[1st Edition] 9780128124680, 9780128124673

Table of contents :
Content:
Series PagePage ii
CopyrightPage iv
ContributorsPages ix-x
Chapter One - New Insights Into the Roles of Retinoic Acid Signaling in Nervous System Development and the Establishment of Neurotransmitter SystemsPages 1-84E. Zieger, M. Schubert
Chapter Two - AMBRA1, a Novel BH3-Like Protein: New Insights Into the AMBRA1–BCL2-Family Proteins RelationshipPages 85-113A. Di Rita, F. Strappazzon
Chapter Three - Rationale for the Combination of Dendritic Cell-Based Vaccination Approaches With Chemotherapy AgentsPages 115-156I. Truxova, M. Hensler, P. Skapa, M.J. Halaska, J. Laco, A. Ryska, R. Spisek, J. Fucikova
Chapter Four - Smac Mimetics to Therapeutically Target IAP Proteins in CancerPages 157-169S. Fulda
Chapter Five - Consequences of Keratin Phosphorylation for Cytoskeletal Organization and Epithelial FunctionsPages 171-225M.S. Sawant, R.E. Leube
Chapter Six - Plastid Protein Targeting: Preprotein Recognition and TranslocationPages 227-294P. Chotewutmontri, K. Holbrook, B.D. Bruce
Chapter Seven - Immunomodulatory Activity of VEGF in CancerPages 295-342A. Lapeyre-Prost, M. Terme, S. Pernot, A.-L. Pointet, T. Voron, E. Tartour, J. Taieb

Citation preview

INTERNATIONAL REVIEW OF CELL AND MOLECULAR BIOLOGY Series Editors GEOFFREY H. BOURNE JAMES F. DANIELLI KWANG W. JEON MARTIN FRIEDLANDER JONATHAN JARVIK LORENZO GALLUZZI

1949–1988 1949–1984 1967–2016 1984–1992 1993–1995 2016–

Editorial Advisory Board KEITH BURRIDGE AARON CIECHANOVER SANDRA DEMARIA SILVIA FINNEMANN KWANG JEON

CARLOS LOPEZ-OTIN WALLACE MARSHALL SHIGEKAZU NAGATA MOSHE OREN ANNE SIMONSEN

Academic Press is an imprint of Elsevier 50 Hampshire Street, 5th Floor, Cambridge, MA 02139, United States 525 B Street, Suite 1800, San Diego, CA 92101-4495, United States The Boulevard, Langford Lane, Kidlington, Oxford OX5 1GB, United Kingdom 125 London Wall, London, EC2Y 5AS, United Kingdom First edition 2017 Copyright © 2017 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. ISBN: 978-0-12-812467-3 ISSN: 1937-6448 For information on all Academic Press publications visit our website at https://www.elsevier.com/books-and-journals

Publisher: Zoe Kruze Acquisition Editor: Alex White Editorial Project Manager: Fenton Coulthurst Production Project Manager: Magesh Kumar Mahalingam Cover Designer: Mark Rogers Typeset by SPi Global, India

CONTRIBUTORS B.D. Bruce Graduate School of Genome Science and Technology, University of Tennessee, Knoxville, TN, United States P. Chotewutmontri Graduate School of Genome Science and Technology, University of Tennessee, Knoxville, TN, United States A. Di Rita IRCCS Santa Lucia Foundation; University of Rome Tor Vergata, Rome, Italy J. Fucikova 2nd Faculty of Medicine and University Hospital Motol, Charles University; Sotio a.s., Prague, Czech Republic S. Fulda Institute for Experimental Cancer Research in Pediatrics, Goethe-University, Frankfurt; German Cancer Consortium (DKTK); German Cancer Research Center (DKFZ), Heidelberg, Germany M.J. Halaska 3rd Faculty of Medicine and Faculty Hospital Kralovske Vinohrady, Charles University, Prague, Czech Republic M. Hensler Sotio a.s., Prague, Czech Republic K. Holbrook University of Tennessee, Knoxville, TN, United States J. Laco Faculty of Medicine and Faculty Hospital in Hradec Kralove, Charles University, Prague, Czech Republic A. Lapeyre-Prost INSERM U970, PARCC (Paris Cardiovascular Research Center), Universit
e Paris-Descartes, Paris, France R.E. Leube Institute of Molecular and Cellular Anatomy, RWTH Aachen University, Aachen, Germany S. Pernot INSERM U970, PARCC (Paris Cardiovascular Research Center), Universit
e Paris-Descartes; Service d’h
epatogastroent
erologie et d’oncologie digestive, H^ opital Europ
een Georges Pompidou, Paris, France

ix

x

Contributors

A.-L. Pointet INSERM U970, PARCC (Paris Cardiovascular Research Center), Universit
e Paris-Descartes; Service d’h
epatogastroent
erologie et d’oncologie digestive, H^ opital Europ
een Georges Pompidou, Paris, France A. Ryska Faculty of Medicine and Faculty Hospital in Hradec Kralove, Charles University, Prague, Czech Republic M.S. Sawant Institute of Molecular and Cellular Anatomy, RWTH Aachen University, Aachen, Germany M. Schubert Sorbonne Universit
es, UPMC Universit
e Paris 06, CNRS, Laboratoire de Biologie du D
eveloppement de Villefranche-sur-Mer, Observatoire Oc
eanologique de Villefranche-surMer, Villefranche-sur-Mer, France P. Skapa 2nd Faculty of Medicine and University Hospital Motol, Prague, Czech Republic R. Spisek 2nd Faculty of Medicine and University Hospital Motol, Charles University; Sotio a.s., Prague, Czech Republic F. Strappazzon IRCCS Santa Lucia Foundation, Rome, Italy J. Taieb INSERM U970, PARCC (Paris Cardiovascular Research Center), Universit
e Paris-Descartes; Service d’h
epatogastroent
erologie et d’oncologie digestive, H^ opital Europ
een Georges Pompidou, Paris, France E. Tartour INSERM U970, PARCC (Paris Cardiovascular Research Center), Universit
e Paris-Descartes; Service d’immunologie biologique. H^ opital Europ
een Georges Pompidou, Paris, France M. Terme INSERM U970, PARCC (Paris Cardiovascular Research Center), Universit
e Paris-Descartes, Paris, France I. Truxova 2nd Faculty of Medicine and University Hospital Motol, Charles University; Sotio a.s., Prague, Czech Republic T. Voron INSERM U970, PARCC (Paris Cardiovascular Research Center), Universit
e Paris-Descartes; Service de chirurgie digestive, H^ opital Europ
een Georges Pompidou, Paris, France E. Zieger Sorbonne Universit
es, UPMC Universit
e Paris 06, CNRS, Laboratoire de Biologie du D
eveloppement de Villefranche-sur-Mer, Observatoire Oc
eanologique de Villefranche-surMer, Villefranche-sur-Mer, France

CHAPTER ONE

New Insights Into the Roles of Retinoic Acid Signaling in Nervous System Development and the Establishment of Neurotransmitter Systems E. Zieger, M. Schubert1 Sorbonne Universit
es, UPMC Universit
e Paris 06, CNRS, Laboratoire de Biologie du D
eveloppement de Villefranche-sur-Mer, Observatoire Oc
eanologique de Villefranche-sur-Mer, Villefranche-sur-Mer, France 1 Corresponding author: e-mail address: [email protected]

Contents 1. 2. 3. 4. 5. 6.

Introduction Developmental Signaling Retinoids as Morphogens, Teratogens, and Drugs RA Signaling Pathway Major Functions of RA Signaling RA Signaling During Nervous System Development 6.1 Posteriorizing Neural Plate 6.2 Setting Up RA Signaling System and Boundaries of Neural Tube 6.3 Patterning Neural Tube Along the Anterior–Posterior Axis 6.4 Patterning Neural Tube Along Dorsal–Ventral Axis 6.5 Orchestrating Proliferation, Differentiation, and Survival of Neural Progenitors 6.6 Inducing and Coordinating Neural Crest Development 6.7 Inducing and Coordinating Cranial Placode Development 6.8 Regulating Formation of Neural Crest and Cranial Placode Derivatives 7. RA Signaling and the Establishment of Neurotransmitter Systems 7.1 Natural Neuroactive Substances: Neurotransmitters, Neuromodulators, and Neurohormones 7.2 Catecholaminergic Neurons 7.3 Serotonergic Neurons 7.4 Glutamate and GABAergic Neurons 8. RA Signaling Outside Vertebrates: Evolutionary Considerations 9. Concluding Remarks Acknowledgments References

International Review of Cell and Molecular Biology, Volume 330 ISSN 1937-6448 http://dx.doi.org/10.1016/bs.ircmb.2016.09.001

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2017 Elsevier Inc. All rights reserved.

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Abstract Secreted chiefly from the underlying mesoderm, the morphogen retinoic acid (RA) is well known to contribute to the specification, patterning, and differentiation of neural progenitors in the developing vertebrate nervous system. Furthermore, RA influences the subtype identity and neurotransmitter phenotype of subsets of maturing neurons, although relatively little is known about how these functions are mediated. This review provides a comprehensive overview of the roles played by RA signaling during the formation of the central and peripheral nervous systems of vertebrates and highlights its effects on the differentiation of several neurotransmitter systems. In addition, the evolutionary history of the RA signaling system is discussed, revealing both conserved properties and alternate modes of RA action. It is proposed that comparative approaches should be employed systematically to expand our knowledge of the context-dependent cellular mechanisms controlled by the multifunctional signaling molecule RA.

1. INTRODUCTION The nervous system can be understood as a means of internal interconnection that enables multicellular animals to coordinate their different physiological activities and interact with their environment (Sherrington, 1906). Depending on the morphology of an organism and the tasks it needs to perform throughout its life history, more or less specialized connections are required to receive, process, and distribute information in a targeted manner. Accordingly, animals have evolved a wide range of different neural cell types and networks (Arendt et al., 2016; Hartenstein and Stollewerk, 2015). Wolpert (1992) described metazoan evolution as “the brilliant result of altering developmental programs.” In order to form different characters, embryos rely on instructive signals that differentially modulate target gene transcription in specific cellular subsets, thus inducing them to commit to different developmental fates and to assume different properties and functions. While the number of cell types varies between different animals, ranging from a maximum of about 16 in sponges to approximately 244 in vertebrates (Niklas et al., 2014), only a surprisingly limited number of well-conserved signaling pathways appears to be required for generating and organizing this cellular diversity. The focus of this review is on one of these pathways, retinoic acid (RA) signaling, and its roles during the formation of the vertebrate nervous system. Special attention is given to the still poorly understood influence of RA signaling on the neurotransmitter identity of specific neuronal subsets, including catecholaminergic, serotonergic,

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glutamatergic, and GABAergic neurons. In addition, the evolutionary background of RA signaling is discussed, which can provide new insights into the conserved properties of this multifunctional morphogen, but still remains elusive outside of the chordate phylum.

2. DEVELOPMENTAL SIGNALING A common feature of intercellular signaling pathways is that they can alter cell physiology and development in disparate ways, depending on a spatial and temporal context. This flexibility is generated by several mechanisms that often act in combination in determining the specificity of cellular responses. One important element is cell competence, which is acquired during the specific developmental history of a cell, and is based on differences in its molecular makeup at the epigenetic, transcriptional, and/or translational levels, e.g., chromatin compaction, expression of transcriptional regulators, presence of signaling pathway components, and other interacting factors (Wang et al., 2015). These differences affect the capability of cells to interpret developmental signals and can be established as early as at the first cleavage, when maternal factors become unevenly distributed between the initial two blastomeres (Heasman, 2006; Kumano and Nishida, 2009; Peng and Wikramanayake, 2013; Wang et al., 2015; Weitzel, 2004). Another way to achieve cell type-specific responses is through the variation of signal intensity, for instance by producing a concentration gradient of signaling molecules (i.e., morphogens) (Kicheva et al., 2012). This process often involves the secretion of a morphogen from a localized source, followed by its nondirectional spread throughout a tissue and its degradation (Kicheva et al., 2012). However, morphogen gradients can also be formed through lineage or directional transport of morphogens (Kicheva et al., 2012). Lastly, different signaling pathways do not function as isolated entities, but are integrated into highly complex molecular frameworks that enable contextual responses (Attisano and Wrana, 2013; Guruharsha et al., 2012). During the course of development, the spatial pattern of cell identities and behaviors is thus dynamically determined through the cross talk between variable levels of morphogen signals and depends recursively on cellular competence. Classical nuclear receptor (NR) pathways function through ligandinduced transcription factors that regulate the expression of specific target genes in response to endocrine signals, such as vitamin D, sex steroids (e.g., progestins, estrogens, androgens), adrenal steroids (glucocorticoids,

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mineralocorticoids), thyroid, and retinoid hormones (McKenna et al., 1999). NRs contain a highly conserved zinc finger-based DNA-binding domain that allows them to associate with specific regulatory sites in the genome, i.e., enhancers or silencers (Escriva et al., 2000; Kininis and Kraus, 2008; McKenna et al., 1999). At their target promoters, NRs directly control transcriptional activity in a ligand-dependent manner, through both the regulation of RNA polymerase binding and chromatin remodeling, which in turn modulates gene expression via epigenetic changes (Evans and Mangelsdorf, 2014; Kininis and Kraus, 2008; McKenna et al., 1999). NR function generally involves the recruitment or release of coregulators, with coactivators mediating the potentiating functions of activated NRs and corepressors transducing the attenuating functions of nonactivated NRs (McKenna and O’Malley, 2002). Owing to posttranslational modification and selective recruitment of coregulators, NR dimerization as well as complex interactions between NRs, coregulators, and other transcription factors, NRs are able to govern a broad array of finely tuned transcriptional responses (Evans and Mangelsdorf, 2014; Kininis and Kraus, 2008; McKenna and O’Malley, 2002). Accordingly, these signaling pathways play very diverse roles in the regulation of physiological, developmental, and metabolic processes (Kininis and Kraus, 2008; McKenna and O’Malley, 2002; McKenna et al., 1999).

3. RETINOIDS AS MORPHOGENS, TERATOGENS, AND DRUGS Retinoids are biologically active derivatives of vitamin A (retinol) that have great impact on vertebrate development and homeostasis (Doll
e and Niederreither, 2015). A disruption of physiological retinoid levels during pregnancy, for instance, can cause abnormal morphological development: an excess of vitamin A can lead to exencephaly, cleft palate, spina bifida, eye defects, hydrocephaly as well as shortening of the mandible and maxilla, whereas vitamin A deficiency has been correlated with malformations in the eye, urogenital tract, diaphragm, heart, and lung (Collins and Mao, 1999). Certain retinoids, such as 13-cis RA (isotretinoin, Accutane®), are well known to act as teratogens in both animals and humans (Collins and Mao, 1999). Therefore, a great environmental threat is posed by cyanobacteria blooms, which can produce significant levels of teratogenic retinoids, contaminating sources of food and drinking water (Jonas et al., 2014; Wu et al., 2012). Moreover, glyphosate-based herbicides have been shown to increase the

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physiological activity of retinoid signaling pathways and are thus considered responsible for the higher incidence of embryonic malformations and spontaneous abortions in populations exposed to pesticides (Lo´pez et al., 2012). Nonetheless, retinoids, such as 13-cis RA (isotretinoin, Accutane®), are also being used for numerous pharmacological applications, including the treatment of skin diseases and various forms of cancer (Camacho, 2003; Collins and Mao, 1999; Doll
e and Niederreither, 2015). In addition, their potential for attenuating neurological diseases is currently under investigation (Doll
e and Niederreither, 2015; Endres et al., 2014; Lerner et al., 2012; Mizee et al., 2014). Taken together, retinoids are important as teratogens, but also for medical applications and it is therefore crucial to improve our understanding of retinoid signaling functions.

4. RA SIGNALING PATHWAY The structurally simple and diffusible molecule RA (Fig. 1) is well known to act as a crucial regulator of gene expression during vertebrate embryogenesis (Blomhoff, 1994; De Luca, 1991; Sporn et al., 1994). Different stereoisomeric forms of RA can be found in vivo, including all-trans, 9-cis, and 13-cis RA. While it has been established that in vertebrates all-trans RA is the physiologically active form, possible biological functions of other RA isomers are still being debated (Campo-Paysaa et al., 2008; Mic et al., 2003). Endogenous RA synthesis requires the cellular uptake of retinol, which is tightly regulated. Due to its lipophilic nature, virtually all retinol present in the bloodstream is bound to serum retinol-binding proteins (RBPs) (Goodman, 1984). The transmembrane transporter STRA6 (stimulated by RA protein 6) functions as RBP receptor and mediates the bidirectional flux of retinol between extracellular RBPs and intracellular RBPs (D’Ambrosio et al., 2011; Kawaguchi et al., 2007, 2012). The activity of STRA6 depends on both extra- and intracellular retinoid levels (D’Ambrosio et al., 2011; Kawaguchi et al., 2012; Sun, 2012). Retinol uptake via STRA6 activity can thus be enhanced through coupling of LRATs (lecithin retinol acyltransferases), which are important enzymes for the metabolism of retinol in the visual system and in storage tissues (Golczak et al., 2008; Isken et al., 2008; Kawaguchi et al., 2007). Moreover, in the bloodstream RBP can form a complex with transthyretin (TTR), a thyroid hormone distributor protein (D’Ambrosio et al., 2011; Richardson, 2009; Yamauchi and Ishihara, 2009) that blocks the ability of RBP to associate with STRA6, thus suppressing STRA6-mediated retinol uptake (Berry et al., 2012). Inside the cell, retinol

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Fig. 1 Schematic representation of the structure, metabolism, and canonical signaling pathway of retinoic acid (RA). Abbreviations: ADH, alcohol dehydrogenase; CRABP, cellular retinoic acid-binding protein; CRBP, cellular retinol-binding protein; CYP26, cytochrome P450 subfamily 26 protein; RA, retinoic acid; RALDH, retinaldehyde dehydrogenase; RAR, retinoic acid receptor; RARE, retinoic acid response element; RBP, retinol-binding protein; RXR, retinoid X receptor; SDR, short-chain dehydrogenase/reductase; STRA6, stimulated by retinoic acid protein 6; TTR, transthyretin. Modified from Laudet, V., Zieger, E., Schubert, M., 2015. Evolution of the retinoic acid signaling pathway. In: Doll
e, P., Neiderreither, K. (Eds.), The Retinoids. John Wiley & Sons, Hoboken, NJ, pp. 75–90.

is bound to cellular retinol-binding proteins (CRBPs), since it is otherwise chemically instable and has a low solubility in aqueous media (Newcomer, 1995). Four types of CRBPs (CRBP I, II, III, and IV) with distinct tissue distributions have been structurally characterized that bind either retinol or

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retinal and play an important part in the absorption, transport, metabolism, and excretion of these retinoids (Conforti et al., 2000; Mezaki et al., 2012; Napoli, 1993; Noy, 2000). The conversion of cellular retinol into RA begins with the reversible oxidation of retinol to retinal (retinaldehyde). This reaction is catalyzed either by cytosolic alcohol dehydrogenases (ADHs) or by short-chain dehydrogenases/reductases (SDRs) (Par
es et al., 2008). Subsequently, retinaldehyde dehydrogenases (RALDHs) irreversibly transform retinal into RA (Par
es et al., 2008). In vertebrates, four RALDHs with distinct tissue-specific expression profiles are known to contribute to endogenous RA production: RALDH1, 2, and 3 are class 1 aldehyde dehydrogenases (ALDH1s), whereas RALDH4 is a class 8 ALDH (Theodosiou et al., 2010). Aside from RALDHs, cytochrome p450 monooxygenases, such as CYP1B1, can also oxidize retinol or retinal to RA (Chambers et al., 2007; Collins and Mao, 1999). As for retinol and retinal, RA is bound by specific cellular RA-binding proteins, known as CRABP-I and CRABPII, which are crucial for the intracellular balance between RA synthesis, degradation, and transportation to RA receptors (RARs) within the nucleus (Cai et al., 2012; Carvalho and Schubert, 2013; Dong et al., 1999; Napoli, 1999; Noy, 2000). CRABP-I is mainly involved in RA catabolism and, through interaction with RA degrading enzymes, even regulates cytoplasmatic levels of various types of RA metabolites (Boylan and Gudas, 1992). In contrast, CRABP-II appears to be more directly involved in the process of RA signaling. Binding of RA induces CRABP-II to dissociate from the endoplasmic reticulum and undergo nuclear translocation, which ultimately leads to the formation of a complex between CRABP-II and the NR of RA, through which RA is transferred from the binding protein to the receptor (Majumdar et al., 2011; Schug et al., 2007). Owing to the great importance of RA as a morphogen, physiological retinoid levels must be tightly controlled and buffered, for instance, against the variable bioavailability of retinol. This is achieved through a complex system of proteins and enzymes that bind, redistribute, synthesize, and degrade RA and its precursors (Doll
e and Niederreither, 2015). The activity of most of these molecules directly depends on retinoid concentration levels and is regulated via multiple positive and negative feedback loops (Doll
e and Niederreither, 2015). For example, RA can repress its synthesizing enzymes and promote its degrading enzymes (Dobbs-McAuliffe et al., 2004; Ross and Zolfaghari, 2011; Wang et al., 2002). For its removal from the cell, RA is oxidized by specific cytochrome P450 enzymes, the CYP26s (White et al., 1997). In mammals, the three CYP26s, namely CYP26A1, CYP26B1, and

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CYP26C1, can degrade RA into 4-oxo-RA, 4-hydroxy-RA, 5,6-epoxy-RA, 5,8-epoxy-RA, and probably other polar metabolites (White and Schilling, 2008). In human fetal liver, RA can also be hydroxylated by CYP3A7, which might be crucial in the embryo for preventing RA-induced toxicity (Chen et al., 2000). In general, RA signals are transduced by heterodimers of two nuclear hormone receptors, the RAR, and the retinoid X receptor (RXR). These RAR/RXR heterodimers bind directly to specific DNA motifs, called RA response elements (RAREs) (Chambon, 1996). Typically, a RARE consists of two direct repeats that have the consensus nucleotide sequence (A/G) G(G/T)(G/T)(G/C)A, but the topology of and spacing between RARE half sites varies significantly (Balmer and Blomhoff, 2005; Moutier et al., 2012). In the absence of a ligand, the association of RAR/RXR heterodimers with RAREs leads to the recruitment of corepressor complexes that mediate chromatin compaction and target gene repression (Vilhais-Neto and Pourqui
e, 2008). However, the ability of the heterodimers to interact with the regulatory regions of certain target genes can also be dependent on ligand binding (Gutierrez-Mazariegos et al., 2014b; Moutier et al., 2012). Fixation of RA by RAR activates the ligand-dependent transcription factor function of the RAR/RXR heterodimer through a conformational change of RAR that causes the dissociation of the corepressor complex and the recruitment of a coactivator complex, which ultimately induces target gene transcription (Gronemeyer et al., 2004; Rhinn and Doll
e, 2012). While the molecular mechanisms of RAR function have been studied extensively (RochetteEgly, 2015; Samarut and Rochette-Egly, 2012; Urvalek and Gudas, 2014; Wei, 2015), the versatile modes of RXR action are less well understood (Ahuja et al., 2003; Evans and Mangelsdorf, 2014; Germain et al., 2002; Lefebvre et al., 2010). In mammals, three RAR (α, β, and γ) and three RXR (α, β, and γ) genes have been identified, which show diverse expression patterns as well as divergent functions and, due to alternative splicing of RAR and RXR gene transcripts, are believed to associate into more than 30 different RAR/RXR heterodimers that transduce signals in the presence of all-trans RA (Gutierrez-Mazariegos et al., 2014b; Mark et al., 2009; Samarut and Rochette-Egly, 2012; Urvalek et al., 2014). The highly gene-specific transcriptional responses to RAR-mediated RA signals are further based upon a plethora of coregulators as well as on the placement or removal of epigenetic marks on histones or on DNA (Gudas, 2013; Urvalek and Gudas, 2014; Wei, 2015). Furthermore, RA can activate RAR/RXR heterodimers independently of DNA binding, as part of nongenomic signaling cascades, or bind and

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activate a number of other NRs, such as testicular receptor 4 (TR4), chicken ovalbumin upstream promoter transcription factor 2 (COUP-TFII) or RAR-related orphan receptor β (RORβ) (Carvalho and Schubert, 2013). Apart from CRABP, intracellular RA can also be bound by fatty acidbinding protein type 5 (FABP5), which shuttles it to another NR, the peroxisome proliferator-activated receptor β/δ (PPARβ/δ), that transduces a different set of RA signaling functions (Berry and Noy, 2009; Schug et al., 2007). For instance, it was shown in mice that RA-induced neuronal differentiation is first mediated through RAR and subsequently through PPARβ/δ activity and that this switch in RA signaling is accomplished through changes in the CRABP-II/FABP5 protein ratio (Yu et al., 2012). In addition, RA plays an important role in the transcriptionindependent activation of the PI3K/Akt (phosphatidylinositol 3-kinase/ v-akt murine thymoma viral oncogene homolog) signaling pathway, which is involved in cell cycle regulation (Garcı´a-Regalado et al., 2013; Lee et al., 2014; Zhang et al., 2015b). RA can also convey developmental signals by modulating the activity of various types of other proteins, such as protein kinase C α (PKCα), cAMP response element-binding protein (CREB), mitogen-activated protein kinase 1/2 (MEK1/2), extracellular signalregulated kinases 1/2 (ERK1/2), or ribosomal s6 kinase (RSK) (Carvalho and Schubert, 2013). The result is a complex cross talk between the genomic and nongenomic effects of RA. For example, by RARs present in cell membrane lipid rafts RA induces kinase cascades, which phosphorylate several targets in the nucleus, including RARs and their coregulators that thus induce the transcription of specific sets of target genes (Carvalho and Schubert, 2013; Rochette-Egly, 2015). Taken together, RA signals exert their biological functions through multiple interacting molecular cascades that can produce finely tuned transcriptional responses in a cell typespecific manner.

5. MAJOR FUNCTIONS OF RA SIGNALING In vertebrates, RA signaling is involved in a great variety of biological processes, during embryogenesis (Fig. 2) as well as in the adult. For example, RA signaling functions are required for initiating and driving meiosis, which leads to the generation of haploid germ cells (Griswold et al., 2012; Koubova et al., 2014; Yu et al., 2013). In addition, RA signals have been implicated in germ cell maturation, survival, and oocyte competence (Best et al., 2015; Endo et al., 2015; Ikami et al., 2015; Paik et al., 2014; Pauli et al., 2013). However, after fertilization, exposure of the developing embryo to

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Fig. 2 Overview of vertebrate tissues and organ systems that require retinoic acid (RA) signaling for their proper development. Modified from Carvalho, J.E., Schubert, M., 2013. Retinoic acid: metabolism, developmental functions and evolution. In: Dakshinamurti, S. (Ed.), Vitamin-Binding Proteins: Functional Consequences. CRC Press, Taylor & Francis Group, Boca Raton.

exogenous RA is known to cause various teratogenic effects (Collins and Mao, 1999). Receptors for RA are maternally expressed and it has been shown that the relative proportions of RAR and RXR subtype transcripts change dramatically during the first 24 h of zebrafish development, coinciding with the transition from maternal to zygotic mRNAs (Blumberg et al., 1992; Oliveira et al., 2013). In response to treatments with exogenous RA, this early RAR/RXR signaling system was further found to upregulate enzymes for the degradation and downregulate enzymes for the synthesis of RA, suggesting a role in protecting the embryo against RA teratology (Oliveira et al., 2013). Nevertheless, exogenous RA still triggers an advancement of the zebrafish developmental program by precociously activating genes that normally remain silent until later developmental stages and by concomitantly inhibiting genes associated with earlier developmental stages (Oliveira et al., 2013).

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Under normal conditions, RA synthesis commences at gastrula stages in trunk paraxial mesoderm and lateral plate mesoderm (Dobbs-McAuliffe et al., 2004; Duester, 2008; Niederreither et al., 1999; Swindell et al., 1999) and creates a linear two-tailed source–sink gradient of the morphogen, which is restricted anteriorly and posteriorly by the activity of RA degrading enzymes (Shimozono et al., 2013). These variable concentrations of RA are essential for axial patterning of embryonic tissues. The neural plate, for instance, is initially anterior-like and increasing levels of RA signaling are required in order to specify the more posterior domains of the hindbrain and spinal cord (Blumberg et al., 1997; Cai et al., 2012; Dup
e and Lumsden, 2001; Grandel et al., 2002; Lara-Ramı´rez et al., 2013). Similarly, graded RA signals provide inductive positional cues in the foregut endoderm and trunk mesoderm that contribute to the correct specification and development of their derivatives, including the posterior pharyngeal arches (Kopinke et al., 2006), lung (Chen et al., 2010), pancreas (Huang et al., 2014; Zhang et al., 2013), somites (Cunningham et al., 2015; Moreno and Kintner, 2004; Retnoaji et al., 2014), kidney (Le Bouffant et al., 2012; Takayama et al., 2014), heart (Zaffran and Niederreither, 2015; Zaffran et al., 2014), and limbs (Cunningham and Duester, 2015; Monaghan and Maden, 2012). RA signaling has also been implicated in skeletogenesis and liver morphogenesis, but its precise roles are still poorly understood (Green et al., 2016; Huang et al., 2009; Ijpenberg et al., 2007; Spoorendonk et al., 2008). Furthermore, RA signaling affects dorsoventral patterning of the neural tube (Pera et al., 2014; Wilson et al., 2004), heart and visceral organ laterality, left–right symmetry during somite formation (Samarut et al., 2015; Vermot and Pourqui
e, 2005), and the termination of body axis extension (Denans et al., 2015; Lara-Ramı´rez et al., 2013). Apart from specifying patterns of cellular identities, RA signals are also important mediators of cell cycle behavior and can, for instance, determine the precise timing of stem cell differentiation by generating heritable epigenetic changes in gene expression, which modulate the ability of stem cells to respond to both retinoid and other signaling pathways (Gudas, 2013; Ikami et al., 2015; Paschaki et al., 2013a; Rochette-Egly, 2015; Urvalek and Gudas, 2014). Moreover, RA has been shown to have context-dependent pleiotropic effects on cell survival (Schug et al., 2007). These functions are not only crucial for coordinating embryogenesis, but also for the regeneration and maintenance of adult tissues, during which RA signals further contribute to cell positional memory, by establishing combinatorial Hox gene expression (Blum and Begemann, 2015; Gudas, 2012; Ikami et al., 2015;

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Lepp and Carlone, 2015; Monaghan and Maden, 2012; Paschaki et al., 2013a). Accordingly, RA signaling fulfills several key roles in the adult, such as epithelial and immune homeostasis (Chanchevalap et al., 2004; Choi and Fuchs, 1990; Raverdeau and Mills, 2014; Sun et al., 2007), adult neurogenesis (Bonnet et al., 2008; Jacobs et al., 2006; Paschaki et al., 2013a), and spermatogenesis (Ikami et al., 2015).

6. RA SIGNALING DURING NERVOUS SYSTEM DEVELOPMENT During vertebrate nervous system development, the RA signaling pathway is involved in the regulation of a multitude of different biological processes including rostrocaudal regionalization (Das et al., 2014; Paschaki et al., 2013b), neural crest induction (Knecht and Bronner-Fraser, 2002; Villanueva et al., 2002), the specification of neural progenitors, and the coordination of neuronal maturation (Janesick et al., 2015). In addition, there is ample evidence that RA signals are important for the differentiation of certain neurotransmitter phenotypes, such as dopaminergic cells (Jeong et al., 2006; Samad et al., 1997; Valdenaire et al., 1998), norepinephrinergic cells (Dupin and Le Douarin, 1995a; Holzschuh et al., 2003; Matsuoka et al., 1997a), cholinergic cells (Berrard et al., 1993; Kim et al., 2001; Nilbratt et al., 2007), serotonergic cells (O’Reilly et al., 2007; Ruiz i Altaba and Jessell, 1991), neuropeptide Y expressing cells (Magni et al., 2000; Mannon and Kaiser, 1997), and GABAergic cells (Chatzi et al., 2011). Moreover, RA signaling has been implicated in synaptic transmission and plasticity, adult neurogenesis, and several neuronal disorders (Bonnet et al., 2008; Bremner et al., 2012; Chen et al., 2014; Jacobs et al., 2006; Paschaki et al., 2013a).

6.1 Posteriorizing Neural Plate According to the activation–transformation model of Nieuwkoop, neural tissue is first specified as anterior neuroectoderm and requires posteriorizing signals that impose posterior identities in a dose-dependent manner (Nieuwkoop, 1952). First experiments using exogenous RA treatments as well as injections of constitutively active and dominant negative RARs into African clawed frog embryos have revealed an important role for RA signaling in patterning the neuroectoderm along its anterior–posterior axis (Blumberg et al., 1997; Durston et al., 1989; Sive and Cheng, 1991). Initial RA production, however, depends on the suppression of anterior genes by early signals that already act at late blastula stages, when the neural plate is still

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forming, such as WNT and FGF factors (Kudoh et al., 2002; Pera et al., 2014). WNTs are secreted by nonaxial mesoderm progenitors at the posterior neural plate (Bang et al., 1999; Green et al., 2014). From there, they diffuse anteriorly, generating a linear caudal-to-rostral WNT gradient that induces dose-dependent gene expression within the neural plate area, which ultimately leads to the segmentation of forebrain, midbrain, and hindbrain primordia (Bang et al., 1999; Green et al., 2014). The major function of WNT signaling during this process is probably to counter TCF3dependent transcriptional repression, which is essential for vertebrate head formation (Dorsky et al., 2003; Green et al., 2014; Kim et al., 2000). In addition, FGFs are expressed in a similar caudal-to-rostral gradient during neural plate posteriorization and several studies have revealed permissive interactions and interdependences between these two signaling pathways, which probably reflect alternative modes of signal integration that have yet to be elucidated (del Corral and Morales, 2014; Green et al., 2014; Pera et al., 2014). The RA synthesizing enzyme RALDH2 is first expressed during early gastrula stages in the paraxial mesoderm of various vertebrate embryos, including zebrafish, African clawed frog, chicken, and mouse (Grandel et al., 2002). In the caudal epiblast (tail end), high levels of FGFs antagonize RA signaling (Fig. 3A) by promoting cyp26 and attenuating raldh2 expression (del Corral and Morales, 2014; Green et al., 2014; Pera et al., 2014). This is necessary for maintaining a pool of axial stem cells that later on give rise to the spinal cord and somites (Green et al., 2014; Pera et al., 2014). However, FGFs also stimulate WNT signaling, which promotes raldh2 expression in the paraxial mesoderm along the prospective trunk and neck region (del Corral and Morales, 2014; Green et al., 2014; Pera et al., 2014). In the rostral ectoderm, cyp26 continues to be expressed, due to low levels or the complete absence of WNT and FGF signals (Kudoh et al., 2002). This protection of the anterior domain against the posteriorizing activities of RA signals is fundamental for enabling forebrain and midbrain development (Kudoh et al., 2002). Accordingly, the combinatorial effects of FGF and WNT signals are required for suppressing anterior fates and for modulating RA signaling, which has the distinct role of activating the expression of early posterior genes, such as homeodomain proteins (Grandel et al., 2002; Kudoh et al., 2002). For instance, work on zebrafish embryos with a mutated raldh2 gene suggested that the loss of RA signaling most severely affects the general expansion of the hindbrain as well as hox gene expression patterns prior to somitogenesis (Grandel et al., 2002). Therefore, RA signaling acts already on

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Fig. 3 Networks of interacting signals that, in vertebrates, control (A) the anterior– posterior regionalization of the neural plate, (B) the anterior–posterior patterning of the hindbrain and spinal cord domains of the neural tube, and (C) the dorsal–ventral patterning of the spinal cord domain. Abbreviations: BMP, bone morphogenetic protein; CDX, caudal-related homeobox protein; CER, cerberus protein; DI1–DI6, dorsal interneurons 1–6; DKK, dickkopf protein; FB, forebrain; FGF, fibroblast growth factor; FP, floor plate; FRZB, frizzled-related protein; HB, hindbrain; HOX, homeobox protein; IGF, insulin-like growth factor; MB, midbrain; MN, motorneurons; RA, retinoic acid; RP, roof plate; SC, spinal cord; SHH, Sonic hedgehog; V0–V3, ventral interneurons 0–3; WNT, wingless-related integration site protein.

the developing neural plate to establish the gross regionalization of the central nervous system (CNS).

6.2 Setting Up RA Signaling System and Boundaries of Neural Tube Suppression of WNT and FGF signals in the anterior neural plate by various factors secreted from the pharyngeal endomesoderm, e.g., Cerberus,

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Dickkopf, FRZB, and IGF, allows the expression of a cascade of transcription factors, including otx, gbx, pax2/5/8, and en1/2, which establish the forebrain–midbrain and the midbrain–hindbrain boundaries (Dworkin and Jane, 2013; Green et al., 2014; Nakamura et al., 2005; Pera et al., 2001). In the developing neural tube, the midbrain–hindbrain boundary (MHB) acts as an important signaling center, also known as the isthmic organizer, which produces an anterior-high WNT and FGF signaling gradient throughout the midbrain and anterior hindbrain territory that mirrors signaling from the caudal end of the embryo (Dworkin and Jane, 2013; Nakamura et al., 2005; Pera et al., 2001). At this point in development, RALDH2 is present in the paraxial/ somitic mesoderm along the presumptive spinal cord domain, up to and including the first somite (Glover et al., 2006). Accordingly, the source of endogenous RA is located at the caudal limit of the presumptive hindbrain domain (Glover et al., 2006). From here, RA diffuses anteriorly, generating a posterior-high concentration gradient across the hindbrain territory that is patterned by localized CYP26 expression domains and opposed by the anterior-high WNT and FGF gradients emanating from the MHB (Fig. 3B) (Cunningham and Duester, 2015; Glover et al., 2006; Hernandez et al., 2007; Shimozono et al., 2013; White and Schilling, 2008). The resulting interactions of RA, WNT, and FGF signaling with CDX, LIM, and TGIF transcription factors as well as their modulation of CYP26 activity in this area define the boundary between the presumptive hindbrain and spinal cord domains (Castillo et al., 2010; Chang et al., 2016a; Cunningham and Duester, 2015; Lara-Ramı´rez et al., 2013). Among other things, this leads to the downregulation of cyp26 expression in the anterior spinal cord territory and to the onset of restricted raldh2 expression within specific areas of the spinal cord territory, including the roof plate, the ventrolateral motor neuron domains, and, transiently, dorsal interneurons (Castillo et al., 2010; Cunningham and Duester, 2015; Lara-Ramı´rez et al., 2013; Wilson et al., 2004). As in the hindbrain, RA signaling in the developing spinal cord is restricted through region-specific CYP26 expression domains and through an opposing posterior-high FGF gradient produced by the tail bud (Cunningham and Duester, 2015; Lara-Ramı´rez et al., 2013). In summary, a complex source and sink system of RA signals is created in the prospective hindbrain and spinal cord during neurulation that is subsequently required to regulate multiple neurogenic processes.

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6.3 Patterning Neural Tube Along the Anterior–Posterior Axis 6.3.1 Hindbrain and Anterior Spinal Cord As described earlier, the combinatorial actions of WNTs, FGFs, CYP26 enzymes, and other factors restrict RA signaling to somitic levels, where it inhibits both FGF and WNT signaling, in order to drive somitogenesis, neural fate specification, and neuronal differentiation (Fig. 3B) (del Corral and Morales, 2014; Green et al., 2014; Pera et al., 2014). Diffusion and localized degradation of the lipophilic RA molecule results in a linear two-tailed morphogen gradient, based on which the different cellular identities along the anterior–posterior axis of the developing neural tube are patterned (Blumberg et al., 1997; Cai et al., 2012; Dup
e and Lumsden, 2001; Grandel et al., 2002; Lara-Ramı´rez et al., 2013; Shimozono et al., 2013). Vertebrate hindbrain development is characterized by the generation of transient, lineage-restricted compartments, called rhombomeres, that can be identified through their expression of distinct sets of genes as well as through differences in their neuronal organization (Clarke and Lumsden, 1993; Lumsden and Krumlauf, 1996; Vaage, 1969). It has been demonstrated in several animal models, e.g., in lampreys (Murakami, 2004), chicken (Dup
e and Lumsden, 2001), and rats (White et al., 2000), that more posterior rhombomeres require progressively higher amounts of RA for their proper specification. This RA-induced patterning is mainly achieved through the regulation of nested, region-specific expression patterns of homeobox genes, which produce a combinatorial “hox-code” capable of assigning distinct positional identities (Glover et al., 2006; Kessel and Gruss, 1991). Typically, vertebrate hox genes are organized into chromosomal clusters that are characterized by temporal and spatial colinearity, which means that a gene located in a more 30 position within its cluster generally becomes activated earlier and has a more anterior expression limit than the gene in the next more 50 position (Duboule and Doll
e, 1989; Gaunt et al., 1989; Izpisu´a-Belmonte et al., 1991). Several hox genes are directly regulated by RA signaling through RAREs in their regulatory sequences and, intriguingly, hox gene colinearity appears to be linked to RA signaling, since genes at the 30 end of a hox complex respond more rapidly to RA signals than genes positioned in more 50 positions (Alexander et al., 2009; Oosterveen et al., 2003; Simeone et al., 1990, 1991; Zhang et al., 2000). Moreover, 30 hox genes tend to become activated in response to RA signals, while 50 hox are more likely to be repressed (Liu et al., 2001). These differences might

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well be associated with the ability of RA signaling to remove repressive modifications from the chromatin domains of the 30 hox genes hox1–hox5 (Mazzoni et al., 2013). Generally, the developing hindbrain expresses hox1–hox5 paralogs, while hox4–hox11 are present in the spinal cord territory (Philippidou and Dasen, 2013). However, this expression is spatially and temporally dynamic, with some hox genes being expressed over a narrow time window during early development and others persisting into postnatal stages (Philippidou and Dasen, 2013). Thus, not only the concentration but also the timing of RA exposure is an important factor in hox gene regulation, which further requires interactions with several competing signals. Simply put, FGFs alone promote a midbrain character, FGFs plus WNTs a rostral hindbrain character and RA plus WNTs a caudal hindbrain and anterior spinal cord character (del Corral and Morales, 2014). Initially, RA signals directly activate hoxa1 and hoxb1 expression throughout the hindbrain and toward the midbrain, but induction of cyp26 by FGF signaling from the anterior midbrain–hindbrain region subsequently limits this expression to the prospective boundary between rhombomeres 4 and 5 (Duester, 2008; Sirbu et al., 2005). FGF8 from the MHB defines the extension of rhombomere 1 by suppressing hox gene expression, while RA signals activate the transcription factor VHNF1 in the posterior hindbrain (Alexander et al., 2009; Terriente and Pujades, 2015). FGF3 and FGF8, which are released from the area of the rhombomeres 3–5, cooperate with VHNF1 to regulate the expression of krox20 in rhombomere 5 and of mafb/kreisler in the rhombomeres 5 and 6 (Alexander et al., 2009; Terriente and Pujades, 2015). The three cisregulatory elements of krox20 integrate inputs from RA, WNT, FGF, and several transcription factors to specifically induce transcription in the rhombomeres 3 and 5, where krox20 is needed for repressing hoxb1 by binding its activator PIASxβ (Alexander et al., 2009). Interestingly, this repression also requires RA signaling via a RARE in the hoxb1 repressor (Studer et al., 1994). In contrast, levels of hoxb1 expression in rhombomere 4 are stimulated by HOXA1 as well as by HOXB1 itself, via an upstream HOX response element and in conjunction with the cofactors PBX and MEIS (Moens and Selleri, 2006; Tvrdik and Capecchi, 2006). In order to prevent disruptive fluctuations in RA availability, HOXA1-PBX-MEIS also binds directly to an enhancer that is required for maintaining normal raldh2 expression

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levels in the mesoderm (Vitobello et al., 2011). In addition, CRABPs have been shown to provide a key feedback mechanism, which compensates for changes in RA production and thus contributes to signaling robustness during hindbrain patterning (Cai et al., 2012). In the region of rhombomere 6, kreisler-dependent induction of hoxa3 and hoxb3 represses hoxb1 through its HOX response element (Wong et al., 2011). As the somites differentiate, they cease RA production and thus stimulation of hoxa1 and hoxb1 expression fades throughout the hindbrain region (Alexander et al., 2009; Glover et al., 2006). Since hoxa1 does not possess an autoregulatory element, its expression rapidly declines, whereas HOXB1 maintains its own expression in rhombomere 4 and activates hoxa2 and hoxb2 expression, which provide positive feedback to hoxb1 expression (Studer et al., 1998; T€ umpel et al., 2007; Tvrdik and Capecchi, 2006). Multiple feedback loops and the gradual retreat of raldh2 expression from anterior maturing somites cause shifting differences in RA, WNT, and FGF signaling levels along the developing hindbrain and anterior spinal cord, which, together with the combinatorial activities of various transcription factors and the direct auto- and cross-regulatory interactions between the hox genes themselves, required for the generation of nested hox gene expression domains (Glover et al., 2006; Sheth et al., 2014; Terriente and Pujades, 2015). Moreover, recent studies on lampreys show that this hox regulatory network is largely conserved to the base of vertebrates (Parker et al., 2014). 6.3.2 Spinal Cord The posterior part of the nervous system has a separate developmental origin than the anterior part and thus the majority of spinal cord progenitors receive different signals and undergo a different transcriptional program than the cells contributing to the fore-, mid-, and hindbrain territories (Gouti et al., 2015). Most of the cells that make up the developing spinal cord originate from a pool of axial stem cells, which is maintained by the antidifferentiation effects of FGF signaling at the caudal epiblast (del Corral and Morales, 2014; Gouti et al., 2015; Pera et al., 2014). These stem cells give rise to neuromesodermal progenitors that express both mesoderminducing and neural transcription factors, including Brachyury (BRA), SOX2, SOX3, NKX1-2, and CDX (Gouti et al., 2015). Accordingly, neuromesodermal progenitors can become either presomitic mesoderm precursors, expressing MSGN1 and TBX6, or preneural tube cells expressing NKX1-2 and SOX2 (Gouti et al., 2015). However, positive feedback between CDX/BRA and WNT/FGF signaling from the caudal epiblast

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is thought to maintain their neuromesodermal progenitor state (Fig. 3B) (Gouti et al., 2015). As cells migrate out from the caudal lateral epiblast, they are exposed to decreasing levels of WNT and FGF signaling, but increasing levels of RA signals, which emanate from the somites that are differentiating in an anterior-to-posterior wave along the forming CNS (Gouti et al., 2015; Lara-Ramı´rez et al., 2013). Owing to their ability to attenuate WNT and FGF signaling and to stimulate the expression of hox genes, RA signals have been implicated in inducing the developmental progression of neuromesodermal progenitors (Gouti et al., 2015). From in vitro experimentation, it has been proposed that a prolonged exposure to WNT and FGF signaling vs RA signaling biases neuromesodermal progenitors to adopt a mesodermal rather than a neural fate (Gouti et al., 2014, 2015). This idea is supported by the fact that mutant mice lacking a functional raldh2 gene for RA synthesis exhibit an expansion of mesodermal markers at the expense of markers of the prospective posterior neuroepithelium (Ribes et al., 2009). However, the molecular mechanisms underlying the neural–mesoderm switch at the caudal epiblast remain to be characterized. Cells assuming a preneural tube fate join the caudal end of the neural plate, which borders at the posterior-most portion of the developing neural tube, and follow the morphogenetic movements involved in neural tube closure (Gouti et al., 2015). Upon entering the spinal cord, preneural tube cells are instructed by increasing levels of RA signaling to transition to a neural progenitor fate, which is marked by a downregulation of NKX1-2 and an upregulation of PAX6 and IRX3 (Gouti et al., 2015). In addition, RA signals have been shown to stimulate neural progenitor proliferation within the forming spinal cord, which initially consists of a single layer of pseudostratified epithelium that expands rapidly so that the subsequent differentiation of neural cells can take place within the lateral mantle layer, while undifferentiated progenitors remain in the ventricular layer that lines the lumen of the neural canal (del Corral and Morales, 2014; England et al., 2011; Mich and Chen, 2011; Wilson et al., 2003). This function of RA signaling is believed to be particularly important at the brachial and lumbar levels of the spinal cord, to increase the number of progenitors within the prospective lateral motor columns, from where the limbs will be innervated (del Corral and Morales, 2014; Sockanathan and Jessell, 1998). These lateral motor columns are absent from other thoracic levels of the spinal cord, where instead two distinct columns of lateral motor neurons are generated: the preganglionic motor column contains

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visceral motor neurons, which innervate the sympathetic ganglia and glandular targets, and the hypaxial motor column contains motor neurons, which innervate the musculature of the body wall (Philippidou and Dasen, 2013). In addition, a group of mammal-specific phrenic motor neurons is formed at rostral cervical levels to supply the diaphragm muscle and a median motor column, comprising motor neurons that project to the axial muscles, spans across the entire length of the spinal cord (Philippidou and Dasen, 2013). While interneuron subclasses are typically generated along the full extent of the spinal cord, the expression of distinct of LIM transcription factors and HOX proteins has been associated with the segregation of motor neurons into different columns as well as into different pools within these columns, each of which exhibiting a separate identity and axonal target (Dasen et al., 2003, 2005, 2008; Kania and Jessell, 2003; Landmesser, 2001; Liu et al., 2001). As in the hindbrain, the hox genes of the spinal cord display nested, colinear expression patterns (Philippidou and Dasen, 2013). RA signaling is required to induce the expression of hox5 at cervical levels and of nolz1 at brachial levels (Ji et al., 2009; Lara-Ramı´rez et al., 2013; Philippidou and Dasen, 2013). The transcription factor NOLZ1 in turn activates hoxc6 at the forelimb level, which is a postmitotic determinant of the brachial lateral motor column (Ji et al., 2009; Lara-Ramı´rez et al., 2013). A particular importance of RA for the specification of the lateral motor columns is reflected by the strong expression of raldh2 in brachial and lumbar, but not thoracic, motor neuron domains (Lara-Ramı´rez et al., 2013). Loss of this raldh2 expression leads to the formation of reduced and disorganized lateral motor columns, while the retroviral mis-expression of raldh2 at thoracic levels was shown to cause an ectopic induction of lateral motor column identities, as marked by the expression of typical transcription factors, such as isl1, isl2, and lim1 (Lara-Ramı´rez et al., 2013; Paschaki et al., 2013b; Sockanathan and Jessell, 1998; Vermot et al., 2005). Initially, ISL1 is present in all lateral motor column neurons, but RA signals subsequently restrict isl1 expression to more medial areas, while instead promoting lim1 expression, thus favoring lateral rather than medial motor neuron fates (Ji et al., 2006, 2009; Paschaki et al., 2012; Sockanathan and Jessell, 1998). This is relevant for establishing the specific topography of the lateral motor column, since LIM homeodomain proteins control EphA receptors and ephrin-A ligands that coordinate cell body positioning and axonal trajectory development (Kania and Jessell, 2003).

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Conversely, RA appears to be dispensable for the induction of more posterior spinal cord hox genes, which instead rely on other graded caudal signals, including WNTs, FGFs, and TGFβ proteins, such as GDF11 (del Corral and Morales, 2014; Liu et al., 2001; Philippidou and Dasen, 2013). However, RA signals are needed for the refinement of the anterior borders of hox gene expression domains along the spinal cord, a process during which they exert rostralizing rather than caudalizing influences (del Corral and Morales, 2014; Liu et al., 2001; Philippidou and Dasen, 2013). This strictly context-dependent manner of RA signaling action is probably achieved through the modulation of neural progenitor responsiveness to WNT/FGF vs RA signals by CDX transcription factors. CDX removes repressive chromatin modifications from posterior 50 hox genes in order to enable their transcriptional activation via FGF/WNT signaling in the caudal spinal cord (Bel-Vialar et al., 2002; Mazzoni et al., 2013; Philippidou and Dasen, 2013). In contrast, RA signaling modifies the chromatin domains of anterior 30 hox genes, in order to induce their expression within the hindbrain (Mazzoni et al., 2013; Philippidou and Dasen, 2013). Interestingly, RA signaling is able to exclude CDX from the prospective hindbrain, but upregulates it in the spinal cord, where cdx expression is further induced and maintained by caudal WNT and FGF signals (Chang et al., 2016a; Houle et al., 2000, 2003; Lee and Skromne, 2014; Philippidou and Dasen, 2013; Skromne et al., 2007). Moreover, CDX appears to stimulate the RA signaling pathway in the hindbrain, but antagonizes it in the posterior spinal cord. For instance, blocking CDX causes a posterior shift of mesodermal raldh2 expression, which creates an overlap of high FGF and RA signaling levels within caudal neural tissues that leads to the ectopic formation of hindbrain and anterior spinal cord structures in the posterior-most neural tube territory (Shimizu et al., 2006; Skromne et al., 2007). Conversely, CDX represses cyp26 expression in the prospective hindbrain domain, allowing for the proper positioning of the boundary between the hindbrain and spinal cord territories (Chang et al., 2016a). Accordingly, RA signals and CDX trigger the activation of hox genes from opposite sites of the cluster and seem to cross-regulate each other in an opposite manner in the hindbrain and in the posterior spinal cord. This might be due to the fact that the FGF/WNT signaling center at the MHB arises much later in development than the one at the caudal epiblast, so that a robust positive feedback loop between FGF/WNT and CDX cannot be established, before high levels of RA

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signals clear CDX from the hindbrain. Alternatively, early activities of RA signaling at the future hindbrain territory of the neural plate, such as the induction of early genes like hox1 and meis3 in zebrafish (Kudoh et al., 2002), might initiate downstream events that prevent positive feedback interactions between FGF/WNT signaling and CDX. In any case, this mirrored use of the same interacting signaling gradients for the formation of two very different parts of the nervous system provides a striking example for the multifunctionality of developmental signaling pathways and highlights the importance of their dynamic integration into a specific temporal and spatial context.

6.4 Patterning Neural Tube Along Dorsal–Ventral Axis The embryonic dorsal–ventral axis is initially defined by a signaling gradient of BMPs, which are inhibited dorsally by secreted factors such as Chordin, Noggin, and Follistatin (Langdon and Mullins, 2011). Low BMP signaling levels allow neuroectodermal tissues to develop from the dorsal ectoderm, while moderate and high BMP levels specify lateral and ventral cell types, respectively (Langdon and Mullins, 2011). In gastrulating embryos of the African clawed frog, RA is synthesized in the dorsal mesoderm and degraded in the adjacent dorsal ectoderm and ventral mesoderm (Pera et al., 2014). Morpholino oligonucleotide-mediated inhibition of RA synthesis in these embryos causes a ventralized phenotype, indicating that RA signaling exerts dorsalizing functions during early embryogenesis (Pera et al., 2014; Strate et al., 2009). Indeed, reduced levels of RA signaling were shown to downregulate the dorsalizing BMPantagonist Chordin, but upregulate ADMP (antidorsalizing morphogenetic protein), an activator of the ventralizing BMP signaling pathway, whereas an increase in RA signaling levels had the opposite effect (Pera et al., 2014; Strate et al., 2009). Thus, early interactions of RA signaling with the BMP pathway likely contribute to setting up the dorsal–ventral axis in embryos of the African clawed frog. As neurulation proceeds, previously lateral epidermis comes to lie dorsally above the closing neural tube (Liu and Niswander, 2005). Paracrine secretion of TGFβ proteins (e.g., BMPs, Dorsalin, Activin) from this new dorsal epidermis specifies the dorsal roof plate of the neural tube, while Sonic hedgehog protein (SHH) from the underlying notochord induces the ventral floor plate (Liu and Niswander, 2005). Subsequently, the dorsal roof plate cells induce a cascade of dorsalizing TGFβ protein signals throughout

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the dorsal neural tube, whereas the ventral floor plate acts as a ventralizing SHH secreting signaling center (Liu and Niswander, 2005). As a result, two opposing signaling gradients are established that provide positional cues to the cells found along the dorsal–ventral axis of the neural tube (Fig. 3C) (Liu and Niswander, 2005). Nevertheless, in order for early dorsal–ventral patterning and neural differentiation to occur, high levels of RA signaling from the paraxial mesoderm first have to extinguish FGF signaling from somitic levels and thus terminate the FGF-mediated repression of homeodomain-containing transcription factors (del Corral and Morales, 2014; Mazzoni et al., 2013; Novitch et al., 2003; Patel et al., 2013). Accordingly, RA signals temporally coordinate dorsal–ventral with anterior– posterior patterning (Hashiguchi and Mullins, 2013). Moreover, a decrease of RA signaling in the developing hindbrain and spinal cord has been shown to cause a dorsal expansion of ventral patterning genes and progenitor domains, whereas an increase in RA signaling triggers a rapid ventral shift of both (Glover et al., 2006; Wilson et al., 2004). Importantly, these effects can be observed during early as well as during late stages of neurogenesis, suggesting that RA signals not only contribute to establishing the dorsal– ventral axis, but also actively maintain it throughout a protracted period of CNS development (Glover et al., 2006; Wilson et al., 2004). The actions and interactions of BMP/SHH, WNT/FGF, and RA signaling with CDX, LIM, and TGIF transcription factors are necessary for the downregulation of cyp26 expression in the anterior spinal cord territory as well as for the generation of restricted domains of raldh2 expression in the roof plate, in the ventrolateral motor neuron domains, and, transiently, in dorsal interneurons throughout the developing spinal cord (Castillo et al., 2010; Cunningham and Duester, 2015; Lara-Ramı´rez et al., 2013; Wilson et al., 2004). Conversely, transcriptomic analyses of mutant mice with impaired RA signaling and studies on retinoid-free VAD (vitamin A deficient) quails have revealed a mis-regulation of several components of the BMP signaling pathway and of other dorsal-specific factors, including PAX3, PAX7, MSX2, and WNTs (Martinez-Morales et al., 2011; Paschaki et al., 2013b; Ribes et al., 2009; Wilson et al., 2004). In particular, MSX2 and WNTs are activated in a rostral-to-caudal progression in normal embryos, but depleted in VAD quails, indicating that their induction might depend on somite-derived RA (del Corral and Morales, 2014). Furthermore, SHH signaling appears to be reduced by both the blocking and the overactivation of the RA signaling pathway (del Corral and Morales, 2014). These results suggest that RA signaling exerts its dorsalizing effects,

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at least in part, through cross talk with the signaling gradients present along the dorsal–ventral axis of the neural tube. From dorsal to ventral, the spinal cord comprises the dorsal roof plate cells, the dorsal interneurons DI1, DI2, DI3, DI4, DI5, and DI6, the ventral interneurons V0, V1, and V2, motor neurons, the ventral interneurons V3, and the ventral floor plate cells, all of which express unique combinations of transcription factors (Catela et al., 2015). Low levels or the absence of RA signaling have been associated with the specification of more ventral interneurons, such as V3, while higher levels of RA signaling are thought to promote more dorsal fates, such as V0, V1, and motor neurons (England et al., 2011; Lara-Ramı´rez et al., 2013; Lupo et al., 2006). A lack of RA signals was thus shown to diminish V0, V1, and V2 interneuron populations and reduce the expression of dbx1 and dbx2 in progenitors of V0 and V1 interneurons as well as of nkx6.2 and irx3 in progenitors of V1 and V2 interneurons (del Corral et al., 2003; Novitch et al., 2003; Pierani et al., 1999; Wilson et al., 2004). Multiple roles of RA signaling have further been described during the specification of motor neurons, which become dramatically decreased under RA-deprived conditions (Novitch et al., 2003; Wilson et al., 2004). Emanating from the paraxial mesoderm, RA signals promote class I homeodomain gene expression (e.g., pax7, pax6, dbx1/2, and irx3), while coincident SHH signaling from the notochord and floor plate counteracts RA signaling activity and induces class II homeodomain gene expression (e.g., nkx6 and nkx2) (Novitch et al., 2003). Class I and class II homeodomain proteins exhibit mutual cross-repressive interactions and define individual progenitor domains in a combinatorial fashion by regulating the expression of neuronal subtype determinants (Novitch et al., 2003). Thus, the motor neuron lineage becomes specified by PAX6 (class I homeodomain) and NKX6 (class II homeodomain) proteins, which downregulate repressors of OLIG2, a known determinant of motor neuron and oligodendrocyte fates, while RA signaling mediates the transcriptional activation of the olig2 gene (Novitch et al., 2003). The requirement of RA signaling for pax6 and olig2 expression seems to be a general feature of vertebrates, as it has been reported in zebrafish, quail, and mice (del Corral et al., 2003; England et al., 2011; Paschaki et al., 2013b; Ribes et al., 2009; Wilson et al., 2004). In addition, RA signaling also acts downstream of olig2 expression to induce pan-neuronal differentiation genes and homeodomain protein effectors involved in motor neuron maturation, such as HB9 (Lee et al., 2009; Novitch et al., 2003).

