The Zebrafish in Biomedical Research: Biology, Husbandry, Diseases, and Research Applications (American College of Laboratory Animal Medicine) 0128124318, 9780128124314

Table of contents :
Cover
American College of Laboratory Animal Medicine Series
The Zebrafish in Biomedical Research
Copyright
Dedication
Contributors
List of reviewers
Preface
Section I: Introduction
1 - History of Zebrafish Research
Setting the Stage
Establishing the Zebrafish Model
Synthesizing Genetics and Embryology
Making a Big Splash
Developing New Tools for Molecular Analyses
Expanding Zebrafish Research Into New Areas
Conclusions
References
2 - Zebrafish Taxonomy and Phylogeny or Taxonomy and Phylogeny
Introduction
The Phylogenetic Position of Zebrafish
Zebrafish and Related Danio Species as an Evolutionary Model System
Cypriniform and Danionid Relationships
The Emerging Danionid Model System
The Zebrafish Model in an Evolutionary Context
Evolutionary Considerations for Zebrafish-to-Human Comparisons
Importance of Genome Duplications for Zebrafish Evolution
Zebrafish and Its Relation to Other Fish Model Species
Medaka
“Evolutionary Mutant” Fish Models
Nonteleost Fish Provide Connectivity From Zebrafish to Human
Conclusion and Outlook
References
3 - Zebrafish Genetics
Introduction
The Zebrafish Karyotype
The Zebrafish Genetic Map
Zebrafish and the Teleost Genome Duplication
Zebrafish Gene Nomenclature Conventions
Chromosome Evolution After the Teleost Genome Duplication
Gene Evolution After the Teleost Genome Duplication
The Zebrafish Genome Sequence Assembly
Genetics of Zebrafish Sex Determination
Conclusions
Acknowledgments
References
4 - Geographic Range and Natural Distribution
Geographic Range
Geographic Range of the Zebrafish
Home Range
Natural Distribution
Floodplains
Rivers
Streams
Natural Still Bodies of Water
Man-Made Habitats
Connectivity Between Habitats
Temporal Distribution
Climate
Seasons
Winter
Summer
Monsoon
Postmonsoon
Regional Microclimates
Altitudes
Temperatures
Water Conditions
Depth
Flow
Turbidity
pH
Salinity
Substrates
Vegetation
Human Impact
Conclusions
References
5 - Behavior of Wild Populations
Introduction
Life History of the Zebrafish
Social Behavior
Group Size
Aggression
Patterns of Aggression in Zebrafish
Dominance Hierarchies
Effects on Lifetime Fitness
Reproductive Behavior
Finding a Mate
Courtship and Spawning
Spawning Strategies
Foraging Behavior
Searching for Food
Prey Selection
Macro and Microhabitats
Antipredatory Behavior
Variations in Behavior
Behavioral Differences Among Individuals
Behavioral Differences Among Populations
Conclusions
References
Section II: Biology
Appearance, development, anatomy, physiology
6 - Zebrafish in Biomedical Research: Head and Body: Anatomy
Introduction
Conclusion
References
7 - Establishing The Body Plan: The First 24 Hours of Zebrafish Development
The Early-stage Embryo
Fertilization and Cleavage
Blastula Stage
Gastrula Stage
Somitogenesis
Early Head Formation
Initial Formation of Differentiated Cell Types: Ectoderm
Initial Formation of Differentiated Cell Types: Mesoderm
Initial Formation of Differentiated Cell Types: Endoderm
References
Tissues and organs associated with special senses
8 - Zebrafish Integumentary System
Introduction
Epidermal Development and Anatomy
Development and Anatomy of the Hypodermis and Collagenous Dermal Stroma
Development and Anatomy of Elasmoid Scales
Zebrafish Skin as a Model for Skin Disease, Wound Healing and Regeneration
Outlook
Acknowledgments
References
9 - Zebrafish Pigmentation
Introduction
Stripes and Their Development
Scales, Fins, and Other Sites of Pigmentation
Physiological and Pathological Effects on Pigmentation
Conclusions
Acknowledgments
References
10 - Respiratory System
Introduction
The Mechanics of Respiration
The Gills
Morphology and Blood Flow
Gill Development
Internal Morphology
Neurobiology
Cutaneous Respiration
Indicators of Stress to the Respiratory System
Hyperventilation
Aquatic Surface Respiration
Conclusion
References
11 - Skeletal System Morphophysiology
The Functions of the Skeleton are Diverse
Bone Develops Through Different Modes of Ossification
Bone Exhibits a Hierarchical Structure
Nanoscale: Matrix and Mineral
Microscale: Cellular Composition and Related Microstructure
Mesoscale and Macroscale: Identifiable Bone Types and Functional Groupings
Summary
Acknowledgments
References
12 - Zebrafish Myology
Embryonic Origins
Head Muscles
Extraocular Muscles
Arch-associated Muscles
Mandibular Arch Muscles
Hyoid Arch Muscles
Posterior Branchial Arch Muscles
Hypobranchial Muscles
Trunk Muscles
Pectoral Fin Muscles
Pelvic Fin Muscles
Median Fin Muscles
Conclusions
References
13 - It Takes Guts: Development of the Embryonic and Juvenile Zebrafish Digestive System
Introduction
Endoderm Formation and Migration to Midline
Organization of the Embryonic Endoderm
Proliferation of the Intestinal Endoderm
Maturation of the Embryonic Intestinal and Esophageal Endoderm
Development of Smooth Muscle and Enteric Neurons
Liver Development
Pancreas Development
Maturation of the Postembryonic Digestive System
Conclusions
Acknowledgments
References
14 - The Zebrafish Cardiovascular System
Introduction
The Cardiovascular System
The Zebrafish as a Cardiovascular Model Organism
Anatomy of the Adult Cardiovascular System
Adult Vascular Anatomy
Adult Heart Anatomy
The Developing Cardiovascular System
Tools and Methods for Visualizing Heart and Vessels in Developing Zebrafish
Microangiography, Dye Fills, Resin Casting
Particle Image Velocimetry (PIV)
Transgenic Zebrafish and Time-lapse Imaging
Heart Development and Regeneration
Blood Vessel Development
Lymphatic Vessel Development
Lymphatic Perivascular Cells
References
15 - Development of The Zebrafish Pronephric and Mesonephric Kidneys
Introduction
Structure of The Pronephros and Mesonephros
Formation of The Pronephros
Formation of The Mesonephros
Concluding Remarks
References
16 - The Reproductive System
Introduction
Primordial Germ Cell Specification and Migration (0–24h Postfertilization, hpf)
The Larval Bipotential/Undifferentiated Gonad (5–25 Dpf)
Sex Determination and Differentiation (20–30 Dpf)
The Adult Gonads
Stages of Oogenesis
Premeiotic Germ Cells
Meiotic Germ Cells
The Testis
Premeiotic Germ Cells (Spermatogonial Cells)
Meiotic Germ Cells (Spermatocytes)
Postmeiotic Germ Cells (Spermatids and Spermatozoa)
Molecular Markers for Specific Spermatogenic Stages (Antibody and in Situ)
Hormonal Regulation of Reproduction
Steroid Hormone Synthesis
Steroid Hormone Receptors
Regulation of Reproduction by the HPG Axis
Summary
References
17 - Endocrine Systems
Introduction
Hypothalamus and Pituitary
Thyroid
Interrenal Cells (Adrenal Equivalent)
Development of Interrenal Cells and Corticosteroid Stress Response
Parathyroid Hormones
Pineal
Development of the Pineal Gland in Zebrafish Embryos and Larvae
Endocrine Pancreas
α-Cells
β-Cells
δ-Cells
Development of the Endocrine Pancreas
Growth
Energy & Metabolism
Conclusion
References
18 - Zebrafish Nervous Systems
The Central Nervous System
Development of the central nervous system
Organization and Function of the Main Brain Regions and Spinal Cord
The Telencephalon and Olfactory Bulb
Diencephalon
Mesencephalon
Metencephalon
Spinal Cord
Adult Neurogenesis and Regeneration
The Peripheral Nervous System
Development and Function of Dorsal Root Ganglia
Development and Function of the Enteric Nervous System
Development and Function of Sympathetic Neurons
Conclusions
References
19 - Immunology
Introduction
Cellular Components of the Immune System and Their Lineage Markers
Hematopoiesis and Immune-Related Tissues
B Lymphocytes
T Lymphocytes
Natural Killer (NK) Cells
Mononuclear Phagocytes
Granulocytes
Molecular Components of the Adaptive Immune System
Immunoglobulin Structure and Genomic Organization
IgHμ
IgHδ
IgHζ
IgL
T Cell Receptor (TCR)
Major Histocompatibility Complex Genes
MHCI “U” Lineage
MHCI “Z” Lineage
MHCI “L” Lineage
MHCII Sequences
Toll-Like Receptors (TLRs)
TLR1 Subfamily
TLR3 Subfamily
TLR4 Subfamily
TLR5 Subfamily
TLR7 Subfamily
TLR11 Subfamily
TLR Signaling
Fish-Specific Families of Innate Immune Receptors
Novel Immune-Type Receptor (NITR) Family
Diverse Immunoglobulin Domain-Containing Protein (DICP) Family
Polymeric Immunoglobulin Receptor-Like (PIGRL) Family
Novel Immunoglobulin-Like Transcript (NILT) Proteins
Leukocyte Immune-Type Receptors (LITRs)
Complement System
Summary
References
20 - Physiology: Hematology and Clinical Chemistry, Gas Exchange, and Regulatory Osmolality
Introduction
Nomenclature
Chapter Contents
Additional Resources
Hematology and Clinical Chemistry
Developmental Hematopoiesis
Adult Hematopoiesis
Normal Values
Leukocytes
Erythrocytes
Thrombocytes
Serum Chemistry
Gas Exchange
Cutaneous Gas Exchange in Zebrafish Embryos and Larvae
Branchial Gas Exchange in Adult Zebrafish
Oxygen Transport
Excretion of Metabolites
Carbon Dioxide
Ammonia
Regulatory Osmolality
Ionocytes and Ion Homeostasis
H+–ATPase–Rich Ionocytes
Na+–K+–ATPase-Rich Ionocytes
NA+–Cl− Cotransporter Expression
K+-Secreting Ionocytes
Ionocyte Function and Differentiation
Control of Sodium Levels
Control of Chloride Levels
Control of Calcium and Phosphate Levels
Acid–Base Regulation
Summary and Conclusion
References
Organ systems: development, anatomy, physiology
21 - Zebrafish in Biomedical Research: the Retina and Vision
Introduction
The Organization and Function of the Zebrafish Retina Organization
Light Detection and Signal Transmission
Regulation and Maintenance of Retinal Neurons
Neuronal Classification in the Zebrafish Retina
Photoreceptors
Diversity and Connectivity of Retinal Circuits
Function
Behavioral Assays
The Optokinetic Response (OKR)
Phototaxis
The Optomotor Response (OMR)
The Adult Escape Response
Physiological Assays
Electroretinogram (ERG)
Single-cell Recordings
Fluorescent Strategies
Genetically Encoded Indicators
Summary
References
22 - The Mechanosensory Lateral Line System
Introduction
Anatomy of the Lateral Line
Hair Cells
Support Cells
Innervating Fibers
Lateral Line Function
Development of the Lateral Line
pLLP Migration
The Roles of Wnt and FGF Signaling During pLLP Migration
Neuromast Maturation
Growth of the Lateral Line
Regenerative Capacity of the Lateral Line
Identity of Hair Cell Progenitors
Genetic Regulation of Regeneration
Conclusion
References
23 - Inner Ear and Hearing
Introduction to the Auditory System
Otic Induction
Macular Development
Otolith Development
Hair Cell Development
VIIIth Nerve Development
Early Development of Hearing
Growth of Sensory Maculae
Hearing Thresholds in Adults
Accessory Auditory Structures
References
Section III: Husbandry
24 - Introduction to Zebrafish Husbandry
References
25 - Aquatics Facility Design Considerations: Incorporating Aquatics into an Animal Facility
Introduction
Species Dependence
Similarities to Rodent Facilities
Unique Features
Program Components
Housing
Procedural Space
Breeding
Behavioral Testing
Support Spaces
Marshalling/Storage
Cagewash
Planning Approaches
Organization
Structural Support
Adjacencies
Separations
Housing System Types
Centralized System
Distributed System
Stand-Alone System
Environmental Parameters
Temperature
Humidity
Lighting
Light Intensity
Light Source
Light Cycle
Housing Room Planning and Design
Sizes
Features
Finishes
Durable
Cleanable
Safe and Maintainable
Repairable
Flooring
Partitions and Walls
Ceilings
Procedure Room Planning and Design
Sizing
Special Features
Procedure Lighting
Snorkel Exhausts
Compressed Air
Dimmable Lighting
Cage/Tank Wash Planning and Design
Equipment types
Flow Considerations
Throughput Considerations
Features
Casework
Doors
Mechanical, Electrical and Plumbing System Design Considerations
HVAC Design
Electrical
Lighting
Plumbing
Special Considerations
Facility Sustainability
Redundancy
Lessons Learned
References
Further Reading
26 - Aquatic Housing
Introduction to Aquatic Housing
Primary Enclosures (Tanks)
Materials Selection
Thermoplastic Polymers
Glass
Acrylic
Polyethylene and Fiberglass
Emerging Materials
Tank Design
Box Tanks
Serial Tanks
Passive Overflow Tanks
Active Overflow Tanks
Specialty Application Tanks
Mating Tanks
Mass Embryo Production Systems
Respirometers and Mazes
Environmental Enrichment
Secondary Enclosure Systems
Static Enclosures
Recirculating Aquaculture Systems
Small to Medium Scale RAS Aquaria
Large Scale RAS Aquaria
Water Loop
Flow-Through Systems
Advanced Applications
Genotyping Racks
Automated Heat Shock Systems
Introduction to Defined Flora
Summary
References
27 - Cleaning and Disinfection of Life Systems
Abstract
Recirculating Aquaculture Systems (RAS) Components Overview
Types of Soils and Dirt Found in and on RAS Equipment
Cleaning Agents and Techniques used in Aquatics Facilities
Cleaning Validation Techniques
Conclusions
Further Reading
28 - Zebrafish Aquatic Systems: Preventative Maintenance and Troubleshooting
Preventative Maintenance
Record Keeping
Redundancy
Monitoring
Prioritization of Maintenance
Fault Tolerances
Monitoring of Specific Devices and Subsystems
Pump Speeds, Water Flow, Water Change Cycles
Biological Filtration Maintenance
Degassing Device
Mechanical Filtration Maintenance
Biological Filtration
Ultraviolet Lamps
System Water Generation
Pump/Motor Monitoring
Automation
Troubleshooting or Diagnosing Nonnormal Events
Human Factor
Pattern Recognition
Flowchart Approach
Case Studies: NIH Examples
References
29 - Water Quality For Zebrafish Culture
Introduction
The Aquatic Environment
Testing, Units, and Measurement
Making Water For Controlled Aquatic Environments
Water Quality For Zebrafish Culture
Temperature
pH
Conductivity/Salinity
Alkalinity
Hardness
Dissolved Gases (O2, CO2, N2)
Nitrogenous Wastes (NH3, NO2−, NO3−)
Ammonia
Nitrite
Nitrate
Biological Filtration
Conclusion
References
Further Reading
30 - Recirculating Aquaculture Systems (RAS) for Zebrafish Culture
Introduction
Required and Optional Components
Racks and Tanks
Pumps and Circulation
Aeration
Mechanical Filtration (Solids Removal)
Rotating Drum Microscreen Filters
Rapid Sand Filters
Floating Bead Filters
Pressurized Bag Filters and Cartridge Filters
Foam Fractionation (Protein Skimming)
Biological Filtration
Moving Bed Bioreactors
Fluidized Sand Filters
Floating Bead Filters
Trickling Media (TM) Filters
Chemical Filtration and Modification of Water Quality
Granular Activated Carbon
Calcium Limestone Reactors
Zeolite Reactors (Ammonia Towers)
Chemical Probes
Dosing Systems
Disinfection
Ultraviolet (UV) Disinfection Units
Ozone (O3)
Additional Apparatus
Temperature Control
Automation Monitoring, Control, and Alarm Systems
Acknowledgments
References
31 - Zebrafish Breeding and Colony Management
Introduction
Spawning and the Sexes
Laboratory Spawning
In Vitro Fertilization
Cryopreservation
Colony Management
References
32 -Zebrafish Larviculture
Introduction
Natural History/Biology of Larvae
Natural Distribution
Natural History and Reproductive Biology
Biological Characteristics and Staging of Zebrafish Larvae
Feeding
Nutrient Requirements
Feeding Behavior/Anatomy
Feed Characteristics
Feed Types
Water Quality
Physical
Chemical
Epigenetic Factors
Parental Condition
Methodology
Basic Approaches
Published Methods
Future Directions
Conclusion
References
33 - Zebrafish Nutrition—Moving Forward
Introduction
Nutrients
Protein and Amino Acids
Lipids
Carbohydrates
Fiber
Minerals
Vitamins
Nonnutritive Components
Energy
Dietary Energy Utilization
Energy Requirements
Feed Additives
Antioxidants
Immunostimulants
Antimicrobial Agents
Natural and Anthropogenic Contaminants
Live Diets
Formulated Diets: Classification and Use
Feed Management
Storage
Ration
Feed Frequency and Timing
Diet-microbiome Interactions
Conclusions
Acknowledgments
References
34 - Anesthesia, Analgesia, and Euthanasia of the Laboratory Zebrafish
Anesthesia
Introduction
Preanesthetic Assessment and Fish Handling
Anesthetic Support and Monitoring
Monitoring Parameters
Opercular Beat Rate
Immobility
Response to Tail Fin Pinch
Response to Sharp Knock/Deep Vibration
Other
Coughing
Color
Aversive Behaviors
Recovery
Personnel Safety
Anesthetic Agents
Characteristics of An Ideal Anesthetic Agent
Tricaine Methanesulfonate (Also Known As MS222, TMS)
Eugenol (Clove Oil, Isoeugenol)
2-Phenoxyethanol (2-PE)
Benzocaine
Metomidate and Etomidate
Lidocaine
Propofol
Gradual Cooling (Hypothermia)
Others: Ketamine, Propoxate, Quinaldine, Isoflurane, Electroanesthesia
Special Anesthetic Considerations
Anesthesia of Embryos
Anesthesia of Larvae
Long-Term Anesthesia
Repeated Anesthesia
Analgesia
Introduction
Assessment of Pain and Discomfort
Analgesic Agents
Lidocaine
Aspirin
Morphine
Euthanasia
Introduction
Methods of Euthanasia
Overdose With Anesthetic Agents
Adult Euthanasia
Larval Euthanasia
Physical Methods of Euthanasia
Euthanasia Guidelines
AVMA Guidelines on The Euthanasia of Animals
National Institutes of Health (NIH) Euthanasia Guidelines
CCAC Guidelines for The Euthanasia of Animals
European Union Directive 2010/63/EU
Conclusion
References
35 - Health Surveillance Programs
Introduction
Development of a Health Surveillance Program
Selection of Agents
Testing Frequency
Health Surveillance Sampling Sources
Live Animals
Sentinel Fish
Colony Fish
Embryos
Environmental Samples
Testing Methods
Health Report
Disease Prevention
References
Further reading
36 - Importation and Quarantine
Introduction
Importation Request and Approval
Receipt
Quarantine Goals
General Quarantine Practices
Quarantine Strategies
Environmental Testing
Embryo Surface Disinfection
Quarantine Facility Design
Facility Workflow
References
37 - Export and Transportation of Zebrafish
Documentation
Health Certificate
Transportation
Materials and Methods for Shipping Cryopreserved Sperm
Materials and Methods for Shipping Adult Zebrafish
Exterior Packaging
Primary and Secondary Containment of Fish
Absorbent Material
Filler
Temperature Control and Monitoring
Labels
Water, Air and Shipping Densities
Methods and Materials for Shipping Zebrafish Embryos
Primary Containment of Embryonic Zebrafish
Water and Stocking Density for Embryonic Zebrafish
Surface Disinfection
References
38 - Regulations, Policies and Guidelines Pertaining to the Use of Zebrafish in Biomedical Research
Introduction
Laws, Policies, and Guidelines (Anderson 2002; Bayne & Anderson, 2015, 2017)
United States Department of Agriculture (USDA)
Department of Health and Human Services (DHHS)
Institute of Laboratory Animal Research (ILAR)
Association of Assessment and Accreditation of Laboratory Animal Care International (AAALACi)
American Veterinary Medical Association (AVMA)
Occupational Safety and Health Administration (OSHA) and Other Responsible Organizations
Institutional Responsibilities for Zebrafish Oversight
Scientists
Veterinary Care and Animal Care Staff
Institutional Official
Institutional Animal Care and Use Committee (IACUC) (Garber, Barbee et al. 2011, OLAW 2015)
Disaster Response Planning
Personnel Qualifications and Training
Specific Challenges for Oversight of Zebrafish Research
Knowledgeable IACUC Members
Performance Based Standards
Transportation
Animal Numbers
Counting of Zebrafish Onto Protocols and Pain Category Assignment
Pain, Distress and Discomfort
Centralization of Zebrafish Facilities
Environmental Enrichment
Primary Housing: Housing Density and Sanitation
Environmental Monitoring
Veterinary Care
Acknowledgment
References
Section IV: Diseases
39 - Water Quality and Idiopathic Diseases of Laboratory Zebrafish
Introduction
Diseases of Water Quality
Ammonia Toxicity
Nitrite Toxicity
Chlorine and Chloramine Toxicity
Heavy Metal Toxicity
Supersaturation and Gas Bubble Disease
Nephrocalcinosis
Idiopathic Diseases
Egg-associated Inflammation
Spinal Deformities
Operculum Malformations
Hepatic Megalocytosis
Pathobiology and Clinical Signs
Cardiac Pathologies
Organ and Tissue Hyperplasia
Fin Lesions/Erosion
References
40 - Important Parasites of Zebrafish in Research Facilities
Microsporidia
Pseudoloma Neurophilia
Pleistophora Hyphessobryconis
Piscinoodinium Pillulare
Ichthyophthirius Multifiliis
Myxidium Streisingeri (Myxozoa)
Pseudocapillaria Tomentosa
Transversotrema Patialense
Metacercariae of Digenea
References
Further Reading
41 - Bacterial and Fungal Diseases of Zebrafish
Introduction
Bacterial Diseases
Mycobacterium Species
Background
Role of Surface Biofilms
Transmission
Clinical Signs and Pathological Changes
Diagnosis
Control and Treatment
Surface Disinfection Eggs
Antibiotic Treatment
Specific Recommendations for Employee Safety
Edwardsiella Ictaluri
Background
Clinical Signs and Pathological Changes
Diagnosis
Control and Treatment
Other Bacterial Infections
Background
Clinical Signs and Pathological Changes
Diagnosis
Control and Treatment
Mycoplasma Species
Background
Clinical Signs and Pathological Changes
Diagnosis
Control and Treatment
Water Molds and Mycotic Diseases
Background
Clinical Signs and Pathological Changes
Diagnosis
Control
References
42 - Viral Diseases
Introduction
Naturally Occurring Oiral Infections
Importance of Natural Viral Infections in Zebrafish to Biomedical Research
Redspotted Grouper Nervous Necrosis Virus
Infectious Spleen and Kidney Necrosis Virus
Zebrafish Picornavirus
Endogenous Viral Elements
Experimental Susceptibility to Viral Infections
The Zebrafish as a Viral Infection Model
Experimental Infection Studies with Fish Viruses
Experimental Infection Studies with Mammalian Viruses
Comments on Detection, Diagnosis, Risk Assessment, and Decision-making
Detection and Diagnosis
Risk Assessment and Decision-making
Restrictions on Zebrafish Movement
Conclusions
References
43 - Nonexperimentally Induced Neoplastic and Proliferative Lesions in Laboratory Zebrafish
Introduction
Seminomas
Description
Pathobiology and Clinical Signs
Diagnosis
Control and Treatment
Intestinal Carcinomas
Description
Diagnosis
Control and Treatment
Ultimobranchial Gland
Description
Pathobiology and Linical Signs
Diagnosis
Control and Treatment
Lymphosarcoma and Other Hematopoietic Tumors
Description
Pathobiology and Clinical Signs
Diagnosis
Control and Treatment
Soft Tissue Srcoma/Peripheral Nerve Sheath Tumors
Pathobiology and Clinical Signs
Diagnosis
Control and Treatment
Optic Pathway Tumor
Description
Pathobiology and Clinical Signs
Diagnosis
Control and Treatment
Melanoma
Description
Pathobiology and Clinical Signs
Diagnosis
Extraspinal Chordoma
Description
Pathobiology and Clinical Signs
Diagnosis
Hepatocellular Tumors
Description
Pathobiology and Clinical Signs
Diagnosis
Control and Treatment
Biliary and Pancreatic Ductal Lesions
Description
Pathobiology and Clinical Signs
Diagnosis
Control and Treatment
Nephroblastoma/Ependymoma
Description
Pathobiology and Clinical Signs
Diagnosis
Hemangioma
Description
Pathobiology and Clinical Signs
Diagnosis
Control and Treatment
Thyroid Tumors
Description
Pathobiology and Clinical Signs
Diagnosis
References
44 - Special Procedures for Zebrafish Diagnostics
Environmental Samples
Clinical examination
Selecting Fish for Diagnostic Evaluation
External Examination and Gross Necropsy
Histopathology
Immunohistochemistry and Diagnostic Antibodies
Bacteriology
Bacteriology With Histopathology
Virology
Molecular Diagnositic Tests
Nonlethal Testing
Treatment
Treatments Delivered in Water
Oral Treatments
References
Further reading
Section V: Scientific Research
45 - Zebrafish as a Model to Understand Vertebrate Development
Introduction to Embryology
Developmental Stages and Conservation
Genes and Development
The Emergence of Zebrafish as a Versatile Vertebrate Model System to Understand Embryology
Zebrafish Research Advances Our Understanding of the Mechanisms Governing Development
Early Development In a “Fish-Shell”
Development of the Central Nervous System
Ectodermal Derivatives
Germ Layer Integration to Build the Craniofacial Skeleton
Segmenting the Trunk for Muscle, Bone and Much More
Development of the Circulatory System
Organogenesis—the Kidney
Organogenesis—Endoderm Derived Organs
Summary
References
46 - Zebrafish as a Model for Revealing the Neuronal Basis of Behavior∗
Introduction
Vertebrate Brains Have Much in Common Across Species
The Challenge of Understanding How Brains Generate Behavior
Behavior
Neurons
Wiring
Models
The Goal
A History of Zebrafish as a Model for Understanding the Generation of Behavior
The Beginning
Mutagenesis and Behavior
The Quantitative Study of Behavior in Larval Zebrafish
The Advent of in vivo Functional Imaging in Zebrafish
Electrophysiology in Zebrafish
Circuit Perturbations
Imaging and Perturbations in Freely Swimming Zebrafish
Advantages and Disadvantages of Zebrafish Compared to Other Model Organisms for the Study of the Neural Basis of Behavior
What is a Model Organism in the Context of Systems Neuroscience?
Nematodes, Flies, and Mice Versus Zebrafish
Major Avenues of Investigation in Zebrafish With a Couple of Case Studies
Short-Latency Escapes
Prey Capture
The Future for Zebrafish in Studies of the Neuronal Basis of Behavior
EM Connectivity on Order
Transynaptic Mapping
The Physiological Holy Grail—Genetically Encoded Voltage Imaging
Direct to Circuits in Zebrafish
Circuits and Behavior From Embryo to Adult
References
47 - Zebrafish as a Model to Understand Human Genetic Diseases
Zebrafish Versus Human: How Similar are They?
An Expanding Toolkit for Generating Zebrafish Models of Human Genetic Disease
Mutagenesis Screens
Morpholino Knockdown
Targeting Induced Local Lesions in Genomes (TILLING)
Gene Editing
Disease Models: What Genes do we Choose to Target?
Acknowledgments
References
48 - Zebrafish as a Model for Investigating Animal–Microbe Interactions
Introduction
The Zebrafish: An Adaptable and Multifaceted Model Organism
Vertebrate Immune System
Natural and Surrogate Microbial Associations
High Fecundity and Small Size
Rapid Ex-utero Development
Optical Transparency and Amenability to Live Imaging
The Zebrafish: Forging and Overturning Paradigms in Animal–Microbe Interaction Research
Immunity and Interhost Dispersal of Intestinal Microbiota
Fighting Mycobacterium Infection is an Inflammatory Balancing Act
Outlook
References
49 - Targeted Editing of Zebrafish Genes to Understand Gene Function and Human Disease Pathology
Background
Why Modify Gene Expression in Zebrafish?
Gain Versus Loss of Gene Function
Forward and Reverse Genetics (Historical and Conceptual Differences in Research Approach)
Transient Methods for Altering Gene Expression
Morpholinos
mRNA Injection and Pharmacological Approaches
Techniques for Genome Editing
Site-specific Nucleases
ZFNs and TALENs
CRISPR
Sequence Integration
Random Insertion of Exogenous DNA
Introduction of Exogenous DNA by Nucleases
Knock-in by HDR and MMEJ Mechanisms
Knock-in by Nonhomologous End Joining
Conclusions
References
50 - Zebrafish as a Platform for Genetic Screening
Introduction and History
Mutagenesis
Recovery
Phenotyping
Genotyping
Validation
Characterization
Concluding Remarks
References
51 - Zebrafish as a Platform for Drug Screening
Introduction
Zebrafish Chemical Screening, a Statistical Overview
Zebrafish Chemical Screens by Category
Metabolic Screens
Behavioral Screens
Selected Tissue-Specific Screens
Heart
Lateral Line
Tissue-Specific Disease Models
Clinical Implications
Limitations and Potential Solutions
Conclusion
References
Index
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American College of Laboratory Animal Medicine Series Steven H. Weisbroth, Ronald E. Flatt, and Alan L. Kraus, eds.: The Biology of the Laboratory Rabbit, 1974 Joseph E. Wagner and Patrick J. Manning, eds.: The Biology of the Guinea Pig, 1976 Edwin J. Andrews, Billy C. Ward, and Norman H. Altman, eds.: Spontaneous Animal Models of Human Disease, Volume 1, 1979; Volume II, 1979 Henry J. Baker, J. Russell Lindsey, and Steven H. Weisbroth, eds.: The Laboratory Rat, Volume I: Biology and Diseases, 1979; Volume II: Research Applications, 1980 Henry L. Foster, J. David Small, and James G. Fox, eds.: The Mouse in Biomedical Research, Volume I: History, Genetics, and Wild Mice, 1981; Volume II: Diseases, 1982; Volume Ill: Normative Biology, Immunology, and Husbandry, 1983; Volume IV: Experimental Biology and Oncology, 1982 James G. Fox, Bennett J. Cohen, and Franklin M. Loew, eds.: Laboratory Animal Medicine, 1984 G. L. Van Hoosier, Jr., and Charles W McPherson, eds.: Laboratory Hamsters, 1987 Patrick J. Manning, Daniel H. Ringler, and Christian E. Newcomer, eds.: The Biology of the Laboratory Rabbit, 2nd Edition, 1994 B. Taylor Bennett, Christian R. Abee, and Roy Henrickson, eds.: Nonhuman Primates in Biomedical Research, Volume I: Biology and Management, 1995; Volume II: Diseases, 1998 Dennis F. Kohn, Sally K. Wixson, William J. White, and G. John Benson, eds.: Anesthesia and Analgesia in Laboratory Animals, 1997 James G. Fox, Lynn C. Anderson, Franklin M. Loew and Fred W. Quimby, eds.: Laboratory Animal Medicine, 2nd Edition, 2002 Mark A. Suckow, Steven H. Weisbroth and Craig L. Franklin, eds.: The Laboratory Rat, 2nd Edition, 2006 James G. Fox, Muriel T. Davisson, Fred W. Quimby, Stephen W. Barthold, Christian E. Newcomer and Abigail L. Smith, eds.: The Mouse in Biomedical Research, 2nd Edition, Volume I: History, Wild Mice, and Genetics, 2007; Volume II: Diseases, 2007; Volume III: Normative Biology, Husbandry, and Models, 2007; Volume IV: Immunology, 2007 David Backer, ed: Flynn’s Parasites of Laboratory Animals, 2007 (Wiley) Richard E. Fish, Marilyn J. Brown, Peggy J. Danneman and Alicia Z. Karas, eds.: Anesthesia and Analgesia in Laboratory Animals, 2nd Edition, 2008 Jack R. Hessler and Noel D.M. Lehner, eds.: Planning and Designing Animal Research Facilities, 2009 Mark A. Suckow, Karla A. Stevens, and Ronald P. Wilson, eds.: The Laboratory Rabbit, Guinea Pig, Hamster and other Rodents, 2011 J. Harkness, P Turner, S VandeWoude, C Wheler, eds: Biology and Medicine of Rabbits and Rodents, 5th Ed, 2012 (Wiley) Christian R. Abee, Keith Mansfield, Suzette Tardif and Timothy Morris, eds.: Nonhuman Primates in Biomedical Research, 2nd Edition, Volume I: Biology and Management, 2012; Volume II: Diseases, 2012 Kathryn Bayne and Patricia V. Turner, eds.: Laboratory Animal Welfare, 2013 James G. Fox and Robert P. Marini eds.: Biology and Diseases of the Ferret, 2E, 2014 (Wiley) M. Michael Swindle, ed.: Swine in the Laboratory: Surgery, Anesthesia, Imaging and Experimental Techniques, 3E, 2015 (Taylor & Francis) James Fox, (Editor-in-Chief), Lynn Anderson, Glen Otto, Kathleen Pritchett-Corning, and Mark Whary, eds.: Laboratory Animal Medicine, 3rd edition, 2015 Trenton Schoeb and Kathryn Eaton, eds.: Gnotobiotics, 2017 Robert Marini, Lynn Wachtman, Suzette Tardif, Keith Mansfield and James Fox, eds.: The Common Marmoset in Captivity and Biomedical Research, 2018 Mark Suckow, F. Claire Hankenson, Ronald Wilson and Patricia Foley, eds.: The Laboratory Rat, 3rd Edition, 2019

