2010; Gertz et al., 2014). Interestingly, even in the mouse SVZ, a similar neuronal progenitor was reported that shares the cellular morphology and marker expression patterns with the human oRG cells (Wang et al., 2011) (or outer VZ progenitors of mouse) (Shitamukai et al., 2011). However, the majority of the human and ferret oRG daughter cells (both basal and apical daughter cells) still become nonneurogenic progenitors

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with proliferative activity (Gertz et al., 2014). In the case of the mouse oRG, most of the apical daughter cells derived from the oRGs become Pax6-negative differentiated cells, whereas the basal daughter cells that inherit the basal processes are Pax6-positive proliferative cells (Wang et al., 2011). From these data, it would be reasonable to assume that the expansion of the SVZ is limited in the mouse neocortex. The migratory behavior of these progenitors is characteristic. The cell nuclei move rapidly up into the basal process before cytokinesis, and most of them divide asymmetrically with a horizontal cleavage plane. The upper (superficial/basal) daughter inherits the basal process and maintains the oRG morphology. This migratory behavior was termed “mitotic somal translocation” (Hansen et al., 2010). It is interesting that not only the morphological features but also the migratory behaviors of the oRG cells/OSVZ progenitors/outer VZ progenitors are similar to the REP cells in the mouse neocortex. Although the original report of Tabata et al. (2009) showed that the REP cells could become the basal progenitors after they retract the basal process, it is also likely that the REP cells have the potential of asymmetric self-renewing cell division with maintaining the basal process (Shitamukai et al., 2011). Future study will be needed to elucidate the molecular mechanisms how the oRG/REP cells are produced from the RGCs in the VZ, how the oRG/REP cells migrate to the SVZ, how the oRG/REP cells are maintained in the SVZ, and how the oRG/REP cells differentiate to neurons and the basal progenitors to understand the molecular basis of human neocortical expansion.

13.3 Molecular mechanisms that regulate the initiation of migration and cell polarity during migration Recently, the molecular mechanisms that underlie the early phase of neuronal migration, including the initiation of migration, the regulation of multipolar migration, and the transition of the migration mode, have been intensely studied. Many key regulators have been identified through the positional cloning of mutations in the human cortical malformation, the analyses of genetically engineered or mutant mice, and the in vivo analysis of neuronal migration by the introduction of RNA interference (RNAi)emediated loss-of-function, dominant-negative, and gain-of-function approaches using in utero electroporation (Fig. 13.5).

MARK2/Par1 Rac1

Lis1 JNK p35 Dcx

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p27

RhoA

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FIGURE 13.5 Molecular mechanisms that regulate the morphological changes and the initial phase of migration.

Mst3

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13.3.1 Coupling between neuronal differentiation and migration The regulation of neuronal differentiation is thought to be associated with the regulation of the cell cycle of the neuronal progenitors. In the developing cerebral cortex, the G1 phase of the cell cycle plays a crucial role in determining when the neuronal progenitors undergo cell cycle exit and neuronal differentiation. During the period of neurogenesis, which peaks at around E14 in mice, the G1 length in the progenitors increases, and this change is correlated with an increase in cell cycle exit (Takahashi et al., 1995a,b). G1 progression is promoted by the cyclin-dependent kinase (CDK) 4/cyclin D and CDK6/cyclin D but is also kept in check by CDK inhibitors (CKIs) (Sherr and Roberts, 2004). Two families of CKIs are known: the Cip/Kip family, including p21Cip1, p27Kip1, and p57Kip2, and the INK4 family, including p15Ink4b, p16Ink4a, p18Ink4c, and p19Ink4d. While the INK4 proteins specifically target CDK4/cyclin D and CDK6/cyclin D during G1, the Cip/ Kip proteins inhibit a broader spectrum of CDKecyclin complexes. In p27Kip1 knockout mice, a decrease in neuronal production occurs during midcorticogenesis, and an increase in the production of late-born neurons is observed, resulting in an enlargement of the upper cortical layers (Goto et al., 2004). Conversely, overexpression of p27Kip1 in the cortical progenitors at E12e14 results in a reduction in the number of upper-layer neurons (Tarui et al., 2005). Furthermore, p27Kip1 is also shown to be involved in neuronal differentiation and migration in the developing cerebral cortex independent of its cyclin-binding capacity. p27Kip1 is the predominant Cip/Kip protein in the developing cerebral cortex and is distributed mainly in the cytoplasm of SVZ/MAZ/IMZ neurons. Nguyen et al. (2006) showed that overexpression of p27Kip1 increased the number of differentiated neurons, whereas knockdown of p27Kip1 impaired neuronal differentiation. The effects of p27Kip1 on neuronal differentiation were independent of its roles in cell cycle regulation, since the cell cycle mutant version of p27Kip1 that could not bind to cyclins and CDKs also promoted neuronal differentiation. The role of p27Kip1 in neuronal differentiation is, at least in part, mediated by the stabilization of the proneural transcription factor neurogenin-2 (Ngn2). Supporting this mechanism, Ngn2 overexpression rescued the neuronal differentiation defect elicited by p27Kip1 knockdown. These data indicate that the cell cycle inhibitor p27Kip1 promotes neuronal differentiation by stabilizing Ngn2 without affecting the cell cycle. p27Kip1 is also reportedly involved in neuronal migration. Kawauchi et al. (2006)reported that Cdk5, which is a key regulator of corticogenesis and is activated in postmitotic neurons (Gilmore et al., 1998, and see Section 13.3.3), bound and phosphorylated p27Kip1 at Ser10 and protected p27Kip1 against degradation. Knockdown of p27Kip1 resulted in neuronal migration failure. p27Kip1 knockdown neurons did not exhibit the typical multipolar morphology in the lower IMZ/MAZ but instead had a relatively round morphology with thin processes. Dominant-negative Cdk5 also caused similar phenotypes. In the upper IMZ, p27Kip1 knockdown neurons transformed to bipolar cells but could not migrate into the CP. The accumulation of p27Kip1 suppressed the phosphorylation of the actin-binding protein cofilin through the suppression of RhoA. These data indicate that the interaction between Cdk5 and p27Kip1 regulates neuronal migration through the regulation of the actin cytoskeleton. Another Cip/Kip family member, p57Kip2, is also expressed in migrating neurons and is involved in neuronal migration (Itoh et al., 2007), further underscoring the overlapping but distinct roles that CKIs play in neuronal differentiation and migration. Concurrent with the final cell division, the neural progenitors differentiate into neurons, and this process is determined by several transcription factors, including the proneural basic helixeloopehelix (bHLH) transcription factor neurogenin-2 (Ngn2), and is discussed extensively (Rubenstein and Rakic, 2013). In Ngn2 knockout mice, the migration of pyramidal neurons is severely affected, and the neurons cannot enter the CP. However, whether this migration failure is directly caused by the loss of Ngn2 or indirectly by other means, such as the upregulation of Mash1 in the dorsal telencephalon (which is normally expressed in the ventral telencephalon), remains unclear. Hand et al. (2005) showed that Ngn2 controlled neuronal migration independently of its proneural transcriptional activities. They showed that the acute ablation of Ngn2 through the introduction of Cre expression vectors to the conditional allele of Ngn2 caused migration failure in the IMZ/SVZ/MAZ. They also found that a tyrosine at position 241 in Ngn2 was phosphorylated in vivo, and the mutation of this tyrosine residue (Ngn2Y241F) preserved its proneural effect. Importantly, the expression of Ngn2Y241F in cortical neurons had a dominant-negative effect and nearly abolished the radial migration of cortical neurons. One of the reasons for this dominant-negative effect on migration was that this mutant Ngn2 could not recruit the Ngn2-associated transcription coactivator because of the missing phosphorylation site. The authors also showed that the cotransfection of dominant-negative RhoA with Ngn2Y241F rescued the migration failure induced by Ngn2Y241F. Ge et al. (2006) showed that expression of Ngn2 sequestered the transcriptional coactivator CBP (CREB-binding protein) from the endogenous RhoA promoter in the neural progenitor cells and inhibited RhoA expression, resulting in the promotion of neuronal migration. On the other hand, the tyrosine 241 mutant of Ngn2 could not suppress expression of RhoA, because this mutant Ngn2 could not recruit the CBP from the RhoA promoter. These data suggest that the redistribution of transcription coactivators seems to be critical for the regulation of migration-related gene expressions by neurogenic bHLH factors. Ngn2 also regulates neuronal migration directly through the induction of the small G protein Rnd2 before the initiation of migration. Heng et al. (2008) showed that Ngn2 directly bound to the consensus-binding sites for Ngn2 (known as E-

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box) in the 30 -enhancer of the Rnd2 gene in cortical cells in vivo. Rnd2 expression was first detected in the VZ, peaked in the SVZ/MAZ/IMZ, and sharply decreased in the CP, suggesting that Rnd2 is transiently expressed in migrating neurons. Knockdown of Rnd2 in the migrating neurons increased the abnormally located neurons in the IMZ, similar to the conditional Ngn2-null mutant neurons, and the Rnd2 knockdown neurons showed an abnormal multipolar morphology with abnormally long neurites. Overexpression of Rnd2 in conditional Ngn2-null mutant neurons partially rescued the migration failure. Rnd2 expression was also maintained by other bHLH transcription factors, such as NeuroD (which is also a direct downstream molecule of Ngn2). These data suggest that a single proneural protein controls the complex cellular behavior of cell migration through a direct pathway involving the transcriptional activation of a small G protein. On the other hand, the transcription repressor RP58 suppresses the Ngn2eRnd2 pathway (Ohtaka-Maruyama et al., 2013). Rp58 itself regulates the multipolar-to-bipolar transition. Migrating neurons in the RP58 knockout mouse showed severe defects in the formation of the leading processes and the multipolar-to-bipolar transition. It was also shown that this migratory defect in the knockout mouse was rescued by knockdown of Ngn2. As Ngn2 is known to activate RP58 (Seo et al., 2007), RP58 functions as a negative feedback regulator of Ngn2.

13.3.2 Controlling the initiation of radial migration Periventricular heterotopia (PH) is an X-linked human brain malformation in which subsets of neurons fail to migrate into the developing cerebral CP and remain as nodules of neurons that line the ventricular surface. Filamin A is known to be the causative gene (Fox et al., 1998). This phenotype suggests that mutations in filamin A disrupt the initiation of radial migration from the VZ. Filamin A is known to regulate the F-actin network. While filamin A protein is distributed in the CP and IMZ, its mRNA is expressed throughout the developing cortex, suggesting that filamin A protein is degraded in the VZ (Nagano et al., 2002). Nagano et al. (2002) searched for molecules that were preferentially expressed in the VZ and that were related to the initiation of migration and found that FILIP (filamin A-interacting protein) was expressed only in the VZ. FILIP interacted with filamin A and induced the degradation of F-actin-associated filamin A in a calcium-dependent manner in the VZ. The degradation of filamin A caused by overexpression of FILIP in the migrating cortical neurons resulted in their round morphology and suppressed radial migration below the CP, suggesting that an adequate amount of filamin A is important for the early phase of migration. Filamin A is also shown to be important for the regulation of cell shape in the early phase of radial migration (Nagano et al., 2004). Overexpression of a mutant filamin A lacking the actin-binding domain affected the stage of multipolar migration and inhibited radial migration. Transfected cells exhibited an abnormal round morphology in the SVZ/MAZ and the lower IMZ and could not transform into the bipolar shape of locomoting cells, similar to the results for overexpression of FILIP. On the other hand, filamin A overexpression by FILIP knockdown changed the multipolar morphology to a radially oriented bipolar morphology in the SVZ/MAZ and IMZ. These data suggest that filamin A helps migrating neurons to determine their mode of migration, either multipolar or bipolar, before entering the CP. Mitogen-activated protein kinases (MAPKs) are intracellular signal transduction molecules expressed in all eukaryotic cells and modulate basic cellular events such as cell proliferation, death, migration, and differentiation (Morrison and Davis, 2003). MAPKs are activated via signaling cascades involving MAPK kinases (MAPKKs), which in turn are activated by MAPK kinase kinases (MAPKKKs). MEKK4 is one of w17 MAPKKKs cloned from mammalian cells (Gerwins et al., 1997). MEKK4 knockout mice develop specific central nervous system phenotypes, including not only severe neural tube closure defects and massive neuroepithelial apoptosis but also PH. Sarkisian et al. (2006) showed that the formation of PH in MEKK4 knockout mice arose as a result of neuronal migration defects. Knockdown of MEKK4 in migrating cortical neurons also resulted in migration failure in the IMZ. One of the downstream molecules of MEKK4, MKK4/SEK1 (an MAPKK), interacts with and phosphorylates filamin A, and the protein level of filamin A was elevated in MEKK4 knockout mice. These results suggest that the level of filamin A is regulated by not only FILIP but also the MEKK4-MKK4/SEK1 pathway and is crucial for the proper initiation of migration.

13.3.3 Regulation of multipolar migration Dynamic morphological changes are associated with the rearrangement of cytoskeletons in migrating neurons. Kawauchi et al. (2003) revealed the important roles of the small G protein Rac1 in the morphological changes of migrating cortical neurons. Dominant-negative forms of Rac1 or the Rac1 guanine nucleotide exchanging factor STEF/Tiam1, both of which are expressed in migrating cortical neurons, caused a defect in radial migration. The transfected cells exhibited a round morphology with short and irregular processes in the IMZ and could not transform into bipolar locomoting neurons. The activation of JNK (Jun-N-terminal kinase), which was shown to regulate

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microtubule dynamics through the phosphorylation of MAP1B, was decreased in the transfected neurons. These data indicate that the reorganization of microtubules plays pivotal roles in the transformation of multipolar migrating neurons to bipolar locomoting neurons. Humans with hemizygous deletions of 17p13.3 have disorders known as isolated lissencephaly sequence (ILS) or MillereDieker syndrome (MDS). These disorders are characterized by type I or classical lissencephaly (agyria/pachygyria), a human brain developmental disorder manifested by smooth brain surfaces and disorganized cortical layering. The causative gene for type I lissencephaly in MDS is Lis1 (or Pafah1b1, the b subunit of the platelet-activating factor acetylhydrolase isoform Ib) (Hattori et al., 1994; Reiner et al., 1993). The cell biology of Lis1 and its roles in neuronal migration have been extensively studied. Lis1 interacts with dynein, dynactin, Ndel1, and Nde1 to regulate nuclear movement and cell migration through the regulation of microtubules. Mouse knockout models show that the graded reduction of the Lis1 encoding gene Pafah1b1 affects developmental processes including cell division and neuronal migration (Hirotsune et al., 1998). Tsai et al. (2005) studied the functions of Lis1 in neuronal migration in the wild-type environment using an RNAi system. As a result, Lis1 knockdown neurons could not transform from multipolar migrating neurons to bipolar locomoting neurons and stopped their migration mostly in the VZ/SVZ/MAZ, suggesting that Lis1 is required for exit from the stage of multipolar migration. The number of processes from the soma of Lis1 knockdown neurons during the multipolar stage was not significantly different from that of the control neurons, but the processes of the Lis1 knockdown neurons were more branched. The axon-like processes observed in the multipolar stage were also shorter and somewhat curved and branched in the Lis1 knockdown neurons. Mutations in the X-linked gene doublecortin (DCX) are the most common genetic cause of subcortical band heterotopia or double-cortex syndrome in females and a major cause of lissencephaly in males. DCX encodes doublecortin (des Portes et al., 1998; Gleeson et al., 1998), which regulates the dynamics of microtubules in a phosphorylation-dependent manner. Dcx knockout mice surprisingly do not show disruptions in neocortical lamination (Corbo et al., 2002), probably because of genetic compensation from doublecortin-like kinases (Deuel et al., 2006; Koizumi et al., 2006). On the other hand, Bai et al. (2003). showed that the acute ablation of Dcx by RNAi in rats disrupted radial migration and caused the formation of subcortical band heterotopia. Dcx knockdown neurons exhibited a multipolar morphology mainly in the IMZ, suggesting that Dcx is required for the transition from multipolar migration to locomotion. Cdk5 is a unique serine/threonine kinase with a close homology to other Cdks, but its kinase activity is mainly detected in postmitotic neurons. The association of Cdk5 with a neuron-specific regulatory subunit, either p35 or its isoform p39, is essential for the kinase activity. Cdk5 and its activator p35 have important roles in the formation of cortical structures in the developing mouse brain, and this is brought about mainly by the regulation of neuronal migration (Chae et al., 1997; Gilmore et al., 1998). Ohshima et al. (2007) found that Cdk5 had important functions in the multipolar-to-bipolar transition during radial migration in a study using live imaging in Cdk5 knockout mice and a dominant-negative approach in a normal environment. Ngn2 also reportedly upregulated the transcription of p35 (Ge et al., 2006), and activated Cdk5 stabilized p27 in a phosphorylation-dependent manner (discussed earlier). Furthermore, p27 regulates the stabilization of Ngn2 in the VZ/SVZ/MAZ (see Section 13.3.1), suggesting a positive feedback loop between Ngn2, Cdk5/p35, and p27. On the other hand, Tao et al. reported that RapGEF2 was phosphorylated downstream of Cdk5 and regulated the multipolar-to-bipolar transition via the Rap1-N-cadherin pathway (Ye et al., 2014). Besides, a serine/threonine kinase, Mst3, regulated the multipolar-to-bipolar transition in a Cdk5-phosphorylation-dependent manner through inhibition of RhoA (Tang et al., 2014). Knockdown of Mst3 in the migrating neurons by in utero electroporation suppressed the migration of the neurons with multipolar morphology in the IMZ, which was recovered by coexpression of the phosphomimic form of Mst3 (at the cdk5 phosphorylation site) or a knockdown vector for RhoA. Cdk5 may regulate neuronal polarity via multiple pathways. The Par (partition defective) family proteins are highly conserved regulators of cell polarity and were discovered while studying Caenorhabditis elegans cell polarity regulation (Kemphues et al., 1988). Directed neuronal migration and the establishment of a mature neuronal morphology are also highly polarized processes. Sapir et al. (2008) studied the function of the polarity kinase MARK2/Par-1 in neuronal migration. Reduced MARK2 levels using in utero electroporation, which affected microtubule dynamics, resulted in the stalling of multipolar migrating neurons at the IMZ border and affected the centrosomal dynamics in locomoting neurons. Increased MARK2 levels also changed the migrating neurons to a round shape. MARK2 is known to phosphorylate Dcx at serine 47 in the leading process of migrating neurons and to affect microtubule dynamics (Schaar et al., 2004), whereas Cdk5/p35 phosphorylates Dcx at serine 297 in the soma (Tanaka et al., 2004). Dcx is also phosphorylated by JNK (Gdalyahu et al., 2004). These data suggest that cytoskeletal rearrangement involving microtubules is crucial for the transition from a multipolar morphology to a bipolar morphology and the establishment of cell polarity.

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13.3.4 Extracellular molecules that affect migrating cells The secreted glycoprotein Reelin is produced mainly from the CajaleRetzius cells in the MZ of the developing cerebral cortex. However, a small amount of Reelin was also observed around the MAZ (Hirota et al., 2015). In addition, expression of one of the Reelin receptors, ApoER2 (low-density lipoprotein receptor-related protein 8 [Lrp8]) was also observed around the MAZ during cortical development. Jossin and Cooper (2011) showed that the ReelineRap1 pathway controlled neuronal polarization in the IMZ through regulating the surface expression of N-cadherin. They demonstrated that the forced expression of Rap1GAP in the migrating neurons in the developing mouse cortex using in utero electroporation caused migratory defect of the neurons below the SP and that this phenotype was rescued by overexpression of N-cadherin. Furthermore, transfection of a C-terminus-deletion mutant of VLDLR, another Reelin receptor, into migrating neurons also resulted in migratory failure, accompanied by defective cellular polarization during multipolar-to-bipolar transition. This defect was rescued by coexpression of both the constitutively active form of Rap1 and wild-type Akt1. Dab1 is known to be phosphorylated in Reelin signaling and functions as a hub protein in this signaling. Recently, it was also reported that DCC (deleted in colorectal carcinoma), a Netrin-1 receptor, also interacted with Dab1 and regulated the multipolar-to-bipolar transition of cortical migrating neurons (Zhang et al., 2018). Netrin-1 indeed induced phosphorylation of Dab1 at tyrosines 220/232 through DCC and Fyn kinase. Knockdown of DCC by in utero electroporation caused suppression of multipolar-to-bipolar transition, which was rescued by coexpression of the phosphomimic form of Dab1 (Y220D/Y232D), although Netrin-1 was mainly detected in the MZ (Stanco et al., 2009). Another Netrin-1 receptor, Unc5D, was expressed in the multipolar neurons in the MAZ/SVZ during cortical development (Sasaki et al., 2008) and bound to the ectodomain of FLRT2 (fibronectin and leucine-rich transmembrane protein2) secreted from neurons in the CP. This interaction functions as a chemorepellent signal, resulting in the delayed migration of multipolar neurons (Yamagishi et al., 2011). Indeed, both FLRT2 and Unc5D knockout mice showed earlier departure of some migrating neurons from the SVZ.

13.4 Conclusion Recent advances in the real-time imaging of migrating neurons have shed new light on classical studies in developmental neuroanatomy, and recent accumulating data strongly support the concept that migrating neurons dynamically change their morphologies during the course of migration. Genetic studies of human brain malformations, such as lissencephaly and PH, and of defects in mutant mouse models have also identified several key molecules that underlie this complex neuronal migration. In vivo studies using in utero electroporation in combination with in vitro studies have uncovered multiple important molecular pathways regulating neuronal migration. Furthermore, these studies have also suggested a potential relationship between neuronal migration failure and human psychiatric and neurological disorders, such as schizophrenia and dyslexia (Galaburda et al., 2006; Kamiya et al., 2005; Niwa et al., 2010). Future studies on the cellular and molecular mechanisms of neuronal migration should contribute to a better understanding of the mechanisms of not only normal development but also neurological disorders of the cerebral cortex.

See also Migration: Neuronal Migration Disorders; Nucleokinesis; Radial Migration in the Developing Cerebral Cortex; Radial Migration of Neurons in the Cerebral Cortex.

List of abbreviations CP cortical plate EM electron microscopy IMZ intermediate zone MAZ multipolar cell accumulation zone MZ marginal zone OSVZ outer subventricular zone REP rapidly exiting population RGC radial glial cell RGF radial glial fiber SEP slowly exiting population SVZ subventricular zone VZ ventricular zone

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Glossary Multipolar cell accumulation zone (MAZ) Located just above the VZ and overlapped with the lower part of the SVZ. Postmitotic multipolar cells are accumulated in the MAZ. Pin-like morphology The morphology of cells in VZ that have contact with the ventricular surface but do not attach to the pial surface. Rapidly exiting population (REP) One of the direct progeny populations of radial glial cells that moves rapidly to the IMZ/SVZ using a migration mode of somal translocation and undergoes mitosis. REP includes basal progenitors for neurons but may also contain some glial progenitors. Slowly exiting population (SEP) One of the direct progeny populations of radial glial cells that remain in the VZ for a long time and slowly enter the MAZ, where they accumulate as multipolar cells.

Supplementary data Supplementary data related to this article can be found online at doi:10.1016/B978-0-12-814407-7.00013-4

References Bai, J., Ramos, R.L., Ackman, J.B., Thomas, A.M., Lee, R.V., LoTurco, J.J., 2003. RNAi reveals doublecortin is required for radial migration in rat neocortex. Nat. Neurosci. 6, 1277e1283. Bellion, A., Baudoin, J.P., Alvarez, C., Bornens, M., Metin, C., 2005. Nucleokinesis in tangentially migrating neurons comprises two alternating phases: forward migration of the Golgi/centrosome associated with centrosome splitting and myosin contraction at the rear. J. Neurosci. 25, 5691e5699. Berry, M., Rogers, A.W., 1965. The migration of neuroblasts in the developing cerebral cortex. J. Anat. 99, 691e709. Boulder Committee, 1970. Embryonic vertebrate central nervous system: revised terminology. Anat. Rec. 166, 257e261. Brittis, P.A., Meiri, K., Dent, E., Silver, J., 1995. The earliest patterns of neuronal differentiation and migration in the mammalian central nervous system. Exp. Neurol. 134, 1e12. Chae, T., Kwon, Y.T., Bronson, R., Dikkes, P., Li, E., Tsai, L.H., 1997. Mice lacking p35, a neuronal specific activator of Cdk5, display cortical lamination defects, seizures, and adult lethality. Neuron 18, 29e42. Corbo, J.C., Deuel, T.A., Long, J.M., et al., 2002. Doublecortin is required in mice for lamination of the hippocampus but not the neocortex. J. Neurosci. 22, 7548e7557. des Portes, V., Pinard, J.M., Billuart, P., et al., 1998. A novel CNS gene required for neuronal migration and involved in X-linked subcortical laminar heterotopia and lissencephaly syndrome. Cell 92, 51e61. Deuel, T.A., Liu, J.S., Corbo, J.C., Yoo, S.Y., Rorke-Adams, L.B., Walsh, C.A., 2006. Genetic interactions between doublecortin and doublecortin-like kinase in neuronal migration and axon outgrowth. Neuron 49, 41e53. Englund, C., Fink, A., Lau, C., et al., 2005. Pax6, Tbr2, and Tbr1 are expressed sequentially by radial glia, intermediate progenitor cells, and postmitotic neurons in developing neocortex. J. Neurosci. 25, 247e251. Fietz, S.A., Kelava, I., Vogt, J., et al., 2010. OSVZ progenitors of human and ferret neocortex are epithelial-like and expand by integrin signaling. Nat. Neurosci. 13, 690e699. Fish, J.L., Dehay, C., Kennedy, H., Huttner, W.B., 2008. Making bigger brainsethe evolution of neural-progenitor-cell division. J. Cell Sci. 121, 2783e2793. Fox, J.W., Lamperti, E.D., Eksioglu, Y.Z., et al., 1998. Mutations in filamin 1 prevent migration of cerebral cortical neurons in human periventricular heterotopia. Neuron 21, 1315e1325. Galaburda, A.M., LoTurco, J., Ramus, F., Fitch, R.H., Rosen, G.D., 2006. From genes to behavior in developmental dyslexia. Nat. Neurosci. 9, 1213e1217. Gdalyahu, A., Ghosh, I., Levy, T., et al., 2004. DCX, a new mediator of the JNK pathway. EMBO J. 23, 823e832. Ge, W., He, F., Kim, K.J., et al., 2006. Coupling of cell migration with neurogenesis by proneural bHLH factors. Proc. Natl. Acad. Sci. USA 103, 1319e1324. Gertz, C.C., Lui, J.H., Lamonica, B.E., Wang, X., Kriegstein, A.R., 2014. Diverse behaviors of outer radial glia in developing ferret and human cortex. J. Neurosci. 34 (7), 2559e2570. Gerwins, P., Blank, J.L., Johnson, G.L., 1997. Cloning of a novel mitogen-activated protein kinase kinase kinase, MEKK4, that selectively regulates the cJun amino terminal kinase pathway. J. Biol. Chem. 272, 8288e8295. Gilmore, E.C., Ohshima, T., Goffinet, A.M., Kulkarni, A.B., Herrup, K., 1998. Cyclin-dependent kinase 5-deficient mice demonstrate novel developmental arrest in cerebral cortex. J. Neurosci. 18, 6370e6377. Gleeson, J.G., Allen, K.M., Fox, J.W., et al., 1998. Doublecortin, a brain-specific gene mutated in human X-linked lissencephaly and double cortex syndrome, encodes a putative signaling protein. Cell 92, 63e72. Goto, T., Mitsuhashi, T., Takahashi, T., 2004. Altered patterns of neuron production in the p27 knockout mouse. Dev. Neurosci. 26, 208e217. Hand, R., Bortone, D., Mattar, P., et al., 2005. Phosphorylation of Neurogenin2 specifies the migration properties and the dendritic morphology of pyramidal neurons in the neocortex. Neuron 48, 45e62. Hansen, D.V., Lui, J.H., Parker, P.R., Kriegstein, A.R., 2010. Neurogenic radial glia in the outer subventricular zone of human neocortex. Nature 464, 554e561. Hatanaka, Y., Hisanaga, S., Heizmann, C.W., Murakami, F., 2004. Distinct migratory behavior of early- and late-born neurons derived from the cortical ventricular zone. J. Comp. Neurol. 479, 1e14.

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Cdk5 phosphorylates and stabilizes p27kip1 contributing to actin organization and cortical neuronal migration. Nat. Cell Biol. 8, 17e26. Kemphues, K.J., Priess, J.R., Morton, D.G., Cheng, N.S., 1988. Identification of genes required for cytoplasmic localization in early C. elegans embryos. Cell 52, 311e320. Kitazawa, A., Kubo, K., Hayashi, K., Matsunaga, Y., Ishii, K., Nakajima, K., 2014. Hippocampal pyramidal neurons switch from a multipolar migration mode to a novel “climbing” migration mode during development. J. Neurosci. 34, 1115e1126. Koizumi, H., Tanaka, T., Gleeson, J.G., 2006. Doublecortin-like kinase functions with doublecortin to mediate fiber tract decussation and neuronal migration. Neuron 49, 55e66. Miyata, T., Kawaguchi, A., Okano, H., Ogawa, M., 2001. Asymmetric inheritance of radial glial fibers by cortical neurons. Neuron 31, 727e741. Miyata, T., Kawaguchi, A., Saito, K., Kawano, M., Muto, T., Ogawa, M., 2004. 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Nagano, T., Yoneda, T., Hatanaka, Y., Kubota, C., Murakami, F., Sato, M., 2002. Filamin A-interacting protein (FILIP) regulates cortical cell migration out of the ventricular zone. Nat. Cell Biol. 4, 495e501. Namba, T., Kibe, Y., Funahashi, Y., Nakamuta, S., Takano, T., et al., 2014. Pioneering axons regulate neuronal polarization in the developing cerebral cortex. Neuron 81 (4). Nguyen, L., Besson, A., Heng, J.I., et al., 2006. p27kip1 independently promotes neuronal differentiation and migration in the cerebral cortex. Genes Dev. 20, 1511e1524. Niwa, M., Kamiya, A., Murai, R., et al., 2010. Knockdown of DISC1 by in utero gene transfer disturbs postnatal dopaminergic maturation in the frontal cortex and leads to adult behavioral deficits. Neuron 65, 480e489. Noctor, S.C., Flint, A.C., Weissman, T.A., Dammerman, R.S., Kriegstein, A.R., 2001. Neurons derived from radial glial cells establish radial units in neocortex. Nature 409, 714e720. Noctor, S.C., Martinez-Cerdeno, V., Ivic, L., Kriegstein, A.R., 2004. Cortical neurons arise in symmetric and asymmetric division zones and migrate through specific phases. Nat. Neurosci. 7, 136e144. Ochiai, W., Minobe, S., Ogawa, M., Miyata, T., 2007. Transformation of pin-like ventricular zone cells into cortical neurons. Neurosci. Res. 57, 326e329. Ohshima, T., Hirasawa, M., Tabata, H., et al., 2007. Cdk5 is required for multipolar-to-bipolar transition during radial neuronal migration and proper dendrite development of pyramidal neurons in the cerebral cortex. Development 134, 2273e2282. Ohtaka-Maruyama, C., Hirai, S., Miwa, A., Heng, J.I.-T., Shitara, H., et al., 2013. RP58 regulates the multipolar-bipolar transition of newborn neurons in the developing cerebral cortex. Cell Rep. 3 (2), 458e471. Okamoto, M., Namba, T., Shinoda, T., Kondo, T., Watanabe, T., et al., 2013. TAG-1-assisted progenitor elongation streamlines nuclear migration to optimize subapical crowding. Nat. 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Chapter 14

Nucleokinesis Orly Reiner1 and Eyal Karzbrun2 1

Department of Molecular Genetics, The Weizmann Institute of Science, Rehovot, Israel; 2Kavli Institute of Theoretical Physics and Department of

Physics, University of California, Santa Barbara, CA, United States

Chapter outline 14.1. Nucleokinesis: introduction 14.2. The nucleus 14.2.1. The nuclear membrane and nuclear pores 14.3. Chromatin 14.4. Membraneless organelles in the nucleus 14.5. Higher order structure of the nucleus 14.6. Diseases 14.6.1. Cohesinopathies 14.6.2. Affecting the nuclear envelope 14.7. Interactions between the nucleus and the cytoskeleton 14.7.1. The LINC complex, structure 14.8. The LINC complex, function 14.9. The LINC complex in nuclear positioning 14.10. The link between the nucleus and the centrosome

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14.11. The LINC complex in nucleokinesis 14.12. Nucleokinesis during interkinetic nuclear movement 14.13. Microtubule binding motors 14.13.1. Dynein 14.13.2. Kinesin Kif1a 14.14. Cytoskeleton dynamics as nuclear drivers 14.15. Collective mechanisms for nuclear migration 14.15.1. Intercellular signaling 14.15.2. Mechanical interactions 14.16. The role of INM during neurodevelopment 14.17. INM summary 14.18. Conclusions and future directions Acknowledgments References

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14.1 Nucleokinesis: introduction Nucleokinesis, literally the movement of the nucleus, may or may not be coupled with cellular motility. In this chapter, we will present our view on the moving nucleus. We initiate with the structure of the nucleus from the exterior side, which is surrounded by a membrane, with specific gates that are the nuclear pores. We briefly discuss the structure of the chromatin and internuclear membraneless structures, such as nucleoli. We then detail the higher order structure of the nucleus, for which major advancements have been progressed during recent years. Underscoring these structural studies, we present several diseases that are known to affect the structure of the nucleus. We then describe how the nucleus is connected with the cytoskeleton and finally discuss one interesting developmental motion of nuclei that is interkinetic nuclear motility. Possible signaling-related as well as mechanical-related mechanisms are presented. Collectively, we aim to introduce the concept that the migrating nucleus is more than just a passive cargo.

14.2 The nucleus 14.2.1 The nuclear membrane and nuclear pores The eukaryotic nucleus is alienated from the rest of the cell by the nuclear envelope, which consists of two nuclear membranes, nuclear pore complexes (NPCs) and in metazoans also of nuclear lamina (schematic presentation in Fig. 14.1). The outer nuclear membrane, which is continuous with the rough endoplasmic reticulum, is separated from the inner nuclear membrane by a breach of 20e50 nm, which is termed the perinuclear space. The two membranes are populated with distinct sets of proteins, even though they are fused to each other next to the NPCs. The nuclear pores are composed of

Cellular Migration and Formation of Axons and Dendrites. https://doi.org/10.1016/B978-0-12-814407-7.00014-6 Copyright © 2020 Elsevier Inc. All rights reserved.

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Cytoplasm

IF

Actin Microtubules KASH domain proteins

NPC

SUN domain proteins INM Heterochromatin

PNS

ONM

Euchromatin Lamina Nucleus

FIGURE 14.1 Schematic presentation of the physical interaction between the nucleus and the cytoskeleton. The metazoan nucleus is separated from the cytoplasm by an outer nuclear membrane (ONM), an inner nuclear membrane (INM), and a mesh of lamins. These components combined with the nuclear pore complexes (NPCs) form the nuclear envelope. The nuclear envelope is in close association with the chromatin fiber. The lamina interacts preferentially with heterochromatin, whereas the NPCs interact preferentially with transcribed euchromatin. The bridge between the cytoskeleton and the nucleus is established by the SUN domain and KASH domain proteins. SUN domain proteins traverse the INM and interact with lamina and heterochromatin inside the nucleus. KASH domain proteins traverse the ONM and interact directly with cytoskeleton fibers (actin fibers) or with intermediate filaments (IF) associated proteins or with microtubules motor proteins in the cytoplasm. Inside the perinuclear space (PNS), the KASH domain proteins bind the SUN domain proteins to form the physical link between the cytoskeleton and the nucleus.

huge multiprotein complexes (about 1000 proteins, 120 MDa in humans) that form channels between the nucleus and the cytoplasm (reviewed by (Hetzer et al., 2005; D’Angelo and Hetzer, 2006; Stewart et al., 2007; Knockenhauer and Schwartz, 2016; Beck and Hurt, 2017)). Key structural elements of the NPC are the cytoplasmic ring, the inner pore ring and the nuclear ring, as well as peripheral elements such as the nuclear basket and the cytoplasmic filaments. A subset of these 1000 proteins is remarkably stable, and the proteins do not turnover during years, whereas other components are highly dynamics, exhibiting a dwell time of minutes (Knockenhauer and Schwartz, 2016). These pores serve as the primary gates for communication between the cytoplasm and the nucleus. These activities include free diffusion of small molecules in both directions and the major active transport routes through the NPC, such as the nuclear import of proteins, the export of RNA and ribonucleoproteins (RNPs), and the bidirectional shuttling of molecules, including RNA, RNPs, and proteins involved in signaling, biogenesis, and turnover dependent on transport receptoremediated import and export routes. Other noncanonical functions of the NPC include transcription-coupled mRNA export, as well as other processes associated with chromatin biology (Beck and Hurt, 2017). Here we will focus on the connection between the nucleus and the cytoskeleton. In metazoans, a nuclear lamina is located next to the inner nuclear membrane from the inside of the nucleus. The nuclear lamina is assembled mainly from A- and B-type lamins. The lamins are filamentous proteins from type-V intermediate filaments that fold into coiled-coil dimers. These dimers interact in a head to tail manner to build polymers that generate a mesh of 10e50 nm in width. The nuclear lamina directly interacts with the inner nuclear membrane and supports the whole structure of the nuclear envelope (reviewed by (Gruenbaum et al., 2005; Dechat et al., 2008; Imamoto and Funakoshi, 2012)). In humans, B-type lamins are ubiquitously expressed, whereas expression of A-type lamins is developmentally regulated: A-type lamins are absent in early embryonic cells but appear during tissue differentiation. A key function of the nuclear lamins is to maintain the shape and mechanical properties of nuclei. In addition, lamins have important roles in many nuclear processes, including transcription, DNA replication, cell cycle control, and DNA repair (reviews (Dechat, Adam et al., 2010; Dechat, Gesson et al., 2010)).

14.3 Chromatin The nuclear envelope serves also as a physical barrier between the cytoplasm and the key constituent of the nucleus, the chromatin fiber. The chromatin fiber is composed of a basic repetitive unit, the nucleosome, which is built from 147 bp of DNA wrapped twice around a histone octamer. Nucleosomes are organized in higher-order structures, which are determined by posttranslational modifications in histones tails, DNA methylation, regulatory factors that bind the different

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modifications and by architectural proteins such as histone H1 (reviewed by (Allis et al., 2007; Bhaumik et al., 2007; Cutter and Hayes, 2015)). During interphase, the chromatin is organized into relatively decondensed euchromatin regions that are transcribed and into condensed heterochromatin regions, which are nontranscribed (reviewed by (Trojer and Reinberg, 2007)). These two regions in the chromatin fiber interact with different components of the nuclear envelope. The inner nuclear membrane and the lamina interact preferentially with heterochromatin, while the NPC interacts with euchromatin. These interactions are important for regulation of transcription and chromosomes position (reviewed by (Akhtar and Gasser, 2007)). A typical mark of constitutive heterochromatin, usually detected in gene-poor regions of pericentromeres, is the trimethylation of histone H3 on lysine 9 (H3K9me3), while H3K27me3 is usually enriched on facultative heterochromatin. Both marks recruit distinct protein machineries and may underlie distinct biological features, although the consequence is chromatin compaction in both cases (Saksouk et al., 2015). Superresolution microscopy allowed the categorization of three groups of chromatin structures based on major types of histone modifications (Xu et al., 2018). Histone acetylation forms spatially segregated nucleosome nanoclusters, active histone methylation forms spatially dispersed nucleosome nanodomains, and repressive histone methylation forms highly condensed large aggregates, thus allowing to identify structures that range in size between tens of nanometers to a few microns.

14.4 Membraneless organelles in the nucleus The nucleus contains several membraneless organelles that vary in size and define highly dynamic macromolecular assemblies, whose components rapidly cycle between the organelle and the surrounding (reviews (Mitrea and Kriwacki, 2016; Shin and Brangwynne, 2017)). These structures represent liquid-phase condensates, which form via a biologically regulated (liquideliquid) phase separation process. The nuclear structures are composed of proteins, RNA and DNA molecules, and their concentrated states are a central aspect of how these structures are formed. The nuclear structures include, for example, nucleoli, Cajal bodies, and nuclear speckles. Here we will discuss the nucleolus as an example for these interesting structures. The nucleolus is the largest of these membraneless organelles and serves a hub for ribosomal RNA synthesis and nascent ribosome assembly (review (Pederson, 2011)). Nucleolar assembly is initiated by RNA Polymerase I (RNA Pol I) transcription of clustered ribosomal RNA (rRNA) genes (rDNA) bound to the transcription factor UBF. Ribosome biogenesis occurs in distinct subcompartments, which can be identified by electron microscopy, starting from the fibrillar centers, where rDNA is transcribed into rRNA. Then the pre-rRNA molecules transit through the dense fibrillar component, where they are spliced and the small ribosomal subunit is assembled, then move into the granular component where the large ribosomal subunit is assembled. Pre-ribosomal particles are then released into the nucleoplasm and subsequently exported into the cytoplasm where functional ribosomes are assembled. The composition and the physical properties of the nucleolus are highly dynamic and exhibit liquid-like behavior in vivo, which is dependent upon phase separation of their molecular components (Brangwynne et al., 2011; Hernandez-Verdun, 2011). The mixture of proteins and nucleic acids separates into liquid and hydrogel-like structures. In the hydrogel, dense-structure, the molecules can reach concentration of 10e100 fold, reaching millimolar ranges. A large proportion of the proteins known to associate with membraneless organelles exhibit multivalency through the display of repeated low complexity motifs (e.g., SR, RGG, or FG motifs) and/or of multiple copies of folded domains, such as RRM domains. It has been proposed that specific protein-nucleic acid or proteineprotein interactions initiate the assembly of these membraneless organelles, and this is followed by phase separation of other components. This concept has been demonstrated in nucleoli, where the transcription factor UBF binds to organized clusters of rRNA genes which are followed by the recruitment of RNA polymerase 1. RNA transcription starts and then the assembly of the nucleolus begins (Yang et al., 2003; Grob et al., 2014). HNRNPU is involved also in regulation of nucleolus formation through its interaction with B23, a nucleolar protein required for rRNA processing (Yao et al., 2010).

14.5 Higher order structure of the nucleus Molecular, biochemical, and microscopy-based techniques have broadened our view on how the chromatin is organized in relation to the nuclear membrane and in relation to subnuclear structures such as nucleoli and nuclear speckles. The molecular techniques to probe chromosome organization, based on chromosome conformation capture (3C) and subsequent advances as 4C, 5C, HiC technologies, enhanced our understanding of the spatial organization of the genome (reviews (Gibcus and Dekker, 2013; Pombo and Dillon, 2015; Yu and Ren, 2017; Eagen, 2018; Roy et al., 2018)). These methods revealed that chromatin is folded by looping, self-association, and compartmentation and allowed to identify topologically associated domains (TADs), some of which are associated with repressed chromatin and some with active chromatin. In the fruit fly, the TADs superimpose on the banding patterns of polytene chromosomes (Eagen et al., 2015;

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Ulianov et al., 2016). A clear segregation of the heterochromatin marker H3K9me3 modification was observed within the TAD boundary regions. In addition, the probability of intrachromosomal contact between pairs of loci in a chromosome decreased consistently with increasing genomic distance. Multiple studies demonstrate that direct contacts occur between promoters and enhancers, resulting in the formation of the transcription preinitiation complex. In addition, interactions between distantly located sequences can be promoted by cohesins and the CCCTC-binding factor (CTCF) (Pombo and Dillon, 2015). The direct role of CTCF in TAD insulation and in instructing chromatin loops has been demonstrated following acute depletion of the protein in cells (Nora et al., 2017). CTCF depletion affected transcriptional activity but not either the spread of H3K27me3 domains or the higher-order of genomic compartmentalization. Similar experiments targeting different subunits of the cohesion complex resulted in similar conclusions ((Haarhuis, van der Weide et al., 2017; Rao et al., 2017; Schwarzer et al., 2017; Wutz et al., 2017) and review (Rada-Iglesias et al., 2018). HNRNPU (heterogeneous nuclear ribonucleoprotein U), also known as scaffold attachment factor A (SAF-A), is also involved in regulating the architecture of the 3D genome (review (Zhang et al., 2018)). Disruption of HNRNPU decreased TAD boundary strengths at borders between A (active) and B (inactive) compartments (Fan et al., 2018). HNRNPU is also involved in formation of chromatin loops through its oligomerization and interactions with chromatin-associated RNAs dependent on its AAAþ ATPase region, and binding of DNA (Romig et al., 1992; Vizlin-Hodzic, Johansson et al., 2011; Vizlin-Hodzic, Runnberg et al., 2011; Nozawa et al., 2017). Long-range chromatin interactions between and within compartments or TADs are also significantly remodeled upon HNRNPU depletion (Fan et al., 2018). The molecular techniques described involve averaging the distances from many chromosomes. Microscopy techniques, especially those involving multiplexing, allowed to investigate the spatial arrangements of TAD compartments in several chromosomes in one study (Wang et al., 2016). Their findings indicated that TADs are largely organized into two compartments spatially arranged in a polarized manner in individual chromosomes. In addition, they noted that active and inactive X chromosomes adopt different folding and compartmentalization configurations, thus suggesting that the spatial organization of chromatin domains can change in response to regulation. Yet, it should be noted that the correlations between contact frequency, measured by Hi-C, and spatial distance, measured by FISH, are not resolved and should be used with caution (Fudenberg and Imakaev, 2017). In most cell types, the nuclear periphery is associated with heterochromatin, whereas euchromatin is more found centrally located. Mapping of lamin Bechromatin interactions was done primarily by the DamID technology, in which bacterial DNA adenine methyltransferase (Dam) is tethered to a nuclear lamina protein (typically Lamin B1) leading to adenine methylation of DNA regions that contact the nuclear lamina protein (Pickersgill et al., 2006). These studies revealed that large genomic regions, termed LADs, are bound to the nuclear lamina (Amendola and van Steensel, 2014). LADs are strongly enriched for H3K27me3, which is a known repressive mark. Therefore, LADs are of particular interest for two broad reasons. First, their nuclear lamina anchoring helps to establish interphase chromosome topology and thus the overall genome spatial organization. Second, most of the several thousands of genes in LADs are expressed at very low levels, suggesting a role in gene repression (reviews (van Steensel and Belmont, 2017; Yanez-Cuna and van Steensel, 2017)). Depletion of the nuclear matrix-associated protein, HNRNPU, resulted in a global condensation of the chromatin and a significant increase in the LADs (Fan et al., 2018). Microscopy-based techniques allowed to demonstrate how specific regions of the genome can organize in relation to the nuclear membraneless organelles. RNA polymerase I (Pol I)-transcribed rDNA genes, which are encoded on several distinct chromosomes, localize within the nucleolus (Pederson, 2011). Specific examples of RNA polymerase II (Pol II)transcribed genes have been shown to localize near the periphery of nuclear speckles (Khanna et al., 2014), a nuclear body that contains various mRNA processing and splicing factors (Spector and Lamond, 2011). Due to the sizes of the nuclear membraneless organelles, the abovementioned interactions may be too far apart to be detected by proximity ligations. To overcome this disadvantage a method that enables genome-wide detection of multiple DNA interactions that occur simultaneously within the nucleus was developed (Quinodoz et al., 2018). They recapitulated known genome structures identified by Hi-C, including chromosome territories, compartments, TADs, and loop structures. In addition, by extending their method to simultaneously measure RNA and DNA interactions, they found that two interchromosomal hubs correspond to DNA organization around the nucleolus and nuclear speckles.

14.6 Diseases 14.6.1 Cohesinopathies Heterozygous mutations in genes encoding for components of the cohesion complex have been associated with human developmental syndromes collectively known as cohesinopathies (reviews (Liu and Krantz, 2008; Singh and Gerton, 2015;

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Watrin et al., 2016)). Homozygous deletions of genes encoding the mitotic cohesin subunits are early embryonic lethal while heterozygous knockout mice have variable phenotypes. For example, Cornelia de Lange syndrome is caused by mutations in several different cohesin genes, and patients exhibit craniofacial and limb deformities, hirsutism, gastroesophageal dysfunction, neurodevelopmental delay, and mild to severe intellectual disability. The majority of the mutations are in the NIPBL gene. Interestingly, it has been shown the mouse ortholog interacts with Zfp609 and the integrator complex to transcriptionally regulate cortical neuronal migration (van den Berg et al., 2017).

14.6.2 Affecting the nuclear envelope Mutations in various genes encoding for nuclear envelope proteins collectively known as laminopathies are associated with several human pathologies such as progeria, muscular dystrophy, axonal neuropathy, leukodystrophy, and lipodystrophy (reviewed by (Burke and Stewart, 2002; Capell and Collins, 2006; Rankin and Ellard, 2006; Worman and Bonne, 2007)). Most known mutations are in A-type lamins, yet also mutations in the nuclear lamin gene LMNB2 have been described and these are associated with epilepsy and early ataxia syndrome (Damiano et al., 2015). Lamin knockout mouse models have shown multiple deficits that include abnormal nuclear structure, changes in gene expression as well as abnormal signaling (Mounkes et al., 2003; Vergnes et al., 2004; Kim et al., 2011; Solovei et al., 2013; Giacomini et al., 2016). It has been shown that Lamin B can be locally translated in the axon and as such functions outside of the nucleus (Yoon et al., 2012). The molecular mechanisms involved in lamin-associated diseases include malfunction of nuclear envelope proteins in mechanical support for the nucleus, in regulation of nuclear architecture and in transduction of signals from the cytoplasm to the transcriptional machinery, as well as nuclear independent functions (Broers et al., 2004). Mutations in genes encoding for lamins, as well as for the inner nuclear membrane protein Emerin, lead to abnormalities in nuclear morphology such as convolutions in the nuclear envelope and nuclear blebbing (Goldman et al., 2004; Lammerding et al., 2004; Lammerding et al., 2005). Mutated forms of the Lamin A/C gene altered genomic LADs, which impacted epigenomic programming of myogenic differentiation (Perovanovic et al., 2016). Mutations in the Lamin A/C gene also result in lower resistance of the nucleus to mechanical pressure (Lammerding et al., 2004). In agreement to these results, knockdown of Lamin A/C in human epithelial cells reduced the stiffness of the nuclei by twofold as measured by resistance of nuclei to aspiration force (Pajerowski et al., 2007). Furthermore, increased condensation of the chromatin fiber enhanced the stiffness of the nuclei by fivefold. Moreover, prolonged exposure of nuclei to aspiration force resulted in irreversible nuclear deformations, which were dependent on the chromatin structure (Pajerowski et al., 2007). Thus, by affecting the shape and stiffness of the nucleus, both the nuclear envelope and the chromatin fiber are expected to affect cellular mechanical processes in which the nucleus is involved. These processes include cell migration, nucleokinesis, and interkinetic nuclear movement. Furthermore, it is also possible that as mechanical pressure is translated into changes in gene transcription (reviewed by (Wang et al., 2009)), morphological changes in the structure of the nucleus that occur during nucleokinesis and cell migration will also affect gene transcription. Together, these evidences imply that the moving nucleus is more than just a passive cargo. Indeed, mutations in different proteins, which are part of the nuclear structure, affect motility. For example, mutations in Lamin A/C or knockout of the outer nuclear membrane protein Nesprin-2 result in inhibited migration of fibroblasts (McClintock et al., 2006; Lee et al., 2007; Luke et al., 2008). Mutations in the outer nuclear membrane protein UNC-83 and in the inner nuclear membrane protein UNC-84 reduce nucleokinesis in Hyp7 cells and P cells in Caenorhabditis elegans (Malone et al., 1999; Starr et al., 2001). In addition, nucleokinesis in photoreceptor cells during the eye development is dependent on Lamin B and the outer nuclear membrane protein Klar in Drosophila melanogaster (MosleyBishop et al., 1999; Patterson et al., 2004) and on the inner nuclear membrane protein Syne2a in Zebrafish (Tsujikawa et al., 2007). Moreover, directed migration of melanocytes is dependent on the function of Histone H1 and on increased condensation level of the chromatin fiber (Gerlitz et al., 2007; Gerlitz and Bustin, 2010). Interestingly, polarized posttranslational changes in histone H1, core histones, and DNA were detected in cells facing the scratch in a wound-healing assay. Migration-associated changes in histone H1 localization were also observed during nucleokinesis in the simple multicellular organism Neurospora crassa (Gerlitz et al., 2007). Collectively these data suggest that dynamic reorganization of the chromatin fiber is an early and evolutionary conserved event in the cellular response to migration cues. The importance of the nuclear envelope and the organization of the chromatin fiber to nucleokinesis and to cell migration suggest that nuclear-associated molecules are connected physically to the cytoskeleton. This connection may enable the cytoskeleton to change the shape of the nucleus and/or to move it inside the cell. Initial experiments of Ingber and colleagues demonstrated the ability of cytoskeleton alterations to cause deformations inside the nucleus (Maniotis et al., 1997). In these experiments a fibronectin-coated micropipette was linked to integrins on the cell membrane. Pulling

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the micropipette led to cellular elongation along with distortion of the cytoskeleton. Simultaneously, the nucleus and even the nucleoli inside the nucleus were distorted toward the pulling source. These results suggest that force is transmitted directly from the cell membrane through the cytoskeleton to the nuclear envelope and the chromatin inside the nucleus (Maniotis et al., 1997). The molecular bridge between the cytoskeleton and the nucleus has been identified and has been termed the linker of nucleoskeleton and cytoskeleton (LINC) complex (Crisp et al., 2006).

14.7 Interactions between the nucleus and the cytoskeleton 14.7.1 The LINC complex, structure The LINC complex is formed by a direct interaction between members of the SUN (Sad1p and UNC-84) domain protein family that are localized to the inner nuclear membrane and members of the KASH (Klarsicht, Anc-1, Syne-1 homology) domain protein family, which are localized to the outer nuclear membrane (see Fig. 14.1). The SUN domain proteins interact with structural elements inside the nucleus, while the KASH domain proteins interact with the cytoskeleton in the cytoplasm. This interaction forms a physical link between the cytoskeleton and the nucleus in all eukaryotic cells (reviewed by (Fridkin et al., 2009; Starr, 2009; Mejat and Misteli, 2010; Horn, 2014; Chang, Worman et al., 2015)). Direct evidence demonstrated that mechanical force is applied to nesprin-2G, a member of the LINC complex, and the tension was reduced in samples from laminopathy patients (Arsenovic et al., 2016). The SUN domain proteins contain transmembrane domains, which are incorporated into the inner nuclear membrane, leaving the N-terminal of the protein in the nucleoplasm and the C-terminal part of the protein inside the perinuclear space. In the nucleoplasm, the N-terminal part of the SUN domain proteins interacts directly with various nuclear factors such as Lamin A and Emerin in mammals (Haque et al., 2010, Crisp et al., 2006, Haque et al., 2006), telomere-localized proteins in Schizosaccharomyces pombe and in Saccharomyces cerevisiae (Chikashige et al., 2006; Antoniacci et al., 2007; Conrad et al., 2007), a centromere-localized protein in S. pombe (King et al., 2008), and even DNA in Dictyostelium discoideum (Xiong et al., 2008). The part of the protein that is inside the perinuclear space binds the KASH domain proteins through the SUN domain, which is a motif of 121 residues that was first identified in the C-terminal of UNC-84 (C. elegans) and of Sad1p (S. pombe) (Malone et al., 1999). The SUN domain protein family has been expanded through evolution, and in mammals four members have been identified. Two of which are ubiquitously expressed (Sun1 and Sun2) (Crisp et al., 2006; Wang et al., 2006) while the other two are expressed mainly in the testis (Sun3 and Spag4) (Shao et al., 1999; Crisp et al., 2006). The KASH domain is a hydrophobic motif of w60 residues, which was first identified in the C-terminus of Klarsicht (D. melanogaster), Anc-1 (C. elegans) and Syne-1,2 in mammals (Starr and Han, 2002). Within this motif, 20 residues form a transmembrane domain, which traverses the outer nuclear membrane, and 30e35 residues are localized into the perinuclear space, where they bind the SUN domain. This interaction holds most of the KASH domain proteins to the outer nuclear membrane and prevents them from reaching the endoplasmic reticulum (Malone et al., 2003; Padmakumar et al., 2005; Crisp et al., 2006; Kracklauer et al., 2007). The N-terminal part of the KASH domain proteins, which exceeds from the outer nuclear membrane to the cytoplasm, varies in length. In most KASH domain family members, the cytoplasmic part contains spectrin repeats or coiled-coil domains, which confirm elasticity, length, and ability to homodimerize as well as to interact with additional partners. Through their N-terminus the KASH domain family members interact with various cytoskeletal elements. Syne-1 and Syne-2 in mammals, Anc-1 (C. elegans) and MSP-300 (D. melanogaster) contain an Factin binding domain (ABD) (Rosenberg-Hasson et al., 1996; Starr and Han, 2002; Zhang et al., 2002; Zhen et al., 2002; Zhang et al., 2005; Wang et al., 2009). In addition, Syne-3a in mammals binds the intermediate filaments interacting protein Plactin (Wilhelmsen et al., 2005). However, the interaction of this group of proteins is not limited to microfilaments and intermediate filaments; KASH domain proteins are able to bind microtubule motor proteins as well. Kinesin binding was shown for Syne-1, Syne-2 (Fan and Beck, 2004; Zhang et al., 2009), and Syne-4 (Roux et al., 2009) in mammals and for UNC-83 in C. elegans (Meyerzon et al., 2009). Association with the Dynein complex was shown for Syne-1 and Syne2 in mammals (Zhang et al., 2009) and for UNC-83 (Fridolfsson, Ly et al.), ZYG-12 (Malone et al., 2003), and Kms1 (Miki et al., 2002) in C. elegans. In mammals, the KASH domain proteins have several names: Nesprin (nuclear envelope spectrin repeat) or Syne (synaptic nuclear envelope) or Myne (myocyte nuclear envelope) or NUANCE. Multiple Syne isoforms are formed from the four mammalian genes by alternative transcription starting sites and by alternative splicing. Syne-1e3 are ubiquitous (Zhang et al., 2001; Mislow et al., 2002; Wilhelmsen et al., 2005), whereas Syne-4 has been found only in secretory epithelial cells (Roux et al., 2009). In summary, the LINC complex is composed of SUN and KASH domain proteins, which are connected on one side to the cytoskeleton and on the other side to the nuclear envelope and the chromatin fiber.

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14.8 The LINC complex, function The LINC complex is essential for viability as has been found using knockout animals. Double knockout mice for the SUN domain proteins SUN1 and SUN2 or for the KASH domain proteins Syne-1 and Syne-2 die soon after birth (Zhang et al., 2007; Lei et al., 2009). In lower eukaryotes, depletion of or mutations in the KASH domain proteins Zyg-12, KDP-1 (C. elegans), and Msp-300 (D. melanogaster) or the SUN domain protein SUN-1 is embryonic lethal (Rosenberg-Hasson et al., 1996; Malone et al., 2003; Fridkin et al., 2004; McGee et al., 2009). Detailed analysis of the knockout animals, as well as of additional mutants revealed the critical role of the LINC complex in positioning the centrosome next to the nucleus as well as in force transmission from the cytoskeleton to the nucleus. When these processes are impaired, defects are found in various cellular mechanical events such as meiosis, cytokinesis, nuclear positioning, centrosome positioning, and nucleokinesis.

14.9 The LINC complex in nuclear positioning The importance of the LINC complex to positioning of nuclei is evolutionary conserved and was found both in C. elegans and in mammals. In C. elegans, the KASH domain proteins ANC-1 and Zyg-12 are necessary for the positioning of nuclei in even spaces in syncytial hypodermal cells and in the gonad, respectively (Starr and Han, 2002; Zhou et al., 2009). Mammalian muscle fibers contain hundreds of nuclei, which are aligned and evenly spaced. The positioning of nuclei in the muscle fiber is dependent on Syne-1. Furthermore, Syne-1 function is also required for the localization of nuclei beneath the postsynaptic membrane at neuromuscular junctions (Zhang et al., 2007; Puckelwartz, Kessler et al., 2009). These cellular phenotypes are also associated with cardiomyopathy when a dominant negative form of Syne-1 is expressed (Puckelwartz, Kessler et al., 2009; Puckelwartz, Kessler et al., 2009).

14.10 The link between the nucleus and the centrosome The centrosome in metazoans, or the spindle pole body (SPB) in yeast are tethered to the nucleus in most cells. This tethering is important for several processes such as mitosis and nucleokinesis. Studies on nuclear envelope proteins and on the LINC complex revealed the role of these components in the positioning of the centrosome. In S. cerevisiae, the SUN domain protein Mps3 interacts with an outer nuclear membrane protein Mps2 to form a bridge, which facilitates incorporation of newly duplicated SPB into the nuclear membrane (Jaspersen et al., 2006). Attachment of the SPB in S. pombe and the centrosome in D. discoideum to the nucleus is dependent on a bridge, which is formed between the chromatin and the SPB/centrosome through the LINC complex. Interfering with the chromatin structure in S. pombe or with the LINC complex in D. discoideum leads to deformations in the nuclear shape (King et al., 2008; Xiong et al., 2008). In D. discoideum, this interference results also in detachment of the centrosome from the nucleus (Xiong et al., 2008). In C. elegans, SUN-1, ZYG-12, the dynein complex, and microtubules are needed for the attachment of the centrosome to the nucleus during early embryonic divisions. In a proposed model, ZYG-12, which is anchored to the outer nuclear membrane by SUN-1, recruits the dynein complex to the nuclear envelope. Dynein generates a pulling force along the microtubule fibers to bring the centrosome and the nucleus close to each other. Once the centrosome is close to the nucleus, it is thought that dimers are formed between ZYG-12 monomers, which are found both on the outer nuclear membrane and in the centrosome to hold the two organelles together (Malone et al., 2003; Minn et al., 2009). Deficiencies in Lamin A/C or in Emerin lead to detachment of the centrosome from the nucleus in mammalian fibroblasts (Lee et al., 2007; Salpingidou et al., 2007). However, it is thought that centrosomal localization in these cells is independent of the LINC complex since members in the LINC complex are not dependent on Lamin A/C or Emerin for their proper localization (Crisp et al., 2006; Salpingidou et al., 2007). Emerin may be exceptional in this regard since some Emerin is found also on the outer nuclear membrane, where it may interact directly with microtubules (Salpingidou et al., 2007). In neurons and glial cells in mice, tethering of the nucleus to the centrosome during nucleokinesis is dependent on the LINC complex. As will be described in more details below, the LINC complex interacts with both dynein and kinesin. These interactions enable movement of the nucleus along the microtubules toward the centrosome (Zhang et al., 2009). In contrast to all the above examples where the LINC complex or additional nuclear envelope proteins help to tether the centrosome to the nucleus, the mammalian KASH domain protein Syne-4, which is found in secretory epithelial cells, has an opposite role. Expression of Syne-4 in HeLa cells leads to an increase in the distance between the centrosome and the nucleus. Since Syne-4 interacts with kinesin, it has been suggested that kinesin molecules bound to the nuclear envelope push the nucleus away from the centrosome. This activity may be the basis for achieving the unusual cellular architecture of secretory epithelial cells (Roux et al., 2009).

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14.11 The LINC complex in nucleokinesis The importance of the LINC complex to nucleokinesis was first identified in C. elegans, where mutations in unc-83 (encodes for a KASH domain protein) and unc-84 (encodes for a SUN domain protein) interfere with nuclear migration in several cell types (Sulston and Horvitz, 1981; Malone et al., 1999; Starr et al., 2001). Only recently it was found that UNC83 interacts with the Kinesin-1 subunit KLC-2 and the dynein-regulating proteins NudE and DLC-1 (Meyerzon et al., 2009; Fridolfsson et al., 2010). Thus, UNC-83 not only links the nucleus to a motor protein but may also switch between forward and backward movements. Kinesin-1 is the major motor that moves the nucleus while movement by the Dynein complex may be used to avoid obstacles. This activity has been observed during nucleokinesis in Hyp7 cells. In a singlecell C. elegans embryo, migration of the pronucleus is dependent on the link between the centrosome and the nucleus, which is formed through ZYG-12 (a KASH domain protein) and SUN-1 (Malone et al., 2003; Minn et al., 2009). Nucleokinesis in developing photoreceptor cells in D. melanogaster is dependent on Klaroid (a SUN domain protein), Klarsicht (a KASH domain protein), lamin, and dynactin (Mosley-Bishop et al., 1999; Patterson et al., 2004; Kracklauer et al., 2007). Therefore it was suggested that the LINC complex, which is anchored to the nuclear envelope by lamins interacts with the dynein complex to facilitate dynein-dependent movement of the nucleus toward the centrosome (Kracklauer et al., 2007). In migrating myoblasts, depleting nesprin-2G (part of the LINC complex) interfered with directed cell migration and centrosome orientation (Chang et al., 2015). In metastatic tumor cells, the ability to invade relays on the secretion of transmembrane membrane-type 1 matrix metalloproteinase (MT1-MMP), which was found to be regulated by the LINC complex molecule LIS1 (Infante et al., 2018).

14.12 Nucleokinesis during interkinetic nuclear movement Interkinetic nuclear movement (INM, IKNM) is a cyclic motion of the nuclei within the cell cytoplasm, and in synchrony with the cell cycle. This motion was first discovered in the neural tube by Shaper in 1897 and rediscovered by Sauer in 1935 (Schaper, 1897; Sauer 1935, 1936). Developing neuroepithelia are composed of elongated neural progenitor cells, each spanning the entire thickness of the epithelium from the ventricular (apical) surface to the laminal (basal) side (Noctor et al., 2001; Taverna et al., 2014). The progenitors are bipolar cells and have distinct apical and basal surface composition. The apical surface faces the ventricle; possesses a primary cilia; and is fenced with contractile actomyosin, tight junctions, and cadherins. The basal side is connected to the extracellular matrix of the lamina through integrins (Haubst et al., 2006). Nuclei undergoing mitosis occupy the apical (ventricular) surface, where centrosomes are localized. This is followed by apical-to-basal motion of the daughters’ nuclei, and progression of G1-phase. S-phase takes place as the nuclei reaches the basal side of the cell, and G2 occurs during basal-to-apical motion. INM is essential for normal brain development, and has been the focus of numerous studies and reviews (Taverna and Huttner, 2010; Reiner et al., 2012; Spear and Erickson, 2012; Miyata et al., 2014; Dantas et al., 2016; Bertipaglia et al., 2017). INM has been observed in other nonneuronal epithelia including the intestine and the epiblast (Grosse et al., 2011; Meyer et al., 2011; Yamada et al., 2013; Carroll et al., 2017). To date, researchers still debate on key questions: What are the molecular forces that lead to nuclear motion? How cell cycle coordinates motion? What are the biological benefits and physical constrains that emerge from INM? Is the motion completely cell autonomous, or are there collective cellular effects? Multiple mechanisms have so far been identified to drive nucleus motion during INM, and these can be classified into three categories: microtubule binding motors, cytoskeletal dynamics, and mechanical interactions with neighboring cells. Imaging studies revealed asymmetry between apical and basal motions, suggesting different mechanisms are involved in different stages of motion. Basal-to-apical motion exhibits fast migration bursts, 0.14 mm/min, whereas the apical-to-basal motion is continuous and slow, 0.06 mm/min (Tsai et al., 2010; Kosodo et al., 2011). In addition, basal-to-apical motion exhibits a linear progression with time, whereas apical-to-basal motion is diffusive and sublinear (Tsai et al., 2010; Okamoto et al., 2013; Okamoto et al., 2014). In the next sections, we describe the main candidates responsible for INM (Fig. 14.2).

14.13 Microtubule binding motors Microtubules span the neuronal progenitor, elongated along the apicalebasal axis. Their minus-end emanates from the centrosome which is tethered to the apical side. The plus-end is growing toward the basal side. As such, they provide a cellular railway for nuclear trafficking. A plus-end anchor has not been yet identified, but it has been suggested to involve IQGAP in a calcium-sensitive fashion (Kholmanskikh et al., 2006). Two microtubule binding motors, Dynein 1 and Kif1a, have been shown to drive nuclear motion during INM.

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FIGURE 14.2 Schematic presentation of interkinetic nuclear motion. Nuclei undergo a cyclic apical-to-basal motion in coordination with cell cycle progression. Nuclear motions are driven by dynein (basal-to-apical) and kinesin Kif1a (apical-to-basal). Additional mechanisms for apical-to-basal motion involve mechanical pressure between nuclei, tension along radial glia fibers. Calcium waves may provide G1 phase cells with an elevation in intercellular calcium that may drive cells toward S phase, rather than cell cycle exit. Exposure to graded Notch activity (blue gradient) along the apicalebasal axis may be partly achieved by the repeated movement of the nuclei.

14.13.1 Dynein Dynein 1, a cytoplasmic protein complex, is involved in transport of vesicular cargos toward the minus end of microtubules. In utero electroporation of rat embryonic brain with shRNA against dynein heavy chain abolished apical migration, though basal migration persisted (Tsai et al., 2010). Thus, it was concluded that dynein pulls the nucleus toward the apically tethered centrosome. This is similar to dynein activity in neuronal migration, where the lagging nucleus is pulled toward the centrosome, which is leading the motion (Tsai et al., 2007). Inhibition of dynein also prevented mitotic entry (Tsai et al., 2010). Thus, basal-to-apical motion is essential for progression of the cell cycle. Three mechanisms have been shown to recruit dynein to the nuclear envelope. First, the LINC and nucleopore protein complexes involve SUN, Syne, and Nesprin proteins as was shown in mouse cortex and retina (Zhang et al., 2009; Yu et al., 2011). In addition, two G2-phase-specific recruitment complexes were shown based on nucleoporins RanBP2 and Nup133 (Hu et al., 2013). RanBP2 binds to BicD2-Dynactin and Nup133 binds to CENPF-NudE/F-LIS1. These complexes are directly regulated by Cdk1 (Baffet et al., 2015). Cdk1 is a cell cycle regulator and plays a role in the transition from the G2 phase into mitosis. Cdk1 phosphorylates RanBP2 and induces export of CENP-F from the nucleus, thus activating both complexes. Dynein regulators LIS1 and NudE were also shown to control INM. Both genes are involved in severe neurodevelopmental disorders, lissencephaly and microcephaly (Reiner et al., 1993; Feng and Walsh, 2004). Reduction in the levels of LIS1, resulted in ectopic mitosis and impairment in INM (Gambello et al., 2003; Tsai et al., 2005; Yingling et al., 2008). Inhibition of NDE1 using shRNA in embryonic rat brains resulted in cell cycle arrest of proliferating neural progenitors and apical-to-basal INM (Doobin et al., 2016). Using optical trap assays, LIS1 and NudE were shown to enable a persistent force producing state of dynein, which is required for the transport of large structures such as the nuclei (McKenney et al., 2010; Reddy et al., 2016).

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14.13.2 Kinesin Kif1a Kinesin Kif1a was shown to have a direct role in apical-to-basal INM (Tsai et al., 2010). Kif1a is also involved in neuronal migration (Carabalona et al., 2016; Wu et al., 2018) and axonal transport (Hung and Coleman, 2016). Kif1a mutations have been implicated in neurodevelopmental disorders in humans (Esmaeeli Nieh et al., 2015). Kif1a binding to microtubules is regulated by the microtubule-associated protein DCX (Liu et al., 2012) and couples to the nuclear envelope through LINC (Wu et al., 2018). No cell cycleespecific recruitment complex has yet been identified. Kif1 inhibition by RNAi and expression of dominant-negative mutant resulted in severe, but not complete, inhibition of apical-to-basal nuclear migration (Tsai et al., 2010; Carabalona et al., 2016). Surprisingly, no interference with cell cycle was observed, and cells can enter S-phase in proximity to the ventricle in the absence of INM. In addition, Kif1 inhibition resulted in a decrease in the frequency of asymmetric neurogenic divisions was observed, which may be a clue to the role of INM in coordinating symmetric and asymmetric divisions during neurodevelopment (Esmaeeli Nieh et al., 2015; Carabalona et al., 2016).

14.14 Cytoskeleton dynamics as nuclear drivers The importance of both the actin and the microtubule cytoskeleton during INM has been shown using chemical inhibitors (Karfunkel, 1972; Messier and Auclair 1973, 1974; Webster and Langman, 1978). In zebrafish brain and retina, actomyosin is the main driver for basal-to-apical motion (Norden et al., 2009; Leung et al., 2011). Furthermore, INM is independent of dynein, microtubule, and centrosomes (Strzyz et al., 2015). Actomyosin is recruited to the basal side of nucleus a few minutes before motion, and pushes the nucleus from behind. In contrast, in mammalian neocortex, the role of actomyosin has been less clear. Several studies reported that low dosage of blebbistatin inhibited apical-to-basal motion (Schenk et al., 2009; Shinoda et al., 2018), yet another study reported no effect of blebbistatin (Tsai et al., 2010). The RhoGTPase Rnd3, a regulator of the actin cytoskeleton, has also been implicated in nuclear motion (Pacary et al., 2013). The difference between zebrafish and the neocortex may be attributed to difference in cell shapes (Lee and Norden, 2013). The zebrafish neuroepithelium cells are about 50 mm in length, which is much smaller than the 120-mm mouse neuronal progenitors. Thus, it is possible that actomyosin activity may be sufficient to drive nuclear motion over short distances, while microtubule motors are required in order to drive the nuclei along the thin and long processes of the mammalian VZ progenitors. Additionally, in the zebrafish, the motion of a single nucleus does not span the entire epithelium, and s-phase nuclei occupy a range of positions along the apical-basal axis, suggesting the apical-to-basal motion is not complete (Baye and Link, 2007).

14.15 Collective mechanisms for nuclear migration 14.15.1 Intercellular signaling The ventricular zone is a region of high cell density. Near the apical surface, cells encompass the entire space, with no extracellular gaps (Shinoda et al., 2018). Furthermore, neuronal progenitors are directly coupled through gap junctions (Weissman et al., 2004). Conventional blockers of gap junctions and transfection of cells with dominant-negative constructs of connexin 43 (Cx43) and Cx43-specific antisense oligodeoxynucleotides (asODNs) all act to slow INM (Pearson et al., 2005). shRNA knockdown of Cx43 retards the basal-to-apical motion, and blockade of gap junctions/hemichannels induces phosphorylation of cell cycle regulator Cdc42 (Liu et al., 2010). In addition, there is some evidence for cell cycle homogeneity and synchronicity in neighboring cells (Cai et al., 1997a; Cai et al., 1997b). Taken together, this suggests that intercellular signaling events may regulate nuclear migration and cell cycle. One candidate for mediating intercellular signaling is calcium. Calcium flux has been shown to regulate key cellular behaviors including proliferation and cell cycle transitions (Berridge, 2001; Webb and Miller, 2003; Humeau et al., 2018). It was shown that calcium waves spontaneously appear and propagate between neuronal progenitors and along radial fibers, in brain slices (Weissman et al., 2004; Liu et al., 2010; Rash et al., 2016). These waves, which can also occur in response to electrical or mechanical stimulation, are mediated by extracellular ATP and involve the release of calcium from IP3-sensitive intracellular stores. Calcium waves controlled the G1 to S transition and prevented cell cycle exit (Weissman et al., 2004; Rash et al., 2016). Additionally, calcium waves along radial fibers regulated neuronal migration rates. Are calcium waves used to synchronize INM, for sensing cellular density or optimizing proliferation (Malmersjö et al., 2013)? More work is required to answer this and to better understand the role of calcium signaling and dynamics during INM.

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14.15.2 Mechanical interactions The slow and diffusive apical-to-basal nuclear progression has led researchers to hypothesize that it is not driven by molecular motors. Instead, it has been suggested that nuclei are passively pushed away from the apical surface, due to the basal-to-apical moving nuclei and the high nuclear density. In this scenario, a G2 nuclei migrating to the apical surface makes room in the dense ventricular zone, by pushing a G1 nuclei away from the surface. To support this hypothesis, microbeads were incorporated into the apical side of mouse brain slices and exhibited apical-to-basal migration similar to G1 nuclei (Kosodo et al., 2011). These results were also supported by a mathematical simulation, showing that an active basal-to-apical flux of nuclei results in a passive apical-to-basal nuclear motion. Additionally, in both zebrafish and mouse, it has been shown that pharmacological inhibition of G2 to M transition completely stalls apical and basal nuclear motion, suggesting a coupling between the two motions (Kosodo et al., 2011; Leung et al., 2011). However, TAG-1 knockdown which results in loss of basal process and leads to “overcrowding” of the apical surface did not enhance apical-to-basal motion (Okamoto et al., 2013). This suggests that passive motion is not sufficient to induce apical-to-basal motion and may work in coordination with active processes. Another possible physical mechanism involved in nuclear motion is the tension along radial glia fibers and across the apical surface (Miyata and Ogawa, 2007; Okamoto et al., 2013; Okuda et al., 2013; Shinoda et al., 2018). Near the ventricle, apical processes occupy 60% of area sections, and somata occupies 40% of the area (Shinoda et al., 2018). During M-phase, the cell soma rounds up, occupying a large volume and compressing neighboring processes. This compression may lead to tension buildup, which can later release and push the daughter cells away from the apical surface during G1. This hypothesis has been supported by mechanical perturbation experiments and mathematical simulations (Shinoda et al., 2018). Finally, the high cellular density of differentiated cells has been recently suggested to mechanically limit nuclear motion on the basal side (Watanabe et al., 2018). During neurogenesis, progenitor cells can reach 300 mm in length, spanning the entire cortex with their long basal fibers, while the nuclear motion is limited to the ventricular zone, about 100 mm in thickness. It is not clear what limits the nuclei motion range, and one possibility is that the presence of differentiated cell layers mechanically limits the motion of nuclei. Indeed, when cells basal to the ventricular zone were removed via local expression of toxins, progenitor nuclei exhibited a large travel range passing the ventricular zone boundaries (Watanabe et al., 2018). Overall, more studies are required in order to demonstrate the collective, or noncell autonomous, nature of nuclear migration.

14.16 The role of INM during neurodevelopment The role of INM during neurodevelopment remains uncertain, but several hypotheses have emerged. The simplest function of INM may be to increase the surface density of mitotic phase nuclei, and thus to increase the surface density of proliferating cells (Smart, 1973; Frade, 2002; Miyata, 2008; Okamoto et al., 2013). Alternatively, it was proposed that INM in the retina determines the exposure of nuclei to apical-basal gradients of Notch and Delta (Murciano et al., 2002; Del Bene et al., 2008). Notch is increased on the apical side and prevents progenitors from leaving the cell cycle. Delta is enriched on the apical side and promotes neuronal differentiation. Thus, the time the nucleus spends in the apical or basal sides during INM, determines its exposure to delta and notch, and thus its fate. How is INM coupled to neurodevelopment? At the onset of neurogenesis (mouse E11) the epithelium is 40 mm thick (Takahashi et al., 1995). The proliferative layer (ventricular zone, VZ) thickness reaches a maximal value of about 80e120 mm, depending on the position along the anterioreposterior axis (Okamoto et al., 2014). The coupling between cell cycle and motion requires that either nuclei velocity or cell cycle phase length will adapt to the increasing travel distance. Indeed, it has been reported that the cell cycle increases from 8 to 18 h from E11 to E16, mainly due to G1 phase increase by fourfold from 3 to 12 h (Takahashi et al., 1995). However, the length of G2þS phase remained 2 h, with no observable adaptations. Thus, it suggests that basalward motion adapts to the increasing travel distance by increasing G1 phase. It has also been suggested, through the use of G1 progression inhibitor Olomoucine, that the lengthening of G1 phase may be the cause and not result of neurogenesis (Calegari and Huttner, 2003). This raises the question whether change in the progenitor layer thickness will extend G1 and affect the onset of neurogenesis.

14.17 INM summary Over the past decade, there has been a big progress in understanding the molecular players in INM. Identification of dynein and Kif1a as main nuclear drivers and regulation of dynein by Cdk1 has been important steps toward understating the

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coupling between cell cycle and nuclear motion. It remains to be seen which migration mechanisms are common and which diverge between different tissues and species. Of special interest is the characterization of INM in humans, and the recent emergence of brain organoids can provide a suitable experimental platform (Karzbrun et al., 2018). Another piece which is still far from being understood is the collective INM behavior. How is nuclear motion carried out in the crowded environment? Does density affect cell cycle, proliferation, or differentiation? What is the role of calcium signaling and are there other modes of intercellular signaling? We will have to patiently wait as the answers to these questions unfold.

14.18 Conclusions and future directions The above studies suggest that the movement of the nucleus requires the tight coordination of multiple cellular elements. Extracellular stimuli trigger intracellular signaling pathways resulting in changes in calcium concentrations, activation of phosphorylation/dephosphorylation cascades, and modulation of the assembly/disassembly kinetics of cytoskeletal elements. In radially migrating neurons, the centrosome has a key position as the hub of microtubule polymerization. The cytoskeleton is tightly linked to the nucleus and the physical forces involved in its mobilization are generated from the cytoskeleton and cytoskeletal attached molecular motors. However, the nucleus is not a passive cargo. Today it is clear that the organization of the nuclear envelope actively influences nucleokinesis. Nevertheless, it is probable that also the organization of the chromatin fiber influences the link between the centrosome and the nucleus. Furthermore, it will be interesting to investigate whether there are transcriptional mediated events occurring within the nucleus that are specifically activated by migration cues.

Acknowledgments We thank current and previous lab members for their contribution, support, and useful comments. Our research has been supported (to O.R.) in part by the Israel Science Foundation (Grant No. 347/15) and the Legacy Heritage Biomedical Program of the Israel Science Foundation (Grant No. 2041/16), ISF-NSFC joint research program (Grant No. 2449/16). This work was carried out with the aid of a grant no. 2397/18 from the Canadian Institutes of Health Research (CIHR), the International Development Research Centre (IDRC), the Israel Science Foundation (ISF), and the Azrieli Foundation, ERA-NET Neuron with support of the IMOH (Grant No. 3-0000-12276), GermaneIsraeli Foundation (GIF) (Grant no. I-1476-203.13/ 2018), United StateseIsrael Binational Science Foundation (BSF) (Grant no. 2017006), Nella and Leon Benoziyo Center for Neurological Diseases, Jeanne and Joseph Nissim Foundation for Life Sciences Research, Wohl Biology Endowment Fund, Lulu P. and David J. Levidow Fund for Alzheimer’s Diseases and Neuroscience Research, the Helen and Martin Kimmel Stem Cell Research Institute, the Kekst Family Institute for Medical Genetics, the David and Fela Shapell Family Center for Genetic Disorders Research. O.R. is an incumbent of the Berstein-Mason professorial chair of Neurochemistry. E.K. acknowledges the support of the Human Frontier Science Program (LT000629/2018-L).

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Calcium signalling during embryonic development. Nat. Rev. Mol. Cell Biol. 4 (7), 539e551. Webster, W., Langman, J., 1978. The effect of cytochalasin B on the neuroepithelial cells of the mouse embryo. Am. J. Anat. 152, 209e221. Weissman, T.A., Riquelme, P.A., Ivic, L., Flint, A.C., Kriegstein, A.R., 2004. Calcium waves propagate through radial glial cells and modulate proliferation in the developing neocortex. Neuron 43, 647e661. Wilhelmsen, K., Litjens, S.H., Kuikman, I., Tshimbalanga, N., Janssen, H., van den Bout, I., Raymond, K., Sonnenberg, A., 2005. Nesprin-3, a novel outer nuclear membrane protein, associates with the cytoskeletal linker protein plectin. J. Cell Biol. 171 (5), 799e810. Worman, H.J., Bonne, G., 2007. Laminopathies": a wide spectrum of human diseases. Exp. Cell Res. 313 (10), 2121e2133. Wu, Y.K., Umeshima, H., Kurisu, J., Kengaku, M., 2018. Nesprins and opposing microtubule motors generate a point force that drives directional nuclear motion in migrating neurons. Development (Camb.) 145. Wutz, G., Varnai, C., Nagasaka, K., Cisneros, D.A., Stocsits, R.R., Tang, W., Schoenfelder, S., Jessberger, G., Muhar, M., Hossain, M.J., Walther, N., Koch, B., Kueblbeck, M., Ellenberg, J., Zuber, J., Fraser, P., Peters, J.M., 2017. Topologically associating domains and chromatin loops depend on cohesin and are regulated by CTCF, WAPL, and PDS5 proteins. EMBO J. 36 (24), 3573e3599. Xiong, H., Rivero, F., Euteneuer, U., Mondal, S., Mana-Capelli, S., Larochelle, D., Vogel, A., Gassen, B., Noegel, A.A., 2008. Dictyostelium Sun-1 connects the centrosome to chromatin and ensures genome stability. Traffic 9 (5), 708e724. Xu, J., Ma, H., Jin, J., Uttam, S., Fu, R., Huang, Y., Liu, Y., 2018. Super-resolution imaging of higher-order chromatin structures at different epigenomic states in single mammalian cells. Cell Rep. 24 (4), 873e882. Yamada, M., Udagawa, J., Hashimoto, R., Matsumoto, A., Hatta, T., Otani, H., 2013. Interkinetic nuclear migration during early development of midgut and ureteric epithelia. Anat. Sci. Int. 88, 31e37. Yanez-Cuna, J.O., van Steensel, B., 2017. Genome-nuclear lamina interactions: from cell populations to single cells. Curr. Opin. Genet. Dev. 43, 67e72. Yang, W., Xu, Y., Wu, J., Zeng, W., Shi, Y., 2003. Solution structure and DNA binding property of the fifth HMG box domain in comparison with the first HMG box domain in human upstream binding factor. Biochemistry 42 (7), 1930e1938. Yao, Z., Duan, S., Hou, D., Wang, W., Wang, G., Liu, Y., Wen, L., Wu, M., 2010. B23 acts as a nucleolar stress sensor and promotes cell survival through its dynamic interaction with hnRNPU and hnRNPA1. Oncogene 29 (12), 1821e1834. Yingling, J., Youn, Y.H., Darling, D., Toyo-Oka, K., Pramparo, T., Hirotsune, S., Wynshaw-Boris, A., 2008. Neuroepithelial stem cell proliferation requires LIS1 for precise spindle orientation and symmetric division. Cell 132 (3), 474e486. Yoon, B.C., Jung, H., Dwivedy, A., O’Hare, C.M., Zivraj, K.H., Holt, C.E., 2012. Local translation of extranuclear lamin B promotes axon maintenance. Cell 148 (4), 752e764. Yu, J., Lei, K., Zhou, M., Craft, C.M., Xu, G., Xu, T., Zhuang, Y., Xu, R., Han, M., 2011. KASH protein Syne-2/Nesprin-2 and SUN proteins SUN1/2 mediate nuclear migration during mammalian retinal development. Hum. Mol. Genet. 20, 1061e1073. Yu, M., Ren, B., 2017. The three-dimensional organization of mammalian genomes. Annu. Rev. Cell Dev. Biol. 33, 265e289. Zhang, L., Song, D., Zhu, B., Wang, X., 2018. The role of nuclear matrix protein HNRNPU in maintaining the architecture of 3D genome. Semin. Cell Dev. Biol. Zhang, Q., Ragnauth, C., Greener, M.J., Shanahan, C.M., Roberts, R.G., 2002. The nesprins are giant actin-binding proteins, orthologous to Drosophila melanogaster muscle protein MSP-300. Genomics 80 (5), 473e481. Zhang, Q., Ragnauth, C.D., Skepper, J.N., Worth, N.F., Warren, D.T., Roberts, R.G., Weissberg, P.L., Ellis, J.A., Shanahan, C.M., 2005. Nesprin-2 is a multi-isomeric protein that binds lamin and emerin at the nuclear envelope and forms a subcellular network in skeletal muscle. J. Cell Sci. 118 (Pt 4), 673e687. Zhang, Q., Skepper, J.N., Yang, F., Davies, J.D., Hegyi, L., Roberts, R.G., Weissberg, P.L., Ellis, J.A., Shanahan, C.M., 2001. Nesprins: a novel family of spectrin-repeat-containing proteins that localize to the nuclear membrane in multiple tissues. J. Cell Sci. 114 (Pt 24), 4485e4498. Zhang, X., Lei, K., Yuan, X., Wu, X., Zhuang, Y., Xu, T., Xu, R., Han, M., 2009. SUN1/2 and Syne/Nesprin-1/2 complexes connect centrosome to the nucleus during neurogenesis and neuronal migration in mice. Neuron 64 (2), 173e187. Zhang, X., Xu, R., Zhu, B., Yang, X., Ding, X., Duan, S., Xu, T., Zhuang, Y., Han, M., 2007. Syne-1 and Syne-2 play crucial roles in myonuclear anchorage and motor neuron innervation. Development 134 (5), 901e908. Zhen, Y.Y., Libotte, T., Munck, M., Noegel, A.A., Korenbaum, E., 2002. NUANCE, a giant protein connecting the nucleus and actin cytoskeleton. J. Cell Sci. 115 (Pt 15), 3207e3222. Zhou, K., Rolls, M.M., Hall, D.H., Malone, C.J., Hanna-Rose, W., 2009. A ZYG-12-dynein interaction at the nuclear envelope defines cytoskeletal architecture in the C. elegans gonad. J. Cell Biol. 186 (2), 229e241.

Chapter 15

Radial migration in the developing cerebral cortex Stephen C. Noctor1, Christopher L. Cunningham2 and Arnold R. Kriegstein3, 4 1

Department of Psychiatry and Behavioral Sciences, UC Davis MIND Institute, Sacramento, CA, United States; 2Solomon H. Snyder Department of

Neuroscience, Johns Hopkins University School of Medicine, Baltimore, MD, United States; 3Department of Neurology, University of California, San Francisco, CA, United States; 4The Eli and Edythe Broad Center of Regeneration Medicine and Stem Cell Research, University of California, San Francisco, CA, United States

Chapter outline 15.1. 15.2. 15.3. 15.4. 15.5. 15.6. 15.7.

Introduction Production of cortical projection neurons Organization of the neocortex Trajectory of migrating neurons in the developing brain Modes of migration Radial migration in the developing human neocortex Factors that regulate the radial migration of cortical neurons 15.7.1. Secreted molecules 15.7.1.1. Reelin 15.7.1.2. Semaphorins 15.7.2. Neurotransmitters 15.7.2.1. GABA 15.7.2.2. Glutamate 15.7.2.3. ATP

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15.7.3. Adhesion molecules 15.7.3.1. Integrins 15.7.3.2. Gap junctions 15.7.4. Cytoskeletal regulators 15.7.4.1. Lis1 15.7.4.2. Doublecortin 15.7.4.3. Filamin A (FLNA/FLN1) 15.7.4.4. Cdk5 15.7.5. Transcription factors 15.7.5.1. Pax6 15.7.5.2. Tbr2 15.7.5.3. Neurogenins 15.8. Summary References

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15.1 Introduction The mature human cerebral cortex comprises approximately 20e25 billion neurons and 30e40 billion glial cells according to stereological estimates (Pelvig et al., 2008), or 16 billion neurons and 60 billion nonneuronal cells as determined through isotropic fractionation (Azevedo et al., 2009). Remarkably, the generation of tens of billions of cells of diverse, complex cell types is accomplished over a span of intense proliferation before birth. During prenatal development the cerebral cortex is initially condensed into a compact laminar structure called the cortical plate (CP) before it expands and develops into its mature form. The CP comprises approximately 1 billion cells at the 13th week of gestation and increases to approximately six billion cells at the 20th week of gestation (Samuelsen et al., 2003). Most newly arrived CP cells are neurons since birthdating studies have shown that cortical neurons are generated prior to glial cells. For example, retrospective 14C dating of human cortical cells showed that the age of cortical neurons matched that of an individual, while glial cells were younger than the individual, indicating a postnatal origin for many glial cells (Spalding et al., 2005). An increase in the number of CP cells by 5 billion between the 13th and 20th weeks of gestation suggests that a similar number of neurons were produced in roughly the same amount of time. Thus, given that 5 billion cortical cells are produced over 7 weeks, more than 1100 new neurons were produced every second during this stage of cortical development. It therefore follows that

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over 1000 cortical neurons begin their migration to the CP each second. Recent studies indicate that cortical neurogenesis continues until 35 weeks of gestation (Arshad et al., 2016). As a result, the intense period of cortical neuron migration continues over a lengthy period lasting several months. Given the high levels of cell production during gestation, even subtle variations in the rate of proliferation have the potential to greatly alter the number of migrating cortical neurons. For example, a reduction in the rate of proliferation equal to just one half of 1% would reduce the number of newborn neurons migrating to the CP that numbers in the millions. The two principal neuronal subtypes, projection neurons and local circuit interneurons, are produced in different regions of the developing brain (Parnavelas, 2000). Excitatory projection neurons are produced in the proliferative zones of the dorsal forebrain and migrate radially to reach the CP. In contrast, work in rodents shows that inhibitory cortical neurons are produced in the ganglionic eminences of the basal forebrain and migrate tangentially to reach the CP in the dorsal neocortex (De Carlos et al., 1996; Anderson et al., 1997; Tamamaki et al., 1997; Wichterle et al., 1999; McKinsey et al., 2013) (see Noctor Fig. 15.1 and Chapter 16). Genesis of interneurons in primates may follow this pattern. While evidence has been reported that in the human and nonhuman primate neocortex some interneurons are produced in the dorsal cortex (Letinic et al., 2002; Fertuzinhos et al., 2009; Petanjek et al., 2009), other studies have found that the majority of primate neocortical GABAergic interneurons originate from ganglionic eminences (Ma et al., 2013). This chapter focuses on the radial migration of excitatory projection neurons in the dorsal cerebral cortex. The newly born projection neurons face the formidable task of migrating substantial distances across the cortical wall from their place of birth in the proliferative zones to their target destination in the CP. The mechanisms that regulate radial migration must be tightly regulated to ensure that the vast numbers of cortical neurons embarking for the CP each second arrive at their proper destination during the several months of cortical neurogenesis in humans. Not surprisingly, this period of development is sensitive to genetic abnormalities and/or environmental interference, and multiple cortical malformations have been linked to errors of proliferation and cellular migration during brain development including epilepsy, lissencephaly, dyslexia, mental retardation, fragile X, and autism.

15.2 Production of cortical projection neurons Advances in the field of molecular biology over the past decade have provided new tools for investigation of the developing brain. Molecular approaches readily allow researchers to control gene expression in the developing neocortex in a regional, temporal, and cell-specific manner. In addition, fluorescent reporter proteins have proven invaluable for detailed characterization of cellular morphology and function in living tissue (Prasher et al., 1992; Shimomura, 2005; Haldar and Chattopadhyay, 2009). Inducing reporter gene expression into discrete populations of mitotic neural precursor cells and migrating neuronal cells has been achieved through injection of retroviral vectors (Palmer et al., 1999), DNA plasmids (Saito, 2006), and mutant mice, e.g., CreeLoxP (Hippenmeyer et al., 2010). Visualizing precursor cells and migratory neurons in live tissue using fluorescent reporter proteins or other fluorescent molecules fundamentally enhanced our understanding of neurogenesis, the patterns of neuronal migration, and the relationships between precursor cells and migrating neurons in the developing neocortex. Cortical projection neurons are produced in two proliferative zones that line the ventricular system of the developing brain. Each proliferative zone is home to a distinct class of precursor cell. The primary proliferative zone, the ventricular zone (VZ) is directly adjacent to the lumen of the ventricle and is present from the moment of neural tube closure. The secondary proliferative zone, the subventricular zone (SVZ), is superficial to the VZ and appears just prior to the peak onset of neurogenesis in the cerebral cortex of vertebrates (Angevine et al., 1970; Bayer and Altman, 1991). The size of the SVZ varies across mammalian species, and at later stages of neurogenesis the SVZ can be subdivided into an inner and outer SVZ (Smart et al., 2002). The outer SVZ is present in some rodents at the end of neurogenesis and is substantially larger in nonhuman primates (Martinez-Cerdeño et al., 2012). VZ precursor cells are specialized neuroepithelial cells that are called radial glial cells during the neurogenic phase of cortical development (Rakic, 2003a). Radial glia are bipolar cells that have a soma in the VZ, a short descending process that contacts the ventricular lumen, and an ascending pial process that spans the wall of the developing brain to contact the pial membrane. The pial processes of radial glia radiate outward from the ventricle to the surface of the brain in a pattern that dominates the appearance of the brain during early stages of development, and to a degree dictates organization of the mature cerebral cortex (Rakic, 1988). Radial glial cells were initially proposed to serve as support cells that maintained the shape of the nervous system during development (see Bentivoglio and Mazzarello, 1999). In the 1970s, Rakic’s fundamental discovery that radial glial cells play a central role in guiding the radial migration of newborn neurons in the cerebral cortex was an important milestone in developmental studies (Rakic, 1971).

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FIGURE 15.1 Scheme depicting proliferative zones during neurogenesis and the formation of minicolumn and cortical column functional units in the prenatal mammalian cerebral cortex. Each minicolumn contains approximately 80e100 neurons. Multiple minicolumns together form a single cortical column. As proposed by Rakic (1988), proliferative units in the ventricular zone (VZ) of the dorsal forebrain produce neurons that are destined for a single minicolumn functional unit (dark blue column) in the CP (CP). Each proliferative unit in the VZ may comprise 5e10 radial glial cells. Radial glial cells in the VZ can be identified by expression of the transcription factor Pax6 (red). In addition, each radial glial cell possesses a single pial fiber that stretches across the cortical wall toward the pial surface of the brain. The pial fiber guides the migration of newborn neurons (blue) to their respective minicolumn functional units in the CP. Neighboring proliferative units in the VZ produce cells that are destined for adjacent minicolumns in the CP (gray columns). Radial glial cells produce neurons (blue) and intermediate progenitor cells (yellow) that migrate away from the ventricle to establish the subventricular zone (SVZ). Intermediate progenitor (IP) cells express the transcription factor Tbr2 (yellow). Intermediate progenitor cells in rodents divide primarily in the SVZ and produce pairs of neurons (blue). Recent work shows that the SVZ is subdivided into an inner SVZ comprising a dense band of Tbr2þ neurogenic IP cell and an outer SVZ (yellow) comprising Pax6þ radial glial cells that have translocated away from the VZ to the oSVZ and retain their pial fiber, and a diffuse band of neurogenic Tbr2þ IP cells. Excitatory projection neurons are derived from the VZ and SVZ of the dorsal forebrain and migrate radially along radial glial cell fibers. Inhibitory interneurons (gray) are derived from the proliferative zones of the medial ganglionic eminence (MGE, gray) in the basal forebrain and migrate tangentially into the overlying cerebral cortex. Mechanisms that direct the migration of interneurons to specific cortical minicolumns remain to be determined.

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Work over the last decade has added to this picture by demonstrating that radial glial cells also produce cells in the neuronal lineage (Malatesta et al., 2000; Miyata et al., 2001, 2004; Noctor et al., 2001, 2002, 2004, 2008; Tamamaki et al., 2001; Anthony et al., 2004; Attardo et al., 2008). SVZ precursor cells are generated by radial glial cells in the VZ (Noctor et al., 2004), and then migrate radially to a position just superficial to the VZ where they establish the SVZ compartment (Bayer and Altman, 1991). SVZ precursor cells are known by several names including subependymal cells, intermediate progenitor cells, and basal progenitors. This review will use the term “intermediate progenitor cell” (Noctor et al., 2004). During prenatal cortical development, SVZ precursor cells produce cortical neurons (Tarabykin, 2001; Letinic et al., 2002; Rakic, 2003b; Haubensak et al., 2004; Miyata et al., 2004; Nieto et al., 2004; Noctor et al., 2004, 2008; Hansen et al., 2010), which migrate radially to the CP along radial glial fibers (Noctor et al., 2008). Unlike radial glia that usually divide asymmetrically to produce single daughter cells, intermediate progenitor cells often divide symmetrically to produce pairs of daughter neurons (Noctor et al., 2004, 2008). The proliferative VZ becomes progressively thinner during development and is not present in the adult brain (Bayer and Altman, 1991; Smart et al., 2002), but the neurogenic SVZ compartment remains in the adult cerebral cortex and produces neurons destined for the olfactory bulb throughout life (Lim and Alvarez-Buylla, 2016). The bulk of research on development of the cerebral cortex has relied upon rodent models to reveal mechanisms that guide neuronal migration. Rodent models are tremendously useful (Martinez-Cerdeno and Noctor, 2019), but recent work has highlighted significant differences between the development of rodent and primate cerebral cortex. For example, the SVZ is a comparatively simple structure in rodents, whereas in primates and some other gyrencephalic mammals, the SVZ is larger and includes discrete functional areas that are called the inner SVZ and the outer SVZ (Smart et al., 2002). Radial glial cells in the rodent VZ can be identified by expression of the transcription factor Pax6, and intermediate progenitor cells in the rodent SVZ can be identified by expression of the transcription factor Tbr2 (Bulfone et al., 1999; Englund et al., 2005; Hevner, 2019). The inner SVZ in primates appears to be similar to the rodent SVZ in cellular density and cellular composition (Martinez-Cerdeño et al., 2012). In contrast, the outer SVZ in primates contains Tbr2þ cells and Pax6þ radial glial cells that have translocated from the VZ to a position above the inner SVZ (Hansen et al., 2010). The outer SVZ is present in embryonic rats at the end of gestation and contains the same sets of precursor cells, but is significantly larger in primates than in rodent model species (Martinez-Cerdeño et al., 2012).

15.3 Organization of the neocortex The mammalian cerebral cortex is a laminated sheet of tissue that is 1e4 mm thick (Martinez Cerdeno et al., 2017) and organized into six horizontally arranged layers. This elaborate organization is further broken down into radial columns, or cortical columns, that span across the six cortical layers and are thought to represent discrete functional units (Mountcastle, 1957, 1997). Each cortical column comprises multiple “minicolumns” of approximately 80e140 radially arranged neurons that span the cortical layers (Rakic, 1988; Jones, 2000) (Noctor Fig. 15.1). The columnar or radial organization of the cerebral cortex is thought to derive from the organization of cells in the embryonic VZ and the radial deployment of newly born migrating neurons. Rakic’s Radial Unit Hypothesis proposed that precursor cells in the VZ (radial glia) are organized into discrete proliferative units that establish a proto-map of cortical columns in the adult cerebral cortex (Rakic, 1988). The Radial Unit Hypothesis predicts that each proliferative unit in the VZ produces the cortical neurons that populate a cortical column (Rakic, 1988) (Noctor Fig. 15.1). Data showing substantial neurogenesis in the rodent embryonic SVZ (Haubensak et al., 2004; Miyata et al., 2004; Noctor et al., 2004), and the inner and outer SVZ of primates (Kriegstein et al., 2006; Fietz et al., 2010; Hansen et al., 2010), have led to modifications of the Radial Unit Hypothesis to specify the contribution of precursor cells in the SVZ and particularly the radial glial-like cells in the outer SVZ of primates (see Fig. 15.1).

15.4 Trajectory of migrating neurons in the developing brain One of the remarkable aspects of cortical development is that neurons are not born in their adult location, but instead must migrate substantial distances, up to 7000 mm or more for neurons generated near the lateral ventricle, to reach their destination. To put this into perspective, the migration of cortical projection neurons from the VZ to the CP would be comparable to a person scaling a wall greater in height than Yosemite National Park’s 3000-foot high El Capitan and arriving at a specific height at the proper time. In the developing neocortex, excitatory cortical neurons are generated in an inside-out sequence such that the deepest layers of the cerebral cortex are born and migrated into the cortical mantle before the superficial layers (Angevine and Sidman, 1961). Thus, as development proceeds neurons must migrate progressively longer distances through an increasing number of cortical cells. Despite the complexity, this feat is achieved with such

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regularity and precision that there is a remarkable degree of uniformity in the architectonic pattern of cortical areas from one person to the next. Experiments using fluorescently labeled cells and/or time-lapse imaging in cultured brain tissue have revealed the complex patterns of neuronal migration in the neocortex. Experiments in the 1990s demonstrated that in rodents cortical inhibitory interneurons are generated in the ventral forebrain and migrate long distances along a tangential trajectory into the overlying dorsal neocortex (De Carlos et al., 1996; Anderson et al., 1997; Tamamaki et al., 1997; Wichterle et al., 1999) (see Chapter 16). Similar findings have been reported in primates (Letinic et al., 2002; McKinsey et al., 2013), but some interneuron subtypes may also be produced in the dorsal neocortex of primates (Letinic et al., 2002), for review see (Jones, 2009). Time-lapse microscopy of fluorescently labeled cells in the dorsal neocortex has since revealed that projection neurons also exhibit complex patterns of migration. Projection neurons do not simply migrate along a direct radial trajectory from the proliferative zones to the CP, but instead undergo distinct stages of migration over a period of days that can be identified based on the morphology, orientation, and position of immature neurons in the cortical wall (Kriegstein and Noctor, 2004; Noctor et al., 2004). The first stage of migration for cortical neurons produced in the dorsal VZ entails detachment of the newborn cell from the ventricular surface and rapid ascension to the SVZ along the pial fiber of the mother radial glial cell. The second stage of migration is marked by a dramatic change in neuronal morphology; cortical neurons acquire a multipolar morphology and remain in the SVZ for up to one day. Multipolar neurons appear to maintain an affiliation with the mother radial glial cell fiber. The migratory pause of young cortical neurons in the SVZ had been noted in previous work and was originally termed “sojourning” (Bayer and Altman, 1991), and the multipolar morphology of radial glial daughter cells in the SVZ has been noted by multiple investigators (Noctor et al., 2001, 2004, 2008; Tabata and Nakajima, 2003; Tabata et al., 2009). Though neurons sojourning in the SVZ do not migrate appreciable distances in the radial direction, the cells are nonetheless very active. The neurons repeatedly extend and retract multiple processes in a manner that suggests a search for environmental cues (Tabata and Nakajima, 2003; Noctor et al., 2004). After remaining in the SVZ for one day or longer, many cortical neurons begin the third stage of migration by extending a process toward the ventricular surface and in most cases exhibiting a retrograde somal movement toward the ventricle (Noctor et al., 2004). Among excitatory cortical neurons, the cellular process that is extended into the VZ is thin, resembles an axon, and often appears to contact the surface of the lateral ventricle. Neurons in the third stage of migration may spend as long as 24 h near the margin of the ventricle before beginning the fourth and final stage of migration. The considerable amount of time that immature neurons spend in the VZ and SVZ before initiating migration to the CP indicates that the cells interact with their environment during stages 1e3 and suggests the possibility that doing so may be a prerequisite for those cells to reach their appropriate target. Interestingly, cortical interneurons have also been shown to make ventricle-directed movements after migrating tangentially into the dorsal cortex and before initiating radial migration toward the CP (Nadarajah et al., 2002). This further suggests a coordination of the migration of excitatory and inhibitory cortical cells that may contribute to their integration into the CP and points to a potentially crucial source of migration guidance molecules located near the ventricular lumen. Indeed, recent findings have shown that upon arriving in the CP, neurons initially form discrete clusters of cells that include both excitatory and inhibitory neurons that remain contact with one another for approximately one day before exiting the clusters and continuing morphological maturation (Shin et al., 2019). The final stage of migration is marked by the growth of a leading process directed toward the pial surface and commencement of radial migration toward the CP (Noctor et al., 2004). The rudimentary axonal-like process that extends from the neuron toward the ventricle during stage 3 remains in the VZ as the neuron migrates radially to the CP (Noctor et al., 2004). The portion of this trailing process that remains in the VZ may be vestigial since collateral processes that branch off and begin growing in the intermediate zone presumably represent the mature trajectory of the axon (Noctor et al., 2004). Throughout the first three stages of migration most newborn daughter neurons appear to remain closely attached to the pial fiber of their mother radial glial cell (Noctor et al., 2004, 2008), highlighting the importance of lineage relationships in the developing brain. However, fidelity between the daughter neuron and the parental pial fiber appears to decrease during the final stage of migration: Approximately, one-third of the daughter cells remain attached to the parental radial glial fiber after they have reached the CP (Noctor et al., 2001). The remaining cells do not stray very far from the parental fiber and migrate radially in parallel with their siblings along neighboring radial glial fibers that are only a few cell diameters distant from the parent fiber. Cortical neurons that are produced by intermediate progenitor cell divisions in the SVZ also exhibit similar changes in morphology, orientation, and position that mark the stages of migration for neurons generated in the VZ. The SVZ-derived neurons first acquire a multipolar morphology, next they extend a thin axonal-like process toward the ventricle and migrate toward the surface of the VZ, and finally commence radial migration toward the CP (Noctor et al., 2004). The pairs of SVZ daughter neurons also appear to maintain a strong affiliation with their parental pial fiber. Time-lapse recordings in the embryonic rodent neocortex have captured intermediate progenitor cell divisions in the SVZ that produce a pair of daughter cells that both migrate toward the CP along their parental pial fiber (Noctor et al., 2008).

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Interneurons that are generated in the basal forebrain migrate perpendicular to the radial glial matrix and may not rely upon an association with radial glial cells for guidance during the long tangential phase of their journey into the dorsal neocortex. But upon reaching their target destination in the neocortex the interneurons appear to associate with radial glial fibers, switch from a tangential to a radial orientation, and migrate radially into the developing CP (Polleux et al., 2002; Poluch and Juliano, 2007; Yokota et al., 2007; Elias et al., 2010). Each successive wave of newborn projection neurons generated in the proliferative zones is thought to go through similar sequential changes in morphology, orientation, and position that mark the migration of projection cells toward the CP (Noctor et al., 2008). This finding suggests that temporal changes in extrinsic signaling in the cortical wall do not drive the changes in neuronal morphology or orientation exhibited by migrating projection neurons in the dorsal cortex, but rather that projection neurons respond to persistent signals that are expressed or perhaps secreted within distinct laminar structures of the neocortex during neurogenic phases of prenatal development. Furthermore, it is likely that newborn neurons follow a maturational program that promotes a sequential or temporal change in responsiveness to signals that originate from different strata of the cortical wall in the following sequence: (1) signals that originate in or near the SVZ during the second multipolar stage of migration; (2) signals that originate near the ventricle during the third stage of retrograde migration toward the ventricle; and finally (3) signals originating from the superficial layers of the cortical wall during the fourth stage of radial migration. This scheme outlines a developmental program that regulates the radial migration in the neocortex. Other important signaling mechanisms have been shown to guide migration and undoubtedly help to shape the response of immature neurons to extrinsic signaling factors. For example, establishment of neuronal polarity (Barnes and Polleux, 2009) and gap junction coupling between migrating neurons and radial glial cells (Fushiki et al., 2003; Elias et al., 2007; Matsuuchi and Naus, 2013) are two factors that have been shown to regulate radial migration that will be discussed further below. The changes in responsiveness to extrinsic signaling factors and the morphological changes exhibited by the immature neurons as they progress through the stages of radial migration most likely rely on different intracellular mechanisms. Neuronal migration is thus a complex interplay between the migrating cell and its environment that relies on intracellular machinery as well as proper responsiveness to extrinsic signaling. The large number of factors that are required for neuronal migration presents greater opportunity for interference with the process and may have exerted evolutionary pressure that selected for redundant signaling molecules or pathways, such as doublecortin and doublecortin-like (Sapir et al., 2000). Not surprisingly, neuronal migration is sensitive to genetic abnormalities and/or environmental interference, and a number of nervous system malformations have been identified that result from defects in neuronal migration (Manzini and Walsh, 2011). Furthermore, multiple disorders have been linked to errors of neuronal migration, including epilepsy, lissencephaly, mental retardation, fragile X, and autism. The cost borne by society for these disorders is enormous. For example, epilepsy affects 1.2% of the population in the United States with total direct costs of approximately $10 billion, and $10,000 to $20,000 annual cost per patient depending on severity (Cramer et al., 2014; Begley and Durgin, 2015; Zack and Kobau, 2017). This estimated cost includes components such as health care and work loss but does not account for the personal difficulties faced by individuals and families coping with developmental disorders on a daily basis.

15.5 Modes of migration In addition to the stages of radial migration outlined above, distinct “modes” of migration, locomotion, multipolar, and somal translocation have been described for radially migrating cortical neurons. The “locomotion” mode of migration refers to freely migrating immature neurons that have a leading process that extends tens of microns to hundreds of microns from the cell body depending on the brain region, the animal model, and the stage of development under study. Freely migrating immature neurons, or neuroblasts as they are sometimes called, have been described in numerous publications including the earliest studies of the developing cerebral cortex (His, 1889; Ramón y Cajal, 1995), later work based on electron microscopy (Rakic, 1971), and work based on fluorescent labeling techniques (O’Rourke et al., 1992; Fishell et al., 1993; Tamamaki et al., 1997; Miyata et al., 2001; Nadarajah et al., 2001; Noctor et al., 2001; Tamamaki et al., 2001; Nadarajah et al., 2002). Locomoting cells migrate through repeated saltatory motions in which the migrating cell extends a leading process approximately 50e100 microns from the soma, and then translocates the nucleus into the leading process as the cell moves forward (Schaar and McConnell, 2005; Shieh et al., 2011). Locomoting cells appear to respond to attractive and repulsive factors to find their target destination. Upon migrating to the SVZ cortical neurons exhibit a “multipolar” mode of migration prior to initiating radial migration to the CP (Tabata and Nakajima, 2003). The “somal translocation” mode of migration refers to migrating cells that have a long leading process that is fixed at the target destination of the migrating cell. This class of cells migrates by transporting the nucleus within the already established

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leading process until it reaches the target destination. Before commencing migration, the translocating cells must either extend a leading process to the target destination through a pathfinding process not unlike that of the leading process for “locomoting” cells, or inherit a preexisting process in contact with the target destination. Some proponents of translocation in the embryonic neocortex propose that newborn neurons inherit the pial fiber of their parent radial glial cell after mitosis and the nucleus of the daughter neuron is then transported to the CP within the pial process. This mode of migration would presumably simplify the migratory process by not requiring radial glial cells for directional guidance. Magini may have been the first to link cells with long radial processes to neuronal migration. He noted varicosities on radial fibers in Golgi-stained preparations of the embryonic cortex and proposed that the varicosities represented the nuclei of neurons migrating along single radial cell processes (Magini, 1888). Berry and Rogers also studied Golgistained tissue and proposed that radial glial cells were binucleate and that the nuclei of newborn neurons translocated to the CP within radial glial pial fibers (Berry and Rogers, 1965). With the advent of new staining and microscopic techniques, it has been shown that radial glia are not multinucleated, and that the pial fiber varicosities visualized by Magini, Berry, and others in Golgi-stained material are not separate cells but rather enlargements of radial glial pial fibers, and are often associated with M-phase radial glial cells (Weissman et al., 2003; Noctor et al., 2007, 2019). Morest examined Golgi-stained cells in the developing opossum cortex and suggested that translocation may represent a distinct mode of migration for cortical neurons (Morest, 1970). In fact, time-lapse imaging studies performed in multiple mammalian species from rodent to primate have readily identified translocating cells in the embryonic neocortex (Miyata et al., 2001, 2004; Nadarajah et al., 2001; Noctor et al., 2004, 2008; Hansen et al., 2010; Wang et al., 2011; Betizeau et al., 2013; Pfeiffer et al., 2016), demonstrating that translocation is an important component in the developing neocortex. However, currently available evidence indicates that while translocating cells are identifiable at many stages of cortical development, most of these cells are not candidates for neuronal identity. For example, Brittis and colleagues described cells with processes that stretched from the ventricle to the pial surface at early stages of cortical development and concluded these cells were migrating neurons (Brittis et al., 1995). However, these authors used a monoclonal antibody against phosphorylated growth-associated protein-43 (GAP-43), which is expressed by both neurons (Meiri et al., 1991) and mitotic precursor cells (Brittis et al., 1995; Esdar et al., 1999), and thus cannot rule out a precursor cell identity for phospho-GAP-43þ cells. It is also worth noting that phosphorylated GAP-43þ cells are morphologically identical to cells labeled with antibodies against phosphorylated vimentin, and the location and distribution of the phospho-GAP-43þ cells in the embryonic cerebral cortex matches the distribution of mitotic cells labeled with other phospho markers including phospho-vimentin and phospho-histone 3: at the surface of the ventricle where radial glial cells divide and in the SVZ where intermediate progenitor cells divide (Weissman et al., 2003; Cunningham et al., 2013). Other markers that have been used to identify newborn neurons in the embryonic neocortex raise similar issues. For example, anti-Hu antibodies label both immature neurons and precursor cells and thus may not positively identify one specific cell type (Miyata et al., 2004). Of interest, translocating cells described in many time-lapse imaging studies match previous morphological descriptions of radial glial cells that transform into GFAP-expressing astrocytes at the end of cortical neurogenesis in ferret and nonhuman primates (Schmechel and Rakic, 1979; Voigt, 1989). In addition, electrophysiological recordings obtained from translocating cells in embryonic rat cerebral cortex showed that translocating cells lack the inward voltage-gated currents that are expressed by immature cortical neurons (Noctor et al., 2001, 2004, 2008) and express nuclear transcription factors, such as Pax6, that are associated with radial glia (MartinezCerdeño et al., 2012). An important caveat concerning whole-cell recordings obtained from translocating cells is that this does not rule out the possibility that these cells could begin expressing neuron-specific membrane channels at a later time. Studies in the developing human neocortex have also reported that translocating cells do not express neuronal specific markers (de Azevedo et al., 2003) and remain mitotically active (Hansen et al., 2010). These data are consistent with results obtained in rodent and support the idea that translocating cells that migrate from the ventricular surface toward the CP are not neurons, but rather precursor cells in the radial glial/astrocyte lineage. Translocation of neurons over shorter distances has been reported in the developing brain. Evidence suggests that upon reaching the CP, some locomoting neurons extend a leading process through the CP and can then migrate to the top of the CP through nuclear translocation. Additionally, neurons in other regions of the central nervous system, such as the E10-E13 mouse hindbrain, have been shown to migrate through somal translocation (Hawthorne et al., 2010). The distances involved in some of these regions, up to 100 mm, are similar in length to that of the leading process for a typical locomoting neuron in the developing neocortex. Therefore, it remains possible that these cells extend a leading process to their target and then migrate away from the germinal zones near the ventricular lumen. In summary, somal translocation as a model of cellular migration remains an important topic. The relative proportions of locomoting versus translocating cells among cortical neurons may vary developmentally and across regions of the developing brain, but this remains to be determined.

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15.6 Radial migration in the developing human neocortex The developing human neocortex contains a very large number of outer subventricular radial glia (oRG cells) that participate in neurogenesis and guide neuronal migration (Hansen et al., 2010). Similar cells have also been observed in the developing cortex of the ferret (Fietz et al., 2010; Reillo and Borrell, 2012; Poluch and Juliano, 2015) and nonhuman primates (Martinez-Cerdeño et al., 2012; Betizeau et al., 2013). The oRG cells are not embedded in the ventricular epithelium and do not have a process in contact with the lateral ventricle, yet they undergo asymmetrical cell division to produce intermediate progenitor cells, that in turn divide multiple times within the SVZ to generate cortical neurons. The oRG cells also have pial-directed fibers that support neuronal migration (Hansen et al., 2010). The presence of oRG cells provides a mechanism for greatly increasing neuronal production while also providing additional radial guides for neuronal migration, thus providing a solution for cortical expansion without requiring a proportionate increase in ventricular size. The presence of a large abventricular neurogenic zone alters the concept of the radial unit hypothesis, as it interposes a large, nonventricular, source of ontogenetic units, and could modify the point-to-point mapping of the ventricular epithelial pattern onto the cortical mantle. In addition to these theoretical issues, the pattern of OSVZ neurogenesis suggests that the four stages of neuronal migration established for ventricular neurogenesis may not apply to neurogenesis in this region. oRG cells undergo a process of mitotic somal translocation during cell division that is characterized by movement of the cell body and nucleus toward the pial surface of approximately 55 mm (Hansen et al., 2010; Gertz et al., 2014). When the oRG cell undergoes cytokinesis, the cleavage plane is orthogonal to the radial axis, with the self-renewed oRG cell in the basal position, leaving the daughter intermediate progenitor cell out of contact with the parental radial fiber. With subsequent oRG cell divisions, and the associated mitotic somal translocations, the parent radial glial cell and fiber move progressively farther away from the intermediate progenitor cells, which undergo multiple rounds of division without significant radial migration. The intermediate progenitor cells will eventually produce neurons that migrate radially, but presumably on nonparental radial glial fibers. This suggests that cells are free to disperse tangentially within the OSVZ prior to radial migration and make a strict radial ontogenetic column unlikely (Ostrem et al., 2017).

15.7 Factors that regulate the radial migration of cortical neurons In the following section, we briefly summarize the roles of select factors that regulate the migration of embryonic cortical neurons.

15.7.1 Secreted molecules 15.7.1.1 Reelin Reelin is a large glycoprotein secreted by CajaleRetzius cells in the marginal zone of the prenatal neocortex that interacts with the extracellular matrix and regulates the radial migration and laminar formation of cortical structures (Chai and Frotscher, 2016). The reeler mouse was first described in 1951 by Falconer (Falconer, 1951). Subsequent work revealed a defect in the laminar organization of the cerebral cortex (Hamburgh, 1963), produced by a near inversion of the cortical layers despite a normal birth order sequence of cortical neurons (Caviness and Sidman, 1973). The reeler cortical phenotype results from a mutation in the Reln gene (D’Arcangelo et al., 1995), which drives expression of Reelin protein by CajaleRetzius cells in the marginal zone of the prenatal neocortex (Ogawa et al., 1995). The human homolog, RELN, was subsequently sequenced (DeSilva et al., 1997), and mutations in the RELN gene were found to be associated with an autosomal recessive form of lissencephaly (Hong et al., 2000), providing further evidence to support the concept that Reelin plays a critical role in cortical lamination and radial migration. Studies in the 1980s reported an increased contact between radial glial cells and migrating neurons in the reeler mouse (Pinto-Lord et al., 1982). Subsequent studies suggested that Reelin regulates the morphology of both radial glial cells and cortical neurons in the dentate gyrus (Forster et al., 2002) and neocortex (Hartfuss et al., 2003; Chai et al., 2015). Evidence shows that one function of Reelin secreted by CajaleRetzius cells in the marginal zone may be to act as a stop signal that induces termination of radial migration at the top of the CP (Frotscher et al., 2017), by activating integrin receptors and N-cadherin (Chai and Frotscher, 2016; Hirota and Nakajima, 2017). Migrating neurons express two receptors that bind Reelin, the very low density lipoprotein receptor (VLDLR) and the apolipoprotein E receptor 2 (ApoER2), which are members of the LDL family of lipoprotein receptors (D’Arcangelo et al., 1999; Hiesberger et al., 1999). Reelin bound to VLDLR or ApoER2 results in phosphorylation of the intracellular adaptor protein Disabled-1 (Dab-1), which interacts with intracellular domains of VLDLR or ApoER2. Dab-1 is phosphorylated by Src family kinases (Arnaud et al., 2003; Kuo et al., 2005), leading to cytoskeletal changes that modulate migratory behavior (Chai et al., 2009). There are many regulatory checkpoints in the Reelin signaling pathways. Notably, activated

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Dab-1 is degraded by the E3-ubiquitin ligase, Cullin-5 (Feng et al., 2007), which regulates Reelin-dependent cortical lamination and migration by modulating Dab1 degradation (Simo et al., 2010). Genetic knockout of Notch signaling in migrating neurons induces defects in migration that mirror those in the reeler mouse, and the cortical lamination defects in reeler mice can be rescued by overexpression of the Notch intracellular domain (Frotscher et al., 2017). The apparent cross talk between these signaling pathways is thought to be mediated by Dab-1, which prevents degradation of the Notch intracellular domain (Hashimoto-Torii et al., 2008). Reelin signaling has also been linked to the dynamics of actin filament polymerization in migrating cells. N-cofilin, a molecule that promotes the formation of new actin filaments, is phosphorylated in the leading process of migrating neurons by Reelin signaling through ApoER2 (Chai et al., 2009). Phosphorylated n-cofilin loses its capacity to depolymerize Factin and thereby exerts a stabilizing effect on the neuronal cytoskeleton, suggesting the possibility that Reelin may act as a stop signal by actions on the neuronal cytoskeleton (Frotscher et al., 2017). Of note, time-lapse imaging of radial migration in reeler mice has revealed defects in the radial trajectory of migrating neurons during initial stages of migration (Britto et al., 2010), providing evidence that Reelin signaling impacts the radial migration of cortical neurons at multiple points in addition to terminal stages of migration in the upper cortical layers (Hirota and Nakajima, 2017).