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Finally, even in the ventral-most progenitors of spinal cord V3 interneurons, RA signals are needed for the stimulation of Notch receptor activity to decrease, but not eliminate, ascl1 expression, thus favoring V3 interneuron identity over serotonin-expressing neurons, which are characteristic of hindbrain levels (Jacob et al., 2013). Accordingly, multiple sources of RA within the spinal cord and from the somitic mesoderm, together with restricted domains of degradation by CYP26, provide crucial cues for the specification of different neural subtypes throughout the neural tube. While some of these effects might be the result of a direct regulation of ventral patterning genes by RA signaling, it is likely that a large part of RA signaling functions during the dorsal–ventral patterning of the spinal cord is achieved indirectly, through the modulation of SOXB1 transcription factors and other components involved in the cross-regulatory networks of ventral patterning genes (del Corral and Morales, 2014; Ribes et al., 2009).

6.5 Orchestrating Proliferation, Differentiation, and Survival of Neural Progenitors Apart from conferring cellular identities during developmental patterning, RA signaling also plays a central part in the decision between stem cell maintenance and the timing of neural progenitor maturation. In anamniote vertebrates (i.e., in cyclostomes, cartilaginous and bony fish, and amphibians), primary neurogenesis begins within the deep layer of the open neural plate, with superficial cells maintaining their proliferative state (Chalmers et al., 2002). However, these primary neurons are thought to be mostly transient and have probably been lost secondarily in amniotes (i.e., in reptiles, birds, and mammals) as a consequence of their direct development, which does not require an early sensory–motor system to guide simple larval behaviors (Wullimann, 2009; Wullimann et al., 2005). In amniotes, the first neurons appear in the rostral brain and give rise to an early axonal scaffold, which is comparable to the fiber tracts generated by the earliest formed anamniote neurons that arise in the cortex during secondary neurogenesis (Ware et al., 2015). Moreover, initial neurogenesis in the amniote and secondary neurogenesis in the anamniote brain are characterized by a fairly similar deployment of neural transcription factors (e.g., NGN1, NEUROD, MASH1/ZASHLA, and PAX6), LIM and HOX proteins (e.g., LHX6/7, TBR2, and DLX2a) as well as GABA/GAD-positive cells (Wullimann, 2009; Wullimann et al., 2005). It thus seems likely that the developmental mechanisms governing the differentiation of the first cerebral neurons are conserved across vertebrates.

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While it is well established that RA signals promote the differentiation of anamniote primary neurons by inhibiting key proliferation factors, such as zic, geminin, notch, and foxd4l1, as well as by stimulating proneural and neurogenic gene expression, studies addressing the potential involvement of RA signaling in anamniote secondary neurogenesis or amniote primary neurogenesis, are relatively scarce (Janesick et al., 2015). So far, RA signaling has been implicated in the differentiation of mammalian photoreceptors, hippocampus, and cortical neurons (Janesick et al., 2015). For the latter, RA secreted from the meninges, which comprises several membranes ensheathing the brain, appears to be a crucial cue that controls cell cycle exit in the cortex (Siegenthaler and Pleasure, 2011; Siegenthaler et al., 2009). Interestingly, RA signals derived from radial glia cells have recently been shown to drive the formation of the blood–brain barrier, which constitutes the innermost layer of the meninges (Mizee et al., 2013). Astroglia cells are known to produce RA in order to maintain retinoid homeostasis in the brain, and it has further been suggested that astroglia-derived RA might stimulate neurogenesis within the neurogenic niches of the postnatal brain (Haskell and LaMantia, 2005; K€ ornyei et al., 2007; Shearer et al., 2012; Wang et al., 2011). However, these findings remain to be corroborated by in vivo analyses, which should also consider the embryonic brain (Shearer et al., 2012). RA generated by RALDH3 in the subventricular zone of the basal ganglia is required for ventral forebrain development, in particular, for the correct expression of the dopamine autoreceptor D2 and GABAergic differentiation (Chatzi et al., 2011; Molotkova et al., 2007). In addition, the cerebrospinal fluid has been identified as an important source of RA signals, which were shown to promote neuroepithelial cell survival and neurogenesis (Alonso et al., 2011, 2014; Chang et al., 2016b; Parada et al., 2008). RA signals that originate from the paraxial mesoderm and from within the posterior neural tube also drive neurogenesis, once the neural identities of the prospective hindbrain and spinal cord have been established (del Corral et al., 2003; Duester, 2013; Janesick et al., 2015). During this process, RA simultaneously acts as a permissive differentiation signal, by repressing the FGFdependent maintenance of progenitor cell function, and as an instructive signal that induces neuronal differentiation (Duester, 2013). A direct target of RA signaling, which is needed for inducing embryonic stem cells to differentiate into neurons, is hoxa1 (Janesick et al., 2015; Makki and Capecchi, 2011). Furthermore, btg2 is believed to contribute to RA-induced differentiation and was shown to enhance RAR activity by

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modulating histone H4 methylation and acetylation (Janesick et al., 2015). Many other genes that facilitate cell cycle exit or inhibit the proliferation of neural progenitors can be regulated either directly or indirectly by RA signaling, but often require specific developmental conditions (Janesick et al., 2015): highly cell- and gene-specific transcriptional responses to RA are mediated by numerous coregulators, by the placement or removal of epigenetic marks as well as through a number of alternative RAR-independent signaling pathways (Carvalho and Schubert, 2013; Gudas, 2013; Urvalek and Gudas, 2014; Wei, 2015). For instance, RA signals can determine the precise timing of stem cell differentiation by generating heritable epigenetic changes in chromatin responsiveness to both retinoid and other signaling pathways (Gudas, 2013) and exert pleiotropic effects on cell survival and cell cycle, depending on the presence of specific binding proteins (Schug et al., 2007; Wolf, 2008). Accordingly, the blocking of RA signaling was shown to reduce neural progenitor proliferation in the telencephalon of mice (Rajaii et al., 2008) as well as in the neural tube of quail and zebrafish embryos (England et al., 2011; Mich and Chen, 2011; Wilson et al., 2003), while RA treatment increases the number of proliferating cells in chicken ventral neural tube explants (Sockanathan and Jessell, 1998). Intriguingly, RA signals seem to attenuate, but not extinguish, the proliferation of precursor cells in the olfactory epithelium in order to prevent a depletion of the progenitor pool (Paschaki et al., 2013a). A similar mode of action has also been described for a distinct population of slowly dividing precursors in the developing and adult forebrain of mice (Haskell and LaMantia, 2005). Taken together, current evidence indicates that RA signals can modulate cell cycle progression as well as cell cycle speed in a precisely timed and context-dependent manner. However, the involvement of RA signaling in amniote neurogenesis, in particular in the anterior CNS, still remains largely unexplored.

6.6 Inducing and Coordinating Neural Crest Development In the dorsal ectoderm of blastula stage embryos, FGF signals together with antagonists of WNT and BMP signaling activate early neural genes, such as erni, otx2, and sox1/3, which specify the neural plate (Streit et al., 2000). However, this dorsal neural plate territory is surrounded by a zone, where BMPs from ventral and WNTs from posterior–lateral levels of the embryo are only partially inhibited (Groves and LaBonne, 2014). The resulting medial-low, lateral-high gradient of WNT/BMP activity defines the neural

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plate border and, probably in conjunction with FGF, RA, and Notch signaling from underlying tissues, induces the expression of a distinct set of transcription factors, known as neural plate border specifiers (Groves and LaBonne, 2014; Martinez-Morales et al., 2011; Milet and MonsoroBurq, 2012; Yardley and Garcı´a-Castro, 2012). Subsequently, mutual cross-regulatory interactions between these neural plate border specifiers, which include distalless5, pax3, pax7, and zic1/3, as well as between the neural and nonneural transcription factors activated in adjacent tissues stabilize and sharpen the neural plate border domain by suppressing neural as well as epidermal fates (Simo˜es-Costa and Bronner, 2015). Within the neural folds that arise from this neural plate border, a transient population of multipotent migratory cells becomes specified during gastrulation, the so-called neural crest. Neural crest cells (NCCs) give rise to various structures, such as the craniofacial skeleton, the peripheral nervous system (PNS), and melanocytes (Marchant et al., 1998; Weinstein and Hemmati-Brivanlou, 1999; Wilson and Hemmati-Brivanlou, 1997). NCCs are, however, absent from the anterior neural folds, which differentiate into the forebrain, suggesting that the specification of the neural crest requires additional posteriorizing signals (Knecht and Bronner-Fraser, 2002; Villanueva et al., 2002). Exposure to FGF, WNT or RA has been shown to transform the anterior neural folds into neural crest, while blocking of these signals inhibits the expression of neural crest marker genes in the medial and posterior neural folds (Knecht and Bronner-Fraser, 2002; Martinez-Morales et al., 2011; Monsoro-Burq et al., 2003; Villanueva et al., 2002). Thus, the induction of neural crest is restricted to the medial and posterior portions of the neural folds and requires low levels of BMP signaling as well as the posteriorizing activities of FGF, WNT and RA signals (Knecht and Bronner-Fraser, 2002; Martinez-Morales et al., 2011; Monsoro-Burq et al., 2003; Villanueva et al., 2002). WNTs secreted from caudal nonaxial mesoderm progenitors diffuse anteriorly to activate gbx1/2 transcription throughout the posterior domain of the neural plate (Green et al., 2014; Milet and Monsoro-Burq, 2012). Next, mutual antagonism between GBX1/2 and the initially widely expressed neural plate marker OTX2 divides the neural plate at the MHB (Milet and Monsoro-Burq, 2012). In the anterior section, OTX2 enables the expression of the WNT antagonist DKK1, which represses posterior fates within the rostral neural plate border (Fig. 4A). This promotes placode instead of neural crest formation, which is crucial for vertebrate head development (Milet and Monsoro-Burq, 2012). However, the distinction

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Fig. 4 Network of interacting signals that, in vertebrates, act along the developing central nervous system (CNS) to govern neural crest and cranial placode formation. (A) Specification and regionalization of the preplacodal region (PPR) at the anterior neural plate border. (B) Coordination of neural crest cell (NCC) specification and emigration along the developing neural tube, posterior to the midbrain–hindbrain boundary (MHB). Abbreviations: BMP, bone morphogenetic protein; CYP26, cytochrome P450 subfamily 26 protein; DKK, dickkopf protein; FGF, fibroblast growth factor; MHB, midbrain– hindbrain boundary; NCC, neural crest cells; PPR, preplacodal region; RA, retinoic acid; RIPPLY3, ripply transcriptional repressor 3; TBX1, t-box 1 protein; WNT, wingless-related integration site protein; ZIC1, zic family member 1.

between the future placodal and NCCs is already prefigured in the neural plate border, where neural crest regulators are located more medially than placodal regulators (Simo˜es-Costa and Bronner, 2015). As the neural plate folds, medial neural plate border specifiers thus induce a suite of genes called neural crest specifiers, which include foxd3, sox9, id, twist, and snail, while lateral neural plate border specifiers induce key placodal regulators, such as six1, eya1/2, and irx1 (Simo˜es-Costa, and Bronner, 2015). This process is thought to require an increase in BMP signaling levels, which is caused by the elevation of the neural folds (Milet and Monsoro-Burq, 2012; Simo˜es-Costa and Bronner, 2015). In addition, neural border and neural crest specifier induction are modulated by WNT, FGF, and RA signals that function posterior to the MHB to activate early posterior neural plate

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markers, such as gbx2 (Martinez-Morales et al., 2011; Milet and MonsoroBurq, 2012). The neural crest specifiers induce various neural crest effector genes that provide unique properties to the NCCs and allow them to undergo an epithelial-to-mesenchymal transition, during which they detach themselves from the dorsal neural tube, migrate along characteristic pathways through the body, and differentiate into a variety of different cell types within their respective target tissues (Martinez-Morales et al., 2011; Sauka-Spengler and Bronner-Fraser, 2008; Simo˜es-Costa and Bronner, 2015). Moreover, neural crest specifiers engage in positive cross-regulatory interactions that stabilize the regulatory state of the neural crest and ensure the maintenance of a number of specifier genes within migrating NCCs, which is thought to be crucial for preserving their plasticity and developmental potential (Groves and LaBonne, 2014; Plouhinec et al., 2014; Simo˜es-Costa and Bronner, 2015). For instance, the progeny of a single NCC can include sensory neurons, melanocytes, adrenomedullary cells, and glia cells (Gilbert, 2010). The generation of these diverse NCC fates might well be controlled through their progressive specification and emigration from the developing neural tube, which occurs in an anterior-to-posterior sequence that is highly coordinated with somitogenesis in the adjacent mesoderm (del Corral and Morales, 2014; Martinez-Morales et al., 2011; Sauka-Spengler and Bronner-Fraser, 2008). Thus, the formation of the PNS and other neural crest derivatives is directly linked to both CNS development and mesodermal segmentation (del Corral and Morales, 2014; Martinez-Morales et al., 2011). The orderly onset of neural crest induction, specification, and emigration along the rostral–caudal axis of the developing neural tube is ensured by several signals that are integrated in a complex and dynamic manner (Fig. 4B). RA signaling, for instance, is first required for expanding the expression of early dorsal patterning genes, such as the key neural border specifier pax7, toward the caudal neural plate (Martinez-Morales et al., 2011). Conversely, a posterior-high FGF gradient prevents the neural border specifiers from inducing early neural crest markers, such as snail2, and maintains high levels of the BMP antagonist Noggin (Martinez-Morales et al., 2011). While FGF signaling and BMP inhibition are necessary for the initial specification of the neural plate border and neural crest domain, further neural crest development depends on an increase of BMP levels and on a decrease of FGF levels during neurulation (Martinez-Morales et al., 2011; Milet and Monsoro-Burq, 2012). Thus, FGFs delay neural crest

Retinoic Acid in the Nervous System

31

specification in caudal regions and prevent BMP signals from stimulating wnt1 expression, which initiates the emigration of specified NCCs (Martinez-Morales et al., 2011). Anteriorly, RA signaling from the maturing somites antagonizes FGF signaling. Lower levels of FGFs permit snail2 expression and the downregulation of Noggin by signals from the paraxial mesoderm (Martinez-Morales et al., 2011; Sela-Donenfeld and Kalcheim, 2000). This enables various positive regulators to augment BMP signaling levels, which drive neural crest specification as well as the WNT1mediated induction of neural crest emigration (Martinez-Morales et al., 2011; Milet and Monsoro-Burq, 2012). In addition, RA signaling also triggers wnt1 expression, albeit only in specified NCCs that already express snail2 (Martinez-Morales et al., 2011). Accordingly, the antagonism between FGF and RA signaling appears to govern the rostral-to-caudal progression of neural crest development in concert with WNT, Noggin/BMP, and various other signals, although their specific roles and interactions have not yet been defined in detail.

6.7 Inducing and Coordinating Cranial Placode Development The cranial placodes arise from a common precursor field in the anterior neural plate border, termed the preplacodal region (PPR) (Steventon et al., 2014). As described earlier, the neural plate border is induced by a combination of intersecting signals, including FGFs from the dorsal ectoderm and intermediate levels of WNTs and BMPs from lateral and ventral regions of the embryo, that activate the expression of neural plate border specifiers, such as distalless5, pax3, pax7, and zic1/3 (Groves and LaBonne, 2014; Milet and Monsoro-Burq, 2012). However, some of these genes are expressed more medially within the neural plate border than others (Simo˜es-Costa and Bronner, 2015). Subsequent cross-regulatory interactions between the neural plate border specifiers and neural as well as nonneural transcription factors from adjacent tissues thus divide the cells of the neural plate border into a medial premigratory neural crest population and a more lateral preplacodal population (Simo˜es-Costa and Bronner, 2015). In addition, the secretion of DKK1 from the anterior pharyngeal endoderm inhibits WNT signaling anterior to where the MHB will be established, thus suppressing posterior fates and promoting placode formation (Fig. 4A) (Milet and Monsoro-Burq, 2012). As the neural plate folds, the more lateral neural plate border specifiers induce key placodal regulators, such as six1, eya1/2, and irx1

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(Simo˜es-Costa and Bronner, 2015). One of the earliest neural border specifiers expressed at the anterior neural plate is the zinc-finger transcription factor ZIC1, which is necessary for promoting placodal fate and regulating six1 and eya1/2 expression (Hong and Saint-Jeannet, 2007). Since zic1 expression does not overlap with the prospective PPR, it affects placode formation in a non cell autonomous manner (Jaurena et al., 2015). Recently, it has been reported that ZIC1 upregulates raldh2 expression in a discrete U-shaped ectodermal domain around the anterior neural plate and that it further induces LPGDS (lipocalin-type prostaglandin D2 synthase), which serves as a secreted lipophilic carrier for RA (Jaurena et al., 2015). This function of ZIC1 was shown to be crucial for the induction of placodal fate markers by RA (Jaurena et al., 2015). In the anterior neural plate, ZIC1 thus activates RA signaling, which acts rostrally to specify the PPR, with the neural plate itself being protected from RA by the expression of CYP26 enzymes (Fig. 4A) (Jaurena et al., 2015). Interestingly, RA signaling appears to exert this role in a RAR-independent manner (Jaurena et al., 2015), while its effects on the anterior–posterior patterning of the preplacodal territory are RAR dependent (Janesick et al., 2012). RA signaling via RAR initially upregulates the TBX1 transcription factor (Fig. 4A), which is required for fgf8 expression and engages in a positive feedback loop with this key preplacodal marker gene (Janesick et al., 2012). However, RAR activation also induces rippley3 expression lateral to the PPR, which is now marked by fgf8 expression (Janesick et al., 2012). Together with its corepressor GROUCHO, RIPPLEY3 represses the ability of TBX1 to maintain fgf8 expression in adjacent tissues, thus setting the posterior–lateral border of the PPR (Janesick et al., 2012). In addition, TBX1 further enhances this process by positively regulating rippley3 expression (Janesick et al., 2012). Accordingly, antagonism between FGF and RA signals is deployed for developmental patterning, even in this anterior-most part of the developing nervous system. As development proceeds, the PPR becomes subdivided into separate territories with distinct identities that can be perceived as transient local thickenings of the embryonic head and neck ectoderm (Steventon et al., 2014). These placodes give rise to a multitude of different cell types and are generally referred to as the olfactory, lens, trigeminal, epibranchial, and otic placodes (Steventon et al., 2014). The proper positioning of the preplacodal territory via RA signaling might be relevant for the subsequent separation of the different placodes, since this process is strongly influenced by the interactions and cellular movements between

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preplacodal and adjacent tissues (Janesick et al., 2012; Saint-Jeannet and Moody, 2014). However, current evidence does not support a direct involvement of RA signals in the regionalization of the PPR, which is mainly achieved through transcription factor interactions and the dynamic integration of FGF, WNT, and BMP signaling (Saint-Jeannet and Moody, 2014).

6.8 Regulating Formation of Neural Crest and Cranial Placode Derivatives The placodes and neural crest give rise to a great variety of different structures, including sensory organs and the PNS (Steventon et al., 2014). RA signals have been shown to affect the behavior of migrating NCCs, for instance, through the direct transcriptional regulation by a complex upstream RARE of nedd9, a scaffolding protein that assembles complexes involved in cell adhesion, migration, division, and survival (Aquino et al., 2009; Knutson and Clagett-Dame, 2015). This interaction likely contributes to at least some of the diverse functions RA signals exert during the morphogenesis of neural crest derivatives. Especially, the NCCs that give rise to craniofacial structures and originate mostly from caudal forebrain, midbrain, and anterior hindbrain levels are well known to be under the control of RA, which at later developmental stages is synthesized locally in the forebrain and facial ectoderm by RALDH2 as well as RALDH3 enzymes (Lee et al., 1995; Minoux and Rijli, 2010). These RA signals shape the facial area together with FGF8 and BMP4 signaling, through the coordination of local cell survival around the nasal placode and patterning of the prenasal and premaxillary skeletal elements derived from of the frontalnasal process (Minoux and Rijli, 2010). By modulating the migration, proliferation, survival, patterning, and/or differentiation of NCCs as well as their interaction with target tissues, RA signals further contribute to the development of many other neural crest derivatives, including, for example, periocular tissues of the eye (Bohnsack et al., 2012; Kumar and Duester, 2010; Matt et al., 2005, 2008), cartilage structures of the posterior pharyngeal arches (Minoux and Rijli, 2010; Niederreither et al., 2003), components of the cardiac outflow tract and heart septum (El Robrini et al., 2015; Zaffran and Niederreither, 2015) as well as teeth (Samarut et al., 2015). In addition, mutant mice lacking a functional RALDH2 enzyme for RA synthesis typically exhibit an aganglionic bowel phenotype (Niederreither et al., 2001, 2003). Therefore, RA signaling is thought to be crucial for

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the morphogenesis of the enteric nervous system from vagal NCCs, which exit the neural tube at the level of the hindbrain and migrate past the somites toward the foregut (Simkin et al., 2013). During this process, RA exposure was shown to stimulate the expression of ret, a receptor for glial-derived neurotrophic factor (GDNF), within the vagal NCCs (Simkin et al., 2013). GDNF is secreted by the gut mesenchyme and can promote survival, migration, proliferation, and differentiation of its target cells (Simkin et al., 2013). Furthermore, RA was shown to stimulate cell interactions that are involved in organizing migrating vagal NCCs into chains (Simkin et al., 2013). Thus, RA signaling enables vagal NCCs to efficiently invade and colonize the foregut and regulates their subsequent differentiation into enteric nervous system (Simkin et al., 2013). The peripheral components of the sensory nervous system often arise from both neural crest and placode progenitors, which tend to interact to drive the complex morphogenetic events that underlie PNS development (Steventon et al., 2014). For instance, NCCs constitute a major part of the frontal–nasal mesenchyme that surrounds and patterns the olfactory placode by releasing RA, which further regulates neural progenitor fate and differentiation within the olfactory epithelium (Bhasin et al., 2003; Paschaki et al., 2013a). Moreover, antagonism between FGF and RA signaling in the olfactory placode was shown to be responsible for the specification of a small population of neuroendocrine cells, which are located in the rostral hypothalamus and basal forebrain, where they integrate inputs from multiple brain areas and secrete gonadotropin-releasing hormone, a key regulator of gonad maturation, puberty, and sexual behavior (Sabado et al., 2012). RA signaling has further been implicated in the development of the adenohypophyseal (Maden et al., 2007) and lateral line (Gibbs and Northcutt, 2004) placodes as well as in the morphogenesis and patterning of the otocyst, which is formed though the invagination of the otic placode and gives rise to components of the inner ear (Hans et al., 2007; Romand et al., 2006; Thiede et al., 2014). Taken together, RA signals are used throughout the embryo and affect the formation of very different neural crest and placode derivatives, often by antagonizing FGF and other signaling pathways. Nonetheless, the precise roles of RA signaling as well as the underlying mechanisms of its actions during neural crest and placode development remain largely unknown, in particular regarding the potential contribution of RA to the formation of the PNS, which is hinted at, for example, by the importance of RA signals during peripheral nerve regeneration (Mey, 2006; Monaghan and Maden, 2012; Zhelyaznik et al., 2003).

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7. RA SIGNALING AND THE ESTABLISHMENT OF NEUROTRANSMITTER SYSTEMS 7.1 Natural Neuroactive Substances: Neurotransmitters, Neuromodulators, and Neurohormones Any chemical agent that is synthesized by a neuron and affects the properties of other neurons or effector cells (e.g., muscle cells and gland cells) can broadly be defined as a natural neuroactive substance (Hoyle, 1985). Three main types of natural neuroactive substances are generally distinguished: neurotransmitters, which are released at a synapse and alter ion permeability of the postsynaptic membrane; neuromodulators, which are released in the vicinity of synapses and affect transmission at the pre- and/or postsynaptic membrane; and neurohormones, which are released into the extracellular space or general circulation and may affect proximal as well as distant targets (Hoyle, 1985). However, for most neuroactive substances these functions overlap to variable degrees (Hoyle, 1985) and often neurons use cotransmission of several neuroactive substances to generate complex disparate outputs (Nusbaum, 2001; Powis and Bunn, 1995). Since every neurotransmitter system usually comprises specific synthesis enzymes as well as a carefully regulated molecular machinery responsible for the transport, packing, storage, release, uptake, activation, inactivation, and reception of a neuroactive substance, it is not surprising that neurons are among the most transcriptionally active cells (Moroz et al., 2006; Nelson et al., 2006). Most hormones are short polypeptides, but can also be derived from amino acids or lipids, such as steroid hormones, like cortisone or estrogen in vertebrates and ecdysone in arthropods, that are synthesized from the lipid cholesterol (Hartenstein, 2006). Important vertebrate neurohormones are the releasing hormones of the hypothalamus, which mainly regulate hormone secretion in the pituitary gland and thus control a multitude of physiological processes (e.g., thyrotropin-, corticotropin-, gonadotropin-, and growth-hormone–releasing hormone, somatostatin, dopamine, and neurotensin) (Besser and Mortimer, 1974; Guillemin, 2005), the neurohormones of the pituitary gland, which are involved in smooth muscle contraction, homoeostasis, neuromodulation, and the mediation of various social behaviors (oxytocin and vasopressin) (Stoop, 2012), the catecholamines of the chromaffin cells of the adrenal medulla, which act, for instance, on the cardiovascular system, muscle contraction, and general metabolism (dopamine, epinephrine, and norepinephrine), and the indolamines of the enterochromaffin-like cells in the gastrointestinal tract, which regulate the

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secretion of hydrochloric acid, digestive enzymes, and various other compounds (histamine and serotonin) (Westheide and Rieger, 2015). A great diversity of both peptidic and lipidic neurohormones can also be found in invertebrates, such as FMRFamides that can control heartbeat, respiration, and egg laying, myomodulins that affect muscle contraction, adipokinetic hormones that stimulate fat metabolism, pigment dispersing, timeless, and period hormones that are involved in generating a circadian rhythm, allatostatins and allatotropins that regulate food intake and inhibit the synthesis of juvenile hormone, tachykinin-related peptides that have diuretic functions and influence endocrine secretion, prothoracicotropic hormones and molt-inhibiting hormones that influence ecdysteroid synthesis, which is crucial for molting and maturation, and many others (Alzugaray et al., 2013; Bendena et al., 1999; Grimmelikhuijzen et al., 1999; N€assel, 1996, 2002; Smith and Sedlmeier, 1990; Veenstra, 2010). Classical neurotransmitter systems include aminoacidergic (e.g., aspartate, GABA, glutamate, glycin, and homocysteine), cholinergic (e.g., acetylcholine), monoaminergic (e.g., catecholaminergic, like dopamine, norepinephrine, and epinephrine, indolaminergic, like tryptamine and serotonin, and others, such as histamine and taurin), and purinergic (e.g., adenosine, AMP, ADP, and ATP) messengers as well as a plethora of neuropeptides and gaseous transmitters (e.g., nitric oxide and carbon monoxide) (Foye et al., 2008; Von Bohlen und Halbach and Dermietzel, 2006). These neurotransmitter systems are widely distributed throughout the animal kingdom, and a surprisingly large set of their molecular components has been detected in the seemingly simple cnidarians as well as in nerveless animals, such as sponges and placozoans (Fig. 5) (Grimmelikhuijzen et al., 1996; J
ekely, 2013; Kass-Simon and Pierobon, 2007; Richards et al., 2008; Riesgo et al., 2014; Smith et al., 2014; Watanabe et al., 2009). Intriguingly, many amino acid-derived neurotransmitters, including catecholamines (epinephrine, norepinephrine, and dopamine), indoles (serotonin and melatonin), histamine, acetylcholine, and nitric oxide, are also present in microorganisms, which exploit them to communicate between themselves as well as with their respective hosts (Hughes and Sperandio, 2008; Roshchina, 2010). This finding has prompted the hypothesis that the acquisition of bacterial genes via horizontal gene transfer played an essential role in the evolutionary elaboration of the existing biochemical pathways involved in animal neurotransmission (Iyer et al., 2004). RA signaling acts as a major regulatory factor during vertebrate neurogenesis (Duester, 2013; Janesick et al., 2015). Treatments with RA have

Fig. 5 Phylogenetic tree depicting a consensual view of metazoan relationships. Dotted lines indicate taxons with uncertain phylogenetic positions. Names of species discussed in this review are indicated. Data were retrieved from Borowiec, M.L., Lee, E.K., Chiu, J.C., Plachetzki, D.C., 2015. Dissecting phylogenetic signal and accounting for bias in whole-genome data sets: a case study of the Metazoa. bioRxiv 013946; Dunn, C.W., Giribet, G., Edgecombe, G.D., Hejnol, A., 2014. Animal phylogeny and its evolutionary implications. Annu. Rev. Ecol. Evol. Syst. 45, 371–395; Telford, M.J., Copley, R.R., 2011. Improving animal phylogenies with genomic data. Trends Genet. 27, 186–195; Westheide, W., Rieger, G. (Eds.), 2013. Einzeller und wirbellose Tiere, 3. Aufl. ed, Spezielle Zoologie. Springer Spektrum, Berlin.

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Table 1 Effects of Retinoic Acid Signaling on Neurotransmitter Phenotype Differentiation Affected Neurotransmitter Location Genes References Dopamine OH

+

NH2

TH D1 and D2 receptor

Jacobs et al. (2007, 2011)

Hindbrain

AP-2

Holzschuh et al. (2003)

Hindbrain

Neurog3 SERT 5-HT1A receptor

Neumeister et al. (2004), O’Reilly et al. (2007), Jacob et al. (2013), and Carcagno et al. (2014)

Spinal cord

Neurog3 Notch/ Delta GLT1

Chan et al. (2012), Jacob et al. (2013), and Carcagno et al. (2014)

OH

Norepinephrine

OH

OH

+

NH2

OH NH2

Serotonin

-

Substantia nigra

OH NH

Glutamate

+

O

O

OH

OH NH2

GABA

+

O NH2

OH

Basal ganglia GAD67 of the subventricular zone Substantia nigra

Chatzi et al. (2011)

ALDH1a1 Jacobs et al. (2011) and Kim et al. (2015)

Overview of different neurotransmitter systems that, in vertebrates, can be induced (+) or suppressed () by retinoic acid (RA) signaling in specific neuronal subsets.

been shown to induce the differentiation of stem cells into neurons with different neural identities in a concentration-dependent manner (Okada et al., 2004). RA can thus be used to generate, for instance, dopaminergic (Cooper et al., 2010; Korecka et al., 2013; Li et al., 2012), cholinergic (Berrard et al., 1993; Handler et al., 2000; Szutowicz et al., 2015), glutamatergic (Urban et al., 2015; Yu et al., 2015) or GABAergic neurons (Addae et al., 2012; Chatzi et al., 2009; Shan et al., 2011). While progress has been made toward uncovering the mechanisms through which RA signals modulate the cell cycle and promote neural fate (Duester, 2013; Ikami et al., 2015; Janesick et al., 2015; Rochette-Egly, 2015), relatively little is known about how RA signaling might contribute to the establishment of distinct neurotransmitter phenotypes (Table 1).

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7.2 Catecholaminergic Neurons Common catecholamine neurotransmitters include dopamine, norepinephrine, epinephrine, and octopamine. While dopaminergic neurotransmission is widespread in all bilaterian animals, norepinephrine and epinephrine are mainly present in vertebrates, and octopamine is typical for protostomes, but can also be found as trace amine in vertebrates (Fig. 5) (Barron et al., 2010; Moret et al., 2004). Dopamine has been shown to modulate motor circuits in representatives of the cnidarians, nematodes, platyhelminthes, annelids, mollusks, arthropods, and chordates (Barron et al., 2010). In addition, dopamine plays a well-conserved role in the regulation of feeding-related behaviors, learning, intrinsic reward, and reinforcement in diverse animal phyla (Barron et al., 2010). In the CNS of vertebrates, dopaminergic neurons can be found throughout the forebrain, the midbrain, in several “dopaminergic nuclei” of the caudal hindbrain, and in the spinal cord (Smeets and Gonza´lez, 2000). A loss of dopamine-secreting neurons in the substantia nigra (midbrain area) is known to cause Parkinson’s disease (Gr€ oger et al., 2014), and altered levels of dopamine neurotransmitter activity have been associated with schizophrenia (Kuepper et al., 2012), attention deficit hyperactivity disorder (del Campo et al., 2011), and restless legs syndrome (Trenkwalder and Paulus, 2010). In the periphery, dopamine is further released by sympathetic nerves (Goldstein and Holmes, 2008), chromaffin cells in the adrenal medulla, neuroendocrine cells in the kidney and pancreas, retinal cells, and peripheral leukocytes (Rubı´ and Maechler, 2010). Within the bloodstream, dopamine inhibits norepinephrine release and acts as a vasodilator, whereas in other regions of the body it is secreted in a paracrine manner to regulate local processes, including sodium excretion, urine output, pancreatic insulin production, gastrointestinal motility, intestinal mucosal tissue oxygenation, and the activity of lymphocytes (Rubı´ and Maechler, 2010; Yamamoto and Vernier, 2011). In contrast to dopamine, norepinephrine and epinephrine are mainly found in vertebrates (and in some protozoan, placozoan, cnidarian, and cephalopod species). Protostomes, in contrast, generally use the closely related neurotransmitter octopamine (Kass-Simon and Pierobon, 2007; Moret et al., 2004; Pfl€ uger and Stevenson, 2005). Within the CNS of vertebrates, norepinephrine is present in only six nuclei named A1, A2, A5, A7, locus coeruleus, and subcoeruleus, which originate from different rhombomeres and show distinct subtype identities (Robertson et al., 2013). The most prominent norepinephrine-containing cell cluster of the

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brain is the locus coeruleus, which sends projections to all cortical regions as well as to thalamic nuclei, septum, hippocampus, and basal lateral amygdala (Loughlin et al., 1986). In comparison, epinephrine is found in three main clusters of neurons known as nuclei C1, C2, and C3 as well as in the rostral medulla of the brainstem (Ziegler et al., 2002). The C1 neurons innervate brain areas that are strongly linked with anxiety and stress responses and provide major adrenergic input to the locus coeruleus. C2 neurons innervate the amygdala, which is an integrative center for emotion, emotional behavior, and motivation, and C3 neurons also innervate the locus coeruleus (Ziegler et al., 2002). Norepinephrine and epinephrine neurotransmission in the CNS have been implicated in the stimulation of attention, arousal, alertness, restlessness, and anxiety as well as in memory formation and retrieval (Berridge et al., 2012; Sara, 2015; Tank and Lee Wong, 2014). Furthermore, norepinephrine-positive neurons are the principal neuronal phenotype found in peripheral sympathetic ganglia, which target smooth and cardiac muscle, cutaneous and glandular structures as well as parenchymal organs (such as liver, kidney, bladder, and reproductive organs) (Robertson, 2004). Upon stimulation by sympathetic preganglionic neurons, sympathetic nerve terminals and most importantly the adrenal medulla, which is located at the center of the adrenal gland, release norepinephrine as well as adrenaline/epinephrine and small amounts of dopamine into the bloodstream (Robertson, 2004). This triggers a multitude of physiological responses, also known as fight or flight response, which include increased blood pressure and cardiac output, relaxation of bronchial, intestinal, and many other smooth muscles, mydriasis, and metabolic changes that increase levels of blood glucose and free fatty acids (Tank and Lee Wong, 2014). Octopamine has first been discovered in the posterior salivary glands of the cephalopod Octopus vulgaris (Erspamer and Boretti, 1951) and octopamine-dependent neurotransmission is typically found in protostomes, but may also occur in vertebrates, where octopamine is used as a trace amine (Jagiełło-Wo´jtowicz, 1978). In most invertebrates, high concentrations of octopamine are present both within neuronal and nonneuronal tissues and modulate nearly all physiological processes by targeting peripheral organs as well as sense organs and processes within the CNS (Roeder, 2005). Dopamine, norepinephrine, and epinephrine are synthesized from the amino acid tyrosine within the cytosol. The rate-limiting enzyme for their synthesis is tyrosine hydroxylase (TH), which catalyzes the hydroxylation of tyrosine to L-DOPA that is then decarboxylated to dopamine by aromatic amino acid decarboxylase (AAAD) (Daubner et al., 2011). Both L-DOPA

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and dopamine can be used as precursors for melanin pigment synthesis and have been shown to play a role in immunity and cuticle hardening in insects (Tang, 2009) as well as in the production of neuromelanin in the substantia nigra of humans (Fedorow et al., 2005; Solano, 2014). Moreover, in some neurons L-DOPA itself acts as a neurotransmitter or neuromodulator, without being transformed into dopamine. This has been reported, for example, for neurons in the hypothalamus and the dorsal vagal complex of vertebrates (Bj€ orklund and Dunnett, 2007; Misu and Goshima, 2006; Misu et al., 2003). Dopamine can further be transformed into norepinephrine by dopamine-β-hydroxylase (DBH) and, through an additional step catalyzed by phenylethanolamine-N-methyl transferase (PNMT), into epinephrine. The DBH enzyme is homologous to tyramine-β-hydroxylase (TBH), which synthesizes the neurotransmitter octopamine from tyramine (Monastirioti et al., 1996). Close relationships of protostome octopamine receptors and vertebrate α-adrenergic receptors suggest that the two neurotransmitter systems have diverged from a common evolutionary origin, with some evidence pointing toward the octopaminergic system being younger than the vertebrate adrenergic system (Barron et al., 2010; Pfl€ uger and Stevenson, 2005). The TH enzyme catalyzes the rate-limiting step in dopamine synthesis (Daubner et al., 2011), and it has been suggested that the gene encoding TH might be a direct downstream target of the RA signaling pathway, since RAR can robustly activate the TH promoter in a ligand-dependent manner (Jeong et al., 2006). However, RA signaling appears to exert opposite effects on TH expression, depending on the cell type (Jeong et al., 2006). For instance, all-trans RA treatment decreases TH expression levels as well as the dopamine content in dopaminergic MN9D cells, but increases th mRNA levels in human neuroblastoma SK-N-BE(2)C cells and induces transcription of the th gene in human SMS-KCNR cells (Jeong et al., 2006). RA signals also directly regulate the expression of the dopamine autoreceptor D2 (Samad et al., 1997; Valdenaire et al., 1998), and the loss of RA synthesis enzymes or receptors causes a specific reduction or elimination of D2 expression, which is associated with impaired locomotion and depressive-like behaviors (Baik et al., 1995; Krezel et al., 1998; Krzyzosiak et al., 2010; Molotkova et al., 2007; Samad et al., 1997; Wolf, 1998). The development of midbrain dopaminergic neurons has been particularly well investigated, since their dysfunction can be linked to movement disorders, such as Parkinson’s disease, as well as to drug abuse and addiction

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(Gr€ oger et al., 2014; H€ oglinger et al., 2004; Volkow et al., 2009; Whalley, 2014). During the development of mesodiencephalic dopamine neurons, the orphan NR NURR1 is responsible for the activation of TH expression and the induction of dopaminergic fate (Jacobs et al., 2011). In addition, the paired-like homeobox transcription factor PITX3 is required for the formation of a specific subpopulation of mesodiencephalic dopamine neurons in the substantia nigra, which selectively express the RA-generating enzyme ADH2 (Jacobs et al., 2007). Both ADH2 and TH expression are lost in response to the mutation of PITX3, which prevents the final differentiation of substantia nigra dopaminergic neurons, but can be rescued by RA treatment (Jacobs et al., 2007). Apart from TH, expression of DELTA-like 1 and D2 can be restored in PITX3-deficient substantia nigra dopaminergic neurons through embryonic RA treatment, whereas a number of other genes appear to be specifically modulated by PITX3 (Jacobs et al., 2011). Moreover, RA signaling was shown to represses NURR1-mediated dlk1 expression in precursors of substantia nigra dopaminergic neurons, which is thought to further promote terminal differentiation (Jacobs et al., 2011). These data demonstrate that, in contrast to other mesodiencephalic regions, the dopaminergic differentiation of neural precursors in the substantia nigra is specifically controlled through RA-dependent as well as RA-independent functions of PITX3 (Jacobs et al., 2007, 2011). Interestingly, dopamine neurons of the substantia nigra and the ventral tegmental area of the midbrain further express high levels of the enzyme RALDH1, which was shown to synthesize GABA for the corelease with dopamine, hence substituting the conventional GABA-synthesis enzymes GAD65 and GAD67 (Kim et al., 2015). Since RALDH1 is also capable of RA synthesis (Molotkov and Duester, 2003) and since RA is known to promote GABAergic neurotransmitter phenotypes (Addae et al., 2012; Chatzi et al., 2011), it might be valuable to investigate, whether RA signaling contributes to the specification and/or maintenance of both neurotransmitter phenotypes in neurons of the substantia nigra. It has further been postulated that RA signaling is involved in the specification and differentiation of late born norepinephrinergic nuclei in the hindbrain, including the locus coeruleus, and potentiates norepinephrine production by inducing AP-2, which regulates the transcription of both TH and DBH (Holzschuh et al., 2003; Kim et al., 2001; Wilson et al., 2007). In fact, regulation by AP-2 has been proposed for a multitude of neural genes, encoding, for example, acetylcholinesterase (AChE), choline acetyltransferase (ChAT), neuron-specific enolase (NES), synapsin II (SYN2),

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presenilin-2 (PSEN2), and pro-enkephalin (PENK) (Kim et al., 2001). It is thus not too surprising that RA treatments can switch neuroblastoma cell lines from a norepinephrinergic to a cholinergic phenotype by reducing TH activity, norepinephrine levels, and the calcium-dependent release of norepinephrine, while enhancing the expression of ChAT and inducing the calcium-dependent release of acetylcholine in conjunction with changes in the cellular distribution of both vesicular monoamine transporter 2 (VMAT2) and vesicular acetylcholine transporter (VAChT) (Handler et al., 2000). In addition, RA was shown to induce the expression of a sodium-dependent norepinephrine transporter (NET) in the adrenergic cells of the superior cervical ganglia of newborn rats, while decreasing th mRNA levels (Matsuoka et al., 1997b) and has further been reported to promote the proliferation of TH-expressing cells in quail neural crest cultures, causing a dose-dependent increase in the number of adrenergic cells (Dupin and Le Douarin, 1995b; Rockwood, 1996). From these studies, it is apparent that RA signals can modulate various components of catecholaminergic as well as other neurotransmitter systems, through both direct and indirect mechanisms, to instruct differentiating neurons to adopt a specific neurochemical identity in a highly context-dependent manner (Table 1).

7.3 Serotonergic Neurons Predating the evolutionary origin of neurons, the indolamine serotonin (or 5-hydroxytryptamine, 5-HT) functions as a signaling molecule in both prokaryotic and eukaryotic microorganisms (Lyte and Freestone, 2010; Tsavkelova et al., 2006), plants (Murch and Saxena, 2002; Murch et al., 2000), protozoans (Csaba et al., 2010), and sponges (Nickel, 2010; Weyrer et al., 1999). Moreover, serotonin neurons that arise early during development have been described in almost all animal groups (Fig. 5), prompting the hypothesis that the role of serotonin as a neurotransmitter might have been a direct consequence of the evolution of synapses (Hay-Schmidt, 2000; Ryan and Grant, 2009). In vertebrates, serotonin modulates a wide range of physiological processes in all major organ systems, including the cardiovascular, pulmonary, gastrointestinal, and genitourinary systems as well as the CNS (Berger et al., 2009). Most serotonin neurons are found in the hypothalamus and hindbrain of vertebrates, where they contribute to the regulation of locomotion, mood, sleep, anxiety, drug abuse, food intake, and sexual behavior (Veenstra-VanderWeele et al., 2000).

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The first rate-limiting step of serotonin synthesis is the hydroxylation of L-tryptophan to L-5-hydroxytryptophan by tryptophan hydroxylase (TPH). Vertebrates possess at least two different TPH proteins, with TPH1 being found in several tissues, and TPH2 being neuron specific (C^ ot
e et al., 2003; Gutknecht et al., 2009; Savelieva et al., 2008; Walther, 2003). Subsequently, AAAD and its coenzyme pyridoxal 50 -phosphate decarboxylate L-5-hydroxytryptophan to serotonin (Lovenberg et al., 1963). In the hindbrain of African clawed frog larvae, exogenous RA induces dose-dependent changes in the number, distribution, and projection patterns of serotonin-positive neurons (Ruiz i Altaba and Jessell, 1991). Moreover, in cells derived from rat embryonic raphe nuclei RA treatments have been shown to increase intracellular levels of serotonin, of the 5-HT1A receptor as well as of the reuptake transporter SERT (Neumeister et al., 2004; O’Reilly et al., 2007). Since both the 5-HT1A receptor and SERT are important negative regulators of serotonin levels within the extracellular cleft, their increased expression probably causes an inhibition of serotonin neurotransmission and might thus contribute to RA-inducible depressionrelated behaviors in mice (Neumeister et al., 2004; O’Reilly et al., 2007). Consistently, it has been postulated that brain serotonin concentrations can be lowered through the oral administration of RA (Smazal and Schalinske, 2013). These data indicate an influence of RA signaling on several components of the serotonin neurotransmitter system. In vertebrates, TGFβ, together with FGFs and SHH, are crucial for establishing the molecular signaling pathways leading to serotonergic differentiation (Farkas et al., 2003; Ye et al., 1998), and the formation of serotonin-producing cells has been shown to depend on a specific transcription factor program involving at least four genes: ascl1, nkx2.2, lmx1b, and pet-1 (Pattyn, 2003; Pattyn et al., 2004). Together, ascl1 and nkx2.2 activate the expression of lmx1b, pet-1, isnm1, and gata3, which induces the serotonin transmitter phenotype (Jacob et al., 2009; Pattyn et al., 2004). LMX1B controls major components of serotonin metabolism, including TPH and SERT. Notably, besides working together with PET-1 and NKX2.2 to specify the vast majority of serotonin-positive neurons, LMX1B may act autonomously or with other factors to control the formation of specific subsets of serotonergic neurons, especially in rostral brain regions (Cheng et al., 2003). Moreover, LMX1B is also necessary for the development and maintenance of dopaminergic neurons (Nakatani et al., 2010; Yan et al., 2011). PET-1, on the other hand, is a precise marker of developing and adult serotonergic neurons and is mainly involved in

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serotonergic neuron maturation, regulating axonal innervation of the somatosensory cortex, expression of appropriate firing properties, and the expression of the 5-HT1A and 5-HT1B receptors (Hendricks et al., 2003). PET-1 is further required in adult serotonin neurons to preserve normal anxiety-related behaviors through direct autoregulated control of serotonergic gene expression (Liu et al., 2010). However, the transcriptional code of serotonin progenitors in the ventral hindbrain is largely shared by the topographically related progenitors of V3 neurons in the ventral spinal cord, which also express NKX2.2, ASCL1, and FOXA2, but acquire a glutamatergic neurotransmitter phenotype (Carcagno et al., 2014; Jacob et al., 2013). Recent studies in mice have revealed that different levels of RA signaling are responsible for the serotonin vs V3 fate decision (Carcagno et al., 2014; Jacob et al., 2013). Around the time neurogenesis commences, RA signaling is weak or absent in the ventral hindbrain domain of serotonin progenitors and ascl1 expression levels are markedly higher than in the ventral spinal cord domain of V3 progenitors, where RA signaling is strong (Jacob et al., 2013). In order to achieve this dose-dependent attenuation of ascl1 expression, RA signals activate the Notch pathway, directly as well as through the induction of neurog3 expression in the spinal cord, which ultimately leads to the repression of ascl1 by HES proteins (Carcagno et al., 2014; Jacob et al., 2013). Thus, since high levels of ASCL1 are sufficient for committing neural tube progenitors to a serotonergic fate, RA signals allocate serotonergic and glutamatergic V3 identities in an ASCL1-dependent manner by inducing quantitative changes in the transcriptional code, via NEUROG3 and Notch function (Carcagno et al., 2014; Jacob et al., 2013). Hindbrain serotonergic neurons are subdivided into a rostral group near the mid–hindbrain boundary and a caudal group in the prospective myelencephalon, with a serotonin neuron-free zone located in between, in rhombomere 4 (Smidt and van Hooft, 2013). Due to the dynamic autoand cross-regulatory interactions of RA, FGF, and WNT signals as well as of various transcription factors and hox genes, the expression of HOXB1 is induced and selectively maintained in rhombomere 4, where it activates the transcription factor PHOX2 that represses the serotonergic developmental program (Glover et al., 2006; Philippidou and Dasen, 2013; Smidt and van Hooft, 2013; Terriente and Pujades, 2015). Therefore, RA signaling is crucial for determining the proper distribution of serotonin-positive neurons during development and potentially affects serotonin signaling directly through the modulation of the 5-HT1A receptor and SERT (Table 1).

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7.4 Glutamate and GABAergic Neurons As for serotonin, glutamate and GABA (γ-aminobutyric acid) are ancient chemical messengers that fulfill a variety of functions in plants, protists, and metazoans (Bouch
e et al., 2003; Bucci et al., 2005; Elliott and Leys, 2010; Ramoino et al., 2010, 2014; Van Houten et al., 2000). Glutamate is the most abundant excitatory amino acid in the vertebrate brain, which participates in most nervous system functions and is crucial for synaptic plasticity, learning, and memory (McEntee and Crook, 1993). GABA, on the other hand, reduces neuronal excitability, which is especially important for the control of motor function (Draper et al., 2014; Wong et al., 1991) and the regulation of anxiety (Lydiard, 2003). In addition, GABA is involved in vision (Herrmann et al., 2011) and, during nervous system development, influences neural migration as well as neurite extension and acts as a neurotrophic factor (Watanabe et al., 2002). Outside the nervous system, glutamate and GABA signaling have further been implicated in the control of numerous physiological processes, ranging from insulin release and bone formation to gastrointestinal motility and hormone secretion (Nedergaard et al., 2002; Watanabe et al., 2002). Glutamate is a nonessential amino acid and thus, for metabolic purposes, found in most cells, albeit at variable concentrations. Since glutamate cannot cross the blood/hemolymph-brain barrier that is present in numerous taxa, including vertebrates, arthropods, and cephalopods (Fig. 5), it has to be synthesized within the CNS from local precursors (Hawkins, 2009; Parpura and Verkhratsky, 2012). However, neurons are not capable of producing glutamate de novo from glucose through the tricarboxylic acid cycle, because they lack the enzyme pyruvate carboxylase (PC), which, in vertebrates, is specifically expressed in astrocytes and, in insects, is found in glia cells (Bak et al., 2006; Freeman et al., 2003; Shank et al., 1985). Instead, neuronal glutamate production is based on a metabolite trade known as the glutamate/glutamine cycle, during which the glutamate precursor glutamine is released from glial cells and subsequently metabolized to glutamate within the presynaptic vesicles of neurons by the mitochondrial enzyme glutaminase (Bak et al., 2006). Astrocytes, in turn, take up glutamate released into the synaptic cleft by neurons and convert it back to glutamine, a reaction catalyzed by the glial glutamine synthetase (GS) enzyme (Bak et al., 2006; Schousboe et al., 2013). GABA is synthesized from glutamate by glutamate decarboxylase (GAD). Mammals possess two GAD enzymes named GAD65 and GAD67 (Erlander et al., 1991). While GAD67 is an active holoenzyme that is present throughout the cytosol of GABAergic neurons, GAD65 is

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enriched in nerve endings and functions predominantly as a dormant apoenzyme, which can rapidly be activated through binding of its coenzyme, pyridoxal phosphate (Walls et al., 2011). In addition, a second evolutionary conserved mechanism for GABA synthesis exists, which involves the oxidative deamination of the precursor putrescine to γ-aminobutyraldehyde and the subsequent conversion into GABA by an aldehyde dehydrogenase (ALDH1a1, also called RALDH1) (Kim et al., 2015; Seiler and Eichentopf, 1975). Metabolization of GABA to succinic semialdehyde by GABA transaminase (GABA-T) occurs only in the presence of α-ketoglutarate, which by accepting the amino group removed from GABA reforms glutamate, while succinic semialdehyde is oxidized to succinic acid that enters the tricarboxylic acid cycle thus completing a metabolic glutamate/GABA loop known as the GABA shunt (Richard and DeLorey, 1999; Schousboe et al., 2013). As described earlier, RA signals are crucial for the specification and differentiation of glutamatergic V3 neurons in the ventral spinal cord, where they suppress serotonin neurotransmitter phenotypes by upregulation of neurog3 and Notch-mediated inhibition of ascl1 (Carcagno et al., 2014; Jacob et al., 2013). In cell culture studies, RA was further shown to promote glutamatergic sensory cell fates (Martinez-Monedero et al., 2008; Urban et al., 2015; Yu et al., 2015), which is particularly interesting, since RA signaling has been implicated in the development and regeneration of sensory hair cells in the inner ear that use glutamate as a major neurotransmitter (Lee et al., 2015; Romand et al., 2006; Rubbini et al., 2015; Thiede et al., 2014). However, it is currently unknown, if RA signaling can also affect the induction and/or functioning of glutamatergic neurotransmitter systems in a nonpermissive manner. In the adult CNS, RA has been reported to decrease levels of astrocytic glutamate transporter-1 (GLT-1) through RXRmediated transcriptional inhibition and the activation of protein kinase C (PKC) activity, which leads to GLT-1 internalization (Chan et al., 2012). Since GLT-1 is responsible for about 90% of glutamate uptake in the forebrain, its deletion from astrocytes causes excess mortality, lower body weight, and spontaneous seizures, due to the defective clearance and neurotoxicity of extracellular glutamate (Petr et al., 2015). Furthermore, GLT-1 deficiency has been linked to dicarboxylic aminoaciduria and obsessive–compulsive disorders in humans as well as neurodegeneration in mice (Petr et al., 2015). Conversely, overexpression of GLT-1 reduces glutamate activity and impairs prepulse inhibition of the startle reflex, a simple form of information processing that is reduced in schizophrenia (Bellesi and Conti, 2010; Bellesi et al., 2009). Thus, RA signaling might contribute to the

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regulation of extracellular glutamate levels, which is essential for CNS function and homeostasis. RA treatment of embryonic stem cells can be used for the efficient and scalable generation of functional GABAergic interneurons in culture (Addae et al., 2012; Chatzi et al., 2009; Shan et al., 2011). In the subventricular zone of mice basal ganglia, RA generated by RADLH3 is required for the specification and differentiation of both GABAergic striatal projection neurons and GABAergic interneurons that migrate to the olfactory bulb and cortex (Chatzi et al., 2011). In this context, RA signaling was shown to activate the expression of the GABA-synthesizing enzyme GAD67, albeit through an unidentified mechanism, since a RARE has not been identified in the promoter region of the mouse gad67 gene (Chatzi et al., 2011; Goodman and Kline, 1996). As previously mentioned, RA signaling is involved in the specification of a population of neurons located in the substantia nigra that corelease dopamine and GABA (Jacobs et al., 2011; Kim et al., 2015). In these cells, GABA is synthesized by RALDH1, a typical RA synthesizing enzyme, instead of by GAD (Jacobs et al., 2011; Kim et al., 2015). In addition, neurons of the ventral tegmental area, which is located in the vicinity of the substantia nigra, also express RALDH1 and have recently been reported to corelease GABA with glutamate (Kim et al., 2015; Root et al., 2014). From these findings, it can be hypothesized that RA signaling might be involved in the induction and/or the maintenance of a GABA cotransmitter phenotype. Apart from influencing neurotransmitter identity, RA is capable of rapidly changing excitatory as well as inhibitory synaptic strength, by acting through a very distinct nongenomic mechanism requiring the binding of RA to RAR, which subsequently acts as RNA-binding protein to regulate translation (Arendt et al., 2015a,b). RA synthesis is induced in response to prolonged neural inactivity, due to a lack of calcium influx in the absence of spontaneous neurotransmission (Arendt et al., 2015a,b). In excitatory synapses, RA subsequently enhances synaptic strength by increasing the abundance of postsynaptic ionotropic glutamate receptors (AMPARs) (Arendt et al., 2015a,b; Chen et al., 2014). In inhibitory synapses, on the other hand, RA induces the rapid internalization of GABAA receptors from the synaptic membrane, thus attenuating inhibitory neurotransmission (Arendt et al., 2015a,b; Sarti et al., 2013). In sum, while the transcriptional regulation by RA signaling can promote distinct neurotransmitter identities in a context-dependent manner, RA activity on the translational level directly interferes with synaptic plasticity and neurotransmission (Table 1).