THE ZEBRAFISH IN BIOMEDICAL RESEARCH Biology, Husbandry, Diseases, and Research Applications Edited by

Samuel C. Cartner Animal Resources Program, University of Alabama at Birmingham Birmingham, AL, United States of America

Judith S. Eisen Institute of Neuroscience, University of Oregon Eugene, OR, United States of America

Susan C. Farmer University of Alabama at Birmingham, Birmingham, AL, United States of America

Karen J. Guillemin Institute of Molecular Biology, University of Oregon, Eugene, OR, United States of America; Humans and the Microbiome Program, CIFAR, Toronto, ON, Canada

Michael L. Kent Departments of Microbiology and Biomedical Sciences, Oregon State University Corvallis, OR, United States of America

George E. Sanders Department of Comparative Medicine, University of Washington Seattle, WA, United States of America

Academic Press is an imprint of Elsevier 125 London Wall, London EC2Y 5AS, United Kingdom 525 B Street, Suite 1650, San Diego, CA 92101, United States 50 Hampshire Street, 5th Floor, Cambridge, MA 02139, United States The Boulevard, Langford Lane, Kidlington, Oxford OX5 1GB, United Kingdom Copyright © 2020 Elsevier Inc. All rights reserved. No part of this publication may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopying, recording, or any information storage and retrieval system, without permission in writing from the publisher. Details on how to seek permission, further information about the Publisher’s permissions policies and our arrangements with organizations such as the Copyright Clearance Center and the Copyright Licensing Agency, can be found at our website: www.elsevier.com/permissions. This book and the individual contributions contained in it are protected under copyright by the Publisher (other than as may be noted herein). Notices Knowledge and best practice in this field are constantly changing. As new research and experience broaden our understanding, changes in research methods, professional practices, or medical treatment may become necessary. Practitioners and researchers must always rely on their own experience and knowledge in evaluating and using any information, methods, compounds, or experiments described herein. In using such information or methods they should be mindful of their own safety and the safety of others, including parties for whom they have a professional responsibility. To the fullest extent of the law, neither the Publisher nor the authors, contributors, or editors, assume any liability for any injury and/or damage to persons or property as a matter of products liability, negligence or otherwise, or from any use or operation of any methods, products, instructions, or ideas contained in the material herein. Library of Congress Cataloging-in-Publication Data A catalog record for this book is available from the Library of Congress British Library Cataloguing-in-Publication Data A catalogue record for this book is available from the British Library ISBN: 978-0-12-812431-4 For information on all Academic Press publications visit our website at https://www.elsevier.com/books-and-journals

Publisher: Andre G. Wolff Acquisition Editor: Andre G. Wolff Editorial Project Manager: Barbara Makinster Production Project Manager: Poulouse Joseph Cover Designer: Greg Harris Typeset by TNQ Technologies

Dedication

We dedicate this book to our late colleague, George Streisinger, PhD, without whom this volume would be unnecessary. Streisinger introduced the world to zebrafish as a genetic model to study development of the vertebrate nervous system. Because of his untimely death in 1984, Streisinger never knew that he founded an entirely new field of research using zebrafish as a genetic model to investigate every aspect of vertebrate development as well as what goes awry in genetic, infectious, and environmental diseases. Streisinger was born in Budapest, Hungary, in 1927 and came to the United States with his family in the late 1930s to escape Nazi persecution. He had a penchant for biology and was the sole author of several publications in the 1940s, while he was still in high school. Streisinger received a Bachelor of Science degree from Cornell University in 1950, received his doctorate in genetics from the University of Illinois in 1953, and conducted postdoctoral research at the California Institute of Technology until 1956, when he joined Cold Spring Harbor. In 1960, Streisinger took a faculty position in the University of Oregon’s nascent Institute of Molecular Biology, where he spent the remainder of his scientific career. As a postdoctoral fellow, Streisinger was a pioneering molecular biologist who made major contributions to the genetics of T-even bacteriophages. At the University of Oregon, he turned his attention to more complex systems, using the same types of mutational analysis to understand vertebrate development. Streisinger was elected to the United States National Academy of Sciences in 1975, during the early stages of his zebrafish research. His publication in 1981 of the methodology for producing clonal lines of homozygous zebrafish broke new ground and enabled scientists around the world to envision incorporating zebrafish into their research programs. Streisinger was a master at balancing his scientific and other interests. For example, he was an extraordinary culinarian, a highly sought-after goat show judge, and a serious antiwar activist. Despite not living to see it come to fruition, Streisinger set the tone for a field in which researchers are truly collegial, sharing resources and reagents, and working across local and international boundaries to move science forward. We imagine that he would be delighted and also completely amazed at how this field he started has progressed. With this volume, we salute George Streisinger and his legacy. Samuel C. Cartner Judith S. Eisen Susan C. Farmer Karen J. Guillemin Michael L. Kent George E. Sanders

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Contributors

John Aliucci Aquatics Systems Engineering Specialist, Charles River Laboratories, Frederick, MD, United States of America Andrew J. Aman Department of Biology, University of Virginia, Charlottesville, VA, United States of America

Peter Currie Australian Regenerative Medicine Institute, Monash University, Clayton, VIC, Australia Louis R. D’Abramo Department of Biology, University of Alabama at Birmingham, Birmingham, AL, United States of America Alan J. Davidson Department of Molecular Medicine and Pathology, School of Medical Sciences, Faculty of Medical & Health Sciences, The University of Auckland, Auckland, AKL, New Zealand

Michael J.F. Barresi Smith College, Northampton, MA, United States of America Carrie L. Barton Zebrafish Colony Manager, Sinnhuber Aquatic Research Laboratory, Corvallis, OR, United States of America