15.7.1.2 Semaphorins The Semaphorin family consists of 20 secreted and membrane-bound proteins that share a 500 amino acideconserved “Sema domain” (Hu and Zhu, 2018)). Semaphorins act as chemoattractants and chemorepulsants, and play an important role in axon guidance. Semaphorin 3A (Sema 3A), a diffusible molecule, was previously shown to promote the migration of neurons from the basal forebrain to the tangential migratory stream (Marin et al., 2001). Sema 3A is expressed in a gradient with the highest levels of expression in the upper layers of the developing neocortex (Polleux et al., 2000). Sema 3A binds to the Neuropilin1 receptor, which is expressed by migrating neurons (Chen et al., 2008). Semaphorins have also been identified as important regulators of radial migration in the embryonic cortex. Sema 3A acts as a chemoattractant for apical dendrites of differentiating cortical neurons (Polleux et al., 2000) and appears to be necessary for the radial migration of layer II/III neurons (Chen et al., 2008). Conditional knockout or knockdown of Neuropilin1 by in utero electroporation impairs the radial migration of cortical neurons. Furthermore, functional blockade of Neuropilin1 with antibodies causes migrating neurons to lose their radial orientation (Chen et al., 2008). Other semaphorins have also been implicated in regulating radial migration in other brain regions. For example, Sema 6A is thought to initiate the radial migration of cerebellar granule cells through modulation of nuclear translocation (Kerjan et al., 2005). In many cases the semaphorins may act as key downstream effectors of transcriptional regulators. For example, the transcription factor Bcl11a negatively regulates Sema 3C, which is necessary for the radial migration of upper cortical layer neurons (Wiegreffe et al., 2015).

15.7.2 Neurotransmitters 15.7.2.1 GABA GABA is the primary inhibitory neurotransmitter in the adult neocortex and is also present in the neocortical proliferative zones during neurogenic stages of development in rodents and primates (Van Eden et al., 1989; Del Rio et al., 1992; Schwartz and Meinecke, 1992). GABA receptor transcripts are present in the embryonic neocortex (Ma and Barker, 1995), physiologically functional GABAA receptors have been demonstrated in precursor cells and immature neurons (LoTurco et al., 1995; Owens et al., 1996), and migrating neurons express both GABAA and GABAB receptors (LoTurco et al., 1995; Behar et al., 2001). Furthermore, GABA appears to increase the migratory movement of embryonic neurons in culture in a dose-dependent manner. Low, femtomolar concentrations of GABA reportedly exert a chemotaxic effect on neurons, while higher micromolar concentrations of GABA may exert a less specific chemokinetic effect that increases the motility of cortical neurons (Behar et al., 1996). GABA receptor subtypes may exert differential effects on migrating neurons that could reflect distinct actions that help to shape the migratory trajectory of cortical neurons during the different stages of migration. Barker and colleagues have shown that GABA, and/or taurine, working through GABAB receptors acts as a chemoattractant for cortical neurons (Behar et al., 2001). Furthermore, Luhmann and colleagues proposed that GABA acts through the GABAC receptor to promote radial migration and through the GABAA receptor, which is expressed at higher concentrations in the upper CP, to act as a stop signal for migrating neurons (Denter et al., 2010). More recent work supports this result by showing that GABA can promote radial migration of cortical neurons in lower cortical layers while acting as a stop signal in uppermost cortical layer (Luhmann et al., 2015). Furthermore, evidence suggests that GABA has distinct effects through ionotropic and metabotropic receptors. GABAB receptors promote the tangential migration of cortical neurons while ionotropic GABAA receptors are crucial for radial migration of excitatory neurons (Luhmann et al., 2015).

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15.7.2.2 Glutamate Glutamate is the main excitatory neurotransmitter in the adult brain, and work has demonstrated a role for glutamate in radial migration in the prenatal neocortex. Endogenous glutamate may be released from CajaleRetzius cells in the marginal zone (del Rio et al., 1995), postmitotic neurons in the CP (Behar et al., 1999), migratory glutamatergic cells, or perhaps even by radial glial cells (Owens and Kriegstein, 2002). Glutamate receptors are expressed by postmitotic neurons migrating to the CP and in the CP (LoTurco et al., 1991; Behar et al., 1999). Glutamate exerts a number of actions in the developing cortex including stimulation of neurite outgrowth (Pearce et al., 1987), modulation of proliferation (Flint et al., 1995; Haydar et al., 2000), and regulation of migration. Glutamate reportedly serves as a chemoattractant that stimulates the radial migration of cortical neurons from the proliferative zones to the CP (Behar et al., 1999). Similar effects are thought to be mediated by NMDA receptors in cerebellar granule neurons (Komuro and Rakic, 1993; Behar et al., 1999), early electrophysiological evidence obtained from whole cell patch clamp recording indicated that migrating cortical neurons do not express functional NMDA receptors (LoTurco et al., 1991). But subsequent work provided evidence for expression of subunits for both ionotropic and metabotropic glutamate receptors in the prenatal cerebral cortex (LopezBendito et al., 2002; Soria and Valdeolmillos, 2002; Lujan et al., 2005), and functional regulation of neuronal migration through glutamate signaling, particularly via the NMDA receptor (Behar et al., 1999; Luhmann et al., 2015).

15.7.2.3 ATP ATP is released through connexin hemichannels expressed by radial glial cells in the embryonic proliferative zones and modulates proliferation in the embryonic neocortex (Weissman et al., 2004). Intermediate progenitor cells are generated in the VZ (Noctor et al., 2008) and migrate to the SVZ and lower IZ in a manner that is nearly indistinguishable from the first stage of radial migration for cortical neurons (Noctor et al., 2004), and thus may utilize common signaling pathways for regulation of cellular movement, such as doublecortin. Rakic and colleagues have demonstrated that intermediate progenitor cells express the ATP receptor P2Y1, and that ATP is essential for the migration of intermediate progenitor cells to the SVZ (Liu et al., 2008). Interestingly, postmitotic neurons appear to lose responsiveness to ATP (Weissman et al., 2004), thus this effect may be specific in its effects on the migration of proliferative intermediate progenitor cells (Liu et al., 2008). More recent work has shown that the tangential migration of cortical interneurons, but not the radial migration of projection neurons, is dependent on mitochondrial production of ATP (Lin-Hendel et al., 2016).

15.7.3 Adhesion molecules Adhesion molecules mediate the interaction between cells and the extracellular matrix, and between adjacent cells. A significant amount of research has examined the importance of adhesion molecules for radial migration. Adhesion molecules are known to provide physical interaction between cells and the ECM, but also are increasingly being investigated for their roles in signal transduction (Juliano, 2002).

15.7.3.1 Integrins Integrins are glycoproteins that are expressed on the cell surface and mediate cellecell and celleextracellular matrix interactions. Integrin receptors are heterodimers of alpha and beta subunits that dimerize in combinations that form more than 20 different integrin receptor subtypes and usually act as cellular receptors for extracellular matrix proteins including collagen, fibronectin, or laminin (Juliano, 2002; Tharmalingam and Hampson, 2016). The actions and specificity of integrin receptors are dependent on the combination of alpha and beta subunits that make up each dimer. Alpha subunits are thought to determine ligand specificity and therefore the responses of integrin receptors (Anton et al., 1999). Several integrins have been implicated in regulating the radial migration of cortical neurons. For example, incubation of dissociated neuron/glial cell cultures obtained from the cortex with function-blocking antibodies for Alpha3 and/or Alpha-v integrin disrupts neuronal migration (Anton et al., 1999). Integrins are also involved in the migration of cortical interneuronsethe Alpha3/Beta1 integrins have been shown to regulate cortical interneuron migration (Stanco et al., 2009). In relation to radially migrating neurons, Alpha3 mutant mice have been shown to display severe defects in radial migration (Schmid et al., 2004). This effect is thought to decrease the gliophilic interactions of migrating neurons, effectively inhibiting the neuroneRG interaction (Anton et al., 1999). The only known subunit that dimerizes with the Alpha3 subunit is the Beta1 subunit. Selective knockout of the Beta1 subunit produces a profound disruption of the radial glial scaffold, but did not produce a migration defect (Graus-Porta et al., 2001). This suggests, perhaps, that the Alpha3 subunit plays a larger role in regulating neuroneglial interactions (Anton et al., 1999; Graus-Porta et al., 2001). On the other hand, Müller and

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colleagues demonstrated that deletion of Beta1 integrin from radial glial cells produces profound disruption of cortical lamination, but that selective Beta1 integrin deletion from cortical neurons does not produce defects in migration or cortical lamination (Belvindrah et al., 2007). This result could suggest that Alpha3 integrin plays a greater role in regulating neuronal migration than Beta1, or perhaps that additional integrin subunits also play important roles in radial migration and exert compensatory influences over each other (Schmid et al., 2004). Consistent with this idea, recent work has demonstrated that the Alpha5/Beta1 receptor is necessary for radial migration. Knockdown of Alpha5 integrin inhibits radial migration, producing an ectopic placement of cortical neurons in lower layers (Marchetti et al., 2010). In addition, recent studies have investigated integrin signaling by manipulating downstream effectors of integrin signaling. For example, knockout of integrin-linked kinase in the embryonic forebrain produces a lamination defect that resembles lissencephaly (Niewmierzycka et al., 2005). Interestingly, Reelin has been shown to bind to the Alpha3/Beta1 integrin receptor that is expressed by radially migrating neurons, and application of Reelin to dissociated neuron/glial cell cultures restricts migration in a manner comparable to that of Alpha3 integrin antibodies (Dulabon et al., 2000). Furthermore, Anton and colleagues showed that the level of Dab1 protein is significantly decreased in Alpha3 mutant mice (Schmid et al., 2005). Together these data suggest a novel signaling pathway through which Reelin may interact with adhesion molecules to regulate radial migration.

15.7.3.2 Gap junctions Gap junctions are large-diameter channels that are composed of two hemichannels on adjoining cells that form a potential aqueous pore that joins the cytoplasm of adjacent cells (Niculescu and Lohmann, 2014; Uhlen et al., 2015). Gap junction pores permit the exchange of ions and molecules up to approximately 1200 Da in size between coupled cells (Simon and Goodenough, 1998). Gap junction proteins are composed of six connexin (Cx) subunits, and at least 20 connexin genes have been identified in rodents and humans. The mature brain gap junction channels can link together adjacent neurons to produce an electrically coupled cluster of neurons that fires synchronous action potentials. In the developing neocortex, electrical coupling between CP neurons has been proposed to presage the development of synaptic coupling (Yuste et al., 1992). Functions associated with gap junction channels in the embryonic rodent neocortex also include regulation of proliferation (Weissman et al., 2004), cytoskeletal elements (Olk et al., 2009), and neuronal migration (Fushiki et al., 2003; Elias et al., 2007; Matsuuchi and Naus, 2013). Multiple connexins are highly expressed in the cerebral cortex and contribute to coupling between neurons and glial cells (Rash et al., 2001). In the embryonic rodent neocortex Connexin 26 (Cx26) and Cx43 have been implicated in radial migration. Neurons that express shRNAs that target Cx43 or Cx26 proteins are unable to migrate radially to the CP (Elias et al., 2007). Conditional knockout mice lacking normal expression of Cx43 gene also display cortical laminar malformations that are consistent with migration defects (Wiencken-Barger et al., 2007; Cina et al., 2009). The role(s) of Cx26 and Cx43 in neuronal migration are thought to be a function of cell adhesion, not gap junction channel formation, since mutant forms of Cx26/43 that do not form functional channels rescue the shRNA-mediated migration defect. This concept is supported by demonstration of the adhesive functions of connexin hemichannels (Cotrina et al., 2008). Connexins appear to be crucial elements for the migration of multiple cell types in a variety of systems, including tumor cells (Naus and Laird, 2010), and cortical interneurons (Elias et al., 2010; Matsuuchi and Naus, 2013).

15.7.4 Cytoskeletal regulators Migration involves a dynamic interplay between the cytoskeleton and cytoskeletal-regulating elements. Migrating cells utilize the dynamics of actin for extension of filopodia and lamellipodia, formation and support of focal adhesions, and detachment of the migrating cell from its substrate. Microtubules and microtubule-associated proteins are also involved in process stabilization and nuclear translocation during migration (Kawauchi and Hoshino, 2008). The regulation of these processes is critical for the proper radial migration of neocortical neurons.

15.7.4.1 Lis1 Microtubule organization in migrating neurons is regulated by several factors including platelet-activating factor acetylhydrolase isoform 1b regulatory subunit 1 (PAFAH1B1, or LIS1) (Moon and Wynshaw-Boris, 2013). Mutations in the LIS1 gene cause lissencephaly in humans (Reiner et al., 1993), a brain malformation characterized by a smooth cerebral cortex, lack of sulci and gyri, and disrupted cortical layers. Mutant Lis1 KO mice present with delayed neuronal migration in heterozygote KO animals and a more dramatic phenotype in homozygote KOs (Hirotsune et al., 1998). Lis1 knockdown through in utero RNA interference (RNAi) induces an accumulation of multipolar neurons in the SVZ of the embryonic

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cortex (Tsai et al., 2005). Lis1 is a microtubule-associated protein that binds tubulin and other microtubule regulatory proteins (Sapir et al., 1997) including Dynein/Dynactin (Faulkner et al., 2000; Smith et al., 2000), Ndel1/NUDEL (Sasaki et al., 2000), and Nde1/mNudE (Feng et al., 2000). Lis1 is thought to induce its effects by promoting dynein ATPase activity (Mesngon et al., 2006). Genetic knockout or RNAi knockdown of Ndel1, which is thought to facilitate the interaction of Lis1 and dynein, prevents many cells from migrating away from the proliferative zones (Sasaki et al., 2005). Lis1 also interacts with actin, as haploinsufficiency of Lis1 in granule neurons leads to reduced F-actin in ends of processes (Kholmanskikh et al., 2003). Evidence shows cross talk between Lis1 and Reelin signaling that regulates cellular migration. The Alpha 1 and 2 subunits of the Pafah1b complex bind to the VLDLR (Assadi et al., 2003; Zhang et al., 2007), which promotes its interaction with Lis1 (Chai and Frotscher, 2016), making this an important node in the regulation of neuronal migration (Moon and Wynshaw-Boris, 2013).

15.7.4.2 Doublecortin Doublecortin (DCX) is mutated in X-linked lissencephaly and is the major genetic cause of subcortical band heterotopia (Dobyns and Truwit, 1995). DCX is a microtubule-associated protein that interacts with and stabilizes polymerized microtubules (des Portes et al., 1998; Gleeson et al., 1998; Francis et al., 1999; Horesh et al., 1999). DCX is expressed in leading processes of migrating and differentiating neurons (Francis et al., 1999). JNK and Cdk5 (see below) are upstream kinases of DCX (Kawauchi et al., 2003, 2006). Knockdown of JNK disturbs leading process formation and migration (Kawauchi et al., 2003). Furthermore, phosphorylation of DCX by JNK localizes DCX to the distal ends of neuronal processes (Gdalyahu et al., 2004). Lis1 has also been shown to interact with DCX (Caspi et al., 2000), and overexpression of DCX rescues the migration defect present in Lis1 mutants (Tanaka et al., 2004). Thus, both molecules may interact to regulate microtubule dynamics that are necessary for migration. Knockdown of doublecortin through RNAi interrupts normal radial migration (Bai et al., 2003). In knockdown cortices, there are both cell autonomous and noncell autonomous effects, which lead to formation of subcortical band heterotopias caused by the premature cessation of neuronal migration in the SVZ/ IZ (Bai et al., 2003). DCX genetic knockout mice do not exhibit alterations in cortical lamination (Corbo et al., 2002), but this is likely due to compensation by doublecortin-like kinase (DCLK) (Deuel et al., 2006; Koizumi et al., 2006), which shares functional similarities with DCX. RNAi of DCLK produces similar defects as DCX (Koizumi et al., 2006). In addition, double DCX and DCLK knockout mice have severe defects in neuronal migration, leading to disruptions in laminar distribution (Kerjan et al., 2009). DCX has also been demonstrated to interact with F-actin (Tsukada et al., 2005), further demonstrating that the signaling components that regulate individual components of the cytoskeleton have significant overlap. Mutations in another X-linked gene, ARX, have also been linked with a migration defect that can produce severe lissencephaly. ARX is expressed by precursor cells in the basal and dorsal telencephalon and by migrating interneurons (Friocourt and Parnavelas, 2010). ARX is thought to be important for the tangential migration and differentiation of GABAergic interneurons (Kato and Dobyns, 2005), but has also been implicated in the radial migration of neurons in the cerebral cortex (Friocourt et al., 2008). Among important finding regarding the use of short hairpin RNA constructs in studies of neuronal migration was the finding that these constructs can have off-target effects by impacting endogenous miRNAs, indicating the use of other approaches to interfere with RNA transcription as well as rescue experiments employing constructs that produce the target protein but are immune to the RNAi tool being used (Baek et al., 2014).

15.7.4.3 Filamin A (FLNA/FLN1) Filamin A is an actin-binding protein produced by the FLNA gene, which is expressed by migrating neurons in the prenatal neocortex (Fox et al., 1998). FLNA mutations in humans lead to periventricular nodular heterotopia, an X-linked dominant condition characterized by an accumulation of abnormally placed cortical neurons located adjacent to the lateral ventricle (Fox et al., 1998). The abnormal location of cortical neurons at the site of neurogenesis indicates that these cells fail to migrate away from the proliferative zones toward the CP. Filamin A interacting protein (Filip1) regulates FLNA expression by inducing Filamin A degradation (Nagano et al., 2002). Filamin A is thought to cross-link F-actin, which increases the viscosity of F-actin network and enhances cell motility (Stossel et al., 2001). Evidence suggests that Filamin A promotes leading process extension in a manner that is regulated by Filip1. In support of this idea, expression of a dominant negative form of Filamin A disrupts leading process formation and prevents the migration of transfected cortical neurons (Nagano et al., 2004), while RNAi knockdown of Filip1 leads to a reduction in the number of multipolar cells in the SVZ and IZ (Nagano et al., 2004). Rac1, a molecule involved in the formation of filopodia and lamellipodia also binds to Filamin A in nonneural cells (Stossel et al., 2001). A dominant negative form of Rac1 induces aberrant leading process formation, similar to defects seen with manipulation of FLNA expression (Kawauchi et al., 2003). Notably, Rac1 also activates JNK, which is an upstream kinase in the DCX signaling pathway (Kawauchi et al., 2003). In addition, mitogen-activated protein

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kinase kinase-4 (Mekk4) interacts with Filamin A and regulates FLNA expression (Sarkisian et al., 2006). MEKK4 acts upstream of JNK, which provides an interesting link between the FLNA and DCX pathways (Hirai et al., 2006).

15.7.4.4 Cdk5 Cyclin-dependent kinase 5 (Cdk5) is a kinase that phosphorylates cytoskeletal proteins, including microtubule- and actinassociated proteins, and cell motility proteins. Cdk5 mutants have laminar defects that are similar to those present in the neocortex of reeler mice (Chae et al., 1997). Cdk5 phosphorylates focal adhesion kinase (FAK), which regulates organization of microtubules in migrating neurons (Xie et al., 2003). Cdk5 activates p27Kip1, which in turn activates the actinbinding protein cofilin (see Reelin section above). This activation is regulated by RhoA, which must be suppressed in order for the Cdk5-mediated activation of cofilin (Kawauchi et al., 2006). Knockdown of p27Kip1 decreases the concentration of F-actin and induces migration defects that can be rescued by the expression of a dominant negative form of RhoA (Kawauchi et al., 2006). Recent work has demonstrated that inhibition of Cdk5 by roscovitine, or inhibition of Src family kinases by PP2, decreases the ability of neurons to migrate radially (Nishimura et al., 2010). P35 activates Cdk5 (Chae et al., 1997), and p35 knockout produces a cortical layering migration defect that is thought to result from impaired interaction of neurons and RG cells (Gupta et al., 2003). PKCdelta stabilizes p35 and Cdk5 activation by phosphorylating Cdk5 (Zhao et al., 2009). PKCdelta knockdown causes severe impairments in radial migration, which can be rescued by cotransfection with wild-type p35 (Zhao et al., 2009). Nishimura and colleagues also demonstrated that inhibition of PKCdelta activity with rottlerin inhibits radial migration of cortical neurons (Nishimura et al., 2010). Cdk5 is also linked to other molecules that are required for radial migration. For example, Cdk5 phosphorylates DCX (Tanaka et al., 2004) and Ndel1 (Niethammer et al., 2000). In addition, Cdk5 and Reelin signaling have been shown to phosphorylate Dab-1 independently (Keshvara et al., 2002). These findings demonstrate the significant cross talk and interplay between molecular pathways and cytoskeletal elements that regulate the mechanisms of radial migration.

15.7.5 Transcription factors 15.7.5.1 Pax6 Recent work has begun to elucidate the important roles that transcription factors play in radial migration. Pax6 is expressed by radial glial cells in the VZ (Gotz et al., 1998), and radial glial cells that have translocated out of the VZ into the SVZ maintain Pax6 expression (Hansen et al., 2010). This pattern of expression is not specific to primates as it has been reported in multiple vertebrates (Martinez-Cerdeño et al., 2012; Martinez-Cerdeno et al., 2016; Martinez Cerdeno et al., 2017). Pax6 has been implicated in neuronal migration, although the specific role that Pax6 plays is not yet clear. Pax6 knockout (Small eyedSey/sey) embryos exhibit a migration defect for late-born cortical neurons that is nonecell autonomous (Caric et al., 1997). Precursor cells lacking Pax6 expression that were obtained from Pax6 KO embryos and transplanted into the cerebral cortex of wild-type embryos exhibited similar patterns of migration compared to transplanted precursor cells obtained from wild-type littermates, suggesting that Pax6 produces an environment that is permissive for the migration of late-born cortical neurons (Caric et al., 1997). Talamillo et al. (2003) used chimeric analysis to demonstrate that Pax6 is important for the radial migration of cortical cells from the SVZ to the overlying CP (Talamillo et al., 2003). Evidence that Pax6 plays a role in migration has also been derived from cell culture work, in which Pax6 overexpression in HeLa cells induces transfected cells to dissociate from cell clusters and exhibit migratory behavior that resemble those exhibited by migrating neurons (Cartier et al., 2006).

15.7.5.2 Tbr2 The Tbr2/Eomes transcription factor is expressed by intermediate progenitor daughter cells that are produced by Pax6expressing radial glia (Englund et al., 2005). The Tbr2þ cells retain Pax6 expression while mitotic (Martinez-Cerdeño et al., 2012) and migrate to a position in the SVZ that is superficial the VZ. Tbr2þ cells are found in the SVZ of the dorsal but not ventral forebrain (Englund et al., 2005). Tbr2 seems to play a role in proper layer formation of the dorsal neocortex (Sessa et al., 2008). Evidence supporting a role for Tbr2 in guiding migration comes from analysis of GABAergic neurons and epiblast cells. Sessa and colleagues demonstrated that Tbr2 attracts the migration of GABAergic neurons produced in the basal telencephalon into the dorsal cortex by inducing expression of the chemoattractant Cxcl12 (Sessa et al., 2010). Tbr2 is required for the delamination of epiblast cells (Arnold et al., 2008). Tbr2 is thought to downregulate the expression of cell adhesion molecules, thereby freeing migratory cells to move toward their destination (Arnold et al., 2008). Thus, intermediate progenitor cells, which are generated by radial glial cells at the surface of the ventricle, may require Tbr2 expression to detach from the ventricular surface and migrate radially to the overlying SVZ (Mihalas and Hevner, 2017).

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15.7.5.3 Neurogenins Neurogenins 1 and 2 (Ngn1/2) are theorized to enhance the migration of projection neurons produced in the dorsal cortex (Nobrega-Pereira and Marin, 2009). In support of this concept, Ngn1/2 is transiently expressed at the onset of radial migration (Hand et al., 2005), increases the expression of DCX and p35, and decreases expression of RhoA (Ge et al., 2006). Conditional knockout of Ngn2 leads to major impairments in radial migration (Hand et al., 2005). Ngn2 must be phosphorylated, as a mutant form of Ngn2 that cannot be phosphorylated at tyrosine 241 leads to a failure of cortical neurons to initiate radial migration into the CP (Hand et al., 2005). These migration defects can be rescued by decreasing RhoA activity using a dominant negative form of RhoA (Hand et al., 2005). In addition, Ngn2 induces the expression of the GTP-binding protein Rnd2 in newly generated neurons. Rnd2 silencing produces a migration defect similar to that seen in Ngn2 mutants, and expression of Rnd2 rescues the cortical migration defect in Ngn2 mutant animals (Heng et al., 2008).

15.8 Summary Cortical projection neurons are produced in the dorsal proliferative zones and migrate to the CP through distinct stages that can be identified based on changes in morphology and orientation of the immature neurons. Different signaling factors are likely involved in regulating distinct stages of migration and/or the transition from one stage of migration to the next. Some of the signaling factors that regulate radial migration are concentrated in different cortical lamina: For example Reelin is produced by CajaleRetzius cells in the marginal zone of the embryonic neocortex. Evidence suggests that sequential generations of projection neurons (i.e., those born on E16, E17, E18, etc. in the rat) undergo the four stages of migration (Noctor et al., 2004, 2008). Thus it seems likely that newborn projection neurons respond to persistent extrinsic signals present within distinct laminar structures, rather than responding to temporal changes in extrinsic signaling. Furthermore, it is likely that the newborn projection neurons follow a maturational program that induces temporal changes in the neuron’s responsiveness to extrinsic signals originating from different cortical strata. Since newborn projection neurons first migrate to the SVZ, it follows that the neurons might initially be responsive to signals that originate from the SVZ. Alternatively, projection neurons may respond to signals originating from more superficial layers, such as the CP or marginal zone, but encounter a stop signal upon reaching the SVZ. During the second multipolar stage of migration the immature neurons remain in the SVZ for periods of one day or longer (Bayer and Altman, 1991; Noctor et al., 2004, 2008). While the multipolar cells do not migrate appreciable distances in the SVZ, they are nonetheless very active, extending, and retracting multiple processes (Tabata and Nakajima, 2003; Noctor et al., 2004). This suggests the possibility that neurons actively search for signals that guide their trajectory and/or destination during stage 2. The third stage of migration involving retrograde movements into the VZ toward the ventricle, and the final fourth stage of radial migration to the CP would likewise result from responsiveness to extrinsic signals originating from different cortical strata. The factors discussed above may play a role in multiple stages of migration. For example, many neurons appear to maintain a close association with their parent radial glial cell throughout the four stages of migration. Therefore, the adhesion between neurons and radial glial cells provided by molecules such as gap junction channels may be required during each stage of migration. A comparison of cortical migration defects produced by single molecules highlights mechanisms that may guide neurons during specific stages of migration. Unique mechanisms may be required for neurons to transition from one stage to the next and specific subsets of neurons may respond differently to similar extrinsic cues. For example, mutations in the FLNA gene produce ectopic clusters of neurons that are located adjacent to the lateral ventricle, while mutations in the doublecortin gene produce subcortical band heterotopias in females (Walsh and Goffinet, 2000). FLNA mutations permit the outgrowth of axonal processes, but prevent the formation of leading processes. Therefore, FLNA mutations may impact initial migration away from the ventricle (Walsh and Goffinet, 2000), but specifically for the subset of neurons that are generated directly by radial glial cells. Alternatively, since FLNA mutations allow axon outgrowth but impede leading process formation, affected neurons may proceed through stage 3 but fail to transition to stage 4. In this scheme, FLNA/ neurons in the SVZ would extend axonal-like processes and approach the ventricle during stage 3 of migration, but lacking the ability to develop a leading process the neurons would fail to transition to stage 4 and remain near the lateral ventricle. Mutations in the doublecortin gene have also been well characterized (des Portes et al., 1998; Gleeson et al., 1998). Doublecortin is expressed at the distal tips of leading processes and axon growth cones (Friocourt et al., 2003; Schaar et al., 2004). Knockdown of doublecortin in rodents through RNAi (Bai et al., 2003), or double knockout of doublecortin and doublecortin-like (Deuel et al., 2006), produces a subcortical band of neurons that can be visualized at the SVZ/IZ border. In contrast, the subcortical band heterotopia present in female carriers of doublecortin mutations appears to be located

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comparatively further from the ventricle than that in the rodent knockdown model. The apparent difference in subcortical band location in rodents and primates most likely results from the differences in the site of neurogenesis. Many neurons are produced in the SVZ of rodents (Haubensak et al., 2004; Miyata et al., 2004; Noctor et al., 2004; Sessa et al., 2008), and in the outer SVZ of primates (Hansen et al., 2010; Martinez-Cerdeño et al., 2012). The subcortical band present at the SVZ/IZ border in rodent models likely represents the cohort of neurons that were generated in the SVZ and failed to migrate any further. These neurons presumably did not progress to the third stage of migration, which is marked by the extension of an axonal process and retrograde movement toward the ventricle, as a result of DCX-linked defects in axonal and neuritic process outgrowth. Cortical neurons generated in the outer SVZ of humans with mutant doublecortin presumably exhibit a similar failure to migrate. However, it should be noted that the four stages of migration described in the rat model of neocortical development have not yet been explored in primate. However, recent work in primates has made great strides in that direction. New reports show that primates possess a greatly expanded SVZ that includes proliferative oRG cells. Together, new data showing substantial neurogenesis in the SVZ of rodents and neurogenic oRG cells in the primate outer SVZ suggest that a modification of the radial unit hypothesis may be warranted. As noted above, a large abventricular neurogenic zone alters the concept of the radial unit hypothesis, as it interposes a large nonventricular source of ontogenetic units between the VZ and the cortex, and could obscure the point-to-point mapping of the ventricular epithelial pattern onto the cortical mantle. Future research should elucidate the trajectory of neurons that are produced in the primate outer SVZ to determine the extent of radial versus tangential deployment of these cells. Furthermore, while the stages of migration identified in rodent may be helpful for elucidating factors that guide radial migration in the developing neocortex, the degree to which radial migration in rodents mirrors that in primates remains to be determined.

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Chapter 16

Mechanisms of tangential migration of interneurons in the developing forebrain Fanny Lepiemme, Carla Silva G.* and Laurent Nguyen* GIGA-Stem Cells / GIGA-Neurosciences, University of Liège, Liège, Belgium

Chapter outline 16.1. Birth of distinct interneuron subtypes and onset of their migration from the subpallium 345 16.2. Molecular cues drawing the path of cortical interneuron migration 347 16.3. Molecular cues controlling the integration of interneurons into the cortical migratory streams 348 16.4. Molecular cues controlling the intracortical dispersion of interneurons 349 16.5. Signals dictating the arrest of interneuron migration within the cortical wall 350 16.6. Role of subpallial transcription factors in the tangential migration of interneurons into the cortex 350

16.7. Cell-intrinsic regulation of cortical interneuron migration 16.8. Dynamic remodeling of the cytoskeleton during interneuron migration 16.9. Regulation of the tangential migration of interneurons in the rostral migratory stream to the olfactory bulb 16.10. Molecular regulation of the migration of striatal interneurons 16.11. Evolutionary perspective of the tangential migration 16.12. Conclusions and perspectives List of acronyms and abbreviations References

351 352 352 353 354 354 355 356

16.1 Birth of distinct interneuron subtypes and onset of their migration from the subpallium Cortical (CT) interneurons have been extensively studied due to their fundamental role in fine-tuning the excitability of cortical circuits (Tremblay et al., 2016). These cells were initially organized in cardinal classes based on their selective expression of molecular markers such as parvalbumin (PV), somatostatin (SST), vasoactive intestinal peptide (VIP), and reelin (RLN) (Wamsley and Fishell, 2017). These broad interneuron classes were further subdivided to accommodate cells with particular synaptic morphology, activity, and function (Wamsley and Fishell, 2017). We currently know that this diversity builds up from its development from the ventral telencephalon until the recruitment and integration into neuronal networks (Wamsley and Fishell, 2017). The majority of CT interneurons are generated in the ganglionic eminences (GEs), whereas a smaller fraction arises in the preoptic area (POA) (Anderson et al., 1997; Gelman et al., 2011; Nery et al., 2002). Pluripotent progenitors located in the medial and caudal domains of the GEs or in the POA sequentially generate distinct interneuron subtypes (Anderson et al., 2001; Gelman et al., 2011). Local morphogens and specific transcription factors (TFs) determine the interneuron output generated by these progenitors. For example, the medial ganglionic eminence (MGE) generates the majority of PV and SST interneurons under the control of Sonic Hedgehog (SSH), NK homeobox 2.1 (NKX2.1), and LIM homeobox 6 (LHX6) TFs (Butt et al., 2008; Liodis et al., 2007; Sussel et al., 1999; Zhao et al., 2008). It has also been demonstrated that progenitors located in different dorsal to ventral regions of the MGE have the potential to generate distinct interneuron subtypes, influenced by differential

* Co-last author

Cellular Migration and Formation of Axons and Dendrites. https://doi.org/10.1016/B978-0-12-814407-7.00016-X Copyright © 2020 Elsevier Inc. All rights reserved.

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morphogen gradients and expression of downstream effectors (Flames et al., 2007; Flandin et al., 2010; Fogarty et al., 2007; Wonders et al., 2008; Xu et al., 2010, 2005). A remarkable effort has also been made to establish the TF network involved in MGE-derived interneuron specification. For example, LHX8, sex-determining region Y-box 6 (SOX6), distal-less homeobox (DLX) 5/6, and ARX (Aristaless-related homeobox) were proposed as downstream effectors or NKX2.1 adjuvants (Sussel et al., 1999; Zhao et al., 2008). Distinctive TFs are at play in the generation and specification of CGE (caudal ganglionic eminence)-derived interneurons. Examples of such TFs are COUPTF1 and COUPTF2, as well as GSH1 and GSH2 (Kanatani et al., 2008; Willi-Monnerat et al., 2008; Xu et al., 2010). The POA generates a smaller fraction of CT interneurons from NKX2.1-expressing progenitors, independently on LHX6 TF (Flames et al., 2007; Gelman et al., 2011). A more specific marker of this structure is NKX5.1 (Gelman et al., 2011). Interestingly, the POA has the ability to generate interneuron subtypes that resemble the CGE-derived interneurons, as suggested by the expression of markers such as RLN or neuropeptide Y (NPY) (Butt et al., 2005; Gelman et al., 2011). The lateral ganglionic eminence (LGE) contributes to the formation of the striatum as well as the generation of olfactory bulb (OB) interneurons (Deacon et al., 1994; Flames et al., 2007; Olsson et al., 1995; Stenman et al., 2003). The molecular signature of LGE also includes TFs mostly enriched in the pallium, such as paired box 6 (PAX6), neurogenin-2 (NGN2), and developing brain homeobox 1 (DBX1), but the development of more ventral portions of this structure requires the action of Gsh genes (Flames et al., 2007). The generation of OB interneurons instead requires the action of Dlx5/6 genes (Waclaw et al., 2006). As previously mentioned, the GEs sequentially generate interneurons that populate diverse brain structures. While the majority of MGE-derived interneurons are generated at E13.5, CGE-derived interneurons are generated later, peaking at E15.5 (Butt et al., 2005; Nery et al., 2002). The initial effort in characterizing the potential of subpallial domains to generate diverse interneuron types has now been complemented with single-cell ribonucleic acid (RNA) sequencing analysis to further elucidate how diversity is achieved. Interestingly, it was recently demonstrated that the transcriptional programs are largely conserved in progenitors located in different locations within the GEs, and only a small set of genes is differentially expressed during development (Mayer et al., 2018; Mi et al., 2018). The conserved group of genes would be, for example, necessary for the acquisition of interneuron-broad characteristics, and the discrete nonconserved set of genes would be sufficient to grant the subtype-specific fate acquisition observed postmitotically (Mayer et al., 2018; Mi et al., 2018). After being generated, interneurons start their migration program toward the brain areas they will integrate. In the past years, a great effort in characterizing, at the cellular and molecular levels, how interneurons move in space has been made. We actually know significantly how interneurons are attracted and which paths are followed during migration. Figs. 16.1e16.2.

Tangential migration Radial migration LGE

VZ SVZ Mantle

Pallium MGE Subpallium

POa

FIGURE 16.1 Schematic depicting a coronal section through the telencephalon. Tangentially migrating cortical interneurons (purple) arise from the subpallium and migrate into the pallium, whereas cortical projection neurons (blue) undergo radial migration along glial processes (orange). Also shown are the sources of tangentially migrating cells: lateral ganglionic eminence (LGE, red), medial ganglionic eminence (MGE, green), and preoptic area (POA, yellow). The ventricular zone (VZ) and subventricular zone (SVZ) comprise a substantial proportion of the MGE and LGE.

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347

CGE

LGE MGE

OB

RMS

FIGURE 16.2 Schematic depicting tangential migration routes of olfactory interneurons from the lateral ganglionic eminence (LGE) (red) through the rostral migratory stream (RMS) into the olfactory bulb (OB) and cortical interneurons from the medial ganglionic eminence (MGE; green) and caudal ganglionic eminence (CGE; blue) that populate the entire cerebral cortex.

16.2 Molecular cues drawing the path of cortical interneuron migration To reach the pallium, CT interneurons have to move and interact with various substrates that grant their spatial progression. They undergo saltatory migration, and during the moving phase, they receive and integrate molecular cues that guide their progression to appropriate forebrain locations (Fig. 16.3). It has been demonstrated that the GEs contain motogenic factors for interneurons. Examples of these factors are the hepatocyte growth factor (HGF), the urokinase-type plasminogen activator receptor (u-PAR), brain-derived neurotrophic factor (BDNF), and neurotrophin-4 (NT-4) acting on tropomyosin receptor kinase B (TrkB) and activating phosphoinositide 3 (PI3) kinase pathway and the glial-derived neurotrophin (GDNF) (Fiumelli et al., 2000; Polleux et al., 2002; Powell et al., 2003, 2001). However, the exact time window, source, and specificity of action of these molecules remain unclear. Interestingly, interneurons are not equally responsive to the same motogens. For example, HGF selectively stimulates the movement of subtypes of interneurons that will populate the anterior cingulate and parietal regions (Powell et al., 2003). Also, GDNF seems to influence primarily PV interneurons (Canty et al., 2009). Moreover, proteins that guide interneuron migration can also function as motogens. One example is EPHA4 transducing reverse signaling involving ephrin-A2 (Steinecke et al., 2014b). Interneurons are repelled from their birthplaces to initiate migration. These cells express the receptor roundabout (ROBO1) that transduces Slit-mediated repulsion to promote their dispersion away from ventricular zone (VZ) and subventricular zone (SVZ) of the GEs and POA (Zhu et al., 1999). Ephrin-A3 and ephrin-A5 are highly expressed in the VZ of the GEs and transduce their effects via EPHA4 (Rudolph et al., 2010; Zimmer et al., 2008). Ephrin-A5/EPHA4 signaling is particularly important to allow CT interneurons scattering into the SVZ and to prevent ventral migration and invasion of the anterior entopeduncular area (AEP). According to their molecular identity, interneurons follow specific migratory pathways inside the subpallium (Zimmer et al., 2011). MGE-derived interneurons undertake an inner path of migration, and POA-derived interneurons by opposition migrate superficially. These two routes of migration circumvent the striatum that is a nontarget area for MGE- or POA-generated interneurons. Striatal avoidance by CT interneurons highly depends on expression of class III semaphorins in the striatal primordium. SemaA and SemaF were identified as the main mediators of the organized interneuron exit toward the cerebral cortex (Tamamaki et al., 2003). Sema3A prevents neuropilin-1 (NRP-1)-expressing interneurons (MGE-derived) entering the cortical plate (CP), directing them toward the dorsal cortex, and Sema3F forced NRP2-expressing interneurons (POA-derived) to navigate into the lower intermediate zone (IZ) (Marín et al., 2001; Tamamaki et al., 2003). Ephrin-A3 expressed in the striatal boundary also contributes to direct MGE-derived neurons in a deeper migratory stream toward the cortex (Rudolph et al., 2010; Zimmer et al., 2008). Striatal cells, expressing EPHB1, are also involved in repelling POA-derived interneurons by reverse signaling via ephrin-B3 (Rudolph et al., 2014). In this type of interaction, repulsion would result from an increase in phosphor-Src and phosphor-FAK (focal adhesion kinase) in CT interneurons (Rudolph et al., 2014). Neuregulin-1 (NRG1) is also crucial to channel ErbB4-expressing interneurons through the LGE toward the palliale subpallial boundary (PSB) (Flames et al., 2004). According to its location and isoform, NRG1 can act as a short- or long-

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MZ IZ/SVZ CP

LGE

Sema3A Sema3F Ephrin-A3 Slit3

MGE

Path of migrating interneurons

Striatum anlage

Ephrin-A5 Slit1

HGF/SF GDNF

Netrin-1 NT4 TAG-1 + fibers

SDF-1 Neuregulin-1

FIGURE 16.3 Schematic of a coronal section through the forebrain depicting the two primary pathways of tangentially migrating interneurons (blue dotted line) through the marginal zone (MZ) and intermediate zone/subventricular zone (IZ/SVZ). Interneurons exit the streams and enter the cortical plate (CP), with many interneurons diving toward ventricular zone (VZ) before ascending into the CP. Also shown are the approximate location of guidance cues and growth factors that guide migrating interneurons along their routes. The membrane-bound neuregulin-1 (þ) is restricted to the lateral ganglionic eminence (LGE) and striatum, whereas the diffusible form of neuregulin-1 is expressed along the SVZ/IZ route. Reproduced from Flames, N., Long, J.E., Garratt, A.N. et al. Short- and long-range attraction of cortical GABAergic interneurons by neuregulin-1. Neuron 44, 251e261.

range molecular cue (Flames et al., 2004). The short-range attraction guiding CT interneuron migration to the PSB is mediated by the isoform NRG1-CRD, a membrane-bound form containing the extracellular cysteine-rich domain (Buonanno and Fischbach, 2001; Falls, 2003). CGE-derived interneurons take different migratory routes during mouse development. The caudal migratory stream (CMS) was the first described to bring interneurons to the caudal cortex and hippocampus (Yozu et al., 2005). More recently, Touzot and collaborators revealed the existence of two additional migratory paths for CGE-derived interneurons, a medial path crossing the MGE (medial migratory stream or MMS) and a lateral path crossing the PSB (LMS) (Touzot et al., 2016). Neurons migrating using the LMS probably also integrate the cortex (Touzot et al., 2016). Interneurons migrating via the MMS probably integrate the amygdala region (Touzot et al., 2016). The POA-derived Dbx1-expressing cells generate CT interneurons and amygdala projection neurons (Hirata et al., 2009). Dbx1-expressing interneurons migrate toward the caudal part of the telencephalon using a preoptic amygdala stream (PAS) (Hirata et al., 2009). Part of the population that downregulates Nrp2 expression is able to cross the Sema3F-expressing striatum and migrate laterally to the cortex (Kanatani et al., 2008; Marín et al., 2001). SST-expressing interneurons from the AEP also use the PASeCMS to reach the PSB (Touzot et al., 2016). After exiting the GEs, interneurons undergo extensive molecular regulation to choose the path of migration in the pallium.

16.3 Molecular cues controlling the integration of interneurons into the cortical migratory streams Once interneurons cross the PSB, they move in the pallium via specific routes named migratory streams (Marín, 2013). At early stages (around E13 in the mouse), CT interneurons progress in two main migratory streams, one localized in the marginal zone (MZ) and another in the upper part of the SVZ that overlaps with the lower IZ (Marín, 2013). A small fraction of interneurons can also be found in the subplate (Lavdas et al., 1999; Wichterle et al., 2001). Although CT interneurons settle in the CP, they are excluded from this region at the onset of corticogenesis. Tangential migration is partly dependent on the interaction with RG cells, and some tangentially migrating cells closely associate with corticofugal axons (Denaxa et al., 2001; Yokota et al., 2007a). However, the accumulation of CT interneurons in axon-sparse regions such as the lower IZ suggests that axons are not the favorite substrate of migration of these neurons (Denaxa et al., 2001).

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Furthermore, it was demonstrated that interneurons reach the cortical wall in appropriate numbers in the absence of the thalamocortical tract (TCA) (Zechel et al., 2016). Molecular cues, localized along pallial-specific regions, are critical to steer and maintain interneurons in the migration streams. For example, chemokine 12 (CXCL12) acting on CXCR4 and/or CXCR7 was identified as an important attractant cue for CT interneurons (Stumm et al., 2007; Tiveron et al., 2006). During corticogenesis, CXCL12 is expressed by the meninges above the MZ and in regions at proximity to the IZ (Daniel et al., 2005; Stumm et al., 2003). Long-range attraction from the secreted NRG1-Ig isoform (containing an extracellular immunoglobulin-like domain) also guides ErbB4-expressing interneurons in pallial regions below the CP (Buonanno and Fischbach, 2001; Falls, 2003; Flames et al., 2004). Tangential migration of CT interneurons seems also to be influenced by local electrical activity, as CT interneurons generate calcium oscillations in response to agonists of glutamate, glycine, and gamma-aminobutyric acid (GABA) receptors (Avila et al., 2013; Métin et al., 2000; Poluch et al., 2001; Poluch and König, 2002; Soria and Valdeolmillos, 2002). It is still largely unknown if different migration streams randomly host different interneuron subtypes or, on the contrary, if there is specificity in the choice of the migration path. A recent study, however, describes that migration and axonal targeting are coupled events to optimize interneuron function. For example, Martinotti cells select the marginal stream as their route of migration, and this choice would facilitate the axonal targeting to layer I (Lim et al., 2018). Interneuron subpallial origin does not seem to be determinant to the choice of the migratory stream, since interneurons born from the MGE or CGE might tangentially migrate in MZ or IZ/SVZ (Miyoshi and Fishell, 2011a). Interestingly, the gene expression profile of interneurons migrating through the MZ was shown to be different from those migrating through the IZ/SVZ, notably for some cell surface receptors (Antypa et al., 2011). Also, the deletion of both netrin-1, a diffusible molecule expressed along interneuron migratory routes, and integrin a3b1, its specific receptor expressed in interneurons, leads to migration defects in the MZ but not in the SVZ (Stanco et al., 2009). In the mouse, lack of expression of glycine receptor a2 subunit (Glya2R) leads to a specific reduction of interneuron migration in SVZ migratory stream (Avila et al., 2013). GABAergic signaling also contributes to the choice of the migratory streams, since specific blockade of GABAB receptors results in accumulation of tangentially migrating interneurons in the VZ/SVZ of the neocortex, decreasing their numbers in the MZ and lower part of IZ (López-Bendito et al., 2003). The next phase of CT interneuron migration involves intracortical dispersion, an event equally ruled by tight molecular regulation.

16.4 Molecular cues controlling the intracortical dispersion of interneurons CT interneurons invade the CP by switching from tangential to radial/oblique migration. Some molecular mechanisms underlying this modification have been proposed. One involves a modification of responsiveness to selective cues such as CXCL12 by the time they exit the migratory streams to invade the CP (Li et al., 2008; Lopez-Bendito et al., 2008). Although CXCL12 and its receptor CXCR4 maintain their expression throughout embryogenesis, the invasion of CP does not require downregulation of CXCR4eCXCL12 signaling (López-Bendito et al., 2008; Stumm et al., 2007). However, since ephrin-B-mediated signaling can block CXCR4 function in a different tissue-specific context (Lu et al., 2001), it is possible that a similar mechanism is at play in the cerebral cortex where the level of ephrin-B is high (Villar-Cerviño et al., 2015). Another strong candidate for CP invasion is neuregulin-3 (NRG3) since CP invasion by interneurons and their intracortical organization requires NRG3 (Bartolini et al., 2017). This molecule is a short-range chemoattractant abundantly expressed in the cortex, and its diffusion is limited by posttranslational modification (Zhang et al., 1997). Interestingly, NRG3 signaling is mediated by ERBB4 receptor (Bartolini et al., 2017). Premature CP invasion by interneurons is also observed in mouse models lacking microglia (Squarzoni et al., 2014). Whether microglia control CP invasion via chemokine/neuregulin signaling and function or by alternative mechanisms remains to be elucidated. TCAs also seem to influence the radial dispersion of CT interneurons not by serving as a migration substrate but instead by releasing glutamate that by its turn regulates potassium chloride cotransporter 2 (KCC2) expression (Zechel et al., 2016). According to the proposed model, glutamate would restrict KCC2 levels in the plasma membrane via calpain action (Zechel et al., 2016). Interestingly, calpain activity is involved in growth cone remodeling and branching, two fundamental cellular events in the shift from tangential to radial migration (Baudoin et al., 2012; Daniel E. Lysko et al., 2014). The meninges are a primordial source of CXCL12 that can signal on CXCR4-expressing TCAs, promoting their growth and projection through the pallium (Abe et al., 2015). Intermediate progenitors (IPs) are an additional source of cues promoting TCA growth (Abe et al., 2015; Sessa et al., 2010; Tiveron et al., 2006). Neurotransmitters/modulators might also be important for the cortical dispersion of CGE-derived interneurons (Murthy et al., 2014). Finally, the mother centriole of migrating CT interneurons assembles a short primary cilium that senses SHH signals arising from the cortical migratory paths (Baudoin et al., 2012; Komada et al., 2008). This ciliar-mediated signaling controls MGE-derived cell in a context-dependent manner and facilitates their reorientation toward CP during migration (Baudoin et al., 2012). The mechanisms by which SHH would influence the shift tangential-to-radial migration are unknown.

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The appropriate positioning of interneurons in the neocortex during their tangential-to-radial switch may also depend on dynamic interactions with RG cells (Yokota et al., 2007b). Consistent with this idea, the migration of interneurons in the CP was altered in ferrets suffering from cortical dysplasia with RG scaffold disruption (Poluch and Juliano, 2007). Moreover, interactions between migrating interneurons and radial glia could be mediated by connexin 43, as shown by its knockdown and by the observation of gap junctions between interneurons and radial processes by electron microscopy (Elias et al., 2010). Interneurons distribute in the cortex occupying a specific laminar distribution. This phase takes place during the first postnatal days in rodents (Miyoshi and Fishell, 2011b). It has been shown that MGE-derived interneurons distribute in an inside-out manner, similar to pyramidal neurons, early-born interneurons populating deep layers, and late-born interneurons targeting superficial layers (Miyoshi et al., 2007). In contrast to MGE-derived interneurons that progressively invade the CP depending on their birthdate, CGE-derived interneurons mostly end their migration in superficial layers irrespective of their birthdate, suggesting that factors other than intrinsic programs contribute to its final distribution (Miyoshi et al., 2010). Indeed, serotoninergic signaling contributes to control superficial layers occupancy of CGE-derived interneurons. The serotonin receptor (5-HT3AR) is upregulated in CGE-derived interneurons during the period of cortical invasion, and the main sources of serotonin in the developing cortex are the raphe fibers spreading in the MZ and IZ (Wallace and Lauder, 1983). 5-HT3AR activation contributes to the generation of early patterns of activity that are fundamental for shaping CGE-derived interneuron networks (De Marco García et al., 2011). Elegant studies further demonstrated that the molecular cues generated by pyramidal cells also shape interneuron distribution in cortical layers (Hevner et al., 2004; Lodato et al., 2011; Pla et al., 2006). For example, Lodato and colleagues showed that the exclusive lack of subcerebral projection neurons in a mutant mice and their replacement by callosal projection neurons impaired the laminar positioning of particular interneuron subtypes (Lodato et al., 2011).

16.5 Signals dictating the arrest of interneuron migration within the cortical wall By the second postnatal week in the mouse, interneurons might have reached the appropriate layers, and they will consequently stop their movements (Inamura et al., 2012). The progressive decrease in interneuron motility and the later settling at a particular place are instructed by extrinsic signals. Calcium waves seem to play an important role in the control of interneuron motility (Kumada and Komuro, 2004; Moya and Valdeolmillos, 2004). Calcium fluctuations are indeed maintained in CT interneurons by GABA depolarizing membrane currents triggered by GABAA receptors (Inada et al., 2011). During the first postnatal days, the inversion of the intracellular chloride gradient arising from the increased expression of KCC2 generates GABAAR-mediated membrane hyperpolarizing currents (Ben-Ari, 2002; Inamura et al., 2012), leading to a decrease in motility and arrest of interneurons in the developing cerebral cortex (Bortone and Polleux, 2009). Interestingly, some studies consisting in the coculture of interneurons with more mature cortical postnatal cells have shown that factors released by cortical cells could also induce arrest of interneuron migration (Inamura et al., 2012). The nature of these signals and the details of the signaling remain unknown.

16.6 Role of subpallial transcription factors in the tangential migration of interneurons into the cortex Beyond their fundamental role in cell specification, TFs also influence interneuron migration. Among them, LHX6 is highly expressed in SST- and PV-expressing interneurons (Liodis et al., 2007), and silencing experiments revealed that it regulates the expression of guidance cues (Alifragis et al., 2004, p. 6). LHX6 also controls the distribution of interneurons in the deep cortical layers regulating the expression of CXCR4 and CXCR7 (Vogt et al., 2014; Zhao et al., 2008). Recent evidence suggests that LHX6 might not act alone to control CT interneuron migration, as this TF can form complexes with other LIM domainecontaining proteins (Vogt et al., 2014; Zhao et al., 2008). Additional TFs control the tangential migration of cortical interneurons. Knockout mouse models for Dlx1/2 and Arx showed major deficit of interneuron migration toward the cortex (Anderson et al., 1997; Colombo et al., 2007). Interestingly, the migration defects observed in Dlx1/2 mutants could be rescued by decreasing GSH1 level or by expressing ARX (Colasante et al., 2008; Wang et al., 2013). A cell-autonomous role for ARX was indeed reported for the migration of ventrally generated interneurons (Friocourt et al., 2008). At the cellular level, LHX6 and DLX1/2 control interneuron migration in multiple ways, notably, by regulating neuritogenesis and the expression of chemokine and neuregulin

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receptors/ligands (Cobos et al., 2007; Zhao et al., 2008). Interneuron migration defects resulting from DLX1/2 knockdown largely result from impaired cytoskeleton dynamic remodeling and involve p21-activated kinase 3 (PAK3), an effector of RhoGTPase and cell division control protein 42 (CDC42) (both modulators of actin polymerization) and microtubuleassociated protein 2 (MAP2) (Cobos et al., 2007; Colasante et al., 2008). NKX2-1 acts upstream LHX6 (Du et al., 2008) and contributes to the migration and recruitment of interneurons to the striatum by repressing Nrp2, thereby making interneurons insensitive to semaphorin-mediated repulsion signaling from the striatum (Nóbrega-Pereira et al., 2008). This mechanism seems particularly important for interneurons born after E13.5 in the MGE and AEP that take deep migration routes toward the cortex (Marín et al., 2001). SP9 acts upstream NKX2-1, LHX6, and ARX as a TF able to indirectly regulate interneuron migration (Liu et al., 2018). Moreover, direct effects of SP9 might involve the regulation of proteins with relevant roles in neuronal migration such as atypical chemokine receptor (ACKR3), EPHA3, and suppression of tumorigenicity 18 (St18) (Liu et al., 2018). Another example of hierarchical TF relationship influencing the migration of ventrally generated cells by direct and indirect mechanisms involves forkhead box protein G1 (FOXG1) (Yang et al., 2017). This TF controls the expression of Dlx1/2 and sustains the expression of Robo1 and EphA4 (Yang et al., 2017). Guidance receptor expression from the EPH/ephrin family in POA-derived interneurons is instead controlled by LHX1 (Symmank et al., 2018). The netrin receptor UNC5b is also indirectly regulated by Smad-interacting protein 1 (SIP1) in migrating CT interneurons (van den Berghe et al., 2013). Overexpression of Unc5b misguides CT interneurons toward ventral regions. At the mechanistic level, UNC5b is a receptor for netrin1 (NTN1) and leucine-rich repeat transmembrane proteins FLRT2 and FLRT3. High levels of UNC5b mediate repulsion of interneurons from NTN1-negative (VZ of LGE, MGE and striatal anlage) and FLRT2/3-positive (LGE) regions, leading to interneuron accumulation in ventral forebrain areas (van den Berghe et al., 2013). COUPTF2 is a TF driving the migration of CGE-derived interneurons, and its upregulation in MGE-derived interneurons is able to divert them toward caudal locations (Kanatani et al., 2008). These observations strongly suggest that fate acquisition and molecular regulation of interneuron migration might be linked and coordinated events.

16.7 Cell-intrinsic regulation of cortical interneuron migration Experiments consisting in culturing CT interneurons derived from the same proliferative structure at different developmental stages and performing comparison of their differential ability to move in a similar environment revealed that cell-intrinsic mechanism might also be at play and influence interneurons distribution in the target tissue (Inamura et al., 2012). Mechanisms underlying cell-intrinsic regulation remain largely unknown, but several reports have indicated that modifying the expression level of the chloride transporter KCC2 during the maturation of neuroblasts has an impact on membrane potential and consequently on calcium fluctuations that participate in the regulation of neuron migration (Bortone and Polleux, 2009; Kumada and Komuro, 2004). The cell-intrinsic regulation component of interneuron migration might also have a fundamental role in shaping the formation of neuronal networks. This is exemplified by the recent discovery that self-regulation of the saltatory migration of CT interneurons shapes cortical morphogenesis by regulating dorsal neurogenesis (Silva et al., 2018). The posttranslational processing of the C-terminal tail of myosin light-chain kinase (MLCK) by the cytosolic carboxypeptidase 1 (CCP1) is a crucial cellular event involved in the regulation of pausing duration during migration cycle of CT interneurons (Bellion et al., 2005). The saltatory migration of CT interneurons is not synchronized among interneurons, and loss of CCP1 reduces pause duration, which has a direct impact at the population level. This results in recruitment of a larger fraction of interneurons invading the cortex. The temporal accumulation of supernumerary interneurons in the SVZ/IZ migratory stream of the cortical wall leads to increased proliferation of dorsal IPs through a cross-talk involving diffusible cues (Silva et al., 2018). These developmental defects lead to the expansion of cortical upper layers that may trigger aberrant cortical circuit formation supporting specific behavior. Interneuron primary cilium was also proposed as a structure that binds and transduces signals from guidance cues in a cell-autonomous fashion (Higginbotham et al., 2012). Interestingly, this structure is highly dynamic, being able to change in length, rotate, or branch (Higginbotham et al., 2012). Ciliar integration of molecular cues would grant cellular changes in morphology and movement (Higginbotham et al., 2012). The absence of the cilium-localized ADP ribosylation factorelike 13B (ARL13B) protein was shown to reduce the dynamics of ciliar movement during the migration cycle, making interneurons move less efficiently and extend multiple processes in multiple directions, probably due to an inefficiency in receiving extracellular signals (Higginbotham et al., 2012). It is still unclear if the intracellular signaling cascades triggered in the primary cilium and in the growth cone to direct interneuron movement are similar.

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16.8 Dynamic remodeling of the cytoskeleton during interneuron migration While migrating, interneurons undergo nuclear saltatory movements and continuously extend and retract neurites from the leading process. The tip of leading process is a growth coneelike structure able to sense environmental cues (Bellion et al., 2005). These cells also possess a membrane protrusion at the rear of the cell called trailing process that allows a change in the direction of migration (Bellion et al., 2005). Interneuron migration is composed of two phases. First, the leading process extends some branches, and as one of the branches stabilizes, a large swelling that contains the centrosome and the Golgi apparatus moves away from the nucleus toward the leading process. The second phase, called nucleokinesis, consists in a very fast nuclear translocation toward the swelling, with retraction of the trailing process (Bellion et al., 2005). All these events require microtubule and actomyosin cytoskeleton remodeling. Migrating neurons have a large perinuclear network of microtubules connecting the nucleus to the centrosome (Rivas and Hatten, 1995; Tanaka et al., 2004). By using a mutant mouse model of lissencephaly-1 (Lis-1), a protein regulating microtubule function, it was shown that the migration rate of interneurons was decreased but not completely abolished, suggesting that other cytoskeletal forces greatly contribute to interneuron migration (McManus et al., 2004; Tanaka et al., 2004). It was demonstrated that actomyosingenerated forces drive nucleokinesis, since this nuclear event is blocked upon blebbistatin treatment, a specific inhibitor of nonmuscle myosin-II (Bellion, 2005). Forces generated by actomyosin contraction take place behind the nucleus where myosin-II and F-actin accumulate (Bellion, 2005; Martini and Valdeolmillos, 2010). Spontaneous and fast intracellular calcium transients seem to be responsible for this actomyosin remodeling (Martini and Valdeolmillos, 2010). Cofilin, for example, is critical for the proper formation of actomyosin filaments that provide forces during motility (Tielens et al., 2016). The choice of the directionality is made by the growth coneelike structures present at the tip of the neurites and by polarity reversion at the trailing process (Kakita and Goldman, 1999; Nadarajah et al., 2002). F-actin is also concentrated in the growth cone and in some small protrusions found along the leading process. These actin protrusions are the first step for leading process branching. These structures will be invaded by microtubules, allowing the emergence of the branch and its stabilization (Lysko et al., 2014). NRG1 and CXCL12 have distinct effects on leading process branching. NRG1 promotes branch extension close to the source of attraction, and CXCL12 reduces branching frequency (Flames et al., 2004; LópezBendito et al., 2008; Lysko et al., 2014). The molecular cascade involved in CXCL12-mediated control over branching involves calpain cleavage of cortactin, favoring actin polymerization and allowing doublecortin (DCX) to bundle and stabilizing microtubules (Lysko et al., 2014). Growth cone remodeling is also modulated by microtubule and actomyosin cytoskeleton (Métin et al., 2006). Interestingly, the fine molecular regulation at the leading process tip involves disrupted in schizophrenia 1 (Disc1), a susceptibility gene for many psychiatric disorders. In the absence of DISC1, Girdin, an actininteracting protein, is not properly targeted to the growth cone, leading to the formation of less stabilized actin filaments (Steinecke et al., 2014a).

16.9 Regulation of the tangential migration of interneurons in the rostral migratory stream to the olfactory bulb In the olfactory bulb (OB), interneurons are generated from embryogenesis throughout life from different proliferative regions. Before birth, OB interneurons are generated from the dorsal region of the LGE (dLGE) and, after birth, from the anterior part of SVZ of the lateral ventricle (Altman, 1969; Corbin et al., 2000; Dellovade et al., 1998; Lois and AlvarezBuylla, 1994; Luskin, 1993; Luskin and Boone, 1994; Wichterle et al., 2001). OB interneurons include not only GABAergic neurons but also dopaminergic tyrosine hydroxylase (TH) cells and glutamatergic juxtaglomerular interneurons (Baker et al., 1983; Brill et al., 2008; Halász et al., 1982; McLean and Shipley, 1988). According to the inputs received and the way they control OB circuitry, OB interneurons were further classified into periglomerular or granule cells (Kosaka and Kosaka, 2016; Lledo and Valley, 2016). To reach the OB, interneurons follow a very defined path termed the rostral migratory stream (RMS) in postnatal stages (Doetsch et al., 1997; Doetsch and Alvarez-Buylla, 1996; Lois et al., 1996; Lois and Alvarez-Buylla, 1994; Rousselot et al., 1995). Strikingly, the RMS is not organized during embryogenesis, and migrating cells were, at these stages, found in large nondefined extracellular spaces (Kishi et al., 1990). Within the RMS, interneurons establish neurophilic interactions, facilitating the interaction between interneurons and the formation of chains of migrating neuroblasts (Wichterle et al., 1997). Interneurons undergo “chain migration” within glial fibrillary acidic protein (GFAP)epositive astrocytes that form glial tube to help and direct interneuron migration toward the OB (Bozoyan et al., 2012; Peretto et al., 2005; Snapyan et al., 2009; Whitman et al., 2009). They display a classical bipolar morphology of interneurons with a trailing and long and branched leading process (O’Rourke, 1996). Chain migration depends on the substrate nature. For example, it was demonstrated that the polysialylatede neural cell adhesion molecule (PSAeNCAM), promoting homo- and heterophilic interactions, is necessary for chain

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migration of interneurons into the OB (Chazal et al., 2000; Cremer et al., 1994; Tomasiewicz et al., 1993). With maturation, PSAeNCAM expression increases in the RMS, correlating to an increased chain migration (Hu, 2000; Murase and Horwitz, 2002; Pencea et al., 2001; Rousselot et al., 1995). Other adhesion molecules have a dynamic expression in the RMS. For example, tenascin-C was found strongly expressed in astrocytes that associate to the RMS, and a- and b-integrins were also found expressed in this route (Jankovski and Sotelo, 1996; Murase and Horwitz, 2002). RMS astrocytes express ephrin-B2 and ephrin-B3, whereas EPHA4, EPHB2, and EPHB3 receptors are expressed by migrating neuroblast within the RMS (Conover et al., 2000). To integrate the RMS, interneurons are guided by various migration cues. For example, during embryogenesis, the septum releases chemorepulsive SLIT1 and SLIT2 that might orient the direction of migration (Itoh et al., 1998; Yuan et al., 1999). These molecules are also able to repel migrating interneurons from SVZ in postnatal stages. The OB itself provides guidance signals that attract migrating neuroblasts. For example, the mitral cells from the OB express NTN1 that is sensed by migrating interneurons via deleted in colorectal cancer (DCC) and neogenin receptors (Murase and Horwitz, 2002). NTN1 is attracting for migrating interneurons only during embryogenesis as its expression decreases with the maturation of the OB30,31. The OB also releases motogenic molecules such as MIA (migration-inducing activity) to stimulate interneuron migration (Mason et al., 2001). RMS astrocytes synthesize and secrete vascular endothelial growth factor (VEGF) necessary for blood vessel formation and growth (Bozoyan et al., 2012). ROBO receptors expressed in astrocytes are also involved in neuroblast-generated SLIT1 repulsion necessary for the formation of glial tunnels necessary for migration (Kaneko et al., 2010). Astrocytes express GABA transporters, whereas migrating neuroblast express GABA. GABA uptake stimulates neuroblast migration, and GABA induces Ca2þ waves in astrocytes, a mechanism involved in trapping BDNF and regulating the frequency of stationary phases (Bolteus and Bordey, 2004; Snapyan et al., 2009). Na-K-Cl cotransporter 1 (NKCC1) was shown to be necessary for neuroblasts to maintain a normal migratory speed by controlling the resting membrane potential independently on GABAA-mediated signaling (Mejia-Gervacio et al., 2011). Astrocytes can also release glutamate to control the survival and migration of neuroblasts expressing N-methyl-D-aspartic acid (NMDA) receptors (Platel et al., 2010). After reaching the OB, interneurons shift from a tangentially oriented to a radial migration and integrate this structure in a way that contributes for its laminar organization. The integration of different subtypes of OB interneurons is sequential. For example, calbindin-positive interneurons, generated during late embryogenesis, are integrated before calretininexpressing interneurons, generated mainly during postnatal periods19. The glycoprotein RLN was shown to be fundamental for the radial migration and the correct organization of OB layers (Hack et al., 2002). OB layering also requires that neuroblasts detach from RMS, and it was demonstrated that tenascin-R and prokineticin-2 are involved in this process (Ng et al., 2005; Saghatelyan et al., 2004). Tangential migration is also adopted by cells that must move for shorter distances. This is the case of striatal (STR) interneurons. We next describe the main observations that improved our understanding of the molecular regulation behind the movement of these cells.

16.10 Molecular regulation of the migration of striatal interneurons STR interneurons migrate tangentially in the presumptive striatum from the MGE or POA/AEP where they are generated from distinct pools of progenitors (Flames et al., 2007; Flandin et al., 2010; Marın et al., 2000). While CT interneurons downregulate Nkx2-1 few hours after being generated, STR interneurons retain the expression of this TF (Nóbrega-Pereira et al., 2008). Transplantation assays demonstrated that the presumptive striatum influences the migration of STR interneurons by releasing attractive cues (Villar-Cerviño et al., 2015). As CT interneurons, STR interneurons express ERBB4 receptor and are attracted to NRG sources from corridor cells surrounding the STR (Villar-Cerviño et al., 2015). ErbB4 expression depends on LHX6 and NKX2-1 transcription factors (Du et al., 2008; Nóbrega-Pereira et al., 2008; Zhao et al., 2008). Different isoforms of ERBB4 have been identified in migrating interneurons and remains unclear if CT interneurons and STR interneurons activate the same intracellular pathway(s) downstream ERBB4 activation to migrate (Rakic et al., 2015). Furthermore, not all STR interneurons express ERBB4, suggesting that STR attraction is also mediated by other molecular cues (Fox and Kornblum, 2005; Yau et al., 2003). Notably, ephrin-BeEPHB-mediated signaling mediates STR interneuron cortical avoidance (Villar-Cerviño et al., 2015). Their arrest in the striatum requires the interaction between ephrin-B3 expressed by STR interneurons with EPHB1. Intracellularly, these interactions result in a decrease in the endogenous levels of phospho-Src and phosphor-FAK (Rudolph et al., 2014). These observations are a demonstration that the same class of molecular cues might organize and direct the migration of distinct cell types, such as CT and STR interneurons to distinct structures depending on binding partners and intracellular signaling cascades.

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16.11 Evolutionary perspective of the tangential migration During the course of evolution, the brain expanded and became a more complex structure (Rakic, 2009). Although not conclusive, several lines of evidence support a multiple origin for cortical interneurons from pallial and subpallial origins in humans (Letinic et al., 2002; Petanjek et al., 2009). Also, a complexification of a transient cellular layer, the subpial granular layer, is involved in the generation of migrating interneurons in the primate (Rakic, 2009; Zecevic et al., 2011). Early during development, primate and human interneurons are exclusively generated from the GEs and are tangentially oriented in the upper SVZ, lower IZ, subplate, and MZ of the dorsal telencephalon (Petanjek et al., 2009). Dorsallygenerated interneurons seem, however, to migrate nontangentially as suggested by their orientation in the dorsal telencephalon (Petanjek et al., 2009). The description of the molecular regulation of human tangentially and nontangentially migrating interneurons remains largely unknown. The requirement of late-generated interneurons from sources other than the GEs might be associated with the emergence of long periods of brain development and the refinement of brain circuitry in primates and humans. For example, the extension of plasticity periods in humans might have been suited by the late migration of CT interneurons from the SVZ and targeting the prefrontal cortex of young children (Paredes et al., 2016). Despite the fact that the nature and origin of interneuron progenitors might differ across species, interneurons generated from subpallial structures conserved a tangential mode of migration already present in more ancestral branches including amphibians and sauropsides (Cobos et al., 2001; Jarvis et al., 2005; Letinic et al., 2002; Moreno et al., 2008; Petanjek et al., 2009; Tanaka and Nakajima, 2012; Tuorto et al., 2003; Veenman and Reiner, 1994). The guidance cues for interneurons were preserved during evolution as demonstrated by interspecies transplantation experiments, showing that mammal interneurons can migrate and integrate the cortical primordium of sauropsides (Métin et al., 2007). With cortical expansion and refinement, however, the complexification of signaling molecules regulating interneuron sorting and network integration specialized, as suggested by transplantation experiments in which nonmammalian interneurons do not successfully integrate the mammalian CP (Tanaka and Nakajima, 2012).

16.12 Conclusions and perspectives Tangential migration has been reported as a strategy used by interneurons to move and integrate distant cortical networks. A significant effort has been made to better characterize interneuron migration, focusing on their morphological remodeling, the guidance cues implicated in the migration, and the mechanisms involved in the formation of the migration paths. Recent technical advancements will soon allow us to more deeply understand if the properties of interneuron migration display specificity regarding the molecular signature, cellular subtype, or evolutionary origin. Clonal analysis of interneuron migration using genetic construct allowing the visualization of cell morphology might be a useful tool to expand to the domain of cell migration (Evrony et al., 2015; García-Moreno et al., 2014; Kornack and Rakic, 1995; Livet et al., 2007; Loulier et al., 2014; Luskin et al., 1988). Indeed, clonal analysis has been used to elucidate if lineage is a good predictor of positioning within the forebrain (Ciceri et al., 2013; Mayer et al., 2016, 2015). Also, genetic toolboxes such as the mosaic analysis with double markers (MADMs) coupled to knockout strategies targeting key proteins for interneuron migration might advance our knowledge on cell-autonomous and nonautonomous mechanisms regulating their progression in the cortical wall (Beattie et al., 2017; Hippenmeyer et al., 2010). Most of our current knowledge on interneuron development arises from rodent models, but the possibility to differentiate interneurons from human-induced pluripotent stem cells (hiPSCs) opens new horizons to shed light on their mechanisms of migration and maturation (Liu et al., 2013; Sun et al., 2016). Moreover, they are useful tools to test the cellular consequences of human mutations identified in patients suffering from interneuropathies (Lancaster et al., 2013). Such approach has recently uncovered a selective migration deficit of cortical interneurons migrating in “assembloids” and derived from patients suffering from Timothy syndrome, a disease associated with autism spectrum disorders and epilepsy (Bagley et al., 2017; Birey et al., 2017; Xiang et al., 2017). The improvement of culture systems such as the recent development of assembloids might allow better understanding the mechanisms of generation, maturation, and differentiation of different subtypes of interneurons, bringing closer the perspective of investigating cell-specific motility properties of human cells. In silico modeling has also been useful to extrapolate the consequences of molecular modifications at the population dynamics and as predictors of the outcome of cell-to-cell interactions (Silva et al., 2018; Villar-Cerviño et al., 2013). The exponential interest that the study of tangential migration received in the past years will undoubtedly help us unraveling the ultimate and more challenging goal of being able to understand, fix, or improve interneuron function in brain malfunction scenarios.

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List of acronyms and abbreviations 5-HT3AR 5-hydroxytriptamine receptor 3A ACKR3 atypical chemokine receptor AEP anterior entopeduncular area ARL13B ADP ribosylation factorelike protein 13B ARX Aristaless-related homeobox BDNF brain-derived neurotrophic factor CCP1 cytosolic carboxypeptidase 1 CDC42 cell division control protein 42 CGE caudal ganglionic eminence CMS caudal migratory stream COUPTF COUP transcription factor CP cortical plate CRD cysteine-rich domain CT cortical DBX developing brain homeobox DCC deleted in colorectal cancer DCX doublecortin DLX distal-less homeobox FAK focal adhesion kinase FLRT fibronectin leucine-rich repeat transmembrane protein FOXG1 forkhead box protein G1 GABA gamma-aminobutyric acid GDNF glial-derived neurotrophin GEs ganglionic eminences GFAP glial fibrillary acidic protein Glya2R glycine receptor a2 subunit GSH GS homeobox GTP guanosine triphosphate HGF hepatocyte growth factor hiPSCs human-induced pluripotent stem cells IP intermediate progenitor IZ intermediate zone KCC2 potassium chloride cotransporter 2 LGE lateral ganglionic eminence LHX LIM homeobox Lis-1 lissencephaly-1 LMS lateral migratory stream MADM mosaic analysis with double markers MAP2 microtubule-associated protein 2 MGE medial ganglionic eminence MIA migration-inducing activity MLCK myosin light-chain kinase MMS medial migratory stream MZ marginal zone NGN neurogenin NKCC1 Na-K-Cl cotransporter NKX NK homeobox NMDA N-methyl-D-aspartic acid NPY neuropeptide Y NRG neuregulin NRP neuropilin NT-4 neurotrophin-4 NTN netrin OB olfactory bulb PAK3 P21-activated kinase 3 PAS preoptic amygdala stream PAX6 paired box 6

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PI3 phosphoinositide 3 POA preoptic area PSAeNCAM polysialylatedeneural cell adhesion molecule PSB pallialesubpallial boundary PV parvalbumin RG radial glial RLN reelin RMS rostral migratory stream RNA ribonucleic acid ROBO roundabout SIP Smad-interacting protein SOX6 sex-determining region Y-box 6 SSH Sonic Hedgehog SST somatostatin St18 suppression of tumorigenicity 18 protein STR striatal SVZ subventricular zone TCA thalamocortical tract TF transcription factor TH tyrosine hydroxylase TrkB tropomyosin receptor kinase B u-PAR urokinase-type plasminogen activator receptor VEGF vascular endothelial growth factor VIP vasoactive intestinal peptide VZ ventricular zone

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The distribution of GABA-containing perikarya, fibers, and terminals in the forebrain and midbrain of pigeons, with particular reference to the basal ganglia and its projection targets. J. Comp. Neurol. 339, 209e250. https://doi.org/10.1002/cne.903390205. Villar-Cerviño, V., Kappeler, C., Nóbrega-Pereira, S., Henkemeyer, M., Rago, L., Nieto, M.A., Marín, O., 2015. Molecular mechanisms controlling the migration of striatal interneurons. J. Neurosci. 35, 8718e8729. https://doi.org/10.1523/JNEUROSCI.4317-14.2015. Villar-Cerviño, V., Molano-Mazón, M., Catchpole, T., Valdeolmillos, M., Henkemeyer, M., Martínez, L.M., Borrell, V., Marín, O., 2013. Contact repulsion controls the dispersion and final distribution of Cajal-Retzius cells. Neuron 77, 457e471. https://doi.org/10.1016/j.neuron.2012.11.023. Vogt, D., Hunt, R.F., Mandal, S., Sandberg, M., Silberberg, S.N., Nagasawa, T., Yang, Z., Baraban, S.C., Rubenstein, J.L.R., 2014. Lhx6 directly regulates Arx and CXCR7 to determine cortical interneuron fate and laminar position. Neuron 82, 350e364. https://doi.org/10.1016/ j.neuron.2014.02.030. Waclaw, R.R., Allen, Z.J., Bell, S.M., Erdélyi, F., Szabó, G., Potter, S.S., Campbell, K., 2006. The zinc finger transcription factor Sp8 regulates the generation and diversity of olfactory bulb interneurons. Neuron 49, 503e516. https://doi.org/10.1016/j.neuron.2006.01.018. Wallace, J.A., Lauder, J.M., 1983. Development of the serotonergic system in the rat embryo: an immunocytochemical study. Brain Res. Bull. 10, 459e479. Wamsley, B., Fishell, G., 2017. Genetic and activity-dependent mechanisms underlying interneuron diversity. Nat. Rev. Neurosci. 18, 299e309. https:// doi.org/10.1038/nrn.2017.30. Wang, B., Long, J.E., Flandin, P., Pla, R., Waclaw, R.R., Campbell, K., Rubenstein, J.L., 2013. Loss of Gsx1 and Gsx2 function rescues distinct phenotypes in Dlx1/2 mutants. J. Comp. Neurol. 521, 1561e1584. https://doi.org/10.1002/cne.23242.

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Chapter 17

Migration in the hippocampus Hirofumi Noguchi, Guangnan Li and Samuel J. Pleasure Department of Neurology, University of California, San Francisco, CA, United States

Chapter outline 17.1. Overview of hippocampal structure and lamination 365 17.1.1. Terminology important for studying hippocampal structure 365 17.2. Developmental specification of hippocampal fields 366 17.2.1. The basic developmental scheme of the hippocampus 366 17.2.2. The cortical hem 366 17.2.3. The cortical hem organizes the hippocampal fields 366 17.2.4. The role of canonical Wnt signaling in hippocampal development 367 17.3. Migration of CajaleRetzius cells in the hippocampus 367 17.3.1. What are CajaleRetzius cells? 367 17.3.2. What are the functions of CajaleRetzius cells? 367 17.3.3. What are the origins of CajaleRetzius cells? 367 17.3.4. The cortical hem is the major source of Cajale Retzius cells for the dorsal telencephalon 368 17.3.5. The extent of the cortex covered by hem-derived CajaleRetzius cells 368 17.3.6. Recruitment of hem-derived CajaleRetzius cells to the meninges 368

17.3.7. Tangential dispersion of CajaleRetzius cells in the marginal zone 17.4. Migration of hippocampal pyramidal neurons 17.5. Migration of hippocampal interneurons 17.5.1. Cellular and distributional diversity of interneurons in the hippocampus 17.5.2. Origins and migration of hippocampal interneurons 17.6. Migration of neural progenitors and granule cells in the dentate gyrus during development 17.6.1. The basic developmental scheme of the dentate gyrus 17.6.2. Migration of granule neurons to form the granule cell layer 17.6.3. Emergence and migration of long-lived neural stem cells and establishment of subgranular zone 17.7. Conclusions References

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17.1 Overview of hippocampal structure and lamination Generating a functional hippocampus requires the assembly of a host of distinct cell types through an ordered developmental program. In this chapter, the current understanding of this developmental program is presented. Data from studies in rodents without particularly dealing with the phylogenetic differences in detail are primarily discussed.

17.1.1 Terminology important for studying hippocampal structure The hippocampal formation includes the dentate gyrus, the hippocampus proper, the subiculum, the presubiculum, the parasubiculum, and the entorhinal cortex. The hippocampus or hippocampal complex includes the hippocampus proper and the dentate gyrus. The hippocampus proper can be further divided into three subregions: CA1, CA2, and CA3 (CA is the abbreviation for cornu ammonis; Fig. 17.1). The principal cell layer in the hippocampus proper is called the stratum pyramidale. Deep to the pyramidal cell layer is the stratum oriens. Superficial to the CA3 is the stratum lucidum, which is occupied by mossy fibers projecting from the dentate granule cells. The stratum radiatum is located superficial to the stratum lucidum in CA3 and the pyramidal cell layer in the CA1 and CA2. The most superficial layer in the hippocampal proper is called the stratum lacunosummoleculare (Fig. 17.1).

Cellular Migration and Formation of Axons and Dendrites. https://doi.org/10.1016/B978-0-12-814407-7.00017-1 Copyright © 2020 Elsevier Inc. All rights reserved.

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CA1

CA2

SO SP SR SLM

CA3 SL

SM SG

F

H

DG FIGURE 17.1 Schema of hippocampal structure and lamination. CA, cornu ammonis; DG, dentate gyrus; F, fimbria; H, hilus; SO, stratum oriens; SP, stratum pyramidale; SR, stratum radiatum; SLM, stratum lacunosum-moleculare; SL, stratum lucidum; SM, stratum moleculare; SG, stratum granulosum.

The principal cell layer in the dentate gyrus is called the stratum granulosum. Superficial to the granule cell layer (GCL) is the stratum moleculare. The GCL and molecular layer are sometimes collectively referred to as the fascia dentata. Deep to the GCL is the polymorphic cell layer, usually referred to as the hilus. The thin layer at the border between the GCL and hilus is called the subgranular zone. Mossy cells are the main projection neurons in the hilus (Fig. 17.1).

17.2 Developmental specification of hippocampal fields 17.2.1 The basic developmental scheme of the hippocampus In adult rodents, the hippocampus is situated at the medialetemporal edge of the neocortex and harbors the neural circuitry essential for cognitive functions such as learning and memory (Lisman, 1999). The closure of the rostral neural tube begins around E8.5 from the presumptive forebrainemidbrain junction and proceeds toward the rostral end, which is completed by E9.0 except for the roof of the myelencephalon. Then it follows the outward expansion of the telencephalic vesicles and the concomitant invagination of the dorsomedial telencephalon. As the medial wall develops, it gives rise to a series of archipallial structures including (1) nonneuronal, secretory epithelium of the choroid plexus, which is adjacent to the roof; (2) the cortical hem, which is called the fimbria later when it is filled with projection fibers; (3) the hippocampal complex composed of the dentate gyrus and CA fields; and the subiculum, which is the transitional structure between the hippocampal complex and the medial limbic neopallium (the cingulate and retrosplenial cortex).

17.2.2 The cortical hem Several signaling centers have been found to govern embryonic development of the central nervous system via secreted signaling molecules (Lumsden and Krumlauf, 1996; Tanabe and Jessell, 1996). In agreement with this general theme, the cortical hem, an anatomical structure located at the edge of the hippocampus, has been revealed to express several members of the Wnt and Bmp families (Furuta et al., 1997; Grove et al., 1998). It has been hypothesized that Wnts and Bmps from the cortical hem can regulate hippocampal development (Grove and Tole, 1999; Grove et al., 1998). Furthermore, beyond being a signaling organizer in hippocampal development, the cortical hem also gives rise to a variety of cell types, including CajaleRetzius cells in the cortical marginal zone, glial cells in the fimbria, and epithelial cells of the choroid plexus, as shown by lineage analysis with the Wnt3a-cre line (Louvi et al., 2007; Yoshida et al., 2006).

17.2.3 The cortical hem organizes the hippocampal fields Several critical patterning events in succession are required for the proper development of the hippocampus, which include how the pallium is specified versus subpallium, how the pallium is divided into the neopallium and archipallium, and how the archipallium is further subdivided into fields. Several transcription factors have been found to be critical for these serial decisions. In Lhx2 mutants, the cortical hem and antihem are expanded at the expense of the pallial structures (Bulchand et al., 2001; Mangale et al., 2008; Monuki et al., 2001; Porter et al., 1997). In Emx1/2 compound mutants, the roof is expanded at the cost of the cortical hem and archipallium (Shinozaki et al., 2004), whereas, in the Foxg1 mutants, the

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cortical hem and archipallium are expanded at the expense of the neopallium (Muzio and Mallamaci, 2005). Therefore, the combined actions of Lhx2, Foxg1, Emx1, and Emx2 play a crucial role in defining the identity of the hippocampus. Analysis of Lhx2 and Foxg1 mutants suggests that suppression of Lhx2 and Foxg1 is essential for the specification of the cortical hem. Interestingly, Lhx2 null patches generated in the pallium from chimeric embryos displayed the marker expression profile characteristic of the cortical hem, and, more strikingly, hippocampal fields with proper order developed adjacent to each ectopic hem (Mangale et al., 2008). This evidence strongly argues that suppression of Lhx2 is sufficient to specify the fate of the cortical hem and that the cortical hem is competent to organize hippocampal development (Grove, 2008; Subramanian and Tole, 2009).

17.2.4 The role of canonical Wnt signaling in hippocampal development Wnt3a is expressed as early as E9.5 and marks the full length of the cortical hem (Louvi et al., 2007), which is followed by a series of Wnts expressed in the hem and medial pallium (Grove et al., 1998; Lee et al., 2000). Meanwhile, the Wnt receptors and signal transducers of the Tcf/Lef1 family are established in the medial pallium (Galceran et al., 2000; Kim et al., 2001). The hippocampus is completely missing in the Wnt3a mutant (Lee et al., 2000), and a knock-in dominant negative Lef1 mutant also displays similar defects (Galceran et al., 2000), whereas the Lef1 and Lrp6 mutants with decreased Wnt activity in the medial pallium show compromised development of the dentate gyrus and hilus only (Galceran et al., 2000; Li et al., 2008b; Zhou et al., 2004). Residual hippocampal field-specific cells are still detectable in the Wnt3a mutants, suggesting that Wnt signaling is important for the expansion of field-specific progenitors but not sufficient for field specification (Lee et al., 2000). However, when an activated form of Lef1 was expressed in the neopallium, a subset of hippocampal markers were ectopically induced (Machon et al., 2007), arguing that canonical Wnt signaling may be sufficient for specification and that compensation mechanisms by other Wnts may come into play in the Wnt3a mutants.

17.3 Migration of CajaleRetzius cells in the hippocampus 17.3.1 What are CajaleRetzius cells? The studies of human infants by Gustav Retzius (1893) and Santiago Ramón y Cajal (1890) led to the identification of a transient population of neurons located in the marginal zone of the developing cerebral cortex, which are commonly called the CajaleRetzius cells in the literature.

17.3.2 What are the functions of CajaleRetzius cells? As one of the earliest neuronal cell types generated in the developing telencephalon, the CajaleRetzius cells have become a subject of intense research in the past several decades, mainly because they are the major cellular source producing the extracellular glycoprotein Reelin (Alcantara et al., 1998) which is essential for cerebral development (D’Arcangelo et al., 1995; Ogawa et al., 1995). In the Reelin-mutant mice Reeler, there are profound defects in cortical lamination (Ogawa et al., 1995). It has been hypothesized that the CajaleRetzius cells, as one of critical cellular components in the marginal zone and secreting Reelin, organize the radial migration of cortical neurons and their laminar distribution via the actions of Reelin (Marin-Padilla, 1998; Rice and Curran, 2001; Stanco et al., 2009). Additionally, there are other functions proposed for CajaleRetzius cells. CajaleRetzius cells serve as pioneer neurons that can facilitate the development of hippocampal connectivity (Del Rio et al., 1997) and modulate the morphology of radial glial cells (Super et al., 2000).

17.3.3 What are the origins of CajaleRetzius cells? There was a long-held assumption that these cells are generated from ventricular progenitors and distributed locally after translocation into the subpial region (Marin-Padilla, 1998). However, this view has been fundamentally changed by recent findings that CajaleRetzius cells originate from focal regions and spread globally to cover the whole cortical surface. These focal sources are located at the borders of the developing pallium, which include the cortical hem (TakiguchiHayashi et al., 2004; Yoshida et al., 2006), the pallial septum, and the palliumesubpallium boundary (PSPB) (Bielle et al., 2005). These cells initially migrate radially to reach the pia, and then spread tangentially. CajaleRetzius cells of different origins display distinct molecular signatures: p73 (Meyer et al., 2004), p21 (Siegenthaler and Miller, 2005), and Zic1-3 (Inoue et al., 2008) for septal- and hem-derived cells; Er81 for septal-derived

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cells (Zimmer et al., 2010); and Ebf2 for cells of ventral pallial origin (Hanashima et al., 2007). These studies suggest that multiple pathways may be involved in the development of CajaleRetzius cells of different origins. It is not yet clear whether the heterogeneity of these CajaleRetzius cells in origin and molecular determinants reflects functional differences.

17.3.4 The cortical hem is the major source of CajaleRetzius cells for the dorsal telencephalon There have been several earlier studies implicating the cortical hem as a source of CajaleRetzius cells. Gdf7 is strictly expressed in the dorsal midline. Fate mapping by Gdf7-cre crossed with a reporter line shows that the roof plate structure can give rise to cells localized in the marginal zone (Monuki et al., 2001). Shortly after that report, Trp73 was found to be exclusively colocalized with Reelin in neocortical CajaleRetzius cells (Meyer et al., 2002). Trp73 was highly expressed in the cortical hem early on and then displayed a mediolateral gradient, which led to the notion that Trp73-positive CajaleRetzius cells in the neocortex are derived from the cortical hem (Meyer et al., 2002). In support of this, Cajale Retzius cells were found to move out from the cortical hem in a green fluorescent protein (GFP) insertional mouse line which serendipitously labels Reelin-positive CajaleRetzius cells in the marginal zone of the developing cortex (TakiguchiHayashi et al., 2004), and in Fzd10-taulacZ transgenic mice, LacZ mRNA is restricted to the cortical hem but the LacZ protein continues to be detectable in CajaleRetzius cells in the cortical marginal zone (Zhao et al., 2006). Further definitive evidence came from cumulative fate mapping analysis and cell ablation experiments with Wnt3a-cre (Yoshida et al., 2006) and DN-Trp73-cre (Tissir et al., 2009) lines. The number of hem-derived CajaleRetzius cells is regulated at their origin by transcription factors Lhx5 (Zhao et al., 1999), Lhx2 (Bulchand et al., 2001; Monuki et al., 2001), Foxg1 (Hanashima et al., 2007; Muzio and Mallamaci, 2005; Zhao et al., 2006), Emx2 (Mallamaci et al., 2000), Pax6 (Stoykova et al., 1997), etc. The balance of Foxg1 expression in the cortical hem is especially critical for the production and fate specification of CajaleRetzius cells (Liu et al., 2018). Loss of Foxg1 leads to expansion of CajaleRetzius cells, whereas forced continued expression leads to dentate granule neuron fate (Muzio and Mallamaci, 2005). Foxg1 is an upstream transcription factor of Lhx2, and high expression of Lhx2 leads to a shift in cell fate from the cortical hem to hippocampal specification (Bulchand et al., 2001; Godbole et al., 2018).

17.3.5 The extent of the cortex covered by hem-derived CajaleRetzius cells The hem-derived CajaleRetzius cells migrate in a caudomedial to rostrolateral direction to cover the hippocampus and the neocortex until they reach the ventrolateral boundary (Tissir et al., 2009; Yoshida et al., 2006). They somewhat penetrate the boundary but tend to avoid invading into the olfactory cortex and olfactory bulb. Those regions seem to be dominated by CajaleRetzius cells derived from the septum and PSPB (Bielle et al., 2005).

17.3.6 Recruitment of hem-derived CajaleRetzius cells to the meninges Explant analysis with the hem and various cortical tissues suggests that the meninges produce a chemoattractant for hemderived CajaleRetzius cells which is important for regulating the marginal zone localization of these cells (Borrell and Marin, 2006). Cxcl12, a chemo-kine secreted by the meningeal fibroblasts, promotes the anchorage of the Cxcr4expressing CajaleRetzius cells to the meninges (Borrell and Marin, 2006; Paredes et al., 2006) after they migrate radially away from the hem ventricular zone. In the Cxcr4 mutant mice, CajaleRetzius cells tend to descend from the meninges and get stuck in the developing cortical plate. Cell displacement is particularly prominent in the regions close to the hem, such as the dentate gyrus and CA3 (Borrell and Marin, 2006; Paredes et al., 2006).

17.3.7 Tangential dispersion of CajaleRetzius cells in the marginal zone Although the meninges provide attractant signals to CajaleRetzius cells, they do not supply directional guidance cues for the migration of CajaleRetzius cells to their final destination. Time-lapse imaging in vitro and in vivo indicates that contact-mediated repulsion between CajaleRetzius cells imposes restriction on their movement and drives the formation of the final distribution pattern (Borrell and Marin, 2006; Villar-Cervino et al., 2013). This repulsion is mediated by a set of Eph/ephrin molecules, EphB1/B2/B3, contributing to the widespread distribution of CajaleRetzius cells on the cortical surface (Villar-Cervino et al., 2013). Indeed, CajaleRetzius cells fail to distribute throughout the cortical surface in EphB1/ B2/B3 mutants. Furthermore, there is evidence that this contact repulsion is important for maintaining boundaries between groups of CajaleRetzius cells from different origins. Decreasing contact repulsion by inhibiting Eph/ephrin signaling

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increases the diffusion of CajaleRetzius cells into other territories and blurs the boundaries (Villar-Cervino et al., 2013). Because the repulsion between CajaleRetzius cells controls the distribution of these cells, the migration speed of CajaleRetzius cells becomes a key parameter influencing the final distribution pattern in the cortical surfaces (Barber et al., 2015). The migration speed of CajaleRetzius cells is independently regulated by different mechanism from that in cellecell repulsion. Vesicle-associated membrane protein 3 (VAMP3), highly expressed in CajaleRetzius cells, negatively controls the migration speed of CajaleRetzius cell in a cell autonomous manner. Deletion of VAMP3 increases the migration speed of CajaleRetzius cells, especially in cortical hem- and stratum-derived CajaleRetzius cell, which results in their ectopic distribution in other CajaleRetzius cell territories and alters the final distribution pattern of CajaleRetzius cells in the cortical surface (Barber et al., 2015). In the hippocampus, CajaleRetzius cells tangentially invade the border between the CA1 and the dentate gyrus along the hippocampal fissure and settle in the suprapyramidal regions of the pial surface in dentate gyrus in the first postnatal week. Meanwhile, Reelin released from CajaleRetzius cells has multiple roles regulating the migration of neurons and morphology of radial glial cells during the development of hippocampus and dentate gyrus (see more discussion in Section 17.6).

17.4 Migration of hippocampal pyramidal neurons The hippocampal pyramidal cells are derived from the radial glia at the germinal ventricular zone of the hippocampal primordium around E14 in mice (Angevine, 1965). Neuronal precursors leaving the ventricular zone display multipolar morphology (Nakahira and Yuasa, 2005). Meanwhile, they sojourn for about 4 days in the subventricular and intermediate zones before settling into the hippocampal plate, the presumptive stratum pyramidale (Altman and Bayer, 1990b; Nakahira and Yuasa, 2005). Analogous to the development in the neocortex, the hippocampal plate splits the preplate into subplate and marginal zone. Subplate corresponds to the prospective stratum oriens, whereas the marginal zone turns into inner and outer marginal zones, which develop into the prospective stratum radiatum and stratum lacunosum-moleculare, respectively. The migration of the pyramidal neurons also follows an “inside-out” sequence as in the neocortex where early-born neurons constitute the deep location of the pyramidal layer and late-born neurons take up the superficial location. However, instead of forming layers of functionally distinct neurons like neocortex, the pyramidal neurons are populated in the hippocampus as one single homogeneous layer. Furthermore, hippocampal pyramidal neurons migrate with distinct patterns compared to cortical neurons. During cortical development, differentiated neurons from radial glia cells migrate toward the pia using the radial fiber of the parent cell (Noctor et al., 2004). Newborn neurons have a multipolar shape in the SVZ when they are first produced. Then, as they start to migrate to the cortical surface, they adopt a bipolar shape by extending their processes along an apical-basal axis with the radial fiber (Noctor et al., 2004). Similarly, pyramidal neurons in the hippocampus also form a multipolar shape just above the ventricular zone and become bipolar as they leave for the stratum pyramidale (Kitazawa et al., 2014). However, the processes of pyramidal neurons, especially the leading directional processes, are highly branched compared to cortical migrating neurons (Kitazawa et al., 2014). Pyramidal neurons simultaneously attach to multiple radial glial fibers with their processes, and use them as a scaffold to migrate to their final destination. While migrating, pyramidal neurons switch between radial fiber scaffolds and as a result, they migrate in a zigzag manner. This migration pattern has been called the “climbing migration mode,” and is quite distinct from the radial straight path that cortical neurons employ to reach their final destination (Kitazawa et al., 2014). Compared with cortical neurons, the duration that pyramidal neurons take to migrate to their final position is longer even though their migration distance is considerably shorter. The signaling pathways that regulate the radial glia-guided migration in the neocortex also play pivotal roles in hippocampal development. The pathways involving Reelin, Cdk5, and Dcx are the most prominent among these. Reelin is secreted largely by the CajaleRetzius cells and to a lesser extent by the GABA (g-aminobutyric acid)-ergic interneurons in both neocortex and hippocampus (Alcantara et al., 1998). The presumptive role of CajaleRetzius cells in cortical lamination (Marin-Padilla, 1998; Rice and Curran, 2001; Soriano and Del Rio, 2005) is challenged by recent studies in which normal lamination was still maintained even with the massive loss of CajaleRetzius cells (Bielle et al., 2005; Tissir et al., 2009; Yang et al., 2000; Yoshida et al., 2006). However, the essential role of Reelin (which is mutated in the Reeler mice) in the cortical and hippocampal lamination has been firmly established by the genetic evidence (D’Arcangelo et al., 1995; Ogawa et al., 1995). Duplication of the pyramidal cell layer in CA1 was seen in Reeler mice as a consequence of malpositioning of the pyramidal neurons (Caviness and Sidman, 1973; Stanfield and Cowan, 1979). Reelin initiates intracellular signaling by engaging the two membrane receptors: the very low-density lipoprotein receptor (Vldlr) and the apolipoproliprotein E receptor 2 (Apoer2) (D’Arcangelo et al., 1999; Hiesberger et al., 1999). Intracellular signaling is further transmitted by the adaptor protein Dab1. Mutants lacking both receptors or Dab1 show phenotypes indistinguishable from the Reeler (Howell et al., 1997; Sheldon et al., 1997; Trommsdorff et al., 1999; Ware et al., 1997).

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Cyclin-dependent kinase 5 (Cdk5) is a homolog of the cyclin-dependent kinase family of serine/threonine kinases. Abnormal stratification was observed in the Cdk5 mutants as manifested by the loose organization of the hippocampal pyramidal layer (Ohshima et al., 1996). Granule cell dispersion and CA1eCA3 pyramidal cell disorganization are also associated with Lis1 haploinsufficiency (Wang and Baraban, 2008). Doublecortin (Dcx) is a microtubule-associated protein encoded by X-linked gene (Kerjan and Gleeson, 2007). Heterozygous females and hemizygous males displayed partial splitting of the pyramidal cell layer in CA3 and loose organization in other areas. In addition, disruptions of pyramidal layer were detected in CA1 and CA2 regions (Corbo et al., 2002). Dcx has two close homologs in mice: doublecortin-like kinase 1 (Dclk1) and doublecortin-like kinase 2 (Dclk2). Dcx; Dclk2 double null mice displayed more severe delamination in the hippocampus, including ectopic neurons in stratum oriens and discontinuous CA1 field (Kerjan et al., 2009). How Reelin mechanistically regulates radial migration is still controversial (Soriano and Del Rio, 2005; Tissir et al., 2009). More intriguingly, the Reelin pathway may cross talk with Cdk5 somewhere downstream of Dab1 (Beffert et al., 2004). Besides, the pyramidal neurons in the hippocampus failed to form a discrete layer in the alpha-N-catenin mutant (Park et al., 2002), raising the possibility that cell adhesion molecules contribute to the compaction of pyramidal cells at the last stage of migration.

17.5 Migration of hippocampal interneurons 17.5.1 Cellular and distributional diversity of interneurons in the hippocampus The GABAergic interneurons in the cerebral cortex (which includes neo-, paleo-, and archicortex) are a population of cells with tremendous diversity. To tackle this, morphological, molecular, and physiological criteria have been proposed for describing the different features of interneurons (Petilla Interneuron Nomenclature et al., 2008). However, these criteria may have limitations under certain circumstances, since for each single interneuron these features are potentially subjected to change due to developmental stages or neural plasticity. Nevertheless, the use of molecular markers is the most common approach to define the interneurons in the analysis of their developmental origins and migratory routes toward different destinations. In this context, the molecular approach has the obvious advantage over morphological or physiological approaches in terms of following a cohort of interneurons over a long developmental period or making quantification over a wide range of anatomical structures. Beyond the cellular diversity, each subtype usually displays a very distinct spatial distribution, as exemplified by a study of the allocation of different molecularly defined interneuron subtypes in the hippocampus (Jinno and Kosaka, 2006). By taking a stereological approach to avoid sampling bias in tissue levels and orientations, this study reveals striking differences in laminar distributions and numerical densities along the dorsoventral axis of the hippocampus among nine neurochemical markers, including GAD67, calcium-binding proteins (parvalbumin, calretinin, and calbindin), neuropeptides (somatostatin, neuropeptide Y (NPY), cholecystokinin, vasoactive intestinal protein), and neuronal nitric oxide synthase.

17.5.2 Origins and migration of hippocampal interneurons The embryonic telencephalon consists of the pallium and subpallium. The pallium gives rise to the neocortex and hippocampus, and subpallium to the basal ganglia. The analysis of mice harboring the mutations in the homeobox genes Distal-less 1 and 2 (Dlx1/2), which are expressed in the subpallium, reveals a dramatic reduction of interneurons in the neocortex (Anderson et al., 1997) and an almost complete loss of interneurons in the hippocampus (Pleasure et al., 2000), thus providing evidence that hippocampal interneurons originate in the subpallial telencephalon and migrate tangentially to their pallial targets. The embryonic ganglionic eminence can be divided into three different domains: lateral ganglion eminence (LGE), medial ganglion eminence (MGE), and caudal ganglion eminence (CGE) (Fig. 17.2). These domains have differential contribution to different interneuron subtypes. MGE-derived interneurons give rise to the majority of parvalbumin (PV) and somatostatin (SOM) expressing subtypes, whereas calretinin (CR) and vasoactive intestinal peptide (VIP) expressing subtypes mainly come from the CGE (Butt et al., 2005; Miyoshi et al., 2007; Xu et al., 2004). A majority of interneurons in the hippocampus are produced from the MGE prior to interneuron production from the CGE (Pelkey et al., 2017; Tricoire et al., 2011). The fact that each region produces a different set of interneuron subtypes raises the possibility that these regions have distinct radial glia cells that are restricted to produce one particular interneuron subtype, or that individual

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FIGURE 17.2 Origins and migratory routes for hippocampal interneurons. (A) A schematic coronal section at the level that contains LGE and MGE at E15.5; (B) a schematic coronal section at the level that contains CGE at E15.5; (C) a schematic horizontal section that contains LGE, MGE, and CGE at E15.5. Hippocampal interneurons originated at LGE, MGE, POA (A), or CGE (B) take a lateral route and course through the neocortex before reaching the hippocampus. Alternatively, some hippocampal interneurons from CGE (C) take a caudal route to enter the hippocampus. CGE, caudal ganglion eminence; hip, hippocampus; LGE, lateral ganglion eminence; MGE, medial ganglion eminence; ncx, neocortex; POA, preoptic area; S, septum.

radial glia cells have the ability to produce multiple subtypes of interneurons. The studies traced the fate of sibling interneurons produced from the same parent cell, reveal that individual radial glia cells in the MGE can produce both PVand SOM-expressing interneurons (Mayer et al., 2015; Petros et al., 2015; Sultan et al., 2016). During cortical development, radial glia cells asymmetrically divide to produce neurons at early stages of cortical development. Subsequently, they start to give rise to the intermediate progenitors which symmetrically produce neurons. SOM-expressing interneurons are asymmetrically produced from radial glia cells directly, whereas PV-expressing interneurons are generated from intermediate progenitors through symmetric cell division (Petros et al., 2015). Clonal analysis also reveals that although the majority of clonally related interneurons are predominantly distributed in one brain structure, a portion of sibling interneurons migrate and widely disperse across both cortex and hippocampus (Mayer et al., 2015; Sultan et al., 2016). In addition to MGE and CGE, the preoptic area (POA) can also produce hippocampal interneurons although its contribution is minor (Gelman et al., 2009, 2011). The POA produces highly diverse subtypes of interneurons, including PV, SOM, NPY, and reelin expressing interneurons (Gelman et al., 2009, 2011; Liodis et al., 2007; Zhao et al., 2008). Alternatively, distinct interneuron subtypes arise from the same region in different temporal waves (Butt et al., 2005; Fogarty et al., 2007; Miyoshi et al., 2007). The development of interneuron subtypes in the hippocampus may involve similar mechanisms, although it needs to be validated by future investigation. The production of interneurons from different origins is regulated by different transcription factors. Nkx2.1 is critical for MGE-derived interneurons. The number of hippocampal neurons in the Nkx2.1 mutant is dramatically reduced as a result of misspecification of MGE into LGE (Pleasure et al., 2000). COUP-TFII is preferentially expressed in the CGE and in the interneurons of the caudal migratory stream. Caudal migration of the CGE-derived cells requires COUP-TFII, and ectopic expression of COUP-TFII in MGE tissue is sufficient to drive the caudal migration once transplanted into CGE (Kanatani et al., 2008). Interneurons of different origins take on distinct migratory routes to leave the subpallium. Interneurons derived from the MGE migrate laterally and bypass the striatum before they enter the cortex and hippocampus (Li et al., 2008a; Marin and Rubenstein, 2001; Marin et al., 2001), whereas the CGE-derived cells migrate into the cortex and hippocampus through either laterally to the cortex or caudally to the caudal-most end of the telencephalon, respectively (Kanatani et al., 2008; Touzot et al., 2016; Yozu et al., 2005). Aside from these migration routes of CGE-derived cells, they also exhibit the medial migratory stream, by which CGE-derived cells migrate into the amygdala through the medial telencephalon (Touzot et al., 2016). Once the interneurons reach the pallium, both MGE- and CGE-derived cells follow a similar migratory route to enter the hippocampus (Tricoire et al., 2011). They migrate tangentially toward their final cortical destination by two different streams: one in the subventricular zone (SVZ) or lower IZ, and the other one in the marginal zone (MZ) (Jimenez et al., 2002; Lavdas et al., 1999; Nery et al., 2002; Polleux et al., 2002). In the hippocampus, Dlx2 positive cells are detected as early as E15.5 in the hippocampal marginal zone and E16.5 in the hippocampal SVZ (Pleasure et al., 2000). Similarly, hippocampal interneurons have been shown to populate the hippocampal primordium at E15 by using the Gad67eGfp knock-in embryos (Manent et al., 2006). At this stage, the sizable superficial stream courses toward the subiculum and the CA1 field, whereas the smaller deep stream approaches

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the border between the neocortex and the subiculum. By E16, the superficial MZ stream already arrives at CA3, whereas the deep stream just passes the CA1 field. Interneurons invade the dentate gyrus primordium via the superficial stream by E17. Tangentially migrating interneurons navigate below the layer of CajaleRetzius cells in the MZ. Interestingly, during late gestation, interneurons are present within the SVZ and the inner MZ (future stratum radiatum), whereas CajaleRetzius cells are located in the outer MZ (future stratum lacunosum-moleculare). Therefore, distinct neuronal populations appear to populate different laminae of the hippocampus by birth. Interneurons of different origins have distinct preference for their hippocampal targets. The MGE-derived interneurons migrate to the hippocampus CA regions and avoid the dentate gyrus (Pleasure et al., 2000; Wichterle et al., 2001), while the CGE-derived interneurons migrate to both the CA and the dentate gyrus regions (Nery et al., 2002). Interneurons destined for hippocampus interact with the trophic factors or basement membrane components secreted from the meningeal cells. MGE-derived interneurons labeled by the Lhx6eGfp line show diminished marginal stream and premature invasion into the hippocampal plate by the end of the gestation in the Cxcr4 mutant (Li et al., 2008a). Interneuron-specific ablation of integrin a3 in the Netrin-1 null background resulted in a significant reduction of marginal interneurons that reached the hippocampus at birth (Stanco et al., 2009). Thus, the meninges can exert its influence on the tangential dispersion of hippocampal interneurons in different ways.

17.6 Migration of neural progenitors and granule cells in the dentate gyrus during development 17.6.1 The basic developmental scheme of the dentate gyrus The formation of the dentate gyrus involves the successive development of the germinative matrices at different locations for distinct stages (Altman and Bayer, 1990a, b; Li and Pleasure, 2005) (Fig. 17.3). Each of these proliferative components gives rise to granule cells that end up at different parts of the ultimate dentate GCL. During the transition from one germinative matrix to the next one, the migration of the proliferative precursors takes a particular route and is regulated by a distinct set of molecules. The granule cells produced from each individual germinative matrix involve a unique developmental plan to settle in the dentate plate. In contrast to the “inside-out” laminar formation of neocortex, the GCL in the dentate gyrus is established in an “outside-in” manner, in which the outer shell of the GCL forms first and newborn granule neurons are continuously added inside of the GCL (Mathews et al., 2010). During early embryonic development, the putative primary dentate neuroepithelium is distinguished from its neighboring regions by its location, shape, and cellular dynamics. It is located adjacent to a ventricular indentation called the dentate notch (Fig. 17.3A and B). This primary dentate neuroepithelium contains fewer mitotic cells near the lumen of the ventricle than the hippocampal neuroepithelium. At the beginning of the last week of murine embryonic development,

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FIGURE 17.3 Formation of the germinative matrices at different locations during the development of the dentate gyrus. (A) The primary germinative matrix (green dots) forms at the ventricular zone and the subventricular zone of the dentate primordium around E13.5; (B) the secondary germinative matrix (blue dots) develops around E15.5 in the junctional area between the fimbria and the developing dentate pole. These cells develop the glial fiber extending to the developing hilus (blue line); (C) the tertiary matrix (orange dots) emerges in the hilus around perinatal ages, while the secondary germinative matrix further extends subpially and wraps around the presumptive dentate pole; and (D) the final germinative matrix (red dots) is established in the subgranular zone by the second postnatal week and develop a secondary radial glial fiber toward to the molecular layer of dentate gyrus (red line). D, dentate primordium; dgl, dentate granular layer; DN, dentate notch; F, fimbria; FDJ, fimbriodentate junction; H, hilus; HF, hippocampal fissure; Hip, hippocampal primordium.

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the primary dentate neuroepithelium gives rise to the first wave of granule cells (Li et al., 2008c; Zhou et al., 2004), which initially settle at the tip of the future dentate upper blade and proceed toward the lower blade, thus forming a suprapyramidal to infrapyramidal morphogenic gradient (Altman and Bayer, 1990a, b; Li et al., 2008c). Meanwhile, a collection of proliferative cells emerge in the subventricular region of the primary dentate neuroepithelium and extend toward the pia, which is defined as the secondary germinative matrix by Altman and Bayer (Altman and Bayer, 1990a). As development proceeds, these cells are then localized subpially all the way toward the hippocampal fissure (Li et al., 2008c; Fig. 17.3C). Subpial neurogenesis produces granule cells that constitute the outer shell of the dentate plate (Altman and Bayer, 1990a; Li et al., 2008c). Subpial proliferation starts to subside in the second half of the first postnatal week and gradually disappears in the second week. At perinatal ages, a population of mitotic cells segregate from the secondary germinative matrix and appear in the hilar region (Altman and Bayer, 1990a; Li et al., 2008c), which is defined as the tertiary germinative matrix by Altman and Bayer (1990a); Fig. 17.3C. The tertiary germinative matrix is highly active during the first postnatal week and generates a very large number of granule cells which are deposited into the inner shell of the dentate plate and integrated in the GCL (Altman and Bayer, 1990a; Li and Pleasure, 2005). Toward the end of the second postnatal week, the tertiary germinative matrix is succeeded by the proliferative subgranular zone (SGZ) located at the border between the GCL and the hilus (Fig. 17.3D), which serves as the major site for persistent neurogenesis in the adult hippocampus (Altman and Bayer, 1990a; Li et al., 2008c; Li and Pleasure, 2005).

17.6.2 Migration of granule neurons to form the granule cell layer Similar to cortical neurons during development, granule neurons also utilize glial fibers as a scaffold for migration. The migration of granule neurons into the primitive dentate gyrus and formation of the GCL in the dentate are governed by the glial fibers of precursors in the secondary and tertiary germinative matrix, respectively (Brunne et al., 2013; Li et al., 2008c). When precursors in the secondary germinative matrix reach the subpial region between fimbria and dentate (fimbria-dentate junction: FDJ), they extend their glial projections to the developing hilus (Li et al., 2008c; Fig. 17.3B). Newborn granule cells at the ventricular zone or the subpial region are likely to utilize these transhilar radial glial fibers to climb into the presumptive upper blade. Precursor cells localized in the FDJ express chemokine receptor Cxcr4 and is regulated by meningeally expressed Cxcr4 ligand, Cxcl12. In Cxcr4 mutants, precursors are dislodged from the subpial region (Li et al., 2008c; Lu et al., 2002), which subsequently affects both the development of transhilar radial glial fibers and the migration of granule cells derived from the secondary germinative matrix (Bagri et al., 2002; Li et al., 2008c). As a consequence, a disorganized upper blade is formed and misplaced granule cells become continuous with the CA3 pyramidal neurons. Reelin signaling is not required for the subpial localization of precursors, but it is critical for their progeny to migrate into the forming dentate plate and to establish the GCL (Li et al., 2008c). In Reeler mice, granule cells are dispersed in the whole dentate pole and the organization of the dentate blade is lost (Li et al., 2008c; Stanfield and Cowan, 1979). A set of studies about the role of reelin signaling in the dentate gyrus demonstrate that the glial processes of precursors in tertiary germinative matrix contribute to the migration of granule neurons to form a V shape of CGL (Brunne et al., 2013; Forster et al., 2002; Frotscher et al., 2003; Hartfuss et al., 2003; Weiss et al., 2003). The precursors in tertiary germinative matrix, which reach hilus, develop the radial glial process, called secondary radial glial scaffold, toward the pial surface (Brunne et al., 2013; Fig. 17.3D). In Reeler or Scrambler mice lacking Dab1, these radial glial fibers are directionless and disorganized. Interestingly, in contrast to the scattered granule neurons in the dentate gyrus of Reeler mice, inducing Dab1 deletion specific to the granule neurons, but not precursors, leads to milder defects of GCL formation, in which the V shape of the GCL is established, albeit with delays in formation and migratory defects in a portion of granule neurons (Brunne et al., 2013; Forster et al., 2002). In this mutant, radial glial fibers are properly established, suggesting that reelin stimulates precursors to develop the glial scaffold, thereby contributing to the migration of the granule neurons for GCL formation. The importance of radial fibers for granule neuron migration is also demonstrated in mutant mice lacking the proper secondary glial scaffold. The “V” shape of the subgranular zone in its mature state mirrors the contour of the basement membrane of the dentate pole such that the dentate radial glial scaffolding is arranged perpendicular to the pia with their end feet attached to the basement membrane. In brain, specific deletion mutants of integrin-b1 (Forster et al., 2002), focaladhesion kinase (Fak) (Beggs et al., 2003), integrin-linked kinase (Ilk) (Niewmierzycka et al., 2005), and dystroglycan (Moore et al., 2002) radial glial fibers are regionally disorganized, in which the GCL adopts an extremely wavy appearance. Reelin plays a role as an attractant signal during the development of the radial glial process (Frotscher et al., 2003; Zhao et al., 2004). Reflecting the direction that radial fibers extend to the pia, expression of reelin is observed along the pial

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surface during the development of the secondary radial glial fiber (Brunne et al., 2013). CajaleRetzius cells occupy the pial surface and function as a source of reelin for precursors. Loss of CajaleRetzius cells resulting from Trp73 mutations leads to a loss of the dentate lower blade and hypoplasia of radial glial fibers (Meyer et al., 2004; Yang et al., 2000).

17.6.3 Emergence and migration of long-lived neural stem cells and establishment of subgranular zone There are two limited regions in which neurogenesis persists into adulthood in the rodent brain; the SVZ of the lateral ventricles and the SGZ of the dentate gyrus in the hippocampus. Both regions inherit neural stem cells (NSCs) during developmental stages, and neurogenesis persists in these NSC pools throughout life. However, the establishment of longlived NSCs in each region is achieved through different developmental processes. NSCs in the SVZ originate from the ventricular zone of lateral ventricles and are maintained well into adulthood in the same region where they exist during the developmental stage (Kriegstein and Alvarez-Buylla, 2009). In contrast, since the dentate gyrus is established in the forebrain during development, NSCs in the SGZ are required to migrate into the dentate gyrus (Altman and Bayer, 1990a; Li et al., 2013). This migration step is the characteristic feature of the establishment of long-lived NSCs in the SGZ. Long-lived NSCs in the SGZ initially emerge in the ventral hippocampus during late gestation in response to Shh signaling activity (Li et al., 2013, Fig. 17.4). The descendants of these cells migrate into the dorsal hippocampus along the longitudinal axis of the hippocampus from the temporal to septal poles and become part of the tertiary germinative matrix. Afterward, these descendant cells eventually settle at the border of the hilus and GCL, forming the SGZ. The emergence of long-lived NSC pools is regulated by Shh signaling activity. Long-lived NSCs are generated from Shh-responding cells in the ventral hippocampus (Li et al., 2013). Shh ligands are highly secreted from the amygdala in prenatal stages, which likely contributes to the production of Shh-responding cells in the ventral hippocampus (Li et al., 2013). The ablation of Shh signaling activity in NSCs leads to deficiency of the SGZ (Breunig et al., 2008; Li et al., 2013). After progeny of Shhresponding cells leave the amygdala and reach the dentate gyrus in postnatal stages, mossy cells located in the hilus function as a continued source of Shh ligands and contribute to the maintenance of NSCs into adulthood (Li et al., 2013). In addition to the amygdala and local neurons in dentate gyrus, epithelial cells are also shown to supply Shh ligands in prenatal and postnatal dentate gyrus (Choe et al., 2015). These Shh ligands are transported into the dentate gyrus through the bloodebrain barrier via platelets and contribute to the proliferation and maintenance of NSCs. It is not clear yet how ventrally derived NSCs move into the primitive dentate gyrus. However, there are several guidance molecules presumably involving in their migration to their final positions. The subgranular localization of NSCs can be reinforced by cellecell interaction between NSCs and granule cells. Interestingly, loss of Ephb1 or Ephb2, which are expressed in NSCs, results in the displacement of NSCs from the presumptive subgranular zone (Chumley et al., 2007).

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FIGURE 17.4 Origin and migratory route of long-lived neural stem cells (NSCs) in dorsal dentate gyrus. The illustration of coronal (A) and sagittal (B) sections of developing mouse brain at P0. Shh-responding cells emerge in the ventral hippocampus in response to Shh ligands from amygdala (blue dots) and contribute to the production of long-lived NSCs. The descendants of Shh-responding cells migrate along the longitudinal axis of the hippocampus from the temporal (blue line) to septal (red line) poles and eventually settle in the subgranular zone as long-lived NSCs. These NSCs are maintained into adult and contribute to the daily production of newborn neurons throughout a life.

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Mutation of Efnb3, which is expressed in the GCL, phenocopies the Ephb1 and Ephb2 mutants. Therefore, Efnb3 signaling mediated by Ephb1 or Ephb2 controls the positioning of NSCs in the SGZ (Chumley et al., 2007). After NSCs settle in the SGZ, they develop secondary radial glial fibers and continuously produce granule neurons, the majority of granule neurons in the GCL. Concomitantly, postnatal NSCs are also shown to contribute to the production of astrocyte in the molecular layer of the dentate gyrus (Brunne et al., 2010). Radial NSCs directly differentiate into astrocytes from the SGZ and migrate to the molecular layer across the GCL. Even after the completion of the dentate gyrus development, NSCs are maintained in the SGZ into adulthood and contribute to hippocampal function by producing new neurons daily (Deng et al., 2010). Newly generated neurons tangentially migrate along the blood vessel to final positions in the GCL (Sun et al., 2015). These newborn neurons are integrated into the preexisting hippocampal neuronal circuit and are implicated in hippocampus-dependent learning and memory (Deng et al., 2010; Imayoshi et al., 2008).