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8. RA SIGNALING OUTSIDE VERTEBRATES: EVOLUTIONARY CONSIDERATIONS Recent estimates suggest that the total number of animal species on earth ranges from 7,400,000 to 10,000,000, albeit it has also been proposed that there might be 5,000,000–6,000,000 species of insects alone (Pimm et al., 2014). Thus far, approximately 1,900,000 extant animal species have been described, but the great majority is still unknown (Pimm et al., 2014). The phylogenetic relationships between these species hold the key to understanding how different body plans and characters have emerged throughout the course of evolution and what fundamental properties animal life forms have in common. However, many critical relationships, especially at the root of the phylogenetic tree, between the sponge, ctenophore, cnidarian, and bilaterian lineages, are still uncertain (Fig. 5) (Borowiec et al., 2015; Dunn et al., 2014; Schierwater et al., 2016; Telford and Copley, 2011). RA signaling was initially described in mammals and, over the past few decades, has been thought to be first vertebrate-, then chordate-, and finally deuterostome-specific, with putative roles in protostomes, where incomplete complements of the classical signaling pathway had been identified (Fig. 6) (Campo-Paysaa et al., 2008). While the vertebrate RAR binds both

Fig. 6 Distribution of retinoic acid (RA) signaling pathway components across the animal kingdom. The colored bars indicate the presence of proteins required for canonical RA signaling in different taxa. Abbreviations: CYP26, cytochrome P450 subfamily 26 protein; CRABP, cellular retinoic acid-binding protein; CRBP, cellular retinol-binding protein; RALDH, retinaldehyde dehydrogenase; RAR, retinoic acid receptor; RXR, retinoid X receptor.

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all-trans and 9-cis RA, RXR is thought to bind only 9-cis RA (GutierrezMazariegos et al., 2014b). However, endogenous levels of both isoforms have been measured in a variety of animals, including species that lack an unequivocal RAR ortholog (Andr
e et al., 2014; Gutierrez-Mazariegos et al., 2014b). Studying how the RA system is deployed in other animal taxa is thus likely to reveal not only the conserved properties of this impotent vertebrate morphogen, but also new functions, signaling cascades, and regulatory mechanisms. This might further allow the identification of new medical targets or strategies to manipulate RA-dependent processes either during disease progression or in regenerative therapy (Camacho, 2003; Collins and Mao, 1999; Doll
e and Niederreither, 2015). ADH and RALDH enzymes for the synthesis of RA are ubiquitously distributed across the animal kingdom (Fig. 6) (Andr
e et al., 2014). Even the demosponge Suberites domuncula possesses a β-carotene dioxygenase (BCO) and a retinol dehydrogenase/reductase (RDH) and it was further suggested that various retinoid precursors are generated by bacteria encapsulated in the bacteriocytes of this animal (M€ uller et al., 2011; Wang et al., 2013; Wiens et al., 2003). Treatment with exogenous RA was shown to upregulate the gene expression of a RXR-like protein, a BMP, and a LIM protein in primmorphs of S. domuncula, while also stimulating the formation of canal-like structures and siliceous spicules (Larroux et al., 2006; M€ uller et al., 2011; Wang et al., 2013; Wiens et al., 2003). Additional evidence for the RA-mediated regulation of a homeobox-containing gene comes from the freshwater sponge Ephydatia muelleri (Nikko et al., 2001). Furthermore, three aldehyde dehydrogenases (ALDHs), a single CYP26, and a single RXR have been identified in the placozoan Trichoplax adhaerens (Albalat and Can˜estro, 2009; Reitzel et al., 2011). Together, these data indicate an ancient origin of RA signaling at the base of metazoans, which might have involved microorganismic contributions. However, the presence of other pathway components, such as CRBPs and CRABPs and true RAR receptors, as well as the structural and functional properties of sponge RXR-like proteins remain to be investigated. Comparable to the situation in sponges, RARs are probably absent from ctenophores and cnidarians (Fig. 6) (Laudet et al., 2015). In fact, only three NRs have been identified in ctenophores so far, all of which belong to the subfamily NR2A (HNF4) and lack an otherwise well-conserved zinc-finger DNA-binding domain (Reitzel et al., 2011). Analyses of the genomes of two anthozoan and one hydrozoan species, all of which do not have a medusa stage in their life cycle, did not detect a bona fide RXR (Chapman et al.,

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2010; Fuchs et al., 2014; Putnam et al., 2007; Shinzato et al., 2011). In contrast, RXRs are present in species possessing a medusoid stage, such as the scyphozoan Aurelia aurita, the hydrozoan Clytia hemisphaerica, and the cubozoan Tripedalia cystophora, in which the RXR was further shown to bind 9-cis RA with high affinity and to interact as a monomer with the promoters of J1A and J1B crystallins (Fuchs et al., 2014; Kostrouch et al., 1998). In the scyphozoan A. aurita, the transition from a sessile asexual polyp stage to a pelagic medusa via strobilation (spontaneous transverse body segmentation) coincides with the differential upregulation of RA synthesizing enzymes and RXR (Fuchs et al., 2014). Moreover, pharmacological blocking of RXR function suppresses strobilation, while incubation in retinol or RA is sufficient to substitute for the temperature shift that normally induces this process (Fuchs et al., 2014). These findings prompted the hypothesis that RA signaling links putative temperature-sensing receptors to downstream genes responsible for the onset of metamorphosis in A. aurita (Fuchs et al., 2014). This notion is further supported by a recent transcriptomic analysis of the different life cycle stages of A. aurita, showing that RXR upregulation during strobilation is accompanied by the upregulation of klf13 (kr€ uppel-like factor 13) and precedes the expression of the paired-like homeobox gene drg (Brekhman et al., 2015). In vertebrates, KLFs are involved in various developmental processes, such as cell cycle regulation, and are known for their regulatory interactions with NRs, including RAR-dependent RA signaling (Knoedler and Denver, 2014; Shi et al., 2012; Zheng et al., 2009). DRG transcription factors, in turn, are know for controlling somatosensory functions in the PNS of a wide variety of animals, including both invertebrates and vertebrates (Brekhman et al., 2015; Nomaksteinsky et al., 2013). Several other studies provide evidence for an involvement of RA signaling in cnidarian development. For instance, in cell cultures of the anthozoan Renilla koellikeri RA positively regulates cell proliferation and differentiation into epithelium-associated cells including sensory cells (Estephane and Anctil, 2010), in the planula larvae of the hydrozoan Clava multicornis RA signaling is required during nervous system patterning and neurogenesis (Pennati et al., 2013), and in two anthozoan species RXR-like immunoreactivity was localized in putative ectodermal sensory neurons (Bouzaiene et al., 2007). In sum, RA signaling functions appear to be well integrated into cnidarian development, with particularly important roles during metamorphosis and neural development. Yet, the classical RA signaling pathway, mediated by RAR/RXR heterodimers, is likely an innovation of bilaterian

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animals, although taxon sampling in nonbilaterian groups is still insufficient to reach a final conclusion (Laudet et al., 2015). In the Ecdysozoa, RARs have most likely been lost secondarily (Fig. 6) and the only documented case of an intracellular RBP, which recognizes both retinol and RA, comes from the shrimp Metapenaeus ensis, where it is expressed in the ovary (together with RXR), in the eyestalk, testis, and epidermis (Cui et al., 2013; Gu et al., 2002; Laudet et al., 2015). However, the proposed phylogenetic affinity between this shrimp RBP and the chordate CRBPs and CRABPs is highly controversial (Gu et al., 2002; Laudet et al., 2015), and it has thus been proposed that intracellular RBPs evolved independently in the Ecdysozoa and Chordata (Albalat et al., 2011). Furthermore, a nematode-specific fatty acid- and retinoid-binding protein (FAR) has been described that is able to bind RA and is involved in nematode development, reproduction, and the infection of host plants (Zhang et al., 2015a). Of note, the phylogenetic relationships between this nematode-specific protein and the chordate CRBPs and CRABPs still remain to be assessed. RXRs, on the other hand, are well known to be widely distributed across all bilaterian phyla and have been cloned from various representatives of the Deuterostomia, Ecdysozoa, and Lophotrochozoa (Fig. 6), including the platyhelminthes Schistosoma mansoni and Schistosoma japonicum (de Mendonc¸a et al., 2000; Laudet et al., 2015; Qiu et al., 2013). While the two RXR homologues of S. mansoni are unlikely to bind RA and probably require specific heterodimeric partners to interact with DNA response elements, it was reported that the RXR of S. japonicum can bind 9-cis RA and is upregulated by RA treatment (de Mendonc¸a et al., 2000; Qiu et al., 2013). Furthermore, in nematodes RXR antagonists as well as other synthetic and natural retinoids have been shown to cause developmental defects and decrease animal viability (Hurst et al., 2014). Arthropod RXRs typically form heterodimers with ecdysone receptors to mediate ecdysteroid hormone functions during development, reproduction, molting, metamorphosis, and regeneration (Andr
e et al., 2014). In crustaceans, multiple RXR isoforms are differentially expressed (Cui et al., 2013; Li et al., 2014), but it is not clear whether they can also transduce RA signals, which have so far only been shown to affect glucose metabolism and negatively interfere with limb regeneration in decapods (Hopkins and Durica, 1995; Sreenivasula Reddy and Srilatha, 2015). In insects, however, the presence of endogenous 9-cis RA and all-trans RA as well as their ability to directly activate RXR orthologs has been confirmed (Nowickyj et al., 2008), and a

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number of physiological responses to RA signals have been described, such as teratogenic effects on larval development (Nakamura et al., 2007) and the inhibition of metamorphosis (Neˇmec et al., 1993), but also positive effects on cell survival and neurite outgrowth in cultures of embryonic locust neurons (Sukiban et al., 2014). Interestingly, RXR immunoreactivity was detected in both the cytoplasm and the neurites of specific neuronal subsets in the embryonic and adult CNS of the locust Locusta migratoria, hinting at a potential role for RA in insect neurogenesis and nervous system function (Bui-G€ obbels et al., 2015; Sukiban et al., 2014). Moreover, RA signals have been reported to regulate neurohormonal gene expression in response to tissue damage in the fruit fly Drosophila melanogaster, thus generating a developmental checkpoint for regeneration (Halme et al., 2010). This is a surprising result considering that the RXR of D. melanogaster, called Ultraspiracle (USP), is probably unable to bind RA (Iwema et al., 2007). Altogether, these findings strongly suggest that RA signaling is present in at least some ecdysozoan species, although the physiological relevance and underlying mechanisms are still largely unexplored. In addition to RXRs, lophotrochozoans and deuterostomes also possess RARs and CYP26 orthologs (Fig. 6), suggesting an increased importance of RA signaling functions in these two clades (Laudet et al., 2015). In gastropod mollusks, rxr and rar are expressed starting from early developmental stages and treatment with either exogenous RA, RXR pan-agonist or vertebrate RAR antagonist disrupts embryogenesis causing, for example, abnormal eye and shell development (Carter et al., 2010, 2015; Creton et al., 1993). Similar to the locust L. migratoria, RXR, but also RAR, of the gastropod Lymnaea stagnalis are localized in the cytoplasm, neurites, and growth cones of CNS neurons (Bui-G€ obbels et al., 2015; Carter et al., 2010, 2015). While RAR antagonist blocks RA-dependent growth cone turning in regenerating neurons of L. stagnalis, exposure to exogenous RA or RXR agonist induces growth cone turning, even in isolated neurites without a cell body or nucleus (Carter et al., 2010, 2015; Farrar et al., 2009). This suggests that a nongenomic RA-dependent mechanism, mediated by both RAR and RXR, controls growth cone guidance in L. stagnalis (Carter et al., 2010, 2015). Electrophysiological experiments have further revealed rapid dose- and isomer-dependent effects of RA on the firing properties of L. stagnalis neurons, which were significantly reduced in the presence of RXR antagonist and are likely mediated through the RA-dependent modulation of calcium signaling (Vesprini and Spencer, 2014; Vesprini et al., 2015). RA is also

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necessary for long-term memory formation in L. stagnalis, but not for learning or intermediate-term memory, and this function of the (light sensitive) RA molecule can be promoted by constant darkness (Rothwell and Spencer, 2014; Rothwell et al., 2014). Altogether, these roles of RA signaling in gastropod mollusks are reminiscent of nongenomic RA actions during vertebrate homeostatic synaptic plasticity. For example, within neuronal dendrites RA has been shown to bind RAR that subsequently functions locally as a RNA-binding protein modulating protein translation (Chen et al., 2014). Concerning RA functions outside of the mollusk nervous system, injection of 9-cis RA into adult gastropod females was shown to trigger the development of male sexual organs, in a RXR-dependent, and likely RAR-independent, manner (Abidli et al., 2013; Castro et al., 2007; Stange et al., 2012). Intriguingly, this phenomenon, also known as imposex, seems to correlate with an inability to store retinoids in the form of retinyl esters in susceptible species (Gesto et al., 2013). Taken together, these studies illustrate multiple roles for both RXR- and RAR-dependent RA signals during snail development and neurogenesis. It is thus tempting to conclude that lophotrochozoan and vertebrate RA signaling systems are quite similar. Conversely, however, the RARs of at least two gastropod species, Nucella lapillus and Thais clavigera, are incapable of binding RA, albeit exhibiting the capacity to form heterodimers with RXRs and to recognize DNA response elements organized in direct repeats (Gutierrez-Mazariegos et al., 2014a; Urushitani et al., 2013). Accordingly, investigations of classical as well as alternative RA signaling pathway components and their functional properties in basal lophotrochozoans, such as lophophorate annelids and platyhelminthes, are required to shed additional light on the mechanisms of action of this morphogen in lophotrochozoans. In deuterostomes, all major components needed for classical RA signaling, i.e., RAR and RXR as well as RALDH and CYP26, appear to be present (Fig. 6), although intracellular RBPs such as CRBPs and CRABPs, have so far only been identified and characterized in vertebrates, with the notable exception of one putative CRBP ortholog in cephalochordates (Laudet et al., 2015). At present, very little is known about the functions and mechanisms of RA signaling in nonchordate deuterostomes. For instance, all-trans RA might either disrupt or merely delay sea urchin embryonic development and probably interferes with the migration of micromerederived cells (Laudet et al., 2015). In contrast, studies in invertebrate chordates, such as cephalochordates and ascidian tunicates, have revealed numerous well-conserved functions of RA signals during chordate development,

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for example, in axial patterning, tissue specification, and organogenesis (Campo-Paysaa et al., 2008; Carvalho and Schubert, 2013; Laudet et al., 2015). In tunicates, an independent loss of the RA machinery has occurred in certain groups, such as the larvaceans, while in other groups RA and RAR have been implicated in tissue regeneration and bud development (Andr
e et al., 2014). However, it has also been shown that the RXR of the ascidian Polyandrocarpa misakiensis induces the expression of transdifferentiation-related genes, such as RXR, ERK, and MYC, in a RA-dependent manner without a major contribution of RAR (Kawamura et al., 2013). Remarkably, despite gene loss and fragmentation of the tunicate hox cluster, RA-driven hox1 expression in the epidermal ectoderm of the ascidian Ciona intestinalis was shown to be essential for organizing the atrial siphon placode, a homolog of the vertebrate otic placode (Sasakura et al., 2012). This hints at an invertebrate chordate origin of RA signaling functions in vertebrate neural crest and placode development. In addition, interactions between the RA, FGF/MAPK, and canonical WNT pathways control anterior–posterior patterning of the tail epidermis and tail PNS in C. intestinalis, in a manner that resembles mechanisms that are typically deployed by vertebrates to coordinate embryonic patterning and body elongation (Pasini et al., 2012). In contrast, RA and FGF signals probably act independently during posterior elongation in the cephalochordate amphioxus and, contrary to vertebrates, are not mandatory for posterior somite formation (Bertrand et al., 2015). Yet, cephalochordates are characterized by a particularly vertebrate-like, albeit less elaborate, retinoid signaling system (Laudet et al., 2015). Since they have not undergone the two rounds of wholegenome duplications that are typical of vertebrates, their rar and rxr are each encoded by a single genomic locus (Andr
e et al., 2014; Laudet et al., 2015). Importantly, it has been demonstrated that the amphioxus RAR/RXR heterodimer can be activated by binding RA, which is necessary, for instance, for establishing the collinear expression patterns of amphioxus hox genes as well as for the specification of different cellular identities throughout the amphioxus larva (Escriva et al., 2002; Schubert et al., 2006). Nevertheless, amphioxus seems to lack several genes encoding retinol storage, transport, and cellular uptake proteins, which might thus be a vertebrate exclusivity, although storage of retinyl esters might also occur in gastropod mollusks (Albalat et al., 2011; Andr
e et al., 2014; Laudet et al., 2015). Moreover, as mentioned above, RA signaling in amphioxus might not engage in the mutual antagonistic cross talk with FGF signaling that governs many developmental processes in vertebrates (Bertrand et al., 2015).

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Fig. 7 Summary of retinoic acid (RA)-dependent processes in different invertebrate taxa. Note that, except for chordates, these functions are likely mediated through alternative signaling mechanisms, i.e., not involving the activation of retinoic acid receptors (RARs) by RA.

In conclusion, RA signaling exhibits a great diversity of biological functions (Fig. 7) and molecular mechanisms that seem to be scattered across the animal kingdom. This impression can be attributed at least in part to the very scarce availability of reliable information from many invertebrate taxa. Future work will thus have to address this problem by focusing on the characterization of the retinoid system and its function in various organisms located at key positions in the animal tree of life, including, but not being limited to, sponges, ctenophores, cnidarians, xenacoelomorphs, basal ecdysozoans and lophotrochozoans, cephalopods, echinoderms, hemichordates as well as additional invertebrate chordate taxa.

9. CONCLUDING REMARKS Throughout vertebrate development, opposing RA and FGF signaling gradients are fundamental for coordinating the formation and patterning of many different tissues and organ systems (Fig. 2). During this process, RA signaling contributes to the proper specification of neural progenitor populations and controls their progression into differentiated neurons in a context-dependent manner, thus matching neurogenesis to other organogenetic processes occurring along the embryonic trunk. However, while considerable progress has been made toward elucidating the molecular

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framework underlying RA-driven neural progenitor specification and differentiation, much less is known about how and under which conditions other functions of RA are mediated. For instance, the role RA plays in promoting neural progenitor proliferation and maintenance, in determining the subtype identity and neurotransmitter phenotype of differentiating neurons, and in modulating neurotransmission and synaptic plasticity are just beginning to be uncovered. In order to better exploit RA for therapeutic purposes, it is thus urgent to expand our knowledge of its context-dependency and to characterize its alternate signaling mechanisms, functions, and interactions. To this end, new techniques that allow the visualization of RA signaling activity in vivo should help to pinpoint regions of particular interest (Shimozono et al., 2013). Furthermore, it will be important to expand studies of RA signaling toward later developmental and adult stages and perform comprehensive stage-, tissue-, and cell-specific assays of its genomic and nongenomic targets. Moreover, the evolution of RA signaling systems should be investigated further, since most invertebrate species appear to deploy this molecule in different ways that could help identify novel targets and tools for the pharmaceutical manipulation of RA-dependent processes in vertebrates. Also, exciting new insights into how a more complex RA signaling system might have contributed to the formation of a more elaborate nervous system could potentially be gained from studying key taxons, such as cephalochordates, which possess a prototypical chordate RA signaling and nervous system, and cephalopods, which are characterized by a highly complex nervous system that has evolved independently from that of vertebrates (Fig. 5).

ACKNOWLEDGMENTS The authors are indebted to Jenifer C. Croce for critical comments on the manuscript. This work was supported by a grant from the Agence Nationale de la Recherche (ANR-11-JSV2002-01) and by funds from the R
eseau Andr
e Picard (ANR-11-IDEX-0004-02, Sorbonne Universite´s). E.Z. is a doctoral fellow of the Studienstiftung der deutschen Wirtschaft (SDW).

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CHAPTER TWO

AMBRA1, a Novel BH3-Like Protein: New Insights Into the AMBRA1–BCL2-Family Proteins Relationship A. Di Rita*,†, F. Strappazzon*,1 *IRCCS Santa Lucia Foundation, Rome, Italy † University of Rome Tor Vergata, Rome, Italy 1 Corresponding author: e-mail address: [email protected]

Contents 1. Introduction 2. AMBRA1 in Autophagy, Selective Autophagy, and Beyond 2.1 AMBRA1 in Autophagy 2.2 AMBRA1 in Mitophagy 2.3 AMBRA1 in Apoptosis, Development, Differentiation, Viral Infection, and Proliferation 2.4 AMBRA1 in Diseases 3. Autophagy and Apoptosis Cross-Talk 3.1 The Antiapoptotic BCL2 Proteins Control the Autophagy Process 3.2 AMBRA1 Contains a BH3 Domain Necessary for BCL2 Binding 4. Autophagy Inhibition by the Oncogenic Function of BCL2 4.1 Autophagy Pathway as a Tumor Suppressor 4.2 BCL2 Inhibits AMBRA1-Dependent Autophagy: Could It Be a Novel Mechanism to Regulate Tumorigenesis? 5. BCL2–AMBRA1 Interaction as a Novel Therapeutic Target 5.1 Autophagy Induction Through BCL2–AMBRA1 Dissociation 5.2 Is AMBRA1-Mediated Mitophagy Regulated by BCL2 Family Proteins? 6. Conclusion Acknowledgments References

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Abstract Cellular homeostasis swings like a pendulum backward and forward between life and death. Two of the main processes, which regulate this equilibrium, are autophagy and apoptosis. While autophagy is a highly conserved self-digestion mechanism that mediates degradation of damaged or surplus components, apoptosis is a programmed cell suicide in which typical death signals induce the elimination of undesired cells. Both International Review of Cell and Molecular Biology, Volume 330 ISSN 1937-6448 http://dx.doi.org/10.1016/bs.ircmb.2016.09.002

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these processes are highly regulated by complex molecular machineries, including some common proteins whose “dual role” favors one process or the other. Among these proteins, the well-known antiapoptotic factor BCL2 downregulates autophagy through interactions with the essential autophagic effectors, BECN1/BECLIN 1 and AMBRA1. Recently, we have demonstrated that the proautophagic protein AMBRA1 contains a BH3 domain necessary for AMBRA1 binding with the antiapoptotic factor BCL2. We found that the AMBRA1–BCL2 couple have a “dual role” in autophagy and apoptosis: the mitochondrial pool of BCL2 is able to inhibit AMBRA1-dependent autophagy, whereas in cell death conditions, the cleaved form of AMBRA1 (AMBRA1CT), resulting from CASP/CASPASES-cleavage, abrogates the prosurvival activity of BCL2 and promotes a proapoptotic amplification loop. The CASP-cleaved form of AMBRA1 bound other antiapoptotic members of the BCL2 family proteins such as MCL1 and BCL2L1/BCL-X; by contrast, no binding could be detected with the proapoptotic-BCL2 factors such as BAK1/BAK and BAX. These findings underline an intricate interplay between autophagy and cell death in which the proautophagic protein AMBRA1 and the antiapoptotic BCL2 family members are the major players. Here, we give an overview of the AMBRA1-BCL2 family proteins interactome and its involvement in controlling life and cell death. We discuss a putative therapeutic target which offers the novel BH3 motif identified in the C-terminal part of AMBRA1.

ABBREVIATIONS ActA actin assembly-inducing protein AMBRA1 autophagy/beclin-1 regulator 1 BAD BCL2-associated agonist of cell death BAK1/BAK BCL2-antagonist/killer 1 BAX BCL2-associated X protein BCL2 B-cell lymphoma 2 BCL2L1/BCL-X BCL2-like 1 BCL2L2/BCL-W BCL2-like 2 BECN1/BECLIN1 beclin 1, autophagy related BH3 BCL2 homology domain 3 BH3-only BCL2 homology domain 3 only BNIP3 BCL2/adenovirus E1B 19 kDa interacting protein 3 BNIP3L/NIX BCL2/adenovirus E1B 19 kDa interacting protein 3-like CASP/CASPASES caspases CAPN/CALPAINS calpains CUL4 cullin 4 CUL5 cullin 5 CYCS/CYTOCHROME C cytochrome C DYNLL1/DLC1 dynein, light chain, LC8-type 1 DYNLL2/DLC2 dynein, light chain, LC8-type 2 ER endoplasmic reticulum ER-BCL2 endoplasmatic reticulum BCL2 FUNDC1 FUN14 domain containing 1

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MAP1LC3B/LC3B microtubule-associated protein 1 light chain 3 beta MCL1 myeloid cell leukemia 1 mito-BCL2 mitochondrial BCL2 MOMP mitochondrial outer membrane permeabilization OPTN optineurin PARK2/PARKIN Parkin RBR E3 ubiquitin protein ligase PIK3C3/VPS34 phosphatidylinositol 3-kinase catalytic subunit type 3 PINK1 PTEN-induced putative kinase 1 PPP2/PP2A protein phosphatase 2A SQSTM1/P62 Sequestosome 1 UVRAG UV radiation resistance associated

1. INTRODUCTION The balance between death and life is at the basis of cellular homeostasis regulation. Autophagy and apoptosis are two well-known processes, which mediate this important equilibrium through the action of several proteins. Autophagy is a cellular self-digestion-mediated by degradative vesicles named autophagolysosomes (Mizushima, 2007; Suzuki and Ohsumi, 2007). This mechanism, in addition to remove randomly cellular components, can also eliminate some substrates “selectively.” The selective autophagy pathways are controlled by specific molecular components named “cargo receptors” or “autophagic receptors” which can interact with specific substrates, thus bringing them into autophagosome vesicles (Stolz et al., 2014; Svenning and Johansen, 2013). On the other side, apoptosis is a tightly controlled physiological pathway in which specific death signals lead to the elimination of undesired, damaged, or infected cells (Bright and Khar, 1994; Kerr et al., 1972; Orrenius et al., 2007; White and McCubrey, 2001). This programmed cell death is controlled by the CASP/CASPASES (caspases)-dependent pathway that consists of three different mechanisms: the intrinsic or extrinsic pathway and the ER-stress induced apoptosis (Kumar, 2007; Li et al., 2006). Among these two important cellular processes, AMBRA1 (the autophagy/beclin-1 regulator 1) is one of the bestcharacterized proteins implicated in both the autophagic machinery and in the regulation of apoptosis. Its proautophagic role, as a regulator of the early autophagic steps and as a scaffold protein in the recruitment of the autophagy-implicated molecules, links the process of autophagy with other cellular pathways such as cellular proliferation, differentiation, development,

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apoptosis, and pathologies (Cecconi et al., 2007; Cianfanelli et al., 2015a,b; Fimia et al., 2007; Heinrich et al., 2013; Nardacci et al., 2014; Skobo et al., 2014; Strappazzon et al., 2011; Va´zquez et al., 2012; Yazdankhah et al., 2014). These multiple functions of AMBRA1 are controlled by posttranslational modifications on AMBRA1 and/or on AMBRA1 interactors, but also by CASP-mediated cleavage (Strappazzon et al., 2016). It is known that the antiapoptotic BCL2 proteins, such as BCL2 (B-cell lymphoma 2), BCL2L1/BCL-X (BCL2-like 1), and MCL1 (myeloid cell leukemia 1), negatively regulate autophagy (Germain et al., 2011; Maiuri et al., 2007; Pattingre et al., 2005; Strappazzon et al., 2011; Wu et al., 2014a). Interestingly, it is emerging that these BCL2 proteins are also AMBRA1-interacting proteins. The interaction between AMBRA1 and the “ostensible outsider” BCL2 proteins highlights the AMBRA1 extensive network of interactions and confirms AMBRA1’s connective role between autophagy and apoptosis. This review describes the prevailing knowledge about the relationship between AMBRA1 and its antiapoptotic BCL2-interacting proteins at the apoptosis/autophagy crossroads.

2. AMBRA1 IN AUTOPHAGY, SELECTIVE AUTOPHAGY, AND BEYOND AMBRA1 was described for the first time as a positive regulator of autophagy during vertebrate’s embryogenesis (Fimia et al., 2007). AMBRA1 is a protein of 1300 amino acids, and the AMBRA1 gene, located on human chromosome 11 and mouse chromosome 2, is composed of 18 exons. According to the whole-chromosomal duplication, after divergent evolution of fish and tetrapod (Meyer and Van de Peer, 2005), AMBRA1 gene of zebrafish is composed of 19 exons and codifies for two paralog nonredundant genes called ambra1a and ambra1b (Benato et al., 2014; Skobo et al., 2014). The difference in the number and in the length of the exons corresponds to the C-terminal region of AMBRA1 low sequence identity among mammals and zebrafish. Instead, the N-terminal region is highly conserved and contains three WD40 domains necessary for organizing a scaffold for protein–protein interactions. In addition, AMBRA1 possesses Serine- and Proline-rich regions with still unknown functions and several consensus sequences which allow the binding with its noted interactors CUL4 (cullin 4)—DDB1 (damage-specific DNAbinding protein 1), CUL5 (cullin 5), PPP2/PP2A (protein phosphatase 2A),

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MAP1LC3B (microtubule-associated protein 1 light chain 3 beta), TRAF6 (TNF receptor-associated factor 6), DYNLL1, DYNLL2, and BCL2 (Antonioli et al., 2014; Cianfanelli et al., 2015b; Di Bartolomeo et al., 2010; Nazio et al., 2013; Strappazzon et al., 2015, 2016). In addition to these famous domains, AMBRA1 is defined as an intrinsically disordered protein. Its plasticity is responsible for its multiple interactions and functions (Cianfanelli et al., 2015c). Its intricate and unknown structure sustains the real nature of AMBRA1 as one of the main players in several physiological and pathological processes. As mentioned earlier, AMBRA1 has a central role in autophagy regulation and in addition a crucial function in selective autophagy: in particular, AMBRA1 is implicated in the mitochondrial selective autophagy hereinafter called mitophagy (Strappazzon et al., 2015; Van Humbeeck et al., 2011). AMBRA1 is also involved in several processes such as proliferation, differentiation, development of the cells, apoptosis and is associated with several pathologies (Cecconi et al., 2007; Cianfanelli et al., 2015b; Fimia et al., 2007; Nardacci et al., 2014; Pagliarini et al., 2012; Skobo et al., 2014; Strappazzon et al., 2016; Va´zquez et al., 2012; Yazdankhah et al., 2014).

2.1 AMBRA1 in Autophagy In basal condition, AMBRA1 interacts with BECN1/BECLIN 1 (beclin 1, autophagy related) and PIK3C3/VPS34 (phosphatidylinositol 3-kinase catalytic subunit type 3) proteins which form the PIK3C3 complex at the dynein light chains DYNLL1/DLC1 (dynein, light chain, LC8-type 1) and DYNLL2/DLC2 (dynein, light chain, LC8-type 2) of the dynein motor complex. Following autophagy induction, the PIK3C3 complex translocates, thanks to an ULK1 (Unc-51 like autophagy activating kinase 1)dependent phosphorylation on AMBRA1, to the endoplasmatic reticulum where it activates the PIK3C3 complex in order to initiate the autophagosome formation. AMBRA1 is thus a positive regulator of autophagy (Di Bartolomeo et al., 2010; Fimia et al, 2007). AMBRA1 also mediates a positive feedback loop of the autophagic pathway by controlling ULK1 stability through a K63-ubiquitylation mediated by the E3 ligase TRAF6 (Nazio et al., 2013). Moreover, AMBRA1 is an essential cofactor for other E3 ubiquitin ligases during autophagy. For instance, it has been demonstrated that AMBRA1 dynamically interacts with the E3 ubiquitin ligases CUL5 and CUL4 in order to regulate, respectively, the early and late stages of autophagy (Antonioli et al., 2014). In basal condition, AMBRA1

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interaction with the E3 ligase CUL4, through DDB1 (damage-specific DNA-binding protein 1), stabilizes AMBRA1 protein level regulating AMBRA1 ubiquitylation. Following autophagy induction, a rapid release of AMBRA1 from DDB1 leads to CUL4/AMBRA1 dissociation which increases the AMBRA1 pool. The CUL4/DDB1/AMBRA1 separation is transient; a consequent reassociation induces AMBRA1 degradation, thus promoting the autophagic process termination. Moreover, upon CUL4 dissociation, AMBRA1 interacts with CUL5 through TCEB2/ELONGIN B (transcription elongation factor B (SIII), polypeptide 2 (18 kDa, elongin B)) in order to suppress the CUL5 E3 ligase activity. This repression allows the DEPTOR (DEP-domain-containing mTOR-interacting protein) stabilization which inhibits mTORC1, so functionality leading to a positivefeedback loop on autophagy regulation. In addition to its cofactorial role for E3 ligases TRAF6, CUL4, and CUL5, AMBRA1 is a direct substrate for two different E3 ligases which control its ubiquitin-mediated proteasomal degradation. As mentioned earlier, in basal conditions, CUL4–DDB1 complex coordinates AMBRA1 ubiquitylation and subsequently its degradation in order to repress autophagy. Moreover, upon autophagy induction, the E3 ubiquitin ligase RNF2 (Ring finger protein 2) interacting with WASH (Wiskott–Aldrich syndrome protein and SCAR homolog) regulates AMBRA1 ubiquitylation on Lys45 by Lys48-linked chains in order to arrest the autophagic pathway (Xia et al., 2014).

2.2 AMBRA1 in Mitophagy It was initially demonstrated that AMBRA1 is a PARK2/PARKIN (parkin RBR E3 ubiquitin protein ligase) interactor and that this binding is strongly increased during prolonged mitochondrial depolarization (Van Humbeeck et al., 2011). These authors found that, following mitophagy induction, AMBRA1 is recruited, like PARK2, in perinuclear mitochondria clusters, named mitoaggresomes, this in order to collaborate in autophagosome formation by activating the PIK3C3 complex. More recently, AMBRA1 was described as a LIR (LC3-interacting region)-containing protein, the mitochondrial pool of AMBRA1 being able to interact, during mitophagy, with MAP1LC3B via this LIR motif (Strappazzon et al., 2015). The interaction between AMBRA1 and MAP1LC3B is essential in order to potentiate the PINK1 (PTEN-induced putative kinase 1)/PARK2-dependent mitophagy. In addition, by expressing a mitochondria-targeted construct of AMBRA1 (AMBRA1–ActA), AMBRA1 can induce a massive PARK2

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and SQSTM1/P62 (Sequestosome 1)-independent mitophagy pathway (Strappazzon and Cecconi, 2015a; Strappazzon et al., 2015). In this context, AMBRA1–ActA interacts with MAP1LC3B protein via its MAP1LC3Binteracting region (LIR) in order to facilitate mitochondrial clearance by transporting damaged mitochondria onto autophagosomes. It is noteworthy that the wild-type form of AMBRA1 is sufficient to restore mitophagy per se in PINK- or PARK2-defective cells, this highlighting a specific AMBRA1dependent mitophagy pathway. It would be of great interest to identify the novel E3 ubiquitin ligase which ubiquitylates mitochondria following AMBRA1–ActA expression, in order to signal them for degradation. Finally, AMBRA1 presents typical characteristics of the mitophagic receptors FUNDC1 (FUN14 domain containing 1), BNIP3 (BCL2/adenovirus E1B 19 kDa interacting protein 3), and BNIP3L/NIX (BCL2/adenovirus E1B 19 kDa interacting protein 3-like), including (1) the outer mitochondrial membrane localization, (2) the presence of a LIR motif, and (3) the power of mitophagy induction following overexpression at the mitochondria. These common features of AMBRA1 with the other “mitophagic receptors” could define AMBRA1 as a novel mitophagic receptor. These findings suggest a key role of AMBRA1 in the regulation of cellular homeostasis through mitophagy. The multitask feature of AMBRA1 in both autophagy and selective autophagy is illustrated in Fig. 1.

2.3 AMBRA1 in Apoptosis, Development, Differentiation, Viral Infection, and Proliferation A central role of AMBRA1 in apoptosis was initially demonstrated by the presence of a large number of apoptotic cells in Ambra1-deficient mice (Cecconi et al., 2007; Fimia et al., 2007). In addition, AMBRA1 role in programmed cell death is confirmed by the observation that AMBRA1 is a direct substrate of CASP and CAPN/CALPAINS (Calpains) mediate a prodeath cleavage on its Aspartate 482 (Pagliarini et al., 2012). In particular, the CASP-resistant mutant (AMBRA1-D482A) preserves cells from death better than AMBRA1’s wild-type form does; this indicates that a tight regulation of its proautophagic activity occurs during the apoptotic process. A dynamic interaction between the mitochondrial pool of AMBRA1 and the antiapoptotic BCL2 protein has also been demonstrated, underlying the involvement of a proautophagic factor in a prodeath pathway (Strappazzon et al., 2011, 2015).

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Fig. 1 AMBRA1 is a proautophagic protein implicated both in macroautophagy and mitophagy. (A) In basal condition, AMBRA1 together with BECN1 and PIK3C3 (PIK3C3 complex) is bound to the dynein motor complex of microtubule. Following autophagy induction, the ULK1-mediated phosphorylation on AMBRA1 leads to the PIK3C3 complex translocation to the endoplasmatic reticulum where the phagophore originates. The AMBRA1 phosphorylation, mediated by ULK1 kinase, is thus fundamental for triggering autophagy. (B) AMBRA1 is a LIR protein: its interaction with the autophagosome marker MAP1LC3B enhances PINK1/PARK2-dependent mitophagy, under mitophagic stress. Moreover the AMBRA1–ActA construct induces mitophagy per se in a PINK1/ PARK2-independent manner and the wild-type form of AMBRA1 can induce mitophagy upon CCCP/FCCP treatment (most likely it is anchored to the mitochondria by linking mito-BCL2).

AMBRA1 has also a fundamental role during embryogenesis in vertebrates, acting as a regulator of neural tube formation and controlling cell proliferation during central nervous system (CNS) development (Fimia et al., 2007). Ambra1 is principally expressed in the neuroepithelium at embryogenesis’s early stage and is subsequently present in the spinal cord,

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encephalic vesicles, neural retina, and dorsal root ganglia (Fimia et al., 2007; Va´zquez et al., 2012). Ambra1-defected mouse embryos show serious neural tube defects correlated with autophagy impairment and subsequent excess in cell proliferation at early stages of embryogenesis; this causes the induction of apoptosis process and the accumulation of ubiquitylated proteins in the neuroepithelium. These developmental anomalies due to Ambra1 deletion lead to the embryonic lethality around stage E16.5 (Fimia et al., 2007). Interestingly, the central role of Ambra1 in controlling CNS dynamics can be observed during adult life. In this context, Ambra1 is highly expressed in the brain subventricular zone, together with BECN1, supporting neural stem cells pool and controls the amount of immature neurons by inducing the survival of neural precursor cells (Yazdankhah et al., 2014). Moreover, it has been observed that the in vitro Ambra1 downregulation in adult neural stem cells leads to a reduction in cell proliferation, as well as to an increase in basal apoptosis and a major sensitivity to cell death induced by DNA damage. AMBRA1 is also fundamental for myogenesis, its expression being crucial for correct morphogenesis and consequently for the development of skeletal muscle in zebrafish (Skobo et al., 2014). The zebrafish’s two paralogous genes of AMBRA1, ambra1a and ambra1b, are both necessary for embryogenesis and larval development. In fact, the lack of both ambra1a and ambra1b causes the reduction of the locomotor activity and induces defects in myofibers and myosepta, which can be reversed by coinjection of human AMBRA1 mRNA. It has been noticed that, although the ambra1a or ambra1b knockdown genes are sufficient to change muscle structure, the double knockdown of the two paralogous genes shows more defects in respect to the single knockdown; this suggests that ambra1a and ambra1b proteins also have distinct roles in zebrafish, besides working in similar molecular systems. In analogy with this evidence, mice embryos that are homozygous for an Ambra1 gene trap mutation (Ambra1gt/gt) present a severe myopathy characterized by anomalies in mitochondria morphology, disorganization of sarcomeres, a high reduction, and abnormal orientation of myofibers (Skobo et al., 2014). Autophagic machinery is also crucial for viral infection. High levels of AMBRA1 expression noticed in the peripheral blood mononuclear cells and in lymph nodes of nonprogressor human immunodeficiency virus-1 (HIV-1)-infected patients contribute to the maintenance of a strong autophagic response. The induced autophagy in HIV-1-infected individuals could be responsible for the clinical stability which has been detected in the absence of therapy (Nardacci et al., 2014).

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Finally, AMBRA1 is defined as a tumor suppressor, acting as a MYC (v-myc avian myelocytomatosis viral oncogene homolog) antagonist. It thus plays a crucial role in cell proliferation and tumorigenesis (Cianfanelli et al., 2015b). Upon growth factor receptor’ activation, the protooncogene MYC is phosphorylated on its Ser 62, thus leading to its stabilization and promoting the increase in cell proliferation (Sears et al., 2000). In this context, AMBRA1 directly interacts with the catalytic subunit of the phosphatase PPP2 induces MYC dephosphorylation, this leading to oncoprotein proteasomal degradation. Interestingly, this AMBRA1-mediated inhibition of cancer cells proliferation is correlated with MTORC1 (MTOR complex 1) repression, which is a signal for MYC dephosphorylation (Cianfanelli et al., 2015b). It is noteworthy that missense, nonsense, and frame-shift mutations of AMBRA1 gene, correlated with AMBRA1 loss of function, have also been associated with cancer in several human tissues (Cianfanelli et al., 2015b). In addition, the levels of AMBRA1 protein are reversed correlated with the levels of Phospho-Ser62 MYC both in lung and breast cancer cell lines. Moreover, the AMBRA1 expression is sufficient to reduce the phosphorylation on the oncogene MYC and thus promoting the decrease in tumorigenesis in cancer cell lines characterized by a low level of endogenous AMBRA1 gene. Instead, the massive disruption of the AMBRA1 locus promotes MYC hyperphosphorylation and hyperproliferation and thus, tumorigenesis (Cianfanelli et al., 2015b). These important results underline that AMBRA1 is the missing link not only in autophagy machinery but also in the regulation of cell proliferation and in susceptibility to cancers. In fact, it has been demonstrated that heterozygous for an Ambra1 gene trap mutation mice (Ambra1+/gt) are three times more likely to generate lung, liver, and kidney tumors, respect to the wild-type mice. Furthermore, cells deficient for AMBRA1 gene have a pronounced capacity to grow once injected into nude mice (Cianfanelli et al., 2015b).

2.4 AMBRA1 in Diseases Since AMBRA1 is involved in multiple processes, it is no surprise that AMBRA1 is correlated to a number of pathological conditions, mainly to the neural-related pathologies. Among these diseases, schizophrenia is a well-characterized neural disturbance associated with AMBRA1 mutations. For instance, it has been demonstrated that the genetic mutation in a limited region of chromosome

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11 in which the AMBRA1 gene is located is implicated in schizophrenia etiology (Rietschel et al., 2012). Schizophrenia is a serious mental disorder characterized by distortion of thought and perception, by abnormalities in social behavior, by hallucinations that contributing to genetic factors promoting the risk of developing the disease. A genetic approach-based study demonstrated a correlation between AMBRA1 and various aspects of impulsivity (Heinrich et al., 2013). In addition, comprehensive behavioral studies of mice which were heterozygous for an Ambra1 gene trap mutation (Ambra1+/gt) show an autismlike phenotype in adult and pup females, including compromised social interactions, a tendency to exhibit repetitive behaviors and impaired cognitive flexibility (Dere et al., 2014). It has been observed that these Ambra1+/gt mice present another pathological phenotype regarding the dramatically enhanced and prolonged neuropathic pain following nerve damage and axonal degeneration. The AMBRA1 haploinsufficiency mimicking the reduction in autophagy response in Schwann cells after nerve injury results in a persistent and severe painful response which induces self-lesioning behaviors in the injured limb (Marinelli et al., 2014). Moreover, it has been demonstrated that AMBRA1 expression is strongly downregulated in mice affected by amyotrophic lateral sclerosis (ALS) (Wakabayashi et al., 2014). ALS is a severe neurodegenerative disorder characterized by muscle twitching, stiff muscles, and progressive weakness due to muscles size decrease (Bendotti et al., 2016). Interestingly, mutations of the mitophagic receptor OPTN (Optineurin) are involved in the onset of ALS (Maruyama et al., 2010). More particularly, the ALSassociated mutation E478G in OPTN determinates a decrease in OPTN recruitment to the mitochondria and a consequent impairment in the mitophagic process. This result highlights a potential function for mitophagic defective proteins in ALS (Wong and Holzbaur, 2014). In line with this evidence, it would be now crucial to investigate whether the AMBRA1 protein, by acting as a “defective mitophagic receptor,” could play a role in the onset of ALS disease.

3. AUTOPHAGY AND APOPTOSIS CROSS-TALK The interplay between autophagy and apoptosis is highly intricate. There are several key proteins implicated in both these processes, indicating a correlation between these two regulator pathways of cellular homeostasis

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(Fimia et al., 2013; Sinha and Levine, 2008). In fact, both autophagy and apoptosis proceed in parallel rails and with several common passengers traveling on distinct routes, although these are often entwined.

3.1 The Antiapoptotic BCL2 Proteins Control the Autophagy Process The involvement of BCL2 family in several cellular processes is due to its large network of interactions. For instance BCL2 is a BECN1-interacting protein, linking directly BCL2 to the central player of the autophagy mechanism (Liang et al., 1998). More particularly, the endoplasmic reticulum pool of BCL2 (ER-BCL2), together with CISD2/NAF-1 (CDGSH iron sulfur domain 2), is able to negatively regulate the BECN1-autophagic pathway (Chang et al., 2010; Pattingre et al., 2005). The interaction between BECN1 and ER-BCL2 leads to a dual antiapoptotic and antiautophagic regulation. The mitochondrial pool of BCL2 (mito-BCL2) exhibits an antiapoptotic function and can also interact with BECN1 (Pattingre et al., 2005; Sinha and Levine, 2008). Moreover, other antiapoptotic factors, such as BCL2L1, MCL1, BCL2L2 (BCL2-like 2), and v-BCL2 (viral homologous of BCL2), negatively regulate autophagy by an inhibiting interaction with BECN1 (Erlich et al., 2007; Feng et al., 2007; Germain et al., 2011; Maiuri et al., 2007; Oberstein et al., 2007; Pattingre et al., 2005; Sinha et al., 2008). The interaction between BECN1 and the antiapoptotic BCL2 proteins occurs through a highly conserved BH3-like domain on BECN1 (Maiuri et al., 2007), defining BECN1 as a BH3-only protein (Sinha and Levine, 2008). Mutations on BECN1 BH3 domain inhibit its interactions with BCL2/BCL2L1/MCL1 and allow the autophagic process to occur (Maiuri et al., 2007; Oberstein et al., 2007). There are four proposed mechanisms that could explain the disruption of the binding between BECN1 and antiapoptotic BCL2 repertoire in order to promote autophagy: (1) posttranslational modification on BECN1 or on the antiapoptotic members; (2) BECN1 dissociation from antiapoptotic BCL2 family mediated by competitive BH3-only proteins; (3) the regulation of the BECN1 and BCL2 proteins expression; (4) the subcellular localization of these proteins (Sinha and Levine, 2008). The MAPK8/JNK (mitogen-activated protein kinase 8) kinase-mediated phosphorylation of BCL2 disrupts the BCL2–BECN1 interaction (Pattingre et al., 2009; Wei et al., 2008a,b). Moreover, BECN1 can be phosphorylated on its BH3 domain and this modification is sufficient for BECN1 release from the BCL2 and BCL2L1 complex (Zalckvar et al., 2009). According to Maiuri (2007) the BH3-only protein BAD

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(BCL2-associated agonist of cell death) is also a competitive inhibitor of BECN1–BCL2L1/BCL2 interactions. In addition, several BH3-only proteins, such as BNIP3L, BBC3/PUMA (BCL2-binding component 3), BIK (BCL-interacting killer), PMAIP1/NOXA (phorbol-12-myristate-13acetate-induced protein 1), BCL2L11/BIM (BCL2-like 11) are implicated in promoting autophagy or mitophagy (Abedin et al., 2007; Rashmi et al., 2008; Yee et al., 2009; Zhang et al., 2008). In a BAD-like manner, these BH3-only proteins could inhibit the BECN1–BCL2L1/BCL2 axis in order to explain their functions. Finally, the BH3 mimetic compound ABT737 inhibits the BECN1–BCL2L1/BCL2 interaction and is responsible for autophagy induction (Maiuri et al, 2007). These findings highlight not only the importance of the BECN1 BH3-like motif for the interaction with the majority of the antiapoptotic members, but also the competitive process involved in these various bindings. Beyond BECN1, further autophagic proteins are BCL2 family-interacting proteins; among them: (1) UVRAG (UV radiation resistance associated) binds the proapoptotic member BAX (BCL2-associated X protein), leading to a block of the apoptosis pathway by limiting BAX activation and its translocation to mitochondria (Yin et al., 2011); (2) the autophagic ubiquitin-like protein ATG12 (Autophagy related 12) interacts with the prosurvival BCL2 and MCL1 members, acting as a positive regulator of apoptosis upstream the MOMP (mitochondrial outer membrane permeabilization) (Rubinstein et al., 2011); (3) AMBRA1, in basal condition, can interact with the mitochondrial pool of BCL2. This binding inhibits the proautophagic activity of AMBRA1 at the mitochondria (Strappazzon et al., 2011). More recently, it has been demonstrated that also MCL1 and BCL2L1 can coimmunoprecipitate with AMBRA1. It would be now necessary to investigate whether the endogenous protein AMBRA1 can form a complex at the mitochondria with MCL1/BCL2L1 and to evaluate, as in the case of BCL2, whether MCL1 or BCL2L1 can inhibit the proautophagic activity of AMBRA1.

3.2 AMBRA1 Contains a BH3 Domain Necessary for BCL2 Binding The intricate relationship between autophagy and death machinery never ceases to amaze: in fact, the AMBRA1–BCL2 interaction, which regulates autophagy, also plays a role in the apoptotic process (Strappazzon et al., 2016). Following MOMP, the apoptotic proteases CASP and CAPN cleave several proautophagic members, such as ATG4D (Autophagy related 4D,

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cysteine peptidase), ATG5 (Autophagy related 5), BECN1, and AMBRA1, thus inhibiting their prosurvival activity (Fimia et al., 2013). These cleaved forms translocate to the mitochondria and increase apoptotic pathway (Betin and Lane, 2009; Cho et al., 2008; Fimia et al., 2013; Rohn et al., 2011; Wirawan et al., 2010). More particularly, the ATG5 cleavage regulated by CAPN produce a truncated protein which contributes to apoptosis by interacting directly with BCL2L1 at the mitochondria (Yousefi et al., 2006). The CASP-mediated cleavage of ATG4D produces a C-terminal fragment which localizes to mitochondria (Betin and Lane, 2009). In addition, BECN1 is cleaved by two CASP, generating a mitochondria target fragment of BECN1, named BECN1CT; this loses its autophagic activity and gains a proapoptotic function (Wirawan et al., 2010). CASP-mediated cleavage of AMBRA1 also generates a C-terminal fragment (483–1300 aa) hereinafter called AMBRA1CT, which disrupts the BCL2 antiapoptotic function through a direct interaction with BCL2. Interestingly, this interaction occurs through a BH3 consensus sequence localized at the C-terminal region of AMBRA1 (Pagliarini et al., 2012; Strappazzon et al., 2016). In this way, the cleaved form of AMBRA1 acts as a BH3-only protein, abrogates the prosurvival activity of BCL2 and amplifies apoptosis. As shown in Fig. 2, the alignment of the BH3 consensus sequence of AMBRA1 with the other known BH3 motifs confirms the similarity of this BCL2-interacting protein sequence and in particular highlights a strong resemblance with the BH3 domain of BCL2L1. Of note, two point mutations on the BH3 domain of AMBRA1 (AMBRA1BH3-AE) reduce AMBRA1–BCL2 interaction and lead to a reduction in the proapoptotic function of AMBRA1CT.

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Fig. 2 AMBRA1 contains a BH3 consensus sequence which allows its binding with BCL2. Sequence-based alignment of the best-characterized BH3 containing proteins highlights the analogy between the known BH3 consensus sequence and AMBRA1 BH3 motif. It is noteworthy that several residues are highly conserved: in fact, the consensus LxEAGDxxx is highly similar between AMBRA1 and BCL2L1 BH3 motifs.

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In analogy with BECN1–BCL2 binding, it would be opportune to investigate whether posttranslational modifications, protein competition, gene expression, or protein subcellular localization could modulate AMBRA1– BCL2 interaction. Interestingly, AMBRA1CT can coimmunoprecipitate with the antiapoptotic proteins MCL1 and BCL2L1 (Strappazzon et al., 2016). The BH3 motif of the C-terminal region of AMBRA1 is also probably important for its interaction with MCL1 and BCL2L1. It would be of great interest to investigate, as in the case of BCL2, whether the antiapoptotic functions of MCL1 and BCL2L1 could be inhibited by the C-terminal part of AMBRA1 following cell death induction. The dual effect between AMBRA1 and the antiapoptotic member BCL2 seems to be restricted to the antiapoptotic category of proteins since no binding can be found between AMBRA1 and the proapoptotic BCL2 members such as BAK1/BAK (BCL2-antagonist/killer 1) and BAX. It is noteworthy that AMBRA1 needs to be cleaved in order to block BCL2 function; furthermore, upon apoptotic induction, MCL1 and BCL2L1 bind AMBRA1CT in preference compared to the full-length form of AMBRA1. The fulllength AMBRA1 could have a conformation state that masks the BH3 region on its C-terminal part and thus could arrest its inhibitory regulation on BCL2. During apoptosis, we could speculate that the AMBRA1 conformational change opens a “door” that favors its interaction with antiapoptotic BCL2 proteins. For instance, the interaction between the proautophagic factor ATG12 and BCL2 family proteins is atypical and unique. ATG12 contains a noncanonical BH3-like motif: its sequence is similar to other BH3 domains, but the structure forms a loop that binds the antiapoptotic BCL2 members’ hydrophobic pocket, instead the normal alpha helix (Rubinstein et al., 2011). In line with this study, it should be very interesting to perform a conformational analysis of AMBRA1 under normal conditions as compared to apoptosis conditions. Since AMBRA1 is a disordered protein, unfortunately to date it has been very difficult to obtain a threedimensional (3D) structure of the protein. In future, resolving the AMBRA1 3D structure would be useful so as to evaluate the importance of its conformation in the regulation of AMBRA1 multiinteractions. In particular, since AMBRA1 and BCL2 family proteins are both at the crossroads between autophagy and apoptosis, resolving the AMBRA1 structure could offer a solution for a better understanding of AMBRA1’s role in the complicated interplay between life and death. These findings support the evidence for a strict interplay between autophagy and apoptosis: indeed, the BCL2 family interaction network with

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proautophagic proteins controls both autophagy and apoptosis. Furthermore, CASP- and/or CAPN-mediated cleaved proteins such as AMBRA1CT, ATG5, BECN1CT, and ATG4DCT are also capable of controlling apoptosis. Of note, only AMBRA1CT and ATG5 have been shown to act directly on the antiapoptotic BCL2 family proteins, as shown in Fig. 3. Proautophagic members

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Fig. 3 Interactions between the BCL2 family members and the proautophagic proteins and their respective effects on autophagy and apoptosis processes. BECN1 interactions with MCL1 and BCL2L1 negatively control both autophagy and apoptosis. The interaction with ER-BCL2 leads to a block in the autophagic and apoptotic processes. The ubiquitin-like ATG12 interacts with MCL1 and BCL2 proteins enhancing apoptosis. AMBRA1 is a mito-BCL2 interacting protein: this interaction inhibits AMBRA1-dependent autophagy by trapping a pool of AMBRA1 at the mitochondria. The proautophagic protein UVRAG interacts with the proapoptotic member BAX in order to block apoptosis. In addition, proautophagic truncated forms, originated from CASP or CAPN cleavage regulate apoptosis: for instance, AMBRA1CT interacts with BCL2 in order to increase apoptosis, whereas cleaved-ATG5 induces apoptosis by interacting with BCL2L1. BECN1CT and ATG4DCT, instead, promote apoptosis, although to date there is no evidence for their interaction with BCL2 family of proteins.