Cuong Q. Diep Department of Biology, Indiana University of Pennsylvania, Indiana, PA, United States of America

Diana P. Baumann Head, Reptile & Aquatics, Stowers Institute for Medical Research, Kansas City, MO, United States of America

Bruce W. Draper University of California Davis, Department of Molecular and Cellular Biology, One Shields Ave., Davis, CA, United States of America

Ingo Braasch Department of Integrative Biology and Program in Ecology, Evolutionary Biology and Behavior, Michigan State University, East Lansing, MI, United States of America

Earle Durboraw Animal Resources Program, University of Alabama at Birmingham, Birmingham, AL, United States of America

Susan E. Brockerhoff Biochemistry, University of Washington, Seattle, WA, United States of America

Judith S. Eisen Institute of Neuroscience, University of Oregon, Eugene, OR, United States of America

Shawn M. Burgess Translational and Functional Genomics Branch, National Human Genome Research Institute, National Institutes of Health, Bethesda, MD, United States of America

Susan C. Farmer University of Alabama at Birmingham, Birmingham, AL, United States of America

Samuel C. Cartner Animal Resources Program, University of Alabama at Birmingham, Birmingham, AL, United States of America

Kay Fischer Oregon Veterinary Diagnostic Laboratory, Oregon State University, Corvallis, OR, United States of America

Daniel Castranova Eunice Kennedy Shriver National Institute of Child Health and Human Development, National Institutes of Health, Bethesda, MD, United States of America

L. Adele Fowler Department of Biology, University of Alabama at Birmingham, Birmingham, AL, United States of America

Joseph R. Fetcho Dept. of Neurobiology and Behavior, Cornell University, Ithaca, NY, United States of America

Dawnis M. Chow Dept. of Neurobiology and Behavior, Cornell University, Ithaca, NY, United States of America

Marina Venero Galanternik National Institute of Child Health and Human Development, National Institutes of Health, Bethesda, MD, United States of America

Whitney M. Cleghorn Biochemistry, University of Washington, Seattle, WA, United States of America

Julia Ganz Department of Integrative Biology, Michigan State University, East Lansing, MI, United States of America

Jason Cockington Australia New Zealand Association of Aquarium Professionals inc., Brisbane, QLD, Australia

Daniel A. Gorelick Center for Precision Environmental Health, Baylor College of Medicine, Houston, TX, United States of America; Department of Molecular and Cellular Biology, Baylor College of Medicine, Houston, TX, United States of America

Allison B. Coffin Integrative Physiology and Neuroscience, Washington State University Vancouver, WA, United States of America Chereen Collymore Animal Care and Veterinary Services, University of Ottawa, Ottawa, ON, Canada

Karen J. Guillemin Institute of Molecular Biology, University of Oregon, Eugene, OR, United States of America; Humans and the Microbiome Program, CIFAR, Toronto, ON, Canada

James D. Cox Director of Animal and Biological Services, HHMI-Janelia Research Campus, Ashburn, VA, United States of America Marcus J. Crim IDEXX BioAnalytics, Columbia, MO, United States of America

Lauren M. Habenicht Office of Laboratory Animal Resources, Center for Comparative Medicine, University of Colorado Denver | Anschutz Medical Campus, Aurora, CO, United States of America

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xiv

Contributors

Hugh S. Hammer University of Pittsburgh, Pittsburgh, PA, United States of America; The University of Alabama at Birmingham, Birmingham, AL, United States of America

Katrina N. Murray Zebrafish International Resource Center, University of Oregon, Eugene, OR, United States of America

Alexandria M. Hudson Integrative Physiology and Neuroscience, Washington State University Vancouver, WA, United States of America

James T. Nichols Department of Craniofacial Biology, University of Colorado Anschutz Medical Campus, Aurora, CO, United States of America

Michael G. Jonz Department of Biology, University of Ottawa, Ottawa, ON, Canada

Lauren Pandolfo National Institute of Child Health and Development, National Institutes of Health, Bethesda, MD, United States of America

Jan Kaslin Australian Regenerative Medicine Institute, Monash University, Clayton, Melbourne, VIC, Australia Michael L. Kent Departments of Microbiology and Biomedical Sciences, Oregon State University, Corvallis, OR, United States of America David Kimelman Department of Biochemistry, University of Washington, Seattle, WA, United States of America Ronald Y. Kwon Department of Orthopaedics and Sports Medicine, University of Washington, Seattle, WA, United States of America; Institute for Stem Cell and Regenerative Medicine, University of Washington, Seattle, WA, United States of America; Department of Mechanical Engineering, University of Washington, Seattle, WA, United States of America David Lains Zebrafish International Resource Center, University of Oregon, Eugene, OR, United States of America Christian Lawrence Boston Children’s Hospital, Boston, MA, United States of America Johan Ledin Department of Organismal Biology, Science for Life Laboratory, Uppsala University, Uppsala, Sweden

David M. Parichy Department of Biology and Department of Cell Biology, University of Virginia, Charlottesville, VA, United States of America Narendra H. Pathak Smith College, Northampton, MA, United States of America Gregory C. Paull Department of Biosciences, University of Exeter, Exeter, Devon, United Kingdom Randall T. Peterson College of Pharmacy, University of Utah, Salt Lake City, UT, United States of America Jennifer B. Phillips Institute of Neuroscience, University of Oregon, Eugene, OR, United States of America John H. Postlethwait Institute of Neuroscience, University of Oregon, Eugene, OR, United States of America Morgan Prochaska Department of Biology, Clarkson University, Potsdam, NY, United States of America David W. Raible Department of Biological Structure, University of Washington, Seattle, WA, United States of America

Carole J. Lee Department of Biosciences, University of Exeter, Exeter, Devon, United Kingdom

Alberto Rissone Translational and Functional Genomics Branch, National Human Genome Research Institute, National Institutes of Health, Bethesda, MD, United States of America

Jianlong Li Department of Biology, Clarkson University, Potsdam, NY, United States of America

Erik Sanders Aquatics Lab Services LLC, St. Peters, MO, United States of America

Christine Lieggi Center of Comparative Medicine and Pathology, Memorial Sloan Kettering Cancer Center and Weill Cornell Medicine, New York, NY, United States of America

George E. Sanders Department of Comparative Medicine, University of Washington, Seattle, WA, United States of America

Christiana Lo¨hr Oregon Veterinary Diagnostic Laboratory, Oregon State University, Corvallis, OR, United States of America; Department of Biomedical Sciences, Oregon State University, Corvallis, OR, United States of America Kimberly L. McArthur Dept. of Neurobiology and Behavior, Cornell University, Ithaca, NY, United States of America; Dept. of Biology, Southwestern University, Georgetown, TX, United States of America Braedan M. McCluskey Department of Biology, University of Virginia, Charlottesville, VA, United States of America Noriko Mikeasky Department of Biology, Indiana University of Pennsylvania, Indiana, PA, United States of America Donna Mulrooney Oregon Veterinary Diagnostic Laboratory, Oregon State University, Corvallis, OR, United States of America

Justin L. Sanders Department Biomedical Sciences, Oregon State University, Corvallis, OR, United States of America; Oregon Veterinary Diagnostic Laboratory, Oregon State University, Corvallis, OR, United States of America Kellee R. Siegfried University of Massachusetts Boston, Biology Department, 100 Morrissey Blvd., Boston, MA, United States of America Natalie L. Smith Department of Biochemistry, University of Washington, Seattle, WA, United States of America Sean T. Spagnoli Biomedical Sciences, Oregon State University College of Veterinary Medicine, Corvallis, OR, United States of America Amber N. Stratman National Institute of Child Health and Human Development, National Institutes of Health, Bethesda, MD, United States of America

Contributors

xv

Eric D. Thomas Graduate Program in Neuroscience, University of Washington, Seattle, WA, United States of America

Stephen A. Watts Department of Biology, University of Alabama at Birmingham, Birmingham, AL, United States of America

David Traver Department of Cellular and Molecular Medicine, Section of Cell and Developmental Biology, Division of Biological Sciences, University of California, San Diego, CA, United States of America

Brant M. Weinstein National Institute of Child Health and Human Development, National Institutes of Health, Bethesda, MD, United States of America

Frank J. Tulenko Australian Regenerative Medicine Institute, Monash University, Clayton, VIC, Australia Charles R. Tyler Department of Biosciences, University of Exeter, Exeter, Devon, United Kingdom Kenneth N. Wallace Department of Biology, Clarkson University, Potsdam, NY, United States of America Chongmin Wang Eunice Kennedy Shriver National Institute of Child Health and Human Development, National Institutes of Health, Bethesda, MD, United States of America

Monte Westerfield Institute of Neuroscience, University of Oregon, Eugene, OR, United States of America Christopher M. Whipps SUNY-ESF, State University of New York College of Environmental Science and Forestry, Environmental and Forest Biology, Syracuse, NY, United States of America Travis J. Wiles Institute of Molecular Biology, University of Oregon, Eugene, OR, United States of America Michael B. Williams Department of Biology, University of Alabama at Birmingham, Birmingham, AL, United States of America

Claire J. Watson Department of Orthopaedics and Sports Medicine, University of Washington, Seattle, WA, United States of America; Institute for Stem Cell and Regenerative Medicine, University of Washington, Seattle, WA, United States of America

Jeffrey A. Yoder Department of Molecular Biomedical Sciences, College of Veterinary Medicine, North Carolina State University, Raleigh, NC, United States of America

Amanda Watts Institutional Animal Care and Use Committee, University of Alabama at Birmingham, Birmingham, AL, United States of America

Jeffrey R. Zynda Principal & Science Practice Leader, Perkins and Will, Boston, MA, United States of America

Tejia Zhang College of Pharmacy, University of Utah, Salt Lake City, UT, United States of America

List of reviewers Bruce Riley Texas A & M University, College Station, TX, United States of America Carrie Carmichael Stowers Institute for Medical Research, Kansas City, MO, United States of America David Janz University of Saskatchewan, Saskatoon, SK, Canada David McLean Northwestern University, Evanston, IL, United States of America Florence Marlow Icahn School of Medicine at Mount Sinai, New York, NY, United States of America John Postlewait

University of Oregon, Eugene, OR, United States of America

Matthew S. Alexander Children’s of Alabama/University of Alabama, Birmingham, AL, United States of America Monte Westerfield University of Oregon, Eugene, OR, United States of America

xvii

Preface

The American College of Laboratory Animal Medicine (ACLAM) was established in 1957 to support education, training, and research in laboratory animal medicine and to recognize specialists in the veterinary field by providing a route for certification. A mission of the ACLAM is to provide scholarly information to ensure consistent standards for laboratory animal care and improve reproducibility of scientific research. To this end, ACLAM has sponsored a series of reference books on the biology, husbandry, diseases, and use of the most frequently studied animal models of human and animal disease. The first one on the laboratory rabbit was published in 1974. Now with over 15 volumes in the set, they offer state-of-the-art reference materials for biomedical researchers. The Zebrafish in Biomedical Research adds to this series and is the first comprehensive publication on this important animal research model. The contents of this book include 51 chapters divided into five main sections: Introduction, Biology, Husbandry, Diseases, and Scientific Research. The Introduction includes a comprehensive review of the history of the zebrafish in biomedical research, followed by chapters reviewing taxonomy, genetics, natural geographical distribution, and behavior of wild populations. The Biology section includes chapters on the development, anatomy, and physiology of organ systems. The Husbandry section provides extensive coverage of facility design, housing systems and their maintenance, water quality, breeding, colony management, nutrition, and other support activities. There is also a brief review of the regulations, policies, and guidelines pertaining to the use of zebrafish in research in the United States. The section on Diseases provides up-to-date information on the idiopathic, parasitic, bacterial, fungal, and viral diseases of zebrafish. Biomedical researchers and laboratory animal professionals will find the chapter on special procedures for disease diagnosis and treatment very helpful. Scientific Research covers the major uses of the zebrafish in biomedical research as models for developmental, behavioral, and genetic diseases. In addition, there is a chapter reviewing the novel uses of zebrafish in understanding the hostemicrobeepathogen interaction. The final two chapters review the zebrafish as a platform for technology development and toxicology and drug screening. All the contributors are experts in their specialties and the application of the zebrafish model to their research. As with all volumes in the ACLAM series, the authors and editors have donated their time and effort to this publication. Permission fees required for the use of figures, tables, etc., were paid by the University of Alabama at Birmingham. All these efforts were made in the name of fostering continuing education in laboratory animal science and promoting responsible use of animals in biomedical research. A special thanks is extended to reviewers of each chapter and the authors’ and editors’ home institutions for allowing them to work so diligently on this book. The authors and editors appreciate the helpful support from the Elsevier staff. Samuel C. Cartner Judith S. Eisen Susan C. Farmer Karen J. Guillemin Michael L. Kent George E. Sanders

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C H A P T E R

1 History of Zebrafish Research Judith S. Eisen Institute of Neuroscience, University of Oregon, Eugene, OR, United States of America understanding developmental processes that occur deep inside a human fetus’s mother, and thus remain mysterious. These invisible processes could be understood if the development of other animals were discovered to mirror that of humans. Finding that this is the case would allow animals to serve as proxies, models that we can investigate in ways in which we cannot study ourselves, and would enable us to discover how human development normally unfolds and how abnormal development might lead to human birth defects. The hypothesis that animal development could serve as a model for human development was established early, for example by comparisons between human and animal embryos made by the Hippocratic school and by Aristotle (Lonie, 1981; Needham, 1963), and reinforced more recently, for example in the late 1800s by Ernst Haeckel whose drawings of developing animals revealed the similarities of their embryonic body plans and how these morphed into related, but distinct, species-specific, adult forms during development (Fig. 1.2). The idea that studying a variety of animals could reveal the principles underlying development suggests that it is reasonable to choose a model to study based on its experimental tractability for addressing a particular research question. In this regard, teleost fish have been particularly useful for many types of observations (Oppenheimer, 1936; Wourms, 1997; Wourms, Whitt, 1981). For embryology, oviparous teleosts have been particularly advantageous because fertilization, and indeed their entire developmental process, unfolds in the water column, and thus, they do not have to be extracted from the mother for observation or experimentation (Oppenheimer, 1936). In addition, some teleost embryos show “striking lucidity,” providing an exceptional platform for microscopic observations of morphogenetic and cellular processes during the unfolding of living embryos (Trinkaus, 1990).

Although PubMed (pubmed.gov) lists papers studying zebrafish embryos as early as the 1950s (Fig. 1.1), zebrafish research did not hit its stride until nearly a decade after publication of the pioneering work by George Streisinger in 1981 (Streisinger, Walker, Dower, Knauber, & Singer, 1981). Why has this small tropical fish become an important biomedical model and what has made its popularity soar beginning from the mid 1990s?

Setting the Stage Scientists have been fascinated with learning how humans and other animals develop since well before the common era. Hippocrates, the renown Greek physician who lived over 2500 years ago and is considered the father of modern medicine, or at least his followers of the Hippocratic school are so considered, provided an overview of human embryology in several volumes including the “Hippocratic Treatises ‘On Generation’ and ‘On the Nature of the Child,’” and noted the similarity to chick embryology (Lonie, 1981; Needham, 1963). Aristotle, the celebrated Greek scientist and philosopher who lived about 200 years later also carried out extensive observations of animal development based on dissections at various developmental stages, which he published in his books “On the Generation of Animals,” “The History of Animals,” “On Respiration,” and “On the Motion of Animals” (Gilbert, Barresi, 2016; Needham, 1963). Fast forward 1700 years to when another historical luminary, Leonardo da Vinci, one of the foremost artists and inventors during the Italian Renaissance, contributed significantly to our understanding of embryonic development through drawings and quantitative measurements of dissected avian and mammalian embryos, including a human fetus (Needham, 1963). Why the enduring fascination with embryology? Part of it undoubtedly comes from the difficulty of The Zebrafish in Biomedical Research https://doi.org/10.1016/B978-0-12-812431-4.00001-4

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© 2020 Elsevier Inc. All rights reserved.

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1. History of Zebrafish Research

FIGURE 1.1 Since the 1990s there has been significant acceleration in publication of papers concerning zebrafish. Data from PubMed (https:// www.ncbi.nlm.nih.gov/pubmed/?term¼zebrafish).

Interestingly, Aristotle contributed not only to our understanding of embryology of terrestrial vertebrates, but he also appears to have been the first contributor to our recorded knowledge of teleost development, an area to which there was little in the way of additional recorded information until the 1700s (Oppenheimer, 1936; Wourms, 1997). Although he was limited to observations that could be made without magnification, Aristotle was still able to describe aspects of teleost egg development and compare them with the development of avian eggs (Oppenheimer, 1936). From the beginning of the 18th century, a variety of teleost species have been studied to uncover aspects of embryonic development (Wourms, 1997); readers are referred to the excellent reviews cited above to learn more about the breadth of contributions made by these species to understanding embryogenesis. Notable among the fish that have contributed to understanding vertebrate embryonic development is the killifish, Fundulus heteroclitus, commonly known as the mummichog or simply Fundulus (Atz, 1986). This fish played a prominent role in the early days of the now world-renowned Embryology Course at the Marine Biological Laboratories in Wood Hole, Massachusetts (Atz, 1986). In the late 1800s and early 1900s, a number of scientists developed exquisitely detailed descriptions of Fundulus embryology. Their drawings provided a foundation for experimental studies exploring the potential of different parts of the developing embryo to respond to a variety of perturbations, for example, extirpation and grafting. A series of biology luminaries published studies of Fundulus embryos during this time, including Thomas Hunt Morgan, the father of genetics of the fruit fly, Drosophila melanogaster, Jacques Loeb, who helped

pioneer experimental biology (https://embryo.asu. edu/pages/jacques-loeb-1859-1924) as well as comparative brain physiology and psychology (http://www. thecrimson.com/article/1965/2/11/jacques-loebbridging-biology-and-metaphysics/), Philip Armstrong and Julie Swope Child at the Woods Hole Marine Biological Laboratories, Jane Oppenheimer, a pioneering professor at Bryn Mawr, John Paul Trinkaus at Yale, and William Ballard at Dartmouth (Loeb, 1916; Morgan, 1895; Oppenheimer, 1936, 1979; Trinkaus, 1990) (https://embryo.asu.edu/featured/1549). These studies provided important insights into the processes occurring during vertebrate embryogenesis and laid the groundwork for research carried out today. During this same time frame, the rediscovery of Mendel’s genetic experiments ushered in an era of concerted effort to learn the genetic basis of inheritance (Gilbert, 1978). Initially, this work included understanding the relationship between genes and development. However, these disciplinesdembryology and geneticsdbecame largely separate in the early 1900s and remained that way for many decades (Gilbert, 1991, 1998). The study of embryology and genetics began to reunite in the 1970s when a number of researchers started to use genetic approaches in fruit flies to dissect the basis of development. For example, as a step toward his dream of building a bridge between developmental genetics and molecular biology, Ernst Hadorn, developmental genetics pioneer of the University of Zurich, organized a conference on this topic in 1972 (Nothinger, 2002). Another developmental genetics pioneer, Edward Lewis at the California Institute of Technology, investigated how genes controlled the development of specialized organs from individual body segments, work for which he

I. Introduction

5

Establishing the Zebrafish Model

(McGinnis & Krumlauf, 1992) and revealed that the basis for the similarities in embryogenesis among disparate animals is that development is orchestrated by orthologs of the same genes in animals as diverse as fruit flies, chickens, mice, and humans (Beddington & Smith, 1993; Carver & Stubbs, 1997; Davidson, 1994; Gellon & McGinnis, 1998; Manak & Scott, 1994). This realization cemented the idea that animals could be investigated not only to unveil the secrets of their development but also as models to reveal the secrets of human development.