17.7 Conclusions The hippocampus is a complex neural structure that depends for its development on a host of distinct cell types that are assembled during embryonic and postnatal development from sources both within and outside the medial pallium. One of the more unique aspects of hippocampal development is the formation of the dentate gyrus, which involves the formation of a specialized neural stem cell niche. The assembly of this structure and all the other principal and nonprincipal cells of the hippocampus utilize molecular and cellular strategies also important elsewhere in the cortex, but they are arranged in unique ways to allow the formation of the specialized hippocampal circuit. In the years to come, there will be major new advances in understanding how the distinct cell types are produced but also how they assemble themselves into a functioning circuit.

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Tricoire, L., Pelkey, K.A., Erkkila, B.E., Jeffries, B.W., Yuan, X., McBain, C.J., 2011. A blueprint for the spatiotemporal origins of mouse hippocampal interneuron diversity. J. Neurosci. 31, 10948e10970. Trommsdorff, M., Gotthardt, M., Hiesberger, T., Shelton, J., Stockinger, W., Nimpf, J., Hammer, R.E., Richardson, J.A., Herz, J., 1999. Reeler/Disabledlike disruption of neuronal migration in knockout mice lacking the VLDL receptor and ApoE receptor 2. Cell 97, 689e701. Villar-Cervino, V., Molano-Mazon, M., Catchpole, T., Valdeolmillos, M., Henkemeyer, M., Martinez, L.M., Borrell, V., Marin, O., 2013. Contact repulsion controls the dispersion and final distribution of Cajal-Retzius cells. Neuron 77, 457e471. Wang, Y., Baraban, S.C., 2008. Aberrant dentate gyrus cytoarchitecture and fiber lamination in Lis1 mutant mice. Hippocampus 18, 758e765. Ware, M.L., Fox, J.W., Gonzalez, J.L., Davis, N.M., Lambert de Rouvroit, C., Russo, C.J., Chua Jr., S.C., Goffinet, A.M., Walsh, C.A., 1997. Aberrant splicing of a mouse disabled homolog, mdab1, in the scrambler mouse. Neuron 19, 239e249. Weiss, K.H., Johanssen, C., Tielsch, A., Herz, J., Deller, T., Frotscher, M., Förster, E., 2003. Malformation of the radial glial scaffold in the dentate gyrus of reeler mice, scrambler mice, and ApoER2/VLDLR-deficient mice. J. Comp. Neurol. 460, 56e65. Wichterle, H., Turnbull, D.H., Nery, S., Fishell, G., Alvarez-Buylla, A., 2001. In utero fate mapping reveals distinct migratory pathways and fates of neurons born in the mammalian basal forebrain. Development 128, 3759e3771. Xu, Q., Cobos, I., De La Cruz, E., Rubenstein, J.L., Anderson, S.A., 2004. Origins of cortical interneuron subtypes. J. Neurosci. 24, 2612e2622. Yang, A., Walker, N., Bronson, R., Kaghad, M., Oosterwegel, M., Bonnin, J., Vagner, C., Bonnet, H., Dikkes, P., Sharpe, A., et al., 2000. p73-deficient mice have neurological, pheromonal and inflammatory defects but lack spontaneous tumours. Nature 404, 99e103. Yoshida, M., Assimacopoulos, S., Jones, K.R., Grove, E.A., 2006. Massive loss of Cajal-Retzius cells does not disrupt neocortical layer order. Development 133, 537e545. Yozu, M., Tabata, H., Nakajima, K., 2005. The caudal migratory stream: a novel migratory stream of interneurons derived from the caudal ganglionic eminence in the developing mouse forebrain. J. Neurosci. 25, 7268e7277. Zhao, C., Guan, W., Pleasure, S.J., 2006. A transgenic marker mouse line labels Cajal-Retzius cells from the cortical hem and thalamocortical axons. Brain Res. 1077, 48e53. Zhao, S., Chai, X., Förster, E., Frotscher, M., 2004. Reelin is a positional signal for the lamination of dentate granule cells. Development 131, 5117e5125. Zhao, Y., Flandin, P., Long, J.E., Cuesta, M.D., Westphal, H., Rubenstein, J.L., 2008. Distinct molecular pathways for development of telencephalic interneuron subtypes revealed through analysis of Lhx6 mutants. J. Comp. Neurol. 510, 79e99. Zhao, Y., Sheng, H.Z., Amini, R., Grinberg, A., Lee, E., Huang, S., Taira, M., Westphal, H., 1999. Control of hippocampal morphogenesis and neuronal differentiation by the LIM homeobox gene Lhx5. Science 284, 1155e1158. Zhou, C.-J., Zhao, C., Pleasure, S.J., 2004. Wnt signaling mutants have decreased dentate granule cell production and radial glial scaffolding abnormalities. J. Neurosci. 24, 121e126. Zimmer, C., Lee, J., Griveau, A., Arber, S., Pierani, A., Garel, S., Guillemot, F., 2010. Role of Fgf8 signalling in the specification of rostral Cajal-Retzius cells. Development 137, 293e302.

Chapter 18

Hindbrain tangential migration Constantino Sotelo and Alain Che´dotal Institut de la Vision, Sorbonne Université, INSERM, CNRS, Paris, France

Chapter outline 18.1. Introduction 381 18.2. Tangential migration: a historical overview 382 18.3. Molecular mechanisms controlling the tangential migration of precerebellar neurons 384 18.3.1. Influence of the midline on tangentially migrating precerebellar neurons 384 18.3.2. Why do PCN neurons migrate near the pial surface? 388 18.4. Molecular mechanisms controlling the tangential migration of facial motor neurons 389 18.4.1. Origin and migration of facial motor neurons 389

18.4.2. The caudal migration of FBM neurons 18.4.2.1. The planar cell polarity pathway 18.4.2.2. Other molecules controlling FBM caudal migration 18.4.3. Role of chemoattraction and chemorepulsion 18.4.4. Role of the meninges in the tangential migration of FBM neurons 18.5. Ending tangential migration 18.6. Conclusion Acknowledgments References

390 390 391 392 393 394 395 395 395

18.1 Introduction The rhombencephalic vesicle or rhombencephalon is at the origin of the hindbrain and generates the pons, medulla oblongata, and most of the cerebellum. The pons and medulla, or lower brainstem, contain cranial nerve nuclei (Lumsden and Keynes, 1989), centers mediating arousal and sleep, centers regulating blood pressure and breathing (Bouvier et al., 2010), many components of the vestibular and auditory systems (Farago et al., 2006) and the majority of noradrenergic (locus coeruleus) (Wang et al., 2005) and serotonergic (raphe nuclei) neurons (Scott, 2005). The lower brainstem also contains precerebellar nuclei (PCN) which include the inferior olive (IO), lateral reticular nuclei (LRN), external cuneate nuclei (ECN), basilar pontine nucleus (PN), and reticulotegmental nucleus (Rodriguez and Dymecki, 2000). During early development, the hindbrain is segmented along the anteroposterior axis into a fixed number of rhombomeres (Kiecker and Lumsden, 2005). Each rhombomere contains clonally related neurons, and there is very little intermixing between neighboring segments. As a result, neurons within nuclei localized at similar anteroposterior location in the brainstem often have a common rhombomeric origin. However, some neurons defy rhombomeric confines and use a tangential mode of migration to settle outside their rhombomere of origin (see below). Here, we will review the molecular mechanisms regulating tangential migration of hindbrain neurons in two of the most studied systems: the facial branchiomotor (FBM) neurons and the PCN neurons. The genetic cascades of transcription factors which orchestrate the specification and differentiation of FBM and PCN neurons have been extensively studied, but will not be touched on here even though some of these transcription factors can also act at later developmental phases to control the responsiveness of migrating FBM and PCN neurons to several guidance cues as well as the expression of these cues (see chapter by Di Meglio and Rijli, this book). In this review, we will mostly focus on the various axon guidance molecules, receptors, and signaling pathways which control the tangential migration of hindbrain neurons.

Cellular Migration and Formation of Axons and Dendrites. https://doi.org/10.1016/B978-0-12-814407-7.00018-3 Copyright © 2020 Elsevier Inc. All rights reserved.

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18.2 Tangential migration: a historical overview In the developing nervous system, two main modes of neuronal migration coexist: radial migration for neurons that move perpendicular to the brain surface along radial glia processes and tangential migration for neurons that migrate parallel to the brain surface without following radial glia but using instead other cellular substrates as scaffolding such as neurons or axons (Métin et al., 2008). Although the sheer volume of systems that use tangential migration was neglected for many years, numerous studies in the past two decades have shown that a variety of forebrain cortical neurons migrate tangentially (De Carlos et al., 1996; O’Rourke et al., 1992): GABAergic interneurons (Anderson et al., 1997), olfactory bulb interneuron precursors (Jankovski and Sotelo, 1996; Lois et al., 1996), gonadotropin-releasing hormone neurons (Casoni et al., 2016; Schwanzel-Fukuda et al., 1996), striatal interneurons (Marín et al., 2001), and even projection neurons forming the globus pallidus (Nobrega-Pereira et al., 2010). Tangential migration has been studied more extensively in the hindbrain where its existence was first noted by Wilhelm His (His, 1891). His observed the existence of a broad germinative neuroepithelium which formed a prominent longitudinal bulge, the “Rautenlippe” or rhombic lip, bordering the dorsal edge of the fourth ventricle in the hindbrain. His proposed that neurons of the pontine nuclei and IO are generated in this dorsal neuroepithelium and then move ventrally to reach their final position. Almost 20 years later, in human embryos, Charles Essick (Essick, 1907) also observed this migratory stream and described it as “a path which corresponds in every detail to the fibro-nuclear mass which I described for the adult as the corpus ponto-bulbare.” During the 1950s, Harkmark (Harkmark, 1954) performed unilateral electrolytic lesions of the rhombic lip in chick embryos and obtained direct experimental evidence demonstrating that PCN neurons originate from the rhombic lip. In addition, Harkmark’s work also showed that IO neurons do not cross the ventral midline during their migration, thereby suggesting that only IO axons cross at the olivary commissure to project in the contralateral cerebellum. The development of radioautographic method and its ability to mark dividing neurons with tritiated thymidine (H3-Thy) provided a tool to label IO neurons and determine that they are born at embryonic day 12 (E12) to E13 in the rat (Altman and Bayer, 1978), E9 and E11 in the mouse (Pierce, 1973). This method also allowed to follow labeled neurons from their origin in the neuroepithelium to their final location, as well as to characterize the sequence, mode, and pattern of their migration. In rat embryo, Ellenberg and colleagues (Ellenberger et al., 1969) described two circumferential migratory streams, the marginal and submarginal streams, which they, erroneously, thought to both contain tangentially migrating IO neurons. Only many years later were Altman and Bayer (Altman and Bayer, 1987b), using that same H3-Thy technique, able to demonstrate that in fact IO neurons only follow the intramural circumferential pathway, which corresponds to the submarginal stream, whereas the marginal stream contains lateral reticular and external cuneatus neurons (Fig. 18.1) (Altman and Bayer, 1987a). The same conclusion was reached using 5-bromodeoxyuridine (BrdU) labeling (Bourrat and Sotelo, 1991), which in addition revealed that there is no a clear correlation between the birthdates of IO neurons and their final location within the IO, except in its caudal part where a perfect lateromedial temporal gradient is established. Using a similar strategy, the migratory pathways followed by LRN/ECN and PN neurons were also described (Fig. 18.1). These PCN neurons were shown to migrate just beneath the pial surface (marginal stream). PN neurons migrate anteriorly in the so-called anterior extramural stream (AES), whereas LRN/ECN neurons follow a caudal route along the posterior extramural stream. More direct support for tangential migration of PCN neurons was obtained using retrogradely HRP labeling in living brain slices (Bourrat and Sotelo, 1988, 1990) which in addition showed the morphology of the migrating neurons. Likewise, chick-quail chimeras demonstrated that cells originating from the rhombic lip contribute to PCN (Tan and Le Douarin, 1991). Finally, by using modern and sophisticated genetic fate-mapping techniques which enable the identification of the neuroproliferative zones where hindbrain neurons originate and their visualization by genetic markers (transcription factors), the pathway and mode of migration that these neurons use to reach their ultimate positions could be followed over time (Ray and Dymecki, 2009). In the hindbrain, tangential migration is not limited to PCN neurons. Three types of cerebellar neurons derived from the rhombic lip also migrate tangentially: the granule cells, the projection neurons of the deep nuclei, and the unipolar brush cells (Fink et al., 2006; Gilthorpe et al., 2002). Moreover, oculomotor neurons cross the midline by migrating tangentially as suggested by in vitro data from the 1970s (Puelles-Lobez et al., 1975; Puelles and Privat, 1977). This is also the case for other hindbrain neurons in chick and cartilaginous fish (Book and Morest, 1990; Fritzsch et al., 1993). Likewise, vestibulo-acoustic efferent neurons also migrate tangentially across rhombomere 4 (r4) midline (Bruce et al., 1997; Simon and Lumsden, 1993). Finally, branchiomotor neurons of the facial nucleus (FBM neurons) do not cross the midline but migrate tangentially and caudally across two rhombomeres, before migrating radially to their final location (Fig. 18.2). The first indications that FBM neurons migrate tangentially in rodent embryos were obtained in the 1980s using H3-Thy labeling studies (Altman and Bayer, 1980). Moreover, in Hoxb-1 knockout mice, r4 identity is changed and

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(B) Dorsal

Lateral Cer

RL FP

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FIGURE 18.1 Schematic representation of the location and pathways followed by tangentially migrating inferior olivary neurons and pontine neurons in the embryonic hindbrain at E13eE14. All originate from the rhombic lip (RL, in purple). (A) Pontine neurons (PN) migrate rostrally until the r2/r3 border (arrowhead) where they turn ventrally toward the floor plate (in gray). Inferior olivary neurons (IO) migrate at a more caudal position. (B) Cross section at the level IO. IO neurons migrate ventrally toward the floor plate. Their axon/leading process crosses it but not their cell bodies.

(A)

E11

E12.5

FBM

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FVM r5 VEGF-A Wnt5a

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FIGURE 18.2 Schematic representation of the location and pathways followed by tangentially migrating branchiomotor neurons of the facial nucleus (FBM, blue) in the embryonic mouse hindbrain. (A) Ventral views and (B) cross sections at similar ages but distinct rostrocaudal positions along r4er6. Left (E11), FBM neurons are born in r4 more medially than visceromotor neurons of the facial nucleus (FVM, green). Both FBM and FVM neurons project axons laterally that exit the brainstem in r4. VEGF and Wnt5 are expressed caudally to r4 and were proposed to attract FBM neurons away from r4. Middle (E12.5), FVM migrate radially and laterally in r5, whereas FBM migrate tangentially and caudally from r4 to r6. FBM also migrate laterally. Right (E15), in a last step, FBM neurons migrate radially from dorsal to ventral in r6.

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FBM neurons migrate laterally within r4 and fail to form a facial nerve (Goddard et al., 1996; Studer et al., 1996), clearly demonstrating that FBM neurons originates from r4 and migrate caudally to r5 and r6 (Altman and Bayer, 1980).

18.3 Molecular mechanisms controlling the tangential migration of precerebellar neurons 18.3.1 Influence of the midline on tangentially migrating precerebellar neurons Most animal species are bilateria with a central nervous system symmetrically organized along the midline that extends throughout the anteroposterior axis. In vertebrates, specialized cells, differentiating during neural tube closure, delineate the dorsal midline, or roof plate, and the ventral midline, or floor plate (Placzek and Briscoe, 2005). At all hindbrain levels, like in the spinal cord and forebrain, there are subpopulations of neurons projecting axons across the midline and other neurons whose axons never cross the midline (Chédotal, 2014). Tangentially migrating hindbrain neurons exhibit distinct behavior when reaching the floor plate: some neurons will cross it, whereas others will stop. Even though the study of Ramón y Cajal (Ramón y Cajal, 1890) on the development of commissural neurons in the chick spinal cord strongly suggested that floor plate cells secrete chemotropic diffusible molecules that attract axons toward the floor plate, it was only in the 1980s, with the development of collagen gel cultures, that direct evidence supporting the chemotactic hypothesis of Cajal was obtained (Lumsden and Davies, 1986). A few years later, it was proposed that a protein called netrin1 can promote the attraction of axons toward the floor plate (Kennedy et al., 1994; Serafini et al., 1994). In parallel, genetic screens in Drosophila and Caenorhabditis elegans led to the characterization of other secreted and membrane-bound proteins, such as slits, semaphorins, or ephrins (Drescher et al., 1995; Kolodkin et al., 1993; Rothberg et al., 1990), which influence midline crossing by developing axons. With a few exceptions, these proteins and their receptors are evolutionarily conserved and found in most phyla (Friocourt et al., 2017; Patthey et al., 2017). We will briefly summarize the current working model explaining how midline crossing is regulated in vertebrates, but restricting it to data relevant for this chapter (Fig. 18.3), further information can be found in other reviews (Chédotal, 2011; Pignata et al., 2016) or other chapters in this book. Classically, it was thought that commissural axons first grow toward the midline in response to the binding of Netrin1 to the receptor deleted in colorectal cancer receptor (DCC), expressed on their growth cone (Fazeli et al., 1997; Serafini et al., 1996). However, it was recently shown, using conditional knockouts, that commissural axons still extend toward the ventral midline in absence of netrin1 at the floor plate and that neural precursors in the ventricular zone (VZ) constitute an alternative and complementary source of netrin1 (Dominici et al., 2017; MorenoBravo et al., 2019; Varadarajan et al., 2017). Moreover, netrin1 seems to act primarily by promoting haptotaxis rather than long-range chemoattraction (Dominici et al., 2017; Moore et al., 2009). In the spinal cord, the morphogen sonic hedgehog (Shh) cooperates with netrin1 to attract axons as they approach the ventral midline (Charron et al., 2003; Okada et al., 2006; Wu et al., 2019), but if this is also the case in the brainstem has not been assessed. Upon reaching the midline, there is evidence suggesting that axons gain responsiveness to Slit repellents (Slit1eSlit3) that activate Robo1 and Robo2, two receptors of the roundabout family (Blockus and Chédotal, 2016; Evans and Bashaw, 2010). This steers axons away from the floor plate to the contralateral side of the brain. In precrossing axons, Slit repulsion was thought to be kept inactive by the third Robo receptor, Robo3, counteracting Robo1/Robo2 activation (Sabatier et al., 2004). However, there is little direct support for this model and more recent data suggest that Robo3 is a divergent Robo receptor that promotes rather than inhibits commissural axon extension to the midline (Zelina et al., 2014). In addition, in vitro data suggest that Slit binding to Robo1 (and possibly Robo2) triggers its direct interaction with DCC that in turn could silences netrin1 signaling (Stein and Tessier-Lavigne, 2001). Whether this mechanisms is at play in vivo remains to be established. Therefore, midline crossing by commissural axons is a two-step process, starting with the activation of attractive/growth-promoting signals and inactivation of repulsive signals, followed by the subsequent activation of repulsive mechanisms and the inactivation of attractive ones. Interestingly, several floor plateederived axon guidance molecules and their receptors play a key role in patterning the tangential migration of hindbrain neurons along the anteroposterior and dorsoventral axes. Pontine neurons following the AES, first migrate rostrally from the rhombic lip of r6er8 (Farago et al., 2006) across r5 and r4 (Cambronero and Puelles, 2000), along a route parallel to the floor plate. Upon reaching the r2/r3 boundary at the level of the root of the trigeminal nerve, they turn 90 degrees ventrally and progress toward the floor plate (Geisen et al., 2008; Kratochwil et al., 2017; Zelina et al., 2014). Conversely, all the other precerebellar neurons migrate directly away from the rhombic lip toward the floor plate. Explant cultures suggest that floor plate releases attractant(s) for IO and LRN neurons (de Diego et al., 2002; Taniguchi et al., 2002) and that floor plate attraction of precerebellar neurons is mediated by netrin1 and its receptor DCC.

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FIGURE 18.3 Regulation of precerebellar migration at the floor plate. Schematic drawings showing cross sections of the mouse embryonic hindbrain at the level of the inferior olive. (A), precerebellar neurons extend leading processes and migrate tangentially toward the floor plate. They express the DCC and Robo3 receptors. Netrin-1 (green) is expressed by both floor plate (FP) and ventricular zone (VZ) progenitors and their radial processes extending to the pial surface (Pia). Slits (red) are also present at the floor plate. (B), In wild-type mice, inferior olivary (IO) neurons migrate tangentially (arrowheads) from the rhombic lip (RL) dorsally toward the ventral midline (yellow). Their axons cross the midline but their cell bodies stop before reaching the midline. (C), In Robo1/Robo2 double knockouts and in Slit1/Slit2 double knockouts, IO neurons are still able to send axons that cross the midline, but their somata migrate from the rhombic lip and some of them can abnormally pass across the midline. (D), By contrast, the IO neurons migrate normally in conditional knockouts lacking all floor plate Slits or Robo1/Robo2 in IO neurons indicating that the migration defects seen in null-knockouts are non-cell-autonomous. (E), IO neurons do not cross the midline, and their axons project ipsilaterally into the cerebellum in Robo3 knockout embryos. Midline crossing is not restored for IO axons in Robo1/2/3 triple knockouts, suggesting that in this system, Robo3 does not modulate Slit/Robo-dependent repulsion. (F), Similar defects are seen in Robo3 conditional knockouts selectively lacking Robo3 in IO neurons indicating that Robo3 acts cell-autonomously. (G), In netrin1 full KO, the vast majority of IO neurons project ipsilaterally and their cell bodies migrate ventrally. Some commissural axons also exit the CNS. This phenotype is recapitulated when netrin1 is ablated from the ventricular zone (DVZ). (H), By contrast, IO neuron migration is normal in mutant mice lacking floor plateederived netrin1. (I), In Dcc knockout the migration of IO neurons is perturbed and not all can reach the midline remaining scattered all along the submarginal stream. IO neurons project ipsilaterally and contralaterally.

Netrin1 is strongly expressed in the floor plate throughout the hindbrain and all migrating precerebellar neurons express its receptors DCC, whereas the other netrin1 receptors, Unc5a-c and Neogenin, are expressed in IO neurons within the rhombic lip but not detected during their migration (Alcántara et al., 2000; Bloch-Gallego et al., 1999; de Diego et al., 2002). Migrating pontine neurons express Unc5c and a subset also express Unc5b (Kim and Ackerman, 2011; Di Meglio et al., 2013). An attractive activity of Netrin1 on all types of migrating PCN neurons was demonstrated using hindbrain explants and collagen gel assay (Alcántara et al., 2000; Causeret et al., 2002, 2004; de Diego et al., 2002; Marcos et al., 2009; Taniguchi et al., 2002; Yee et al., 1999; Zelina et al., 2014). In such assays, Netrin1 attraction is inhibited by addition of anti-DCC antibodies (Marcos et al., 2009; Taniguchi et al., 2002; Yee et al., 1999) and does not occur when using PCN from Dcc knockout mice (Marcos et al., 2009). The phenotypic analysis of the development of PCN nuclei in netrin1 and Dcc knockout embryos, also supported that this ligand/receptor complex guides PCN neurons toward the midline (Fig. 18.3). In netrin1 and Dcc knockout mice, all PCN neurons leave the rhombic lip but their migration toward the floor plate is affected. In these mutant mice, LRN/ECN and IO neurons are still able to migrate

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ventrally, but their migration is perturbed (Bloch-Gallego et al., 1999). The size of the LRN is significantly reduced (Marcos et al., 2009). In both DCC and netrin1 KOs, the IO nucleus is fragmented with small subsets of neurons scattered along the distal two-thirds of the submarginal migratory stream, deeper in the hindbrain parenchyma, and only a subset of IO neurons is able to reach the midline. These and the ectopic IO neurons still project to the cerebellum but on the ipsilateral side. This ipsilateral projection is not due to a migration of IO neurons across the midline (Marcos et al., 2009). In DCC and netrin1 KOs, PN neurons maintain a dorsal migratory trajectory (Yee et al., 1999; Zelina et al., 2014) explaining the absence of pontine nucleus (Fazeli et al., 1997; Serafini et al., 1996). Moreover, PN neurons also migrate abnormally in mice lacking the polycomb protein Ezh2 in which netrin1 expression extends more dorsally than in wildtype embryos (Di Meglio et al., 2013). Although these data suggest that a gradient of netrin1 attracts precerebellar neurons to the floor plate, other studies challenge this conclusion. PCN migration defects were observed in netrin1 full knockouts, but it was found that their migration is normal in conditional mutants selectively lacking floor plateederived netrin1. First, the main source of netrin1 for PCNs is not the floor plate but VZ precursors which appear to secrete netrin1 at the pial surface and promote the growth of the leading process of PCNs close to basal lamina (Dominici et al., 2017). It was also found that the preference of PCN for netrin1 constrains their migration to the CNS. In netrin1 knockouts, PN migration is randomized and a significant fraction escape the CNS through the trigeminal and vestibular nerve roots (Moreno-Bravo et al., 2018; Yung et al., 2018) (Fig. 18.3). Interestingly, the PN migratory stream contains different populations of neurons originating from different rhombomeres and expressing distinct levels of Unc5b receptors in addition to Unc5c. In both Unc5c and Unc5b knockouts, subsets of PN neurons migrate prematurely to the midline (Kim and Ackerman, 2011; Di Meglio et al., 2013). A few studies have also tried to characterize the signaling pathways downstream of DCC activated by Netrin1 in migrating PCN neurons. Not surprisingly, the identified downstream effectors are known to act on the cytoskeleton which is essential to neuronal migration. Two distinct pathways have been studied, one involving the Rho family of small GTP-binding proteins (Rho GTPases) and the second the microtubule associated protein MAP1B. Rho GTPases are major modulators of the actin cytoskeleton involved in axonal growth and cell migration. In several cell lines, netrin1 binding to DCC activates the Rho GTPases Cdc42 and Rac1 (Li et al., 2002). Cdc42, Rac1, RhoA, and RhoB are expressed by migrating PCN neurons (Causeret et al., 2004). In hindbrain explants, the pharmacological inhibition of Rac1 and Cdc42 perturb the extension of the leading process of IO/LRN neurons induced by Netrin1, while blocking RhoA/B or its effectors, Rock kinases, inhibits nucleokinesis (Causeret et al., 2004). Rock kinases are Serine-threonine kinases playing a role in actomyosin assembly (Causeret et al., 2004). It has not yet been shown that these molecules control PCN migration in vivo. The second pathway that seems to operate downstream of netrin1/DCC in PCN involves MAP1B. In cortical neuron cultures, netrin1 induces/stimulates MAP1B phosphorylation. This effect is dependent on two kinases, glycogen synthase kinase 3 (GSK3) and cyclin-dependent kinase 5 (CdK5) (Xie et al., 2006), which are activated by netrin1. PN neurons from MAP1B KO fail to be attracted by Netrin1 (Del Rıo et al., 2004). Accordingly, MAP1B phosphorylation is reduced in Netrin1 KO, and the development of IO and PN neurons is altered in MAP1B KO (Del Rıo et al., 2004) or in MAP1B: MAP2 double KO (Teng et al., 2001). Last, the IO appears absent in Cdk5 KO (Ko et al., 2001), but the PN still forms. As in the spinal cord and other commissural systems, Slit proteins and their receptor Robos also influence the tangential migration of PCN neurons (Fig. 18.3). All Slits are expressed by floor plate cells (Causeret et al., 2002; Dominici et al., 2018; Marillat et al., 2002) during PCN migration. In addition, Slit2 and Slit3 expressed by motor neurons of the facial nucleus (Geisen et al., 2008) and in the rhombic lip represent other potential sources of repulsive forces for PCN neurons. PCN neurons express the three Robo receptors (Robo1eRobo3) during their migration (Marillat et al., 2004; Di Meglio et al., 2008). Robo3 is only expressed until PCN neurons either reach the midline (IO, PN) or cross it (LRN/ECN). There is also no clear evidence of Robo1 or Robo2 upregulation in PCN neurons after midline crossing (Tamada et al., 2008). As for netrin1, in vitro assays and organotypic cultures showed that Slits can repel migrating neurons from the PN (Gilthorpe et al., 2002), IO, and LRN/ECN (Causeret et al., 2004; Tamada et al., 2008). The analysis of Slit and Robo full knockouts brought in vivo support to these findings, but also revealed some differences in the manner in which Slit/Robo affect the different types of PCN. In Slit1/Slit2 and Robo1/Robo2 double knockouts, a significant fraction of IO neurons freely cross the floor plate to settle in the contralateral side (Fig. 18.3), supporting a classic midline gate keeper function of Slit/Robo and suggesting that the floor plate provides a stop signal to migrating IO neurons (de Diego et al., 2002). However, a conditional knockout strategy showed that IO neurons still stop at the midline in absence of all Slits at the floor plate or when Robo1/2 receptors are selectively silenced in IO neurons (Dominici et al., 2018). These results, show that the migratory defects reported in the full knockouts are non-cell-autonomous and that Slit/Robo repulsion does not play a direct role in the arrest of IO neuron at the midline. Hence, the molecular mechanisms which stop IO neurons at the midline are still unknown. Ephrin/Eph repulsion could be involved as in EphA4 receptor knockouts some ION abnormally cross the floor plate which express its ligand ephrinB3 (Hashimoto et al., 2011).

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The case of LRN/ECN neurons is more complex as they all migrate across the floor plate in wild type. So far no defects in the migration of these neurons were detected in Slit1/Slit2 or Robo1/Robo2 double knockouts, and they still cross the floor plate and move away from it. As described in the spinal cord (Long et al., 2004), it may be necessary to simultaneously remove the three slits to affect the migration of LRN/ECN neurons. The case of the PN is also puzzling as migration is perturbed well before reaching the midline in Robo1/Robo2 and Slit1/Slit2 double mutants. In these mice, cohorts/chains of pontine neurons fail to initiate rostral migration and immediately progress toward the floor plate (Geisen et al., 2008). In Hoxa2 knockout, where Slit2 expression in the facial nucleus and Robo1/2 expression in pontine neurons are downregulated (Geisen et al., 2008), many pontine neurons migrate too early toward the floor plate. Altogether these results suggested that Slits might repel pontine neurons from the rhombic lip and force them to migrate above the facial nucleus before being allowed to respond to floor plate attraction. However, neither the conditional ablation of Slits from the facial nucleus nor the selective silencing of Robo1/Robo2 in pontine neurons perturb their migration, suggesting that as for the ION, Slit/Robo signaling might not directly control the tangential migration of pontine neurons (Dominici et al., 2018). The third Robo receptor, Robo3 (Friocourt and Chédotal, 2017; Sabatier et al., 2004) emerged as a major regulator of PCN migration. Robo3 is expressed by all migrating PCN until their leading process crosses the floor plate and the migration of all PCN neurons is perturbed in Robo3 null mice in a fully penetrant manner. Midline crossing of LRN/ECN neurons is entirely prevented, and they form a nucleus on the ipsilateral side of the hindbrain. IO neurons still migrate to the floor plate and aggregate in a clear nucleus, albeit disorganized, but their axons do not cross and project into the ipsilateral cerebellum. This shows that tangential migration is not blocked in the absence of Robo3 but that the leading process of IO axons does not cross the midline and gives rise to an axon on the wrong side of the brain. Last, PN neurons are unable to migrate ventrally and continue rostrally when reaching the r2/r3 border and form two ectopic masses in the dorsolateral side of the medulla oblongata (Marillat et al., 2004; Zelina et al., 2014) Interestingly, human patients suffering from a rare neurological disease named horizontal gaze with progressive scoliosis (HGPPS) carry mutations in the ROBO3 gene (Friocourt and Chédotal, 2017). High-resolution magnetic resonance (MR) imaging of the brain showed that they have a hypoplastic butterfly-shaped pons and an enlarged fourth ventricle. This suggests that in HGPPS patients, pontine neurons either do not migrate properly to the midline and remain ectopic or they may even partially degenerate (Jen et al., 2004). What is the mechanism of action of Robo3? First, IO neurons also project ipsilaterally in Robo3 conditional knockout (Robo3loxlox) that were crossed to mice expressing Cre recombinase under the Ptf1a promoter (Ptf1a::cre) (Badura et al., 2013; Renier et al., 2010). In these mice, Cre is highly expressed in IO neurons and not in other PCN neurons. This demonstrates that Robo3 acts cell-autonomously in IO neurons. This was confirmed for PN neurons using other Cre driver lines (Zelina et al., 2014). The analysis of the Robo3 null mice shows that the projection of PCN neurons and that of the other hindbrain and spinal cord commissural neurons is ipsilateral (Marillat et al., 2004; Sabatier et al., 2004). This initially suggested that, as for spinal cord commissural axons, Robo3 is required in PCN to counteract Slit/ Robo repulsion and to allow netrin1 attraction to occur. In Robo3 KO, Slit repulsion mediated by Robo1 and Robo2 would unavoidably be activated before midline crossing, thus forcing the axons and leading processes of PCN neurons to stay ipsilaterally. Accordingly, in Robo1/Robo2/Robo3 triple KO, in which repulsive Robo1 and Robo2 are also absent, midline crossing is partially rescued (Fig. 18.3) (Di Meglio et al., 2008). However, more recent studies suggest that Robo3 is a DCC coreceptor promoting the migration of PN neurons toward the floor plate and mediating netrin1 function (Zelina et al., 2014). Likewise, there is also genetic evidence supporting an attractive role of Robo3 during ION migration (Di Meglio et al., 2008). What could be the signaling pathways downstream of Robo receptors in migrating PCN neurons? This has not yet been studied, but Rho GTPases could be involved as they are highly expressed in migrating PCN neurons and control their migration in vitro (see above). Rho GTPases are activated upon GTP binding and inactive when bound to GDP. The switch from their active to their inactive state is controlled by two families of proteins: the guanine nucleotide exchange factors (GEFs) and the GTPase activating proteins (GAPs). GEFs activate Rho GTPAses, while GAPs inactivate them by inducing GTP hydrolysis. Many studies have shown that Rho GTPases play an important role in the modulation of Slit function. For instance, in vertebrate neurons, a constitutively active Cdc42 blocks the repulsive effect of Slit (Wong et al., 2001). Robo also controls the activity of Rho GTPases through a family of Slit-/Robo-specific GAPs, SrGAP1-3 (Endris et al., 2002). Therefore these studies suggest that Rho GTPases could be common effectors of Netrin/DCC and Slit/Robo signaling in PCN neurons. Another study showed that Src-kinases are also acting downstream of Robo3 (Zelina et al., 2014). Although these data indicate that netrin1/DCC and Slit/Robo influence PCN migration, other molecules could be involved. In netrin1, DCC, Slit, and Robo KO, IO and LRN neurons still migrate away from the rhombic lip, suggesting that they are able to respond to repellents from the dorsal hindbrain that could play a role similar to BMPs (Butler and Dodd, 2003) and Draxin (Islam et al., 2009) in the spinal cord. However, PCN migration was reported to be unaffected in

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Draxin knockout mice (Asrafuzzaman Riyadh et al., 2014). Alternatively, another and nonexclusive explanation could be the existence of additional floor plate attractants for PCN neurons, such as is the case for Shh that cooperates with Netrin1 to guide spinal cord commissural axons toward the midline (Charron et al., 2003; Wu et al., 2019). There are also results suggesting that after crossing, PCN neurons and axons become unresponsive to floor plate and become attracted by unknown factors from the alar plate and cerebellum (Taniguchi et al., 2002; Zhu et al., 2003). In the spinal cord, the floor plate secretes semaphorins and cell adhesion molecules (CAMs), which are also involved in the control of midline crossing. For instance, the gain in responsiveness of commissural axons to semaphorin 3b repulsion after midline crossing (Nawabi et al., 2010) is stimulated by a soluble form of the Neural cell adhesion molecule (NrCAM). Although precerebellar neurons express at least some components of the receptor complex for semaphorins, such as Plexind1, Plexina2 (Gesemann et al., 2001; Vilz et al., 2005), and neuropilin2 (Chen et al., 1997) and also several CAMs (Backer et al., 2002) the function of these molecules in the control of neuronal migration is unclear. For instance, NrCAM is expressed by migrating IO neurons (not by LRN/ECN and PN) but the IO is normal in NrCAM KO (Backer et al., 2002). Transient axonal glycoprotein 1 (TAG-1), also known as Contactin 2, is expressed by migrating PN and LRN/ECN neurons but is absent from IO neurons (de Diego et al., 2002; Horie et al., 2003; Kyriakopoulou et al., 2002; Wolfer et al., 1994; Yee et al., 1999). TAG-1 can mediate homophilic or heterophilic interactions and binds to other CAMs such as L1/ NgCAM, NrCAM, and NCAM. The migration of LRN/ECN neurons is perturbed in TAG-1 knockout mice (Denaxa et al., 2005) and in hindbrain explants treated with anti-TAG-1 blocking antibodies (Kyriakopoulou et al., 2002) suggesting that the protein could mediate homophilic interaction between migrating LRN/ECN neurons. However, TAG-1 has also been shown to modulate the response of sensory axons to Sema3A repulsion (Law et al., 2008), and this could also be the case in PCN neurons. Migrating PN and ECN neurons also strongly express another CAM named MDGA1 (Litwack et al., 2004) of unknown function.

18.3.2 Why do PCN neurons migrate near the pial surface? For years, the integrity of the subpial basement membrane has been considered essential for a normal migration in cortical structures, especially the cerebral cortex (Lambert de Rouvroit and Goffinet, 2001) and cerebellum (Sievers et al., 1994). Moreover, meninges were thought to constitute a mechanical barrier to migration which would prevent migrating cells from leaving the nervous parenchyma. More recently, it was revealed that meninges secrete molecules which create permissive or nonpermissive environments to migrating neurons (Lu et al., 2002) such as PCN neurons. One of the characteristic features of migrating PCN neurons is that they follow a superficial route right underneath the pial surface (LRN/ECN and PN) or at a short distance (IO) from it. Electron microscopy studies showed that PN and LRN neurons migrate in close contact with radial glia endfeet attached to the subpial basal membrane (Ono and Kawamura, 1989, 1990). Two main mechanisms may explain the preferential subpial migration PN and LRN/ECN neuron: adhesion to radial glia endfeet and/or attraction by meningeal cell-derived factors. Alpha-dystroglycan (a-DG) is a transmembrane protein linking the extracellular matrix to the actin cytoskeleton (Yurchenco and Wadsworth, 2004). a-DG is localized at the endfeet of radial glia, and it is also expressed by PN neurons (Ohtsuka-Tsurumi et al., 2004) suggesting that it may regulate the interaction between PN neurons and the basal lamina. Several enzymes such as Large, POMT1/POMT2, POMGnT1, and fukutin act to modify and glycosylate a-DG; they are often mutated in human patients with congenital muscular dystrophy. Interestingly, many of these patients also present pontine heterotopia (Saito et al., 2003) and, in addition, the pontine nucleus is hypoplastic and ectopic in Large mutant mice (Litwack et al., 2006; Qu and Smith, 2005). By contrast, the IO and the LRN/ECN nuclei develop normally in Large mutants (Litwack et al., 2006). In these mutants, ectopic PN neurons are found in regions where the radial glia is not properly anchored and the glia limitans is interrupted (Qu and Smith, 2005), suggesting that the migration defects are secondary to radial glia disorganization. However, a-DG binds laminin and is thought to control Netrin1 binding and distribution in the extracellular matrix (Yurchenco and Wadsworth, 2004). a-DG also interacts with several ECM proteins containing a so-called Laminin G module that is found in Slits (Yurchenco and Wadsworth, 2004). Therefore, the abnormal glycosylation of a-DG in Large mutants could alter the distribution of axon guidance molecules such as Netrin1 and Slits and thereby perturb the migration of PN neurons. Accordingly, in Large mutants, as in Robo3 knockouts, PN neurons fail to turn ventrally and project ipsilaterally (Qu and Smith, 2005). It is not known if PN neurons migrate normally in a-DG KO (Satz et al., 2008). CXCL12 (also known as Stromal cellederived factor-1, SDF-1) is a chemokine expressed by meninges, which attracts cerebellar granule cell progenitors in the cerebellum and CajaleRetzius cells in the neocortex. Meninges overlying the AES express CXCL12 and migrating PN neurons express its receptor, CXCR4 (Vilz et al., 2005; Zhu et al., 2009). In collagen gel assay, CXCL12 expressing cells attract neurons migrating from rhombic lip explants and PN neurons migrate

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deeper in hindbrain explants deprived from meninges (Zhu et al., 2009). The attractive activity of CXCL12 on PN neurons has been confirmed in vivo as in Cxcr4 and Cxcl12 knockouts, ectopic PN neurons are found inside the brainstem (Vilz et al., 2005) and migrate toward the midline and more posteriorly in several streams (Zhu et al., 2009). CXCL12 attractive activity is again specific for PN neurons as the LRN/ECN and IO neurons migrate normally in absence of CXCL12/ CXCR4. The CXCR4 receptor was shown to interact with myosin II, which is a major component of the migration machinery (He et al., 2010; Rey et al., 2002; Vicente-Manzanares et al., 2002). Interestingly, PN neurons fail to migrate ventrally in mice carrying mutation in the myosin-IIB heavy chain (Ma et al., 2004; Solecki, 2012), indicating that in PCN as in other neurons, myosin II motors are essential for tangential migration. Retinoic acid is another meningeal derived factor influencing the migration of PCN neurons. Developing PN and IO nuclei were found to be very sensitive to the teratogenic action of retinoic acid (RA) (Yamamoto et al., 2003), the metabolic product of vitamin A (retinol). RA activates gene transcription and plays a role in CNS patterning, in part by controlling hox gene expression pattern in the hindbrain. Using an RA reporter mouse, it was shown that RA signaling is active in migrating IO and PN (Yamamoto et al., 2003). RA synthetizing enzyme retinaldehyde dehydrogenase-2 (RALDH2) is enriched in the meninges (Niederreither et al., 1997; Zhang et al., 2003) surrounding the hindbrain, while RA nuclear receptors such as CRABPI (cellular RA binding protein I (Zhang et al., 2003) (Dollé et al., 1990) and downstream effector are expressed at high level in all migrating PCN neurons (Yamamoto et al., 2003). PN and IO fail to form after in utero exposure to high doses of RA. Interestingly, in these RA-treated mice, PN neurons do not follow the AES but migrate in small chains toward the midline a phenotype somehow reminiscent of the PN migration defects in HoxA2 and Slit/Robo KO; this suggests that RA could act upstream of these molecules in migrating PN neurons. Likewise, in RA-treated mice, PN neurons fail to express Pax6 which is also known to control the migration of PN neurons (Engelkamp et al., 1999; Horie et al., 2003). Despite these recent developments, it is still unknown why IO neurons migrate more deeply in the hindbrain than mossy fiber projection neurons and their migration substrate remains elusive. The reason for which the molecular mechanisms controlling the superficial position of migrating PN neurons differ from those of LRN/ECN neurons is also unclear especially since, to a large extent, the two types of neurons seem to express many common receptors and transcription factors. However, recent studies suggest that some transcription factors, such as NfIb, are expressed only by one type of mossy fiber projection neurons (Zhu et al., 2009). Likewise, the RNA-binding protein Musashi 1 controls the translation of Robo3 in IO neurons but not in PN neurons (Kuwako et al., 2010). Interestingly, recent studies showed that the PN neurons might be confined to the CNS by netrin1 which accumulates at the pial surface and is absent from the PNS. In netrin1 knockout embryos, a large fraction of PN neurons escape the main migratory stream to invade the trigeminal and auditory nerves in the PNS (Moreno-Bravo et al., 2018; Yung et al., 2018).

18.4 Molecular mechanisms controlling the tangential migration of facial motor neurons 18.4.1 Origin and migration of facial motor neurons Cranial motor neurons can be classified into three types: somatomotor, visceromotor, and branchiomotor (Guthrie, 2007). Somatomotor neurons (basal plate derivative) belong to the motor nuclei of the III, IV, VI, and XII cranial pairs and provide innervation to the muscles derived from the somatic mesoderm. They occupy a bilateral column on both sides of the floor plate and close to the ventricle. Visceromotor neurons belong to the dorsal nucleus of the vagus (X), the salivary inferior nucleus (IX), the facial nucleus (FVM), and the accessory nucleus of the III. They all innervate parasympathetic ganglia and are located close to the ventricle more laterally than somatomotor neurons) and close to the limit between the basal and alar plates. Branchiomotor neurons belong to the nucleus ambiguous, the trigeminal nucleus (V), and the facial nucleus (FBM). They are disposed ventrolaterally and provide innervation to the striated muscles derived from the mesoderm of the branchial arches. The migration of facial motor neurons has been mostly studied in three vertebrate model systems: the zebrafish, the chick, and the mouse. As stated above, facial motor neurons innervate either branchial muscles (FBM neurons) derived from the second pharyngeal arch or parasympathetic ganglia (FVM neurons). These two different classes of facial motor neurons originate from different rhombomeres: FVM neurons develop exclusively from r5 (Auclair et al., 1996; Guthrie, 2007; Jacob and Guthrie, 2000), whereas FBM neurons have origins and locations that vary between species. In the mouse and zebrafish, all FBM neurons are generated in r4 but migrate caudally, to r6 or r7, respectively (Chandrasekhar, 2004; Jacob and Guthrie, 2000; Wanner et al., 2013). In the chick, most FBM neurons are also generated in r4, and the others in r5, but caudal migration does not occur (Fig. 18.2). The vast majority of motor neurons in the vertebrate hindbrain follow

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relatively simple migratory pathways, moving in a ventral to dorsal direction within the same rhombomere they are born (Kiecker and Lumsden, 2005; Lumsden and Keynes, 1989). By contrast, FBM neurons migrate and follow complex routes which combine different modes of migration. The migration of FBM neurons can be subdivided into three successive phases (Chandrasekhar, 2004; Chandrasekhar et al., 1997; Garel et al., 2000; Song et al., 2006). In mouse embryos, r4-derived FBM neurons first migrate tangentially, parallel to the floor plate and close to the ventral midline (between E10 and E12.5), following a caudal direction through r5 to reach r6, while projecting their axons rostrally through an exit point located in dorsal r4 (Chandrasekhar, 2004; Guthrie, 2007; Song, 2007; Studer et al., 1996). This caudal migration is not observed in chick where FBM neurons remain within r4. Chick r5 and r6 have been considered to lack specific attractive extrinsic cues required to initiate FBM neurons caudal migration (Studer, 2001). Time-lapse and genetic studies in zebrafish showed that during this first phase, late-born FBM neurons migrate along the axons of pioneer FBM neurons and also at older stages along axons of the medial longitudinal fasciculus (MLF) (Wanner et al., 2013). In a second phase, described in mouse and zebrafish, FBM neurons undertake, upon reaching r6, a mediolateral migration away from the basal plate to reach their dorsolateral position. During these first two phases (caudal and lateral), FBM neurons adopt a tangential mode of migration, and their collective migration involves the cell-adhesion molecule Cadherin-2 (the fish N-cadherin) (Rebman et al., 2016; Stockinger et al., 2011). Cadherin-2 also regulates the axonophilic migration of FBM neurons on MLF axons (Wanner and Prince, 2013). In a third and last phase, FBM neurons migrate radially along the radial glia scaffold to reach their final destination near the pial surface in ventral r6er7, where they condense/cluster on both sides of the midline in two symmetrical nuclei. Overall, FBM migration lasts about 4 days in the mouse and is completed by E14 (Fig. 18.2). The phenotypic analysis of a large variety of genetically modified mice and zebrafish has led to the identification of many molecules regulating each phase of FBM migratory process. Although most studies have focused on the first phase of FBM migration, they also showed that the molecular mechanisms controlling the tangential migration (from r4 to r6) of FBM neurons differ from those controlling their radial migration.

18.4.2 The caudal migration of FBM neurons 18.4.2.1 The planar cell polarity pathway Planar cell polarity (PCP) genes encode proteins which control the establishment of epithelial cell polarity in many species and tissues. In vertebrates, major components of the PCP pathways (Fig. 18.4) are transmembrane proteins such as the Wnt receptor Frizzled (Fz), van gogh-like 2 (vangl2), the atypical Cadherins Celsr1-3, and their cytoplasmic effectors such as Prickle1, the scaffold protein Scribbled1 (Scrb1), and Disheveled (Dvl). In developing epithelia, PCP proteins are often asymmetrically distributed at the cell surface (Wallingford, 2010). Many studies in zebrafish and mouse have shown that components of the PCP pathways are all broadly expressed in the hindbrain during FBM migration both in migrating neurons and along their pathway. In these species, the analysis of multiple mutants of knockouts affecting PCP genes has demonstrated that PCP signaling regulates the direction of migration of FBM neurons. Therefore, in mice and fish, the inactivation of prickle1, scrb1, vangl1, vangl2, fz3, celsr1-3 disrupts the caudal migration of FBM neurons which are unable to reach r6 and either stay in r4 or migrate at ectopic locations although their differentiation and proliferation do not seem to be affected (Bingham et al., 2002; Carreira-Barbosa et al., 2003; Chandrasekhar, 2004; Glasco et al., 2016; Jessen and Solnica-Krezel, 2004; Jessen et al., 2002; Park and Moon, 2002; Qu et al., 2010; Rohrschneider et al., 2007; Vivancos et al., 2009; Wada et al., 2005, 2006; Wang and Nathans, 2007). Interestingly, in Celsr1-deficient mice, many FBM neurons follow a rostral as well as lateral direction into r2 and r3 instead of moving caudally (Qu et al., 2010). The use of Celsr1 conditional knockouts suggests that Celsr1 acts non-cell-autonomously and that its expression in the ventricular zone of r2-r5 represses the attractive activity of anterior rhombomeres for FBM neurons (Glasco et al., 2016). This masked attractive signal could involve Wnts, which are secreted glycoproteins which bind to Fz receptors. During FBM migration, Wnt5a is expressed in a high-caudal to low-rostral gradient in the rhombencephalon (Fig. 18.4) (Vivancos et al., 2009). In hindbrain explants, Wnt5a and Wnt7a, soaked beads can attract FBM neurons, whereas inhibitors of Wnt signaling block their caudal migration (Glasco et al., 2016; Vivancos et al., 2009). The migration of FBM neurons is also perturbed in Wnt5a knockout. These data suggest that Wnt5a could be one of the factors which attract FBM neurons to r5 and r6. PCP signaling is known to control the cellular distribution of motile cilia in multiciliated cells and of the primary cilia in other ciliated cells. Several components of the PCP pathways such as Dsv, Fzd, and Vangl2 are often associated with primary cilia which are microtubule-based organelles that protrude into the extracellular space and, through intercellular signaling, participate in the perception and integration of environmental cues such as morphogens (Wallingford, 2010). PCP genes also control the asymmetric localization and alignment of motile cilia in multiciliated cells (Borovina et al., 2010) and as such were shown to influence the flow of diffusible signals (such as morphogens and slits) in embryos.

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(A) Celsr1-3

wnt5 Ryk? Vangl2/Stbm

Fzd3

Pk1a

Dsh?

Rock JNK

(B)

Scrb1

Cytoskeleton

E15

E15

r3 r4 r5 r6

Celsr1 KO

Fz3 KO, Vangl2 KO, Wnt5a KO

FIGURE 18.4 The PCP pathway and FBM migration. (A), The major components of the PCP pathway and their interactions. The wnt receptor Ryk may interact with Fz and Wnt5 binding to Fz3 may activate the PCP pathway. Dvl function is unknown in migrating FBM neurons. (B), The PCP pathway controls the tangential migration of mouse FBM neurons. In Celsr1 knockout, some FBM neurons migrate rostrally and invade r3. In Fz3, Vangl2, and Wnt5a knockouts, the caudal migration of FBM neurons is perturbed (at variable level) and many remain at ectopic position in r4 and in R5.

Likewise, ciliogenesis is perturbed in zebrafish lacking Dvl or Fzd proteins or in Celsr2/Celsr3 knockout mice (Qu et al., 2010). The perturbed layout or abnormal growth of cilia consecutive to mutations in PCP genes could perturb the anteroposterior and dorsoventral diffusion or distribution of the guidance cues which allow FBM neurons to identify and follow their migratory pathway. Accordingly, in zebrafish, FBM neurons graft experiments have shown that many PCP genes (vangl, srcb1, Fz3, Celsr2, prickled 1) act in a non-cell-autonomous manner, and this has also been shown for Celsr1 in mouse (Qu et al., 2010). In addition, PCP signaling downstream of Dvl can influence both actin cytoskeleton (via RhoA/ Rho kinase and actomyosin) and microtubules (via Rac and c-Jun amino-terminal kinase) and thereby control FBM neuron motility. In hindbrain explants, the caudal migration of FBM neuron is perturbed in the presence of JNK or Rho kinase inhibitors (Vivancos et al., 2009). However, Dvl does not seem involved in FBM migration in zebrafish. One should also remember that, several PCP components (such as Celsr3 and Wnt5a) are known to activate other signaling pathways independently of PCP that could also be involved in FBM migration. For instance, in the spinal cord and cortex, the chemotropic activity of Wnts on postcrossing axons is mediated by the Ryk receptor and not by Fz (Lyuksyutova et al., 2003). Likewise, Clsr3 has homophilic properties and, in mouse, it acts cell-autonomously in migrating FBM neurons (Qu et al., 2010).

18.4.2.2 Other molecules controlling FBM caudal migration The cell adhesion molecule TAG-1 is expressed by premigratory FBM neurons and required for their migration (Garel et al., 2000; Sittaramane et al., 2009). TAG-1 expression in FBM is positively regulated by the transcription factor Ebf1 (Garel et al., 2000). In zebrafish, TAG-1 loss-of-function using tag-1 morpholinos blocks the migration of FBM neurons

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(Sittaramane et al., 2009). The use of suboptimal doses of morpholino provided evidence in favor of a genetic interaction between tag-1, laminin1 (lama1), and strabismus (Vangl2 protein) to control the migration of FBM. This suggests that tag1, lama1, and vangl2 cooperate to control the migration of FBM (Sittaramane et al., 2009). Interestingly, the expression of tag1 is switched off during dorsal migration in r6, whereas the expression of Cadherin-8 (Cad8) is turned on (Garel et al., 2000). These results suggest that during their caudal migration, FBM neurons dynamically regulate the expression level of CAMs at their surface from r4 to r6 in response to constant molecular changes occurring in their environment. Presenilin1 is a transmembrane protein mostly known for its role in Alzheimer disease as the mediator of “g-secretase” processing of the b-amyloid precursor protein (Reiner et al., 2009). Presenilin1, is an aspartyl protease which can also process many other proteins such as Notch, cadherins (Marambaud et al., 2002) Erb4, and DCC, which are all involved in neuronal migration (Louvi et al., 2004; TanIguchi et al., 2006). The phenotypic analysis of mouse embryos with targeted disruption of Psen1 showed that lack of Psen1 perturbs neuronal migration in cerebral cortex. This is most likely due to abnormal signaling downstream of Psen1 partners, such as Notch and Reelin (Reiner et al., 2009). Psen1 also plays a role in the tangential migration of a variety of brainstem neurons including IO and PN neurons (Louvi et al., 2004). In Psen1 knockout, the identity of FBM neurons is mostly preserved but a majority fails to migrate and remains at ectopic positions in r4 (Louvi et al., 2004). Interestingly, FBM neuron migration defects are remarkably similar in Psen1 mutant and Cdk5 mutants (see below). It is known that p35/cdk5 phosphorylation of b-catenin regulates its interaction with Presenilin-1 (Kesavapany et al., 2001). Therefore, the abnormal FBM migration in Cdk5 mutant could be due to altered b-catenin/ Presenilin-1 signaling. However, other Cdk5 downstream partners, such as the dynein heavy chain, could also be involved. In legs at odd angles (Loa/Loa) mutant mice, which carry a missense mutation in dynein heavy chain, only a fraction of FBM neurons migrate to their correct caudal position, while the remaining ones migrate and settle into more anterior nuclei (Hafezparast et al., 2003). It has also been proposed that the lack of Presenilin-1 may perturb FBM migration by altering the activity of GSK3 activity and kinesin-1 (Pigino et al., 2003). All these cytoplasmic molecules play key roles in the regulation of cytoskeleton dynamics and in the control of cell migration in many systems. In conclusion, Psen1 is likely to influence the tangential migration of both PCN and FBM neurons by modulating the activity of axon guidance receptors and CAMs (DCC, N-Cadherins, Erb4,.) and also of molecules acting on the cytoskeleton (Cdk5, b-catenin, Tau,.). Interestingly, b-catenin binding to Psen1 requires Cadherin (Serban et al., 2005) and mediates b-catenin ubiquitination suggesting that to some extent the neuronal migration defects in Psen1 mutants are due to abnormal Cadherin/catenin signaling.

18.4.3 Role of chemoattraction and chemorepulsion Direct evidence supporting the existence of environmental cues being able to guide the caudal migration of FBM neurons was first obtained using transplantation experiments. As previously mentioned, unlike rodent or fish FBM, chick FBM do not migrate caudally. To determine whether this was due to the lack of attractive guidance cues in chick rhombomeres caudal to r4 or whether chick FBM was unable to respond to such cues, Studer (Studer, 2001) performed homotopical in ovo transplantations of mouse r5 and/or r6. In these chick/mouse chimeras, chick ventral r4 neurons tangentially migrated caudally through r5 and r6 toward the transplanted mouse rhombomeres just like mouse FBM neurons. Further, when mouse r4 was homotopically transplanted in chick, mouse FBM neurons failed to migrate caudally and remained in r4. These experiments clearly indicate that local chemotropic cues present in mouse r5 and r6, but not in chick, attract r4-derived FBM neurons caudally and trigger their tangential migration. Several diffusible chemotropic molecules are expressed in mouse r5 and r6 and were shown to control the migration of FBM neurons. Netrins, secreted semaphorins, Slits are known to control the tangential migration of many classes of forebrain interneurons and precerebellar neurons (see references above). In the hindbrain, these molecules are all enriched at the ventral midline, along which FBM neurons migrate. Moreover, receptors for these molecules such as DCC and UNC5A (netrin receptors), plexins and neuropilin1 (semaphorin receptors), Robo1 and Robo2 (Slit receptors) are expressed by FBM neurons from the time they leave the neuroepithelium, to the end of their migration (Barrett and Guthrie, 2001; Hammond et al., 2005; Meléndez-Herrera and Varela-Echavarría, 2006; Schwarz et al., 2008). Although there is strong evidence supporting a role for these molecules in the patterning of motor projections (Varela-Echavarría et al., 1997), so far, there are no reports of abnormal FBM migration or positioning in mice deficient for these molecules or their receptors (Geisen et al., 2008). The lack of reported FBM phenotypes in these mice is intriguing and merits to be more carefully studied. Vascular endothelial growth factor (VEGF-A) is a proangiogenic factor playing a key role in angiogenesis in normal and pathological conditions (Ruiz de Almodovar et al., 2009). Several VEGF-A isoforms are generated by alternative splicing and bind to several receptors such as the receptor tyrosine kinase VEGFR1 and VEGFR2 and to the semaphorin

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receptor Neuropilin1. It was shown that VEGF-A164 isoform is expressed in the developing hindbrain in the floor plate and along the pathway followed by tangentially migrating FBM neurons which highly express Neuropilin1 (Fig. 18.2) (Schwarz et al., 2004). The VEGF-A164 and Sema3A binding sites on neuropilin1 are distinct and have thus rendered it possible to generate neuropilin1 knock-in mice in which the binding of only one of the two ligands is prevented (Gu et al., 2003). Interestingly, it was shown that in neuropilin1 full knockout but not in the neuropilin1 knockin in which the Sema3A binding site was selectively mutated, most FBM neurons are unable to reach r6 and stay at ectopic locations in more rostral rhombomeres. The same migratory defects are observed in hindbrain explants treated with an antibody which blocks VEGF-A164 function. In addition, VEGF-A164-soaked beads attract FBM neurons in explants (Schwarz et al., 2004; Vivancos et al., 2009). These results show that VEGF-A164 is one of the factors that mediate the caudal migration of FBM neurons. In contrast, Sema3A, and not VEGF, guides FBM axons but does not influence FBM migration (Schwarz et al., 2004). These results emphasize that VEGF is not only essential to pattern the vasculature but is also important for guiding the migration of some neuronal populations. Finally, it is important to note that VEGF offers one of the rare examples of a factor involved in axon guidance and not in neuronal migration.

18.4.4 Role of the meninges in the tangential migration of FBM neurons Like PCN neurons, tangentially migrating FBM neurons are influenced by meningeal-derived factors. The first evidence for a role of CXCL12 in neuronal migration was obtained in zebrafish where this chemokine was shown to guide the migration of cells from the cephalic placode during the formation of the neuromasts of the lateral line (David et al., 2002). A few years later it was found that CXCL12 is also involved in the caudal migration of FBM neurons. In zebrafish, embryos injected with CXCL12 morphants, r4-derived FBM neurons do not reach r6 and remain at ectopic locations. CXCL12 is highly expressed in the hindbrain meninges, and its G-protein-coupled receptor CXCR4 is expressed by the migrating facial motor neurons (Sapède et al., 2005). More recently, it has been demonstrated that CXCL12 can bind two different receptors: CXCR4 and CXCR7. The analysis of their expression pattern revealed that CXCR4 is expressed by FBM neurons migrating from r4 to r6, whereas CXCR7 is restricted to motor neurons in ventral r5. To determine the role played by each receptor (Cubedo et al., 2009), the expression of CXCR7b or CXCR4b was silenced in zebrafish. In both cases, the tangential migration of FBM neurons was altered albeit differently: in CXCR7b morphants, ectopic FBM neurons accumulated in r5, whereas in CXCR4b morphants, they remained scattered within r4 and r5. Therefore, CXCR7 is necessary for proper tangential migration, most probably by providing a permissive pathway through r5. As mentioned previously, retinaldehyde deshydrogenase-2, the enzyme responsible for the synthesis of retinoic acid, is highly expressed in the meninges and mesenchyma surrounding the hindbrain during FBM migration (Zhang et al., 2003). The study of the zebrafish mutant neckless (nls), in which the retinaldehyde deshydrogenase-2 gene is disrupted, has shown that many FBM neurons do not migrate out of r4, thereby suggesting that RA plays a role not only in the specification of rhombomere identity but also in the caudal migration of FBM neurons. Although direct evidence is still lacking, it is therefore possible that meninges also exert a guidance function on FBM neuron tangential migration through the secretion of RA. In zebrafish, postmitotic FBM neurons start moving ventrally in r4 until they reach the laminin-containing basement membrane that delimits r4 and r5. At this boundary, FBM neurons change velocity and centrosome orientation and initiate their caudal migration using the basal lamina as a substrate (Grant and Moens, 2010). While PCP signaling set up the planar polarity of epithelia, their dorsoventral polarity requires members of the PAR-aPKC complex. The genetic inactivation of aPKCl and aPKCz disrupts the basement distribution and its laminin distribution (Grant and Moens, 2010). Having lost its sealing capacity, the basement membrane allows some FBM neurons to escape the hindbrain and invade the meninges. This defect is also observed when laminin1 expression is silenced suggesting that, in these mutants, FBM neurons respond to the attractive action of a meningeal-derived attractants but also that the basement membrane acts as a barrier for FBM neurons. This is very reminiscent of what has been described in the cerebellum and cerebral cortex where interfering with basal membrane integrity and integrin signaling allows migrating neurons to migrate out of the brain parenchyma (Belvindrah et al., 2006; Li et al., 2008; Sievers et al., 1986). In conclusion, a plethora of factors are involved in the regulation of the caudal tangential migration of FBM neurons from r4 to r6er7. Many of them are similar to those at play in other cases of tangential migration (chemotropic or adhesion molecules, meningeal factors). Some act through cell-autonomous mechanisms, such as the transcription factors required to generate either the rhombomeric identity of r4 (Hoxb1 and Pbx4) or FBM neuronal identities (Ebf1-3, Nkx6.1, Hdac1, and Tbx20). These factors control the ability of motor neurons to interact with local environmental signals and play a cellautonomous role in the migratory process. In contrast, other factors such as Krox20 and Kreisler, or proteins of the

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PCP pathway, exert non-cell-autonomous role and modulate the expression of guidance cues in the territories present across the migratory route of FBM neurons (Bingham et al., 2010).

18.5 Ending tangential migration In the final phase of their migratory journey, FBM neurons change direction again and migrate in a dorsoventral direction along radial glia to the ventrolateral surface of the medulla oblongata, beneath the pial surface. Electroporation is a method which allows introducing plasmids in proliferating cells. This can be applied in ovo to chick embryos or in utero to mouse embryos. Although PCN neurons primarily migrate tangentially at the hindbrain surface, several studies using electroporation of GFP plasmids have shown that PCN neurons switch to a radial mode of migration upon reaching their destination and migrate inward inside the hindbrain. As detailed below, genetic studies strongly suggest that distinct signaling pathways are involved in the tangential and radial migration of FBM neurons. In particular, radial migration is Reelin-dependent while caudal and lateral tangential migrations are not. In adult reeler mutant mice, the cytoarchitectonic of the facial nucleus is severely perturbed (Goffinet, 1984). Reeler mice are mutated in the gene encoding Reelin, a large extracellular matrix protein containing several EGF-like repeats (D’Arcangelo et al., 1995). The facial nucleus of the reeler contains a similar number of neurons than in wild-type mice, but it is less clearly organized, its main body is wedge-shaped and spread out more widely in a radial direction (Goffinet, 1984). These defects suggest that Reelin can control the interaction between migrating FBM neurons and radial glial processes. Moreover, similar FBM migration defects were reported in mice deficient for the intracellular adaptor protein disabled-1 (Dab1; scrambler and yotari mice) which acts downstream of the reelin receptors VLDLR (very low-density lipoprotein receptor) and ApoER2 (apolipoprotein E receptor type2) (Sheldon et al., 1997; Tissir and Goffinet, 2003). Therefore, the integrity of the Reelin signaling pathway is necessary for FBM neurons to complete their migration. Similarly, in reeler mice the migration of IO neurons is also partially perturbed. In reeler, postmitotic IO neurons are still able to migrate from the rhombic lip to their proper ventral location, but the cytoarchitecture of the ION, in particular its characteristic lamellate shape, is altered. It therefore appears that, as for FBM neurons, Reelin controls the last phase (radial) of IO neuron migration, although in this case, the source of Reelin is still unclear (Tissir and Goffinet, 2003). This is also the case for FVM neurons, whose radial migration is disrupted in reeler and prevents them from reaching their normal subpial location (Rossel et al., 2005). Cyclin-dependent kinase 5 (Cdk5), a serine/threonine kinase only expressed in postmitotic neurons, plays many roles in the development and plasticity of the central nervous system. Cdk5 is a major regulator of cytoskeleton dynamics and modulates the migration of many classes of neurons. Monomeric Cdk5 lacks kinase activity and requires, for its activation, the association with regulatory partners such as p35 or p39 (Xie et al., 2006). In Cdk5 knockout mice, the lamellation of the IO never occurs, and the migration of FBM neurons is also altered. FBM neurons condense in two ectopic bilateral masses in the dorsal pons, at a more rostral position than the normal location of the facial nuclei. Interestingly, in compound mutants for Cdk5 and its activator, p35, FBM defects are more severe than in Cdk5 single mutant (Ohshima et al., 2002). Indeed, the cell bodies of FBM neurons do not initiate their tangential migration and are unable to leave r4. However, the facial nerve still exits the brainstem at its correct position in r4 (Ohshima et al., 2002). The IO and FBM migratory defects described in newborn Cdk5 knockout mice (the mutation is lethal) are reminiscent of those found in Reelin/Dab1 mutant mice, suggesting that Cdk5/p35 and Reelin/Dab1 might interact in a common signaling pathway to control neuronal migration. However, the migration defects in cortical structures in p35 and Reelin/Dab1 double mutants are more severe than in the corresponding single mutants, suggesting that Cdk5/p35 and Reelin/Dab1 are more likely to act in parallel or synergistic signaling pathways (Beffert et al., 2004; Ohshima et al., 2002). This conclusion is supported by the fact that in reeler, early tangential migration does occur, and only radial migration is affected. Indeed, even though a large fraction of FBM neurons reach their final location, some of them remain ectopic along their radial pathway (Tissir and Goffinet, 2003). FBM migration defects in Ckd5 knockout mice and in reeler mutants are thus quite different (Goffinet, 1984). The reeler model demonstrates that, in the IO and FBM, the last phase of radial migration is essential for the acquisition of the characteristic anatomical subdivisions of these nuclei. In the rostral migratory stream, reelin was shown to act as a mitral cell-derived signal which triggers the detachment of olfactory bulb interneurons from their migratory chain and allows radial migration to proceed (Hack et al., 2002). Reelin could exert a similar function on tangentially migrating FBM and IO neurons. As PCN neurons are among the last to be born in the hindbrain, they can be selectively labeled by electroporation of expression constructs encoding GFP. Examination of fixed samples and time-lapse on hindbrain slices show that the morphology of many PCN neurons change when they switch from tangential to radial migration (Kawauchi et al., 2006; Okada et al., 2007) and that a new

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leading process emerges on the side opposite to the pial surface. After a delay of up to 32 h (Watanabe and Murakami, 2009), the soma then moves inward along radial glia processes to its final position (Kawauchi et al., 2006). ln some neurons, the tangential leading process is kept but the neurons nonetheless switch their migratory path to a radial direction (Watanabe and Murakami, 2009). Interestingly, the electroporation of a dominant-negative N-cadherin in LRN neurons induces premature switch to radial migration (Kawauchi et al., 2006), but the exact mechanism is unknown. However, in zebrafish, it was shown that the homophilic tangential migration of cerebellar granule cell precursors requires N-cadherin (Rieger et al., 2009), suggesting that switching N-cadherin expression off in PCN neurons may contribute to the initiation of their radial migration. Interestingly, the final positioning of PCN and FBM neurons is perturbed in mice deficient for the GEF Trio (Backer et al., 2007) which is a putative partner of cadherin-11, a molecule that is expressed in these neurons during their radial migration.

18.6 Conclusion Altogether studies have shown that although both FBM and IO neurons migrate tangentially in the hindbrain, their migratory pathways are to a large extent shaped by distinct guidance cues. In the case of PCN neurons, the floor plate and the various attractants and repellents that it secretes play a major role in influencing tangential migration, whereas there is yet no evidence that this is the case for FBM neurons. Likewise, VEGF has a major influence on tangentially migrating FBM neurons but is not known to guide IO neurons. However, these apparent differences could in effect be due to the fact that, in most cases, the involvement of a particular guidance cue has only been studied in only one of these nuclei. Still, there are also some striking similarities between both systems such as the crucial role of meningeal-derived factors that keep most of these neurons close to the pial surface and the role of reelin in the initiation of radial migration.

Acknowledgments We thank A. Ypsilanti for critical reading of the manuscript. This work is supported by the Agence Nationale pour la Recherche (ANR), the Fondation pour la Recherche Médicale (Equipes FRM), and the Spanish Ministry of Education and Science (Grant BFU2008-00588/BFI).

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Neurosci. Res. 64, 20e29. Wolfer, D.P., Henehan-Beatty, A., Stoeckli, E.T., Sonderegger, P., Lipp, H.P., 1994. Distribution of TAG-1/axonin-1 in fibre tracts and migratory streams of the developing mouse nervous system. J. Comp. Neurol. 345, 1e32. Wong, K., Ren, X.R., Huang, Y.Z., Xie, Y., Liu, G., Saito, H., Tang, H., Wen, L., Brady-Kalnay, S.M., Mei, L., et al., 2001. Signal transduction in neuronal migration: roles of GTPase activating proteins and the small GTPase Cdc42 in the Slit-Robo pathway. Cell 107, 209e221. Wu, Z., Makihara, S., Yam, P.T., Teo, S., Renier, N., Balekoglu, N., Moreno-Bravo, J.A., Olsen, O., Chédotal, A., Charron, F., et al., 2019. Long-range guidance of spinal commissural axons by Netrin1 and sonic hedgehog from midline floor plate cells. Neuron 101, 635e647. Xie, Z., Samuels, B.A., Tsai, L.-H., 2006. Cyclin-dependent kinase 5 permits efficient cytoskeletal remodeling–a hypothesis on neuronal migration. Cerebr. Cortex 16 (Suppl. 1), i64ei68. Yamamoto, M., Zhang, J., Smith, D., Hayakawa, Y., McCaffery, P., 2003. A critical period for retinoic acid teratogenesis and loss of neurophilic migration of pontine nuclei neurons. Mech. Dev. 120, 701e709. Yee, K.T., Simon, H.H., Tessier-Lavigne, M., O’Leary, D.D., 1999. Extension of long leading processes and neuronal migration in the mammalian brain directed by the chemoattractant netrin-1. Neuron 24, 607e622. Yung, A.R., Druckenbrod, N.R., Cloutier, J.-F., Wu, Z., Tessier-Lavigne, M., Goodrich, L.V., 2018. Netrin-1 confines rhombic lip-derived neurons to the CNS. Cell Rep. 22, 1666e1680. Yurchenco, P.D., Wadsworth, W.G., 2004. Assembly and tissue functions of early embryonic laminins and netrins. Curr. Opin. Cell Biol. 16, 572e579.

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Zelina, P., Blockus, H., Zagar, Y., Péres, A., Friocourt, F., Wu, Z., Rama, N., Fouquet, C., Hohenester, E., Tessier-Lavigne, M., et al., 2014. Signaling switch of the axon guidance receptor Robo3 during vertebrate evolution. Neuron 84, 1e15. Zhang, J., Smith, D., Yamamoto, M., Ma, L., McCaffery, P., 2003. The meninges is a source of retinoic acid for the late-developing hindbrain. J. Neurosci. 23, 7610e7620. Zhu, Y., Khan, K., Guthrie, S.C., 2003. Signals from the cerebellum guide the pathfinding of inferior olivary axons. Dev. Biol. 257, 233e248. Zhu, Y., Matsumoto, T., Mikami, S., Nagasawa, T., Murakami, F., 2009. SDF1/CXCR4 signalling regulates two distinct processes of precerebellar neuronal migration and its depletion leads to abnormal pontine nuclei formation. Development 136, 1919e1928.

Chapter 19

Neuronal migration in the developing cerebellar system Christophe Laumonnerie and David J. Solecki Department of Developmental Neurobiology, St. Jude Children’s Research Hospital, Memphis, TN, United States

Chapter outline 19.1. Introduction 19.1.1. Part I. Diverse migration pathways and guidance cues during cerebellar system development 19.1.1.1. Distinct cerebellar germinal zones: the ventricular zone and rhombic lip 19.1.1.2. Migration of purkinje cells 19.1.1.3. Migration of minor ventricular zone derivatives: Pax2-positive interneurons, basket cells, golgi cells, and stellate cells 19.1.1.4. Migration of precerebellar nuclei 19.1.1.5. Migration of upper rhombic lip derivatives 19.1.2. Part II. The cytoskeletal organization of cerebellar granule neurons 19.1.2.1. Cerebellar granule neuron migration diversity after the establishment of the secondary germinal zone 19.1.2.2. The road to the two-stroke motility paradigm

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19.1.2.3. The roles of the microtubule cytoskeleton and associated motors 19.1.2.4. The role of the actin cytoskeleton 19.1.2.5. The role of microtubule-actin cross talk 19.1.3. Part III. The facets of cerebellar granule neuron polarity: timing cell recognition, differentiation, germinal zone exit, and morphogenesis 19.1.3.1. Cerebellar granule neuron recognition/ adhesion: the contribution of astrotactins and the siah2ePard3eJamC pathway 19.1.3.2. The Zeb1-Pard6/3A transcriptional pathway 19.1.3.3. The foxo polarization pathway 19.1.4. Part IV. Migration deficits in cerebellar medulloblastomas: the effects of perturbed migration pathways are no longer limited to cognitive deficiency Acknowledgments References

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19.1 Introduction The cerebellum is a structure of the central nervous system that is located on the dorsal hindbrain on top of the fourth ventricle and participates in the fine regulation of motor control (Ito, 1984), cognition (Leiner et al., 1986; Schmahmann, 1996), and visceral processes (Haines et al., 1990). The cerebellum is composed of various cell types exhibiting a wide range of morphology that reflects tightly regulated ontological processes. The neuroscientist Santiago Ramón y Cajal, through his microscopic studies at the end of the 19th century, was the first to highlight the diversity of cellular morphology in the cerebellum. Morphologically, the cerebellum is a foliated structure consisting of a laminated cortex, white matter, and paired sets of four cerebellar nuclei. The cerebellar cortex organizes itself with a central region called the vermis and two lateral hemispheres. The vermis has three main lobes that are distributed along the rostrocaudal axis and further subdivide into 10 lobules. This organization is well conserved in avian and mammalian species, with some variation in the size and number of the lobules. The cortex includes three laminae: the molecular layer is the most superficial of these, followed by the Purkinje layer, and then the internal granular layer (IGL) (Fig. 19.1). The laminae are each composed of a well-

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FIGURE 19.1 The cerebellum and the precerebellar system. (A) Schematic representation depicting parasagittal section of the hindbrain of an adult mouse depicting the interconnectivity between the precerebellar structure and the cerebellum. The cerebellum receives two principal inputs that are the mossy fibers (MFs) and the climbing fibers (CFs). MFs originate from the reticulotegmental nucleus (RTN) the basal pontine nucleus (BPN), the external cuneate nucleus (ECN) and lateral reticular nucleus (LRN), and project principally onto the granule neurons (GN) in the internal granule layer (IGL) of the cerebellar cortex (Cb Cx). (B) GNs in return form local excitatory synapses in the molecular layer (ML) with different GABAergic interneurons and more importantly with the Purkinje cell (PC) which resides in the Purkinje cell layer (PCL), in between the ML and the IGL. PC also receives CF afferents originating from the inferior olivary nucleus (ION) in the ventral hindbrain. (A) PCs send their inhibitory projections out of the Cb Cx toward the cerebellar nuclei (CN). The CN, only output of the cerebellum, project in return glutamatergic projections toward different hindbrain, midbrain, and thalamic nuclei. CN as well send GABAergic nucleo-olivary (NeO) projections to the ION.

characterized set of neurons. Stellate and basket cells are GABAergic inhibitory interneurons that populate the molecular layer, which otherwise consists mostly of parallel fiber axons. Candelabrum cells and Purkinje cells (PCs) are also GABAergic neurons and reside in the Purkinje layer. The granular layer is more diverse and contains two types of GABAergic inhibitory interneurons, Golgi and Lugaro neurons, as well as cerebellar granule neurons (CGNs) and unipolar brush cells (UBCs), which are glutamatergic excitatory neurons. With the exception of Purkinje cells, all cortical neurons project locally. In addition to the cerebellar cortex, deep cerebellar nuclei (CNs) are located in the deep white matter of the cerebellum adjacent to the roof of the fourth ventricle. They receive their main afferent projections from the Purkinje cells and are the only output of the cerebellum. The CNs consist of several neuronal cells types: large glutamatergic projection neurons, midsized GABAergic inhibitory projection neurons (Mugnaini, 1985; Ruigrok, 1997), and small GABAergic and glycinergic interneurons (Uusisaari et al., 2007). The cerebellum and, by extension, the precerebellar system that provides the principal input to the cerebellum, has been a model of choice for investigating neuronal development and, specifically, neuronal migration, as well as how complex circuitry is assembled during development (Fig. 19.1). The complex flow of information processed by the cerebellum results from the precise interconnectivity of the cerebellum and the precerebellar nuclei. The precerebellar nuclei provide the two principal inputs to the cerebellar cortex: climbing fibers (CFs), originating from the inferior olivary nucleus, and mossy fibers (MFs), projecting from the pontine nuclei, external cuneate nucleus, and lateral reticular nucleus (Altman and Bayer, 1987). Through the integration resulting from the laminar organization of the cerebellar cortex, the information is transferred to the CNs by the PCs. Finally, the CN projection neurons send afferents back outside the cerebellum. This chapter will cover the developmental processes that give rise to this complex architecture, from early patterning events to migration, highlighting the interdependence of many of these events. Later sections will take the example of CGN molecular events during the cytoskeletal rearrangement and polarity regulation involved in the migration process. Finally, examples of how knowledge of the regulatory events involved in precise migration control is relevant to a better understanding of human disease pathology will be presented, such as the development of medulloblastoma (MB).

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19.1.1 Part I. Diverse migration pathways and guidance cues during cerebellar system development 19.1.1.1 Distinct cerebellar germinal zones: the ventricular zone and rhombic lip Cerebellar studies have a long history (Ramon y Cajal, 1911), but only recently have researchers begun to understand the underlying molecular machinery responsible for cell fate decisions, seeking to confirm the validity of developmental models based on the earlier anatomical descriptions. In the last two decades, thanks to advances in molecular biology and, specifically, the generation of transgenic mouse lines allowing the precise study of the cellular lineage of distinct cell types by using the Cre/Lox recombination system, much progress has been made toward unraveling the origin of the different cerebellar cells. 19.1.1.1.1 Early patterning The different types of cerebellar neuron each have a distinct morphology and location in the mature cerebellum. Recently, it has become clear that all neurons and glial cells that contribute to the cerebellum develop from progenitors residing in the neuroepithelium of rhombomere 1 (Rh1)(Millet et al., 1996; Wingate and Hatten, 1999; Chizhikov and Millen, 2003; Zervas et al., 2004). The cerebellum arises from a specialized region at the midbrain/hindbrain boundary (Hallonet et al., 1990; Hallonet and Le Douarin, 1993; Hallonet and Alvarado-Mallart, 1997). Here, at embryonic day 8.5, the mutually exclusive expression of the homeobox genes Otx2 (anteriorly) and GBX2 (posteriorly) defines the isthmic organizer region (IsO) (Broccoli et al., 1999; Baudoin et al., 2008). The IsO then orchestrates the patterning of cerebellar and mesencephalic structures by releasing secreted morphogenic factors such as Fgf8 and Wnt1 (Martinez et al., 1999; Sotelo, 2004). Ventral expression of sonic hedgehog (Shh) by the ventral midline also promotes Fgf8 expression by the IsO from E8.5 to E12 (Epstein et al., 1999; Fuccillo et al., 2006). Hox gene expression domains delimit the caudal expansion of the cerebellar territory, as observed in a Hoxa2 mutant in which the cerebellar territory expands caudally (Gavalas et al., 1997). This posterior boundary of Rh1 is further defined by Fgf8 expression, which represses the expression of Hox genes (Irving and Mason, 2000). After initial patterning of the presumptive cerebellar territory, cerebellar histogenesis starts at E9. At this stage, the cerebellar anlage comprises two protuberances separated by the opening of the fourth ventricle. Subsequently, these two parts grow as a result of intense proliferation and eventually undergo a 90 degrees rotation, leading to the unitary cerebellar plate consisting of the medial vermis and the two cerebellar hemispheres (Altman and Bayer, 1997; Sgaier et al., 2005). It should be noted that this movement converts the embryonic rostrocaudal axis into the mediolateral axis of the cerebellar primordium (Sgaier et al., 2005). Cell fate decisions are determined in specific subregions of the neuroepithelium at the beginning of cerebellar morphogenesis and, thus, by the expression of a specific set of transcription factors (Wang and Zoghbi, 2001; Sotelo, 2004). Often in neurodevelopment, the same neuroepithelium can produce different cell types, neurons, and glia, in successive temporal waves or even produce a secondary germinative pool that can export the proliferation to another site. In the cerebellum, similar to what is seen in other structures such as the forebrain, neuronal fate specification is compartmentalized according to the neurotransmitter fate; more specifically, between glutamatergic and GABAergic neurons (Fig. 19.2). Neuroepithelia that produce the cerebellar neurons can be divided into two principal molecularly defined regions that reflect this dichotomy: the rhombic lip (RL), a germinative neuroepithelium lining the dorsal edge of the fourth ventricle in the hindbrain, and the ventricular zone (VZ), which lines the dorsolateral aspect of the fourth ventricle, respectively. Cerebellar development is characterized by primary and secondary germinal zones (GZs). The secondary zones are the external granular layer (EGL), formed at the surface of the developing cerebellar surface originating from the RL, and the prospective white matter (PWM), containing progenitors from GABAergic interneurons and astrocytes, delaminated from the VZ, as well as progenitors of oligodendrocytes that arise from an extracerebellar GZ (Hashimoto et al., 2016). Other cells, such as PCs and neurons from the cerebellar nuclei (CN), appear to delaminate from the neuroepithelium as postmitotic neurons. 19.1.1.1.2 The rhombic lip and Atoh1 domain define the glutamatergic lineage The RL extends along the full length of the anteroposterior axis of the rhombencephalon and is subdivided into the upper and lower RL. The glutamatergic neurons of the cerebellum derive exclusively from the upper RL, whereas the posterior RL contributes to the formation of different hindbrain nuclei, including some of the precerebellar system (see below). BenArie and colleagues were the first to report that a basic helix-loop-helix (bHLH) type of transcription factor, Atoh1 (also known as Math1), is expressed in the developing RL and is involved in the production of CGNs (Ben-Arie et al., 1997).

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FIGURE 19.2 The cerebellum origin and migration. Schematic representation depicting parasagittal section of the cerebellum at different developmental time-point. Around E12 neurons originating from the Atoh1 positive rhombic lip (RL) have initiated their subpial migration through the rhombic lip migratory stream (RLS) and started to accumulate rostrally forming the NTZ. The ventricular zone (VZ) is subdivided into rostral interneuron progenitors (PIP) giving rise to GABAergic projection neurons joining the forming NTZ, while the posterior VZ contains Purkinje cell progenitors (PCPs) producing Purkinje cells (PCss) that migrate radially along the radial glia. Around E14.5, neurons of the NTZ have progressed inwards forming the cerebellar nuclei (CN). The surface of the cerebellar anlage is now occupied by granule neuron progenitors (GNPs) coming from the RL thus forming the EGL. The VZ is now solely producing interneuron progenitors that populate the PWM. PCs pile below the EGL in a several cell thick layer: the Purkinje cell plate (PP). Radial glial cells start retracting their apical process and translocate beneath the PP where they will differentiate into Bergmann glia (BG). Around E17.5, GNPs in the EGL proliferate and contribute to expand the surface of the cerebellar cortex while the PCs rearrange into a 1 cell layer. Some GNPs start differentiating and extend processes in between the EGL and the PC layer to form the molecular layer (ML). INs in the PWM keep proliferating while some differentiate and populate the CN or settle beneath the PC layer. Finally, the RL produces unipolar brush cells (UBCs) that will migrate in the PWM to settle in the future granule layer (GL). On the right-hand side a timeline recapitulate the successive wave of neurogenesis coming from the Atoh1 positive RL and the Ptf1a positive VZ that contribute to the cerebellum formation.

Molecular fate maps of the Atoh1 lineage have revealed different waves of neurogenesis, each contributing to a specific lineage (Machold and Fishell, 2005; Ozdamar et al., 2005; Englund et al., 2006). Cells born between E10.5 and E12.5 give rise to the glutamatergic neurons of the CNs. These neurons are postmitotic, downregulate Atoh1, and express Tbr1, Irx3, Meis2, and Lhx2/9 (Morales and Hatten, 2006). From E12.5 to E16.5, the RL produces granule neuron progenitors (GNPs) that remain proliferative as they delaminate from the RL. They characteristically maintain their expression of Atoh1 (Aruga et al., 1998). Finally, around E13.5, the RL, as it widens, begins to produce postmitotic UBCs, characterized by Tbr2 expression, and continues to do so until E18.5 (Englund et al., 2006). 19.1.1.1.3 The ventricular zone and Ptf1a domain define the GABAergic lineage In the cerebelless mutant mouse strain, the loss of expression of the bHLH transcription factor pancreatic transcription factor 1a (Ptf1a) results in the total loss of GABAergic neurons, showing that Ptf1a is essential for the determination of

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these different lineages (Hoshino et al., 2005). Analysis of a Ptf1a null mutant, showed that the fate of cells produced by the VZ was changed to that of granule cells (Pascual et al., 2007). Interestingly, a recent experiment showed that substituting Ptf1a and Atoh1 expression in their respective domains resulted in a switch from GABAergic to glutamatergic neurons in the VZ and from glutamatergic to GABAergic neurons in the RL (Seto et al., 2014). Surprisingly, Ptf1a expression was recently shown to be largely restricted to nonmitotic cells of the cerebellar anlage (Huang et al., 2010). This finding, together with the characterization of the Ptf1a-null phenotype in which GABAergic precursors migrate to the rhombic lip and acquire a GNP-like phenotype (Pascual et al., 2007) and the observations suggesting that Ptf1a displays antiproliferative activity by downregulating cyclin D2 (Rodolosse et al., 2004), argues in favor of Ptf1a having a function in locking down the phenotype of this lineage. Birthdating experiments of the Ptf1a lineage have revealed a temporal sequence for the generation of the different GABAergic neurons contributing to the cerebellum (Miale and Sidman, 1961; Altman and Bayer, 1997; Sekerkova et al., 2004; Hoshino et al., 2005; Ayala et al., 2007) (Fig. 19.2). The position of the progenitors within the dorsoventral axis has enabled the definition of some subdomains in the VZ (Chizhikov et al., 2006). At E10.5, a ventral domain, pc2v (also defined as a Pax2þ interneuron progenitor (PIP)) is characterized by Neph3 expression and a low level of E-Cad. These progenitors give rise to intermediate progenitors expressing the transcription factors Pax2 and Gsx1, which delaminate to populate the PWM. The dorsal domain pc2d (also referred to as a Purkinje cell progenitors (PCPs)) expresses Neph3 and a high level of E-Cad and gives rise to postmitotic PCs expressing Corl2 and Olig2 (Maricich and Herrup, 1999; Ayala et al., 2007; Gardel et al., 2010; Mizuhara et al., 2010; Seto et al., 2014). From E10.5 to E13.5, the ventral phenotype progressively extends dorsally at the expense of the dorsal domain (Seto et al., 2014). The different GABAergic neurons are produced in the following sequence: From E10.5 to E13.5, the ventral domain produces GABAergic projection neurons from the CNs, followed by the GABAergic interneurons from the CNs, while the dorsal domain produces PCs (Seto et al., 2014). From E13.5 to the perinatal stages, the VZ exclusively produces intermediate progenitors giving rise to the parenchymal astrocytes and GABAergic interneurons found in the cerebellar cortex (Seto et al., 2014). GABAergic interneurons are produced in an inside-out sequence from the granular layer to the molecular layer in the following order: Golgi, Lugaro, candelabrum, stellate, and basket cells (Seto et al., 2014) (Fig. 19.2). Other transcription factors have been shown to be involved in defining the subtype of GABAergic neurons: the bHLH transcription factor Ascl1 is involved in regulating interneurons but not PCs (Grimaldi et al., 2009), whereas an Lhx1/5 mutant specifically reduces PCs but not Pax2-positive interneurons (Zhao et al., 2007). More recently, differential expression of two members of the AP-2 transcription factor family, Tfap2A and Tfap2b, has been proposed in the specification of GABAergic interneurons (Zainolabidin et al., 2017). 19.1.1.1.4 Other Rh1 derivatives The roof plate, the most dorsal part of the neuroepithelium on top of the RL, does not produce neurons and is defined by early expression of the transcription factor Lmx1a. The roof plate gives rise to the choroid plexus (Chizhikov et al., 2006) and ultimately contributes to the fate specification of the neuroepithelium by secreting morphogens such as BMPs, Notch/ Delta, and Gdf7 (Machold et al., 2007; Broom et al., 2012). The choroid plexus is also a source of the Shh secreted in the cerebrospinal fluid that influences the proliferation of progenitors in the VZ, as well as the radial glia (Huang et al., 2010). Atoh1 is also expressed early in a supplementary domain in Rh1 close to the IsO. This rostral isthmic domain is dependent on Fgf8 (Green et al., 2014) and contributes to the formation of isthmic nuclei such as the parabigeminal nucleus and the dorsal nucleus of the lateral lemniscus, previously reported to arise from an Rh1-derived Atoh1-positive domain (Machold and Fishell, 2005; Rose et al., 2009).

19.1.1.2 Migration of purkinje cells Of all the cell types that arise during the ontogeny of the cerebellum, PCs are probably the most important in orchestrating the development of the entire structure and especially in regulating proliferation and the processes leading to the cortical layering and compartmentalization of the cerebellar cortex. Purkinje cells are the iconic cell type of the cerebellum and are easily identified by their distinctive morphology in the mature cerebellum. In the cerebellar cortex, they are arranged in a monolayer between the molecular layer and the granular layer. Their morphological distinctiveness lies in the extensive treelike arborization of their dendrites in a parasagittal plane. This cell type was discovered by the anatomist Jan Evangelista Purkinje in 1837 (Lewis et al., 2004), and its morphology was described in further detail 50 years later by Santiago Ramón y Cajal (Ramón y Cajal, 1888), using the silver impregnation method developed by Camillo Golgi (Golgi, 1883). Purkinje cells are generated from a specific pool of progenitors within the ventricular neuroepithelium, and they belong, along with the GABAergic interneurons, to the Ptf1a þ lineage (see above (Hoshino et al., 2005)). The essential

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developmental progression can be recapitulated in culture, suggesting that these steps are partly regulated through the unfolding of a cell-autonomous program. Around E12.5, newborn PCs initiate corticogenesis by moving radially from the VZ toward the pial surface beneath the superficial region of the cerebellar anlage, which is occupied by the CN neurons (Fig. 19.2). Here, around E14.5, they gather to form a layer several cells thick called the Purkinje cell plate (Miale and Sidman, 1961; Altman and Bayer, 1997; Miyata et al., 2010). The radial migration of PCs is supported by the physical scaffold provided by radial glia (Altman and Bayer, 1997). This migration step is positively regulated by and dependent on reelin signals secreted initially by RL-derived CN neurons in the nuclear transitory zone (NTZ) and later by GNPs from the EGL (Miyata et al., 1996; Schiffmann et al., 1997). Migrating PCs maintain close contact with the radial glia, which is mediated by several adhesion molecules (Yuasa et al., 1991; Yuasa, 1996; Bentivoglio and Mazzarello, 1999; Hatten, 1999; Redies et al., 2011). A recent study described a subpopulation of PCs originating at E10.5 from the most posterior periventricular part of the lateral cerebellum that initially follow a rostral tangential migration path, with an elongated morphology, through the PWM to interact only with Bergmann glia in the later phase. This initial tangential path was not affected in a reelin mutant (Miyata et al., 2010). Successive waves of PCs accumulate through the process of radial migration to form the Purkinje plate. After birth, together with the expansion of the cerebellar surface due to the intense proliferation in the EGL, the plate gradually evolves into the final monolayer arrangement, which is achieved by the end of the first postnatal week in rodents (Altman and Bayer, 1997) (Fig. 19.2). Although this arrangement has been assumed to be a consequence of mechanical constraint (Altman and Bayer, 1997), it is likely to be influenced by other factors, such as the expression of reelin by the EGL (Miyata et al., 1997; Magdaleno et al., 2002; Jensen et al., 2004), and there are indications that the contact with the molecular layer also prevents the movement of PCs and preserves their correct alignment (Carletti et al., 2008). Finally, PC dendrites develop in a well-defined sequence over the first few weeks of postnatal life under the influence of signals secreted by granule cells (Altman and Bayer, 1997; Sotelo, 2004). There is also evidence that the glial processes of Bergmann glia provide physical and trophic support, as Purkinje dendrites are in tight contact with the radially oriented Bergmann fibers (Yamada et al., 2000; Lordkipanidze and Dunaevsky, 2005; Ango et al., 2008). Both the fibers and the dendritic tree are oriented along the parasagittal axis of the cerebellum, acting as palisades that partition bundles of parallel fibers in the molecular layer (Palay and Chan-Palay, 1974). The adult cerebellum displays a certain degree of compartmentalization, with a pattern of transverse zones and parasagittal stripes, that is believed to support the integration and processing of information (reviewed in (Sillitoe et al., 2005; Marzban and Hawkes, 2011)). This compartmentalization is revealed by intrinsic differences between subsets of PCs, such as the expression patterns of zebrin II in the PCs of postnatal mice (Brochu et al., 1990). This specific distribution appears to reflect an early subtype specification in the VZ; PCs born between E10 and E11.5 are destined to become zebrin II positive, whereas PCs born later (E11.5 to E13.5) belong to zebrin IIenegative stripes (Larouche and Hawkes, 2006). To date, however, little is known about the molecular determinants of this pattern because clonal analysis has revealed that the same PC progenitor contributes to both stripes (Larouche and Hawkes, 2006), and the only mutation to profoundly disturb these patterns is the deletion of the bHLH transcription factor Ebf2 (Croci et al., 2006). Nevertheless, there is some evidence of a molecular lineage (Chung, Sillitoe et al., 2009). It could also be interesting to study whether there is any relation to the early born PCs that follow tangential migration (Miyata et al., 2010). The implications of this compartmentalization of PCs in stripes lie beyond the scope of this chapter; nevertheless, it is worth mentioning that this pattern directly influences the distribution of interneurons and their dendritogenesis (Consalez and Hawkes, 2012; Leto et al., 2016), as well as UBCs (Chung, Marzban et al., 2009).

19.1.1.3 Migration of minor ventricular zone derivatives: Pax2-positive interneurons, basket cells, golgi cells, and stellate cells GABAergic interneurons comprise multiple subsets of morphologically and neurochemically distinct phenotypes that integrate in the cerebellar cortex and the DCN. In addition to GABA, most interneurons in the CN and granular layer also use glycine as a neurotransmitter, and a few are exclusively glycinergic (Simat et al., 2007; Husson et al., 2014). These cells are produced from a secondary proliferative zone in the PWM from late embryonic life to the second postnatal week (Weisheit et al., 2006). This pool of secondary progenitors of inhibitory interneurons has been identified as a population of Pax2-positive cells that are born from the VZ around E12, delaminate, and later migrate to populate the PWM (Zhang and Goldman, 1996; Maricich and Herrup, 1999; Weisheit et al., 2006; Ayala et al., 2007; Gardel et al., 2010) (Fig. 19.2). There, they produce the various interneuron phenotypes according to an inside-out progression (see above). Sonic hedgehog from the choroid plexus contributes to expanding the early generated pool of GABAergic interneurons (De Luca, Parmigiani et al., 2015). As they migrate inwards into the PWM, this source of Shh is replaced by the PC layer lying above, which continues to stimulate proliferation (Fleming et al., 2013).

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As mentioned above, the different classes of GABAergic interneurons can be distinguished by their “birthdate.” Nevertheless, data suggest that they are all derived from a single population of Pax2þ intermediate progenitors and that their morphological differences are established under the influence of local cues provided by the microenvironment in which they eventually settle (Ayala et al., 2007; Gardel et al., 2010). Surprisingly, little is known about the migration of postmitotic interneurons from the PWM to the different layers of the cerebellar cortex. It has been proposed that interneurons in the cerebral cortex are dispersed through contact inhibition (Magyar-Lehmann, Suter et al., 1995). If a similar mechanism operates in the cerebellum, that could explain, at least in part, the progressive dispersal of interneurons to more superficial layers in conjunction with cerebellar growth. More details have now been obtained concerning the migration of molecular layer interneurons. Young stellate and basket cells migrate radially to the interface between the molecular layer and EGL and become progressively integrated into the forming molecular layer (Yamanaka et al., 2004). Direct apposition of migrating interneurons to Bergmann glia has been observed in vivo and would be consistent with the Bergmann glia guiding migration (Guijarro et al., 2006; Simat et al., 2007). However, time-lapse microscopy has revealed more complex migration behavior, with successive steps of radial and tangential migration (Cameron et al., 2009), raising the possibility that the axons of granule cells also act as a substrate of migration (Cameron et al., 2009). Whereas basket cells axons develop along PC dendrites under the influence of the cell adhesion protein neurofascin 186 (Ango et al., 2004), stellate axons and dendrites grow along the Bergmann glia scaffold, an interaction mediated by Chl1 (Ango et al., 2004).

19.1.1.4 Migration of precerebellar nuclei The cerebellum receives afferents mainly from the MF and CF systems that originate mostly from precerebellar nuclei in the hindbrain. MFs originate from the basal pontine nucleus (BPN) (also known as the pontine gray nucleus) and the reticulotegmental nucleus (RTN), the external cuneate nucleus, and the lateral reticular nucleus (plus some from the spinal trigeminal nuclei and Clarke’s column in the dorsal spinal cord) (Altman and Bayer, 1987), whereas CFs originate from the inferior olivary nucleus (ION) that sends glutamatergic inputs to PCs (Ruigrok et al., 1995). Both systems also send branches to the CN (Fig. 19.1). All MF neurons originate from a high-Wnt1, Atoh1-positive progenitor domain in the posterior part of the lower RL (Ben-Arie et al., 2000; Rodriguez and Dymecki, 2000; Landsberg et al., 2005; Ozdamar et al., 2005) that spans the region from rhombomere6 to pseudo-rhombomere8 in the mouse (Di Meglio et al., 2013). However, CF neurons have their origin in a low-Wnt1, Ptf1a-positive domain (Landsberg et al., 2005) that is slightly ventral to the lower RL (Hoshino et al., 2005). Overall, these neurons all come from dorsal domains and will ultimately take one of three paths of migration (Fig. 19.3A). Inferior olivary nucleus neurons are born from E9.5 to E11.5 (Pierce, 1973), and it has been observed that a mutation in Ptf1a also prevents the formation of the ION (Hoshino et al., 2005). Neurons follow an intramural circumferential pathway within the medulla toward the ventral hindbrain, where their cell bodies stop before crossing the floor plate, while their axons cross the midline to project to the contralateral cerebellum (Altman and Bayer, 1987) (Fig. 19.3B). The migration of ION neurons requires netrin-1 signaling. Ntn-1 mutant showed aberrant formation and ventral reduction of the ION, which also failed to send out contralateral projections (Bloch-Gallego et al., 1999). Neurons of the external cuneate nucleus and lateral reticular nucleus, born from E10.5 to E12.5 and from E11.5 to E12.5 respectively (Ozdamar et al., 2005), delaminate from the RL around E13.5 and migrate tangentially toward the ventral midline, forming the posterior extramural stream (Altman and Bayer, 1987) (Fig. 19.3B). There is evidence that these neurons cross the midline around E15.5 and start migrating dorsally to the contralateral side, finally reaching their prospective location around E17.5 (Okada et al., 2007). Here, they switch from tangential to radial migration and enter the superficial hindbrain (Kawauchi et al., 2006; Okada et al., 2007) (Fig. 19.3B). The BPN and RTN neurons form the anterior extramural stream (AES) that gives rise to the future pontine nuclei. These neurons migrate anteriorly and ventrally toward the ventral midline, but most neurons remain on the ipsilateral side. The RTN neurons are generated from E12.5 to E13.5 (Ozdamar et al., 2005), and upon reaching the prospective pontine territory around E14.5, they initiate a radial migration, leaving the surface of the ventral hindbrain (Kawauchi et al., 2006; Okada et al., 2007). The BPN neurons are born from E13.5 to E16.5 and also follow the AES (Ozdamar et al., 2005). These neurons remain on the surface and pile up in an inside-out fashion (Altman and Bayer, 1987). Both populations also exhibit some degree of lateral migration (Shinohara et al., 2013). Interestingly, the rhombomeric patterning of the progenitors, which translates into differential Hox gene expression, is conserved during the migration process and translates into rostrocaudal patterning in the pontine nuclei (Di Meglio et al., 2013). This stream of migrating neurons is guided under the influence of a ventral netrin-1 attractive signal from the ventral midline (Yee et al., 1999) and by precocious ventral attraction that drives the AES anteriorly and is somehow prevented by Robo-Slit signaling (Geisen et al., 2008). A recent study suggested that Robo-Slit acts here in an uncharacterized non-cell-autonomous way (Dominici et al., 2018).

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FIGURE 19.3 The precerebellar nuclei origin and migration. Sagittal view of the developing hindbrain depicting the anterior (AES) and posterior extramural stream (PES) emanating from the Atoh1 positive rhombic lip (RL). Neurons in the AES migrate anteriorly and ventrally to form the reticulotegmental nucleus (RTN) and the basal pontine nucleus (BPN). (A) Upon reaching the midline cells switch to a radial migration and form the pontine nuclei, a limited number of neurons cross the midline to settle in the contralateral nuclei. Neurons in the PES migrate ventrally, cross the midline to settle on the contralateral side. (B) Upon reaching their respective presumptive positions, cells initiate a switch to radial migration to form the external cuneate nucleus (ECN) and lateral reticular nucleus (LRN). Cells originating from the Ptf1a lineage follow the inferior olivary migratory stream to settle ventrally and form the inferior olivary nucleus. On the right, a timeline illustrates the Atoh1 and Ptf1a lineage and the corresponding birthdates.

It is interesting to note that the loss of Ptf1a resulted in the loss of the ION and a fate change of CF neurons to MF neurons, as shown by the expression of Mbh2 (Ooshio et al., 2007). In the cerebellum, the expression of Ptf1a versus Atoh1 appears to be the main determinant of the GABAergic versus glutamatergic phenotype; however, in the precerebellar system, Ptf1a-positive neurons of the ION are glutamatergic. This suggests that Ptf1a versus Atoh1 is not the only determinant and that additional factors, such as Lbx1 and Olig3 are involved (Ooshio et al., 2007; Storm et al., 2009). Nevertheless, it is interesting that CFs that are Ptf1a positive mostly project onto PCs that are also Ptf1a positive, whereas MFs, which derive from Atoh1-positive progenitors, project onto CGNs that are also of Atoh1 origin. This suggests that these transcription factors regulate a matching identity, potentially through homophilic adhesion, that is useful for targeting in circuit building.

19.1.1.5 Migration of upper rhombic lip derivatives 19.1.1.5.1 Deep cerebellar nuclei The CNs develop in parallel with the cerebellar cortex and are important as they constitute the principal output of the cerebellum toward rostral and caudal structures (from lateral and medial nuclei, respectively). They receive some afferents from the CFs and MFs, but their principal afferents come from the PCs of the cerebellar cortex (Fig. 19.1). As described earlier, deep cerebellar nuclei are composed of glutamatergic and GABAergic neurons that arise, respectively, from the RL and the ventral VZ. Both populations are generated quite early in development, from E10.5 to E12.5, while the cerebellar anlage is still relatively thin (Fig. 19.2). Glutamatergic neurons delaminate from the RL and move rostrally along a subpial stream, following what is described as nongliophylic tangential migration (Machold and Fishell, 2005; Fink et al., 2006). Upon reaching the anterior boundary of the cerebellum, the cells condense into a NTZ. There, they are segregated into a series of mediolaterally distributed nuclei as they move inward. This inward motion is thought to occur through passive

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displacement due to the expansion of the cerebellar cortex with the arrival of newborn GNPs or through active migration (Altman and Bayer, 1985). RL-derived CN neurons are characterized by a transcriptional expression sequence that influences their lateromedial distribution in the forming CNs (Fink et al., 2006; Morales and Hatten, 2006). Briefly, the first neurons to be born express Lhx9 and migrate more laterally, followed by Tbr1-positive neurons that migrate more medially (to the fastigial nucleus). This progressive distribution from lateromedial to more dorsal positions is believed to reflect a decreased attractive sensitivity to the ventrally guidance molecule netrin-1 (Alcantara et al., 2000; Gilthorpe et al., 2002). However, the migration of GABAergic neurons contributing to the CNs is poorly described, and as they are born from the most ventral region of the cerebellar VZ, they may simply migrate from the PWM into the forming CNs as they move ventrally. 19.1.1.5.2 Granule neuron progenitors and cerebellar granule neurons Similar to the CN glutamatergic neurons, GNPs are produced in and delaminated from the RL and then undergo anterior subpial migration. Unc5c/netrin-1 signaling mediates a repulsive signal that prevents ectopic rostral migration (Przyborski et al., 1998). As they migrate, GNPs maintain expression of Atoh1, as well as Meis1 and Pax6 (Jereb et al., 2018). The GNPs spread evenly on the surface of the cerebellum to form a uniform layer, the EGL, at the surface of the cerebellar anlage (Fig. 19.2). There, they constitute a secondary GZ, where they continue to divide with peak proliferation until the end of the first week of postnatal development. This proliferation is sustained under the influence of mitogenic signals such as Shh secreted by the underlying PCs (Dahmane and Ruiz i Altaba, 1999; Nicot et al., 2002; Lewis et al., 2004; Choi et al., 2005). During amplification and after subsequent migrations, granule cells that are clonally related can spread over relatively long distances, with a preference for the anteroposterior versus the mediolateral axis. This preference probably contributes to the massive postnatal expansion of the cerebellum on the anteroposterior axis (Legue et al., 2015). Nevertheless, this anteroposterior expansion is limited by anchoring centers from Bergmann glia in the future grooves of the folia (see below), and this may contribute to the folding of the cerebellar surface (Legue et al., 2015). However, mediolateral expansion stems mostly from postmitotic tangential migration, similar to that in the proposed chick model (Ryder and Cepko, 1994). The diversity of migratory behavior of GNPs and CGNs after the formation of the EGL has been well characterized and will be described in further detail below to illustrate how this migratory behavior varies upon differentiation. 19.1.1.5.3 Unipolar brush cells UBCs are glutamatergic interneurons characterized by a single thick dendrite that ends in a “brush” of dendrioles (Harris et al., 1993; Mugnaini et al., 1994). At least three subtypes of UBC have been described based on their immunoreactivity for calretinin or mGluR1a and phospholipase C beta 4 (Nunzi et al., 2002; Chung et al., 2009). In the cerebellum, UBCs are unevenly distributed along the anteroposterior axis and are most abundant in lobules IX and X, which are associated with vestibular functions (Dino et al., 1999; Sekerkova et al., 2014). The precise origin of UBCs was debated until recently when, by using Tbr2 as a marker, Englund and colleagues precisely located the origin of UBCs within the core of the RL at E13.5 (Englund et al., 2006). The UBC progenitor pool is different from the pool providing GNPs, and these cells proliferate in the rhombic lip until birth (Englund et al., 2006). Most UBC progenitors express Wnt1 early in development, but the expression is downregulated before the UBCs leave the RL (Hagan and Zervas, 2012). As a consequence of their more medial location in the RL, their migratory route differs from that of the GNPs and CN neurons (Fig. 19.2). They migrate out of the RL via two streams: a dorsal pathway whereby the cell disperse into the forming cerebellar white matter, avoiding the future mass of CNs before invading the granular layer, and a rostral pathway toward the brainstem along the fourth ventricle (Englund et al., 2006). Most UBCs reach the granular layer by P10.

19.1.2 Part II. The cytoskeletal organization of cerebellar granule neurons 19.1.2.1 Cerebellar granule neuron migration diversity after the establishment of the secondary germinal zone The unraveling of the full scope of the intricate developmental migrations of rhombic lipederived neurons has required the use of cutting-edge intersectional lineage tracing and complex genetic gain- or loss-of-function studies and represents the newest addition to our global understanding of how cell migrations build the cerebellar system. However, the study of the so-called radial migrations of CGNs has deep historical roots in the developmental neurobiology field. Radial migrations and their contributions to CNS lamination were postulated as early as the work of the anatomists Wilhelm His,

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Albert von Kölliker, and Gieseppe Magini, who were credited with many of the pioneering histological examinations of developing brain tissue and theories on the nature of neuronal positioning during lamination (Bentivoglio and Mazzarello, 1999). Contemporaries of these early histologists focused on the cerebellar cortex proper and the CGNs because the simple laminar structure, the apparently lower cell diversity, and the relative abundance of CGNs in the cortex were deemed advantageous for descriptive studies at that time. Among these early investigations, the Golgi impregnation studies of Santiago Ramón y Cajal set the stage for our modern understanding of how CGNs transition from the EGL secondary GZ to a final destination in the IGL (Ramon y Cajal, 1911). Ramón y Cajal noted that simple, rounded cells in the EGL transformed into bipolar-shaped cells that were subtly displaced at various depths from the cerebellar surface. He also speculated that these cells might be initially displaced on an axis parallel to the surface and noted that bipolar cells then sprouted a third process oriented perpendicular to the cerebellar surface, which he presciently labeled a leading process because the “protoplasm and nucleus” of the CGN was eventually similarly displaced perpendicular to its long, parallel-fiber axons before acquiring vertical bipolarity and appearing to descend to positions in the IGL while trailing an axonal process. The cell bodies of displaced CGNs then acquired dendritic projections as they appeared to fully mature. Surprisingly, the state of understanding of how CGNs became displaced from their secondary GZ niche remained frozen for many years after Ramón y Cajal’s work. The advent of advanced autoradiographic techniques in the 1960s confirmed that radial migrations occurred and extended the concept that displacement only took place after the final division of proliferative cells. However, the field remained divided on the question of how cells accomplished this task, with one group holding that nuclei and cytoplasm were transported in preformed processes to accomplish displacement (similar to interkinetic nuclear migration (Sauer, 1935; Sauer and Walker, 1959) in the cortical VZ (Morest, 1970)) and another group holding that neuroblasts physically migrated along cellular elements, such as glial fibers or axonal tracts, to their final laminar positions. Eventually, a series of microscopy, tissue culture, and other technological advances enabled significant progress that bolstered the arguments in favor of radial migration along glial fibers. First, Pasko Rakic applied a combination of Golgi staining and electron microscopy (EM) to the developing macaque cerebellum to address his previous puzzling EM observation that CGNs in human cerebellar tissue could traverse the molecular layer, which was, in reality, a thicket of previously differentiated CGN parallel fibers (Rakic and Sidman, 1970). This unique application of Golgi staining to highlight the silhouettes of individual cells in densely packed tissue revealed that the polarized cell bodies and thickened leading processes engorged with cytoplasmic organelles of nearly every descending CGN were in close association with at least one and sometimes multiple Bergmann glial fibers and that, intriguingly, the Bergmann glial cell membrane in the vicinity of CGNs was morphologically specialized in comparison to membrane regions free of such close contact (Rakic, 1971). Second, Mary Beth Hatten established purification methods for various types of cerebellar cell and recombined pure CGNs with cerebellar glia to generate the first reconstituted model to permit direct observation of migration events in the brains of developing mammals (Hatten 1984, 1985). The use of high-resolution Nomarski optics revealed that CGNs are not only closely associated with cerebellar glia, directly migrating along their fibers, but can also navigate past stalled CGNs or from one glial fiber to another (Edmondson and Hatten, 1987) (Fig. 19.4A). In addition, these studies hinted at aspects of cell biology that could not be imagined from static histological EM micrographs, e.g., that glialguided migration occurs in a saltatory fashion whereby pauses along the glial fiber punctuate movement, that the motility of the tip of the leading process and the CGN cell body occur independently, and that the polarized leading process undergoes apparent contractions before cell body movement. Komuro and Rakic further filled in details of CGN migration diversity in more physiologically relevant milieus by using lipophilic dyes to label individual cells in acute cerebellar slices and following their migration using confocal laser scanning microscopy (Komuro and Rakic 1993, 1995, 1998; Komuro et al., 2001). These studies confirmed that a saltatory migration cadence occurred in all stages of CGN migration in a tissue slice, and the tangential migration of newly differentiated CGNs that was highlighted by the retroviral labeling studies of Cepko and colleagues (Ryder and Cepko, 1994). Moreover, these studies revealed several surprising new aspects of CGN motility: (1) tangentially migrating CGNs in cerebellar hemispheres are attracted to midline structures; (2) CGNs accelerate their migration across a thickening molecular layer to ensure a constant transit time for this layer during development; (3) CGNs pause to round up their leading process and cell body at defined layer boundaries; and (4) CGNs migrate long distances in the IGL in the absence of glial guides. Taken together, these studies filled in and dramatically extended Ramón y Cajal’s initial vision for the movement of CGNs to their final destination. The unexpected findings prompted new hypotheses regarding the reasons for the saltatory nature of the migration (addressed in the remainder of Part II of this chapter) and how polarized morphologies were linked to the timing of CGN maturation or the timing of changes in migratory behavior, such as the shift from tangential to radial migration (addressed in Part III).

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FIGURE 19.4 CGN glial-guided migration. (A) Dynamics of CGN migration along a glial fiber in an in vitro microculture. Note the CGN cell body and leading process are always juxtaposed to the underlying glia fiber. lp, leading process. (B) Schematic representation of the two-stroke motility cycle. The tan CGN maintains close contact with the gray glial fiber at all phases on the movement cycle (red springs ¼ neuron-glial adhesions). In Phase I, the centrosome (blue circle) is located in the cell body and the nucleus (gray outline) is located in the rear of the cell body. In Phase IIs, the centrosome moves into the leading process. In Phase III, the cell body and nucleus catch up to the centrosome as the CGN inches forward along the glial guide. (A) Figure adapted with permission from a time lapse video from Edmondson, J.C., Hatten, M.E., Glial-guided granule neuron migration in vitro: a highresolution time-lapse video microscopic study. J. Neurosci. 1987;7(6):1928e1934.

19.1.2.2 The road to the two-stroke motility paradigm Perhaps the most exciting and surprising finding of Edmondson and Hatten’s time-lapse microscopy analysis of migration was the observation that during migration periods, CGN cell body advancement was saltatory, with a cyclic shift from movement to rest phases occurring over the course of the migration, and that cell body movement was asynchronous with the motile tip of the leading process (Edmondson and Hatten, 1987). Moreover, high-resolution Nomarski optics revealed that intracellular vesicles within migrating granule neurons flowed into the leading process with similar periodicity to the overall movement cycle, whereas in stationary neurons, this flow halted with the cessation of migration. The application of EM imaging after time-lapse microscopy by Gregory and colleagues revealed a deeper specialization of the leading process by showing that organelles such as the Golgi apparatus or the centrosome microtubule organizing center, as well as a higher abundance of cytoskeletal polymers such as microtubules and microfilaments, were specific to the leading process (Gregory et al., 1988). Taken together, the results of these studies suggested, for the first time, that migrating CGNs were highly polarized with respect to their subcellular organization and that the saltatory motion cycle was a cell biological entry point to unravel the molecular mechanisms governing neuronal motility.

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The excitement regarding the discovery of the saltatory motion was buttressed by a series of ex vivo imaging studies that showed saltatory motion in neurons within various migration pathways and brain regions, including radially migrating excitatory neurons and tangentially migrating interneurons (O’Rourke et al., 1992; Komuro and Rakic, 1993; Komuro and Rakic, 1995; Komuro and Rakic, 1998; Komuro et al., 2001; Nadarajah et al., 2001; Nadarajah et al., 2002; Nadarajah et al., 2003). These time-lapse experiments showed that migrating neurons extend a thin leading process in the direction of migration, that the nucleus remains in the rear of the neuronal cell body during migration, that extensive interstitial adhesion junctions form between glial cells and neurons, and that nuclear/cell body movement occurs in a saltatory series of steps called nucleokinesis that is accompanied by the release of the adhesion site followed by forward movement of the cell body. These findings spurred active research in the tractable CGN model system to uncover the source and substrate of saltatory motion or nucleokinesis. Komuro and Rakic noted that calcium oscillations occurred near the time of cell body advance (Komuro and Rakic, 1996). Rivas and Hatten used single-cell labeling techniques to discern that the leading process was contractile, whereas the thin axonal trailing processes were markedly less dynamic (Rivas and Hatten, 1995). Kawaji and colleagues noted that the migration dynamics could be linked to the leading process type (Kawaji et al., 2004). The leading processes of tangentially migrating CGNs possessed axonal characteristics that correlated with discontinuous movement between the leading-process tip and cell body, with enhanced rates of cell displacement. In contrast, the leading processes of radially migrating CGNs possessed dendritic characteristics that correlated with the synchronized advancement of the leading-process tip and cell body, with lower rates of overall displacement. Despite the intriguing nature of these findings, their contribution to the understanding of saltatory motion was mainly correlative. A more advanced framework for understanding saltatory motion and nucleokinesis was fashioned by dynamic imaging of the large organelles that were localized in a polarized fashion in the leading process, as reported by Gregory and colleagues (Gregory et al., 1988). Labeling of the CGN centrosome with genetically encoded fluorescent proteins confirmed the findings of the limited number of EM observations that centrosomes remain just forward of the nucleus in the leading process. Surprisingly, however, high-resolution spinning-disk confocal microscopy showed that this organelle displayed “two-stroke” motility in the direction of its movement along the glial guide, predominantly before the forward movement of the nucleus (Solecki et al., 2004) (Fig. 19.4B). Interestingly, this peculiar motility was observed only in migratory CGNs, as the centrosomes of stationary neurons were either stalled or followed a random orbit within the cell body. Although connections between nuclear and centrosome positioning have been described in invertebrate model systems (Reinsch and Gonczy, 1998; Mosley-Bishop et al., 1999; Malone et al., 2003; Archambault and Pinson, 2010; Burakov and Nadezhdina, 2013) and the centrosome is mildly attached to the nucleus in homogenates of mammalian cells grown in culture (Nadezhdina et al., 1979; Mitchison and Kirschner, 1984), a distinctive feature of mammalian neurons is that the centrosomes are not physically tethered at the nuclear membrane and must use complex cell biological mechanisms to ensure station-keeping between organelles that can be separated by tens of microns (Bellion et al., 2005; Tsai et al., 2007; Umeshima et al., 2007). Not long after the initial reports of the saltatory advance of centrosomes appeared, a similar configuration for Golgi positioning was proposed, and this was ultimately confirmed by long-term time-lapse imaging of fluorescently labeled Golgi proteins in in vitro cultures and ex vivo slice preparations (Bellion et al., 2005). Like saltatory motion itself, two-stroke motility has been observed in the major migration models in the mouse brain (Bellion et al., 2005; Higginbotham et al., 2006; Tsai et al., 2007; Renaud et al., 2008; Hao et al., 2009; Baudoin et al., 2012; Shinohara et al., 2012; Hutchins and Wray, 2014; Silva et al., 2018) and in migrating neurons purified from developing human brain tissue (Ostrem et al., 2014). Thus, two-stroke motility has become a framework by which to dissect the basic mechanisms of how neurons move to their final position, as the dynamics of motility have been found to be perturbed in many models of diseases in which lamination is affected. The next three sections will consider the contributions of the two main cytoskeletal polymers (microtubules and actin), their molecular motors, and possible points of integration during the two-stroke motility cycle.