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4. AUTOPHAGY INHIBITION BY THE ONCOGENIC FUNCTION OF BCL2 The BCL2 protein has been initially identified as an oncoprotein, blocking apoptotic cell death without increasing cell proliferation (McDonnell et al., 1989; Vaux et al., 1998). From another viewpoint, AMBRA1 protein is not only a scaffold protein regulating autophagy but also a point of convergence of both autophagy and cell proliferation. Indeed, AMBRA1 could function as a tumor suppressor, acting as oncogene MYC antagonist and regulating cell proliferation and tumorigenesis (Cianfanelli et al., 2015b). In this context, AMBRA1 interacts directly with the phosphatase PPP2 which consequently induces MYC dephosphorylation and thus promotes its proteasomal degradation.

4.1 Autophagy Pathway as a Tumor Suppressor The controversial role of autophagy in cancer divides the scientific community. The dual nature of autophagy could modulate prosurvival of cells so acting as a cell defender or, alternatively it could control prodeath mechanisms in tumor onset and progression, as a cell hangman. There is a large body of evidence to support the autophagy-mediated inhibition of cancer development and to describe autophagy as a tumor suppressor mechanism. Among proautophagic genes, BECN1 was initially identified as a tumor suppressor (Liang et al., 1999; Qu et al., 2003); it has been observed that the BECN1 locus is deleted in 50–70% of breast cancers and in up to 75% of ovarian cancers. In addition, the heterozygocity for BECN1 increases the frequency of spontaneous neoplasia and aids the development of lesions induced by hepatitis B virus. More recently, AMBRA1 has been described as acting as a tumor suppressor regulating MYC oncoprotein proteasomal degradation, directly interacting with PPP2. AMBRA1 mutations are associated with tumor development in various human tissues (Cianfanelli et al., 2015b). In addition, Atg4C-deficient mice showed augmented susceptibility to chemical carcinogene-induced fibrosarcoma (Marin˜o et al., 2007). Furthermore, frameshift mutations in Atg2B (Autophagy related 2B), Atg5, Atg9B (Autophagy related 9B), and Atg12 genes are correlated with gastric and colorectal cancers (Kang et al., 2009). Mutations in UVRAG result in low autophagic levels and in an increased on probability of colorectal and gastric carcinoma (Kim et al., 2008). Finally, the gene codified for the autophagosome factor MAP1LC3B

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is often deleted in breast, prostate, liver, and ovarian cancers (Jin, 2006). Beyond the involvement of several proautophagic factors in tumor suppressor regulation, the autophagic process limits tumorigenesis by inducing senescence regulating oncogenes (Young et al., 2009). In fact, autophagy induces the RAS oncogene-mediated senescence, while the overexpression of ULK3 (Unc-51 like kinase 3) induces both autophagy and senescence at the same time. Autophagy-mediated inhibition of tumors occurs through the intracellular reactive oxidative species (ROS) levels regulation by removing damaged mitochondria and protein aggregates. Autophagydefective cells with accumulated damaged mitochondria and with unfolded proteins show increased ROS levels, so causing DNA damage and SQSTM1 accumulation (Karantza-Wadsworth et al., 2007). It has been observed that, in autophagy-deficient cells, SQSTM1 accumulation activates the NF-kB pathway in order to protect damaged cells from oxidative stress, thus promoting tumorigenesis (White, 2012). Likewise, the accumulation of DNA damage and also the increase in DNA level before the onset of tumorigenesis, both favor the onset of cancer (Matheu et al., 2007). These findings define autophagy as a tumor suppressor system. Its role, however, is not limited to cellular defense: the intricate functionality of the autophagic pathway in tumorigenesis is still controversial, and further studies are necessary to clarify its role. Several studies suggest that an autophagy-mediated multiple-steps regulation occurs in carcinogenesis (Galluzzi et al., 2015). In fact, at the early stage of cancer development, autophagy mainly provides a tumor suppression function by maintaining cell homeostasis, whereas at the late stage, autophagy favors tumor progression and spread metastasis by supporting thus tumor resistance.

4.2 BCL2 Inhibits AMBRA1-Dependent Autophagy: Could It Be a Novel Mechanism to Regulate Tumorigenesis? Of note, BCL2 overexpression is a common occurrence in human cancers and can suppress the physiological activation of autophagy both in cultured cells and in mice, by acting on BECN1 (Liang et al., 1998; Pattingre et al., 2005). In addition, BCL2 localized to the mitochondria (mito-BCL2) binds AMBRA1 and inhibits its proautophagic function (Strappazzon et al., 2011). BCL2 can directly or indirectly control autophagy: its direct association with BECN1 negatively regulates the autophagic process as well as the BCL2mediated sequestration of the proautophagic AMBRA1. Consequently we could hypothesize that cellular BCL2 overexpression disrupts the tumor suppressor activity of the autophagy pathway by blocking AMBRA1

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function (Fig. 4). Since it has been demonstrated that KSHV (Kaposi’s Sarcoma-associated Herpesvirus) v-BCL2 binds to BECN1 and functions as a constitutive inhibitor of autophagy (Wei et al., 2008a), one issue to address would be investigating whether AMBRA1 can interact and be inhibited by KSHV c-BCL2. Also, bearing in mind, that AMBRA1 mutations are associated with human cancers (Cianfanelli et al., 2015b), it would be interesting to investigate whether in such cancers, BCL2 expression could be inversely correlated to AMBRA1 expression. Such putative findings would contribute to early diagnosis and to predicting prognosis for cancer. It would be of great interest to investigate whether in such cancers, cellular BCL2 and maybe viral BCL2, might exert their oncogenic activity, at least in part through autophagy inhibition. We have recently demonstrated that AMBRA1 contains a BH3-like motif along its C-terminus sequence necessary for AMBRA1 interaction with BCL2 protein and that the mutant form AMBRA1BH3-AE partially loses its capability to interact with the antiapoptotic factor. We found that AMBRA1 needs to be cleaved in order to inhibit the antiapoptotic function of BCL2. This fact opens up the question about why the full-length form is not able to exert such an effect. It is quite possible that the N-terminal part of the protein, by binding other

mito-AMBRA1 BH 3

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Fig. 4 High levels of cellular BCL2 regulate autophagy by sequestering a pool of mitochondrial AMBRA1: is it an oncogenetic mechanism? BCL2 ensures sequestration of a pool of AMBRA1 on the mitochondrial outer membrane, thus negatively regulating the autophagic process and, most likely, oncogenesis.

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Fig. 5 Reciprocal interaction between AMBRA1 and BCL2 in the regulation of apoptosis and autophagy. (A) In normal condition, full-length AMBRA1 is sequestrated by BCL2 on the outer mitochondrial membrane, thus inhibiting its proautophagic activity. In this context, it would be interesting to investigate whether MCL1 and BCL2L1 inhibit AMBRA1-dependent autophagy. Induction of apoptosis results in CASP-mediated cleavage of BECN-1. (B) AMBRA1 cleavage inactivates its autophagic function and induces a strong binding between the C-terminal fragment of AMBRA1 and mito-BCL2. This binding between BCL2 and C-terminal part of AMBRA1 accelerates the release of CYCS from mitochondria, amplifying cell death. Another point of interest would be to test whether the C-terminal part of AMBRA1 also acts on the antiapoptotic function of both BCL2L1 and MCL1.

proteins, induces a particular conformation which blocks its inhibitory action on BCL2. In conclusion, according to its full-length form or cleaved form, AMBRA1 is at the crossroad between autophagy and apoptosis (Fig. 5).

5. BCL2–AMBRA1 INTERACTION AS A NOVEL THERAPEUTIC TARGET 5.1 Autophagy Induction Through BCL2–AMBRA1 Dissociation Human cancers are “alive” due, in part, to their strong inhibitory action on cell death. In fact, some cancers, in particular, breast cancer, express high levels of the antiapoptotic factor BCL2. Proteins from the BCL2

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family are involved in MOMP, a phenomenon mediating CYCS (cytochrome C) release from mitochondria (essential in cell death induction). For this reason, MOMP is a good target in cancer therapy. Indeed, BH3 mimetics are already used in chemotherapy and have provided beneficial insights into the regulation of the BCL2–BECN1 complex as well as in identifying additional pathways involved in autophagic cell death. ABT-737 and HA14-1 also stimulate other proautophagic pathways and hence activate the nutrient sensors SIRT1/SIRTUIN1 (sirtuin 1) and PRKAA1/AMPK (protein kinase, AMP-activated, alpha 1 catalytic subunit), inhibit MTOR, deplete cytoplasmic TP53/P53 (tumor protein 53), and trigger the CHUK/IKKα (conserved helix-loop-helix ubiquitous kinase) and IKBKB/IKKβ (inhibitor of kappa light polypeptide gene enhancer in B-cells, kinase beta) kinases (Maiuri et al., 2009; Malik et al., 2011; Oltersdorf et al., 2005). We recently proposed a novel mechanism of action for ABT-737, disrupting the interaction between AMBRA1 and mito-BCL2, and thus contributing most likely to autophagy induction. Consequently, exploiting AMBRA1–BCL2 interaction could be used in order to develop novel anticancer therapies. An unanswered question is what controls the dissociation of the AMBRA1–BCL2 interaction. Further studies are needed so as to discover binding partners which can alter the reciprocal affinity of BCL2 and AMBRA1 at the mitochondria. It should be interesting, as in the case of BECN1 (Shi and Kehrl, 2010), to check whether ubiquitylation of the BH3 domain of AMBRA1 can disrupt its binding to BCL2. It should also be exciting to investigate whether posttranslational modifications on AMBRA1 (such as its phosphorylation by kinases) regulate this binding. ULK1 kinase phosphorylates AMBRA1 to dissociate it from microtubules (Di Bartolomeo et al., 2010); it would be thus necessary to check whether ULK1 is responsible for the dissociation between AMBRA1 and BCL2. Developing combination therapies between BH3 mimetics and kinase activators could provide a powerful effect on the AMBRA1–BCL2 complex. In conclusion, identification of the mechanisms, which maintain or disrupt this complex, may allow us to develop additional drugs to target it, thus ensuring an improved therapeutic outcome. Compounds which disrupt the complex would lead to increased autophagy and may be useful for the prevention and/or treatment of certain diseases, while compounds favoring AMBRA1–BCL2 binding could be interesting in contexts where autophagy needs to be delayed (for instance, in established tumors in which autophagy is considered a prosurvival mechanism).

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5.2 Is AMBRA1-Mediated Mitophagy Regulated by BCL2 Family Proteins? In the last decade, more evidence is emerging to suggest that loss of mitophagy proteins can promote tumorigenesis, underlying an oncosuppressor function of mitophagy (Cesari et al., 2003; Koop et al., 2009; Strappazzon and Cecconi, 2015b). An interesting study (Maji et al., 2015) indicated that Sabutolax, a novel BH3 mimic, and MCL1 antagonist induces a toxic mitophagy leading to an intensive cell death. Whether MCL1 inhibits AMBRA1-mediated mitophagy is an intriguing possibility. In this sense, Sabutolax could be used to induce AMBRA1/MCL1 disruption and most likely mitophagy induction, which could lead to cell death. Additionally, a common feature of all mitophagic proteins is the negative regulation by BCL2 family members: in fact, BCL2 inhibits PARK2-mediated mitophagy (Hollville et al., 2014), whereas BCL2L1 inhibits FUNDC1mediated mitophagy (Wu et al., 2014b). We demonstrated that BCL2 could inhibit the proautophagic activity of AMBRA1 (Strappazzon et al., 2011). A major unanswered question is whether BCL2 homologs can control AMBRA1-mediated mitophagy. Despite the complex nature of the role of autophagy/mitophagy in cancer, the stimulation of mitophagy appears to play a pivotal role in fighting cancer. Consequently, a better understanding of whether and how BCL2 family proteins control AMBRA1-mediated mitophagy is required.

6. CONCLUSION The intricate relationship between autophagy and apoptosis never ceases to amaze. These two processes implicated in physiological and pathological conditions controlled cellular welfare and preserve cells from undesired conditions. These two mechanisms occur via extremely controlled pathways. Although their modes of action, one degrading general or selective damaged substrates, the other inducing cellular suicide are different, both autophagy (including its several selective forms) and apoptosis proceed in order to preserve cellular homeostasis. Beyond previous studies revealing different points of convergence between apoptosis and autophagy (Fimia et al., 2013; Marin˜o et al., 2014), we, on our part, have recently discovered that the proautophagic protein AMBRA1, thanks to a switch between its full-length- or cleaved-form, plays a central role in the autophagy–apoptosis crossroads. Indeed, the scaffold autophagic protein AMBRA1 contains a

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BH3 domain with a conserved molecular mechanism for binding to the antiapoptotic BCL2 protein and most likely the antiapoptotoic BCL2 homologs MCL1 and BCL2L1. AMBRA1, in addition to its roles in macroautophagy and mitophagy (Cianfanelli et al., 2015c; Strappazzon et al., 2015), can act as a positive mediator of mitochondrial apoptosis (Strappazzon et al., 2016). The C-terminal part of AMBRA1, AMBRA1CT, generated by CASP-mediated cleavage, is able to inactivate the BCL2 antiapoptotic factor, by a direct binding via its BH3-like domain, so defining the existence of a reciprocal regulation between AMBRA1 and BCL2 at the mitochondria. Other studies will be necessary to investigate the relationship between AMBRA1 and the other members of the BCL2 family proteins like MCL1 and BCL2L1. The BH3 domain of AMBRA1 appears to have a complex and crucial role in the switch between autophagy and apoptotic cell death. Pushing or repressing the AMBRA1–antiapoptotic BCL2 family protein interactions could lead to the discovery of new and more effective anticancer therapies. Further studies are necessary to better investigate the AMBRA1 interaction with the antiapoptotic BCL2 family in order to elucidate its activity in cellular life and death switching.

ACKNOWLEDGMENTS We thank Mrs M. Acun˜a Villa and Dr. M. Bennett for secretarial and proof-reading work. This work was supported in part by a grant from the Italian Ministry of Health (Progetto Giovani Ricercatori GR2011-2012-02351433 to F.S.). Conflict of Interest: The authors declare no conflict of interest.

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CHAPTER THREE

Rationale for the Combination of Dendritic Cell-Based Vaccination Approaches With Chemotherapy Agents I. Truxova*,†, M. Hensler†, P. Skapa{, M.J. Halaska§, J. Laco¶, A. Ryska¶, R. Spisek*,†, J. Fucikova*,†,1 *2nd Faculty of Medicine and University Hospital Motol, Charles University, Prague, Czech Republic † Sotio a.s., Prague, Czech Republic { 2nd Faculty of Medicine and University Hospital Motol, Prague, Czech Republic § 3rd Faculty of Medicine and Faculty Hospital Kralovske Vinohrady, Charles University, Prague, Czech Republic ¶ Faculty of Medicine and Faculty Hospital in Hradec Kralove, Charles University, Prague, Czech Republic 1 Corresponding author: e-mail address: [email protected]

Contents 1. Introduction 2. Immunological Aspects of Anticancer Chemotherapy With Regards to DC Functions 2.1 Direct DC Stimulatory Effects of Anticancer Agents 2.2 Chemotherapeutic Agents Affecting DC Functions by Inducing ICD in Tumor Cells 3. Clinical Trials of Combined DC-Based Immunotherapy and Chemotherapy Affecting DC Functions 4. Conclusions Acknowledgments References

117 119 119 125 137 147 147 147

Abstract Owing to their central role in the initiation and regulation of antitumor immunity, dendritic cells (DCs) have been widely tested for use in cancer immunotherapy. Despite several encouraging clinical applications, existing DC-based immunotherapy efforts have yielded inconsistent results. Recent work has identified strategies that may allow for more potent DC-based vaccines, such as the combination with antitumor agents that have the potential to synergistically enhance DC functions. Selected cytotoxic agents may stimulate DCs either by directly promoting their maturation or through the induction of immunogenic tumor cell death. Moreover, they may support DC-induced adaptive immune responses by disrupting tumor-induced immunosuppressive mechanisms via selective depletion or inhibition of regulatory subsets, such as myeloid-derived suppressor

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cells and/or regulatory T cells (Tregs). Here, we summarize our current knowledge on the capacity of anticancer chemotherapeutics to modulate DC phenotype and functions and the results of ongoing clinical trials evaluating the use of DC-based immunotherapy in combination with chemotherapy in cancer patients.

ABBREVIATIONS ANXA1 annexin A1 APC antigen-presenting cell ATP adenosine triphosphate ATRA all-trans retinoic acid BLM bleomycin CCR7 chemokine (C–C motif ) receptor 7 CIK cytokine-induced killer CRT calreticulin CTX cyclophosphamide DAMP damage-associated molecular pattern ER endoplasmic reticulum FDA Food and Drug Administration FPR1 formyl peptide receptor 1 5-FU 5-fluorouracil GM-CSF granulocyte–macrophage colony-stimulating factor HMGB1 high mobility group box 1 HSP heat-shock protein hTERT human telomerase reverse transcriptase ICD immunogenic cell death iDC immature dendritic cell IFN interferon IFNAR type I IFN receptor LOX-1 lectin-type oxidized LDL receptor-1 mDC myeloid or mature dendritic cell MDSC myeloid-derived suppressor cell MHC major histocompatibility complex MM multiple myeloma MMP matrix metalloproteinase NSCLC nonsmall cell lung carcinoma PAP prostatic acid phosphatase PBMC peripheral blood mononuclear cell pDC plasmacytoid dendritic cell PD-L2 programmed death receptor-ligand 2 PGE2 prostaglandin E2 PRR pattern recognition receptor PS phosphatidylserine RAGE receptor for advanced glycation end-products SCLC small cell lung carcinoma SREC-1 scavenger receptor associated with endothelial cells TAA tumor-associated antigen TLR toll-like receptor

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TNF-α tumor necrosis factor-α Treg regulatory T cell VEGF vascular endothelial growth factor

1. INTRODUCTION Dendritic cells (DCs) are relatively heterogenous population of bone marrow-derived cells that are seeded in all tissues with the tendency to accumulate at sites of intense antigenic challenge (lymphoid organs, mucosal surfaces, the skin, etc.). DCs are recognized as the most potent antigenpresenting cells (APCs) regulating both innate and adaptive arms of the immune system. Both mice and humans have two major subsets of DCs: myeloid DCs (mDCs; also known as conventional or classical DCs) and plasmacytoid DCs (pDCs) (Stockwin et al., 2000). Upon differentiation from common bone marrow progenitors, immature DCs (iDCs) migrate to different tissues where they engulf and process extracellular material including tumor-associated antigens (TAAs) and become activated by various stimuli (microbial or endogenous). Mature DCs (mDCs) are very potent in presenting the captured antigen to T cells in lymphoid tissue leading to the induction of antitumor immune response (Palucka and Banchereau, 2012). DCs are known to have a significant impact on oncogenesis, tumor progression, and response to therapy. This has been demonstrated in various preclinical tumor models (Kroemer et al., 2013) and also in clinical studies (Senovilla et al., 2012). Owing to considerable advances related to their biology and function and development in manufacturing techniques, DCs became a promising tool for cancer immunotherapy (Stockwin et al., 2000). The main strategies to develop DC-based anticancer interventions are (1) ex vivo generation of DC vaccines, (2) targeting DC receptors by TAAs in vivo, and (3) DC-derived exosomes (Boczkowski et al., 2000; Galluzzi et al., 2014; Klechevsky et al., 2010; Viaud et al., 2010). Ex vivo DCs are mainly generated through in vitro differentiation of peripheral blood mononuclear cells (PBMCs) or previously separated monocytes in the presence of granulocyte–macrophage colony-stimulating factor (GMCSF) and IL-4 (Curti et al., 2004). Autologous DCs can be then expanded, matured, and loaded with tumor antigens and administered back to cancer patients to generate antitumor immunity. The matured status of DCs and their ability to produce IL-12p70 are critical for the induction of CD8+ T cell responses (Czerniecki et al., 2007) and the clinical responses in cancer patients treated by DC-based vaccines (Draube et al., 2011).

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DCs have been tested in cancer immunotherapy clinical trials for two decades. The early clinical trials of DC-based cancer immunotherapy established the general safety and tolerability of this cancer treatment and demonstrated the low risk of toxicity compared to other treatment options (e.g., chemotherapy and radiotherapy). Since the publication of the first DC vaccination trial in melanoma patients in 1995 (Nestle et al., 1998), autologous DCs have been employed in immunotherapy for several tumor types, including melanoma, lymphoma, prostate cancer, renal cell carcinoma (RCC), glioblastoma, etc. (Anguille et al., 2014; Cao et al., 2014; Hsu et al., 1996; Nakai et al., 2010; Podrazil et al., 2015; Vacchelli et al., 2013b). Recently, some clinical trials have yielded encouraging clinical outcomes. The treatment of metastatic prostate cancer with sipuleucel-T (Provenge), cellular product based on enriched blood APCs that are cultured with a fusion protein of prostatic acid phosphatase (PAP) and granulocyte–macrophage colony-stimulating factor (GM-CSF), resulted in a 4-month prolonged median survival in phase III trials. Sipuleucel-T was in 2010 approved by Food and Drug Administration (FDA) for the treatment of metastatic prostate cancer. Additional strategies exploring the potential benefit of DC-based immunotherapy in metastatic prostate cancer are in advanced stages of the clinical testing (Podrazil et al., 2015). Despite some encouraging clinical applications, clinically effective DC-based immunotherapy as a standalone treatment remains a distant goal. The limited success of DC-based immunotherapy approaches in advanced cancer patients might be due to the establishment of tumorinduced immunosuppression (de Visser et al., 2006). Experimental evidence supports the fact that the goal of the immunotherapy in the late stages is not necessarily the complete eradication of tumor cells but rather the establishment of an equilibrium between the host immune system and the tumor cells (Dunn et al., 2004). Therefore, there is an emerging evidence that the optimal benefit of DC-based immunotherapy might be achieved by combinations with other antitumor therapies that enhance DC function. Treatment of metastatic cancer essentially relies on cytotoxic drugs that kill tumor cells or hinder their proliferation. Although a primary goal of anticancer chemotherapy is the tumor mass reduction, it is now clear that antitumor activities of chemotherapy also rely on several other on-target and also off-target effects, especially directed to the host immune system, that

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reduce the immunosuppressive activity of malignant cells and cooperate for successful tumor eradication. Clinically employed anticancer agents may stimulate immunosurveillance by acting on tumor cells (1) by increasing the expression or presentation of TAAs on the surface of tumor cells (Jackaman et al., 2012), (2) by increasing the susceptibility of malignant cells to the cytotoxic activity of immune cells (Vandenabeele et al., 2010), and (3) by causing tumor cells to emit danger signals during cell death that are sensed by DCs as adjuvants leading to the stimulation of antitumor immune responses (Kroemer et al., 2013). Moreover, the anticancer chemotherapeutics may mediate immunostimulatory effects by directly affecting the immune system (1) by influencing the homeostasis of the hematopoietic compartment through transient lymphodepletion (Finn, 2012), (2) by subverting tumor-induced immunosuppressive mechanisms through selective inhibition or killing of suppressive immune populations, such as myeloidderived suppressor cells (MDSCs) and regulatory T cells (Tregs) (Ghiringhelli et al., 2007; Vincent et al., 2010), and (3) by exerting direct stimulatory effects on immune effectors (Zitvogel et al., 2013). Here, we discuss the impact of anticancer chemotherapy on DC phenotype and functional characteristics with a focus on indirect DC stimulatory mechanisms mediated by distinct danger signals released from dying tumor cells treated by chemotherapeutics known as immunogenic cell death (ICD) inducers. The immunomodulatory side effects of anticancer agents could potentially increase the therapeutic efficacy of DC-based interventions. Therefore, we will summarize the results of current clinical trials evaluating the use of DC-based immunotherapy in combination with chemotherapy in cancer patients.

2. IMMUNOLOGICAL ASPECTS OF ANTICANCER CHEMOTHERAPY WITH REGARDS TO DC FUNCTIONS 2.1 Direct DC Stimulatory Effects of Anticancer Agents The administration of some cytotoxic drugs can directly target DCs leading to an improved DC phenotype (Table 1b). Low doses of paclitaxel, vinblastine, etoposide, and methotrexate directly upregulate the expression of CD40, CD80, CD86, and MHC class II molecules and induce the production of proinflammatory cytokines by DCs (Shurin et al., 2009; Tanaka et al., 2009a). The injection of vinblastine into the skin of mice triggers in situ maturation of skin-resident DCs and boosts humoral and cellular

Table 1a An Overview of ICD-Inducing Anticancer Chemotherapeutics Affecting DC Functions Indirect Effects on DC Functions Through the Emission of ICDDirect Effects on DC Associated DAMPs From Tumor Cells Functions Agent Class

ICD Doxorubicin, inducers idarubicin, epirubicin, daunorubicin

Mitoxantrone

CTX

Indications

References

32, 33, 41, 46, 67, 69–71, 93, 94, 110

Anthracyclines ATP-driven recruitment of DCs into the tumor; activation of NLRP3 inflammasome; CRT-mediated " of phagocytosis; HSP-dependent " of maturation-associated markers and release of proinflammatory cytokines; " processing and crosspresentation of TAAs by DCs

Induce various levels of DC maturation—" CD40, " IL-1β, IL-6, IL-12, and TNF-α; doxorubicin enhances antigen presentation by DCs

Leukemia, lymphoma, uterine, ovarian, and breast malignancies

Topoisomerase ATP-driven recruitment of DCs inhibitors into the tumor; activation of NLRP3 inflammasome; CRT-mediated " of phagocytosis; HSP-dependent " of maturation-associated markers and release of proinflammatory cytokines; " processing and crosspresentation of TAAs by DCs

Induce various levels of DC maturation—" CD40, " IL-1β, IL-6, IL-12, and TNF-α

Multiple sclerosis, 4, 33, 48, 60, 61, 71, acute leukemia, breast carcinoma, 97, 98 non-Hodgkin’s lymphoma, prostate cancer

Alkylating agents

Induces preferential Lymphoma, expansion of CD8α+ DCs; leukemia, solid tumors stimulates tumor and regional LN infiltration by DCs; DCs isolated from CTX-treated mice in vitro inhibit suppressive functions of Tregs

ATP-driven recruitment of DCs into the tumor; activation of NLRP3 inflammasome; CRT-mediated " of phagocytosis; HSP-dependent " of maturation-associated markers and release of proinflammatory cytokines; " processing and crosspresentation of TAAs by DCs

4, 42, 43, 48, 60, 61, 97, 98

Oxaliplatin

Platinum compounds

Inhibits the expression of ATP-driven recruitment of DCs into the tumor; activation of NLRP3 PD-L2 on DCs inflammasome; CRT-mediated " of phagocytosis; HSP-dependent " of maturation-associated markers and release of proinflammatory cytokines; " processing and crosspresentation of TAAs by DCs

Colorectal cancer 4, 44, 48, 50, 60, 61, 97, 98

Bortezomib

Proteasome inhibitors

X ATP-driven recruitment of DCs into the tumor; activation of NLRP3 inflammasome; CRT-mediated " of phagocytosis; HSP-dependent " of maturation-associated markers and release of proinflammatory cytokines; " processing and crosspresentation of TAAs by DCs

4, 48, 52 Multiple myeloma, mantle cell lymphoma

Bleomycin

X Glycopeptides ATP-driven recruitment of DCs into the tumor; activation of NLRP3 inflammasome; CRT-mediated " of phagocytosis; HSP-dependent " of maturation-associated markers and release of proinflammatory cytokines; " processing and crosspresentation of TAAs by DCs

4, 48, 60, Hodgkin’s lymphoma, penile 61, 97, 98 cancer, testicular cancer, squamous carcinomas of the head and neck, cervix, and vulva

Abbreviations: ATP, adenosine triphosphate; CRT, calreticulin; CTX, cyclophosphamide; DAMPs, damage-associated molecular patterns; DC, dendritic cell; HSP, heat-shock protein; ICD, immunogenic cell death; LN, lymph node; TAAs, tumor-associated antigens; TNF-α, tumor necrosis factor-α; Treg; regulatory T cell.

Table 1b An Overview of Anticancer Chemotherapeutics Directly Affecting DC Functions Indirect Effects on DC Functions Through the Emission of ICDAssociated DAMPs From Tumor Cells Direct Effects on DC Functions Agent Class

Indications

References

Paclitaxel NonICD inducers

Taxanes

X

32, 33, 36, " CD40, CD80, CD86, and MHC class II; Kaposi’s sarcoma, lung, ovarian, breast, 37 " proinflammatory cytokines; and head and neck " DC-mediated antigen presentation cancers

Docetaxel

Taxanes

X

" Cell cycle regulator p21wafl1/clip1 35 NSCLC, breast, associated with a favorable DC phenotype ovarian, and prostate and correlates with increased expression of cancers CD83 and CD86 on DCs

Vinblastine

Antimicrotubule agents

X

32, 33, 34 " CD40, CD80, CD86, and MHC class II; Hodgkin’s lymphoma, " proinflammatory cytokines; in situ NSCLC, breast, maturation of skin-resident DCs testicular, and head, and neck cancers

Vincristine

Antimicrotubule agents

X

" DC-mediated antigen presentation; increases the abundance of CD83 + and type I DCs

Etoposide

Topoisomerase inhibitors

X

" CD40, CD80, CD86, and MHC class II; Kaposi’s and Ewing’s 32, 33 " proinflammatory cytokines sarcomas, leukemia, glioblastoma, lung, ovarian, and gastrointestinal cancers

Non-Hodgkin’s lymphoma

32, 36, 37, 38

Methotrexate Antimetabolites

X

" CD40, CD80, CD86, and MHC class II; Several solid and hematopoietic " proinflammatory cytokines; " tumors DC-mediated antigen presentation

Gemcitabine Antimetabolites

X

" cross-presentation of tumor antigens to CD8+ T cells; " CD14+ monocytes, CD11c+ myeloid cells, and CD123+ plasmacytoid DCs

Decitabine

X DNA methyltransferase inhibitors

32, 33, 36, 37

NSCLC, pancreatic, 32, 36, 37, 39 bladder, and breast cancers

Promotes the differentiation of tumorLeukemia and derived CD11b + cells into mature F4/80/ myelodysplastic CD11c/MHC class II+ APCs producing # syndrome amounts of IL-13, IL-10, PGE2, and proangiogenic mediators (VEGF, MMP-9)

40

Abbreviations: APC, antigen-presenting cell; DAMPs, damage-associated molecular patterns; DC, dendritic cell; ICD, immunogenic cell death; MHC, major histocompatibility complex; MMP-9, matrix metalloproteinase-9; NSCLC, nonsmall cell lung cancer; PGE2, prostaglandin E2; VEGF, vascular endothelial growth factor.

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immune responses (Tanaka et al., 2009b). Furthermore, docetaxel-based chemotherapy enhances the expression of the cell cycle regulator p21wafl1/cip1 which is associated with a favorable DC phenotype (Liu et al., 2013). The positive effect on DC-mediated antigen presentation has been documented for paclitaxel, methotrexate, vincristine, and gemcitabine (Galluzzi et al., 2012; Nowak et al., 2003; Shurin et al., 2009; Tanaka et al., 2009b). Another aspect of DC biology affected by applied anticancer chemotherapy is DC differentiation and the distribution of different DC subsets. Vincristine-based chemotherapy, in combination with doxorubicin and glucocorticoids, increases the abundance of CD83+ and type I DCs in patients with multiple myeloma (MM) (Kovarova et al., 2007) and an overall increase of CD14+ monocytes, CD11c+ myeloid cells, and CD123+ pDCs was observed in patients with advanced pancreatic cancer receiving gemcitabine (Soeda et al., 2009). A culture of tumorinfiltrated CD11b+ myeloid cells from tumor-bearing mice in the presence of DNA methyltransferase inhibitor 50 -aza-20 deoxycytidine (decitabine) and GM-CSF promotes their differentiation into mature F4/80/CD11c/MHC class II+ APCs. These tumor-derived myeloid APCs produce reduced amounts of immunosuppressive (IL-13, IL-10, prostaglandin E2 (PGE2)) and proangiogenic (vascular endothelial growth factor (VEGF), matrix metalloproteinase-9 (MMP-9)) mediators (Daurkin et al., 2010). Moreover, cytotoxic drugs endowed with the capacity to stimulate DCs through the induction of ICD-associated DAMP (damage-associated molecular pattern) release from dying tumor cells (discussed in details in the next section) (Table 1a), have been also shown to affect DC functions directly. Intriguingly, anthracyclines and mitoxantrone induce DC maturation by upregulating the expression of CD40 (doxorubicin, idarubicin) and by stimulating the production of proinflammatory cytokines, such as IL-1β (daunorubicin, epirubicin, idarubicin, mitoxantrone), IL-6 (daunorubicin, epirubicin, mitoxantrone), IL-12 (idarubicin, mitoxantrone), and tumor necrosis factor-α (TNF-α) (idarubicin, mitoxantrone) (Tanaka et al., 2009a). Doxorubicin was also shown to promote the ability of DCs to present antigens to T cells in vitro by upregulating antigen-processing machinery gene components, costimulatory molecules, and IL-12p70 (Kaneno et al., 2009; Shurin et al., 2009). Recently, the immune response against anthracycline-treated tumor cells driven by intratumoral DCs was found to be dependent on ANXA1 (annexin A1)–FPR1 (formyl peptide receptor 1) signaling. FPR1 and its ligand, ANXA1, promote stable interactions between dying tumor cells and human or murine DCs. Lack or inhibition

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of FPR1 signaling leads to defective intratumoral DC maturation and weak antitumor T cell responses, abolishing the efficacy of anthracycline-based chemotherapy (Vacchelli et al., 2015). The application of cyclophosphamide (CTX) in vivo affects DC homeostasis leading to the preferential expansion of CD8α+ DCs, the main subset involved in the cross-presentation of cell-derived antigens (Schiavoni et al., 2011). Moreover, CTX induces the expansion of DCs in peripheral blood and stimulates tumor and regional lymph node infiltration by DCs. DCs isolated from CTX-treated mice in vitro inhibit suppressive functions of Tregs leading to the enhancement of effector T cell functions (Nakahara et al., 2010). This is in line with the fact that CTX is well known for its ability to decrease the number and immunosuppressive functions of Tregs, especially when used at relatively low, so-called metronomic, doses (Ghiringhelli et al., 2007). Suppressive immune cells present within the tumor (including not only Tregs, but also MDSCs, tumor-associated macrophages (TAMs), etc.) contribute largely to tumor suppressive microenvironment and tumor escape, thus hampering the desired effect of various antitumor therapies (Gabrilovich and Nagaraj, 2009). Strategies targeting these cell populations that may increase the benefit of cancer treatment have been discussed elsewhere (Galluzzi et al., 2012; Zitvogel et al., 2008) and they are beyond the scope of this review which focuses primarily on the combination of DC-based immunotherapy and cytoreductive chemotherapy with secondary immunostimulatory activity. Recent study showed that in vitro exposure to oxaliplatin reduces the expression of T cell inhibitory molecule programmed death receptor-ligand 2 (PD-L2) on both human DCs and tumor cells. Downregulation of PD-L2 resulted in enhanced antigen-specific T cell proliferation and Th1 cytokine secretion as well as enhanced recognition of tumor cells by T cells (Lesterhuis et al., 2011). Taken together, these results illustrate that the immunomodulatory side effects of anticancer chemotherapeutic agents can increase the therapeutical potential of immune-based interventions and they should be considered in the design of effective immunotherapy protocols.

2.2 Chemotherapeutic Agents Affecting DC Functions by Inducing ICD in Tumor Cells 2.2.1 Concept of ICD The anticancer benefits of chemotherapy treatment have long been considered to be a consequence of direct cytotoxicity or permanent arrest of the cell cycle. The chemotherapy was thought to nonspecifically target rapidly

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dividing cells leading to the assumption that this therapy can also target proliferating immune cells and result in immunosuppression. It has to be mentioned that many chemotherapeutic drugs used in the clinic are low/nonimmunogenic or even have notable immunossupressive side effects—either directly, by inhibiting or killing effector cells, or indirectly, by provoking anergy or immune paralysis. Therefore, the role of the immune system in anticancer therapy has been widely neglected and overlooked (Gebremeskel and Johnston, 2015). However, it is now widely accepted that several chemotherapeutics and anticancer agents can activate the tumor immunosurveillance. One of the mechanisms participating in eliciting an antitumor immune response is the ability of some chemotherapeutics to enhance the immunogenicity of tumor cells by inducing ICD, a cell death modality that stimulates an antitumor immune response, as documented by many experiments in immunocompetent mice vaccinated with tumor cells succumbing to ICD (Galluzzi et al., 2012; Kroemer et al., 2013; Mikyskova et al., 2016). Only a few stimuli are able to trigger bona fide ICD. These include some chemotherapeutics that are employed in the clinic, including anthracyclines (doxorubicin, epirubicin, idarubicin) (Casares et al., 2005; Fucikova et al., 2011; Obeid et al., 2007), mitoxantrone (Casares et al., 2005; Obeid et al., 2007), oxaliplatin (Martins et al., 2011; Tesniere et al., 2010), CTX (Ziccheddu et al., 2013), bortezomib (Spisek et al., 2007), and bleomycin (BLM) (Bugaut et al., 2013). Furthermore, ICD can be triggered by certain physical modalities such as UV-C irradiation, hypericin-based photodynamic therapy, and high hydrostatic pressure (Fucikova et al., 2014; Galluzzi et al., 2013; Garg et al., 2012a) and finally some oncolytic viruses are also intrinsically endowed with the capacity to initiate ICD (Vacchelli et al., 2013a). The ability of these inducers to stimulate antitumor immune response depends on the establishment of adaptive stress responses that promote the coordinated emission of endogenous danger signals from dying cells (Zitvogel et al., 2010). These molecules are known as DAMPs and operate on a series of receptors expressed by DCs to stimulate the adaptive arm of the immune system (Krysko et al., 2013). Several DAMPs have been identified as a hallmark features of ICD, namely, (1) the preapoptotic surface exposure of the endoplasmic reticulum (ER) chaperone calreticulin (CRT) and heat-shock proteins (HSPs) HSP70 and HSP90; (2) passively released molecules such as high mobility group box 1 (HMGB1); and (3) the secretion of adenosine triphosphate (ATP). Moreover, certain ICD-inducing chemotherapeutic agents can stimulate tumor cells to produce type I interferons (IFNs). Althought, type I IFNs

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do not represent DAMPs per se, they are endowed with strong immunostimulatory activity and they appear to be critical for chemotherapy-induced cell death to be perceived as immunogenic (Garg et al., 2015; Kroemer et al., 2013; Krysko et al., 2012). The role of DAMPs in the activation of immune system has been demonstrated in numerous in vitro tumor cell line models (Garg et al., 2012b; Obeid et al., 2007), in vivo mice immunization experiments (Krysko et al., 2012), and the latest reports indicate that monitoring DAMPs or DAMP-associated stress responses in cancer patients may have prognostic or predictive value (Fucikova et al., 2015, 2016). 2.2.2 Anthracyclines (Doxorubicin, Idarubicin, Epirubicin) and Mitoxantrone Anthracyclines, including doxorubicin, idarubicin, and epirubicin, were initially described in 1963 as antibiotics. For their strong efficacy in anticancer therapy, they have been used for many years for the treatment of multiple solid and hematological malignancies (leukemia, lymphoma, uterine, ovarian, and breast cancer) (Zhang et al., 2015). Anthracyclines and anthracenediones (mitoxantrone) exert their cytostatic and cytotoxic functions by intercalating between base pairs of the DNA/RNA strand and inhibiting topoisomerase II, thus preventing DNA/RNA synthesis and replication of rapidly growing cancer cells which finally leads to the cell death (Rabbani et al., 2005). This type of cell death exhibits the key characteristics of ICD including preapoptotic CRT surface exposure (ecto-CRT) (Obeid et al., 2007), the production of type I IFNs (Sistigu et al., 2014), the secretion of ATP, and the passive release of HMGB1 during late stage apoptosis (Apetoh et al., 2007a). Ecto-CRT on dying tumor cells acts as “eat me” signal for APCs promoting the intensive phagocytosis of apoptotic bodies which subsequently leads to the activation of DCs. Fully mature DCs can induce T cell-mediated immune responses against dying tumor cells (Obeid et al., 2007). HMGB1 released from dying tumor cells binds to pattern recognition receptors (PRRs) on DCs and is required for optimal antigen presentation by DCs (Apetoh et al., 2007a). Moreover, anthracycline-based chemotherapy can promote the recruitment of myeloid cells, including neutrophils, monocytes, macrophages, and cells with a DC-like phenotype into the tumor bed depending on the production of urokinase, IL-8, and monocyte chemoattractant protein 1 (MCP-1) (Niiya et al., 2003), the CCL2/CCR2 signaling axis (Ma et al., 2014), or ATP released from dying tumor cells (Elliott et al., 2009).

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2.2.3 Cyclophosphamide CTX is a nitrogen mustard alkylating agent from the oxazophorine group. In 1958, CTX was first assayed in clinical trials for the treatment of cancer and in 1959, it became the eighth cytotoxic anticancer agent approved by FDA. CTX still remains one of the most widely used drugs for the treatment of a variety of diseases, including hematological and solid malignancies and autoimmune disorders. CTX has antiangiogenic properties and induces the activation of immune system (Sistigu et al., 2011). The immunostimulatory effects of CTX are partly mediated by its ability to induce ICD. Immunogenic properties of CTX are characterized by rapid preapoptotic translocation of CRT to the cell surface, type I IFN production, ATP secretion, and passive release of HMGB1 from tumor cells (Bezu et al., 2015). CTX represents the oldest and most extensively studied example of chemotherapy exhibiting potent immunomodulatory effects. However, its biological activities are highly dose dependent. 2.2.4 Oxaliplatin Oxaliplatin is platinum-based antineoplastic agent which has shown antitumor activity in a wide range of murine and human tumor cell lines in preclinical studies (Di Francesco et al., 2002). It has been approved for use in combination with 5-fluorouracil (5-FU) and folinic acid (FOLFOX regimen) for the therapy of advanced colorectal carcinoma (Tesniere et al., 2010). Platinum-based chemotherapeutic agents (oxaliplatin, cisplatin, carboplatin) exert their anticancer effects via the formation of similar DNA platinum adducts or intrastrand and interstrand cross links. At the site of bound platinum adducts, DNA denaturates which leads to the formation of strand breaks and ultimately to the inhibition of DNA/RNA synthesis and transcription followed by the reduction of tumor cell replication and the activation of apoptotic pathways (Stojanovska et al., 2015). Unlike cisplatin and carboplatin, oxaliplatin is regarded as a potent stimulator of ICD and the presentation of DAMPs (Lesterhuis et al., 2010; Tesniere et al., 2010). Oxaliplatin prompts the translocation of CRT and HSP70 to the surface of dying tumor cells, the release of HMGB1, the secretion of ATP, and the production of type I IFNs, thereby promoting their recognition by APCs for eventual presentation to effector T cells (Krysko et al., 2012; Tesniere et al., 2010). Oxaliplatin treatment increases HLA I expression in tumor cells which helps immune system to recognize and efficiently remove these cells (Liu et al., 2010).

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2.2.5 Bortezomib In 2003, bortezomib (Velcade) was approved by FDA for therapy of MM and represents the first proteasome inhibitor to be approved for clinical use. Bortezomib is now used as a front-line treatment for newly diagnosed MM patients and for the treatment of relapsed/refractory MM and mantle cell lymphoma (Chen et al., 2011). The ubiquitin–proteasome pathway is responsible for the degradation of misfolded and mutated proteins and plays an essential role in regulating homeostatic and various cellular events including those involved in tumorigenesis (Adams, 2004). Bortezomib inhibits a key regulator of intracellular protein degradation, the 26S proteasome, leading to the induction of cell cycle arrest and subsequently apoptotic cancer cell death. Bortezomib has been decribed as a bona fide ICD inducer and its immunogenic properties have been demonstrated in mice with B16 tumor treated with DC vaccine. Tumor outgrowth was significantly decreased when the mice were treated with the combination of bortezomib and DCs. Further studies revealed that the therapeutic effect was dependent on NK cells and CD8+ T cells (Schumacher et al., 2006). As an ICD inducer, bortezomib triggers the exposure of CRT and HSPs (HSP60, HSP70, and HSP90) on the surface of dying tumor cells, the release of HMGB1, and the production of type I IFNs (Bezu et al., 2015; Chang et al., 2012; Cirone et al., 2012; Spisek et al., 2007). Tumor cells treated by bortezomib are efficiently phagocytosed by DCs and induce cell contact-dependent activation of DCs. Activated DCs then stimulate the IFN-γ production by antitumor-specific T lymphocytes (Spisek et al., 2007). These effects are predominantly mediated by HSPs and CRT present on the surface of bortezomib-treated tumor cells. Despite the immunogenic effects mentioned earlier, the application of bortezomib downregulates the expression of major histocompatibility complex (MHC) molecules on DCs (Schumacher et al., 2006) and reduces the viability of some immune cells, such as myeloid and pDCs and activated T cells. 2.2.6 Bleomycin BLM is an antitumor antibiotic glycopeptide produced by the bacterium Streptomyces. BLM causes breaks in DNA, similar to those obtained with radiotherapy. This DNA damage has been demonstrated to be mediated by the induction of oxidative stress. BLM is indicated for the paliative treatment for Hodgkin’s lymphoma, penile cancer, testicular cancer, and squamous

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carcinomas of the head and neck, cervix, and vulva (Bugaut et al., 2013; Pol et al., 2015). It was recently shown that BLM is able to trigger ICD. In particular, it is able to induce reactive oxygen species (ROS)-mediated ER stress and autophagy, which results in the surface exposure of CRT and ERp57, the release of HMGB1 and ATP. The antitumor immunity triggered by BLM relies on CRT, CD8+ T cells, and IFN-γ (Bugaut et al., 2013). 2.2.7 Modulating DC Functions by Immunogenic Chemotherapy-Induced DAMPs 2.2.7.1 Sensing ICD-Associated Danger Signals by DCs

Activation of antitumor immune response must be preceded by recognition of tumor cells as foreign elements. Tumor cells differ from normal cells by the expression of altered-self or neoantigens that arise as a consequence of genetic instability and high mutation rate in transformed cells (Linnemann et al., 2015). TAA uptake is followed by the maturation of DCs and the migration of large number of mature DCs to draining lymph nodes. The mature DCs express peptide–MHC complexes on their surface as well as appropriate costimulatory molecules (CD80, CD86, CD40). This allows the priming of CD4+ T helper and CD8+ cytotoxic T lymphocytes, the activation of B cells and the initiation of an adaptive immune response. DCs play a crucial role in sensing danger signals released or exposed on dying, stressed, or injured cells including tumor cells (DAMPs). Thus they have a central position in danger model of the immune response proposed in 1994 by Polly Matzinger which states that the immune system can distinguish between dangerous and innocuous endogenous signals (Matzinger, 1994). As mentioned earlier, the release or exposure of DAMPs (CRT, HSPs, HMGB1, ATP) and the production of type I IFNs can be triggered by distinct chemotherapeutics (and modalities) via the induction of specific type of cell death called ICD. The common feature of emitted DAMPs is to trigger several PRRs expressed on DCs and their type, duration, and timing regulate DC functions, such as maturation (Table 2). Consequently, these signals determine the quality and quantity of costimulation provided by DCs and thus have the power to define the outcome of T cell immunity. DCs recognize ICDassociated DAMPs by distinct PRRs: (1) toll-like receptors (TLRs) (namely TLR2 and TLR4) (Garg et al., 2012a,b; Krysko et al., 2012; van Eden et al., 2012); (2) scavenger receptors (scavenger receptor associated with endothelial cells (SREC-1) (Murshid et al., 2014) and lectin-type oxidized LDL receptor 1 (LOX-1)); (3) type I IFN receptor (IFNAR) (Zitvogel et al., 2015);

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Table 2 An Overview of DAMPs Associated With Chemotherapy-Induced ICD and Their Impact on DC Functions ICD-Associated DAMPs DC Receptors Effect on DC Functions References

CRT

CD91

A potent “eat me” signal crucial for antitumor immunity; increases susceptibility to phagocytosis by DCs

48, 60–63, 95, 96

CD91, TLR2, HSP70, TLR4, LOX-1, HSP90, SREC-1, CD14 HSP60, HSP72, GP96

Mediate DC maturation and 52, 60, 61, 87, 89, activation (" maturationassociated molecules—CD83, 98–101, 112 CD80, CD86, CD40, " production of TNF-α, IL-1β, IL-6, and IL-12); form complexes with tumor antigens facilitating antigen processing and presentation by DCs

ATP

P2Y2 and P2X7

60–62, 90, Act as “find me” signal and enhances the accumulation of 92, 93 DCs in the tumor; causes activation of NLRP3 inflammasome leading to IL-1β release from DCs

HMGB1

TLR2, TLR4, RAGE

56, 60–62, “Antigen processing” signal 86, 102, 103, crucial for efficient crosspresentation of TAAs by DCs; 110, 111 stimulates the production of TNF-α, IL-1, IL-6, and IL-8 from myeloid cells; can cause DC maturation

Type I IFNs

IFNAR

42, 60, 61, Stimulate DC maturation, 67, 88, 104 antigen processing, and presentation by DCs; generate DCs with cytotoxic activity

Abbreviations: ATP, adenosine triphosphate; DAMPs, damage-associated molecular patterns; DC, dendritic cell; CRT, calreticulin; HMGB1, high mobility group box 1; HSP, heat-shock protein; ICD, immunogenic cell death; IFN, interferon; IFNAR, type I interferon receptor; LOX-1, lectin-type oxidized LDL receptor 1; RAGE, receptor for advanced glycation end-products; SREC-1, scavenger receptor associated with endothelial cells; TAAs, tumor-associated antigens; TNF-α, tumor necrosis factor-α; TLR, toll-like receptor.

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Fig. 1 Schematic representation of the impact of DAMPs on DC functions. DAMPs emitted from tumor cells during ICD include CRT, HSPs, ATP, HMGB1, and type I IFNs. These molecules bind to respective cognate receptors like CD91 (for CRT and HSPs), TLR2/TLR4 (for HSPs and HMGB1), RAGE (for HMGB1), P2Y2/P2X7 (for ATP), SREC-1 and LOX-1 (for HSPs), and IFNARs (for type I IFNs) on the surface of immature DCs. This leads to an enhanced engulfment of tumor cells and antigen processing by DCs, DC maturation characterized by upregulation of costimulatory molecules such as CD80, CD83, CD86, HLA-DR, and CD40, and by an increase in proinflammatory cytokine production. Matured DCs efficiently present TAAs in the context of MHC molecules to T cells inducing antitumor T cell responses. Abbreviations: ATP, adenosine triphosphate; CRT, calreticulin; DAMP, damage-associated molecular patterns; DC, dendritic cell; HMGB1, high mobility group box 1; HSP, heat-shock protein; ICD, immunogenic cell death; IFN, interferon; IFNAR, type I IFN receptor; LOX-1, lectin-type oxidized LDL receptor 1; MHC, major histocompatibility complex; RAGE, receptor for advanced glycation endproducts; SREC-1, scavenger receptor associated with endothelial cells; TAA, tumorassociated antigen; TLR, toll-like receptors.