Establishing the Zebrafish Model

FIGURE 1.2 This cover of the August 1989 edition of Trends in Genetics shows the similarity among vertebrate animals in a modified drawing from Ernst Haeckel’s 1879 book “The Evolution of Man.” The top row shows fish, salamander, tortoise, and chicken embryos at an early stage of embryonic development when they are very similar. The middle and bottom rows show the same animals at increasingly later developmental stages; the last stage shows species-specific characteristics. It is important to note that although the picture shown here has been widely reproduced, it remains controversial because not all features of embryos illustrated in the top row are conserved, as described in more detail by Richardson et al. (1997). Cover image of Trends in Genetics volume 5 issue 56 from August 1989 used with permission from Elsevier.

was awarded a Nobel Prize in Physiology or Medicine in 1995 (https://www.nobelprize.org/nobel_prizes/ medicine/laureates/1995/press.html). Christiane Nusslein-Volhard and Eric Wieschaus conducted a genetic screen in fruit flies to learn the mechanisms underlying body patterning (Nusslein-Volhard & Wieschaus, 1980) and shared the 1995 Nobel Prize with Lewis. Subsequent experiments in the late 1980s and early 1990s uncovered the molecular nature of the mutations isolated in the Nusslein-Volhard/Wieschaus screen

It is against this backdrop that George Streisinger, a professor in the nascent Institute of Molecular Biology at the University of Oregon, chose zebrafish, Danio rerio, a small, tropical, fresh-water fish, as a model in which to explore the genetic basis of vertebrate neural development. Streisinger was a pioneering molecular biologist who contributed significantly to the dawn of the modern molecular era through his historic work on bacteriophage in the 1950s. Among his major contributions were an understanding of the DNA triplet code, including deciphering the first in vivo codon assignments, as well as uncovering the structure of the bacteriophage genome (Grunwald & Eisen, 2002; Stahl, 1995). One of Streisinger’s close University of Oregon colleagues, and an avid supporter of his zebrafish endeavors, was Franklin Stahl, another molecular biology pioneer who discovered that DNA replication is semiconservative (Streisinger, 2004). By the mid-1960s, many of the initial questions of molecular biology were nearing resolution and attention was turning toward using genetic approaches to understand the mechanisms underlying more complex systems, such as the “almost mystical” questions of nervous system function and the basis of behavior and cognition (Grunwald & Eisen, 2002). In parallel with Streisinger’s choice to use zebrafish to dissect nervous system function and development, two other molecular biology pioneers established other genetic models to investigate development (Grunwald & Eisen, 2002). Sidney Brenner, codiscoverer of messenger RNA, who also showed that the amino acid sequence of a protein is determined by the RNA nucleotide sequence from which it is translated, developed the roundworm, Caenorhabditis elegans as a new genetic model, work for which he was awarded a Nobel Prize in Physiology or Medicine in 2002 (http://www. nobelprize.org/nobel_prizes/medicine/laureates/ 2002/). Seymour Benzer, who used bacteriophage to reveal the structure of genes and how they are aligned within the genome, began studying nervous system function and behavior using fruit flies.

I. Introduction

6

1. History of Zebrafish Research

FIGURE 1.3 This cover of the July 1995 issue of Developmental Dynamics shows a portion of the zebrafish staging series, a crucial advance for standardizing conditions between different experiments and laboratories. A fertilized egg is shown at the top, and development progresses clockwise. All of the fish were imaged live (Kimmel, Ballard, Kimmel, Ullmann, & Schilling, 1995). The fish in the middle is 3 days postfertilization (3 dpf) (an enlarged view of a fish of this stage is also shown separately, below the cover image). By three dpf, zebrafish have hatched and are referred to as larvae, whereas, at earlier stages, before hatching, they are called embryos. The head of the three dpf larva in the cover image is at the top; the head of the larva below the cover image is to the left. The dark region of the head is the eye. The heart appears red because it contains red blood cells. The large structure under the heart is the yolk, which will nourish the larva until it is able to eat, starting about 1 day later than when this photograph was taken. The embryo at the top of the cover image was just fertilized and is shown within its chorion or eggshell. The chorions were removed prior to taking the other photographs. The next embryo, going clockwise, is at the eight-cell stage. All 8 cells sit on top of the large yolk cell, although only four of them are visible; four of them are hidden behind the visible cells. The next embryo is at the 256-cell stage, followed by an embryo at the 50% epiboly stage. During epiboly, the cells move from sitting on top of the yolk to completely surrounding it. The next embryo, at five o’clock in the cover image, is at the bud stage; “bud” refers to the tailbud, the beginning of the tail, which is pointed toward the bottom of the photograph. The embryo at six o’clock in this cover image is at the four-somite stage; somites are the precursors of the segmented body muscles that fish use for swimming. The embryo at

Why did Streisinger choose zebrafish? Research on Fundulus had already provided many insights into vertebrate development, so Fundulus could have been an obvious choice, despite lacking any previous history of genetic research. There were, however, several other fish species with considerable histories of genetic research, including medaka (Shima & Mitani, 2004), goldfish (Fu, 2016) and Xiphophorus (Kazianis, 2006). Streisinger initially brought a number of tropical fish species, including medaka, into his laboratory; however, no written records describing the basis of his choice have been uncovered (Grunwald & Eisen, 2002). Although it is only speculation, perhaps the difficulty of removing Fundulus embryos from the protective eggshell, or chorion (Atz, 1986; Trinkaus, 1990), or the inclusion within Fundulus and medaka embryos of oil droplets that obscure visibility (Wourms, 1997), contributed to Streisinger’s ultimate choice of zebrafish. During the late 1950s and early 1960s, several other laboratories also began investigating zebrafish embryology (Hisaoka, 1958), including muscle and nervous system development (van Raamsdonk, Mos, Smit-Onel, van de Laarse, & Fehres, 1983; van Raamsdonk, Tekronnie, Pool, & van de Laarse, 1980; Weis, 1968a, 1968b), as well as using zebrafish in the aquaculture trade to study toxicology (Laale, 1977). However, none of these studies incorporated the type of genetic analysis that was central to Streisinger’s approach to obtaining a mechanistic understanding, and whether any of this work contributed to Streisinger’s choice of zebrafish is unknown. Whatever the reasons for Streisinger’s choice of zebrafish, it is now clear that zebrafish is an excellent research model, based on several characteristics of its development. First, in contrast to many fish, zebrafish are not seasonal breeders; thus, embryos can be obtained for study throughout the year. Second, like most fish, fertilization, and all subsequent developmental processes occur within the water column, not inside of the mother; thus, zebrafish are amenable to both observation and manipulation throughout the developmental period. Third, a single female zebrafish can produce hundreds of eggs in a single spawning, making it possible to have statistically significant numbers of closely related embryos for even a single experiment. Fourth, zebrafish embryos and early larvae are nearly transparent, thus using appropriate microscopic methods, it is possible to image essentially every cell during development (Fig. 1.3). Fifth, zebrafish embryos develop rapidly, hatching as predatory larvae in 3 days

=

seven o’clock is at the fifteen-somite stage, and the embryo at nine o’clock is at the twenty five-somite stage; this embryo is nearly 22 h postfertilization. The embryo at 11 o’clock is about 28 h postfertilization. Cover image of Developmental Dynamics volume 203 number three from July 1995 used with permission from Wiley.

I. Introduction

Synthesizing Genetics and Embryology

(Fig. 1.3), and thus development of individual cells or structures deep inside a zebrafish embryo can be observed in real time, or by time-lapse videography, as they occur. Streisinger’s focus on using genetics to uncover developmental processes necessitated establishing tools appropriate for this purpose. Vertebrate genetics is cumbersome because, after mutagenesis, heterozygous carriers need to be crossed to wild types to establish a stock that will contain both wild types and heterozygotes, and then individuals within that stock need to be crossed to one another to identify those that carry mutations of interest. This identification is accomplished

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by screening the offspring of the mated heterozygotes, only one-quarter of which will show a phenotype when a mutation is recessive. Streisinger reasoned that he could facilitate the production of homozygous offspring by circumventing the need to breed heterozygous male and female partners (Grunwald & Eisen, 2002). He achieved this goal by adapting methods to activate eggs without fertilization, as well as by methods to produce gynogenotes, homozygous diploid embryos whose entire genetic contribution derives from the mother. These gynogenotes were useful not only for genetic screens, but Cyrus Levinthal at Columbia University obtained some gynogenotes from Streisinger (Charline Walker, personal communication) to investigate axon trajectories of isogenic neurons (http:// www.nasonline.org/publications/biographicalmemoirs/memoir-pdfs/levinthal-cyrus.pdf). Streisinger’s seminal work describing the cloning of zebrafish was published in 1981 in Nature (Streisinger et al., 1981) and heralded on the cover (Fig. 1.4), just 7 months after publication in the same journal of the seminal work on fruit flies (Nusslein-Volhard & Wieschaus, 1980) for which Nusslein-Volhard and Wieschaus were awarded the 1995 Nobel Prize in Physiology or Medicine that they shared with Edward Lewis. Streisinger’s article received national attention, including an editorial cartoon published in the Chicago Tribune (Fig. 1.5).

Synthesizing Genetics and Embryology Streisinger’s ability to develop zebrafish as a model was fostered by the atmosphere of the University of Oregon Institute of Molecular Biology. Founded in 1959, the Institute nurtured a handful of the brightest young molecular biologists, in part by pooling resources for equipment, facilities, and administrative support to help

FIGURE 1.4

This issue of Nature from MayeJune 1981 published the groundbreaking paper from George Streisinger showing that zebrafish are amenable to genetic analysis (Grunwald & Eisen, 2002; Streisinger et al., 1981). This cover illustrates zebrafish embryos, enclosed within their chorions. Some of the embryos in this picture have normal, black pigment cells in their skin and eyes, but some lack black pigment cells in their skin and eyes, and thus, appear golden in color. The difference in these embryos is the result of a mutation in a single gene, which is called golden, after the appearance of the mutant embryos. The discoverers of the molecular nature of the zebrafish golden gene showed that the human ortholog is responsible for 25%e40% of the difference in skin color between Europeans and West Africans (Lamason et al., 2005). Cover image of Nature volume 291 issue 5813 from MayeJune 1981 used with permission from Springer Nature.

FIGURE 1.5 This cartoon was published in the Chicago Tribune as a response to George Streisinger’s seminal paper describing the cloning of zebrafish (Streisinger et al., 1981). Wayne Stakyskal cartoon from 1981 used with permission from the Chicago Tribune.

I. Introduction

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1. History of Zebrafish Research

underwrite long-term projects, including Streisinger’s zebrafish research program, which took over 10 years to come to fruition (Grunwald & Eisen, 2002; Streisinger, 2004). Oregon also provided an extremely collegial atmosphere and even before zebrafish was ready for prime time as a genetic model, Charles Kimmel, one of Streisinger’s Oregon colleagues who made important early contributions to understanding clonal selection in the immune system (Tauber & Podolsky, 2000), realized the potential of the zebrafish embryo for understanding the cellular basis of neural development and initiated this as a new direction in his research program. Kimmel started his work in this area by describing the development of an identified neuron, the Mauthner cell (Kimmel, Sessions, & Kimmel, 1981), made famous by work in other teleosts and in amphibians (Korn & Faber, 2005). Kimmel’s work revealed that morphologically similar neurons were repeated in each hindbrain segment (Metcalfe, Mendelson, & Kimmel, 1986), leading him to ask whether these neurons could be related by cell lineage (Grunwald & Eisen, 2002). The optical clarity and rapid development of the zebrafish embryo make it ideally suited to carry out lineage studies. So in the summer of 1982 Kimmel visited Michael Bennett’s laboratory at the Marine Biological Laboratory (Kimmel, personal communication) to learn how to inject individual cells, called blastomeres, in young Fundulus embryos, using a fluorescent dye called Lucifer Yellow, developed to the study passage of molecules between cells via gap junctions (Kimmel, Spray, & Bennett, 1984; Stewart, 1981). Kimmel was later joined in his zebrafish lineage tracing endeavors, by Oregon colleagues Monte Westerfield and Judith Eisen, both of whom had backgrounds using Lucifer Yellow to label and record from neurons, and who worked with Kimmel to establish the use of another fluorescent dye, fluorescein dextran, that did not pass through gap junctions. Kimmel’s lineage studies, many carried out with the partnership of undergraduate students, were completely captivating. No one knew the time course of development, and with the instrumentation available at the time, it was not possible to make the kind of computer-controlled, time-lapse movies that are routine today. Even if it were possible, no forum existed at that time in which such movies could be published. So days were divided up into manageable segments of four or more hours, and teams of people sat during each segment in a completely dark room, except for instrument lights and what would be by today’s standards a miniscule video monitor, and watched and took notes, as cell movements and divisions were recorded onto a videocassette recorder, and in many cases traced with different colored markers onto sheets of acetate taped to the video monitor screen. These studies provided

unprecedented insights into vertebrate development and enabled Kimmel to describe lineage contributions to essentially all of the major cell types in the embryo (Kimmel & Warga, 1988). During some of these sessions, Eisen and Westerfield watched as the earliestdeveloping spinal motoneurons extended axonsd something never seen before in vertebrate embryosd and were able to classify these neurons into distinct, identifiable subtypes that could later be studied using a genetic approach (Beattie, Raible, Henion, & Eisen, 1999; Eisen, Myers, & Westerfield, 1986). This was a heady time for those involved. The clarity of the zebrafish embryo was stunning and the application of fluorescent dye technology, previously used only for studies of gap junctional coupling in amphibians and Fundulus (Kimmel et al., 1984; Spray, Harris, & Bennett, 1979), lineage tracing and neuronal morphology in the medicinal leech (Stewart, 1981; Weisblat, Zackson, Blair, & Young, 1980), and laser-ablation in crustaceans (Marder & Eisen, 1984; Selverston & Miller, 1980), meant that researchers learned something new from every gorgeous experiment. And every experiment revealed phenotypes that could be investigated by combining these modern embryological and neurobiological approaches with Streisinger’s newly developed genetic tools. Casting a pall over the nascent field of zebrafish developmental genetics, Streisinger died completely unexpectedly in August 1984, and thus, he never saw the blossoming of this new and exciting era. Streisinger’s death could have been the end of the new beginning for zebrafish research. However, his Oregon colleagues, already deeply immersed in their studies of zebrafish lineage and neural development, took up the baton and began promoting zebrafish as an outstanding new model in which to investigate the genetic basis of developmental processes. A critical aspect of support for zebrafish research came from the National Institutes of Health, which allowed Kimmel to become Principal Investigator of Streisinger’s grant to study zebrafish genetics. David Grunwald, at that time a postdoctoral fellow in Streisinger’s laboratory, and Charline Walker, the research assistant who had worked with Streisinger to develop his genetic tools, were instrumental in helping keep up the momentum of zebrafish research. The Oregon group’s studies had already provided important new insights into the commonality of the developmental processes among vertebrates, leading Kimmel to publish an influential article, in the issue of Trends in Genetics shown in Fig. 1.2, declaring “the fish is a frog . ..is a chicken . ..is a mouse” that can be studied “for both detailed . ..embryogenesis and . ..genetic analysis” (Grunwald & Eisen, 2002; Kimmel, 1989). By this time, a number of researchers prominent for their studies of fruit flies, including Jose Campos-

I. Introduction

Synthesizing Genetics and Embryology

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FIGURE 1.6 This issue of Development from December 1996 was devoted entirely to papers describing the first large scale genetic screens for zebrafish mutations carried out in the Nusslein-Volhard laboratory at the Max-Planck-Institute fur Entwicklungsbiologie in Tubingen Germany and the Driever and Fishman laboratories at Harvard in Boston Massachusetts. There were 37 papers describing the characterization of about 4000 mutations (Eisen, 1996; Grunwald & Eisen, 2002). Cover image of Development volume 123 from December 1996 used with permission from Development and Robert Kelsh.

Ortega at the University of Cologne, as well as NussleinVolhard, had taken note of the work being done in Oregon and had begun to consider adding zebrafish studies to their repertoire (Grunwald & Eisen, 2002). In 1990, the Oregon group facilitated this possibility by hosting a small meeting that brought together researchers from a variety of fields and highlighted the utility of zebrafish as a model. They also established an informal course on zebrafish husbandry, genetics, and embryology, and hosted numerous visiting scientists from around the world who came to learn this new model system (Grunwald & Eisen, 2002). Westerfield created the “Zebrafish Book” (http://zfin.org/zf_info/zfbook/

zfbk.html), a primer on zebrafish methodology, and later established the Zebrafish Information Network (zfin; http://zfin.org/), which hosts the Zebrafish Model Organism Database. Westerfield also founded ZIRC, the Zebrafish International Resource Center, the first zebrafish genetic repository. In 1994, Cold Spring Harbor hosted the first open international meeting on zebrafish development and genetics, and in 1998, a course on zebrafish development and genetics was initiated at the Marine Biological Laboratory in Woods Hole, Massachusetts. In the ensuing years, there has been a proliferation of zebrafish meetings, as well as courses held in a variety of venues around the world.

I. Introduction

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1. History of Zebrafish Research

Making a Big Splash Although the Oregon group continued to use Streisinger’s methodology to conduct mutational screens, uncovering genes involved in a variety of different developmental processes, Nusslein-Volhard developed a more ambitious plan, to essentially recapitulate in zebrafish the screen she had carried out to discover embryonic patterning mutants in fruit flies (Mullins, Hammerschmidt, Haffter, & Nusslein-Volhard, 1994; Mullins & Nusslein-Volhard, 1993). As she was developing new mutagenesis and fish-rearing procedures for this endeavor in her laboratory at the Max Planck Institute in Tubingen, Germany, her talented former graduate student, Wolfgang Driever, who had spent a year as a postdoctoral fellow with Westerfield in Oregon, was recruited to Massachusetts General Hospital by Marc Fishman, to establish a parallel screening effort. In similar fashion to the earlier fruit fly mutational screen, the “Big Screen” in zebrafish revealed mutations that affected nearly every aspect of the embryonic body plan and provided an exceptional window into the genetic processes governing vertebrate development. This was the largest forward mutational screen ever undertaken in a vertebrate and a tour de force by the Nusslein-Volhard and Driever laboratories. The screen involved 65 people, including Nusslein-Volhard’s colleague Friedrich Bonhoeffer and Driever’s colleague Fishman, as well as many brilliant young students and postdoctoral fellows who went on to establish their own laboratories. Participants examined more than a million and a half embryos over about a 2-year period, resulting in the isolation of over 4000 mutations, about half of which were characterized over the next year (Driever et al., 1996; Eisen, 1996; Haffter et al., 1996). Rather than publish these characterizations piecemeal, as a series of papers describing mutations affecting specific aspects of embryonic development, the entire set of 37 screen papers was published as a cohesive collection that constituted an entire, ectopic volume of the journal Development (December 1996, volume 123; Fig. 1.6), complete with a flipbook that revealed the process of embryogenesis as viewed by time-lapse microscopy (Karlstrom & Kane, 1996). This was a historic decision because it facilitated the assessment of the entire range of mutant phenotypes. The brilliance of both the earlier fruit fly screen and also the zebrafish “Big Screen” was categorizing mutants into phenotypic groups. This organization allowed gene functions to be ascribed to temporal and spatial pathways even before the genes themselves had been identified, and greatly facilitated elucidation of the molecular pathways underlying developmental processes. The “Big Screen” catapulted zebrafish to the

forefront as an outstanding new model in which to investigate the genetic underpinnings of essentially every aspect of vertebrate development. Importantly, these mutants were made available so that other laboratories could utilize zebrafish to begin or enhance investigations focused on particular developmental processes.