19.1.2.3 The roles of the microtubule cytoskeleton and associated motors Microtubules are cylindrical polymers of the a- and b-tubulin proteins that form a cytoskeletal system providing structural support in the cell cytoplasm, as well as an architectural network by which molecular machines called motor proteins can perform physical tasks (e.g., segregation of chromosomes during mitosis or material and organelle transport) (Watanabe et al., 2005; Etienne-Manneville, 2013; Borisy et al., 2016). Although EM analyses noted microtubules in the CGN leading process in vivo and after correlated light and EM microscopy (Rakic, 1971; Gregory et al., 1988), the acquisition of an overall picture of the microtubule cytoskeleton in migrating CGNs had to await the development of improved microtubule labeling techniques. Tubulin antibody staining revealed that bundled microtubules are located in a central region of the leading process and in a “cage-like web of filaments” surrounding the CGN nucleus (Rivas and Hatten, 1995) (Fig. 19.5A) that sometimes appears as a forklike microtubule structure, depending on the method of fixation used and/or the neuronal

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FIGURE 19.5 The CGN microtubule cytoskeleton and the roles of microtubule motors. (A) Tubulin antibody staining reveals a cage-like lattice of microtubules that surrounds the CGN nucleus. Inset: shows a higher magnification view of the CGN cage. (B) Dynamics of the CGN microtubule cytoskeleton. A Venus-Tubulin construct was introduced into CGNs and high-speed time-lapse imaging used to examine morphological changes. In Phase I the cage is compact, in Phase II it stretches, and in Phase III the cage is deformed as the nucleus advances in the cell body. lp, leading process. (C) Diagram of the role of microtubule motors in CGN migration. Increased expression of KIF11 prevents microtubule slide to lock CGNs in a stationary phase of migration. In Phase II of the CGN migration cycle, cytoplasmic dynein motors in the leading process help guide the centrosome into the leading process. In Phase III of the cycle, nuclear associated cytoplasmic dynein motors transport the nucleus forward to the centrosome while kinesin motors provide additional force to rotate the nucleus during forward movement. (A) and (B) adapted with permission from Solecki D.J., Model, L., Gaetz, J., Kapoor, T.M., Hatten, M.E., Par6alpha signaling controls glial-guided neuronal migration. Nat. Neurosci. 2004;7(11):1195e1203).

cell type (Xie et al., 2003). Labeling microtubules with a genetically encoded fluorescent protein tag highlighted alterations to the microtubule cage during two-stroke motility: During forward movement, perinuclear microtubules undergo dynamic shape fluctuations, but they continually encircle the nucleus as the cage and nucleus move as a unit (Solecki et al., 2004). Just before cell body translocation, the cage stretches and a group of microtubules physically enter the proximal leading process (Fig. 19.5B). During translocation, the microtubules in the cage are compressed because of the forward nuclear movement, resulting in a compact profile that perdures until the next saltatory round of two-stroke motility. Umeshima et al. confirmed that the nuclei of CGNs migrating in ex vivo slice preparations are surrounded by a cage of microtubules (Umeshima et al., 2007). They further showed that the cage is composed of different populations of microtubules: (1) a tyrosinated population representing the significant cage fraction that is unstable and (2) an acetylated population around the anterior surface of the nucleus that is stable. Surprisingly, acetylated microtubules extended from the nucleus toward the proximal leading process and were not associated with the centrosomes as previously thought. The dissolution of these microtubule-based structures by pharmacological manipulation with drugs such as nocodazole or colchicine blocks the movement of neurons throughout the brain by destabilizing their leading and trailing processes and ultimately perturbs two-stroke motility, illustrating the importance of the microtubule-based cytoskeleton for neuronal migration.

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Insights gained from the positional cloning of genes in which mutations cause the human neuronal migration disorder called lissencephaly and from their subsequent genetic manipulation in mice have provided a compelling entry point through which to understand the roles of microtubules and some microtubule motor proteins in neuronal migration during brain lamination (Gleeson and Walsh, 2000; Ross and Walsh, 2001; Kato and Dobyns, 2003; Metin et al., 2008). Isolated lissencephaly syndrome and MillereDieker syndrome are rare brain disorders characterized by a smooth-appearing brain with defective lamination, intractable epilepsy, and apparent defects in neuronal migration that are associated with spontaneous deletions in the vicinity of the PAFAH1B1 gene on the short arm of chromosome 17 (Dobyns et al., 1993; Reiner et al., 1993). PAFAH1B1 encodes the Lis1 protein, which is an evolutionarily conserved, essential cofactor for the cytoplasmic dynein minus endedirected microtubule motor protein (Morris et al., 1998; Efimov and Morris, 2000; Faulkner et al., 2000; Smith et al., 2000; Morris, 2003; Shu et al., 2004; Tsai et al., 2005; Tsai et al., 2007). The genetic deletion of PAFAH1B1 (Hirotsune et al., 1998) or a series of genetically and biochemically interacting proteins (Umeshima et al., 2007) leads not only to profound inhibition of all microtubule minus endedirected dynein transport but also to CGN migration defects, including stalled motility of CGNs in microculture and in their movement to the IGL in vivo. Interestingly, the loss of Lis1 function perturbed two-stroke nucleokinesis in both CGNs and cortical pyramidal neurons in a unique fashion: in both cell types, nuclear displacement was considerably slowed and the progressive centrosome/cell body advance was affected, although centrosome movement was randomized, as opposed to being halted completely (Tsai et al., 2007; Umeshima et al., 2007). These results led to the formulation of a model of a dual role for dynein activity, whereby dynein is tethered at the cell cortex in the leading process and, as in the yeast model system (Dujardin and Vallee, 2002), helps guide the centrosome forward in the first stage of the two-stroke cycle, whereas dynein on the nuclear envelope serves to move the nucleus toward the centrosome in the final stage of the two-stroke nucleokinesis model (Tsai et al., 2007; Vallee et al., 2009) (Fig. 19.5C). In contrast to the single dynein motor that moves cargos in a minus endedirected manner along cytoplasmic microtubules, scores of kinesins move cargos in the opposite direction to the plus end (Vallee et al., 2004; Hook and Vallee, 2006; Hirokawa et al., 2009; Vallee et al., 2009; Hirokawa et al., 2010; Bachmann and Straube, 2015). Given the assortment of kinesins that have been identified, it is not surprising that various kinesin-related functions concern the twostroke nucleokinesis system (Fig. 19.5C). For example, KIF11 is expressed in CGNs that have reached their final destination in the IGL in various cell biological models, including the mitotic spindle and extending axons. KIF11 is a slow kinesin motor that acts as an antagonist of dynein-dependent transport (Falnikar et al., 2011). Pharmacological inhibition of KIF11 and gene silencing stimulate CGN migration both in vitro and in ex vivo slices, potentially by stimulating microtubule sliding. This suggests that high KIF11 activity stalls migration once CGNs have reached the IGL. As a second example, KIF6 loss of function strongly diminished the directional persistence and boosted the randomized migration directions of CGNs while simultaneously stimulating the appearance of multiple leading processelike extensions, implicating the defective polarized leading-process morphology in random motility (Falnikar et al., 2013). Remarkably, KIF6 cooperates with the MgcRacGAP protein in CGNs as it does in the centralspindlin complex of cells undergoing cytokinesis, suggesting that the well-characterized cell biological mechanisms of the centralspindlin complex in cytokinesis are recycled for leading-process morphogenesis in migrating neurons. In a final example, kinesin motors cooperate with dynein to regulate the efficiency of the nucleokinesis phase of the two-stroke motility. Interestingly, Wu and colleagues discovered that the nuclei of migrating CGNs tend to rotate on an axis parallel to their migration direction before or during the last phase of the movement cycle, whereas stationary or postmigratory nuclei display no such rotation (Wu et al., 2018). Moreover, nuclear rotations were accompanied by the formations of “sharp peaks” at the leading edge of the nucleus, indicating that localized pulling forces were applied to the moving nuclei. The individual inhibition of dynein and KIF5B not only strongly affected nuclear rotation but also decreased the overall displacement of migrating CGNs and abrogated the apparent pulling forces applied to the leading edge of the nuclei, suggesting that coordination between plusand minus-end motors is required for the optimal application of force to the large neuronal nucleus during CGN motility. Taken together, the results of these studies implicate the motor proteins that drive microtubule-dependent organelle transport as playing key roles in driving the two-stroke motility of migrating CGNs.

19.1.2.4 The role of the actin cytoskeleton Actin is one of the most abundant proteins in eukaryotic cells. It polymerizes to form microfilaments that, along with microtubules, represent the second classical cytoskeletal structure required for a host of cellular activities (Pollard and Cooper, 2009; Gardel et al., 2010; Svitkina, 2018). The balance between soluble G-actin and polymerized F-actin, the cellular locations of actin polymerization, and the motor proteinedependent movement of microfilament-containing structures all play critical roles in cellular motility across cell types and species. Early EM studies noted that the CGN

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leading process was a site of microfilament enrichment. The first systematic analysis of actin distribution revealed that phalloidin-labeled microfilaments were located in a subcortical rim in the CGN cell body, with heavy staining over the length of the thickened migratory process (Rivas and Hatten, 1995). Although confocal microscopy lacks sufficient resolution to fully address the relation between microfilaments and microtubules, these initial studies found that the microfilaments appeared to be displaced closer to the plasma membrane. Remarkably, very few microfilaments were observed along the shaft of the CGN axonal process that did not display the contractile oscillations that occur in the leading migratory process during saltatory movement (Rivas and Hatten, 1995). Treatment with cytochalasin B, a pharmacological agent that dissolves microfilaments, simultaneously halted leading-process contractions and the saltatory advance of the neuronal cell body, providing the earliest evidence of the importance of the microfilament system in CGN migration along glial fibers. The following sections describe advanced genetic and cell biological studies that provide further mechanistic insights into how actin filaments propel neuronal motility. Active actin polymerization is thought to create protrusive forces that “push” the advance of the leading edges or tips of migratory cells (Pollard and Borisy, 2003; Ridley et al., 2003; Danuser and Waterman-Storer, 2006; Skau and Waterman, 2015). By a tour de force combination of biochemistry, cell biological studies, and genetics, the basis of actin-based protrusive activity has been conceptualized in what is dubbed the dendritic nucleation model (Zigmond, 1998; Goley and Welch, 2006; Pollard, 2007; Soderling, 2009). In brief, the activation of actin-related protein 2/3 (Arp2/3) by WiskotteAldrich syndromeerelated complex proteins (WASP, WAVE/SCAR, and WASH) leads to branched actin assembly of preexisting microfilaments. This, in turn, creates a host of growing, barbed actin filament ends that are synchronously oriented toward the periphery of the leading edge. The combined action of the sum of barbed-end growth progressively leads to leading-edge advancement. Biochemical regulators of this process include members of the Rho family of small guanosine triphosphatases (GTPases) (Rac, Cdc42, and RhoA), which interact with multiple target proteins that control the activity of Arp2/3, WASP-related complexes, and factors that bundle actin filaments into higher-order structures called stress fibers (Hall and Nobes, 2000; Ridley et al., 2003; Govek et al., 2011; Hall, 2012). The functions of the three main Rho GTPases (Rac, Cdc42, and RhoA) have been genetically examined in the CGN model system to elucidate the mechanistic impact of actin polymerization and organization on the movement of CGNs from the EGL to the IGL. Rac1 was the first GTPase to be examined in CGNs. It acts as a classic upstream regulator of Arp2/3 activity in lamellipodial structures by controlling the activity or localization of WASP and WAVE proteins (Tahirovic et al., 2010). The genetic deletion of Rac1 in CGNs leads to the accumulation of mature neurons in the molecular layer. Rac1-deficient CGNs are not only immotile and unable to extend the long leading process required for migration to the IGL; they also have mislocalized WASP and WAVE proteins. The subsequent genetic deletion of RhoA (Mulherkar et al., 2014), a key regulator of the contractile assembly of Arp2/3-polymerized actin filaments into stress fibers, also revealed critical features of actin regulation of CGNs. RhoA-deficient CGNs could not efficiently migrate to the IGL and showed cell-autonomous alterations in morphology that ultimately led to gross defects in cerebellar foliation, highlighting the importance of contractile-fiber assembly in CGNs (Mulherkar et al., 2014). Finally, the genetic deletion of Cdc42, the classic WASP/WAVE activator in filopodia structures, also perturbs CGN movement to the IGL (Govek et al., 2018). Interestingly, the loss of Cdc42 causes CGNs in the EGL to lose their pseudo-columnar arrangement and transform into a multipolar morphology that is misaligned to glial-guide migration, leading to a unique foliation defect with thinning of the vermis. Like microtubules motors, actin-based molecular motors play critical roles in cell migration. Perhaps the bestunderstood actin-based motor protein is myosin II, which creates contractile actin microfilament sliding just forward of the nucleus and at the rear of the cell to ensure steady forward motility in nonneuronal cells (Vicente-Manzanares et al., 2009; Ma and Adelstein, 2014). The CGN is a preeminent model system for understanding the role of actomyosin contractility in neuronal migration (Fig. 19.6). The use of time-lapse imaging of live CGNs migrating along glial fibers, combined with novel methods for analyzing real-time cytoskeletal dynamics, enabled an examination of the precise spatiotemporal coordination of actin dynamics during this specialized mode of migration. The proximal portion of the leading process is a region of high actin turnover, and during forward movement, there is a cycle of F-actin accumulation in which F-actin also flows from the proximal to the distal regions of the leading process in the direction of migration (Solecki et al., 2009; He et al., 2010) (Fig. 19.6A). Fluorescence recovery after photobleaching analysis also showed that the proximal portion of the leading process is the region of highest actomyosin turnover in migrating CGNs. Four-dimensional volumetric analysis and pulse-chase photoactivation experiments have shown that this flow represents an apparent transfer of actin between the cell body and the proximal leading process before cell body translocation. Myosin II motors are present along the length of the leading process, and oscillations in the myosin II fluorescence signal during the motility cycle suggest the occurrence of actomyosin contractile events in this region. Whereas the initial time-lapse imaging studies relied on partially correlative arguments based on the locations of the polymer or myosin II motor to determine the

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FIGURE 19.6 The CGN actin cytoskeleton and the roles of actomyosin contractility. (A) Dynamics of the CGN actin cytoskeleton its relationship to the centrosome. A Venus-Centrosome reporter (green) and an RFP-Actin reporter (red) were introduced into CGNs and high-speed time-lapse imaging used to examine morphological changes. Note the leading process is a site of major F-actin enrichment and the centrosome is located close to actinenriched domains in the leading process. lp, leading process. (B) Myosin heavy chain antibody staining reveals leading process accumulation of this critical actin-based motor. (C) Diagram of the actomyosin cytoskeleton throughout the CGN motility cycle. In Phase I the centrosome and actin is located in the cell body. In Phase II both centrosome and actin are shuttled down the leading process and actomyosin microtubule coupling are needed for centrosome, cilia, and surface receptor movements coupled to actomyosin flow. In Phase III the cell body advances due to contraction of leading process actomyosin.

locations of contractile force generation, recent examinations have been carried out with a traction-force imaging modality that directly reports the traction force in pico newtons, accompanied by an accurate assessment of the force vector (Jiang et al., 2015; Umeshima et al., 2018). These studies have revealed two significant areas of traction force in the leading process: (1) an area at the leading tip of the leading process, where force vectors extend away from a migrating CGN, that is associated with leading-process extension and (2) an area in the proximal portion of the leading process, just forward of the nucleus, that contracts during the cell body translocation phase of the two-stroke migratory cycle. The administration of cytoskeletal drugs that inhibit myosin II and F-actin flow had profound effects on CGN two-stroke motility. The application of blebbistatin, a myosin II ATPase inhibitor, or jasplakinolide, a drug that locks F-actin in a stable state, immediately halts the cell body/nuclear translocation phase of the movement, indicating that myosin II plays a pivotal role in the last phase of the movement cycle (Solecki et al., 2009). Given that myosin II had been reported to accumulate at the rear of migrating SVZa neurons and interneurons from the medial ganglionic eminence, further studies were needed to confirm that the leading process was the site of actomyosin contractility (Bellion et al., 2005), despite the dynamic live-cell localization measurements and the subsequent direct assessment of contractile force via traction-force microscopy. Focal

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blebbistatin or jasplakinolide application with a micropipette revealed that the inhibition of leading-process actomyosin activity halted migration, whereas inhibition in the CGN cell body or rear had no effect (He et al., 2010). Actomyosin inhibition also had unexpected effects on organelle positioning events before cell body translocation (Solecki et al., 2009; Trivedi et al., 2014). Interestingly, both myosin II and F-actin accumulate near centrosomes, primary cilia, and the Golgi, as actin-based flow occurs in the first phase of the two-stroke motility cycle. Remarkably, actomyosin inhibition halts the forward flux of microtubules, as well as the entry of the centrosome, primary cilia, and Golgi into the proximal leading process, thereby effectively arresting the first phase of the two-stroke motility cycle. The block in organelle movement was not due merely to a lack of cellular translocation because blebbistatin or jasplakinolide treatment also halted the nondirected movement of the centrosome, cilia, and Golgi. Thus, like cytoplasmic dynein, leading-process myosin II motors contracting actin microfilaments play a dual role in controlling movement in both phases of the two-stroke motility cycle. Moreover, actin flow represents an unappreciated vector of polarity in migrating CGNs. Besides organelle positioning, are any other functions associated with leading-process actomyosin flow? Based on their pioneering work, Bray and White proposed that the linkage of the inner face of the plasma membrane with actomyosin constituted an actin-rich cortex that flowed to control the localization of cell surface components and cytoskeletal coordination during cell migration (Bray and White, 1988). Consistent with this hypothesis, Wang and colleagues detected forward flow of neurotrophin receptors toward and through to the distal tip of the leading process of migrating neurons, but not of stationary neurons, that was halted by the addition of blebbistatin or jasplakinolide (Wang et al., 2012). Moreover, Trivedi and colleagues noted that adhesion receptors that mediate neuroneglial interaction were localized to leadingprocess domains with high actomyosin content, and they also found that actomyosin inhibition prevented their turnover or transport to new sites of cell contact (Trivedi et al., 2014) (Fig. 19.6C). Thus, not only does actomyosin control both phases of the two-stroke motility cycle, but the cortical flow driven by myosin II also regulates the appropriate positioning of cell surface receptor proteins within the leading-process plasma membrane.

19.1.2.5 The role of microtubule-actin cross talk Although the potential for microtubuleemicrofilament interactions in neuronal cell migration has been postulated since the earliest EM studies in the field (Rakic, 1971; Gregory et al., 1988), the assessment of the precise mode of such interactions has been slowed by the lag in applying cell biological tools to the primary nervous system models. One of the first wellcharacterized examples of a regulatory interaction between the microtubule and actin cytoskeletons came from a careful analysis of CGNs from animals that were haploinsufficient for the PAFAH1BI gene (e.g., lacking one PAFAH1BI allele, PAFAH1BIþ/) (Kholmanskikh et al., 2003). Unexpectedly, phalloidin staining revealed that PAFAH1BIþ/ CGNs possessed shorter neurites and had fewer actin-labeled structures in their leading process than did normal CGNs, and this was associated with diminished Rac and Cdc42 activity but elevated levels of RhoA activity. Surprisingly, RhoA inhibition normalized Rac and Cdc42 activity and migration to near wild-type levels, indicating that PAFAH1BIþ/ deficiency unexpectedly fed back onto the Rho GTPases that have been shown through genetic studies to be critical regulators of actin polymerization levels in migrating neurons. Further functional studies showed that Lis1 forms a complex with IQGAP1, a protein that maintains Rac and Cdc42 in the active state at the leading edges of fibroblasts and the tips of extending neurites, and Clip170, a protein that tethers microtubules to the leading edge (Kholmanskikh et al., 2006). Taken together, these results suggest that Lis1 maintains leading-process extension by boosting Rac1 and Cdc42 activity and maintains appropriate levels of actin polymerization through leading-edge IQGAP1 recruitment. At the same time, Lis1eClip170 interactions maintain appropriate levels of microtubule recruitment at the leading edge of growing neurites. Despite the definition of the individual contributions of the microtubule- and actin-based motor proteins to two-stroke motility, our knowledge of how the two systems function together in migrating neurons was fragmentary for an extended period. Pharmacological studies showing that the simultaneous inhibition of myosin II and cytoplasmic dynein caused a complete halt of all two-stroke motility parameters, as compared to the slowed movement with single inhibition, led to a search for a physical linkage between the microtubule and actin cytoskeletons in the proximal portion of the leading process (Trivedi et al., 2017). The drebrin protein that binds the sides of F-actin filaments and microtubule plus ends through direct interaction with the neuronal þ TIP protein end-binding protein 3 (EB3) proved a useful reporter for establishing the sites of microtubuleeactomyosin interaction in migrating CGNs, as it is enriched in these cells during their motile phase (Trivedi et al., 2017). Super-resolution microcopy examination of drebrin localization revealed that (1) drebrin dynamically localizes to the proximal leading process before or during cell body translocation in a myosin IIe dependent manner and (2) a layer of drebrin and F-actin intervenes between the plasma membrane and elements of the microtubule cytoskeleton during CGN migration, making drebrin the most specific reporter for anterograde leading-process actomyosin flow yet found (Fig. 19.6B). Inhibiting drebrin function or blocking drebrin-microtubule end binding resulted

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in a lack of microtubule advance into the proximal leading process, the randomization of the direction of centrosome and cell body motility, and ultimately, the prevention of CGN migration to the IGL. Thus, the physical interaction between microtubules and microfilaments driven by the leading-process flow of drebrin coordinates the activities of these two cytoskeletal elements to produce the saltatory, polarized movement of cytoplasmic organelles and the neuronal cell body seen in two-stroke migration.

19.1.3 Part III. The facets of cerebellar granule neuron polarity: timing cell recognition, differentiation, germinal zone exit, and morphogenesis Whereas examination of the cytoskeleton has clarified the molecular basis of the saltatory movement of CGNs along their glial guides, examination of how CGNs polarize during differentiation, migration, or morphological development has shed light on how they navigate the complex temporal transitions in their migratory behavior, such as the shift from tangential to radial migration, the exit of GNPs from the outer EGL, and the temporal control of polarized morphogenesis.

19.1.3.1 Cerebellar granule neuron recognition/adhesion: the contribution of astrotactins and the siah2ePard3eJamC pathway Early ultrastructural (Rakic, 1971) and correlated high-resolution time-lapse EM (Gregory et al., 1988) studies revealed that during CGN migration, CGNeBergman glial contacts were favored over CGNeparallel fiber interactions, thereby prompting a host of studies designed to understand how migrating neurons recognized or adhered explicitly to their appropriate migration substrates at the expense of other cellular elements (Rakic, 1985). Significant advances in our understanding of CGN recognition of glial substrates have come from the pioneering work on Astrotactins and the factors regulating junctional adhesion molecules (Fig. 19.7). Astrotactin1 (ASTN1), the first CGNeBergmann glial adhesion molecule containing EGF and complement-like perforin domains to be discovered (Fig. 19.7A), was isolated from a clever screen for antibodies against CGN surface antigens that blocked CGNeglial interactions and CGN motility in microcultures (Edmondson et al., 1988; Zheng, 1996). Immunostaining with anti-ASTN antibodies revealed a protein expressed on the CGN surface that is localized to sites of CGNeglial apposition (Fishell and Hatten, 1991). Gene targeting in mice showed that ASTN1 functions in CGN migratory events in vivo during the assembly of the cerebellum cortical layers (Adams NC, Tomoda T et al., 2002). Interestingly, ASTN1-driven CGNeglial contact plays critical roles in generating the polarized morphology of migrating CGNs, as ASTN1 loss of function not only blocks polarized leading-process formation but also disrupts the intricate polarization of the microtubule and microfilament cytoskeletons (Fishell and Hatten, 1991). A recent study using fluorescent-labeled ASTN1 and CGN live-cell imaging demonstrated that ASTN1 trafficking represents a new aspect of polarity in the saltatory movement of neurons along their glial guides. Wilson et al. found that during early CGN migration, when the neuron is presumably attached to the glial fiber, ASTN1-Venus was located in the leading aspect of the neuronal cell body, where a specialized “interstitial junction” had previously been observed in correlated time-lapse microscopy and EM studies (Wilson et al., 2010). As CGNs translocated along the glial fiber and the cell body junction was released, ASTN1Venus flowed from the cell body to the proximal domain of the leading process, where a new adhesion site formed (Fig. 19.7B). Wilson and colleagues also found that the newly identified Astrotactin2 (ASTN2) protein is not a neuron-glial adhesion molecule like its homolog ASTN1; instead, it functions in CGNeglial junction formation by forming a complex with ASTN1 to regulate ASTN1 cell surface recruitment, thus providing a trafficking-based mechanism that controls polarized ASTN1 flow in the leading process. Interestingly, ASTN2 appears to be a general regulator of membrane trafficking events critical for synaptic function as it controls the surface expression of multiple synaptic proteins; such as Neuroligins, the KCC2 potassium channel or Olfactomedins, by regulating their retrieval and intracellular degradation (Behesti et al., 2018). Patients with intellectual disabilities have mutations or deletions in the ASTN2 fibronectin type III domain which compromises ASTN2 receptor trafficking functions, suggesting that ASTNs are central players in neuronal developmental disorders affecting circuit function beyond the previously established role in neuronal migration (Behesti et al., 2018). Recent examination of the regulation of the evolutionarily conserved partitioning defective (Pard) complex has revealed intimate connections between cell polarity signaling and adhesive recognition control during the shift from tangential to radial migration (Famulski et al., 2010) (Fig. 19.8). The Pard complex, containing the PDZ domain adaptor proteins Pard3 and Pard6 plus atypical protein kinase C, is the best-characterized polarity signaling complex and is essential for tightjunction formation, cell migration, and axon morphogenesis. Early studies showed that Pard6 is required for organizing two-stroke motility and actomyosin regulation required for actin cytoskeletal components that coordinate CGN

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FIGURE 19.7 ASTN1 and JAM-C are major CGN recognition/adhesion molecules. (A) ASTN1 domain structure. ASTN1 has three membrane spanning domains, and its extracellular carboxy-terminus has three epidermal growth factor-like domains (EGFs), a mac-perforin-like domain (MACPF), a fibronectin type III domain (FNIII), and an Annexin-like domain (Anx) (diagram kindly provided by Mary Beth Hatten). (B) Dynamics of the ASTN1 dynamics in a migrating CGN. A Venus-ASTN1 construct was introduced into CGNs and high-speed time-lapse imaging used to examine morphological changes. Note ASTN1 initially is located in the CGN cell body and courses into the leading process during migration. lp, leading process. (C) Rationale of JAM-C-pHluorin time-lapse imaging probe. (D) Dynamics of the CGN JAM-C adhesions and its relationship to the actin cytoskeleton. A JAM-Cadhesion reporter (green) and an RFP-Actin reporter (red) were introduced into CGNs and high-speed time-lapse imaging used to examine morphological changes. Note the leading process is a sif-actin enrichment, and the centrosome is located close to actin-enriched domains in the leading process. lp, leading process. Newly formed JAM-C-pHluorin puncta appear in actin-enriched regions of the CGN leading process. (B) Figure adapted with permission from a time lapse video from (Wilson PM., RH.Fryer, Y.Fang, ME.Hatten. Astn2, a novel member of the astrotactin gene family, regulates the trafficking of ASTN1 during glial-guided neuronal migration. J. Neurosci. 2010;30(25): 8529e8540). (C) and (D) reproduced with permission of Famulski, J.K., Trivedi, N., Howell, D., Yang, Y., Tong, Y., Gilbertson, R., Solecki, D.J., 2010. Siah regulation of Pard3 controls neuronal cell adhesion during germinal zone exit. Science.

nucleokinesis during migration along Bergman glial fibers (Solecki et al., 2009). The role of other Pard-complex components and the identity of upstream regulators of polarity during neuronal migration have been relatively unexplored. Examination of the regulation of the Pard complex during CGN differentiation revealed that Pard3 expression is low in immature CGNs and increases as CGNs terminally differentiate, suggesting that Pard3 plays a role in GZ exit or in the switch from tangential to radial migration (Famulski et al., 2010). Indeed, systematic gain- or loss-of-function analyses in ex vivo cerebellar slices confirmed that Pard3 activity is necessary and sufficient for CGN GZ exit. What controls Pard3 levels in differentiating CGNs? A two-hybrid screen identified the seven in absentia homolog (Siah) family of E3 ubiquitin ligases as Pard complexebinding proteins and critical regulators of Pard3. Siah ubiquitin ligases interact with Pard3 via a

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FIGURE 19.8 Polarity regulation in GNPs. (A) Diagram of the early postnatal cerebellar cortex where position of morphologically simple GNPs in the EGL (blue tint) is shown relative to tan, neurite bearing maturing CGNs in the inner EGL, ML, and IGL. Dashed box shows inset blown up in panel B. (B) Siah2 and Zeb1 form a complementary polarity inhibition pair in GNPs (round cell, blue tint) that reduces Pard complex mRNA expression and targets Pard3 or Drebrin proteins for degradation, which arrests GNPs is a morphologically simple premigratory state. As Siah2 and Zeb1 expression recedes, CGNs become morphologically differentiated and the previously inhibited polarity or cytoskeletal proteins facilitate the cell biological activities needed for CGNs to migrate to the final destination.

peptide sequence termed a Siah degron sequence (House et al., 2003). Siah overexpression can induce proteasomal degradation of Pard3 without affecting Pard6 or atypical protein kinase levels, suggesting that Siah ligases antagonize Pard3 function at a posttranslational level. Siah expression in vivo is complementary to Pard3 expression: Siah2 is high in GNPs but is extinguished in premigratory CGNs. Siah activity is necessary and sufficient to maintain immature CGNs within the EGL, as Siah gain of function promotes GZ occupancy, whereas Siah loss of function induces precocious migration toward the IGL. Interestingly, Siah inhibition of GZ exit is dependent on its targeting of Pard3 for degradation, as Pard3 overexpression rescues any Siah-dependent inhibition of migration. Long-term time-lapse imaging of cerebellar slices revealed that excess Siah activity does not deter the motility of CGNs but restricts their movement to the EGL, blocking radial migration to the IGL. Thus, Siah/Pard3 antagonism controls the shift from tangential to radial migration in developing CGNs. The regulation of cell adhesion through junctional adhesion molecule (JAM-C) is a primary downstream function of Siah and Pard3. JAM-C, an immunoglobulin superfamily member (Ebnet et al., 2004), is an epithelial tight-junction component that requires Pard3 binding to the cytoplasmic domain to aid tight-junction formation (Ebnet et al., 2001). Not only is JAM-C expressed in differentiating CGNs but JAM-C-mediated adhesion is necessary and sufficient for CGN GZ exit. Interestingly, JAM-C undergoes leading processedirected polarized trafficking and exocytosis similar to what has been described for ASTN1 (Fig. 19.7C and D). Pard3 activity is essential for recruiting JAM-C to neuroneneuron or neuroneglial cell contacts, and disruption of the Pard3/JAM-C binding blocks CGN migration. Thus, the activity of Pard3 in promoting CGN GZ exit is through JAM-C recruitment to the plasma membrane, which facilitates a posttranslational, polarity-dependent recognition of CGNs, enabling them to navigate along glial fibers to their final destination in the IGL.

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19.1.3.2 The Zeb1-Pard6/3A transcriptional pathway The Siah-Pard3 posttranslational antagonism described in the previous section represents a model of GZ exit in which polarity regulates cell recognition events in the absence of changes to neuronal differentiation and with changes only in the timing of tangential-to-radial migration choices. Interestingly, a parallel mechanism exists whereby the Zeb1 transcription factor regulates CGN polarization timing in a manner that is linked to the overt differentiation status of GNPs and has deep parallels with mesenchymal epithelial transitions (METs) during epithelial cell differentiation (Singh et al., 2016) (Fig. 19.8). As a prelude to these studies, expression analyses revealed that the Pard-complex components Pard6 and atypical protein kinase C are bona fide CGN differentiation markers, as their expression increases as GNP differentiation proceeds. A bioinformatics screen revealed highly conserved Zeb1 transcription factor binding sites in the genes encoding both of these components. Zeb1, a critical regulator of epithelial polarity that locks epithelial cells in the undifferentiated mesenchymal state (Liu et al., 2008; Wellner et al., 2009), is highly expressed in unpolarized GNPs, and its expression diminishes as these cells become polarized CGNs. Systematic necessity/sufficiency testing shows that Zeb1 activity is required to maintain GNPs in the outer portion of the EGL and to arrest their differentiation into postmitotic CGNs in ex vivo cerebellar slices. The loss of Zeb1 in vivo at E18.5 (before the perinatal lethality associated with Zeb1 knockout (Liu et al., 2008)) leads to precocious CGN differentiation marker expression in the EGL. Because Zeb1 inhibits GNP differentiation and downstream events linked to CGN polarization, identification of Zeb1 targets by a functional genomics screen was used to determine how sustained Zeb1 expression maintained GNPs. Zeb1 inhibits the expression of a group of genes whose expression increases during the P0 to P15 GNP differentiation window, including apical or basolateral polarity pathway genes (Pard6, Pard3, Dlg2, and Lin7a) and the genes encoding the E-cadherin (Cdh1) and Close homolog of L1 (Chl1) adhesion molecules, all of which are expressed in cerebellar laminae occupied by postmitotic CGNs at P7 and not in the EGL, where Zeb1 is expressed. Zeb1 regulation of polarity-gene expression is direct, as both ChIP-seq and ChIP PCR show Zeb1 binding of the promoters of Pard6a, Pard3, Chl1, Cdh1, Lin7a, and Limk2. In classic METs, restoring the polarity-gene expression is sufficient to bypass the mesenchymal state induced by factors such as Zeb1 (Aigner et al., 2007). A functional genomics screen was used to identify the Zeb1-target genes that might similarly bypass Zeb1 in CGNs. Restoring the expression of Pard6a, Pard3, and Chl1 enabled GNPs to acquire mature CGN status, characterized by long neurites, the expression of the P27 cell cycle inhibitor, the absence of Ki67 labeling or EdU incorporation, and GZ exit with subsequent migration to the IGL. These findings have not only established a parallel between CGN differentiation and MET-like events in epithelial cells but have also established the Pard complex as a critical regulator of the timing of GNP residency within the outer EGL GZ niche.

19.1.3.3 The foxo polarization pathway The Zeb1ePard complex transcriptional pathway provides one glimpse of polarity control in CGNs, in which Pardcomplex inhibition in GNPs restricts the timing of polarity acquisition in their CGN progeny. However, there are parallel pathways that spur gene expression networks that transcriptionally promote polarization in CGNs after they differentiate (de la Torre-Ubieta and Bonni, 2011) (Fig. 19.9). De la TorreeUbieta and colleagues noted that FOXO transcription factors were enriched in developing neurons across the brain at stages where neuronal polarization was occurring (de la Torre-Ubieta et al., 2010). Not only do rat CGNs express increasing levels of FOXO-1,-3, and -6 during their early postnatal differentiation, but FOXO-6 is highly expressed in the inner EGL, molecular layer, and IGL, where differentiating CGNs undergo morphogenesis to shape their axons and dendrites. FOXO loss of function prevented the maturation of CGN processes into defined axonal and dendritic domains without overtly affected the timing of terminal differentiation. Analysis of FOXO target genes revealed that although FOXO is required for maximal expression of Pard6, Pak1 kinase, CRMP-2, kinesin 5a, and Disc1, the Pak1 kinase is the target responsible for the lack of neurite polarization in FOXOdeficient CGNs, and its expression rescues all FOXO polarization defects. Interestingly, FOXO1 also regulates the expression of the DCX microtubule-binding protein in cooperation with the SnoN-family transcriptional regulators to control parallel fiber branching and gate the timing of CGN migration to the IGL (Huynh et al., 2011). Thus, FOXO transcription factors reveal transcriptional networks of polarity-inducing genes that constitute a neuronal morphogenesis program that gates the timing of migration with the morphological maturation of axonal or dendritic processes that is essential for synaptic connectivity of neurons that have reached their appropriate laminar positions in developing cerebellar circuits. Consistent with this principle, additional studies have shown that the transcription factors NeuroD1 (Gaudilliere et al., 2004), NeuroD2 (Yang et al., 2009), SP4 (Ramos et al., 2007), and MEF2a (Shalizi et al., 2006) also regulate distinct stages of dendritic formation in CGNs later in their development.

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FIGURE 19.9 Transcriptional activation of distinct polarity and morphogenesis pathways in differentiating CGNs. CGN morphogenesis occurs in discreet stages controlled by stage specific transcription factor expression and their transcriptional targets. FOXO transcription factors trigger granule neuron polarization by activating Pak1 kinase expression. SnoN1 repression of a FOXO1 transcriptional complex directly regulates timely onset of Dcx expression modulating the CGN positioning to the IGL. Dendrite morphogenesis involves growth, pruning, and maturation stages. NeuroD promotes the initiation of dendrite growth and branching, while Sp4 regulates dendrite pruning, and the sumoylated repressor form of the transcription factor MEF2A drives the maturation of postsynaptic dendritic claws. Concurrent with dendrite pruning and maturation, development of presynaptic structures in parallel fiber axons is regulated by the transcription factor NeuroD2, which is regulated by the ubiquitin ligase Cdc20-APC. Image depicts a coronal rat cerebellar section overlaid with CGN development as drawn by Ramo n y Cajal.

19.1.4 Part IV. Migration deficits in cerebellar medulloblastomas: the effects of perturbed migration pathways are no longer limited to cognitive deficiency For more than three decades, the impact of defective neuronal positioning has been appreciated to be a significant factor contributing to a spectrum of birth defects that perturb brain lamination and circuit function. The positional cloning of genes that are mutated in neuronal migration disorders, such as lissencephaly (Reiner et al., 1993; Chang et al., 2007; Keays et al., 2007), schizencephaly (Hehr et al., 2010), or pachygyria (Schaffer et al., 2018), has provided critical insights into the molecular cellular mechanisms governing brain lamination. Surprisingly, the paradigm of defective migrations, i.e., arrested GZ exit, has been expanded to include pediatric brain cancers, another group of diseases that afflict children and can trace their origins to defective developmental processes. Medulloblastomas (MBs) are cancerous tumors that develop at the base of the skull within the cerebellar territory. They are the most common malignant neuronal tumors in children, although some MBs appear in adolescence or adulthood (Northcott et al., 2012; Robinson et al., 2012; Taylor et al., 2012). Gene expression, epigenetic profiling, and wholegenome sequencing analyses have proved critical to subdividing MBs into four molecularly and clinically distinct subtypes. Two of these subtypes can be traced back to well-defined neuronal progenitor populations in the developing cerebellum: (1) Sonic hedgehog (Shh)-subtype MBs, so called because of the presence of activating mutations in the Shh signaling pathway, which are derived from GNP-like cells, as the gene expression signature of tumor cells of this subtype is nearly identical to that of proliferating GNPs from the developing cerebella of humans and mice (Goodrich et al., 1997; Wechsler-Reya and Scott, 2001; Lee et al., 2003; Oliver et al., 2003; Uziel et al., 2005) and (2) Wnt-subtype MBs, so called because of mutations activating the Wnt signaling pathway, which have a gene expression signature similar to that of lower RL progenitors that seed brainstem nuclei such as the pons (Gibson et al., 2010). The results of early histological

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studies examining the locations of ectopic “rests” of apparently immature neurons in postmortem human tissues suggested that MBs could be derived from progenitor-like cells from the developing cerebellum. Consistent with this early idea, MBs of the Shh and Wnt subtypes demonstrate anatomic differences at diagnosis, with Shh-subtype tumors predominantly appearing at the dorsal aspect of the cerebellum and Wnt-subtype tumors being located in the fourth ventricle and infiltrating the brainstem (Gibson et al., 2010) (Fig. 19.10A and B). Mouse models of the Shh- and Wnt-subtype MBs that recapitulate the driver mutations of each tumor type have provided evidence of apparent GZ occupancy defects in both tumors (Goodrich et al., 1997; Oliver et al., 2005; Yang et al., 2008; Gibson et al., 2010). Mice that harbor mutations in or are haploinsufficient for the Patched Shh receptor (Goodrich et al., 1997) exhibit chronic activation of the Shh signaling pathway, a potent mitogenic signal (WechslerReya and Scott, 1999), in GNPs, leading to GNP transformation and MB tumorigenesis on the dorsal aspect of the cerebellum because Patched deficiency acts as a mitogenic stimulus to these cells. The proliferating preneoplastic GNPs remain in the dorsally located EGL for long periods, leading to a perduring EGL even at stages in which the GNP population is extinguished in wild-type animals (Goodrich et al., 1997; Oliver et al., 2005). Similarly, activation of the Wnt signaling pathway in the lower RL leads to abnormal collections of preneoplastic progenitor cells in the brainstem anatomical location where Wnt tumors frequently reside (Gibson et al., 2010). Indeed, in utero electroporation studies demonstrate that lower RL progenitors do not efficiently transit from the RL to the brainstem when Wnt signaling is activated, providing direct evidence that the migration of this cell population is affected (Fig. 19.10C, left panel). Similarly, activation of the Shh pathway in EGL resident GNPs also create a cohort of cells that display GZ exit and radial migration defects (Fig. 19.10C, right panel). Studies in the GNP system have suggested mechanisms to explain how a lack of GZ exit can contribute to tumorigenesis. Choi and colleagues used in vivo and in vitro approaches to examine whether the EGL represented a mitogenic niche in the developing cerebellum, and they found that a delay in GZ exit in vivo caused by the loss of chemotactic activity for CGN migration to the IGL resulted in additional GNP proliferation (Choi et al., 2005). A clever application of in vitro seeding of GNPs on cerebellar slices followed by careful analysis of the cell proliferation revealed that GNPs incorporated into the EGL of the slice continued to proliferate in an Shh-dependent manner, whereas GNPs incorporated into the IGL quickly differentiated and became postmitotic. EGL-resident GNPs may be the only cells of the CGN lineage that can respond to Shh, as gene expression analyses show that Shh-responsive genes are restricted to the EGL in vivo. Interestingly, the CXCR4/SDF1 system, which forms a chemoattractant that maintains GNPs in the EGL, synergized with Shh signaling to boost greater GNP proliferation than was obtained with either factor alone (Klein et al., 2001). Thus, perduring GZ occupancy in the EGL or brainstem can expose preneoplastic progenitor cells to extended temporal windows of proliferative signaling that ultimately could render the cells susceptible to additional genetic hits that would further transform them into MB subtypes, depending on their lineage and GZ location. Surprisingly, analysis of the Zeb1-Pard6/3 transcriptional pathway revealed that GZ occupancy pathways that restrict GZ exit and the acquisition of polarity may also be relevant to MB (Singh et al., 2016). Not only is Zeb1 expression activated by Shh in GNPs; Zeb1 is also overexpressed in mouse and human MBs in which Shh signaling is activated and Zeb1 targets, including the Pard6 and Pard3 polarity genes, are only weakly expressed. A functional genomic screen in ex vivo slices revealed Zeb1 repression of its targets, which is relevant to the aberrant GZ exit of Ptcdeficient GNPs. Acute deletion of Ptc leads to a cohort of GNPs that do not exit their GZ ex vivo, which is identical to what occurs in in vivo MB models (Fig. 19.10C, left panel), but Zeb1 silencing or elevated expression of Pard6, Lin7a, or Chl1 restores GZ exit. This finding indicates that Zeb1 and its targets are functionally downstream of the Shh pathway during GNP GZ exit and shows that defective polarity signaling underlies GZ-exit defects in MB. The finding that SHH maintains Zeb1 expression and that Zeb1 target expression is reduced in MB reveals an antagonism between the main GNP mitogen and the polarity required for GZ exit. This antagonism suggests that SHH inhibits the MET-like event that controls GNP GZ exit and that preneoplastic GNPs or MB cells are inherently polarity deficient. The possibility that Zeb1 controls an active program to block polarization is particularly relevant to MB. These tumor cells express high levels of the FOXO, NeuroD1, and Mef2c transcription factors that promote CGN polarization, but they are insufficient to induce polarization of transformed GNPs (Robinson et al., 2012). Thus, Zeb1 is a candidate factor that may act downstream of SHH in MB to counteract the polarization program. Further studies are required to determine whether restoring the polarity balance in MB will yield a therapeutic benefit as a complement to existing first-line or targeted therapies.

FIGURE 19.10 Altered GZ exit and migration in cerebellar tumorigenesis. (A) Mice that harbor mutations that activate the Wnt signaling pathway in the lower rhomic lip (left) or are haploinsufficient for the Patched Shh receptor in GNPs (right) exhibit cerebellar tumor formation. (B) Wnt- and Shhtumor locations are not only highly stereotyped in heat maps delineating tumor location incidence but also overlap anatomically with the brainstem and dorsal cerebellar EGL that are the GZs for LRL or EGL progenitors. (C) In utero and ex vivo cerebellar electroporations that activate Wnt signaling in the LRL (left) or Shh signaling in the EGL create cohorts of progenitors that do not initiate migration in a timely manner. Panel A, B and the left panel in C were reproduced with permission of Gibson P., Tong, Y., Robinson, G., Thompson, M.C., Currle, D.S., Eden, C., Kranenburg, T.A., Hogg, T., Poppleton, H., Martin, J., Finkelstein, D., Pounds, S., Weiss, A., Patay, Z., Scoggins, M., Ogg, R., Pei, Y., Yang, ZJ., Brun, S., Lee, Y., Zindy, F., Lindsey, J.C., Taketo, M.M., Boop, F.A., Sanford, R.A., Gajjar, A., Clifford, SC., Roussel, MF., McKinnon, P.J, Gutmann, D.H., Ellison, D.W., Wechsler-Reya, R., Gilbertson, R.J., Subtypes of medulloblastoma have distinct developmental origins. Nature 2010:468(7327): 1095e1099, the right panel was reproduced with permission of Singh S, Howell, D., Trivedi, N., Kessler, K., Ong, T., Rosmaninho, P., Raposo, A.A., Robinson, G., Roussel, M.F., Castro, D.S., Solecki, D.J., Zeb1 controls neuron differentiation and germinal zone exit by a mesenchymal-epithelial-like transition. Elife 2016 5.

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Acknowledgments Keith A. Laycock, Ph.D., ELS edited the manuscript. The Solecki Laboratory is funded by the American Lebanese Syrian Associated Charities (ALSAC) and by grants 1R01NS066936 and R01NS104029 from the National Institute of Neurological Disorders (NINDS).

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Chapter 20

Neuronal migration of guidepost cells Franck Bielle1 and Sonia Garel2, 3, 4 1

Institut du Cerveau et de la Moelle Epinière, Paris, France; 2Institut de Biologie de l’École Normale Supérieure (IBENS), PSL Université, Paris,

France; 3Institut National de la Santé et de la Recherche Médicale (INSERM) U1024, Paris, France; 4Centre National de la Recherche Scientifique (CNRS) UMR 8197, Paris, France

Chapter outline Chapters cited 435 20.1. An introduction to guidepost cells 436 20.1.1. Neuronal migration in the context of axonal tracts formation 436 20.1.2. Defining the notion of guidepost cells 436 20.2. Role of neuronal migration in the formation of the lateral olfactory tract 437 20.2.1. Anatomy and development of the lateral olfactory tract 437 20.2.2. Diffusible guidance cues in the pathfinding of lateral olfactory tract axons 438 20.2.3. Roles of guidepost “lot cells” 439 20.2.4. Tangential migration of lot cells: specification, routes, and molecular mechanisms 439 20.2.5. Fate of lot cells 440 20.3. Hippocampal Cajaleretzius cells in the formation of axonal connections 440 20.3.1. Anatomy and development of the hippocampus and entorhinohippocampal projections 440 20.3.2. CajaleRetzius cells as putative guidepost neurons for the formation of entorhinal projections 440 20.3.3. Toward a more generic role of CajaleRetzius cells as guideposts? 442 20.4. Migration of neuronal guidepost cells in the formation of thalamocortical connections 442 20.4.1. Anatomy and development of thalamocortical and corticofugal axons 442 20.4.2. Pioneer cortical subplate axons in the pathfinding of thalamocortical projections 443 20.4.3. Origin and migration of subplate neurons 444

20.4.4. The subpallium is a major intermediate target for thalamocortical axons 444 20.4.5. Guidepost cells in the diencephalic and subpallial pathfinding of thalamocortical projections 445 20.4.6. Migration of guidepost corridor cells: routes and guidance cues 446 20.4.7. Fate of guidepost cells for thalamocortical projections 447 20.5. Neuronal migration of guidepost cells in the formation of the corpus callosum 447 20.5.1. Anatomy and development of the corpus callosum 447 20.5.2. Roles of glial cells in the development of the corpus callosum 447 20.5.3. Tangentially migrating neurons in the development of the corpus callosum 448 20.6. Neuronal migration of guidepost cells and evolution of brain wiring 449 20.6.1. Tangential migration of guidepost neurons: a hallmark of the telencephalon? 449 20.6.2. Neuronal migration of guidepost cells in the evolution of the internal capsule 450 20.7. Towards an integration of migrating guidepost neurons in normal and pathological brain development 451 20.7.1. Guidepost neurons in the shaping of axonal tract organization and topography 451 20.7.2. Integrating tangential neuronal migration of guideposts in normal and pathological brain development 452 20.8. Conclusions 452 References 453

Chapters cited 4 Axon guidance: netrins 5 Axon guidance: semaphoring/Npn/Plexin signaling 6 Ephrin/Eph signaling in axonguidance

Cellular Migration and Formation of Axons and Dendrites. https://doi.org/10.1016/B978-0-12-814407-7.00020-1 Copyright © 2020 Elsevier Inc. All rights reserved.

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7 8 15 16 17 18 19 25 26 21 22

Axon guidance: slit/Robo signaling Nonconventional axon guidance cues: hedgehog, TGF-b/BMP, and Wnts in axon guidance Radial migration in the developing cerebral cortex Mechanisms of tangential migration of interneurons in the developing forebrain Migration in the hippocampus Hindbrain tangential migration Neuronal migration in the developing cerebellar system The impact of different modes of neuronal migration on brain evolution Neuronal migration disorders Striatal circuit development and synapse maturation (Book title: Synapse Development and Maturation) Cajal-Retzius and subplate cells: transient cortical neurons and circuits with long-term impact (Book title: Synapse Development and Maturation)

20.1 An introduction to guidepost cells 20.1.1 Neuronal migration in the context of axonal tracts formation The functioning of the mammalian brain relies on the formation of specific axonal connections between neurons located in distinct regions. As overall presented in this series of chapters, cell migration is essential to control the positioning of all the neurons, as they are generated by a spatially restricted proliferative niche and subsequently migrate to their final destination. Neuronal cells migrate radially or tangentially and, either as they migrate or after completing their migration, extend an axon toward their synaptic targets. While conserved molecular and cellular mechanisms control the orientation of neuronal migration and the pathfinding of axonal connections (Chapters 4-8, 15-19 and 25, Tessier-Lavigne and Goodman, 1996), they are often studied independently as two events occurring sequentially in the life of individual neurons. However, as brain development is a continuous process, at a given developmental stage, neurons migrate while axons of other neuronal subpopulations navigate toward their targets, raising the intriguing question of how these two processes are coordinated to ensure the functional emergence of neural circuits. This issue is particularly important in brain regions that host extensive neuronal migrations such as the embryonic mammalian telencephalon, which contains the cerebral cortex (pallium) and basal ganglia (subpallium) (Chapters 15-17 and 47). Studies over the past decade have revealed that the development of the telencephalon relies on multiple streams of tangential migration that redistribute neuronal populations over long distances. For instance, the subpallium produces interneurons that populate the entire telencephalon via distinct migratory streams (Chapters 15-17), and early-born pallial neurons, including CajaleRetzius cells, migrate superficially to cover the entire primordium of the cerebral cortex (Chapters 15-17, 25 and 48). These neuronal migrations are essential to build the complex architecture of neural circuits that underlies the functioning of the cerebral cortex and basal ganglia. Accordingly, developmental defects in telencephalic neuronal migrations have been directly associated with the etiology of several neurologic and psychiatric diseases in humans (Chapter 26). The relative recent discovery and characterization of these migratory streams have also revealed that the embryonic telencephalon constitutes a constantly evolving cellular landscape for navigating axons. While dissecting interactions between neuronal migration and axon guidance appears thus essential to gain a global perspective on how the cellular architecture and the wiring of neural circuits are established during brain development, we are just beginning to address these issues. In this chapter, we will review recent studies revealing that tangential neuronal migration, in addition to its wellcharacterized function in redistribution and neural morphogenesis, acts as a dynamic “guidepost” system to position cues important for the formation of major axonal tracts. To this aim, we will first briefly introduce the notion of “guidepost” cells and present four main examples in which neuronal migration has been shown to participate in the formation of axonal tracts in the mouse telencephalon. We will further discuss the implications of these findings regarding morphogenesis, the evolution of brain wiring, and pathological cerebral development.

20.1.2 Defining the notion of guidepost cells Axonal projections often travel over long distances along a highly stereotyped pattern to reach their final synaptic partners. During this process, pioneer axons are guided by intermediate targets located along their path, which produce secreted or membrane-bound environmental signals required for the further and appropriate navigation of growing axons (Fig. 20.1). Complex and long axonal trajectories are thus the result of simpler and shorter individual steps in which growing axons navigate from one intermediate target to another, until they reach their final destination. For instance, the ventral midline of the central nervous system in both invertebrates and vertebrates constitutes a major intermediate target for organizing the pathfinding decision of growing axons to cross toward the contralateral side of the body (Goodman and Shatz, 1993; Tessier-Lavigne and Goodman, 1996).

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FIGURE 20.1 Intermediate targets and guidepost cells are major players in axon guidance. (A,B) During their progression toward their final target (blue), axonal growth cones (green) successively reach different intermediate targets (yellow). Each intermediate target attracts the axonal growth cone and orients its pathfinding. (C) At a cellular scale, guidepost cells (pink) provide short-range guidance activities of intermediate targets by acting on the growth cone (green) via either secreted or transmembrane factors.

The term “guidepost cells” was originally introduced to describe a particular intermediate target in the limb bud of the grasshopper embryo (Bate, 1976; Bentley and Keshishian, 1982; Ho and Goodman, 1982). In this system, pioneer axons follow a stereotyped path (Bate, 1976; Thomas et al., 1984) along which specialized nonadjacent guidepost cells constitute essential intermediate targets required for the further progression of axons (Bentley and Caudy, 1983). These guidepost cells are immature neurons that have not extended an axon yet and have a high affinity for the growth cones of pioneer axons: Once a single filopodia contacts the cell body of guidepost cells, the filopodia is stabilized and the growth cone reorients to establish a direct membrane apposition (Bentley and Keshishian, 1982; Taghert et al., 1982; Bastiani and Goodman, 1984; O’Connor et al., 1990; Sabry et al., 1991). These original studies have defined guidepost cells as noncontinuous cellular landmarks located along the future path of growing axons, which are required for their correct navigation by providing highaffinity substrates allowing the crossing of low-affinity intervals (Hammarback and Letourneau, 1986). From these pioneering studies, the concept of guidepost cells has been extended to specialized discrete cell populations that are located at critical decision points along the trajectory of pioneer axons and that participate in their guidance (Fig. 20.1). They can be isolated or form a continuous domain and eventually send an axonal extension along the future path of the tract (Fig. 20.1). In contrast to some intermediate targets that have a long-range activity, guidepost cells act at shortrange or via cellecell contact. In vertebrates, most of the guideposts cells identified so far are glial cells, such as floor plate cells (Tessier-Lavigne et al., 1988; Bovolenta and Dodd, 1990; Placzek et al., 1990; Bovolenta and Dodd, 1991; Campbell and Peterson, 1993; Kennedy et al., 1994; Serafini et al., 1994; Serafini et al., 1996), boundary cap cells (Golding and Cohen, 1997; Fraher et al., 2007), radial glia of the optic chiasma (Misson et al., 1988; Guillery et al., 1995; Marcus et al., 1995; Wang et al., 1995) for review (Mason and Sretavan, 1997), glial bridges of the anterior commissure and postoptic commissure (Silver et al., 1982; Pires-Neto et al., 1998; Barresi et al., 2005; Lent et al., 2005), or glial subpopulations of the corpus callosum (Silver et al., 1982; Silver and Ogawa, 1983; Silver et al., 1993; Shu and Richards, 2001; Shu et al., 2003b; Shu et al., 2003c). However, studies conducted over the past decades have revealed that tangentially migrating neuronal cells participate in the guidance of several major axonal tracts of the mammalian telencephalon (for review, Marin et al., 2010).

20.2 Role of neuronal migration in the formation of the lateral olfactory tract 20.2.1 Anatomy and development of the lateral olfactory tract The first system in which migrating neuronal guidepost cells have been proposed to orient axonal pathfinding is the lateral olfactory tract, which is a major component of the circuit conveying olfactory information to the cerebral cortex. Sensory olfactory neurons from the nasal cavities project onto the dendrites of mitral and tufted cells of the olfactory bulb, which in turn send their axons to higher-order olfactory centers such as the anterior olfactory nucleus, olfactory tubercle, piriform cortex, entorhinal cortex, and cortical nuclei of the amygdala (Fig. 20.2A; Schwob and Price, 1984; Shipley and Ennis,

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E12 + + + − + −+ + − − + − + − + − Sema3F

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FIGURE 20.2 Lot cells in the development of the lateral olfactory tract (LOT). (A) The LOT (green) contains axons of the olfactory bulb projecting to the anterior olfactory nucleus (AONs), the olfactory tubercle (Tub), the piriform cortex (Pir), the entorhinal cortex (EC), and the amygdala (Am). (B) Lot cells migrate ventrally from the pallium and stop at the pallium/subpallium boundary (PSPB, dashes) where they reorientate along a rostrocaudal direction. They are attracted by Netrin1, and their migration is limited by the repellent Sema3F. Their distribution extends from the amygdala at E12, to the zone posterior to the olfactory bulb at E12.5. (C) At E12.5, LOT axons extend along a caudal direction in close contact with lot cells and in a superficial position.

1996; Zou et al., 2001; Diodato et al., 2016). The primary axons of mitral cells form a tight axonal bundle called the lateral olfactory tract (LOT), which navigates superficially over the olfactory cortices. From this primary axonal highway, collaterals extend into the underlying pallial structures to target secondary olfactory neurons, such as layer III pyramidal neurons of the piriform cortex (Price, 1973; Leonard, 1975; Schwob and Price, 1984). Mitral cells are among the first neurons to develop in the telencephalon and start extending their axons into the LOT from embryonic day 12.5 (E12.5) in the mouse (Cajal, 1890; Hinds and Ruffett, 1973; Pini, 1993; Sugisaki et al., 1996; Mombaerts, 2006). Ventral mitral cells pioneer the LOT, and between E13 and E14, most of mitral cells extend their axon, giving to the LOT its definitive arch shape that runs from the olfactory bulb across the piriform cortex and the entorhinal cortex (Fig. 20.2A). By E15, collaterals start to extend into the piriform cortex and progressively into other secondary targets, including the amygdala, entorhinal cortex, and olfactory tubercles (Hinds and Ruffett, 1973; Mombaerts, 2006).

20.2.2 Diffusible guidance cues in the pathfinding of lateral olfactory tract axons Both long-range and local environmental cues have been implicated in the guidance of LOT axons (de Castro, 2009). Indeed, the septum produces a diffusible repulsive activity, which is essential for the lateral pathfinding of mitral cells axons (Pini, 1993), and involves the secreted proteins Slit1 and Slit2 (Nguyen Ba-Charvet et al., 1999, 2002) mediated by their receptors, Robo1 and Robo2 (Fouquet et al., 2007). In addition, two secreted semaphorins (Sema) have been proposed to further orient the growth of LOT axons: Sema3B, which is expressed in the mesenchymal tissue of the prospective skull, promotes the growth of olfactory bulb axons, whereas, Sema3F which is expressed in the cortical plate and the subpallium,

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repels LOT axons (de Castro et al., 1999 or for review, de Castro, 2009). While these activities could delineate the subpial longitudinal path of LOT axons, their in vivo function remains to be characterized. Finally, a panel of additional proteins have been shown or proposed to regulate later aspects of LOT morphogenesis. In particular, the secreted protein anosmin1, which is expressed in the piriform cortex, has an attractive activity on LOT axons and promotes their branching (SoussiYanicostas et al., 2002). Similarly, activation of the Nogo signaling pathway has been shown to promote the branching of LOT axons (Iketani et al., 2016).

20.2.3 Roles of guidepost “lot cells” Aside from these diffusible guidance activities, the group of Fujisawa showed that the future path of the LOT contains a short-range permissive activity (Sugisaki et al., 1996). The lot1 monoclonal antibody allowed the identification of a specific population of “lot cells” (Sato et al., 1998), which could correspond to previously described horizontal neurons of the developing piriform cortex (Derer et al., 1977). They are among the first neurons generated in the telencephalon as they are born between E9.5 and E11.5 and appear in the caudoventral telencephalon at E12.0 and their distribution extends to the olfactory bulb before LOT axons start growing at E12.5 (Sato et al., 1998). Lot cells are densely packed on the future path of the LOT, and they constitute a cellular substrate for growing axons (Fig. 20.2B and C). As axons grow, lot cells are displaced along the internal border of the bundle; they are associated with collaterals once they are formed (Hirata and Fujisawa, 1999) and are not detected after E18.5, as lot1-immunostaining disappears (Sato et al., 1998). Lot1 immunostaining was shown to recognize a specific isoform of the metabotropic mGluR1 receptor and constitutes a specific marker of lot cells at early stages (Hirata et al., 2012). Lot cells share a common location and some molecular markers with reelinpositive CajaleRetzius cellsdanother group of early-born superficial neurons; however, they constitute a specific neuronal subpopulation, as most of them express lower levels of reelin (Sato et al., 1998; Dixit et al., 2011; de Frutos et al., 2016). Remarkably, lot1/mGluR1-expressing cells were also shown to respond to the activity of LOT axons (Hirata et al., 2012), indicating that lot cells could be part of an early olfactory protocircuit. Both the location and the timing of lot cells distribution suggested that they could constitute the cellular substrate of the permissive activity previously described (Sugisaki et al., 1996; Sato et al., 1998). To directly test this putative role, Sato et al. proceeded to toxic ablation of lot cells by local exposure to 6-OHDA, thereby revealing that the growth of LOT axons was stalled at the position where lot cells were eliminated (Sato et al., 1998). These experiments strongly supported a shortrange activity of lot cells on LOT axonal growth, thereby fulfilling the definition of guidepost cells.

20.2.4 Tangential migration of lot cells: specification, routes, and molecular mechanisms Because lot cells have a short-range activity, their positioning is crucial to their guidepost function. Explant cultures and tracing experiments suggested that lot cells derive from all regions of the dorsal pallium and migrate tangentially from E10.5 to settle ventrally in the future path of the LOT (Fig. 20.2B; Tomioka et al., 2000). Mutant mice analyses have shown that two transcription factors, Gli3 and Lhx2, are required for the proper development of lot cells, supportive of a dorsal pallium origin (Tomioka et al., 2000; Kawasaki et al., 2006; Saha et al., 2007; Amaniti et al., 2015). However, as lot cells are identified at the end of their migration by lot1 immunostaining, deciphering their precise origins has constituted a long-lasting challenge. Recent evidence indicates that lot cells are not generated in the dorsal pallium but produced by the thalamic eminence, which is also an identified generation site for both CajaleRetzius cells and additional cells that migrate toward the olfactory bulb (Huilgol et al., 2013; Barber and Pierani, 2016; Huilgol and Tole, 2016; Ruiz-Reig et al., 2017). Several structures and guidance cues participate in the guidance of lot cells migration. Organotypic cultures showed that lot cells are attracted ventrally, but the subpallium constitutes a nonpermissive territory for their migration: When lot cells arrive to the pallium/subpallium boundary, they change their orientation as they make a 90 degrees turn and align rostrocaudally (Fig. 20.2B, Kawasaki et al., 2006). Netrin1 has been shown to attract migrating lot cells and could participate in the ventral attractive activity (Kawasaki et al., 2006). However, Netrin1 invalidation, or the invalidation of its receptor Dcc, only affects the positioning of ventralmost lot cells (Kawasaki et al., 2006). Consistently, this altered positioning of guidepost cells could account for the specific pathfinding defect of ventralmost LOT axons in Netrin1 KO embryos (Kawasaki et al., 2006). Sema3F, which is expressed in the subpallium and cortical plate, is repulsive for migrating lot cells via the receptor Neuropilin2 (Nrp2) (Ito et al., 2008). Loss of Sema3F or Nrp2 leads to ectopic lot cells in the depth of the piriform cortex, whereas they are normally restricted to a subpial position (Ito et al., 2008). Finally, ventralmost lot cells are not properly positioned in double Slit1; Slit2 mutants and LOT axons originating from the medial bulb are misguided toward the midline, whereas dorsalmost lot cells are normally positioned and axons of the lateral bulb follow a normal path along them (Fouquet et al., 2007). Thus, in general, the altered positioning of lot cells after the

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invalidation of guidance cues or their corresponding receptors could participate or mediate the associated guidance defect of LOT axons. However, in the case of Sema3F, Slit1 and Slit2, LOT axons are also directly guided by the same cues, and thus the axon guidance defects observed in mutant embryos could result from a combination of direct and lot cellse mediated mechanisms. Future works using conditional mutants or local known-down for receptors will be required to fully understand the relative function of guidance cues in lot cell positioning and axon guidance (Fig. 20.2B and C).

20.2.5 Fate of lot cells Because lot cells are identified by their location and lot1/mGluR1 immunostaining (Sato et al., 1998), it has been difficult to trace these cells overtime. It has been established that lot1/mGluR1-positive cells disappear from the LOT region, raising the possibility that these cells are eliminated by programmed cell death (Sato et al., 1998). However, several studies provided evidence that at least some lot cells persist. On the one hand, experimental tracing supported the hypothesis that some cells in the LOT region give rise to mitral cells migrating to the olfactory bulb (Huilgol et al., 2013; Huilgol and Tole, 2016; Ruiz-Reig et al., 2017). On the other hand, using DNp73-cre transgenic lines backcrossed with tdTomato reporters, which label 80% of all CajaleRetzius cells (Tissir et al., 2009), it was established that lot cells are part of a shared lineage, at least for a vast majority of this guidepost population (de Frutos et al., 2016). Such lineage tracing confirmed that lot cells respond to neuronal stimulation of LOT axons and enable to trace these cells over time, since cre-driven recombination is permanent (de Frutos et al., 2016). Strikingly, a large population of lot cells was shown to reenter migration after E14, a timepoint at which lot cells have performed their guidepost functions, thereby providing an additional source of Cajale Retzius cells in the growing adjacent neocortex (de Frutos et al., 2016). It thus appears that guidepost lot cells are part of an early olfactory circuit and that subset of these cells migrate out of the olfactory cortex to perform additional functions in the brain. These findings collectively support the notion that guidepost cells can have multiple roles in brain development, which is regulated overtime by their migratory behavior and positioning (Squarzoni et al., 2015).

20.3 Hippocampal Cajaleretzius cells in the formation of axonal connections 20.3.1 Anatomy and development of the hippocampus and entorhinohippocampal projections Entorhinal pyramidal neurons are the main excitatory input of the hippocampus, which is subdivided into the CA1 to CA4 and the dentate gyrus (DG) (van Groen et al., 2002; van Groen et al., 2003; for review, Turner et al., 1998). They send their axons through the perforant path to apical dendrites of hippocampal projection neurons (Fig. 20.3A). Whereas layer II entorhinal neurons project onto granule cells of the dentate gyrus, layer III entorhinal neurons project onto pyramidal neurons of CA1. These entorhinal afferences target the apical dendrites of hippocampal pyramidal neurons and granule cells located in the stratum lacunosum moleculare (SLM) or the outer molecular layer (OML), respectively (Fig. 20.3). Noteworthy, the SLM and the OML are adjacent due to the curve described by the hippocampal structure and are only separated by the hippocampal fissure (Blackstad 1956, 1958; Hjorth-Simonsen and Jeune, 1972; Amaral and Witter, 1995). In addition to this entorhinal input, hippocampal projection neurons receive associational and commissural afferences, sent by the ipsilateral and contralateral hippocampal projections neurons, respectively (Fig. 20.3B). These inputs are restricted to specific layers, the stratum radiatum (SR) and stratum oriens (SO) for hippocampal neurons and the internal molecular layer (IML) for granule cells. During mouse development, entorhinal axons are first detected in the hippocampus at E15, arborize specifically in the SLM from E17, and invade the OML by postnatal day 0 (P0) (Fig. 20.3B). Commissural axons invade specifically the contralateral SR at E18 and IML at P2 (Super and Soriano, 1994; Super et al., 1998; Deng and Elberger, 2001; Deng et al., 2006). While hippocampal afferences are restricted to their final target layers prenatally, their targets develop postnatally. Indeed, hippocampal pyramidal neurons and granule cells are generated locally with a peak of production at E14e16 (Caviness, 1973; Soriano et al. 1986, 1989), but their apical dendrites are not detected in the SLM before P2 (Bayer and Altman, 1987; Super et al., 1998). Thus, entorhinal axons arrive to and restrain their growth to the position where their definitive target will form, but before these targets are developed.

20.3.2 CajaleRetzius cells as putative guidepost neurons for the formation of entorhinal projections While the functioning of the hippocampus relies on both projection neurons and GABAergic interneurons, its development is additionally linked to a transient population of glutamatergic neurons located in the marginal zone of the hippocampus, CajaleRetzius cells (Chapters 17 and 48; Gorski et al., 2002; Radnikow et al., 2002; Hevner et al., 2003). CajaleRetzius cells

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FIGURE 20.3 Hippocampal CajaleRetzius cells are guidepost neurons for entorhinohippocampal axons. (A) The mature entorhinohippocampal projection is visible in a coronal section plane. Neurons of the entorhinal cortex (EC) provide the main excitatory input to the hippocampus (CA1e3) and to the dentate gyrus (DGs). Entorhinal axons originating from layer II (light green) target the stratum lacunosum moleculare (SLM) of CA1e2, and those originating from layer III (dark green) target the outer molecular layer (OML) of the DG. (B) The inputs of CA1e3 and DG respect a laminated pattern since their early development. This lamination is prefigured at E15.5 by a distribution of migrating cells delineating the future layers: CajaleRetzius cells (pink) are distributed in SLM and OML, and GABAergic neurons (yellow) are distributed in stratum oriens (SOs), stratum radiatum (SR), and inner molecular layer (IML). Between E16.5 and P0, entorhinal axons invade specifically the SLM and OML in close association with CajaleRetzius cells although their future finale targets, the apical dendrite neurons of the pyramidal layer (sp, dark blue) and of the granule cell layer (g, cyan), develop later. Commissural/associational axons (light gray) will develop in layers containing GABAergic cells. At P15, entorhinal axons have established synapses on the apical dendrites of their definitive targets, and CajaleRetzius cells have mostly disappeared. g, granule cells; hf, hippocampal fissure; sp, stratum pyramidale.

are early-born pallial neurons that cover the entire cortical sheet and hippocampal primordium and regulate cortical lamination via the production of the extracellular protein reelin (Chapter 15; D’Arcangelo et al., 1995; Hirotsune et al., 1995; Ogawa et al., 1995; D’Arcangelo et al., 1997; for review, Tissir and Goffinet, 2003). During mouse development, both CajaleRetzius cells and GABAergic neurons migrate tangentially into the hippocampus (Chapters 17 and 48): GABAergic interneurons are generated in the subpallium (Anderson et al., 1997a, 1997b; Anderson et al., 2001; Wichterle et al., 2001; Xu et al., 2008; Gelman et al., 2009), whereas CajaleRetzius cells are generated in focal pallial territories and migrate tangentially in the marginal zone to cover the entire pallium (Takiguchi-Hayashi et al., 2004; Bielle et al., 2005; Yoshida et al., 2006). In the developing hippocampus, CajaleRetzius cells distribution overlaps with the future SLM and OML, whereas the GABAergic interneurons distribution overlaps with the SO, SR, and IML (Fig. 20.3; Nowakowski and Rakic, 1979; Soriano et al., 1994; del Rio et al., 1995, 1996, 1997; for review, Forster et al., 2006). Standard and electron microscopy have shown that pioneer entorhinal axons transiently establish synaptic contacts with CajaleRetzius cells in SLM and OML and conversely pioneer commissural axons with GABAergic neurons in SO, SR, and IML (Super et al., 1998). The role of CajaleRetzius cells in entorhinal axons pathfinding was examined in organotypic slice cultures, which recapitulate the development of entorhinalehippocampal projections (Frotscher and Heimrich, 1993; Li et al., 1993; Frotscher et al., 1995; del Rio et al., 1997). In this system, the toxic ablation of CajaleRetzius by local 6OHDA treatment ex vivo prevented the growth of entorhinal axons in the hippocampus, whereas the growth of commissural axons still occurred showing a specific association between the presence of CajaleRetzius cells and the growth of entorhinal axons (del Rio et al., 1997). These experiments thus strongly support a role of hippocampal CajaleRetzius cells as guidepost neurons (Forster et al., 1998), although this remains to be demonstrated in vivo.

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What could be the molecular mechanism underlying CajaleRetzius cells activity? An obvious candidate is the extracellular protein reelin (D’Arcangelo et al., 1995; Hirotsune et al., 1995; Ogawa et al., 1995; D’Arcangelo et al., 1997; Di Donato et al., 2018; for review, Tissir and Goffinet, 2003). However, in spite of a disorganized laminar organization, reeler mutants show no major pathfinding defects of entorhinal projections to their target layers (Frotscher and Heimrich, 1993; Li et al., 1993; Frotscher et al., 1995; del Rio et al., 1997; Borrell et al., 1999; Deller et al., 1999). Still, in vivo analyses and ex vivo loss-of-function experiments revealed that reelin favors the branching, collateral formation, and synaptogenesis of entorhinal axons in the hippocampus (Frotscher and Heimrich, 1993; Li et al., 1993; Frotscher et al., 1995; del Rio et al., 1997; Borrell et al., 1999; Deller et al., 1999). CajaleRetzius cells thus likely act via the production of other factors that remain to be characterized. An alternative nonexclusive hypothesis is that CajaleRetzius cells may provide an axonal scaffold for ingrowing axons. Indeed, in contrast to other hippocampal axons, CajaleRetzius cells of the SLM and OML establish an early projection to the entorhinal cortex (Fig. 20.3B; Ceranik et al., 1999; Ceranik et al., 2000; Deng et al., 2007). Several guidance cues have been shown to regulate either entorhinal axons guidance, such as Netrin1, ephrinA3 and A5, and SDF-1/CXCL12 (Stein et al., 1999; Barallobre et al., 2000; Ohshima et al., 2008), or the migration of Cajale Retzius cells, such as SDF-1/CXCL12 (Borrell and Marin, 2006; Paredes et al., 2006). However, as CajaleRetzius cells influence the early morphogenesis of the hippocampus and guidance cues have pleiotropic effects, the relative role of CajaleRetzius cells as potential guideposts in vivo and the environmental cues that control their positioning still remain to be fully investigated.

20.3.3 Toward a more generic role of CajaleRetzius cells as guideposts? It is noteworthy that in both the hippocampus and the olfactory cortex, CajaleRetzius cells and related lot cells have been proposed to act as guidepost cells, raising the question of whether these cells may exert similar functions in the neocortex. This possibility is intriguing since CajaleRetzius cells are located in the developing layer 1, which comprises long-range axonal inputs, axons from local interneurons as well as the apical dendrites of principal cortical neurons (Llinas et al., 2002; Rubio-Garrido et al., 2009; Chen and Kriegstein, 2015; Harris and Shepherd, 2015; Ibrahim et al., 2016; Roth et al., 2016). Reducing the density of neocortical CajaleRetzius cells in the somatosensory neocortex by a third during embryonic and postnatal life is sufficient to induce a decrease in the excitatory inputs targeting the dendrites of principal cortical neurons (de Frutos et al., 2016). Whether this phenotype is linked to an axonal guidepost function of these cells remains entirely to decipher.

20.4 Migration of neuronal guidepost cells in the formation of thalamocortical connections 20.4.1 Anatomy and development of thalamocortical and corticofugal axons The six-layered cerebral cortex is specific to mammals and is connected to the rest of the brain by a unique circuitry. Particularly, the internal capsule gathers most of the afferences and efferences of the cerebral cortex. On the one hand, it contains thalamocortical axons (TCAs), which convey sensory and motor information from the thalamus to the different cortical areas. On the other hand, it contains corticofugal axons, which project either reciprocally to the thalamus (corticothalamic axons, CTAs) or to other subcortical targets (for instance, corticostriatal and corticospinal axons). Importantly, these projections show a topographic organization as axons of distinct principal thalamic nuclei target specific cortical areas, which reciprocally connect onto the same thalamic nuclei (Crandall and Caviness, 1984; Jones, 1985). The development of the internal capsule was much studied not only because of the major functional relevance of this projection but also because it is a suited system to address axon guidance in the vertebrate brain due to its long path through the developing brain and its relative easy observation due to its important size (Fig. 20.4; Miller et al., 1993; Metin and Godement, 1996; Richards et al., 1997; Molnar et al., 1998; Auladell et al., 2000). TCAs extend from E12 to E13.5, cross the prethalamus, diencephalon/telencephalon boundary, and enter the subpallium. Within the subpallium, they navigate internally through the medial ganglionic eminence (MGE), through the lateral ganglionic eminence (LGE), and into the pallium (Fig. 20.4A; Chapter 16). Conversely, the first corticofugal axons, which are sent by subplate neurons from E12.5, cross the pallium/subpallium boundary (PSPB) at E13.5 and pause in the proximal part of the LGE (Fig. 20.4A). Corticofugal and TCAs meet at E14 in the LGE where they follow the similar paths in opposite directions. On the one hand, TCAs cross the PSPB and progress to the subplate of their target cortical area from E14.5 to E18.5, synapse onto subplate neurons, and, after a “waiting period,” extend collaterals into the cortical plate to reach layer IV pyramidal

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FIGURE 20.4 Corridor cells migration in the development of thalamocortical projections. The top panels (A,B,C) represent schematic hemicoronal sections of a mouse embryonic telencephalon, whereas the bottom panels (D,E) represent median sagittal views of a mouse embryonic telencephalon. (A) Thalamocortical axons (TCAs) extend from the dorsal thalamus (DTs) to the neocortex (Ncx) and cross different structures: the prethalamus (PTs), the diencephalonetelencephalon boundary (DTB, red dashes), the subpallium (SuP), the palliumesubpallium boundary (PSPB, red dashes). TCAs pioneer the internal capsule until they meet subplate axons (purple) in the SuP near the PSPB. Different cellular and molecular actors were involved in TCA guidance: PT and SuP cells sending an axon to the DT, the repellent Slit2 in the hypothalamus, the attractant Netrin1 in the SuP, Nrg1-Ig at the PSPB. (B,D) At E12, a tangential cell migration (pink cells) from the LGE to the MGE is repelled by Slit2 and builds a permissive corridor for TCA between the MGE subventricular zone and the globus pallidus (GPs). The corridor extends toward the caudomedial side of the SuP. TCAs are repelled by the hypothalamic source of Slit2 and grow toward the PSPB. (C) At E14, pioneer TCAs grow through the permissive corridor (pink cells) and striatum. The permissivity of these structures relies on the membrane-bound cue Nrg1-CRD. C, caudal; d, dorsal; DT, dorsal thalamus; DTB, diencephalone telencephalon boundary; GP, globus pallidus; l, lateral; LGE, lateral ganglionic eminence; m, medial; MGE, medial ganglionic eminence; Ncx, neocortex; PSPB, palliumesubpallium boundary; PT, prethalamus; r, rostral; St, striatum; v, ventral.

neurons (Lund and Mustari, 1977; De Carlos and O’Leary, 1992; Allendoerfer and Shatz, 1994; Herrmann et al., 1994). On the other hand, corticofugal axons grow along the path of TCA into the diencephalon, where CTAs reach the thalamus from E15.5 and other corticofugal axons turn ventrally toward subcortical targets (Jones et al., 1982; McConnell et al., 1989; De Carlos and O’Leary, 1992; Terashima, 1995; Auladell et al., 2000). Thus, while TCAs pioneer the internal capsule throughout the subpallium, corticofugal axons pioneer the internal capsule in the cerebral cortex and across the pallium/subpallium boundary.

20.4.2 Pioneer cortical subplate axons in the pathfinding of thalamocortical projections Structures and transient axonal populations located along the path of TCAs have been shown to delineate the complex trajectory of these projections. Pioneer cortical subplate neurons have first been proposed to act in this process, by acting as a scaffold for the further progression of TCA into and inside the cerebral cortex (Fig. 20.4; McConnell et al., 1989; Kostovic and Rakic, 1990; for review, Hoerder-Suabedissen and Molnar, 2015; Luhmann et al., 2018). These axoneaxon interactions have been further proposed to participate in the general formation of the internal capsule as well as the

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topographic organization of these projections (“handshake hypothesis”) (Blakemore and Molnar, 1990; Molnar and Blakemore, 1991; reviewed in Molnar and Blakemore, 1995). While topographic ordering of TCAs has been shown to rely on positional information located in the subpallium (Dufour et al., 2003; Bonnin et al., 2007; Wright et al., 2007; Powell et al., 2008; Demyanenko et al., 2011a; Demyanenko et al., 2011b) and the cerebral cortex (Barbe and Levitt, 1992; Gao et al., 1998; Mann et al., 1998; Mackarehtschian et al., 1999; Mann et al., 2002; Takemoto et al., 2002; Uziel et al., 2002; Bolz et al., 2004; Shimogori and Grove, 2005), several lines of experiments are consistent with a role of subplate axons in the pathfinding of TCA within the neocortex as well as along the internal capsule trajectory. First, the local ablation of subplate neurons in the visual cortex prevents the invasion of corresponding thalamic geniculocortical axons (Ghosh et al., 1990; Ghosh and Shatz, 1993). Consistently, in mutant mice that have an impaired preplate splitting and thus subplate cells in the marginal zone, such as reeler, cdk5
/
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mutant mice, TCAs form oblique bundles across the cerebral cortex to reach the ectopic subplate (Caviness and Rakic, 1978; Caviness, 1982; Caviness and Frost, 1983; Yuasa et al., 1994; Ohshima et al., 1996; Chae et al., 1997; Gilmore et al., 1998; Molnar et al., 1998; Molnar and Cordery, 1999; Rakic et al., 2006). In addition to a role in the guidance of TCAs inside the neocortex, subplate cortical axons were also shown to guide TCAs across the pallial/subpallial boundary (for review, Hoerder-Suabedissen and Molnar, 2015). Initial studies on Tbr1, Gbx2, and Pax6 mutant mice have defective correlated corticothalamic and thalamocortical pathways suggesting that the formation of these axonal projections is interdependent (Hevner et al., 2002). Further on, conditional mutants of Apc in the neocortex, which largely disrupts neocortex development and the formation of descending cortical projections, were used to graft back wild-type cortical explants to assess the roles of pioneer cortical axons in the guidance of TCAs (Chen et al., 2012). These ex vivo experiments reveal that the outgrowth of pioneer cortical axons is sufficient to regulate the entrance of TCAs into the neocortex, supporting a model of axoneaxon interactions in the formation of the internal capsule (Deck et al., 2013; for review, Garel and Lopez-Bendito, 2014; Squarzoni et al., 2015; Anton-Bolanos et al., 2018; LopezBendito, 2018). At the molecular level, several adhesion and guidance molecules have been proposed to regulate the interactions between pioneer cortical axons and TCAs. The transmembrane protein Linx is highly expressed by cortical axons, promotes the growth of thalamic axons and cortex-specific inactivation of the Linx gene (Mandai et al., 2014), and prevents TCAs from entering into the neocortex primordium. Similarly, mutant analyses and in vivo rescue experiments showed that the guidance molecule Draxin produced by cortical neurons is required for the entry of TCA into the neocortex (Shinmyo et al., 2015). Genetic interactions reveal that Draxin acts through the receptors deleted in colorectal cancer (Dcc) and neogenin (Neo), by promoting the outgrowth of TCAs (Shinmyo et al., 2015). Collectively, these experiments put forward a role for pioneer cortical subplate axons in regulating the ingrowth of TCAs across the pallial/subpallial boundary, by the local production of adhesion or secreted factors (Garel and Lopez-Bendito, 2014; Anton-Bolanos et al., 2018; Lopez-Bendito, 2018).

20.4.3 Origin and migration of subplate neurons Subplate neurons, which constitute the earliest born neurons in the cerebral cortex together with CajaleRetzius cells, exert essential functions during corticogenesis (for review, Hoerder-Suabedissen and Molnar, 2015; Luhmann et al., 2018). As aforementioned, they control not only TCA pathfinding but also the migration and polarity of cortical neurons (Namba et al., 2014; Ohtaka-Maruyama et al., 2018) and constitute a major early relay of incoming thalamic activity (for review, Allendoerfer and Shatz, 1994; Luhmann et al., 2018). The subplate is formed by diverse neuronal populations, including early-born excitatory cells as a vast majority, some inhibitory interneurons originating from the ganglionic eminences, as well as cells undergoing migration (Hoerder-Suabedissen and Molnar, 2015). In contrast to their assumed local origin in the neocortex, in vivo tracing experiments revealed that a large population of subplate neurons is generated in the proliferative zone of the rostromedial telencephalic wall (RMTW), near the septum, and reaches their position by tangential migration (Pedraza et al., 2014). Intriguingly, the RMTW also produces CajaleRetzius cells (Pedraza et al., 2014), supporting the idea that these two early-born populations share common features. This pallial albeit extracortical origin indicates that at least part of subplate guidepost cells is positioned by a timely controlled migratory process. The molecular and cellular pathways regulating this remarkable process remain, however, entirely to be explored.

20.4.4 The subpallium is a major intermediate target for thalamocortical axons While subplate neurons participate in the guidance of TCAs into and inside the cerebral cortex, TCA pathfinding toward and throughout the subpallium has been shown to rely on direct interactions with “local” structures (Fig. 20.4). First, the hypothalamus exerts a repulsive activity on TCA that prevents axons from entering this territory (Braisted et al., 1999),

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which is mediated by the repellent Slit2 via Robo1 and Robo2 receptors (Bagri et al., 2002; Lopez-Bendito et al., 2007; Braisted et al., 2009; Bielle et al., 2011). Channeling of TCAs into the subpallium also relies on early axonal projections between the prethalamus and the subpallium (Feng et al., 2016). Further along the axonal trajectory, the subpallium is shown to exert an attraction for both TCAs and corticofugal axons and constitutes a main intermediate target for these connections (Metin and Godement, 1996; Braisted et al., 1999). Analyses of mutant mice affecting the regionalization and development of the subpallium (Marin, 2002) have revealed the relative importance of the LGE and MGE in guiding TCAs. In particular, mutants affecting more specifically the development of the LGE, such as Ebf1 or Gsh1;Gsh2 doubles mutants, severely impair TCA pathfinding, in contrast to mutants that perturb MGE development, such as Nkx2.1 mutants (Garel et al., 1999; Sussel et al., 1999; Marin et al., 2002; Yun et al., 2003). Thus, the LGE and its derivatives seem to play an essential role in the guidance of TCAs. In addition, ex vivo experiments in embryonic brain slices have additionally revealed that the pallium produces a growth-promoting activity for TCAs, which is in part mediated by the secreted isoform of Neuregulin1 and its receptor ErbB4 (Lopez-Bendito et al., 2006). In parallel to the identification of the structures regulating TCA pathfinding through the subpallium, several studies were conducted to determine the molecular nature of the guidance activities involved. In particular, analyses of mice carrying mutations for guidance cues or their receptors have implicated Netrin1, Slit1 and Slit2 and their receptors Robo1 and Robo2 as well as Sema6A in the general pathfinding of TCAs in the subpallium (Braisted et al., 2000; Leighton et al., 2001; Bagri et al., 2002; Bonnin et al., 2007; Lopez-Bendito et al., 2007; Powell et al., 2008; Braisted et al., 2009; Little et al., 2009). In addition, members of the protocadherin family were shown to play essential roles in TCA guidance and internal capsule formation (Tissir et al., 2005; Uemura et al., 2007; Zhou et al., 2008, 2009). In particular, Celsr3 is a seven-pass cadherin ortholog to drosophila flamingo, which acts in both the planar cell polarity pathway and neurite outgrowth and is widely expressed in the mantle of the telencephalon and forebrain. Its specific inactivation in the subpallium and prethalamus severely impairs the formation of the thalamocortical connections: TCAs stall in the ventral subpallium just after crossing the telencephalic/diencephalic boundary, whereas cortical axons stall in the proximal part of the LGE (Zhou et al., 2008, 2009). These studies revealed an absolute requirement of Celsr3 expression by an intermediate target that acts at short range and demonstrated in vivo the major function of such intermediate target. Noteworthy, constitutive mutants in the Frizzled3 gene that is known in drosophila to participate in the planar cell polarity pathway with flamingo have a very similar phenotype in the pathfinding of the internal capsule, suggesting that the two genes also cooperate in mice during this major axonal wiring event (Wang et al., 2002, 2006). Finally, Netrin1, Sema3A and its receptor Npn1, ephrinA5 and its receptors EphA4 and EphA7, as well as L1 and CHL1, which are members of the immunoglobulin superfamily of cell adhesion molecules, have been shown to regulate the topography of TCAs inside the internal capsule and orient distinct thalamic axons toward different cortical domains (Dufour et al., 2003; Bonnin et al., 2007; Wright et al., 2007; Powell et al., 2008; Demyanenko et al., 2011a, 2011b). Thus, while these molecules do not modify the overall pathfinding of the internal capsule, they act on TCAs to precisely orient their positioning inside the axonal bundle. In sum, the subpallium is a major intermediate target for TCA, and the LGE is particularly involved in this guidance activity (Fig. 20.4A). At the molecular level, a series of secreted and transmembrane molecules expressed in the subpallium were shown to participate in TCA pathfinding and have provided definitive evidence for the in vivo requirement of this intermediate target.

20.4.5 Guidepost cells in the diencephalic and subpallial pathfinding of thalamocortical projections In parallel, different populations of guidepost neuronal cells have been proposed long ago to act in the pathfinding of thalamocortical projections. In particular, a group of neurons called “perireticular cells” or “internal capsule cells” are located in the future path of the internal capsule and send a cellular process to the thalamus at E12.5 in mouse embryos (Fig. 20.4). By analogy with the transient subplate cells, perireticular cells were suggested to provide a scaffold for CTAs and TCAs (Mitrofanis 1992, 1994; Mitrofanis and Baker, 1993; Mitrofanis and Guillery, 1993; Adams and Baker, 1995; Metin and Godement, 1996; Molnar et al., 1998; Braisted et al., 1999; Molnar and Cordery, 1999). The embryological origin, the molecular identity, the fate, and the potential role in TCA guidance of these back-labeled cells remain to be determined, as their widespread distribution and the lack of molecular markers have so far limited their characterization and specific manipulation. Nevertheless, their guidepost role for thalamocortical axons is compatible with the phenotype observed in several mutants: the absence, decrease in number, and displacement of perireticular or back-labeled cells in Mash1, Lhx2
/
, and Emx2
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mutants, respectively, lead to correspondent guidance defects of TCAs (Tuttle et al., 1999; Bishop et al., 2000; Lopez-Bendito et al., 2002; Bishop et al., 2003; Lakhina et al., 2007).

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In addition, a distinct population of TCA guidepost cells has been identified. These cells are GABAergic neurons that migrate tangentially from the LGE into the MGE and form a cellular “corridor” between the proliferative zones of the MGE and the globus pallidus (Fig. 20.4BeE; Lopez-Bendito et al. 2006). Accordingly, they are located in the MGE, express molecular markers of LGE-derived neurons, such as Islet1, Ebf1, and Meis2, and do not express MGE molecular markers such as Nkx2.1. These neurons, named “corridor cells,” migrate from E11.5 to E14 in the superficial mantle of the subpallium in a ventral direction, superficially to the large stream of MGE-derived interneurons that migrate toward the cerebral cortex. Ex vivo analysis in embryonic brain slices has shown that corridor cells constitute a permissive territory for the internal growth of TCAs through MGE-derived territories, which are overall nonpermissive for TCAs. While the factors controlling the nonpermissive activities of MGE-derived territories remain to be determined, corridor cells were shown to express a membrane-bound isoform of Neuregulin1 (Fig. 20.4). As previously mentioned, TCAs express the Neuregulin1 receptor ErbB4, and gain-of-function experiments in embryonic slices as well as telencephalic conditional Neuregulin1 mutants and constitutive ErbB4 mutants indicate that this signaling pathway regulates the pathfinding of TCAs throughout the corridor. The migration of corridor neurons thus generates a Neuregulin1 permissive domain essential for the internal pathfinding of TCAs within the subpallium. As such, corridor cells are immature neurons that act via contact or a short-range activity to position an axonal tract and constitute bona fide guidepost cells. Additional cues located on corridor neurons have been more recently shown to regulate TCA progression, in particular membrane-bound factors including the transmembrane molecule Linx (Mandai et al., 2014), Celsr3, or Fzd3 (Feng et al., 2016). Notably, Celsr3 and Fzd3 expression is required in Islet1-expressing guideposts that are located not only in the corridor but also on adjacent prethalamic cells, which both extend an axonal scaffold across the diencephalic/telencephalic junction and guide TCAs (Feng et al., 2016). This finding raises the possibility that some of these neurons are perireticular cells, which are located in the prethalamus and vicinity in a distinct position from corridor cells (Fig. 20.4AeC). These guidepost populations could act at distinct steps: Perireticular cells would regulate the entrance of TCAs into the subpallium, whereas corridor cells orient the internal pathfinding of TCAs inside the MGE (Lopez-Bendito et al., 2006). Further analyses will determine whether perireticular and corridor cells may overlap to some extent and thereby reveal whether some corridor cells may additionally act by axon-mediated contact. In addition to channel TCAs along an internal path, corridor neurons express gradients of guidance cues along the rostrocaudal axis that enable the initial topographic addressing of subsets of TCAs to rostral, intermediate, or caudal cortical regions (Bielle et al., 2011). In particular, they express Netrin1, Slit1, Sema3A, and ephrinA5 that were all shown to orient TCAs along the rostrocaudal axis. Intriguingly, combination of the guidance cues Netrin1 and Slit1 in this context triggered emergent unpredictable responses (Bielle et al., 2011; Leyva-Diaz et al., 2014; Dupin et al., 2015), putting forward the corridor as a major guidance structure (for review, Garel and Lopez-Bendito, 2014). Consistently, transient defects in corridor migration and organization in conditional Ebf1 mutants impair the transfer of a precise topographic map between the thalamus and neocortex (Lokmane et al., 2013). Thus, corridor cells constitute key guidepost cells, which enable not only the channeling of TCAs in the internal capsule but also their initial topographic arrangement en route to the neocortex (Garel and Lopez-Bendito, 2014).

20.4.6 Migration of guidepost corridor cells: routes and guidance cues Given the short range of their guidepost action, the positioning of corridor cells by tangential migration is at the core of their action. Indeed, the cellular movement from the LGE to the MGE territory allows the precise localization of a permissive environment in the range of TCA growth cones. The identification of corridor cells as LGE-derivatives raises the question of the mechanisms governing their specification in respect to radially LGE-derived striatal neurons, as well as of the cues that orient their migration and arrest. We have so far no information on the molecular mechanisms governing differential cell specification among radially migrating and tangentially migrating LGE derivatives. However, striatal neurons share with corridor cells the expression of several transcription factors (Islet1, Ebf1, Meis2, Ctip2), whereas others appear more specific to striatal neurons (Foxp1, Foxp2) (Bielle et al., 2011), thereby providing grounds for further studies. In regard to the migratory routes of corridor neurons, static three-dimensional analyses of the corridor indicate that this structure has a funnel shape that runs from the LGE and converges toward the diencephalic/telencephalic boundary (Fig. 20.4BeE; Bielle et al., 2011). The corridor is thus likely formed by a tangential migration generated from different rostrocaudal domains of the LGE, but precise in utero fate mapping will be required to address this issue. While the positioning of corridor cells relies on the transcription factor Pax6 (Simpson et al., 2009) and on the OL protocadherin (Uemura et al., 2007), the factors that attract the ventral migration of corridor cells are still unknown. Recently, a repulsive activity from the ventral subpallium has been shown to limit the ventral migration of these cells and

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participates in shaping the corridor (Fig. 20.4) (Bielle et al., 2011). At the molecular level, in vitro, in vivo, and ex vivo experiments show that the repellent Slit2, via Robo1 and Robo2 receptors, mediates this repellent activity. The abnormal migration of corridor cells induced by Slit2 inactivation leads to a malformation of the corridor shape that does not extend up to the ingrowing TCAs. Ex vivo rescue experiments indicate that the defects in corridor cells migration are largely responsible for the abnormal ventral misrouting of TCAs in the subpallium of Slit2 mutants. Thus, Slit2 regulates the migration of corridor cells, which in turn delineate the internal navigation of TCAs. If these experiments implicate Slit2 in corridor cells migration, they also show that guidance cues have multiple and complex activities in the formation of axonal tracts via a direct effect on the axonsdSlit2 in the hypothalamus repels axonsdand via the positioning of migrating guidepost cells (Fig. 20.4DeE). Furthermore, they open the way for future studies aimed at unraveling how guidance cues previously implicated in TCA pathfinding act either directly on axons or via guidepost cells positioning.

20.4.7 Fate of guidepost cells for thalamocortical projections Subplate neurons, such as CajaleRetzius cells of the neocortex, are eliminated during early postnatal life in mice, but some might be persistent in adults, in particular in humans (Hoerder-Suabedissen and Molnar, 2015; Meyer and GonzalezGomez, 2018). Fate mapping of corridor cells, however, recently revealed that these neurons are part of a larger stream of migration generated in the LGE, which gives rise to components of the extended central amygdala, including the bed nucleus of the stria terminalis (Tinterri et al., 2018). While it is technically impossible so far to specifically mark guidepost cells and trace their derivatives, combination of cre-driven fate mapping and intersectional genetics support the hypothesis that corridor cells are part of a larger population of GABAergic projection neurons of the extended amygdala, a structure conserved across vertebrates (Tinterri et al., 2018). As such, it implies that corridor-like neurons should exist in nonmammalian vertebrates, which has previously been reported (Bielle et al., 2011), even though they do not act as guidepost in these species that lack an “internal” capsule. Thus, the role of guideposts for TCAs would constitute an evolutionary more recent function of corridor neurons. Regardless of evolutionary perspectives, these findings reveal that transient guidepost interactions in the context of axon guidance can occur not only with transient cells but also with neuronal populations that will contribute to adult brain structures.

20.5 Neuronal migration of guidepost cells in the formation of the corpus callosum 20.5.1 Anatomy and development of the corpus callosum The corpus callosum is a large dorsal axonal commissure that interconnects the two cerebral hemispheres and allows the communication between the right and left dorsal regions of the cerebral cortex (Fig. 20.5). In contrast to the anterior commissure, which interconnects ventral cortical regions and is present in all the vertebrates, the corpus callosum is a hallmark of eutherian mammals. If callosal neurons constitute a heterogeneous population (Molyneaux et al., 2009; for review, Fame et al., 2011), they are all pyramidal neurons specified by the transcription factor Satb2 and located in upper and lower layers of the cerebral cortex (Yorke and Caviness, 1975; Porter and White, 1983; Alcamo et al., 2008; Britanova et al., 2008). The development of the corpus callosum has been shown to rely on a series of cellular events (Silver et al., 1982; Hankin et al., 1988; Hankin and Silver, 1988; Shu and Richards, 2001; Shu et al., 2003a, 2003b, 2003c; Gobius et al., 2016; for review, Lindwall et al., 2007; Chedotal, 2011): (1) the fusion of the midline of the two cerebral hemispheres; (2) the attraction toward the midline of pioneer axons of the cingulate cortex; (3) the channeling of these axons superficially to the septum toward the contralateral hemisphere; (4) the ingrowth and pathfinding across the dorsal midline of the rest of callosal axons; and (5) the guidance of axons in the contralateral hemisphere toward their cortical target. During mouse embryogenesis, axons of the cingulate cortex pioneer the tract at E15.5 (Koester and O’Leary, 1994; Ozaki and Wahlsten, 1998; Rash and Richards, 2001; Piper, Plachez et al., 2009), followed by frontal callosal neurons and, finally, parietal cortical callosal neurons. At E18.5, callosal axons have a dorsoventral topographic organization within the corpus callosum: Cingulate and frontal axons are located in a dorsal pathway, whereas parietal axons occupy a more ventral domain of the axonal bundle (Fig. 20.5; Niquille et al., 2009).

20.5.2 Roles of glial cells in the development of the corpus callosum During the sequential phases of corpus callosum formation, distinct subpopulations of glial cells have been shown to play essential roles (Silver et al., 1993; Shu and Richards, 2001; Shu et al., 2003a, 2003c, 2003d; Lent et al., 2005; Smith et al., 2006). First, the “midline glial zipper” and astroglial cells have been proposed to regulate midline fusion (Gobius et al.,

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FIGURE 20.5 Guidepost neurons in callosal axons guidance. (A) The corpus callosum (cc) is the largest commissure of the brain and allows callosal axons from the cerebral cortex (CTX) of one hemisphere to project onto the controlateral cerebral cortex. The corpus callosum is a median structure located above the septum (Se). (B) Different callosal axons cross successively the corpus callosum: first cingulate axons (Cing, dark gray), second frontal axons (Front, dark green), and finally parietal axons (Par, light green). Glutamatergic neurons (pink) and later GABAergic neurons (yellow) migrate to the corpus callosum and build a network through which callosal axons grow. Glutamatergic neurons located at the setumecortex boundary form the sling that migrates from the lateral ventricles (pink arrows). Glutamatergic neurons release Sema3C (þ), which attracts callosal axons and guides them in combination with other guidance cues secreted by two glial structures, the indusium griseum (IG) and the glial wedge (GW). Cc, corpus callosum; Cing, cingulate callosal axons; CTX, cerebral cortex; Front, frontal callosal axons; GW, glial wedge; IG, indusium griseum; Par, parietal callosal axons; Se, septum; SuP, subpallium.

2016), in particular at the level of the septum (for review, Silver, 1993; Lindwall et al., 2007). More dorsally, at the corticoseptal boundary, the “glial sling,” which is composed of migratory cells moving converging toward the midline, has been shown to be essential to corpus callosum formation by providing a cellular substrate for callosal axons (Fig. 20.5; Silver et al., 1982; Shu et al., 2003b; Lent et al., 2005; Ren et al., 2006). In parallel, the “glial wedge” of radial glia fibers channels axons into the corpus callosum by secreting a diffusible repellent that prevents callosal axons from entering the septum (Fig. 20.5; Shu and Richards, 2001; Shu et al., 2003c, 2003d). And finally, glial cells of the indusium griseum have been shown to control the positioning of different cell types in the corpus callosum region, including pioneer cingulate neurons (Shu et al., 2003c; Smith et al., 2006). At the molecular level, these glial cells have been proposed to act on axon guidance by expressing short-range and long-range guidance cues (Silver and Ogawa, 1983; Silver et al., 1993; Shu and Richards, 2001; Richards, 2002; Shu et al., 2003c, 2003d; Lent et al., 2005; Smith et al., 2006; for review, Lindwall et al., 2007), including Slits (Erskine et al., 2000; Plump et al., 2002; Shu et al., 2003d), Wnts (Keeble et al., 2006), ephrins (Williams et al., 2003; Mendes et al., 2006), and draxin (Islam et al., 2009). In particular, Slit2 expression in the glial wedge repels callosal axons mainly via Robo1 receptors, and this activity is essential to channel axons across the midline and prevent them from entering the septum (Shu et al., 2003d; Andrews et al., 2006). Analyses of mutant mice have revealed that several transcription factors, including NFIA, NFIB, and JSAP1 (das Neves et al., 1999; Shu et al., 2003a; Ha et al., 2005; Steele-Perkins et al., 2005; Piper et al., 2009a), as well as Fgf signaling, are required for the proper development of these glial cell populations (Smith et al., 2006; Tole et al., 2006). However, the mechanisms governing their specification, positioning, and relative functions in callosal guidance still remain to be fully unraveled.

20.5.3 Tangentially migrating neurons in the development of the corpus callosum As aforementioned, the subcallosal “glial” sling was initially described as a glial structure, generated by a medial cell migration from the adjacent ventricular zones (Fig. 20.5B). Its absence is associated with an acallosal phenotype, which can be rescued by a “sling graft,” supporting a major role for this structure in the guidance of callosal axons (Silver et al., 1982; Katz et al., 1983; Silver and Ogawa, 1983; Hankin and Silver, 1988; Silver et al., 1993; Shu and Richards, 2001).

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While the presence of neurons in the corpus callosum was reported in the cat, mouse, and human (Riederer and Innocenti, 1992; Riederer et al., 2004; Ren et al., 2006; Jovanov-Milosevic et al., 2010), recent studies have revealed that the “glial” sling is largely composed of glutamatergic immature neurons expressing NeuN, b-tubulin, calretinin, Emx1, and NFIA (Shu et al., 2003b; Ren et al., 2006; Niquille et al., 2009). They are generated from E14.5 to P2 and have been proposed to migrate medially from the ventricular zone next to the sling (Fig. 20.5; Shu et al., 2003b; Niquille et al., 2009) or from retrobulbar regions (Meyer and Goffinet, 1998; Meyer et al., 1998). While the sling is a transient structure disappearing at P10, the lack of detected apoptotic events raises the possibility that some of these neurons might migrate to another location (Hankin et al., 1988; Shu et al., 2003b). The diversity of neurons of the corpus callosum and their close relationship with callosal axons development were further characterized (Niquille et al., 2009). Above the calretinin-positive glutamatergic neurons of the sling, two other “stripes” of neurons with the same molecular identity are detected in the developing corpus callosum: One is located in the middle of the axonal bundle, whereas the other is located at its dorsal boundary (Fig. 20.5B). Intriguingly, these stripes of neurons delineate distinct topographic domains of the corpus callosum, as the “middle stripe” delineates the ventral parietal pathway and the dorsal frontocingulate pathway. While the specific function of this cellular organization remains to be investigated, further studies have revealed that calretinin-positive glutamatergic neurons regulate callosal axons pathfinding by secreting Sema3C. In vitro and ex vivo, Sema3C attracts cingular and neocortical callosal axons, and this activity is mediated by Npn1 expression on callosal axons (Bagnard et al., 1998, 2000; Niquille et al., 2009; Piper et al., 2009b). Attraction of callosal axons by Sema3C is silenced in postcrossing axons by the upregulation of EphB1 (Mire et al., 2018). Consistently, Sema3C or Npn1 invalidation leads to partial or complete agenesis of the corpus callosum (Niquille et al., 2009; Piper et al., 2009b), suggesting that calretinin-immunoreactive neurons play a major function in the formation of the corpus callosum via Sema3CeNpn1 signaling. A second neuronal population of GABAergic neurons intermingles with calretinin-immunoreactive neurons (Fig. 20.5; Niquille et al., 2009). These neurons likely originate in the ganglionic eminences and migrate by tangential migration into the corpus callosum anlage. Electron microscopy and confocal imaging showed that axons are in close apposition with both glutamatergic and GABAergic neurons of the corpus callosum and form a three-dimensional complex network (Niquille et al., 2009). The invalidation of Mash1 drastically reduces the number of GABAergic neurons in the corpus callosum, disorganizes calretinin-immunoreactive neurons, and leaves glial cells unaffected (Niquille et al., 2009). Collectively, these experiments suggest that GABAergic neurons might participate in the formation of a cellular scaffold important to support midline crossing of callosal axons (Fig. 20.5). While the exact function of GABAergic neurons remains to be characterized, these experiments show a role of two distinct tangentially migrating neuronal guidepost cells in the formation of the corpus callosum. In addition, they further reveal the complexity of the cellular network of neuronal and glial cells that emerges during callosum development and is required for its pathfinding. How these guidepost neurons are generated and positioned and how their function is coordinated with the one of glial cells will constitute a true challenge to understand the mechanisms governing corpus callosum formation.

20.6 Neuronal migration of guidepost cells and evolution of brain wiring 20.6.1 Tangential migration of guidepost neurons: a hallmark of the telencephalon? In the light of the experimental evidence presented, the positioning of guidance cues by tangentially migrating guidepost neurons appears as a major event in the formation of functionally essential telencephalic tracts. Indeed, the LOT and TCA convey all the sensory information to the cerebral cortex, the corpus callosum allows the intercommunication between the two cortical hemispheres, and the entorhinohippocampal pathway is the principal afference to the hippocampus. Is this phenomenon characteristic of this structure? The mammalian telencephalon is the site of several well-described tangential migration, and their number and characterization increases as scientists examine its development (Anderson et al., 1997a; Anderson et al., 1997b; Marin et al., 2000; Anderson et al., 2001; Wichterle et al., 2001; Stenman et al., 2003; Takiguchi-Hayashi et al., 2004; Bielle et al., 2005; Waclaw et al., 2006; Yoshida et al., 2006; Garcia-Moreno et al., 2008; Garcia-Moreno et al., 2010; for review, Marin and Rubenstein, 2003). Coherently, identified guidepost cells are localized in known sites of tangential migrations, such as the subpallium and the marginal zone of the pallium. Therefore, the fact that tangentially migrating streams are well documented in the mammalian telencephalon has likely been a decisive factor in the identification of migrating guidepost cells. As such, it will be of interest to examine whether known guidepost cells in the vertebrate brain, such as neurons of the optic chiasma, are also positioned by tangential migration (Sretavan and Reichardt, 1993; Sretavan et al., 1994; Marcus et al., 1995; Marcus

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and Mason, 1995; Sretavan et al., 1995; Jeffery, 2001; for review, Mason and Sretavan, 1997). Nevertheless, it is also possible that the large extent of tangential neuronal migration in the mammalian telencephalon has favored the emergence or the general “use” of such an axonal guidance mechanism. Conversely, tangential neuronal migration has both quantitatively and qualitatively increased during the evolution of the telencephalon in tetrapods, with a particular expansion in mammals and furthermore in primates (Letinic et al., 2002; Rakic and Zecevic, 2003; Bystron et al., 2005, 2006; Petanjek et al., 2009). The identification of migrating guidepost cells in the mammalian telencephalon thus raises the intriguing possibility that this particular mechanism of axon guidance has participated in the evolution of brain wiring.

20.6.2 Neuronal migration of guidepost cells in the evolution of the internal capsule The requirement of migrating guidepost cells to guide an axonal projection can seem at first glance a far from parsimonious way to build a new circuitry during evolution. Indeed, a dedicated cell type has to be specified, and positioned at the correct developmental time and place and then has to be endowed with an axon guidance activity. The whole process constitutes a series of steps unlikely to happen together by chance. However, if we consider the two examples of this guidance mechanism, the corpus callosum and internal capsule, their evolutionary context is very particular. In parallel to the functional emergence of the six-layered neocortex, these projections appeared in the mammalian telencephalon. Indeed, the corpus callosum is absent in noneutherian mammalian vertebrates, and the two pallial hemispheres are only connected by the anterior commissure, which travel through the subpallium. It is believed that the evolutionary addition of the corpus callosum allowed the fast transfer of information between the two cortical hemispheres as its volume grew. More “ancestrally”, thalamic projections underwent a change in their trajectory from ventral pallial and subpallial targets to the functionally emerging neocortex. Indeed, the development of an internal capsule, versus an external telencephalic peduncle in reptiles and birds, provides a short and extensive track for conveying sensory information directly to the neocortex. As such, both the trajectories of the mammalian internal capsule and the corpus callosum are evolutionary recent in tetrapods. Could we elaborate an evolutionary scenario through which these changes occurred? While additional studies on eutherians and marsupials will be required to address the mechanisms of corpus callosum evolution, comparative analysis on the migration of guidepost corridor neurons of the internal capsule has shown that these cells are evolutionary conserved (Bielle et al., 2011). Indeed, cells with a corridor immunophenotype (Islet1-positive, Nkx2.1-negative, FoxP2-negative) localized in the MGE adjacently to striatal projection neurons of the LGE (Islet1-positive, Nkx2.1-negative, FoxP2positive) are observed not only in mammals (human, sheep and mouse) but also in other amniotes (turtle, snake, and chick). Thus, corridor-like cells also exist in birds and reptiles that do not have an “internal” capsule. These observations indicate that this tangentially migrating neuronal population probably appears in stem amniotes and likely has an ancestral function independent of the guidepost one that it acquired in mammals (Bielle et al., 2011). Based on expression patterns and genetic tracings using Islet1-cre and Nkx2.1-cre lines (Srinivas et al., 2001; Xu et al., 2008), we found that corridorlike neurons are integrated in the central division of the extended amygdala, a basal ganglia subdivision involved in aversive response to negative stimuli (Tinterri et al., 2018). How corridor neurons may have acquired their guidepost function? Experiments in organotypic cultures using mouse and chicken embryonic brain slices indicate that chicken corridor-like cells have a guidepost activity on both mouse and chicken thalamic axons. However, in vivo, chicken corridor-like cells follow a different migratory route toward the midline and are never in contact with ingrowing thalamic axons (Fig. 20.6). The differential positioning of corridor cells by migration in chicken and mouse embryos was related to a repulsive activity located in the ventral MGE and subpallial preoptic area. Accordingly, comparative expression studies, in ovo gain-of-function experiments and mouse mutant analyses, indicated that a differential expression of the secreted repellent Slit2 participates in the specific positioning of corridor cells in these two species (Fig. 20.6). Thus, the “gain” of a guidepost role by the corridor cells population in amniotes seems to require principally a different direction of migration, and a change in the expression pattern of the repulsive factor Slit2 could participate in this process. These results show that minor modifications in the orientation of the positioning of a preexisting neuronal population is able to induce a large-scale long-range modification in neural wiring, by switching TCAs from an external to an internal path. These findings reveal that the local modulation of tangential neuronal migration can provide unique spatially and temporally regulated opportunities for the formation of novel axonal connections during brain evolution. They also open the way for further studies on the formation of the corpus callosum as well as on the general role of guidepost neurons distribution in the evolution of brain wiring.

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External External FIGURE 20.6 Corridor cells migration shapes the development of thalamic projections in different species. (AeE) Schematic median sagittal views of the left hemisphere of a mouse embryonic telencephalon (AeD) and of the chicken embryonic telencephalon (E,F). (A,B) At E12, corridor cell migration (pink cells) extends from the LGE to the MGE and is repelled by Slit2 allowing the corridor to extend toward the caudomedial side of the subpallium. TCAs are repelled by the hypothalamic source of Slit2 and grow toward the PSPB. At E14, TCAs have met the corridor cells and take an internal path. (C,D) In Slit2
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mutant, corridor cell migration (pink cells) abnormally extends toward the midline. At E14, TCAs have not met the corridor cells at the caudomedial side of the subpallium and take an external path. (E,F) In the chicken at E5.5, Slit2 expression is reduced compared with the mouse embryo, and corridor cell migration (pink cells) extends from the LGE to the midline. At E6, TCAs have not met corridor cells at the caudomedial side of the subpallium and take an external path. C, caudal; d, dorsal; DT, dorsal thalamus; DTB, diencephalonetelencephalon boundary; l, lateral; LGE, lateral ganglionic eminence; m, medial; MGE, medial ganglionic eminence; Ncx, neocortex; PSPB, palliumesubpallium boundary; PT, prethalamus; r, rostral; St, striatum; v, ventral.

20.7 Towards an integration of migrating guidepost neurons in normal and pathological brain development 20.7.1 Guidepost neurons in the shaping of axonal tract organization and topography Guidepost neurons delineate the future path of axonal tracts, raising the possibility that they could not only provide a permissive substrate for their pathfinding but also regulate the overall shape of the axonal bundle, the formation of collaterals, or the internal organization of tracts. In particular, the LOT, corpus callosum, and TCA projections are all compact axonal tracts that show an internal topographic organization (Molnar et al., 1998; Inaki et al., 2004; Niquille et al., 2009). Indeed, axons of distinct origins are presorted and grouped inside the axonal bundle. For instance, thalamic axons that relay different sensory input to specific cortical areas are located in a specific domain of the internal capsule, whereas callosal axons of frontal and parietal regions are located in dorsal or ventral paths of the corpus callosum, respectively. Pioneering studies of Sperry have showed that the topography of projections is controlled by graded labels present in the axons and in the final target. More recent studies have further indicated that the presorting of axons in intermediate targets participates in this topography (Garel et al., 2002; Dufour et al., 2003; Garel et al., 2003; Bonnin et al., 2007; Wright et al., 2007; Powell et al., 2008; Imai et al., 2009; Demyanenko et al., 2011a, 2011b). While axoneaxon interactions may participate in this presorting, corridor cells/TCA interactions were shown to regulate the local topography of axons (Bielle et al., 2011; Lokmane et al., 2013; Garel and Lopez-Bendito, 2014), raising the possibility that guidepost cells might be more generally involved in the axonal sorting and organization of the tract.

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20.7.2 Integrating tangential neuronal migration of guideposts in normal and pathological brain development Brain development is like a castle of cards in which all the cellular events occurring at the same time and place are integrated to build a remarkable and complex architecture that will enable its functioning. As such, tangential migration constitutes a remarkable process to coordinate cell distribution, morphogenesis of structures, and the guidance of axonal tracts. The identification of migrating guidepost cells constitutes in that regard the tip of the iceberg. Indeed, these studies reveal that morphogenetic events ensuring the development of the telencephalon require a global and systemic analysis and that guidance cues acting on cell migration and axon guidance can have interregulated activities. For instance, Neuregulin1 expression in the pallium and subpallium regulates the guidance of both GABAergic interneurons and incoming TCAs to the neocortex (Flames et al., 2004; Lopez-Bendito et al., 2006). Unraveling the complex cellular interactions in brain morphogenesis is particularly important to progress in our comprehension of pathological brain development. Indeed, in front of a functionally or anatomically abnormal brain, identifying the deficient steps and the causal processes constitutes a huge task. As many neurological and psychiatric diseases have been linked to abnormal neuronal migration (Chapter 38), the description of migration of guidepost cells opens novel perspectives. Indeed, it provides not only as a new possibly deficient step but also as a hub of development where neurogenesis, migration, morphogenesis, and axon guidance need to be all successfully integrated to achieve the building a functional connectivity. For the time being, the alteration of this mechanism in neuropathology remains completely hypothetical, but some speculations can be drawn from available data. Guidepost corridor cells are required for the development of the internal capsule in the mouse and are conserved in humans (Bielle et al., 2011). Alterations of corridor cells migration are thus expected to result in an abnormal internal capsule also in humans. Mutations of tubulin-a and tubulin-b isoforms TUBA1A and TUBB2B were identified in forebrain development defects in humans (Keays et al., 2007; Fallet-Bianco et al., 2008; Morris-Rosendahl et al., 2008; Jaglin et al., 2009; Kumar et al., 2010), for review (Jaglin and Chelly, 2009). The phenotypes associate corticogenesis defects (lissencephaly, heterotopia, polymicrogyria) with axonal tract defects (internal capsule and corpus callosum). Interestingly, the internal capsule was observed in one fetus with mutation of TUBA1A stopping just medial to the external globus pallidus (Fig. 5I and J in Fallet-Bianco et al., 2008), which corresponds to the position of the corridor in embryos. As TUBA1A mutations alter neuronal migration as shown by the observation of lissencephaly and heterotopia, corridor cells migration could be also altered so that corridor cells could not play their short-range guidepost role explaining the internal capsule defect. Although TUBA1A mutation could also directly affect axon growth and guidance, TCAs seem to achieve the pathfinding steps preceding the corridor crossing. The previous example addressed very dramatic morphological phenotypes related to early severe developmental defects. At the other side of the pathological spectrum, abnormal corridor cells could be involved in disease with very subtle morphological phenotypes such as schizophrenia. Indeed, in addition to their guidepost function, corridor cells are neurons involved in the basal ganglia emotion processing circuitry (Tinterri et al., 2018), and middle disturbance of corridor cells could hence lead to abnormal sensory information (due to TCA defects) and emotional information (due to basal ganglia defects) circuitries, both observed in schizophrenia. Therefore, the defects of guidepost cells migration could be predicted to lead to syndromes associating axonal defect and migration defect involving several cerebral structures. Corpus callosum agenesia associated with cortical interneuron defects or neuronal migration defects could constitute such a syndrome. Some mutations of Aristaless-related homeobox gene (ARX) cause the XLAG syndrome (X-linked lissencephaly with abnormal genitalia) with lissencephaly associated with corpus callosum agenesia and severe epilepsy associated with a reduction in cortical interneurons (Berry-Kravis and Israel, 1994; Dobyns et al., 1999a; for review, Okazaki et al., 2008; Shoubridge et al., 2010). Mutations in LIS1 and DCX also associated cortical interneuron defects and callosal abnormalities (Reiner et al., 1993; des Portes et al., 1998; Gleeson et al., 1998; Dobyns et al., 1999b; Deuel et al., 2006; Koizumi et al., 2006; Kappeler et al., 2007; Marcorelles et al., 2010). Further neuropathological studies and careful examination of mouse models for these mutations will allow to establish whether defects in cell migration affect in a coordinated manner the formation of structures and brain wiring, thereby providing novel grounds to our understanding of complex syndromes.

20.8 Conclusions With the recent discovery that tangential migration is a major actor in the morphogenesis of the mammalian telencephalon came the intriguing observation that it is additionally important to position cellular guideposts that control the pathfinding of major axonal tracts. These migrating guidepost cells are glutamatergic or GABAergic and have been associated with

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previously described sites of tangential migrations. While we are just beginning to understand the cellular and molecular pathways controlling these events and our knowledge is still rather fragmentary, these findings reveal that telencephalic development has to be considered as a dynamic process in which all cellular actors can interact and provide novel perspectives on their evolution and pathological development.

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Chapter 21

Neuronal migration in the postnatal brain Masato Sawada1 and Kazunobu Sawamoto1, 2 1

Department of Developmental and Regenerative Neurobiology, Institute of Brain Science, Nagoya City University Graduate School of Medical

Sciences, Nagoya, Japan; 2Division of Neural Development and Regeneration, National Institute for Physiological Sciences, Okazaki, Japan

Chapter outline 21.1. Introduction 465 21.2. Regulation of neuronal migration in the normal brain 466 21.2.1. Migratory scaffolds 466 21.2.1.1. Neighboring cells in the neuronal chain 466 21.2.1.2. Astrocytes 468 21.2.1.3. Blood vessels 469 21.2.2. Directional control from the V-SVZ toward the OB 469 21.2.3. Migration termination in the OB 470 21.3. Regulation of neuronal migration in the injured brain 470 21.3.1. Migratory scaffolds in the injured brain 470

21.3.1.1. Neighboring cells in the neuronal chain 470 21.3.1.2. Astrocytes 471 21.3.1.3. Blood vessels 472 21.3.1.4. Radial glial cells 472 21.3.2. Directional control toward a lesion 472 21.3.3. Enhancement of neuronal migration as a strategy for endogenous neuronal regeneration 473 21.4. Postnatal neuronal migration in primates 473 21.5. Summary 474 References 474

21.1 Introduction During mammalian brain development, neural stem cells (NSCs) residing in the periventricular region actively generate immature neurons, which travel to their final destinations in the brain through various migratory paths. After birth, the NSCs are restricted to two regions and generate neurons throughout life (Kempermann, 2011). The larger of these regions is the ventricular-subventricular zone (V-SVZ), which lines the lateral walls of the lateral ventricles, and the second region is the subgranular zone of the dentate gyrus in the hippocampus (Kempermann, 2011). Neurons are continuously produced in the V-SVZ. Under physiological conditions, these immature neurons migrate rapidly through a specialized migratory path called the “rostral migratory stream” (RMS) toward the olfactory bulb (OB), a primary processing center for odor information (Doetsch and Alvarez-Buylla, 1996; Lois and Alvarez-Buylla, 1994; Lois et al., 1996) (Fig. 21.1). After arriving at the OB, the migrating neurons differentiate into mature olfactory interneurons (Lois and Alvarez-Buylla, 1994; Luskin, 1993), which elicit various olfactory functions (Alonso et al., 2012; BretonProvencher et al., 2009; Moreno et al., 2009; Muthusamy et al., 2017; Sakamoto et al., 2011, 2014). After a brain injury, the V-SVZ-derived neurons migrate toward the lesion, where they differentiate into mature neurons (Arvidsson et al., 2002; Parent et al., 2002; Yamashita et al., 2006) (Fig. 21.1). These observations suggest that neurogenesis in the postnatal V-SVZ contributes to neuronal regeneration and functional recovery after brain injury. Notably, neuronal migration from the periventricular region is widely conserved among vertebrates, from fish to primates, including humans (Garcia-Verdugo et al., 2002; Paredes et al., 2016b; Sawada and Sawamoto, 2013). Thus, neurons generated in the V-SVZ have a remarkable, evolutionarily conserved potential to migrate for a long distance through the complex neuronal circuits of the postnatal brain. This postnatal migration is distinct from the neuronal migration occurring during brain development. In this chapter, we focus on the migration of neurons generated in the postnatal V-SVZ and summarize the unique regulatory mechanisms of this migration under physiological and pathological conditions.