(4) CD14 (Asea et al., 2000); (5) receptor for advanced glycation end-products (RAGE) (Krysko et al., 2012); (6) CD91 (known as low-density lipoprotein receptor-related protein-1 (LRP-1)) (Garg et al., 2012b); and (7) purinergic receptors P2Y2 and P2X7 (Garg et al., 2012b; Ghiringhelli et al., 2009) (Fig. 1). The role of these receptors and their activation by different immunogenic chemotherapy-induced DAMPs in regulating DC functions will be discussed in next section. 2.2.7.2 Recognition and Phagocytosis of Tumor Cells by DCs

Phagocytic removal of apoptotic cells is a standard proces occuring in vivo. This is thought to be due to the presence of “eat me” signals like

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phosphatidylserine (PS) on the surface of dying cells that recruit phagocytes to the sites of cell death leading to their clearance. However, PS is rather a tolerogenic signal which does not induce the immune response (Steinman et al., 2000). In contrast, ICD inducers, including some chemotherapeutics, have the capacity to trigger the release of specific “find me” signals and the exposure of “eat me” signals from dying tumor cells that are sensed by DCs resulting in the recruitment of DCs to the sites of cell death and the recognition and phagocytosis of such “immunogenic” tumor cells. The recruitment of myeloid cells, icluding DCs, to the sites of extensive apoptosic cell death is potentiated by ATP, a well-known “find me” signal representing one of the hallmarks of ICD. Tumor cells exposed to chemotherapeutic agents inducing ICD secrete ATP during the phase of PS exposure on the plasma membrane in an autophagy-dependent manner and ATP in the extracellular space is required for the generation of an effective chemotherapy-elicited anticancer immune response (Martins et al., 2014). Ma et al. demonstrated that anthracycline-based chemotherapy stimulates the ATP-driven and purinergic P2Y2 receptor-dependent accumulation of CD11c+CD11b+Ly6Chigh DCs in the tumor and that this subset of cells is particularly efficient at presenting tumor antigens to CD8+ T cells both in vitro and in vivo (Ma et al., 2013). The recognition and efficient engulfment of dying tumor cells by phagocytes is mediated by “eat me” signals present on the surface of tumor cells. It was shown that immature murine bone marrow-derived DCs were capable of engulfing anthracycline-treated tumor cells in vitro and cross-present tumor-derived antigens to tumor antigen-specific MHC class I and II-restricted T cells (Casares et al., 2005). In line with these data, another study documented that the treatment with doxorubicin or epirubicin increased the uptake of tumor cells by human PBMCs (Brusa et al., 2008). One of the most important “eat me” signals is ecto-CRT. The presence or absence of ecto-CRT was identified as a major biochemical difference between immunogenic and non-ICD. Ecto-CRT binds CD91 on phagocytes leading to induction of phagocytosis and efficient clearance of dying tumor cells (Gardai et al., 2005). Importantly, CRT–CD91 interaction leads to the signaling through NF-κB in DCs and to the release of inflammatory cytokines, such as TNF-α, IL-6, IL-1β, and IL-12. This cytokine milieu induced by CRT exposure leads to Th17 priming in immunosuppressive microenvironment (Pawaria and Binder, 2011). Cells deficient in CRT undergo apoptosis but are not efficiently removed by phagocytes, suggesting that CRT exposure is a major molecular determinant for

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phagocytosis (Obeid et al., 2007). Recently, high expression of CRT on tumor cells in nonsmall cell lung carcinoma (NSCLC) patients was shown to be associated with a higher density of infiltrating mature DCs and effector memory T cells subsets, suggesting that CRT triggers the activation of adaptive immune response in vivo in the tumor microenvironment (Fucikova et al., 2016). The role of an “eat me” signal was documented also for HSP90 as the expression of HSP90 on the surface of bortezomib-treated myeloma cells contributed to the adhesion of tumor cells to DCs in vitro (Spisek et al., 2007). 2.2.7.3 DC Activation

In response to various microbial and endogenous stimuli (such as DAMPs), iDCs undergo a process of maturation which is accompanied by several events including (1) changes in morphology, (2) loss of phagocytic capacity, (3) upregulation of costimulatory molecules, such as CD80, CD86, CD40, and OX40L, (4) upregulation of MHC class II molecules, (5) enhanced secretion of cytokines and chemokines, and (6) the acquisition of chemokine receptor expression allowing them to migrate to lymph nodes, such as chemokine (C–C motif ) receptor 7 (CCR7). Finally, mature DCs (mDCs) are able to efficiently present tumor antigens to T cells leading to the induction of antitumor-specific immune response. The maturation of DCs can be induced by HSPs, the ICD-associated DAMPs belonging to a family of highly conserved chaperone proteins that play an important role in folding newly synthesized proteins as well as refolding of proteins affected due to various stress conditions (Garg et al., 2010). Although located primarily in intracellular compartments, HSPs can translocate to the cell surface in response to stress signals triggered by ICD inducers. Ecto-HSP70 and HSP90 exhibit potent immunostimulatory activity and determine the immunogenicity of dying tumor cells in some models (Tesniere et al., 2008; Udono and Srivastava, 1994). This is due to their ability to bind various receptors on DCs, such as CD91, TLR2, and TLR4 leading to the phenotypical and functional maturation of DCs. It was documented that coculture of immature DCs with bortezomibtreated tumor cells expressing high levels of ecto-HSP90 led to a contactdependent upregulation of maturation-associated molecules like CD83, CD80, and CD86 on DCs (Spisek et al., 2007). Similarly, the interaction of DCs with anthracycline (idarubicin, doxorubicin)-treated tumor cells induces the upregulation of costimulatory molecules and MHC class II

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molecules on DCs resulting in enhanced ability to stimulate tumor-specific CD4+ and CD8+ IFN-γ-producing T cells (Fucikova et al., 2011). Moreover, the members of the HSP70 family have the capacity to stimulate TLR2/TLR4- and the CD14-associated pathway in monocytes and DCs leading to the activation of the NF-κB and IRF signaling pathways. This results in the release of proinflammatory cytokines such as TNF-α, IL-1β, IL-6, and IL-12 (Asea et al., 2000, 2002). HSP70 can also upregulate CD86 and CD40, thereby promoting DC maturation (Singh-Jasuja et al., 2000). All together these data support the concept that ecto-HSPs on tumor cells might be a crucial event for initiating effective antitumor immune response on the level of APCs by increasing the immunogenicity of tumor cells. ATP released from tumor cells succumbing to ICD not only attract myeloid cells into the tumor as mentioned in the previous section but it also facilitates myeloid cells to differentiate into inflammatory DCs. ATP activates purinergic P2X7 receptors on DCs, thereby activating the NLRP3 inflammasome leading to IL-1β release. This cytokine, together with antigen presentation, is required for the polarization of IFN-γ-producing CD8+ T cells and for an adaptive immune response to tumor cells (Ghiringhelli et al., 2009). Furthermore, DC maturation can be also induced by HMGB1, a nonhistone chromatin-binding protein, released from dying cells during the course of ICD (Dumitriu et al., 2007). HMGB1 can induce an inflammation by stimulating the production of proinflammatory cytokines, such as TNF-α, IL-1, IL-6, and IL-8, from myeloid cell populations (Chen et al., 2004). Tumor cells dying by ICD in response to anthracyclines secrete type I IFNs which trigger danger signaling through TLR3 activation and this is required for the initiation of adaptive immunity (Sistigu et al., 2014). It was also shown that by binding a common receptor, IFNAR (composed of two subunits, IFNAR1 and IFNAR2), type I IFNs can promote crosspriming by stimulating the maturation of DCs and their capacity to process and present antigens to CD8+ T cells (Zitvogel et al., 2015). Moreover, lowdose CTX boosts the differentiation of mature DCs from bone marrowresident DC precursors in vivo and ex vivo and CTX-induced mobilization and maturation of DCs is mediated by endogenous type I IFNs (Schiavoni et al., 2011). Of note, IFN-α is able to generate highly potent antigenpresenting DCs with cytotoxic activity (IFN-DCs). IFN-DCs induce tumor antigen-specific CD8+ T cells and acquire partial phenotypic and functional characteristics of NK cells, such as the expression of CD56, a NK cell

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marker, IFN-γ production, and cytolytic functions. Tumor cell killing activity was shown to be dependent on TNF-related apoptosis-inducing ligand (TRAIL) (Papewalis et al., 2008). Another ICD-associated DAMP that has been shown to activate DCs is uric acid which is released from dying tumor cells during late stage apoptosis. Hu et al. reported that the depletion of uric acid by using allopurinol decreases CD8+ T cell activity and inhibits tumor regression. Tumor regression is associated with high tumor levels of uric acid and its intratumoral injections accelerates tumor regression (Hu et al., 2004). Uric acid has the ability to engage the NLRP3 inflammasome, a complex of proteins aimed at activating caspase-1, leading to the cleavage of pro-IL-1 and pro-IL-18 to bioactive IL-1 and IL-18 (Martinon et al., 2006). It was also reported that uric acid crystals stimulate the maturation of DCs by increasing the expression of costimulatory molecules (CD80, CD86) and leads to the enhancement of T cell response to tumor antigens. Administration of uric acid as an adjuvant significantly enhances the ability of tumor lysate-pulsed DC vaccine in delaying the tumor growth (Wang et al., 2015). 2.2.7.4 Antigen Processing and Presentation

After internalization by DCs, most exogenous antigens, including TAAs, are processed through an endosomal and lysosomal pathway in which the proteins are cleaved into peptides and loaded into MHC class II molecules (Lennon-Dumenil et al., 2002). Alternatively, exogenous antigens can be released into the cytosol where they are processed by the proteasome, the main nonlysosomal protease, and transferred to the ER for loading into MHC class I molecules. This exogenous MHC class I presentation pathway is also referred as cross-presentation (Delamarre et al., 2003). The regulation of antigen uptake, processing, and presentation is under tight control. One of the important antigen-processing signals is HMGB1. This protein is normally found in the nucleus but it can be released from stressed cells during immunogenic apoptosis or secondary necrosis and mediates its extracellular activities by binding to various PRRs expressed predominantly by APCs, such as TLR2, TLR4, and RAGE (Apetoh et al., 2007b; Demaria et al., 2005). Treatment of tumor cells by ICD-inducing cytotoxic agents leads to postapoptotic release of HMGB1 which is recognized by TLR4 on DCs. TLR4 regulates the tumor antigen processing and is indispensable for efficient cross-presentation of tumor antigens by DCs. In the absence of TLR4 signaling, phagosomes fuse with lysosomes resulting in degradation of

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dying cells and consequently inefficient antigen presentation by DCs to CD4+ and CD8+ T cells. Importantly, the depletion of HMGB1 from CT26 cells used for mice immunization compromised the ability of mice to resist rechallenge with live CT26 tumor cells showing that the eradication of tumors by immunogenic chemotherapy (anthracyclines) requires the binding of HMGB1 to TLR4 (Apetoh et al., 2007b). In similar line, cancer patients bearing a defective TLR4 allele experienced increased incidence of tumor metastasis and reduced capacity of DCs to present antigens from dying tumor cells (Krysko et al., 2012). HSPs released in the extracellular space upon ICD bind tumor antigens and facilitate their cross-presentation by interacting with a number of receptors found on APCs like CD91, CD14, and scavenger receptors, such as LOX-1 and SREC-1 (Calderwood et al., 2007). After endocytosis, the complexes of HSPs and tumor antigens are degraded and tumor peptides are cross-presented to CD8+ T cells via MHC class I molecules (Doody et al., 2004). These complexes can along with MHC class I molecules activate antigen processing and presentation in DCs through TLR4 receptors and CD14 (Asea et al., 2000, 2002). Recently, another mechanism facilitating the uptake of HSP–peptide complexes by APCs and leading to the stimulation of CD4+ T cells was reported. This study showed that antigenic peptides associated with HSP90 are taken up into MHC class II molecules by a mechanism dependent on binding the scavenger receptor SREC-1 resulting in the internalization and processing of HSP90–antigen complexes by APCs. Finally, it leads to the antigen presentation to CD4+ T cells and their stimulation by APCs (Murshid et al., 2014).

3. CLINICAL TRIALS OF COMBINED DC-BASED IMMUNOTHERAPY AND CHEMOTHERAPY AFFECTING DC FUNCTIONS As mentioned in previous sections, DC-based immunotherapy has produced inconsistent clinical results leading to the development of combinatorial strategies. One such approach is the combination of DC vaccine with chemotherapeutic agents endowed with DC stimulatory properties improving DC phenotype and function and allowing to generate more potent DC-based immunotherapies. In this section, we summarize the latest advances in the use of ex vivo generated DC vaccines in combination with chemotherapeutic agents activating DC directly or inducing bona fide ICD, focusing on studies and clinical trials that have been launched during last

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15 years (Tables 3a and 3b). Since February 2001, there have been 35 clinical trials initiated to evaluate the safety and efficacy of such anticancer intervention (source http://www.clinicaltrials.gov). Of these 35 studies, 15 involve DCs pulsed with TAAs or TAA-derived peptides (NCT02018458, NCT00228189, NCT02503150, NCT02548169, NCT00722098, NCT00197912, NCT00683670, NCT00083538, NCT02705703, NCT00478452, NCT02224599, NCT02223312, NCT02479230, NCT01241162, NCT02332889), 9 DCs exposed to tumor lysate or inactivated allogeneic tumor cell line (NCT02215837, NCT00313235, NCT00197912, NCT01241682, NCT00083538, NCT02033616, NCT02105675, NCT02111577, NCT01803152), 6 DCs transfected with mRNA encoding for TAAs or tumor RNA (NCT00228189, NCT00978913, NCT02662634, NCT00617409, NCT01446731, NCT00049218), 5 DCs combined with cytokine-induced killer (CIK) cells (NCT02539017, NCT02504229, NCT02215837, NCT01691625, NCT02651441), and 4 involve not specified DC vaccine (NCT02423928, NCT00753220, NCT02419170, NCT00547144). The clinical studies testing the combination of DC-based immunotherapy and the category of chemotherapeutics with direct impact on DCs involve six studies with paclitaxel administered either as a single cytostatic agent (NCT02033616, NCT02018458) or in combination with all-trans retinoic acid (ATRA) (NCT00617409), FOLFIRINOX (folinic acid, oxaliplatin, irinotecan, 5-FU) + gemcitabine (NCT02548169), carboplatin, carboplatin/Alimta or cisplatin/Alimta (NCT02662634), or cisplatin + radiotherapy (NCT01691625); four studies with docetaxel administered either alone (NCT01446731, NCT02105675) or in combination with prednisone (NCT02111577) or CTX and epirubicin (NCT02539017); five studies with gemcitabine alone (NCT01803152, NCT00547144, NCT02479230) or combined with FOLFIRINOX, FOLFIRINOX + paclitaxel (NCT02548169) or cisplatin (NCT02651441); two studies with decitabine as a single chemotherapeutic agent (NCT01241162, NCT02332889), and two studies with etoposide combined with dexamethasone, thalidomide, cisplatin, adriamycin, and CTX (DT-PACE) (NCT00083538) or carboplatin (NCT00049218). Of the studies testing the application of DC-based immunotherapy and the chemotherapeutics classified as ICD inducers, 16 involve CTX either alone (NCT00722098, NCT00313235, NCT02423928, NCT00683670, NCT00753220, NCT00978913, NCT02419170, NCT02018458, NCT02705703, NCT00478452, NCT02224599, NCT02223312) or combined with

Table 3a Clinical Trials Recently Launched to Evaluate the Therapeutic Profile of DC-Based Immunotherapy Combined With Chemotherapy Inducing ICD Drug Indication Reference Phase Status Immunotherapy Specification Chemotherapy Specification

Doxorubicin Breast cancer epirubicin oxaliplatin Breast cancer

CTX

NCT02018458 I/II

Recruiting Cyclin B1/WT-1/CEFloaded DC vaccine

CTX, doxorubicin, paclitaxel, or carboplatin

NCT02539017 II

Not yet recruiting

Epirubicin, CTX, docetaxel

DC- and CIK cell-based immunotherapy

Gastric cancer NCT02504229 II

Recruiting DC- and CIK cell-based immunotherapy

Colorectal cancer

NCT00228189 I/II

Completed Mature DCs pulsed with CEA Standard adjuvant oxaliplatin/ petide or electroporated with capecitabine therapy (arm C) CEA mRNA

Colorectal cancer

NCT02503150 III

Not yet recruiting

Pancreatic cancer

NCT02548169 I

Recruiting Antigen-pulsed DCs

Antigen-pulsed DCs

Oxaliplatin + S-1 regimen

Modified FOLFOX6 therapy FOLFIRINOX alone or followed by 5-FU or gemcitabine; or gemcitabine + nab-paclitaxel

Gastric cancer NCT02215837 II

Active, not Tumor lysate-pulsed DCs and FOLFOX regimen recruiting CIK cells

Melanoma

NCT00722098 II

Terminated DCs loaded with TAAs

CTX alone

Melanoma

NCT00313235 I/II

Completed DCs pulsed with irradiated allogeneic melanoma cells (Colo 829)

CTX alone

Continued

Table 3a Clinical Trials Recently Launched to Evaluate the Therapeutic Profile of DC-Based Immunotherapy Combined With Chemotherapy Inducing ICD—cont’d Drug Indication Reference Phase Status Immunotherapy Specification Chemotherapy Specification

Prostate cancer

NCT02423928 I

Recruiting DCs injected into the prostate CTX (+ ipilimumab for last 10 patients) following prostatic cryoablation

Melanoma

NCT00197912 I/II

CTX (+ Celecoxib and IL-2) Completed Mature DCs pulsed with tumor antigens (p53, survivin, telomerase peptides) or allogeneic tumor lysate

Melanoma

NCT00683670 I

CTX alone Recruiting Mature DCs loaded with gp100 melanoma peptides (G209-2M and G280-9V) + 10 unique melanoma tumorspecific peptides

Prostate cancer

NCT00753220 I/II

Terminated DCs injected into the prostate CTX alone following prostatic cryoablation

Breast cancer and melanoma

NCT00978913 I

Completed DCs transfected with mRNA CTX alone encoding for hTERT, survivin, and p53

NSCLC

NCT02419170 0

Not yet recruiting

Mature DC-based vaccine

CTX alone

Breast cancer

NCT02018458 I/II

Recruiting Cyclin B1/WT-1/CEFloaded DC vaccine

CTX, doxorubicin, paclitaxel, or carboplatin

Mesothelioma NCT01241682 I

Completed Tumor lysate-pulsed autologous DCs

CTX + Alimta chemotherapy

Multiple myeloma

NCT00083538 II

Completed Myeloma antigen or myeloma DT-PACE chemotherapy for cell-loaded DCs high risk patients

Solid malignancies

NCT02705703 I/II

Not yet recruiting

Ovarian cancer

NCT00478452 I/II

Completed Peptide-pulsed mature DCs

CTX alone

Solid malignancies

NCT02224599 I/II

Not yet recruiting

TAA peptide-pulsed DCs + GM-CSF

CTX alone

Hematologic malignancies

NCT02223312 I/II

Not yet recruiting

TAA peptide-pulsed DCs + GM-CSF

CTX alone

Breast cancer

NCT02539017 II

Not yet recruiting

DC- and CIK cell-based immunotherapy

Epirubicin, CTX, docetaxel

TAA peptide-pulsed DCs + GM-CSF

CTX alone

Abbreviations: CEA, carcinoembryonic antigen; CIK, cytokine-induced killer cell; CTX, cyclophosphamide; DC, dendritic cell; DT-PACE, dexamethasone, thalidomide, cisplatin, adriamycin, CTX, etoposide; 5-FU, 5-fluorouracil; FOLFIRINOX, folinic acid, oxaliplatin, irinotecan, 5-FU; FOLFOX, oxaliplatin, leucovorin, 5-FU; GM-CSF, granulocyte monocyte-colony stimulating factor; NSCLC, nonsmall cell lung cancer; S-1, tegafur, gimeracil, oteracil potassium; TAA, tumor-associated antigen.

Table 3b Clinical Trials Recently Launched to Evaluate the Therapeutic Profile of DC-Based Immunotherapy Combined With Chemotherapy Directly Affecting DC Functions Drug Indication Reference Phase Status Immunotherapy Specification Chemotherapy Specification

Paclitaxel

NSCLC

NCT02662634 II

Recruiting Tumor RNA-loaded and electroporated mature autologous DCs

Paclitaxel prior to immunotherapy followed by carboplatin/Abraxane, carboplatin/Taxol, carboplatin/Alimta, or cisplatin/Alimta

Breast cancer

NCT02018458 I/II

Recruiting Cyclin B1/WT-1/CEFloaded DC vaccine

Standard preoperative chemotherapy—paclitaxel, doxorubicin, CTX, or carboplatin

Pancreatic cancer NCT02548169 I

Recruiting Antigen-pulsed DCs

FOLFIRINOX alone or followed by 5-FU or gemcitabine; or gemcitabine + nab-paclitaxel

SCLC

NCT00617409 II

Active, not DCs transduced with an recruiting adenoviral vector containing p53

Paclitaxel alone or in combination with ATRA

Ovarian, fallopian, and peritoneal carcinomas

NCT02033616 II

Not yet recruiting

Esophageal carcinoma

NCT01691625 NS

Recruiting DC- and CIK cell-based immunotherapy

Autologous DCs loaded with Adjuvant chemotherapy— autologous irradiated tumor paclitaxel alone cells + GM-CSF Chemoradiotherapy with paclitaxel and cisplatin

Docetaxel

Etoposide

Docetaxel alone

Prostate cancer

NCT01446731 II

Active, not IL-1β, TNF-α, IL-6, and recruiting PGE2-matured DCs transfected with PSA, PAP, survivin, and hTERT mRNAs

Prostate cancer

NCT02105675 II

Active, not DCs loaded with killed tumor Docetaxel alone recruiting cell line

Prostate cancer

NCT02111577 III

Recruiting DCs loaded with killed tumor Docetaxel + prednisone cell line

Breast cancer

NCT02539017 II

Not yet recruiting

DC- and CIK cell-based immunotherapy

Docetaxel, epirubicin, CTX

Multiple myeloma NCT00083538 II

Completed Myeloma antigen or myeloma DT-PACE chemotherapy cell-loaded DCs including etoposide for high risk patients

SCLC

NCT00049218 I/II

Completed DCs transduced with an adenoviral vector containing p53

Etoposide, carboplatin

NCT01803152 I

Recruiting Mature DCs loaded with autologous sarcoma tumor lysates

Gemcitabine alone

Completed Intratumoral DC-based immunotherapy

Gemcitabine alone

Gemcitabine Soft tissue and bone sarcomas

Pancreatic cancer NCT00547144 I

Continued

Table 3b Clinical Trials Recently Launched to Evaluate the Therapeutic Profile of DC-Based Immunotherapy Combined With Chemotherapy Directly Affecting DC Functions—cont’d Drug Indication Reference Phase Status Immunotherapy Specification Chemotherapy Specification

Recruiting Tumor blood vessel antigen peptide-pulsed alpha-type-1 polarized DCs

Gemcitabine alone

Pancreatic cancer NCT02548169 I

Recruiting Antigen-pulsed DCs

FOLFIRINOX alone or followed by 5-FU or gemcitabine; or gemcitabine + nab-paclitaxel

NSCLC

NCT02651441 I/II

Not yet recruiting

Gemcitabine, cisplatin

Neuroblastoma and various sarcomas

NCT01241162 I

Active, not Hiltonol-matured DCs pulsed Decitabine alone recruiting with CT antigens

Breast cancer

Decitabine

NCT02479230 I

NCT02332889 I/II Gliomas, medulloblastoma, neuroectodermal tumors

D-CIK cells

Recruiting Hiltonol-matured DCs pulsed Decitabine alone with peptide mixes derived from MAGE-A1, -A3, and NY-ESO-1 + GM-CSF

Abbreviations: ATRA, all-trans retinoic acid; CEF, peptide pool derived from cytomegalovirus, Epstein–Barr virus and influenza; CIK, cytokine-induced killer cell; CT, cancer testis; CTX, cyclophosphamide; DC, dendritic cell; D-CIK, DC-activated cytokine-induced killer cell; DT-PACE, dexamethasone, thalidomide, cisplatinum, adriamycin, CTX, etoposide; 5-FU, 5-fluorouracil; FOLFIRINOX, folinic acid, oxaliplatin, irinotecan, 5-FU; GM-CSF, granulocyte monocyte-colony stimulating factor; hTERT, human telomerase reverse transcriptase; MAGE, melanoma-associated antigen; NS, not specified; NSCLC, nonsmall cell lung cancer; PAP, prostatic acid phosphatase; PGE2, prostaglandin E2; PSA, prostate-specific antigen; SCLC, small cell lung cancer; TNF-α, tumor necrosis factor-α; WT-1, Wilms’ tumor 1.

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Alimta (NCT01241682), DT-PACE (dexamethasone, thalidomide, cisplatin, adriamycin, etoposide) (NCT00083538), or Celecoxib + IL-2 (NCT00197912); 5 oxaliplatin administered in combination with capecitabine (NCT00228189), S-1 regimen (Tegafur, Gimeracil, Oteracil Potassium) (NCT02504229), 5-FU + Leucovorin (FOLFOX) (NCT02503150, NCT02215837), or FOLFIRINOX + gemcitabine or gemcitabine + paclitaxel (NCT02548169); 1 doxorubicin as a single cytostatic agent (NCT02018458); and 1 study involve epirubicin combined with CTX and docetaxel (NCT02539017). In most studies, the primary objective was to determine the safety and tolerability of DC vaccines employed along with chemotherapy as a standard of care and the clinical impact of these interventions is not mentioned. Moreover, only limited number of randomized studies were reported including a control group consisting of patients treated either by chemotherapeutic agents or DC-based vaccine only. Furthermore, 17 clinical trials was initiated after 2014, thus limiting the number of trials already reporting clinical results. In a limited group of 10 studies with reported clinical data, only 3 included a control group with patients receiving only chemotherapy, thus comparing combined chemoimmunotherapy with monotherapy only. In a pilot study, Punt and colleagues investigated the effect of treatment with oxaliplatin and capecitabine on nonspecific and specific DC vaccine-induced adaptive immune responses in stage III colon cancer patients. Although the antitumor immune response in the treated group was comparable to that of patients treated by DC vaccine only, oxaliplatin administration significantly enhanced nonspecific T cell reactivity (Lesterhuis et al., 2010) documenting the role of this platinum derivative in augmenting the immunostimulatory potential of DC vaccines. Combinatorial chemoimmunotherapy regimens involving CTX-based chemotherapy and DCs loaded with human epidermal growth factor receptor 2 (Her2/neu), human telomerase reverse transcriptase (hTERT), and pan-DR epitope (PADRE) peptides have been tested in phase I/II trial enrolling ovarian cancer patients in advanced stage. Althought the therapy turned out to be well tolerated, it does not lead to the improvement of the clinical parameters (overall survival (OS) and progression-free survival (PFS)) and to the enhancement of tumor antigenspecific T cell response when compared to DC-based therapy only (Chu et al., 2012). Another study investigated the effect of p53-modified adenovirus-transduced DC vaccine and second-line chemotherapy (paclitaxel) on adaptive immune responses in small cell lung carcinoma (SCLC) patients with extensive-stage disease. In general, more than half

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of patients developed anti-p53-specific immune response after DC vaccination. Patients with a positive immune response had a trend toward improved survival suggesting that DC vaccine sensitizes SCLC patients to subsequent chemotherapy (Chiappori et al., 2010). Recently, Zhang et al. published the results from a study investigating the effect of gemcitabine/cisplatin treatment combined with DC- and CIK cell-based vaccine on the recurrence and survival rate of patients with nonsmall cell lung carcinoma (NSCLC). They observed that the percentage of CD4+ T cells, NK cells, and the CD4/CD8 ratio were significantly increased after the course of treatment. Of note, the percentage of CD4+ T cells, NK cells, and the CD4/CD8 ratio in the peripheral blood were identified to be significantly increased in the group of patients receiving both immunotherapy and chemotherapy compared to the control arm of patients treated with standart chemotherapy regimen only. In the similar line, the synergistic function of chemoimmunotherapy has been demonstrated by different clinical parameters monitored within the study. Firstly, the median disease free survival time of patients in the observation groups was significantly increased up to 28 months compared to control group (22 months). And secondly, 3-year cumulative survival rate was significantly increased in the chemoimmunotherapy-treated group of patients compared to control group (Zhao et al., 2014). Based on promising results coming from preclinical testing the DC + CIK-based vaccine combined with gemcitabine/cisplatinbased chemotherapy is now being tested in phase I/II clinical trial. In the vast majority of clinical studies evaluating the potential of DC-based immunotherapy and chemotherapy with DC stimulatory capacity, the details about the control arm of patients cohort are missing. However, the majority of these studies documented the induction of specific antitumor immune response after the administration of combinatorial therapy (Carreno et al., 2015; Ellebaek et al., 2012; Hegmans et al., 2010; Krishnadas et al., 2015; Podrazil et al., 2015; Svane et al., 2007; Yi et al., 2010). Three studies have demonstrated the clinical stabilization of disease (Ellebaek et al., 2012; Svane et al., 2007; Yi et al., 2010) or complete response following chemoimmunotherapy (Krishnadas et al., 2015). Of note, in phase I/II trial, patients with metastatic prostate cancer treated by DC-based immunotherapy DCVAC/PCa composed of autologous poly I:C-activated DCs pulsed with killed prostate cancer cell line LNCap and combined with docetaxel lived 6 months longer than predicted by standard nomograms and Kaplan–Meier analyses showing that patients treated by this regimen had a lower risk of death compared to predicted risk (Podrazil et al., 2015). In general, DC-based vaccines were well tolerated in all reported

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studies and the vast majority of severe adverse events and toxicity was related to applied chemotherapy. Taken together, the results of these studies (the majority of which was phase I or II trials) indicate that DC-based anticancer interventions and chemotherapy are generally safe and elicit anticancer immune responses that, at least in a fraction of patients, underpin objective clinical responses.

4. CONCLUSIONS The long-standing perception that the administration of chemotherapy has predominantly negative effect on the cells of the innate and adaptive arm of the immune system has been recently contested. Certain cytotoxic agents can potentiate antitumor immunity by various mechanisms and some of them kill tumor cells through ICD. This provides the rationale for combining chemotherapy with immune-based therapies to explore the synergies and possibly enhance their efficacy. Over the past decade, the increased knowledge of DC biology and their role in the coordination of the immune responses caused DCs to be perceived as a promising vehicle for cancer immunotherapy. Although many trials of DC-based immunotherapy reported the induction of antitumor immune responses, the clinical success of this treatment has been suboptimal pointing out to the need for additional interventions that would enhance DC functions and improve the efficacy of DC vaccines. As mentioned earlier, distinct chemotherapeutic agents have the potential to induce the maturation and activation of DCs either directly or indirectly through eliciting the emission of DAMPs from dying tumor cells. Although the clinical studies indicate that DC vaccines applied in combination with such chemotherapeutics are in general well-tolerated, larger, well-designed studies are needed to test the hypothesis of synergistic effect between DC-based immunotherapy and chemotherapy in clinical settings.

ACKNOWLEDGMENTS This project was supported by Ministry of Health of Czech Republic—conceptual development of research organization, University Hospital Motol, Prague, Czech Republic, 00064203, the research grant PRVOUK P37/11 and BBMRI_CZ LM2015089 and student research grant GAUK 682214.

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CHAPTER FOUR

Smac Mimetics to Therapeutically Target IAP Proteins in Cancer S. Fulda1 Institute for Experimental Cancer Research in Pediatrics, Goethe-University, Frankfurt, Germany German Cancer Consortium (DKTK), Heidelberg, Germany German Cancer Research Center (DKFZ), Heidelberg, Germany 1 Corresponding author: e-mail address: [email protected]

Contents 1. Introduction 2. IAP Proteins 3. Mechanisms of Action of Smac Mimetics 4. Smac Mimetics as Single Agents 5. Smac Mimetic-Based Combination Therapies 6. Conclusions Acknowledgments References

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Abstract Inhibitor of Apoptosis (IAP) proteins are overexpressed in a variety of human cancers. Therefore, they are considered as promising targets for the design of therapeutic strategies. Smac mimetics mimic the endogenous mitochondrial protein Smac that antagonizes IAP proteins upon its release into the cytosol. Multiple preclinical studies have documented the ability of Smac mimetics to either directly induce cell death of cancer cells or to prime them to agents that trigger cell death. At present, several Smac mimetics are being evaluated in early clinical trials. The current review provides an overview on the potential of Smac mimetics as cancer therapeutics to target IAP proteins for cancer therapy.

1. INTRODUCTION Resistance to programmed cell death is a characteristic feature of human malignancies (Hanahan and Weinberg, 2011) that contributes to tumorigenesis as well as to treatment resistance (Fulda, 2009). There are different modes of programmed cell death including apoptosis (Galluzzi et al., 2012). The extrinsic (death receptor) and the intrinsic (mitochondrial) International Review of Cell and Molecular Biology, Volume 330 ISSN 1937-6448 http://dx.doi.org/10.1016/bs.ircmb.2016.09.004

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pathways represent the two major signaling cascades that eventually lead to activation of caspases (Fulda and Debatin, 2006). In the intrinsic (mitochondrial) pathway, apoptotic stimuli trigger the release of mitochondrial intermembrane space proteins including Smac into the cytosol (Fulda et al., 2010). There also exist additional modes of programmed cell death besides apoptosis (Galluzzi et al., 2012). For example, necroptosis is a regulated form of necrosis that involves defined molecular mechanisms including receptorinteracting protein (RIP)1, RIP3, and mixed lineage kinase domain-like (MLKL) as core signaling molecules (Vanden Berghe et al., 2014). Cell death pathways are tightly regulated by various mechanisms to avoid their accidental activation, and a large variety of positive and negative regulators have been identified in the last decades (Fulda, 2009). For example, Inhibitor of Apoptosis (IAP) proteins belong to the group of antiapoptotic proteins (Fulda and Vucic, 2012). Since mechanisms that block programmed cell death including antiapoptotic proteins favor the evasion of programmed cell death that is typically observed in human cancers, a better understanding of the underlying mechanistic details can open new perspectives for therapeutic interventions. The current review will focus on Smac mimetics as cancer therapeutics to target IAP proteins for cancer therapy.

2. IAP PROTEINS Mammalian IAP proteins comprise eight family members, including X chromosome-linked IAP (XIAP), cellular IAP1 (cIAP1), and cIAP2 (reviewed in Fulda and Vucic, 2012). Among other functions, XIAP can antagonize apoptosis by inhibiting caspases (Fulda and Vucic, 2012). cIAP proteins act as E3 ubiquitin ligases that mediate ubiquitination of target proteins via their really interesting new gene (RING) domain (Fulda and Vucic, 2012). Human cancers typically express high levels of IAP proteins which favor their evasion of programmed cell death (Fulda and Vucic, 2012). As aberrantly high expression of IAP proteins has also been linked to treatment resistance and poor prognosis, IAP proteins are widely viewed as suitable targets for the development of experimental therapies (Fulda and Vucic, 2012). Since endogenous Smac protein promotes cell death upon its release from the mitochondrial intermembrane space into the cytosol by antagonizing IAP proteins, the most extensively exploited strategy is based on the design of small-molecule inhibitors that mimic the endogenous IAP antagonist Smac and are therefore called Smac mimetics.

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3. MECHANISMS OF ACTION OF SMAC MIMETICS Smac mimetics are small molecules that mimic the N-terminal portion of Smac protein that encompasses a four-amino acid stretch (i.e., Ala-Val-ProIle) that is critical for the binding of Smac to the Baculovirus IAP repeat domain (BIR)3 and BIR2 domains of IAP proteins (Liu et al., 2000; Wu et al., 2000). Similar to the endogenous protein Smac, Smac mimetics bind to IAP proteins and antagonize their functions (Fulda and Vucic, 2012). Since Smac protein has been reported to form a homodimer under physiological conditions, bivalent compounds in addition to monovalent (monomeric) Smac mimetics have been developed that are composed of two monomeric elements which are linked by a chemical bridge. Since one bivalent Smac mimetic can bind to two IAP proteins and is better at triggering dimerization and autodegradation of cIAP proteins (Darding et al., 2011; Dueber et al., 2011; Feltham et al., 2011), bivalent compounds exhibit a higher cytotoxicity than monomeric ones. Binding of Smac mimetics to XIAP disrupts the interaction of XIAP to caspase-3, -7, and -9, leading to caspase activation and caspase-dependent apoptosis (Fulda and Vucic, 2012). Binding of Smac mimetics to cIAP1 and cIAP2 stimulates their E3 ubiquitin ligase activity and engages their proteasomal degradation (Darding et al., 2011; Dueber et al., 2011; Feltham et al., 2011; Varfolomeev et al., 2007; Vince et al., 2007). This in turn activates non-canonical nuclear factor-kappa B (NF-κB) signaling by stabilizing NF-κB-inducing kinase (NIK), which is constitutively ubiquitinated by cIAP proteins (Varfolomeev et al., 2007; Vince et al., 2007). This leads to transcriptional activation of NF-κB target genes, for example, tumor necrosis factor (TNF)α (Oeckinghaus et al., 2011; Zarnegar et al., 2008). TNFα then mediates Smac mimetic-triggered cancer cell death in an autocrine or paracrine fashion via the TNF receptor 1 (TNFR1) (Varfolomeev et al., 2007; Vince et al., 2007). Depletion of cIAP proteins also leads to reduced ubiquitination of RIP1, which promotes the interaction of RIP1 with cell death signaling proteins such as caspase-8 and Fas-associated protein with death domain (FADD), resulting in the formation of a signaling platform composed of several proteins (Fulda and Vucic, 2012).

4. SMAC MIMETICS AS SINGLE AGENTS Smac mimetics as single agents have been shown to trigger cell death in several cancers, e.g., chronic lymphocytic leukemia (CLL), acute

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lymphoblastic leukemia (ALL), and acute myeloid leukemia (AML) (Brumatti et al., 2016; Lalaoui et al., 2016; McComb et al., 2016; Opel et al., 2015; Schirmer et al., 2016; Steinhart et al., 2013; Steinwascher et al., 2015). Smac mimetic-stimulated engagement of auto- or paracrine TNFα signaling has been shown to play a critical role for the induction of cell death upon treatment with Smac mimetic monotherapy (Varfolomeev et al., 2007; Vince et al., 2007). Accordingly, interfering with TNFα/TNFR1 receptor–ligand interaction, either by pharmacologically using TNFα-blocking antibodies or by genetically using RNA interference technology, has been described to inhibit Smac mimetic-induced cell death (Varfolomeev et al., 2007; Vince et al., 2007). Besides TNFα, also other death receptor/ligand systems such as the tumor necrosis factor-related apoptosis-inducing ligand (TRAIL) have been reported to be involved at least in certain cancers (Eckhardt et al., 2013). Activation of transcription factors such as NF-κB or interferon-regulatory factor 1 (IRF1) by Smac mimetics has been demonstrated to contribute to apoptosis, since genetic blockage of NF-κB or IRF1 rescued cells from Smac mimetic-mediated apoptosis (Eckhardt et al., 2013, 2014). Smac mimetics as single agents have been shown to trigger cell death in several cancers including primary tumor samples, for example, from patients with CLL (Opel et al., 2015).

5. SMAC MIMETIC-BASED COMBINATION THERAPIES In addition to the use of Smac mimetics as single agents, a number of strategies have been developed to combine Smac mimetics with other cytotoxic principles. These combination treatments include conventional chemotherapeutic drugs as well as different types of molecular targeted agents. The combination of Smac mimetics with agents that trigger death receptors such as TRAIL, CD95 ligand, or TNFα turned out to be one of the most effective strategies to induce cell death in cancer cells in multiple different cancer types. As far as leukemia is concerned, Smac mimetics have been shown to act in concert with TRAIL to induce apoptosis in ALL cell lines as well as primary blasts (Fakler et al., 2009). In addition, the Smac mimetic/TRAIL therapy exerted antileukemic activity in vivo in a patient-derived mouse model (Fakler et al., 2009). Also, Smac mimetics sensitized CLL cells derived from primary patient samples to TRAILinduced apoptosis even in poor prognostic subgroups of the disease,

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e.g., in patient samples with deletion of chromosome p17, mutation in p53, resistance to fludarabine of unmutated variable heavy-chain genes (Loeder et al., 2009). In pancreatic cancer, Smac mimetics in combination with antibodies directed against agonistic TRAIL receptors or with soluble TRAIL-triggered apoptosis in vitro and suppressed tumor growth in vivo (Stadel et al., 2010; Vogler et al., 2009). In colorectal carcinoma cells, EGFR-targeted TRAIL synergized with Smac mimetics to overcome apoptosis resistance (Moller et al., 2014). The combination of Smac mimetics and TRAIL not only exhibited antitumor activity against the primary tumor but has also been described to prevent metastasis, for example, in preclinical models of cholangiocarcinoma (Fingas et al., 2010). Besides TRAIL, agents that target CD95 as another death receptor of the TNFR superfamily, e.g., CD95 ligand or agonistic CD95 antibodies, have been shown to act in concert with Smac mimetics to induce cancer cell death (Geserick et al., 2009; Loeder et al., 2010). Moreover, cooperative induction of cell death has been reported for Smac mimetics and TNFα (Laukens et al., 2011; Roesler et al., 2016; Schenk and Fulda, 2015; Wang et al., 2008). Smac peptides and Smac mimetic compounds have been shown to act in concert with various anticancer drugs in haematological malignancies as well as in solid tumors in preclinical in vitro and in vivo studies (Bockbrader et al., 2005; Carter et al., 2005; Chauhan et al., 2007; Cheng et al., 2010; Chromik et al., 2014; Dean et al., 2010; Dineen et al., 2010; Fandy et al., 2008; Fulda et al., 2002; Guo et al., 2002; Loeder et al., 2012; Metwalli et al., 2010; Probst et al., 2010; Servida et al., 2011; Stadel et al., 2011; Tenev et al., 2011; Wagner et al., 2013; Ziegler et al., 2011). For example, in childhood ALL, Smac mimetics synergized with chemotherapeutic drugs such as AraC, gemcitabine, cyclophosphamide, doxorubicin, etoposide, vincristine, or taxol to engage apoptotic cell death (Loeder et al., 2012; Schirmer et al., 2016). A synergistic interaction to trigger apoptosis in childhood ALL has also been reported for Smac mimetics in combination with glucocorticoids including dexamethasone and prednisolone, both in vitro and also in a patient-derived xenograft model in vivo (Belz et al., 2014). In glioblastoma, Smac mimetics have been shown to increase the sensitivity toward the commonly used chemotherapeutic drug temozolomide (Wagner et al., 2013). This Smac mimetic-imposed chemosensitization occurred without the requirement of death receptors or their ligands (Wagner et al., 2013). By comparison, Smac mimetic-stimulated

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upregulation of interferon-β (IFNβ) and IFN-induced proapoptotic signaling turned out to contribute to the induction of cell death (Marschall and Fulda, 2015). In another study TNFα has been reported to be necessary for Smac mimetic-mediated sensitization to chemotherapeutic agents in vitro and in vivo (Probst et al., 2010), pointing to a context-dependent involvement of TNFα. Also, cotreatment with Smac mimetics and chemotherapeutics has been documented to upregulate TNFα production in tumor tissue in a pancreatic cancer model in vivo (Dineen et al., 2010). In AML, Smac mimetics primed cells for cytarabine-mediated cell death (Chromik et al., 2014). When caspase activation was blocked, Smac mimetic/cytarabine cotreatment triggered necroptosis as an alternative mode of cell death (Chromik et al., 2014). Besides DNA-damaging chemotherapeutic drugs, also γ-irradiation has been reported to cooperate with Smac mimetics to elicit cell death in a variety of cancer types (Berger et al., 2011; Giagkousiklidis et al., 2007; Vellanki et al., 2009; Vucic et al., 2005). In glioblastoma, Smac mimetics sensitized not only glioblastoma cell lines but also primary glioblastoma cells and glioblastoma-initiating stem-like cancer cells to γ-irradiation-imposed apoptosis (Berger et al., 2011; Vellanki et al., 2009). Also, Smac mimetics increased the antiglioma activity of a combination of radio/chemotherapy, using temozolomide in an in vivo mouse model of glioblastoma (Ziegler et al., 2011). Targeting IAP proteins by Smac mimetics also enhanced the radiosensitivity of pancreatic, breast, colorectal, or head and neck squamous cell carcinoma cells (Giagkousiklidis et al., 2007; Hehlgans et al., 2015; Yang et al., 2011, 2012). In addition to death receptor agonists, chemotherapeutics and γ-irradiation, Smac mimetics have also been applied in combination with various signal transduction inhibitors. For example, epigenetic drugs including demethylating agents or histone deacetylase (HDAC) inhibitors have been shown to synergize with Smac mimetics. In AML, Smac mimetics have been shown to sensitize cells, including CD34-positive AML stem– progenitor cells, to demethylating drugs such as 5-azacitadine or 5-Aza-2-deoxycytidine (5-Aza) (Carter et al., 2014; Steinhart et al., 2013). Also, HDAC inhibitors such as MS273 or SAHA turned out to act in concert with Smac mimetics to trigger cell death in AML cells (Steinwascher et al., 2015). In addition, Smac mimetics cooperated to induce cancer cell death in combination with proteasome inhibitors, e.g., in multiple myeloma (Chauhan et al., 2007) or in melanoma (Lecis et al., 2010). Furthermore, Smac mimetics have been reported to potentiate the antitumor activity of various

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tyrosine kinase inhibitors, for example, the BCR-ABL inhibitor nilotinib, the FLT3 inhibitor PKC412 (Weisberg et al., 2007, 2010), or PDGFR inhibitors (Ziegler et al., 2008). In breast cancer, Smac mimetics have also been combined with Her2 antagonists to suppress proliferation and to elicit apoptosis (Foster et al., 2009). Moreover, Smac mimetics have been shown to act in concert with recombinant type I and type II IFNs. In AML, the Smac mimetic BV6 and IFNα have been described to synergistically trigger cell death in AML cells, whereas no synergistic toxicity was found against normal peripheral blood lymphocytes (Bake et al., 2014). Smac mimetic/IFNαmediated cell death depended on a TNFα/TNFR1-stimulated signaling and IRF1 in AML cells, since pharmacological or genetic inhibition of TNFα, TNFR1, or IRF1 rescued cell death upon treatment with Smac mimetic (Bake et al., 2014). However, a differential requirement of TRAIL or TNFα as mediators of IFNα/Smac mimetic-induced cell death has also been documented depending on the cellular context (Roesler et al., 2016). Accordingly, cooperative TRAIL production has been demonstrated to mediate IFNα/Smac mimetic-induced cell death in A172 glioblastoma cells (Roesler et al., 2016). Besides IFNs, also other immune stimuli have been reported to potentiate the antitumor activity of Smac mimetics, for example, oncolytic viruses or non-infectious immunostimulatory molecules including poly(I:C) or CpG oligonucleotides (Beug et al., 2014). Also, Smac mimetics have been described to enhance the efficacy of Bacillus Calmette–Guerin (BCG) immunotherapy in a preclinical model of bladder cancer (Jinesh et al., 2012).

6. CONCLUSIONS Smac mimetics are currently being tested in phase I/II clinical trials in a number of human cancers. While administration of Smac mimetics in principle proved to be well-tolerated (Amaravadi et al., 2011; Infante et al., 2010; Sikic et al., 2011), the first in-human phase I dose escalation study in which LCL161 has been given to patients with advanced stages of solid tumors reported a dose-limiting toxicity, i.e., cytokine release syndrome (Infante et al., 2014). Since preclinical studies have shown that Smac mimetic-stimulated non-canonical NF-κB activation results in increased production of inflammatory cytokines, for example, TNFα (Varfolomeev et al., 2007; Vince et al., 2007), and since increased levels of inflammatory cytokines have been detected in the circulation of Smac mimetic-treated

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patients (Infante et al., 2014), this side effect can be explained by an on-target mechanism. It also implies that Smac mimetic-stimulated upregulation of TNFα, which has also been shown to act as an essential mediator of Smac mimetic-induced cell death, represents a double-edged sword. Thus, it will likely be a challenge to maximize Smac mimetic-mediated antitumor activity while minimizing potential unwanted effects. One approach may reside in the design of rational combination regimens of Smac mimetics together with other cytotoxic principles, given the promising preclinical results showing synergistic drug interactions. Taken together, Smac mimetics may open new perspectives for the treatment of cancer, which warrant further investigation.

ACKNOWLEDGMENTS The expert secretarial assistance of C. Hugenberg is greatly appreciated. This work has been partially supported by grants from the Deutsche Forschungsgemeinschaft, the BMBF (German Cancer Consortium (DKTK), German Cancer Research Center (DKFZ) and CI3 (131A029B)), the Deutsche Krebshilfe, IUAP VII, and Wilhelm Sander-Stiftung.

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emricasan combines with the SMAC mimetic birinapant to induce necroptosis and treat acute myeloid leukemia. Sci. Transl. Med. 8, 339ra369. Carter, B.Z., Gronda, M., Wang, Z., Welsh, K., Pinilla, C., Andreeff, M., Schober, W.D., Nefzi, A., Pond, G.R., Mawji, I.A., Houghten, R.A., Ostresh, J., Brandwein, J., Minden, M.D., Schuh, A.C., Wells, R.A., Messner, H., Chun, K., Reed, J.C., Schimmer, A.D., 2005. Small-molecule XIAP inhibitors derepress downstream effector caspases and induce apoptosis of acute myeloid leukemia cells. Blood 105, 4043–4050. Carter, B.Z., Mak, P.Y., Mak, D.H., Shi, Y., Qiu, Y., Bogenberger, J.M., Mu, H., Tibes, R., Yao, H., Coombes, K.R., Jacamo, R.O., McQueen, T., Kornblau, S.M., Andreeff, M., 2014. Synergistic targeting of AML stem/progenitor cells with IAP antagonist birinapant and demethylating agents. J. Natl. Cancer Inst. 106, djt440. Chauhan, D., Neri, P., Velankar, M., Podar, K., Hideshima, T., Fulciniti, M., Tassone, P., Raje, N., Mitsiades, C., Mitsiades, N., Richardson, P., Zawel, L., Tran, M., Munshi, N., Anderson, K.C., 2007. Targeting mitochondrial factor Smac/DIABLO as therapy for multiple myeloma (MM). Blood 109, 1220–1227. Cheng, Y.J., Jiang, H.S., Hsu, S.L., Lin, L.C., Wu, C.L., Ghanta, V.K., Hsueh, C.M., 2010. XIAP-mediated protection of H460 lung cancer cells against cisplatin. Eur. J. Pharmacol. 627, 75–84. Chromik, J., Safferthal, C., Serve, H., Fulda, S., 2014. Smac mimetic primes apoptosisresistant acute myeloid leukaemia cells for cytarabine-induced cell death by triggering necroptosis. Cancer Lett. 344, 101–109. Darding, M., Feltham, R., Tenev, T., Bianchi, K., Benetatos, C., Silke, J., Meier, P., 2011. Molecular determinants of Smac mimetic induced degradation of cIAP1 and cIAP2. Cell Death Differ. 18, 1376–1386. Dean, E.J., Ward, T., Pinilla, C., Houghten, R., Welsh, K., Makin, G., Ranson, M., Dive, C., 2010. A small molecule inhibitor of XIAP induces apoptosis and synergises with vinorelbine and cisplatin in NSCLC. Br. J. Cancer 102, 97–103. Dineen, S.P., Roland, C.L., Greer, R., Carbon, J.G., Toombs, J.E., Gupta, P., Bardeesy, N., Sun, H., Williams, N., Minna, J.D., Brekken, R.A., 2010. Smac mimetic increases chemotherapy response and improves survival in mice with pancreatic cancer. Cancer Res. 70, 2852–2861. Dueber, E.C., Schoeffler, A.J., Lingel, A., Elliott, J.M., Fedorova, A.V., Giannetti, A.M., Zobel, K., Maurer, B., Varfolomeev, E., Wu, P., Wallweber, H.J., Hymowitz, S.G., Deshayes, K., Vucic, D., Fairbrother, W.J., 2011. Antagonists induce a conformational change in cIAP1 that promotes autoubiquitination. Science 334, 376–380. Eckhardt, I., Roesler, S., Fulda, S., 2013. Identification of DR5 as a critical, NF-kappaBregulated mediator of Smac-induced apoptosis. Cell Death Dis. 4.e936. Eckhardt, I., Weigert, A., Fulda, S., 2014. Identification of IRF1 as critical dual regulator of Smac mimetic-induced apoptosis and inflammatory cytokine response. Cell Death Dis. 5.e1562. Fakler, M., Loeder, S., Vogler, M., Schneider, K., Jeremias, I., Debatin, K.M., Fulda, S., 2009. Small molecule XIAP inhibitors cooperate with TRAIL to induce apoptosis in childhood acute leukemia cells and overcome Bcl-2-mediated resistance. Blood 113, 1710–1722. Fandy, T.E., Shankar, S., Srivastava, R.K., 2008. Smac/DIABLO enhances the therapeutic potential of chemotherapeutic drugs and irradiation, and sensitizes TRAIL-resistant breast cancer cells. Mol. Cancer 7, 60. Feltham, R., Bettjeman, B., Budhidarmo, R., Mace, P.D., Shirley, S., Condon, S.M., Chunduru, S.K., McKinlay, M.A., Vaux, D.L., Silke, J., Day, C.L., 2011. Smac mimetics activate the E3 ligase activity of cIAP1 protein by promoting RING domain dimerization. J. Biol. Chem. 286, 17015–17028.

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CHAPTER FIVE

Consequences of Keratin Phosphorylation for Cytoskeletal Organization and Epithelial Functions M.S. Sawant, R.E. Leube1 Institute of Molecular and Cellular Anatomy, RWTH Aachen University, Aachen, Germany 1 Corresponding author: e-mail address: [email protected]

Contents 1. Introduction 2. Characteristics of Keratin Phosphorylation 2.1 Keratin Phosphorylation Is Complex, Fast, and Linked to Other Posttranslational Modifications 2.2 Phosphorylation Occurs Preferentially in the Keratin End Domains 2.3 Phosphorylation Increases Keratin Solubility 2.4 Common Guidelines Determine Keratin Phosphorylation 3. Regulation of Protein Binding to Keratins by Phosphorylation 3.1 Phosphorylation-Dependent Keratin-14-3-3 Protein Association Enhances Keratin Solubility and Affects mTOR Signaling 3.2 Sequestration of Raf Kinase Is Regulated by Keratin Phosphorylation 3.3 Association of Cytolinkers With Keratin Is Influenced by Phosphorylation 3.4 Phosphorylation-Dependent Association of the Ubiquitin Ligase Pirh2 With Keratin Affects Cell Survival 4. Role of Keratin Phosphorylation in Cell Physiology 4.1 Mitosis Is Linked to Keratin Phosphorylation and Keratin Network Remodeling 4.2 The Epithelial Stress Response Is Coupled to Altered Keratin Phosphorylation 4.3 Keratin Phosphorylation Protects Against Ubiquitin-Dependent Degradation and Reduces Sensitivity to Apoptosis 5. Disease Relevance of Keratin Phosphorylation 5.1 Keratin Phosphorylation Affects Keratin Network Organization and Function in Simple Epithelia 5.2 Keratin Phosphorylation Is Altered in Skin Diseases 5.3 Keratin Phosphorylation Is Altered in Cancer

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172 173 173 178 179 180 181 181 182 182 183 183 183 187 194 196 196 200 204

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6. Concluding Remarks: Significance of Keratin Phosphorylation Acknowledgments References

210 210 211

Abstract Intermediate filaments are major phosphoproteins. The complex patterns of intermediate filament phosphorylation make up a poorly understood code reflecting cytoskeletal properties and cellular function through an intense crosstalk with multiple signaling pathways. This review focuses on the epithelial keratin intermediate filaments highlighting the tight-knit relationship of keratin phosphorylation and network organization during cell division and apoptosis, and the importance of keratin phosphorylation during epithelial stress responses. The occurrence of keratin phosphorylation in genetic skin diseases and acquired diseases of simple epithelial tissues in liver, pancreas, and colon will be discussed. Finally, we will review the role of keratin phosphorylation in cancer with an emphasis on migration.

1. INTRODUCTION Intermediate filament proteins are abundant cellular phosphoproteins. Individual polypeptides carry multiple phosphorylation sites that are regulated by numerous kinase and phosphatase activities resulting in complex contextdependent phosphorylation patterns that reflect cellular differentiation and function (Binukumar et al., 2013; Omary and Ku, 2006; Sihag et al., 2007; Snider and Omary, 2014). Clarifying how the interplay of different phosphorylation sites affects the structure and function of intermediate filaments is a challenging task because properties of the intermediate filament cytoskeleton may be altered only incrementally in certain situations and different phosphorylation sites may have opposing effects. To further complicate matters, the effects of phosphorylation may occur locally or globally and may manifest rapidly or only after some lag period. Given the complexity of intermediate filament phosphorylation, we limit the review to the epithelial keratin intermediate filaments. Keratins are among the most abundant cellular proteins in epithelial cells and therefore provide a large buffer reservoir to cope with physical, chemical, and microbial stresses imposed by the hostile exterior environment (Pan et al., 2013; Pekny and Lane, 2007; Toivola et al., 2010, 2015). Keratins are obligatory heteropolymers that are held together by hydrophobic interactions through their α-helical rod domain forming stable coiled-coils (Herrmann et al., 2003, 2009). Filamentous keratin is composed of

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equimolar amounts of types I and II keratin polypeptides, which are encoded by 28 keratin type I (KRT9, 10, 12–20, 23–28, 31, 32, 33a, 33b, 34–40) and 26 keratin type II genes (KRT1–5, 6a–c, 7, 8, 71–86) (www.interfil.org). Since the expression of specific keratin pairs is a reliable indicator of epithelial differentiation and function, detection of keratins has found wide application in tumor and histodiagnosis (Bragulla and Homberger, 2009; Moll et al., 2008). Our understanding of the contribution of individual keratins to particular epithelial properties, however, is still limited. Yet, it is generally accepted that these functions involve posttranslational modifications including keratin phosphorylation (Kim et al., 2015; Omary et al., 2006; Snider and Omary, 2014). The precise details remain to be elucidated. We first review current knowledge on the modes of keratin phosphorylation and its consequences for the structure and function of the keratin cytoskeleton. We especially emphasize the importance of keratin phosphorylation during cellular stress response. We then summarize the role of keratin phosphorylation in diseases that are caused by toxins, gene mutations, and malignant transformation. All of this is based on our firm belief that the broad range of keratin phosphorylation and its multiple modes of regulation are ideally suited to modulate cytoskeletal plasticity in the context of complex cellular functions and thereby contribute to epithelial physiology and pathology.

2. CHARACTERISTICS OF KERATIN PHOSPHORYLATION The complex patterns of keratin phosphorylation can be considered as a code, which not only reflects the specific status of cellular differentiation and function in a given context but also determines structural and functional properties of the keratin cytoskeleton itself. To understand the basis of these properties, it is important to elucidate the underlying rules governing keratin phosphorylation, some of which will be described in the following paragraphs.