Developing New Tools for Molecular Analyses Understanding the mechanisms underlying mutant phenotypes revealed by the “Big Screen” as well as other, smaller screens (for example (Beattie et al., 1999)), required learning which genes were affected and the molecular nature of the defects. Several years earlier, John Postlethwait had anticipated the need for molecular landmarks spread out across the genome and created the first genetic linkage map for zebrafish (Johnson, Midson, Ballinger, & Postlethwait, 1994). Consistent with the Oregon way of carrying out science, much of the mapping was done by a team of exceptional undergraduate students. The techniques used to establish the map (Postlethwait & Talbot, 1997) made it relatively straightforward to begin to clone the genes defined by mutant phenotypes, and thus, to understand not only what had gone awry, but also to learn how these mutated genes and resulting phenotypes corresponded to the genotypes and phenotypes of other organisms. In addition, these studies revealed that the zebrafish lineage had experienced a genome duplication event and that this event was shared by all teleosts, an understanding that allowed accurate connection of the zebrafish genome to the human genome (Amores et al., 1998; Force et al., 1999). Perhaps not surprisingly by this time, the cloning of zebrafish mutations again reinforced the idea that similarities in embryogenesis among different animals are governed by orthologs of the same genes, opening the path to use zebrafish to investigate genome architecture and evolution (Postlethwait et al., 1998), as well as to serve as a model for human genetic diseases (Zon, 1999). The momentum of zebrafish research was already accelerating, and it was clear that additional resources would enable many more laboratories to embrace this model. The nascent zebrafish research community, represented by John Postlethwait and Marc Fishman, as well as Len Zon from Harvard Medical School and Nancy Hopkins from the Massachusetts Institute of Technology, worked with the then National Institutes of Health Director, Harold Varmus, and in 1998, the NIH established the “Trans-NIH Zebrafish Initiative” in which many participating NIH institutes solicited applications “to increase our support of the zebrafish as an animal model for development, organ formation,

I. Introduction

Expanding Zebrafish Research Into New Areas

behavior, aging, and disease research” (https://grants. nih.gov/grants/guide/pa-files/PA-01-095.html). It was now evident that zebrafish had arrived as a model! As more laboratories adopted the zebrafish model, new resources were developed, and following in the tradition of establishing an interactive research community, these resources were readily shared. For example, at one of the early Cold Spring Harbor Zebrafish Development and Genetics Meetings, researchers from Harvard brought and shared reagents, and later Chi-Bin Chien from the University of Utah also shared reagents at a Strategic Conference of Zebrafish Researchers. In a similar fashion, Steve Ekker at the Mayo Clinic developed and shared the antisense methods for knocking out the function of nearly any gene (Nasevicius & Ekker, 2000), and more recently a number of laboratories have developed and shared gene-editing strategies (Varshney, Sood, & Burgess, 2015), as well as new strategies for cloning existing mutations (Hill et al., 2013; Miller, Obholzer, Shah, Megason, & Moens, 2013).

Expanding Zebrafish Research Into New Areas Zebrafish has continued to increase in popularity as a model to investigate the cellular, molecular, and genetic mechanisms underlying developmental processes (see Chapter 45 by Pathak and Barresi). As of 2019, there were over 1300 laboratories worldwide listed in ZFIN (http://zfin.org/) that utilize zebrafish to investigate many aspects of animal biology, including human disease mechanisms. Below I describe some of the new directions for zebrafish research that extend beyond understanding the types of developmental mechanisms illustrated above. Learning how the nervous system orchestrates behavior remains a significant challenge that is being addressed by the international “Brain Initiative” (https://www.braininitiative.nih.gov/). Zebrafish are ideally suited for investigating the basic biological mechanisms underlying neural function because of their accessibility for behavioral measurements, in concert with genetic and neural activity manipulation and imaging studies (see Chapter 46 by Mcarthur and colleagues). Zebrafish exhibit a variety of complex behaviors, including, among many others, sleep and social interactions (Orger & de Polavieja, 2017; Stednitz et al., 2018), that should provide new insights into the underlying mechanisms that can then be translated into understanding brain mechanisms and behavior in humans under normal conditions and in disease states. Understanding the molecular mechanisms of human genetic disorders is a prerequisite for developing new tools to diagnose and treat these diseases. The

11

outstanding experimental attributes of zebrafish are being increasingly leveraged to establish functional models of human genetic diseases and to develop new clinical tools for their diagnosis and treatment (Phillips & Westerfield, 2014) (see Chapter 47 by Phillips and Westerfield and Chapter 49 by Rissone and colleagues). The list of human disease models to which zebrafish is making important contributions includes cancer, tuberculosis, neurodevelopmental, neurodegenerative and psychiatric disorders, kidney diseases, cardiovascular diseases, skeletomuscular diseases, gastrointestinal tract dysfunctions; this list continues to expand at a dizzying pace, as exemplified by many of the chapters within this volume. Discovering new drugs is critical for developing therapeutic approaches to human diseases. Zebrafish has become an excellent model for drug discovery because it is straightforward to use chemical screens, in which zebrafish, typically embryos or larvae, are exposed to libraries containing many different molecules with pharmaceutical potential, to identify novel therapeutic agents (Peterson, Link, Dowling, & Schreiber, 2000) (see Chapter 51 by Zhang and Peterson). In fact, numerous chemical screens have been carried out and are already providing important insights that have implications for understanding human health (Rennekamp & Peterson, 2015). This effort has been augmented by the recent realization that human cancer cells can be implanted into zebrafish, thus serving as “avatars” that present highly sensitive platforms for developing and testing pharmacological therapies matched to specific tumor behaviors (Fior et al., 2017; Leslie, 2017). Another exciting new direction for zebrafish research is as a model in which to study host-microbe interactions (see Chapter 48 by Wiles and Guillemin). In many contexts, microbes are thought of as infectious agents that cause disease. The optical transparency and genetic tractability of zebrafish have enabled it to become an outstanding model for two infectious human diseases, tuberculosis (Ramakrishnan, 2013) and leprosy (Madigan, Cameron, & Ramakrishnan, 2017), that have historically been difficult to investigate in mammalian models. These new studies are providing surprising insights that are likely to lead to new therapeutic approaches. A related aspect of understanding host-microbe relationships is to learn about host interactions with the microbial organisms associated with essentially every plant and animal on the planet. A tenet of animal development is that initial patterning of the body depends on a maternal contribution of molecules packaged into the egg, and then unfolds based on the regulation of zygotic genes and their products (Chan et al., 2009; Li, Lu, & Dean, 2013; Schier, 2007). It is increasingly clear that after an embryo leaves the protective environment either inside its mother or inside its eggshell, it

I. Introduction

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1. History of Zebrafish Research

becomes associated with communities of microbes, often referred to as microbiota that influence many aspects of its subsequent development and physiology. However, the extent to which host-associated microbiota modulate the normal development and/or physiology of any animal species remains unknown. Zebrafish is ideally suited as a model in this burgeoning new field, because it is straightforward to rear zebrafish germ-free, in the absence of any microbes, or gnotobiotically, with exposure to only very specific microbes (Melancon et al., 2017). At the same time, both host and microbial genetics can be manipulated, and the exquisite optical properties of zebrafish larvae enable real-time visualization of microbe biogeography and reciprocal responses of both the host and associated microbes, including normal residents (Wiles et al., 2016), as well as invasive pathogens (Ramakrishnan, 2013). In addition to the many features conserved with other animals, zebrafish are able to accomplish something that mammals, such as mice and humans, are unable to achieve, which is to regenerate nearly any damaged body part. Thus far, zebrafish have been shown to regenerate damaged limbs, hearts, retinas, brains, and spinal cords (Gemberling, Bailey, Hyde, & Poss, 2013). This feat is not unique to zebrafish, because other nonmammalian vertebrates, such as salamanders and axolotls, can also regenerate limbs (Dall’Agnese & Puri, 2016). However, the ease of studying zebrafish and manipulating its genome has established it as an important model that holds out the promise of discovering mechanisms that might be exploited to aid regeneration in humans (Mokalled & Poss, 2018). As zebrafish research has expanded, there have been a number of efforts to standardize zebrafish husbandry. Beginning in the early days of zebrafish research, several resources were developed, including “The Zebrafish Book,” dedicated to George Streisinger. This book was originally published by Monte Westerfield in 1993 and printed by the University of Oregon Press. This book is now in its fifth edition (Westerfield, 2007); the fourth edition is available online from ZFIN (zfin. org). Since that time, there have been additional books detailing zebrafish husbandry methods for research laboratories (Harper & Lawrence, 2011; NussleinVolhard & Dahm, 2002). The importance of standardizing husbandry was made very clear recently when the journal Zebrafish devoted an entire issue to health and husbandry (http://online.liebertpub.com/toc/ zeb/13/S1). Zebrafish also regularly publishes a section on this topic. The many chapters on health and husbandry in this volume complement the chapters on zebrafish research and provide state-of-the-art information for those who rear zebrafish that will enhance the utility of zebrafish as an outstanding research model.

Conclusions Streisinger’s groundbreaking paper describing genetic tools for zebrafish marked a turning point for developmental biology. Since that time, this small fish has become established as a model for uncovering genetic mechanisms underlying many aspects of development, differentiation, physiology, regeneration, and evolution. Zebrafish has also become a model in which to investigate mechanisms by which environmental factors, such as animal-associated microbes, can influence developmental and physiological processes by modulating host genes. Research utilizing the zebrafish model continues to accelerate, making the future bright for even more important discoveries that will expand our understanding of the genetic mechanisms that transform a single cell, the fertilized egg, into a multicellular animal with a backbone, as well as how vertebrate organs function, how these mechanisms contribute to human health, and how they may result in human diseases when they go awry.

References Amores, A., Force, A., Yan, Y. L., Joly, L., Amemiya, C., Fritz, A., et al. (1998). Zebrafish hox clusters and vertebrate genome evolution. Science, 282(5394), 1711e1714. Atz, J. W. (1986). Fundulus-heteroclitus in the laboratoryea history. American Zoologist, 26(1), 111e120. Beattie, C. E., Raible, D. W., Henion, P. D., & Eisen, J. S. (1999). Early pressure screens. Methods in Cell Biology, 60, 71e86. Beddington, R. S., & Smith, J. C. (1993). Control of vertebrate gastrulation: Inducing signals and responding genes. Current Opinion in Genetics and Development, 3(4), 655e661. Carver, E. A., & Stubbs, L. (1997). Zooming in on the human-mouse comparative map: Genome conservation re-examined on a highresolution scale. Genome Research, 7(12), 1123e1137. Chan, T. M., Longabaugh, W., Bolouri, H., Chen, H. L., Tseng, W. F., Chao, C. H., et al. (2009). Developmental gene regulatory networks in the zebrafish embryo. Biochimica et Biophysica Acta, 1789(4), 279e298. https://doi.org/10.1016/j.bbagrm.2008.09.005. Dall’Agnese, A., & Puri, P. L. (2016). Could we also be regenerative superheroes, like salamanders? BioEssays, 38(9), 917e926. https:// doi.org/10.1002/bies.201600015. Davidson, E. H. (1994). Molecular biology of embryonic development: How far have we come in the last ten years? BioEssays, 16(9), 603e615. https://doi.org/10.1002/bies.950160903. Driever, W., Solnica-Krezel, L., Schier, A. F., Neuhauss, S. C., Malicki, J., Stemple, D. L., et al. (1996). A genetic screen for mutations affecting embryogenesis in zebrafish. Development, 123, 37e46. Eisen, J. S. (1996). Zebrafish make a big splash. Cell, 87(6), 969e977. Eisen, J. S., Myers, P. Z., & Westerfield, M. (1986). Pathway selection by growth cones of identified motoneurones in live zebra fish embryos. Nature, 320(6059), 269e271. https://doi.org/10.1038/ 320269a0. Fior, R., Povoa, V., Mendes, R. V., Carvalho, T., Gomes, A., Figueiredo, N., et al. (2017). Single-cell functional and chemosensitive profiling of combinatorial colorectal therapy in zebrafish xenografts. Proceedings of the National Academy of Sciences of the United States of America. https://doi.org/10.1073/pnas.1618389114.

I. Introduction

References

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1. History of Zebrafish Research

Oppenheimer, J. M. (1936). Historical introduction to the study of teleostean development. Osiris, 2, 124e148. Oppenheimer, J. M. (1979). 50 Years of Fundulus. Quarterly Review of Biology, 54(4), 385e395. https://doi.org/10.1086/411443. Orger, M. B., & de Polavieja, G. G. (2017). Zebrafish behavior: Opportunities and challenges. Annual Review of Neuroscience. https:// doi.org/10.1146/annurev-neuro-071714-033857. Peterson, R. T., Link, B. A., Dowling, J. E., & Schreiber, S. L. (2000). Small molecule developmental screens reveal the logic and timing of vertebrate development. Proceedings of the National Academy of Sciences of the United States of America, 97(24), 12965e12969. https://doi.org/10.1073/pnas.97.24.12965. Phillips, J. B., & Westerfield, M. (2014). Zebrafish models in translational research: Tipping the scales toward advancements in human health. Disease Models and Mechanisms, 7(7), 739e743. https:// doi.org/10.1242/dmm.015545. Postlethwait, J. H., & Talbot, W. S. (1997). Zebrafish genomics: From mutants to genes. Trends in Genetics, 13(5), 183e190. Postlethwait, J. H., Yan, Y. L., Gates, M. A., Horne, S., Amores, A., Brownlie, A., et al. (1998). Vertebrate genome evolution and the zebrafish gene map. Nature Genetics, 18(4), 345e349. https:// doi.org/10.1038/ng0498-345. van Raamsdonk, W., Mos, W., Smit-Onel, M. J., van de Laarse, W. J., & Fehres, R. (1983). The development of the spinal motor column in relation to the myotomal muscle fibers in the zebrafish (Brachydanio rerio). I. Posthatching development. Anatomy and Embryology, 167(1), 125e139. van Raamsdonk, W., Tekronnie, G., Pool, C. W., & van de Laarse, W. (1980). An immune histochemical and enzymic characterization of the muscle fibres in myotomal muscle of the teleost Brachydanio rerio, Hamilton-Buchanan. Acta Histochemica, 67(2), 200e216. Ramakrishnan, L. (2013). The zebrafish guide to tuberculosis immunity and treatment. Cold Spring Harbor Symposia on Quantitative Biology, 78, 179e192. https://doi.org/10.1101/sqb.2013.78.023283. Rennekamp, A. J., & Peterson, R. T. (2015). 15 years of zebrafish chemical screening. Current Opinion in Chemical Biology, 24, 58e70. https://doi.org/10.1016/j.cbpa.2014.10.025. Richardson, M. K., Hanken, J., Gooneratne, M. L., Pieau, C., Raynaud, A., Selwood, L., et al. (1997). There is no highly conserved embryonic stage in the vertebrates: Implications for current theories of evolution and development. Anatomy and Embryology, 196(2), 91e106. Schier, A. F. (2007). The maternal-zygotic transition: Death and birth of RNAs. Science, 316(5823), 406e407. https://doi.org/10.1126/ science.1140693. Selverston, A. I., & Miller, J. P. (1980). Mechanisms underlying pattern generation in lobster stomatogastric ganglion as determined by selective inactivation of identified neurons. I. Pyloric system. Journal of Neurophysiology, 44(6), 1102e1121. Shima, A., & Mitani, H. (2004). Medaka as a research organism: Past, present and future. Mechanisms of Development, 121(7e8), 599e604. https://doi.org/10.1016/j.mod.2004.03.011.

Spray, D. C., Harris, A. L., & Bennett, M. V. (1979). Voltage dependence of junctional conductance in early amphibian embryos. Science, 204(4391), 432e434. Stahl, F. (1995). George streisinger (Vol. 68). Washington DC: National Academy Press. Stednitz, S. J., McDermott, E. M., Ncube, D., Tallafuss, A., Eisen, J. S., & Washbourne, P. (2018). Forebrain control of behaviorally driven social orienting in zebrafish. Current Biology, 28(15), 2445e2451. https://doi.org/10.1016/j.cub.2018.06.016. e2443. Stewart, W. W. (1981). Lucifer dyesdhighly fluorescent dyes for biological tracing. Nature, 292(5818), 17e21. Streisinger, L. (2004). From the sidelines. Eugene, Oregon: University of Oregon Press. Streisinger, G., Walker, C., Dower, N., Knauber, D., & Singer, F. (1981). Production of clones of homozygous diploid zebra fish (Brachydanio rerio). Nature, 291(5813), 293e296. Tauber, A. I., & Podolsky, S. H. (2000). The generation of diversity clonal selection theory and the rise of molecular immunology. Cambridge MA: Harvard University Press. Trinkaus, J. P. (1990). In H.-J. Marthy (Ed.), Some contributions of research on early teleost embryogenesis to general problems of development. New York: Plenum. Varshney, G. K., Sood, R., & Burgess, S. M. (2015). Understanding and editing the zebrafish genome. Advances in Genetics, 92, 1e52. https://doi.org/10.1016/bs.adgen.2015.09.002. Weis, J. S. (1968a). Analysis of the development of nervous system of the zebrafish, Brachydanio rerio. I. The normal morphology and development of the spinal cord and ganglia of the zebrafish. Journal of Embryology and Experimental Morphology, 19(2), 109e119. Weis, J. S. (1968b). Analysis of the development of the nervous system of the zebrafish, Brachydanio rerio. II. The effect of nerve growth factor and its antiserum on the nervous system of the zebrafish. Journal of Embryology and Experimental Morphology, 19(2), 121e135. Weisblat, D. A., Zackson, S. L., Blair, S. S., & Young, J. D. (1980). Cell lineage analysis by intracellular injection of fluorescent tracers. Science, 209(4464), 1538e1541. Westerfield, M. (2007). The zebrafish book (5th ed.). Eugene, Oregon: University of Oregon Press. Wiles, T. J., Jemielita, M., Baker, R. P., Schlomann, B. H., Logan, S. L., Ganz, J., et al. (2016). Host gut motility promotes competitive exclusion within a model intestinal microbiota. PLoS Biology, 14(7), e1002517. https://doi.org/10.1371/journal.pbio.1002517. Wourms, J. P. (1997). The rise of fish embryology in the nineteenth century. American Zoologist, 37, 269e310. Wourms, J. P., & Whitt, G. S. (1981). Future directions of research on the developmental biology of fishes. American Zoologist, 21(2), 597e604. Zon, L. I. (1999). Zebrafish: A new model for human disease. Genome Research, 9(2), 99e100.

I. Introduction

C H A P T E R

2 Zebrafish Taxonomy and Phylogeny or Taxonomy and Phylogeny Braedan M. McCluskey1, Ingo Braasch2 1

Department of Biology, University of Virginia, Charlottesville, VA, United States of America;2Department of Integrative Biology and Program in Ecology, Evolutionary Biology and Behavior, Michigan State University, East Lansing, MI, United States of America

Introduction

terms, however, all lineages of living vertebrates are fish, with tetrapods, including ourselves, being a specialized type of (lobe-finned) fish capable of living on land (Long, 2011). Evidence for our own piscine evolutionary past is found all over the human body and genome. Table 2.1 provides a phylogenetic classification of zebrafish. Focusing on living representative of vertebrates, Fig. 2.1 shows the phylogenetic position of zebrafish, a ray-finned fish, within the larger vertebrate tree of life and its relation to our own human species. Fig. 2.2 illustrates zebrafish’s phylogenetic relation to other ray-finned fish groups. The subphylum of vertebrates emerged within the chordate phylum more than 500 million years ago (Pough & Janis, 2019). Their closest living relatives are the subphyla urochordates (tunicates), which include, for example, the developmental research organisms Ciona (Kourakis & Smith, 2015) and Oikopleura (Marti-Solans et al., 2015) and, more distantly, the cephalochordates with the classic evo-devo model organism amphioxus (Branchiostoma) (Escriva, 2018) (Fig. 2.1). Among the vertebrates, zebrafish belongs to the lineage of jawed vertebrates or gnathostomes. As the name implies, the emergence of the jaw apparatus was a key innovation of gnathostomes that enabled the exploration of novel food sources, thereby, contributing to the evolutionary success of jawed vertebrates (Liem, Bemis, Walker Jr., & Grande, 2001). Another major morphological innovation leading to the gnathostomes was the acquisition of two sets of paired fins, that is, the pectoral and pelvic fins (Liem, Bemis, Walker, & Grande, 2001), as we can find them in a more derived form in zebrafish. Cyclostomes, that is, lampreys and hagfishes, remain as the only living group of jawless or agnathan

In 1822, the Scottish physician Francis Hamilton scientifically described the zebrafish as Danio rerio along with a few other species of the genus from Eastern India (Hamilton, 1822). Danio rerio remains the correct scientific name for zebrafish, despite recurring changes since the description by Hamilton. In 1916, the Danio genus was divided into two subgenera: Danio and Brachydanio, the latter of which included zebrafish (Weber & de Beaufort, 1916). In 1991, Danio and Brachydanio were synonymized (Barman, 1991) and in 2003 Danio was again separated into the genera Devario and Danio, including zebrafish (Fang, 2003). The zebrafish is one of more than 20 species within the genus Danio and one of more than 4,000 species within the order of Cypriniformes (Parichy, 2015; Stout, Tan, Lemmon, Lemmon, & Armbruster, 2016). In the following, we will briefly describe the phylogenetic position of zebrafish within the broader scale of the vertebrate tree of life and discuss important evolutionary considerations for the use of zebrafish in biomedical research. Along the way, we will discuss zebrafish’s relation to other classic and emerging fish model systems for developmental, genomic, evolutionary, and biomedical research to illustrate the diverse evolutionary framework into which zebrafish research is placed.