Cellular Migration and Formation of Axons and Dendrites. https://doi.org/10.1016/B978-0-12-814407-7.00021-3 Copyright © 2020 Elsevier Inc. All rights reserved.

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FIGURE 21.1 Neuronal migration in the postnatal brain. (A) Neuronal migration toward the olfactory bulb (OB) under normal physiological conditions. Neurons generated in the ventricular-subventricular zone (V-SVZ) migrate in chains toward the OB through the rostral migratory stream (RMS), a specialized migratory path. (B) Neuronal migration after brain injury. V-SVZ-derived neurons migrate in chains toward the lesion.

21.2 Regulation of neuronal migration in the normal brain Neurons generated in the V-SVZ form linear cell aggregates called “chains,” and migrate at a high speed toward the OB through the RMS (Doetsch and Alvarez-Buylla, 1996; Lois and Alvarez-Buylla, 1994; Lois et al., 1996; Wichterle et al., 1997) (Fig. 21.2). After arriving at the OB, each neuron detaches from the chain (Fig. 21.2) and starts to migrate individually toward its final destination layer in the OB, where it differentiates into one of several kinds of mature olfactory interneurons (Lois and Alvarez-Buylla, 1994; Luskin, 1993) (Fig. 21.3). Recent evidence suggests that the long-distance neuronal migration in the postnatal brain is achieved by the appropriate control of scaffolds, directionality, and migration termination.

21.2.1 Migratory scaffolds 21.2.1.1 Neighboring cells in the neuronal chain Immature neurons generated in the V-SVZ form chain-like cell aggregates and move along each other during their migration toward the OB, suggesting that the neurons use their neighbors as a migratory scaffold in chain migration (Doetsch and Alvarez-Buylla, 1996; Lois and Alvarez-Buylla, 1994; Lois et al., 1996; Wichterle et al., 1997). These neurons adhere to each other by forming electron-dense junction complexes called “adherens junction (AJ)-like structures,” which are observable by electron microscopy (Doetsch et al., 1997), suggesting that the control of intercellular adhesion between neurons contributes to the coordination of chain migration. Although the molecular identity of the AJ-like structures in chain-forming neurons is presently unknown, several cell adhesion molecules are reported to regulate the chain migration of neurons in the RMS. For example, polysialylated form of neural cell adhesion molecule (PSA-NCAM) is expressed in migrating neurons in the RMS. The PSA moiety is negatively charged and is thought to weaken the adhesion between NCAM molecules (Gascon et al., 2008; Rutishauser, 2008). Both enzymatic elimination of the PSA moiety and deletion of the NCAM gene disrupt neuronal migration from the V-SVZ to the OB, suggesting that the PSA moiety is involved in efficient neuronal chain migration (Chazal et al., 2000; Cremer et al., 1994; Ono et al., 1994). In addition, N-cadherin is a calcium-dependent cell adhesion molecule that is expressed in migrating neurons in the VSVZ and RMS (Yagita et al., 2009). N-cadherin-deficient mice showed disorganized neuron chains in their V-SVZ (Porlan et al., 2014), suggesting that N-cadherin contributes to the neuronal chain migration. Although cadherins are the central components of the AJs in epithelial cells, it remains unclear whether N-cadherin is involved in the formation and maintenance of the AJ-like structures observed between chain-forming neurons.

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FIGURE 21.2 Regulation of neuronal migration in the normal brain. (A) Chain migration of ventricular-subventricular zone (V-SVZ)-derived neurons in the rostral migratory stream (RMS). Migrating neurons (red) form chain-like cell aggregates and move along each other within the glial tube (blue) toward the olfactory bulb (OB) (left, green). Blood vessels provide a scaffold for the migrating neurons. The long-distance directional movement of the migrating neurons from the V-SVZ to the OB is determined by a combination of V-SVZ-derived repulsive factors (yellow arrow) and OB-derived attractive factors (green arrow). After arriving at the OB, neurons detach from the chains in response to detachment signals (red arrow) and start to migrate individually. Green lines, serotonin fibers. (B) Neuronal chains along blood vessels. Chain-forming neurons (red) adhere to each other by electrondense junction complexes called AJ-like structures. These neuronal chains are enwrapped in a glial tube (blue). Chain-forming neurons attach to blood vessels (pink) via the astrocytic endfeet.

In addition to the intercellular adhesions between neurons, interactions between neurons and the extracellular matrix (ECM) also control neuronal chain migration. ECM proteins such as laminin and Tenascin-C are concentrated within the RMS (Peretto et al., 2005), whereas a6b1-integrin and b8-integrin are expressed in migrating neurons (Belvindrah et al., 2007; Emsley and Hagg, 2003; Mobley and McCarty, 2011). The neuronal chain formation and migration in the RMS are disrupted in knockout (KO) mice lacking any of these proteins (Belvindrah et al., 2007; Mobley and McCarty, 2011). Moreover, cultured neurons from wild-type but not from b1-integrin KO mice form chains in a laminin coatingedependent manner, suggesting that laminin-integrin signaling controls neuronal chain migration (Fujioka et al., 2017). In addition, an ECM modulator, Disintegrin and metalloproteinase domain-containing protein 2 (ADAM2), promotes neuronal migration in the RMS (Murase et al., 2008), indicating that ECM remodeling may also be involved in efficient neuronal chain migration. Although the precise mechanisms of ECM-mediated neuronal chain formation and migration are still unknown, the chain-forming neurons might adhere to each other via thin layers of ECM proteins such as laminin. Alternatively, the ECM signaling might affect other adhesion molecules, like N-cadherin. For neurons to migrate efficiently in chains, their morphology and movement need to be coordinately controlled. During their migration in the RMS, neurons move in a saltatory manner, executed by the repeated extension of their leading process and subsequent somal translocation (Hikita et al., 2014; Schaar and McConnell, 2005). Consequently, the neurons exhibit resting and migratory phases, which are controlled by RhoA signaling (Ota et al., 2014; Shinohara et al., 2012). The resting neurons form an indentation in their soma in a Rac1-dependent manner, to make a path for migrating neurons (Hikita et al., 2014). Moreover, the neuronal chain formation and migration toward the OB are also affected by cytoskeletal regulation (Hirota et al., 2007; Khodosevich et al., 2009; Koizumi et al., 2006; Nakamuta et al., 2017; Sonego et al., 2013, 2015). These findings suggest that the spatiotemporal control of cell morphology and movement is critical for efficient neuronal chain migration occurring within the narrow migratory path in the RMS.

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FIGURE 21.3 Termination of neuronal migration in the olfactory bulb (OB). Neurons stop migrating either in the granule cell layer (GCL) or in the glomerular layer (GL) of the OB and differentiate into granule cells or periglomerular cells, respectively. The neuronal migration in the rostral migratory stream (RMS) and OB is negatively regulated by PlexinB2 and Gmip (blue). During the termination process of radial migration in the OB, the neurons transiently extend an filopodium-like lateral protrusion (FLP) from the proximal part of the leading process. Repulsive Sema3E-PlexinD1 signaling suppresses FLP formation and maintains the migratory morphology and radial migration of neurons until they reach their final position in the OB (red and green text). After they stop migrating, mature neurons begin to secrete Sema3E (green circles), to help maintain the migration capacity of later-born neurons. Neurotransmitters such as serotonin and acetylcholine promote the radial migration and subsequent survival of neurons in the OB (orange).

Although several adhesion molecules are reported to be involved in the neuronal chain migration, how these molecules function in chain migration is still largely unknown. In particular, how a large number of neurons, tightly packed within chains, can migrate simultaneously and smoothly without stacking in the RMS largely remains a mystery. More studies on the spatiotemporal control of the adhesion, movement, and morphology of migrating neurons are still needed to clarify the mechanisms of neuronal chain migration in the RMS.

21.2.1.2 Astrocytes Chain-forming neurons migrating in the RMS are surrounded by astrocytic tunnel-like structures called a “glial tube” (Fig. 21.2A) (Lois et al., 1996). The glial tube structure develops during the first 3 weeks of postnatal development in rodents (Bozoyan et al., 2012; Peretto et al., 2005). These astrocytes not only act as NSCs that produce neurons (Alonso et al., 2008; Gritti et al., 2002) but also physically separate the migrating neurons from the surrounding brain parenchyma. These astrocytes extend their processes in parallel with the neuronal chain and make direct contacts with the neurons (Kaneko et al., 2010; Lois et al., 1996). Glial tube-forming astrocytes in the RMS produce various ECM proteins that promote neuronal migration. Notably, when neurons that have arrived in the OB are transplanted back into the V-SVZ, they migrate again through the RMS toward the OB (Sawada et al., 2018). This finding suggested that the RMS provides migrating neurons with a special environment that maintains them in an immature state. Neuronal migration within the glial tube depends on repulsive Slit-Robo signaling between the neurons and the astrocytes (Kaneko et al., 2010). While migrating neurons secrete Slit1, the surrounding astrocytes express its receptor Robo2. Slit1 KO mice show disorganized glial tubes and defective neuronal migration in the RMS, suggesting that SlitRobo signaling maintains the glial tube structure by controlling the astrocytic distribution and morphology, thereby providing the specialized migratory path for neurons (Kaneko et al., 2010). Astrocytes in the RMS also help create an advantageous microenvironment for neuronal migration. The migrating neurons secrete GABA, which negatively regulates their migration speed through the receptor GABAAR on neurons (Bolteus and Bordey, 2004). Astrocytes express the GABA transporter GAT4, with which they take up excessive extracellular GABA within the RMS, to promote neuronal migration (Bolteus and Bordey, 2004). In addition,

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astrocyte-derived glutamate promotes the survival of migrating neurons in the RMS (Platel et al., 2010). Astrocytes also express ephrin-B2/B3, which promotes neuronal proliferation in the V-SVZ and neuronal migration in the RMS through their receptors EphA and EphB on neurons and astrocytes (Conover et al., 2000; Todd et al., 2017). Although the mechanisms for glial-tube formation and maintenance have been reported, the significance of the glial tube in neuronal migration remains poorly understood. Migrating neurons are surrounded by astrocytic tunnels in rodents (Lois et al., 1996) and primates (Gil-Perotin et al., 2009; Kornack and Rakic, 2001; Pencea et al., 2001), but not in zebrafish (Adolf et al., 2006; Grandel et al., 2006; Kishimoto et al., 2011a) or rabbits (Ponti et al., 2006). Therefore, comparisons of the neuronal migration with or without the glial tube in various vertebrates should help clarify the role of this structure in neuronal migration in the RMS.

21.2.1.3 Blood vessels During the neonatal stage, astrocytes in the RMS secrete vascular endothelial growth factor, which promotes the development of blood vessels that extend tangentially along the RMS (Bozoyan et al., 2012). These blood vessels are associated with the neurons in the RMS and act as a scaffold that promotes their migration (Snapyan et al., 2009; Whitman et al., 2009). During this blood vesseleguided migration, the neurons attach directly to the astrocytic endfeet enwrapping the blood vessels rather than to the vascular endothelial cells (Fig. 21.2) (Le Magueresse et al., 2012). Brain-derived neurotrophic factor (BDNF), which is secreted by vascular endothelial cells and presented by glial tube-forming astrocytes, promotes blood vesseleguided neuronal migration in the RMS (Snapyan et al., 2009). The mechanisms for blood vesseleguided neuronal migration have just started to be clarified. To understand the scaffold role of blood vessels, it is important to examine the cellular and molecular interactions between neurons and perivascular cell types, including astrocytes, pericytes, and smooth muscle cells. It is also possible that the perivascular space helps to create an advantageous microenvironment for neuronal migration, including guidance molecules derived from the blood flow and perivascular cerebrospinal fluid (CSF) flow. Given that well-developed vascular networks run throughout the brain, blood vesseleguided migration may be an important strategy used by neurons to reach their destinations.

21.2.2 Directional control from the V-SVZ toward the OB The direction of migrating neuron movement from the V-SVZ toward the RMS is determined by repulsive Slit-Robo signaling. Slit, produced from the choroid plexus and septum (Hu, 1999; Wu et al., 1999), forms a concentration gradient within the V-SVZ, as a result of CSF flow generated by the ciliary beating of ependymal cells (Sawamoto et al., 2006). Since migrating neurons express Robo, the Slit1 concentration gradient orients neurons rostrally, determining their migratory directionality within the V-SVZ (Sawamoto et al., 2006). Although Slit can explain the neurons’ direction of movement from the V-SVZ toward the RMS, it may not be sufficient to guide neurons toward the OB, due to the long distance between the V-SVZ and OB. Co-culture experiments of OB tissues and neurons suggested that attractive factors for neurons reside in the OB (Liu and Rao, 2003). Several OBderived growth and neurotrophic factors, including hepatocyte growth factor (HGF), BDNF, glial-derived neurotrophic factor, Netrin-1, and Prokineticin-2 (PK2), have been proposed to attract migrating neurons toward the OB (Chiaramello et al., 2007; Garzotto et al., 2008; Murase and Horwitz, 2002; Ng et al., 2005; Paratcha et al., 2006; Puverel et al., 2009). On the other hand, neurons still migrate rostrally in the RMS even after surgical elimination of the OB (Kirschenbaum et al., 1999). Thus, neurons are guided toward the OB by mechanisms that are both dependent on and independent of OBderived attractive factors. In addition to the long-distance guidance, local factors that promote neuronal migration also contribute to the directional neuronal migration in the RMS. Migrating neurons express ErbB4, whose ligands Neuregulin-1/2 are expressed in the RMS (Anton et al., 2004). ErbB4-deficient neurons move in altered directions in the RMS (Anton et al., 2004). In addition, vascular endothelial cell-derived BDNF acts as a local guidance cue for blood vesseleguided neuronal migration in the RMS (Snapyan et al., 2009). A recent study showed that serotonin secreted from serotonergic fibers running along the RMS also promotes neuronal migration (Garcia-Gonzalez et al., 2017). Notably, the serotonergic fiber-guided neuronal migration is observed not only in mice but also in zebrafish, zebra finches, rhesus monkeys, and humans (Garcia-Gonzalez et al., 2017), indicating that this mechanism is widely conserved among vertebrates. Thus, the direction of the long-distance movement of migrating neurons from the V-SVZ toward the OB is determined by a combination of V-SVZ-derived repulsive factors, OB-derived attractive factors, and local factors that promote neuronal migration.

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21.2.3 Migration termination in the OB After arriving at the OB, neurons detach from the chains and start to migrate individually. These neurons terminate their migration in either the granule cell layer or the glomerular layer of the OB and differentiate into granule cells or periglomerular cells, respectively (Lois and Alvarez-Buylla, 1994; Luskin, 1993). Altering the neurons’ migration affects their final positioning, dendritic patterns, and functions in the OB (Belvindrah et al., 2011; Petri et al., 2017; Sawada et al., 2018), suggesting that appropriate control of the maintenance and termination of the radial migration of neurons is critical for the proper formation and function of neuronal circuits in the OB. Reelin, PK2, and Tenascin-R show activity for the detachment of cultured chain-forming neurons (Hack et al., 2002; Ng et al., 2005; Saghatelyan et al., 2004). In mouse mutants of these molecules, neurons in chains accumulate in the core of the OB, indicating that Reelin, PK2, and Tenascin-R act as detachment signals for the chain-forming neurons in vivo (Hack et al., 2002; Ng et al., 2005; Saghatelyan et al., 2004). Interestingly, the expression levels of Reelin and Tenascin-R are affected by olfactory activity (Okuyama-Yamamoto et al., 2005; Saghatelyan et al., 2004), suggesting that olfactory input regulates the process of neuronal detachment from chains. During the detachment process, the intercellular adhesion between chain-forming neurons decreases. In this process, the chain-forming neurons show decreased expressions of NCAM and b1-integrin, regulated by a downregulation of sphingosine-1-phosphate receptor 1 (Alfonso et al., 2015). Electron microscopic analyses revealed that the density of AJlike structures in chain-forming neurons in the anterior RMS is lower than that in the posterior one (Fujikake et al., 2018). Reelin-Dab1 signaling and Fyn, a nonreceptor tyrosine kinase, coordinately promote neuronal detachment from the RMS by decreasing the AJ-like structures (Fujikake et al., 2018). These findings suggest that neurons detach from the chains by decreasing their intercellular adhesions in response to various detachment signals. Neuronal migration in the RMS and OB is negatively regulated by PlexinB2 (Saha et al., 2012) and the RhoGAP protein Gmip (Ota et al., 2014). In the termination process of radial migration in the OB, the neurons transiently extend a long protrusion called a filopodium-like lateral protrusion (FLP) from the proximal part of their leading process during the resting phase of migration (Fig. 21.3) (Sawada et al., 2018). Repulsive Sema3E-PlexinD1 signaling suppresses the FLP formation and maintains the neurons’ migratory morphology and radial migration until they reach their final position in the OB (Sawada et al., 2018). Neurotransmitters such as serotonin (Garcia-Gonzalez et al., 2017) and acetylcholine (Kaneko et al., 2006) also control the radial migration and subsequent survival of neurons in the OB. Thus, both morphological/cytoskeletal regulation and neuronal activity in the migrating neurons contribute to their appropriate migration and maturation in the postnatal OB. In contrast to the chain-forming neurons migrating in the RMS, which is a permissive microenvironment for migration surrounded by specialized astrocytes, neurons move individually in the OB tissue, which contains a high density of mature neurons and their processes. Therefore, the neurons may require scaffolds such as blood vessels during radial migration to efficiently reach their destination in the OB (Bovetti et al., 2007). The OB continuously reorganizes its neuronal circuitry in response to changes in the external olfactory environment. Such activity-dependent regulation of the migration and maturation of V-SVZ-derived neurons may contribute to the plasticity of the olfactory neuronal circuitry.

21.3 Regulation of neuronal migration in the injured brain Neurogenesis in the postnatal V-SVZ is affected by pathological conditions such as brain injury and neurodegeneration. After a stroke, neurons generated from NSCs in the V-SVZ migrate toward the lesion, where they differentiate into mature neurons (Arvidsson et al., 2002; Parent et al., 2002; Yamashita et al., 2006) (Fig. 21.4), suggesting that these neurons contribute to endogenous neuronal regeneration and functional recovery. It has been also reported that some of these neurons are generated locally in the injured striatum (Magnusson et al., 2014). Recent studies have revealed commonalities and differences in the mechanisms for neuronal migration under physiological and pathological conditions. Moreover, several strategies using artificial scaffold-mimetic materials and other biomaterials help to promote neuronal migration toward a lesion and thus have potential applications in treating brain injuries, using endogenous neuronal regeneration (Fig. 21.4).

21.3.1 Migratory scaffolds in the injured brain 21.3.1.1 Neighboring cells in the neuronal chain After a stroke, neurons migrating from the V-SVZ toward the lesion form chain-like cell aggregates and adhere to each other by forming AJ-like structures, similar to those in the RMS. In the injured brain, the neuronal chains are often dissociated and then reorganized (Zhang et al., 2009). Since individually migrating neurons show less directional

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FIGURE 21.4 Regulation of neuronal migration in the injured brain. (A) Scaffolds and the directional control of migrating neurons. After brain injury, neurons (red) generated from neural stem cells in the ventricular-subventricular zone (V-SVZ) migrate along radial glial fibers (light blue) and blood vessels (pink) toward the lesion (brown), where they differentiate into mature neurons (yellow). Factors secreted from reactive astrocytes (blue circles), reactive microglia (green circles), and blood vessels (pink circles) attract the migrating neurons. (B) Enhancement of neuronal migration using biomaterials. The persistent and controlled release of growth/trophic factors from biomaterials is a potential strategy for promoting efficient neuronal migration in the injured brain. Providing appropriate scaffolds for migrating neurons (radial glial fiber-mimetic and blood vessel-mimetic materials) in the injured brain is another strategy for promoting neuronal migration.

movement compared with neuronal chains, chain migration is a likely strategy for efficiently supplying large numbers of neurons from the V-SVZ to a lesion after brain injury. These chain-forming neurons in the injured brain express b1integrin, and the chain formation and AJ-like structures are disrupted in migrating neuron-specific b1-integrin-KO mice (Fujioka et al., 2017), suggesting that b1-integrin is involved in the neuronal chain migration in the injured brain. Although PSA-NCAM (Chazal et al., 2000; Cremer et al., 1994; Ono et al., 1994) and N-cadherin (Porlan et al., 2014; Yagita et al., 2009) are critical regulators of chain migration in the RMS, whether they also regulate the neuronal chain migration toward a lesion remains unknown.

21.3.1.2 Astrocytes In the injured brain, astrocytes rapidly change their morphology and gene expression. These reactive astrocytes, which contribute to glial scar formation and inflammation, have both neurotoxic and neuroprotective roles in the injured brain (Sofroniew, 2015). After cortical injury, neurons are observed along astrocytic processes (Saha et al., 2013), suggesting that the astrocytes act as a scaffold for migrating neurons in the injured brain. While astrocytes in the RMS provide a preferential environment for migrating neurons by physically separating them from the brain parenchyma, those in and around a lesion do not form an apparent RMS-like glial tube. Further studies on the detailed functions of these reactive astrocytes will improve our understanding of their role in the neuronal migration involved in regeneration processes.

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21.3.1.3 Blood vessels In the injured brain, neurons form chain-like cell aggregates and attach to both preexisting and newly generated blood vessels via the endfeet of astrocytes enwrapping the vessel (Kojima et al., 2010; Yamashita et al., 2006). Slice culture experiments revealed that the neurons migrating along blood vessels frequently make a U-turn when they reach a branch point of the vessel and jump from one blood vessel to another (Grade et al., 2013; Kojima et al., 2010; Zhang et al., 2009). Together, these observations indicate that blood vessels are an important scaffold for neurons migrating toward a lesion. The neuronal migration along blood vessels in the ischemic striatum is promoted by BDNF, similar to that in the RMS (Grade et al., 2013; Snapyan et al., 2009). Reactive astrocytes present the vascular endothelial cell-derived BDNF, which promotes neuronal migration toward a lesion (Grade et al., 2013). While b1-integrin regulates neuronal chain migration in the RMS (Belvindrah et al., 2007; Emsley and Hagg, 2003), it also regulates the blood vesseleguided neuronal chain migration in the ischemic striatum (Fujioka et al., 2017). Further morphological and molecular characterizations of these blood vessels are needed to better understand their role as a migratory scaffold.

21.3.1.4 Radial glial cells Radial glial cells are evolutionarily conserved embryonic NSCs that extend thin fibers toward the pia from their soma, which is located in the ventricular zone (VZ) (Rakic, 1972). During brain development, neurons generated from radial glial cells migrate along radial glial fibers toward the cortical surface. Radial glial cells persist in the VZ and act as a scaffold for migrating neurons, even in the adult brain, under physiological and pathological conditions in lower vertebrates such as fish (Adolf et al., 2006; Kishimoto et al., 2011b), reptiles (Font et al., 2001; Perez-Canellas and Garcia-Verdugo, 1996; Ramirez-Castillejo et al., 2002), and birds (Alvarez-Buylla and Nottebohm, 1988; Scott et al., 2012). On the other hand, radial glial cells are transformed into multiciliated ependymal cells, parenchymal glial cells, and adult NSCs during the neonatal period in mammals (Fuentealba et al., 2015; Ge et al., 2012; Merkle et al., 2004, 2007; Spassky et al., 2005; Tramontin et al., 2003) and during aging in fish (Ogino et al., 2016). In the rodent brain, unlike the physiological condition, the radial glial cells persist after a neonatal brain injury such as neonatal trauma or hypoxia/ischemia (Jinnou et al., 2018). Neurons generated in the V-SVZ then use N-cadherin for radial glial fibereguided migration toward a cortical lesion. Expressing a dominant-negative form of N-cadherin in radial glial cells inhibits the radial fibereguided migration and maturation of neurons in the injured cerebral cortex (Jinnou et al., 2018). These findings suggest that the mammalian brain has a latent potential for neuronal regeneration in response to brain injury using radial glial fibereguided neuronal migration. Even in the adult mammalian brain, radial glia-like cells appear after ischemia (Zhang et al., 2007) and overexpress a constitutively active form of ErbB2 (Ghashghaei et al., 2007), and thus could act as a migratory scaffold for neurons. Further study on the mechanisms for the injury-induced maintenance of radial glial fibers and the interactions between migrating neurons and fibers is needed to better understand the role of radial glial cells as a migratory scaffold.

21.3.2 Directional control toward a lesion Brain injury induces the death of mature neurons and other cell types including vascular cells, and the activation of immune cells in and around the lesion. While neurons do not migrate toward the cerebral cortex and striatum in the normal adult brain, they migrate toward these brain regions after an injury. This injury-induced neuronal migration toward a lesion is controlled by attractant factors secreted from inflammatory cells, vascular cells, and glial cells in and around the lesion. While vascular endothelial cells secrete stromal cellederived factor-1 (SDF-1) and Angiopoietin-1 (Ang-1) after an ischemic stroke, migrating neurons express their receptors, CXCR4 and Tie-2, respectively. Inhibiting these signals decreases neuronal migration toward a lesion (Imitola et al., 2004; Kojima et al., 2010; Ohab et al., 2006; Thored et al., 2006), suggesting that they act as attractant factors for migrating neurons. Similarly, activated astrocytes and microglia secrete MCP-1 and CXCL13, whereas migrating neurons express their receptors CCR-2 and CXCR-5, respectively. Neuronal migration toward a lesion is disrupted in CCR-2- and CXCR-5-KO mice (Chapman et al., 2015; Yan et al., 2007), suggesting that these molecules also have attractant roles in neuronal migration. Compared with the migration in the RMS, migrating neurons in the injured brain show a less stable direction of movement and a longer resting phase (Grade et al., 2013; Kojima et al., 2010; Zhang et al., 2009). Activated astrocytes, microglia, and macrophages are not uniformly distributed in and around a lesion, resulting in uneven concentration gradients of various attractant factors, which may be one reason for the directional instability and sustained pauses of the migrating neurons.

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21.3.3 Enhancement of neuronal migration as a strategy for endogenous neuronal regeneration Neuronal migration toward a lesion is controlled by combinations of migratory scaffolds and attractive cues. Considering that the neuronal migration in the injured brain is less efficient than that in the RMS, the provision of appropriate scaffolds and attractant factors could be an effective strategy for promoting neuronal migration. The ability to produce a persistent, effective concentration of attractant factors in the injured brain may be key to promoting neuronal migration toward a lesion. However, intravenous injection is not effective for delivering factors that promote neuronal migration to the brain parenchyma, due to the bloodebrain barrier (BBB), and the continuous injection of these factors into the brain parenchyma is technically difficult. In contrast, recent studies suggest that implanted gelatin hydrogels containing growth/trophic factors can continuously release these factors into the brain. For example, injecting GDNF-containing gelatin hydrogels into the V-SVZ promoted neuronal migration (Fon et al., 2014a). Furthermore, in a stroke model, injecting HGF-containing gelatin hydrogels into the ischemic striatum recruited more neurons to migrate toward the lesion (Nakaguchi et al., 2011). These studies indicated that the persistent and controlled release of growth/ trophic factors from biomaterials may be an efficient strategy for promoting neuronal migration in the injured brain. Providing appropriate scaffolds for migrating neurons in the injured brain is another potential strategy for promoting neuronal migration. Given that migrating neurons use their neighboring neurons, astrocytes, radial glial fibers, and blood vessels as endogenous scaffolds for migration, the effects of materials that mimic these scaffolds on neuronal migration in the injured brain have been investigated. Nanofibrous scaffolds conjugated with BDNF (Fon et al., 2014b) and microfiber scaffolds coated with self-assembled colloidal graphene (Zhou et al., 2016) have the potential to facilitate neuronal migration. Transplanted blood vessel mimics, including laminin-conjugated porous sponge (Ajioka et al., 2015) and injectable gels (Fujioka et al., 2017), act as scaffolds to promote neuronal migration into injured cerebral cortex and striatum. Furthermore, transplanting a radial glial fiber-mimic, N-cadherin-Fc-conjugated porous sponge, into injured neonatal cerebral cortex, promotes the migration and maturation of V-SVZ-derived neurons, leading to improved gait behaviors in mice (Jinnou et al., 2018). Although less invasive methods for delivering scaffold biomaterials need to be developed before their clinical application, artificial scaffolds represent a promising strategy for promoting the migration and regeneration of V-SVZ-derived neurons in the injured brain.

21.4 Postnatal neuronal migration in primates Although most of our knowledge about neurogenesis in the V-SVZ has been obtained using rodent brains, neurogenesis in the V-SVZ is observed in various vertebrates, from fish to primates, including humans (Garcia-Verdugo et al., 2002; Paredes et al., 2016b; Sawada and Sawamoto, 2013). The conservation of neurogenesis in the V-SVZ throughout evolution suggests not only that it is an important biological phenomenon, but also that it has potential as a source for endogenous neuronal regeneration after human brain injury. Macaque monkeys (Old World monkeys) show neuronal chain formation within astrocytic tunnel-like structures (GilPerotin et al., 2009; Kornack and Rakic, 2001; Pencea et al., 2001). Although the chains of migrating neurons gradually disperse into small neuronal clusters when they approach the OB in these animals, BrdU-incorporating mature neurons are observed in the OB 3 months after BrdU injection, suggesting that V-SVZ-derived neurons differentiate into mature neurons in the OB (Kornack and Rakic, 2001). Common marmosets, which belong to the New World monkeys, were recently established as a new model animal for studying neuroscience, and they have been used in several studies on neurogenesis in the postnatal V-SVZ (Bunk et al., 2011; Leuner et al., 2007; Sawamoto et al., 2011). The V-SVZ cytoarchitecture in common marmosets is similar to that in the human brain (Sawamoto et al., 2011). Continuous chain-like neuronal structures that migrate from the V-SVZ toward the OB through the RMS are clearly observed in neonatal but not adult primates. Moreover, cultured neurons dissected from the neonatal common marmoset V-SVZ migrate in chains (Sawamoto et al., 2011), similar to those in rodents. Together, these results suggest that neuronal chain migration from the V-SVZ toward the OB is well conserved, even in primates. Immature neurons have been also observed in the adult human OB (Bedard and Parent, 2004; Curtis et al., 2007). However, recent studies indicate that the numbers of proliferating cells and migrating neurons in the human V-SVZ decrease rapidly during the first few months after birth, thereby leading to the low neurogenic potential of the adult VSVZ and low neuronal migration toward the OB (Kam et al., 2009; Morton et al., 2017; Quinones-Hinojosa et al., 2006; Sanai et al., 2004, 2011), which are similar to those in common marmosets. Neurons expressing immature neuronal markers were observed in the poststroke striatum of adult human brains (Jin et al., 2006; Macas et al., 2006; MartiFabregas et al., 2010; Minger et al., 2007). It remains unknown whether these neurons are derived from the V-SVZ, or generated locally, as reported in the normal human brain (Ernst et al., 2014).

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The neonatal human brain appears to have unique migratory pathways, which may be related to the plasticity in prefrontal cortical circuits, in addition to the RMS that is conserved in many animal species. In the neonatal human brain, NSCs reside in the V-SVZ and actively generate neurons, which migrate toward the OB and the prefrontal cortex through the RMS and medial migratory stream (MMS), respectively (Sanai et al., 2004, 2011). A recent study suggests that the periventricular brain regions in human infants harbor a large number of migrating neurons, some of which show chain-like cell aggregates and associate with blood vessels (Paredes et al., 2016a). Furthermore, some of these neurons are oriented toward the cerebral cortex, similar to the observation in neonatal rodents (Inta et al., 2008). Considering the contribution of neurogenesis in the mouse V-SVZ to neuronal regeneration and functional recovery after neonatal brain injury (Jinnou et al., 2018; Yang et al., 2007, 2008), the human V-SVZ could be an effective source for endogenous neuronal regeneration after a perinatal brain injury such as neonatal hypoxia/ischemia.

21.5 Summary Neurons generated in the postnatal V-SVZ form chain-like aggregates and migrate rapidly within a glial tube for a long distance toward the OB. These morphological features are not observed during the embryonic period, suggesting that the chain formation and glial tube have roles specific to the neuronal migration in the postnatal brain. Neuronal migration in the postnatal brain may be affected by changes in the physiological condition of the brain, such as neuronal activity, development, and aging. Furthermore, recent evidence suggests that neuronal migration also depends on nonneural systems such as immune and vascular systems. Further analyses of the spatiotemporal interactions between migrating neurons and other cell types, and identification of the molecules mediating these interactions will be necessary to understand the regulatory mechanisms of neuronal migration in the postnatal brain. Neuronal migration in the postnatal brain appears to be a remarkable strategy for modifying postnatal brain functions and not just a delivery system of newborn neurons from the germinal zone to their destination. Further studies aimed at clarifying the adaptive potential of migrating neurons in the postnatal brain are needed to establish novel strategies for endogenous neuronal regeneration.

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Ramirez-Castillejo, C., Nacher, J., Molowny, A., Ponsoda, X., Lopez-Garcia, C., 2002. PSA-NCAM immunocytochemistry in the cerebral cortex and other telencephalic areas of the lizard Podarcis hispanica: differential expression during medial cortex neuronal regeneration. J. Comp. Neurol. 453, 145e156. Rutishauser, U., 2008. Polysialic acid in the plasticity of the developing and adult vertebrate nervous system. Nat. Rev. Neurosci. 9, 26e35. Saghatelyan, A., de Chevigny, A., Schachner, M., Lledo, P.M., 2004. Tenascin-R mediates activity-dependent recruitment of neuroblasts in the adult mouse forebrain. Nat. Neurosci. 7, 347e356. Saha, B., Peron, S., Murray, K., Jaber, M., Gaillard, A., 2013. Cortical lesion stimulates adult subventricular zone neural progenitor cell proliferation and migration to the site of injury. Stem Cell Res. 11, 965e977. Saha, B., Ypsilanti, A.R., Boutin, C., Cremer, H., Chedotal, A., 2012. Plexin-B2 regulates the proliferation and migration of neuroblasts in the postnatal and adult subventricular zone. J. Neurosci. 32, 16892e16905. Sakamoto, M., Ieki, N., Miyoshi, G., Mochimaru, D., Miyachi, H., Imura, T., Yamaguchi, M., Fishell, G., Mori, K., Kageyama, R., et al., 2014. Continuous postnatal neurogenesis contributes to formation of the olfactory bulb neural circuits and flexible olfactory associative learning. J. Neurosci. 34, 5788e5799. Sakamoto, M., Imayoshi, I., Ohtsuka, T., Yamaguchi, M., Mori, K., Kageyama, R., 2011. Continuous neurogenesis in the adult forebrain is required for innate olfactory responses. Proc. Natl. Acad. Sci. USA 108, 8479e8484.

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Sanai, N., Nguyen, T., Ihrie, R.A., Mirzadeh, Z., Tsai, H.H., Wong, M., Gupta, N., Berger, M.S., Huang, E., García-Verdugo, J.M., et al., 2011. Corridors of migrating neurons in the human brain and their decline during infancy. Nature 478, 382e386. Sanai, N., Tramontin, A.D., Quinones-Hinojosa, A., Barbaro, N.M., Gupta, N., Kunwar, S., Lawton, M.T., McDermott, M.W., Parsa, A.T., ManuelGarcia Verdugo, J., et al., 2004. Unique astrocyte ribbon in adult human brain contains neural stem cells but lacks chain migration. Nature 427, 740e744. Sawada, M., Ohno, N., Kawaguchi, M., Huang, S.H., Hikita, T., Sakurai, Y., Bang Nguyen, H., Quynh Thai, T., Ishido, Y., Yoshida, Y., et al., 2018. PlexinD1 signaling controls morphological changes and migration termination in newborn neurons. EMBO J. 37, e97404. Sawada, M., Sawamoto, K., 2013. Mechanisms of neurogenesis in the normal and injured adult brain. Keio J. Med. 62, 13e28. 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Sonego, M., Oberoi, M., Stoddart, J., Gajendra, S., Hendricusdottir, R., Oozeer, F., Worth, D.C., Hobbs, C., Eickholt, B.J., Gordon-Weeks, P.R., et al., 2015. Drebrin regulates neuroblast migration in the postnatal mammalian brain. PLoS One 10, e0126478. Spassky, N., Merkle, F.T., Flames, N., Tramontin, A.D., Garcia-Verdugo, J.M., Alvarez-Buylla, A., 2005. Adult ependymal cells are postmitotic and are derived from radial glial cells during embryogenesis. J. Neurosci. 25, 10e18. Thored, P., Arvidsson, A., Cacci, E., Ahlenius, H., Kallur, T., Darsalia, V., Ekdahl, C.T., Kokaia, Z., Lindvall, O., 2006. Persistent production of neurons from adult brain stem cells during recovery after stroke. Stem Cells 24, 739e747. Todd, K.L., Baker, K.L., Eastman, M.B., Kolling, F.W., Trausch, A.G., Nelson, C.E., Conover, J.C., 2017. EphA4 regulates neuroblast and astrocyte organization in a neurogenic niche. J. Neurosci. 37, 3331e3341. 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Chapter 22

Transcriptional and posttranscriptional mechanisms of neuronal migration Zeljka Krsnik2, D, Tatiana Popovitchenko1, D and Mladen-Roko Rasin1 1

Department of Neuroscience and Cell Biology, Rutgers University, Robert Wood Johnson Medical School, New Brunswick, NJ, United States;

2

Croatian Institute for Brain Research, School of Medicine, University of Zagreb, Zagreb, Croatia

Chapter outline 22.1. Introduction to neuronal migration 479 22.1.1. Different ways to migrate: “I did it my way” 480 22.2. Transcriptional and posttranscriptional control of neuronal migration 480 22.2.1. Radial migration 480 22.2.1.1. Radial migration: locomotion 480 22.2.1.2. Radial migration: translocation 481 22.2.1.3. Subtypes of neocortical radial glia; outer radial glia and the somal translocation mode of migration 481

22.2.1.4. RNA-binding proteins 22.2.1.5. lncRNAs 22.2.1.6. MicroRNAs 22.2.2. Tangential migration: transcriptional and posttranscriptional control 22.2.2.1. Interplay of transcription factors and tangential migration guidance cues 22.3. Conclusion and future directions List of acronyms and abbreviations References

488 492 493 495 498 503 503 503

22.1 Introduction to neuronal migration Neocortical functions come together in pitch perfect harmony thanks to exquisitely assembled neural circuits. Neocortical neural circuits are composed of diverse cell types. A medley of neuronal diversity allows for impressive neocortical functions such as higher cognition, voluntary motor behavior, and conscious sensory perception (Rakic, 2007; Rakic et al., 2007). Two neuronal populations are supported by glia cells called astrocytes and oligodendrocytes, and a macrophage cell type called the microglia. This is all among a vast vasculature made of epithelial cells/blood vessels. The two main neocortical neuronal subtypes are excitatory glutamatergic projection neurons (also called pyramidal) and inhibitory gamma-aminobutyric acid-containing (GABAergic) interneurons (also called locally projecting neurons) (Jones, 1986; DeFelipe and Farinas, 1992; Rakic, 2007; Kwan et al., 2012a; DeBoer et al., 2013; Evsyukova et al., 2013). Glutamatergic neurons are highly polarized cells that constitute 70%e80% of all neocortical neurons, and their placement defines the six layers of the neocortex (Rakic, 2007; Kwan et al., 2012a; DeBoer et al., 2013; Evsyukova et al., 2013; Custo-Krieg et al., 2013; Popovitchenko and Rasin, 2017). These neurons not only connect the neocortex with other regions of the central nervous system, including the thalamus, brain stem, and spinal cord, but also connect distal neocortical regions and even layers within the same region. GABAergic interneurons are not polarized and constitute the remaining 20%e30% of neocortical neurons. They project locally and, thus, were first recognized by Cajal as neurons with local axons that keep glutamatergic neurons in check. These two main subpopulations have been molecularly and functionally distinguished through the outstanding efforts of many neuroscientists. Remarkably, both glutamatergic and GABAergic neurons have specialized locations within the six-layered neocortex. Despite this stringent positioning, a series of discoveries have confirmed that no neocortical neurons þ

these authors contributed equally.

Cellular Migration and Formation of Axons and Dendrites. https://doi.org/10.1016/B978-0-12-814407-7.00022-5 Copyright © 2020 Elsevier Inc. All rights reserved.

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are born within the neocortex. They are rather born at distant regions and migrate, as development proceeds, to take their final layer-specific positions and integrate into appropriate circuits (Rakic, 1974, 2009; McConnell, 1995a, 1995b; Anderson et al., 1997a, 1997b; Corbin et al., 2001; Tarabykin et al., 2001; Hatten, 2002; Nadarajah and Parnavelas, 2002; Kwan et al., 2012a; Marin and Rubenstein, 2001, 2003; Bielas et al., 2004; Kriegstein and Noctor, 2004; Cooper, 2008; Huang, 2009; Rakic, 2009; Evsyukova et al., 2013; Marin, 2013; Guo and Anton, 2014; Luhmann et al., 2015; Honda et al., 2003). Even more fascinating is that the two main subtypes, inhibitory and excitatory, have distinct origins. Glutamatergic neurons originate from neural stem cells in both the ventricular zone (VZ) and subventricular zone (SVZ), which are found in the dorsal part of the telencephalon called the pallium. GABAergic interneurons originate from neural stem cells found in distinct regions of the ventral telencephalon called the subpallium. Given the complex coordination needed to guide both excitatory and inhibitory neurons to their places in the neocortex, it is not surprising that migration is one of the most vulnerable stages of development in the neocortex. Indeed, disrupted neocortical migration is associated with many neurodevelopmental disorders (Ross and Walsh, 2001), including encephalopathic epilepsy, autism spectrum disorders, and schizophrenia (Sheen and Walsh, 2003; Mcmanus and Golden, 2005; LoTurco and Bai, 2006; Dobyns et al., 1996; Kahler et al., 2008; Guerrini and Parrini, 2010; Wegiel et al., 2010; Verotti et al., 2009; Liu et al., 2011; Evsyukova et al., 2013; Muraki and Tanigaki, 2015; Reiner et al., 2016; Guerrini and Filippi, 2005; Kanatani et al., 2005). It is therefore crucial to understand the mechanisms guiding migration. Interestingly, many of the same molecular mechanisms that are critical for neuronal migration will also determine the growth of the neurites, dendrites, and axons or even regulate proliferation of neural precursors. To cover all the possible roles of each molecule associated with neural and glial migration within the brain development is out of the scope of this chapter. Instead, this chapter highlights the current state of knowledge of molecular and cellular underpinnings of neuronal migration. These molecular processes lead to the fascinating harmony of neocortical structure and function.

22.1.1 Different ways to migrate: “I did it my way” Neurons migrate into the neocortex’s six layers using two main movement strategies: radial and tangential. Radial migration proceeds vertically and superficially from deeply positioned birth sites, whereas tangentially migrating neurons migrate parallel to the surface of the neocortex and perpendicular to the radial migration. This section briefly describes the basic differences between the two strategies. It should be noted that neuronal migration is a mostly conserved process between rodents and primates. For example, in mice, glutamatergic neurons travel through distinct radial migration stages that will be described in the following section. While migrating radially, they need to cross the SVZ, intermediate zone (IZ), and the single-layered subplate (SP) to take their final place in the cortical plate (CP). Travel routes are similar, but more extensive in the brains of primates. There, glutamatergic neurons migrate through the inner SVZ, periventricular fiber reach zone (PVFRZ), outer SVZ (oSVZ), IZ, an evolutionary expanded SP, and then to their final position in the CP. An even more fascinating evolutionary fact is that neuronal migration in mice takes course over about 1e2 weeks, whereas neuronal migration in humans takes place over almost 3 months. Though migration of immature neurons in primates is more complex and advanced than that of their murine counterparts, the molecular mechanisms we will describe in this chapter were largely elucidated in the mouse and other rodents.

22.2 Transcriptional and posttranscriptional control of neuronal migration 22.2.1 Radial migration 22.2.1.1 Radial migration: locomotion As the predominate neuronal population of the neocortex, glutamatergic neurons give the neocortex its six-layered structure. Each layer will have a distinct set of functions determined by the birthdate of its glutamatergic neurons (Angevine and Sidman, 1961; Sidman and Rakic, 1973; Berry and Rogers, 1965; Rakic, 1974; McConnell, 1995a, 1995b). The earliest glutamatergic neurons will be born in the VZ of the dorsal pallium from neural stem cells called radial glia (RG). Described in the 19th century by Giuseppe Magnini, RG became the focus of intense research and were found to be stem cells with curious neuroanatomic features not seen in stem cells outside of the brain. RG are highly polarized and have two main processes: a short one that reaches the apical surface of the dorsal pallium and a longer one that will extend from the VZ to attach at the basal surface membrane (Schmechel and Rakic, 1979). The basal surface is covered by the thinnest of the three meninges called the pia; thus, the long fiber is accurately called either the basal or the pial process.

Molecular mechanisms of neuronal migration Chapter | 22

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The basal process can best be understood as a rope that glutamatergic neurons use to climb up to the CP (a developmental region that will ultimately be compartmentalized into the six neocortical layers). Migration that utilizes the radial process is called radial migration (Rakic, 1972, 1974, 1978; Evsyukova et al., 2013; Edmondson and Hatten, 1987; Mason et al., 1988; O’Rouke et al., 1992; Gadisseux et al., 1990; Misson et al., 1988; Miyata et al., 2001; Noctor et al., 2001, 2004; Hatten and Mason, 1990; Hatten, 2002). Radial migration consists of several steps. The migrating neuron will first extend a leading process, followed by the movement of its nucleus upward (nucleokinesis), and finally the retraction of the trailing process; this process continues to cycle until the neuron reaches its final destination. As we can see, RG have multiple roles: they begin as neural stem cells of glutamatergic neurons and continue, as every good parent does, to guide their progeny to the location of their future maturation.

22.2.1.2 Radial migration: translocation Another method of radial migration in the CP is somal translocation, which was first posited in the 1970s (Morest, 1970; Schmechel and Rakic, 1979; Parnavelas, 2000; Nadarajah et al., 2001, 2003). All migration begins from the division of a RG into itself and a daughter cell or into two daughter cells. In the case of somal translocation, a daughter cell actually inherits the RG basal process that is still attached to the pial surface. The cell body (soma) of this newly born neuron will rapidly ascend (translocate; the cell body almost seems as if it is being pulled up by the process) radially to take its ultimate position in the CP. Thus, somal translocation has been demonstrated to be used by early glutamatergic neurons that will migrate to the lower layers of the neocortex (Miyata et al., 2001; Nadarajah et al., 2001), or by neurons migrating to upper layers at the end stages of neurogenesis after the final neurogenic RG division takes place (Noctor at al. 2004). The fact that somal translocation is also used after the last division of a RG is expected because the basal process is no longer needed by the RG (Berry and Rogers, 1965; Nadarajah et al., 2001; Miyata et al., 2001; Noctor et al., 2004; Miyata and Ogawa, 2007).

22.2.1.3 Subtypes of neocortical radial glia; outer radial glia and the somal translocation mode of migration There is a subtype of RG called the outer radial glia (oRG), named for their placement outside of the VZ/SVZ in the socalled oSVZ. The oSVZ expanded evolutionarily with the acquisition of excessive numbers of oRG. oRG also have a basal process attached to the pial surface, but they lack attachment to the ventricular surface. oRG are proliferative cells that are capable of potentiating the number of neurons produced by the total RG pool. RG and oRG are known to share some markers (e.g., PAX6) but not others (e.g., HOPX is only expressed in oRG) (Liu et al., 2011; Pollen et al., 2015). Remarkably, oRG use somal translocation to take their position in the oSVZ and will continue to produce neurons once there. oRG migration by somal translocation has been observed to increase in length and frequency, as cortical evolution has advanced from developing brains in mice to humans do they also inherit the process? (Ostrem et al., 2014; Betizeau et al., 2013; Gertz et al., 2014). 22.2.1.3.1 Interplay of transcription factors and radial migration guidance cues Time-lapse imaging studies coupled to histological analysis have proven indispensable to understanding migration in the neocortex (Sidman and Rakic, 1973; Rakic et al., 1974; Noctor et al., 2001, 2002, 2004, Tabata and Nakajima, 2003; Kriegstein and Noctor, 2004; LoTurco and Bai, 2006; Rakic, 1974; Anderson et al., 1997; Corbin et al., 2001; Hatten, 2002; Nadarajah and Parnavelas, 2002; Kwan et al., 2012a; Marin and Rubenstein, 2001, 2003; Tabata and Nakajima, 2003; Bielas et al., 2004; Kriegstein and Noctor, 2004; Ayala et al., 2007; Cooper, 2008; Huang, 2009; Merot et al., 2009; Rakic, 2009; Marin et al., 2010; Marin, 2013; Guo and Anton, 2014; Luhmann et al., 2015; Cooper, 2014). There are at least six stages of radial migration: (1) birth of a neuron from an RG in the VZ; (2) bipolar migration to the SVZ; (3) arrival at the SVZ/IZ where migrating neurons become multipolar and have random movements up, down, and sideways; (4) reacquisition of the bipolar stage to migrate via RG basal process to the CP; (5) radial locomotion toward the CP using the RG basal process; and (6) detachment and terminal translocation once the final position within the CP is reached. The first-born cortical neurons use somal translocation to migrate into the preplate (PP) to start establishing the CP. As these first glutamatergic neurons arrive and accumulate within the PP, the PP will be split into the marginal zone (MZ) and the subplate (SP) (Sidman and Rakic, 1973; Rakic, 1972; Bystron et al., 2008; Allendoerfer and Shatz, 1994; Kostovic and Rakic, 1990). An extracellular protein called reelin is secreted by pioneer neurons in the neocortex. These neurons are called CajaleRetzius cells and are found in the MZ. They are among the earliest neurons to arrive into the PP by, surprisingly, tangential migration from the subpalluium (Ramón y Cajal, 1890; Retzius, 1893; D’Arcangelo et al., 1995;

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Ogawa et al., 1995; Soriano and Del Rio, 2005; Frotscher, 1998; Meyer and Goffinet, 1998; Meyer and González-Gómez, 2018; Tissir and Goffinet, 2003). Removing reelin from neocortical development results in a disorganization of neocortical layering (Caviness, 1982-83; D’Arcangello et al., 1995; Ogawa et al., 1995; D’Arcangelo, 2014). It was demonstrated that the RG scaffolds made of basal processes were disrupted in reelin-depleted mice (D’Arcangello et al., 1995; D’Arcangelo and Curran, 1998; Ogawa et al., 1995; D’Arcangelo, 2014). Reelin acts through the receptors ApoER2 and Vldlr, which will phosphorylate the downstream protein disabled-1 (Dab1). Dab1 was established as the downstream effector of reelin signaling in migration. Disruption of Dab1 or the receptors, ApoER2 and Vldr, results in a disorganization of glutamatergic neurons layers. Yet, the disorganization phenotype is strongest when reelin itself is not functioning (D’Arcangelo, 2014). Surprisingly, despite the mislocalization of neurons in reelin-deficient mice, misplaced neurons have normal connectivity and postnatal functionality (Polleux et al., 1998; Guy et al., 2015). For example, even though subcortically projecting neurons were found in upper layers, they still extended their axons to the appropriate subcortical terminal targets. Disrupted functions of cyclin kinase 5 (Cdk5), linked to Dab1, also result in migratory defects similar to that observed in reelindeficient mice (Gilmore et al., 1998). Reelin disruption of neuronal migration has also been found to involve notch. Reeler deletion mice lack the intracellular domain of notch-1 (NICD) in nuclei, even though full-length notch-1 is expressed at normal levels in these mice (Hashimoto-Torii et al., 2008). Elegant in vivo experiments demonstrated that the mutation of notch results in a failure of neurons to migrate properly into the CP. Furthermore, this effect was associated with the Dab1enotch-1 interaction and selective for postmitotic and migratory neurons (Hashimoto-Torii et al., 2008). In addition to extracellular factors, transcription factors are in a strong position to influence many processes because of their potential impact on diverse genes, and indeed, radial migration has been shown to be dependent on their proper expression. Deletions of the class III POU domain transcription factors, Brn-1 (also known as Pu3f3) and Brn-2 (also known as Pou3f2), are required for the initiation of radial migration (Sugitani et al., 2002; McEvilly et al., 2002). Brn-1 and Brn-2 are expressed in upper layers IIeIV at postnatal day 0 (P0). Single Brn-1 knockouts (KOs) do not have severe layering defects but do exhibit disrupted neuronal organization with the CP (McEvilly et al., 2002). However, cortical migration is reversed in the Brn-1//Brn-2/ double knockout (dKO). This was confirmed by finding markers of layers IIeV, TLE3 and TLE1, expressed in deeper regions of the neocortex. Additionally, using an elegant series of birthdate labeling experiments with bromodeoxyuridine (BrdU), authors were able to follow migrating neurons. Neurons in Brn1//Brn-2/ mice interestingly do not express Dab1 and have been linked to the p39/p35 components of Cdk5 (Sugitani et al., 2002; McEvilly et al., 2002). The paired box protein Pax6 transcription factor is widely used to identify RG within the developing neocortex, and it has been suggested that Pax6 deletion affects both radial and tangential migration. If Pax6 is depleted, regional neocortical organization is disrupted, and neurons are stalled in migratory arrest. The stalled cells form heterotopias at the SVZ/VZ border. Interestingly, Pax6 depleted neurons express Map2, but not NeuN. Whereas both Map2 and NeuN are mature neuron markers, NeuN expression is characteristic of neurons that have reached the IZ during normal migration (Talamillo et al., 2003). Acquired expression of NeuN is a fascinating demonstration that localization of cells during migration corresponds to an internal molecular program. In contrast to Pax-6, T-box brain factor 1 (TBR1) is not normally expressed in the VZ and SVZ and as such is widely used as marker of early-born neurons. When deleted, neocortical layer organization is disrupted due to improper neuronal migration (Hevner et al., 2001; Han et al., 2011; McKenna et al., 2011). Here, the SP ends up in the middle of the CP; the effect of this is that upper layers and deep layers are split and are instead above and below the abnormally positioned SP, respectively. The Satb2 transcription factor is expressed in most intracortically projecting neurons and can be found in both upper and lower layers, and as it turns out, Satb2 also plays a role in intracortical migration (Alcamo et al., 2008; Britanova et al., 2008). Intriguingly, Satb2þ neurons destined for lower layers will arrive normally, but ones destined for upper layers will be stuck in the IZ and do not enter the CP. The basic helixeloopehelix (bHLH) transcription factor neurogenin-2 (Ngn-2) plays roles in the acquisition of neuronal features, neuronal subtype specification, and dendritic growth. It also plays a role in radial migration (Schuurmans et al., 2004; Hand et al., 2005; Heng et al., 2008; Nguyen et al., 2006a; Ohtaka-Maruyama et al., 2013). Ngn-2 is transiently expressed in glutamatergic neurons within the SVZ and in the IZ when radial migration is initiated. Ngn-2 deletion results in an accumulation of neurons in both the SVZ and VZ, indicating halted migration (Schuurmans et al., 2004). Additionally, conditional deletion of Ngn-2 through ex vivo electroporation of a Cre-containing construct resulted in cells that failed to migrate out of the VZ (Hand et al., 2005). These experiments demonstrate that Ngn-2 affects radial migration cell autonomously. To elucidate the mechanism of Ngn-2 in radial migration, authors mutated tyrosine 241, in the C-terminus of Ngn-2, to an unphosphorylatable phenylalanine (Y241F). This mutation resulted in impaired polarity and nucleokinesis,

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demonstrating its necessity in radial migration. Surprisingly, Y241F mutation does not interfere with Ngn-2 DNA-binding capability; in other words, this mutation preserves the transcription factor status of Ngn-2 (Hand et al., 2005). Doublecortin (Dcx) is an Ngn-2 target gene and a known proneuronal gene. Dcx promotes radial migration (Bai et al., 2003; Tanaka et al., 2004; LoTurco et al., 2006) and modulates actin and microtubule cytoskeleton dynamics. Dcx is increased not only by the activity of Ngn-2 but also by other bHLH transcription factors expressed in developing neocortices, such as neurogenin-1 and NeuroD (Ge et al., 2006). bHLH transcription factors enact a proneuronal fate by displacing the transcriptional coactivator CBP from the promoter region of the Rho family small-GTPase RhoA, thereby inhibiting RhoA. In line with this mechanism, the migratory defect in Ngn-2 KOs was rescued by a dominant-negative form of RhoA. This demonstrates that RhoA inhibits radial migration unless Ngn-2 enacts disinhibition. It was later demonstrated that Ngn-2 can directly affect expression of the small GTP-binding protein Rnd2 (Heng et al., 2008). Silencing of Rnd2 mimics the radial migration defect found when Ngn-2 is depleted. Amazingly, expressing Rnd2 in Ngn-2-deficient cells corrects radial migration defects (Heng et al., 2008). Together, these studies revealed that Rnd2 is the main effector of Ngn-2-driven radial migration through RhoA inhibition. Further studies reinforced the coordination of Ngn-2 and RhoA in radial migration. The cyclin-dependent kinase inhibitor p27Kip1 stabilizes Ngn-2 protein levels and blocks RhoA signaling. This promotes neural migration through added enforcement of the Ngn2 signaling axis in radial migration (Nguyen et al., 2006b). In addition, inhibition of RhoA by other molecules such as Mst3/Stk25 kinases also regulates migration (Tang et al., 2014). Collectively, there is an abundance of support for Ngn2’s regulatory role in radial migration as well as the need to control the GTPase activity of RhoA. Transcriptional repression is likewise a critical mechanism of radial migration. The zinc finger transcription factor RP58 (also known as ZNF238 or ZFP238), a member of the BTB/POZ-domain family, is an upstream transcriptional repressor of Ngn-2 that can cell-autonomously regulate the multipolar to bipolar transition of glutamatergic neurons during migration (Ohtaka-Maruyama et al., 2013). Deletion of RP58 stops neuronal migration at the SP where migratory cells get stuck in the multipolar stage and fail to go back to the bipolar one. This phenotype is fully rescued by Ngn-2 silencing and partially rescued by Rnd2 silencing. Authors elucidated that RP58 directly binds to both Ngn-2 promoter and an enhancer element in the exon encoding 30 untranslated region (UTR) DNA (Ohtaka-Maruyama et al., 2013). Interestingly, RP58 expression can be induced by Ngn2 itself (Seo et al., 2007), whereas overexpression of Ngn-2 results in the migratory ectopy similar to RP58 deletion (Ohtaka-Maruyama et al., 2013), indicating the utmost importance of the spatiotemporal interplay between transcriptional repression and derepression. Deletion of RP58 resulted in increased Rnd2 mRNA levels; authors demonstrated that this was through direct RP58 binding to specific 30 UTR exon enhancer element E-box and effectively repressing Rnd2 (Heng et al., 2015), suggesting multiple downstream RP58 targets. RP58-depleted neurons had impaired multipolar-to-bipolar transition during their radial migration from the IZ to the CP. The RP58 suppression of Rnd2 is counteracted by bHLH transcription activators such as Ngn-2, NeuroD1, and NeuroD2. The proneuronal bHLH transcription factor Ngn-2 promotes radial migration through Rnd2, and another proneuronal bHLH transcription factor, achaeteescute family bHLH transcription factor 1 (Ascl1), promotes neural migration through the control of second Rho GTPase member, Rnd3 (Pacary et al., 2011). Similar to Rnd2, Rnd3 also inhibits RhoA activity to promote the migration. However, they are localized to partially distinct subcellular compartments, which results in them acting on distinct stages of radial migration. While Rnd2 is important for the multipolar to bipolar transition in the IZ, Rnd3 is a membrane-associated protein in migrating neurons. There, it inhibits RhoA signaling and is important for locomotion specifically in the cortical plate. Interestingly, even though both Rnd2/3 inhibit RhoA, they cannot rescue each other’s phenotypes unless Rnd2 is targeted to the membrane. This elegant set of experiments further supports the finding that Rnd2 and Rnd3 act on distinct stages of neural migration by different subcellular compartmentalization. Collectively, proneuronal bHLH transcription factors Ngn-2 and Ascl1 are in position to be master regulators of successive stages of radial migration. Ascl1 itself is also subject to upstream control. Using an unbiased proteomics approach, another zinc finger transcription factor, Zfp609, was shown to interact with the cohesion loading factor Nipped-B-like (NIPBL). Zfp609 was also found to reside at active promoter and enhancer regions of transcripts that control neuron migration, such as Sema3a, Plxnd1, Gabbr2, and Ascl1 (van den Berg et al., 2017). In particular, Zfp609 and Nipbl associate with the integrator complex that releases RNA Pol-II from a paused state. Silencing of either Zfp609, Nibpl, or integrator subunits Ints1 and Ints11 results in halted migrating neurons in the IZ 3 days after silencing at E17.5. Interestingly, when Zfp609 or Nibpl was silenced and cells analyzed at a later stage (postnatal day 2, P2), a heterotopic accumulation was found in the white matter (van den Berg et al., 2017). Increased levels of repressor element 1 (RE1) silencing transcription factor (REST) lead to premature stoppage of radially migrating cells at the VZ/SVZ border and within the IZ (Mandel et al., 2011). Interestingly, the REST-expressing

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halted cells will eventually differentiate into neurons, suggesting that REST overexpression does not ablate neuronal differentiation, but rather delays it. Migratory arrest was rescued partially by the expression of Dcx, which promoted later stages of migration. These findings further suggested that early stages of migration are DCX independent, whereas later ones require Dcx. It seems that Dcx mRNA is repressed by REST in the VZ progenitors before migration is initiated. Indeed, ablation of REST in adult neural progenitors results in increased DCX levels (Gao et al., 2011). The COUP transcription factor 1, COUP-TFI, was shown to repress Rnd2 at the postmitotic level to regulate radial migration. Repression was limited to later-born, intracortically (or callosally) projecting, glutamatergic neurons (Alfano et al., 2011). Earlier-born, subcortically projecting, neurons are generated and migrate normally in COUP-TFI knockouts but have altered neurite development. Upper-layer neurons are also normally specified but migrate abnormally. The migration defect was rescued by Rnd2 silencing. COUP-TF1 repression of Rnd2 was independent of Ngn-2; DNA binding occurred not at, but rather between, the Ngn-2-binding sites within the Rnd2 enhancer. Interestingly, COUP-TFI repression shows a low to high gradient from rostral to caudal parts of the developing neocortex, following the pattern of radial migration, but contrasting Rnd2 expression levels. Overall, it is clear that tight transcriptional control in the form of repression and derepression allows radial migration to proceed in the correct cell types and at the correct time. The downregulation of a zinc fingerecontaining transcription factor, Kruppel like factor 4 (KLF4), is required for normal migration (Qin and Zhang, 2012). KLF4 is expressed in RG (Qin et al., 2011) and forms a feed-forward loop with the LIF-JAK-STAT3 pathway (Hall et al., 2009; Qin and Zhang, 2012). Overexpression of Klf4 resulted in a majority of cells stuck at the VZ/SVZ border. This migratory defect was rescued by a dominant-negative form of STAT3, suggesting that Klf4-induced overexpression of STAT3 is responsible for the Klf4 migratory pathology. Conversely, silencing of Klf4 resulted in excessive migration to the CP and more cells with a bipolar morphology; this suggests that Klf4 has an important repressive role in the transition from the multipolar to the bipolar stage. However, Klf4 silencing migratory pathologies were transient. It would be interesting to understand if later processes such as dendrite maturation or circuit integration were affected by the delayed migration of Klf4-deficient cells. Once glutamatergic neurons are born, the majority, if not all, will quickly transition from a bipolar shape in the VZ to a multipolar one in the SVZ/IZ. While in the multipolar stage, the somas of some migrating neurons have been observed to become static or even migrate randomly (to the side, basally toward pia, or apically toward the VZ) for some time while multiple processes are extended and retracted. The multipolar stage cells that made their final decision in the VZ to be postmitotic neurons were also named the “slowly exiting population” due to their relatively slower exit from VZ in comparison with the “rapidly exiting population.” This latter “rapidly exiting” population uses somal translocation and is also multipolar, but the cells in this group can undergo divisions basal progenitors (also known as Tbr2þ intermediate progenitors [IPs]; Tabata et al., 2009). These movements are reminiscent of filapodial exploration and continue until one apically oriented process at the site of the centrosome extends as a thin protrusion to become a developing axon. Then, another process becomes thicker and is defined by Golgi and centrosome movements toward the basal surface of the soma. The thicker process becomes the dominant so-called “leading process” of the migrating neuron. Bcl11a is a zinc finger transcription factor that regulates transcription through interaction with COUP-TF and also has a migration-specific phenotype. Deletion of Bcl11a disrupts the switch from the multipolar to bipolar stage of radial migration, and as a result, the arrival of cells to their destined layer is delayed. Interestingly, this effect was limited to laterborn upper-layer neurons. Mechanistically, this is due to increased expression of a member of the semaphorin family of signaling molecules, Sema3c. Silencing of Sema3c in the Bcl11a-deficient neurons results in rescued migratory abnormalities (Wiegreffe et al., 2015). This suggests that Sema3c is downstream of Bcl11a. Silencing of PR domain-containing 8 (Prdm8) protooncogene member of zinc finger C2H2 superfamily containing the PRD1-BF1 and RIZ homology domain (PRDM) also results in delayed multipolar to bipolar transition. In addition, Prdm8 silencing ultimately results in accumulation of migrating cells within the deep layers of developing neocortex. In contrast, when Prdm8 was overexpressed, transfected cells were found in superficial layers. A number of downstream targets were revealed through an unbiased microarray screen (Inoue et al., 2014). Silencing of one of the putative Prdm8 downstream targets, Ebf3 helixeloopehelix transcription factor, mimics the Prdm8 silencing, and multipolar to bipolar transition was disrupted (Iwai et al., 2018). Ebf3 was further shown to regulate expression of Neurod1 and that NeuroD1 is sufficient to rescue migratory phenotype from Ebf3-silenced neurons. A transcription factor that has been associated with intellectual disability is a member of the AFF transcription factors, the lymphoid nuclear protein related to AF4/AFF member 3 (LAF4/AFF3). Similarly to Pgf6, Laf4 has also been implicated in radial migration. Silencing led to more than 80% of transfected cells accumulating below the CP (Moore et al., 2014). When Laf4 was overexpressed, neurons migrated normally into the CP, suggesting that a minimum threshold of Laf4 is necessary and sufficient to neocortical radial migration. A direct downstream transcriptional target of Laf4 is the

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cell adhesion protein MAM domainecontaining glycosylphosphatidylinositol anchor 2 (Mdga2). Importantly, when Mdga2 is overexpressed in migrating neurons that are Laf4 deficient, neocortical migratory defects are rescued. Silencing of filamin A (Flna), platelet-activating factor acetylhydrolase 1b, regulatory subunit 1 (Pafah1b1; also known as lissencephaly 1 (LIS1)), and doublecortin (Dcx) in developing neocortices leads to pathologies in bipolar to multipolar to bipolar ratios. Overexpression of filamin A (Flna) reduced migratory cells in the multipolar stage in the SVZ and the IZ (Nagano et al., 2004). Filip1 can degrade Flna protein. Silencing of Filip1 led to increased Flna expression; in essence, this is a recapitulation of Flna overexpression. Again, there were less migratory neurons in the multipolar stage. However, when Filip1 was overexpressed, resulting in less Flna, the result was reversed: there were more cells in the multipolar stage (Nagano et al., 2002, 2004). In humans, mutations in the FLNA gene result in an accumulation of cells close to the lateral ventricles called periventricular nodular heterotopias. These heterotopias correlate with epilepsy and reading disorders (LoTurco and Bai, 2006; Dobyns et al., 1996; Sheen and Walsh, 2003). While Flna seems to be important for negatively controlling the transition to the multipolar stage, another heterotopiaassociated protein, doublecortin (Dcx), was shown to be required for cells to leave the multipolar stage (Bai et al., 2003). Importantly, manipulations of these two genes will affect cells at different distances from the VZ. While Flna will have an effect on migrating neurons closer to the VZ, Dcx will have an effect on cells further away in the IZ. This is in line with expression patterns: Dcx expression follows FlnA expression as neurons migrate away from the VZ. These findings bisected the multipolar stage into substages (LoTurco and Bai, 2006): an initial stage, affected by Flna, when glutamatergic neurons migrate between the VZ and SVZ, and a second one, affected by Dcx, when neurons migrate away from the SVZ and move to the IZ. Experiments with Flna and Dcx demonstrate that correctly timed transitions between multipolar and bipolar morphologies, in the correct compartments, during radial migration are necessary for the progression of radial migration. Additionally, a surface glycoprotein member, ephrin/A5 protein/receptor protein tyrosine phosphatase Mu domain glycosylphosphatidylinositol anchor, Mdga1 (Takeuchi and O’Leary, 2006), F-actin depolymerizing factor, N-cofilin (Bellenchi et al., 2007), and the a3b1 and a5b1 integrins (Dulabon et al., 2000; Marchetti et al., 2010) were all found to be important in the transition to bipolarity after a multipolar state in the SVZ and the IZ. Overall, understanding of the multipolar stage of migration is of outmost importance. For example, it is easy to envision that if the multipolar stage is significantly prolonged, prolonged random movements would occur before radial migration, resulting in misplaced neurons. Amazingly, the functional importance of morphological transition during migration was recently reinforced by observing transient neuronal communication. Authors found that the final transition between multipolar and bipolar shapes occurred at the SP among some of the earliest neurons in the cortex (Kostovic et al., 2015). These early-born neurons formed NMDAR-based transient synapses. After instruction from these early SP neurons, the multipolar to bipolar transition was achieved, and radial migration could proceed (Ohtaka-Maruyama et al., 2018). It is tantalizing that a specialized activity such as synaptic communication could guide neurons in a process that many cells undergo. The platelet-activating factor acetylhydrolase 1b, regulatory subunit 1 (Pafah1b1; also known as lissencephaly 1, Lis1), is also required for cells to leave the multipolar stage (Tsai et al., 2005; Hatten, 2005). Failure to undergo this transition again results in heterotopias in humans. In addition to heterotopias, lissencephaly (smooth brain) and pachygyria (few gyri) are also human disease manifestations of neuronal migration defects; mutations in either Lis1 and Dcx can cause lissencephaly or pachygyria. Human genetic studies have revealed other genes important for neuronal migration disorders, including the tubulin subunits TUBA1A and TUBB3. Mutation of these led to heterotopias (Liu, 2011; Manzini and Walsh, 2011). The loss of F-actin depolymerizing factor n-cofilin resulted in aberrant radial migration (Belenchi et al., 2007). Collectively, these findings identify microtubules, centrosome, and actin polymerization/depolymerization as cellular locations of interest in neuronal radial migration. Once bipolarity is assumed through structural reorganization of the Golgi body and the centrosomes, the neuron starts to migrate along the RG basal process: first by moving the leading process, then by nuclear movement toward the leading process, and finally with the axon trailing behind. It is believed that radially migrating neurons hold tight to the RG process by either or both integrins and gap junctions (Elias et al., 2007). Indeed, gap junction proteins connexin-26 and connexin43 are required for radial migration of neurons (Elias et al., 2007). In addition to attachment, the intrinsic centrosome movement dictates proper migration (Umeshima et al., 2007). Overall, cytoskeletal coordination between microtubules, myosin, and cellular adhesion proteins is critical during migration (Schaar and McConnel, 2005). Mutations in genes encoding proteins involved in cytoskeletal organization often lead to migratory failures (Fig. 22.1). Neurons migrate from the proliferative zones to the CP through the VZ/SVZ, the IZ, and SP. The X-linked intellectual disability protein PHD finger protein 6 (PHF6) has two zinc finger PHD-type domains and localizes to the nucleus. Silencing of Phf6 leads to an excessive number of transfected cells in the IZ. Interestingly, fewer cells reached the upper

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FIGURE 22.1 Transcriptional and posttranscriptional control of neuronal radial migration. (A) Newly born glutamatergic neurons (NNs) from radial glia (RG) in the ventricular zone (VZ) migrate into subventricular zone (SVZ). Within the SVZ, NNs become multipolar neurons (MNs) and undergo so-called multipolar migration. This set of events is followed by the acquisition of bipolar shape, and thereafter the neuron starts to use radial locomotion (RLN) to migrate through the intermediate zone (IZ) to reach the cortical plate (CP). Once the destined layer within the CP is reached, the neuron detaches and becomes a postmigratory neuron (PMN). (B) Transcriptional and posttranscriptional pathways within migrating neurons are regulated by transcription factors, RNA-binding proteins (RBPs), microRNAs (miRNAs), and long noncoding RNAs (lncRNAs). (C) List of known transcriptional and posttranscriptional regulators of radial migration. MZ, marginal zone; SP, subplate; oSVZ, outer SVZ.

parts of the CP in the Phf6-depleted cells, whereas the lower CP and VZ had comparable numbers to the control (Zhang et al., 2013). The extent of migratory pathology corresponded to the level of the knockdown induced by each of the three shRNAs. To dissect the mechanism through which PHF6 acts upon radial migration, coauthors identified that all four members of the PAF1 transcriptional elongation process (Paf1, Leo1, Ccd37, and Ctr9) are associated with PHF6. Silencing of Paf1 in developing neocortices phenocopied Phf6 silencing. Furthermore, neuroglycan C/chondroitin sulfate proteoglycan 5 (NGC/CSPG5) was identified as one of the downstream transcriptional targets of Phf6. NGC/CSPG5 is a human schizophrenia susceptibility gene, and knockdown in mice impaired radial migration in a similar way to Phf6 and Paf1 silencing, suggesting that all three operate in the same pathway or in redundant ones. Expressing NGC/CSPG5 in Phf6-silenced migrating cells rescued the migratory pathology. All these phenotypes were observed in E19 brain, 5 days after transfecting developing neocortices. Remarkably, when transfected cells with silenced Phf6 were analyzed at P6, authors found them within heterotopias in white matter. Cells expressed Cux1, a marker of upper-layer neurons, and were hyperexcitable, suggesting that these were mature, mislocalized neurons. Collectively, these results suggest that the intellectual disability gene Pgf6 regulates neocortical migration through transcriptional control of the schizophrenia risk gene NGC/CSPG5. There is accumulating evidence that members of the forkhead box containing (FOXP) transcription factors also contribute to neocortical radial migration. One of these, Foxp1, contains an autism-related mutation that hinders radial migration (Li et al., 2018). Another member of the FOX transcription factors, Foxp2, will delay radial migration in vivo when overexpressed due to accumulation of neurons in the IZ (Clovis et al., 2012). Proper spatiotemporal protein levels of both Foxp1 and Foxp2 in developing neocortices seem to be under the control of the posttranscriptional regulators described in detail later. Similarly to Foxp2, FoxO6 deficiency in developing neocortices results in an accumulation of neurons in the IZ and fewer upper-layer Cux1þ neurons that reach their final destination (Paap et al., 2016). It was also demonstrated that FoxO6 positively regulates the transcription of Plxna4 through direct binding of specific Daf-16-binding elements within the Plxna4 promoter. Accordingly, when Plxna4 is expressed in FoxO6 knockouts, migratory arrest is reversed (Paap et al., 2016). Neurons will continue their locomotion until the final position in the CP is reached where they can terminally translocate and start differentiating. This critical point is when the transition to a mature neuron can proceed; recently, the

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transcription factor sex-determining region Y-box 11 (Sox11) was identified as necessary for this transition. Authors originally set up a screen for regulators of dendritic development and found that when Sox11 was mutated, the result was an intracortical migratory defect (Hoshiba et al., 2016). When Sox11 was silenced, late-born neurons migrated through the SVZ and IZ, reached the CP, but accumulated in layer 5 instead of their destined places in layers IIeIII. This was attributed to excessively complex dendrites, suggesting that dendrite maturation must be suppressed by Sox11 during migration. Indeed, Sox11 expression is high in neurons during radial migration, but decreases as radial migration ends and neurons start to develop dendrites. Once neurons reach the CP, how do they know to let go of the RG basal process to position themselves in the sixlayered neocortex? Even though this question is relatively unexplored, experiments have revealed the contribution of several pathways. For glutamatergic neurons to terminate their migration, they must stop and detach from the RG fibers that they are so faithfully climbing. Reelin is considered to be one such “stop” signal and indeed was shown to regulate detachment through Dab1 (Dulabon et al., 2000; Olson et al., 2006; Sanada et al., 2004). Recent studies have also uncovered that the antiadhesion matrixesecreted protein SPARC-like 1 (SPARCL1; also known as Sc1 and RG antigen Rgrs1) is involved in RG detachment. Sparcl1 is localized to the upper parts of the RG basal process, within the CP, where neurons need to terminate their migration, and incidentally where Sparcl1 inhibits neuroneRG attachment (Gongidi et al., 2004). Another receptor, PlxnD1, interacts with the extracellular factor Sema3E and was shown to be involved in migration termination in the olfactory bulb (Sawada et al., 2018), though it is also likely expressed in pyramidal neurons and blood vessels (GENSAT). Intrinsic changes in actin have also been shown to impact neuronal migration termination. In particular, the Nckassociated protein (Nap1) is an adaptor protein thought to modulate actin nucleation and also plays a critical role in the switch from neuronal migration to differentiation (Yokota et al., 2007). Briefly, Nap1 overexpression leads to premature migratory termination and promotes neurite differentiation, whereas Nap1 silencing results in disrupted postmigratory differentiation. Weakening of adhesion molecules like N-cadherin by the introduction of dominant-negative constructs also results in premature termination of radial migration (Wakimoto et al., 2015). N-cadherin was reported to have critical role in the multipolar to bipolar transition during neuronal migration (Jossin and Cooper, 2011). Interestingly, the Reelin receptor apolipoprotein E receptor 2 (ApoER2) is required not only for the early stage of radial migration, but also for the termination of neuronal migration in the developing neocortex (Hirota et al., 2018). Lastly, reelin itself seems to downregulate adhesive molecules such as integrins to allow neuronal detachment and stop migration (Dulabon et al., 2000; Sanada et al., 2004). Collectively, the understudied, but exciting, area of migration termination research has begun to be unraveled. Future studies will certainly further reveal the role of transcription factors in this step. The first neurons that will populate the CP become members of the deepest layer VI of the neocortex. The next wave of neurons will use radial locomotion and undergo the same stages of migration to surpass the layer VI neurons and position themselves above to form layer V. Layers V and VI are characterized by glutamatergic neurons that project subcortically to connect the neocortex with the thalamus, brain stem, and spinal cord. The waves of generation and migration continue to form upper layers II-IV that contain glutamatergic neurons which project intracortically to connect different neocortical regions and layers. Interestingly, some late-born intracortically projecting neurons are also located within the deep layers. Curiously, even though early-born cells and late-born cells undergo the same phases of migration, the transcriptional control mechanism active in these cells is different. Take, for example, the case of the maternally imprinted transcription factor PLAG1-like zinc finger 1 (Zac1). When Zac1 was overexpressed in progenitors that give rise to lower-layer, subcortically projecting neurons, cells migrated normally. In contrast, when Zac1 was upregulated in progenitors of lateborn, intracortically projecting neurons, cells stalled in the IZ (Adnani et al., 2015). Zac1 overexpression also delayed progenitor cell, maturation to neurons within the IZ. Neuronal migration was also disrupted in Zac1 KO mice. Cux1þ (also known as Cdp) upper-layer neurons were exclusively affected; in the KO, they stayed in the VZ/SVZ by E18.5 when they would normally be in the upper layers of the CP. Migration of subcortically projecting neurons expressing Ctip2 seemed unaffected. The migration phenotype was shown to be at least partially dependent on Zac1 transcriptional control. When its downstream target, Pac1 (the receptor of the neuropeptide PACAP), was overexpressed, cell stalling in the IZ was again observed. Meanwhile, Pac1 silencing in Zac1-overexpressed cells provided a partial migratory rescue. These findings suggest that both Zac1 and Pac1 are important in radial migration. However, the partial migratory rescue suggests that Zac1 has other, unexplored, functions that contribute to its stalled migration phenotype. Remarkably, when a Sox transcription factor is deleted, Sox5, migration of intracortically projecting neurons is again disrupted. In this case, upper layers migrate normally, but deep layers are reversed, with layer VI thalamocortical glutamatergic neurons positioned above layer V corticospinal neurons. Authors determined that this stems from disrupted CP formation as in Sox5 mutants and the PP is not split into the MZ and SP (Kwan et al., 2008; Lai et al., 2007). Sox5 is

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expressed only in postmitotic neurons of the SP and layer VI, whereas its expression is undetectable in VZ, SVZ, and IZ. These studies presented the first example of intracortical-specific layer inversion. There is some evidence that migration of glutamatergic neurons and of interneurons is regulated through overlapping pathways. Aristaless-related homeobox (ARX) results in X-linked lissencephaly in humans. In mouse models, deletion of ARX led to disrupted radial migration and also affected the migration of interneurons (Liu et al., 2011; Friocourt et al., 2008; Marsh et al., 2009; Marcorelles et al., 2010). Arx silencing prevented neurons that started migration to become multipolar in the SVZ and IZ; while overexpression of Arx causes migrating cells to have tangentially oriented processes. Arx manipulation in either form disrupted migration of pyramidal neurons (Friocourt et al., 2008). Furthermore, deletion of a member of the MIG-10/RIAM/Ldp protein family member lamellipodin (Lpd) induced a tangential migratory mode in radially destined glutamatergic neurons (Pinheiro et al., 2011). Lpd deletion reduced the activity of the serum response factor (SRF) transcription factor. Decreasing the SRF activity in vivo results in tangential migration of glutamatergic neurons, a phenotype mimicking the Lpd deletion. In addition, Lpd deletion phenotype was rescued by restoring the SRF activity (Pinheiro et al., 2011). Interestingly, a portion of glutamatergic neurons will indeed travel tangentially from the lateral cortex to reach the subiculum, a transitional cortical area situated in front of the allocortex (“old cortex”), the hippocampus (Britanova et al., 2006). These studies demonstrate that there may be an overlap in both process and control of neuronal migration between pyramidal neurons and interneurons (Table 22.1). 22.2.1.3.2 Posttranscriptional events in radial migration: the role of RNA-binding proteins, microRNA, and long noncoding RNA The role of posttranscriptional regulation has been an active focus of research over the past decade. During this time, many, if not all, neuronal processes have been shown to be regulated at the level of the transcript, and migration is no exception (Evsyukove et al., 2013; Popovitchenko and Rasin, 2017; Lennox et al., 2018; Rajman and Schratt, 2017). There are many kinds of molecules that act posttranscriptionally and include proteins such as RNA-binding proteins (RBPs) and nucleic acids such as microRNA (miRNA) and long noncoding RNA (lncRNA). Here, we will summarize recent studies to demonstrate how these three groups of posttranscriptional regulators are crucial to migration, and we will reveal how much work remains to elucidate the full contribution of this level of control in migration.

22.2.1.4 RNA-binding proteins RBPs can act on every step of posttranscriptional processing: splicing, localization, stability, translation, and decay (Keene, 2007). These proteins contain RNA-binding domains, such as the RNA recognition motif (RRM) or KH domain, that allow them to bind mRNA transcripts. Extensive studies of the fragile X mental retardation protein (FMRP) RBP have shown it to be associated with attention-deficit disorder and autism spectrum disorder (Bagni et al., 2012; Kwan et al., 2012b; Darnell et al., 2011). FMRP also has a critical role in radial migration (La Fata et al., 2014). More specifically, FMRP, through its directly bound mRNA target N-cadherin (also known as cadherin-2, Cad2), regulates the multipolar to bipolar transition during radial migration. In particular, migrating cells were observed to accumulate in the SVZ and the IZ of the developing neocortex in Fmr1 knockout mice (Fmr1 encodes FMRP). Migrating cells that lack Fmr1 have a prolonged multipolar stage and have delayed formation of the leading process. N-cadherin is important for the multipolar to bipolar transition (Jossin and Cooper, 2011). It was shown that FMRP directly binds N-cadherin mRNA; correspondingly, in Fmr1 knockout mice, both protein and steady-state mRNA levels of N-cadherin were decreased. Follow-up experiments suggested that FMRP prevents N-cadherin decay. The migration pathology observed in Fmr1 knockouts was rescued by reintroducing N-cadherin. Another RBP, a member of embryonic lethal abnormal vision-like (ELAVL) family, Hu antigen R was shown to regulate Foxp1 and Foxp2 translation in the developing neocortex. This was also shown to be dependent on HuR’s phosphorylation state (Popovitchenko et al., 2016). Both of these Fox transcription factors are known to affect neocortical migration, and indeed, there was a neuronal placement pathology in HuR conditional knockouts (Kraushar et al., 2014). Translational reinitiation has also been implicated in neuronal migration through the density-regulated protein (DENR, Haas et al., 2016). DENR mutations have been associated with autism spectrum disorder (p.C37Y) and Asperger syndrome with comorbid epilepsy (p.P121L); these mutations notably lay in domains important for chromatin remodeling and transcription (SWIB/MDM2 domain) or adjacent to a domain enabling translation (eIF1-like/SUI1 domain). DENR is involved in specific stages of translation: translation initiation, translation reinitiation, and ribosome recycling. When Denr is silenced or overexpressed (overexpression with either the mouse or human variant) in developing neocortices, fewer transfected neurons reach the CP. Additionally, there was increased retention of cells across the VZ and IZ, but not a significant amount. Interestingly, silencing a partner of DENR, malignant T cell amplified sequence 1 (Mcts1), in

TABLE 22.1 Radial migration: transcriptional regulators. Radial migrationdtranscriptional regulator (synonyms)

Transcription factor type/class

Key downstream gene In migration (confirmed or *proposed)

Arx (Arx1)

Homeobox domain

Not reported

Friocourt et al. (2008), Marsh et al. (2009)

Ascl1 (Mash1, ASH1, bHLHa46)

Basic helixeloopehelix

Rnd3

Pacary et al. (2011)

Bcl11a (2810047E18Rik, COUP-TF interacting protein 1, CTIP1, D930021L15Rik, Evi9, Evi9a, Evi9b, Evi9c, mKIAA1809, mouse myeloid leukemia gene)

Zinc finger

Sema3c

Wiegreffe et al. (2015)

Brn-1 (Brn1, Pou3f3, HST011, Otf8, urehr2)

Class III POU domain

*Dab1, p39/p35 components of Cdk5

McEvilly et al. (2002), Sugitani et al. (2002)

Brn-2 (Brn2, Pou3f2, 9430075J19Rik, A230098E07Rik, Otf7)

Class III POU domain

*Dab1, p39/p35 components of Cdk5

McEvilly et al. (2002), Sugitani et al. (2002)

COUP-TFI (Nr2f1, COUP-TFI, Erbal3, Tcfcoup1)

Chicken ovalbumin upstream promoter transcription factor

Rnd2

Alfano et al. (2011)

Ebf3 (3110018A08Rik, O/E2, Olf-1/Ebf-like 2)

Helix-loop-helix

NeuroD1

Iwai et al. (2018)

FoxO6

Fork head domain

Plxna4

Paap et al. (2016)

References

Fork head domain

Not reported

Li et al. (2018)

Fork head domain

Not reported

Clovis et al. (2012)

Klf4 (EZF, Gklf, Zie)

Zinc finger protein

Stat3

Qin and Zhang (2012)

Laf4/Aff3 (Alf4, Laf4l)

AFF (AF4/FMR2) family of putative transcription factors

Mdga2

Moore et al. (2014)

Neurogenin-2 (Ngn-2, Neurog2, Atoh4, bHLHa8, Math4A, Ngn2)

Basic helixeloopehelix

Dcx, RhoA, Rnd2, RP58

Tanaka et al. (2004), Ge et al. (2006), LoTurco et al. (2006), Nguyen et al. (2006a,b), Seo et al. (2007), Heng et al. (2008), Hand et al. (2005)

Dab1

Hashimoto-Torii et al. (2008)

Pax6 (1500038E17Rik, AEY11, Dey, Dickie’s small eye, Gsfaey11, Pax-6)

Homeobox domain

Not reported

Talamillo et al. (2003)

Phf6; interacts with Paf1

Zinc finger, PHD type

Proteoglycan 5 (Ngc/Cspg5)

Zhang et al. (2013)

Notch1, intracellular domain

Continued

Molecular mechanisms of neuronal migration Chapter | 22

Foxp1 (3110052D19Rik, 4932443N09Rik) Foxp2 (2810043D05Rik, D0Kist7)

489

490 PART | II Migration

TABLE 22.1 Radial migration: transcriptional regulators.dcont’d Radial migrationdtranscriptional regulator (synonyms)

Transcription factor type/class

Key downstream gene In migration (confirmed or *proposed)

Prdm8

References

Zinc finger C2H2 superfamily

Many putative targets assessed through microarrays

Inoue et al. (2014), Inoue et al. (2015)

REST (2610008J04Rik, NRSF)

Repressor element

Dcx

Mandel et al. (2011), Gao et al. (2011)

RP58 (Zbtb18, Znf238, Sfp238)

BTB/POZ domain family

Neurogenin-2, Rnd2

Ohtaka-Maruyama et al. (2013), Heng et al. (2015)

Satb2 (mKIAA1034)

Special AT-rich sequence binding protein

Not reported

Alcamo et al. (2008), Britanova et al. (2008)

Sox5 (A730017D01Rik)

Sex-determining region Y-box

Not reported

Kwan et al. (2008), Lai et al. (2007)

Sox11 (1110038H03Rik, 6230403H02Rik, end1)

Sex-determining region Y-box

Not reported

Hoshiba et al. (2016)

SRF

MADS-box

Not reported

Pinheiro et al. (2011)

Tbr1

T-box

Not reported

Hevner et al. (2001), Han et al. (2011) McKenna et al. (2011)

Zac1 (Plagl1, LOT1)

Pleiomorphic adenoma gene zinc finger protein

Pac1

Adnani et al. (2015)

Zfp609; interacts with Nibpl

Zinc finger protein

Ascl1, Sema3a, Plxnd1, Gabbr2

van den Berg et al. (2017)

Molecular mechanisms of neuronal migration Chapter | 22

491

developing neocortices did not alter radial migration, suggesting that remaining amounts of MCTS1 are sufficient for normal migration. However, introducing Mtcs1 shRNAs into the Denr overexpression experiment rescued migration defects. Overall, these findings suggest that the cooperation of both factors is important for neocortical neuronal migration. Interestingly, while the disease-associated mutations, p.C37Y and p.P121L, disrupted interaction with Mcts1, their individual overexpression results in migratory defects similar to the Denr wild-type overexpression. Several RBPs regulate radial migration through alternative splicing. Nova2 is KH-type RBP shown to regulate intracortical migration of late-born upper-layer neocortical and Purkinje cerebellar neurons (Yano et al., 2010). BrdU tracing at E14 and E16, along with IHC experiments in Nova2 KO, revealed in a portion of upper-layer neurons positive for Brn-2 (layer II/III) and Cux1 (layer II/IV) were mislocalized under lower-layer neurons positive for ER81 (layer V) and Foxp2 (layer VI) by postnatal day 10. BrdU tracing at E12 confirmed that cells at this stage migrated properly into deep layers by P0. An unbiased screening approach using Nova2 HITS-CLIP to pull down all RNAs associated with Nova2 revealed a cluster of mRNAs bound by Nova2 that are associated with reelin and other migratory pathways. Further analysis revealed that Nova2 regulates migration through the precisely timed inclusion/exclusion of alternative Dab1 exons 7b and 7c. Specifically, it seems that Nova2 blocks the inclusion of Dab1 exons 7b and 7c specifically during midneocortical neurogenesis (E14.5eE16.5). An elegant set of experiments further confirmed that the presence of these exons hinders neuronal migration in a dominant-negative fashion. Dab1 is prone to alternative splicing through many pathways, as demonstrated by a recent study examining the RBPs RNA-binding motif 4 (RBM4) (Dhananjaya et al., 2018). Rbm4 deficiency resulted in migratory defects of upper-layer neurons. Mechanistically, Rbm4 directly promotes the inclusion of Dab1 exons 7 and 8; splicing is antagonized by another RBP, the polypyrimidine tract-binding protein 1 (PTBP1). Rbm4a knockout mice had an increased presence of Satb2þ neurons in lower layers and the IZ, whereas Foxp2þ neurons were correctly placed within lower layers. In utero electroporation of Rbm4a knockouts and Rbm4 shRNAs further confirmed that the migratory defect was specific to lateborn upper-layer neurons. Full-length Dab1, but not Dab1 with deleted exons 7 and 8, rescued deficits in radial migration caused by silencing of Rbm4. Furthermore, alternative splicing of Dab1 regulates the multipolar to bipolar transition of migrating neurons (Zhang et al., 2017). In particular, Dab1 isoforms lacking exons 7 and 8 or retaining exons 9bc halt the multipolar stage and result in aberrant leading process extension, which was rescued by introducing full-length Dab1. Collectively, these data present strong evidence that an RBP can efficiently regulate the sensitivity of a migrating neuron to reelin signaling. Nova was also shown to regulate migration of spinal cord interneurons by regulating Dcc splicing (Leggere et al., 2016), solidifying its role in migration overall. The zinc finger protein unkempt is known to bind DNA, but a CCCH domain also allows for RNA binding (Hall TM, 2005). Silencing of unkempt results in an accumulation of transfected cells in the IZ. Remarkably, the migration defect was rescued by wild-type unkempt, but not by a variant lacking the RNA-binding CCCH domain (Murn et al., 2015). Specifically, translation repression of unkempt-bound mRNA targets was identified to be responsible for appropriate migration. Another example of translational repression in migration was discovered through study of the eukaryotic initiation factor 4E1 (eIF4E1). eIF4E1 silencing in developing neocortices induces migratory halt of Satb2þ neurons in the IZ (Yang et al., 2014). Silencing of an eIF4E-associated protein, eIF4G, rescued the migratory pathology. Meanwhile, silencing of another eIF4E1 partner, 4E-T, resulted in many transfected cells continuing their journey by moving into the CP. This phenotype was rescued with wild-type 4E-T, but not a 4E-T variant missing the eIF4E-binding domain. The eIF4E-4E-T translational complex suppresses translation of bHLH transcription factors that were shown to control radial migration, like neurogenin-2. Collectively, spatiotemporal translational repression and derepression has come into focus as a key mechanism in neocortical development (Kraushar et al., 2014, 2015; Yang et al., 2014; Popovitchenko et al., 2016; Popovitchenko and Rasin, 2017; Chau et al., 2018; Zahr et al., 2018; Lennox et al., 2018). Silencing of the RBP Staufen2, partner of translational repressor RBP Pumilio-2 (Vessey et al., 2008; Zahr et al., 2018) and expressed in RG, results in accumulated Satb2þ cells in the IZ and correspondingly fewer cells in both the CP and VZ. The Klf4 zinc finger transcription factor, which regulates radial migration (Qin and Zhang, 2012), interacts with another member of the Staufen family, Staufen1. Klf4 interaction with Staufen1 promotes mRNA decay to regulate cortical neurogenesis (Moon et al., 2018). Cytoskeletal genes are regulated in radial migration at the transcriptional level, and they are likewise subject to regulation at the posttranscriptional level. A recent screen compared neural progenitors and neurons in mice and humans. There, authors found that cytoskeleton genes were differentially spliced between the two cell types and in both species (Zhang et al., 2016). The alternatively spliced cytoskeleton genes have enriched association with proliferation, neuronal differentiation, and neuronal migration. For example, when the neuronal splice variant of ninein is misexpressed in progenitors, the result is migratory arrest in the SVZ and IZ. In another case, abnormal splicing and inclusion of an extra exon of Flna in progenitors is associated with periventricular nodular heterotopia in humans. Several RBPs were identified

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as regulators of the splicing balance in the developing cortex. In particular, the interplay of a heterogeneous nuclear ribonucleoprotein (hnRNP), polypyrimidine tractebinding protein (Ptbp1), and RNA-binding Fox proteins (Rbfox1/2/3) were shown to act oppositely on neuronal-specific exons between progenitors and neurons. Ptbp1 suppresses exon inclusion in progenitors, whereas Rbfox promotes exon inclusion in neurons. The Ptbp1 knockout mouse exhibits periventricular nodular heterotopias. Similarly, silencing with shRNA resulted in defective radial migration where more transfected cells were found in the SVZ and IZ. In contrast, overexpression of Rbfox proteins resulted in a more transfected cells found in the SVZ and IZ. Migratory defects were amplified when Ptbp1 silencing was combined with ectopic expression of Rbfox. The opposing effects of Ptbp1 and Rbfoxs dictate neocortical neurogenesis and migration through their alternative splicing activity. Interestingly, silencing of Rbfox2 or Rbfox3 in developing neocortices did not disrupt radial migration (Hamada et al., 2016). However, silencing of a specific isoform of Rbfox1, isoform 1 (Rbfox1iso1), leads to migratory defects that result in an accumulation of cells in the lower part of the CP and IZ (Hamada et al., 2015, 2016). This was found to be the result of disrupted nucleokinesis of migrating cells. Rbfox1iso1 silencing did not affect the multipolar to bipolar transition but hindered radial migration and terminal translocation from radial processes within the CP. Rbfox1iso1-deficient neurons were not able to enter the uppermost part of the CP even though the leading process reached the MZ. As in Rbfox1iso1 silencing, silencing of Rbfox1iso2 resulted in disrupted terminal translocation (Hamada et al., 2015). Neither isoforms effected RG proliferation, but both effected nucleokinesis. Mutations in Rbfox1 have been associated with neurodevelopmental disorders, including autism spectrum disorders, intellectual disability with epilepsy, attention-deficit hyperactivity disorders, and schizophrenia (Hamada et al., 2015, 2016). Given the frequency of alternative splicing events and their roles in fate-determining processes, these results demonstrate the necessity of careful examination of protein function at the isoform-specific level. Alternative splicing is also important for the last stages of radial migration. Neocortical deletion of a splicing regulator, the neural-specific Ser/Arg repeaterelated protein of 100 kDA (nSR100/SRRM4), resulted in a lamination phenotype where more lower-layer Tbr1þ neurons and fewer upper-layer Satb2þ neurons were observed in the CP (Quesnel-Valieres et al., 2015). Just as RBPs themselves can be subject to posttranscriptional regulation, they too can regulate other regulators. Take, for example, the case of the RBP, PIWIL1, that binds piwi RNAs (piRNAs), a subset of small RNAs (DeBoer et al., 2013; Zhao et al., 2015; Viljetic et al., 2017). It was revealed that PIWIL1 has a role in radial migration by controlling the multipolar to bipolar transition. The PIWI/Argonaute/Zwille and PIWI domains are required for proper radial migration. This was further found to be due to regulation of microtubule-associated proteins (Zhao et al., 2015). Adenomatous polyposis coli (APC) also deserves mention in this section of the chapter. APC’s role in neuronal migration (but also in RG polarity, neuronal generation, and axonal tract development) (Yokota et al., 2009) was shown before it was revealed that APC can act as an RBP (Preitner et al., 2014). Nevertheless, loss of APC in the developing neocortex resulted in ectopic accumulations of deep layers within upper layers and vice versa, indicating complete loss of radial migration and totally disrupted neocortical development. In addition to expansive roles in transcription factor regulation and regulation of posttranscriptional factors, RBPs interact with epigenetic factors as well. The RBP splicing factor proline- and glutamine-rich (SPFQ) associates with the epigenetic regulators histone demethylase LSD1 (also known as KDM1A) and histone deacetylase CoREST (Saud et al., 2017). This proteineprotein interaction was confirmed in N2a cells and E17.5 neocortices. Silencing of SPFQ results in an accumulation of transfected neurons in the VZ/SVZ with fewer cells reaching the CP, whereas the number of cells in the IZ was unchanged. Cells that failed to migrate to the CP expressed upper-layer marker Cux1 (Saud et al., 2017), further suggesting the vulnerability upper-layer cell migration to precise intrinsic signaling. The histone deacetylase CoREST was found to control the multipolar to bipolar transition during radial migration (Fuentes et al., 2012). CoREST functions require the histone demethylase LSD1. When CoREST is silenced at E14.5, fewer cells reached the CP and accumulated in the VZ by E17.5. However, when analyzed at P0, there were fewer cells in upper layers, but an accumulation of cells across all other layers VZ, SVZ, IZ, and deep layers V and VI. Through elegant coelectroporation experiments, authors found that CoREST function in radial migration depends on the interaction with LSD1. Other domains that were important for interaction with REST/NRSF, SUMO1, or SUMO2/3 were irrelevant to radial migratory deficits. Silencing of LSD1 on its own phenocopied the migratory defect of silenced CoREST.

22.2.1.5 lncRNAs lncRNAs are RNA species that are longer than 200 nucleotides. They have been found to regulate transcription and chromatin states (Vance and Pontig, 2014). Though lncRNAs share many features with mRNAs, such as promoters, UTRs,

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and exons/introns, they are never translated. lncRNAs are highly expressed in the developing brain with remarkable spatiotemporal specificity, including within the neocortex (Aprea et al., 2013; Molyneaux et al., 2015; Johnson et al., 2015; Liu et al., 2016; Elsen et al., 2018; Lipovich et al., 2014). Therefore, lncRNAs were hypothesized to play significant roles in intricate developmental processes. Indeed, several lncRNAs are required for normal development, including of the neocortical layers (Sauvageau et al., 2013). lncRNAs were shown to regulate expression of well-studied neocortical transcription factors, including Neurog1. lncRNA also regulate transcripts specifically associated with neuronal migration. The lncRNA Paupar regulates expression and activity of Pax6 (Vance et al., 2014), whereas linc-Brn1b, Dali, and Pnky (Savageau et al., 2013; Chalei et al., 2014) regulate Brn-1. In particular, Dali silencing decreased Brn-1 (also known as Pou3f3) expression (Chalei et al., 2014). Dali was found to control gene expression by binding to promoters and by interacting with chromatin-modifying proteins to inhibit DNA methylation. Interestingly, Dali is recruited toward the DNA sites through binding to Brn-1. The lncRNA Pnky binds the splicing factor Ptbp1, but not Ptbp2 (Ramos et al., 2015; Zhang et al., 2016). Silencing of Pnky in developing neocortices results in more transfected cells in the IZ and fewer in the VZ, with no change in the portion of cells that reached the CP (Ramos et al., 2015). One recent study sought to determine the transcriptomes of proliferative progenitors (Btg2/Tis1 negative, Tubb3 negative), differentiating progenitors (Btg2/Tis1 positive, Tubb3 negative), and neurons (Btg2/Tis1 negative, Tubb3 positive) based on expression of predetermined and specific markers. After multifactor identification and verification of their isolated cell populations, authors isolated uncharacterized genes that were enriched in the “differentiating progenitors” group. lncRNAs were overrepresented in this group. Authors chose protein coding and lncRNA transcripts to further analyze, seemingly at random, with one condition that the gene does not have a previously reported role in cell fate specification of cortical progenitors (Aprea et al., 2013). Two lncRNAs were selected and both had an effect on migration to the CP: Gm17566 (w84.3%) and Miat (w38%). The genomic regions of both were investigated, and Gm17566 was found to lie antisense to Prox1 (Aprea et al., 2013). Prox1 is a transcription factor expressed by lateral ganglionic eminence (LGE)/caudal ganglionic eminence (CGE) interneurons (Rubin and Kessaris, 2013) and then only by CGE interneurons (Miyoshi et al., 2015). Prox1 is required for the migration of interneurons from the lateral migratory steam into the CP (Miyoshi et al., 2015; Touzot et al., 2016). It is unknown why a lncRNA (Gm17566) that is antisense to an interneuron lineage tracer (Prox1) has such dramatic effects on pyramidal neuron positioning. Despite its subtle effects on cell migration, Aprea et al. chose to further characterize Miat. Curiously, Miat overexpression and knockdown yielded similar, not opposite, phenotypes; Miat-depleted or Miat-enriched cells failed to reach the CP by about half of unaffected control neurons. Authors explained that either manipulation was akin to a loss of function (Aprea et al., 2013), possibly suggesting a dose-sensitive function of the lncRNA Miat. lncRNAs are transcripts that can act in both cis and trans to regulate their targets. For example, the most famous lncRNA, Xist, acts in cis to inactivate the X chromosome as part of a three-dimensional transcription complex. It is only through careful and arduous experimentation that the determination of a cis/trans mechanism can be confirmed (Kopp and Mendell, 2018). Recent attempts, and successes, to accomplish this at scale have been reported in other systems; one study notably used a phenotype-based cell culture assay to screen for lncRNAs of interest (Joung et al., 2017). Another focused on identifying lncRNAs in intergenic regions (lincRNAs) and generated an astonishing 18 knockouts, with one of these knockouts (linc-Brn1b) affecting lamination of upper layers (Sauvageau et al., 2013). In the cases of Gm17566 and Miat, no further attempts at characterization have been made. Inferences can be made based on genomic positioning; however, only more experiments would be truly informative of targets and mechanism.

22.2.1.6 MicroRNAs While mechanisms that lncRNAs utilize to posttranscriptionally regulate their targets require much unraveling, miRNA mechanisms are more straightforward. However, they are no less varied in the pathways that they target. For example, let-7 miRNA was identified to regulate radial migration by impacting autophagy of migrating neurons (Petri et al., 2017). However, the studies that solidified miRNAs as critical in development were a series of experiments with Dicer mutants. The intracellular maturation of miRNAs and their journey with Dicer and Argonaut proteins as part of the RISK complex has been well described and reviewed (Tonelli et al., 2008; Kawase-Koga et al., 2009; McNeill and Van Vactor, 2012; Barca-Mayo and Tonelli, 2014; Davis et al., 2015; Sun and Shi, 2015). Mature miRNAs have been associated with almost every step of neocortical development, including a number of miRNAs that have been implicated in neuronal migration. The expansive literature on Dicer deletions is informative to the broad scale of miRNA involvement in cortical development and migration (Kawase-Koga et al., 2009; McNeill and Van Vactor, 2012; Sun et al., 2011). Several mouse

494 PART | II Migration

mutants with conditional deletions of Dicer have been generated and exhibited migration defects. The effect was specific to prenatal deletion as Camk2-Cre (deletion starts in postnatal postmitotic neurons) Dicer deletions did not result in migration defects (Hebert et al., 2010; Davis et al., 2008). The earliest Dicer deletion (with FoxG1-Cre, deletion starts at E8.5-E9.5) results in a migration defect in early glutamatergic neurons. In particular, there was a misplacement of Tuj1þ, Tbr1þ, and/or HuD/HuCþ neurons throughout the early developing neocortex at E11.5, whereas these cell groups should normally form the layer that demarcates the CP (Nowakowski et al., 2011). Interestingly, authors found an expanded CajaleRetzius population with an excess of calretininþ CajaleRetzius cells, but not reelinþ ones, within Dicer-depleted forebrains. Later, conditional deletion of Dicer resulted in excess Cux1þ neurons. Conditional deletion with in utero electroporation at E13.5 did not result in an early migratory defect; however, many transfected cells were found in layer I at P14 (Nowakowski et al., 2013). These effects contrasted previous reports where Dicer was deleted more extensively using Emx1-Cre (Kawase-Koga et al., 2009). Authors used Emx1-Cre/Dicer (deletion starts at E9.5) conditional knockouts and suggested that both early- and late-born glutamatergic neurons showed premature differentiation and aberrant migration. However, when Dicer was depleted by Nestin-Cre (deletion starts at E11.5), cortices were thinner, but migration of only the late-born neurons was disrupted. This series of early Dicer deletions is not indicative of differential interpretation or experimentation, rather the results reflect the heterogenous neocortical cell population and the necessity to study cell typeespecific effects. Dicer deletions in early postmitotic neurons that are still migrating have also been examined (Volvert et al., 2014). The Nex-Cre-Dicer conditional knockout and also acute NeuroD-Cre (another driver specific for postmitotic neurons) electroporation exhibited similar migration phenotypes. In Nex-Cre-Dicer neocortices, more neurons were found in lower layer V and fewer in upper layers at P2. Meanwhile, acute deletion of Dicer using NeuroD-Cre electroporations resulted in an accumulation of neurons in the IZ by E17, with fewer present in the CP. When analyzed at P2, there were more transfected cells in layers V/VI and fewer in upper layers IIeIII (Volvert et al., 2014). This suggests a delayed migration of neurons from the IZ into the CP. It may be fruitful to examine postmitotic deletion at a later stage and determine if there is also a delayed or a halted migration of neurons into upper layers. CoREST was previously identified as upregulated in Dicer deletions, and it was found to regulate radial migration (Fuentes et al., 2012). Silencing of CoREST in Dicer-depleted neurons rescued the migratory pathology, whereas overexpressing CoREST disrupted migration. Using microarrays, authors demonstrated that miRNAs targeting CoREST (miR22, miR-124, and miR-185) were also reduced in Dicer-deficient cells. These miRNAs target the CoREST 30 UTR to reduce its expression. As expected, cotransfection of these miRNAs into Dicer-depleted cells rescued radial migration. When these miRNAs were silenced, the result was disrupted multipolar to bipolar transition during radial migration and an accumulation of CoREST mRNA, suggesting that CoREST is also subject to translational control. In this study, CoREST was found to inhibit the transcription of Dcx (Volvert et al., 2014). Thus, the pathway in a wild-type situation is that Dicer processes miRNA-22&124, which results in decreased CoREST, allowing expression of Dcx that then drives neuronal radial migration onward and upward. miR-134 was further described to target other neocortical genes such as Dcx (Gaughwin et al., 2011). In line with the above findings, miR-134 reduces cell migration in vitro and in vivo. This is due to control of its target Dcx. miR-134 decreases Dcx expression by targeting its 30 UTR. When pre-miR-134 is overexpressed in developing neocortices in vivo, significantly, fewer cells were found in the CP with more cells across VZ, SVZ, IZ, and SP. This pre-miR-134-induced pathology was restored by coelectroporating the Dcx. The histone demethylase, Lsd1, also plays a role in CoREST-regulated radial migration (Fuentes et al., 2012; Suad et al., 2017). miR-137 decreases Lsd1 expression in vivo and promotes differentiation and radial migration of transfected neurons. However, when LSD1 lacks its 30 UTR in the presence of overexpressed in miR-137, the effects were reversed, indicating that migration is dependent upon Lsd1 repression by the miRNA. In addition, authors identified nuclear receptor TLX as a repressor of miR-137 through recruitment of Lsd1 to DNA regions responsible for miR-137 expression. Silencing of TLX mimicked migratory effects of miR-137 overexpression (Sun et al., 2011; Li et al., 2008a). Correspondingly, silencing of miR-137 rescued the migratory defects seen when TLX was silenced. This series of experiments detailed a fascinating feedback regulatory loop between MiR-137 and Lsd1/TLX. miR-9 has been studied extensively and has received significant attention in neurodevelopment (Coolen et al., 2013; Radhakrishnan and Anand, 2016). miR-9 regulates the production of reelin-producing CajaleRetzius neurons by suppressing Foxg1 expression at its 30 UTR; interestingly, this effect was apparent in the medial pallium but not the dorsal one (Shibata et al., 2008). Silencing of miR-9 resulted in less CajaleRetzius neurons, whereas overexpression resulted in excessive amounts beyond the marginal zone where they normally reside. miR-9 regulates expression of Map1b, and several transcription factors, Foxp1 and Foxp2, that are known to regulate radial migration (Dajas-Bailador et al., 2012; Clovis et al., 2012; Shibata et al., 2011; Radhakrishnan and Anand, 2016). Indeed, laminar distribution of layer-specific markers was aberrant in the midst of miR-9 silencing; however, it needs to be established whether this is due to aberrant

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protein expression, loss of proliferation, or disrupted migration. miR-132 collaborates with miR-9 at the 30 UTR of Foxp2 to regulate its expression and migration (Clovis et al., 2012). Foxp2 mRNA and protein expression is increased and ectopic in the Emx1-Cre/Dicer knockout mouse. Ectopic expression of Foxp2 results in disrupted radial migration, which is reflected in an accumulation of transfected cells in the IZ. Though wild-type Foxp2 rescued the cell arrest, mechanistically, this was due to miR-9 and miR-134 acting on the 30 UTR of Foxp2 to precisely repress Foxp2. This was verified as mutation of the miRNA-binding sites in the Foxp230 UTR results in a significant migratory defect. Collectively, these results further confirm significance of miRNAs in radial migration and show that multiple miRNAs can cooperatively regulate the same transcript. Another miRNA that has been implicated in neuronal migration is miR-338 (Kos et al., 2017). miR-338 is upregulated at wE14.5 in the neocortex. To investigate the role of this miRNA in corticogenesis, authors in utero electroporated miR338 overexpression and corresponding sponge constructs. Overexpression at E14.5 resulted in an increase of cells in the IZ and the VZ, and the sponge decreased cells in both regions. Authors confirmed that miR-338 manipulations were not affecting neuronal viability. In addition, Kos et al. correlated migration defects with changes in morphology. Overexpression produced more multipolar neurons, and the sponge induced more nonpolar morphologies (Kos et al., 2017). Three miRNAs as part of the miR379-410 cluster can regulate radial migration by controlling the expression of N-cadherin (Rago et al., 2014). When either miR543 or miR369-3p and miR496 are overexpressed in developing neocortices, transfected cells rush out from the VZ. This was rescued by N-cadherin lacking its 30 UTR. When miR369-3p and miR496 were coexpressed in postmitotic neurons under the NeuroD1 promoter, more transfected cells localized to the lower CP and IZ and fewer to the upper CP. As expected, contrasting results were revealed when these miRNAs were silenced. miRNAs can be found in intergenic regions or in intragenic regions. Two miR-128 variants are found intragenically: miR-128-1 is located in an intron of R3hdm1, and miR-128-2 is located in an intron of Arpp21. Mature miR-128 is strongly expressed in the developing CP; the precursor of mIR-128-2 (pre-miR-128-2) is enriched in the VZ and SVZ, whereas the precursor of miR-128-1 (pre-miR-128-1) was not detectable in developing neocortices (Franzoni et al., 2015). These findings suggest that posttranscriptional processing of miR-128 and its precursors are spatiotemporally defined. To test this, authors prematurely expressed miR-128. When analyzing cortices at P7, overexpression of pre-miR-128-1 did not have a visible effect on radial migration. However, overexpression of pre-miR-128-2 disrupted migration of upper-layer Cux1þ neurons, which were dispersed throughout lower layers. Conversely, blocking miR-128 using a sponge resulted in more neurons migrating into upper layers. Thus, proper amounts of miR-128 at given times during radial migration are critical. Further work identified Pard6b and Phf6 as downstream targets of miR-128. Authors decided to overexpress Phf6 and a molecule important in migration, neuropilin-2, which is importantly not target of miR-128. Only Phf6 overexpression rescued the migratory defects caused by pre-miR-128-2 overexpression. It was suggested that miR-128 may contribute to terminal translocation of migrating neurons at the onset of differentiation. Notably, Phf6 is an intellectual disability gene (Franzoni et al., 2015; Zhang et al., 2013), and global miR-128 deletion or postmitotic neuron-specific deletion results in hyperactive motor behavior and severe epileptic seizures (Tan et al., 2013) (Table 22.2).

22.2.2 Tangential migration: transcriptional and posttranscriptional control Though interneurons are not the majority neuronal group in the neocortex, they play a crucial role in regulating excitatory activity generated by pyramidal neurons. Indeed, an imbalance of inhibition is hypothesized to result in severe cortical diseases such as epilepsy. As if migration of glutamatergic neurons and associated mechanisms were not fascinating enough, let us delve into the mechanisms behind the migration of neocortical interneurons. Interneurons are not even born in the developing neocortex. The GABAergic inhibitory neurons are born at a significant distance in the ganglionic eminences. They use a mode of migration called tangential migration (Anderson et al, 1997, 2001; 2002; Corbin et al., 2001; de Carlos et al., 1996; Lavdas et al., 1999; Main and Rubenstein, 2001; Evsyukova et al., 2013; Tamamaki et al., 1997; Wichterle et al., 1997, 1999, 2001; Letinic et al., 2001; De Carlos et al., 1996; Fertuzinhos et al., 2009; Petanjek et al., 2009; Jimenez et al., 2002; Nery et al., 2002; Guo and Anton, 2014; Xu et al., 2004; Sultan et al., 2013, 2018; Corbin et al., 2001; Faux et al., 2012). Once born, neocortically e destined GABAergic neurons will somehow know that they are different from any other neuron that is supposed to stay in close proximity to their progenitors. Cells will execute their journey to the developing neocortex with seemingly incredible precision down to the specific layer that corresponds to their birthdate. Interneuron migration seems doomed to be an odyssey, as an even more long distance migration is present in evolutionarily higher species. In mice, interneurons exclusively come to the cortex from the ganglionic eminences and

Posttranscriptional factor type/class

Key downstream gene In migration (confirmed or *proposed)

References

RNA-binding protein, KH type domains

Cadherin-2

La Fata et al. (2014)

HuR (Elavl1, HuA, 2410055N02Rik, W91709)

RNA-binding protein, RNA recognition motif domain

Foxp1, Foxp2

Kraushar et al. (2014), Popovitchenko et al. (2016)

DENR (1500003K04Rik), interacts with Mcts1

Associated with translation initiation, reinitiation, and ribosome recycling

Not reported

Haas et al. (2016)

Radial migrationdposttranscriptional regulator (synonyms) FMRP (Fmr1, Fmr-1)

Nova2 (LOC384569)

RNA-binding protein, KH-type domain

Dab1, dCC

Yano et al. (2010)

RBM4 (Lark1, Rbm4a)

RNA-binding protein, RNA recognition motif

Dab1

Dhananjaya et al. (2018)

Ptbp1 (hnRNP I, Ptb); acts opposingly to Rbfox RNA binding proteins

Heterogeneous nuclear ribonucleoprotein

Dab1

Dhananjaya et al. (2018) Zhang et al. (2016)

Unkempt (B230379M23Rik, mKIAA1753, Zc3h5)

RNA-binding zinc finger protein

iCLIP identified a number of putative Unkempt bound mRNA targets

Hall (2005)

eIF4E1; interacts with eIF4G and 4E-T

Initiation factor

bHLH transcription factors (e.g., neurogenin-2)

Yang et al. (2014)

Staufen-2; interacts with Pumilio-2

Double-stranded RNA-binding protein

Not reported

Vessey et al. (2008)

Rbfox1 (A2bp, A2bp1, FOX1, HRNBP1) isoforms 1 and 2

RNA-binding protein, RNA recognition motif

Not reported

Hamada et al. (2016)

nSR100/SRRM4 (1500001A10Rik, B230202K19Rik, bv, flopsy, fp, mKIAA1853, nSR100)

Splicing regulator

Not reported

Quesnel-Valieres et al. (2015)

PIWIL1 (MIWI)

Piwi RNA (piRNA)ebinding protein

Not reported

Zhao et al. (2015), Viljetic et al. (2017)

APC (CC1, Min)

RNA-binding protein

HITS-CLIP identified a number of putative APC-bound mRNA targets

Yokota et al. (2009), Preitner et al. (2014)

SFPQ (1110004P21Rik, 2810416M14Rik, 5730453G22Rik, 9030402K04Rik, D4Ertd314e, PSF, REP1); interacts with epigenetic regulators like KDM1A and CoREST

Splicing factor

Not reported

Saud et al. (2017)

Paupar

Long noncoding RNA

Pax6

Vance et al. (2014)

Dali

Long noncoding RNA

Brn-1

Chalei et al. (2014)

Pnky

Long noncoding RNA

Ptbp1

Ramos et al. (2015), Zhang et al. (2016)

Gm17566

Long noncoding RNA

Prox1

Aprea et al. (2013)

496 PART | II Migration

TABLE 22.2 Radial migration: posttranscriptional regulators.