2.1 Keratin Phosphorylation Is Complex, Fast, and Linked to Other Posttranslational Modifications The difficulty in understanding and characterizing the consequences of keratin phosphorylation is that it involves multiple sites, which are recognized by different kinases (Table 1). Some of these sites may even be targeted by multiple kinases depending on the cell type and its functional state (see also Steinert, 1988). These activities are counteracted by phosphatases with a

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Table 1 Different Kinases Known to Phosphorylate Keratins Keratin/Residue and Kinase Tissue/Cell Type References

cAMP-dependent protein kinase

49–69 kD keratins from calf Ikai and McGuire (1983) snout epidermis K8/K18 from rat liver

Yano et al. (1991) and Velasco et al. (1998)

60 kD keratin from guinea Inohara and Sagami (1983) pig epidermis K8-S12/S23/S36/S50 Ando et al. (1996) (major) K8-S8/S33/S42/S416/ S423/S425 (minor) from rat liver Ca2+/calmodulindependent protein kinase

K8/K18 from rat liver

Yano et al. (1991)

Cdc2 kinase

K18-S33 in HT29a cells

Ku et al. (1998a)

ERK1/2

K8-S431 in Panc-1 cellsb and AGS cellsc

Ku and Omary (1997)

Herpes simplex virus2-US3 kinase

K17 in A431 cellsd

Murata et al. (2002)

JNK

K8-S431 in Panc-1b cells

Park et al. (2011)

a

K8-S73 in HT29 cells

He et al. (2002) and Ku et al. (2002a)

MAPK-activated protein kinase (MK2)

K18-S52 in HT29a cells

Menon et al. (2010)

p38 kinase

K8-S73 in HT29a cells and Ku et al. (2002a), Menon et al. A431d cells (2010), and Woll et al. (2007)

p42 kinase

K8-S73 in HT29 cellsa

Ku et al. (2002a)

p90 ribosomal protein S6 kinase 1

K17-S44 in murine keratinocytes and HeLa cellse

Pan et al. (2011)

PKC

K20-S13 in HT29 cellsa

Zhou et al. (2006)

a

K20-S13 in HT29 cells

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Table 1 Different Kinases Known to Phosphorylate Keratins—cont’d Keratin/Residue and Kinase Tissue/Cell Type References

PKCδ

K8-S73 in A549f and rat alveolar epithelial cells

PKCε

K8/K18 in HT29 cellsa

Ridge et al. (2005) Omary et al. (1992) g

K8-S8/S23 in GH4C1 cells Akita et al. (2007) K18-S52 (recombinant peptide)

Tao et al. (2006a)

PKCζ

K18-S33 in A549 cellsf

Sivaramakrishnan et al. (2009)

Raf kinase

K18-S52 (major)

Ku et al. (2004)

K18-S33 (minor) in BHK cellsh Src

K19-Y391 in HT29a and NIH3T3 cellsi

Zhou et al. (2010)

a

Human colorectal adenocarcinoma-derived cell line. Human pancreatic carcinoma-derived cell line. c Human gastric cancer-derived cell line. d Human vulva squamous cell carcinoma-derived cell line. e Human cervix adenocarcinoma-derived cell line. f Human lung adenocarcinoma-derived cell line. g Rat pituitary tumor-derived cell line. h Syrian golden hamster kidney fibroblast-derived cell line. i Murine embryonic fibroblast-derived cell line. b

different spectrum of recognition sites and cell-type specificity (Toivola et al., 1997). Another layer of complexity is added by the different kinetics of the various enzymes. With this highly specialized and adjustable toolbox, slow and fast alterations can be accomplished in defined subcellular locations and in different functional contexts. This is also evident from the variable effects of a large number of reagents on keratin phosphorylation (Table 2). The remarkable speed and efficiency of phosphorylation-dependent processes are exemplified in the observation that disassembly of the keratin cytoskeleton could be induced within less than a minute by an overall inhibition of tyrosine phosphatase activity (Strnad et al., 2002). Even more, short exposure to normal room light sufficed to efficiently prevent keratin network disassembly in this situation (Strnad et al., 2003; Woll et al., 2007). The linkage of phosphorylation to other modifications such as acetylation, glycosylation, ubiquitination, and sumoylation adds another degree of complexity. For example, K8 phosphorylation regulates K8 transamidation

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Table 2 Different Reagents Known to Induce Keratin Phosphorylation Keratin/Residue and Reagent Tissue/Cell Type References

Acetaminophen

K8-S73 and K8-S431 in liver Guldiken et al. of K8-G62C and K8-R431C (2015) transgenic mice

Acetone extract of Bupleurum scorzonerifolium

K8-S73 from A549a cells

Chen et al. (2005)

Acrylamide

PtK1 cellsb

Eckert (1985) and Eckert and Yeagle (1980)

K8/K18 in rat hepatocytes AICAR (5aminoimidazole-4-carboxamide ribonucleoside)

Velasco et al. (1998)

Calyculin A

Keratins from rat parotid acinar cells

Takuma et al. (1993)

cAMP

Types I and II keratins in Deery (1993) canine thyroid epithelial cells

EGF

K8-S431 in HT29 cellsc

Ku and Omary (1997)

K8-S73 in A431 cellsd

Moch et al. (2013)

Estradiol-17 beta

Keratins in rat vaginal epithelium

Gupta et al. (1990)

Ethanol

K8/K18 from rat liver

Sanhai et al. (1999)

MH1C1 cellse

Negron and Eckert (2000)

Forskolin

K8/K18 in CaCo-2 cellsc

Baricault et al. (1994)

Griseofulvin

K8-S79/S436 and K18-S33 in Mallory–Denk bodies of murine liver

Fortier et al. (2010)

Pervanadate

K8/K19 tyrosine in HT29 cellsc and normal mouse colon epithelium K8-S73 from HT29 cellsc

Feng et al. (1999)

K19-Y391 in colon epithelium of human K19-overexpressing mice

Zhou et al. (2010)

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Table 2 Different Reagents Known to Induce Keratin Phosphorylation—cont’d Keratin/Residue and Reagent Tissue/Cell Type References

K19-Y394 in colon epithelium of nontransgenic mice K19-Y391 in HT29c and BHK-21 cellsf K8-S73 in A431 cellsd

Woll et al. (2007)

K8/K18 in HT29 cellsc Phorbol ester 12-Otetradecanoylphorbol-13-acetate (TPA) K8-S431 in Panc-1 cellsb

Chou and Omary (1991)

Prostaglandins

K20-S13 in HT29-MTX cellsg

Menon et al. (2010)

Sorbitol, H2O2, UV light, phorbol ester TPA

K17-S44 in murine keratinocytes, basaloid skin tumors and HeLa cellsh

Pan et al. (2011)

Sorbitol, heat, urea

K8-S73 in A431 cellsd

Woll et al. (2007)

Sphingosylphosphorylcholine

K8-S73 and K8-S431 in Panc-1 cellsi

Park et al. (2011)

Thioacetamide

K8-S431 and K18-S33 in mice liver

Strnad et al. (2008)

Thyrotropin releasing hormone K8-S8/S23 in GH4C1 cellsj

Lee et al. (2014)

Akita et al. (2007)

a

Human lung adenocarcinoma-derived cell line. Rat kangaroo kidney epithelium. Human colorectal adenocarcinoma-derived cell line. d Human vulva squamous cell carcinoma-derived cell line. e Rat hepatoma-derived cell line. f Syrian golden hamster kidney fibroblast-derived cell line. g Goblet cells differentiated from HT29 cell line using methotrexate. h Human cervix adenocarcinoma-derived cell line. i Human pancreatic carcinoma-derived cell line. j Rat pituitary tumor-derived cell line. b c

(Kwan et al., 2012), favors K8 sumoylation (Ku et al., 2010), coincides with changes in K8/K18 glycosylation (Budnar et al., 2010), and also has a counter-regulatory relationship with the neighboring glycosylation residues (Tao et al., 2006a). Conversely, K8 acetylation modulates K8 phosphorylation (Snider et al., 2013).

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2.2 Phosphorylation Occurs Preferentially in the Keratin End Domains Keratin polypeptides share basic structural features with all other cytoplasmic intermediate filament polypeptides: A conserved α-helical rod domain of
310 amino acids is flanked by highly variable end domains (Steinert et al., 1983). The rod domain is involved in heterodimerization of a type I and a type II keratin polypeptide forming the very stable parallel coiled-coil structure through strong hydrophobic interactions (Coulombe and Fuchs, 1990; Hatzfeld and Weber, 1990). The vast majority of phosphorylation sites have been identified in the end domains which may be most accessible to kinases and phosphatases in the cellular environment (Steinert et al., 1982; Steinert, 1978, 1988). As an example, the distribution of phosphorylation sites is depicted for K8 in Fig. 1. At the structural level, NMR studies revealed a conformational change in bovine hoof keratin that was phosphorylated by cAMP-dependent protein kinase (Yeagle et al., 1990). An increase in the rigidity of the otherwise flexible keratin head and tail domains may be responsible for this effect. Furthermore, phosphorylation-induced increased conformational motility as observed in the helical rod domains of ethanol-fed rat liver keratins as determined by circular dichroism and NMR analyses (Sanhai et al., 1999). The end domains of the type II keratins contain the highly conserved subdomain H1 in the head region and the conserved subdomain H2 in

Fig. 1 Scheme depicting identified phosphorylation sites in the human type II keratin K8. Kinases that have been shown to target the respective sites are provided alongside. The scheme highlights the domain structure of K8 consisting of (i) the 90 amino acidlong head domain encompassing the H1 subdomain, (ii) the 308 amino acid-long central rod domain that is subdivided into coils 1A, 1B, and 2 with connecting linkers L1 and L12, and (iii) the 85 amino acid-long tail domain including the H2 subdomain. Note that most phosphorylation sites are located in the head and tail domains.

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the tail region (Fig. 1; Steinert et al., 1985). K5 and K8 mutants lacking the H1 subdomain showed severe filament assembly defects (Hatzfeld and Burba, 1994; Wilson et al., 1992). Not only the protein sequence of the H1 domain but also its secondary conformation consisting of turn, β-strand, Ω-loop, and α-helical segments is highly conserved (Chipev et al., 1992). Thus, phosphorylation within the H1 subdomain may disrupt its structural arrangement. Furthermore, surface lattice models show that the H1 subdomain is critically involved in the alignment of neighboring molecules in keratin filament assembly (Parry and Steinert, 1992; Steinert, 1991a,b).

2.3 Phosphorylation Increases Keratin Solubility Since keratins assemble rapidly and efficiently into higher order structures in vitro, mechanisms must exist to disassemble keratin filaments in vivo during keratin network remodeling. A prime mechanism appears to be phosphorylation. Phosphorylation is expected to prevent assembly of tetramers, which are the major soluble keratin species in cultured cells (Chou et al., 1993). They are composed of two keratin heterodimers that align in an antiparallel and partially staggered fashion and are held together through ionic interactions (Coulombe and Fuchs, 1990; Herrmann et al., 2003, 2009). Phosphorylation presumably prevents the lateral alignment of the nonpolar tetramers into the
60 nm unit length filament (ULF) (Herrmann et al., 2009). In addition, phosphorylation may also interfere with the next assembly stage, i.e., the longitudinal annealing of ULFs. It is assumed that both assembly stages are regulated by the head and tail domains (Hatzfeld and Burba, 1994; Wilson et al., 1992), although a precise molecular understanding is still lacking. Besides involving keratin modification, it may also rely on accessory factors that are recruited or released. It has been known for a long time that the equilibrium between the soluble and filamentous keratin pool is determined by phosphorylation (Feng et al., 1999; Strnad et al., 2002; Woll et al., 2007; Zhou et al., 1999). It is assumed that phosphorylation induces release of soluble tetrameric subunits from keratin filaments. An extreme example was observed in Xenopus oocyte maturation, where phosphorylation of type II keratins led to disassembly of cortical keratin filaments and the generation of keratin oligomers (Klymkowsky et al., 1991). However, another possibility is that phosphorylation preferably targets depolymerized keratin subunits which form the soluble pool of keratins (He et al., 2002). The adjustability in the equilibrium between the soluble and the filamentous state of keratins

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plays an important role in maintaining cytoskeletal pliability. It is rapid, reversible, and can be temporally and spatially regulated. The phosphorylationdependent dynamic properties of the keratin cytoskeleton may thus contribute to cellular plasticity.

2.4 Common Guidelines Determine Keratin Phosphorylation Common principles determine keratin phosphorylation. They act at different levels and are influenced by multiple factors: • Phosphate is mainly incorporated in serine residues of the nonhelical end domains of the keratin molecule (Fig. 1; Gilmartin et al., 1980; Ikai and McGuire, 1983; Steinert, 1988; Sun and Green, 1978). • Increased keratin phosphorylation is the result of more keratin molecules being phosphorylated (Steinert, 1988). • Phosphorylated keratin species are heterogeneous consisting of a mixture of molecules with single site and multiple site phosphorylations (Liao and Omary, 1996). • Individual phosphorylation sites can be phosphorylated by more than one kinase (Fig. 1; Steinert, 1988). • Each kinase phosphorylates keratins in a specific manner. For example, cAMP-dependent protein kinases phosphorylate K8/K18 exclusively at serine residues, whereas Ca2+/calmodulin-dependent protein kinase II targets serine as well as threonine residues (Yano et al., 1991). • Phosphorylation of type II keratins is preferred over phosphorylation of type I keratins (Liao et al., 1995a; Yano et al., 1991). For example, on average four phosphate molecules are incorporated into K8 and two phosphate molecules are incorporated into K18 by cAMP-dependent protein kinase whereas 1 and 0.8 phosphate molecules are incorporated into K8 and K18, respectively, by Ca2+/calmodulin-dependent protein kinase II (Yano et al., 1991). Also, the stoichiometry of phosphorylation is more for K8 than K18 as determined by pulse-chase experiments using radiolabeled phosphate (Liao et al., 1995a). • Although phosphorylation of type I and type II keratins often increases in similar proportion (e.g., phosphorylation of K8/K18 in heat stress or G2/M arrest), it is not directly linked to each other (Chou and Omary, 1993; Liao et al., 1995a). • Keratin phosphorylation is highly dynamic. For example, the increase of K18-S52 phosphorylation by a factor of three in S-phase and four in the G2/M-phase, occurs in a reversible fashion (Liao et al., 1995a).

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3. REGULATION OF PROTEIN BINDING TO KERATINS BY PHOSPHORYLATION Intermediate filament-associated proteins regulate the state of keratin assembly in multiple ways. Thus, they sequester nonfilamentous keratin in the soluble cytoplasmic pool, they increase filament bundling and aggregation, they affect transport along microtubules and they cross-link keratin filaments to the other filament systems and to cell junctions (Listwan and Rothnagel, 2004; Snider and Omary, 2014; Sonnenberg and Liem, 2007). In addition, keratins bind to signaling molecules. The following sections present examples which support the notion that the interactions between keratins and these partners are regulated by phosphorylation.

3.1 Phosphorylation-Dependent Keratin-14-3-3 Protein Association Enhances Keratin Solubility and Affects mTOR Signaling Among the proteins associating with keratins in a phosphorylationdependent manner 14-3-3 proteins have received most attention. They are a ubiquitous family of proteins that play an important role in signal transduction and cell-cycle progression (Aitken et al., 1995; Burbelo and Hall, 1995; Gardino and Yaffe, 2011). 14-3-3 proteins associate preferentially with the hyperphosphorylated cytosolic but not with the less phosphorylated cytoskeletal K8/K18 fraction during the S/G2/M phase of the cell cycle (Liao et al., 1996). It was therefore suggested that 14-3-3 proteins act as solubility cofactors for keratins. It was further shown that 14-3-3 proteins bind to the phosphorylated K18-S33 residue and that the K18S33–14-3-3 interaction contributes to normal mitotic progression (Liao et al., 1996). A complex of solubilized K5/K17/actin was recently shown to be stabilized by 14-3-3σ, enhancing its “bioavailibility” and contributing to polarized assembly during migration, thus, making it highly relevant for breast tumor invasion (Boudreau et al., 2013). Complex formation was shown to be promoted by PKCζ, known to phosphorylate K18 at a 14-3-3 binding site. It was antagonized by phosphatase PP2A, which has been linked to keratin dephosphorylation (Park et al., 2011; Tao et al., 2006b). Hence, keratin phosphorylation likely contributes to regulating the switch between the filamentous and soluble keratin pool.

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In case of K17, whose expression is upregulated in stratified epithelia upon wounding, phosphorylation of K17-T9/S44 is essential for its interaction with 14-3-3σ (Kim et al., 2006). The K17–14-3-3σ interaction ensures the cytosolic retention of 14-3-3σ, which concomitantly stimulates Akt/mTOR activity to support proper cell growth and size. Thus, keratin phosphorylation-regulated 14-3-3 association may affect tissue repair and homeostasis in skin development (Kim et al., 2006).

3.2 Sequestration of Raf Kinase Is Regulated by Keratin Phosphorylation K8 directly associates with Raf kinase, sequestering it during basal conditions to the K8/K18 complex (Ku et al., 2004). During oxidative or chemical stress, however, phosphorylation of K18-S33 and of Raf S338 and S621 occured which led to the release of Raf from the keratin-Raf complex and induced Raf-kinase activation. Moreover, the K18-S52 residue served as a major physiologic substrate of the activated Raf-kinase. The released and activated Raf kinase bound preferentially to 14-3-3 instead of K8 (Ku et al., 2004). This example illustrates how keratin phosphorylation modulates kinase function by sequestration and activation and how the released and activated kinase reciprocally affects keratin phosphorylation.

3.3 Association of Cytolinkers With Keratin Is Influenced by Phosphorylation It was observed that keratin aggregates associate with different plakin protein family members upon phosphatase inhibition. This became evident in studies demonstrating that K13 granules, which were formed by treatment with the tyrosine phosphatase inhibitor orthovanadate in A431 cells, contained the large plakin domain-containing cytolinker plectin (Strnad et al., 2002). In contrast, treatment of A431 cells with the threonine–serine phosphatase inhibitor okadaic acid resulted in keratin granules that were devoid of plectin immunoreactivity. Interestingly, the plectin-positive orthovanadate-induced keratin granules are reversible giving rise to a novel keratin filament network, whereas okadaic acid-induced keratin granules lack this capacity (Strnad et al., 2002). This suggests that plectin may serve as an anchoring or chaperone-like remodeling factor for the keratin network. An association of K8 granules with periplakin, another plakin domaincontaining cytolinker, was observed in the scratch-wound edges of MCF-7 monolayers that were treated with okadaic acid (Long et al., 2006). It was further demonstrated that the periplakin–K8 interaction was involved in

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efficient collective epithelial sheet migration which went along with K8 reorganization and hyperphosphorylation of K8-S431 (Long et al., 2006). In addition, keratin granules induced by either orthovanadate or okadaic acid in murine keratinocytes contain the cytolinker epiplakin (Spazierer et al., 2008). Remarkably, epiplakin only partially colocalized with the keratin filament network in untreated primary mouse keratinocytes but showed a near perfect colocalization after keratin hyperphosphorylation and in stressful conditions such as UV irradiation and osmotic shock, both of which were also linked to increased keratin phosphorylation (Long et al., 2006; Spazierer et al., 2008). Taken together, phosphorylation of keratins appears to regulate their interaction with the different cytolinker proteins.

3.4 Phosphorylation-Dependent Association of the Ubiquitin Ligase Pirh2 With Keratin Affects Cell Survival Phosphorylation-mediated granular aggregate formation of the keratin filament network is known to be a consequence of an altered association of keratin with Pirh2, a p53-induced RING-H2 type-ubiquitin E3 ligase. This was demonstrated in the human lung cancer cell lines H1299 and A549 (Duan et al., 2009). JNK- and p38-mediated phosphorylation of the K8/K18 filamentous network in response to UV irradiation resulted in dissociation of the Pirh2-K8/K18 complex and led to keratin aggregation accompanied by microtubule-dependent mitochondrial redistribution. This resulted in an increased sensitivity to apoptosis (Duan et al., 2009).

4. ROLE OF KERATIN PHOSPHORYLATION IN CELL PHYSIOLOGY Stress–response, mitosis, and apoptosis are major events regulating epithelial cell fate. Each of these events involves climactic situations threatening cell and tissue homeostasis. Phosphorylation-mediated keratin reorganization is common to all these processes and serves diverse purposes in each of these situations as detailed below.

4.1 Mitosis Is Linked to Keratin Phosphorylation and Keratin Network Remodeling Mitosis involves substantial rearrangements of all major structural elements leading to a reversible rounding of the cell. Given the role of phosphorylation in modifying the arrangement of keratin filaments, an elevated level of

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keratin phosphorylation is to be expected during mitosis. This notion was fully supported by experimental observations that are summarized in Table 3 and exemplified in Figs. 2 and 3. Specifically, of the type II keratins K4, K5, K6, and K8 are phosphorylated by mitogen-regulated kinases in proliferating epithelial cells (Table 3) at the highly conserved LLS/TPL motif in the H1 head domain suggesting an evolutionary conserved function of keratin phosphorylation in mitosis (Liao et al., 1997; Toivola et al., 2002). The higher level of phosphorylated keratins in the soluble fraction as compared to the insoluble fraction during mitosis further indicated a role of phosphorylation in shifting the solubility equilibrium of the keratin pool (Baribault et al., 1989; Celis et al., 1983; Chou and Omary, 1993; Fey et al., 1983; Lane et al., 1982). Additionally, keratin filament reorganization into granules is often observed during mitosis (Figs. 2 and 3; Aubin et al., 1980; Baribault et al., 1989; Fey et al., 1983; Horwitz et al., 1981; Jones Table 3 Keratin/Keratin Residues Known to Undergo Phosphorylation During Mitosis Keratin/Keratin Residue Source References

K55/K49 K8S431

Rat hepatocytes a

HT29 cells BHK cells

K18S52

b

Baribault et al. (1989) Liao et al. (1997) Ku and Omary (1997)

a

Liao et al. (1995a)

a

HT29 cells

K18S33

HT29 cells

Ku et al. (1998b) and Ku et al. (2002b)

K4T133c

Human esophageal epithelium

Toivola et al. (2002)

K5T150c

KC cellsd

K6T145c

Human epidermis

c

K8S73

HT29 cellsa

Liao et al. (1997)

Murine mitotic basal crypt cells of Toivola et al. (2002) the intestine Murine regenerating hepatocytes A431 cellse a

Woll et al. (2007)

Human colorectal adenocarcinoma-derived cell line. Syrian golden hamster kidney fibroblast-derived cell line. These residues are located in the conserved LLS/TPL motif of the type II keratins. d Human foreskin primary keratinocytes. e Human vulva squamous cell carcinoma-derived cell line. b c

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Fig. 2 Detection of keratin phosphorylation in mitosis. (A–C) show fluorescence micrographs of human keratin 5-enhanced yellow fluorescent protein chimeras K5-YFP in methanol-acetone fixed human epidermal keratinocyte-derived HaCaT B10 cells (Moch et al., 2013) and a corresponding indirect immunofluorescence micrograph using antibody LJ4 (Toivola et al., 2002) detecting phosphorylated T150 of keratin 5 together with the corresponding nuclear Hoechst staining. (D–F) The fluorescence micrographs depict the distribution of human keratin 13 chimera HK13-enhanced green fluorescent protein (K13-EGFP) in methanol-acetone fixed vulva carcinoma-derived AK13-1 cells (Windoffer and Leube, 1999) and the corresponding indirect immunofluorescence of antibody LJ4 reacting with phosphorylated S73 of K8 (Liao et al., 1997) together with the corresponding nuclear DAPI staining. Note the appearance of the respective keratin phosphoepitopes in granules that are detected in the mitotic cells (labeled by *) of both €ll. Scale cell lines. The images D, E and F were kindly provided by Dr. Stefan Wo bars ¼ 10 μm.

et al., 1985; Lane et al., 1982; Schwarz et al., 2015; Windoffer and Leube, 2001). Mitotic human vulva carcinoma A431 and cervix carcinoma HeLa present both a diffuse and granular cytoplasmic keratin pattern, whereas the keratin network of rat kangaroo kidney-derived PtK2 cells does not disassemble into granules but is only redistributed (Horwitz et al., 1981). Another report showed that the percentage of transformed human epithelial amnion cells containing mitotic keratin granules increased from prophase onwards, reaching a peak during late anaphase/early telophase and plummeting during late telophase. In contrast, normal mitotic amnion cells did not show this characteristic keratin reorganization during mitosis, in spite of similar keratin phosphorylation levels as the transformed cells (Celis et al., 1983). This suggests that additional factors such as mitosis duration

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Fig. 3 Keratin filament network disassembly and reassembly during mitosis. The images in (A–F) show Hoechst stains and corresponding fluorescence of human keratin 5-enhanced yellow fluorescent protein chimeras in methanol-acetone fixed keratinocyte-derived cell line HaCaT B10 (Moch et al., 2013) during different phases of mitosis (A0 –F0 ). Note the reversible keratin granule formation that is induced at late prophase. Accumulation of keratin granules is typically seen in the cleavage furrow (arrows in D0 and E0 ). The structural reorganization is coupled to keratin phosphorylation (see Fig. 2). Scale bars ¼ 10 μm.

or overall kinase activity levels influence keratin reorganization during mitosis. Moreover, preferential phosphorylation of K8 over K18 was observed during mitosis, in rat hepatocytes. However, in this case, phosphorylation of both K8 and K18 upon triton X-100 permeabilization suggested that keratin phosphorylation during mitosis may also depend on the accessibility of their phosphorylation sites (Baribault et al., 1989). Mitotic keratin restructuring may influence the kinetics and efficiency of cell division as has been suggested for vimentin phosphorylation (Ikawa et al., 2014). Moreover, in vivo monitoring of K8 restructuring during the trophoectodermal cell divisions in blastocysts of K8-YFP knockin mice provided direct proof

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for the significance of mitotic keratin reorganization in the native tissue context (Schwarz et al., 2015). Interestingly, A431 and HeLa cells show a concentration of keratin granules in the cleavage furrow during mitosis. Fig. 3D–E0 highlights the enrichment of keratin granules in this region of dividing immortalized human HaCaT keratinocytes. The cleavage furrow harbors several kinases such as Rho-kinase and Aurora B which phosphorylate and thus regulate intermediate filament reorganization, which is essential for efficient cytokinesis (Horwitz et al., 1981; Kawajiri et al., 2003; Kosako et al., 1999; Yasui et al., 1998). Furthermore, p38 mitogen-activated protein kinase, which plays an important role in cell proliferation, colocalizes with keratin granules during mitosis in A431 cells (Woll et al., 2007). This indicates the necessity of a kinase-enriched environment for keratin reorganization during mitosis. It is likely also relevant for other cell types that do not form keratin granules during mitosis. In these instances, the locally increased kinase activities could sever the keratin network at the cleavage furrow for distribution of filamentous keratin into both daughter cells. Moreover, keratins are substrates of mitotic kinases as shown by in vitro phosphorylation of K8-S431 through mitogen-activated protein kinase and cdc2 kinase (Ku and Omary, 1997). The hyperphosphorylation of keratins during mitosis also modulates their binding to associated proteins. This is well demonstrated by the requirement of K18-S33 phosphorylation for an interaction between K18 and 14-3-3 family of proteins, facilitating normal mitotic progression (Liao et al., 1996). Although K18-S33A mice did not have defects in liver regeneration, they displayed abnormal mitotic arrest-related figures such as tripolar and angular mitotic bodies and anomalous proportions of mitotic stages. Furthermore, retention of 14-3-3-ζ in the nucleus of K18-S33A hepatocytes suggested a role of keratin phosphorylation in modulating 14-3-3 distribution, which is an important sequestering and compartmentalizing factor for multiple proteins (Ku et al., 1998a, 2002b; Liao et al., 1996). Thus, keratin phosphorylation during mitosis may not only contribute to cytoskeletal reordering but may also impact the function of keratinassociated proteins.

4.2 The Epithelial Stress Response Is Coupled to Altered Keratin Phosphorylation Stress-activated protein kinase signaling is a hallmark feature of the cellular stress response. Common phosphorylation targets are proteins known for

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their abundance and cytoprotective functions. Because of these characteristics, keratins have been classified as stress proteins (Toivola et al., 2010). Keratins buffer the impact of the activated kinase signaling by acting as a “phosphate sponge,” and actively contribute to molecular defense mechanisms against diverse forms of stress as discussed in the following sections. 4.2.1 Chemical Stress and Keratin Phosphorylation Contribute to Mallory–Denk Body Formation Alcohol, heavy metals, N,N0 -dicyclohexylcarbodiimide (DDC), microcystin-LR (MC-LR) and griseofulvin are hepatotoxic. They induce the formation of Mallory–Denk bodies (MDBs) that are keratin-rich cytoplasmic granular inclusions in hepatocytes (Denk et al., 1979; Franke et al., 1979; Zatloukal et al., 2007). MDBs are also observed in several chronic liver diseases such as alcoholic and nonalcoholic steatohepatitis, chronic cholestasis, metabolic disorders, and hepatocellular neoplasms (Denk et al., 2000; Fausther et al., 2004; Fickert et al., 2003; Stumptner et al., 2000; Zatloukal et al., 2004). MDBs comprise misfolded hyperphosphorylated K8/K18 with phospho-serines at positions 73 and 431 in K8 and at position 33 in K18. Several experimental observations provided evidence that keratin phosphorylation is tightly linked to MDB formation. Thus, treatment of primary cultured rat hepatocytes with the MDB-inducing hepatotoxin, MC-LR, led to a dose-dependent increase in K8/K18 phosphorylation (Guzman and Solter, 2002; Ohta et al., 1992). Similarly, the phosphatase inhibitor okadaic acid increased keratin phosphorylation and induced inclusion body formation in murine liver within 15 min. These inclusions contained keratin aggregates, which stained positive for phosphothreonine and ubiquitin as is the case for typical MDBs (Ohta et al., 1988; Yuan et al., 1998). This observation indicated that keratin hyperphosphorylation plays a role during the early phase of MDB formation. In support, p38MAPK, which phosphorylates K8-S73, was a prerequisite for MDB formation in DDC-fed mice (Nan et al., 2006). Furthermore, autophagy, which leads to clearance of inclusion bodies, was inversely related to K8-S73 phosphorylation (Harada et al., 2008; Kongara et al., 2010), thus, explaining the role of keratin phosphorylation not only in the initiation but also in the continued maintenance of MDBs. Prevalence of MDBs in hepatocyte injury is associated with better tolerance to toxic stress. In accordance, the lack of MDBs in K8 null mice led to increased sensitivity to DDC toxicity (Zatloukal et al., 2000). Furthermore, mice overexpressing the phosphodeficient K18-S52A

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mutant were more susceptible to griseofulvin- and MC-LR-induced liver lesions (Ku et al., 1998b). Hence, keratin hyperphosphorylation is not only a consequence of hepatotoxicity but is part of a cellular defense mechanism during liver intoxication. 4.2.2 Chemical Stress and Phosphorylation Alter the Assembly State of Keratins Another consequence of increased levels of hyperphosphorylated keratins in MDB-containing hepatocytes is an elevation in keratin solubility (Fortier et al., 2010). Although keratin filaments isolated from MDBs assembled in the same manner as normal hepatocellular keratins, the soluble pool of keratins from griseofulvin-treated mouse livers was polymerization incompetent. This was linked to a rise in the acidic isoelectric variants of keratins in the soluble pool, presumably due to increased phosphorylation (Pollanen et al., 1994; Salmhofer et al., 1994; Toivola et al., 1998). A prominent feature of MDB persistence is transglutaminase-mediated covalent crosslinking of K8 (Strnad et al., 2007). The consensus recognition sequence for transglutaminases is QXXφDP (Sugimura et al., 2006). The preferred transamidation site in K8 is Q69, which lies in proximity to the phosphorylation site K8-S73 (69QSLLSP74). The phosphorylation of K8-S73 may mimic an aspartic residue and thereby creates a near-perfect transglutaminase recognition site. This is supported by the observed reduced K8 crosslinking in human colon carcinoma-derived HT29 cells expressing phosphodeficient K8-S73A when treated with the phosphatase inhibitor okadaic acid. Moreover, K8-S73A mice have a reduced ability to form MDBs (Kwan et al., 2012). Since K8 is a better substrate for transamidation than K18, phosphorylation-regulated K8 crosslinking may be an important mechanism to maintain the high K8–K18 ratio observed in Mallory bodies (Nakamichi et al., 2005). It is assumed that phosphorylation ensures the maintenance of the aberrantly folded keratins in the inactive state. Moreover, the association of phosphorylated K8-S79 (corresponding murine residue to human K8-S73) with activated p38MAPK suggested that keratin phosphorylation plays a role in sequestering excessive kinase activity (Fortier et al., 2010). Interestingly, p38MAPK activity has been directly implicated in keratin aggregate formation in cultured cells (Fig. 4; Woll et al., 2007). It will be interesting to find out why keratin phosphorylation has two fundamentally different effects on keratin organization, namely increasing keratin solubility on one hand and increasing crosslinking, on the other hand, and how these opposing effects are regulated.

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Fig. 4 Inhibition of p38 MAPK prevents vanadate-induced keratin granule formation. Confocal laser scanning microscopy of methanol/acetone-fixed AK13-1 cells producing fluorescent human keratin 13-enhanced green fluorescent protein chimera HK13-EGFP (Windoffer and Leube, 1999). Cells were treated with orthovanadate (OV) at 15 mM for 15 min after a preincubation with the p38 inhibitor SB202190 in (A) and at 10 mM for 5 min in the absence of SB202190 in (B). Note that the typical interphase morphology of the keratin network is preserved in (A) but not in (B) despite the lower vanadate concentration and shorter treatment period. The images were kindly provided by Dr. Stefan €ll. Scale bars ¼ 10 μm. Wo

4.2.3 Heat Stress Leads to Recruitment of Chaperones to Phosphorylated Keratin Resulting in Keratin Aggregate Formation Administration of heat stress to cell lines producing mutant keratins that are known to cause the skin blistering disease epidermolysis bullosa simplex (EBS) led to an increased stimulation of the stress-activated protein kinases p38, JNK1/2, and ERK activity (Chamcheu et al., 2011a). This heatinduced kinase overload is accompanied by increased K5 phosphoepitopes, which colocalise with HSP70 (Chamcheu et al., 2011a). The heat-shock response went along with an elevation in keratin aggregation, the extent of which depends on disease severity (Chamcheu et al., 2011a). Likewise, a heat stress of 42°C for 16 h in HT29 cells or of 60°C for 5 min in A431 cells induced the formation of hyperphosphorylated K8-S73 in granular aggregates (Liao et al., 1995b, 1997; Woll et al., 2007). This provided further evidence for the role of hyperphosphorylated keratins in cellular stress responses. Moreover, the hyperphosphorylated keratins associated with heat-shock proteins and up to 50% of the cells contained clumped keratin filaments (Liao et al., 1995b). But purified K8/18 proteins from heatstressed cells were able to form normal filaments in vitro (Liao et al., 1995b)

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indicating that cellular factors are responsible for the altered keratin organization in heat-shocked cells. It was reported that HSP70 acts as a cochaperone for K8/K18 filaments along with HSP40 (Izawa et al., 2000). Moreover, HSP27 has the potential to modulate K8/K18 assembly dynamics (Kayser et al., 2013). In addition to their innate ability of supervising protein folding, the heat-shock proteins also have a potential role in regulating keratin (K5, K14, and K10) ubiquitination (Yamazaki et al., 2012) thereby affecting the half life of keratins during heat stress. This also appears to be relevant for the clearance of cytoplasmic aggregates containing EBS-mutant keratins (Loffek et al., 2010). Furthermore, the heat-induced architectural rearrangement of the keratin cytoskeleton also coincided with translational inhibition, polysome disruption, and organelle translocation in immortalized mouse mammary epithelial cells (Shyy et al., 1989) suggesting functions of keratins beyond providing a phosphate sink during stress. 4.2.4 Mechanical Stress Is Linked to Keratin Phosphorylation and Keratin Network Reorganization Phosphorylation of keratins displayed a direct relationship with mechanical stress. Upsurge in K8-S73 and K18-S33 phosphorylation due to PKCδ and PKCζ activation, respectively, was shown to be an outcome of shear stress in human and rat alveolar epithelial cells (AECs) (Sivaramakrishnan et al., 2008). Moreover, the extent of phosphorylation in these residues showed a shear force-dependent increase. The resistance to shear stress increased from the perinuclear to the peripheral region in AECs as determined by storage modulus measurements using particle tracking microrheology. This correlated with decreasing keratin phosphorylation and increasing mesh size (Flitney et al., 2009; Sivaramakrishnan et al., 2008). Phosphorylation of keratins in shear stress resulted in increased bundling of the keratin network. Moreover, shear stress of 7.5–30 dynes/cm2 for 0–24 h induced the formation of ubiquitin-positive aggregates in the perinuclear space of the AECs. Furthermore, an increase of the triton X-100 soluble keratins was observed. This was accompanied by phosphorylation of K8-S73 which served as a recognition factor for the ubiquitin-mediated keratin degradation that occurred under shear stress (Ridge et al., 2005; Sivaramakrishnan et al., 2009). Thus, keratin phosphorylation during shear stress may not only act as a keratin reorganizing factor, but also as a mechanism to prevent the subsequent accumulation of ubiquitinated aggregates (Johnston et al., 1998; Kopito, 2000; Zatloukal et al., 2007). Furthermore, K18-S33 phosphorylation was shown

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to be required for association of K18 with 14-3-3ζ which led to keratin network rearrangement during shear stress. The reduced resistance of K18S33A-containing AECs to shear force further confirmed the need for keratin phosphorylation in providing a resilient cytoskeletal defense against mechanical force (Flitney et al., 2009; Sivaramakrishnan et al., 2008, 2009). A 10% increase in the cell surface area (CSA) by mechanical stretch was shown to lead to considerable straightening of the keratin filaments in primary cultures of pulmonary alveolar type II (AII) cells. This was accompanied by a widening of the extracellular gap of desmosomes (Felder et al., 2008). A 20% increase in CSA, however, restored keratin filament waviness and desmosomal gap width. This paradox was explained by increased keratin phosphorylation, notably of K8-S431, and the observed decrease in keratin bundles with concurrent increase in thinner filaments as determined by transmission electron microscopy. It remains to be shown whether this is a general keratin-dependent mechanoprotective mechanism. 4.2.5 Osmotic Stress Induces Kinase-Dependent Alterations of the Keratin Cytoskeleton Similar to the various types of stress discussed so far, osmotic shock also induces hyperphosphorylation of keratins. As demonstrated in human colorectal carcinoma HRT18 and CaCo-2 cells, hyper- and hypoosmotic stress increased phosphorylation of K8-S73 and K8-S431 (Tao et al., 2006b). Furthermore, increased K8–73 phosphorylation was linked to formation of keratin aggregates that contain phosphorylated active p38 MAPK in osmotically challenged human vulva carcinoma A431 cells (Woll et al., 2007). A more complex situation was encountered in HT29 cells. While K8-S431 was hyperphosphorylated upon hyperosmosis, it was dephosphorylated in hypoosmosis. This may be explained by the association of the phosphorylated K8-S431 with protein phosphatase 2A (PP2A) which occurs uniquely under hypoosmosis in these cells (Tao et al., 2006b). Furthermore, 605 mM sorbitol induced K17-S44 phosphorylation in murine skin keratinocytes by p90 ribosomal S6 kinase1, which is downstream to the MAPK cascade (Pan et al., 2011). 4.2.6 Microbes Elicit Keratin Phosphorylation Leading to Keratin Network Reorganization Formation of a kinase-rich environment is a common consequence of viral infection. Hepatitis C virus and coxsackievirus B4 infections in liver and pancreas, respectively, have been associated with K8/K18 hyperphosphorylation

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(Toivola et al., 2004, 2009). Also, rotavirus infection in human colonic cell lines showed phosphorylation-mediated K8/K18 network alterations leading to a more hazy K8/K18 staining pattern and keratin network rearrangement in differentiating CaCo-2 cells (Brunet et al., 2000; Liao et al., 1995b). Besides, association of high viral titers with partially disrupted pancreatic keratin network was observed upon coxsackievirus B4 infection (Toivola et al., 2009). Similarly, Herpes simplex virus-2 US3 (HSV-2 US3) protein kinase expression resulted in a decrease in filamentous K17 along with an increase in K17 phosphorylation (Hertel, 2011; Murata et al., 2002). It still has to be sorted out in each case whether the phosphorylation-mediated keratin reorganization during viral infection is a cellular strategy to override the infectious attack or, contrariwise, a viral strategy to enhance pathogenicity. The following reports portrayed another scenario of virus-induced keratin network reorganization. It was observed that human papilloma virus HPV-16 invasion led to a complete collapse of the keratin network in human HaCaT keratinocytes, in human SiHa cervical epithelial cells, and in low grade cervical neoplasia in vivo (Doorbar et al., 1991; McIntosh et al., 2010). The HPV16 E1–E4 proteins not only bind strongly to K18 and weakly to K8 through their YPLLKLL amino terminal peptide but also bind to each other through their carboxytermini (Wang et al., 2004). They therefore function as efficient keratin crosslinkers. Moreover, the filamentous pool of K18 in the HPV 16-infected SiHa cells showed a high degree of K18-S33 phosphorylation but was presumably unable to enter the soluble pool because of the E1–E4 crosslinking. In accordance, expression of HPV16 E1–E4 in SiHa cervical epithelial cells restricted K18 to the insoluble pool even in G2/M arrested and okadaic acid-treated cells both of which favor keratin solubilization and association of phosphorylated keratin with 14-3-3 protein (Wang et al., 2004). Similarly, it was reported for HaCaT cells that persistence of HPV16 E1–E4 protein induced a timedependent increase of p38 MAPK and JNK activation as well as an elevation of keratin phosphorylation at K8-S73, K8-S431, K18-S33, and K18-S52 (McIntosh et al., 2010). Furthermore, time-lapse recordings of K13 in HaCaT cells showed a dramatic reduction in keratin dynamics in the presence of HPV16 E1–E4 (McIntosh et al., 2010). Another situation of phosphorylation-dependent keratin restructuring has been described for the nerve parasite Spraguea lophii (Weidner and Halonen, 1993). Its outer envelope is stabilized by K4/K13. The keratin envelope undergoes phosphorylation and disassembly at the time of spore activation in a calmodulin- and calcium-dependent manner. When spore

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activation was blocked, keratin phosphorylation and disassembly were also inhibited. Thus, keratin phosphorylation appears to enhance discharge of infective sporoplasm into the target cells.

4.3 Keratin Phosphorylation Protects Against Ubiquitin-Dependent Degradation and Reduces Sensitivity to Apoptosis Keratin phosphorylation is an early event in apoptosis (Liao et al., 1997). This becomes evident in cells treated with the apoptosis-inducing drug anisomycin. Hyperphosphorylated K8-S73 was detected in A431 cells within 5 min of anisomycin treatment (Woll et al., 2007) and hyperphosphorylated K8-S73, K8-S431, and K18-S52 were identified in HT29 cells after 30 min of anisomycin addition (Ku et al., 1997). Interestingly, hyperphosphorylation of K8-S73 and K8-S431 was also observed in response to Fas- and TNF-induced apoptosis in mice (Ku et al., 2003). Furthermore, K20-S13 phosphorylation was noted during apoptosis as a result of chemically-induced mouse colitis (Ku et al., 1997; Zhou et al., 2006). Fig. 5 presents an example of spontaneously occurring apoptosis in cultured human HaCaT keratinocytes depicting the granule formation of a fluorescent keratin reporter. The newly formed keratin granules become positive for phosphorylated K5-T150. The execution of apoptosis requires caspase-mediated cleavage of multiple cellular proteins. Keratins K14, K18, K19, and K20 harbor caspase target sites (VEV/MDA/S) in their rod domain. K18 contains the additional caspase cleavage sequence DALDS in its tail domain (Ku and Omary, 2001; Ku et al., 1997; Zhou et al., 2006). This explains why caspase activation leads to a collapse of the keratin filament network which is followed by the formation of large cytoplasmic inclusions. It was observed in TRAIL-induced apoptosis in the breast carcinoma MCF cell line that these cytoplamsic inclusions contain hyperphosphorylated keratins (K18-S52) together with caspase-cleaved keratin fragments and catalytic caspase subunits (MacFarlane et al., 2000). Although phosphorylation of keratins preceded caspase activation and phosphorylated keratins colocalized with caspasecontaining apoptotic aggregates, keratin phosphorylation did not increase its propensity to be digested by caspases (Schutte et al., 2004). In agreement, induction of apoptosis in the absence of K8-S73 phosphorylation by simultaneous treatment with the apoptosis-inducing cyclin-dependent kinase inhibitor roscovitine and the broad-reactive protein kinase inhibitor staurosporine did not prevent the appearance of apoptotic inclusions

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Fig. 5 Detection of keratin phosphorylation during apoptosis. (A–D) show corresponding phase contrast (A) and fluorescence images (B–D) of methanol-acetone fixed immortalized human epidermal keratinocyte-derived HaCaT B10 cells (Moch et al., 2013). (B) Depicts nuclear Hoechst staining, (C) the distribution of human keratin 5-enhanced fluorescent protein chimeras K5-YFP, and (D) the distribution of phosphorylated K5-T150 as detected by immunofluorescence using antibody LJ4 (Toivola et al., 2002). Note the altered morphology of the apoptotic cell marked by * along with nuclear condensation (B) and formation of keratin granules (C) that are enriched in keratin phosphoepitope K5-T150p (D). Scale bar ¼ 10 μm.

containing caspase-cleaved keratin fragments (Schutte et al., 2004). Yet, keratin phosphorylation may enhance the caspase-mediated disintegration of keratin filaments into aggregates. Ubiquitination, which is closely related to apoptosis, is influenced by keratin phosphorylation. N-[N-(N-acetyl-L-leucyl)-L-leucyl]-L-norleucine (ALLN)-mediated proteasome inhibition in HT29 cells revealed the presence of low amounts of phosphorylated K8-S73, K8-S431, and K18-S52 in the ubiquitinated keratin fractions (Ku and Omary, 2000). The extremely low stoichiometric ratio of phosphorylated K8-S73/S431 to the total

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ubiquitinated K8, however, indicated that keratin phosphorylation and ubiquitination are inversely related. This notion was supported by the observation that keratin degradation was increased in HT29 cells producing phosphodeficient K8-S73A mutants (Ku and Omary, 2000). K8-S73 is contained in the sequence motif 69QSLLSPL75 that is conserved in type II keratins. Mutation of K8-P74 to K8-L74, which prevents phosphorylation of K8-S73 by the proline-directed kinase p38, elevated K8 ubiquitination. On the other hand, mutation of K8-L71 to K8-P71, which creates a new potential phosphorylation site at K8-S70, led to decreased K8 ubiquitination (Ku and Omary, 2000). Interestingly, the antiapoptotic protein P-elementinduced wimpy testis (PIWI)-like 2-induced K8-S73 phosphorylation through p38MAPK and led to a decrease in ubiquitin-mediated K8 degradation (Jiang et al., 2014). Taken together, these observations demonstrated that keratin phosphorylation protects keratins from degradation during apoptosis. Furthermore, it was proposed that the maintenance of keratin levels reduces cellular sensitivity to Fas-mediated and TNF-induced apoptosis (Caulin et al., 2000; Gilbert et al., 2001; Lee et al., 2013).

5. DISEASE RELEVANCE OF KERATIN PHOSPHORYLATION 5.1 Keratin Phosphorylation Affects Keratin Network Organization and Function in Simple Epithelia K8/K18 is the major keratin pair of the glandular epithelia in liver, exocrine pancreas, and intestine. Animal model systems expressing mutant versions of these simple epithelial keratins have been successful in defining the functional role of keratins in these organs (Ku et al., 1995, 2002b; Ku and Omary, 2006; Toivola et al., 2008). They also helped to elucidate aspects of keratin phosphorylation which will be described in the following sections. Besides, animal models of renal tubular epithelial cell injuries have also shown K8/K18 phosphorylation at K8-S73 and K18-S33 (Djudjaj et al., 2016). 5.1.1 Altered Keratin Phosphorylation Affects Keratin Network Organization and Stress Resilience in Hepatocytes Induction of extensive liver damage by administration of serine/threonine phosphatase inhibitors serves as a paradigm for serious liver injury (Fujiki and Suganuma, 1993; Holmes and Boland, 1993). It has been demonstrated that the phosphatase inhibitor MC-LR increased hyperphosphorylation of

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K8/K18 which resulted in perturbation of the keratin filament network in hepatocytes. It led to a time-dependent solubilization of keratin filaments. Furthermore, phosphopeptide maps of the phosphorylated keratins implicated Ca2+/calmodulin-dependent kinase (Toivola et al., 1997). Similarly, K8-S73 and K8-S431 were hyperphosphorylated in acetaminopheninduced acute liver failure, which was accompanied by enhanced keratin solubility and perturbed keratin network formation (Guldiken et al., 2015). K18-R89 is a highly conserved residue among type I keratins. Mutation of the corresponding residue in K14, K10, and K9 is associated with EBS, epidermolytic hyperkeratosis (EH), and epidermolytic palmoplantar keratoderma (EPPK), respectively (Fuchs, 1994; McLean and Lane, 1995). It is generally assumed that these mutations, which are located at the beginning of the helical rod domain within the helix initiation motif, directly interfere with proper keratin polymerization. Mutation of K18-R89 to K18-C89 alters the consensus sequence 89RXXS92 for calmodulin-dependent protein kinase and protein kinase C to 89CXXS92 thereby inactivating the potential phosphorylation site S92 (Pearson and Kemp, 1991). But so far there is no experimental evidence that K18-S92 is targeted by kinases (Ku et al., 1995). Yet, expression of K18-R89C led to altered phosphorylation at other sites of K18 and of other keratin polypeptides (Ku et al., 1995). Thus, transgenic mice expressing K18R89C presented elevated phosphorylation of K18-S33 and K8-S73 with extensive alteration of the keratin filament network in hepatocytes (Ku et al., 1995). This was accompanied by hepatocyte fragility, inflammation, and necrosis whereas the pancreas showed no histopathological changes. Phosphoglycosylated K8/K18 was detected in the soluble fraction of the cultured cells and mouse tissues producing K18-R89C mutants but not in control cells and tissues expressing wild-type K8/K18. Moreover, K18-R89C mice exhibited a higher sensitivity to MC-LR-induced liver toxicity in comparison to mice overexpressing wild-type K18. The MC-LR treatment of K18-R89C mice was associated with a time-dependent increase of phosphorylated K18 in the soluble fraction. The keratin-solubilizing effect correlated with keratin phosphorylation and may play a major role in modifying the effects of K18-R89C by influencing its binding to associated proteins (Ku et al., 1995). Furthermore, hepatocytes of K18-R89C mice were more susceptible to Fas-mediated apoptosis than hepatocytes from mice overexpressing wild-type K18 (Ku et al., 2003). As compared to nontransgenic mice and mice overexpressing wild-type K18, the K18-R89C mice also showed increased susceptibility to thioacetamide-induced liver injury and fibrosis (Strnad et al., 2008).

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5.1.2 The Pancreas Is Sensitive to Changes in Keratin Expression Levels Leading to Altered Keratin Phosphorylation Similar to the situation in liver, cytoplasmic keratin filament reorganization was observed in the exocrine pancreas of K18-R89C mice. But no significant histopathological or functional alterations were detectable (Toivola et al., 2008). This may be due to the persistence of intact apicolateral keratin bundles in the pancreas of the K18-R89C mice in contrast to the network disruption in hepatocytes (Ku et al., 1995). Yet, pancreatic keratins also react to pharmacological stress by hyperphosphorylation as demonstrated in mice treated with caerulein, which induced pancreatic injury and increased phosphorylation of keratins 8 and 18 (Toivola et al., 2008). Elevating K8 or K18 alone in the pancreas did not change the histology in comparison to the wild type. Yet, pancreatic K8 overexpression was accompanied by hyperphosphorylation of K18–S33, a 14-3-3 binding site that is crucial for mitotic progression (Liao and Omary, 1996). In addition, K8-overexpressing pancreas displayed elevation of K18, thus differing from pancreas overexpressing K18 which does not show any alterations in K8 levels (Liao and Omary, 1996). Furthermore, pancreatic overexpression of both K8 and K18 exhibited prominent pancreatic alterations with ageenhanced vacuolization and atrophy of the exocrine pancreas. In addition to phosphorylation of K18-S33 increased phosphorylation of K8-S73 was also noted in these mice (Liao and Omary, 1996). As described above, K8-S73 is phosphorylated in stress, mitosis, and apoptosis (Omary et al., 2006). Increased mitosis and apoptosis are unlikely reasons for the increase in K8-S73 phosphorylation in these mice, because no change in 14-3-3 distribution, in proliferating nuclear antigen expression or in the amount of caspase-cleaved K18 or K19 fragments was detected. This leaves stress as the most likely cause of keratin hyperphosphorylation in mice overexpressing K8 and K18 and suggests that elevated keratin is by itself a stressor. In K8/K18-overexpressing mice K18-S33 phosphorylation is not restricted to the apicolateral keratin filaments of pancreatic acinar cells as is the case in the wild type but extends to cytoplasmic filaments (Ku et al., 2002b). Also, K18-S33A-overexpressing mice show more dispersed distribution of keratin filaments in pancreatic acini than in wild-type K18 transgenic mice (Ku et al., 2002b). Also, the K8/K18-overexpressing murine pancreata show a decrease in size but an increase in the number of zymogen granules. Similar pancreatic secretion defects were observed in mice lacking syncollin, a zymogen granule-interacting protein (Wasle et al., 2005). The function of keratin phosphorylation is not limited to

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exocrine pancreas. Thus, phosphorylation of a 60 kDa keratin was described in hamster insulinoma cells during insulin release concurrent with depolarization-induced calcium influx (Schubart et al., 1980, 1982). This observation hints toward a contribution of Ca2+-activated protein kinasemediated keratin phosphorylation to regulated insulin release. 5.1.3 Keratin Phosphorylation Is Linked to Liver Disease Progression The link between keratin phosphorylation and liver disease progression was investigated in transgenic mice expressing K8-G61C, a K8 variant that is associated with cryptogenic and noncryptogenic liver disease in humans (Ku et al., 2001, 2002a; Ku and Omary, 2006). K8-G61 lies in the highly conserved H1 subdomain of the K8 head region. The K8-G61 ! K8-C61 mutation leads to more crosslinking of K8 upon oxidative challenge (Ku and Omary, 2006). The aberrant crosslinking of K8 reduces the accessibility of K8-S73 to p38MAPK, which is responsible for K8 hyperphosphorylation during apoptosis, stress, and mitosis (Ku and Omary, 2006; Liao et al., 1997). Therefore, K8-G61C transgenic mice exhibited increased predisposition to MC-LR-induced liver injury and Fas-mediated apoptosis as compared to mice expressing only wild-type K8 (Ku and Omary, 2006). Although the keratin filament collapse during Fasmediated apoptosis in K8-G61C transgenic mice is comparable to that seen in wild-type K8 mice, the elevation of K8-S73 phosphorylation upon Fas stimulation was
65% less than that of wild-type K8-expressing mice (Ku and Omary, 2006). This confirms the inhibitory effect of K8-G61C mutation on K8-S73 phosphorylation. The attractive phosphate sponge concept put forward by the Omary Lab suggests that the keratin cytoskeleton neutralizes surplus kinase activity (Ku and Omary, 2006). Thus, inhibition of K8-S73 phosphorylation by p38 MAPK in K8-G61C mice may shunt the kinase activity to other targets such as cJun, cAMP response element-binding protein (CREB), or p90RSK which have been shown to arouse apoptosis upon phosphorylation (Baines and Molkentin, 2005; Deak et al., 1998; Roux and Blenis, 2004). The significance of site-specific keratin phosphorylation for progression of liver disease was delineated in an investigation of hepatocytic events in chronic hepatitis patients (Shi et al., 2010). The analyses revealed elevated phosphorylation of K18-S33 and K18-S52 in cirrhotic and noncirrhotic hepatitis B correlating with increasing liver lesions (Shi et al., 2010). While K18-S52 phosphorylation increased parallel to liver injury progression, K18-S33 phosphorylation did not show such a tight correlation. Furthermore, K18-S52 phosphorylation increased in parallel to alanine aminotransferase (ALT)

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activity, whereas K18-S33 phosphorylation upsurged irrespective of ALT activity. It was suggested that K18-S52 serves as a hepatocytic protective factor in hepatitis B whereas K18-S33 phosphorylation may serve as an early indicator of hepatitis B virus infection. Yet, both K18-S33 and K18-S52 phosphorylation are dependable markers of chronic hepatitis B progression (Shi et al., 2010). 5.1.4 Keratin Phosphorylation Affects Keratin Network Organization and Function of Intestinal Epithelial Cells Treatment of human colon carcinoma-derived CaCo-2 cells with forskolin, which is known to increase cAMP levels, led to the redistribution of keratin filaments from the cell periphery to the cell interior (Baricault et al., 1994). This change in the spatial organization of the keratin network correlated with keratin hyperphosphorylation. 2D gel electrophoresis of the 32Plabeled keratin extracts revealed increased phosphorylation of K8, K18, and K19 and appearance of additional isoelectric variants of K8. Morphologically, forskolin treatment resulted in reduction of intercellular spaces and shortening of microvilli. In addition, decreased expression of hydrolases was observed in the apical brush border of forskolin-treated confluent CaCo-2 monolayers (Baricault et al., 1994). The following studies shed further light on the functional role of keratin phosphorylation in the intestine. K20-S13 phosphorylation, which positively responds to PKC activation, was shown to be essential for keratin filament reorganization. Thus, introduction of K20-S13A together with wild-type K8 into fibroblast-derived NIH-3T3 cells prevented okadaic-induced keratin filament network breakdown (Zhou et al., 2006). Additionally, the observed hyperphosphorylation of K20-S13 in apoptosis, starvation, and DSS-induced colitis can be taken as an indication of its physiological relevance. Besides, the preferential phosphorylation of K20-S13 in goblet cells of the murine small intestine makes it a unique intestinal goblet cell marker (Zhou et al., 2006). Furthermore, K20-S13 is also a physiological target of the MAPK-activated protein kinase (MAPKAP) MK2. Pharmacological blockade of MK2/3 or p38 MAPK inhibited prostaglandin-mediated mucin secretion in differentiated HT29-MTX cells, a mucus-secreting HT-29 subpopulation. Thus, p38/MK2/3-dependent K20 phosphorylation may be involved in mucin secretion of intestinal epithelia (Menon et al., 2010).