The Phylogenetic Position of Zebrafish In everyday language, the term “fish” applies to many different lineages of aquatic vertebrates that have gills and fins (Nelson, 2006). In strict phylogenetic The Zebrafish in Biomedical Research https://doi.org/10.1016/B978-0-12-812431-4.00002-6

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© 2020 Elsevier Inc. All rights reserved.

16

Phylogenetic classification of zebrafish (Danio rerio). Classification below the gnathostome level is following Betancur et al. (2017).

Kingdom

Metazoa (Animalia)

Superphylum

Deuterostomia

Phylum

Chordata

Subphylum

Vertebrata

Infraphylum

Gnathostomata

Megaclass

Osteichthyes (Euteleostome)

Superclass

Actinopterygii

Class

Actinopteri

Subclass

Neopterygii

Infraclass

Teleostei

Supercohort

Clupeocephala

Cohort

Otomorpha

Subcohort

Ostariophysi

Superorder

Cypriniphysae

Order

Cypriniformes

Suborder

Cyprinoidae

Family

Danionidae

Genus

Danio

Species

Danio rerio

vertebrates as outgroups to gnathostomes. Cyclostomes lack jaws and paired fins and feature many other morphological as well as genomic differences, but also important shared characteristics with the jawed vertebrates. Thus, these jawless fish are central taxa in comparative studies to reconstruct ancestral conditions during the early phase of vertebrate evolution and the evolutionary changes that led to the emergence of jawed vertebrates (Nikitina, Bronner-Fraser, & Sauka-Spengler, 2009; Shimeld & Donoghue, 2012) (Fig. 2.1). Gnathostomes are further subdivided into two main groups: the cartilaginous fishes (Chondrichthyes), that is, sharks, skates, and chimeras, and the bony vertebrates (Euteleostome or Osteichthyes), to which zebrafish and human belong (Pough & Janis, 2019) (Fig. 2.1). Several cartilaginous fish species are being used in developmental studies (reviewed in Onimaru, Motone, Kiyatake, Nishida, & Kuraku, 2018) and the recent availability of chondrichthyan genomes (Hara et al., 2018; Venkatesh et al., 2014) promises a rich future for cartilaginous fishes as piscine model organisms for the investigation of gnathostome biology and outgroups to bony vertebrates. Within bony vertebrates, the lineages leading to zebrafish and human then parted around 400e450 million years ago, with the ray-finned fishes (Actinopterygii) that include zebrafish diverging from the lobe-finned fishes (Sarcopterygii) that gave rise to the tetrapods. Only three groups of lobe-finned fishes have survived until today: the coelacanths and the lungfishes, the latter of which are, in turn, the closest living relatives

Cartilaginous fishes

Chordates

Tetrapods

VGD1/2

Vertebrates

Lungfishes

Lobe-finned

Coelacanths

Jawed vertebrates

Ray-finned fishes

LCEA

Bony vertebrates

TABLE 2.1

2. Zebrafish Phylogeny and Evolution

Cyclostomes Urochordates Cephalochordates 700 600

500

400 300

200 100

0

million years ago

FIGURE 2.1 Phylogeny of the vertebrate lineage. Zebrafish belongs to the ray-finned fishes, with their phylogeny further detailed in Fig. 2.2. Reconstructing the last common euteleostome (i.e., bony vertebrate) ancestor (LCEA ¼ red dot) is essential for the comparison of ray-finned and lobe-finned vertebrates, especially for the biomedical link of zebrafish to human. VGD1 and VGD2 indicate the likely occurrences of two rounds of vertebrate genome duplication at the base of vertebrates. The timescales in Figs. 2.1 and 2.2 are based on data obtained from www.timetree.org.

I. Introduction

Zebrafish and Related Danio Species as an Evolutionary Model System

Teleost fishes are divided into three main lineages: the clupeocephalans that include zebrafish and most other fish model systems (see below); the osteoglossomorphs, which include, for example, arowana, African butterflyfish, and mormyrid electric fish; and the elopomorphs to which eels and tarpons belong (Fig. 2.2). The interrelationships of these three major lineages have been difficult to resolve and are a matter of ongoing investigation (Betancur et al., 2017). Within the clupeocephalans, two main radiations are recognized: the ostariophysans with more than 10,000 species and the percomorphs with more than 14,000 species (Alfaro et al., 2009; Chakrabarty et al., 2017; Near et al., 2013). Ostariophysans include fishes as diverse as catfishes, characins, electric knifefishes, and the cypriniforms with zebrafish (Chakrabarty et al., 2017).

Zebrafish Danio genus

Cypriniforms

Carps/Goldfish Cyprininae

Ostariophysans

Blind cavefish Astyanax mexicanus Livebearers platyfish, swordtail, guppy

Medaka Oryzias latipes Percomorphs

Clupeocephalans

Annual killifishes e.g. Nothobranchius furzeri

Cichlids Stickleback Gasterosteus aculeatus Antarctic icefish Notothenioids

Teleosts TGD

17

Pufferfishes

Neopterygians Salmonids

Zebrafish and Related Danio Species as an Evolutionary Model System

Osteoglossomorphs bony tongues, mooneyes Elopomorphs eels, tarpons

Cypriniform and Danionid Relationships

Holostei Spotted gar Lepisosteus oculatus

Ray-finned fishes

Acipenseriforms sturgeons, paddlefish Polypteriforms bichirs, reedfish 400

300

200

100

0

million years ago

FIGURE 2.2 Phylogeny of the ray-finned fish lineage TGD indicates the occurrence of the Teleost Genome Duplication at the base of teleosts. Black pin symbols show the occurrence of additional, lineagespecific genome duplication events.

of the third group, the tetrapods that include us humans (Pough & Janis, 2019) (Fig. 2.1). With the advent of next-generation sequencing techniques and the acquisition of large-scale genomic data across the tree of life, major strides have been made in phylogenomics (i.e., the use of genome-wide sequence information to infer phylogenetic relationships) to reconstruct the evolution of ray-finned fishes (e.g., Betancur et al., 2013; Hughes et al., 2018; Near et al., 2012). Within the ray-finned fishes, the zebrafish belongs to their most species-rich clade, the teleost fishes (Teleostei) (Fig. 2.2). With more than 25,000 species, teleosts make up almost 50% of all living vertebrates (Helfman, Collette, Facey, & Bowen, 2009; Nelson, 2006). Together with their sister lineage, the holostean fishes that consist of bowfin and gars, teleosts are grouped into the Neopterygii. More distantly, teleosts are related to the acipenseriforms (sturgeons and paddlefishes) and the polypteriforms (bichirs and reedfish), the earliest branching lineage among living ray-finned fishes (Betancur et al., 2013; Near et al., 2012) (Fig. 2.2).

Zebrafish is surrounded by cypriniform biodiversity. Cypriniforms represent the largest group of freshwater fishes and besides minnows and suckers also includes species important for aquaculture, such as common carp and grass carp, as well as many popular ornamental species, such as goldfish and rasboras (Stout et al., 2016). The taxonomy of this diverse group has experienced numerous revisions as more species have been described and phylogenetic inference methods have shifted from morphological to molecular methods. This change is particularly relevant to zebrafish, which is referred to as Brachydanio rerio in much of the scientific literature from prior to 1995. Only recently have phylogenomic approaches proved sufficient to provide strong support for relationships between families within Cypriniformes and between species within Danio (McCluskey & Postlethwait, 2015; Stout et al., 2016). The most extensive molecular phylogenomic study of Danio to date supports D. aesculapii as the closest relative of D. rerio (Fig. 2.3). Zebrafish is also closely related to D. kyathit with evidence of gene flow between those lineages during speciation (McCluskey & Postlethwait, 2015). Relationships within Danio will be better resolved as the diversity within this group is further described.

The Emerging Danionid Model System The dozens of Danio species are phenotypically diverse, but all share the major advantages of zebrafish as a model vertebrate making danios ideal for studying interspecific evolution (Irion, Singh, & Nusslein-Volhard, 2016; Parichy, 2006). All danios have

I. Introduction

18

2. Zebrafish Phylogeny and Evolution

are siphoned from the substrate prior to hatching. If embryos are needed at a specific time, in vitro fertilization is often a better option than natural spawning.

The Zebrafish Model in an Evolutionary Context Evolutionary Considerations for Zebrafish-toHuman Comparisons

FIGURE 2.3 Phylogenetic tree showing the relation of zebrafish to other Danio species. Zebrafish strains include two lab strains (AB and TU), as well as two wild strains (NA and WIK). According to this phylogeny following McCluskey and Postlethwait (2015), D. rerio is most closely related to D. aesculapii within the larger danionid clade.

externally fertilized, transparent embryos, and high fecundity, making them immediately accessible for developmental studies. In addition, transgenesis methods used for zebrafish also function in other danios, allowing the use of reporter constructs to investigate differences in gene expression across development (Eom, Bain, Patterson, Grout, & Parichy, 2015). Furthermore, interspecific Danio hybrids can be readily produced by in vitro fertilization and are viable, though many interspecific hybrids are sterile (Parichy & Johnson, 2001). The ability to make hybrids with zebrafish allows for the investigation of phenotypic differences via complementation tests with the huge array of mutants available for D. rerio (Quigley et al., 2005; Spiewak et al., 2018). These approaches have been thus far applied to the analysis of pigmentation, the most striking difference between these species, but are also applicable to understanding the evolution of other phenotypes. Importantly, Danio species can be raised under the laboratory conditions used for zebrafish. They benefit, however, from modest accommodations to the care for Danio rerio. Environmental enrichment in the form of opaque tank bottoms, substrate, and/or plastic plants is beneficial to many species. Some species may also require larger tanks and high water flow. Dietary enrichment with brine shrimp, bloodworms, or rotifers helps condition fish during development and prior to breeding. Unlike laboratory-adapted zebrafish, danios do not breed well in small, clear crossing tanks. For many danios, breeding in the lab can be induced as described in the home aquarium community. Selecting only conditioned fish, use a static tank with freshwater, a substrate to collect eggs, and plastic plants. Embryos

For a meaningful comparative approach that makes use of zebrafish as biomedical model for investigating human biology and disease, we need to be able to define those shared characteristics of the zebrafish and human genome and phenotype that are derived from their last common bony vertebrate, that is, euteleostome, ancestor (LCEA in Fig. 2.1) and distinguish them from the lineage-specific, secondary changes that have occurred independently in the ray-finned and lobe-finned lineages. Clearly, significant evolutionary changes have impacted the zebrafish and human lineages since their last common bony vertebrate ancestor. For example, considering the lobe-finned lineage leading to human, the water-to-land transition and the emergence of tetrapods involved major changes at the morphological level, such as turning fins into limbs, concomitantly with changes at the genomic level, for example the loss of genes encoding some structural components of fins (Amemiya et al., 2013; Wood & Nakamura, 2018). On the other hand, morphological changes within ray-finned fishes, such as the emergence of a homocercal caudal fin structure in early teleosts or the evolution of the Weberian apparatus within the ostariophysans illustrate important phenotypic changes leading to zebrafish (Metscher & Ahlberg, 1999) that need to be taken into account when aiming to compare zebrafish and human anatomy. Besides morphological innovations, it might also be secondary reductions or losses of structures, tissues, and cell types that can diversify and thus complicate the connectivity of zebrafish-to-human comparisons. For example, mammals have reduced their repertoire of neural crest cell-derived pigment cell types to the melanocytes, while teleosts like zebrafish possess a whole suite of different chromatophore types (melanophores, xanthophores, iridophores, leucophores, etc.) (Parichy & Aman, 2019), with many of these pigment cell types likely inherited from a bony vertebrate ancestor. A detailed comparison of zebrafish and human morphology and physiology is thus essential for best practices in the translation of biomedical relevant data across lineages, motivating the development and annotation of phenotypic ontologies as used by The Zebrafish Information Network ZFIN (http://zfin.org/) that can then be applied across species (Van Slyke, Bradford, Westerfield, & Haendel, 2014).

I. Introduction

Zebrafish and Its Relation to Other Fish Model Species

Importance of Genome Duplications for Zebrafish Evolution At the genomic level, an important aspect of vertebrate evolution that needs to be considered in humanto-zebrafish comparisons is the role of three rounds of whole-genome duplications (or polyploidizations) that have impacted the zebrafish and human lineages in several ways (Figs. 2.1 and 2.2). Polyploidizations are rare events in animals but have occurred comparatively frequently in fishes (Braasch & Postlethwait, 2012) (Fig. 2.2). The amplification and diversification of vertebrate gene families imposed by the three vertebrate genome duplications have led to complex scenarios of gene function evolution that sometimes are difficult to disentangle. All living teleost species, including zebrafish, are derived from an ancestor that underwent a wholegenome duplication event at the dawn of teleost evolution. This Teleost Genome Duplication (TGD) occurred within the neopterygian lineage after the separation of teleosts from the holostean fishes (gars and bowfin), but before the divergence of the three major teleost lineages of clupeocephalans, osteoglossomorphs, and elopomorphs (Fig. 2.2) (reviewed in Braasch & Postlethwait, 2012). For details on the genomic impacts of the TGD see Postlethwait and Braasch (2019) in this volume. Following the TGD, the teleost genome originally was tetraploid, but it has secondarily returned to the diploid state through the process of rediploidization. Importantly, depending on methods of TGD gene duplicate inference, it has been estimated that between 1,200 and 3,400 pairs of TGD gene duplicates have remained in the zebrafish genome (Howe et al., 2013; Pasquier et al., 2017; Roux, Liu, & Robinson-Rechavi, 2017). Thus, for a significant portion of human genes, there will be two TGD-derived gene duplicates (also called co-orthologs or paralogs) in the zebrafish genome. This can complicate comparisons between zebrafish and human as one may have to take two genomic regions in the zebrafish into account, potentially harboring two TGD co-orthologs of the gene under investigation. For example, there are two sonic hedgehog (shh) genes present in the zebrafish genome, shha and shhb, that are co-orthologs to the single SHH gene in human. Since their fixation in the teleost genome, TGD duplicates will have diverged in function in often complex patterns, following evolutionary fates such as subfunctionalization (the distribution of ancestral gene functions among duplicates) and neofunctionalization (emergence of novel gene functions) (Force et al., 1999). Comparing zebrafish to other teleosts, one will further find differences in terms of TGD duplicate retention and loss among species. Although the overall

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retention rate of TGD duplicates appears to be relatively similar across teleosts, differences in terms of the specific genes retained in duplicate within individual teleost lineages have contributed to the genomic diversification of teleost lineages. Even if different teleost groups have retained both TGD duplicates, lineage-specific divergence in the functional roles among duplicates can be observed (Braasch & Postlethwait, 2012). In addition to the TGD, the vertebrate lineage is derived from likely two additional, earlier ancient rounds of whole-genome duplication at the base of the vertebrate lineage, that occurred after their separation from nonvertebrate chordates. The evolutionary sequence of these Vertebrate Genome Duplications, VGD1 and VGD2 (Fig. 2.1), remains a matter of ongoing debate (Sacerdot, Louis, Bon, Berthelot, & Roest Crollius, 2018; Smith et al., 2018). Differential loss of VGD1/VGD2 gene duplicates after the divergence of the ray-finned and lobe-finned lineages has led to situations in which no directly orthologous genes remain in the zebrafish and human genomes (Postlethwait, 2007). This can complicate the transition of genetic information from one system to the other. For example, the intensively studied human stem cell factor POU5F1 (OCT4) has no direct ortholog in the zebrafish genome because of secondary loss from the ray-finned genome following VGD1/VGD2; its VGD1/VGD2 duplicate Pou5f3, on the other hand, is present in the rayfinned, and thus, zebrafish genome, while having been secondarily eliminated from the eutherian mammal and hence human genome (Frankenberg et al., 2014). In summary, a careful examination of gene family history evolution is paramount for the transfer of genetic information between zebrafish and human. Gene orthology predictions provided, for example, by the ZFIN and Ensembl (www.ensembl.org) databases are good starting points for in-depth phylogenetic investigations of gene family relationships across vertebrates.

Zebrafish and Its Relation to Other Fish Model Species Zebrafish is the most commonly used fish species in biomedical research, but it is important to remember that zebrafish is just one of tens of thousands of teleost fish species. It, therefore, provides a snapshot of the tremendous genotypic and phenotypic biodiversity of teleosts, and comparisons to other phylogenetically diverse fish models are essential to inform the evolution of zebrafish in relation to the teleost, ray-finned, and bony vertebrate ancestors.

I. Introduction

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2. Zebrafish Phylogeny and Evolution

Medaka The other main fish species used in biomedical research is the Japanese rice fish, medaka (Oryzias latipes), that has been cultivated in Japan for centuries. The medaka is very similar in husbandry and laboratory use to zebrafish (Kinoshita, Murata, Naruse, & Tanaka, 2009), has a sequenced genome (Kasahara et al., 2007), and numerous laboratory and natural strains, mutant and transgenic lines, and other resources that can be received through the NBRP Medaka Resource Center (https://shigen.nig.ac.jp/medaka/). Comparative tables of developmental stages of zebrafish and medaka have been developed (Furutani-Seiki & Wittbrodt, 2004; Tena et al., 2014). Medaka is embedded in a larger Orzyias species complex with largely unexplored potential for evolutionary research (Hilgers & Schwarzer, 2019). Furthermore, medaka is a percomorph teleost, and its lineage diverged from the lineage leading to zebrafish early during clupeocephalan teleost evolution (Fig. 2.2). Thus, from an evolutionary point of view, the use of medaka in biomedical research is highly complementary to that of zebrafish (Furutani-Seiki & Wittbrodt, 2004). Comparison of gene expression and gene functions between the two are useful to evaluate the extent of lineage-specific divergence in either species, which is particularly important given that both lineages diverged relatively soon after the TGD and have hence undergone, for example, separate paths in TGD gene duplicate retention/loss, as well as TGD gene duplicate function divergence (Furutani-Seiki & Wittbrodt, 2004). For example, while the two TGD gene duplicates shha and shhb are retained in zebrafish, medaka has only kept the shha copy of the sonic hedgehog gene. In contrast, there are two sox10 TGD duplicates in the medaka genome, while zebrafish retained only a single sox10 gene.

“Evolutionary Mutant” Fish Models Beyond zebrafish and medaka, there are numerous other teleost species utilized in biomedically-focused research, often because they feature specific adaptations that resemble maladaptive conditions and diseases in human. These “evolutionary mutant models” (Albertson, Cresko, Detrich, & Postlethwait, 2009) have become powerful tools for the study of vertebrate development, evolution, and human disease (Braasch et al., 2015; Schartl, 2014). They benefit from recent genome sequencing initiatives, as well as from the transfer of advances in the genetic-developmental analytical toolbox, such as genome-editing, transgenics, and other functional approaches first developed and/or optimized in zebrafish. Below we highlight a few examples of these biomedical fish models, but this list is far from exhaustive.