Miat

Long noncoding RNA

Not reported

Aprea et al. (2013)

Let-7

microRNA

Not reported

Petri et al. (2017)

Dicer

Endoribonuclease, helicase with RNase motif

CoREST

Nowakowski et al. (2011), Kawase-Koga et al. (2009), Fuentes et al. (2012), Volvert et al. (2014)

microRNA

CoREST

Fuentes et al. (2012)

microRNA

CoREST

Fuentes et al. (2012)

miR-134

microRNA

Dcx

Gaughwin et al. (2011)

miR-137

microRNA

LSD1

Fuentes et al. (2012), Suad et al. (2017)

miR-9

microRNA

Foxg1, Map1b, Foxp1, Foxp2

Shibata et al. (2008), Clovis et al. (2012), Radhakrishnan and Anand (2016)

miR132; collaborates with miR-9

microRNA

Foxp2

Clovis et al. (2012)

miR-338

microRNA

Not reported

Kos et al. (2017)

miR379-410 complex; mir369-3p and miR496

microRNAs

Cadherin-2

Rago et al. (2014)

miR-128; pre-miR-128-2

microRNA; pre-microRNA

Pard6b, Phf6

Franzoni et al. (2015)

Molecular mechanisms of neuronal migration Chapter | 22

miR-22 miR-124

497

498 PART | II Migration

predominantly from the medial ganglionic eminence (MGE) and CGE but also from the LGE. Meanwhile, some interneurons in humans do exclusively originate from ventral progenitors but may emerge from the dorsal pallium (Letinic et al., 2001; Petanjek et al., 2009; Yu and Zecevic, 2011) and the ventral pallium (Fertuzhinos et al., 2009; Petanjek et al., 2009). Other interneurons of the forebrain emerge from the preoptic area (POA) and septal anlage. Distinct origins also mean different paths to arrive to their destined different brain regions: ventrolaterally to the striatum, caudally to hippocampus, and rostrally to the olfactory bulb. In addition, neocortical oligodendrocytes also originate from the ventral subpallium (Spassky et al., 1998). Traditionally, neocortical interneuron migration is divided into six stages: start of migration, joining a migratory route, crossing the corticostriatal boundary, joining the migratory stream within the developing neocortex, intracortical placement, and termination of migration. Once interneurons arrive in the developing neocortex, some interneurons will migrate below the CP, through the IZ, SVZ, and even the SP, whereas others will take their path above the CP. Upon reaching their final regional destination, interneurons will switch to radial migration to take their layer-specific positions (Nadarajah and Parnavelas, 2002; Miyoshi and Fishell, 2011; Ang et al., 2003; Yokota et al., 2007). Remarkably, recent evidence shows that glutamatergic neurons can dictate the ultimate layer position of interneurons (Lodato et al., 2011). Indeed, interneurons will not take their final positions until glutamatergic neurons acquire their identities (Lopez-Bendito et al., 2008; Pla et al., 2006; Miyoshi et al., 2007). The final destination of an interneuron is determined not only by their birthday, but also by their molecular signature and origin site (Miyoshi et al., 2010; Miyoshi and Fishell, 2011). The first interneurons to migrate to the neocortex come from the MGE and are followed by those that come from the CGE (Miyoshi et al., 2010; Pla et al., 2006; Valcanis and Tan, 2003). Migrating GABAergic interneurons born in the MGE follow the insideeout pattern of layer placement: earlier-born somatostatinþ (SSTþ) and parvalbuminþ (PVþ) interneurons migrate to the deeper layers, whereas later-born calretininþ (CR) neurons move above SST and PV neurons to upper layers (Miller, 1985; Fairen et al., 1986; Hammond et al., 2006; Yozu et al., 2004; Rymar and Sandikot, 2007; Ang et al., 2003; Butt et al., 2005; Cobos et al., 2006; Fogarty et al., 2007; Miyoshi et al., 2007, 2010; Nery et al., 2002; Xu et al., 2004; Rakic, 2009; Valcanis and Tan, 2003). Curiously, interneurons born in the CGE that use the caudal migratory route will not follow insideeout placement as the MGE-derived neurons do (Miyoshi et al., 2010). CGE-derived interneurons predominantly express reelin or vasoactive intestinal polypeptide (VIP) and are destined to mostly migrate to upper neocortical layers (Miyoshi et al., 2010). By Defining the basic subpopulations, we can distinguish some features of interneurons. However, recent studies have revealed the complexity of interneurons at an increasing pace. GABAergic interneuron heterogeneity is based on transcriptomic differences, shape, and electrophysiological differences that may even change in adult brains (DeFelipe et al., 2013; Mi et al., 2018). A number of detailed reviews have been written on the fascinating behavior of interneurons as they migrate (Evsyukova et al., 2013; Yokota et al., 2007; Kwan et al., 2012a; Manent et al., 2011; Nadarjah and Parnavelas, 2002; Kriegstein and Noctor, 2004; Imamura et al., 2012; Tan and Shi, 2013; Huang, 2009; Chu and Anderson, 2015; Wonders and Anderson, 2006; Sultan et al., 2013; Hu et al., 2017; Bandler et al., 2017; Corbin et al., 2001; Marin, 2013; Higginbotham et al., 2012; Guo and Anton, 2014; Marin and Rubenstein, 2001, 2003; Bartolini et al., 2013; DeBoer and Anderson, 2017; Ayala et al., 2007). Upon reflection, we would do poor justice in describing the minute details of tangential migration. Instead, we aim to summarize the most important transcriptional and posttranscriptional regulators of tangential migration.

22.2.2.1 Interplay of transcription factors and tangential migration guidance cues The laminar distribution of interneurons was recently shown to depend upon glutamatergic excitatory neurons (Lodato et al., 2011). Deletion of the deep layer transcription factor Fezl (also known as Fezf2 or Zfp312) results in loss of subcerebral projections (Chen et al., 2005a, 2005b; Molyneaux et al., 2005). Neocortical neurons lacking Fezf2 acquire upper-layer identities and callosal projections. Ultimately, this affects the intracortical distribution of GABAergic neurons as fewer interneurons were found in deep layers. This phenotype was paralleled by an increased accumulation of GABAergic neurons in upper layers. Authors confirmed that the total number of GABAergic neurons within the neocortical layers did not decrease. Misplacement did not disrupt fate specification of cortical interneurons, but SSTþ and PVþ interneurons were misplaced into upper layers and CRþ interneurons were unaffected. When Fezf2 is misexpressed in upper layers and ectopic subcortical projections are induced, GABAergic SSTþ and Lhx2þ interneurons in deep layers aggregate within these corticofugal ectopic projections, whereas reelinþ interneurons of upper layers were absent. The homeobox genes Dlx1/Dlx2 are important for interneuron differentiation. Deletion of both results in failed migration of interneurons and formation of heterotopias close to the ventricles of the subpallium (Anderson et al., 1997a).

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Additionally, fewer interneurons reach the neocortex (Anderson et al., 1997b). These deletions affect late-born interneurons. Furthermore, Dlx1/Dlx2 directly regulates the expression of Zfhx1b (also known as Sip1 or Zeb2) to generate MGE-derived cortical interneurons. Deletion of Zfhx1b leads to steady expression of Nkx2.1, which results in a switch from cortical to striatal interneurons. In contrast to Dlx1/Dlx2, the deletion of the bHLH transcription factor Ascl1 (previously known as Mash1) depleted early-born GABAergic interneurons (Casarosa et al., 1999). In Ascl1-depleted brains, interneurons destined to use the MZ as the migratory stream was more significantly depleted than ones that migrated under the CP, SP, or within the IZ. Ascl1 is expressed at low levels in migrating interneurons within the neocortical SVZ and IZ (Miyoshi et al., 2010). Overexpression of either Ascl1 or Dlx2 into the ventral telencephalon increased the number of transfected cells that migrated tangentially into the neocortex (Liu et al., 2017). In particular, Ascl1 increased the presence of migrating neurons in VZ, SVZ, and IZ, whereas Dlx2-transfected neurons migrated through the VZ and SVZ. Amazingly, when either Ascl1 or Dlx2 was overexpressed in RG, transfected glutamatergic neurons appeared to change their mode of migration from radial to tangential. When Dlx2 was silenced in cells that were transfected with Ascl1 overexpression, more transfected cells were found in the IZ, but significantly fewer in the VZ and SVZ (Liu et al., 2017). These findings suggest that Dlx2 induces tangential migration through VZ and SVZ as a downstream Ascl1 target, whereas Ascl1 itself promotes tangential migration through the IZ. Members of ephrin signaling, Eph receptor B1 and B2 (Ephb1 and Ephb2), were found as additional Ascl1 downstream targets in regulating the tangential migration. In Ascl1 null mutant, the expression of different members of notch signaling was lost in ventral telencephalon, Delta-like 1 (Dll1), Delta-like 3 (Dll3), and Hes5 (Casarosa et al., 1999), suggesting that Ascl1 is required for notch signaling within the ventral telencephalon. Another homeobox transcription factor, NK2 homeobox 1 (Nkx2.1), is expressed in the MGE and regulates MGEderived interneuronal tangential migration (Sussel et al., 1999). Expression levels of Nk2.1 distinguish subpallial neurons destined for the neocortex (low expression) versus ones that will populate the basal ganglia (high expression) (Nobrega-Pereira et al., 2008). In this way, Nkx2.1 serves as a sorting factor that decides to which pool interneurons belong. Nkx2.1 specifies the fate of interneurons through its downstream target, another transcription factor, the LIMhomeodomain family member, Lhx6 (Du et al., 2008). Lhx6 is undetectable in Nkx2.1 knockouts, whereas Nkx2.1 directly promotes the expression of Lhx6 through its promoter. Furthermore, Lhx6 rescues the MGE-derived interneuron migratory defects of the Nkx2.a knockouts. Nkx2.1 knockouts lack SSTþ and PVþ interneurons. The rescue of this phenotype by overexpression of Nkx2.1 is dampened by silencing Lhx6 (Du et al., 2008). Similarly, Lhx6 expression in the MGE of the Nkx2.1 knockout induced GABA expression and rescued SSTþ and PVþ interneurons. Lhx6 regulates tangential migration of MGE-derived interneurons (Alifragis et al., 2004; Liodis et al., 2007; Zhao et al., 2008; Neves et al., 2013; Vogt et al., 2014; Flandin et al., 2011). Briefly, disrupted Lhx6 reduces SSTþ and PVþ interneurons, slows tangential migration, and induces abnormal intracortical distribution of interneurons. Interneurons in Lhx6 mutants abnormally take a very deep SVZ migratory route instead of the superficial migratory stream (Liodis et al., 2007). Lhx6 is expressed in SSTþ and PVþ tangentially migrating interneurons and marks a subset of Dlxþ migrating neurons (Liodis et al., 2007; Zhao et al., 2008). In Lhx6-PLAP mice with Lhx6 that lacks exons 3e6 and part of exon 2 (these exons contain the N-terminus, both LIM1 and LIM2 domains, and 49e62 amino acid residues of the homeodomain), interneurons reach the neocortex, but will not populate the MZ or the CP. Most notably, cells did not migrate into layers IIeIV, with some reduction in layer V. Interestingly, interneurons migrating through the SVZ were possibly increased. It has been suggested that the migratory defects in Lhx6PLAP/PLAP mice occur via a decrease of ErbB4, the receptor for neuregulin-1 (Flames et al., 2004), expression in Lhx6-mutated brains. In addition, levels of a chemokine receptor CXCR4 were reduced. The chemokine CXCL12 (also known as SDF1) and its receptor, CXCR4, regulate laminar positioning of interneurons (Stumm et al., 2003; Li et al., 2008b; Lopez-Bendito et al., 2008). Reduced levels of CXCL12 and CXCR4 are also present in Lhx6PLAP animals, but to more severe effect (Zhao et al., 2008). Overall, disrupted Lhx6 resulted in decreased SSTþ, PVþ, and calretininþ neocortical interneurons, whereas NPYþ interneurons were spared. Further studies showed that in Lhx6 knockouts, some MGE-derived cells acquire the CGE-like fate, with expression of Sp8 in the MGE of Lhx6PLAP/PLAP mice. As reelin is expressed in superficial layer interneurons derived from the CGE and in deep layer interneurons derived from the MGE (Alcantara et al., 1998; Miyoshi et al., 2010), more reelinþ interneurons were found in upper layers of Lhx6PLAP/PLAP. This further suggests that MGE neurons partially acquire a CGE fate (Vogt et al., 2014). The cell-autonomous migratory defects of Lhx6PLAP/PLAP cells were further confirmed through elegant transplantation and rescue experiments. In addition, the Lhx6PLAP/PLAP phenotypes were rescued by Lhx6 downstream targets Lhx8 and CXCR7. Lhx6 promotes expression of Arx through direct binding of its enhancer, which is within the adjacent PolA1 gene on chromosome X and intronic

500 PART | II Migration

Cxcr7 enhancer (Vogt et al., 2014). The chemokine receptor CXCR7 and its ligand CXCL12 were further revealed to regulate the placement of interneurons within layer V and some PVþ neuronal differentiation or maturation. Lhx6 was found to coordinate gene expression associated with interneuron development and migration (e.g., Shh, Nkx2.1) together with Lhx8 (Flandin et al., 2011). The special AT-rich DNA-binding protein 1 (SATB1) was revealed to act downstream of Lhx6 to induce the transition of tangentially migrating immature interneurons into terminally differentiated SSTþ interneurons (Denaxa et al., 2012). Expression of Satb1 is induced once migrating neurons enter the cortical plate, and expression is activity dependent. Overexpressing Satb1 halts tangential migration, and deletion results in abnormal differentiation of SSTþ interneurons (Denaxa et al., 2012; Close et al., 2012). While Lhx6 does not induce expression of Satb1, Lhx6 needs Satb1 to properly differentiate interneurons. Another transcription factor, Sox6, was revealed as a downstream effector of Lhx6 (Batista-Brito et al., 2009). Sox6 and Lhx6 colocalize in migrating interneurons. Deletion of Sox6 resulted in loss of PVþ and SSTþ interneurons, whereas VIPþ interneurons were unaffected. Migrating interneurons that lack Sox6 were able to reach the neocortex but were not capable of switching from tangential to radial migration. This results in abnormal accumulation of cells within layers I and VI, at the expense of other layers. Removal of Sox6 from MGE also results in encephalopathic epilepsy. Another zinc finger transcription factor that regulates the migration of MGE-derived interneurons is Sp9 (Liu et al., 2018). Sp9 deletion via Nkx2.1-Cre results in a retardation of tangential migration through the IZ, SVZ, and importantly the MZ. This suggests that Sp9 is required to promote tangential migration of MHE-derived interneurons. When P30 adult neocortices were analyzed, Sp9 deletion resulted in 70% fewer PVþ and around 30% fewer SSTþ neurons, whereas NPYþ interneurons were increased. Similarly, when Sp9 was selectively deleted from postmitotic migratory interneurons, PVþ interneurons were reduced by about 65%. However, MGE-derived interneurons accumulated as Nkx2.1þ heterotopias within the ventral telencephalon of the Sp9 conditional knockouts. Using an unbiased RNAseq approach, authors identified that the expressions of Lhx6, Lhx8, Shh, and Arx were reduced in Sp9-null mutants. Sp6 was shown to directly bind to the Lhx6 promoter and putative enhancers of Lhx6 and Lhx8, but was only able to functionally activate one putative Lhx6 enhancer. This suggests a possible mechanism of Lhx6 expression driven by Sp6. Another LIM homeodomain transcription factor, Lhx1, regulates tangential migration of cortical interneurons from the preoptic area (POA) (Symmank et al., 2018). Conditional deletion of Lhx6 using Hmx3-Cre resulted in a switch from normal superficial migration to a deep route through the basal telencephalon. Once they crossed the pallial/subpallial boundary, Lhx1-deficient cells accumulated within the deep VZ and SVZ compared with control, and neurons spread out mostly through the IZ, CP, and MZ. These migratory defects were associated with changes in the expression levels of the bidirectional repulsive ephrin signaling members EFNB3/EPHA4. Lhx1 deletion resulted in an increase of EPHNA4, which is decreased in normal migratory POA cortical interneurons. In contrast, when Lhx1 was silenced, levels of EFNB3 decreased (Zimmer et al., 2011). Once they reached the neocortex, Lhx1-deficient interneurons will accumulate in upper neocortical layers at a higher rate than normal. Interestingly, many of the POA-derived interneurons expressed Ctip2, a widely used marker of layer V glutamatergic neurons. The chicken ovalbumin upstream promoter transcription factor II (COUP-TFII; also known as Nrf2f2 and characterized as an orphan nuclear receptor) plays role in radial migration and also regulates migration of interneurons (Kantani et al., 2008). In particular, COUP-TFII is required for the caudal migratory stream (Kantani et al., 2008). It is expressed not only in the CGE and in about 85% of CGE-derived interneurons, but also in about 10% of MGE-derived interneurons (Kanatani et al., 2008; Willi-Monnerat et al., 2008; Miyoshi et al., 2010). COUP-TFII, via Sp1, binds the promoter of neuropilin-2 and induces its expression to regulate migration of POA-derived interneurons (Kanatani et al., 2008, 2015). Overexpression of COUP-TFII directs interneurons to migrate into the amygdala and to the caudal end of the neocortex. Yet, it resulted in a failure of interneurons to migrate into the dorsal neocortex. When Coup-tfII was silenced, more interneurons migrated to rostral neocortical regions. Overexpression of neuropilin-2 in the COUP-TFIIsilenced cells rescued the migratory defects. In contrast, COUP-TFI (also known as Nrf2f1) mutant mice have an altered laminar distribution of reelinþ, VIPþ, CRþ, and VIPþCRþ interneurons (Touzot et al., 2016). It was suggested that COUP-TFI regulates tangential migration from CGE by regulating transcription factor Sp8 and COUP-TFII expression. Transcriptional repression was also implicated as a mechanism controlling interneuron migration from the POA (Pensold et al., 2017). Deletion of DNA methyltransferase (Dnmt1) results in defective migration and a decrease of interneurons within the neocortex. This is regulated by the downstream Dnmt1 target Pak6. Dnmt1 represses Pak6 expression to keep migrating neurons undifferentiated and thereby capable of migrating. Suppression of Pak6 expression is independent of methylation of Pax6 or its known upstream regulators, suggesting a de novo regulatory site or new Dntm1 functions (Fig. 22.2).

Molecular mechanisms of neuronal migration Chapter | 22

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FIGURE 22.2 Transcriptional and posttranscriptional control of neuronal tangential migration. Top left: GABAergic neurons are born predominantly in subpallial ganglionic eminences and use multiple tangential migratory streams to reach the neocortex. Bottom left, right: List of known transcriptional and posttranscriptional regulators of tangential migration.

A number of extracellular factors have also been associated with transcriptional and posttranscriptional control that may guide interneurons toward the neocortex. This includes trophic factors, such as brain-derived neurotrophic factor (BDNF) and neurotrophin-4 (Brunstrom et al., 1997). In addition, hepatocyte growth factor/scattered factor and its cleavage regulator urokinase-type plasminogen activator protein have been associated with interneuron migration, specifically calbindinþ neurons (Powell et al., 2001). Guidance molecules such as semaphorin signaling (e.g., neuropilin-1, neuropilin2, Sema3A/3F, ephrin e.g., (Ephb2, EFNB3, EPHA4) and Slit (Slit1/2) have been reported to regulate tangential migration (Marin et al., 2001; Tamamaki et al., 2003; Zimmer et al., 2011). Finally, posttranslational control via deglutamynation by carboxypeptidase 1 regulates tangential migration of cortical interneurons (Silva et al., 2018). Other modes of tangential migration of noncortical interneurons are out of the scope of this chapter (Table 22.3). 22.2.2.1.1 Posttranscriptional events in tangential migration: the role of RNA-binding proteins and microRNA Very little is known about posttranscriptional processes in interneuron migration. In mice, miR-9 has three precursor variants. In the double pre-miR9-2/3 mutant mice, Foxg1 expression was increased. This double mutation results in suppressed production of CajaleRetzius cells and early-born neurons (Shibata et al., 2011). These phenotypes were rescued by overexpression of an RBP, Elavl2 (Hu antigen B). In addition to these malformations, tangential migration of interneurons was disrupted. Interestingly, miR-9 regulates Gsh2 and thus tangential migration of olfactory bulb neurons. Overall, miR-9 was implicated as a master regulator of many types of neuronal migration. Postmitotic MGE-derived interneurons express several miRNAs, suggesting their importance in tangential migration (Tuncdemir et al., 2015). Dicer conditional deletion from MGE-derived interneurons that express Nkx2.1 resulted in an overall decrease of Gad67- and Lhx6-expressing interneurons (Tuncdemir et al., 2015). Even though interneurons with depleted Dicer expressed GABAergic features, there was a decreased proportion of interneurons expressing PV or SST. Furthermore, the number of interneurons using the MZ to migrate tangentially in the developing neocortex was significantly reduced, whereas mild reduction was observed in the deeper route through the IZ and SVZ. Interestingly, within the SVZ/IZ route, Dicer deletion results in more SSTþ neurons to be contained when compared with the wild-type control. Even though the migrating interneurons took these pathways at lower rate, their entrance into the CP was further decreased. Microarray analysis revealed that Dicer deletion resulted in increased expression of apoptotic machinery members, including Igf1. lncRNAs such as evf2 were associated with interneuron migration through regulation of the homeodomain box containing transcription factors Dlx5 and Dlx6 (Feng et al., 2006; Wang and Chang, 2011; Wang et al., 2017). Absence of the Evf2 (also known as Dlx6os) lncRNA disrupts tangential migration (Bond et al., 2009; Kohtz, 2014) (Table 22.4).

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TABLE 22.3 Tangential migration: transcriptional regulators. Tangential migration - transcriptional regulator (synonyms)

Transcription factor type/class

Key downstream gene in migration

Ascl1 (ASH1, bHLHa46, Mash1)

Basic helixeloopehelix

Dlx2, Ephb1, Ephb2, Dll1, Dll3, Hes5

Casarosa et al. (1999), Miyoshi et al. (2010), Liu et al. (2017)

COUP-TFI (Nrf2f1, COUP-TF1, Erbal3, Tcfcoup1)

Chicken ovalbumin upstream promoter transcription factor

Not reported

Touzot et al. (2016)

COUP-TFII (Nrf2f2, 9430015G03Rik, Aporp1, ARP-1, COUP-TF2, EAR3, Tcfcoup2); interacts with Sp1

Chicken ovalbumin upstream promoter transcription factor

Neuropilin-1

Kantani et al. (2008), Kantani et al. (2015)

Dlx1 (DII B, Dlx, Dlx-1)/Dlx2 (DII A, Dlx-2, Tes-1)

Homeobox domain

Zfhx1b

Anderson et al. (1997),

Dnmt1 (Cxxc9, Dnmt1o, MommeD2, MTase)

DNA methyltransferase

Pak6

Pensold et al. (2017)

Fezf2 (Fez, Fezl, forebrain embryonic zinc finger, Zfp312)

Zinc finger protein

Not reported for migration

Lodato et al. (2011)

Lhx1 (Lim1)

LIM homeodomain

Efnb3, Ephna4

Symmank et al. (2018)

Lhx6

LIM homeodomain

Erbb4, Satb1, Sox6

Alifragis et al. (2004), Liodis et al. (2007), Zhao et al. (2008), Batista-Brito et al. (2009), Neves et al. (2013), Vogt et al. (2014), Flandin et al. (2011), Denaxa et al. (2012), Flames et al. (2004)

Nkx2.1 (T/EBP, thyroid-specific enhancer-binding protein, thyroid transcription factor-1, tinman, Titf1, Ttf-1)

Homeobox domain

Lhx6

Nobrega-Pereira et al. (2008), Du et al. (2008),

Lhx6 together with Lhx8 (L3, Lhx7)

LIM homeodomain

Shh, Nkx2.1

Flandin et al. (2011)

Satb1 (2610306G12Rik)

Special AT-rich DNAbinding protein 1

Not reported

Denaxa et al. (2012), Close et al. (2012)

Sox6

Sex determining region Y-box

Not reported

Batista-Brito (2009)

Sp9

Zinc finger protein

Unbiased RNAseq approach revealed Lhx6, Lhx8, Shh, and Arx to be putative downstream targets

Du et al. (2008)

Reference(s)

TABLE 22.4 Tangential migration: posttranscriptional regulators. Radial migrationdposttranscriptional regulator (synonyms)

Posttranscriptional factor type/class

Key downstream gene In migration

Evf2 (Dlx6os)

Long noncoding RNA

Dlx5, Dlx6

Feng et al., (2006); Wang and Chang, 2011; Wang et al., (2017), Bond et al. (2009), Kohtz (2014)

miR-9; pre-miR9-2/3

microRNA

Elavl2, Gsh2

Shibata et al. (2011)

Dicer

Endoribonuclease, helicase with RNase motif

Unbiased microarray analysis revealed a number of putative targets, including Igf1

Tuncdemir et al. (2015)

References

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22.3 Conclusion and future directions Tremendous progress has been achieved in understanding the roles of transcription factors and posttranscriptional regulators in neuronal migration. The main question still remains: How do these two major gene expression steps collaborate in space and time to perfectly orchestrate the fascinating journey of both glutamatergic and GABAergic neurons? Importantly, these intrinsic regulators are known to respond to extracellular factors, which were also associated with the control of neuronal migration, as we described previously. Thus, it will be interesting to reveal how extracellular factors dictate collaborative (or competitive) dynamics of transcriptional and posttranscriptional events in neuronal migration. Remarkably, while transcriptional regulators are well studied, we have scarce knowledge on posttranscriptional regulators of neural migration. Posttranscriptional events may give rapid plasticity to vast demands, as neurons go through distinct migratory stages and regions. Nevertheless, we do have a somewhat better understanding of posttranscriptional players in radial than in tangential migration. Indeed, posttranscriptional regulators of interneuronal tangential migration are practically unknown and are just starting to be revealed. Thus, it will be an important field to rise and to reveal the impact of posttranscriptional events in tangential migration.

List of acronyms and abbreviations CP VZ GE IPs IZ KO lncRNA miRNA MZ POA PP RBP RG oRG SP SVZ oSVZ PVFRZ VZ

cortical plate ventricular zone ganglionic eminence intermediate progenitors intermediate zone knockout long noncoding RNA microRNA marginal zone preoptic area preplate RNA-binding protein radial glia outer radial glia subplate subventricular zone outer subventricular zone periventricular reach zone ventricular zone

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Cerebr. Cortex 25 (10), 3535e3546. Wang, K.C., Chang, H.Y., 2011. Molecular mechanisms of long noncoding RNAs. Mol. Cell 43 (6), 904e914. Wang, A., Wang, J., Liu, Y., Zhou, Y., 2017. Mechanisms of long non-coding RNAs in the assembly and plasticity of neural circuitry. Front. Neural Circuits 11, 76. Wegiel, J., Kuchna, I., Nowicki, K., Imaki, H., Wegiel, J., Marchi, E., Ma, S.Y., Chauhan, A., Chauhan, V., Bobrowicz, T.W., de Leon, M., Louis, L.A., Cohen, I.L., London, E., Brown, W.T., Wisniewski, T., 2010. The neuropathology of autism: defects of neurogenesis and neuronal migration, and dysplastic changes. Acta Neuropathol. 119 (6), 755e770. Wichterle, H., Garcia-Verdugo, J.M., Alvarez-Buylla, A., 1997. Direct evidence for homotypic, glia-independent neuronal migration. Neuron 18, 779e791. Wichterle, H., Garcia-Verdugo, J.M., Herrera, D.G., Alvarez-Buylla, A., 1999. Young neurons from medial ganglionic eminence disperse in adult and embryonic brain. Nat. Neurosci. 2 (5), 461e466. Wichterle, H., Turnbull, D.H., Nery, S., Fishell, G., Alvarez-Buylla, A., 2001. In utero fate mapping reveals distinct migratory pathways and fates of neurons born in the mammalian basal forebrain. Development 128 (19), 3759e3771. Wiegreffe, C., Simon, R., Peschkes, K., Kling, C., Strehle, M., Cheng, J., Srivatsa, S., Liu, P., Jenkins, N.A., Copeland, N.G., Tarabykin, V., Britsch, S., 2015. Bcl11a (Ctip1) controls migration of cortical projection neurons through regulation of Sema3c. Neuron 87, 311e325. Willi-Monnerat, S., Migliavacca, E., Surdez, D., Delorenzi, M., Luthi-Carter, R., Terskikh, A.V., 2008. Comprehensive spatiotemporal transcriptomic analyses of the ganglionic eminences demonstrate the uniqueness of its caudal subdivision. Mol. Cell. Neurosci. 37 (4), 845e856. Wonders, C.P., Anderson, S.A., 2006. The origin and specification of cortical interneurons. Nat. Rev. Neurosci. 7, 687e696. Xu, Q., Cobos, I., De La Cruz, E., Rubenstein, J.L., Anderson, S.A., 2004. 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Birth-date dependent alignment of GABAergic neurons occurs in a different pattern from that of non-GABAergic neurons in the developing mouse visual cortex. Neurosci. Res. 49, 395e403. Yu, X., Zecevic, N., 2011. Dorsal radial glial cells have the potential to generate cortical interneurons in human but not in mouse brain. J. Neurosci. 31 (7), 2413e2420. Zahr, S.K., Yang, G., Kazan, H., Borrett, M.J., Yuzwa, S.A., Voronova, A., Kaplan, D.R., Miller, F.D., 2018. A translational repression complex in developing mammalian neural stem cells that regulates neuronal specification. Neuron 97 (3), 520-537.e6. Zhang, B., Wang, W., Zhang, Z., Hu, Y., Meng, F., Wang, F., Lou, H., Zhu, L., Godbout, R., Duan, S., Gao, Z., 2017. Alternative splicing of disabled-1 controls multipolar-to-bipolar transition of migrating neurons in the neocortex. Cerebr. Cortex 22, 1e11. 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Novel function of PIWIL1 in neuronal polarization and migration via regulation of microtubule-associated proteins. Mol. Brain 8, 39. Zimmer, G., Rudolph, J., Landmann, J., Gerstmann, K., Steinecke, A., Gampe, C., Bolz, J., 2011. Bidirectional ephrinB3/EphA4 signaling mediates the segregation of medial ganglionic eminence- and preoptic area derived interneurons in the deep and superficial migratory stream. J. Neurosci. 31 (50), 18364e18380.

Chapter 23

Migration of myelin-forming cells in the CNS Fernando de Castro1 and Bernard Zalc2 Instituto Cajal-CSIC, Spanish Research Council/Consejo Superior de Investigaciones Científicas-CSIC, Madrid, Spain; 2Université Pierre & Marie

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Curie-Paris 6, Centre de Recherche de l’Institut du Cerveau et de la Moelle épinière, UMR_S 975, Inserm U 975, CNRS UMR 7225, Hôpital de la Salpêtrière, Paris, France

Chapter outline 23.1. Introduction 23.1.1. Genesis of myelin-producing cells during development 23.1.2. Oligodendrocyte precursor cells: born to migrate 23.2. Migratory paths followed by oligodendrocyte progenitor and precursor cells 23.3. Chemokinetic factors: the motility of oligodendrocyte precursors

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23.4. Adhesion and chemotactic mechanisms: how the movement of oligodendrocyte precursors is guided? 23.4.1. Adhesion and surface molecules 23.4.2. Secreted factors 23.5. Concluding remarks Acknowledgments References

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23.1 Introduction Myelin, the membranous sheath wrapping large diameter axons, has been a fantastic acquisition of the vertebrates nervous system, an acquisition that can be approximately dated 425 million years ago (Zalc and Colman, 2000; Zalc et al., 2008). In addition to save space and energy as well as to protect the axons, the principal function of the myelin sheath is to increase the velocity of electrical conduction along the axons: depending on the species and the tract, for axons which have the same diameter the myelin sheath increases the speed of conduction by a factor of at least 50 (Huxley and Stampfli, 1949). Two cell types of different embryologic origin can synthesize the myelin: the oligodendrocyte, in the CNS, and the Schwann cell, in the PNS. The former is produced by the neural tube, while the latter emerged from the neural crest. (A third cell type, the olfactory ensheathing glia, originating from the olfactory placode, does not myelinate under physiological conditions, but has the potential to form myelin when transplanted into the CNS; Franklin and Barnett, 2000). As a consequence, although the function of myelin is similar in the two parts of the nervous system, the molecular composition of the myelin sheath is very different whether in the CNS or the PNS (Raine, 1984). Morphologically, myelination is also strikingly different in the PNS and in the CNS: Schwann cell myelinates a single internode on a single axon, and in mammals, the myelinated internode length ranges between 700 and 1000 mm; in contrast, in the CNS, each oligodendrocyte myelinates several internodes (for example, in the rat, the number varies between 2e3 in the spinal cord, 10 and 20 in the corpus callosum, and up to 40 and 50 in the optic nerve), and the length of each internode varies between 100 and 200 mm (Bjartmar et al., 1994; Peters et al., 1991). The discovery that very subtle differences of normal myelin can drastically influence in the physiology of CNS projection neurons opens new avenue to study the role of myelin in normal brain physiology (Tomassy et al., 2014, 2016; Remaud et al., 2017).

Cellular Migration and Formation of Axons and Dendrites. https://doi.org/10.1016/B978-0-12-814407-7.00023-7 Copyright © 2020 Elsevier Inc. All rights reserved.

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23.1.1 Genesis of myelin-producing cells during development Based on either morphological or biochemical criteria, CNS myelin appears identical all along the caudo-rostral axis, from the spinal cord to the olfactory bulb (Morell, 1984). As a consequence, it has for long been assumed that oligodendrocytes were identical and had a similar unique origin all along the neural tube. This assumption was held for a long time despite the fact that based on various criteria, either their morphology (del Rio-Hortega, 1928; Butt et al., 1994; Butt et al., 1997; Fanarraga and Millward, 1997; Hildebrand et al., 1993), their spike-firing physiology (Fröhlich et al., 2011), or their expression of biochemical markers (Butt et al., 1995) or their myelinating plasticity (Belichenko and Celio, 1997; Fanarraga et al., 1998), oligodendrocytes appear as an heterogenous cell population. Furthermore, the initial studies on the origin of oligodendrocytes had pointed to a unique ventral source induced by sSonic hedgehog Hedgehog (Warf et al., 1991; Noll and Miller, 1993). It had notably been shown that, in the spinal cord, oligodendrocytes originate within the pMN domain, i.e., the same domain from where motoneurons originate. The finding that the bHLH transcription factors Olig-1 and Olig-2 are required for motor neuron and oligodendrocyte specification in the spinal cord had even led to speculate the existence of a common bi-potent Olig-specified progenitor, giving rise to both motor neurons and oligodendrocytes (Lu et al., 2001; Zhou and Anderson, 2002). However, all these simple views on the origin of oligodendroglial lineage have been challenged and reconsidered. Different markers have been proposed to identify early progenitors of oligodendrocyte. (Fig. 23.1). The most commonly used is the platelet-derived growth factor receptor alpha (PDGFRa), which is in line with the dependence of most oligodendrocyte precursor cells (OPCs) on PDGF-AA for their proliferation and survival (Richardson et al., 1988; Pringle and Richardson, 1993). Another population of oligodendrocyte precursor cellOPCs has been characterized by the expression of plp/dm-20 transcripts (Timsit et al., 1995; Spassky et al., 1998). The plp gene encodes the proteolipid protein (PLP) and its alternatively spliced product DM-20, which are major proteins of CNS myelin in mammals (PLP and DM-20 account for 50% of the total protein content of myelin; Greenfield et al., 1971). It has been shown that plp-expressing oligodendrocyte precursor cellOPCs do not depend on signal transduction mediated by PDGFRa, and therefore that they belong to a different lineage than the PDGFRa-expressing precursors (Spassky et al., 2001). Another dogma was the strict ventral origin of the oligodendrocytic lineage (Warf et al., 1991) (Fig. 23.2). This has been contested in the spinal cord by the demonstration that the dorsal Dbx1-expressing ventricular domains also give rise to oligodendrocytes, and independently from Shh signalling (Cameron-Curry and Le Douarin, 1995; Cai et al., 2005; Fogarty et al., 2005; Vallstedt et al., 2005). These dorsal domains of oligodendrogenesis are not a sole curiosity of the spinal cord: it It has similarly been shown that in the telencephalon, in addition to a ventral Nkx2.1-expressing domain, oligodendrocyte precursor cellOPCs are also generated from dorsal and alar domains characterized by the expression of Emx1 and Gsh2, respectively (Kessaris et al., 2006). Altogether, it appears that oligodendrocytes are generated all along the caudo-rostral axis of the ventricular neuroepithelium of the neural tube in multiple restricted foci distributed along the dorso-ventral axis. Oligodendrogenesis is not a chronologically coordinated event: vVentral oligodendrocyte precursor cellOPCs emerge earlier than the dorsal ones (E12.5 vs. E18.5, respectively, in the mouse). A similar gradient is observed along the caudal to

(A)

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FIGURE 23.1 Migrating oligodendrocyte precursors. (A): An oligodendrocyte precursor cell isolated from the cerebral cortex of a P0 mouse, in culture for 72 h, shows the typical bipolar morphology, with a long leading process (to the right) and a very short trailing process. The cell has been stained with A2B5 mAb, and the cell nucleus was visualized with DAPI. (B): Migrating oligodendrocyte precursor cells in the hindbrain of an E15.5 mouse embryo (flat-mount), immunostained with anti-PDGFRa Ab. The cells observed at a short distance of the ventricular layer where they originate present a unipolar morphology, reminiscent of the cells observed in culture as shown in (A).

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FIGURE 23.2 Regions of oligodendrogliogenesis and routes of migration. (A): Scheme representing a sagittal section of an E14.5 mouse embryo. The oligodendrogenic regions are represented in colors, depending on their origin within the neural tube. (B): In the telencephalic subventricular zone, three oligodendrogenic domains can be identified: Nkx2.1þ Gsh2þ and Emx1þ. During embryonic development, Nkx2.1þ oligodendrocyte progenitor cells are the first to appear, although the cells originated in this ventromedial domain die later in developmentdrepresented as y-). While the oligodendrocyte precursors generated in the first two domains colonize the basal and lateral telencephalon, those generated in the Emx1þ domain will myelinate the entire neocortex. (C): Oligodendrocyte precursor cells generated in the basal plate of prosomer 1 (BP P1) pursue alembicated routes of migration to colonize the alar prosencephalon, from the optic tectum (OT) to the prethalamus. (D): Almost 80% of the oligodendrocyte precursor cells generated in the spinal cord emerge from basal domains (in red), whereas merely 20% originate from dorso/alar domains (in blue). Vectors on the bottom right of panels B and C indicate the orientation of the three-D distribution of the migratory routes followed by OPCs in the telencephalon and the diencephalon, respectively (D, dorsal; L, lateral; R, retro). Other abbreviations used in this figure are GE (ganglionic eminence), EPA (entopeduncular area), ZLI (zona limitans intrathalamica), DBP (diencephalic basal plate), DD (dorsal diencephalons), P1 and P2 (prosomeres 1 and 2), M (mesencephalon), R1eR7 (rhombomeres from 1 to 7), PT (pretectum), Th (thalamus), Hy (hypothalamus). (A) Basal in red, and alar/dorsal in blue; adapted from Spassky N., Goujet-Zalc C., Parmantier E., Olivier C., Martinez S., Ivanova A., Ikenaka K., Macklin W., Cerruti I., Zalc B., Thomas JL., 1998. Multiple restricted origin of oligodendrocytes. J. Neurosci. 18 8331e8343. (B) Together with a minor contingent entering from the upper Gsh2þ domain; adapted from: Kessaris N., Fogarty M., Iannarelli P., Grist M., Wegner M., Richardson W.D., 2006. Competing waves of oligodendrocytes in the forebrain and postnatal elimination of an embryonic lineage. Nat. Neurosci. 9 (2), 173e179. (C) PTh; Adapted from Garcion E., Faissner A., Ffrench-Constant C., 2001, Knockout mice reveal a contribution of the extracellular matrix molecule tenascin-C neural precursor proliferation and migration. Development 128 2485e2496. (D) Adapted from Richardson W.D., Kessaris N., Pringle N., 2006, Oligodendrocyte wars. Nat. Rev. Neurosci. 7 (1), 11e18.

rostral axis: iIn rodents, for example, oligodendrocyte precursor cellOPCs are generated about 4 days earlier in the spinal cord than in the telencephalon (Temple and Raff, 1986; Spassky et al., 2001). As a result, the proliferation and differentiation of oligodendroglia is not synchronous: while While in birds the process of myelin formation follows a caudal-torostral chronological gradient from the lumbar spinal cord to the forebrain, in rodents, myelination follows a caudo-rostral gradient in the brain, but rostral to caudal in the spinal cord (B. Zalc, personal observation). It is noteworthy that the amount of oligodendrocyte precursor cellOPCs that are physiologically present in the adult CNS are is able to give rise to new myelin-forming oligodendrocytes that become new part of the sheaths in myelinated tracts (Young et al., 2013).

23.1.2 Oligodendrocyte precursor cells: born to migrate As mentioned earlier, oligodendrocytes are distributed nearly homogeneously throughout the adult CNS, including both the white and gray matter, even though their density is much higher in the myelinated tracts than in cortical areas (Mallon et al., 2002; Miller et al., 1992). In the adult, it is self-evident that a myelinating oligodendrocyte, attached to 20 or more

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axons, is not prone to migrate. In contrast, during development, OPCs need to migrate from the multiple, but discrete, foci from where they are generated along the neural tube to colonize the entire CNS. Although the morphology of these migrating OPCs varies across species, often they are monopolar cells, but bipolar or tripolar morphologies are also frequent, and it has been reported that these cells are more polarized in the rat than in the mouse (Milner and FfrenchConstant et al., 1994; Niehaus et al., 1999; Spassky et al., 2001; Spassky et al., 2002). The tip of the migrating OPC process resembles the axonal growth cone of developing neurons (Schmidt et al., 1997; Simpson and Armstrong, 1999; Zhang et al., 1999; Miller et al., 1997; Grabham and Goldberg, 1997), and occasionally, OPCs show very short trailing processes (a typical morphology is illustrated in Fig. 23.1). The extraordinary ability of OPCs to migrate is perfectly reflected by the fact that both in vivo and after transplantation, they are the most dispersive and migratory cells of the entire oligodendroglial lineage (Warrington et al., 1993; Noble et al., 1992).

23.2 Migratory paths followed by oligodendrocyte progenitor and precursor cells In the adult, myelinating oligodendrocytes are distributed, not only in the white matter tracts but also throughout the brain and spinal cord in the gray matter. Indeed, the axons from long projection neurons, and generally speaking all myelinated axons, are myelinated very close from the neuronal soma just after the axon initial segment, i.e., more often in the cortical gray matter (Hill et al., 2008). Therefore, OPCs have to migrate more or less long distances from their ventricular site of origin, localized on the midline of the neural tube, to reach, for example, the most distant heminode at the distal limit of the axon initial segment of neurons distributed in the most marginal layers. The first experimental evidence that OPCs have to migrate long distance was the demonstration that these cells, generated in the preoptic area (a focal region at the ventral midline of the third ventricle), invade the optic nerve from the optic chiasm (Small et al., 1987; Ono et al., 1997). In the spinal cord, OPCs, immunolabeled with the O4 mAb, have been shown to disperse from their ventricular foci of origin, both lateroventrally and ventrodorsally (Ono et al., 1995; Miller and Ono 1998). Retroviral labeling experiments have demonstrated that OPCs in the spinal cord can also migrate tangentially along the rostrocaudal axis, even moving from the level in which they were generated to another (Leber et al., 1996; Leber and Sanes 1995). In the telencephalon, homotopic homochronic quail-chick chimeras have demonstrated that OPCs emerging from the medial subpallium and the entopeduncular areas migrate tangentially to colonize the entire telencephalon (Olivier et al., 2001). Similarly, homotopic homochronic transplants of diencephalic neuroepithelium have demonstrated that OPCs originated in the parabasal domain (p1 prosomere) adopt tangential routes of migration rostrally in the three diencephalic prosomeres toward alar and dorsal mantle layers following pioneer axons (García-López and Martínez, 2010). A similar quail-chick chimera approach conducted in the hindbrain has shown that, in contrast to the diencephalon and telencephalon, in the rhombencephalon, a segmentally restricted population of OPCs developed from metameric basal foci colonizes only their rhombomere of origin (Olivier et al., 2001). The strikingly different migratory properties between rostral versus caudal brain could be due to intrinsic features of each population of OPCs or due to environmental cues. This question was addressed using heterotopic grafts. Rhombencephalic oligodendrocyte precursors adopt long tangential routes of migration when transplanted into the ventral telencephalon, and telencephalic oligodendrocytes show limited tangential migrations when grafted into the hindbrain (Olivier et al., 2001). These data strongly suggested the importance of environmental cues to control and guide the migration of OPCs during CNS development. More recently, it has been reported that migrating OPCs use vasculature surface to migrate. However, since pattern of migration OPCs does not match the disposition pattern of vessels, it has been postulated that migrating OPCs crawl along and jump between vessels. OPC migration in vivo was disrupted in mice with defective vascular architecture but was normal in mice lacking pericytes. Wnt-Cxcr4 (chemokine receptor 4) has been identified as the molecular signaling in regulation of OPCeendothelial interactions (Tsai et al., 2016).

23.3 Chemokinetic factors: the motility of oligodendrocyte precursors As for any migratory cells (including other neural cells), OPCs need chemokinetic (or motogenic) factors to move (see Chapter 12). Since the 1990s, different growth factors have been identified to act as motogenic to induce the migration of OPCs. Among these chemokinetic growth factors, the most convincing evidence concerns basic fibroblast growth factor (FGFb, also known as FGF-2) and platelet-derived growth factor A (PDGF-A). These molecules have been shown to also regulate proliferation survival and differentiation of neural cells (Calver et al., 1998; Fruttiger et al., 1999). Despite controversial studies, it seems now clear that PDGF-A and FGF-2 cooperate to promote the migration of OPCs (Armstrong et al., 1990; Dubois-Dalcq and Murray, 2000; McKinnon et al., 1993; Milner et al., 1997a,b; Rogister et al., 1999; Simpson and Armstrong, 1999). Both factors act as motogenic cues separately in a nonadditive manner (Simpson and Armstrong, 1999; Bribián et al., 2006). Regarding the motogenic function of FGF-2, it has been shown that the extracellular matrix

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glycoprotein anosmin-1 antagonizes the effect. The inhibition by anosmin-1 of the chemokinetic action of FGF-2 is most probably mediated by a competition of the two factors for FGFR1 sites (Bribián et al., 2006). At least in the forebrain subventricular zoneegenerated OPCs that colonize the corpus callosum, in vivo overexpression of anosmin-1 results in higher OPC proliferation, faster migration and differentiation, both during development and in the adult and, as a summary, hypermyelination (Murcia-Belmonte et al., 2016). FGF-2 can be partially substituted by Sonic Hedgehog, at least in a subpopulation of OPCs (Merchán et al., 2007). Although OPCs simultaneously express both PDGF receptors and FGF receptors (Redwine et al., 1997), both signaling pathways are clearly differentiated, and FGF-2 does not exert its effect via PDGF receptors (Milner et al., 1997a,b). In fact, even though cells of the oligodendrocyte lineage express FGFR1, FGFR2, and FGFR3, only FGFR1 mediates the effects of FGF-2 on OPC migration, whereas FGFR2 and FGFR3 presumably promote the proliferation and differentiation of these cells (Bansal et al., 1996). Thus, both PDGF-A and FGF-2 provide highly specific signaling pathways for the development of oligodendroglia, with subtle interactions and combinations of functions. The main evidence regarding the relevance of growth factors in the migration of OPCs arises from the in vivo studies carried out in the PDGF-A null mutant mice, in which low numbers of oligodendrocytes are observed postnatally in cerebral cortex, in brain stem, and specially in structures characterized by low number of PDGFRaþeOPCs during embryonic development, such as the optic nerves, cerebellum, and spinal cord (Fruttiger et al., 1999). In this study, myelin defects in knockouts were more prominent away from the periventricular germinal zones of the neural tube (e.g., the optic nerve), which suggests that PDGF-A might act as a long-range motogenic factor for OPCs. Interestingly, no significant differences in neuronal development have been described in PDGF-A knockouts (Fruttiger et al., 1999). Growth factors are known to interact with molecules from the extracellular matrix (Jones and Jones, 2000), and fibronectin or merosin (but not tenascin-C) have been shown to promote the migration of OPCs, either by themselves or in combination with motogenic PDGF-A (Frost et al., 1996; Garcion et al., 2001). The relevance of interactions between extracellular matrix molecules (such as PSA-NCAM) and growth factors (PDGF-A, FGF-2, BDNF, CNTF) on OPC migration is complex and remains to be fully understood (Muller et al., 2000; Vutskits et al., 2001, 2003; Zhang et al., 2004; Bribián et al., 2006, 2008). It is generally accepted that, as happens with cell differentiation of oligodendroglia lineage, the motility of OPCs results from the combinatorial action of molecules from the extracellular matrix, secreted growth factors, mitogens, hormones, and neurotransmitters (Decker et al., 2000; Dubois-Dalcq and Murray, 2000; Rogister et al., 1999). The role of other growth factors on the motility of OPCs has been less explored. The intraventricular administration of epidermal growth factor (EGF) induces the proliferation of SVZ neural progenitors and the subsequent migration of postmitotic cells away from the ventricular surface concerns mostly astrocytes and little (if at all) OPCs (Fricker-Gates et al., 2000). Tissue-type plasminogen activator has been very recently demonstrated to be chemokinetic for OPCs through its EGF-like domain, both during development and in (re)myelination (Leonetti et al., 2017). Finally, hepatocyte growth factor (HGF), an established motogenic factor for neuronal precursors (Powell et al., 2001), induces a significant 2.5-fold increase in the migration rate of OPCs in vitro, as well as correlative changes in the cytoskeleton of these cells (Yan and Rivkees, 2002). More complete studies on the effect of HGF on OPC motility are certainly needed. Endothelin-1 (but not endothelin-2 or endothelin-3) has been shown to arrest postnatal oligodendroglial cells of the cerebral cortex in a less mature and promigratory phenotypic stage (OPC), promoting the migration of these cells and also acting as a chemoattractant (Gadea et al., 2009). Some data suggest that cytokines such as interleukin-1a (IL-1a) or tumor necrosis factor-a (TNF-a), which are secreted by astrocytes, modulate the FGF-2-induced OPC motility. The combination of IL-1a and FGF-2 was reported to be promigratory, an effect that was neutralized by the addition of transforming growth factor-alpha (TGF-a) to this cocktail (Fok-Seang et al., 1998). Of note, the effects of these cytokines are not correlated with the expression of the contactmediated molecules reviewed in the following.

23.4 Adhesion and chemotactic mechanisms: how the movement of oligodendrocyte precursors is guided? 23.4.1 Adhesion and surface molecules Before reaching the axonal segment they will myelinate, OPCs migrate a more or less long distance starting from the ventricular zone where they are generated. During this migration, whether tangential or radial, OPCs are guided by secreted factors as well as by adhesion molecules expressed by the astrocytes populating the territories they are traveling in and, of course, by axons forming the fiber tracts along which they migrate before stopping and forming the myelin sheath. During development, electrical activity along axons and probably expression of surface axonal molecules are crucial in the choice

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and decision-making of when and where myelination is to occur (Lubetzki et al., 1993; Demerens et al., 1996; Charles et al., 2000; Doyle and Colman, 1993; Pfeiffer et al., 1993). Contact with a preformed axonal tract is a major factor controlling this migratory process, and different adhesion molecules from the extracellular matrix or expressed at the surface of astrocytes and axons have been shown to control the movements of OPCs. Several components of the extracellular matrix take part in the migration of OPCs. Laminin, fibronectin, merosin, and anosmin-1 promote the adhesion and directly or indirectly the migration of OPCs (Frost et al., 1996; Bribián et al., 2008; Murcia-Belmonte et al., 2016), either alone or in combination with growth factors such as PDGF-A or FGF-2 (see earlier). The relative strength of adhesion of OPCs to components of the extracellular matrix correlates with the speed reached by these cells when in motion. For instance, it has been shown in culture that the velocity of OPCs is higher on laminin than on poly-L-lysine substrate (Niehaus et al., 1999; Bribián et al., 2008). N-Cadherins expressed at the cell surface of both oligodendrocytes and astrocytes increase the adhesion of OPCs. Blocking N-cadherin with a specific antiserum reduces the adhesion of oligodendroglia, resulting in an enhanced dispersion of early-postnatal OPCs cultured over astrocytic monolayers (Schnadelbach et al., 2000). Under these experimental conditions, the speed of migration of OPCs doubles when they are grown on merosin (11.2 mm/h) as compared with migration on N-cadherin (5.6 mm/h; Schnadelbach et al., 2000). Tenascin-C has been shown to selectively inhibit the migration of OPCs that respond to fibronectin, but not to merosin (Frost et al., 1996). Tenascin-C is considered as a general inhibitor of the migration of OPCs in both an adhesiondependent and adhesion-independent manner (Kiernan et al., 1996). For example, it has been shown that it is the tenascin-C expressed by astrocytes located at the most distal portion of the optic nerve (the lamina cribrosa) that prevents optic nerve OPCs from invading the retina (Morcos and Chan-Ling, 2000). In the rabbit, the optic nerve is not demarcated by a tenascin-C boundary at the entry of the retina, and retinal ganglion cell axons are myelinated in their initial retinal portion (Bartsch et al., 1994). It has been postulated that, due to the absence of a tenascin-C plug at the level of the retinal end of the optic nerve, OPCs in the rabbit can freely cross the lamina cribrosa and therefore myelinate the retina (Berliner, 1931; Schnitzer, 1985). However, it is of note that no abnormalities of myelination or OPCs distribution have been reported in null-mutant mice for tenascin-C. It is therefore likely that local concentrations of tenascin-C are not the sole stop cue for OPCs, as suggested by the repulsive effect reported for netrin-1 in the optic nerve (see the following text; Garcion et al., 2001; Kiernan et al., 1999; Saga et al., 1992; Spassky et al., 2002). Similarly, due to its high adhesive properties, anosmin-1 has also been suggested as a putative stop signal for OPCs, allowing their final differentiation into myelinforming oligodendrocytes (Bribián et al., 2008; Clemente et al., 2008; Murcia-Belmonte et al., 2016). During late embryonic and early-postnatal stages, both PSA-NCAM and polysialic acid interfere with the migration of OPCs (Wang et al., 1994); for example, PSA-NCAM has been shown to be essential for PDGF-A to attract these migrating cells (Zhang et al., 2004). In support of the relevance of PSA-NCAM in the migration of OPCs, it has been reported that, in response to a demyelinating insult in the olfactory bulb, PSA-NCAMþ cells are detected in the rostral migratory stream and that these cells differentiate into oligodendrocytes and astrocytes at the site of the lesion and also in other structures of the brain, including the corpus callosum (Nait-Oumesmar et al., 1999). Interestingly, and in contrast to neurons, during normal pre- and postnatal development, the forebrain SVZ does not contribute to populate the olfactory bulb with oligodendrocytes (Hu and Rutishauser, 1996; Spassky et al., 2001). Transmembrane semaphorins and some of their receptors have been identified by RT-PCR in different cells of the oligodendroglial lineage in postnatal rats and at a lower level in adult animals, except for Sema4C and 4D (Cohen et al., 2003). In vivo, Sema4D controls the final number of oligodendrocytes, but its exact role in the migration of OPCs remains to be elucidated (Taniguchi et al., 2009), whereas Sema4F is known to stop this migration and would be involved in oligodendroglial differentiation (Armendáriz et al., 2012). It is noteworthy that different components of oligodendroglial cells have been shown to be involved in the migration of OPCs. This is the case for the surface molecule chondroitin sulfate proteoglycan NG2 (also known as AN2), expressed by proliferating and migratory OPCs and downregulated when the cells begin their final differentiation. Immunoblocking of NG2 results in a drastically reduced migration of embryonic and early-postnatal OPCs to approximately one half of their total number estimated under control conditions, without affecting their proliferation. It has been postulated that NG2 may interact with cytoskeleton or integrins, but the exact role of NG2 protein in OPC migration still remains unclear (Niehaus et al., 1999). Blocking antiserum directed against OSP/claudin-11 (also known as OAP-1), a major component of tight junctions maintaining the successive paranodal loops tightly attached to each other, inhibits the migration of neonatal OPCs from rodents in vitro (Tiwari-Woodroof et al., 2001). A similar effect is observed if a1-integrin is blocked (an effect selectively observed with a-v-1-integrin, but not other isoforms), which suggests the direct (or indirect) involvement of integrins in the migration of OPCs, at least in vitro (Tiwari-Woodroof et al., 2001; Milward et al., 2000). However, the complete blockade of integrins does not completely prevent the migration of OPCs (Milner et al., 1996; Niehaus et al., 1999), suggesting the

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implication of other major actors. OPCs express glutamate receptors (a-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid [AMPA] and kainate). Glutamate acting on AMPA receptor was shown to stimulate a-v integrin-mediated migration of this cell type by increasing both the rate of cell movement and the frequency of calcium transients. This increase in OPC migration is mediated by a complex formed by a-v integrin and PLP, the most abundant myelin protein (Gudz et al., 2006). The integrity of G protein and the extracellular ligand for G proteinecoupled receptors sphingosine-1-phosphate 5(S1P5), which in the CNS is exclusively expressed by OPCs and mature oligodendrocytes, is needed for the AMPA-stimulated OPC migration, but not for the basal migratory process (Gudz et al., 2006; Novgorodov et al., 2007). Among the most relevant contact-mediated molecules involved in the migration of neuronal precursors during development are ephrins and their Eph receptors (Marín and Rubenstein, 2001, 2003; see also Chapters 15, 18 and 20). While the membrane-anchored ephrins B2 and B3, acting via EphB receptors, reduce the adhesion and migratory properties from embryonic diencephalic OPCs (including those colonizing the optic nerves), ephrin A5 does not have an effect (Prestoz et al., 2004). In addition, they show that coated EphB2-Fc receptors, which are specific for ephrinB2/B3 ligands, induce dramatic changes in the contact and migratory properties of OPCs, indicating that axonal EphB receptors activate ephrinB signaling in the migrating cells. Based on these findings, it was proposed that OPCs are characterized by an ephrin code and that Epheephrin interactions between axons and OPCs control their distribution in the optic axonal tracts, by modulating the progress and arrest of their migration (Prestoz et al., 2004). The null mutation of EphA4 does not result in any relevant change for oligodendrocytic development, at least in the murine optic nerve (Petros et al., 2006), which seems to confirm that only EphB receptors (and not EphAs) would be involved in the development of oligodendroglia. In the forebrain subventricular zone, some glial precursors undergo radial migration in the first postnatal week (Kakita and Goldman, 1999; Mallon et al., 2002). This observation supports the hypothesis that radial glia would be involved in the migration not only of neurons but also of OPCs, which was proposed in the spinal cord during development, where migrating OPCs are observed in close association with radial glia (Hirano and Goldman, 1988). Other authors propose that radial glia may facilitate the final dispersion of OPCs once in the white matter of the spinal cord, with which they are closely aligned, but this dispersion does not intervene in the early ventral-to-dorsal spread of OPCs because the observed spreading movements are not consistent with the distribution pattern of the radial glia (Noll and Miller, 1993; Ono et al., 1995; Tsai et al., 2003). Moreover, as it is the case for some populations of neurons (for a review, see Marín and Rubenstein, 2003; see also Chapters 19 and 20), OPCs have been shown to combine tangential and radial migration. OPCs can switch from one pattern to the other, at least in some regions of the CNS. Although the molecular basis underlying this OPC/radial glia interaction during their migration remains to be clarified, the described effects of netrin-1 on OPC migration (see the following text) suggest a coordinate role of molecules expressed at the surface of radial glia and of chemotactic factors for the migration and dispersion of OPCs (Tsai et al., 2006).

23.4.2 Secreted factors Since the beginning of the century, a bulk of experimental results support the idea that the migration of OPCs depends not only on contact-mediated mechanisms but also on secreted factors, in particular chemotropic cues such as those involved in the guidance of axonal growth cones and the migration of neuronal precursors. In this respect, it is relevant that the transection of rat optic nerve affects neither proliferation nor migration nor programmed death of OPCs (Ueda et al., 1999). Along the same line, when optic nerves are maintained as explant in culture and provided that the culture medium contains FGF-2, there is a massive migration out of the explant and almost all the cells leaving the explant are OPCs (Sugimoto et al., 2001; Spassky et al., 2002). These data show that even though OPCs often migrate along fiber tracts, axonal contact is not compulsory for these cells to migrate. Furthermore, the set of data showing that OPCs are able to move in the absence of axons strongly suggest that these cells may respond to other signals. We have already described the importance of soluble motogenic (or chemokinetic) factors, such as growth factors PDGF-A and FGF-2. In the next section, we review the data showing the importance of secreted molecules to control the guidance of the navigation in the parenchyma of OPCs. Although their main role in the migration of OPCs is to induce the motility of the cells as motogenic cues (see earlier), PDGF-A and FGF-2 have also been shown to act as chemoattractants (Zhang et al., 2004; Bribián et al., 2006; MurciaBelmonte et al., 2014. This effect is specific, since anosmin-1, which antagonizes FGF-2 motogenesis via FGFR1, does not play any chemotropic effect on OPCs (Bribián et al., 2006; Murcia-Belmonte et al., 2016). Vascular endothelial growth factor A (VEGF-A) acts also as a chemoattractant on ill-defined newborn/early-postnatal neural progenitors and, among them, OPCs; this chemoattractant effect of VEGF-A is FGF-2 dependent and mediated by interaction with VEGF-R2, but not VEGF-R1, the other VEGF-A receptors (Zhang et al., 2003). A second group of secreted molecules involved in the migration of OPCs are the chemotropic (or chemotactic) molecules, a heterogeneous family of molecules known to play relevant roles in the guidance of axonal growth cones and

522 PART | II Migration

the migration of neuroblasts during development (for reviews on this topic, see de Castro, 2003; Marín and Rubenstein, 2001; see also Chapters 3 and 13). Netrins and secreted class III semaphorins are the most studied to date. Netrin-1 can act either as a chemorepellent or chemoattractant for OPCs, depending on the type of receptors that these cells express. In the spinal cord, netrin-1 is produced by the floor plate and the ventral midline cells. OPCs emerging from the ventral ventricular zone in the spinal cord express two netrin-1 receptors: deleted in colorectal cancer (DCC) and Unc5A. It has been shown that, when receptors from the Unc5 family are associated with DCC, netrin-1 acts as a repulsive cue (for a review, see de Castro, 2003). This is indeed what has been observed for the ventral OPCs in the spinal cord, where netrin-1 induces the initial dispersion of these cells out of the ventricular layer where they are produced and their extensive radial migration and expansion once in the white matter (Tsai et al., 2003, 2006) (Fig. 23.3). These data have been confirmed in vivo using the netrin-1 and DCC null mutants, which show an impaired migration of OPCs from their ventral origin to their final destinations in terms of both, the number of cells and their trajectories, but not in the migration rates (Jarjour et al., 2003; Tsai et al., 2009). In the optic nerve, netrin-1 is expressed by astrocytes mostly at the optic papilla (retinal end) and in the temporal quadrant of the nerve. In the mouse, OPCs start to colonize the optic nerve at E14.5, and by E17.5eE18.5, they are distributed all along the nerve including the retinal segment. Interestingly, when OPCs enter the optic nerve, they express DCC only. Later in development, some OPCs, and in majority those present at the retinal end close to the lamina cribrosa, express both DCC and Unc5A. The number of cells expressing both DCC and unc5A increases significantly between E18.5 and P5. This suggests that the effect of netrin-1 might be modulated by a developmentally regulated change in the expression of its receptors DCC and Unc5A. DCC receptor alone might be involved mainly at the time when OPCs enter the nerve, whereas, later in the course of development, DCC in cooperation with Unc5A might participate in the signaling to stop migration, therefore avoiding all OPCs to accumulate at the retinal end. This developmentally regulated change in the type of receptors expressed by OPCs explains the dual attractant and repellent effect of netrin-1 on OPCs migration (Spassky et al., 2002; Sugimoto et al., 2001). This situation is reminiscent of that described for migrating neurons and navigating axon growth cones (for a review, see de Castro, 2003; see also Chapters 3, 19 and 20). Of note, in multiple sclerosis (MS) lesions, astrocytes upregulate netrin-1 expression early during demyelination, and netrin-1 receptors are expressed by OPCs; therefore, netrin-1 expression within demyelinating MS plaques blocks OPC recruitment, which with repeated demyelinating episodes contributes to permanent remyelination failure (Tepavcevic et al., 2014). The semaphorins comprise a large family of secreted and transmembrane proteins. Secreted semaphorins 3A and 3F act as axonal guidance cues in the developing nervous system (Chisholm and Tessier-Lavigne, 1999), and they have been shown to act as chemotactic factors for OPCs during development (Sugimoto et al., 2001; Spassky et al., 2002; Tsai and Miller, 2002). OPCs colonizing the optic nerve have been shown to express neuropilin-1 and neuropilin-2, the receptors for semaphorins 3A and 3F, respectively. Semaphorin 3A, which has been shown to be repellent for OPCs, is expressed by mesenchymal cells forming a ring around the optic nerve. This pattern of semaphorin 3A expression delineates a clear boundary between the outside and the inside of the nerve, which could force the OPCs to stay within the nerve and migrate along its length. Semaphorin 3F is expressed in the retina, notably by retinal ganglion cells, which axons form the optic nerve. It has been shown that chemosecreted semaphorin can be transported along the axons (de Wit et al., 2006), and it can therefore be assumed that semaphorin 3F is also expressed along retinal ganglion cell axons. Semaphorin 3F has been shown to attract migrating OPCs (Spassky et al., 2002). In the brain and spinal cord of patients with MS, semaphorins 3A and 3F have been shown to be overexpressed, and it has been suggested that their increased expression in MS tissue may favor the recruitment of OPCs toward the demyelinated area if attractive guidance cues such as semaphorin 3F are predominantly expressed and thus favor remyelination (Piaton et al., 2011). In contrast, OPC migration may be impaired if inhibitory guidance cues such as Sema3A are predominant, hence inhibiting remyelination (Williams et al., 2007). Other strong chemoattractant for early plp-dm20þ OPCs is neuregulin-1 via its ErbB4 receptor, but this cue is not physiologically relevant for the postnatally generated PDGFRaþ population of cells (Ortega et al., 2012a). Chemokines were initially described as potent chemoattractants for leukocytes (Baggiolini, 1998); they have been extensively reported to arrest migration and immobilize other cells, too. The chemokine CXCL1 and its receptor CXCR2 have been shown to play an important role for the development and maintenance of the oligodendrocyte lineage in the vertebrate CNS. CXCL1 has been proposed as a clear stop signal for migrating OPCs during development of the spinal cord (Tsai et al., 2002). CXCL1 is expressed by astrocytes in the white matter at the time when OPCs colonize this structure (Miller et al., 1997; Robinson et al., 1998). CXCL1 inhibits the migration of neonatal OPCs and enhances their interaction with substrates. CXCL1 inhibits the migration of neonatal OPCs by antagonizing the PDGF-A-induced chemotactic effect on these cells (Tsai et al., 2002). This effect is rapid and reversible and is mediated via the chemokine receptor CXCR2, which is expressed by 85% of OPCs in the spinal cord (Tsai et al., 2002). In the CXCR2/ adult mice, the density of oligodendrocytes is decreased in the cortex and anterior commissure but increased in the corpus callosum and spinal cord. CXCR2/ mice exhibit a significant decrease in spinal cord white matter area, reduced thickness of

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(A)

Sema-3F Netrin-1 Neuropilin-1+/Neuropilin-2+/Unc5A+/DCC+

Sema-3A

(B)

FGFR1 PDGFR-D

Anosmin-1 FGF2 PDGF-A

(C)

PDGF-A CXCL-1 Netrin-1 PDGFR-D+/DCC+/ Unc5A+/CXCR2+

FIGURE 23.3 Secreted cues guide the migration of oligodendrocyte precursors. (A): Oligodendrocyte precursor cells generated in the preoptic area invade the optic chiasm to colonize the optic nerve and progress toward the eye (on the left). These cells express a battery of receptors (neuropilin-1, neuropilin-2, DCC, and Unc5Adthe latter appears only late in development), which allow them to be guided by chemoattractants such as Sema 3F (produced in the retina and probably transported along the axons of retinal ganglion cells) and netrin-1 (expressed in the temporal quadrant of the nerve and in the papilla) and the chemorepellent Sema 3A (expressed by the mesenchyme surrounding the optic nerve). In late development, migrating oligodendrocyte precursors start to express other netrin receptor, Unc5A, in addition to DCC, which turns netrin-1 signal into repellent, and impede these cells to invade the retina (B): While PDGF-A and FGF-2 display motogenic effects (via PDGFR-a and FGFR1, respectively) on migrating oligodendrocyte precursor cells purified from the optic nerve, anosmin-1 interferes with FGFR1 to finally reduce motogenic activity. (C): The oligodendrocyte precursor cells originated in the ventrolateral domain of the spinal cord (see Fig. 23.2D) disperse toward dorsal regions of the structure due to the combined effect of a motogenic cue, such as PDGF-A (via PDGFR-a) and the chemorepulsion due to netrin-1 (given that migrating oligodendrocyte precursor cells coexpress DCC and Unc5A) and CXCL1 (via CXCR2). Altogether, the combination of these diverse factors results in the colonization of the entire spinal cord, although a higher density of oligodendroglial cells is located in the periphery (white matters tracts) than in the center. (A) Adapted from: Spassky N., Goujet-Zalc C., Parmantier E., Olivier C., Martinez S., Ivanova A., Ikenaka K., Macklin W., Cerruti I., Zalc B., Thomas J.L., 1998. Multiple restricted origin of oligodendrocytes. J. Neurosci. 18, 8331e8343 and de Castro F., Bribián A., 2005, The molecular orchestra of the migration of oligodendrocyte precursors during development. Brain Res. Rev. 49 (2), 227e241. (B) Adapted from Bribián A., Barallobre MJ., Soussi-Yanicostas N., de Castro F., 2006. Anosmin-1 modulates the FGF-2-dependent migration of oligodendrocyte precursors in the developing optic nerve. Mol. Cell. Neurosci. 33 (1), 2e14. (C) Adapted from de Castro F., Bribián A., 2005. The molecular orchestra of the migration of oligodendrocyte precursors during development. Brain Res. Rev. 49 (2), 227e241.

myelin sheaths, and a slowing in the rate of central conduction of spinally elicited evoked potentials without significant changes in axonal caliber or number (Padovani-Claudio et al., 2006). These authors point that the relevance of putative compensatory effects seems more likely to occur in other developmental phases than during the migration of OPCs, although they admit that the migration of these cells must be regulated in a combinatorial manner, as occurs for axonal guidance and neuronal migration (Lu et al., 2001; Ma et al., 1998; Winberg et al., 1998). They suggest a model in which

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the widespread uniform expression of PDGF-A, combined with the transient presence of CXCL1, regulates the final spatial distribution of oligodendrocytes in the developing CNS (Tsai et al., 2002; de Castro and Bribián, 2005). In addition to the role of CXCL1/CXCR2, it has been postulated that CXCR4 signaling (mainly activated by CXCL12) on OPCs regulates their survival and attracts their migration toward dorsal regions of the spinal cord during embryonic and postnatal development (Dziembowska et al., 2005). The morphogen Sonic Hedgehog has also been shown to be a potent attractant of OPCs in the early stages of their colonization of the optic nerve, both in mammals and in chick. This effect is not entirely due to the canonical Sonic Hedgehog receptor patched-1 (Merchán et al., 2007), and Lrp-2/megalin has been shown to be the noncanonical receptor involved in this, contributing to homeostasis of Sonic Hedgehog concentration during development (Ortega et al., 2012b). Sonic Hedgehog combines this chemoattraction with a proliferative effect on OPCs, which would explain why both cease by E18.5 in the mouse and do not result in an excessive number of these cells.

23.5 Concluding remarks During development, oligodendrogenesis is delayed compared with neurogenesis. Migration of neuronal precursors, axonal growth, and axon pathfinding are tightly controlled by series of secreted or adhesion molecules. At the time when OPCs start to migrate out of the germinative neuroepithelium, they encounter a nervous parenchyme impregnated with gradients of these secreted chemotactic factors as well as axons expressing contact surface molecules that have directed their migration toward their final position or the axonal pathfinding toward their target. OPCs emerging from different domains express different panels of receptors interacting with these guidance molecules. As a consequence, migration of these cells does not occur at random but appears to be rather predetermined: OPCs emerging from a given domain will colonize the territory corresponding to the factor/s they are responding to. This code of migration, however, is not absolutely strict and univocal, probably because these cells express a relatively large panel of receptors (or ligands) guiding their migration. This is unambiguously illustrated by the heterotopic graft experiments showing the complete change in the mode of migration depending on the territory where the graft is positioned (Olivier et al., 2001). Similarly, ablation of one oligodendrogenic domain is followed by the colonization of this territory by myelin-forming cells originating from a neighboring domain (Kessaris et al., 2006). Along the same line is the observation that, under pathologic conditions, OPCs recruited to colonize a lesion to be remyelinated express both neuropilin-1 and neuropilin-2 and could therefore being repulsed by sema3A or attracted by semaphorin 3F, suggesting that remyelination will be possible only if OPCs respond to the right balance of concentration of these guidance molecules (Williams et al., 2007; Wang et al., 2008; Piaton et al., 2011; de Castro et al., 2013; Lubetzki, C. unpublished data, personal communication). It is noteworthy that the formal demonstration of the influence of guidance cues on myelin repair would represent a fascinating therapeutic target in demyelinating diseases such as MS.

Acknowledgments We are indebted with Dr. Diego García-González for his fundamental contribution in the preparation of figures, Dr. F. Javier Arenzana and Dr. Andrés Mínguez for the photographs included in Fig. 23.1, and Dr. Eva Medina-Rodríguez for her help with bibliography: All of them helped us in our chapter for the first edition of this book. Our groups are supported by grants from the Spanish Ministerio de Ciencia, Innovación y Universidades (SAF2016-77575-R and RD16/0015/0019dpartially funded by F.E.D.E.R./European Union “Una forma de hacer Europa”), Comunidad de Madrid (IND2018 BMD-9751), and Fundación Ramón Areces (Spain) to FdC and ANR OLGA and Investissement d’Avenir ANR-11-INBS-0011dNeurATRIS to BZ.

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Tsai, H.H., Macklin, W.B., Miller, R.H., 2009. Distinct modes of migration position oligodendrocyte precursors for localized cell division in the developing spinal cord. J. Neurosci. Res. 87 (15), 3320e3330. Tsai, H.H., Tessier-Lavigne, M., Miller, R.H., 2003. Netrin 1 mediates spinal cord oligodendrocyte precursor dispersal. Development 130, 2095e2105. Ueda, H., Levine, J.M., Miller, R.H., Trapp, B.D., 1999. Rat optic nerve oligodendrocytes develop in the absence of viable retinal ganglion cell axons. J. Cell Biol. 146, 1365e1374. Vallstedt, A., Klos, J.M., Ericson, J., 2005. Multiple dorsoventral origins of oligodendrocyte generation in the spinal cord and hindbrain. Neuron 45, 55e67. Vutskits, L., Djebbara-Hannas, Z., Zhang, H., Paccaud, J.P., Durbec, P., Rougon, G., Muller, D., Kiss, J.Z., 2001. PSA-NCAM modulates BDNFdependent survival and differentiation of cortical neurons. Eur. J. Neurosci. 13, 1391e1402. Vutskits, L., Gascon, E., Kiss, J.Z., 2003. Removal of PSA from NCAM affects the survival of magnocellular vasopressin- and oxytocin-producing neurons in organotypic cultures of the paraventricular nucleus. Eur. J. Neurosci. 17, 2119e2126. Wang, C., Rougon, G., Kiss, J.Z., 1994. Requirement of polysialic acid for the migration of the O-2A glial progenitor cell from neurohypophyseal explants. J. Neurosci. 14, 4446e4457. Wang, Y., Imitola, J., Rasmussen, S., O’Connor, K.C., Khoury, S.J., 2008. Paradoxical dysregulation of the neural stem cell pathway sonic hedgehog-Gli1 in autoimmune encephalomyelitis and multiple sclerosis. Ann. Neurol. 64 (4), 417e427. Warf, B.C., Fok-Seang, J., Miller, R.H., 1991. Evidence for the ventral origin of oligodendrocyte precursors in the rat spinal cord. J. Neurosci. 11 (8), 2477e2488. Warrington, A.E., Barbarese, E., Pfeiffer, S.E., 1993. Differential myelinogenic capacity of specific developmental stages of the oligodendrocyte lineage upon transplantation into hypomyelinating hosts. J. Neurosci. Res. 34, 1e13. Williams, A., Piaton, G., Aigrot, M.S., Belhadi, A., Théaudin, M., Petermann, F., Thomas, J.L., Zalc, B., Lubetzki, C., 2007. Semaphorin 3A and 3F: key players in myelin repair in multiple sclerosis? Brain 130, 2554e2565. Winberg, M.L., Noordermeer, J.N., Tamagnone, L., Comoglio, P.M., Spriggs, M.K., Tessier-Lavigne, M., Goodman, C.S., 1998. Plexin A is a neuronal semaphorin receptor that controls axon guidance. Cell 95, 903e916. Yan, H., Rivkees, S.A., 2002. Hepatocyte growth factor stimulates the proliferation and migration of oligodendrocyte precursor cells. J. Neurosci. Res. 69, 597e606. Young, K.M., Psachoulia, K., Tripathi, R.B., Dunn, S.J., Cossell, L., Attwell, D., Tohyama, K., Richardson, W.D., 2013. Oligodendrocyte dynamics in the healthy adult CNS: evidence for myelin remodeling. Neuron 77 (5), 873e885. https://doi.org/10.1016/j.neuron.2013.01.006. Zalc, B., Colman, D.R., 2000. Origins of vertebrate success. Science 288 (5464), 271e272. Zalc, B., Goujet, D., Colman, D., 2008. The origin of the myelination program in vertebrates. Curr. Biol. 18 (12), R511eR512. Zhang, H., Vutskits, L., Calaora, V., Durbec, P., Kiss, J.Z., 2004. A role for the polysialic acid-neural cell adhesion molecule in PDGF-induced chemotaxis of oligodendrocyte precursor cells. J. Cell Sci. 117, 93e103. Zhang, H., Vutskits, L., Pepper, M.S., Kiss, J.Z., 2003. VEGF is chemoattractant for FGF-2-stimulated neural progenitors. J. Cell Biol. 22, 1375e1384. Zhang, S.C., Ge, B., Duncan, I.D., 1999. Adult brain retains the potential to generate oligodendroglial progenitors with extensive myelination capacity. Proc. Nat. Acad. Sci. U. S. A. 96, 4089e4094. Zhou, Q., Anderson, D.J., 2002. The bHLH transcription factors OLIG2 and OLIG1 couple neuronal and glial subtype specification. Cell 109 (1), 61e73.

Chapter 24

Coordination of different modes of neuronal migration and functional organization of the cerebral cortex Holden Higginbotham Department of Biology, Brigham Young University, Rexburg, ID, United States

Chapter outline 24.1. Introduction 531 24.1.1. Arealization of the cortex 531 24.1.2. Cortical columns constitute cortical areas 532 24.1.3. Minicolumns constitute columns 532 24.2. Migration of related projection neurons into the same minicolumn 533 24.2.1. Early lack of evidence that sister projection neurons migrate into the same minicolumn 534 24.2.2. Sister projection neurons migrate into the same minicolumn and intersynapse 535 24.3. Integration of projection neurons into cortical minicolumns 535 24.3.1. Migratory scaffolds restrict tangential movement of projection neurons 535 24.3.2. Molecular signaling limits tangential movement of projection neurons during multipolar stage 537

24.4. Integration of interneurons into cortical columns 24.4.1. Interneuron subtypes areally distribute via tangential migration 24.4.2. Do sister interneurons migrate into the same minicolumn? 24.4.3. Sister interneurons preferentially intersynapse 24.4.4. Regulating the timing of the shift from tangential to radial migration 24.4.5. Projection neurons attract migrating interneurons into cortical plate 24.4.6. Radial glial cells trigger a shift in migration mode 24.5. Genetic and cellular mechanisms controlling shifts in migratory modes 24.6. Conclusion List of abbreviations References

538 538 540 540 541 542 543 543 544 545 545

24.1 Introduction 24.1.1 Arealization of the cortex During mammalian neocortical development, regions along the tangential axes (i.e., the rostrocaudal and mediolateral axes) of the neocortex differentiate into distinct functional cortical areas (e.g., somatosensory, visual, motor, memory, association, etc.) in a process called arealization. The boundaries of these areas, identified histologically by Brodmann (Zilles and Amunts, 2010), are initially delineated by complementary and opposing expression patterns of morphogens secreted from signaling centers, which trigger the expression of distinct transcription factors in the neural progenitor cells (NPCs) of the primordial cortical areas (Alfano and Studer, 2013; Grove and Fukuchi-Shimogori, 2003; O’Leary and Sahara, 2008). The transcription factors expressed in the NPCs are specific to the cortical area for which the NPCs supply daughter neurons (O’Leary and Sahara, 2008). The NPCs’ molecular programming drives three events in its daughter neurons that contribute to the unique features and functions of each cortical area (and other noncortical areas (Hevner et al., 2006)): neurogenesis, which includes producing the right number and proportion of neuronal subtypes (Dwyer et al., 2016; Farkas and Huttner, 2008; Gao et al., 2014; Govindan and Jabaudon, 2017; Joseph and Hermanson, 2010); migration and placement of daughter neurons into the four or six layers of the cortex (Kwan et al., 2012; Mihalas and Hevner, 2017;

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Nobrega-Pereira and Marin, 2009); and differentiation of neurons, including the elaboration of dendritic and axonal projections (which can be local, interhemispheric, or extracortical) (Greig et al., 2013; Leone et al., 2008; Mukhtar and Taylor, 2018). The types and numbers of neurons produced, their placement, and the connection patterns they form are unique to each area and thus give each area its unique cytoarchitectonic features first observed by Brodmann (Zilles and Amunts, 2010) but, more importantly, are crucial for the unique functions each area subserves (Rash and Grove, 2006). Disruptions in these three processes (neurogenesis, migration, differentiation) prevent the normal functioning of cortical areas and also underlie many neurocognitive disorders such as autism and mental retardation (Casanova and Trippe, 2009; Diaz and Gleeson, 2009).

24.1.2 Cortical columns constitute cortical areas The basic functional module of each cortical area is thought to be the cortical column (Maruoka et al., 2017; Mountcastle, 1997), although this has been debated (Horton and Adams, 2005). Columns form when excitatory projection neurons (PNs) and inhibitory interneurons (INs) migrate to the same areal region and radially (vertically) distribute into four or six horizontal layers of the allocortex (which includes the olfactory cortex and hippocampus) or neocortex (which includes the sensory, motor, and cognition areas), respectively. The neurons then preferentially connect with each other radially across the layers in the column, and all the cells within the column respond concurrently to stimulation of the same peripheral receptive field using the same touch modality (Mountcastle, 1997), in the case of somatosensory cortex, or respond to the same orientation of a light stimulus (Hubel and Wiesel, 1962; Kondo et al., 2016; Maruoka et al., 2017), in the case of the visual cortex. Columns in motor areas would be expected to control activity in related muscle groups or to control similar patterns of movement. This means that neurons within a column are thought to operate as a functional unit. Evidence suggests that there exists relatively independent subnetworks of excitatory PN microcircuits within columns (Yoshimura et al., 2005) and that, within these micronetworks, only certain types of INs participate (Yoshimura and Callaway, 2005), suggesting that the placement of PNs and specific kinds of INs within these columns is important for the formation of the functional circuitry.

24.1.3 Minicolumns constitute columns Anatomically, columns repeat iteratively across the surface of multiple cortical areas (Maruoka et al., 2017) and, presumably, neurons in a column are of the same type and proportion, are placed in similar relative locations, and preferentially synapse with each other to form their own microcircuit (Bureau et al., 2004). Thus, a given areal region contains multiple adjacent and morphologically similar columns with the same basic internal circuitry and external connectivity patterns (e.g., ocular dominance columns, barrel columns) (Jones and Rakic, 2010; Sansom and Livesey, 2009). Each column is further subdivided into minicolumns (or microcolumns) that contain interconnected PNs and INs, and the PNs in the column are thought to arise from the same NPC (Rakic, 1988; Sansom and Livesey, 2009) (Fig. 24.1). It is currently debated whether INs in a given minicolumn arise from the same NPC, as will be reviewed later. Within each minicolumn, the microcircuitry seems to follow a canonical organization, with afferents arriving at layer IV, from which information flows to layers II and III, and then to deeper layers V and VI, and from there to the other hemisphere or subcortical structures (Douglas and Martin, 2004) (Fig. 24.1). Across areas, the fine details of the microcircuitry can vary (Kätzel et al., 2011), but it seems the general rule that the transcription factor molecular logic of NPCs in a certain area dictates the specific arrangement of their daughter neurons in the minicolumn and supports the unique functional characteristics of each minicolumn, column, and cortical area (Ohki et al., 2006). For example, NPCs that express the transcription factor Ctip1 produce minicolumns in sensory areas that have a circuitry that supports sensory input and not motor output (Greig et al., 2016). Thus, cortical arealization arises as adjacent neural NPCs sharing similar molecular programs elaborate minicolumns with the same laminar arrangement, cell composition, and connectivity pattern (Buxhoeveden and Casanova, 2002). This chapter will focus on how the process of neuronal migration contributes to the arealization of the cortex, focusing specifically on the elaboration of cortical minicolumns. This chapter begins with a discussion of how PNs’ tangential movement is limited so that progeny of single NPCs is confined within a single minicolumn. Then, our current understanding of how migrating INs become distributed within specific cortical areas via tangential migration and then integrate within minicolumns via a shift to radial migration is summarized. In the final section, what intracellular mechanisms might control the shift between the two modes of migration, i.e., tangential and radial in both PNs and INs, is discussed, and it has been debated that functional minicolumn formation depends on precise control of the timing and location of migration mode shifts. Disruptions in the formation of columns has been associated with several disorders,

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FIGURE 24.1 Ontogenetic radial units develop into minicolumns. Rakic’s Radial Unit Hypothesis asserts that radial glial neural progenitor cells (RGs; dark red, blue, yellow) produce clones of excitatory projection neurons (light red, blue, yellow) that migrate into the cortical layers (II/III, IV, V/VI) where they receive sensory input, intersynapse, and project to the contralateral hemisphere and/or subcortical structures (arrows indicate direction of signal transmission). Sister neurons of a vertical radial unit (three units are shown, separate by dashed vertical lines) are more likely to intersynapse and simultaneously respond to the same kind of sensory input, making them a functional unit called a minicolumn. Inhibitory interneurons (I), which arise from progenitors in the subpallium (not shown), also migrate to and integrate into minicolumns. Whether interneurons in the same column descend from the same progenitor (both purple) or not (one orange, one green) is still debated.

including schizophrenia, dyslexia, Alzheimer’s disease, and autism (Casanova and Tillquist, 2008; Donovan and Basson, 2017), all of which have also been associated with defects in neuronal migration (Galaburda, 2005; Reiner et al., 2016; Valiente and Marín, 2010).

24.2 Migration of related projection neurons into the same minicolumn How does neuronal migration contribute to the formation of minicolumns? Whatever the mechanism, it would have to allow for neurons that are stacked vertically in the laminae to be preferentially connected to each other and be able to form an independent microcircuit. One simple mechanism that could produce this arrangement would be to elaborate minicolumns using neurons from the same NPC cell. Since the daughter neurons of a single NPC are already spatially close, the columns would readily form if these neurons were kept close during migration, and there was some mechanism to drive their preferential synapsing with each other during migration or after migration finishes. Rakic’s Radial Unit Hypothesis fits these requirements well. According to the Radial Unit Hypothesis, the tangential coordinates of PNs in the mature cortex topologically match the tangential coordinates of the NPCs from which the PNs arise from because the newborn neurons migrate radially into the laminae attached to the processes of the NPCs from which they arise, with little tangential movement into surrounding areas (Rakic, 1988, 2007). The PN NPCs, also known as radial glial cells, act as a migratory scaffold not only for their daughter PNs but also for INs (Malatesta et al., 2000; Rakic, 1972; Tamamaki et al., 2001; Yokota et al., 2007), supporting the idea that radial glia help restrict the tangential movement of migrating neurons. The Radial Unit Hypothesis predicts that neurons in a radial unit or minicolumn that span across the radial axis of the cortex

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would be more closely related and more likely to synapse than neurons spanning across multiple columns along the tangential axis (Costa and Hedin-Pereira, 2010). In other words, the ontogenetic logic of neurogenesis and neuronal placement would dictate the connectivity logic within the minicolumns (Shi et al., 2017). What is the evidence that sister neurons (neurons that arise from the same NPC) have a limited tangential distribution within cortical areas or within minicolumns and preferentially synapse with each other? The evidence for PNs and then for INs will be reviewed later.

24.2.1 Early lack of evidence that sister projection neurons migrate into the same minicolumn Early examination of the restricted tangential distribution of sister PNs used delivery of low-titer, replication-defective retroviral vectors expressing the LacZ gene to NPCs lining the ventricles, so that the placement of the NPC’s neuron progeny after they migrate could be visualized. These initial studies showed that clusters of neurons that presumably came from the same NPC were seen in all areas of the developing cortex, with a concentration of clusters in the dorsomedial areas (Price and Thurlow, 1988; Walsh and Cepko, 1992, 1993). Interestingly, in the migratory intermediate zone (IZ), clustered cells were often tightly radially aligned, but in the differentiating cortical plate, the radial clusters had more tangential spread, and neurons in some clusters seemed to have migrated radially along separate radial glial processes into different columns (Walsh and Cepko, 1988). Neurons migrating tangentially in the ventricular zone (VZ), subventricular zone (SVZ), and IZ were also observed (called horizontal cells (Fishell et al., 1993; O’Rourke et al., 1995; Price and Thurlow, 1988)) and were initially proposed to be tangentially migrating NPCs (Walsh and Cepko, 1993) but were later shown to be postmitotic INs migrating tangentially (Anderson et al., 1997; O’Rourke et al., 1997). In the neonatal cortex, this tangential spread of sister neurons was also observed, with fewer than 25% of LacZ-labeled sister neurons being found together within a single or adjacent radial column (Walsh and Cepko, 1988). In fact, astroglial cells were more likely to be found in radial arrays than neurons were (Walsh and Cepko, 1988). This wide distribution across multiple columns comported with a study that looked at random clonal expression of LacZ through genetic recombination and found sister neuron clonal clusters widely distributed, even across hemispheres (Wilkie et al., 2004). Although many of those clones were likely the result of independent or very early recombination events, some of the clones could be widely dispersed sister neurons. In contrast, other groups found a more consistent radial clustering of neuronal clones, with PN clones being distributed in a narrow vertical band (Kornack and Rakic, 1995; Soriano et al., 1995; Tan and Breen, 1993) and, in fact, less radial clustering of astroglial cells (Luskin et al., 1988). These findings were supported by another study, which saw radial stacking of clones (Cai et al., 1997). All together, these studies were unclear in their support of the Radial Unit Hypothesis and the restricted tangential migration of sister PNs. One explanation for the conflicting findings described earlier might be that those early studies used low titer virus to label sister neurons with the assumption that clustered, infected cells arose sequentially from the same NPC. However, the possibility remained that progeny from two or more different NPCs could have been labeled and had migrated to different areas or, coincidentally, to the same area. And in fact, the viral vectors used in those early studies labeled NPCs of the pallium and subpallium, and so they were labeling both PN and IN progeny, which are derived from separate NPC pools and could either disperse widely or cluster, since INs spread widely across the cortex but also associate closely with PNs (Parnavelas et al., 1991). The possibility of having labeled multiple NPCs was reduced with the introduction of retroviruses that contain unique genetic barcodes that are extracted in situ after migration to confirm that the clustered cells are indeed clones, arising from the same NPC (because they all share the same barcode) (Walsh and Cepko, 1992). Using this technique, one group found that, in clustered clones, the cells were of the same type (PN or IN), and unclustered clones were composed of multiple cell types (Reid et al., 1995), suggesting that earlier studies had in fact simultaneously labeled pallial and subpallial NPCs and had conflated them as clones. The explanation for the unclustered clones was that multiple NPCs may have all received the same barcode by chance. So, had the clustered, clonal neurons of the same cell type migrated into a columnar orientation? They had not, but rather resided near each other in the same one or two laminae (Walsh and Cepko, 1992), a finding that has been replicated using a different technique (Maruoka et al., 2011). Moreover, the clustered clones spanned tangentially across barrel columns (Walsh and Cepko, 1992), and across areas in the hippocampus (Grove et al., 1992), which suggested that sister neurons do not migrate into the same column and countered the proposed connection between radial units, minicolumn formation, and cortical arealization. Using another technique where the LacZ gene on the X chromosome was randomly expressed in a subset of NPCs, neurons clustered in radial columns, but the columns did not seem to coincide with areal boundaries in the barrel cortex (Tan et al., 1995), suggesting the possibility not only that neuronal progeny from a single NPC migrates into the same column but also that a single NPC can contribute neurons to different areal regions. In the same study, several other clonal neurons were spread widely across the cortex, suggesting that these sister neurons had migrated tangentially over large

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distances and across functional areas. What might account for such a wide tangential dispersion of the neurons? Walsh and Cepko hypothesized that neurons might disperse tangentially as they follow radial glial guides that defasciculate as they extend through the cortical plate (Walsh and Cepko, 1992). They claimed that “areal specification occurs after neurogenesis” (Walsh and Cepko, 1992), and thus would not be a product of migration but of some postmigratory organizational process.

24.2.2 Sister projection neurons migrate into the same minicolumn and intersynapse In 2001, Noctor et al. used live imaging to trace the proliferation of a single pallial NPC and the migration of its PN progeny by using GFP-expressing retrovirus at low titer to label NPCs in cultured cortical slices (Noctor et al., 2001). They found that neuronal progeny from a single NPC maintained physical contact with the radial process of the NPC during their migration, and once they reached the cortical plate, they became displaced from the radial process by only 1e2 cell body diameters. This suggested that PNs show very limited tangential distribution and establish radial units, providing convincing evidence that sister neurons’ migration is tangentially restricted after all and supporting the idea that radial units might underlie minicolumns. Do the neurons in these radial units form functional connections? Yes. When labeled radial clones were analyzed for synaptic connections in cultured postnatal cortical slices, Yu et al. found that sister PNs were more likely to synapse with each other, even when their dendritic arbors overlapped with nonsister, neighboring PNs (Yu et al., 2009). The synaptic connections were unidirectional, glutamatergic, and the directionality of the connections matched that seen in mature brains (ascending from layer IV to layers II/III and descending from II/III to V/VI). Interestingly, the likelihood of chemical synaptic connections being formed between sister neurons increased as the brain matured, suggesting that there was a prior signal between sister neurons that preceded their chemical synapsing. The same group later found that sister neurons are first electrically coupled before they become chemically coupled (Yu et al., 2012), and the electrical coupling is required for neurons in the same column of visual cortex to respond to the same type of visual stimulus (Li et al., 2012b). This supports the idea that functional minicolumns can only form if sister neurons migrate in close enough proximity to maintain connections. Further work will need to show what the limits are for tangential dispersion of PNs while still being able to maintain synaptic connections.

24.3 Integration of projection neurons into cortical minicolumns What processes during migration help maintain the proximity and connection between sister neurons in radial units that become minicolumns? It is likely that chemical signals, adhesion to migratory scaffolds, and geometrical/mechanical factors constrain migrating neurons to remain within a narrow tangential spread. Here, the focus is on the adhesion to scaffolds and chemical signals that keep neurons within a narrow migratory path and facilitate their connectivity throughout migration (Fig. 24.2), but it will be interesting to pursue how mechanical and geometric forces determine the pathway of migrating neurons, as has been shown in vitro (Chen et al., 2013; Sun et al., 2011). For example, do migrating neurons express mechanoreceptors, and if so, does their activation at the plasma membrane transduce into changes in cell polarity and migration direction (Cui and Yuan, 2007)? Do mechanical interactions with other neurons and the surrounding tissue constrain migration trajectories? The evidence for a role for blood vessels, radial glia, and molecular signaling in keeping PNs tangentially constrained is discussed in the following.

24.3.1 Migratory scaffolds restrict tangential movement of projection neurons Does cortical vasculature provide a migratory substrate for PNs that keeps sister neurons constrained within a minicolumn? The congruence of the timing and location of angiogenesis, neurogenesis, and neuron migration and the sharing of signaling molecules between these three processes suggest the possibility that blood vessels might act as migratory scaffolds that keep PNs within a narrow radial band as they migrate radially (Javaherian and Kriegstein, 2009; Stubbs et al., 2009). After all, blood vessels act as a migratory scaffold for cells in postnatal cortex. For example, tangentially migrating neuroblasts moving toward the adult olfactory bulb migrate along blood vessels, as do radially migrating neurons moving through the corpus callosum into the neonatal cortex (Bovetti et al., 2007; Le Magueresse et al., 2012; Snapyan et al., 2009). In the embryo, oligodendrocyte precursors migrate along vessels (Tsai et al., 2016), and intermediate PN progenitors in the SVZ associate with vasculature as they divide (Javaherian and Kriegstein, 2009), but there is little evidence that embryonic PNs actually use vessels as a migratory substrate. Furthermore, vasculature patterns do not align with barrel

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Ephrins Rac1 FLRTs FoxG1 Unc5D

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FIGURE 24.2 The tangential spreading of migrating projection neurons (PNs) is tightly restricted within forming minicolumns. For sister neurons to incorporate into the same minicolumn, their radial migration path must be tangentially restricted. There are four points along a migrating PN’s path where its tangential movement is regulated: As it attaches to and moves along the radial glial fiber (1 and 3), during the multipolar migration stage (2), and in the cortical plate when the neuron detaches from the glial fiber and moves tangentially to possibly contact and migrate along an outer radial glial fiber (yellow) or another radial glia (4). Several factors (pink arrows) control the migrating neuron’s attachment to radial glia, including astrotactin and others. Other factors, such as ephrins and FLRTs, regulate how long a neuron remains in the multipolar stage and how far it tangentially moves away from the glial fiber (green arrows). For example, FoxG1, a transcription factor that controls the expression of the FLRT receptor Unc5D, controls the timing of a neuron’s escape from the multipolar phase and resumption of radial migration along the glial fiber. The length of a neuron’s multipolar phase might determine the likelihood of it remaining in the same radial unit as its sister neurons. FLRT, fibronectin and leucine-rich transmembrane proteins.

columns or columns in the visual cortex, reducing the likelihood that angiogenesis patterns constrain the tangential spread of PNs or facilitate minicolumn formation (Adams et al., 2015; Blinder et al., 2013). IN incorporation into columns may rely on vasculature, however, because IN NPCs in the ventral telencephalon closely associate with vessels, unlike dorsal PN NPCs (Tan et al., 2016), and INs depend on vascular endothelial growth factor (VEGF) signaling from vessels for their tangential and radial migration (Barber et al., 2018). It remains to be seen whether disruption in VEGF signaling changes the placement of INs. More evidence supports the idea that sister PNs are kept within the same column by close association to the radial processes of radial glial NPCs (Fig. 24.2). Newborn neurons migrate along the radial process of the NPC from which they arise (Noctor et al., 2001; Rakic, 1972), or along a neighboring radial process (Rakic et al., 1974; Walsh and Cepko, 1988). A number of factors have been revealed to maintain the adhesion between neuron and radial glia, including integrins (Schmid and Anton, 2003), neuregulin (Anton et al., 1997), astrotactin (Fishell and Hatten, 1991), profilin (Kullmann et al., 2011), connexins (Elias et al., 2007), and focal adhesions (Valiente et al., 2011) (reviewed in (McGrath et al., 2001; Solecki, 2012)). The adhesion is broken, however, at two points in PN migration where PNs detach from their radial glial scaffold and spread tangentially: during the multipolar stage and at the end of migration when the neurons enter the cortical plate and begin to differentiate. In the multipolar stage, migrating PNs in the SVZ detach from the radial glial process, extend multiple neurites, send out an axon, move tangentially, and sometimes divide as intermediate progenitors (Kowalczyk et al., 2009; Noctor et al., 2004; Tabata and Nakajima, 2003). PNs remain in the multipolar stage for at least a day and then leave it by resuming a bipolar morphology, reattaching to the glial process, and moving radially up into the cortical plate (Cooper, 2014). Once they reach the laminae of the cortical plate, factors such as reelin and SPARC-1 cause PNs to detach from the radial glia where they again can spread tangentially (Dulabon et al., 2000; Gongidi et al., 2004; Noctor et al., 2001). This suggests that if either the multipolar phase is prolonged or the detachment from glial processes is premature, the construction of a column of connected sister PNs is at risk because of the increased possibility of tangential dispersion. This means the control of the initiation, duration, and ending of the multipolar stage and the timing of radial glial detachment are likely going to control how well sister neurons can maintain the proximity that allows synaptic connections with each other. In fact, in contexts where migrating neurons become prematurely detached from their parent radial fiber, they migrate more tangentially (Ji et al., 2017; Valiente et al., 2011). In addition to constraining the tangential dispersion of sister neurons, radial glial progenitors also might directly support PNs’ physical interaction with each other during migration, which seems to be an important prerequisite for sister neurons subsequently intersynapsing in the cortical plate. Electrical synapses develop between sister PNs embryonically, but they

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can only develop if the younger sister neurons migrate past their older siblings along radial glia in the stereotypical “insideeout” fashion (Berry and Rogers, 1965; Rakic, 1974), because in the Reeler cortex, even though daughter neurons can attach normally to the radial process of the same radial glial cell, they fail to migrate past each other and then fail to develop electrical coupling (He et al., 2015). It may be that physical contact between sister neurons during migration is required for their synapsing. Electrical synapses are also found between the radial glial NPC and its daughter neurons, and this coupling is surprisingly maintained even when the daughter neuron becomes multipolar (He et al., 2015) and apparently detaches from the radial fiber (Tabata and Nakajima, 2003), but it has not yet been tested whether electrical coupling of sister PNs to their mother radial glial NPC is required for the sister PNs’ subsequent electrical synapsing.

24.3.2 Molecular signaling limits tangential movement of projection neurons during multipolar stage Electrical coupling between sister neurons also depends on the migrating neurons’ behavior during the multipolar phase. If ephrin-A or its receptor, which regulates the tangential dispersion of radially migrating PNs during the multipolar stage (Dimidschstein et al., 2013; Torii et al., 2009), is overexpressed in sister PNs, their tangential dispersion is increased, and their electrical coupling is decreased (He et al., 2015). Ephrin-B1 controls the number of neurites that multipolar neurons form and the extent of their movement away from the mother radial glia. It does this by regulating a guanine exchange factor for Rac1, which is a Rho GTPase involved in cell polarity (Dimidschstein et al., 2013). This suggests that restriction of the tangential distribution of PNs so that they migrate in close contact with each other, in part mediated by ephrins and their receptors, is critical for the formation of the microcircuitry of cortical columns. The additional mechanisms that regulate the duration and tangential distribution of PNs during the multipolar stage are beginning to be uncovered (Fig. 24.2). One proposed mechanism utilizes fibronectin and leucine-rich transmembrane proteins (FLRTs), which are transmembrane proteins that can mediate both cellecell adhesion or repulsion signaling, depending on what kind of receptor they bind to (Seiradake et al., 2014). In migrating PNs, both types of signaling seem to occur. Embryonically, FLRT2 is expressed by PNs residing in the cortical plate, where its ectodomain is cleaved by metalloproteinases (Yamagishi et al., 2011). The FLRT2 ectodomains diffuse down into the SVZ where they act as repulsive cues to Unc5D-expressing PNs in the multipolar stage of migration. The normal repulsion by FLRT2 ectodomains prolongs the multipolar phase and delays Unc5D þ PNs’ entry into the cortical plate until Unc5D expression is downregulated, after which the PNs can enter the FLRT2þ regions. Loss of either the Unc5D receptor or FLRT2 ligand causes premature PN entry into the cortical plate, whereas overexpression of Unc5D prolongs the multipolar phase (Yamagishi et al., 2011). Whether disruptions in FLRT2 or Unc5D expression cause multipolar PNs to move sufficiently tangentially to hinder the formation of radial units or minicolumns remains to be tested. The tangential distribution of PNs can be affected, however, by the disrupted expression of another FLRT, FLRT3, which is expressed in migrating PNs. Conditional knockout of FLRT3 in PNs does not affect the radial migration of PNs but instead limits their tangential movement and causes them to cluster in tighter radial columns in the cortical plate than controls (Seiradake et al., 2014). Interestingly, overexpression of FLRT3 leads to clustering also, but the clustering occurs in the IZ, where the neurons become arrested in their radial migration. This clustering depends on the homophilic interactions between FLRT3 expressed on neighboring PNs (Seiradake et al., 2014). These findings suggest that, during migration, the timing of the shift between multipolar/tangential migration and radial migration is controlled in part by repulsive FLRT2 ectodomains released from differentiating PNs residing in the cortical plate, whereas PN tangential spread is controlled by FLRT3-mediated adhesive interactions between PNs in the multipolar phase or just exiting the phase. If FLRT3 levels are too high or too low, the disrupted adhesive attraction between adjacent PNs causes irregular tangential spread in the multipolar phase, and the irregularities are maintained as the PNs enter the cortical plate, indicating that PNs require a fine balance of repulsive and attractive cues to enter minicolumns with the correct timing and tangential distribution (Solecki, 2012). More recent investigations into the role of FLRTs in the tangential movement of PNs during migration suggest the intriguing possibility that the extent of tangential PN movement may underlie the formation of gyri and sulci in gyrencephalic species. In mice (which are lissencephalic), double knockout of FLRT1 and FLRT3, which are both expressed in migrating PNs, limits PN tangential distribution and causes PNs to aberrantly cluster in tight radial columns in the cortical plate (Toro et al., 2017). Fascinatingly, the clusters underlie gyri resembling those of gyrencephalic species (Toro et al., 2017). In the double knockout, the proliferation of NPCs was normal, as was the structure of radial glial fibers (Toro et al., 2017). Furthermore, in the gyrencephalic ferret, FLRT1 and FLRT3 expression is lower in cortical areas that become sulci than in areas that become gyri, and in humans, FLRT1 and FLRT3 expressions are overall lower than those in mouse (Toro et al., 2017). These studies suggest that rather than progenitor proliferation being the only driver of cortical

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gyrification, PN tangential distribution also has a substantial role. This would fit with observations that, in ferret, the beginning of cortical folding happens at the same time as the lateral dispersion of PNs that migrate sequentially along several radial glial fibers (Gertz and Kriegstein, 2015). All of this suggests that the extent of tangential spread during PN migration may differ between lissencephalic species and gyrencephalic species and that difference may underlie gyrification. How this relates to the formation of cortical minicolumns is unknown, but it will be interesting to test if disruptions in FLRT signaling also alter the formation of radial units or connectivity between sister PNs. Another difference between gyrencephalic and lissencephalic species in terms of tangential dispersion of PNs is the role of outer radial glia in the tangential expansion of supragranular (superficial to layer IV) neurons. In human embryonic cortex, midway through neurogenesis, the radial glia found at the ventricle become truncated and no longer extend basal processes all the way to the pial surface (Nowakowski et al., 2016). A new population of radial glia arises from the ventricular radial glia, and the cell bodies of these new radial glia are found in the outer SVZ. The outer radial glia give rise to most supragranular neurons, which presumably use them as a migratory scaffold. As the outer radial glia divide, they expand tangentially, and so the supragranular neurons migrating along them also spread tangentially. This suggests that, in humans at least, there might be a difference in the tangential expansion of supragranular neurons relative to infragranular neurons, and the two populations may incorporate into connected radial columns differently (Nowakowski et al., 2016). In rodents, a more limited tangential expansion of neurons destined for upper layers may occur via the proliferation of intermediate NPCs found in the SVZ (Vasistha et al., 2015) or may occur in medial areas of the cortex that have more bends in the tissue (Gao et al., 2014). All of these data suggest that the tangential movement of PNs is controlled by adhesive factors between neurons and radial glia or other neurons, tangential movement during the multipolar stage, and movement along radial glia that are themselves spread tangentially. Based on this, we might expect to see in gyrencephalic species either broader columns or narrow columns more often comprising neurons that arise from different radial units. If the latter were the case, then in gyrencephalic species the circuitry of minicolumns would become established by mechanisms less dependent on PNs’ tangential migratory distribution. In other areas of the brain are sister neurons more likely to be part of the same microcircuit or minicolumn? In the thalamus, sister neurons form radial clusters inside some thalamic nuclei, as do GABAergic interneurons (Shi et al., 2017). In the hippocampus, sister neurons organize into tangential, rather than radial, columns and rarely form chemical or electrical synapses with each other (Xu et al., 2014). There is currently no evidence that neurons in the cerebellum or spinal cord migrate into columns, but it is possible considering that both are organized somatotopically like the cortex (Kaas, 2012). In Xenopus tectum, sister neurons are more likely to share the same receptive field properties (Muldal et al., 2014), again supporting the view that sister neurons establish a similar logic of connections and may do so by first following similar migratory paths.

24.4 Integration of interneurons into cortical columns Of course, an integral part of functional minicircuits in cortical columns is having an appropriate balance of excitation and inhibition contributed by excitatory PNs and inhibitory INs, respectively. The ratio of INs to PNs remains relatively constant throughout life, with about 20% of all cortical neurons being INs (Sahara et al., 2012). So how do INs become incorporated into cortical columns, and how is IN migration regulated to ensure a proper number of INs are placed into the proper columns, migrate to the correct layers, and establish the appropriate connections? The origin and migration of cortical interneurons have been reviewed extensively here (Petros and Anderson, 2013) and elsewhere (Bartolini et al., 2013; Faux et al., 2012; Guo and Anton, 2014; Laclef and Métin, 2018; Marín, 2013; Métin et al., 2006; Ruiz-Reig and Studer, 2017; Sultan et al., 2013; Tanaka and Nakajima, 2012), as has the role that disrupted IN migration plays in the development of neuropsychiatric disorders (Chu and Anderson, 2015; Volk and Lewis, 2014). Rather than try to summarize the extensive literature on this, I will focus here specifically on what we know about how the areal distribution of INs is controlled, including whether certain IN subtypes preferentially migrate to specific regions along the rostrocaudal or mediolateral axes. Recent evidence that examines whether sister INs migrate to the same cortical columns and/or establish electrical and chemical connections with each other will also be reviewed.

24.4.1 Interneuron subtypes areally distribute via tangential migration Cortical GABAergic INs can be divided into many subtypes, based on their molecular profile, morphology, and electrophysiology (Petilla Interneuron Nomenclature Group et al., 2008; Faux et al., 2012); however, almost all of them fit within three groups: INs that either express the calcium-binding protein parvalbumin (PVþ), the neuropeptide somatostatin

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(SSTþ), or the ionotropic serotonin receptor (5HT3aRþ) (Rudy et al., 2011). INs can be further subdivided based on their expression of other markers such as calbindin, calretinin, and neuropeptide Y, but these still largely fall within the three larger groups. The subtypes arise from different areas of the subpallium (ventral cortex). SSTþ and PV þ INs arise mainly from different zones of the medial ganglionic eminence (MGE) and preoptic area (POA) (Butt et al., 2005; Fogarty et al., 2007; Gelman et al., 2011; Miyoshi et al., 2007; Sultan et al., 2016; Wonders et al., 2008; Xu et al., 2008, 2004), whereas 5HT3aR þ INs arise from the caudal ganglionic eminence (CGE) (Lee et al., 2010). The further division of INs into subsubtypes likely begins in the 18 different subpallial neurogenic domains that express different combinations of transcription factors (Flames et al., 2007). Interneurons leave their neurogenic zones and move tangentially into the pallium (dorsal cortex), crossing the pallialesubpallial boundary. Once they enter the pallium, they segregate into distinct migration streams in the IZ/SVZ and marginal zone (MZ) (reviewed in (Hatanaka et al., 2016)). How do INs in these streams areally distribute? INs in the MZ and IZ/SVZ streams behave differently, even though they contain similar proportions of IN subtypes (Miyoshi and Fishell, 2011). While INs in the IZ/SVZ migrate predominantly in a medial to lateral direction (Anderson et al., 2001; Jiménez et al., 2002; Tanaka et al., 2003), INs in the MZ and VZ migrate in multiple directions (Ang et al., 2003; Tanaka et al., 2003, 2006; Yokota et al., 2007), with a bias toward moving rostrally and caudally more than medially (Tanaka et al., 2006), or even in random directions (Tanaka et al., 2009) (Fig. 24.3). Because migrating INs move from the VZ and IZ/ SVZ to the MZ (Nadarajah et al., 2002; Tanaka et al., 2003), it is possible that the IZ/SVZ is the location for many INs to

Lateral

Medial

MZ

* CP

*

SVZ/IZ

Radial glia

Differentiated PN

Cxcl12 (SDF-1) GABA

Multipolar PN/Intermediate progenitor

Neuregulin

Interneuron Cajal-Retzius cell VZ

FIGURE 24.3 Factors controlling the shift from tangential to radial migration in cortical Ins. Left: INs migrating through the SVZ/IZ stream move from lateral (left) to medial (right) cortex and often display branched leading processes. Some INs in the SVZ/IZ stream move toward the MZ stream. INs in the MZ stream move medially and laterally. Right: Once INs in both streams arrive at their target minicolumn or radial unit, they will shift migration modes and dive radially into the cortical plate (*). Several factors regulate the timing of this shift. Cxcl12 released from CajaleRetzius cells and intermediate progenitors attracts migrating INs and maintains them in the streams. GABA is initially motogenic to INs but then, in preparation to invade the cortical plate, IN express KCC2 (not shown), which causes GABA to be inhibitory to the migrating INs, and so they pause their tangential migration and shift radially. Neuregulin, secreted by differentiating PNs in the cortical plate, attracts INs into the cortical plate. Far right: Radial glial fibers can also trigger a shift from tangential to radial migration and can act as a migratory substrate. IN, interneuron; IZ, intermediate zone; MZ, marginal zone; PN, projection neuron; SVZ, subventricular zone.

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enter the pallium and distribute medially, after which they enter the MZ to become distributed areally (Tanaka et al., 2003). In other words, INs may move through the IZ/SVZ stream to reach their gross areal coordinates and then move to the MZ to achieve their finer coordinates (Ang et al., 2003) that correlate with cortical columns. If this is true, one would expect that INs’ capacity to respond to cues that regulate gross versus fine movements would change as they move from one stream to another, and as expected, differential gene expression in the two streams has been found (Antypa et al., 2011). Among the genes differentially expressed are cell surface receptors and components of intracellular signaling pathways (Antypa et al., 2011). Do IN subtypes preferentially migrate to different areal regions? So far, there is no evidence that regions of the cortex contain different profiles of INs. In fact, the evidence suggests that their multidirectional migration patterns scatter IN subtypes evenly throughout the cortex (Marín, 2013). So, it may be that the control of the areal segregation of IN subtypes migrating in the MZ is subtle, or it may be that areal distribution depends more on clonal identity than on subtype.

24.4.2 Do sister interneurons migrate into the same minicolumn? Do clonally related sister INs migrate to the same areal location or minicolumn along the tangential axis? If so, this would suggest that there is an intrinsic program within individual interneurons that drives them to migrate to a very specific areal location. Live imaging of migrating INs has never revealed cortical INs migrating in pairs or groups, and so sister INs could only arrive at the same tangential coordinate by either running an intrinsic program they both share or following a very precise migratory trajectory that unrelated INs do not follow. Two initial studies addressed whether sister neurons migrate to the same areal region by labeling single MGE and POA NPCs and their IN progeny using low-titer retrovirus expressing GFP, similar to the clonal analysis done in dorsal cortex PN NPCs (Noctor et al., 2001; Yu et al., 2009), except that the fluorescent tag’s expression was limited to neurons arising exclusively from the MGE and POA (Brown et al., 2011; Ciceri et al., 2013). Both studies showed nonrandom clustering of sister interneurons in the cortical plate, with some clustering in vertical columns (Brown et al., 2011), but most of them clustering horizontally within the same or two adjacent layers (Brown et al., 2011; Ciceri et al., 2013). One group observed that if sister neurons were found in the same layer, they were always born at the same time (Ciceri et al., 2013), likely as a result of intermediate progenitors dividing and producing the sister neurons. Surprisingly, the clustered, presumptive sister INs did not always share the same neurochemical profile (25%e50% of clustered INs shared the same profile) (Ciceri et al., 2013), which is surprising considering they presumably came from the same NPC. These findings suggest that sister INs do cluster but not in a strictly vertical orientation as might be expected if they migrated to the same radial unit or minicolumn. Although both of the aforementioned studies provided strong evidence that they had labeled single NPCs and their progeny, they were limited by the possibility that the labeled INs in clusters came from more than one NPC transduced independently by different viral particles. Two subsequent studies attempted to overcome this limitation by expressing in MGE NPCs both a fluorescent tag and a unique DNA barcode, which would unambiguously distinguish IN progeny coming from different NPCs (Harwell et al., 2015; Mayer et al., 2015). These studies seemed to contradict the first studies, finding that clonally related INs distribute very widely in the cortex, across functional areas, and even across other structures in the same hemisphere such as the hippocampus, striatum, and globus pallidus (Harwell et al., 2015; Mayer et al., 2015). In fact, the findings suggested that unrelated INs were more likely to cluster than sister INs. These conclusions were themselves disputed (Sultan et al., 2016) and reargued (Mayer et al., 2016; Turrero García et al., 2016), but ultimately the question of whether clonally related sister INs cluster in minicolumns is unresolved and depends on (1) how “clustering” and “columns” are defined and (2) whether the clustered INs seen in Ciceri et al. and Brown et al. are truly clonally related. What all groups agreed on, however, is that migrating INs do not migrate to random destinations, and sister INs do not often tightly cluster in vertical columns as PNs do. Considering the promiscuity of IN connections to PNs (Fino et al., 2013), it may be that IN placement does not need to be as precise as PN placement does. In vivo imaging of INs at clonal density as has been done in PNs (Noctor et al., 2001) will be the best way to resolve whether sister neurons are more likely to migrate to the same tangential coordinates.

24.4.3 Sister interneurons preferentially intersynapse If we consider a more liberal definition of clustering (residing within 400e500 mm of each other), do clustered sister INs preferentially synapse with each other? Surprisingly, it appears that they do. About one-third of sister INs are electrically coupled by postnatal week 2 and one-fourth are chemically coupled, but the formation of either type of synapse (chemical or electrical) is not dependent on formation of the other type (Zhang et al., 2017). Importantly, electrical synapses, but not chemical ones, are more likely to develop between sister INs. This preference is specific to sister INs and is not simply a

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result of having arisen from the same MGE/POA NPC region. Surprisingly, even as the intersomal distance between two sister INs increases, the likelihood of them being coupled is still significantly higher than two more closely apposed, unrelated INs. However, the electrical coupling does not become established until the sister INs have finished their migration (Zhang et al., 2017). This suggests that INs may be genetically programmed to migrate to a specific cortical area and establish preferential contacts with INs that come from the same NPC, but it is unlikely that sister neurons will, as a rule, be located in the same minicolumn. How would this specific placement and connectivity be controlled? The answer may be as simple as INs becoming placed in a lateral to medial fashion that corresponds to their birthdate; that is, early-born INs stop their tangential migration in more lateral areas, and later-born INs migrate past them to more medial locations (Moore and Price, 1992). The clustering of sister INs could be then simply be a result of them both being born temporally close (like from an intermediate NPC) and migrating to the same location along the lateral to medial axis. The answer, however, may turn out to be more complicated. It turns out that sister INs that are electrically coupled are also more likely to synapse onto the same PN (Zhang et al., 2017). This raises the possibility that sister IN placement and “clustering” might be controlled by signaling from PNs, which distribute a signal that is specific for INs coming from a single progenitor.

24.4.4 Regulating the timing of the shift from tangential to radial migration When INs migrating through the streams home in on their areal position, they next dive radially into the cortical plate. The timing of this shift from a tangential to a radial trajectory is important for both the tangential and radial positioning of INs, and it appears that PNs contribute to the signal to INs about where and when to initiate this shift (Fig. 24.3). In rodents, this shift begins in late embryonic stages and completes during the first postnatal week, with INs in both the MZ and IZ/SVZ streams turning radially into the cortical plate (Hevner et al., 2004; Polleux et al., 2002; Tanaka et al., 2003; Yokota et al., 2007). It has been suggested that while INs are moving through the IZ/SVZ streams, they are responsive to guidance cues that direct them medially, but once they move to the MZ stream, they become unresponsive to global guidance cues, move randomly, and become more widely dispersed along the tangential axes (Tanaka et al., 2009). However, it may be that the apparent random behavior by INs in the MZ is actually INs refining their location in response to local cues given by CajaleRetzius cells (Ang et al., 2003) or the underlying PNs in the cortical plate. What factors control the timing of the shift by maintaining INs in the tangential migration streams of the IZ/SVZ and MZ? Chemokine signaling seems to play an important role. Multipolar PNs and PN intermediate progenitors found in the SVZ, and CajaleRetzius cells in the MZ, secrete the chemokine Cxcl12 (SDF-1), and the chemokine interacts with its Cxcr4 receptor expressed on INs as an attractive cue to prevent them from prematurely exiting the IZ/SVZ/MZ streams and entering the cortical plate (Li et al., 2008; Stumm et al., 2003; Tiveron et al., 2006) (Fig. 24.3). Cxcl12 expression in the IZ increases in a lateral to medial direction and this correlates with the migration of INs medially (Daniel et al., 2005). Just before birth, however, INs become unresponsive to Cxcl12, and this allows them to exit the streams and enter radially into the cortical plate (Li et al., 2008; López-Bendito et al., 2008). Transgenic deletion of Cxcr4 expression in INs causes a premature entry into the cortical plate, a misdistribution of INs across areal regions and across cortical layers, and disruptions in the inhibitory tone (Li et al., 2008; López-Bendito et al., 2008; Meechan et al., 2012). Loss of the Cxcl12 expressed in intermediate progenitors in the SVZ causes the leading processes of INs in the SVZ to be abnormally oriented toward the cortical plate and MZ (Abe et al., 2015). Interestingly, this loss of Cxcl12 in intermediate progenitors also reduces the extension of thalamocortical axons into the cortical plate, but whether the axon misrouting causes the IN defect is not clear. Removal of CajaleRetzius cells, which also express Cxcl12, causes premature entry of INs into the cortical plate (Caronia-Brown and Grove, 2011). Apparently, Cxcl12 to Cxcr4 signaling is not necessary for the tangential migration of INs per se but rather regulates the timing of their tangential to radial shift, which indirectly affects IN areal distribution by altering how long the INs continue to migrate tangentially. Interestingly, Cxcl12 may mechanistically prevent the shift to radial migration by reducing branching of IN leading processes and increasing their migratory speed (Lysko et al., 2014), in part by activating doublecortin, which bundles microtubules in the leading process (Bielas et al., 2007) and is necessary for tangential, but not radial, migration in rodent (Corbo et al., 2002; Kappeler et al., 2006; Koizumi et al., 2006). Neurotransmitters such as GABA also seem to play a role in controlling the timing of the shift from tangential to radial migration (Fig. 24.3). GABA stimulates Caþþ transients and thus the motility of tangentially migrating INs (because GABA is depolarizing to immature INs) (Behar et al., 1998, 1996). This has the effect of increasing the multidirectional motility of INs in the migratory streams (Inada et al., 2011). In preparation to turn and migrate radially, expression of the chloride/potassium transporter KCC2 increases in INs, which changes their response to GABA from depolarization to hyperpolarization. Hyperpolarization reduces migration speed (presumably by lowering Caþþ transients) and increases

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pausing, which occurs before INs turn radially and enter the cortical plate (Bortone and Polleux, 2009). The timing of KCC2 upregulation and the subsequent pausing seem to be controlled in part by thalamocortical afferent axons (TCAs), which course through the same areas and with the same timing as migrating INs. TCAs release glutamate onto INs, which maintains their KCC2 expression at levels that promote pausing but then allow subsequent entry into the cortical plate (Zechel et al., 2016). Disruption of KCC2 expression or blocking GABA signaling in migrating INs changes the timing and ability of INs to radially invade the cortical plate (López-Bendito et al., 2003). Supporting the idea of neurotransmitters controlling the timing of the shift in migration mode (Luján et al., 2005), an interesting study in zebrafish hindbrain suggests the possibility that different neurotransmitters found along a migrating neuron’s path control different phases of migration, specifically at points where the neurons turn perpendicularly (Theisen et al., 2018). If this also holds true for mammalian cortical INs, it will also likely to apply to migrating PNs, which express a variety of neurotransmitter receptors, and signaling through those receptors can change migratory behavior (Hurni et al., 2017). Is there any evidence that cellecell adhesion or cellecell contacts play a role in maintaining INs in the IZ/SVZ and MZ streams? In the rostral migratory stream of the adult cortex, immature neuroblasts arising from the SVZ tangentially migrate to the olfactory bulb as chains of closely apposed cells (Lois et al., 1996) held together by cellecell adhesion molecules such as integrins, neural cell adhesion molecule (NCAM), and neuregulins (reviewed in (Lalli, 2014; Sun et al., 2010)). Once the neuroblasts enter the olfactory bulb, they exit the chains, turn perpendicularly, and migrate radially into the olfactory bulb. In this system, the switch from tangential to radial migration seems to depend on the downregulation of the cellecell adhesion molecules (Alfonso et al., 2015; Fujikake et al., 2018) and expression of reelin (Hack et al., 2002). Although embryonic cortical INs express NCAM (Daniel et al., 2005), integrin (Stanco et al., 2009), and the neuregulin receptor ErbB4 (Yau et al., 2003), in the tangential streams, they migrate as separate cells and do not seem to adhere to each other (Ang et al., 2003; Yokota et al., 2007), but yet those molecules are required for IN migration. The adhesion molecules in cortical INs seem to instead control cell motility, branching dynamics, and channeling INs within certain paths (Krocher et al., 2014; Li et al., 2012a; Luccardini et al., 2013; Lysko et al., 2014; Martini et al., 2009; Stanco et al., 2009; Valiente and Martini, 2009), which appear to be controlled independently in each IN. Like PNs, cell-to-cell ephrin signaling also seems to play a role in IN placement. Migrating INs express ephrins-Bs and ephrins-B receptors, and a conditional deletion of ephrin-B1e3 causes INs to accumulate in the lateral cortex (Talebian et al., 2017). It appears that INs are guided both by reverse signaling from unidentified guide cells expressing the ephrin-B receptors and by forward signaling from radial glial cells. Reminiscing the effect of the loss of ephrin signaling in PNs, a loss of ephrin-A2 leads to patches of cortex lacking both PNs and INs (Homman-Ludiye et al., 2017). Whether these patches are correlated to altered cortical columns remains to be investigated.