5.2 Keratin Phosphorylation Is Altered in Skin Diseases Keratin gene mutations are linked to many skin diseases (Chamcheu et al., 2011b; Stevens et al., 2000). A hallmark feature of these diseases is the

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Fig. 6 Detection of keratin phosphorylation in aggregates containing mutant keratin. Human keratinocyte-derived cells of line HaCaT were transiently transfected with a cDNA construct coding for keratin mutant K14-R125C, which is known to cause the skin blistering disease epidermolysis bullosa simplex. The mutant keratin was tagged at its aminoterminus with enhanced yellow fluorescent protein. (A) The micrograph depicts the fluorescence elicited by the fluorescence-tagged mutant keratin (YFP-K14-R125C) together with a nuclear Hoechst stain. (B) Presents a corresponding fluorescence micrograph showing the distribution of phosphorylated K5 T150 as detected by primary antibody LJ4 (Toivola et al., 2002) together with the nuclear Hoechst stain. Note the preferential localization of the keratin phosphoepitope in granules and thickened filament bundles. Scale bar ¼ 10 μm.

formation of keratin aggregates (e.g., Fig. 6), which aggravates upon mechanical stress. In addition, there is accumulating evidence that these structural alterations are linked to changes in keratin phosphorylation. We will describe this aspect in detail in the following sections. 5.2.1 Increased Keratin Phosphorylation Is Related to Keratin Network Dynamics in Skin Disease EBS, an autosomal dominant skin disorder, manifests itself in the form of trauma-induced epidermal basal cell lysis leading to skin blisters (Coulombe et al., 1991b). Several studies hint toward the contribution of keratin phosphorylation in the pathogenesis and progression of EBS (Chamcheu et al., 2011a; Chan et al., 1993; Chipev et al., 1992). Weber–Cockayne EBS (EBS-WC) is a mild form of EBS which has been linked to missense mutations in the H1 subdomain of the K5 head region and the nonhelical linker segments of the K5 coiled-coil domain (Albers and Fuchs, 1992). The K5-I161 to K5-S161 mutation in the K5 H1

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subdomain of EBS-WC patients leads to subtle blistering of the palmar and plantar skin upon physical trauma (Chan et al., 1993). The substitution of isoleucine by serine creates the new potential PKC phosphorylation site S-Q-R (Woodgett et al., 1986). PKC phosphorylation-mediated keratin remodeling upon mechanical stretch has been discussed in Section 4.2.4. Furthermore, it has been suggested that structural alterations due to the additional phosphorylation site in the H1 subdomain may interfere with the lateral association of keratin dimers during filament assembly (Chipev et al., 1992; Parry and Steinert, 1992; Steinert, 1991a,b). Besides, K5-I161 is next to the helix initiation motif of the coiled-coil rod domain, which is crucial for proper filament formation (Stewart et al., 1989). In line with these theoretical assumptions, ultrastructural analyses of EBS-WC patient-derived keratinocytes revealed mild aggregation and waviness of keratin filaments in basal cells along with nuclear distortions (Chan et al., 1993). Taken together, these observations illustrate how a single keratin phosphorylation site impacts keratin network organization with consequences for cellular stability. Dowling-Meara EBS (EBS-DM) is more severe than EBS-WC. It is known to be caused by missense mutations in either the helix initiation motif or the helix termination motif of the K5 or K14 rod domains. EBS-DM patient-derived epidermal keratinocytes display keratin aggregates and withdrawal of the keratin network from the plasma membrane (Coulombe et al., 1991a; Fine et al., 1991; Ishida-Yamamoto et al., 1991; Letai et al., 1993). Epithelial cell lines expressing the DM-EBS YFP-labeled mutant K14-R125C showed that a high percentage of the mutant K14 was in the soluble pool (Werner et al., 2004). The abundance of phosphoepitopes may safeguard the soluble mutant keratin against ubiquitinmediated degradation, as discussed earlier (Jiang et al., 2014). In addition, K14-R125C-positive aggregates increased upon proteasome inhibition. Conversely, they were reduced by chaperone-associated ubiquitin ligase CHIP/STUB1 through an elevated degradation of the mutant K14 (Loffek et al., 2010). Besides, K14-R125C aggregates (see Fig. 6 as a typical example) turned over rapidly with a half time of less than 15 min (Werner et al., 2004). This was interpreted as a consequence of perturbed keratin assembly (Windoffer et al., 2011). While the mutant keratin polypeptides initiate the assembly process in the cell periphery, in close proximity to focal adhesions as is the case for wild-type keratins (Windoffer et al., 2006), they do not elongate properly. Instead of forming filamentous structures, they generate growing

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granules (Windoffer et al., 2004). Both, elongating filaments and growing granules are transported toward the cell interior with the help of actin filaments and microtubules (Kolsch et al., 2009; Werner et al., 2004). Yet, wildtype filaments are integrated into the peripheral keratin network to complete their prolonged turnover cycle (Windoffer et al., 2011), whereas the mutant keratin granules are incapable of such integration and, instead, dissolve to enter another futile cycle of assembly and disassembly (Kolsch et al., 2010; Moch et al., 2013; Windoffer et al., 2004). 5.2.2 Keratin Phosphorylation Is Increased in Stressed Keratinocytes Producing Mutant Keratins Mutant keratin filaments in immortalized keratinocytes from patients with EBS-DM (K5 E475G and K14-R125P) and EBS-WC (K14 V270M) exhibit thermoinstability with progressive keratin filament disassembly and aggregate formation after 15 min exposure to 43°C unlike the keratin filaments in control cell lines derived from unaffected individuals (Morley et al., 2003). Furthermore, heat stress not only increased keratin aggregate formation in EBS-DM patient-derived keratinocyte cell lines carrying a K5-E475G mutation but also led to an increased level of K5 phosphoepitopes (Chamcheu et al., 2011a). Moreover, the phosphorylated EBS-DM keratins were associated with HSP70. Treatment of the mutant keratinocytes with the chemical chaperone trimethylamine-N-oxide (TMAO) reduced keratin granules as did kinase inhibition. This suggests a potential role of keratin phosphorylation in the misfolded protein response (Chamcheu et al., 2011a). Furthermore, osmotic shock of the immortalized keratinocyte cell lines, that were derived from DM-EBS and WC-EBS patients, showed more pronounced keratin aggregation than the control cell line NEB-1, that was derived from a healthy individual (Beriault et al., 2012; D’Alessandro et al., 2002). Moreover, cell lines with clinically severe EBS mutations were more sensitive to osmotic stress and showed faster activation of the SAPK/JNK pathway (D’Alessandro et al., 2002). Also, gene expression profiles of the KEB-7 cell line derived from a DM-EBS patient with K14-R125P mutation revealed reduction in the levels of dual specificity MAP kinase-associated phosphatases along with higher and longer activation of the p38 and ERK pathways as compared to NEB-1 cells under hypoosmotic stress (Liovic et al., 2008). This suggests the presence of an intrinsic stress, thereby compromising the ability of EBS cells to deal with additional stress. It may in part explain the unexpected reduction of phosphorylated K5 in the triton-X100 soluble pool of osmotically stressed KEB-7 cells which is completely contrary to the increased phosphorylation

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of K5 in the soluble pool of NEB-1 cells (Liovic et al., 2008). Also, rheological assays demonstrated a reduced resilience of EBS mutant K14-R125C filaments to large deformations (Ma et al., 2001). 5.2.3 Increased Keratin Phosphorylation Is Observed in Hyperkeratotic Skin Diseases Links to keratin phosphorylation have also been found in EH, another autosomal dominant skin disorder which involves disruption of the structural integrity of the suprabasal layers of the epidermis. K1 L160P (157NQSLLQPL164 ! 157NQSPLQPL164) is a mutation in the H1 subdomain of K1 of patients with EH which leads to the generation of a new potential phosphorylation site in the adjacent serine (Chipev et al., 1992). In support, p38 MAPK mediated in vitro hyperphosphorylation of K8-L71P mutants, which also contain a novel phosphorylation site (K8S70) in the H1 subdomain corresponding to K1-S159. Besides, K8-L71Ptransfected cells showed increased keratin filament collapse in the presence of okadaic acid (Ku et al., 2002a). The significance of the K1-L160P mutation for keratin filament assembly is also evident from peptide inhibition experiments. While wild-type H1 peptide efficiently disassembled preformed K1/ K10 filaments, a mutant H1 peptide encompassing the K1-L160P mutation was much less efficient (Chipev et al., 1992).

5.3 Keratin Phosphorylation Is Altered in Cancer The biology of cancer is characterized by distinct hallmarks. These include increased cell migration needed for tissue invasion and metastasis, tumor progression that may be linked to dedifferentiation, and loss of growth control all of which have been linked in some way to altered keratin phosphorylation in epithelial carcinomas. We will describe some of these findings in the following paragraphs. They are also summarized in Fig. 7 highlighting the cell-type specificity of the observed phenomena. An explanation for the highly diverse and often opposing effects of keratin phosphorylation may be the complex nature of the phosphorylation patterns and the different overall cellular differentiation status in each instance. 5.3.1 Phosphorylation-Dependent Keratin Network Plasticity Correlates With Tumor Cell Migration Both, cell invasion through a connective tissue and its directional migration require the presence of a flexible leading edge. Increased keratin phosphorylation speeds up keratin cycling and increases network plasticity inducing

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cell shape changes that are advantageous for cell migration and invasiveness (Chung et al., 2013; Windoffer et al., 2011). This may also explain the unique mechanical properties of cancer cells such as softness and elasticity (Cross et al., 2007). Sphingosylphosphorylcholine (SPC), a bioactive lipid that is elevated in blood and ascites of ovarian cancer patients, affects cell growth and cell migration (Boguslawski et al., 2000; Seufferlein and Rozengurt, 1995; Xiao et al., 2000). SPC causes keratin network reorganization into perinuclear, ring-like structures accompanied by an increase in phosphorylation of K8-S431 and K18-S52 in pancreatic cancer-derived Panc-1 cells (Beil et al., 2003). In vivo micromechanical assays and Boyden chamber assays revealed an increase in the viscoelastic and migratory behavior of Panc-1 cells upon SPC treatment. This suggested that the phosphorylation-induced keratin reorganization may be directly linked to decreased mechanical resilience and increased invasiveness of SPC-treated Panc-1 cells and may be relevant for other tumor types (Beil et al., 2003).

Fig. 7 Anomalous K8/K18 phosphorylation in human cancer and its consequences. The scheme shows hyperphosphorylation (upper part) and hypophosphorylation (lower part) of K8/K18 in the context of various cancer conditions. AE-BS: acetone extract of Bupleurum scorzonerifolium; LTB: leukotriene B4; SPC: sphingosylphosphorylcholine; PKP3: plakophilin3; PRL-3: phosphatase of regenerating liver-3; TPA: 12-Otetradecanoylphorbol-13-acetate. Further details are in the text.

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Additional investigation showed that SPC triggers ERK and JNK activity, phosphorylating K8-S431 (Park et al., 2011). SPC-induced keratin reorganization in human pancreatic Panc-1 and gastric cancer AGS cells was accompanied by ERK-mediated keratin phosphorylation (Beil et al., 2003; Mesecke et al., 2011). Considering the importance of ERK signaling for cell migration (Bove et al., 2008; Huang et al., 2004; Rajalingam et al., 2005), the above findings provide another mechanism for enhanced cell motility acting via increased keratin phosphorylation. The exocyst complex component Sec8 has been identified recently as upstream regulator of ERKand p38-mediated phosphorylation of K8 in migrating oral squamous cell carcinoma (OSCC)-derived HSC3 cells (Tanaka and Iino, 2015). Furthermore, leukotriene B4 (LTB 4) is a component that is also elevated in pancreatic cancer. It induces keratin rearrangements in Panc-1 cells similar to those observed upon SPC treatment but acts via protein phosphatase 2A downregulation, which in turn induced ERK activation (Park et al., 2012). SPC-mediated keratin phosphorylation and reorganization depends on transglutaminase-2 (Tg-2) whose cross-linking activity leads to the formation of a triple complex of K8, Tg-2, and p-JNK (Park et al., 2011). Remarkably, transglutaminase-2 is associated with cell survival, invasiveness, metastasis, and chemoresistance in pancreatic, lung, and ovarian tumors (Chhabra et al., 2009; Park et al., 2010a; Verma et al., 2006). Moreover, 12-O-tetradecanoylphorbol-13-acetate (TPA), a potent tumor promoter, also stimulates K8-S431 phosphorylation, keratin reorganization, and Panc-1 cell migration via transglutaminase-2 (Lee et al., 2014). Conversely, ethacrynic acid inhibits transglutaminase-2 and hence obstructs SPCinduced keratin phosphorylation and perinuclear keratin reorganization (Byun et al., 2013). Taken together, multiple players may be involved in keratin network reorganization via altered keratin phosphorylation to adjust viscoelastic and migratory properties of cancer cells. 5.3.2 Keratin Dephosphorylation Is Linked to Tumor Progression In contrast to the above-mentioned observations, a gain of motility was observed in wound healing scratch assays of human OSCC-derived AW13516 cells, which produce either K8-S73A or K8-S431A phosphorylation-deficient keratins (Alam et al., 2011). Furthermore, subcutaneous injection of K8-S73A and K8-S431A OSCC cell clones into NODSCID mice demonstrated increased tumor formation as compared to the wild-type K8-producing OSCC cell clones (Alam et al., 2011). The relevance of these observations for human tumors is underscored by the

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observed absence of phosphorylation at K8-S73 and K8-S431 in 46% and 58% of human OSCC tissue samples, respectively (Alam et al., 2011). A clinicopathological analysis of strongly K8-positive OSCC patient samples revealed a close correlation between loss of K8-S73 and K8-S431 phosphorylation and tumor size, tumor stage, and lymph node metastasis. Moreover, loss of K8-S73 phosphorylation correlated negatively with patient survival (Alam et al., 2011). Thus, keratin phosphorylation may serve as a prognostic marker in certain cancer types such as OSCC. Keratin 8 is recognized as a physiological substrate of phosphatase of regenerating liver (PRL-3), which is consistently overexpressed in liver metastases derived from colorectal cancer (Saha et al., 2001). PRL-3 directly associates with K8 and dephosphorylates it at S73 and S431 residues. PRL-3 and K8 colocalization was observed in lamellipodia and ruffles of colorectal cancer-derived DLD-1 cells. Treatment of DLD-1 cells with PRL-3 inhibitor not only ablated the dephosphorylation at K8-S73 and K8-S431 but also ablated the colocalization of PRL-3 and K8 leading to redistribution of K8. In addition, the observed overexpression of PRL-3 and dephosphorylation of K8-S73 and K8-S431 at the invasive front of primary human colorectal carcinomas supports the role of PRL-3-mediated K8 dephosphorylation in invasion (Mizuuchi et al., 2009). Thus, PRL-3 activity, which has been shown to accelerate cancer cell motility, invasion, and metastasis, also modulates keratin phosphorylation, an important switch regulating keratin dynamics (Fiordalisi et al., 2006; Guo et al., 2004; Zeng et al., 2003). Upregulation of PRL-3 is an essential factor for transformation and metastasis caused by loss of plakophilin 3 (PKP3) in human colon carcinoma-derived HCT116 cells (Khapare et al., 2012). PKP3 is a desmosomal plaque protein whose loss has been implicated in neoplastic progression and metastasis (Aigner et al., 2007; Kundu et al., 2008; Papagerakis et al., 2003; Schwarz et al., 2006). Moreover, PRL-3-dependent K8-S431 dephosphorylation leading to K8 overexpression was observed in PKP3 knockdown cells. In accordance, loss of K8 in PKP3 knockdown cells led to the reversal of the transformed phenotype (Khapare et al., 2012). Upregulation of K8 is a common observation associated with increased cell invasiveness, migration, tumorigenesis, and poor prognosis of many tumors (Fillies et al., 2006; Ku et al., 1996; Raul et al., 2004; Schaafsma et al., 1993). Spheroid perinuclear inclusion bodies containing K8 aggregates are often observed in rhabdoid cells of malignant rhabdoid tumors (MRT) and other malignant neoplasms (Chetty and Asa, 2004; Shiratsuchi et al., 2001).

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The presence of rhabdoid cells is an indication of high aggressive malignancy and poor prognosis of epithelial and mesenchymal tumors (Beckwith and Palmer, 1978; Oda et al., 1993; Oshiro et al., 2000; Tsuneyoshi et al., 1987). Sequence analyses of the entire human K8 gene in samples obtained from frozen MRT tissues and MRT-derived cell lines revealed several missense mutations. Of these, codon 89 (CGC to TGC [R to C]) and codon 290 (AGC to ATC [S to I]) may be of relevance for keratin phosphorylation (Shiratsuchi et al., 2001). K8-R89 is located in the H1 subdomain of the K8 head which is a target of phosphorylation. K8-S290 is a reported phosphorylation site of the K8 tail domain (Ku and Omary, 1997). In addition, both of these mutations may have significance for lateral interactions during keratin assembly. Thus, mutation-induced alterations in keratin phosphorylation may contribute to the aberrant accumulation of keratin in MRT (Chetty and Asa, 2004; Shiratsuchi et al., 2001). The above reports suggested the association of K8 dephosphorylation with tumor progression and aggressiveness. A similar link was suggested from investigations of the proapoptotic effects of the crude acetone extract of Bupleurum scorzonerifolium (AE-BS) on human lung cancer-derived A549 cells (Chen et al., 2005). AE-BS, known for its anticancer properties, led to ERK1/2 activation causing K8-S73 hyperphosphorylation in A549 cells. This observation indicated that K8-S73 phosphorylation, which was also induced by stimulating the proapoptotic receptors Fas and TNF, was linked to diminished cancer cell viability (Gilbert et al., 2001; Ku et al., 2003). Similarly, overexpression of the tumor–suppressor parkin in human cervical cancer-derived HeLa cells was associated with increased phosphorylation of K8/K18 (Song et al., 2013). Loss or mutations of parkin have been linked to acute lymphoblastic leukemia, chronic myeloid leukemia, and colorectal carcinoma (Agirre et al., 2006; Cesari et al., 2003; Poulogiannis et al., 2010). In accordance with these studies, overexpressing parkin in HeLa cells caused growth-inhibitory effects, which were accompanied by phosphorylation of K8 and K18 (Song et al., 2013). However, there are contrary reports such as the occurrence of K8-Y267 phosphorylation in human cholangiocarcinoma tissues and the presence of phosphotyrosine peptides in lung cancer associated with high expression of the protooncogene tyrosine kinase ROS (Gu et al., 2011; Rikova et al., 2007; Snider et al., 2013). Furthermore, RSKmediated phosphorylation of K17-S44 has been implicated in the interaction of K17 with hnRNPK, a partnership leading to CXCR3-dependent tumor growth and invasion as demonstrated in vulva carcinoma-derived A431 cells (Chung et al., 2015; Lo et al., 2010).

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5.3.3 Keratin Phosphorylation Is Linked to Epithelial–Mesenchymal Transition Carcinogenesis is often characterized by epithelial to mesenchymal transition (EMT), a critical mechanism leading to acquisition of invasiveness and malignancy in epithelial cancer cells (Brabletz et al., 2001; Fidler and Poste, 2008; Thiery, 2002). The EMT program is a biological process that converts the phenotype of an epithelial cell to that of a mesenchymal cell (Kalluri and Neilson, 2003). The oncogenic kinase Src, known to phosphorylate K19-Y391, is elevated in highly metastatic cancer cells resulting from loss of epithelial differentiation (Rikova et al., 2007; Zhou et al., 2010). Srcmediated phosphorylation of K19 at Y391 residue leads to its increased partitioning into the soluble fraction upon pervanadate treatment (Zhou et al., 2010). Additionally, the Src-K19 interaction is accompanied by facilitation of EGF-stimulated oncogenesis, which is critical for Src activation (Li et al., 2010). 5.3.4 Keratin Hyperphosphorylation Is a Consequence of Altered Growth Control in Cancer Cells Activated growth signaling pathways in cancer cells have been shown to induce keratin hyperphosphorylation. Thus, increased phosphorylation of K8-S23, K8-S73, and K8-S431 was reported for colorectal cancer-derived CaCo-2 cells upon activation of the EGFR pathway (Arentz et al., 2011). Conversely, blockade of the EGFR pathway not only decreased keratin phosphorylation but also elevated apoptosis in CaCo-2 cells (Arentz et al., 2011). Similar mechanisms may be the reason for the high survival benefits of sorafenib in advanced hepatocellular carcinoma (HCC), which is a malignancy resistant to conventional therapies (Bruix et al., 2005; Cheng et al., 2009; Forner et al., 2012; Llovet et al., 2008; Schwartz et al., 2007). Sorafenib is an inhibitor of serine/threonine and receptor tyrosine kinases. The molecular action of sorafenib involves the generation of endoplasmic reticulum stress by decreasing protein ubiquitination and the unfolded protein response (Honma and Harada, 2013). Additionally, being a multikinase inhibitor, sorafenib blocks keratin phosphorylation at major phosphorylation sites such as K8-S73, K18-S33, and K18-S52 in hepatoma and human hepatocyte-derived OUMS29 cells (Honma and Harada, 2013; Wilhelm et al., 2004). In this way, sorafenib may prevent a defensive response against hepatocellular stress (Harada et al., 2007; Ku et al., 1998b; Ku and Omary, 2006; Kwan et al., 2012; Omary et al., 2004; Toivola et al., 2004; Zatloukal et al., 2007). The effects of sorafenib manifest

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as an increase in autophagy and a decrease in proteasome-mediated MDB formation in HCC-derived Huh7 and OUMS29 cells (Bareford et al., 2011; Honma and Harada, 2013; Park et al., 2010b; Shimizu et al., 2012; Ullen et al., 2010). Both, induction of autophagy and loss of MDB formation are characterized by a reduction in phosphorylated keratin (Harada et al., 2007, 2008; Kongara et al., 2010; Kwan et al., 2012; Zatloukal et al., 2007). The cytotoxic action of sorafenib results in an antiproliferative effect in hepatoma cells, reduced tumor angiogenesis, and increased apoptosis (Honma and Harada, 2013; Liu et al., 2006; Wilhelm et al., 2004). Hence, administration of sorafenib reduces tumor cell viability through its pleiotropic antitumor effects, which exploit the role of phosphorylated keratins as stress proteins.

6. CONCLUDING REMARKS: SIGNIFICANCE OF KERATIN PHOSPHORYLATION Based upon the various aspects discussed above, overall significance of keratin phosphorylation can be summarized as follows: (1) Modulation of cellular plasticity. Phosphorylation affects keratin solubility and keratin filament turnover with consequences on network organization and mechanics during the cell cycle, in response to various types of stress and during carcinogenesis. (2) Regulation of keratin association with other proteins. Phosphorylation regulates binding of keratins to each other and other proteins affecting keratin assembly state, crosslinking to other cell components and signal transduction. (3) Role as a “phosphate sponge.” Keratin phosphorylation sequesters kinase and phosphatase activities. (4) Protection of keratins from degradation. Phosphorylation prevents keratin ubiquitination with consequences for cell viability. Similar to the posttranslational modifications in other intermediate filaments, phosphorylation endows the keratins with a wide gamut of functional properties. Due to its rapid kinetics, phosphorylation of the expansive keratin network serves as a suitable emergency system, mobilizing the wellconnected keratin network to trigger appropriate switches for adaptation to the changing environment.

ACKNOWLEDGMENTS The work was supported by the German Research Council (LE566/18-1 and LE566/22-1) and the IZKF and Boost Fund of RWTH Aachen University. We thank Drs. Nicole Schwarz and Reinhard Windoffer for helpful discussions throughout. We are also grateful to Dr. Stefan Wo¨ll for providing the fluorescence micrographs in Figs. 2D–F and 4 and thank Adam Breitscheidel for graphic design of Figs. 1 and 7.

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CHAPTER SIX

Plastid Protein Targeting: Preprotein Recognition and Translocation P. Chotewutmontri*,1,2, K. Holbrook†,1,3, B.D. Bruce*,†,4 *Graduate School of Genome Science and Technology, University of Tennessee, Knoxville, TN, United States † University of Tennessee, Knoxville, TN, United States 4 Corresponding author: e-mail address: [email protected]

Contents 1. Introduction 2. Origin of Plastids 3. Plastid Protein Targeting Routes 3.1 General Import Pathway 3.2 Roles of General Import Pathway Components 3.3 Noncanonical Trafficking 3.4 Dual Targeting to Plastids and Mitochondria 4. Regulation of Plastid Protein Import 4.1 Organism-Specific Recognition of TPs 4.2 Expression Control 4.3 Precursor-Specific Import Pathways 4.4 Redox Regulation 4.5 Phosphorylation Regulation 4.6 Potential Role of Proline Isomerization 4.7 Regulation by Ubiquitin–Proteasome System 5. Concluding Remarks Acknowledgments References

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Abstract Eukaryotic organisms are defined by their endomembrane system and various organelles. The membranes that define these organelles require complex protein sorting and molecular machines that selectively mediate the import of proteins from the cytosol to 1 2

3

Contributed equally to the manuscript. Current address: Institute of Molecular Biology, University of Oregon, Eugene, OR 97403, United States. Current address: Department of Chemistry and Biochemistry, University of California at Los Angeles, Los Angeles, CA 90095, United States.

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their functional location inside the organelle. The plastid possibly represents the most complex system of protein sorting, requiring many different translocons located in the three membranes found in this organelle. Despite having a small genome of its own, the vast majority of plastid-localized proteins is nuclear encoded and must be posttranslationally imported from the cytosol. These proteins are encoded as a larger molecular weight precursor that contains a special “zip code,” a targeting sequence specific to the intended final destination of a given protein. The “zip code” is located at the precursor N-terminus, appropriately called a transit peptide (TP). We aim to provide an overview of plastid trafficking with a focus on the mechanism and regulation of the general import pathway, which serves as a central import hub for thousands of proteins that function in the plastid. We extend comparative analysis of plant proteomes to develop a better understanding of the evolution of TPs and differential TP recognition. We also review alternate import pathways, including vesicle-mediated trafficking, dual targeting, and import of signal-anchored and tail-anchored proteins.

1. INTRODUCTION With rapid advances in genome sequencing, transcriptional profiling, and proteomics, a comprehensive snapshot of how cells differentially express their genome is now possible. In plants, systems biology approaches have led to the generation of a variety of large datasets (Gaudinier and Brady, 2016). These approaches have revealed the complexity of gene expression and moving the field forward will require an understanding of the subcellular localization of the expressed proteins. In plants, these protein dynamics involve a delicate interplay between not only the nuclear genome and the genomes of the semiautonomous organelles, the plastids, and the mitochondria. Understanding these new challenges comes nearly 40 years after it was first established that some proteins are targeted to the plastid via a posttranslation mechanism that required a specific N-terminal targeting sequence (Chua and Schmidt, 1978; Highfield and Ellis, 1978). Despite the functional similarities, the translocation machineries found in the endoplasmic reticulum (ER), mitochondria, and plastids show no similarity in their core components (Schatz and Dobberstein, 1996). Thus, these import processes appear to have evolved independently. Surprisingly, in plants cells, these processes can coexist with little to no missorting. In this review, we provide a detailed update on our understanding of the structure, function, and regulation of plastid import pathways, with a focus on the chloroplast translocons functioning in the two envelope membranes, outer envelope of chloroplasts (TOC) and inner envelope of chloroplasts (TIC). We also

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look the evolutionary diversity of these molecular machines across a wide range of phototrophic organisms. We take detailed new look the size and diversity of the transit peptides (TPs) (Bruce, 2000). Finally, we start to build a detailed understanding of how TPs can function without sequence homology yet perform a common universal function (Bruce, 2001).

2. ORIGIN OF PLASTIDS Over century ago, Mereschkowsky proposed that plastids derived from cyanobacteria (Martin and Kowallik, 1999). It is now widely accepted based on phylogenetic analysis that the primary plastids originated from endosymbiosis of a cyanobacterial ancestor (McFadden, 2014). Interpretation of the fossil record and molecular clock analysis date the endosymbiosis event to over 1.2 billion year ago (Butterfield, 2000; Yoon et al., 2004). The comparisons of genome organization and sequences of the primary plastids from Archaeplastida which includes Viridiplantae (green algae and plants), rhodophytes, and glaucophytes suggest a single endosymbiosis event (Keeling, 2010; Price et al., 2012; Turner et al., 1999). Aside from the advantage of photosynthetic capability, the only other endosymbiosis event resulting in primary plastids was reported in a protist Paulinella (Reyes-Prieto et al., 2010). To explain the incredibly rare occurrence of primary plastid endosymbiosis, Ball et al. (2013) showed that the key enzymes required for the host to utilize photosynthetic carbon were derived from another bacterium, the pathogenic Chlamydiales. They further proposed that the endosymbiosis occurs only in the infected host where the carbon utilization becomes beneficial. Many attempts to pinpoint the plastid ancestor suggest unicellular nitrogen-fixing cyanobacteria based on 16S RNA, photosynthetic, and metabolic gene sequences (Criscuolo and Gribaldo, 2011; Falcon et al., 2010; Hackenberg et al., 2011; Kern et al., 2011; Pascual et al., 2011). The most comprehensive study using a large-scale comparison of 241 complete genomes including 9 cyanobacteria and 4 photosynthetic eukaryotes indicates that the nitrogen-fixing heterocyst cyanobacteria share the most genes with the photosynthetic eukaryotes (Deusch et al., 2008). Although modern heterocyst cyanobacteria, Nostoc sp. PCC7120 and Anabaena variabilis ATCC29143 harbor around 5500 genes that encode proteins (Kaneko et al., 2001; Markowitz et al., 2012), plastid genomes of land plants only encode around 80 proteins (Timmis et al., 2004). This reduction of plastid genomes was found to occur mainly by the transferring of plastid DNA to the nuclear genome (Kleine et al., 2009). It was proposed

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Plant cell Nucleus Other locations 2700 Cyanobacterial ancestor

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Fig. 1 Endosymbiont gene transfer and protein targeting to the plastid. Bioinformatic analysis of Arabidopsis genome sequence (Martin et al., 2002) indicated that around 4500 nuclear genes are transferred from the cyanobacterial ancestor and only 87 genes remain in the plastid genome. Although the majority of the proteins encoded by the endosymbiont-to-nucleus genes are targeted to other locations in the cell, about 1800 of these proteins are targeted to the plastid. In addition, around 750 proteins encoded by nonendosymbiont nuclear genes are also targeted to the plastid.

that the ability of plastids to import nuclear-encoded proteins enabled the plastid ancestry to transfer its genes to the nucleus without compromising its metabolic capacity (Allen, 2003). During the process of endosymbiosis, the import of proteins encoded by endosymbiont-to-nucleus genes permits the endosymbiont gene copies to undergo pseudogenization and later loss (Martin et al., 1993). In Arabidopsis, bioinformatic analysis estimated that around 4500 protein-encoding genes in the nucleus are originated from the endosymbiont as shown in Fig. 1 (Martin et al., 2002). Less than half of these endosymbiont-to-nucleus proteins (about 1800) are targeted to plastids and the rest functions elsewhere in the cell (Martin et al., 2002). Interestingly, nonendosymbiont-derived proteins (about 750) also localize to plastids (Martin et al., 2002) and function in photosynthesis, respiration, and metabolic pathways (Kleine et al., 2009). While plastid protein import was crucial in facilitating the endosymbiosis event in the past, it becomes indispensable to the function of the cells in Arabidopsis where over 2500 proteins in Arabidopsis must gain access to plastids.

3. PLASTID PROTEIN TARGETING ROUTES Proteins targeted to plastids can be delivered posttranslationally to six plastid locations: the outer envelope membrane (OM), the intermembrane

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Fig. 2 Plastid protein targeting routes. Proteins targeted to plastids are delivered to six locations: outer envelope, intermembrane space, inner envelope, stroma, thylakoid membrane, and thylakoid lumen. The outer envelope proteins use multiple pathways for targeting while the interior proteins containing TP pass through the envelope(s) via the General Import Pathway. Some of the proteins contain a second targeting signal for thylakoid lumen targeting. In addition, experimental evidence and proteomic analysis reveal that many proteins utilize noncanonical pathway.

space (IMS), the inner envelope membrane (IM), the stroma, the thylakoid membrane, and the thylakoid lumen as shown in Fig. 2 (Keegstra and Cline, 1999; Li and Chiu, 2010). Most of these processes involve protein translocation across the plastid membranes: the outer envelope, the inner envelope, and the thylakoid membranes. In general, protein translocation across or insertion into membrane is mediated by oligomeric membrane complexes termed translocons (Walter and Lingappa, 1986). The majority

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of plastid-targeted proteins is synthesized as precursors containing the N-terminal targeting sequences called TPs (Dobberstein et al., 1977; Kleffmann et al., 2004). Similar to the signal hypothesis (Blobel, 1980), TPs act as an intrinsic signal on the precursor protein that is recognized by the targeting receptors associated with the translocons. TPs direct translocation of precursor proteins across the double membranes of plastids via the translocon at the TOC and TIC in a process described as the general import pathway (Cline and Henry, 1996; Schnell et al., 1997; van’t Hof and de Kruijff, 1995a). After the precursor is translocated into the stroma, the TP is readily cleaved allowing the mature domain to fold into its native conformation or to be further targeted to the thylakoid (Richter and Lamppa, 2002). So far, the TP has been utilized in targeting precursor proteins to all of the six locations of plastids and is considered to function in the general import pathway. This pathway recognizes diverse TP sequences from various functional groups of proteins. In summary, the general import pathway functions similar to a central transit hub where the majority of plastid-targeted proteins pass through before reaching their final locations.

3.1 General Import Pathway The general import pathway is a working model describing a TP-directed translocation of proteins into the stroma of plastids (Bruce, 2000). Fig. 3 illustrates the sequential steps in the pathway that reflect the processes that take place in the cytosol, in both the OM and IM, stromal-localized components, and finally the cleavage and processing of the TP. We have not included steps that are involved in the subsequent targeting to the thylakoid nor do we address the targeting to the OM, IM, or IMS. Fig. 3 represents a somewhat selective model that has consolidated many years of work, but emerging research continues to develop a more complete picture of the import system. The OM of plastids has been shown to recruit cytosolic precursors using multiple pathways. Work performed using isolated chloroplasts shows that the precursors are able to directly bind to TOC on the surface of chloroplasts (Fig. 3, step 1a). The binding is reversible when lacking ATP/GTP and denoted as energy-independent binding (Jarvis, 2008; Kouranov and Schnell, 1997; Perry and Keegstra, 1994). Addition of GTP also promotes binding (Inoue and Akita, 2008; Young et al., 1999). TP interaction with chloroplast lipids suggests that the precursor can directly bind to the lipid surface (Fig. 3, step 1e) before being transferred to the membrane receptors TOC159 and

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Fig. 3 The general import pathway. Cytosolic precursors can be targeted to plastids directly by interacting with TOC (1a), or by interacting with the lipids (1e) before transferring to membrane Toc159 (2f ) and reaching Toc34 (3a). The precursors can also interact with cytosolic Toc159 (1b) before transferring to Toc34 (2a). TPs can interact with Hsp90 (1d) before being targeted to Toc64 (2d) and further transferred to Toc34 (3b). Phosphorylated TPs can interact with the guidance complex (1c) composed of 14-3-3 and Hsp70c in cytosol. The precursors from guidance complexes can be transferred to membrane receptors Toc159 (2c) or Toc34 (2b). The binding state of precursor to the chloroplast can be subdivided based on GTP/ATP level and temperature of the system (4–5). When the ATP level is greater than 1 mM, the translocation process is initiated by Hsp93/cpHsp70 (6). When the precursors emerge into the stroma, stromal processing peptidase (SPP) will cleave TP from precursor proteins releasing the mature domain (7). TP will be further degraded by PreP1/PreP2 peptidases (8).

TOC34 (Fig. 3, steps 2f and 3a) (Pilon et al., 1995; Pinnaduwage and Bruce, 1996). Cytosolic factors were shown to interact and recruit precursors to the membrane receptors. Cytosolic Hsp90 captures precursor (Fig. 3, step 1d) and delivers it to the membrane receptor TOC64 (Fig. 3, step 2d) (Qbadou et al., 2006). Phosphorylated TPs interact with 14-3-3 and form the guidance complex with cytosolic Hsp70 Fig. 3 (step 1c) (May and

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Soll, 2000). The guidance complex was proposed to dock to membrane receptors TOC159 (Fig. 3, steps 2c and 3a) or TOC34 (Fig. 3, step 2b) (May and Soll, 2000; Qbadou et al., 2006). Another pathway utilizes the cytosolic TOC159 pool to deliver precursor to the membrane (Fig. 3, steps 1b and 2a) (Hiltbrunner et al., 2001; Smith et al., 2004). When low levels of ATP ( L > P > R/K in the logo plot, occurring nearly equally on both sides of the trans Pro. It is difficult to discern the causality of these differing amino acid distributions. Do the flanking sequences simply influence the Pro isomeric form; or does recognition of the cis Pro isomeric form require some additional residues for its maximum specificity or recognition by the translocon(s)? Although how processing is coupled to translocation is not known, it is not surprising that this may be an intimate interaction that requires subtle spatial control between the emerging preprotein and SPP in the stroma. Investigating the potential role of cis Pro racheting “kinks” in the TP sequence will be a topic of future study.

4.7 Regulation by Ubiquitin–Proteasome System The level of precursor proteins in cytosol has been shown to be regulated by the ubiquitin–proteasome system, specifically through the cytosolic Hsc70 and the C-terminus of Hsc70-interacting protein (CHIP) E3 ubiquitin ligase pathway (Lee et al., 2009c; Shen et al., 2007). Interestingly, other evidence also indicated that a putative C3HC4-type really interesting new gene (RING) E3 ubiquitin ligase SP1 interacts with all of the TOC components and initiates their degradation via proteasome (Ling et al., 2012). It was proposed that E3 ubiquitin ligase SP1 regulates the turnover of TOC components through ubiquination and degradation pathways. In combination with the differential expression of TOC components, this results in modulation of the composition of TOC components (Ling et al., 2012). SP1-mediated TOC composition change was also suggested to control the transition between plastid types (Ling et al., 2012). Clearly, regulation of both the complex components and precursor proteins work together to modulate import to the plastid. This is a highly complex system and additional analysis of regulation pathways are still required to fully understand how these pathways interplay.

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5. CONCLUDING REMARKS Significant research has focused on the mechanisms that control import of nuclear-encoded precursor proteins. The general import pathway represents a central hub for recognition, regulation, and membrane translocation of precursors; this system plays a key role in organelle function and homeostasis. However, it is clear that noncanonical pathways work in concert with the TOC/TIC system to respond to developmental and physiological responses in the cell. Although we have developed a strong basis for understanding plastid import, more detailed analysis of the components mediating cytosolic recognition, TOC assembly, and regulation of the import process are required. Furthermore, understanding how diverse TPs mediate the selective targeting and translocation of precursors will reveal important regulation and quality control elements of this pathway. In depth understanding of TP domain architecture may allow for the design of novel synthetic TPs in the future. In summary, emerging studies that integrate the assembly and regulation of the plastid import pathways will contribute to uncovering the mechanisms that mediate the plasticity of organelle development.

ACKNOWLEDGMENTS We would like to thank Mr. Will Crenshaw and Mr. Jordan Taylor for sharing their unpublished data. We would like to thank Nate Brady for critically reading the manuscript. Support has been provided from the Gibson Family Foundation, TN-SCORE, the Tennessee Plant Research Center, and National Science Foundation to B.D.B. (MCB-0628670, MCB-0344601, and EPS-1004083). JT was supported as an NSF-supported REU by EPS-1004083. This work was supported in part by a UTK Science Alliance grant to K.H. and P.C. and a UTK SARIF grant to K.H.

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CHAPTER SEVEN

Immunomodulatory Activity of VEGF in Cancer A. Lapeyre-Prost*, M. Terme*,1, S. Pernot*,†, A.-L. Pointet*,†, T. Voron*,{, E. Tartour*,§, J. Taieb*,†,1 *INSERM U970, PARCC (Paris Cardiovascular Research Center), Universit
e Paris-Descartes, Paris, France † Service d’h
epatogastroent
erologie et d’oncologie digestive, H^ opital Europ
een Georges Pompidou, Paris, France { Service de chirurgie digestive, H^ opital Europ
een Georges Pompidou, Paris, France § Service d’immunologie biologique. H^ opital Europ
een Georges Pompidou, Paris, France 1 Corresponding authors: e-mail address: [email protected]; [email protected]

Contents 1. Introduction 2. The VEGF Family 2.1 Properties of VEGF Family Members 2.2 Isoforms of VEGF-A and Their Particularities 2.3 Production of VEGF-A 3. VEGF Receptors 4. The VEGF-A/VEGFR Axis 5. Antitumor Immunity 5.1 Tumor Immunosurveillance 5.2 Escape From the Antitumor Immune Response 6. Immunosuppressive Roles of VEGF 6.1 Inhibition of DC Maturation 6.2 Accumulation of MDSCs 6.3 Development of Tumor-Induced Macrophages 6.4 Increase in Tregs 6.5 Effector T Cells 7. Immunomodulatory Properties of Therapeutics Targeting VEGF 7.1 Development of Major Antiangiogenic Molecules 7.2 Immunomodulatory Activity of Antiangiogenic Therapies in Animals 7.3 Immunomodulatory Activity of Antiangiogenic Therapies in Humans 8. Medical Applications 8.1 Antiangiogenic Drugs as Ideal Companions for Anticancer Immunotherapy: Preclinical Studies 8.2 First Clinical Results 8.3 Other Potential Applications 9. Concluding Remarks Acknowledgments References

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Abstract The ability of tumor cells to escape tumor immunosurveillance contributes to cancer development. Factors produced in the tumor microenvironment create “tolerizing” conditions and thereby help the tumor to evade antitumoral immune responses. VEGF-A, already known for its major role in tumor vessel growth (neoangiogenesis), was recently identified as a key factor in tumor-induced immunosuppression. In particular, VEGF-A fosters the proliferation of immunosuppressive cells, limits T-cell recruitment into tumors, and promotes T-cell exhaustion. Antiangiogenic therapies have shown significant efficacy in patients with a variety of solid tumors, preventing tumor progression by limiting tumor-induced angiogenesis. VEGF-targeting therapies have also been shown to modulate the tumor-induced immunosuppressive microenvironment, enhancing Th1-type T-cell responses and increasing tumor infiltration by T cells. The immunomodulatory properties of VEGF-targeting therapies open up new perspectives for cancer treatment, especially through strategies combining antiangiogenic drugs with immunotherapy. Preclinical models and early clinical studies of these combined approaches have given promising results.

1. INTRODUCTION The vascular endothelial growth factor (VEGF) family is involved in blood vessel development and homeostasis, and also in lymphatic vessel formation. The main member of this family, VEGF-A, is a major driver of angiogenesis and vasculogenesis. Angiogenesis is a physiological or pathological process defined as the sprouting and remodeling of small new capillaries from the preexisting blood vasculature, whereas vasculogenesis involves the migration, differentiation, and association of endothelial precursor cells to form primitive blood vessels. In physiological conditions, VEGF-A promotes angiogenesis during embryonic development and is necessary for tissue repair. In cancer patients, VEGF-A production by the tumor results in an “angiogenic switch” required for tumor growth and metastasis (Bergers and Benjamin, 2003). Indeed, as the tumor mass increases, oxygen availability is reduced (hypoxia), and the tumor thus produces proangiogenic factors such as VEGF-A, a process mediated by the transcription factor hypoxia-inducible factor (HIF)-1. The new vasculature formed under the influence of VEGF family members enables the tumor to meet its nutrient requirements. However, tumor vessels are both structurally and functionally abnormal: they are immature, irregularly shaped, dilated, and tortuous (Bergers and Benjamin, 2003).

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They also have lower pericyte coverage (Benjamin and Keshet, 1999) and an unstructured organization lacking well-defined venules, arterioles, and capillaries. The tumor vascular network is also prone to hemorrhage and necrosis, partly owing to an increase in interstitial fluid pressure related to overproduction of VEGF-A (or “vascular permeability factor”). Tumor vessels also differ from their normal counterparts by their dynamic properties. Unlike normal blood vessels, they never become quiescent, thus allowing ongoing tumor development and production of new vessels. The importance of neovascularization in tumor growth depends on the tumor type. About 10 years ago, the first evidence that tumor-derived VEGF-A might also affect cells other than endothelial and tumor cells, and particularly some immune cells, was published (Marigo and Bronte, 2008; Zou, 2005). It now appears that proangiogenic growth factors such as VEGF-A may be involved in tumor escape from antitumor immunity. Here, we review the many overlapping mechanisms involving VEGF family members, and especially the role of VEGF-A in tumor escape from immunosurveillance. These effects include inhibition of dendritic cell (DC) maturation, accumulation of myeloid-derived suppressor cells (MDSCs), an imbalance between effector T cells and regulatory T cells (Tregs), development of tumor-associated macrophages (TAMs), and promotion of T-cell exhaustion. We then analyze how VEGF/VEGFR pathway blockade modulates antitumor immune responses, and finally how recent discoveries may lead to the development of new therapeutic strategies combining antiangiogenic agents with immunotherapeutic approaches.

2. THE VEGF FAMILY 2.1 Properties of VEGF Family Members Six proteins of the VEGF family have been described: VEGF-A, VEGF-B, VEGF-C,-VEGF-D, VEGF-E, and PlGF (placental growth factor). VEGFs are mainly antiparallel homodimeric polypeptides, although heterodimers of VEGF-A and PlGF have also been described (Di Salvo and Thomas, 1995). VEGF is a potent and specific mitogen for endothelial cells, coming from arteries, veins, and lymphatic vessels. VEGF-A is one of the most extensively studied members of the VEGF family. It has a molecular weight of approximately 45 kDa (Ferrara and

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Davis-Smyth, 1997) and is a key regulator of angiogenesis. VEGF-A induces angiogenesis both in vitro (Pepper and Montesano, 1992) and in vivo (Nicosia and Qian 1997). VEGF-A induces the expression of plasminogen activator (PA), PA inhibitor 1 (PAI-1) (Pepper and Montesano, 1991), and interstitial collagenase in endothelial cells (Unemori and Amento, 1992). PA and collagenase production favor the creation of a “prodegradative” environment that allows endothelial cells to migrate and “sprout” into the extracellular matrix. Conversely, PAI-1 is a negative regulator of proteolysis. Normal vessel morphogenesis, including endothelial cell invasion and capillary lumen formation, requires a strictly controlled balance between proteases and protease inhibitors (Pepper and Montesano, 1990). VEGF-A is also called vascular permeability factor for its ability to increase microvascular permeability, a necessary step in the angiogenic process both in tumors and at wound sites (Dvorak and Dvorak, 1995). VEGF-A can also induce the formation of fenestrations in endothelial cells (Roberts and Palade, 1995). VEGF-A can bind directly to tumor cells in an autocrine manner and can promote tumor growth and motility independently of angiogenesis (Cao and Mukhopadhyay, 2012; Hamerlik and Bartek, 2012; Samuel and Ellis, 2011). Serum VEGF-A levels are elevated in cancer patients (Kondo and Suzuki, 1994), and VEGF-A can be produced by tumor cells of various histologies (Berger and Fiebig, 1995).

2.2 Isoforms of VEGF-A and Their Particularities The human VEGF-A gene is composed of eight exons separated by seven introns. Nine isoforms of VEGF-A have been described (VEGF121, VEGF145, VEGF148, VEGF162, VEGF165, VEGF165b, VEGF183, VEGF189, and VEGF206), resulting from alternative RNA splicing (Lange and Neufeld, 2003). The different isoforms share the first 115 N-terminal amino acids and, in most cases, the six C-terminal residues. VEGF165 is the predominant isoform in humans and is commonly known as VEGF-A. VEGF-A isoforms differ in their angiogenic properties: for example, VEGF165b is an endogenous inhibitory form of VEGF-A (Woolard and Bates, 2004). VEGF-A isoforms also vary in their diffusion capacity and their ability to bind heparan sulfate in the extracellular matrix and neuropilin coreceptors on the cell surface (Suarez and BallmerHofer, 2006).

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2.3 Production of VEGF-A VEGFs, particularly VEGF-A, are produced by various cell types, including macrophages, endothelial cells, tumor cells, and tumor-associated stroma (Ferrara and Davis-Smyth, 1997; Freeman and Klagsbrun, 1995; Melter and Briscoe, 2000). VEGF-A mostly remains bound to the cell surface and extracellular matrix. It can be released by heparin or heparinase, or by proteolytic cleavage of the longer isoforms by plasmin (Houck and Ferrara, 1992) or urokinase at the COOH terminus (Ploue¨t and Bayard, 1997), generating a bioactive fragment (Park and Ferrara, 1993). VEGF-A can also be secreted. Production and release of VEGFs, notably VEGF-A, are regulated by a variety of stimuli such as hypoxia, and by oncoproteins such as epidermal growth factor and K-ras (Perrotte and Dinney, 1999; Rak and Kerbel, 2000). The transcription factor HIF-1 plays a key role in hypoxia-driven VEGF-A production. HIF-1-alpha and HIF-1-beta are both constitutively produced, but HIF-1-alpha is degraded in aerobic environments. Therefore, when tumor volume exceeds the critical size of several cubic millimeters, resulting in hypoxia, HIF-1-alpha is not degraded but forms a complex with HIF-1-beta, thereby inducing the transcription of several genes, including VEGF (Takahashi and Shibuya, 2005). VEGF-A is also regulated at the mRNA level. Levy et al. found a crucial role of the 3-UTR RNA-binding protein HuR, a member of the Elav-like protein family, in stabilizing VEGF-A mRNA in hypoxic conditions (Levy and Levy, 1998). Another VEGF-A mRNA 3-UTR-interacting protein called poly(A)-binding protein-interacting protein 2 (PAIP2) has been identified as a coregulator of VEGF-A mRNA stabilization, cooperating with HuR to control VEGF-A at the posttranscriptional level in hypoxic situations (Onesto and Page`s, 2004).

3. VEGF RECEPTORS VEGFs can act in paracrine manner on adjacent endothelial cells and also in autocrine manner to ensure endothelial cell homeostasis (Lee and Iruela-Arispe, 2007). The biological functions of VEGFs are mediated by high-affinity tyrosine kinase receptors. Structurally, VEGF receptors (VEGFRs) have seven extracellular immunoglobulin-like domains (IgG-like), a single

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transmembrane region, and an intracellular portion that contains the tyrosine kinase domain. VEGFs bind and activate three types of cell surface receptor: VEGFR-1/ Flt-1, VEGFR-2/Flk1/KDR, and VEGFR-3 (Fig. 1). The VEGF165 isoform also binds neuropilin NP1, which acts as a specific coreceptor for VEGFR-1 and VEGFR-2 signal transduction (Fuh and de Vos, 2000; Soker and Klagsbrun, 1998; Whitaker and Rosenbaum, 2001). VEGFRs are essential for embryonic vasculogenesis, physiological and pathological angiogenesis, and lymphangiogenesis. VEGFR-1 is a 180-kDa receptor for VEGF-A, VEGF-B, and PlGF. It binds VEGF-A with higher affinity than does VEGFR-2 (Ferrara and DavisSmyth, 1997) but has lower kinase activity. VEGFR-1 is expressed by various cell types, including vascular endothelial cells, macrophages, monocytes (Sawano and Shibuya, 2001), and hematopoietic stem cells (Hattori and Rafii, 2002). VEGFR-1 seems to play a dual role, negatively controlling

Fig. 1 Impact of VEGF-targeting therapies on the VEGF/VEGFR axis. Sunitinib, sorafenib, and regorafenib target VEGF receptor tyrosine kinases. Aflibercept targets VEGF-A, VEGF-B, and PlGF. Bevacizumab targets VEGF-A, and ramucirumab targets VEGFR-2.

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vessel formation during early development, and positively regulating pathological angiogenesis in adulthood. Homozygous deletion of VEGFR-1 in mice results in embryonic lethality due to excessive growth of endothelial cells that disorganize the vascular network (Fong and Breitman, 1995). VEGFR-1 is also involved in endothelial cell migration (Kanno and Sato, 2000). VEGFR-2 is a 200–230-kDa receptor for VEGF-A, VEGF-C, VEGF-D, and VEGF-E. VEGFR-2 is expressed by vascular and lymphatic endothelial cells, and by other cell types such as megakaryocytes and hematopoietic stem cells (Katoh and Satow, 1995). It largely mediates the proangiogenic effects of VEGF-A but is also involved in lymphangiogenesis (Nagy and Dvorak, 2002). VEGFR-2 is the key receptor mediating both physiological and pathological angiogenesis, particularly in the angiogenic switch and tumor neoangiogenesis, involving small vessels. VEGFR-2 gene knock-out mice lack blood island and differentiated endothelial cells, and die between day 8.5 and 9.5 of embryonic life, suggesting that VEGFR-2 signaling is required for the differentiation and proliferation of endothelial precursor cells (Shalaby and Schuh, 1995). VEGFR-3 is a 195-kDa receptor for VEGF-C and VEGF-D and has a key role in lymphangiogenesis (Veikkola and Alitalo, 2001). The signal mediated by VEGFR-3 also seems to play a role in adaptative immunity, by modulating the recruitment of antigen-presenting cells (Chen and Dana, 2004).

4. THE VEGF-A/VEGFR AXIS The role of VEGF-A and its receptors in physiological angiogenesis is well known (Ferrara, 2001). VEGF-A is involved in embryonic and early postnatal development. The role of VEGF-A in vasculogenesis and angiogenesis during embryogenesis was demonstrated in two major studies (Carmeliet and Nagy, 1996; Ferrara and Moore, 1996), showing that mouse embryos in which a single VEGF allele was inactivated had a number of developmental anomalies, including cardiovascular malformations (rudimentary dorsal aorta, thickness of ventricular wall, defective vasculature in other tissues) and died between day 11 and 12 of gestation. Similarly, partial inhibition of VEGF-A during postnatal development results in higher mortality and failed organ development (Gerber and Ferrara, 1999a). VEGF-A also plays a key role in wound healing. Following injury, VEGF-A is released by activated platelets, recruits circulating neutrophils and

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monocytes, increases the permeability of endothelial cells, and induces endothelial cell proliferation and migration (Dvorak and Dvorak, 1995). VEGF-A is also involved in endochondral bone formation, a process required for longitudinal bone growth. VEGF-dependent blood vessels are essential for coupling cartilage resorption to bone formation (Gerber and Ferrara, 1999b). Finally, in physiological conditions, VEGF-A plays a role in angiogenesis in the female reproductive tract, inducing the formation of new capillary vessels required for luteal function (Fraser and Bicknell, 2000). VEGF-A is involved in various nonneoplasic disorders such as rheumatoid arthritis, diabetes, ischemic retinopathies, and psoriasis (Byrne and Harmey, 2005) but is also required for tumor neoangiogenesis, a critical step in tumor growth and survival. Beyond a volume of 2–3 mm3, a lack of oxygen and nutrients in the tumor induces the secretion of proangiogenic factors such as VEGF-A, an effect mediated by activation of the transcription factor (HIF)-1. This results in the creation of new tumor vessels, which facilitates tumor growth and tumor cell dissemination, resulting in the formation of distant metastases (Folkman and Williams, 1971). Several studies of patients with solid malignancies such as gastric cancer (Fondevila and Pera, 2004), non-small cell lung cancer (Mineo and Tonini, 2004), and advanced-stage hepatocellular carcinoma (Tseng and Hu, 2008) have shown a correlation between VEGF overexpression and poorer overall and/or disease-free survival.

5. ANTITUMOR IMMUNITY 5.1 Tumor Immunosurveillance 5.1.1 Basic Principles In 1909, Paul Erhlich postulated that the immune system might be able to recognize and eliminate tumor cells. Subsequently, Sir Burnet (1957) and Thomas (1982) established the concept of immunosurveillance. Mice lacking T and B lymphocytes (RAG 2-deficient mice) or interferon gamma (IFNγ) receptor signaling had a higher rate of spontaneous and carcinogeninduced malignant tumors (Shankaran and Schreiber, 2001). Epidemiological studies, mainly involving organ transplant recipients, showed that cancers of nonviral origin (colon, lung, pancreas, melanoma) were more frequent in immunodeficient or immunosuppressed patients than in immunocompetent subjects (Birkeland and Pukkala, 1995; Gatti and Good, 1971; Penn, 2000). Moreover, several cases of donor-derived melanoma were described in immunosuppressed transplant recipients, further confirming

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the importance of the immune system in preventing tumor development (MacKie and Junor, 2003; Strauss and Thomas, 2010). Finally, the presence in the tumor microenvironment, cytotoxic (CD8+) and memory (CD45RO+) T lymphocytes, and Th1 markers was linked to a lower incidence of tumor recurrence and to increased survival (Galon et al, 2006; Tosolini and Galon, 2011). Cancer cells share oncogenic alterations defined by Hanahan and Weinberg (2000) as hallmarks of cancer, including sustained proliferative signaling, evasion of growth suppressors, cell death resistance, replicative immortality, angiogenesis induction, and activation of invasion and metastasis. In 2011, the ability of tumor cells to evade immune destruction was identified as an emerging hallmark of cancer (Hanahan and Weinberg, 2011). The immune system exerts a form of selection pressure called “immunoediting,” leading from immune surveillance to immune escape and, finally, to the emergence of resistant tumor variants shaped by the host immune system. Tumor-induced immune subversion results from a complex interplay between tumor-derived factors and host cells (including immune cells) present in the tumor microenvironment. In addition to being involved in tumor survival and dissemination, the tumor microenvironment creates “tolerizing” conditions through the production of tumor-derived soluble factors, inhibition of DC maturation, accumulation of immunosuppressive cells (MDSCs, Tregs, TAMs), and expression of checkpoint inhibitors on effector T cells (Marigo and Bronte, 2008; Zou, 2005). 5.1.2 Main Effectors of the Antitumor Immune Response The antitumor immune response is largely dependent on the actions of cytokines that modulate tumor immunogenicity. IFNγ plays an important role in tumor immune surveillance. It affects both adaptative antitumor immune responses by promoting antigen processing and presentation via MHC class I and class II molecules, and innate antitumor responses by activating macrophages (Kaplan and Schreiber, 1998). Shankaran et al. have shown in mouse models that IFNγ and lymphocytes collaborate to prevent tumor development (Shankaran and Schreiber, 2001). IL-12 is another cytokine involved in the antitumor immune response, showing activity in a large variety of murine tumor models (Brunda and Gately, 1993; Robertson and Ritz, 1996). Finally, a lack of IL-12 or IFNγ favors the creation of an immunosuppressive microenvironment which prevents cancer cell immune destruction. Innate immune cells such as natural killer T (NKT) cells and NK cells also contribute to tumor immunosurveillance. NKT lymphocytes are

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characterized by a restricted directory and recognition pattern of glycolipids presented by CD1d, which is related to MHC class I molecules. Some of these cells express markers of NK cells. In 2000, Smyth et al. created a genetargeted and lymphocyte subset-depleted mouse model in order to establish the relative importance of NK and NKT cells in protecting against tumor initiation and metastasis. NKT cells were found to have a protective effect on spontaneous tumors initiated by chemical carcinogens (Smyth and Godfrey, 2000). An artificial ligand of NKT cells, α-galactosyl-ceramide (α-GalCer), was found to induce the regression of metastases. Two major mechanisms are responsible for NKT cell antitumor effects, namely, direct tumor cell lysis and secondary activation of NK cells. NK cells are particularly effective in destroying tumor cells that do not express MHC class I molecules (K€arre and Kiessling, 1986). They therefore have complementary roles to CD8 T cells, which require their targets to express MHC class I molecules. Tumor cells can express ligands like H60, Rae, and MIC-A that bind to activating receptors such as NKG2D or NCR (NKp46, NKp44, NKp30) on NK cells, resulting in NK cell activation and tumor cell lysis. NK cells exert their lytic functions through the perforin/granzyme, Fas Ligand, or TNF-related apoptosis-inducing ligand pathway (Smyth et al., 2000). Finally, NK cells can destroy tumor cells either directly or indirectly by producing cytokines such as IFNγ. Effector T cells also play a key role in tumor immunosurveillance through their cytotoxic activity toward antigen-bearing tumor cells (Russell and Ley, 2002). In colorectal cancer, Page`s and Galon found that abundant tumor-infiltrating effector memory CD45RO+ CD8+ T cells correlated with the absence of pathological signs of early metastatic invasion (vascular emboli, lymphatic invasion, and perineural invasion), and also with better overall and disease-free survival (Galon et al. 2006; Page`s et al. 2005). Increased tumor infiltration by cytotoxic CD8-positive T cells has been shown to correlate with prolonged survival in a variety of other epithelial cell cancers, including hepatocellular carcinoma (Wada and Kojiro, 1998), small-cell lung carcinoma (Eerola and P€a€akk€ o, 2000), esophageal cancer (Schumacher and Schlag, 2001), extrahepatic bile duct carcinoma (Oshikiri and Katoh, 2003), endometrial carcinoma (Kondratiev and Resnick, 2004), pancreatic adenocarcinoma (Fukunaga and Katoh, 2004), and urothelial carcinoma (Sharma and Sato, 2007). More recently, in colorectal cancer, Mlecnik et al. demonstrated that a high density of tumor

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infiltration by cytotoxic CD8+ and memory CD45RO+ T cells is a prognostic marker independent of the TNM stage (Mlecnik and Galon, 2011).