Goldfish and carp: Within the cypriniforms, goldfish (Carassius auratus) and common carp (Cyprinus carpio) are important aquaculture species with a centurieslong history of domestication. Particularly goldfish is a promising model for biomedical investigation with its multitude of variants and strains that feature extreme morphologies of eyes, fins, skeleton, body shape, coloration, etc., some of which are resembling human diseases (Omori & Kon, 2019). Importantly, goldfish and carp are both derived from an additional, carp lineagespecific genome duplication event that occurred in their common ancestor within the last few million years (Fig. 2.2). Using zebrafish as “unduplicated” outgroup with respect to the carp genome duplication, analyzing the evolutionary aftermath of the carp genome duplication is becoming an important avenue to understand the genomic and morphological impact of genome duplications in vertebrates (Chen et al., 2019; Xu et al., 2014). Blind cavefish: Cave populations of the Mexican tetra (Astyanax mexicanus), usually referred to as “blind cavefish,” have become popular research organism to study traits that evolved in response to the constant darkness and seclusion of cave environments and that have resemblances to human disease phenotypes, including eye regression, albinism, obesity, sleep loss, behavioral changes, and others (Jeffery, 2008; Rohner, 2018). The Mexican tetra is a characiform that belongs to the ostariophysan lineage and thus shares a comparatively close relationship to zebrafish (Fig. 2.2). Hence, zebrafish has been used as surrogate species to study the functional consequences of genetic differences between cave versus surface populations of Astyanax (e.g, Gross, Borowsky, & Tabin, 2009; Riddle et al., 2018). Besides medaka, the percomorph clade of teleosts offers a plethora of fish species gaining popularity in biomedical, evolutionary, and genomic research (Fig. 2.2). Livebearers: Livebearing poeciliids (guppies, platies, swordtails) are common model species in evolutionary research, and among them, the genus Xiphophorus has a long tradition to investigate the genomic basis of skin cancer formation because melanoma formation can be generated by different interspecific crossing schemes (Meierjohann & Schartl, 2006). Poeciliids use internal fertilization and development, and genetic manipulations are thus challenging. Therefore, medaka and zebrafish have been used to study aspects of the Xiphophorus oncogenic signaling cascades leading to cancer formation in vivo (Li et al., 2012; Regneri, Klotz, & Schartl, 2016). Annual killifishes: Annual killifishes are a major model system to investigate rapid aging and developmental arrest (diapause), which are adaptations to their unique life cycle in seasonally desiccating water bodies. The turquoise killifish (Nothobranchius furzeri) has

I. Introduction

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References

emerged as the most popular annual killifish species in aging research as it represents the most short-lived known vertebrate with a few months of lifespan and sexual maturity within a few weeks after hatching (Cellerino, Valenzano, & Reichard, 2016; Platzer & Englert, 2016). Cichlids: With several thousand species, cichlids are a prime example for adaptive radiations and phenotypic diversification (Salzburger, 2018) that feature complex social behaviors (Fernald, 2017) and trophic adaptations of the jaw apparatus, which can be used to model human craniofacial diseases (Powder & Albertson, 2016). Again, comparative functional analyses using zebrafish have been instrumental in enlightening the underlying genetic basis of such biomedically relevant phenotypes (Cooper, Wirgau, Sweet, & Albertson, 2013; Powder, Cousin, McLinden, & Albertson, 2014). Sticklebacks: Three-spined stickleback (Gasterosteus aculeatus) and related species are major model organisms for evolutionary and ecological genetics (Peichel & Marques, 2017). Studies on the genetic basis of phenotypic differences among stickleback populations, for example, in the pigmentary and skeletal systems, found parallels to phenotypic diversity among human populations, yet also pointed to human-specific changes during vertebrate evolution, respectively (Indjeian et al., 2016; Miller et al., 2007). Furthermore, sticklebacks often serve as percomorph comparators to zebrafish in developmental-genetic studies (e.g., Askary et al., 2016; Jovelin et al., 2007). Antarctic icefish: Notothenioid icefish are uniquely adapted to the extreme cold of Antarctic waters: they possess antifreeze proteins, many species are characterized by reduced bone formation, and they lack functional hemoglobin and red blood cells. Thus, they can serve as models for human diseases, such as osteopenia and osteoporosis and anemia (Albertson et al., 2009). Due to their unique ecology, Antarctic icefish are difficult to study in captivity, and thus, zebrafish have been used for example to functionally investigate genes with a role in erythrocyte formation that emerged from investigating the unique blood cell development in icefish (Yergeau, Cornell, Parker, Zhou, & Detrich, 2005).

Nonteleost Fish Provide Connectivity From Zebrafish to Human Given the derived nature of both teleost phenotypes and the teleost genome as a result of the TGD, zebrafish-to-human comparisons benefit from the inclusion of morphological and genomic data from nonteleost fishes. In recent years, the holostean spotted gar (Lepisosteus oculatus; Fig. 2.2) has emerged as a “bridge species” that provides connectivity among bony vertebrate

lineages. Due to its “unduplicated” nature with regard to the TGD and its comparatively slow rates of genomic and morphological evolution, gar provides important reference points for the transition of biomedically relevant morphological and genomic information across bony vertebrates and to link zebrafish to human biology (Braasch et al., 2015, 2016). This has helped, for example, to clarify the evolution of joint development in bony vertebrates, enabling to then develop an arthritis model in zebrafish (Askary et al., 2016).

Conclusion and Outlook Zebrafish is a unique representative of ray-finned teleost fish biodiversity that diverged from human more than 400 million years ago. While being a powerful model system to study vertebrate biology and human disease, it is important to keep in mind that zebrafish neither is a “prototypic” fish that stopped evolving after separation from the human lineage nor that findings in zebrafish are necessarily generalizable across fish lineages. A meaningful utilization of the zebrafish model in biomedical research, therefore, calls, on the one hand, for the detailed elucidation of shared and divergent characteristics of zebrafish and human at both the phenotypic and genetic levelsdas exemplified by the information provided in the chapters of this volume. On the other hand, the inclusion of information from the phylogenetically expanding swarm of fish model systems will put zebrafish research into an enriched evolutionary context. With the relentless technical improvements for its investigation, zebrafish will continue to lead the charge in illuminating the genotype-to-phenotype map in the fish world.

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I. Introduction

References

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Platzer, M., & Englert, C. (2016). Nothobranchius furzeri: A model for aging research and more. Trends in Genetics, 32(9), 543e552. https:// doi.org/10.1016/j.tig.2016.06.006. Postlethwait, J. H. (2007). The zebrafish genome in context: Ohnologs gone missing. Journal of Experimental Zoology Part B: Molecular and Developmental Evolution, 308(5), 563e577. https://doi.org/ 10.1002/jez.b.21137. Postlethwait, J. H., & Braasch, I. (2019). Chapter 26: Genetics. In S. Cartner, S. Farmer, & K. Guillemin (Eds.), Zebrafish in biomedical research. Elsevier. Pough, F. H., & Janis, C. M. (2019). Vertebrate life (10th ed.). New York: Oxford University Press. Powder, K. E., & Albertson, R. C. (2016). Cichlid fishes as a model to understand normal and clinical craniofacial variation. Developmental Biology, 415(2), 338e346. https://doi.org/10.1016/ j.ydbio.2015.12.018. Powder, K. E., Cousin, H., McLinden, G. P., & Albertson, R. C. (2014). A nonsynonymous mutation in the transcriptional regulator lbh is associated with cichlid craniofacial adaptation and neural crest cell development. Molecular Biology and Evolution, 31(12), 3113e3124. https://doi.org/10.1093/molbev/msu267. Quigley, I. K., Manuel, J. L., Roberts, R. A., Nuckels, R. J., Herrington, E. R., MacDonald, E. L., et al. (2005). Evolutionary diversification of pigment pattern in Danio fishes: Differential fms dependence and stripe loss in D. albolineatus. Development, 132(1), 89e104. https://doi.org/10.1242/dev.01547. Regneri, J., Klotz, B., & Schartl, M. (2016). Genomic and transcriptomic approaches to study cancer in small aquarium fish models. Advances in Genetics, 95, 31e63. https://doi.org/10.1016/ bs.adgen.2016.04.001. Riddle, M. R., Aspiras, A. C., Gaudenz, K., Peuss, R., Sung, J. Y., Martineau, B., et al. (2018). Insulin resistance in cavefish as an adaptation to a nutrient-limited environment. Nature, 555(7698), 647. https://doi.org/10.1038/nature26136. Rohner, N. (2018). Cavefish as an evolutionary mutant model system for human disease. Developmental Biology, 441(2), 355e357. https://doi.org/10.1016/j.ydbio.2018.04.013. Roux, J., Liu, J., & Robinson-Rechavi, M. (2017). Selective constraints on coding sequences of nervous system genes are a major determinant of duplicate gene retention in vertebrates. Molecular Biology and Evolution, 34(11), 2773e2791. https://doi.org/10.1093/molbev/ msx199. Sacerdot, C., Louis, A., Bon, C., Berthelot, C., & Roest Crollius, H. (2018). Chromosome evolution at the origin of the ancestral vertebrate genome. Genome Biology, 19(1), 166. https://doi.org/ 10.1186/s13059-018-1559-1. Salzburger, W. (2018). Understanding explosive diversification through cichlid fish genomics. Nature Reviews Genetics, 19(11), 705e717. https://doi.org/10.1038/s41576-018-0043-9. Schartl, M. (2014). Beyond the zebrafish: Diverse fish species for modeling human disease. Disease Models and Mechanisms, 7(2), 181e192. https://doi.org/10.1242/dmm.012245. Shimeld, S. M., & Donoghue, P. C. (2012). Evolutionary crossroads in developmental biology: Cyclostomes (lamprey and hagfish). Development, 139(12), 2091e2099. https://doi.org/10.1242/ dev.074716. Smith, J. J., Timoshevskaya, N., Ye, C., Holt, C., Keinath, M. C., Parker, H. J., et al. (2018). The sea lamprey germline genome provides insights into programmed genome rearrangement and vertebrate evolution. Nature Genetics, 50(2), 270e277. https://doi.org/ 10.1038/s41588-017-0036-1. Spiewak, J. E., Bain, E. J., Liu, J., Kou, K., Sturiale, S. L., Patterson, L. B., et al. (2018). Evolution of Endothelin signaling and diversification of adult pigment pattern in Danio fishes. PLoS Genetics, 14(9). https://doi.org/10.1371/journal.pgen.1007538. e1007538.

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2. Zebrafish Phylogeny and Evolution

Stout, C. C., Tan, M., Lemmon, A. R., Lemmon, E. M., & Armbruster, J. W. (2016). Resolving Cypriniformes relationships using an anchored enrichment approach. BMC Evolutionary Biology, 16(1), 244. https://doi.org/10.1186/s12862-016-0819-5. Tena, J. J., Gonzalez-Aguilera, C., Fernandez-Minan, A., VazquezMarin, J., Parra-Acero, H., Cross, J. W., et al. (2014). Comparative epigenomics in distantly related teleost species identifies conserved cis-regulatory nodes active during the vertebrate phylotypic period. Genome Research, 24(7), 1075e1085. https://doi.org/ 10.1101/gr.163915.113. Van Slyke, C. E., Bradford, Y. M., Westerfield, M., & Haendel, M. A. (2014). The zebrafish anatomy and stage ontologies: Representing the anatomy and development of Danio rerio. Journal of Biomedical Semantics, 5. doi:Unsp 1210.1186/2041-1480-5-12. Venkatesh, B., Lee, A. P., Ravi, V., Maurya, A. K., Lian, M. M., Swann, J. B., et al. (2014). Elephant shark genome provides unique

insights into gnathostome evolution. Nature, 505(7482), 174e179. https://doi.org/10.1038/nature12826. Weber, M., & de Beaufort, L. F. (1916). The fishes of the Indo-Australian Archipelago. Leiden: E. J. Brill. Wood, T. W. P., & Nakamura, T. (2018). Problems in fish-to-tetrapod transition: Genetic expeditions into old specimens. Frontiers in Cell and Developmental Biology, 6, 70. https://doi.org/10.3389/ fcell.2018.00070. Xu, P., Zhang, X., Wang, X., Li, J., Liu, G., Kuang, Y., et al. (2014). Genome sequence and genetic diversity of the common carp, Cyprinus carpio. Nature Genetics, 46(11), 1212e1219. https://doi.org/ 10.1038/ng.3098. Yergeau, D. A., Cornell, C. N., Parker, S. K., Zhou, Y., & Detrich, H. W., 3rd (2005). bloodthirsty, an RBCC/TRIM gene required for erythropoiesis in zebrafish. Developmental Biology, 283(1), 97e112. https://doi.org/10.1016/j.ydbio.2005.04.006.

I. Introduction

C H A P T E R

3 Zebrafish Genetics John H. Postlethwait1, Ingo Braasch2 Institute of Neuroscience, University of Oregon, Eugene, OR, United States of America; 2Department of Integrative Biology and Program in Ecology, Evolutionary Biology and Behavior, Michigan State University, East Lansing, MI, United States of America

1

Introduction

studies to human biology requires an understanding of a massive genomic event, a whole-genome duplication (WGD), in the ancestors of all living teleosts, the Teleost Genome Duplication (TGD). As a result of this ancient polyploidization event, a significant portion of human genes have two duplicate gene copies in the zebrafish genome; this situation requires researchers to take special care in making human-to-zebrafish gene comparisons. Here, we discuss the zebrafish genome in terms of the TGD and its significance for zebrafish research.

Teleost fish present an amazing variety of forms, display a wondrous diversity of color patterns, vary enormously in size, occupy environments with remarkably different temperatures and pressures, and represent half of all vertebrate species. Additionally, many teleosts thrive with phenotypes that would be pathogenic in a human, including the degenerated eyes of cavefish, the profound anemia of Antarctic icefish, and the reduced bone mineralization of acidic bogdwelling fish. Understanding the origin of this impressive biodiversity and linking the molecular and cellular basis of phenotypic traits among speciesdincluding, importantly, connecting teleost biology to human health and diseasedrequires an understanding of genetics and genomes. Biologists have focused on the genetics of a few model teleost species in-depth, including medaka (Oryzias latipes), stickleback (Gasterosteus aculeatus), swordtails and platyfish (Xiphophorus species), Mexican tetra cavefish (Astyanax mexicanus), the cichlid radiation, and zebrafish (Danio rerio). Here, we focus on the genetics of zebrafish, a teleost fish model system for investigating the genetic basis of development, physiology, reproduction, evolution, cell biology, and behavior. Zebrafish are optically translucent, have externally developing embryos, a rapid life cycle, small adult size, and large clutches; features that make them amenable to large-scale and high-throughput forward genetic investigations, studies that propelled our early understanding of zebrafish genetics. Clear, external embryos provided zebrafish researchers access to the cellular basis of early development that is more difficult to extract from opaque mammalian embryos huddled inside the womb. The connection of zebrafish genetic

The Zebrafish in Biomedical Research https://doi.org/10.1016/B978-0-12-812431-4.00003-8

The Zebrafish Karyotype The zebrafish genome is organized into 25 chromosome pairs (Amores & Postlethwait, 1999; Daga, Thode, & Amores, 1996; Gornung, Gabrielli, Cataudella, & Sola, 1997; Lee & Smith, 2004; Pijnacker & Ferwerda, 1995; Schreeb, Groth, Sachsse, & Freundt, 1993; Sola & Gornung, 2001; Traut & Winking, 2001). The zebrafish karyotype is within the typical range for actinopterygian (ray-finned) fishes, which show remarkable conservation of chromosome numbers: the majority of teleosts have 48 or 50 chromosomes as diploids (Mank & Avise, 2006; Naruse et al., 2004). Zebrafish has predominantly submetacentric and metacentric chromosomes (Amores & Postlethwait, 1999; Phillips, Amores, Morasch, Wilson, & Postlethwait, 2006). Karyotype investigations of laboratory or pet store zebrafish strains have generally found no evidence for sex chromosomes, although one study of wild zebrafish from northwest India reported heteromorphic sex chromosomes with females (WZ) having a submetacentric Z chromosome and a metacentric W chromosome and males (ZZ) having two submetacentric Z chromosomes (Sharma, Sharma, & Tripathi, 1998).

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© 2020 Elsevier Inc. All rights reserved.

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3. Zebrafish Genetics

(A) Centromeres

(B) Idiogram

Cyto 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 LG 5 7 4 3 9 1 21 2 6 20 18 16 14 17 13 19 8 10 12 23 15 11 24 25 22

FIGURE 3.1 The karyotype of laboratory strain zebrafish. (A) Centromeres labeled yellow with type I satellite DNA (Phillips et al., 2006). (B) The zebrafish ideogram based on replication banding, with black segments corresponding to early replicating bands (Amores & Postlethwait, 1999). Cyto, cytogenetic chromosome number; LG, linkage group number.

By convention, cytogenetic studies ordered zebrafish chromosomes by their length (Fig. 3.1), with chromosome 1 substantially larger than the others, chromosome 3 replicating late in the cell cycle, and chromosomes 7 and 15 as the only large chromosomes that are exactly metacentric (Amores & Postlethwait, 1999).

The Zebrafish Genetic Map In 1974, George Streisinger submitted an application to the US National Science Foundation to make mutations that block early zebrafish development (Varga, 2018). Over the course of the next few years, Streisinger and his colleagues, including Charline Walker and David Grunwald, developed efficient methods for mutagenizing zebrafish and isolating mutants, publishing the first induced mutation in 1988 (Chakrabarti, Streisinger,

Singer, & Walker, 1983; Grunwald, Kimmel, Westerfield, Walker, & Streisinger, 1988; Walker & Streisinger, 1983). The induction of other mutations followed (Felsenfeld, Walker, Westerfield, Kimmel, & Streisinger, 1990; Kimmel, Kane, Walker, Warga, & Rothman, 1989), and 1996 saw the publication of hundreds of mutants from two large-scale mutagenesis screens (Driever et al., 1996; Haffter et al., 1996). Although the first induced mutation was cloned in 1994 by a candidate gene approach (no tail ¼ brachyury, or T-box transcription factor Ta (tbxta) (Schulte-Merker, van Eeden, Halpern, Kimmel, & Nusslein-Volhard, 1994)), it became clear that genetic mapping would be essential for the molecular identification of most zebrafish mutations. The first zebrafish genetic map used 401 random amplified polymorphic DNAs (RAPDs) and simple sequence repeats (SSRs) to construct a genetic linkage map and simultaneously position nine induced

I. Introduction

27

The Zebrafish Genetic Map

(A) Linkage group vs. karyotype

Linkage group

25 20 15 10 5 5 10 15 20 Cytogenetic chromosome number

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(B) Genome sequence length vs. karyotype 80 70 Length (MB)

mutations on the map (Postlethwait et al., 1994). Subsequent genetic maps used other types of DNA markers, including single-nucleotide polymorphisms (SNPs), to create maps with more markers (Fornzler et al., 1998; Gates et al., 1999; Kelly et al., 2000; Knapik et al., 1996, 1998; Postlethwait et al., 1998; Shimoda et al., 1999; Woods et al., 2000, 2005). The application of Streisinger’s techniques to perform half-tetrad genetic analysis with zebrafish (Streisinger, Walker, Dower, Knauber, & Singer, 1981) linked molecular genetic markers to centromeres, and thus, consolidated linkage groups into chromosomal-length constructs (Johnson et al., 1996; Kane, Zon, & Detrich, 1999; Kauffman et al., 1995; Mohideen, Moore, & Cheng, 2000). The use again of Streisinger’s zebrafish cloning methods to produce fully homozygous gynogenetic doubled-haploid zebrafish from the AB and the Tu¨bingen (TU) strains, followed by the production and sequencing of 459 of their F2 progeny for 201,917 SNP loci produced the SATmap, with the greatest resolution of any animal genetic mapping panel (Howe et al., 2013; Patowary et al., 2013). The rate of genetic recombination is not uniform throughout the genome or between the sexes. The recombination rate, in general, is higher near telomeres and lower near centromeres (see, e.g. Howe et al., 2013, Supplementary Fig. 3.3). Sex-specific recombination rates were obtained using androgenetic haploid embryos produced by destroying the egg nucleus with gamma-rays followed by fertilization with normal sperm to provide recombinant offspring derived solely from the male parent (Corley-Smith, Brandhorst, Walker, & Postlethwait, 1999; Singer et al., 2002) to compare with gynogenetic maps based on female meiosis. Results showed that the recombination rate in male meiosis is about half that in female meiosis, especially near the centromere. Consequently, using female meiosis in a positional cloning project gives a greater ratio of genetic map distance in centiMorgans to physical distance in Megabases, but using male meiosis tends to minimize recombination, and thereby, maintain haplotypes intact. Genetic linkage maps ordered linkage groups according to length, with linkage group (LG) 1 being the longest and LG25 the shortest (Fornzler et al., 1998; Gates et al., 1999; Johnson et al., 1996; Kelly et al., 2000; Knapik et al., 1996, 1998; Postlethwait et al., 1994; Shimoda et al., 1999; Woods et al., 2000, 2005). Due to the variable ways that chromatin coils during metaphase of mitosis and chromosomes recombine during prophase I of meiosis, a chromosome’s physical length does not scale one-to-one with its genetic length as determined by recombination rate; thus, the numbering system of cytogenetically defined chromosomes (mitosis) and the numbering convention of linkage groups (meiosis) are not directly synonymous

60 50 40 30 20 10 5 10 15 20 Cytogenetic chromosome number

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FIGURE 3.2 Lengths of chromosomes, linkage groups, and genome sequences. (A) Linkage group numbers (proportional to lengths in centiMorgans based on recombination rates) plotted versus the cytogenetic chromosome numbers (proportional to physical lengths in metaphase chromosome preparations). (B) Lengths of chromosomes measured in numbers of base pairs in the zebrafish genome assembly (GRCz11, http://uswest.ensembl.org/Danio_rerio/ Location/Genome) plotted against cytogenetic chromosome numbers.