24.4.5 Projection neurons attract migrating interneurons into cortical plate Once INs begin their shift from tangential to radial migration, what types of cues from PNs in the cortical plate attract them to invade the cortical plate? INs invade the cortical plate only after PNs have settled into their laminar positions (Pla et al., 2006), suggesting the possibility that they are waiting for a signal that only more mature PNs can provide before they invade. Because MGE-derived INs come to reside in the same layers as PNs that are born at a similar time (Marín, 2013), INs end up positioned in an insideeout fashion like PNs (Miyoshi et al., 2007). However, IN radial migration is not strictly insideeout as it is for PNs, nor is IN lamination as tightly restricted as that of PNs, but rather the INs are initially widely distributed prenatally throughout the cortical plate and then become sorted to layers containing coetaneous PNs in the first postnatal week (Miyoshi and Fishell, 2011). One exception is CGE-derived INs, which are widely distributed prenatally, but then only settle in supragranular layers (layers II/III), no matter their birthdate (Miyoshi et al., 2010). This suggests that IN subclasses, which are specified at birth (Fogarty et al., 2007), migrate toward certain cues from PNs that they specifically respond to because of their birthdate. Supporting this, in the cortex of the reeler mouse, where the insideeout lamination of PNs is inverted, the layering of INs is inverted also (Hevner et al., 2004), but the IN inversion is directly a result of PN mislamination, rather than loss of reelin signaling to the INs (Pla et al., 2006). Also, subcerebral PNs and callosal PNs segregate into different columns (Maruoka et al., 2017) and chemoattract and recruit different subtypes of INs (Lodato et al., 2011), meaning that certain types of interneurons are likely to be positioned specifically within different types of columns. Neuregulin 3 (Nrg3) is at least one cue, expressed in the cortical plate but not in the migration streams, that is an attractant for INs, which respond to Nrg3 via the ErbB4 receptors they express (Bartolini et al., 2017; Flames et al., 2004). When Cxcl12 expression is still high in the migration streams, INs prefer the Cxcl12 attractant and are not pulled into the cortical plate by Nrg3, but once Cxcl12 expression decreases, Nrg3 can then attract INs into the cortical plate (Bartolini et al., 2017). The prolonged settling of INs in the CP during the first postnatal week may also be a time when INs establish transient circuits with PNs (Anastasiades et al., 2016) and may “test the waters” in this way to verify they are synapsing with the correct partners.

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24.4.6 Radial glial cells trigger a shift in migration mode Radial glial cells might also control the shift in migration modes (Fig. 24.3). PN expression of the gap junction protein connexins are necessary for PN adhesion to RGCs and radial migration of PNs, in part by stabilizing a radially oriented leading process (Elias et al., 2007). Similarly, in INs, expression of connexins is not required for tangential migration, but it is required for the switch from tangential to radial migration (Elias et al., 2010). The gap junctions seem to promote the switch by enabling the adhesion of INs to radial glia rather than through any communication between the cells through the gap junction channels. It also seems that the mere contact of INs with radial glial endfeet or soma may change their migration mode (Yokota et al., 2007).

24.5 Genetic and cellular mechanisms controlling shifts in migratory modes If properly executed neuron migration is essential for functional minicolumn formation, then what was reviewed earlier suggests that changes in the location and timing of switches between different modes of migration (tangential and radial) might disrupt minicolumn formation. In both PNs and INs, a shift in migratory mode from radial to tangential or vice versa is accompanied by changes in gene expression. The question now arises: Are there master regulatory genes that dictate the migratory mode (radial or tangential) of PNs and INs? If one does exist, its activity would be expected to change as neurons shift between modes. FoxG1 may be one such regulator. Expressed in radially migrating PNs and INs, FoxG1 acts as a transcriptional repressor that represses genes that are active during tangential migration (Fig. 24.3). For example, during the multipolar phase of PN migration, when PNs move tangentially, FoxG1 expression is downregulated, which allows the netrin receptor Unc5D to be expressed (Miyoshi and Fishell, 2012). Unc5D may confer on the PNs a repulsive response to the netrin or FLRT2 ectodomains secreted from the cortical plate, preventing them from entering the cortical plate and also possibly directing the outgrowth of an axon, which occurs during the multipolar phase (Noctor et al., 2004). Reexpression of FoxG1 is required for PNs to leave the multipolar phase and resume radial migration (Miyoshi and Fishell, 2012). Importantly, FoxG1, which regulates the expression of the Cxcl12 receptor Cxcr4 in migrating INs (Shen et al., 2018; Yang et al., 2017), is also downregulated in INs during tangential migration and is reexpressed as they shift to radial migration into the cortical plate (Miyoshi and Fishell, 2012). In a similar fashion, the transcription factor Sip1 suppresses the expression of another netrin receptor Unc5B and allows interneurons to enter the cortex (van den Berghe et al., 2013). If the formation of minicolumns depends on the duration and timing of tangential migration of both PNs and INs, then disruption in FoxG1 or Sip1 expression or the genes they induce should also alter the cortex’s columnar organization. This question has yet to be tested. This leads to another fundamental question: What are the intracellular changes that must occur in the neuronal cytoskeleton to change migratory modes? And is the shift between migration modes a mechanistic bottleneck? In other words, are mutations underlying neurodevelopment disorders more likely to affect migrating neurons at stages where they shift migration modes? In the case of PNs, the multipolar stage (comprising a shift from radial to tangential and then back to radial) of migration seems to be a vulnerable phase for neurons, being specifically disrupted in several neurodevelopmental disorders (LoTurco and Bai, 2006). Why are neurons seemingly so susceptible to arrest at that stage of migration? First, these shifts require a fundamental change in the constitution and dynamics of the cytoskeleton, including dramatic shifts in the position in intracellular organelles such as the centrosome (Cooper, 2013; Higginbotham and Gleeson, 2007), actin, and microtubules. Some evidence suggests that radially migrating neurons rely more on the actin cytoskeleton, whereas tangentially migrating neurons rely more on microtubules (Eom et al., 2014). For example, when lamellipodin, a protein that integrates signaling from Ras and phosphoinositides to regulate actin dynamics and cell adhesion, is knocked down in radially migrating PNs, they adopt a tangential migration mode and cannot escape the SVZ (Pinheiro et al., 2011). The shift to tangential migration results from a reduction in the F-actin to G-actin ratio and a silencing of the transcription factor SRF, which controls expression of actin (Pinheiro et al., 2011). Also, deletion of n-cofilin, an actin depolymerizing factor, impairs radial migration but only partially impairs IN tangential migration (Bellenchi et al., 2007). Furthermore, loss of Rac1, a RhoGTPase that regulates actin dynamics (Jaffe and Hall, 2005), has no effect on the tangential migration of INs (Vidaki et al., 2012). Loss of the microtubule regulator APC in migrating INs causes overbranching and misorientation of the leading processes (Eom et al., 2014) due to disruptions in microtubule severing, but PN migration is not affected with the loss of APC. In fact, loss of APC does not even affect microtubule severing in PNs (Eom et al., 2014), suggesting that there are entirely different genetic programs controlling radial and tangential migration. Destabilizing F-actin disrupts the shift from tangential to radial migration in neuroblasts moving from the rostral migratory stream into the olfactory bulb and in

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granule neurons in the cerebellum (Kajita et al., 2017; Trivedi et al., 2017), but this switch depends on the coordination between the actin and microtubule cytoskeleton (Trivedi et al., 2017). Recent evidence shows that disrupting posttranslational modification of cytoskeletal proteins such as myosin light chain can shift an IN’s migration mode from a saltatory mode to a smoother, treadmill-like mode (Silva et al., 2018). All this supports the view that switches in migration mode are mechanistically complicated events. The second reason for neuronal vulnerability at migratory mode shift points comes from the different mechanisms that guide neurons in these two modes. Because tangentially migrating neurons do not rely on migratory scaffolds such as radial glia (although they may rely on blood vessels (Magueresse et al., 2012; Sun et al., 2015; Won et al., 2013)), a shift from radial to tangential migration forces the cell to become more reliant on guidance cues, which requires a change in receptor expression and a more microtubule-dependent branching morphology to be able to probe the environment for the cues (Nadarajah et al., 2003). Two studies highlight this dependence that tangentially migrating neurons have on guidance cues. Eliminating the ability to transduce extracellular signals by disabling the primary cilium, an important cellular signaling center that expresses receptors for many extracellular cues (Berbari et al., 2009; Goetz and Anderson, 2010; Guemez-Gamboa et al., 2014; Lee and Gleeson, 2010; Whitfield, 2004) including ErbB4, Cxcr4, and Cxcr7 (Higginbotham et al., 2012), severely disrupts IN, but not PN, migration (Higginbotham et al., 2012). IN migration is slowed, branching is increased, and the shift to radial migration into the cortical plate is disrupted (Baudoin et al., 2012; Higginbotham et al., 2012). The primary cilium is required in INs for their response to cues secreted by PNs (Higginbotham et al., 2012), and the shift from tangential to radial migration specifically depends on sonic hedgehog signaling (presumably from lower layer PNs that express Shh (Harwell et al., 2012)), which alters MT dynamics and facilitates the shift in migration mode (Baudoin et al., 2012). Mutations in the ciliary gene Arl13b that are found in people with Joubert syndrome also disrupt IN migration in mice (Higginbotham et al., 2012). These findings suggest that cognitive disorders such as Joubert syndrome may result from the inability of migrating INs to shift migration modes, thus eliminating the cell’s ability to make correct decisions about their point of entry into particular cortical columns in the cortical plate. In sum, points of shift in migratory modes are points of high susceptibility to environmental and genetic perturbations, and those perturbations could have important implications for the proper formation of minicolumns and a functional cortical circuitry.

24.6 Conclusion This chapter described how the mechanisms governing PN and IN migration may contribute to the development of the columnar structure of the cortex. One pattern that seems to have emerged, for PNs especially, is that neurons arising from the same progenitor migrate to the same location, as the Radial Unit Hypothesis suggests, and are more likely to synapse with each other than with other nearby unrelated neurons. It would seem, then, that minicolumn formation depends on the spatially constrained migration of related neurons, although it is not clear what is the actual advantage to having sister neurons preferentially synapse. Also, minicolumn circuitry is plastic, and so the initial functional connections that are made possible by the constrained migration may be only preliminary to the mature columnar configuration. For example, orientation columns in the visual cortex can reconfigure during adaptation, suggesting that mature columns are initially anatomically derived via migration but then later functionally refined (Bachatene et al., 2015). Since there seems to be a minimum number of synapses that need to form between two neurons before they will cement a connection (Fares and Stepanyants, 2009), and assuming the number of initial connections two cells can form is constrained by the intercellular distance, then migration may simply serve to bring neurons to a “ballpark” location where they can establish initial connections that will later be refined by experience. Most of our understanding of the regulation of the modes of migration to form the columnar architecture of the cortex comes from studies in rodent models. It remains to be seen how much of what have been discussed applies to human cortical development. Some relevant questions are as follows: How do outer radial glia, which are more prominent in humans, contribute to the formation of cortical columns (Fietz et al., 2010; Hansen et al., 2010; Nowakowski et al., 2016; Reillo et al., 2011; Shitamukai et al., 2011; Wang et al., 2011)? Are sister PNs arising from and migrating along the same outer radial glia more likely to form synaptic connections with each other? How do neurons migrating into the cortex postnatally integrate into the already forming/formed columns (Paredes et al., 2016)? Is tangential migration of PNs in the multipolar stage and as they move tangentially from one outer radial glial to another a more recent evolutionary development? Are the transient tangentially migrating PNs (García-Moreno et al., 2018; Griveau et al., 2013) that appear very early in rodent development also found in human, and, if so, how do they contribute to the columnar structure? To what

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extent does the early tangential dispersion of radial glia occur in humans, and how does it contribute to the formation of columns (Wilkie et al., 2004). And crucially, to what extent are psychiatric disorders such as schizophrenia, autism, and bipolar disorder, where cytoarchitectonic defects are subtle, due to alterations in neuronal migration into cortical columns?

List of abbreviations IN interneuron IZ intermediate zone MGE medial ganglionic eminence MZ marginal zone NPC neural progenitor cell PN projection neuron POA preoptic area PV parvalbumin SST somatostatin SVZ subventricular zone VZ ventricular zone

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Chapter 25

The impact of different modes of neuronal migration on brain evolution Fernando Garcı´a-Moreno1, 2, a and Zolta´n Molna´r3 1

Achucarro Basque Center for Neuroscience, Parque Científico UPV/EHU Edif. Sede, Leioa, Spain; 2Ikerbasque Foundation, Bilbao, Spain;

3

Department of Physiology, Anatomy and Genetics, University of Oxford, Oxford, United Kingdom

Chapter outline 25.1. Types of neuronal migration in vertebrate brain developmentdradial and tangential migration shaping vertebrate brains 25.2. The impact of radial migration on brain evolution 25.2.1. Evolution of radial migration 25.2.2. Radial migration on laminar brains 25.2.3. Radial migration on elaborated brains 25.2.4. The influence of radial migration on pallial internal circuitry 25.2.4.1. Somal translocation 25.2.4.2. Glial-guided locomotion 25.2.4.3. Evolutionary origin of glial-aided locomotion 25.3. The impact of tangential migration on brain evolution 25.3.1. Pallial interneurons and the modulation of brain circuits

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25.3.1.1. Conserved features of tangential migration of pallial interneurons in vertebrates 25.3.1.2. Divergence in tangential migratory routes of pallial interneurons 25.3.1.3. Diversifying complexity of GABAergic subtypes 25.3.2. Glutamatergic tangential contributions as developmental scaffolds 25.3.3. Tangential migration shaping brain connectionsdguidepost neurons in evolution 25.3.4. Tangential migrations along the central nervous system 25.4. Conclusions Glossary References

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25.1 Types of neuronal migration in vertebrate brain developmentdradial and tangential migration shaping vertebrate brains One of the main features of the vertebrate nervous system is the disparity between where neurons are born and where these cells eventually reside and function in the mature tissue. Most neurons migrate from their site of birth to a final position elsewhere to integrate into circuits with other neurons. This cellular movement can be from just a short distancedwhen young neurons are passively pushed away by newborn neuronsdor it can span considerable distances with journeys throughout the embryonic braindwhen actively moving neurons navigate to for a definite settlement. The way that the pattern of neuronal migration changed over vertebrate evolution has had a major impact on brain structure and diversity. Cellular diversity in the central nervous system (CNS) varies among species; there are hundreds of different neuronal subtypes in mammals (Zeisel et al., 2018). To generate this diversity, progenitors first change their neuronal production output according to the location and timing of neurogenesis. And secondly, the subsequent neuronal migration accounts as one of the major sources of neuronal diversity and brain complexity. Evolution is the evolution of development. And

a Senior author.

Cellular Migration and Formation of Axons and Dendrites. https://doi.org/10.1016/B978-0-12-814407-7.00025-0 Copyright © 2020 Elsevier Inc. All rights reserved.

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development found in migration an efficient method to exponentially increase the number of neurons in the brain and, at the same type, mix different cell types within the same volume of neural tissue. Changes in neurogenesis or neuronal migratory patterns enabled the emergence of novel neuronal circuits that boosted the computational capabilities in more complex brains and enabled more varied behaviors that provided bases for selection and further evolution. In this chapter, we will describe the evolutionary impact of the two main cellular movements on brain development, i.e., radial and tangential migrations. We review the cellular mechanisms that enabled different forms of radial migrations (somal translocation and glial-guided locomotion) and how these influenced brain elaboration. We also review the variety of functions displayed by tangentially migrating neurons in vertebrate brains, spanning from the orchestration of neurogenesis, setting up polarity of the developing cortex, and the wiring of distant brain areas. Radial migration (Fig. 25.1A) allows the separation of where neurons are born to their final location so that a higher number of neurons can be generated within a given sector of neuroepithelium (Rakic, 1972). This capacity can be further increased by pseudostratification and compartmentalization of the germinal zone with different progenitors aligning at different depths. Ian Smart and Henry Kennedy regarded this compartmentalization an important evolutionary mechanism to make the germinal zone more productive without “jamming” the various movements associated with neuronal generation (Smart et al., 2002). As neurons are born, neurons deriving from the same progenitors tend to maintain gap junctions that enable the early communication between them, allowing clonally related neurons to assemble into synaptically connected networks more likely than unrelated siblings (Yu et al., 2009, 2012; Ohtsuki et al., 2012). The resulting clustering of sibling neurons within the same neurogenic region potentiated the appearance of novel internal circuits. Therefore, elaborated mechanisms of radial migration paved the way to the production of larger and much more diverse and elaborately connected brains. Tangential migration, on the other hand, enabled the evolutionary expansion of cell type variability, boosting neuronal diversification (Fig. 25.1B). The embryonic brain hosts a variety of proliferative regions, demarcated by the singular expression of homeobox genes and other transcription factors (Puelles and Rubenstein, 2003). These combinatorial transcriptional codes provide genetic signatures that dictate the fate of neurons generated in a particular location and developmental stage (Lupo et al., 2006). Tangential migration opens new mechanisms to produce additional neuronal fates by allocating neurogenesis and differentiation to diverse locations at different periods of development (Marin and Rubenstein, 2003). The convergence and divergence of various neuronal and glial cells in particular locations enabled new and unique cellular interactions. Immigrant neurons find a different environment where they can differentiate and end up maturing into a new neuronal type different to both the resident neuron of its birth of place and the inhabitant neuron of the differentiating region (Wichterle et al., 2001). As such, tangential migration mixes neurons from several embryonic sources and increases cellular diversity. This diversification strongly impacts on circuit formation and complexity, changing their computational capabilities and making changes in behaviors possible. But, in addition, tangential migration also enables novel forms of cellular interactions not directly involved in circuit formation or information processing. Tangentially

FIGURE 25.1 The impact of neuronal migration on brain evolution. Radial and tangential migrations sculpt brain morphology during development. (A) Radial migration enabled neurons to leave the vicinity of the periventricular area. Top example (gray) shows the development of a brain area in which no radial migration occurs. Neurons remain clustered next to the ventricular area. Bottom diagram (black) shows how, when radial migration occurs, neurons occupy a thicker postmitotic zone, up to the pia matter. (B) Tangential migration enabled the mixture of different neuronal types within a given brain region. Top diagram (gray) shows a FMU occupied only by neurons born within the FMU extension. Bottom example shows the same locally born neurons (gray) intermingled with other neuronal type (black) developed upon arrival by tangential migration from a different FMU. FMU, fundamental morphogenic unit; GZ, germinative zone; MtZ, mantle zone.

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migrating, early-generated neurons could change early patterns of development and thus open up new developmental avenues, such as the promotion of neurogenesis by setting up polarity of inside-firsteoutside-last migratory pattern or providing the early template for the wiring of brain (Garcia-Moreno et al., 2018). We hypothesize that the contemporary vertebrate brains would assemble into more modest and less elaborate circuits if it were not for the evolutionary innovations brought by tangential migration.

25.2 The impact of radial migration on brain evolution Radial migration is defined as the process by which a cell moves away from the ventricular lumen toward the external portion of the neural tube. The ventricular zone can be divided into discrete subdivisions based on the unique combination of transcription factors. These subdivisions were termed fundamental morphogenic units (FMUs). Each FMU generates the neurons radially located above its germinative zone (Nieuwenhuys and Puelles, 2015) (Fig. 25.1A). Through radial migration, neurons end up in the same FMU where they were born and can be defined as locally born neurons. Remarkably, all locally born neurons share a common branding, a key genetic signature inherited from their original germinative zone. For instance, all neurons born in the pallium; the dorsal area of the alar plate in prosomere 4 (p4) is derived from pallial progenitors, which typically express a combination of Emx (Hevner et al., 2006) and Ngn genes (Gulisano et al., 1996; Frowein et al., 2002), among many others. This drives these neurons toward a glutamatergic differentiative pathway (Rakic, 1972; Mattar et al., 2008). Radial migration relies on the proliferative and guiding capabilities of a specific type of progenitor cell that is termed radial glia cell (RGC Rakic, 1971; Smart et al., 2002). These progenitors are established from neuroepithelial cells and become neurogenic. The early neuroepithelial cells and RGCs span the entire depth of the neuroepithelium from ventricular (apical) to pial (basal) surface. RGCs give risedeither directly or indirectlydto all neurons of the brain (Noctor et al., 2004; Kriegstein et al., 2006; Yu et al., 2009, 2012; Ohtsuki et al., 2012).

25.2.1 Evolution of radial migration Newly generated neurons leave the germinal zone in one of two modes: either by relocating their soma while they maintain their processes (somal translocation) or by using radial glia progenitors to climb to their intended location (glia-guided locomotion) as described in more detail in the following paragraphs. The relation of these two processes with RGCs puts these cells in a central position to understand the impact of radial migration on brain evolution, as they are major players in both the overproduction of neurons and their organization into functional clusters. In a simplistic view, the evolution of RGC moved brain complexity forward. The developmental influence of radial migration during brain development varies highly between regions of the CNS. As a rule of thumb, radial migration is more prevalent in the brain than in the spinal cord, its impact is deeper on rostral regions than caudal portions and accentuated in the alar plate versus the basal plate (Butler and Hodos, 2005). The impact of radial migration on brain morphology and physiology ranges across different groups of vertebrates. In some species, there are very limited migratory patterns, whereas others have very elaborated processes, which enable the delivery of neurons across long distances. Amniote brains usually display a higher degree of radial migration than nonamniote brains (Butler and Hodos, 2005). Among amniotes, the largest influence of radial migration is observed in mammalian and avian brains, whose neurons settle in brain areas far from the ventricular lumen. Among non-amniote species, teleost fish show the most extended use of radial migration. Chondrichthyan and amphibian brains, on the other hand, generally present a fairly reduced morphological impact of radial migration, although there are some exceptions in both vertebrate taxa (Puelles and Rubenstein, 2003; Nieuwenhuys et al., 2014). We will follow Butler and Hodos’ terminology in reference to a broad anatomical classification of vertebrate brains, i.e., laminar and elaborated brains (Butler and Hodos, 2005).

25.2.2 Radial migration on laminar brains Laminar brains are characterized by displaying most of their neurons in the vicinity of the ventricular lumen (Fig. 25.2), resulting from limited or absent neuronal radial migration during development (Lupo et al., 2006; Nieuwenhuys et al., 2014). Neurons in these brains extend their dendrites toward the meninges (i.e., basal surface), generating an outer fibrous layer rich in afferents and axodendritic connections (Fig. 25.2B). Two other main features define laminar brains: reduced number of neurons and lack of secondary neuronal organization (Fig. 25.2C). The developmental production of neurons is low and slow throughout the whole neuraxis. Consequently,

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FIGURE 25.2 Lack of radial migration in laminar brains. (A) During embryonic development, RGCs move their nuclei following interkinetic nuclear movement. Newborn neurons are born next to the ventricular lumen. These neurons are then passively pushed away. (B) As a development result, the neurons in the adult brain reside just on top of the germinative zone, in the stratum griseum. Their dendrites form the stratum album, near the pia matter. (C) A stereotypical laminar brain: Nissl staining on a coronal section of the urodele Pleurodeles waltl at the level of the telencephalon, dorsal at the top. RGC, radial glia cell. From Moreno, N., González, A., 2007. Regionalization of the telencephalon in urodele amphibians and its bearing on the identification of the amygdaloid complex. Front. Neuroanat. 1:1. Abbreviatures from the reference.

laminar brains host the lowest number of neurons among vertebrate brains. And due to the lack of secondary neuronal organization, most of the gray matter in laminar brains is not arranged into cortices or nuclei. These two features are directly related to the limited action of radial migration. However, there are structural exceptions to this simplicity within laminar brains. These exceptions follow the general increase in radial migration taking place in rostral and alar regions of the CNS across vertebrates. For instance, the tectum mesencephali of anurans presents substantial neuronal lamination, with its neurons covering about two-thirds of the neuroepithelial thickness. Different authors distinguish from 6 to 15 neuronal layers in such areas, depending on neuronal density, pattern of afferents, or peptide immunohistochemistry (Roth et al., 1990; Nieuwenhuys et al., 2014). This complex organization of the mesencephalic alar plate can only be achieved by well-developed, stereotyped, and organized radial migration mechanisms from the germinative zones in contact with the fourth ventricle. Among vertebrates, the taxa that typically present laminar brains are among cyclostomes (lampreys), chondrichthyans (squalomorph and squantinomoprh sharks), non-teleost actinopterygians, and amphibians (Butler and Hodos, 2005). The most studied and representative species for laminated brain species are amphibians, specifically, the clawed toad Xenopus laevis and the green frog Rana esculenta among anurans, and the salamander Ambystoma tigrinum as a typical urodele (for review, see Wichterle et al., 2001; Nieuwenhuys et al., 2014). It is in the latter group where the most exacerbated features of laminar brains appear, and the structural austerity is more prominent. In fact, urodelian brain cytoarchitecture seems simpler than the position of this taxon within the vertebrate radiation predicts (Roth et al., 1993; Garcia-Moreno et al., 2018). Most teleost fish brains display higher complexity, increased neuronal numbers, and elaborated neuronal distribution when compared with amphibian brains. This led to suggest that the amphibian brain has undergone an evolutionary secondary simplification. A typical urodelian brain presents a dense central gray layer closely associated with the ventricular surface (Fig. 25.2C). This is a common feature throughout its entire CNS caused by a near absence of radial migration of neuroblasts that leads to its characteristic layering. When embryonic neurons are born, these do not move beyond the ventricular area of influence. As originally described by Herrick (1948), starting from the ventricular lumen, cells are distributed in the ependymal zone, the subependymal zone, the aforementioned stratum griseum or central gray layer and the stratum album (Fig. 25.2B). Apart from those classic descriptions, NeuN immunostaining specific of neuronal bodies shows the same neuronal distribution (Nieuwenhuys and Puelles, 2015; Saito et al., 2017). And the same pattern of staining, crowded at the periventricular neighborhood, is found when other classic markers are studied, such as calretinin, calbindin, GABA, or Islet1 (Hevner et al., 2006; Moreno and González, 2007; Morona and González, 2008). A lack or disruption of radial glia cannot account for the described diminished radial migration in laminar brains as they all derive from brains densely populated by RGCs, including amphibians. It is interesting to note that in some of these species, such as Xenopus, RGCs are morphologically identical to those observed in mammals. The similarities extend beyond the shared protoplasmic extension from ventricular to pial surfaces as the expression of crucial markers for the function of RGCs, such as vimentin and glial fibrillary acidic protein (Dent et al., 1989; Messenger and Warner, 1989), is conserved. Delta-Notch signaling necessary for the formation of the radial glia seems conserved among vertebrates, as overactivation of Notch signaling in Xenopus larvae stimulated the quantitative and qualitative development of radial glia (Kiyota et al., 2008). This conservation of RGCs suggests that the reduced radial migration in laminar brains is not a

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consequence of a lack or divergent formation of the radial glia scaffold. On the contrary, the periventricular distribution of laminar brain neurons might be accounted for by either (1) a diminished neuronal production and/or (2) a simpler RGC-toneuron molecular communication that does not lead new neurons away from their site of origin. Another relevant difference in amphibian radial glia is its persistence throughout the animal’s life (Dent et al., 1989; Gervasi et al., 2000). However, being this feature shared by other vertebrate taxa and by some elaborated brains, it is not possible to link a longlasting radial glial population to reduced radial migration brain architecture. Reduced neuronal production accompanied by a lack of or diminished radial migration generates the simplest brains across the vertebrate phylum. As an anatomical consequence of this formation program, laminar brains host their neurons without any secondary structure, a poorly developed configuration that restricts neuronal circuitry complexity and diversity. These circuits appear computationally limited, eventually explaining the simple behavioral repertoire of laminar brain species.

25.2.3 Radial migration on elaborated brains Elaborated brains are characterized by a complex distribution of neuronal bodies away from the periventricular region (Fig. 25.3). Neurons are found along the entire radial axis, from the ventricular lumen up to the meningeal surface as a consequence of the active neuronal radial movement that takes place during embryonic development: Neurons differentiate and move away from the periventricular surface to positions that cannot be predicted from the site of origin (Butler and Hodos, 2005). During the differentiative stage, neurons extend axonal and dendritic communications in multiple directions, not restricted by the polarity of the layered tissue. Two other major cytoarchitectonic features distinguish elaborated brains from laminar brains: increased number of neurons and substantial aggregation of neurons in secondary structures such as cortical layers or nuclei. Elaborated brains host the largest amount of neurons across the tree of life, resulting from increasing neurogenic gradients from spinal cord toward anterior regions of the brain and from basal to alar plate derivatives. Remarkable examples are found in the telencephala of passerine birds and primates, which accommodate huge amounts of densely packed neurons, or in the astonishing cerebellum of mammals. The cerebellum of the African elephant Loxodonta africana is formed by up to 250 billion neurons, a total of 97.5% of the neurons in the elephant brain (Herculano-Houzel et al., 2014).

FIGURE 25.3 The impact of radial migration in elaborated brains. (A) During embryonic development, newborn neurons (black) are born next to the ventricular lumen. These neurons move radially and abandon the germinative zone, generating an embryonic postmitotic stratum, the mantle zone. (B) In the adult, the mantle zone has further developed into a dense neuronal tissue, whose neurons occupy the entire radial depth of the neuroepithelium. (C) A stereotypical elaborated brain: Nissl staining on a sagittal section of the bird Gallus gallus, anterior at the left (from brainmaps.org). Note the abundant secondary structures of neuronal organization: multiple nuclei at the dorsal ventricular ridge (DVR) of the telencephalon, cortical layers at the cerebellum (Cb).

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In addition, elaborated brains display neurons in a wide range of structures, spanning from multilayered cortices to complex associations of brain nuclei. These structures are present throughout the CNS and cover the whole depth of the neuroepithelium, from the inner ventricular lumen to the most distant brain surface (Nieuwenhuys et al., 2014). These two features that characterize elaborated brainsdoverproduction of neurons and secondary structural organizationdrely on the action of radial migration. Elaborated brains can be found in species across the entire spectrum of vertebrates. Hagfishes among cyclostomes, some groups of chondrichthyans and teleosts present brains elaborated to a variable degree across their neuraxes (Butler and Hodos, 2005). However, it is in amniotes, and especially birds and mammals, where the highest degree of elaboration can be found. A representative example can be found in the telencephalon of birds, which reaches one of the highest neuronal densities across the animal kingdom (Olkowicz et al., 2016) (Fig. 25.3C). Millions of neurons are distributed in a densely populated mantle zone that obliterates the ventricular lumen during embryonic development. These neurons organize their soma in a number of derived structures, mainly nuclei that vary in size, density, and position within the mantle zone (Puelles et al., 2007). Columnar arrangements of neurons, as described in the hyperpallium or in laminated structures such as the auditory field L of the dorsal ventricular ridge, are other examples of the elaboration of neuronal bodies within the avian telencephalon (Reiner et al., 2005). These structural features ultimately help determine the connectivity and function of a given brain area, connecting it to other regions in a stereotyped fashion. Another striking example of neuronal elaboration is the mammalian dorsal thalamus. This relatively small brain region comprises a set of nuclei, up to 40 according to some authors, each of them hosting several neuronal types. Other significant singular neuronal aggrupation are the Calleja islands of the mammalian olfactory tubercle, the highly laminated cortices of the cerebral cortex, cerebellum, and tectum mesencephali across vertebrates, the glomeruli in the mammalian olfactory bulb, among others. The most elaborated vertebrate brains of avian and mammalian taxa lack RGCs in their adult form, as these cells disappear at the end of brain development (Kriegstein et al., 2006). At the final stages of brain formation, once neurons and astrocytes have been generated, RGCs retract their basal processes, indicating the end of the radial migration process. However, as already mentioned, there is no direct correlation between the absence of radial glia in the adulthood and the elaboration of brains. On the contrary, adult teleost fish brains maintain radial glial populations throughout the CNS. Reptilian brains of testudine and lepidosaurian species (turtles and lizards) also keep some subpopulations of RGCs in selected regions of the brain (Garcia-Verdugo et al., 2002). Although RGCs in adult brains seem to play a role in adult neurogenic processes to a limited extent, their persistence is linked to neuroregeneration events (Garcia-Verdugo et al., 2002; Zupanc and Clint, 2003). In any case, the persistence of RGC in adulthood is unrelated to the general structure of the brain. The structural products of radial migration during development are highly populated brains whose neurons aggregate and show a high level of connectivity. The functional capabilities of these complex circuits have hugely diversified during vertebrate evolution, enabling the expansion of the behavioral portfolios observed in amniotes. As a consequence, elaborated brain species have become capable of adapting to and conquering every possible environment. At the brain level, it was an augmented and structured process of radial migration that set the first stone for this biological success.

25.2.4 The influence of radial migration on pallial internal circuitry The mammalian neocortex is perhaps the most studied brain structure. Anatomically, it is the main structure derived from the mammalian pallium. The neocortex lies at the top of the mammalian CNS hierarchy and is responsible for our higher cognitive functions such as language, retaining our memory of the past, planning for the future, and appreciating or producing science and art. Neocortical computational efficacy is based on a canonical circuitry that acts as a functional unit and is seen across cortical areas (Geschwind and Rakic, 2013; Harris and Shepherd, 2015), alongside countless variations on this common theme and endless intracortical communications. The canonical circuit connects neurons within a radial column spanning from the deep cortical white matter to the outermost cortical layers beneath the pial surface. Remarkably, the existence and construction of this unitary cortical brick is intimately related to radial migration (Noctor et al., 2001). Here, we review how the complex mechanisms of radial migration evolved to enable the appearance of the current neocortex. The unique neocortex of mammals exemplifies the major impact that radial migration can have on brain structure and function. It derives during embryonic development from the dorsal region of the pallium, one of its four FMUs (Medina and Abellán, 2009). Locally born dorsal pallial neurons will eventually become the glutamatergic pyramidal neuron population. Once they are born in the periventricular region, at either the ventricular or subventricular zones, these neurons

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FIGURE 25.4 The consequences of somal translocation and glial-aided locomotion. (A) Somal translocation. A newborn neuron inherits a basal process in contact with the meningeal surface (1). After mitosis, the newborn neuron (black) is released from the apical contact at the ventricular surface. The soma is pulled up from the pia mater contact by somal translocation (2). The neuron abandons the germinative zone and occupies the mantle zone stratum (3). (B) Glial-aided locomotion. Later in development, the newborn neuron (black) does not inherit the basal process but passes through a transient multipolar stage when it is detached from any superficial contact (1). After a multipolar-to-bipolar transition, the neuron associates to a radial glial extension (2). Actively climbing the radial process, the neuron ascends to the mantle zone, skips over the previously born neurons, and settles the outermost region of the neuroepithelium (3). (C) In species such as sauropsids (top scheme), somal translocation is the only radial movement. As a consequence, the neurons locate the tissue following and outside-in gradient of neurogenesis. In selected regions of the mammalian brain (bottom scheme), glial-aided locomotion inverts the neurogenic gradient, and late-born neurons occupy the outermost regions of the neuroepithelium.

migrate radially toward the meningeal surface (Kriegstein and Noctor, 2004) (Fig. 25.4). This migration achieves two major advantages: first, it releases the germinative zone of cellular pressure (the “jamming” of the ventricular zone (Smart et al., 2002) (Miyata, 2015), and second, it separates neuronal progeny from the periventricular space. Combined, these features enable a larger production of cortical neurons and their subsequent organization into the stereotypical cortical layers. Current models of cortical neuroblast development describe two stages of radial migration, i.e., somal translocation and glial-aided locomotion (Fig. 25.4). Somal translocation seems to be a conserved mechanism among elaborated brain species (Nadarajah et al., 2001; Nadarajah and Parnavelas, 2002). But it is possible that neuronal locomotion is a mammalian novelty, and it could account for the structural and functional singularities observed in the mammalian dorsal pallium (Hatanaka et al., 2004).

25.2.4.1 Somal translocation During the early stages of neocortical development, the pallial neuroepithelium hosts RGCs that maintain contact with both inner (ventricular or apical) and outer (basal or pial) extremes of the tissue (Nadarajah and Parnavelas, 2002). As it divides, RGCs maintain ventricular contact after mitosis, but the daughter neuron or the intermediate progenitor does not (Fig. 25.4A). Thus, the newly born neuron inherits a basal process in contact with the pia but is free from the ventricular constraint (Nadarajah and Parnavelas, 2002). In a cellular process termed somal or nuclear translocation, the anchored basal process pulls the neuronal soma up from the periventricular area so that the neuron migrates radially to colonize the postmitotic area of the neuroepithelium (Fig. 25.4A). Somal translocation is responsible for the formation of a distinct secondary stratum in the neural tube, the mantle zone. During embryonic neurogenesis, the mantle zone is populated only by postmitotic neurons, and it becomes the mature CNS tissue in the adult brain. Somal translocation is the predominant mode of migration of early-born populations of cortical neurons such as those destined to the preplate and the deepest cortical layers. At these early stages, the mantle zone is a thin neuronal layer, and it is common for newborn neurons to remain attached to the pia shortly after mitosis. At later stages of neurogenesis, somal translocation appears to occur less frequently (Miyata et al., 2001; Nadarajah and Parnavelas, 2002; Miyata and Ogawa, 2007). Somal translocation has been suggested as an evolutionary ancient mode of radial migration given the development of pallium in sauropsids occurs in a homologous fashion (Tsai et al., 1981; Goffinet et al., 1986; Nadarajah and Parnavelas, 2002). Newborn neurons in the developing pallium of birds and reptiles accumulate outside germinative areas after a brief radial movement that resembles the process of somal translocation described earlier. Waves of newly generated neurons

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gather at the apex of the mantle zone, lying deeper to previously born neurons (Tsai et al., 1981; Goffinet et al., 1986). Each new generation of neurons pushes the preexisting mantle zone away slightly. As a consequence, the earliest populations occupy positions near the pial surface in the adult brain, a long distance from the ventricular lumen in some regions of the pallium, as is the case with the ventral pallium (García-Moreno et al., 2018). However, their active migratory movement covers only a small portion of that distance. Neurons born at midneurogenic stages tend to occupy an intermediate position within the mantle zone. And the latest born populations, which are not pushed away by any other new neuronal batch, differentiate near the periventricular area, close to their origin site (Fig. 25.4C). Therefore, the location of a given neuron within the tissue reveals the gradient of neurogenic timing, termed outside-firsteinside-last, or the outside-in gradient of neurogenesis (Tsai et al., 1981; Goffinet et al., 1986). This gradient not only reflects the time when neurons are born but also dictates the internal circuitry the neurons communicate through. The outside-in gradient of neurogenesis impacts directly onto the function of the sauropsidian brain. Somal translocation, or homologue type of cellular movement, is responsible for the elaboration of more complex brains. Once a neuron is born, it detaches from the ventricle and, regardless of an active pulling from a potential pial contact, the newborn neuron naturally leaves the ventricular zone. This germinative area is densely occupied by cycling nuclei, and thus, a newborn cell that no longer needs ventricular contact finds it more attractive to occupy the looser mantle zone. Therefore, we hypothesize that somal translocation enabled the increase of neuronal numbers and their organization in numerous secondary structures and consequently enabled the appearance of modern vertebrate brains. However, a second-type radial migration movement would come to expand these elaborate structures and promote the creation of more intricate and complex neuronal circuitry, pushing them a step further in evolution.

25.2.4.2 Glial-guided locomotion The embryonic neocortex becomes thicker as new waves of cortical neurons are added to the mantle zone. Consequently, the radial process of RGCs extends far away from the thin germinative zone where their nuclei cycle. This major extension of the radial process (Nowakowski et al., 2016) makes it impossible for a newborn neuron to inherit a contact with the superficial meninges after its generation. In addition, many neurons are born in the subventricular zone during the second half of corticogenesis (Hansen et al., 2010; Kelava et al., 2012; Nonaka-Kinoshita et al., 2013; Vasistha et al., 2015). Many progenitors in this secondary proliferative stratum do not make contact with either apical or basal surfaces. Altogether, a typical somal translocation seems impossible for the majority of cortical neurons originated at intermediate and late stages of neurogenesis. If that is the case, how newborn neurons travel away from the germinative zones then? Shortly after mitosis, a newly generated neuron extends a short process directed toward the basal surface (reviewed by Cooper, 2013). This process develops in close contact with one or several RGCs’ basal processes. Following consecutive minor nuclear stretching and process extensions, the cell literally climbs up the glial rope toward the pia (Fig. 25.4B), following the path of the glial scaffolddthis movement is referred to as glial aided. And as the neuron is not merely pushed away by progenitors but actively climbs up, this neuronal movement is termed glial-aided locomotion (Ayala et al., 2007; Metin et al., 2008). At the neurogenic stages in which glial-aided locomotion occurs, there already is a developing mantle zone populated by the earliest born neurons and through which new neurons need to passethe glial-aided movement enables them to move beyond those previously established layers of postmitotic cells. Unlike somal translocation, cells migrating by locomotion skip over older neuronal populations, moving radially to the vicinity of the meningeal surface and settling in their destined position to differentiate (Nadarajah and Parnavelas, 2002). This true innovation inverts the gradient of neurogenic timing, which becomes inside-out (Angevine and Sidman, 1961). The neocortex of mammals displays the earliest born neurons at the deeper layers (inside-first) and the latest generated ones at the cortical surface (outside-last) (Fig. 25.4C). This new position of the neurons within the mantle zone provided a substrate for a change of the internal circuitry, a major impact of radial migration on brain evolution. Clonally related sibling neurons tend to connect more likely between themselves than nonrelated ones (Yu et al., 2009, 2012; He et al., 2015). Glial-aided locomotion promoted a significant increase in the production of neurons within the radial sector, allowing these neurons to communicate more extensively. Thus, this new mechanism of radial migration led to the elaboration of the canonical cortical circuit (Bosman and Aboitiz, 2015). The trisynaptic canonical circuit is considered the basic fundamental processing microcircuit of the neocortex and the basis for neocortex computational efficiency (Kaas, 2012; Harris and Shepherd, 2015). And it is remarkable that this substantial innovation was aided by glial-guided locomotion, a new type of cellular movement. Although similar RGCs are found in avian and reptilian brains, glial-guided locomotion has never been observed, and only somal translocation has been attested in the pallium of sauropsids. Given this, their pallium shows an outside-in distribution of neurons. This gradient follows the convoluted shape of RGCs, which in these brains typically span from

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internal positions in the caudal brain toward the meningeal surface at more rostral levels (Striedter and Beydler, 1997). By means of clonal analysis of cellular lineages in the chick, it was shown that cells derived from the same early RGC distribute in caudorostral columns (Szele and Cepko, 1996). Consequently, pallial neurons build their internal circuitry following a pattern different to that of the mammalian neocortex. They still form a trisynaptic circuit, analogous to the canonical cortical circuit (Karten, 1969, 2013; Dugas-Ford et al., 2012), but this might not be a by-product of radial migration given that its constituent neurons are unlikely to be developmentally or clonally related (Garcia-Moreno et al., 2018). Glial-aided locomotion is unique to mammalian pallial development, leading to the rearrangement of cortical circuitry to perform novel functions. This arrangement allowed the establishment of a platform below the developing cortical plate, the subplate. This neuronal population receives early afferents at the time when the cortical plate neurons are not even born, allowing the assembly of a transient circuitry that later helps set up mature circuits once the cortical plate is formed (Hoerder-Suabedissen and Molnar, 2015). Therefore, understanding how glial-guided locomotion evolved in mammalian development is key to comprehend neocortical evolution.

25.2.4.3 Evolutionary origin of glial-aided locomotion The developmental mechanisms driving radial locomotion have been the object of intense research focus for decades (Cooper, 2013). Comparative approaches aiming to describe the divergences of these mechanisms between vertebrates pointed very early toward a single molecule, reelin, as crucially responsible for the mammalian novelties. Reelin is a secreted and diffusible protein, which in the mammalian developing pallium is highly expressed at the brain surface (Ogawa et al., 1995; Meyer et al., 1999). In reelin-deficient, “reeler” mice, newborn cortical plate neurons fail to perform locomotion. Migratory neurons are unable to climb over the intervening neurons, resulting in a grossly inverted cortex with an outside to inside pattern (Katsuyama and Terashima, 2009). This arrangement resembles what is observed in the sauropsidian pallium. The migratory mechanism triggered by reelin requires the receptors VLDLR and ApoER2 and the intracellular adapter Dab1 by cortical plate neurons (reviewed by Chai and Frotscher, 2016). Mutant mice deficient for each of these molecules suggested the relevant roles of reelin for glial-aided locomotion at several levels: It helped cortical plate neurons attach to the glial scaffold, promoted their radial movement, and maintained RGC morphology and function. Comparative aspects of reelin expression turned its developmental interest into evolutionary relevance (Tissir et al., 2002). Reelin is expressed in the brain of all vertebrate species studied so far, including lamprey, teleost fish, amphibians, etc. (Bar et al., 2000). However, the magnitude of its expression at the pallial pial surface is increased several fold in mammalian brains, the only species where radial locomotion is observed (Bernier et al., 1999, 2000). This pointed toward a straightforward hypothesis (Fig. 25.5) that tried to explain cortical evolutionary origin by means of a novel substantial increase in reelin signaling at the brain surface (Bar et al., 2000). However, the appearance of the six-layered neocortex cannot be accounted for by just one single change at the level of expression of any molecule. As shown by avian in ovo experiments, an indirect experimental increase of reelin signaling in quail embryos did not modify the neurogenic distribution of pallial neurons (Nomura et al., 2008a; b). No glial-aided locomotion was found, and pallial neurons remained located following the usual outside-in gradient of distribution. Reelin overexpression, however, led to the straightening of radial glial fibers and the conversion of multipolar pallial neuroblasts into mammalian-like bipolar ones (Nomura et al., 2008b). These data strongly support a relevant role for reelin in cortical origin, but several crucial aspects of such a remarkable evolutionary event seem unrelated to reelin; for instance, the abrupt increase of pallial neurogenesis observed in the mammalian pallium compared with reptilian brains, the appearance of intracortical callosal connections (Suárez and Richards, 2014; Garcia-Moreno and Molnár, 2015), or the novel reception of collothalamic afferences. We still miss a global hypothesis that can explain the several singularities of the neocortex, which, most likely, all appeared in a short period of evolutionary time.

25.3 The impact of tangential migration on brain evolution Tangential migration describes the process by which a cell departs from the FMU where it was born, moves a variable distance, crosses developmental boundaries, and reaches a different FMU where to differentiate and settle (Fig. 25.1B). Tangential movement occurs parallel to the ventricular and meningeal surfaces and in every possible direction (following rostrocaudal and/or dorsoventral orientations). In terms of development, neurons following tangential migration are found in a different FMU to which they were originated, and therefore, we term them as foreigner neurons. As every other neuron, tangentially migrating neurons inherit a key genetic signature from their original environment (Flames et al., 2007; Anastasiades and Butt, 2011). As neurons travel tangentially carrying this birthplace information, tangential neurons

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FIGURE 25.5 Reelin in the evolution of the neocortex. (A) During the embryonic development of the pallium of sauropsids, there is a low reelin signaling at the marginal zone (gray shade). Newborn neurons do not respond to this signaling and generate the dorsal pallium through somal translocation. Therefore, late-born neurons (black) end up at the deeper mantle zone, near the ventricular surface. (B) Scheme of the nuclei and circuitry within the avian dorsal ventricular ridge. Neurons from entopallium (EPall), dorsal nidopallium (dNPall), and arcopallium (APall) communicate through a neuronal circuit, which is not directly associated to the neuronal birthdate (Tsai et al., 1981, unpublished data). (C) On the other hand, a strong, marginal reelin signaling (gray shade) rules cortical development in mammalian species. The radial glial palisade straightens due to reelin signaling. Newborn neurons at the multipolar stage are capable to sense and respond to the secreted reelin. As a consequence, these neurons climb radially by glial-aided locomotion and invert the neurogenic gradient. Late-born neurons (black) occupy the outermost layer of the neocortex. (D) The inverted location of the pallial neurons, and the presence of the subplate population, reassembled the pallial circuit. In mammalian brains, the cortical canonical circuit is a direct by-product of its columnar development. 1e6, cortical layers; dTh, dorsal thalamus; SP, subplate layer.

display a different key genetic signature to that of the neurons locally born at the differentiative FMU. This is a crucial feature of tangential migration. Due to this intermingle of different neuronal sources, the tangential neuron differentiates into a new cellular type, dissimilar to both the locally born resident neurons, and those differentiated in its original FMU that underwent radial migration. There are examples of tangential migration for glutamatergic and GABAergic neuronal populations. For instance, many GABAergic neurons born in the subpalliumdthe ventral area of the alar plate in prosomere 4 (p4)dmigrate into the pallium by tangential migration (de Carlos et al., 1996; Marin and Rubenstein, 2003). Subpallial progenitors typically express a combination of Mash1, Dlx, and Gsh genes, among many others, leading the derived neurons toward a GABAergic differentiative pathway (Anastasiades and Butt, 2011; Mayer et al., 2018). Once in the pallium, these tangential neurons become pallial interneurons, a cellular type clearly different to both the glutamatergic projecting neurons locally born in the pallium and the GABAergic projecting neurons radially differentiated in its original subpallium (Wichterle et al., 2001). Populations of tangential migratory neurons have been described all along the CNS (Cooper, 2013), and the influence of tangential migration on brain development varies between CNS regions. However, it remains difficult to describe a general trend for this variation. Although a larger number of these movements have been observed in the telencephalon, it is likely that this is only due to a deeper focus by researchers on this area. The reality is that some form of tangential

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migration has been discovered in almost every brain region where it is looked for. And potentially, this type of cellular movement is the developmental cause behind the emergence of most interneuronal cell types of the brain, if not all. The influence of tangential migration on brain morphology and physiology varies across groups of vertebrates. Based on a complexity criterion, we suggest that elaborated brains are formed largely under the developmental influence of tangential migration, as opposed to laminar brains. Indeed, a larger number of tangential populations are known in mammalian brains, the paradigm example of elaborated brain species. However, it is possible that the general emphasis on mammalian research generates a false impression that tangential migrations influence more strongly mammalian brain development. In fact, most instances of tangential migrations first described in mammalian brains were later found to occur in the development of other vertebrate brains. Accordingly, we support a general view of vertebrate brain development in which all taxa and brain regions host foreigner populations that migrated tangentially, but with the degree in which tangential migration has influenced them varying between them. The promotion of new cellular types triggered by tangential migration provides invaluable raw material for evolutionary tinkering. As evidenced in the paragraphs in the following, tangential migration allowed novel cellular types to perform new cellular functions such as the modulation of neuronal circuits, developmental regulators, and axonal pathfinders. But, more importantly, it led to a boost in neuronal diversity: the ultimate factor for brain network complexity.

25.3.1 Pallial interneurons and the modulation of brain circuits The telencephalic alar plate is subdivided into two principal FMUs: pallium and subpallium, clearly separated by a morphogenic boundary termed the pallialesubpallial boundary (Puelles et al., 2017). Pallial and subpallial gene expression programs are divergent and lead to the production of glutamatergic and GABAergic neurons, respectively (Lodato and Arlotta, 2015). Neurons born in each of these regions migrate radially and generate the mature pallial and subpallial derivatives. But also, a substantial number of subpallial neurons migrate away from the subpallium, crossing the pallialesubpallial boundary and settling in the pallium (Fig. 25.6A). These subpallial-derived cells become GABAergic neurons in an overall glutamatergic environment. And as a functional novelty, these foreigner neurons establish short-range connections with local neurons, thus becoming pallial interneurons (Wichterle et al., 2001; Mayer et al., 2018; Tosches et al., 2018).

25.3.1.1 Conserved features of tangential migration of pallial interneurons in vertebrates Tangential migration of cortical GABAergic interneurons was first described in mammals following in vivo injections of cellular trackers (de Carlos et al., 1996; Anderson, 1997). Given all vertebrate brains host GABAergic neurons in the pallium, there were early suggestions that the tangential origin of pallial interneurons was a conserved feature of all

FIGURE 25.6 Tangential migration of pallial interneurons. (A) Schematic coronal section of the telencephalon of a stereotypical vertebrate brain. GABAergic neurons (black) born at the subpallium (SPall) migrate tangentially toward the pallium. These migratory neurons avoid entering the striatal primordium (St) and, after a tangential journey, differentiate in all areas of the pallium. (B) The observed migratory pattern of pallial interneurons (black) through the mantle zone of the avian pallium. GABAergic neurons occupy the entire depth of the postmitotic zone during migration. (C) The migratory trajectory of pallial interneurons in mammalian brains is restricted to two segregated corridors at the boundaries of the mantle zone: the lower intermediate zone (IZ) and the marginal zone (MZ). Later on, migratory neurons are capable of finding molecular cues to enter radially in the cortical plate (CP) and assemble to cortical circuits. (D) The canonical columnar circuit in the mammalian neocortex, modulated by stereotypical pallial interneurons (black). The diagram exemplifies modulation by feedforward inhibition (FFI), feedback inhibition (FBI), and multiscale inhibition (MSI). dTh, dorsal thalamus; GZ, germinative zone; MtZ, mantle zone; PSB, pallialesubpallial boundary.

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vertebrate taxa. Indeed, tangential migration of subpallial neurons has been found in every vertebrate species studied so far. A few years after the first studies in mouse and rat embryos, equivalent cell tracking experiments and xenograft quail-tochick in ovo transplants showed homologous tangential migration in avian species (Cobos, 2001; Tuorto et al., 2003). Later on, in ovo experiments based on cell trackers depicted the same GABAergic neuron movement in Emys orbicularis turtle embryos (Metin et al., 2007). And eventually, the same results were described in amphibian, teleost fish, and even chondrichthyan species (Carrera et al., 2008; Moreno et al., 2008). Altogether, these results strongly suggest the homology of pallial interneurons across species given the wide conservation of pallial interneuron tangential migration among the vertebrate phylum. The pallium of the vertebrate common ancestor, who lived over 500 mya, already possessed subpallial-originated GABAergic interneurons. Evidence for homology goes beyond the histological location and neurotransmitter fate of the pallial interneurons. The singular germinative zone within the subpallium that gives rise to the interneurons has also been observed homologous in every species studied in the necessary detail (Puelles et al., 2000; Flames et al., 2007; GarciaMoreno et al., 2018). In the case of mammalian brains, interneurons are born in the ventricular and subventricular zones of the pallidal subpallium, namely, the medial and caudal ganglionic eminences (MGEs and CGEs, respectively) (Flames et al., 2007; Anastasiades and Butt, 2011). These structures are termed eminences due to their protrusion toward the ventricle. Non-mammalian species display less prominent subpallial protrusions, but the same molecular domains appear during embryonic neurogenesis. These pallidal domains in the ventral subpallium express crucial combinations of the transcription factors NKX2-1, DLX2, GSH2, LHX6, COUPTF1, among many others in all vertebrate species (Moreno et al., 2009). Therefore, interneurons are born at the homologous brain regions in all vertebrates, confirming that pallial interneurons are homologous within the phylum.

25.3.1.2 Divergence in tangential migratory routes of pallial interneurons The origin and fate of pallial interneurons, and part of their migratory mechanisms, are deeply conserved from the vertebrate roots. Transplanting turtle ganglionic eminence cells into embryonic mammals and, conversely, mammalian ganglionic eminence cells into embryonic turtle revealed that mammals and reptiles share the same molecular machinery for tangential migration (Métin et al., 2007). Also, when young interneurons from several vertebrate species were transplanted into the murine ganglionic eminence, they migrated following a conserved tangential path toward the dorsal pallium (Tanaka et al., 2011). However, the migratory route and mechanisms followed by these cells show some interesting divergences. Only the mammalian neuroblasts were capable of invading the postmitotic layers of the murine dorsal palliumdnamely, the cortical plate. Chick and turtle interneurons found the way up to the neocortex but rarely entered within the mantle zone. This finding led to the suggestions that additional cues for interneuronal migration evolved within the mammalian lineage. This suggestion fits an alternative finding regarding the precise corridors followed by interneurons. Mammalian interneurons migrate toward the dorsal pallium by two main corridors flanking the cortical plate: the fibrousrich intermediate zone and the superficial marginal zone (Jimenez et al., 2002). In the case of birds and reptiles, interneurons migrate through the pallium covering the whole depth of the mantle zone and find no molecules preventing their contact to pallial maturing neurons (Fig. 25.6B and C) (Garcia-Moreno et al., 2018). Altogether, these findings depict an evolutionary scenario in which all vertebrate interneurons are born within the pallidal subpallial germinative zones, and most of them migrate dorsally through the postmitotic mantle zone. In the singular case of mammalian brains, the sole exception within vertebrates, the dorsal pallial mantle zone communicates to the interneurons by means of a novel and yet unknown mechanism. This prevents the early interneuronal invasion of the cortical plate and ultimately helps with the time and spatial even distribution of interneurons within the pallium.

25.3.1.3 Diversifying complexity of GABAergic subtypes Beyond the elaboration of the migratory mechanism, mammalian interneurons show other signs of diversification. Cortical interneurons have been traditionally classified in three main types based on their birthdate, cortical location, and expression of marker genes (Tremblay et al., 2016; Mayer et al., 2018). Single-cell RNA sequencingdcurrently the most powerful technique to reveal cellular typesehas revealed the same three main types of pallial interneurons in the pallium of two reptiles: the turtle Trachemys scripta and the lizard Pogona vitticeps (Tosches et al., 2018). This finding provides a strong argument in favor of the conservation of interneuron types across amniotes. But Tosches et al. (2018) also show that the multiple subtypes of cortical interneuron cell types are absent in the reptilian pallial derivatives. Accordingly, mammalian brains evolved a larger diversification of pallial interneurons from the same three main types as a starting point. And so, the influence of tangential migration caused a deeper impact on mammalian pallial evolution.

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The existence of pallial interneurons, and specially their stunning diversity, caused a major shift in pallial function: Interneurons modulate pallial circuits to achieve superior networking capabilities (reviewed by (Tremblay et al., 2016)). All pallial neurons connect to each other by excitatory glutamatergic neurotransmission. When interneurons added inhibition to the pallial circuitry, it prompted three main computational advantages: a graded modulation of the pallial output, and increased internal complexity, and a brake to the excitatory response (Fig. 25.6D). A good example of these modulation mechanisms is the feedforward inhibition (Simons and Carvell, 1989). When thalamocortical input reaches a pyramidal neuron, it also reaches a GABAergic interneuron, which subsequently inhibits the thalamic-recipient pyramidal neuron by GABA communication. The pallial neuron finds its activation modulated as it is inhibited shortly after the original activation. Other examples of GABAergic modulation are the feedback inhibition and the disinhibition (reviewed in Tremblay et al., 2016). Especially in elaborated brains where pallial interneurons further diversified over evolution, the modulation of pallial circuits had a remarkable impact on brain evolution. Altogether, the pallium of vertebrates got a significant boost in its processing capabilities thanks to the arrival of foreigner subpallial neurons by tangential migration. Remarkably, the pallium orchestrates the high-order sensory information processing of the vertebrate CNS. The modulated improved circuitry promoted further this top hierarchical position, so telencephalic development was enhanced over vertebrate evolution. As a biological circle of virtue, such a benefit as the sensational one caused by interneuronal modulation was easily translated into an evolutionary advantage. Interneuronal tangential migration further triggered the expansion of the vertebrate behavioral repertoire and, as a crucial consequence, enabled vertebrate ecological success.

25.3.2 Glutamatergic tangential contributions as developmental scaffolds The pallium of vertebrates is subdivided into four FMUs, which differ in their expression of transcription factor encoding genes such as Emx1, Dbx1, and Lhx5, among many others (Medina et al., 2004). From the midline toward the lateral portions of the neural tube, these pallial subdivisions are termed the medial, dorsal, lateral, and ventral pallia (Puelles et al., 2017). The vertebrate dorsal pallium processes diverse types of information, including sensation from visual, auditory and somatosensory modalities, motor control, and high-order cognitive processes. And as a major divergence within the mammalian lineage, the dorsal pallium derives an exquisite laminated structure termed the neocortex or isocortex. Due to the astonishing relevance of neocortical lamination for mammalian brain functioning (Harris and Shepherd, 2015), great efforts are conducted to the understanding of how this layering occurred in development and evolution. Both radial and tangential migration impacted brain evolution; in fact, early tangential migrations of glutamatergic neurons that are not seen in sauropsids might have played a key role in the origin of mammalian neocortex. During the early stages of neocortical development, a series of tangentially migrating glutamatergic neuronal populations contribute to neocortical development. The ventral and medial pallia, which flank the dorsal pallium at both of its extremes, give rise to migratory neurons that tangentially colonize the dorsal pallium (Fig. 25.7; also see summary of the tangentially migrating neuronal populations in Table 25.1). These are pallial-derived neurons, therefore glutamatergic

FIGURE 25.7 Glutamatergic tangential migration orchestrated neocortical evolution. (A) Several populations of glutamatergic neurons reach the developing dorsal pallium (DPall) by tangential migration in the mouse brain. Neurons born at the cortical hem, rostral medial telencephalic wall (RMTW, depicted in the coronal slide, but actually present at more anterior telencephalic levels), and ventral pallium (VPall) migrate tangentially toward the DPall. The developing neocortex contains CajaleRetzius cells, subplate neurons, and transient pyramidal neurons (black), which play essential roles in neocortical formation. (B) The equivalent neurogenic regions of the avian telencephalon do not give rise to tangential migrations. The chick DPall does not receive external glutamatergic contributions. As a consequence, crucial populations for neocortical development are missing in the developing avian hyperpallium, whose glutamatergic neurons only derive from DPall progenitors. CP, cortical plate; GZ, germinative zone; IZ, intermediate zone; MtZ, mantle zone; MZ, marginal zone; SP, subplate layer.

A brief summary of the origins and destinations of the best researched tangential migrations during CNS development. The table also includes relevant molecular markers of each population, the species or taxa on which the migration has been described and a few key references.

Tangential migratory cellular type

Origin

Destination

Markers

Species described

Pallial interneurons

Pallidal subpallium

Pallium

GABA, Dlx, Lhx6

Vertebrates

de Carlos et al., 1996; Cobos, 2001; Metin et al., 2007; Carrera et al., (2008); Moreno et al., (2008)

CajaleRetzius cells

Cortical hem (MPall) and ventral pallium

Dorsal palliumdlayer 1

Reelin, p75, Lhx5

Mammals

Bielle et al., (2005); Garcia-Moreno et al., (2007); Miquelajauregui et al., 2010

Subplate neurons

Rostralemedial telencephalic wall (MPall)

Dorsal palliumdlayer 6b

Lpar1

Mouse

Pedraza et al. (2014)

Transient pyramidal neurons

Ventral pallium

Dorsal pallium

Dbx1 (at origin)

Mouse

Teissier et al., (2010), 2012

Thalamocortical corridor cells

Striatal subpallium

Pallidal subpallium

Isl1, Ebf1

Amniotes

Lopez-Bendito et al., (2006); Bielle et al., (2011)

Hindbrain diversity of neurons

Rhombic lip (at the rhombomere 1 roof plate)

Precerebellar, pontine, lateral reticular and cuneate nuclei, locus coeruleus

Noradrenaline (locus coeruleus)

Birds and mammals

Aroca et al., (2006); Wingate, 2001; Qu et al., 2006

Adult olfactory bulb interneurons

Telencephalic subventricular zone

Olfactory bulb

GABA, TH

Mammals

Lois and Alvarez-Buylla, 1994; Fuentealba et al., 2015

Cerebellar granule cells

Rhombic lip

External germinal layer (cerebellum)

Atoh1

Tetrapods and teleost fish

Butts et al. (2014)

Glutamatergic neurons of the medial amygdala

Paraventricular hypothalamic zone

Bed nucleus of the stria terminalis and medial amygdaloid nucleus

Otp, Foxg1, Meis2

Tetrapods

Bardet et al., 2008; Garcia-Moreno et al., 2010; Moreno et al., 2010

Olfactory cortex neurons

Several telencephalic subdomains

Piriform cortex and olfactory tubercle

No specific marker known

Mouse

Garcia-Moreno et al., 2008; Ceci et al., 2012

Lateral olfactory tract guidepost neurons

Dorsal pallium and thalamic eminence

Lateral olfactory tract

mGluR1

Mouse

Sato et al., (1998); Ruiz-Reig et al., (2017)

Accessory olfactory bulb mitral cells

Thalamic eminence

Accessory olfactory bulb

Lhx5,NP2, AP2a

Mouse

Huilgol et al., 2013

Neurons of nLOT2

Caudal dorsal pallium

Amygdaloid nucleus of the lateral olfactory tract (VPall)

NeuroD2, Lmo3

Mouse

Remedios et al., 2007

Key references

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TABLE 25.1 Tangential migrations on development and evolution.

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populations, and exert crucial roles in early cortical development. These populations include the CajaleRetzius (CeR) cells, the subplate neurons, and the transient subpopulation of pyramidal neurons (Barber and Pierani, 2015). Originated in the cortical hem of the medial pallium and at the ventral pallium, CeR cells are the first neurons settling the dorsal pallial mantle zone (Meyer et al., 1999) (Fig. 25.7A). CeR cells migrate tangentially in direct contact with the pial lamina and colonize the neocortical primordium in about 24 h in mouse (Garcia-Moreno et al., 2007). Once in the dorsal pallium, these “foreigner” neurons play essential roles for the proper formation of the neocortex. As major examples, CeR cells maintain RGC morphology, and function and stimulate the cortical layering by guiding the radial migration of pallial neuroblasts (Soriano and del Rio, 2005). Experimental paradigms in which CeR cells are missing during early development, or the expression of their crucial genes (reelin, apoER2, Cdk5) is abolished, all lead to malformations of the neocortex, which usually displays an aberrant layering and hosts fewer neurons (Ogawa et al., 1995; Katsuyama and Terashima, 2009). The deeper layer of the neocortex contains subplate neurons (Hoerder-Suabedissen and Molnár, 2015). At least a subpopulation of these cells reaches the dorsal pallium tangentially, from the most anterior regions of the medial pallium (García-Moreno et al., 2008; Pedraza et al., 2014). Early during neocortical neurogenesis as well, subplate neurons from this anterior region migrate tangentially, enter the dorsal pallium, and differentiate below CeR cells (Fig. 25.7A). Radially migrating glutamatergic neurons settle in between both early populations, this way leaving subplate neurons at the bottom of the postmitotic region. Because inputs to the neocortex enter through this deep region, the first inputs contact subplate neurons earlier than any other population. Subplate neurons, then, regulate when and where thalamic axons enter the maturing cortical plate. When these key regulators of cortical wiring are missing, the thalamus establishes an aberrant connectivity with the neocortex, leading to functional abnormalities. A final example of tangential intrapallial contributions to the developing neocortex is the transient pyramid population originated at the ventral pallium from a region expressing Dbx1 transcription factor (Teissier et al., 2010). These cells are born at the same brain region and believed to migrate similarly to the ventral CeR cells (Bielle et al., 2005), which enter the neocortex at earlier stages. The progressive reporter gene expression in the Dbx1-cre line suggested that these pyramidal neurons migrate tangentially to the developing neocortex and colonize the entire cortical plate (with a bias at lower layers). Experiments leading to the abolition of the gene Dbx1 associated with these migratory populations led to a decrease in the number of cortical neurons (Teissier et al., 2010), These observations led to the suggestion that they modulate cortical neurogenesis. However, the existence of this tangential migration remains disputed (Rueda-Alaña et al., 2018). These three populations enter the neocortex from external sources, interfere with the internal developing program, and ultimately help produce the mammalian neocortex (Fig. 25.7A). Recent work demonstrated that these early tangential migrations are distinctive of mammals and therefore could have a potential impact on the evolution of the mammalian neocortex (Fig. 25.7B). Several experiments suggest the mammalian specificity of these migratory movements. In ovo cell tracking experiments in chick led to label the lineage of the homologous germinative populations of the medial and ventral pallia (Garcia-Moreno et al., 2018). No tangential migrations were found within pallial boundaries (Fig. 25.7B), showing the absence of tangential contributions toward the development of the dorsal palliumdexcept the well-known subpallial tangential migration of GABAergic neurons. Therefore, the developing chick hyperpallium not only hosts a poorer neuronal diversity compared with mammals but also diverges by missing the important developmental instructions carried by the tangential populations. A similar scenario is expected in the reptilian dorsal pallium, although no direct experiments have been conducted on reptilian embryos to discriminate the existence of these glutamatergic migrations to date. However, indirect evidence suggests that these early glutamatergic tangential migratory streams are lacking in turtle, similarly as it was found in chick. This evidence includes the distribution of pallial markers, levels of expression of selected proteins, and the abundance of neurons at each pallial subdivision. Subplate markers, for instance, appear scattered throughout the turtle pallium, which rejects the nature of the population as a layer crucial in development (Montiel et al., 2011). Reelin expression at the pallial external surface is very low, indirectly suggesting a diminished population of reelin-expressing cells (Bernier et al., 1999, 2000). The reelin-expressing cells in mammals are the tangentially CeR cells. And the great neuronal numbers at the big protrusion at the ventral pallium, termed dorsal ventricular ridge, suggest that most if not all ventral pallial neurons remain locally and do not migrate toward the dorsal pallium (Aboitiz, 1999). Altogether, these data lead us to suggest that tangential migrations of glutamatergic neurons directed toward the dorsal pallium are also absent in reptilian brains. These migrations most likely only happen in mammalian early developing brains (Garcia-Moreno et al., 2018). And similar to the external scaffolds on a building under construction, tangential contributions of glutamatergic neurons supported and instructed the evolutionary organization of the neocortex. Tangential migration impacted the pallial development of ancient mammals and shaped the neocortex as we see it nowadays. Interestingly, CeR cells exemplify a developmental example of the influence of cell migration in the evolution of the brain. These neurons migrate tangentially from the medial and ventral pallia and constitute the principal neurons of cortical

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layer 1 (Bielle et al., 2005; Garcia-Moreno et al., 2007). But remarkably, CeR cells are principal actors in ruling radial glial migration toward the cortical plate; through reelin secretion and signaling, they set up the polarity of the mammalian cerebral cortex. This might be the most evident link between the two modes of neuronal migration: tangential migratory neurons triggering radial migration and neuronal positioning, ultimately influencing the structure and function of the mammalian dorsal pallium.

25.3.3 Tangential migration shaping brain connectionsdguidepost neurons in evolution The main input arriving to the vertebrate pallium departs from the thalamus. Whereas this axonal path is internal in mammalian brains, where the axons cross the different subpallial structures up to the neocortex, in other vertebrates, this path runs following a more superficial trajectory, therefore avoiding the subpallium (Fig. 25.8) (Tosa et al., 2014). The reason for this divergence lies directly on a seemingly indirect neuronal migration. Subpallial neurons have already been described as highly motile and are known to migrate tangentially all over the pallium and to subpallial FMUs different to where they were born. One of the latter examples is the corridor cell population. The dorsalmost region of the subpallium, the lateral ganglionic eminence (LGE), generates tangential cells that usually migrate to olfactory pallium but also to the medially adjacent globus pallidus (Lopez-Bendito et al., 2006). LGEderived neurons shortly migrate to medial subpallial regions and intercalate within the MGE postmitotic zone. This way, the MGE is split into two regions, inner and outer portions, with a corridor-shaped population of LGE neurons positioned in between (Fig. 25.8A). Navigating thalamic axons are capable of recognizing this corridor, permissive for their growth, and enter the telencephalon by crossing the postmitotic MGE. Their ingress into the pallium happens internally, near the periventricular zones (Lopez-Bendito et al., 2006). This internal entrance is linked to the singular wiring program of the mammalian dorsal pallium: Thalamic afferences to the cortex enter the neuroepithelium by the apical side, and the first neurons these axons find are subplate neurons, which are known to regulate the reciprocal corticothalamic connectivity (Hoerder-Suabedissen and Molnár, 2015). An aberrant tangential migration of LGE-derived corridor cells misguides thalamic afferences to other regions of the pallium, generating a defective cortical wiring. Corridor cells have been described in other amniote species, including chick and turtle (Bielle et al., 2011). In these species, corridor cell population is also capable of allowing the growth of thalamic axons. However, its position diverges when compared with mammals: Sauropsidian guidepost neurons migrate different and reach a medial position, which is adjacent to the MGE germinative zone (Fig. 25.8B). And importantly, this position is distant to the point on the diencephalo-telencephalic junction where the axons first invade from the thalamus. Corridor cells in sauropsid brains do not guide thalamic axons (Bielle et al., 2011). Because of the chemorepulsive action of the globus pallidus, thalamic axons

FIGURE 25.8 Corridor cells in the evolution of the thalamopallial connectivity. (A) Scheme representing a coronal section through the embryonic mouse brain. LGE-derived neurons (black) migrate tangentially into the MGE territory, splitting the globus pallidus (GP) area into two halves. The migratory cells reach the telencephalo-diencephalic junction and generate so a permissive corridor for the growth of thalamic axons (dark gray). As a consequence, thalamic axons arrive to the pallium following an internal path, and the first neuronal population they contact in the cortex is the subplate (SP). (B) Scheme depicting an equivalent chick coronal section but representative of the corridor cells in sauropsids. LGE-derived cells split the GP area by tangential migration, as in the case of mouse. However, the cells do not reach the telencephalo-diencephalic junction due to a divergent expression of Slit2. The thalamic axons cannot find the permissive corridor cells and surround the GP when entering the telencephalon. Following this path at the lateral forebrain bundle (lfb), thalamic axons reach the pallium superficially. DPall, dorsal pallium; dTh, dorsal thalamus; PSB, pallialesubpallial boundary; St, striatum.

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travel the basal telencephalon externally near its surface. They avoid the accumulating mass of dorsal ventricular ridge (DVR) at the pallialesubpallial boundary and reach the pallium superficially. As a developmental consequence, the thalamic axons do not find the homologous populations of the subplate, and the pallial wiring is set up following a divergent program (Fig. 25.8B). There are other examples of tangentially migrating guidepost neurons, such as those guiding the lateral olfactory tract (lot), which runs by the ventrolateral surface of the telencephalon. The axons of the mitral cells in the olfactory bulb innervate so the olfactory cortex, including piriform cortex, olfactory tubercle, and olfactory-associated regions of the amygdala. To pathfind through the entire rostrocaudal extension of the telencephalon, mitral axons need guidance from lot cells, a neuronal population that early settles the topological position where the axons navigate. This population reaches its ventrolateral position tangentially, from several germinative sources including the dorsal pallium and the thalamic eminence (Sato et al., 1998; Kawasaki et al., 2006; Ruiz-Reig et al., 2017). For this migration, a complex molecular mechanism guides the cells toward their final location: Robo/Slit communication between mitral axons and lot cells; Sema3F expression in the telencephalic mantle zone; and ephrin-A5 and netrin-1 expression at the ventral telencephalon among others (Nomura et al., 2006; Ito et al., 2008) help guide the migratory pattern of lot cells. Slit-deficient mice fail the migration of lot cells, and consequently, mitral axons from the olfactory bulb get lost at the ventrolateral telencephalon. The tract develops aberrantly, and the olfactory network is poorly established. Whether homologous pathfinding events occur in other vertebrate species remains to be uncovered yet. Other examples in which tangentially migratory neurons play a relevant function in axon pathfinding are the corpus callosum guidepost neurons (Shu, 2003; Niquille et al., 2009), the CD44-positive neurons for the retinal axons at the optic chiasm (Sretavan et al., 1995), and CeR cells in the guidance of hippocampal projections (Del Rio et al., 1997). Unfortunately, very little if anything has been researched to date on the comparative aspects of these tangential migrations. But it is an emerging idea that tangential migration increased brain complexity at several levels throughout vertebrate evolution, being one of them the establishment of long-range connectivity. New tangential populations will be soon related to the development of many other axonal tracts and in a wide variety of species.

25.3.4 Tangential migrations along the central nervous system We have succinctly depicted so far a few populations that settle their final destinations in the brain by tangential migration, but there are increasing numbers of them in the literature. Tangential migration is uncovered in every brain region and species in which it is sought (Table 25.1). The neurons tangentially conquering new brain regions execute a growing variety of roles. The locus coeruleus in the brain stem is the most important noradrenergic population in the brain and plays relevant homeostatic control roles. Its neurons connect and excite most brain regions, from the most anterior telencephalon down to spinal cord. Locus coeruleus neurons are born early in the alar region of rhombomere 1 (r1). By means of ventralward tangential migration, these noradrenergic neurons reach their final position in the basal plate of r1 (Aroca et al., 2006). This noradrenergic population is well conserved among vertebrates, and as such, its tangential migration has been described in at least mouse and chick embryonic brains. Also in the brain stem, the rhombic lip is the origin of several tangential streams. The main evolutionary impact of these numerous tangential migrations was the increasing diversity of brain stem neuronal types. As such, tangential migration contributes to the formation and neuronal diversification of the pontine nuclei, inferior olive, reticular nuclei, and cuneatus nucleus at least (Wingate, 2001; Qu et al., 2006). Undoubtedly, tangential migration shaped brain stem structure and function early in vertebrate evolution. Most likely, these early tangential events comprise a general feature of vertebrates and will be conserved among species. Tangential migratory neurons also sustain circuit plasticity in continuously forming circuits. The olfactory epithelial neurons communicate to the brain at the olfactory bulbs. These cells, highly exposed to the external environment, are renewed throughout life. Their connections with the olfactory bulb experience a constant growth and refurbishment. A particular type of pallial interneurons migrates from the striatal region of the subpallium rostrally into the olfactory bulb and supports the continuous circuit formation (reviewed by Lledo et al., 2008). This relevant population generates at the subventricular zone of the telencephalon, covers long tangential distances by the rostral migratory stream, and settles the olfactory bulb. These neuroblasts become GABAergic and dopaminergic interneurons. But their most interesting feature is that these neurons are born and migrate during the entire life of the animal. Most mammals show this case of adult neurogenesis followed by tangential migration, which correlates with the mammalian specialization on olfactory reception. Other vertebrate species display adult neurogenesis, in brain regions most likely equivalent to that of the mammalian subventricular zone. However, it remains unclear whether homologous migration and circuit homeostasis happen in other vertebrates. A final example to emphasize the essential role of tangential migration in brain evolution happens at the cerebellum. Whereas Purkinje cells are born locally and settle the cerebellum by radial migration, the GABAergic granule cells reach

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their positions by tangential migration (Komuro et al., 2001). Granule cell precursors are born at the germinative zone of the fourth ventricle, at the rhombic lib during embryonic stages. They undergo tangential migration in direct contact with the meningeal surface, by the surface of the cerebellar primordium, and create the external granule cell layer. This layer comprises a premitotic population of neuroblasts that remain cycling for weeks after the rest of the brain has ended up the neurogenesis period. Eventually, the newborn granule cells migrate radially toward the ventricular surface and settle at their final location in the internal granule cell layer. Because granule cell neuroblasts vastly proliferate for days and weeks, the adult population of granule cells is the most numerous neuronal population in the brain of birds of mammals (Surchev et al., 2007). Up to 75% of all human brain neurons are granule cells. Therefore, tangential migration provides a definitive substrate for the generation and amplification of a massive neuronal population in the adult cerebellum. Granule cell tangential migration evolved differently throughout the vertebrate radiation. All vertebrate species possess granule cells at the cerebellum, and these are all born at the rhombic lip (Butts et al., 2010). However, the extent of this population is bigger in tetrapods, in which granule cells migrate externally and massively in birds and mammals, where these cells migrate at a premitotic state. Remarkably, tangential migration of neuroblasts not only promoted an unprecedented increase of neuronal numbers but also modified the internal connectivity within cerebellar circuits. Tangential migration brought together granule cells from the rhombic lip and Purkinje cells (Butts et al., 2014). These two neuronal types overlap in the tetrapod cerebellum, achieving so a most efficient connectivity, with eventually impacted on the adeptness of the cerebellar processing circuit.

25.4 Conclusions The numerous developmental programs of vertebrate brains reflect the divergent solutions produced by evolution to respond to the environment. Especially at the neuronal level, neuronal types are mainly a product of their cellular development (Arendt et al., 2016; Tosches et al., 2018; Zeisel et al., 2018). Understanding how neurons develop provides unequaled information on brain evolution and function. And one of the key developmental events shaping neuronal types and brain circuits is neuronal migration. Current vertebrate brains were molded by the evolving patterns of neuronal migration. Our brain would be utterly different if other migratory events had happened in the past 400 mya. A deep understanding on the evolution of neuronal migrations is a major step to build the knowledge on the underlying principles of brain function.

Glossary l

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Amniotes: The group of tetrapod vertebrate species that lay their eggs on land or retain the fertilized egg within the mother. Amniotes encompass reptiles (lepidosaurs, testudines, and archosaurs, which include birds) and mammals. CajaleRetzius cells: Early-born glutamatergic neurons residing the superficial layer of the neocortex. Most of them die by apoptosis soon after corticogenesis. They were termed “special neurons” by Cajal due to their rare feature of possessing two or more axons. Contribute to cortical development mainly by the expression of the glycoprotein reelin. Fundamental morphogenic unit (FMU): A discrete subdivision of the embryonic brain, encoding a unique combination of transcription factors. Each FMU generates the neurons radially located above its germinative zone. Germinative zone: The apical neuroepithelium is in contact with the ventricular basal lamina. Neurons are born in the germinative zone. In some brain regions and species, the germinative zone is subdivided into a ventricular zone (progenitors remain in contact with the ventricular surface) and a subventricular zone (progenitors do not contact the ventricular surface). Mantle zone: The external, basal portion of the neuroepithelium, where neurons arrive after migration, differentiate, and assemble mature circuits. The mantle zone is a postmitotic region, meaning that no neurogenesis occurs within its boundaries. Pallial interneurons: GABAergic neurons originated at the subpallium and that integrate into pallial circuits after tangential migration. Most of these neurons contribute to the pallial circuitry with short-range local axons and are so considered interneurons. Although a minority of them establishes long-range connectivity within the mammalian neocortex, for the purpose of our revision, we encompass them all into the general term pallial interneuron. Pallium: The dorsal-most area of the telencephalic alar plate, which produces all glutamatergic neurons of the telencephalon. According to genoarchitectonics and adult derivatives, the pallium can be subdivided into four independent units, the so-called medial, dorsal, lateral, and ventral pallia.

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Dbx1-Derived Pyramidal Neurons Are Generated Locally in the Developing Murine Neocortex. Front. Neurosci. 12, 1e10. https://doi.org/10.3389/fnins.2018.00792. Ruiz-Reig, N., Andrés, B., Huilgol, D., Grove, E.A., Tissir, F., Tole, S., et al., 2017. Lateral Thalamic Eminence: A Novel Origin for mGluR1/Lot Cells. Cereb. Cortex. 27, 2841e2856. https://doi.org/10.1093/cercor/bhw126. Saito, S., Shiina, T., Shimizu, Y., 2017. Tissue and cell. Tissue Cell 49, 514e519. Sato, Y., Hirata, T., Ogawa, M., Fujisawa, H., 1998. Requirement for early-generated neurons recognized by monoclonal antibody lot1 in the formation of lateral olfactory tract. J. Neurosci. 18, 7800e7810. Shu, T., 2003. The glial sling is a migratory population of developing neurons. Development 130, 2929e2937. Simons, D.J., Carvell, G.E., 1989. Thalamocortical response transformation in the rat vibrissa/barrel system. J. Neurophysiol. 61, 311e330. Smart, I.H.M., Dehay, C., Giroud, P., Berland, M., Kennedy, H., 2002. Unique morphological features of the proliferative zones and postmitotic compartments of the neural epithelium giving rise to striate and extrastriate cortex in the monkey. Cerebr. Cortex 12, 37e53. Soriano, E., del Rio, J., 2005. The cells of cajal-retzius: still a mystery one century after. Neuron 46, 389e394. Sretavan, D.W., Pure, E., Siegel, M.W., Reichardt, L.F., 1995. Disruption of retinal axon ingrowth by ablation of embryonic mouse optic chiasm neurons. Science 269, 98e101. Striedter, G.F., Beydler, S., 1997. Distribution of radial glia in the developing telencephalon of chicks. J. Comp. Neurol. 387, 399e420. Suárez, R., Richards, L.J., 2014. Evolution and development of interhemispheric connections in the vertebrate forebrain. Front. Hum. Neurosci. 1e14. Surchev, L., Nazwar, T.A., Weisheit, G., Schilling, K., 2007. Developmental increase of total cell numbers in the murine cerebellum. Cerebellum 6, 315e320. Szele, F.G.F., Cepko, C.L.C., 1996. 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Chapter 26

Neuronal migration disorders Joseph J. LoTurco1 and Jean-Bernard Manent2 1

University of Connecticut, Mansfield, CT, United States; 2Inmed Inserm, Marseille, France

Chapter outline 26.1. Introduction 26.2. Types of malformations 26.2.1. Pachygyria 26.2.2. Lissencephaly 26.2.3. Cobblestone lissencephaly 26.2.4. Subcortical band heterotopia 26.2.5. Periventricular heterotopia 26.2.6. Polymicrogyria 26.2.7. Mammalian target of rapamycin complex pathwayerelated malformations 26.2.8. Microcephaly 26.3. Identified mutations and mechanisms in neuronal migration disorder 26.3.1. Mutations in microtubule-associated proteins (LIS1, DCX, KIF5C, KIF2A, DYNC1H1, and EML1)

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26.3.2. Tubulin mutations (TUBA1A, TUBB2B, TUBB3, TUBG1, TUBA8, and TUBB5) 581 26.3.3. Periventricular heterotopia and mutations in FLNA, ARFGEF2, C6orf70, FAT4, DCHS1, and MOB2 582 26.3.4. Variant lissencephalies and mutations in ARX and RELN 583 26.3.5. Cobblestone malformations and mutations in dystroglycan genes 584 26.3.6. Focal cortical dysplasias and dysplastic megalencephaly and mutations in mTOR, PIK3CA, DEPDC5, AKT3, NPRL3, and PIK3R2 584 26.4. Summary and concluding remarks 585 References 585

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26.1 Introduction Malformations of brain development can arise from disruptions in any number of interconnected developmental processes including alterations in cell fate, cell proliferation, cell death, and cell migration. Disruptions in mechanisms involved primarily in neuronal cell migration cause neuronal migration disorders (NMDs), and these disorders encompass multiple syndromes characterized by aberrations in the cellular architecture of the brain including alterations in sulci and gyri in cerebral cortex and in lamination patterns of the neocortex, cerebellum, and other structures. Alterations associated with NMDs are diverse in terms of type and severity. Such disruptions range from subtle, only seen with microscopic analysis in isolated regions, to disruptions covering large regions of the brain that are easily apparent with MRI imaging. Genetic studies have now revealed many mutations in several genes that cause syndromes associated with disorders in neuronal migration. Functional studies of the products of these genes have in turn revealed proteins and pathways required for distinct cellular processes including those involved primarily in neuronal migration and those involved in processes that indirectly alter neuronal migration when altered. Although NMDs may involve many parts of the brain, they are most apparent in the neocortex, hippocampus, and cerebellum. The enrichment of migration disorders involving these areas is likely due to the highly patterned and regular structure of these brain areas. Any disruption in cell placement is more obvious in such structures. The ordered pattern of cell migration required to establish the precise architectures of cortex and cerebellum requires coordinated molecular and cellular processes in multiple cell types, and disruption in any of these or in their coordination can result in a migration disorder. The required mechanistic complexity may render these structures more susceptible to both genetic and environmental insults. Consistent with the correlation between complex and ordered cellular architecture and susceptibility to neuronal migration disorders, the human brain shows a greater variety of distinguishable NMD types compared with the rodent brain.

Cellular Migration and Formation of Axons and Dendrites. https://doi.org/10.1016/B978-0-12-814407-7.00026-2 Copyright © 2020 Elsevier Inc. All rights reserved.

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NMDs in cerebral cortex are a subset of the more general set of disorders of development, malformations of cortical development (MCD). MCD includes a number of developmental disruptions that cause a displacement of neurons and glia from their typical pattern in cerebral cortex. As neuronal migration involves multiple interactions between cells in the developing brain, disruptions in neuronal positioning can occur by any developmental disruption that may change the cellular environment through which neurons normally migrate. For example, changes in cell proliferation, cell death, cellular differentiation, or disruptions in vascularization, while not primarily disruptions in cellular mechanisms required for neuronal migration, can nonetheless cause disruptions in normal cellular patterns in the brain.

26.2 Types of malformations NMDs, initially categorized by descriptive pathological features and inheritance patterns, are now divided into an increasing number of defined syndromes. These syndromes are characterized by combinations of clinical features, malformation types, genetic causes, and predominant locations of malformations. The descriptive pathological terms remain important components of the newer categories. It is important to recognize, however, that each pathological feature does not represent a single disease or syndrome. For example, distinct categories of NMDs can contain pachygyria to varying degrees, lissencephaly with or without microcephaly, or lissencephaly with or without cerebellar hypoplasia. In fact, the comorbidities of malformation types, even when caused by a single gene mutation, reveal shared and distinct functions of single genes between and within different regions of the brain. The aim of this section is to provide general definitions of the types of major neuropathological features associated with NMDs.

26.2.1 Pachygyria Pachygyria literally means thick or fat gyri. It is characterized by a marked thickening of the gray matter of the neocortex such that the normal cross-sectional thickness is increased typically by 50% or more. The underlying white matter is also often more diffuse and/or thinner than normal. Pachygyria is usually associated with a reduction in the number of gyri and sulci, or agyria, to varying degrees. It is most commonly associated with lissencephaly but can also occur in isolation.

26.2.2 Lissencephaly Type 1 or “classical” lissencephaly (LIS), meaning smooth brain from the Greek root “lisso,” is defined by a lack of gyri and sulci in the cerebral neocortex (Fig. 26.1C). In terms of comparative neuroanatomy, “lissencephalic” refers to species such as mice with neocortices that are smooth and lack sulci or gyri. It is important to distinguish this evolutionary

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lissencephaly from the human pathology. Human lissencephaly, in addition to a lack of sulci and gyri, is characterized by a simplification of neocortical lamination. Typically, the neocortex of lissencephalic patients has three to four layers instead of six. Lissencephaly, like pachygyria, does not necessarily occur across the entire cortex and is often restricted to more frontal or occipital regions, depending on the underlying genetic cause.

26.2.3 Cobblestone lissencephaly Cobblestone lissencephaly, also known as type 2 lissencephaly, is different from classical or type 1 lissencephaly. The lack of sulci and gyri in this malformation type is the result of neurons migrating past the outer limits of normal gray matter to form discrete and interconnected patches. This results in a characteristic bumpy, granular, or “cobblestone” appearance to the surface of the cortex, lacking deep sulci (Fig. 26.3C). At the cellular level, the normal laminar appearance of the cortex is replaced by variable aggregations of cells. There is usually disruption in white matter and myelination, enlarged ventricles, brain stem hypoplasia, and cerebellar hypoplasia. This type of malformation is typically associated with retinal dysplasia and muscular dystrophy.

26.2.4 Subcortical band heterotopia Subcortical band heterotopia (SBH), or band heterotopia, are characterized by contiguous groupings of neurons forming a band of gray matter within the white matter just below the neocortical lamina (Fig. 26.2A). In the most extreme version of SBH, double cortex syndrome, bands of neocortical neurons form bilateral and elongated structures that are as thick or thicker in cross section than the more superficial “normotopic” cortex. SBH are either partially or completely surrounded by cortical white matter. Utilizing enhanced imaging technologies, small band heterotopia have been identified in the brains of some epileptic patients and nonepileptic individuals.

26.2.5 Periventricular heterotopia Periventricular heterotopia (PH), similar to SBH, are also formed by ectopic aggregates of neurons located medial to or underneath the normal neocortical gray matter (Fig. 26.2C). Unlike SBH, however, PH abut and extend into the ventricles. They are also known as subependymal heterotopia and can appear in combinations of single to multiple nodes along the surface of the lateral ventricles. When numerous, the nodes merge to form a wavy contiguous mass along the ventricular surface.

26.2.6 Polymicrogyria Polymicrogyria (PMG) is a malformation whereby a part of or all of the cortex contains many more sulci and gyria than normal, and these additional convolutions are smaller than usual (Fig. 26.3A). PMG can be highly localized and bilaterally

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WM WM FIGURE 26.2 Subcortical band heterotopia (SBH) and periventricular heterotopia (PH). (A) Illustration of a coronal section from a brain with SBH. Note the band of heterotopic neurons in the white matter below the cortical layers. (B) Magnified schematic of the cortical layers showing normal sixlayered organization, with the addition of heterotopic neurons sequestered within the white matter. (C) Drawing depicting a brain with PH that harbors multiple nodules lining the surface of the ventricles. (D) Schematic showing cortical layers with normal organization.

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FIGURE 26.3 Polymicrogyria (PMG) and cobblestone lissencephaly. (A) Illustration of a coronal section from a brain with PMG whose sulci and gyri are more numerous and smaller than normal. (B) Schematic enlargement of (A) showing disorganized cortex with fewer layers than normal. (C) Drawing of a coronal section of a brain with cobblestone lissencephaly. The surface of the brain is smooth but has small extrusions of neurons that dimple the cortical surface. (D) Magnified schematic of the cortex depicting a protrusion of neurons through the normal cortical boundaries giving rise to the cobblestone appearance of the brain surface.

symmetric, asymmetric, or unilateral. The cortex is thinned as in cobblestone lissencephaly, but unlike cobblestone lissencephaly, the cortex retains a degree of regular structure and lamination. At the cellular level, the cortical lamination is reduced to two to three layers, and the cell-sparse marginal zone is typically intact.

26.2.7 Mammalian target of rapamycin complex pathwayerelated malformations Both genetic evidence and certain common cellular pathologies group a subset of MCDs caused by mutations in mammalian target of rapamycin complex (mTOR) pathwayerelated genes. These MCDs include dysplastic megalencephaly (DMEG), or hemimegalencephaly, type II focal cortical dysplasias (FCD II, a and b), and brain malformations observed in tuberous sclerosis (TSC). DMEG, FCD type II, and dysplasias in TSC are causes of intractable epilepsy in children (Palmini et al., 2004; Fauser et al., 2006; Krsek et al., 2008). All are generally characterized by thickened and lost distinctions between cortical lamina, involving large portions of cortex in DMEG, and smaller, often focal, regions in FCD II. These malformations are associated with somatic and germline mutations in genes in the mTOR pathway required for the control of cellular growth and contain abnormally oriented neurons, hypertrophic astrocytes and neurons, and dysmorphic neurons.

26.2.8 Microcephaly Microcephaly results from development disruptions that cause abnormally smaller brains, typically 2e3 standard deviations smaller than normal. Microcephaly is not due primarily to defects in neuronal migration but often occurs in brains that also contain neuronal migration disruptions. Migration disruption in microcephalies can arise either because the same underlying genetic mutation independently affects both cell migration and cell division, such as mutations in the LIS1 gene, or from disruptions in cell migration secondary to changes in cell proliferation or increased cell death.

26.3 Identified mutations and mechanisms in neuronal migration disorder 26.3.1 Mutations in microtubule-associated proteins (LIS1, DCX, KIF5C, KIF2A, DYNC1H1, and EML1) The class of malformation that is most clearly a direct consequence of stalled migration is SBH (Barkovich et al., 1994; Dobyns et al., 1996; Palmini et al., 1991). In its extreme, a large band of cells can form a separate mass of gray matter throughout the cortex, and the presence of such SBH has been referred to as “double cortex syndrome” (Fig. 26.2A; Palmini et al., 1991). During the formation of SBH malformations, many neurons fail to migrate through the intermediate zone and into normally forming cortical lamina and thereby become embedded within the white matter of neocortex

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(Fig. 26.2B). Heterotopia of any type in which subsets of neurons attain abnormal positions while others migrate and pattern normally within the same area suggests somatic mosaicism in migration disruption. How such mosaicism occurs is clear for two major genetic causes of heterotopia: PH (see the following text) and SBH. Heterozygous mutation in the DCX gene in females is the most common cause of SBH. Random X-inactivation in somatic cells naturally creates a cellular mosaic. Heterozygous DCX mutations then combined with X-inactivation will result in some cells with an active copy of the mutant gene, and these cells fail to migrate into normotopic cortex (des Portes et al., 1998b; Gleeson et al., 1998). Somatic mosaicism may also occur from somatic mutations in early stages of embryonic development that creates a mixture of cells with functional and nonfunctional copies of genes critical to migration. Evidence for this type of mosaic pattern has been found by genotyping hair follicles in some SBH cases and finding a mosaic pattern of DCX or LIS1 mutation (Gleeson et al., 2000; Sicca et al., 2003). Such somatic mosaicism in DCX mutations can be a cause of SBH in males (Poolos et al., 2002). The clinical features of double cortex syndrome show a wide range of severity, but typical deficits are mental retardation with a high incidence of seizures (Barkovich et al., 1994; Guerrini and Carrozzo, 2001a, b). The size of heterotopia is generally correlated with the severity of phenotypes (Guerrini and Carrozzo, 2001a, b; Bahi-Buisson et al., 2013). However, some cases of subcortical heterotopia have been discovered in nonsymptomatic mothers of sons with XLIS. The primary reason for a range in the size of SBH in different cases may be due to the natural variation of random Xinactivation. However, the severity of the SBH malformation also correlates with the specific DCX mutation, with some mutations generally causing larger malformations than others. Some milder mutations can even cause SBH in males instead of lissencephaly, which would typically be the result of a DCX mutation. Thus, some migrating cortical neurons may be able to migrate normally even with a defective copy of the DCX gene. Animal models of SBH have been developed and, similar to the human condition, the animal models display seizures and more excitable neocortex (Ackman et al., 2009; Bai et al., 2003; Lapray et al., 2010; Manent et al., 2009; Moustaki et al., 2019; Petit et al., 2014; Plantier et al., 2018; Sahu et al., 2018). Mice mutant for the Dcx gene or related Dclk gene do not show the development of SBH, though these mice do show disruptions in neuronal lamination and migration (Deuel et al., 2006; Kerjan et al., 2009; Koizumi et al., 2006). This indicates a species difference in the DCX-dependent mechanisms and SBH formation in mice, rats, and humans (Ramos et al., 2006). This species difference is further shown by the difference in the effects of RNAi knockdown of Dcx in mice and rats. In rats, but not in mice, targeting of Dcx expression in a subpopulation of migrating neocortical neurons results in formation of SBH (Bai et al., 2003; Ramos et al., 2006). The variability in SBH sizes and the correlation between malformation size and severity of seizure phenotypes in humans inspired an effort to regress heterotopia sizes in an animal model. In studies using RNAi of Dcx to induce formation of heterotopia, and then subsequent conditional reexpression of Dcx, it was shown that reexpression of Dcx in stalled neurons is sufficient to restart migration out of SBH malformations (Manent et al., 2009). The resulting decrease in SBH size is associated with a decrease in susceptibility to pharmacologically induced seizures (Manent et al., 2009). Mutations in microtubule-associated proteins other than LIS1 or DCX were also found in patients with cortical malformations. A germline mosaic mutation in KIF5C, encoding ATP-dependent molecular motors involved in intracellular transport along microtubules, was found in a family with four affected male individuals showing severe MCD and microcephaly (Poirier et al., 2013a,b). De novo missense variants in KIF2A, encoding a protein involved in the ATPdependent depolymerization of microtubules, were found in a female patient with frontal band heterotopia, posteriorbiased pachygyria, and severe microcephaly and in a female patient with congenital microcephaly and posterior predominant pachygyria (Poirier et al., 2013a,b). De novo variants in DYNC1H1 encoding heavy chain of dynein were identified in eight sporadic cases and in a family with two brothers and their mother, associated with a wide range of brain malformations, including abnormal gyral patterns, microcephaly, and callosal, cerebellar, and/or brain stem dysgenesis (Poirier et al., 2013a,b). Mutations in EML1, an MT binding protein, were identified in two unrelated families with giant bilateral periventricular and ribbonlike subcortical heterotopia, polymicrogyria, and agenesis of the corpus callosum (Kielar et al., 2014). Consistent with a causal role for this mutation across species, Eml1 mutations have been discovered in two different rodent mutants that have bilateral SBH: the spontaneous HeCo mouse mutant (Croquelois et al., 2009; Kielar et al., 2014), as well as in the Tish mutant rat (Chen et al., 2000; Lee et al., 1997, 1998; Kielar et al., 2014; Grosenbaugh et al., 2017).

26.3.2 Tubulin mutations (TUBA1A, TUBB2B, TUBB3, TUBG1, TUBA8, and TUBB5) Most cases of type 1 lissencephaly are caused by mutation in either of two microtubule-associated proteins LIS1 or DCX. From this observation, one prediction would be that there should be corresponding mutations in tubulin subunits, which cause malformations of cerebral cortex and cerebellum. In the past few years, mutations in three tubulins, TUBA1A,

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TUBB2B, and TUBA8, have been identified that cause a range of malformation types, including lissencephaly with cerebellar hypoplasia and PMG (Abdollahi et al., 2009; Jaglin and Chelly, 2009; Tischfield and Engle, 2010). The malformation types created by mutations in tubulin genes are surprisingly diverse. Both lissencephaly with pachygyria and PMG occur with mutations in a- and b-subunits. This coincidence of two seemingly opposite phenotypes, a lack of sulci and gyria in LIS and an abundance of extra sulci and gyri in PMG, both originating from mutations in the core components of microtubules, indicates that these types of malformation may be mechanistically related at the molecular level. One possibility for the differences in malformation after mutations in different tubulins may be cell-specific functions and expression of different tubulin isoforms. For example, in one region, the loss of function of one tubulin in radial glia could result in PMG, whereas decreased function of a different tubulin in migrating neurons could cause lissencephaly (Abdollahi et al., 2009; Jaglin and Chelly, 2009). Mutations in the gene TUBA1A, tubulin a-subunit 1 A, cause malformations of the brain that include lissencephaly, cerebellar hypoplasia, agenesis of the corpus callosum, and brain stem anomalies (Keays, 2007; Poirier et al., 2007). Mice with mutations in the same gene show a similar impairment in migration of neurons in the neocortex and hippocampus (Keays, 2007). Mutations in the TUBA1A gene in humans cause a wide spectrum of neuronal malformations that range in severity from isolated pachygyria in the perisylvian cortex to widespread lissencephaly with cerebellar hypoplasia (LCH) (Bahi-Buisson et al., 2008; Kumar et al., 2010; Morris-Rosendahl et al., 2008; Poirier et al., 2007). Moreover, mutations in TUBA1A account for 1%e4% of lissencephaly cases and nearly 30% of LCH cases (Kumar et al., 2010; Morris-Rosendahl et al., 2008). At the level of protein function, missense mutations in TUBA1A interfere with the ability of microtubulebinding proteins (MAPs) to associate with microtubules (Kumar et al., 2010). De novo mutation in TUBB2B causes asymmetrical PMG, which can occur in different locations in each hemisphere. Neurons in TUBB2B-related PMG migrate past the normal limits of the most superficial layer through breaks in the basement membrane (Jaglin et al., 2009). This pattern of migration past the basement membrane is an abnormal migration pattern more typical of type 2 than type 1 lissencephaly. TUBG1 mutations were identified in three patients with complex cortical malformation phenotypes, combining pachygyria and microcephaly in the two most severe cases and combining a posterior-biased pachygyria, laminar heterotopia, and a dysmorphic corpus callosum in the less severe case (Poirier et al., 2013). In a recent study, Bahi-Buisson et al. (2014) investigated a large cohort of 80 patients with tubulin-related cortical malformations and found 45 patients with mutations in TUBA1A (42.5%), 18 with mutations in TUBB2B (16.9%), 11 with mutations in TUBB3 (10.4%), 3 in TUBB5 (2.8%), and 3 in TUBG1 (2.8%). However, no mutation in TUBA8 was identified, suggesting that TUBA1A mutations account for the majority tubulin-related cortical malformations. An earlier study suggested that autosomal recessive mutations in the TUBA8 gene create a syndrome with generalized PMG and hypoplasia of the optic nerve (Abdollahi et al., 2009). Because Tuba8 is expressed at low levels in the developing mouse and human brain (Braun et al., 2010), and because no histological or immunohistochemical alterations were found in the cortex of Tuba8 / mice (Diggle et al., 2017), two of the original TUBA8-mutated patients were subjected to further genetic analyses. Whole exome sequencing allowed identifying an additional homozygous loss-of-function mutation in SNAP29 (Diggle et al., 2017), suggesting that SNAP29 deficiency, rather than TUBA8 deficiency, may be involved. Identification of additional TUBA8 mutations is thus needed to definitively support a causal role of TUBA8 in tubulinrelated cortical malformations.

26.3.3 Periventricular heterotopia and mutations in FLNA, ARFGEF2, C6orf70, FAT4, DCHS1, and MOB2 PH is a heterogeneous disorder characterized by heterotopic organizations of cells and nodules at the surface of the ventricles that may be isolated or occur in combination with other neuronal malformations (Fig. 26.2C) (Dobyns et al., 1997; Fox and Walsh, 1999; Guerrini and Carrozzo, 2001b). In postmortem human PH brains, periventricular nodules are composed of later-born neurons that would normally migrate into upper layers. Also these brains show disruption in the neuroependyma (Ferland et al., 2009). These late-born neurons in nodules are thought to be neurons that failed to start migration away from the ventricular zone at the point of their origin. The clear misplacement of neurons in the neocortex places PH in the category of NMD, but the malformation may result primarily from a deficit in proliferation and cell adhesion in neural progenitors. Bilateral PH malformations, similar to SBH malformations, represent cellular mosaics with most cells migrating normally and others not. The cellular mosaicism and skewed incidence in females indicated a pattern consistent with a causative mutation location of the X chromosome (Dubeau et al., 1995; Fink et al., 1997; Guerrini and Dobyns, 1998). In fact, the first mutant gene related to bilateral periventricular nodular heterotopia (BPNH) was the gene FLNA on Xq28 (Fox et al., 1998). More than 40 different mutations in FLNA have been identified in individuals with bilateral PH, and such

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mutations in FLNA account for 50% of cases with classical bilateral PH (Parrini et al., 2006). Mutations in other regions of the same gene cause a variety of developmental disruptions, many of which have no apparent neurological involvement. The high degree of specificity for the type of mutation in FLNA in terms of the organ and cell type affected suggests that different domains of the filamin A protein are engaged by distinct signaling mechanisms in different cell types (Robertson, 2004, 2005). The FLNA gene codes for the protein filamin A, an actin-binding protein that promotes the formation of branched networks of actin filaments. Actin filaments when bound to filamin A can interact with a large number of proteins including ion channels and membrane proteins involved in cell adhesion (Nakamura et al., 2011). Knockdown by RNAi resulting in a loss of filamin a and b expression interferes with the initiation of migration and cell spreading in nonneuronal cells (Baldassarre et al., 2009). Mouse mutants of the Flna gene have not been produced that recapitulate the PH phenotype; however, a mouse mutant of the Mekk4 gene, a regulator of FLNA protein function, forms PH in the fetal mouse brain. Similarly, either RNAi knockdown of Flna or expression of Flna dominant-negative protein in the mouse ventricular zone impairs proliferation and movement away from the ventricular surface (Ferland et al., 2009). A rare autosomal recessive form of PH with microcephaly is caused by mutation of the ARFGEF2 gene (Sheen et al., 2004). Patients with such mutations in ARFGEF2 show severe developmental delay, seizures beginning in infancy, and microcephaly. The microcephaly phenotype is the primary feature that distinguishes PH caused by ARFGEF2 mutations from PH caused by FLNA mutations, suggesting that the ARFGEF2 gene may play a larger role in the control of proliferation than the FLNA gene. Arfgef2 is highly expressed in neural progenitors in the mouse neocortex. The Arfgef2 gene codes for a protein known as BIG2, which is essential to vesicle trafficking in cells. BIG2 catalyzes guanine diphosphate (GDP) to guanine triphosphate (GTP) and activates ADP ribosylation factors that trigger vesicle trafficking. BIG2 function is inhibited by BREFELDIN A, and expression of BREFELDIN A inhibits cellular proliferation and the transport of Ecadherin and b-catenin to the cell surface (Sheen et al., 2004). Biallelic mutations in the atypical cadherin receptor ligands FAT4 and DCHS1 have been identified in patients with Van Maldergem syndrome, an autosomal recessive disorder characterized by intellectual disability, multiple craniofacial, skeletal and limb malformations, auditory and renal deficits, and a partially penetrant PH phenotype (Cappello et al., 2013). PH phenotype varies from patient to patient, from thin sheets of gray matter to large neuronal masses resembling laminar heterotopia in most severe cases. Although no PH were found in the brains of Fat4 / and Dchs1 / mice, brain embryonic knockdown of either Fat4 or Dchs1 resulted in neuronal mispositioning with electroporated cells accumulating in proliferative areas and fewer cells reaching the cortical plate (Cappello et al., 2013). Functional studies suggested that the ectopic neuronal positioning is not solely due to impaired migratory capabilities but instead arises as a consequence of disrupted proliferation of progenitors and blockade of their subsequent differentiation. FAT4 and DCHS1 are upstream regulators of the canonical Hippo signaling pathway controlling cell proliferation and polarity via MST1/2 kinase-mediated phosphorylation of YAP and NDR1/2 kinases. NDR1/2 kinases are modulated by MOB1/MOB2 and regulate phosphorylation of MEKK proteins, known modulators of FLNA phosphorylation. Interestingly, a female patient with bilateral nodular PH and biallelic mutations in MOB2 was recently described (O’Neill et al., 2018). Brain embryonic knockdown of Mob2 was found to alter neuronal positioning in the developing mouse cortex and nucleusecilia coupling. Strikingly, in vitro knockdown of Mob2 also resulted in enhanced FLNA phosphorylation. This patient phenotype and functional studies in mice suggest a potential link between FLNA and Hippo signaling pathways, which may contribute to the etiologies of PH. A de novo missense mutation in the C6orf70 gene, mapping the minimal critical deleted 6q27 region, was identified in a patient with developmental delay, epilepsy, and PH. Brain embryonic knockdown of C6orf70 resulted in altered neuronal positioning with electroporated cells accumulating along the walls of the lateral ventricles and fewer cells reaching the cortical plate (Conti et al., 2013).

26.3.4 Variant lissencephalies and mutations in ARX and RELN X-linked lissencephaly with abnormal genitalia (XLAG) is a rare genetic disorder that is caused by mutations in the gene ARX (Kato et al., 2004). It is the only known transcription factor to be causative to lissencephaly. XLAG is also distinct from classical LIS in that affected males display abnormal genitalia and a neocortical lamination pattern that is clearly different from aberrant cortical lamination associated with mutations in either LIS1 or DCX. Specifically, the neocortex in individuals with ARX mutations appears to show a three-layered cortex with an abnormal number of neurons in the marginal zone, lacking an internal fiber layer seen in the brains of individuals with DCX or LIS1 mutations (Forman et al., 2005). ARX mutations are associated with a wide spectrum of neurodevelopmental disorders with striking pleiotropy (Kato et al., 2004). The clinical features associated with mutation of ARX range from severe lissencephaly with microcephaly and seizures, to mild mental retardation and autism without lissencephaly (Kato et al., 2004). The phenotypes of mice with

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different Arx mutations, similar to the human phenotypes, reveal disorders in neuronal migration with a range of phenotype severity depending on the specific mutation in the gene (Kitamura et al., 2002, 2009; Marsh et al., 2009). The phenotypes range from embryonic death to learning and seizure disorders with only subtle disruption in neurodevelopment. Migration of GABAergic interneurons is consistently affected by mutations in Arx (Marsh et al., 2009). RNAi knockdown of Arx in both migrating neocortical pyramidal neurons and migrating GABAergic interneurons further shows that Arx has none cell-autonomous functions, as predicted for a transcription factor (Friocourt et al., 2006, 2008). The genes that are downstream of the Arx transcription factor include genes important to neurogenesis, migration, and axonal growth. Genes previously associated with clinical features identified in patients with ARX mutations including autism, epilepsy, and mental retardation are also downstream of ARX (Fulp et al., 2008). As presented earlier, in Chapter 24, patterned migration in the cortex and cerebellum critically depends upon signaling in the reelin pathway. Consistent with this role identified in mouse mutants, autosomal recessive mutations in the RELN gene in humans have been linked to a lissencephaly (Hong et al., 2000). Lissencephaly caused by RELN mutations is rare and causes LCH. LCH can also be caused by some, but not all, DCX and LIS1 mutations (Ross et al., 2001), indicating that LCH is a genetically heterogeneous class of NMD and that DCX and LIS1 may act partially in a shared genetic pathway with reelin signaling. Consistent with this interaction, reelin and Lis1 have been shown to genetically interact with mouse mutants (Assadi et al., 2003; Zhang et al., 2007).

26.3.5 Cobblestone malformations and mutations in dystroglycan genes Cobblestone lissencephaly, also known as type 2 lissencephaly, is caused almost exclusively by breaks in the basement membrane at the pial surface (Fig. 26.3C). As a consequence, neurons migrate into and beyond the marginal zone of the cortex. The result of this overmigration is a near complete loss of normal cortical lamination with heterotopic organizations of cells in the marginal zone (Fig. 26.3D; Muntoni and Voit, 2004; Olson and Walsh, 2002). Because the surface of the cortex loses sulci and gyri as a result of this malformation, it is classified as a type of lissencephaly. Cobblestone lissencephaly is typically associated with hypoplasia and/or malformation of the cerebellum. Cobblestone lissencephaly is a feature of a set of syndromes that include dysgenesis of the retina and muscular dystrophy including muscle eye brain disease, WalkereWarburg syndrome, and Fukuyama-type congenital muscular dystrophy (Muntoni and Voit, 2004; Olson and Walsh, 2002). Patients with cobblestone lissencephaly typically have severe psychomotor retardation, seizures, visual loss, and congenital muscular dystrophy. The genetic causes of cobblestone lissencephaly are typically autosomal recessive mutations in any of six genes involved in the glycosylation of a-dystroglycan: POMT1, POMT2, POMGNT1, FCMD, FKRP, and LARGE (Manzini et al., 2008; Satz et al., 2010). Studies using mouse models show that during cortical development a-dystroglycan stabilizes the glial limitans basement membrane (Satz et al., 2010) and thereby prevents neuronal migration into the marginal zone.

26.3.6 Focal cortical dysplasias and dysplastic megalencephaly and mutations in mTOR, PIK3CA, DEPDC5, AKT3, NPRL3, and PIK3R2 FCD II and DMEG have been shown to be the result of both germline, but most commonly, mosaic somatic mutations in genes that encode proteins in the mTOR pathway. To date, the mutated genes identified in these cortical malformation disorders include mTOR, RPS6, PIK3CA, DEPDC5, AKT3, NPRL3, and PIK3R2 (Poduri et al., 2012; Riviere et al., 2012; Lee et al., 2012; Jansen et al., 2015; D’Gamma et al., 2015; Mizraa et al., 2016; Lee et al., 2019; Pelorosso et al., 2019; Baldassari et al., 2019). All of these converge on the mTORC1 protein complex that is a responsive regulator of cellular anabolic and catabolic processes that regulate cellular growth in response to changes in nutrients and growth factor signaling. The sufficiency of somatic mutation in these genes to cause focal disruptions in migration and cellular growth in developing neurons and glia has been demonstrated in mouse models where somatic introduction of human mutant genes into embryonic mouse neural progenitors leads to the development of dysplastic neurons, altered neuronal migration, and spontaneous seizures (Poduri et al., 2012; Riviere et al., 2012; Lee et al., 2012). Importantly, studies in mouse models and in human samples from FCD II tissue resected in epilepsy surgery indicate that a relatively small percentage of mutationbearing cells in a focal dysplastic region are sufficient to cause seizures that spread throughout the cortex. A direct relationship between mutations in the mTOR pathway and brain malformations that cause epilepsy was first appreciated from the discovery of mutations that cause TSC. Autosomal dominant mutations in the TSC1 and TSC2 genes result in an overactivation of the mTOR. Aberrant activation results from decreased suppression of mTOR signaling due to a decrease in either hamartin or tuberin function, the two proteins coded by the TSC1 and TSC2 genes, respectively. With constitutive activation of mTOR comes a dysregulation in cell growth control and aberrant phosphorylation of S6 kinase

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and S6 (Baybis et al., 2004; Crino, 2007). Sporadic-type II FCD has also been linked to changes in the same activation of mTOR pathways seen in TSC-related FCD (Orlova et al., 2010; Crino, 2007). The similarity in both the lamination and cellular disruptions and in the molecular signatures of cells in all type II FCDs initially indicates that they may arise from similar developmental disruptions (Crino, 2007). The mechanism leading from constitutive activation of mTOR to type II FCD is not entirely clear. One hypothesis is that FCD and DMEG represent clonal expansions of an aberrant neural progenitor with dysregulated growth, and support for such a model comes from the observation that cells within FCD are more likely to be clonally related (Hua and Crino, 2003). Such clonally related cells originating from progenitors with mutations in genes in the mTOR signaling pathway are likely dysregulated in many aspects of their development, not just in neuronal migration, as mTOR signaling is an integrator of several signaling pathways. The underlying causes of another type of FCD, FCD type I, a, and b, are the most enigmatic. A variety of forms, essentially any subtle focal disruption in lamination, without the associated dysmorphic neurons and glia observed in FCD II, make type I FCDs difficult to categorize accurately (Chamberlain et al., 2009). Surgical resection of type I FCD may also have a poorer outcome in terms of relieving seizures (Fauser et al., 2006; Krsek et al., 2009). The mechanism(s) that cause type I FCD may be varied and do not seem related to those mutations responsible for type II FCD. Cells in type I FCD do not share the molecular hallmarks of mTOR signaling shared by cells in type II FCD (Orlova et al., 2010), and it has been suggested that FCD type I malformations may also result from prenatal and perinatal brain injury (Krsek et al., 2010).

26.4 Summary and concluding remarks NMD, and MCD more broadly, has been associated with several untreatable or difficult to treat neurological deficits in children. The most common and nearly universal symptom of these disorders is pharmacologically intractable epilepsy. The causes of NMD vary significantly by the type of malformation predominating in a particular syndrome, but there are significant similarities in gene functions associated with specific NMD types, and mutations in common pathways have mechanistically linked different NMDs and MCDs that display shared cellular pathologies. Genetic disruption in microtubule-associated proteins or microtubules commonly causes lissencephaly and polymicrogyria. Mutations in genes related to the extracellular matrix involving a-dystroglycan almost uniformly cause cobblestone lissencephaly malformations, and mutations required for normal function of mTOR pathway are commonly linked to the now mechanistically related DMEG and FCD II malformations. PH, on the other hand, are related to disruptions in proteins linked to the actin cytoskeleton and cell adhesion. PMG malformations are associated with genetic mutations in the most varied class of genes, and this may be partially due to this malformation type sharing features and causes of many MCDs including lissencephaly, simplified lamination, cobblestone lissencephaly, and overgrowth syndromes, in which increased physical disruption of the pial surface can occur. Future challenges in the field of NMD include increasing the number of identified gene mutations that now appear to occur both as germline and, possibly more commonly, as de novo somatic mutations (Poduri et al., 2013). This will require additional efforts and improvements in single cell sequencing from human tissue. In addition, existing and new animal models need to be developed and used to provide new insights into possible treatment of NMD by identifying the specific defects in neural circuitry and physiology that could serve as therapeutic targets.

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588 PART | II Migration

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̀

Index ‘Note: Page numbers followed by “f” indicate figures, “t” indicate tables and “b” indicate boxes.’