5.2 Escape From the Antitumor Immune Response 5.2.1 Cytokines Cytokines are at the heart of complex cross talk between cancer cells and immune cells. Cytokines can inhibit tumor development by controlling inflammation and immunity, but tumor cells can also use cytokines to promote their development and dissemination. Cytokines can influence tumor growth by acting directly on tumor cells or indirectly by attracting inflammatory cell types and by affecting angiogenesis (Dranoff, 2004). Immunosuppressive cytokines such as TGF-β and IL-10 can be generated both by tumor cells and by the tumor stroma, and can exert their action on both innate and adaptive immune cells. These factors can have a broader immunosuppressive effect in local lymph nodes and spleen, thereby favoring tumor spread and metastasis (Kim and Arihiro, 2006). Cells able to produce TGF-β include tumor cells, Tregs, and stromal cells, and TGF-β production has been associated with the growth of a large variety of tumors (Pasche, 2001). TGF-β is the most potent immunosuppressive factor able to reduce the proliferation, activation, and differentiation of immune cells (Chouaib and Blay, 1997). TGF-β inhibits T-cell proliferation by inhibiting IL-2 production (Brabletz and Serfling, 1993). In addition, TGF-β can regulate antigenpresenting cells either by inhibiting the activation of macrophages and their production of proinflammatory molecules (Bogdan and Nathan, 1993) or by preventing DC maturation (Geissmann and Durandy, 1999). Another immunosuppressive property of TGF-β is its ability to induce the conversion of conventional T cells into Tregs, by inducing expression of the transcription factor Foxp3 (Chen and Wahl, 2003; Ghiringhelli and Zitvogel, 2005a). TGF-β can also inhibit class II transactivator (CIITA) expression (Lee and Benveniste, 1997) and thereby prevent MHC class II upregulation on antigen-presenting cells. Various cell types, including tumor cells, T cells, B cells, and macrophages produce IL-10. IL-10 is another important immunosuppressive cytokine able to suppress cytokine expression by type 1 helper (Thl) T cells and NK cells (Fiorentino and O’garra, 1991), and also to inhibit IL-2 production and T-cell proliferation (Taga and Tosato, 1992). IL-10 also reduces the

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antigen-presenting capacity of monocytes by downregulating class II MHC expression (de Waal Malefyt and de Vries, 1991). Local secretion of IL-10 protects tumor cells from the lytic effects of cytotoxic T lymphocytes (CTLs) (Matsuda and Kiessling, 1994). IL-10 also inhibits nitrogen oxide production by IFNγ-activated macrophages (Gazzinelli and Sher, 1992). Il-10 has been found in tumor biopsies, serum, and ascitic fluid of cancer patients (Gotlieb and Martı´nez-Maza, 1992; Heckel and Platsoucas, 2011; Pisa and Kiessling, 1992). Recently, molecular analyses of head and neck tumors showed that the IL-10 pathway is associated with cancer progression (Bornstein and McWeeney, 2016). We will detail further the role of VEGF in the escape of immune system in tumor situation.

5.2.2 Maturation of DCs DCs are professional antigen-presenting cells. They are able to collect antigens, process them into immunogenic peptides, and then present them to T cells, in association with MHC class I and II molecules. In addition, they express costimulatory molecules such as B7-1 (CD80) and B7-2 (CD86), which are necessary for T-cell activation. B7-expressing antigen-presenting cells, and notably DCs, are the main cell population involved in tumorspecific immune responses (Huang and Levitsky, 1996). DCs can produce cytokines and stimulate the differentiation of effector T and NK cells (Banchereau and Steinman, 1998). DC dysfunction is observed both in tumor-bearing mice and in cancer patients (Gabrilovich and Carbone, 1996a). Indeed, in tumor-bearing mice and cancer patients, tumor-associated DCs fail to express costimulatory molecules like B7 (Chaux and Martin, 1997; Gabrilovich and Carbone, 1997; Nestle and Nickoloff, 1997), which may contribute to their poor capacity to stimulate allogeneic and tumor antigen-specific cytotoxic T-cell responses. Finally, immature DCs can induce Tregs, thus promoting tumor immune tolerance (Moser, 2003).

5.2.3 Accumulation of MDSCs MDSCs are a heterogeneous population (Melani and Colombo, 2003) characterized by their myeloid origin, immature status, and ability to suppress immune responses. MDSC numbers are elevated in tumor-bearing mice and in cancer patients (Almand and Gabrilovich, 2001) and correlate

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with the clinical stage and metastatic tumor burden (Diaz-Montero and Montero, 2009). In tumor-bearing mice, MDSCs produce reactive oxygen species such as the superoxide radical, which interact with NO, leading to the formation of peroxynitrite that induces conformational changes in TCR or CD8 molecules and consequently alters antigen-specific CD8+ T-cell responses (Nagaraj and Gabrilovich, 2007). MDSC express arginase 1, which depletes the medium of arginine, an amino acid involved in T-cell proliferation (Bronte and Zanovello, 2005; Zea and Ochoa, 2004), and also iNOS, which allows NO release in the tumor microenvironment and is involved in the inhibition of T-cell proliferation and induction of T-cell apoptosis (Mazzoni and Segal, 2002). MDSC are thus involved in tumor-associated T-cell tolerance (Huang and Chen, 2006; Kusmartsev and Gabrilovich, 2005). MDSCs have been shown to modulate macrophage cytokine production in animal models (Sinha and Ostrand-Rosenberg, 2007), and to inhibit NK cells in both animal models (Li and Cao, 2009) and patients with hepatocellular carcinoma (Hoechst and Korangy, 2009). In vivo, administration of 5-fluorouracil (5FU) to tumor-bearing mice reduces the number of MDSC in the spleen and tumor, suggesting that the antitumor effect of 5FU is partly mediated by its selective cytotoxic effects on MDSC (Vincent and Ghiringhelli, 2010). 5.2.4 Tumor-Associated Macrophages Macrophages are a major component of immune/inflammatory cells surrounding the tumor (Bingle and Lewis, 2002). Tumor-derived cytokines released into the tumor microenvironment, such as IL-4 and IL-10, are able to polarize tumor-infiltrating macrophages toward an “M2-like” phenotype characterized by IL-10high IL-12low expression. TAMs are preferentially located in hypoxic tumor areas, where they produce VEGF and other proangiogenic factors (Lewis and Lewis, 2000). TAMs have a poor antigenpresenting capacity and a reduced cytotoxic capability due to their weak NO production (Dinapoli and Lopez, 1996). They are able to suppress T-cell activation and proliferation by releasing prostaglandins, IL-10, and TGF-β (Balkwill and Mantovani, 2001). TAMs play a crucial role in tumor-induced inflammation, which promotes tumor growth, progression, invasion, and metastasis (Balkwill and Mantovani, 2001; Coussens and Werb, 2001; Mantovani and Ruco, 1992). Higher macrophage numbers in a variety of human tumors are associated with a worse prognosis (Bingle and Lewis, 2002).

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Additionally, in hypoxic areas of tumors, TAMs can increase MMP-7 expression (Burke and Lewis, 2003), leading to the cleavage of Fas ligand from the surface of tumor cells and thus making them less sensitive to lysis by NK and T cells (Fingleton and Matrisian, 2001). 5.2.5 Increase in Tregs Natural Tregs are involved in the control of self-reactive T cells, ensuring the maintenance of immunologic self-tolerance and thus preventing autoimmune disorders (Sakaguchi, 2004). In tumor situations, Tregs are characterized by CD4, CD25, and forkhead box protein 3 (FOXP3) expression and exhibit a particular phenotype associated with the expression of inhibitory receptors such as PD-1, Tim-3, CTLA-4, and Lag-3 (Huang and Vignali, 2004; Park and Ha, 2012; Wing and Sakaguchi, 2008). Tregs can prevent the development of an effective antitumor immune response by inhibiting CD8 T lymphocytes and NK cells. In murine tumor models, Treg depletion with an anti-CD25 antibody-induced tumor rejection (Casares and Lasarte, 2003). In tumor-bearing mice and patients with colorectal cancer, an increase in Tregs is observed in peripheral blood (Wolf and Grubeck-Loebenstein, 2003) and mesenteric lymph nodes (Clarke and Godkin, 2006), as well as inside the tumor (Chaput and Taieb, 2009). This increase is classically explained by four mechanisms: preferential recruitment of Tregs by the tumor, conversion of memory and naive T cells into Tregs (Mougiakakos, 2011), increased resistance of Tregs to oxidative stress (Mehrotra and Kiessling, 2009), and increased Treg proliferative capacity (Vukmanovic-Stejic and Akbar, 2006). Studies of different solid tumor types, including ovarian, pancreatic, and hepatocellular carcinoma, have shown a correlation between high numbers of intratumoral FOXP3-positive T regulatory cells and poor host outcomes (Curiel and Zou, 2004; Gao and Tang, 2007; Hiraoka and Hirohashi, 2006; Sato and Odunsi, 2005). 5.2.6 Immune Checkpoints and T-Cell Exhaustion As described earlier, the adaptive immune system, and particularly effector T cells, plays a major role in antitumor effects and cancer outcomes. T-cell exhaustion can thus help the tumor to escape the host immune system. T-cell exhaustion is characterized phenotypically by coexpression of immune inhibitory receptors known as immune checkpoints, such as

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program cell death-1 (PD-1), cytotoxic T-lymphocyte-associated protein 4 (CTLA-4), and T-cell immunoglobulin mucin-3 (Tim-3), and by a gradual loss of function (proliferation, cytokine secretion, and cytolytic activity). Little is known about the regulation of these receptors. PD-1–PD-L1 (PD-1 ligand) and Tim-3–Galectin 9 pathway blockade can partially restore T-cell function and improve antitumor immune responses in tumor-bearing mice (Sakuishi and Anderson, 2010). In heavily pretreated patients with various tumor types (non-small cell lung cancer, melanoma, renal cell cancer), anti-PD-1 antibody administration gave a 20–25% response rate and was associated with an increase in CD8+ T-cell infiltration (Hamid and Ribas, 2013; Topalian et al, 2012).

6. IMMUNOSUPPRESSIVE ROLES OF VEGF 6.1 Inhibition of DC Maturation In animal models, continuous VEGF-A infusion drastically reduces the number of mature DCs in the spleen, lymph nodes, and peripheral blood (Gabrilovich and Carbone, 1998). Likewise, in cancer patients, high plasma levels of VEGF-A correlate with a marked reduction in mature DCs and with an increased proportion of immature DCs in peripheral blood (Almand and Gabrilovich, 2000; Osada and Morse, 2008). The numbers of these immature cells decreases after surgical removal of the tumor (Almand and Gabrilovich, 2000), suggesting that defective DC maturation is due to a tumor-derived factor, possibly VEGF-A. VEGF-A prevents activation of the transcription factor nuclear factor-κB (NF-κB) via VEGFR-1 signaling, which affects the normal process of DC differentiation (Dikov and Carbone, 2005; Gabrilovich and Carbone, 1996b; Oyama and Gabrilovich, 1998), as shown in Fig. 2A. PlGF, a specific VEGFR-1 ligand, can also prevent DC differentiation (Dikov and Carbone, 2005). In an in vitro model of myeloid DC differentiation from murine embryonic stem cells exposed to VEGF-A, it was found that VEGFR-1 mainly inhibits the latter steps of DC maturation, whereas VEGFR-2 signaling rather affects DC differentiation from early hematopoietic progenitors (Dikov and Carbone, 2005). In a model of mature human monocyte-derived DC, Mimura et al. demonstrated that VEGF-A inhibits the allostimulatory capacity of DC through VEGFR-2 signaling (Mimura and Fujii, 2007). Using a VEGF-A infusion osmotic pump, Huang et al. developed a mouse model that mimics

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VEGF-A concentrations observed in patients with advanced cancer. They found that VEGF-A impaired allogeneic T-cell stimulation by DCs via both VEGFR-1 and -2 signaling. However, VEGFR-2 blockade was sufficient to restore DC function in this model (Huang and Carbone, 2007).

6.2 Accumulation of MDSCs VEGF-A administration promotes the accumulation of immature myeloid cells in the spleen and lymph nodes of tumor-free mice (Gabrilovich and Carbone, 1998). Similarly, elevated plasma VEGF levels in cancer patients are significantly related to increased MDSC numbers (Almand and Gabrilovich, 2000; Nakamura and Takenoshita, 2013; Osada and Morse, 2008). This MDSC accumulation mediated by VEGF-A is dependent on VEGFR-2 signaling (Huang and Carbone, 2007), and on activation of the Janus kinase2 (JAK2)/signal transducer and activator of transcription 3 (STAT 3) pathway (Nefedova and Gabrilovich, 2004), as shown in Fig. 2B. By inducing MDSC accumulation, VEGF indirectly suppresses both adaptive and innate antitumor immune responses.

6.3 Development of Tumor-Induced Macrophages VEGF-A induces monocyte/macrophage recruitment to the tumor and thereby promotes tumor-associated macrophage development (Linde and Mueller, 2012), as shown in Fig. 2C. However, VEGF-A alone is not sufficient to induce M2 polarization of macrophages: other cytokines produced by tumor cells, such as IL-4 and IL-10, are also necessary. Indeed, inhibition Fig. 2 (A) Impact of VEGF-A on dendritic cell (DC) maturation. VEGF inhibits DC maturation by preventing activation of the transcription factor nuclear factor-κB (NF-κB) via VEGFR-1 signaling. (B) Impact of VEGF-A on myeloid-derived suppressor cells (MDSCs). VEGF-A promotes MDSC accumulation, an effect dependent on VEGFR-2 signaling and on activation of the Janus kinase2 (JAK2)/signal transducer and activator of transcription 3 (STAT 3) pathway. (C) Impact of VEGF-A on tumor-associated macrophages (TAMs). VEGF-A induces tumor-associated macrophage development. (D) Impact of VEGF-A on regulatory T cells (Tregs). VEGF-A fosters Treg accumulation, an effect directly dependent on VEGFR-2 signaling. (E) Impact of VEGF-A on T-cell exhaustion. VEGF-A favors T-cell exhaustion by modulating the expression of the inhibitory checkpoints PD-1, Tim-3, and CTLA-4 and, to a lesser extent, Lag-3 on CD8+ T cells in a VEGFR-2-dependent manner. (F) Immunomodulatory properties of VEGF-A. Tumor-derived VEGF-A blocks DC maturation, favors MDSC and Treg accumulation, and induces TAM recruitment to the tumor. VEGF-A increases the expression of the inhibitory checkpoints PD-1, Tim-3, CTLA4, and Lag-3 on CD8+ T cells in a VEGFR-2-dependent manner. VEGF-A inhibits T-cell proliferation both directly and indirectly via Tregs, TAM, and MDSC.

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of the IL-4 receptor in vivo blocks M2 polarization, resulting in a less aggressive tumor phenotype (Lewis and Lewis, 2000).

6.4 Increase in Tregs A correlation between the Treg percentage and VEGF concentration has been observed in patients with stage IV colorectal cancer and melanoma, suggesting a role of VEGF-A in inducing and/or maintaining Tregs in vivo (Wada and Katano, 2009). VEGF-A can favor Treg accumulation indirectly through its ability to induce immature DC or immature myeloid cells such as MDSC (Ghiringhelli and Zitvogel, 2005b; Serafini and Borrello, 2008). A population of Tregs that express VEGFR-2 has been identified in tumor-bearing mice and cancer patients (Suzuki and Katano, 2010; Terme and Taieb, 2013). Suzuki et al. found that VEGFR-2 was selectively expressed by human CD4+FOXP3high Tregs and that this Treg population had a significantly stronger inhibitory effect on T-cell proliferation than did FOXP3+VEGFR2 cells (Suzuki and Katano, 2010). We subsequently found that VEGF-A could directly induce Treg proliferation via VEGFR-2 signaling (Fig. 2D) both in a mouse model of colorectal cancer and in colorectal cancer patients (Terme and Taieb, 2013). VEGF-A enhances the proliferation of Tregs from tumor-bearing mice but not from tumor-free mice. In addition, the specific population of FOXP3+VEGFR2+ cells in tumors is an independent predictive marker of recurrence and of poorer overall and disease-free survival in patients with colorectal cancer who have undergone curative resection (Suzuki and Katano, 2013). Interestingly, intratumoral FOXP3+ and FOXP3+VEGFR2 cell numbers did not correlate with clinical outcome in this study. Neuropilin-1 (NRP1), a membrane-bound VEGFR coreceptor expressed on Tregs, mediates Treg infiltration of tumors in a VEGFdependent manner (Hansen, 2013). In colorectal cancer liver metastases, it was recently shown that NRP1 is strongly expressed on CD3+CD4+ tumor-infiltrating lymphocytes by comparison with peripheral blood mononuclear cells (Chaudhary and Elkord, 2015).

6.5 Effector T Cells VEGF-A administration reduces both splenic T-cell numbers and the T-cell/B-cell ratio in the lymph nodes and spleen of tumor-free mice, and suppresses their functions (Gabrilovich and Carbone, 1998). T-cell

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defects and premature thymic atrophy occur in tumor-bearing animals and cancer patients. Ohm et al. showed that VEGF-A inhibits the production of T cells by modulating their thymic maturation. Indeed, in a mouse model, recombinant VEGF administration at concentrations similar to those observed in advanced-stage cancer patients led to a decrease in the earliest thymic hematopoietic progenitors (Ohm and Carbone, 2003). Later, Huang et al. showed that VEGFR-1 blockade aggravated thymic atrophy and that VEGFR-2 blockade in mice treated with VEGF normalized T-cell populations in both the spleen and the thymus, suggesting that inhibition of thymic T-cell development occurs through VEGFR-2 signaling and that VEGFR-1 counters this effect (Huang and Carbone, 2007). In vitro experiments show that VEGF also modulates T cells from ascitic fluid and peripheral blood of ovarian cancer patients via VEGFR-2 signaling, after their activation by anti-CD3 antibody and recombinant human interleukin (IL)-2 (Gavalas and Bamias, 2012; Ziogas and Bamias, 2012). More recently, in a mouse model of colorectal cancer (CT26) expressing high levels of VEGF-A, we demonstrated that VEGF-A modulates the expression of the inhibitory checkpoints PD-1, Tim-3, and CTLA-4 and, to a lesser extent, Lag-3, on CD8+ T cells in a VEGFR-2-dependent manner (Fig. 2E). Expression of these inhibitory checkpoints is associated with T-cell exhaustion. We also demonstrated the involvement of the transcription factor NFAT, which was known to be involved in VEGFR-2 signaling (Schweighofer and Hofer, 2009), and in the regulation of PD-1 and CTLA-4 expression (Gibson and Wong, 2007; Oestreich and Boss, 2008). All these findings demonstrate that VEGF-A increases the expression of inhibitory receptors involved in T-cell exhaustion, by activating the VEGFR-2–PLCg–calcineurin–NFAT pathway (Voron and Terme, 2015). VEGF thus has an immunosuppressive role in cancer (Fig. 2F).

7. IMMUNOMODULATORY PROPERTIES OF THERAPEUTICS TARGETING VEGF 7.1 Development of Major Antiangiogenic Molecules As tumors require new blood vessels for growth and metastasis, it was postulated that inhibition of angiogenesis could reduce tumor growth and thus improve mortality and morbidity in cancer patients (Folkman and Williams, 1971). In contrast to the physiological situation, the imbalance between proangiogenic and antiangiogenic factors persists in tumors. Consequently,

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antiangiogenic molecules targeting VEGF might restore the balance between pro- and antiangiogenic factors, deprive the tumor of oxygen and nutrients by destroying its blood supply, and normalize the tumor vasculature (Jain, 2005). Administration of VEGF-A or VEGF-R inhibitors reduces the tumor vasculature and prevents tumor growth both in animal models (Ferrara and Alitalo, 1999; Kim and Ferrara, 1993; Strawn and Shawver, 1996) and in cancer patients (Presta and Ferrara, 1997). In tumorbearing mice, treatment with a VEGF-specific monoclonal antibody reduced intratumoral blood vessel density and drastically inhibited tumor growth, suggesting that VEGF-targeting therapies might be effective in human cancers (Kim and Ferrara, 1993). Proof of concept was obtained in metastatic colorectal cancer, where the addition of an anti-VEGF antibody (bevacizumab) to standard first-line chemotherapy was found to significantly improve overall survival (Hurwitz and Kabbinavar, 2004). Three subclasses of antiangiogenic molecule targeting the VEGFA/VEGFR pathway have now been developed (1) tyrosine kinase inhibitors (TKIs) targeting VEGFRs, such as sorafenib, regorafenib, and sunitinib; (2) monoclonal antibodies such as bevacizumab (targeting VEGF-A) and ramucirumab (targeting VEGFR-2); and (3) aflibercept, a fusion protein composed of extracellular domains from VEGFR-1 and R-2 (targeting VEGFA, B, and PlGF) (Fig. 1). TKIs target the receptors of proangiogenic molecules under the cell membrane, whereas monoclonal antibodies and fusion proteins directly target circulating proangiogenic factors or their receptors present on the cell membrane. Antiangiogenic therapies have shown significant efficacy on various solid tumors in clinical trials (Table 1) and are now part of the oncologic armamentarium. Sunitinib targets VEGFR1–3, PDGFR, c-kit, and Flt3 (Chow and Eckhardt, 2007) and has been approved for the treatment of various tumor types, including gastrointestinal stromal tumors (Demetri and Casali, 2006) and renal cell carcinoma (Motzer and Basch, 2007). Sorafenib is an orally active TKI that targets VEGFR1–3, PDGFR, c-kit, Raf-kinases, and RET (Wilhelm and Kelley, 2006) and is widely approved for standard care of patients with hepatocellular carcinoma (Cheng and Guan, 2009) and advanced renal cell carcinoma (Escudier and Bukowski, 2007). Bevacizumab specifically targets VEGF-A and has also proved effective on various tumors, including metastatic colorectal cancer (Giantonio and Benson, 2007; Hurwitz and Kabbinavar, 2004), breast cancer (Miller and

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Table 1 Targets and Current Indications of VEGF-Targeting Therapies Antiangiogenic Target Indication Tyrosine kinase inhibitors

Sunitinib

VEGFR 1–3/PDGFR/ cKit/Flt3

GIST, mRCC (1st line), pancreatic neuroendocrine tumors

Sorafenib

VEGFR 1–3/PDGFR/ cKit/Raf-kinases/RET

HCC (1st line), advanced RCC (1st line), thyroid cancer

Regorafenib

VEGFR 1–3/PDGFR/ FGFR/Kit/RET/BRAF

Refractory CRC, refractory GIST

Monoclonal antibodies

Bevacizumab

VEGF-A

Ramucirumab VEGFR-2

mCRC (1st and 2nd line), breast cancer, NSCLC, ovarian cancer, RCC, glioblastoma, cervical cancer mCRC (2nd line), advanced gastric or gastroesophageal adenocarcinoma (2nd line), NSCLC

Fusion protein

Aflibercept

VEGF-A/VEGF-B/PlGF

mCRC (2nd line)

GIST, gastrointestinal stromal tumor; HCC, hepatocellular carcinoma; mCRC, metastatic colorectal cancer; NSCLC, non-small cell lung cancer; RCC, renal cell carcinoma.

Davidson, 2007), and lung cancer (Herbst and Hainsworth, 2011; Sandler and Johnson, 2006) when combined with standard chemotherapy. Antiangiogenic therapies have also proven beneficial after tumor progression. Several preclinical studies show that sustained VEGF inhibition induces and maintains tumor regression in several malignancies, including prostate, ovary, and breast cancer (Klement and Kerbel, 2002; Melnyk and Shuman, 1999; Mesiano and Jaffe, 1998). In a model of multidrug resistance using human breast cancer xenografts, Klement et al. reported a significant and durable response to various continuous low-dose chemotherapy regimens combined with an anti-VEGR-2-neutralizing antibody (Klement and Kerbel, 2002). The CORRECT study demonstrated that antiangiogenic treatment with regorafenib was beneficial in patients with colorectal cancer that had progressed despite all standard treatments, including several antiangiogenic agents (Grothey and Laurent, 2013).

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In patients with metastatic colorectal cancer previously treated with oxaliplatin, the addition of aflibercept to FOLFIRI as a second line of treatment significantly improved survival when compared to FOLFIRI plus placebo (Van Cutsem and Allegra, 2012). A subgroup analysis of this study showed benefits whether or not the patients had already received bevacizumab. The TML study, a multicenter phase III trial in patients with colorectal cancer already treated with bevacizumab plus standard first-line chemotherapy showed a survival benefit when bevacizumab was continued with a second-line chemotherapy after disease progression (Bennouna and Kubicka, 2013). Ramucirumab, another antiangiogenic drug targeting VEGFR-2, was recently shown to be effective in a randomized, double-blind, placebocontrolled phase III trial in patients with metastatic colorectal carcinoma who progressed during or after first-line treatment including bevacizumab. Overall survival and progression-free survival were significantly enhanced in patients treated with ramucirumab plus FOLFIRI as compared to placebo plus FOLFIRI (Tabernero and Nasroulah, 2015). Ramucirumab is also an approved second-line treatment for gastric cancer (Wilke and Ohtsu, 2014). Thus, continuous antiangiogenic pressure exerted by VEGF inhibition, while changing chemotherapy beyond progression, improves patient survival. Several studies in patients with other types of cancer, such as metastatic breast and non-small cell lung cancer, confirm this benefit.

7.2 Immunomodulatory Activity of Antiangiogenic Therapies in Animals As VEGF-A modulates immunity, the off-target effects of antiangiogenic drugs have been studied in mouse tumor models and in cancer patients. Anti-VEGF antibody administration enhanced DC numbers and functions in the lymph nodes and spleen of tumor-bearing mice (Gabrilovich and Carbone, 1999). Alfaro et al. studied the effects of anti-VEGF therapies on the transition from myeloid immature precursors to DCs. Using VEGF and supernatants of renal carcinoma cell lines cultured in hypoxic conditions, they created a model of altered human monocyte differentiation into DCs. They found that bevacizumab and sorafenib, but not sunitinib, reversed the inhibitory effects of recombinant VEGF on DC differentiation. Indeed, DC matured under the influence of VEGF expressed less human

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leukocyte antigen-DR (HLA-DR) and CD86, and this effect was blocked by bevacizumab and sorafenib (Alfaro and Perez-Gracia, 2009). In sunitinib-treated tumor-bearing mice, the proportion of MDSC decreased in a dose-dependent manner in the spleen, bone marrow, and tumor. Inhibition of STAT 3 phosphorylation by sunitinib could be responsible for this decrease (Xin and Yu, 2009). In addition, sunitinib reduces the immunosuppressive activity of MDSC in tumor-bearing mice (Draghiciu and Daemen, 2015; Ozao-Choy and Chen, 2009). Two different mechanisms could explain this action on MDSC: first, sunitinib inhibits the proliferation of the monocytic subset of MDSC (Gr1lo), and second, it induces apoptosis of the granulocytic subset (Gr1hi) (Ko and Cohen, 2010). Sorafenib also reduced the proportion of MDSC in an animal model of liver carcinoma (Cao and Liu, 2011). In a mouse model of renal cancer, antiVEGF antibody treatment reduced the number of CD11b+VEGFR1+ myeloid cells (Kusmartsev and Vieweg, 2008). In various tumor models, a reduced fraction of Tregs among CD4 T cells has been observed in peripheral blood, spleen, and tumor after treatment with sunitinib or sorafenib (Cao and Liu, 2011; Ozao-Choy and Chen, 2009; Xin and Yu, 2009). The ability of sunitinib to inhibit STAT 3 could partly explain its effect on Treg accumulation, by blocking the conversion of conventional CD4+Foxp3 T cells into tumor-associated CD4+FOXP3+ Tregs (Kujawski and Yu, 2010). In addition, a decrease in the number of MDSC induced by sunitinib could contribute to reducing the proportion of Tregs. Sorafenib reduces the Treg percentage both by inhibiting proliferation and inducing apoptosis, and also impairs Treg suppressive functions in vivo (Chen and Cheng, 2014). In a mouse model of colorectal cancer, administration of sunitinib, or an anti-VEGF-A antibody prevented Treg accumulation in the spleen and tumor, whereas masitinib (a TKI not targeting VEGFR) had no impact on the Treg proportion or number, demonstrating that specific blockade of the VEGF-A/VEGFR pathway is sufficient to prevent Treg accumulation (Terme and Taieb, 2013). In mice, sunitinib increases the proportion of CD4+ T and CD8+ T cells among tumor-infiltrating lymphocytes (Ozao-Choy and Chen, 2009). Shrimali et al. have also shown that agents targeting the VEGF/VEGFR-2 pathway enhance the antitumor activity of adoptively transferred antitumor T cells, a therapeutic strategy that has proved its efficacy in both murine models and in patients with metastatic melanoma (Shrimali and Rosenberg, 2010). In addition, a monoclonal antibody specific for VEGFR-2 increased tumorspecific CD8+ T cells (Manning and Emens, 2007). The ability of angiogenesis

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inhibitors to increase leukocyte/vessel-wall interactions by modulating the expression of adhesion molecules on tumor endothelial cells and, thus, leukocyte infiltration of tumor tissue could be one of the mechanisms explaining these results (Dirkx and Griffioen, 2006). Sunitinib also prevents tumor antigenspecific T-cell anergy by stimulating IFNγ production and cytolytic activity against the tumor (Ozao-Choy and Chen, 2009). In contrast, in vitro studies have shown that sorafenib reduces the proliferation and activation of effector T cells by inhibiting Raf/MEK/ERK signaling and LCK phosphorylation (Zhao and Xu, 2008). In tumor-bearing mice treated with sorafenib, the IFNγ production and activation status of CD8+ effector T cells were both improved. The cytotoxic activity of effector T cells was not significantly impaired. All these findings suggest that sorafenib differentially affects the diverse effector mechanisms of CD8+ T cells (Chen and Cheng, 2014). Recently, in a mouse colorectal cancer model (CT26) treated with an anti-VEGF-A antibody or with TKIs targeting (sunitinib) or not targeting (masitinib) VEGFR, we found that targeting VEGF-A–VEGFR reduced the coexpression of inhibitory receptors associated with T-cell exhaustion (PD-1, Tim-3, CTLA-4, Lag-3) and restored IFNγ production by intratumoral CD8+ T cells (Voron and Terme, 2015). Finally, sunitinib reduces the expression of immunosuppressive cytokines like IL-10 and TGF-β in the tumor microenvironment (Ozao-Choy and Chen, 2009).

7.3 Immunomodulatory Activity of Antiangiogenic Therapies in Humans 7.3.1 Bevacizumab Bevacizumab, which directly and specifically targets VEGF-A, induces a slight increase in the DC population in peripheral blood, restores their maturation, and enhances their allostimulatory and cytokine production capabilities when administered to patients with lung, breast, or colorectal carcinoma (Osada and Morse, 2008). However, the impact of bevacizumab on MDSC in cancer patients is unclear. Osada et al. reported a reduced number of immature myeloid progenitor cells after bevacizumab treatment in patients with several types of solid cancer (Osada and Morse, 2008). Nevertheless, bevacizumab did not reduce the accumulation of MDSC in peripheral blood of patients with metastatic renal cell carcinoma (mRCC) (Rodriguez and Ochoa, 2009). The discrepancies between these results could be elucidated by using markers to identify MDSC population. Indeed, Osada et al. defined MDSC by CD45+ lin HLA-DR staining, whereas

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Rodriguez characterized granulocytic MDSC by their CD66b expression, suggesting that immunomodulation by bevacizumab only affects MDSC with an immature phenotype. Wada et al. observed a reduction in the Treg percentage in peripheral blood of metastatic colorectal cancer patients treated with anti-VEGF-A (Wada and Katano, 2009). Surprisingly, in another study of previously untreated mRCC patients, bevacizumab plus low-dose IL-2 increased Treg cell proportion without affecting DC activation (Garcia and Rini, 2011). These contradictory results could be explained by the ability of low doses of IL-2 to promote Treg expansion. Indeed, Koreth et al. have shown in patients with active chronic graft-vs-host disease that daily subcutaneous low-dose IL-2 injection increases Treg counts and also the regulatory T-cell/conventional T-cell ratio (Koreth and Soiffer, 2011). In colorectal cancer patients, we have demonstrated that VEGF-A/VEGFR pathway blockade can prevent Treg accumulation by inhibiting their VEGF-induced proliferation (Terme and Taieb, 2013). Finally, in metastatic colorectal cancer patients, bevacizumab increases CD4, CD8, and CD3 lymphocyte numbers (Manzoni and Danova, 2010) as well as IL-2 and IFNγ production (Tsavaris and Baxevanis, 2012). 7.3.2 Sorafenib Sorafenib reduced the proportion of circulating and tumor-infiltrating Tregs, respectively, in patients with hepatocellular carcinoma (Nagai and Sumino, 2012) and renal cell carcinoma (Busse and Keilholz, 2011; Desar and de Vries, 2011). Molhoek et al. found that sorafenib promoted apoptosis of CD4+CD25high T cells in vitro (Molhoek and Slingluff, 2009). As we have already seen, sorafenib can inhibit the maturation of DCs, which is involved in Treg differentiation. In two studies of mRCC patients and cirrhotic patients with advanced hepatocellular carcinoma, sorafenib had no impact on Th1 responses (Busse and Keilholz, 2011; Nagai and Sumino, 2012). Sorafenib has also been shown to modulate innate immune cells such as NK cells. In an in vitro study, sorafenib impaired granule mobilization and, thus, NK cell reactivity, by downregulating PI3Kinase and Erk1/2 phosphorylation (Krusch and Salih, 2009). Nevertheless, the concentrations used in this study did not correspond to those observed in cancer patients treated with these TKIs. In nasopharyngeal carcinoma cell lines, Huang et al. found that sorafenib and sunitinib upregulated NKG2D ligands on tumor cells and

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thereby increased tumor sensitivity to NK cell lysis (Huang and He, 2011). Kohga et al. reported that ADAM9, a metalloproteinase overexpressed in hepatocarcinoma (HCC) cell lines but not in normal tissues, is involved in the shedding of MHC class I-related chain A (MICA). ADAM9 knockdown enhanced the sensitivity of human HCC cells to NK cells, an effect that involved increased expression of membrane-bound MICA on ADAM9 / HCC cells. Sorafenib reduced ADAM9 expression, increased membrane-bound MICA expression, and reduced soluble MICA levels in HCC cells. Sorafenib thus enhanced the NK sensitivity of HCC cells (Kohga and Hayashi, 2010). 7.3.3 Sunitinib In patients with kidney cancer, sunitinib significantly decreases the number of MDSC in peripheral blood (Ko and Finke, 2009). Sunitinib also reduces Treg numbers in the peripheral blood and tumors of mRCC patients. Finke et al. have shown that the largest decline in Tregs occurs after the end of the first treatment cycle with sunitinib, and functional studies suggest that Treg suppressive activity is impaired after sunitinib treatment (Finke and Bukowski, 2008). In mRCC patients, neoadjuvant treatment with sunitinib reduces intratumoral Treg numbers (Adotevi and Tartour, 2010). Sunitinib enhances type-1 cytokine responses but reduces type-2 responses when peripheral blood mononuclear cells from patients with mRCC are stimulated in vitro with anti-CD3/anti-CD28 antibodies (Finke et al., 2008; Ko and Dreicer, 2009). This effect on T-cell responses may be explained by a decrease in the number of Tregs (Finke et al., 2008) and in the MDSC population (Ko and Finke, 2009), both phenomena being induced by sunitinib. Thus, sunitinib and sorafenib seem to modulate T-cell functions differently.

8. MEDICAL APPLICATIONS 8.1 Antiangiogenic Drugs as Ideal Companions for Anticancer Immunotherapy: Preclinical Studies As described earlier in this review, besides their well-known antiangiogenic properties, VEGF-targeting therapies modulate the immunosuppressive tumor microenvironment, enhance the Th1 response of T cells, and increase tumor-infiltrating T-cell numbers (Finke and Bukowski, 2008). Nevertheless, as shown in a mouse model of colorectal cancer expressing

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carcinoembryonic antigen, these agents do not seem to restore a specific T-cell response to tumor antigens (Farsaci and Hodge, 2012). Immunotherapy seeks to boost natural antitumor immunity (Escors, 2014). Cytokines such as IL-2 and IFNγ show modest efficacy in metastatic melanoma and renal carcinoma patients. Sipuleucel-T, an autologous active cellular vaccine, enhances overall survival in patient with castration-resistant metastatic prostate cancer (Kantoff and Schellhammer, 2010). Other immunotherapeutic strategies are designed to counterbalance tumor-induced immunosuppression. Antibodies targeting immune checkpoint inhibitors have proved their interest in cancer treatment. Several phase III studies demonstrate that immune checkpoint blockade can enhance objective and durable responses in previously treated patients with advanced or metastatic solid cancers. Anti-CTLA-4 administration increased overall survival in patients with metastatic melanoma (Hodi and Urba, 2010). AntiPD1 antibodies enhanced survival in previously treated patients with metastatic melanoma, advanced non-small cell lung carcinoma (NSCLC) (Borghaei and Brahmer, 2015; Hamid and Ribas, 2013), or advanced renal cell carcinoma (Motzer and Sharma, 2015). These antibodies have been approved for the treatment of metastatic melanoma and advanced non-small cell lung cancer. However, not all patients benefit from immunotherapy. Interestingly, VEGF-targeting therapies only restore Treg numbers to a physiological level and do not deplete activated T cells, unlike other strategies targeting Tregs (anti-CD25, cyclophosphamide) (Pere and Tartour, 2011; Taieb and Zitvogel, 2006). They also inhibit other immunosuppressive pathways, such as MDSC. As described earlier, antiangiogenic agents transiently normalize tumor vascularization (Matsumoto and Krishna, 2011), which could enhance CD8+ T-cell infiltration into the tumor after vaccination (Manning and Emens, 2007) and reduce immune checkpoint expression by tumor-infiltrating CD8+ T cells. Antiangiogenic molecules have been combined with vaccination strategies in preclinical models. In murine tumor models, VEGF blockade with adenoviral vectors expressing soluble VEGFR-1 and -2, combined with a GM-CSF-secreting tumor cell vaccine, significantly increased the ratio of effector T cells to Tregs in the tumor microenvironment, thereby enhancing antitumor immune protection (Li and Jooss, 2006). Antiangiogenic molecules have also been combined with recombinant viral vectors expressing tumor antigens or costimulatory molecules. Thus, sunitinib combined with an immunotherapeutic protocol associating an adenoviral vector expressing IL-12 with an activated costimulatory molecule (4.1BB ligand) improved

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antitumor efficacy and long-term survival in a MCA26 colorectal tumor model by counteracting intratumoral immune suppression (Ozao-Choy and Chen, 2009). In woodchucks bearing hepatocellular carcinoma, concurrent administration of adenovirus-delivered genes encoding IL-12 and GM-CSF plus antiangiogenic gene therapy with endostatin and pigment epithelium-derived factor genes led to greater regression of large tumor burdens when compared with either treatment alone. This combination increased NK cell numbers and markedly reduced expression of the immunosuppressive molecules CTLA-4 and PD-1 in the tumor microenvironment (Huang et al., 2010). In a breast cancer model, low doses of an anti-VEGFR-2 antibody lead to vascular normalization and improved the efficacy of a whole-cancer-cell vaccine through CD8+ T-cell recruitment and activation (Huang and Poznansky, 2012). Hamzah et al. found that in mice lacking the regulator of G-protein signaling 5 (Rgs5) gene, tumors showed vascular normalization, more mature pericytes, and massive infiltration by CD8+ and CD4+ T cells, leading to better survival than in wild-type mice (Hamzah and Ganss, 2008). Similarly, histidine-rich glycoprotein (HRG) gene gain of function, which downregulates the expression of PlGF by TAMs, normalized the aberrant vasculature, and tumors that overexpressed HRG also contained higher numbers of CD8+ CTLs (Rolny and Claesson-Welsh, 2011). All these findings suggest that vascular normalization helps to lift tumor-induced immunosuppression. Bose et al. showed that a 7-day course of sunitinib made the tumor microenvironment less immunosuppressive and more sensitive to the recruitment and prolonged action of CD8+ T cell induced by vaccination with OVA peptide-pulsed DCs, leading to superior antitumor effectiveness. In this B16.OVA melanoma model, enhancement of OVA-specific CD8+ T-cell responses correlated with an increase in chemotactic signals that may influence antigen-specific CD8 T lymphocyte recruitment to the tumor (Bose and Storkus, 2011). Similarly, in mice immunized with DCs pulsed with p53 peptides, concurrent administration of an anti-VEGF antibody increased the intensity and duration of the antitumoral effect (Gabrilovich and Carbone, 1999). More recently, a combination of immune checkpoint inhibitors and VEGF blockade gave promising results in vivo. Indeed, in a CT26 mouse model of colorectal cancer, a combination of anti-VEGF-A and anti-PD-1 significantly reduced tumor growth as compared with either antibody alone (Voron and Terme, 2015). Anti-PD-1-induced antitumor responses in mice bearing VEGF KO tumors but not their wild-type counterparts producing

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high levels of VEGF. These results suggest that VEGF may induce resistance to anti-PD-1 treatment. The addition of an anti-VEGF antibody restored anti-PD-1 efficacy against VEGF-producing tumors. These preclinical results suggest that a combination of anti-VEGF and anti-PD-1 may be beneficial in patients with tumors that secrete large amounts of VEGF, such as colorectal and renal cell cancers. Together, these preclinical data demonstrate a synergy between VEGF inhibition and immunotherapy, suggesting that VEGF-targeting therapies may enhance immunotherapy in cancer patients. One important issue is the best schedule of drug administration. Farsaci et al. reported that sunitinib administration before poxvirus-based vaccination yielded superior antitumor efficacy, longer survival, and more efficient tumor infiltration by antigen-specific T lymphocytes in mice. However, no modification of intratumoral immune cell infiltration and no antitumor benefit were found when sunitinib was administered after or concurrently with vaccination (Farsaci and Hodge, 2012). Another study suggested that concomitant administration of sunitinib with a vaccine against a tumor antigen inhibited T lymphocyte priming due to a dramatic decrease in CD11b+CD11c+ antigen-presenting cells, leading to vaccination failure (Jaini and Tuohy, 2014). The better antitumor responses observed when sunitinib administration precedes immunotherapy are in keeping with the fact that CD8+ T lymphocyte numbers fall rapidly after the beginning of sunitinib therapy, before expanding again as treatment continues, and that the proportion of Tregs and MDSC is reduced only after 5–7 days of sunitinib treatment. In order to further improve combined antiangiogenic therapy and immunotherapy, the choice of antiangiogenic drug and its dosage are also important factors. As they act both on Tregs and on MDSC, VEGFA/VEGFR-2-targeting therapies seem to be the best choice among the antiangiogenic therapies.

8.2 First Clinical Results In 2010, sunitinib was evaluated in combination with an MVA-based cancer vaccine encoding the tumor-associated antigen 5T4 (MVA-5T4; TroVax) in a randomized phase III study in renal cell carcinoma. No difference in survival was found between patients receiving sunitinib alone and patients receiving sunitinib plus the vaccine. However, the patients were vaccinated before receiving sunitinib, which, according to preclinical observations, may

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explain this disappointing result (Amato and Harrop, 2010). Later, two renal cancer patients experienced a regression of metastatic lesions and disease stabilization after nephrectomy and DC vaccination preceded by sunitinib treatment, suggesting that the synergy between sunitinib and DC vaccine observed in animal models may also occur in cancer patients (Dall’Oglio and Srougi, 2011). Recently, in an open-label phase II trial, Amin et al. evaluated the combination of AGS-003, an autologous DC-based immunotherapy, with sunitinib in 25 patients with intermediate- or poor-risk mRCC. In this study, sunitinib was administered before AGS-003 immunotherapy, at a dose of 50 mg daily for 4 weeks, followed by 2 weeks of treatment. At the end of the first sunitinib cycle, patients were treated with AGS-003 every 3 weeks for 5 doses and then every 12 weeks. Treatment continued until disease progression, toxicity, or the end of the study. Among the 21 assessable patients, 13 patients (62%) experienced a clinical benefit, with 9 partial responses (43%) and 4 disease stabilizations (19%) (Amin and Figlin, 2015). The median overall survival times were 30.2 months for all patients, 61.9 months for intermediate-risk patients and 9.1 months for poor-risk patients, compared to, respectively, 22, 27, and 8.8 months in patients treated with sunitinib alone (Heng and Choueiri, 2009). No complete responses were observed. The most common adverse events were those expected with sunitinib, while AGS-003 was well tolerated with only mild injection-site reactions. In the same study, immunomonitoring of 14 patients revealed that, in 10 patients, the absolute number of CD8+ CD28+ CD45RA effector/memory T cells (CTLs) in peripheral blood was enhanced between baseline and the fifth dose of AGS-003 and that it correlated with overall survival (Amin and Figlin, 2015). This phase II trial established the interest of a therapeutic schedule combining vaccine immunotherapy with antiangiogenic therapy, at least in mRCC. These results justified an international phase III trial which is currently evaluating autologous DC immunotherapy (AGS-003) plus standard sunitinib therapy in patients with advanced renal cell carcinoma. It is planned to randomize 450 patients and the primary outcome measure is overall survival (NCT01582672; Figlin, 2015). Several other studies are currently testing combinations of antiangiogenic drugs with immunotherapies (Table 2). An international phase III trial in mRCC patients (NCT01265901) is evaluating overall survival in patients treated with a multipeptide cancer vaccine called IMA-901 plus GM-CSF, preceded by a single low dose of cyclophosphamide, in

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Table 2 Current Clinical Trials of Combined VEGF-Targeting Drugs and Immunotherapies VEGF-Targeted Registration Therapy Immunotherapy Number Cancer Location

Phase

Sunitinib

AGS-003 vaccine

NCT01582672 Advanced or metastatic RCC

III

IMA901 + GM-CSF

NCT01265901 Advanced or metastatic RCC

III

Nivolumaba Nivolumab

NCT02400385 Advanced melanoma II

a

NCT01472081 mRCC

I

NCT00389285 mRCC

II

Sorafenib

Interleukin-21

Bevacizumab

TG4010 vaccine NCT01383148 Stage IV NSCLC

IIB/III

DC vaccine

NCT02010606 Glioblastoma

I

Nivolumaba

NCT01454102 Stage IIIb/IV NSCLC

I

Pembrolizumaba NCT02337491 Glioblastoma a

Pembrolizumab NCT02039674 NSCLC

I/II

Pembrolizumaba NCT02348008 RCC

I/II

b

NCT01950390 Unresectable stage III/IV melanoma

II

Ipilimumabb

NCT00790010 Unresectable stage III/IV melanoma

I

Atezolizumabc

NCT01633970 Locally advanced or metastatic solid tumors

I

Ipilimumab

Ramucirumab Pembrolizumaba NCT02443324 Gastric or gastroesophageal adenocarcinoma, NSCLC, TCCU a

II

I

Anti-PD-1 antibody. Anti-CTLA-4 antibody. Anti-PDL1 antibody. CRC, colorectal cancer; mCRC, metastatic colorectal cancer; (m)RCC, (metastatic) renal cell carcinoma; NSCLC, non-small cell lung carcinoma; TCCU, transitional cell carcinoma of the urothelium.

b c

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combination with sunitinib. A phase II trial is testing sorafenib in combination with recombinant IL-21 (NCT00389285). Another phase II study is recruiting patients with resected hepatic metastasis of colorectal carcinoma in order to evaluate bevacizumab in combination with DC vaccination (Robert and Hu-Lieskovan, 2016). Phase I/II clinical trials are also testing the safety and tolerability of combinations of antiangiogenic drugs targeting VEGF or their receptors (sunitinib or bevacizumab) with immunomodulatory drugs, especially antibodies targeting inhibitory immune checkpoints (anti-PD-1 and anti-CTLA-4) in patients with mRCC (NCT01472081; NCT02348008) and other types of solid cancer, including non-small cell lung cancer (NCT02039674), unresectable stage III or IV melanoma (NCT02400385), and newly diagnosed or recurrent glioblastoma (NCT02337491). A phase I study is evaluating the combination of ramucirumab plus pembrolizumab (an anti-PD-1 antibody) in patients with locally advanced and unresectable or metastatic gastric or gastroesophageal junction adenocarcinoma, non-small cell lung cancer, or transitional cell carcinoma of the urothelium (NCT02443324). Finally, an open-label phase Ib trial is designed to assess the safety, pharmacology, and preliminary efficacy of atezolizumab (anti-PDL1) administered with bevacizumab or with bevacizumab plus oxaliplatin, leucovorin and 5-fluorouracil, or other chemotherapy regimens, in patients with locally advanced or metastatic solid tumors (NCT01633970).

8.3 Other Potential Applications The adverse effects of antiangiogenic drugs include cardiovascular, renal, gastrointestinal, and skin disorders, leading to a search for predictive biomarkers of efficacy/resistance and tolerability in order to identify those patients most likely to benefit. In addition, predictive biomarkers could help to anticipate antiangiogenic drug refractoriness during treatment. Currently, no biomarker has been validated for routine use in antiangiogenic therapy. As plasma levels of VEGF, PlGF, and sVEGFR-2 are modulated by VEGF-targeting therapies, the value of these molecules as pharmacodynamic markers of antiangiogenic drug responsiveness has been evaluated. However, no correlation with clinical outcomes has been found either at baseline or during therapy (Motzer and Rini, 2006; Norden-Zfoni and Heymach, 2007).

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Biomarkers of tumor metabolism studied with positron emission tomography (Prior and Leyvraz, 2009) or vessel architecture studied with MRI, for example, (Emblem and Sorensen, 2013) may be useful for predicting clinical outcomes. Indeed, pretreatment measurements of capillary permeability by means of dynamic contrast-enhanced MRI, and early pharmacodynamic reduction in tumor enhancement and density, could be predictive of outcomes in mRCC and glioma patients treated with VEGF inhibitors (O’Connor and Jayson, 2012), although this has to be confirmed in larger randomized studies. Proteomic analysis identified mesothelin, fms-like tyrosine kinase-4 (FLT4), and α1-acid glycoprotein as three potential predictors of outcome in ovarian cancer patients treated with bevacizumab (Collinson and Banks, 2013). In a study of bevacizumab-treated patients with ovarian cancer, Backen et al. also found that the plasma concentration of two angiogenesis-associated proteins called angiopoietin 1 (Ang1) and its receptor Tie-2 could predict progression-free survival (Backen and Jayson, 2014), but this too needs to be validated in prospective patient cohorts. Other research has focused on circulating endothelial cells (CEC) and their bone marrow-derived precursors called circulating endothelial progenitors (CEP). Indeed, elevated CEC counts are observed in the peripheral blood of some cancer patients at diagnosis, and CEC numbers normalize in patients who enter complete remission. These cells that depend on high local levels of VEGF for their survival could potentially be helpful for monitoring antiangiogenic therapies. Indeed, higher baseline CEC counts (Dellapasqua and Colleoni, 2008) and a reduction in CEP numbers during therapy (Bertolini and Kerbel, 2006; Farace and Soria, 2007) have been found to correlate with a clinical response in patients receiving antiangiogenic therapy. Immune cells and cytokines are other potentially interesting biomarkers. Elevated circulating levels of the proangiogenic cytokine IL-8 have been observed in several tumor models resistant to sunitinib. In a model of HIF-1α knockdown colon cancer cells, resulting in partial blockade of hypoxia-induced VEGF, Mizumaki et al. observed an upregulation of IL-8 signaling by comparison with wild-type cells. Additionally, inhibition of IL-8 by a neutralizing antibody prevents angiogenesis and tumor growth in HIF-1α-deficient colon cancer cells but not in wild-type xenografts (Mizukami and Chung, 2005). Using xenograft models that mimic clinical resistance to sunitinib, Huang et al. found that elevated secretion of IL-8 by

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tumors correlated with the sunitinib resistance phenotype, which was reversed by coadministration of an IL-8-neutralizing antibody. In this study, intratumoral IL-8 levels were also enhanced in mRCC patients who did not respond to sunitinib (Huang and Teh, 2010). IL-8 signaling appears to compensate for inhibition of the VEGF/VEGFR pathway in order to preserve tumor angiogenesis. In a phase II study of patients treated with a bevacizumab-containing regimen for metastatic colorectal cancer, changes during treatment in plasma levels of cytokines and angiogenic factors (CAF) were studied for their roles in the treatment response and therapeutic resistance. Interestingly, elevated plasma bFGF, PlGF, HGF, and myeloid recruitment factor levels were observed in patient subsets before radiographic progression. However, baseline levels of VEGF and VEGFR-2 were not associated with differences in overall or progression-free survival (Kopetz and Heymach, 2010). In early-stage NSCLC patients treated with pazopanib, which targets VEGFR, PDGF-R, and c-kit, baseline levels of 11 CAF correlated significantly with tumor shrinkage. High CAF levels correlated with better outcomes. The plasma concentration of IL-12 showed the strongest association with clinical outcome. Multivariate analysis revealed that a baseline CAF profile that included hepatocyte growth factor and IL-12 identified responding patients with 81% accuracy (Nikolinakos and Heymach, 2010). Cellular immune parameters could also be useful to predict the response to antiangiogenic drugs. Sunitinib induces a reduction in absolute monocyte numbers. After the first 2 weeks of treatment, monocytes undergo the largest reduction among all peripheral blood mononuclear cells. In addition, patients who derived a clinical benefit from sunitinib treatment had a significantly smaller decrease in monocyte numbers after the first 2 weeks of treatment in cycle 1, as compared to patients with progressive disease (Norden-Zfoni and Heymach, 2007). A decrease in circulating Tregs has also been observed after sunitinib treatment of mRCC patients and correlated with outcomes. Indeed, overall survival was significantly better in patients who experienced a 10% decrease in Treg numbers after 2 cycles of sunitinib. Nevertheless, the decrease in the number of Tregs was not related to the change in tumor burden (RECIST criteria) or to progressionfree survival (Adotevi and Tartour, 2010). Thus, immune biological parameters may be interesting markers that could allow a balance to be struck between the efficacy, toxicity, and cost of antiangiogenic drugs. Nevertheless, the clinical relevance of these potential biomarkers must first be validated in large prospective trials.

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9. CONCLUDING REMARKS VEGF is a key regulator of tumor angiogenesis and modulates tumorinduced immunosuppression. VEGF-targeting therapies have proven their immunomodulatory capacities in animal models and cancer patients. Combining antiangiogenic drugs with immunotherapeutic strategies is a promising way of optimizing antitumor responses, as demonstrated in several preclinical models and by early studies in patients with solid cancers. A better understanding of the mechanisms involved in antiangiogenic druginduced immune changes is required to optimize the synergy of combined antiangiogenic and immunotherapeutic strategies. Finally, reliable immune biomarkers that can predict the efficacy and toxicity of antiangiogenic drugs could help to identify those patients who will benefit most from these therapies.

ACKNOWLEDGMENTS This work was supported by Ligue contre le Cancer and Association des Gastroent
erologues Oncologues. A.L.P. was supported by Assistance Publique des H^ opitaux de Paris (Ann
ee Recherche). T.V. received financial support from ITMO Cancer AVIESAN (Alliance Nationale pour les Sciences de la Vie et de la Sant
e, National Alliance for Life Sciences & Health) within the framework of the Cancer Plan. S.P. was supported by Fonds d’Etudes et de Recherche du Corps M
edical des h^ opitaux de Paris (FERCM). Conflict of Interest: J.T. has an advisory role with Roche. No potential conflicts of interest were disclosed by the other authors.

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