A A-type lamins, 306 A1. See Auditory cortex (A1) A2B. See Adenosine receptor 2B (A2B) AAD. See Acute axon degeneration (AAD) AAV1. See Adeno-associated virus 1 (AAV1) ABD. See F-actin binding domain (ABD) Abelson tyrosine kinase, 164e165 abGCs. See Adult-born granule cells (abGCs) Abnormal chemosensory jump 6 (Acj6), 239e241 abNs. See Adult-born neurons (abNs) abPGNs. See Adult-born periglomerular neurons (abPGNs) ABPs. See Actin-binding proteins (ABPs) AC. See Anterior commissure (AC) aCC. See Anterior corner cell (aCC) Accessory olfactory bulb (AOB), 163, 163f Acetylation, 32e33, 62 ACh. See Acetylcholine (ACh) Achaeteescute family bHLH transcription factor 1 (Ascl1), 483 AChRs. See Acetylcholine receptors (AChRs) Acj6. See Abnormal chemosensory jump 6 (Acj6) ACKR3. See Atypical chemokine receptor (ACKR3) ACM. See Astrocyte-conditioned medium (ACM) Actin, 24e27, 59e60 a-actin, 25 b-actin, 25 g-actin, 25 actin-based motility, 260e261 actin-based motor myosin, 43 actin-binding protein Ena, 158e159 cytoskeleton, 225, 416e419, 418f regulators, 260e261 depolymerization, 9e10 dynamics, 25b during axon formation, 27e30 nucleator, 60 rearrangements, 260 Actin filaments. See Filamentous actin (F-actin) Actin-binding proteins (ABPs), 24, 30 Actin-depolymerizing factor (ADF), 25 Actin-interacting proteins, 159

Active transport mechanisms, 40 Activity-dependent exon. See Exon IV mechanisms, 264e265, 264f orientation of dendrite growth, 266, 267f synaptic plasticity, 45 Acute axon degeneration (AAD), 227 AD. See Alzheimer’s disease (AD) ADAMTS. See A disintegrin and metalloproteinase with thrombospondin motifs (ADAMTS) Adenine nucleotides, 25 Adenomatous polyposis coli (APC), 10e11, 15, 33, 184, 492 Adenosine receptor 2B (A2B), 90 Adenosine triphosphate (ATP), 332 ADF. See Actin-depolymerizing factor (ADF) Adherens junction (AJ), 466 Adhesion molecules, 332e333 gap junctions, 333 integrins, 332e333 and surface molecules, 519e521 ADP ribosylation factorelike 13B (ARL13B), 351 Advanced live-cell imaging techniques, 38e39 AEP. See Anterior entopeduncular area (AEP) Aequorea victoria. See Jellyfish (Aequorea victoria) AES. See Anterior extramural stream (AES) Afferent-derived neurotrophins limit size, 263 African elephant (Loxodonta Africana), 559 AIS. See Axon initial segment (AIS); Axon initiation segment (AIS) AIS markers, 37 AJ. See Adherens junction (AJ) AKT, 8e9 AKT/protein kinase B, 14 AL. See Antennal lobe (AL) ALM. See Anterior lateral microtubule (ALM) Alpha-dystroglycan (a-DG), 388 Alpha3 integrin, 332e333 Alpha3/Beta1 integrin receptor, 332e333 ALS. See Amyotrophic lateral sclerosis (ALS) Alternative splicing, 491e492 of Dab1, 491

Alzheimer’s disease (AD), 116, 220, 227 Ambystoma tigrinum. See Salamander (Ambystoma tigrinum) Amino acid motifs, 44 a-Amino-3-hydroxy-5-methyl-4isoxazolepropionic acid (AMPA), 520e521 Amontillado (Amon), 149 AMPA. See a-Amino-3-hydroxy-5-methyl-4isoxazolepropionic acid (AMPA) AMPARs. See a-Amino-3-hydroxy-5methyl-4-isoxazole propionic acid receptors (AMPARs) Amphid sheath glial cell (AMsh glial cell), 244e245 AMsh glial cell. See Amphid sheath glial cell (AMsh glial cell) Amyloid precursor protein (APP), 90, 149, 220 Amyloid-b (Ab), 220 Amyotrophic lateral sclerosis (ALS), 46, 116, 221 Ang-1. See Angiopoietin-1 (Ang-1) Angelman syndrome, 278 Angiopoietin-1 (Ang-1), 472 Anosmin, 68 Antennal lobe (AL), 233 Anterior commissure (AC), 134e135 axons, 131f Anterior corner cell (aCC), 243 Anterior entopeduncular area (AEP), 347e348 Anterior extramural stream (AES), 382, 409 Anterior lateral microtubule (ALM), 247 Anterior ventral microtubule (AVM), 149, 151f Anterioreposterior axis, mapping mechanisms along, 136e138 Anterograde transport of mitochondrion, 222f Anteroposterior axis (AeP axis), 178 Antisense oligodeoxynucleotides (asODNs), 314 AOB. See Accessory olfactory bulb (AOB) AeP axis. See Anteroposterior axis (AeP axis) APC. See Adenomatous polyposis coli (APC) Apical dendrite initiation and outgrowth of cortical pyramidal neurons, 265e266 Apical progenitor, 294 aPKC. See Atypical protein kinase C (aPKC)

589

590 Index

Aplysia, 57e58 Apolipoproliprotein E receptor 2 (ApoER2), 330e331, 369, 394, 487 APP. See Amyloid precursor protein (APP) Arbor size control by neurotransmission, 263e264 Arborization, cytoskeletal dynamics during, 35e37 Arc. See Activity-regulated cytoskeletal protein (Arc) Arealization of cortex, 531e532 Aristaless-related homeobox (ARX), 345e346, 452, 488 ARL13B. See ADP ribosylation factorelike 13B (ARL13B) Arp2/3 activity, 261 complex, 260 proteins, 28e29 Artificial scaffold-mimetic materials, 470 ARX. See Aristaless-related homeobox (ARX) AS4. See Alternatively spliced segment 4 (AS4) Ascl1. See Achaeteescute family bHLH transcription factor 1 (Ascl1) ASD. See Autism spectrum disorder (ASD) asODNs. See Antisense oligodeoxynucleotides (asODNs) ASTN1. See Astrotactin1 (ASTN1) Astrocytes, 204e205, 208e209, 479 in injured brain, 471 in normal brain, 468e469 Astrotactin1 (ASTN1), 420 Atoh1 domain, 405e407 ATP. See Adenosine triphosphate (ATP) Attraction, 96e97 by Netrins in Drosophila, 98 by UNC-6 in C. elegans, 97e98 Attractive Ephrin/Eph signaling, 125 ATUM-SEM. See Automated tape-collecting ultramicrotome SEM (ATUM-SEM) Atypical chemokine receptor (ACKR3), 350e351 Atypical protein kinase C (aPKC), 6e7, 11e12, 188e189 Auditory system, 135 Autism spectrum disorder (ASD), 116, 164, 278 Autonomic dysreflexia, 202 Autophagy, 223 Autosomal recessive disorder, 99e100 Autosomal-dominant mutations, 223 AVM. See Anterior ventral microtubule (AVM) Axed mutant, 228 Axon degeneration, 217, 225e229 axon pruning and, 225e229 defects in motor proteins cause, 221 pathological, 227 molecular mechanisms, 227e229 Axon formation actin dynamics during, 27e30 microtubules dynamics during, 32e35

Axon growth and branching cell biological mechanisms, 58e65 extracellular regulation during neural development, 65e71 Intracellular signaling mechanisms mediates, 71e73 Axon guidance, 147, 157e158, 162 in C. elegans, 153 nerve cord, 151f commissural, 150e154 commissural axon guidance defects, 150 ephrin/Eph signaling in Cis interactions, 127 membrane distribution, 126e127 trafficking, 127e128 midline, 156e157 molecules, 147e148, 156e157 Netrin function in, 92e98 proteins, 109 signal transduction of forward signaling, 128e130, 129f of reverse signaling, 129f, 130 sonic hedgehog in, 176e182 TGF-b superfamily members in, 183e184 vertebrate midline, 152e153 Wnts in, 183e184 Axon guidance cues cross-talk between axon guidance cues Shh induces response of commissural axons to semaphorin repulsion, 192 Shh regulates Wnt signaling in postcrossing commissural axons, 193 TGF-b family member unc-129 regulates Unc6/Netrin signaling, 193 morphogens as, 176 in neural tube, 178f Axon initial segment (AIS), 30, 45 Axon initiation cytoskeletal dynamics role in, 9e10 major signaling pathways in AKT/protein kinase B, 14 LKB1 and downstream kinases SAD-A/B and MARK1e4, 10e11 PAR3ePAR6eAPKC, 11e12 PI3K and PTEN signaling during axon specification, 13e14 Ras-and Rho-family of small GTPases, 12e13 in vitro, 4e5 in vivo, 5 Axon initiation segment (AIS), 3e4 Axon maintenance, 217 autophagy/lysosome pathway role in maintaining axonal homeostasis, 223e224 glial cells role in, 224 membrane transport and insertion, 222e223 mitochondria transport role in, 221e222 regulations of microtubule stability and organization during, 220e221 track positions and other structural features, 224e225 UPS in, 223 Axon pruning, 217, 225e229

developmental, 225e227 large-scale, 225f Axon regeneration extrinsic mechanisms, 208, 210t growth factors, 205e207 inhibitors of CNS, 204e205 immune-mediated neurorepair mechanisms, 208e209 in injured CNS vs. PNS, 203e204 intrinsic mechanisms of dorsal root ganglion neuron, 203e204, 208 in retinofugal system, 207, 210t Axonal branching, 161 Axonal homeostasis, proteasome and autophagy pathways in autophagy/lysosome pathway role in maintaining, 223e224 UPS in axon maintenance, 223 Axonal microtubules, 61 Axonal outgrowth and branching, 161e162 Axonal transport, 63e64, 221 Axonogenesis, 23 Axons, 3e4, 57, 134, 138, 176, 218, 226, 232 of mammalian VCN, 135 motor proteinebased transport in, 40e43 organelles and structural components, 218f Ab. See Amyloid-b (Ab)

B B-type lamins, 306 BAC. See Bacterial artificial chromosome (BAC) BAPTA. See 1,2-Bis(2-aminophenoxy) ethane-N,N,N0 ,N0 -tetraacetic acid (BAPTA) BAR domain, 63 Basal pontine nucleus (BPN), 409 Basal process, 481 Basal progenitors, 289e290, 295, 484 Basic helixeloopehelix (bHLH), 298, 405e406, 482 bHLHePAS transcription factor, 155e156 transcription activators, 483 Basket cells, 403e404, 408e409 Basket in Drosophila, 220 BBB. See Bloodebrain barrier (BBB) Bcl11a (zinc finger transcription factor), 484 BD. See Binding domain (BD) BDNF. See Brain-derived neurotrophic factor (BDNF) BDSC. See Bloomington Drosophila Stock Center (BDSC) Bergmann glia (BG), 407e408 Beta-amyloid (Ab). See Amyloid-b (Ab) bFGF. See Basic fibroblast growth factor (bFGF) BG. See Bergmann glia (BG) bHLH. See Basic helixeloopehelix (bHLH) Bilateral periventricular nodular heterotopia (BPNH), 582e583 Binary Ephrin/Eph signaling, 132e135 Binary mode, 125 Biochemical analysis, 92

Index

Bipolar cells, 5 Bipolar migrating neurons along radial glial fibers, 290 Blood vessels blood vesseleguided neuronal migration, 469 in injured brain, 472 in normal brain, 469 Bloodebrain barrier (BBB), 202e203, 473 BMI. See Bicuculline methiodide (BMI) BMPRs. See Bone morphogenetic protein receptors (BMPRs) BMPs. See Bone morphogenetic proteins (BMPs) Bone morphogenetic protein receptors (BMPRs), 183 BMPRIA, 184 BMPRIB, 184 Bone morphogenetic proteins (BMPs), 176, 183 BMP7, 183e184 BMP7:GDF7 repels spinal cord commissural axons, 183e184 BPN. See Basal pontine nucleus (BPN) BPNH. See Bilateral periventricular nodular heterotopia (BPNH) Brain circuits modulation, 565e567 Brain development, 452 normal, 451e452 pathological, 451e452 Brain evolution, neuronal migration on neuronal migration types in vertebrate brain development, 555e557 radial migration impact on brain evolution, 557e563 tangential migration along CNS, 571e572 impact on brain evolution, 563e572 Brain stem, pathfinding in, 135 Brain-derived neurotrophic factor (BDNF), 7, 65, 130, 261e262, 347, 469, 501 Branching, 161e162 BrdU. See Bromodeoxyuridine (BrdU) Brn-1, 482 Brn-2, 482 Bromodeoxyuridine (BrdU), 382, 482 BTB domain containing 3 (BTBD3), 266 BTBD3. See BTB domain containing 3 (BTBD3)

C c-Jun N-terminal kinase (JNK), 11, 220, 299e300 C-terminal domain (CTD), 111 Ca2+-dependent mechanism, 187e188 Ca2+/calmodulin-dependent protein kinase II (CaMKII), 184 Cad8. See Cadherin-8 (Cad8) Cadherin-2 (Cad2). See N-cadherin Cadherin-8 (Cad8), 391e392 Cadherin-related clustered protocadherins (cPcdhs), 262 Caenorhabditis elegans. See Roundworm (Caenorhabditis elegans)

CajaleRetzius cells (CeR cells), 330, 367, 440e441, 567e569 for dorsal telencephalon, 368 extent of cortex covered by hem-derived, 368 functions of, 367 origins of, 367e368 putative guidepost neurons, 440e442 recruitment of hem-derived, 368 tangential dispersion of, 368e369 toward more generic role of, 442 Calcium (Ca), 72 Ca++ ions, 72 fluctuations, 350 Calcium-responsive transactivator (CREST), 263 Calcium/calmodulin-dependent protein kinase II (CaMKII), 263 CaMKIIa, 262 Callosal commissure axons (CC axons), 131f Calmodulin-dependent protein kinases (CaMK), 262 Calmodulin-regulated spectrin-associated proteins family (CAMSAP1e3), 61 Calretinin (CR), 370e371 CaMK. See Calmodulin-dependent protein kinases (CaMK) CaMKII. See Ca2+/calmodulin-dependent protein kinase II (CaMKII); Calcium/ calmodulin-dependent protein kinase II (CaMKII) cAMP. See Cyclic adenosine-30 , 50 -monophosphate (cAMP) cAMP response element-binding protein (CREB), 203, 263 CAMs. See Cell adhesion molecules (CAMs) CAMSAP1e3. See Calmodulin-regulated spectrin-associated proteins family (CAMSAP1e3) Candelabrum cells, 403e404 Canoe (Cno), 159 Canonical bone morphogenetic protein signaling, 183 Canonical Rho GTPases, 24 Canonical Shh signaling, 176, 177f Capulet (capt), 158e159 Cargo transport, 260 Casein kinase Ia (CKIa), 184 CASK. See Ca2+/calmodulin-dependent serine protein kinase (CASK) Caspases, 251 Catastrophies, 31 Caudal ganglionic eminence (CGE), 345e346, 370e371, 538e539, 566 interneuron, 493 Caudal migratory stream (CMS), 347e348 CC. See Corpus callosum (CC) CC axons. See Callosal commissure axons (CC axons) CCCTC-binding factor (CTCF), 307e308 CCP1. See Cytosolic carboxypeptidase 1 (CCP1) CDC42. See Cell division control protein 42 (CDC42)

591

Cdh1-APC, 64e65 CDK. See Cyclin-dependent kinase (CDK) CDK inhibitors (CKIs), 298 Cdk5. See Cyclin kinase 5 (Cdk5) Cell displacement, 368 fate decisions, 405 migration, 111, 160e161 Netrin function in, 92e98 polarity, 21e23, 160e161 molecular mechanisms regulating, 297e301, 297f Cell adhesion molecules (CAMs), 7, 68e69, 388 TAG-1, 391e392 Cell biology biological mechanisms growth cones, 58 regulation of cytoskeleton assembly, 58e62 of dendritic growth, 235e237 dynein-dependent trafficking controls dendritic branching, 235e236, 236f microtubule polarity differs between dendrites and axons, 235 role of secretory pathway and Golgi outposts in dendritic elaboration, 236e237 of dendritic growth, 259e263 actin cytoskeleton regulators, 260e261 dendrite elaboration requires satellite secretory pathway, 261e262 intracellular cascades role, 262e263 microtubule network regulators, 259e260 powering dendrite growth, 262 RNA translation in dendrites, 262 of dendritic pruning local endocytosis and compartmentalized calcium transients, 252 microtubule disassembly, 251e252 Cell division control protein 42 (CDC42), 24, 350e351 Cellecell adhesion, 542 contacts, 542 Cellular “cargoes”, 219 diversity, 555e556 mechanisms controlling shifts in migratory modes, 543e544 polarization mechanisms, 23 Cellular RA binding protein I (CRABPI), 389 Central nervous system (CNS), 3e4, 57e58, 201, 233, 515, 555e556 axon regeneration in injured CNS vs. PNS, 203e204 inhibitors, 204e205 tangential migrations along, 571e572 Central pathfinding C. callosum and anterior commissure, 135 optic chiasm, 135 Centrosome-dependent “nuclear pulling” model, 292, 295

592 Index

Centrosomes, 292 centrosomal-microtubule nucleation model, 35 nucleus and, 311 Centrosomin, 236e237 Cerebellar germinal zones, 405e407 early patterning, 405 Rh1 derivatives, 407 RL and Atoh1 domain, 405e406 ventricular zone and Ptf1a domain, 406e407 Cerebellar granule neurons (CGNs), 5, 23, 403e404, 411, 417e418 actin cytoskeleton, 416e419 cytoskeletal organization of, 411e420 glial-guided migration, 413f microtubule cytoskeleton and associated motors, 414e416 microtubule-actin cross talk, 419e420 migration diversity after establishment of secondary germinal zone, 411e412 two-stroke motility paradigm, 413e414 Cerebellar MBs, migration deficits in, 424e425 altered GZ exit and migration in cerebellar tumorigenesis, 426f Cerebellar nuclei (CNs), 403e405, 410e411 Cerebellar system developing cytoskeletal organization of CGNs, 411e420 diverse migration pathways and guidance cues, 405e411 cerebellar germinal zones, 405e407 cerebellum origin and migration, 406f migration of minor ventricular zone derivatives, 408e409 migration of precerebellar nuclei, 409e410 migration of Purkinje cells, 407e408 migration of upper RL derivatives, 410e411 facets of CGNs polarity, 420e423 ASTN1 and JAM-C, 421f CGNs recognition/adhesion, 420e422 foxo polarization pathway, 423 transcriptional activation, 424f Zeb1-Pard6/3A transcriptional pathway, 423 migration deficits in cerebellar MBs, 424e425 Cerebellum, 403 and precerebellar system, 404f Cerebral cortex arealization of cortex, 531e532 cortical columns constitute cortical areas, 532 genetic and cellular mechanisms controlling shifts in migratory modes, 543e544 interneurons integration into cortical columns, 538e543 migration of related projection neurons into same minicolumn, 533e535 lack of evidence for sister projection neurons migration, 534e535

sister projection neurons migrates into same minicolumn and intersynapse, 535 minicolumns constitute columns, 532e533 projection neurons integration into cortical minicolumns, 535e538 Cerebrospinal fluid (CSF), 469 CF. See Climbing fiber (CF) CGE. See Caudal ganglionic eminence (CGE) cGMP. See Cyclic guanosine-30 , 50 -monophosphate (cGMP) CGNs. See Cerebellar granule neurons (CGNs) Chain migration of neurons, 466, 470e471 CharcoteMarieeTooth disease (CMT2), 221 Chemoattractants, 96 Chemokine 12 (CXCL12), 348e349, 352, 388e389, 393, 472 Chemokines, 522e524 Chemokinetic factors, 518e519 Chemotropic cues, 521 Chicken ovalbumin upstream promoter transcription factor II (COUP-TFII), 371, 500 a-Chimaerin, 128e129 Chimeric receptors, 89 Chl1. See Close homolog of L1 (Chl1) Chondroitin sulfate proteoglycans (CSPGs), 204 Choroid plexus (CP), 160, 407 Chromatin, 306e307 fiber, 306e307 remodeling factors and dendritic development, 241 structure, 241 Chromosome conformation capture (3C), 307e308 Chronic axon degeneration, 227 CI. See Conditioning injury (CI) Cis interactions, 127 CKIs. See CDK inhibitors (CKIs) CKIa. See Casein kinase Ia (CKIa) Class A plexins (PlexinAs), 111 Class I dendritic arborization neurons, 266 Class Ia PI3-kinase (PI3KcIa), 13 Clawed toad (Xenopus laevis), 10e11, 558 Climbing fiber (CF), 115e116, 404 Climbing migration mode, 369 CLIP-170, 260e261 Clip115/Clip170 protein, 33e34 Close homolog of L1 (Chl1), 423 Clubfoot, 130 Clustered Pcdhs, 69 Clutch hypothesis, 27 CMMs. See Congenital mirror movements (CMMs) CMS. See Caudal migratory stream (CMS) CMT2. See CharcoteMarieeTooth disease (CMT2) Cnidarians, 126 CNS. See Central nervous system (CNS) CNs. See Cerebellar nuclei (CNs) Cntn5. See Contactin-5 (Cntn5)

Cntnap2, 278 Coadaptation, 128, 137e138 Cobblestone LIS, 579 lissencephaly, 584 Cognitive disorders, 544 Cohesinopathies, 308e309 Collapsin response mediator protein-2 (CRMP-2), 15, 34, 61, 117 Collapsin-1. See Semaphorin 3A (Sema3A) comm mutant phenotype, 153 comm;robo double mutant, 153 comm;slit double mutant, 153 Commissural axons, 152, 180e181 Shh binding to Boc attracts, 179e180 Commissural neurons, 87, 176, 178e180, 182e184 Commissures, 87 Compartmentalized calcium transients, 252 Computational modeling, 138e139 Conditional knock-in, 259 Conditional knockouts, 259 of Ngn2, 336 Conditioning injury (CI), 203 Congenital disorders, schizophrenia, 116 Congenital mirror movements (CMMs), 99 Connexin (Cx), 333 Cx26, 333 Cx43, 314 Contactin 2. See Transient axonal glycoprotein 1 (TAG-1) Contactin-5 (Cntn5), 271e273 Contralateral (nasal) axons, 139 Contralateral retinal ganglion cell axons, 181e182, 181f Controlling initiation of radial migration, 299 Cordon-bleu (Cobl), 60, 260 CoREST, 494 Cornelia de Lange syndrome, 309 Corpus callosum (CC), 135, 447e449 callosal axons guidance, 448f roles of glial cells in development of, 447e448 tangentially migrating neurons in development of, 448e449 Corridor cells, 570e571 migration, 446e447 Cortical columns constitute cortical areas, 532 interneurons integration into, 538e543 Cortical GABAergic INs, 538e539 Cortical neuroblast development, 561 Cortical neurons, 327 Cortical plate (CP), 289e290, 323e324, 325f, 327, 347e348, 480 Cortical projection neurons, 324e326 Corticocollicular projections, 140 Corticospinal tract axons (CST axons), 99, 134, 186 Corticothalamic axons (CTAs), 442 COUP transcription factor 1 (COUP-TFI), 484

Index

COUP-TFII. See Chicken ovalbumin upstream promoter transcription factor II (COUP-TFII) Coupling between neuronal differentiation and migration, 298e299 CP. See Choroid plexus (CP); Cortical plate (CP) cPcdhs. See Cadherin-related clustered protocadherins (cPcdhs) CR. See Calretinin (CR) CeR cells. See CajaleRetzius cells (CeR cells) CRABPI. See Cellular RA binding protein I (CRABPI) Cranial motor neurons, 389 Cre recombinaseebased technologies, 259 CREB. See cAMP response element-binding protein (CREB) CREST. See Calcium-responsive transactivator (CREST) CRISPR/Cas9, 25 CRMP-2. See Collapsin response mediator protein-2 (CRMP-2) CRNs. See CajaleRetzius neurons (CRNs) Cross-talk between axon guidance cues Shh induces response of commissural axons to semaphorin repulsion, 192 Shh regulates Wnt signaling in postcrossing commissural axons, 193 TGF-b family member unc-129 regulates Unc6/Netrin signaling in C. elegans, 193 CSF. See Cerebrospinal fluid (CSF) CSPGs. See Chondroitin sulfate proteoglycans (CSPGs) CST axons. See Corticospinal tract axons (CST axons) CTAs. See Corticothalamic axons (CTAs) CTCF. See CCCTC-binding factor (CTCF) CTD. See C-terminal domain (CTD) Cue-induced local protein synthesis, 262 CXCL12. See Chemokine 12 (CXCL12) Cyclic adenosine-30 , 50 -monophosphate (cAMP), 72, 203 Cyclic guanosine-30 , 50 -monophosphate (cGMP), 72 Cyclic nucleotides as second messengers and modulators, 72e73 Cyclin kinase 5 (Cdk5), 300, 335, 370, 386, 394, 481e482 Cyclin-dependent kinase (CDK), 298 Cytokinesis, 330 Cytoplasm, 218f Cytoplasmic CC2 sequence (LPPPP), 158e159 Cytoplasmic proteins, 64 Cytoskeletal dynamics, 35e37, 261 role in axon initiation and growth, 9e10 Cytoskeletal genes, 491e492 Cytoskeletal regulators, 333e335 Cdk5, 335 DCX, 334 filamin A, 334e335 Lis1, 333e334

Cytoskeleton, 158e159, 161 assembly regulation, 58e62 actin, 59e60 interaction between F-actin and microtubules, 62e63 membrane trafficking and axonal transport, 63e64 microtubules, 60e62 protein translation and stability, 64e65 dynamic remodeling of, 352 dynamics as nuclear drivers, 314 neuronal, 259 role in neuronal polarity establishment, 24e39 actin, 24e27 actin dynamics during axon formation, 27e30 cytoskeletal dynamics during dendritic growth and arborization, 35e37 (membrane) trafficking role during neuronal polarization, 39e46 microtubules, 30e32 microtubules dynamics during axon formation, 32e35 neuronal cytoskeleton and the growth cone, 26fe27f subcellular cytoskeletal specializations, 37e39 trafficking during early neuronal development, 39e40 barriers for functional domains segregation, 45 motor proteinebased transport in axons and dendrites, 40e43 protein stabilization and degradation, 46 RNA transport and local translation, 44e45 secretory and endosomal pathway, 43e44 trafficking during axonogenesis, 41fe42f Cytosolic carboxypeptidase 1 (CCP1), 351

D D-APV. See D-(e)-2-Amino-5phosphonopentanoic acid (D-APV) Dab1, 301, 491 Dab1. See Disabled-protein 1 (Dab1) Dam. See DNA adenine methyltransferase (Dam) Damaged mitochondria, 224 Dar1. See Dendritic arbor reduction 1 (Dar1) DBX1. See Developing brain homeobox 1 (DBX1) DCC. See Deleted in colorectal cancer (DCC) DCLK. See Doublecortin-like kinase (DCLK) DCs. See Dorsal columns (DCs) DCX. See Doublecortin (DCX) Dectin-1, 209 Deficits, 278 Deleted in colorectal cancer (DCC), 89e90, 301, 352e353, 444, 521e522 receptor, 384, 392 Delta-Notch signaling, 558e559

593

Dendrites, 3e4, 232 compartmentalization emergence, 277e278 patterning membrane excitability of dendritic compartments, 278 subcellular patterning of synaptic inputs along dendritic domains, 277e278 dendrite-specific attraction, 265e266 elaboration requires satellite secretory pathway, 261e262 laminar targeting in retina, 269 morphogenesis, 261 morphology, 115 in Drosophila, 232e233 motor proteinebased transport in, 40e43 regeneration after injury, 252 sampling, 265 self-avoidance, 258f, 275e277, 276f targeting, 269, 271e273, 272f Dendritic arbor reduction 1 (Dar1), 237e238 Dendritic arborization neurons, 271 Dendritic field formation control, 242e245 coarse and specific control of PN dendritic targeting, 244 combinatorial ligandereceptor complex guides dendritic branches, 244 Glial control of dendritic targeting, 244e245 Slit and Netrin signaling during midline dendritic guidance, 242e244 Dendritic field formation I control, 263e265 activity-dependent mechanisms, 264e265, 264f afferent-derived neurotrophins limit size, 263 arbor size control by neurotransmission, 263e264 Dendritic field formation II control, 245e248, 265e267 activity-dependent orientation of dendrite growth, 266, 267f apical dendrite initiation and outgrowth of cortical pyramidal neurons, 265e266 interactions between dendrites generate evenly covered territories dendritic self-avoidance, 245e247, 246f Dendritic tiling, 247, 248f positional cues shape asymmetric dendritic arbors in mouse retina, 266e267 scaling growth of arbors and maintenance of evenly covered territor, 248 Dendritic field formation III control, 268e273 integrated, multistep model for synaptic wiring in retina IPL, 273 laminar targeting of retinal dendrites, 269e271 local recognition mechanisms to control synapse selectivity, 273 proto-IPL formation by retinal amacrine cells, 269 transcriptional control of laminar-specific targeting of dendrites in retina, 271e273

594 Index

Dendritic growth, cytoskeletal dynamics during, 35e37 Dendritic mitochondria, 262 Dendritic morphology, transcriptional control of, 237e241 Dendritic outgrowth and branching, 161e162 Dendritic pruning cell biology of, 251e252 signaling mechanisms for, 251 similarities between injury-induced axon degeneration and, 252 Dendritic remodeling, 249e252 cell biology of dendritic pruning, 251e252 signaling mechanisms for dendritic pruning, 251 similarities and differences in dendrite development, dendrite regrowth after pruning, and dendrite regeneration after injury, 252 similarities between dendrite pruning and injury-induced axon degeneration, 252 Sox14 and mical function downstream of ecdysone receptor, 249e251 transforming growth factor-b signaling and ecdysone receptor expression, 249 Dendritic self-avoidance, 245e247 Dendritic tiling, 247, 248f Dendritogenesis, 264e265 Dentate gyrus (DG), 440 Departure from ventricular zone, 294e296 Depolymerization dynamics of microtubules, 31 Derailed function (Drl function), 186e187 Desert hedgehog (Dhh), 176 Developing brain homeobox 1 (DBX1), 345e346 Developing cerebral cortex migratory behaviors during radial migration in, 290e297 radial migration cortical projection neurons, 324e326 factors regulating radial migration of cortical neurons, 330e336 migrating neurons trajectory in developing brain, 326e328 migration modes, 328e329 neocortex organization, 326 radial migration in developing human neocortex, 330 Developmental axon pruning, 225e227 DG. See Dentate gyrus (DG) Dhh. See Desert hedgehog (Dhh) DIAP1. See Drosophila inhibitor of apoptosis (DIAP1) Dictyostelium discoideum, 310e311 Diffusion, 39e40 Disabled-protein 1 (Dab1), 278, 330e331, 481e482 Dishevelled (Dvl), 184, 390 Dishevelled1 (Dvl1), 11 Dissociated neuronal culture systems, 23 Distal axonopathy, 227 Distal-less homeobox (DLX), 345e346

Dlx1/2, 370 dKO. See Double knockout (dKO) dLGE. See Dorsal region of LGE (dLGE) dLGN. See Dorsal lateral geniculate nucleus (dLGN) DLX. See Distal-less homeobox (DLX) DMEG. See Dysplastic megalencephaly (DMEG) DNA adenine methyltransferase (Dam), 308 DNA methyltransferase (Dnmt1), 500 Dock/ELMO complex, 180 Dorsal columns (DCs), 202f, 203 Dorsal lateral geniculate nucleus (dLGN), 139 Dorsal region of LGE (dLGE), 352e353 Dorsal root ganglion (DRG), 28, 60, 202f, 203 Dorsal ventricular ridge (DVR), 570e571 Dorsoventral axis (DeV axis), 176 mapping mechanisms along, 138 Double knockout (dKO), 482 Double-stranded RNA (dsRNA), 242 Doublecortin (DCX), 34, 300, 334, 352, 370, 482e483, 485 Doublecortin-like kinase (DCLK), 334, 370 Down syndrome, 278 Down syndrome cell adhesion molecule (Dscam), 66e67, 69, 90 Dscam1, 245e247 Dreadlocks (Dock), 158 Drebrin, 62 DRG. See Dorsal root ganglion (DRG) Drifter (Dfr), 155e156 DRN. See Dorsal raphe nuclei (DRN) Drosophila, 57e58, 68e69, 132, 152e153, 226e228, 233 Abrupt, 266 CNS, 186 commissural axon guidance, 150e151 dendrite, 260 dendrite morphology, 232e233 dendritic arborization neurons, 260 sensory neurons, 233 ephrin, 132 genetic screens, 147e148 genetics, 158 midline crossing in, 151f motoneurons, 233 motor neurons, 156 Netrins attraction and repulsion in, 98 olfactory projection neurons, 233 PNS neurons, 237e239, 238f Robo1, 149 Wnt5 repels commissural axons from Drosophila posterior commissure, 186e187, 186f Wnt5, complexed with derailed, repels Drosophila mushroom body axons, 187 Drosophila inhibitor of apoptosis (DIAP1), 251 dSarm mutant, 228 dSarm/Sarm1 gene, 227e228 DSBs. See Double-stranded breaks (DSBs)

Dscam. See Down syndrome cell adhesion molecule (Dscam) dsRNA. See Double-stranded RNA (dsRNA) Dutt1, 152 DeV axis. See Dorsoventral axis (DeV axis) Dvl. See Dishevelled (Dvl) Dvl1. See Dishevelled1 (Dvl1) DVR. See Dorsal ventricular ridge (DVR) “Dying back”. See Distal axonopathy Dynactin, 221, 300 Dynamic assembly of F-actin, 60 Dynamic instability, 31 Dynamic remodeling of cytoskeleton, 352 Dynein, 63e64, 219e220, 300, 313 dynein-dependent trafficking controls dendritic branching, 235e236, 236f Dyrk1A, 278 Dysplastic megalencephaly (DMEG), 580, 584e585

E E-box, 298e299 E3-ubiquitin function, 64e65 Earlier-born neurons, 290 Early-born neurons, 294 EBs. See End binding proteins (EBs) EC. See Entorhinal cortex (EC) Ecdysone receptor expression during dendritic remodeling, 249 ECM. See Extracellular matrix (ECM) ECN. See External cuneate nuclei (ECN) EFF-1. See Epithelial fusion failure-1 (EFF-1) EGF. See Epidermal growth factor (EGF) EGL. See External granular layer (EGL) eIF4E1. See Eukaryotic initiation factor 4E1 (eIF4E1) Elaborated brains, radial migration on, 559e560 ELAVL family. See Embryonic lethal abnormal vision-like family (ELAVL family) Electron microscopy (EM), 289e290, 412 Electroporation, 394 EM. See Electron microscopy (EM) Embryonic ganglionic eminence, 370e371 Embryonic lethal abnormal vision-like family (ELAVL family), 488 Embryonic telencephalon, 370 Emerin, 309 EMT. See Epithelial-to-mesenchymal transition (EMT) Emys orbicularis, 565e566 Ena-VAS-Phomology (EVH1), 158e159 Ena/VASP proteins, 29, 260 End binding proteins (EBs), 24, 33e34, 61 EB3, 419e420 Endoplasmic reticulum (ER), 39, 63, 222, 234e235, 261 Endosomal sorting complex required for transport (ESCRT), 128 Endothelin, 71 endothelin-1, 519

Index

Engrailed transcription factors (Ens), 130 Ens. See Engrailed transcription factors (Ens) Enteric axons, 182 Enteric neurons, 182 Entorhinal cortex (EC), 115 Entorhinal pyramidal neurons, 440 Environmental cues, 23e24 Eph(s), 124, 127 EphAs, 124e125 EphA1, 124 EphA4, 134 EphBs, 124e125 fundamental action modes, 125 in invertebrate axon guidance C. elegans, 131e132 insects, 132 phylogeny, 125e126 receptors, 130 kinase activity, 128 in regeneration, 140 rules of interaction, 124e125 Ephrin-As, 124e125, 125f, 132 Ephrin-A2,3,5 triple mutants, 139 Ephrin-A5, 130 Ephrin-Bs, 124e125, 125f Ephrin-B2 expression, 133f expression, 133f Ephrin-B3, 134, 140 Ephrin/Eph signaling, 125 in axon guidance Cis interactions, 127 membrane distribution, 126e127 signal transduction of forward signaling, 128e130, 129f signal transduction of reverse signaling, 129f, 130 trafficking, 127e128 binary, 132e135 fundamental modes, 126f perspectives and open questions, 140e141 proportional, 136e140, 137f structures and binding affinities, 124, 125f Ephrins, 66e68, 124, 127, 132, 156e157, 521 fundamental action modes, 125 in invertebrate axon guidance C. elegans, 131e132 insects, 132 phylogeny, 125e126 in regeneration, 140 rev signaling, 130 rules of interaction, 124e125 Epidermal growth factor (EGF), 88e89, 519 Epilepsy, 116 Epistasis analyses, 159 Epithelial fusion failure-1 (EFF-1), 237 Epithelial-to-mesenchymal transition (EMT), 7e8 ER. See Endoplasmic reticulum (ER). See Estrogen receptor (ER) ER-to-Golgi intermediate compartment (ERGIC), 261 ErbB4-deficient neurons, 469

ERK/MAP kinases. See Extracellular signaleregulated/mitogen-activated protein kinases (ERK/MAP kinases) ERM proteins. See Ezrin/radixin/moesin proteins (ERM proteins) ESCRT. See Endosomal sorting complex required for transport (ESCRT) Estrogen, 278 Eukaryotic initiation factor 4E1 (eIF4E1), 491 Eukaryotic nucleus, 305e306 EVH1. See Ena-VAS-Phomology (EVH1) Evoked neural activity, 70 Excitatory glutamatergic projection neurons. See Pyramidal neurons (PNs) External cuneate nuclei (ECN), 381, 387 External granular layer (EGL), 405 Extracellular cues, 3e4, 6e7 regulating neuronal polarization and axon initiation Netrin-1 and Wnt control axon initiation in C. elegans, 6 polarized emergence of axon in Xenopus RGC, 6e7 underlying emergence of axon and dendrites, 7e8, 8f Extracellular matrix (ECM), 27, 60, 68, 92, 131, 204, 244e245, 467 molecules, 519 Extracellular molecules affecting migrating cells, 301 Extracellular regulation of axon growth and branching, 65e71 additional axon branching molecules, 71 CAM, 68e69 Glial cells and myelination, 69e70 guidance molecules, 66e68 neural activity, 70e71 NGF and neurotrophic factors, 65e66 Extracellular signaleregulated/mitogenactivated protein kinases (ERK/MAP kinases), 262 Extrinsic mechanisms, 208, 210t growth factors, 205e207 inhibitors of central nervous system, 204e205 Ezrin/radixin/moesin proteins (ERM proteins), 30

F F-actin, 58 dynamics, 60 interaction between F-actin and microtubules, 62e63 network, 27 F-actin. See Filamentous actin (F-actin) F-actin binding domain (ABD), 310 F-RGC, 266e267 Facial branchio motor (FBM), 381, 392 caudal migration of, 390e392 PCP, 390e391 role of meninges in tangential migration, 393e394 FAK. See Focal adhesion kinase (FAK)

595

Fak. See Focaladhesion kinase (Fak) FARP2. See FERM domain-containing guanine nucleotide exchange factor (FARP2) Fasciclin IV. See Collapsin-1 FBM. See Facial branchio motor (FBM) FCD. See Focal cortical dysplasias (FCD) FDJ. See Fimbria-dentate junction (FDJ) FedEx, 40e42 FERM domain-containing guanine nucleotide exchange factor (FARP2), 114 Fezf2 transcription factor, 134 FGF. See Fibroblast growth factor (FGF) FGF-2. See Fibroblast growth factor (FGFb) FGFb, 518e519 Fiberepath interactions. See Fiberetarget interactions (FT interactions) Fiberetarget interactions (FT interactions), 125 Fibronectin (FN), 155 Fibronectin and leucine-rich transmembrane proteins (FLRTs), 89, 537 FLRT2, 301 FLRT3, 537 Fibronectin type III (FNIII), 89 Fidgetin, 62 Filament capping, 60 Filamentous actin (F-actin), 25e27, 259e260, 485 Filamin A, 299, 334e335 Filamin A-interacting protein (FILIP), 299 Filopodia, 264e265 Filopodium-like lateral protrusion (FLP), 470 Flp-FRT system, 259 Fimbria-dentate junction (FDJ), 373 Fish-hook (Fish), 155e156 FLN1 gene, 334e335 Flna. See Filamentous actin (F-actin) FLNA gene, 334e335 FLP. See Filopodium-like lateral protrusion (FLP) FLRTs. See Fibronectin and leucine-rich transmembrane proteins (FLRTs) FMRP. See Fragile X mental retardation protein (FMRP) FMUs. See Fundamental morphogenic units (FMUs) FN. See Fibronectin (FN) FNIII. See Fibronectin type III (FNIII) Focal adhesion kinase (FAK), 115e116, 335 Focal cortical dysplasias (FCD), 584e585 Focaladhesion kinase (Fak), 373 Follistatin, 183e184 Forkhead box protein G1 (FOXG1), 350e351, 543 Formin, 60, 260 Förster resonance energy transferebased conformation sensor, 128 FOXG1. See Forkhead box protein G1 (FOXG1) Foxo polarization pathway, 423 FoxO6 deficiency, 486 Foxp1, 486 Foxp2, 486

596 Index

Fragile X mental retardation protein (FMRP), 262, 488 Free tubulin-binding proteins, 33e34 Frizzled receptors (Fz), 390 Fz-type receptors, 190e191 Wnt binding to, 188e190 Frontotemporal dementia with parkinsonism17 (FTDP-17), 220 FT interactions. See Fiberetarget interactions (FT interactions) FTDP-17. See Frontotemporal dementia with parkinsonism-17 (FTDP-17) Fuel suppliers, 219 Full-length Gli2, 176 Functional neuronal connections, 109 Fundamental morphogenic units (FMUs), 557, 565 Fusogens, 237

G

G protein a Z (Gnaz), 182 G-actin. See Globular actin (G-actin) GA. See Golgi apparatus (GA) GABA. See Gamma-amino-butyric acid (GABA) GABAA receptors (GABAARs), 331 GABAC receptor. See GABAA-rho receptors GABAergic interneurons, 406e407, 479 transmission, 264 GAG. See Glycosaminoglycan (GAG) Gain of function (GoF), 126e127, 159e160 GAL4eupstream activation sequence (GAL4eUAS), 232, 259 GAL80 suppression, 232 Gamma-amino-butyric acid (GABA), 331, 348e349, 369 GAN. See Giant axonal neuropathy (GAN) Ganglionic eminence (GE), 160e161, 345e346 GAP. See GTPase-activating protein (GAP) Gap junctions, 333 GAP-43+ cells, 329 Gas1, 182 Gastrulation brain homeobox 2 transcription factor (Gbx2 transcription factor), 156 Gbx2 transcription factor. See Gastrulation brain homeobox 2 transcription factor (Gbx2 transcription factor) GCL. See Granule cell layer (GCL) GCs. See Granule cells (GCs) GDFs. See Growth and differentiation factors (GDFs) GDNF. See Glial cell lineederived neurotrophic factor (GDNF) GDP. See Guanosine diphosphate (GDP) GE. See Ganglionic eminence (GE) GEFs. See Guanine nucleotide exchange factors (GEFs) Gene knockout technology, 5e6 Gene transcription, 241

Genetic analysis in vivo, 93 Genetic mechanisms controlling shifts in migratory modes, 543e544 Genetically modified organisms, 47 Genome-wide association studies (GWAS), 117 Germinal zones (GZs), 405, 425 GFAP. See Glial fibrillary acidic protein (GFAP) GFP. See Green fluorescent protein (GFP) Giant axonal neuropathy (GAN), 223 Gigaxonin, 223 Gli3R, 176 Glia-guided locomotion, 557 Glial cell lineederived neurotrophic factor (GDNF), 65, 347 Glial cells, 69e70 role in axon maintenance, 224 Glial control of dendritic targeting, 244e245 Glial fibrillary acidic protein (GFAP), 352e353 Glial tube, 468 Glial-aided locomotion. See Glial-guided locomotion Glial-guided locomotion, 561e563 evolutionary origin of, 563 Global thalamocortical topography, 139 Globular actin (G-actin), 25 Glutamatergic tangential contributions as developmental scaffolds, 567e570 Glutamination, 62 Glycine receptor a2 subunit (Glya2R), 348e349 Glycogen synthase kinase 3 (GSK3), 10e11, 34, 386 GSK3b, 184 Glycoprotein, 68 130/STAT3 pathway, 209 Glycosaminoglycan (GAG), 204 Glycosylphosphatidylinositol (GPI), 111, 124 Glypican1 (GPC1), 180 Gnaz. See G protein a Z (Gnaz) GNPs. See Granule neuron progenitors (GNPs) GoF. See Gain of function (GoF) Goldfish, 201e202, 208 Golgi apparatus (GA), 35, 261 Golgi cells, 408e409 Golgi outposts, 261e262 in dendritic elaboration, 236e237 Golgi staining, 289e290 GPC1. See Glypican1 (GPC1) GPI. See Glycosylphosphatidylinositol (GPI) GPI-anchored ephrin-As, 126e127 GR. See Glucocorticoid receptor (GR) Granule cell layer (GCL), 366, 403e404 Granule cells (GCs) migration in dentate gyrus, 372e375 developmental scheme of, 372e373 emergence and migration of long-lived NSCs and establishment of SGZ, 374e375 formation of germinative matrices, 372f

migration of granule neurons to form granule cell layer, 373e374 Granule neuron progenitors (GNPs), 405e406, 411 polarity regulation in, 422f Green fluorescent protein (GFP), 292, 368 Green frog (Rana esculenta), 558 Growth and differentiation factors (GDFs), 183 Growth cones, 58, 141 cytoskeleton, 179e180 targeting, 138e139 Growth factors, 205e207 GSK3. See Glycogen synthase kinase 3 (GSK3) GTP. See Guanosine triphosphate (GTP) GTPase-activating protein (GAP), 11, 71, 111, 128e129, 158, 261, 387 GAP-43, 329 GTPases. See Guanosine triphosphatases (GTPases) Guanine diphosphate. See Guanosine diphosphate (GDP) Guanine nucleotide exchange factors (GEFs), 12, 71, 114, 128e129, 158, 180, 387 Guanine triphosphate. See Guanosine triphosphate (GTP) Guanosine diphosphate (GDP), 12, 71, 583 Guanosine triphosphatases (GTPases), 417 Guanosine triphosphate (GTP), 11e12, 71, 583 hydrolysis, 31b Guidance cue, 63e65 molecules, 66e68 receptors, 262 Guidepost cells, 436e437 in context of axonal tracts formation, 436 hippocampal CeR cells in formation of axonal connections, 440e442, 441f hippocampus and entorhinohippocampal projections, 440 intermediate targets, 437f lot cell roles, 439 neuronal migration, 442e447 corpus callosum formation, 447e449 and evolution of brain wiring, 449e450 in evolution of internal capsule, 450 neuronal migration role in LOT formation, 437e440 normal and pathological brain development, 451e452 axonal tract organization and topography, 451 integrating tangential neuronal migration, 452 notion of, 436e437 Guidepost corridor cells, 452 GWAS. See Genome-wide association studies (GWAS) GZs. See Germinal zones (GZs) Gai proteins, 182

Index

H HATs. See Histone acetyl-transferases (HATs) HCN1 channels, 278 HD-PTP. See His domainecontaining protein tyrosine phosphatase (HDPTP) HDACs. See Histone deacetylases (HDACs) HDB. See Horizontal limb of diagonal band (HDB) Hebbian principle, 138e139 Heparan sulfate (HS), 149e150 Heparan sulfate proteoglycans (HSPGs), 67, 92, 149e150 Hepatocyte growth factor (HGF), 347, 469, 519 Hereditary spastic paraplegia (HSP), 221 Hermaphrodite-specific neurons (HSNs), 158 Heterogeneous nuclear ribonucleoprotein (hnRNP), 491e492 Heterogeneous nuclear ribonucleoprotein U (HNRNPU), 307e308 HGF. See Hepatocyte growth factor (HGF) HGPPS. See Horizontal gaze palsy with progressive scoliosis (HGPPS) Hhip expression, 180 High-mobility group (HMG), 249e251 hiPCS. See Human-induced pluripotent stem cells (hiPCS) Hippocampal interneurons, migration of, 370e372 cellular and distributional diversity, 370 origins and migration of, 370e372, 371f, 374f Hippocampal pyramidal cells, 369 Hippocampal pyramidal neurons, 277 migration of, 369e370 Hippocampus developmental specification, 366e367 canonical Wnt signaling in hippocampal development, 367 cortical hem, 366e367 scheme of hippocampus, 366 migration in, 367 of CajaleRetzius cells in, 367e369 of hippocampal interneurons, 370e372 of hippocampal pyramidal neurons, 369e370 of neural progenitors and granule cells in dentate gyrus, 372e375 structure and lamination, 365e366, 366f terminology important for studying, 365e366 His domainecontaining protein tyrosine phosphatase (HD-PTP), 128 Histone deacetylases (HDACs), 241 HMG. See High-mobility group (HMG) hnRNP. See Heterogeneous nuclear ribonucleoprotein (hnRNP) HNRNPU. See Heterogeneous nuclear ribonucleoprotein U (HNRNPU) Homotypic interactions, 269 Hopping gait, 134

Horizontal gaze palsy with progressive scoliosis (HGPPS), 99e100, 164, 387 Hox gene expression, 405 hPSCs. See Human pluripotent stem cells (hPSCs) HS. See Heparan sulfate (HS) HSNs. See Hermaphrodite-specific neurons (HSNs) HSP. See Hereditary spastic paraplegia (HSP) HSPGs. See Heparan sulfate proteoglycans (HSPGs) 5-HT. See Serotonin (5-HT) 5HT3aR+. See Ionotropic serotonin receptor (5HT3aR+) htt. See Huntingtin gene (htt) Human cerebral cortex, 323e324 Human-induced pluripotent stem cells (hiPCS), 278 Huntingtin, 223 Hyephrin-B1 expression, 126

I Ig. See Immunoglobulin (Ig) Ig-SF. See Immunoglobulin superfamily (Ig-SF) IGFR1. See Insulin-like growth factor 1 receptor (IGFR1) IGL. See Internal granular layer (IGL) IHCs. See Inner hair cells (IHCs) Ihh. See Indian hedgehog (Ihh) IKNM. See Interkinetic nuclear movement (INM) Ilk. See Integrin-linked kinase (Ilk) ILS. See Isolated lissencephaly sequence (ILS) IML. See Internal molecular layer (IML) Immature neurons, 473 Immune-mediated neurorepair mechanisms, 208e209 Immunoglobulin (Ig), 89 Ig-CAMs, 69 Immunoglobulin superfamily (Ig-SF), 66e67, 269e271 IMZ. See Intermediate zone (IMZ) In vivo neuronal polarization of pyramidal neurons, 23e24 Indian hedgehog (Ihh), 176 Inferior nucleus (IX), 389 Inferior olivary nucleus (ION), 409 neurons, 409 Inferior olive (IO), 381 Infrapyramidal bundle (IPB), 226 Injured brain, neuronal migration regulation in, 470e473 Injury-induced axon degeneration, 252 INL. See Inner retinal layer (INL) INM. See Interkinetic nuclear movement (INM) Inner hair cells (IHCs), 135 Inner plexiform layer (IPL), 268e269, 268f Inner retinal layer (INL), 268f INs. See Interneurons (INs)

597

Insects, 132 “Inside-out” arrangement of projection neurons, 290 Insulin-like growth factor 1 receptor (IGFR1), 43e44, 222e223 Integrated, multistep model for synaptic wiring, 273 Integrating tangential neuronal migration, 452 Integrin-linked kinase (Ilk), 373 Integrins, 90, 151, 332e333 Intercellular signaling, 314 Interkinetic nuclear movement (INM), 312, 313f, 315e316 nucleokinesis during, 312 role during neurodevelopment, 315 Intermediate progenitors (IPs), 289e290, 349, 484 cells, 325f, 326 Intermediate zone (IMZ), 289e290, 292, 294, 347e348, 480, 534 Internal capsule cells, 445 Internal granular layer (IGL), 403e404 Internal molecular layer (IML), 440 Interneurons (INs), 328, 347e348, 532. See also Tangential migration; of interneurons in INL, 268f integration into cortical columns, 538e543 PN attract migrating interneurons into cortical plate, 542 radial glial cells trigger shift in migration mode, 543 regulating timing of shift from tangential to radial migration, 541e542 migration, 352 dynamic remodeling of cytoskeleton during, 352 primary cilium, 351 sister interneurons migrates into same minicolumn, 540 preferentially intersynapse, 540e541 subtypes areally distribute via tangential migration, 538e540 Intracellular cascades role, 262e263 Intracellular pathways underlying neuronal polarization, 8e15 Intracellular signaling, 113e115, 369 Intracellular signaling mechanisms calcium, 72 cyclic nucleotides as second messengers and modulators, 72e73 Rho family small GTPases, 71e72 Intrinsic mechanisms of dorsal root ganglion neuron axon regeneration, 203e204, 208 Invertebrate, 57e58 dendrite development cell biology of dendritic growth, 235e237 dendritic field formation control, 242e245 dendritic field formation II control, 245e248

598 Index

Invertebrate (Continued ) dendritic remodeling, 249e252 methods for studying dendrite morphology in Drosophila, 232e233 posttranscriptional control of dendritic development, 241e242 structure and anatomy of, 232 transcriptional control of dendritic morphology, 237e241 key model systems in dendritic morphogenesis C. elegans PVD neurons, 234e235 Drosophila dendritic arborization sensory neurons, 233 Drosophila motoneurons, 233 Drosophila olfactory projection neurons, 233 systems, 97e98 attraction and repulsion by netrins in Drosophila, 98 attraction and repulsion by UNC-6 in C. elegans, 97e98 IO. See Inferior olive (IO) ION. See Inferior olivary nucleus (ION) Ionotropic serotonin receptor (5HT3aR+), 538e539 IPB. See Infrapyramidal bundle (IPB) IPL. See Inner plexiform layer (IPL) IPs. See Intermediate progenitors (IPs) Ipsilateral (temporal) axon, 139 Ipsilateral retinal ganglion cell axons, 181e182, 181f Ipsilateral RGCs, 182 IsO. See Isthmic organizer region (IsO) Isolated lissencephaly sequence (ILS), 300 Isthmic organizer region (IsO), 405 IX. See Inferior nucleus (IX) IZ. See Intermediate zone (IMZ)

J J-RGC, 258f, 266e267 Janus kinase/signal transducers and activators of transcription pathway (JAK/STAT pathway), 207 JIP1, 220 JIP3, 63e64 JM. See Juxtamembrane (JM) JNK. See c-Jun N-terminal kinase (JNK) Jnk1e/emutant mice, 220 Joubert syndrome, 544 Juxtamembrane (JM), 156e157

K Kainate, 520e521 KAL-1 gene, 68 Kallmann’s syndrome (KS), 116e117 KASH domain protein. See Klarsicht, Anc-1, Syne-1 homology domain protein (KASH domain protein) Katanin, 35, 62 KCC2. See Potassium chloride cotransporter 2 (KCC2) KIF2A, 63e64

KIF5, 45 Kinesins, 219e221 Kif1a, 314 superfamily proteins, 63 Klarsicht, Anc-1, Syne-1 homology domain protein (KASH domain protein), 310 KLF4. See Kruppel like factor 4 (KLF4) Knockout mice (KO mice), 467 Kruppel like factor 4 (KLF4), 484 KS. See Kallmann’s syndrome (KS) Kuzbanian (Kuz), 156e157

L L-AP4. See 2-Amino-4-phosphonobutyric acid (L-AP4) L-APB. See L-2-Amino-4-phosphonobutyric acid (L-APB) Labile domain, 60e61 LacZ gene, 534 LAF4/AFF3. See Lymphoid nuclear protein related to AF4/AFF member 3 (LAF4/AFF3) Lamellipodia, 23 Lamellipodin (Lpd), 488, 543e544 Lamin A/C gene, 309 Lamin knockout mouse models, 309 Laminar brains, radial migration on, 557e559 Laminar targeting of retinal dendrites, 269e271 Laminin, 34 Laminopathies, 309 LAR. See Leukocyte common antigenrelated protein (LAR) Large-scale axon pruning, 225f Late-born neurons, 294 Late-onset brain neurodegenerative diseases, 116e117 Later-born neurons, 290 Lateral ganglionic eminence (LGE), 345e346, 370e371, 442e443, 570 interneuron, 493 Lateral geniculate nucleus (LGN), 70 Lateral LMC axons, 133f Lateral motor column (LMC), 132, 133f Lateral olfactory tract (LOT), 163, 437e438, 449e450, 570e571 formation role in, 437e440 anatomy and development, 437e438, 438f diffusible guidance cues in pathfinding, 438e439 Lateral positioning, 159e160 Lateral reticular nuclei (LRN), 381, 387 LCH. See Lissencephaly with cerebellar hypoplasia (LCH) LECT2. See Leukocyte cellederived chemoeaxin-2 (LECT2) LEF. See Lymphoid enhancerebinding factor (LEF) Leucine-rich repeat domains (LRR domains), 147e148, 244 Leukocyte cellederived chemoeaxin-2 (LECT2), 244

Leukocyte common antigen-related protein (LAR), 204 Leukocyte immunoglobulin-like receptor family (LILRB3/PirB), 204 LGN. See Lateral geniculate nucleus (LGN) Lhx1 transcription factor. See LIM homeodomain transcription factor (Lhx1 transcription factor) Ligandereceptor signaling, 156e157 Ligands, 127 Light-sheet fluorescence microscopy. See Selective plane illumination microscopy LILRB3/PirB. See Leukocyte immunoglobulin-like receptor family (LILRB3/PirB) LIM homeobox 2. See LIM homeodomain 2 transcription factor (Lhx2 transcription factor) LIM homeobox 6 (LHX6), 345e346 LIM homeodomain 2 transcription factor (Lhx2 transcription factor), 156, 266 LIM homeodomain transcription factor (Lhx1 transcription factor), 500 LIM kinase (LIMK), 8e9 Limb bud innervations, 132e134 LIMK. See LIM kinase (LIMK) Lin-17 receptor, 6 Lin-44 receptor, 6 LINC complex function, 311 in nuclear positioning, 311 in nucleokinesis, 312 structure, 310 lincRNAs. See lncRNAs in intergenic regions (lincRNAs) Lissencephaly (LIS), 578e579 LIS1. See Platelet-activating factor acetylhydrolase 1b. , regulatory subunit 1 (Pafah1b1) Lissencephaly with cerebellar hypoplasia (LCH), 582 Lizard (Pogona vitticeps), 566 LKB1, 10e11 LMC. See Lateral motor column (LMC) LMC neurons. See Ephrins lncRNA. See Long noncoding RNA (lncRNA) lncRNAs in intergenic regions (lincRNAs), 493 Local circuit interneurons, 324 Local endocytosis, 252 Local recognition mechanisms to control synapse selectivity, 273 Local translation, 44e45 Locally projecting neurons. See Gammaaminobutyric acid-containing interneurons (GABAergic interneurons) Locomotion, 289e290, 292 mode of migration, 328e329 LoF. See Loss of function (LoF) Long noncoding RNA (lncRNA), 488, 492e493

Index

Long-lived NSCs, 374e375 Loss of function (LoF), 126e127 analysis, 159 in vivo phenotypes in motor axons, 131f LOT. See Lateral olfactory tract (LOT) Lot cells, 439 fate of, 440 tangential migration, 439e440 Low-density lipoprotein receptor-related protein 8 (Lrp8), 301 Loxodonta Africana. See African elephant (Loxodonta Africana) Lpd. See Lamellipodin (Lpd) LRN. See Lateral reticular nuclei (LRN) LRP6, 188 Lrp8. See Low-density lipoprotein receptorrelated protein 8 (Lrp8) LRR domains. See Leucine-rich repeat domains (LRR domains) Lsd1, 494 Lymphoid enhancerebinding factor (LEF), 184 Lymphoid nuclear protein related to AF4/ AFF member 3 (LAF4/AFF3), 484e485

M Macaque monkeys, 473 Mag, 204 MAG. See Myelin-associated glycoprotein (MAG) Magic Roundabout, 149 Magnetic resonance (MR), 387 MAGUKs. See Membrane-associated guanylate kinases (MAGUKs) Malformations of cortical development (MCD), 577 Malignant T cell amplified sequence 1 (Mcts1), 488e491 MAM domainecontaining glycosylphosphatidylinositol anchor 2 (Mdga2), 484e485 Mammalian cerebral cortex, 326 Mammalian DCC, 89 Mammalian hindbrain, 95e96 Netrin-1 from ventricular zone, 96 neuronal cell migration control by Netrin-1 in hindbrain, 96 Mammalian neocortex, 560 Mammalian slits, 149 Mammalian spinal cord, 92e95 guidance by midline-derived Netrin-1, 92e93 guidance by ventricular zoneederived Netrin-1, 93 interpreting guidance defects caused by loss of Netrin-1, 93e95 synergy between Netrin-1 from floor plate and from ventricular zone, 93 Mammalian target of rapamycin (mTOR), 13, 64, 207 Manduca sexta, 249 MAP1b, 223 MAP1B light chain (MAP1B-LC), 223

MAP1B-LC. See MAP1B light chain (MAP1B-LC) MAP8, 223 MAPK kinase kinases (MAPKKKs), 228, 299 MAPK kinases (MAPKKs), 299 MAPK/Nmnat2/Sarm1-dependent NAD+depletion model, 228 MAPKKKs. See MAPK kinase kinases (MAPKKKs) MAPKKs. See MAPK kinases (MAPKKs) MAPKs. See Mitogen-activated protein kinases (MAPKs) Mapping, 136e140 corticocollicular projections, 140 olfactory wiring, 136 retinogeniculate projections, 139 retinotectal/retinocollicular projection, 136e139 thalamocortical projections, 139 MAPs. See Microtubule-associated proteins (MAPs) MARCM technique. See Mosaic analysis with a repressible cell marker technique (MARCM technique) Marginal zone (MZ), 289e290, 348e349, 371, 539e540 MARK1e4 kinase, 10e11 MARK2 kinase, 10e11 Math1. See Atoh1 MAZ. See Multipolar cell accumulation zone (MAZ) MB. See Medulloblastoma (MB) MCD. See Malformations of cortical development (MCD) Mcts1. See Malignant T cell amplified sequence 1 (Mcts1) md-da neurons. See Multidendritic dendritee arborization neurons (md-da neurons) Mdga2. See MAM domainecontaining glycosylphosphatidylinositol anchor 2 (Mdga2) mDNs. See Midbrain dopaminergic neurons (mDNs) MDS. See MillereDieker syndrome (MDS) Mechanistic target of rapamycin. See Mammalian target of rapamycin (mTOR) MECP2 gene. See Methyl-CpG-binding protein 2 gene (MECP2 gene) Medial ganglionic eminence (MGE), 325f, 345e346, 370e371, 442e443, 495e498, 538e539, 566 Medial LMC axons, 133f Medial longitudinal fasciculus (MLF), 389e390 Medial migratory stream (MMS), 348, 474 Medial nucleus of trapezoid body (MNTB), 135 Medulloblastoma (MB), 404, 424e425 MEF2, 263 MEKK4 knockout mice, 299 Membrane

599

distribution, 126e127 proteins, 222e223 trafficking, 63e64 transport and insertion, 222e223 Membrane-associated periodic actin skeleton, 225 Membrane-bound cell adhesion molecules, 5 Membrane-type 1 matrix metalloproteinase (MT1-MMP), 312 Mesenchymal epithelial transitions (METs), 423 Metalloprotease cleavage, Sliterobo signaling regulation by, 156e157 Metamorphosis, 226e227, 249 Methyl-CpG-binding protein 2 gene (MECP2 gene), 263 N-Methyl-D-aspartate (NMDA), 352e353 NMDA-dependent transmission, 264e265 METs. See Mesenchymal epithelial transitions (METs) MFs. See Mossy fibers (MFs) MGE. See Medial ganglionic eminence (MGE) mGluRs. See Metabotropic glutamate receptors (mGluRs) MIA. See Migration-inducing activity (MIA) MICAL. See Molecule Interacting with CasL (MICAL) Mical function downstream of ecdysone receptor, 249e251 Microcephaly, 580 Microglia, 479 MicroRNAs (miRNAs), 65, 241, 488, 493e495, 501 in dendritic development, 242 miR-124, 65 miR-126e5p, 117 miR-132, 65 miR-16, 65 miR-9, 65, 494e495 translation control in dendritic development, 241e242 Microtubule plus endetracking proteins (+Tips), 33 Microtubule-associated proteins (MAPs), 10, 24, 61e62, 159, 220, 259e261, 582 MAP2, 350e351 Microtubule-organizing centers (MTOCs), 30e31 Microtubules (MTs), 30e32, 42, 58, 60e62, 188, 218, 220, 259e260 affinity regulating kinase-2, 10e11 binding motors, 312e314 dynein, 313 Kinesin Kif1a, 314 cytoskeleton and associated motors, 414e416, 415f disassembly, 251e252 dynamics, 31b during axon formation, 32e35 interaction between F-actin and, 62e63 microtubule-actin cross talk, 419e420 microtubule-based transport, 43 microtubule-binding proteins, 31

600 Index

Microtubules (MTs) (Continued ) microtubule-severing proteins, 35, 62 network regulators, 259e260 polarity difference between dendrites and axons, 235 polymerization, 260 regulatory scheme, 10e11 Midbrain dopaminergic axons, Shh chemoattractant for, 182 Midbrain dopaminergic neurons (mDNs), 182 Midline axon guidance, 156e157 crossing, 192 in Drosophila and mice, 151f midline-derived Netrin-1 in spinal cord, 92e93 SliteRobo function in midline crossing, 150e155 SliteRobo signaling for exiting midline, 155 Migration migrating neurons trajectory in developing brain, 326e328 modes, 328e329 termination in OB, 470 Migration-inducing activity (MIA), 352e353 Migratory behaviors during radial migration, 290e297 bipolar migrating neurons along radial glial fibers, 290 departure from ventricular zone, 294e296 multipolar migration, 292e293 progenitor cell behavior in subventricular zone, 296e297 radial glial fibereindependent mode of migration, 290e292 transformation from multipolar migrating neurons to bipolar locomoting neurons, 293e294 Migratory scaffolds in injured brain, 470e472 astrocytes, 471 blood vessels, 472 neighboring cells in neuronal chain, 470e471 radial glial cells, 472 in normal brain astrocytes, 468e469 blood vessels, 469 neighboring cells in neuronal chain, 466e468 restrict tangential movement of PN, 535e537 MillereDieker syndrome (MDS), 300 Minicolumns constitute columns, 532e533 ontogenetic radial units, 533f Minor ventricular zone derivatives migration, 408e409 miR-9, 65, 494e495 miR-16, 65 miR-124, 65 miR-126e5p, 117 miR-132, 65

miRNAs. See MicroRNAs (miRNAs) Mitochondria transport role in axon maintenance, 221e222 Mitochondrion, 64 Mitofusins, 221 Mitogen-activated protein kinases (MAPKs), 220, 228, 299 Mekk4, 334e335 Mixed microtubule polarity, 260 MLCK. See Myosin light-chain kinase (MLCK) MLF. See Medial longitudinal fasciculus (MLF) MMS. See Medial migratory stream (MMS) MNTB. See Medial nucleus of trapezoid body (MNTB) Modulation, 96e97 Molecular cues controlling integration of interneurons into cortical migratory streams, 348e349 controlling intracortical dispersion of interneurons, 349e350 drawing path of cortical interneuron migration, 347e348 Molecular mechanisms controlling tangential migration of facial motor neurons, 389e394 FBM caudal migration, 391e392 origin and, 389e390 of precerebellar neurons, 384e389 role of chemoattraction and chemorepulsion, 392e393 regulating initiation of migration and cell polarity, 297e301, 297f controlling initiation of radial migration, 299 coupling between neuronal differentiation and migration, 298e299 extracellular molecules affecting migrating cells, 301 multipolar migration regulation, 299e300 Molecular signaling limits tangential movement of PN, 537e538 Molecule Interacting with CasL (MICAL), 114e115 Morphogens, 68, 176 as axon guidance cues, 176 Morphological changes, 289e290, 292, 296fe297f, 299e300 Mosaic analysis with a repressible cell marker technique (MARCM technique), 232 Mosaics, 273e275, 274f Mossy cells, 366 Mossy fibers (MFs), 404 Motor proteinebased transport in axons and dendrites, 40e43 MP. See Multipolar stage (MP) mPFC. See Medial prefrontal cortex (mPFC) MPTP. See 1-Methyl-4-phenyl-1,2,3,6tetrahydropyridine (MPTP) MR. See Magnetic resonance (MR) mRNA. See Messenger RNA (mRNA) mRNA translation machineries, 262

MS. See Multiple sclerosis (MS) MSNs. See Medium spiny neurons (MSNs) MT1-MMP. See Membrane-type 1 matrix metalloproteinase (MT1-MMP) MTOCs. See Microtubule-organizing centers (MTOCs) mTOR. See Mammalian target of rapamycin (mTOR) mTORCs. See Mammalian target of rapamycin complexes (mTORCs) MTs. See Microtubules (MTs) Multidendritic dendriteearborization neurons (md-da neurons), 162 Multiple sclerosis (MS), 521e522 Multipolar cell accumulation zone (MAZ), 292e293, 295 Multipolar cells, 289e290, 292e293, 293f Multipolar IMZ/SVZ/MAZ cells, 293e294 Multipolar migration, 289e290, 292e293 morphology of multipolar migrating cells, 293f regulation, 299e300 Multipolar neurons, 327 Multipolar stage (MP), 23e24 Myelin, 515 Myelin-associated glycoprotein (MAG), 69e70, 201e202, 224 Myelin-forming cells migration. See also Neuronal migration adhesion and chemotactic mechanisms adhesion and surface molecules, 519e521 secreted factors, 521e524 chemokinetic factors, 518e519 genesis of myelin-producing cells during development, 516e517 migratory paths followed by oligodendrocyte progenitor and precursor cells, 518 oligodendrocyte precursor cells, 517e518 regions of oligodendrogliogenesis and routes of migration, 517f Myelinating glial cells, 224 Myelination, 69e70 Myocyte nuclear envelope (Myne), 310 Myosin II-mediated actin contractility, 29e30 Myosin light-chain kinase (MLCK), 351 MZ. See Marginal zone (MZ)

N N-cadherin, 27, 68e69, 466, 487e488, 519e520 N-cofilin, 331, 485 Na-K-Cl cotransporter 1 (NKCC1), 352e353 Nanos protein (Nos), 241e242 Nap1. See Nck-associated protein (Nap1) nBAFs. See Neuronal-specific Brahmarelated gene/Brahma-associated factor complexes (nBAFs) NCAM. See Neural cell adhesion molecule (NCAM) Nck-associated protein (Nap1), 487 NDD-associated dendrite pathology, 278 Nde1, 300

Index

Ndel1, 300, 334 Neighboring cells in injured brain, 470e471 in neuronal chain, 466e468 in normal brain, 466e470 NELL2, 154 Neocortex organization, 326 Neocortical interneuron migration, 498 Neocortical neural circuits, 479 Neogenin (Neo), 89, 444 Neonatal human brain, 474 Nerve growth factor (NGF), 65e66, 226 Nervous systems, 87 Netrin signaling involvement in nervous system disorders, 99e100 sliterobo signaling roles in, 162e163 Nesprin. See Nuclear envelope spectrin repeat (Nesprin) Nestin-cre, 182 Netrin-G1, 89 Netrin-G2, 89 Netrin-G2/NGL-2 interactions, 277 Netrin(s), 66e68, 87, 100e101, 156e157 axon and cell guidance, 99 discovery and structure, 88e89, 88f function in axon guidance and cell migration, 92e98 guidance of other classes of mammalian axons and cells, 96e97 invertebrate systems, 97e98 mammalian hindbrain, 95e96 mammalian spinal cord, 92e95 interactions with other signaling systems, 89e90 effects of floor plate and Netrin-1, 91fe92f Netrin functional domains and interactions with receptors, 92 in nervous system, 99 Netrin-2, 88 Netrin-4, 71, 89 Netrin1, 88, 96, 152, 178, 301, 350e353, 385e386 axon initiation in C. elegans, 6 receptors, 89 signaling during midline dendritic guidance, 242e244 signaling involvement in nervous system disorders, 99e100 Networked axon growth, 222 Neural activity, 70e71, 263 Neural cell adhesion molecule (NCAM), 69, 388, 542 Neural progenitor cells (NPCs), 205, 531e532 migration in dentate gyrus, 372e375 Neural stem cells (NSCs), 205, 374, 465 Neural tissue damage, 202e203 Neuregulins (NRGs), 542 NRG1, 347e348 NRG3, 349, 542 Neurexins (NRXs), 264e265 Neuroblasts, 160 Neurocognitive disorders, 531e532

NeuroD, 263, 298e299 Neurodegenerative diseases, 46 Neurodevelopmental disorders, 278, 480 Neurofilaments, 218 Neurogenesis, 531e532 Neurogenins (Ngns), 336 Ngn genes, 557 Ngn1/2, 336 Ngn2, 298e299, 345e346, 482, 491 signaling, 482e483 Neuroglycan C/chondroitin sulfate proteoglycan 5 (NGC/CSPG5), 485e486 Neuroinflammation, 208e209 Neuroligin (NL) NL1, 264e265 Neurological disorders, 116 semaphorins, plexins, and neuropilins in, 116e117 ALS, 117 ASD, 116 KS, 116e117 late-onset neurodegenerative diseases, 117 Neuromuscular junction (NMJ), 117 Neuronal migration, 161, 289e290, 442e447, 468, 479e480, 532e534. See also Myelin-forming cells migration corridor cells migration, 443f, 446e447, 451f cortical subplate axons in pathfinding of thalamocortical projections, 443e444 diencephalic and subpallial pathfinding of thalamocortical projections, 445e446 enhancement as strategy for endogenous neuronal regeneration, 473 fate of guidepost cells for thalamocortical projections, 447 methods for migration, 480 migratory behaviors during radial migration, 290e297 bipolar migrating neurons along radial glial fibers, 290 departure from ventricular zone, 294e296 multipolar migration, 292e293 progenitor cell behavior in subventricular zone, 296e297 radial glial fibereindependent mode of migration, 290e292 transformation from multipolar migrating neurons to bipolar locomoting neurons, 293e294 molecular mechanisms regulating initiation, 297e301, 297f controlling initiation of radial migration, 299 coupling between neuronal differentiation and migration, 298e299 extracellular molecules affecting migrating cells, 301 multipolar migration regulation, 299e300 in postnatal brain, 466f postnatal neuronal migration in primates, 473e474

601

regulation in injured brain, 470e473 regulation in normal brain, 466e470, 467f subplate neurons, 444 thalamocortical and corticofugal axons, 442e443 transcriptional and posttranscriptional control of, 480e501, 486f radial migration, 480e495 tangential migration, 495e501 types in vertebrate brain development, 555e557 Neuronal migration disorders (NMDs), 577 mutations and mechanisms, 580e585 cobblestone malformations and mutations in dystroglycan genes, 584 FCD and DMEG, 584e585 mutations in microtubule-associated proteins, 580e581 tubulin mutations, 581e582 variant lissencephalies and mutations, 583e584 types of malformations, 578e580 cobblestone LIS, 579 LIS, 578e579 microcephaly, 580 MTOR, 580 normal and lissencephalic brain sections, 578f pachygyria, 578 PH, 579 PMG, 579e580, 580f SBH, 579, 579f Neuronal polarity in vivo axon initiation in vitro, 4e5 axon initiation in vivo, 5 distinction between cues regulating axon specification vs. axon growth, 5e6 extracellular cues regulating neuronal polarization and axon initiation, 6e8 intracellular pathways underlying neuronal polarization, 8e15 Neuronal polarity theory, 21e23 Neuronal polarization, 5, 21e23 cytoskeleton role in establishment, 24e39 extracellular cues regulating, 6e8 future work on neuronal morphogenesis, 47e48 intracellular pathways underlying, 8e15 cytoskeletal dynamics role in axon initiation and growth, 9e10 major signaling pathways in axon initiation and growth, 10e15 role of protein degradation and local translation in axon specification and axon growth, 8e9, 9f maintaining neuronal polarity, 46e47 (membrane) trafficking role during, 39e46 morphology and development stages, 26fe27f Neuronal-specific Brahma-related gene/ Brahma-associated factor complexes (nBAFs), 241

602 Index

Neuronal-specific microtubule-binding proteins, 32 Neuronal/neuron, 21, 57, 160, 176, 257e259, 273e277, 465 cell migration control by Netrin-1 in hindbrain, 96 types, 109e110 connections, 115, 217 cytoskeleton, 259 differentiation protocols, 48 features in developing central nervous system, 289e290 morphogenesis, 47e48 neuronal cell types, 258f g neurons, 226e227 translocation of, 329 Neuropeptide somatostatin (SST+), 538e539 Neuropeptide Y (NPY), 345e346, 370 Neuropilin (Nrp), 111 function in neural circuit development, 115e116 mechanisms of intracellular signaling, 113e115 in neurological disorders, 116e117 Nrp-1, 67, 111e112, 347e348 receptor, 331 Nrp-2, 111e112, 192, 226, 439e440 structural features, 111e113 Neuropsychiatric disorders, 116 Neurorepair, anatomical substrate of, 206e207 Neurospora crassa, 309 Neurotransmission, arbor size control by, 263e264 Neurotransmitters, 470, 541e542 ATP, 332 GABA, 331 glutamate, 332 Neurotrophic factors, 65e66 Neurotrophins (NTs), 10, 65e66 NT-3, 65, 263 NT-4, 347, 501 NGC/CSPG5. See Neuroglycan C/ chondroitin sulfate proteoglycan 5 (NGC/CSPG5) NGF. See Nerve growth factor (NGF) Ngns. See Neurogenins (Ngns) NgR. See Nogo receptor (NgR) NIPBL. See Nipped-B-like (NIPBL) Nipped-B-like (NIPBL), 483 Nissl bodies, 218f Nitric oxide synthase (NOS), 226e227 NK homeobox 2. 1 (NKX2.1), 345e346, 371 NKCC1. See Na-K-Cl cotransporter 1 (NKCC1) NKX2.1. See NK homeobox 2. 1 (NKX2.1) NL. See Neuroligin (NL); Nucleus laminaris (NL) NMDA. See N-Methyl-D-aspartate (NMDA) NMDAR. See N-Methyl-D-aspartate receptors (NMDAR)

NMDs. See Neuronal migration disorders (NMDs) NMJ. See Neuromuscular junction (NMJ) Nogo receptor (NgR), 69e70, 204 “Non-self” neurons, 275, 276f Noncanonical signaling pathway, 179e180 Nonclustered Pcdhs, 69 Normal brain development, 451e452 neuronal migration regulation in, 466e470, 467f Nos. See Nanos protein (Nos) NOS. See Nitric oxide synthase (NOS) NPCs. See Neural progenitor cells (NPCs); Nuclear pore complexes (NPCs) NPY. See Neuropeptide Y (NPY) NrCAM. See Neural cell adhesion molecule (NCAM) Nrf2f2. See Chicken ovalbumin upstream promoter transcription factor II (COUP-TFII) NRGs. See Neuregulins (NRGs) Nrp. See Neuropilin (Nrp) Nrxns. See Neurexins (NRXs) NRXs. See Neurexins (NRXs) NSCs. See Neural stem cells (NSCs) NTs. See Neurotrophins (NTs) NTZ. See Nuclear transitory zone (NTZ) NUANCE, 310 Nuclear envelope spectrin repeat (Nesprin), 310 Nuclear migration, collective mechanisms for intercellular signaling, 314 mechanical interactions, 315 Nuclear pore complexes (NPCs), 305e306 Nuclear transitory zone (NTZ), 407e408 Nuclear translocation. See Somal translocation Nucleic acids, 488 Nucleokinesis, 305 chromatin, 306e307 collective mechanisms for nuclear migration, 314e315 cytoskeleton dynamics as nuclear drivers, 314 diseases affecting nuclear envelope, 309e310 cohesinopathies, 308e309 higher order structure of nucleus, 307e308 during INM, 312 INM role during neurodevelopment, 315 interactions between nucleus and cytoskeleton LINC complex, function, 311 LINC complex structure, 310 LINC complex in, 312 link between nucleus and centrosome, 311 membraneless organelles in nucleus, 307 microtubule binding motors, 312e314 nucleus, 305e306 Nucleus, 305e306 and centrosome, 311 higher order structure of, 307e308 membraneless organelles in, 307

nuclear membrane and nuclear pores, 305e306 presentation of physical interaction between nucleus and cytoskeleton, 306f Nucleus laminaris (NL), 263e264

O OAP-1. See OSP/claudin-11 OB. See Olfactory bulb (OB) Odorant receptor genes (OR genes), 136 OE. See Olfactory epithelium (OE) OHCs. See Outer hair cells (OHCs) Olfactory bulb (OB), 160, 163f, 345e346, 352e353, 465 migration termination in, 470 Olfactory epithelium (OE), 162, 163f Olfactory PNs, dendritic targeting of, 239e241 Olfactory receptor neurons (ORNs), 233 Olfactory sensory neurons (OSNs), 136, 162 Olfactory wiring, 136 Oligodendrocyte precursor cells (OPCs), 205, 516e518 migratory paths followed by, 518 motility, 518e519 Oligodendrocyte-myelin glycoprotein (OMgp), 69e70, 201e202, 204 Oligodendrocytes, 479, 515 Oligodendrogenesis, 516e517 OMgp. See Oligodendrocyte-myelin glycoprotein (OMgp) OML. See Outer molecular layer (OML) ON-alpha (ONa), 273 ONL. See Outer nuclear layer (ONL) ONeOFF direction-selective retinal ganglion cells (ooDSGCs), 258f, 268f, 269, 270fe271f, 271e273 ONa. See ON-alpha (ONa) ONa-RGCs, 275 ooDSGCs. See ONeOFF direction-selective retinal ganglion cells (ooDSGCs) OPCs. See Oligodendrocyte precursor cells (OPCs) OPL. See Outer plexiform layer (OPL) Optic chiasm, 135 Optic nerve injury, 203, 207e208 OR genes. See Odorant receptor genes (OR genes) Orbit/MAST, 159 oRG cells. See Outer subventricular radial glia cells (oRG cells) Organotypic cultures, 439e440 ORNs. See Olfactory receptor neurons (ORNs) Orphan nuclear receptor, 500 OSNs. See Olfactory sensory neurons (OSNs) OSP/claudin-11, 520e521 OSVZ. See Outer subventricular zone (OSVZ) Otx1, 226 Outer hair cells (OHCs), 135 Outer molecular layer (OML), 440 Outer nuclear layer (ONL), 268f

Index

Outer plexiform layer (OPL), 268f Outer plexiform layer, 268e269 Outer subventricular radial glia cells (oRG cells), 330, 481 Outer subventricular zone (OSVZ), 296e297, 480e481

P p21-activated kinase (Pak), 158 PAK3, 350e351 p27Kip1, 298, 335 Pachygyria, 578 Pafah1b1. See Platelet-activating factor acetylhydrolase 1b, regulatory subunit 1 (Pafah1b1) Paired box 6 (PAX6), 345e346 Paired immunoglobulin-like receptor B (PirB), 69e70 Pak. See p21-activated kinase (Pak) Pallial internal circuitry, radial migration influence on, 560e563 evolutionary origin of glial-aided locomotion, 563 glial-guided locomotion, 562e563 somal translocation, 561e562 Pallial interneurons, 565e567 conserved features of tangential migration of, 565e566 divergence in tangential migratory routes of, 566 diversifying complexity of GABAergic subtypes, 566e567 Palliale subpallial boundary (PSB), 347e348 Pallium, 479e480 Palliumesubpallium boundary (PSPB), 367, 442e443 Pancreatic transcription factor 1a (Ptf1a), 406e407 Par genes, 10 Par1b. See Microtubule affinity regulating kinase-2 Par6 genes, 11 Parkinson’s disease (PD), 116 Parvalbumin (PV), 345e346, 370e371 Parvalbumin (PV+), 538e539 Passive pushing out model, 295e296 Pathfinding in brain stem, 135 central, 135 peripheral pathfinding, 132e134 in spinal cord, 134 Pathological axon degeneration, 227 molecular mechanisms, 227e229 Pathological brain development, 451e452 Patterning membrane excitability of dendritic compartments, 278 Pax2-positive interneurons, 408e409 Pax2+ interneuron progenitor (PIP), 406e407 Pax6 protein, 329, 335, 482 PAX6. See Paired box 6 (PAX6) PC2. See Pheromone convertase 2 (PC2) Pcdhs. See Protocadherins (Pcdhs) PCN. See Precerebellar nuclei (PCN)

PCP. See Planar cell polarity (PCP); Purkinje cell progenitor (PCP) PCs. See Purkinje cells (PCs) PCZ. See Primitive cortical zone (PCZ) PD. See Parkinson’s disease (PD) PDGF-A. See Platelet-derived growth factor A (PDGF-A) PDGFRa. See Platelet-derived growth factor receptor alpha (PDGFRa) Peripheral nervous system (PNS), 57e58, 203, 237 axon regeneration in injured CNS vs., 203e204 Peripheral pathfinding, 132e134 Perireticular cells, 445 Periventricular fiber reach zone (PVFRZ), 480 Periventricular heterotopia (PH), 299, 579 and mutations, 582e583 Periventricular nodular heterotopias, 485 PH. See Periventricular heterotopia (PH); Pleckstrin homology (PH) PHD finger protein 6 (PHF6), 485e486 Phenotype-based cell culture assay, 493 Pheromone convertase 2 (PC2), 149 PHF6. See PHD finger protein 6 (PHF6) Phosphatase and tensin homolog (PTEN homolog), 64, 207 signaling during axon specification, 13e14 Phosphatidylinositol (3,4,5)-triphosphate (PIP3), 13 Phosphatidylinositol 3-kinase. See Phosphoinositide-3 kinase (PI3K) Phosphatidylinositol-3 kinase class III (PI3Kc3), 13 Phosphatidylinositol-3 monophosphate (PI(3) P), 13 Phospho-GAP-43+ cells, 329 Phosphoinositide 3 (PI3), 347 Phosphoinositide-3 kinase (PI3K), 6, 13, 130, 184 signaling during axon specification, 13e14 Phosphoinositide-30 kinase/AKT (PI3K/ AKT), 262 Phospholipase C (PLC), 184 Phospholipids, 222e223 phosphoPKCz, 188e189 Phosphorylated n-cofilin loses, 331 Phosphorylation reactions, 262 Phr1 gene, 64e65 Phylogeny of ephrins and ephs, 125e126 PI-3K. See Akt-phosphatidylinositol-3-OH kinase (PI-3K) PI(3)P. See Phosphatidylinositol-3 monophosphate (PI(3)P) PI3. See Phosphoinositide 3 (PI3) PI3K. See Phosphoinositide-3 kinase (PI3K) PI3K/AKT. See Phosphoinositide-30 kinase/ AKT (PI3K/AKT) PI3K/AKT pathway, 207 PI3KeAKTeRhebemTOR pathway, 207 PI3Kc3. See Phosphatidylinositol-3 kinase class III (PI3Kc3) PI3KcIa. See Class Ia PI3-kinase (PI3KcIa)

603

Pinlike morphology, 295e296, 296f PIP. See Pax2+ interneuron progenitor (PIP) PIP3. See Phosphatidylinositol (3,4,5)triphosphate (PIP3) PirB. See Paired immunoglobulin-like receptor B (PirB) Piwi RNAs (piRNAs), 492 PIWIL1, 492 PK2. See Prokineticin-2 (PK2) PKA. See Protein kinase A (PKA) PKB. See Protein kinase B (PKB) PKC. See Protein kinase C (PKC) Planar cell polarity (PCP), 184, 390e391, 391f signaling, 186, 188e190, 390e391 Plasmalemma, 222 Plasticity, 139 Platelet-activating factor acetylhydrolase 1b, regulatory subunit 1 (Pafah1b1), 300, 333e334, 485 Platelet-derived growth factor A (PDGF-A), 518e519 Platelet-derived growth factor receptor alpha (PDGFRa), 516e517 PLC. See Phospholipase C (PLC) Pleckstrin homology (PH), 13 Plexin(s), 111, 262 function in neural circuit development, 115e116 mechanisms of intracellular signaling, 113e115 in neurological disorders, 116e117 Plexin1B receptors, 261 PlexinA1, 149 PlexinA3, 67, 226 PlexinA4, 67, 226, 277e278 PlexinC1. See Virus-encoded semaphorin protein receptor (VESPR) PlexinD1, 117 structural features, 111e113 PlexinAs. See Class A plexins (PlexinAs) Plexinesemaphorineintegrin (PSI), 111 PLM. See Posterior lateral microtubule (PLM) PLP. See Proteolipid protein (PLP) Plus endetracking proteins (+TIPs), 61, 259e260 PMG. See Polymicrogyria (PMG) PN. See Pontine nucleus (PN) PNS. See Peripheral nervous system (PNS) PNs. See Projection neurons (PNs); Pyramidal neurons (PNs) POA. See Preoptic area (POA) Pogona vitticeps. See Lizard (Pogona vitticeps) Polarization regulate PI3 kinase activity, 24 Polarization signaling pathways, 34 Polycomb repressor complexes 1 (PRC1), 241 Polycomb repressor complexes 2 (PRC2), 241 Polymer transport model, 35 Polymerization dynamics of microtubules, 31 Polymicrogyria (PMG), 579e580, 580f

604 Index

Polypyrimidine tractebinding protein (Ptbp1), 491e492 Polysialylatedeneural cell adhesion molecule (PSAeNCAM), 352e353, 466 Pontine neurons, 384 Pontine nucleus (PN), 381, 387 coarse and specific control of PN dendritic targeting, 244 NPCs, 533e534 Positional cues shape asymmetric dendritic arbors, 266e267 Postcrossing commissural axons, 193 Posterior lateral microtubule (PLM), 247 Postnatal neuronal migration in primates, 473e474 Postsynaptic density (PSD), 39 Posttranscriptional control of dendritic development, 241e242 miRNAs in dendritic development, 242 mRNA translation control in dendritic development, 241e242 of neuronal migration, 480e501 Posttranscriptional events in radial migration, 488 lncRNAs, 492e493 MicroRNAs, 493e495 RBPs, 488e492 in tangential migration, 501 Posttranscriptional regulators radial migration, 496te497t tangential migration, 502t Posttranscriptional Robo regulation, 152 Posttranslational modification (PTM), 30, 62 Posttranslational processing, 351 Potassium chloride cotransporter 2 (KCC2), 206e207, 349 Powering dendrite growth, 262 PR domain-containing 8 (Prdm8), 484 PRC1. See Polycomb repressor complexes 1 (PRC1) Prdm8. See PR domain-containing 8 (Prdm8) Precerebellar nuclei (PCN), 381, 388, 393, 404 migration, 409e410, 410f neurons, 394 migrate near the pial surface, 388e389 Preoptic area (POA), 345e346, 371, 500, 538e539 Presenilin1, 392 Primate neocortical GABAergic interneurons, 324 postnatal neuronal migration in, 473e474 Primitive cortical zone (PCZ), 291 Profilins, 29 Progenitor cell behavior in subventricular zone, 296e297 Progressivemotor neuron diseases, 117 Projection neurons (PNs), 233, 324, 532, 536f attracts migrating interneurons into cortical plate, 542 integration into cortical minicolumns, 535e538

migratory scaffolds restrict tangential movement, 535e537 molecular signaling limits tangential movement, 537e538 migration into same minicolumn, 533e535 Prokineticin-2 (PK2), 469e470 Proportional Ephrin/Eph signaling, 136e140, 137f Proportional mode, 125 Prosomere 4 (p4), 563e564 Prospective white matter (PWM), 405 Proteasome and autophagy pathways in axonal homeostasis autophagy/lysosome pathway role, 223e224 UPS in axon maintenance, 223 Protein kinase A (PKA), 7, 180e181, 203 Protein kinase B (PKB), 13 Protein kinase C (PKC), 262 PKCa, 181e182 PKCz, 188e189 Protein(s), 33 degradation, 8e9, 9f, 46, 94e95 phosphatase 1espinophilin pathway, 34 stabilization and degradation, 46 translation and stability, 64e65 14e3-3 proteins regulating cell-intrinsic switch, 180e181 Proteoglycans, 68, 204 Proteolipid protein (PLP), 516e517 Proto-IPL formation by retinal amacrine cells, 269 Protocadherins (Pcdhs), 68e69, 246e247 Protrudin, 63 PRRG4 (WAGR syndromeeassociated protein), 154 PSAeNCAM. See Polysialylatedeneural cell adhesion molecule (PSAeNCAM) PSB. See Palliale subpallial boundary (PSB) PSD. See Postsynaptic density (PSD) PSI. See Plexinesemaphorineintegrin (PSI) PSPB. See Palliumesubpallium boundary (PSPB) Ptbp1. See Polypyrimidine tractebinding protein (Ptbp1) Ptch1, 176 PTEN homolog. See Phosphatase and tensin homolog (PTEN homolog) Ptf1a. See Pancreatic transcription factor 1a (Ptf1a) PTM. See Posttranslational modification (PTM) Pumilio protein (Pum), 241e242 Purkinje cell progenitor (PCP), 406e407 Purkinje cells (PCs), 115e116, 403e404 migration, 407e408 PV. See Parvalbumin (PV) PVFRZ. See Periventricular fiber reach zone (PVFRZ) PWM. See Prospective white matter (PWM) Pyramidal dendrite branching, 262 Pyramidal neurons (PNs), 3e4, 265e266, 289e290, 479

R RA. See Retinoic acid (RA) RabGDI, 154 Rac, 24 Rac1 regulatory protein, 13 Rac1-specific GEF Dock1, 261 Rac1-specific GEF Tiam1, 261 Radial glia (RG), 480 Radial glial cells (RGCs), 290, 324, 472, 557 trigger shift in migration mode, 543 Radial glial fibers (RGFs), 290 RGF-independent migration, 290e291 RGF-independent neuronal migration, 291 RGFeindependent mode of migration, 290e292 Radial migration, 480e481, 537. See also Tangential migration in developing cerebral cortex cortical projection neurons, 324e326 migrating neurons trajectory in developing brain, 326e328 migration modes, 328e329 neocortex organization, 326 on elaborated brains, 559e560 evolution, 557 factors regulating radial migration of cortical neurons, 330e336 adhesion molecules, 332e333 cytoskeletal regulators, 333e335 neurotransmitters, 331e332 secreted molecules, 330e331 transcription factors, 335e336 impact on brain evolution, 557e563 influence on pallial internal circuitry, 560e563 on laminar brains, 557e559 locomotion, 480e481 posttranscriptional regulators, 496te497t radial migration in developing human neocortex, 330 regulating timing of shift from tangential migration to, 541e542 subtypes of neocortical radial glia, 481e488 posttranscriptional events in radial migration, 488 transcriptional regulators, 489te490t translocation, 481 Radial unit hypothesis, 326, 328e329, 533e534 RAGs. See Regeneration-associated genes (RAGs) “Rails”, 219 RALDH2. See Retinaldehyde dehydrogenase-2 (RALDH2) Rana esculenta. See Green frog (Rana esculenta) Rap1b, 13 RapGEF2, 300 Rapidly exiting population (REP), 295 Ras-family of small GTPases, 12e13 “Rautenlippe”, 382 RBD. See Rho GTPase binding domain (RBD)

Index

Rbfox1, isoform 1 (Rbfox1iso1), 492 Rbfox1/2/3. See RNA-binding Foxs (Rbfox1/ 2/3) Rbfox1iso1. See Rbfox1, isoform 1 (Rbfox1iso1) RBM4. See RNA-binding motif 4 (RBM4) Rbm4a knockout mice, 491 RBPs. See RNA-binding proteins (RBPs) RE1 silencing transcription factor (REST), 483e484 RE1. See Repressor element 1 (RE1) Receptor tyrosine kinase (RTK), 124, 262 Reeler mice, 394 Reelin (RLN), 330e331, 345e346, 470, 563, 564f signaling, 373 Regeneration-associated genes (RAGs), 203e204 RELN gene, 330 REP. See Rapidly exiting population (REP) Repressor element 1 (RE1), 483e484 Repulsion, 96e97, 274f, 276f, 277e278 contact-dependent, 275 homotypic, 275 by Netrins in Drosophila, 98 by UNC-6 in C. elegans, 97e98 Repulsive axon guidance molecules, 226 Repulsive Ephrin/Eph signaling, 125 Repulsive Netrin receptors, 89 REST. See RE1 silencing transcription factor (REST) Ret receptor, 65 Reticulon family member 4A (RTN4A/ Nogo-A), 201e202 Reticulotegmental nucleus (RTN), 409 Retina, 207 Retinal ganglion cells (RGCs), 5, 58, 135, 176, 181e182, 203e204, 207e209, 226, 258f, 265e267, 269 Retinaldehyde dehydrogenase-2 (RALDH2), 389 Retinofugal axons, 139 Retinofugal system, axon regeneration in, 207, 210t Retinogeniculate projections, 139 Retinoic acid (RA), 389 Retinotectal/retinocollicular projection computational modeling, 138e139 mapping mechanisms along anterioreposterior axis, 136e138 along dorsoventral axis, 138 Retinotopic mapping, 138 Retrograde diffusion of cytosolic proteins, 40 flow, 59 “Reverse signaling” pathways, 115 RG. See Radial glia (RG) RGCs. See Radial glial cells (RGCs); Retinal ganglion cells (RGCs) RGFs. See Radial glial fibers (RGFs) Rgrs1 antigen. See SPARC-like 1 (SPARCL1) Rh1. See Rhombomere 1 (Rh1) Rho, 24

Rho-family of small GTPases, 12e13, 71e72, 158 RhoA, 12, 335 Rho GTPase binding domain (RBD), 111 Rho kinase (ROCK), 186 Rho-GTPases, 180, 261 signal through, 262 Rhombencephalic vesicle, 381 Rhombencephalon. See Rhombencephalic vesicle Rhombic lip (RL), 405e406 Rhombomere 1 (Rh1), 405 Rhombomere 4 (r4), 382e384 Ribonucleic acid (RNA), 345e346 Pol I, 307e308 Pol II, 308 translation in dendrites, 262 transport, 44e45 Ribonucleoproteins (RNPs), 45, 305e306 Ribosomal RNA (rRNA), 307 RL. See Rhombic lip (RL) RLN. See Reelin (RLN) RMS. See Rostral migratory stream (RMS) RMTW. See Rostromedial telencephalic wall (RMTW) RNA interference (RNAi), 297, 333e334 RNA-binding Foxs (Rbfox1/2/3), 491e492 RNA-binding motif 4 (RBM4), 491 RNA-binding proteins (RBPs), 262, 488e492, 501 RNAi. See RNA interference (RNAi) Rnd2, 298e299 Rnd3 protein, 483 RNPs. See Ribonucleoproteins (RNPs) “Robo code”, 159 Robo family. See Roundabout family (Robo family) Robo protein expression regulation at midline, 152e154 C. elegans midline, 153e154 Drosophila and vertebrate midlines, 152e153 structure, 147e148, 148f Robo signaling downstream, 157e159 abelson tyrosine kinase, 164e165 actin-interacting proteins, 159 Rho family of small GTPases, 158 regulation at midline in vertebrates, 154e155 ROCK. See Rho kinase (ROCK) Rodent models, 326 Rodent somatosensory system, 266 Ror-type receptor CAM-1, 192 Rostral migratory stream (RMS), 160, 352e353, 465 Rostromedial telencephalic wall (RMTW), 444 Roundabout family (Robo family), 89, 262 homologs, 148e149 receptors, 147e148, 148f, 386e387 Robo gene, 148e149 ROBO1 receptor, 347e348 Robo1, 152

605

Robo2, 152 Robo3, 90, 96, 99e100, 152, 164 SliteRobo interactions, 149e150 regulation, 149e150 Roundworm (Caenorhabditis elegans), 5, 35, 57e58, 88e89, 131e132, 233, 384 attraction and repulsion by UNC-6 in, 97e98 axonal guidance in C. elegans nerve cord, 151f cell polarity regulation, 300 dendrite, 260 midline, 153e154 Netrin-1 and Wnt control axon initiation in, 6 PVD neurons, 234e235 TGF-b family member unc-129 regulates Unc6/Netrin signaling in, 193 Wnt ligand CWN2 regulates C. elegans motor neuron axon guidance, 192 Wnts guide axons of C. elegans mechanosensory neurons and D-type motoneurons, 190e191, 191f RP58 (zinc finger transcription factor), 483 rRNA. See Ribosomal RNA (rRNA) RTK. See Receptor tyrosine kinase (RTK) RTN. See Reticulotegmental nucleus (RTN) RTN4A/Nogo-A. See Reticulon family member 4A (RTN4A/Nogo-A) Ryk, 187, 190 repels axons, 187e188 Ryke/emice, 187e188

S S1P5. See Sphingosine-1-phosphate 5 (S1P5) S1PR2. See Sphingosine 1 phosphate receptor 2 (S1PR2) Saccharomyces cerevisiae, 310 SACs. See Starburst amacrine cells (SACs) SAD-A kinases, 10e11 SAD-A/B kinase, 10e11 SAD-B kinases, 10e11 Sad1p and UNC-84 domain protein (SUN domain protein), 310 SAF-A. See Scaffold attachment factor A (SAF-A) Salamander (Ambystoma tigrinum), 558 Saltatory migration, 351 Satb homeobox 1 (Satb1), 271e273 Satb1. See Satb homeobox 1 (Satb1) SATB1. See Special AT-rich DNA-binding protein 1 (SATB1) Satb2 transcription factor, 482 Satellite secretory systems, 262 SAX-3, 153e155 SAX-7, 224e225 Sax3/Robo activation in C. elegans, 159 SBH. See Subcortical band heterotopia (SBH) SCa axonal transport. See Slow component a axonal transport (SCa axonal transport) Scaffold attachment factor A (SAF-A), 307e308

606 Index

Scar tissue, 204e205 Schizophrenia (SCZ), 278 Schizosaccharomyces pombe, 310 Schwann cells, 225e226, 515 SCI. See Spinal cord injury (SCI) SCZ. See Schizophrenia (SCZ) SDF-1. See Stromal cellederived factor-1 (SDF-1) Sdk-1+. See Sidekick-1 (Sdk-1+) Second-order neurons, 162 Secondary damage, 202e203 Secondary zones, 405 Secreted factors, 521e524 Secreted Frizzled-related protein 2 (sFRP2), 188 Secreted molecules reelin, 330e331 semaphorins, 331 Sema 6A, 331 “Sema domain”, 331 Sema3A. See Semaphorin 3A (Sema3A) Sema5c, 109e110 Sema6A knockout mice, 115 Sema6B, 112e113 Sema7A. See Semaphorin in class 7 (Sema7A) Semaphorin 3A (Sema3A), 7, 67, 109e110, 265e266, 331, 522 Semaphorin 3F, 522 Semaphorin in class 7 (Sema7A), 111, 116e117 Semaphorins, 66e68, 109e111, 269e271, 277e278, 331, 522 function in neural circuit development, 115e116 mechanisms of intracellular signaling, 113e115 in neurological disorders, 116e117 repulsion, 192 signaling, 501 structural features, 111e113 Sensory axons, 97 cortex axonal sprouting, 140 discrimination, 136 olfactory neurons, 437e438 SEP. See Slowly exiting population (SEP) Septin genes, 63 SER. See Smooth ER (SER) Serotonin (5-HT), 470 5-HT+ neuron, 189e190 Serotonin receptor (5-HT3AR), 350 Serum response factor (SRF), 488 Sex-determining region Y-box (Sox) HMG domain protein, 155e156 Sox5, 487e488 Sox6, 345e346 Sox11, 265, 486e487 Sox14, 249e251 SFKs. See Src family kinases (SFKs) sFRP2. See Secreted Frizzled-related protein 2 (sFRP2) SGNs. See Spiral ganglion sensory neurons (SGNs)

SGZ. See Subgranular zone (SGZ) SH3 and multiple ankyrin repeat domains 3 (SHANK3), 116 SH3. See Src-homology 3 (SH3) SHANK3. See SH3 and multiple ankyrin repeat domains 3 (SHANK3) Shh. See Sonic hedgehog (Shh) Shootin1 proteins, 13e14 Short hairpin RNA (shRNA), 5e6 shRNA. See Short hairpin RNA (shRNA) Sidekick-1 (Sdk-1+), 273 Sidekick-2 (Sdk-2+), 273 Siglec-4, 201e202 Signal transduction of forward signaling, 128e130, 129f of reverse signaling, 129f, 130 Signaling mechanisms for dendritic pruning caspases, 251 ubiquitineproteasome system, 251 Silencing effect of Slit on Netrin-mediated attraction, 155 Single-cell RNA sequencing, 566 Single-layered subplate (SP), 480 Single-minded (Sim) transcription factor, 155e156 Single-molecule localization microscopy. See Localization-based microscopy SIP1. See Smad-interacting protein 1 (SIP1) Six-layered cerebral cortex, 442 Slit-N, 149 Slit(s), 66e68, 147, 160e161 discovery and structure, 147e148 protein structure, 148f signaling during midline dendritic guidance, 242e244 slit receptor Robo identification, 148e149 SliteRobo interactions, 149e150 slit2 mutant mice, 160 SLIT3, 164 SliteRobo contribution to axon targeting in complex target field, 162e163 SliteRobo function in midline crossing, 150e155 posttranscriptional Robo regulation, 152 Robo protein expression regulation at midline, 152e154 C. elegans midline, 153e154 Drosophila and vertebrate midlines, 152e153 Robo signaling regulation at midline in vertebrates, 154e155 spatial expression patterns, 152 SliteRobo GTPase-activating proteins (srGAPs), 158, 161 srGAP3, 164 SliteRobo interactions, 163 SliteRobo involvement in nervous system disorders, 164 SliteRobo signaling, 157f for exiting midline, 155 modulation, 155e157 transcriptional control, 155e156 regulation

by metalloprotease cleavage, 156e157 by ubiquitination, 157 roles in nervous system cell migration and cell polarity, 160e161 dendritic and axonal outgrowth and branching, 161e162 lateral positioning, 159e160 SLM. See Stratum lacunosum moleculare (SLM) sLNv. See Small lateral neurons (sLNv) Slow component a axonal transport (SCa axonal transport), 218, 219f Slowly exiting population (SEP), 295 SMA. See Spinal muscular atrophy (SMA) Smad-interacting protein 1 (SIP1), 350e351 Small GTPases, 13, 114 Small lateral neurons (sLNv), 153 Small-scale axon terminal arbor pruning, 225e226 SMN1. See Survival motor neuron 1 (SMN1) Smooth ER (SER), 223 SNARE. See Soluble NSF attachment protein receptor (SNARE) SO. See Stratum oriens (SO) SOCS3. See Suppressor of cytokine signaling 3 (SOCS3) Sojourning, 327 Soluble NSF attachment protein receptor (SNARE), 44 Soluble tubulin, 30 SOM. See Somatostatin (SST) Somal translocation, 289e292, 291f, 557, 561e562 Somatomotor neurons, 389 Somatosensory cortex, 266, 267f Somatostatin (SST), 345e346, 370e371 Son of Sevenless (Sos), 158 Sonic hedgehog (Shh), 92e93, 176, 345e346, 384, 405, 424e425, 524 in axon guidance canonical Shh signaling, 176, 177f chemoattractant for midbrain dopaminergic axons, 182 chemoattractant for spinal cord commissural axons, 178e179 noncanonical Shh signaling pathways, 177f 14e3-3 proteins regulating cell-intrinsic switch, 180e181 binding to Boc attracts commissural axons, 179e180 to Gas1 repels enteric axons, 182 guides axons along longitudinal axis of spinal cord, 180 contralateral and ipsilateral retinal ganglion cell axons, 181e182, 181f induces response of commissural axons to semaphorin repulsion, 192 regulates Wnt signaling in postcrossing commissural axons, 193 Sos. See Son of Sevenless (Sos)

Index

SP. See Single-layered subplate (SP) Space-filling mechanisms to optimize dendritic field distribution, 273e277 dendrite self-avoidance, 275e277, 276f tiling and mosaics, 273e275, 274f SPARC-like 1 (SPARCL1), 487 Spastin, 35, 62 Spatial expression patterns, 152 Spatiotemporal control of cytoskeletal dynamics, 261 Special AT-rich DNA-binding protein 1 (SATB1), 500 Sphingosine 1 phosphate receptor 2 (S1PR2), 204 Sphingosine-1-phosphate 5 (S1P5), 520e521 Spinal cord anatomy, 202 pathfinding in, 134 Spinal cord commissural axons, Shh chemoattractant for, 178e179 Spinal cord injury (SCI), 201, 206e207 repair, 202e203 Spinal muscular atrophy (SMA), 116 Spiral ganglion sensory neurons (SGNs), 135 SPNs. See Subplate neurons (SPNs) “Springlike” pia-attached process, 295e296 SR. See Stratum radiatum (SR) Src family kinases (SFKs), 130, 133f, 179 GoF axon guidance phenotypes, 129 Src-homology 3 (SH3), 115, 158 SRF. See Serum response factor (SRF) srGAPs. See SliteRobo GTPase-activating proteins (srGAPs) SST. See Somatostatin (SST) SST+. See Neuropeptide somatostatin (SST+) St18. See Suppression of tumorigenicity 18 (St18) Stabilization, 265 Starburst amacrine cells (SACs), 112e113, 270fe271f Starburst cell dendrites, 275e277 Stathmin/Op-18 activity, 34 STEF/Tiam1, 299e300 Stellate cells, 403e404, 408e409 “Stop and Go” model, 218 “Stop-growing” signals, 264e265 Stratum lacunosum moleculare (SLM), 440 Stratum oriens (SO), 440 Stratum pyramidale, 369 Stratum radiatum (SR), 365, 440 “Stressed” MTs, 220 Stripe assays, 136 Stromal cellederived factor-1 (SDF-1). See Chemokine 12 (CXCL12) Subcellular cytoskeletal specializations, 37e39 Subcellular patterning of synaptic inputs, 277e278 Subcortical band heterotopia (SBH), 579e581, 579f Subgranular zone (SGZ), 373e374 establishment, 374e375 Subpallial neurons, 570

Subpallium, 479e480 Subplate neurons (SPNs), 444, 567e569 “Subunit heterogeneity” strategy, 219e220 a-Subunit of calcium/calmodulin-dependent kinase II (aCaMKII), 45 Subunit transport model, 35 Subventricular zone (SVZ), 23e24, 158, 289e290, 292, 294e295, 324, 325f, 326e327, 347e348, 371, 479, 534 precursor cells, 326 SUN domain protein. See Sad1p and UNC84 domain protein (SUN domain protein) Superresolution imaging methods, 38e39 Suppression of tumorigenicity 18 (St18), 350e351 Suppressor of cytokine signaling 3 (SOCS3), 207 Survival motor neuron 1 (SMN1), 45, 117 SVZ. See Subventricular zone (SVZ) Synapse selectivity, 273 Synaptic nuclear envelope (Syne), 310 Synaptogenesis, 115, 264e265 Syne. See Synaptic nuclear envelope (Syne) Syne-4 protein, 311 Synergy between Netrin-1 from floor plate and from ventricular zone, 93 Syntaphilin, 64 a-Synuclein, 223

T T cell factor (TCF), 184 T-box brain 1 (Tbr1), 271, 482 TADs. See Topologically associated domains (TADs) TAG-1. See Transient axonal glycoprotein-1 (TAG-1) Talipes equinovarus, 130 Tangential jump, 293 Tangential migration, 353, 382e384, 383f, 495e501. See also Radial migration ending, 394e395 impact on brain evolution, 563e572 and evolution, 568t glutamatergic tangential contributions as developmental scaffolds, 567e570 guidepost neurons in evolution, 570e571 pallial interneurons and brain circuits modulation, 565e567 of interneurons. See also Interneurons (INs) birth of distinct interneuron subtypes and onset of migration, 345e346 cell-intrinsic regulation of cortical interneuron migration, 351 coronal section through telencephalon, 346f dynamic remodeling of cytoskeleton, 352 evolutionary perspective of, 354 factors role in, 350e351 molecular cues controlling integration of interneurons, 348e349 molecular cues controlling intracortical dispersion of interneurons, 349e350

607

molecular cues drawing path of cortical interneuron migration, 347e348 molecular regulation of migration of STR interneurons, 353 regulation in RMS to OB, 352e353 signals dictating arrest of interneuron migration within cortical wall, 350 interplay of transcription factors and guidance cues, 498e501 molecular mechanisms controlling tangential migration of precerebellar neurons, 384e389 influence of midline on, 384e388 regulation of precerebellar migration, 385f posttranscriptional events, 501, 502t routes, 347f shaping vertebrate brain, 555e557 transcriptional regulators, 502t Tangentially migrating neurons in development of, 448e449 of guidepost neurons, 449e450 TAOK2, 278 Target of rapamycin complex 2 (TORC2), 247 Targeting and synapse selectivity, 268e273 Tau, 220, 223 expression, 221 TBCB. See Tubulin folding cofactor B (TBCB) Tbr1. See T-box brain 1 (Tbr1) Tbr2, 335 Tbr2+ Ips, 484 TCAs. See Thalamocortical axons (TCAs) TCE. See Translational control element (TCE) TCF. See T cell factor (TCF) Tenascin-C (TN-C), 520 Tenascin-R, 470 Terminal transferase dUTP nick end labeling (TUNEL), 293e294 Terminal translocation, 290e292 Terminal zones (TZs), 67 Tertiary germinative matrix, 373 TFs. See Transcription factors (TFs) TG. See Trigeminal ganglion (TG) TGF-b. See Transforming growth factor-b (TGF-b) TGN. See Trans-Golgi network (TGN) TH. See Tyrosine hydroxylase (TH) Thalamic axons, 266 Thalamic relay nuclei, 139 Thalamocortical axons (TCAs), 115, 442, 444, 449e450, 541e542 subpallium is major intermediate target for, 444e445 Thalamocortical projections, 139 Thalamocortical tract, 348e349 Tiling, 273e275, 274f Timothy syndrome, 278 Tissue-type plasminogen activator (tPA), 519 TK. See Tyrosine kinase (TK) TLR2. See Toll-like receptor 2 (TLR2) TLR4. See Toll-like receptor 4 (TLR4) TN-C. See Tenascin-C (TN-C)

608 Index

Toll-like receptor 2 (TLR2), 209 Toll-like receptor 4 (TLR4), 209 Topographic map formation, 70 Topographic mapping mechanism, 137 Topologically associated domains (TADs), 307e308 TORC2. See Target of rapamycin complex 2 (TORC2) tPA. See Tissue-type plasminogen activator (tPA) TPX2 protein, 61 Trachemys scripta. See Turtle (Trachemys scripta) TRACT method, 202 Trafficking, 127e128 long-range trafficking to dendrites, 261 mechanisms, 23 Trans, 124e125 Trans-Golgi network (TGN), 261 Transcription factors (TFs), 345e346 encoding genes, 567 neurogenins, 336 Pax6, 335 Tbr2, 335 Transcriptional control of dendritic morphology, 237e241 chromatin remodeling factors and dendritic development, 241 identity control of Drosophila PNS neurons, 237e239, 238f of dendritic targeting of olfactory PNs, 239e241, 240f of laminar-specific targeting, 271e273 of neuronal migration, 480e501 Transcriptional regulators radial migration, 489te490t tangential migration, 502t Transcriptional repression, 483, 500 Transcytosis, 43e44, 46 Transforming growth factor-b (TGF-b), 249 family member unc-129 regulates Unc6/ Netrin signaling in C. elegans, 193 signaling, 7 signaling during dendritic remodeling, 249 superfamily members in axon guidance BMP7:GDF7 repels spinal cord commissural axons, 183e184 canonical bone morphogenetic protein signaling, 183 Transient axonal glycoprotein-1 (TAG-1), 293, 388, 391e392 Translational control element (TCE), 242 Transmembrane protein Linx, 444 semaphorins, 520 Transplanted blood vessel mimics, 473 Trigeminal ganglion (TG), 65 TrkB. See Tropomysin-related kinase B (TrkB) TrkRs. See Tropomyosin-related kinase receptors (TrkRs) Tropomyosin-related kinase receptors (TrkRs), 65

Tropomysin-related kinase B (TrkB), 347 TSC. See Tuberous sclerosis complex (TSC) Tuberous sclerosis complex (TSC), 10e11, 580 Tsc2, 130 Tubulin, 30, 61, 220e221 Tubulin folding cofactor B (TBCB), 223 g-Tubulin ring complex (gTuRC), 30 TUNEL. See Terminal transferase dUTP nick end labeling (TUNEL) Turtle (Trachemys scripta), 566 Twin-spot MARCM, 232 “Twist-plus-stretch” model, 292, 295e296 Two-stroke motility paradigm, 413e414 Type 2 lissencephaly. See Cobblestone; lissencephaly Type 2 SGNs, 135 Type-specific neuronal morphology, 271 Tyrosinated tubulin, 32e33 Tyrosination, 62 Tyrosine hydroxylase (TH), 352e353 Tyrosine kinase (TK), 114e115, 125f, 131f TZs. See Terminal zones (TZs)

U u-PAR. See Urokinase-type plasminogen activator receptor (u-PAR) UBCs. See Unipolar brush cells (UBCs) Ubiquitin, 46 Ubiquitin proteosome system (UPS), 46, 64e65, 223, 251 Ubiquitin-specific protease 33 (USP33), 157 Ubiquitination, sliterobo signaling regulation by, 157 UBP. See (S)-1-(2-Amino-2-carboxyethyl)-3(2-carboxybenzyl) pyrimidine-2,4dione (UBP) UNC-129, 193 UNC-34/Enabled protein, 159 unc-40 gene, 89 UNC-6, 193 in C. elegans, 97e98 UNC5 family, 89 receptors, 90 unc-5 gene, 89 UNC-5 receptor, 193 Unc5D, 301 Unipolar brush cells (UBCs), 403e404, 411 United Parcel Service (UPS), 40e42 Untranslated regions (UTRs), 242, 483 Upper RL derivatives migration, 410e411 CNs, 410e411 GNPs and CGNs, 411 UBCs, 411 UPS. See Ubiquitin proteosome system (UPS); United Parcel Service (UPS) Urodelian brain cytoarchitecture, 558 Urokinase-type plasminogen activator receptor (u-PAR), 347 USP33. See Ubiquitin-specific protease 33 (USP33) UTRs. See Untranslated regions (UTRs)

V V-SVZ. See Ventricular-subventricular zone (V-SVZ) Valosin-containing protein (VCP), 251 Van goghlike 2 (vangl2), 390 VAMP3, 368e369 vangl2. See Van goghlike 2 (vangl2) Vascular endothelial growth factor (VEGF), 352e353, 535e536 VEGF-A, 392e393, 521 Vasoactive intestinal peptide (VIP), 345e346, 370e371 VCN. See Ventral cochlear nucleus (VCN) VCP. See Valosin-containing protein (VCP) VEGF. See Vascular endothelial growth factor (VEGF) Ventral cochlear nucleus (VCN), 135 Ventral telencephalon, 139 Ventricular zone (VZ), 23e24, 289e292, 294e295, 324, 325f, 347e348, 384, 405, 472, 479e480, 534 precursor cells, 324 VZederived Netrin-1 in spinal cord, 93 Ventricular-subventricular zone (V-SVZ), 465 directional control from, 469 Vertebrate, 57e58 L1 family members, 224e225 midlines, 152e153 neurons, 275 spinal nerves, 132 Vertebrate brain development, neuronal migration types in, 555e557 Vertebrate dendrites cell biology of dendritic growth, 259e263 actin cytoskeleton regulators, 260e261 dendrite elaboration requires satellite secretory pathway, 261e262 intracellular cascades role, 262e263 microtubule network regulators, 259e260 powering dendrite growth, 262 RNA translation in dendrites, 262 dendrite compartmentalization emergence, 277e278 patterning membrane excitability of dendritic compartments, 278 subcellular patterning of synaptic inputs along dendritic domains, 277e278 dendritic field formation I control, 263e265 activity-dependent mechanisms, 264e265, 264f afferent-derived neurotrophins limit size, 263 control of arbor size by neurotransmission, 263e264 dendritic field formation II control, 265e267 activity-dependent orientation of dendrite growth, 266, 267f apical dendrite initiation and outgrowth of cortical pyramidal neurons, 265e266

Index

positional cues shape asymmetric dendritic arbors in mouse retina, 266e267 dendritic field formation III control, 268e273 integrated, multistep model for synaptic wiring in retina IPL, 273 laminar targeting of retinal dendrites, 269e271 local recognition mechanisms to control synapse selectivity, 273 proto-IPL formation by retinal amacrine cells, 269 transcriptional control of laminar-specific targeting of dendrites, 271e273 neurodevelopmental disorders, 278 space-filling mechanisms to optimize dendritic field distribution, 273e277 dendrite self-avoidance, 275e277, 276f tiling and mosaics, 273e275, 274f structure and function of, 257e259 methods for manipulating and studying dendrite morphology, 259 Vertebrate retina development, synapse formation in Very low-density lipoprotein receptor (VLDLR), 330e331, 394 Vesicular trafficking, 127 VESPR. See Virus-encoded semaphorin protein receptor (VESPR) VIP. See Vasoactive intestinal peptide (VIP) Virus-encoded semaphorin protein receptor (VESPR), 110e111 VLDLR. See Very low-density lipoprotein receptor (VLDLR) VNO project. See Vomeronasal organ project (VNO project) Vomeronasal organ project (VNO project), 163 Vomeronasal sensory neuron (VSN), 163, 163f

VSN. See Vomeronasal sensory neuron (VSN) VZ. See Ventricular zone (VZ)

W Wallerian degeneration, 227, 229 Wallerian degeneration slow mutant mouse (Wlds mutant mouse), 227e228 conserved protective effects, 228f WASP. See WiskotteAldrich syndrome protein (WASP) WAVE regulatory complex (WRC), 261 White matter (WM), 202f WIRS motifs, 261 WiskotteAldrich syndrome protein (WASP), 29, 261 Wlds mutant mouse. See Wallerian degeneration slow mutant mouse (Wlds mutant mouse) WM. See White matter (WM) Wnt control axon initiation in C. elegans, 6 Wnt signaling, 66e68 Wnt1-Cre mice, 178 Wnt3a protein, 367 Wnt5a, 187e188 Wnts in axon guidance, 176 canonical and noncanonical Wnt signaling, 184e186, 185f Wnt binding to Fz attracts postcrossing commissural axons, 188e189 to Fz regulates dopaminergic axon guidance, 189e190 to Ryk repels axons of corticospinal tract and corpus callosum, 187e188 Wnt ligand CWN2 regulates C. elegans motor neuron axon guidance, 192 Wnt3 mediates mediolateral retinotectal topographic mapping, 190

609

Wnt5 repels commissural axons from Drosophila posterior commissure, 186e187, 186f Wnt5, complexed with derailed, repels Drosophila mushroom body axons, 187 Wnts guide axons of C. elegans mechanosensory neurons and D-type motoneurons, 190e191, 191f WRC. See WAVE regulatory complex (WRC)

X X-linked lissencephaly with abnormal genitalia (XLAG), 452, 583e584 X-ray crystallography, 127 Xenopus, 5e6, 90 RGC, 6e7 polarized emergence of axon in, 6e7 Xenopus laevis. See Clawed toad (Xenopus laevis) XLAG. See X-linked lissencephaly with abnormal genitalia (XLAG)

Z Zac1, 487 ZBP1. See Zipcode-binding protein 1 (ZBP1) Zeb1-Pard6/3A transcriptional pathway, 423 Zebrafish, 201e202, 208e209 Zinc finger protein, 491 Zipcode-binding protein 1 (ZBP1), 45, 179e180, 262 Zona pellucida domain protein (ZP domain protein), 244e245 Zonadhesin, 244e245 ZP domain protein. See Zona pellucida domain protein (ZP domain protein) Zymosan, 209