(Fig. 3.2). For correcting this problem, genes mapped to LGs were associated with cytogenetic chromosomes by fluorescence in situ hybridization (Phillips et al., 2006). Note that the zebrafish genome assembly (http://uswest.ensembl.org/Danio_rerio/Info/Index? db¼core) mixes nomenclature, using the numbers of linkage groups adopted from genetic maps, but the symbol “chr”; thus, cytogeneticists call the longest chromosome “chromosome 1,” but gene mappers call it LG 5, and the genome assembly calls it “chr5” or “Chromosome 5” (Figs. 3.1B and 3.2A). Similarly, the late-replicating cytogenetic chromosome 3, which is LG4, becomes “chr4” or “Chromosome 4” in the genome assembly. Note also that this chromosome has more DNA than expected from its physical length, perhaps because of the tight packing of repetitive DNA as chromosomes condense in mitosis (Fig. 3.2B). Current literature uses the genome assembly nomenclature when referring to specific chromosomes or linkage groups. An alternative chromosome nomenclature that minimizes confusion when considering chromosomes

I. Introduction

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3. Zebrafish Genetics

of several species is to use the first letter of the genus and the first two letters of the second term in the binomial name of the species to designate chromosomes, so the zebrafish cytogenetic chromosome 3 (LG4) becomes Dre4 (Danio rerio LG4) and human chromosome 7 becomes Hsa7 (Homo sapiens chromosome 7).

Zebrafish and the Teleost Genome Duplication Exploration of the zebrafish genetic map provided an explanation for the early conundrum that “zebrafish has too many genes,” for example, three engrailed-related genes rather than two as in mammals (Ekker, Wegner, Akimenko, & Westerfield, 1992),: five msx-family genes rather than three in mammals (Akimenko, Johnson, Westerfield, & Ekker, 1995; Postlethwait, 2006): and many “extra” hox’ cluster genes compared to human (Prince, Joly, Ekker, & Ho, 1998). When these “extra” genes were placed on the emerging zebrafish genetic maps, conserved gene content, also known as conserved synteny analyses, showed that they usually resided in pairs of duplicated chromosome segments that represent two fragmented copies of portions of individual human chromosomes (Amores et al., 1998), as would be expected from a genome duplication event. Comparing the seven zebrafish hox clusters to the five that had been cloned in fugu pufferfish (Aparicio et al., 1997) showed that each fugu hox cluster was orthologous to one zebrafish cluster, including duplicated hoxa clusters. The simplest explanation was that the hox cluster duplication event occurred before the divergence of zebrafish and pufferfish lineages early in the teleost radiation. Because orthologs of segments of human chromosomes appeared to be present in two copies in both zebrafish and pufferfish, the simplest hypothesis was a wholegenome duplication event shared by clupeocephalan teleosts (Postlethwait, Amores, Force, & Yan, 1998; Taylor, Braasch, Frickey, Meyer, & Van de Peer, 2003; Taylor, Van de Peer, Braasch, & Meyer, 2001), which includes the last common ancestor of zebrafish and fugu and all other living teleosts except the elopomorphs (eels) and osteoglossiforms (bony tongues). Recent data shows that these two basally diverging teleost groups also share the duplication event (Bian et al., 2016; Du et al., 2019; Gallant, Losilla, Tomlinson, & Warren, 2017; Henkel et al., 2012; Jansen et al., 2017; Vialle et al., 2018), which is now called the teleost genome duplication (TGD). The TGD occurred after the divergence of teleosts from their sister group, the Holostei (including gars and bowfin), because spotted gar has just one chromosome segment orthologous to each human chromosome segment (Amores, Catchen, Ferrara, Fontenot, & Postlethwait, 2011; Braasch et al., 2014, 2015, 2016).

Hox gene clusters provide a typical example for the evolution of duplicate chromosome segments after genome duplication. The human genome contains four HOX gene clusters called HOXA, HOXB, HOXC, and HOXD on four different chromosomes (Fig. 3.3A). These four paralogons can be explained by the hypothesis of two rounds of whole-genome duplication early in the vertebrate lineage, called the vertebrate genome duplications (VGD1 and VGD2), in which a chromosome with one ancestral hox cluster duplicated twice to produce four chromosomes, each with one hox-cluster and its surrounding genes (Dehal & Boore, 2005; Nakatani, Takeda, Kohara, & Morishita, 2007; Sacerdot, Louis, Bon, Berthelot, & Roest Crollius, 2018, but see Smith et al., 2018). Then the third round of genome duplication, the TGD, resulted in eight hox clusters. The eight zebrafish hox clusters (hoxaa, hoxab, hoxba, hoxb, hoxca, hoxcb, hoxda, hoxdb) reside on eight different chromosomes, but only seven of the zebrafish hox clusters contain proteincoding genes (Amores et al., 1998). The eighth cluster, hoxdb, lacks any protein-coding gene, but contains a microRNA gene found in the HOXD complex of mammals, other vertebrates, and even Drosophila hox genes, and is flanked by orthologs of genes that border the human HOXD complex (Woltering & Durston, 2006). Hox clusters of percomorph fish, including medaka (Kurosawa et al., 2006), generally possess, like zebrafish, seven protein-encoding clusters, but the percomorph hoxdb cluster contains protein-coding genes, and reciprocally, percomorphs lack the hoxcb cluster.

Zebrafish Gene Nomenclature Conventions Zebrafish gene nomenclature conventions attempt to reflect gene histories, including evolutionary relationships to human genes and the TGD (see https://wiki. zfin.org/display/general/ZFINþZebrafishþNomencla tureþConventions). First, the zebrafish gene name is generally a lower-case version of its human ortholog, reflecting evolution from a single gene in their last common ancestor 450 million years ago. Second, when zebrafish has two genes that are TGD co-orthologs of the human gene, the gene name has an “a” or “b” appended to it, for example, hoxa9a and hoxa9b are coorthologs of the human HOXA9 gene. Zebrafish genes that are co-orthologs of a single human gene are ohnologs of each other, where ohnologs are paralogs derived from a genome duplication event honoring Susumu Ohno, who suggested the importance of genome duplication in vertebrate evolution (Ohno, 1970). Understanding the TGD and gene map locations showed that, for example, genes formerly called shh and twhh (Ungar & Moon, 1996) are actually TGD co-orthologs of the human SHH (sonic hedgehog) gene, and so are

I. Introduction

Hsa Chromosomes

(A) Hox cluster ohnologons in human X Y 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22

HOXD9

IHH

centromere

HOXA9

HOXC9

DHH

HOXB9

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20Mb

40Mb

60Mb

80Mb 100Mb Hsa7 Paralogs

(B) HOXA9 ohnologs in zebrafish

Dre Chromosomes

SHH

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25

Hsa7

0Mb

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shhb

shha

hoxa9b hoxa9a

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centromere 40Mb

(C) SHH ohnologons in zebrafish C1. Dre7

120Mb

nom1 dnajb6b wdr60 ube3crnf32 esyt2b mnx1 ncapg2 lmbr1 ncapg2 ptprn2

SLC4A2 CHPF2 C2. Hsa7 FASTK ABCF2 SMARCD3 dnajb6a rbm33b en2b htr5aa C3. Dre2 paxip1 shhb

60Mb

rbm33a htr5ab insig1 en2a

SHH 80Mb 100Mb Hsa7 Orthologs

120Mb

140Mb

smarcd3b abcf2b

shha

chpf2 PTPRN2 DNAJB6 INSIG1 RBM33 NOM1 UBE3C HTR5A EN2 RNF32 WDR60 PAXIP1

SHH LMBR1 MNX1

NCAPG2 ESYT2

slc4a2a fastk abcf2a smarcd3a

FIGURE 3.3 Evidence for genome duplication. (A) A dot plot of Hsa7 paralogs on other human chromosomes. Black dots represent genes along Hsa7; their paralogs on other chromosomes are plotted directly above or below the gene’s position on Hsa7. Genes on the left arm of Hsa7 have paralogs, mainly on Hsa2, Hsa12, and Hsa17, including HOX9 paralogy group genes (HOXA9, HOXB9, HOXC9, and HOXD9) in the four human HOX clusters, as would be expected after two rounds of whole-genome duplication, VGD1 and VGD2. Parts of the right arm of Hsa7 are also present in three or four copies, including three hedgehog gene paralogs, SHH, DHH, and IHH. (B) A dot plot of Hsa7 paralogs on zebrafish chromosomes. Black dots represent genes along Hsa7, at the bottom; their orthologs or paralogs on zebrafish chromosomes are plotted directly above the gene’s position on Hsa7. Genes surrounding HOXA9 on Hsa7 have zebrafish orthologs and coorthologs mainly on Dre16 and Dre19, including hoxa9a and hoxa9b in two of the eight zebrafish hox clusters, as would be expected after the TGD. Parts of the rest of Hsa7 also tend to be present in two copies, including two sonic hedgehog gene paralogs, shha and shhb. (C) Double conserved synteny in the zebrafish. C1: The portion of Dre7 containing shha. C2: A part of Hsa7 around SHH. C3: The neighborhood of shhb on Dre2. Lines connect orthologs.

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now known as ohnologs, shha, and shhb, respectively. This nomenclature scheme reflects history and connects zebrafish genetics more appropriately to human biology. Third, genes derived by tandem duplication have a “.1” or “.2” appended to the name, like alpi.1 and alpi.2 (Yang, Wandler, Postlethwait, & Guillemin, 2012). From a practical standpoint, gene names sometimes change over time; for example, the hedgehog genes discussed above; in addition, different authors sometimes use different names for the same gene. Thus, it is essential that, before zebrafish researchers assign a name to a new gene or change a gene name, they contact ZFIN (Zebrafish Information Network) at [email protected] to confirm appropriate usage. In addition, various software applications for RNA-seq or single-cell RNA-seq or whole-genome sequence annotation can use different sources to assign gene names, which can lead to data loss or misapplication. Fortunately, the ZFIN gene search function recognizes and displays gene synonyms. Tracking stable ENSEMBL Gene identifiers (e.g., ENSDARG00000068567 for shha) will likely improve a project’s long-term data integrity.

Chromosome Evolution After the Teleost Genome Duplication The chromosome neighborhood that includes the human SHH gene illustrates several general principles concerning chromosome evolution in the zebrafish lineage. First, the region shows double conserved synteny with zebrafish: two regions of the zebrafish genome (Fig. 3.1A and C, parts of Dre7 and Dre2) contain genes that are orthologs and co-orthologs of the human genes surrounding SHH (Fig. 3.1B). Second, several genes in the human segment are often present in duplicate on both zebrafish ohnologons, for example, besides duplicates of SHH, this region has duplicates of ABCF2, SMARCD3, HTR5A, EN2, and DNAJB6. On the other hand, many genes in the human region are present in a single copy on Dre7 and Dre2 due to gene loss after the TGD (Fig. 3.1). Third, chromosomal rearrangements, especially inversions and transpositions in one or both lineages, have caused gene orders to often differ in human and zebrafish genomes. For example, the orthologs of 11 genes at the left end of the chromosome segment shown in Fig. 3.1A are in the same order in humans but are transposed relative to three genes at the right end of the chromosome segment. Fourth, many human genes in the portion of Hsa7 shown in Fig. 3.1B that do not have orthologs in the portions of Dre2 and Dre7 shown in Fig. 3.1A and C have orthologs on other parts

of these two zebrafish chromosomes. Fifth, other genes in the SHH region of Hsa7 may be on zebrafish chromosomes other than Dre2 and Dre7 due to chromosomal translocations. Inversions and translocations were essential after the TGD to rediploidize the genome, to return a tetraploid genome to a diploid state. In a tetraploid, four homologous chromosomes (two pairs) form chiasma in prophase I of meiosis and segregate irregularly during anaphase I, thus, giving rise to aneuploid gametes, many of which can lead to lethal aneuploid offspring. Chromosome rearrangements that make the two pairs of original chromosomes fail to pair with each other would reduce recombination between them, and thus, increase the likelihood of forming euploid “diploid” gametes, which can lead to euploid progeny with twice as many chromosomes as before the TGD. In addition to local rearrangements by inversions and translocations, the zebrafish lineage experienced multichromosomal changes after its divergence from the spotted gar lineage. Spotted gar, whose lineage diverged from teleosts before the TGD, has 58 chromosomes, many of which are conserved intact with some tetrapods for 450 million years. If the last common ancestor of zebrafish and gar had a similar number of chromosomes, and the TGD doubled the number of chromosomes, then teleosts should have far more chromosomes than the 25 possessed by zebrafish and most other teleosts; evidently, chromosome fusions reduced chromosome numbers. A comparison of genes on chromosomes of gar, chicken, and teleosts suggests that these chromosome fusions occurred in the teleost lineage after it diverged from gar but, somewhat surprisingly, before the TGD (Braasch et al., 2016). Genome-wide chromosome fusions like this have occurred recently in some lineages (e.g., the Antarctic bullhead notothen (Amores, Wilson, Allard, Detrich, & Postlethwait, 2017)).

Gene Evolution After the Teleost Genome Duplication The zebrafish shh ohnologons (Fig. 3.3C) reflect the finding that fewer genes were retained as duplicates in the zebrafish genome after the TGD than relapsed to a single copy. Estimates suggest that about 20%e25% of genes in the preduplication ancestor are currently retained in duplicate (Braasch & Postlethwait, 2012). In contrast, the duplicate retention rate for miRNA genes is substantially higher, at about 40% (Braasch et al., 2016). After genome duplication, all regulatory elements and all protein-coding domains are present

I. Introduction

31

Gene Evolution After the Teleost Genome Duplication

in duplicate, causing the two genes to be totally redundant except for dosage issues (Fig. 3.4A and B). Tandem gene duplication events, on the other hand, are not as predictable, with some regulatory elements or coding domains at the ends of duplicated regions perhaps left uncopied. Gene loss, which can occur by mutation to a pseudogene, say by mutation to a premature stop codon (Fig. 3.4C, also called nonfunctionalization (Force et al., 1999)), is the most frequent fate of gene duplicates. To explain the retention of both duplicates, Susumu Ohno suggested that one copy preserves the original gene functions while the other copy assumes a new, positively selected function (Ohno, 1970). Allan Force suggested a revised framework for viewing the preservation of duplicated genes (Force et al., 1999). If one copy, say the “a” copy, retained all original functions and in addition evolved a new advantageous function, then the other copy, the “b” copy that simply preserved the original functions, would be redundant and would usually be lost. If both copies are to be permanently preserved after the origin of a new function in one of them, then the “a” copy would have to both evolve a new beneficial function (the star in Fig. 3.4D) AND lose an original, essential function that the “b” copy maintains (the pink regulatory element in Fig. 3.4D) so that the “a” copy is preserved by its new advantageous function, and the “b” copy persists because it possesses an essential ancestral function that the “a” copy lacks, a situation called duplicate gene preservation by neofunctionalization (Force et al., 1999). Alternatively, duplicated genes could be preserved by

the reciprocal loss of ancestral essential functions, where the “a” copy loses a function that the “b” copy maintains (the orange regulatory element in Fig. 3.4E), and reciprocally, the “b” copy loses a function that the “a” copy retains (the pink regulatory element in Fig. 3.4E), called duplicate gene preservation by subfunctionalization (Force et al., 1999). Subfunctionalization is important because it occurs solely by degenerative evolutiondthe reciprocal knockout of ancestral essential functionsdand does not require the “invention” of any previously nonexistent function as required by neofunctionalization (duplication, degeneration, complementation, DDC). Note that in all three types of duplicate gene evolution, additional functions can be lost or gained over time. For example, after nonfunctionalization, the entire gene with all of its coding domains and regulatory elements could become unrecognizable due to mutation. After neofunctionalization, all of the ancestral functions except the new positively selected function could be lost on one copy while the other copy could retain all ancestral functions. After subfunctionalization, either or both preserved copies could reciprocally lose additional ancestral functions or could gain new functions. Thus, it is important to distinguish between the initial force that preserves both duplicates, and subsequent evolutionary processes that can occur given that both copies still exist (Braasch, Bobe, Guiguen, & Postlethwait, 2018; Sandve, Rohlfs, & Hvidsten, 2018). Although the relative frequency of duplicate gene preservation by neofunctionalization versus subfunctionalization is not yet known, evidence suggests that

regulatory elements protein coding

(A) Ancestral gene

(B) After genome duplication

a b

(C) Nonfunctionalization

(D) Neofunctionalization

(E) Subfunctionalization

a

a

a

b

b

b

FIGURE 3.4 Duplication, degeneration, complementation model for the preservation of gene duplicates. (A) The ancestral, single-copy gene with regulatory elements and protein-coding domains. (B) Two identical gene copies after genome duplication. (C) Nonfunctionalization destroys gene activity, for example, by an early stop codon mutation. (D) Neofunctionalization involves the gain of a beneficial function (star) and loss of an ancestral essential function in one gene copy (gray circle) and the maintenance of that lost gene function in the other gene copy (pink circle) so that both copies are essential. (E) Subfunctionalization involves the reciprocal loss of ancestral functions (gray square and gray circle) so that the two gene copies complement each other, and both are essential for survival. Once preserved, gene copies can further evolve new functions or reciprocally lose additional functions. Functions can be regulatory elements or protein-coding domains.

I. Introduction

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3. Zebrafish Genetics

both mechanisms play an important role in zebrafish genetics. Some examples include en2a and en2b (Fig. 3.3C) (Force et al., 1999; Postlethwait, Amores, Cresko, Singer, & Yan, 2004) and atp1a1 paralogs (Serluca, Sidow, Mably, & Fishman, 2001), where TGD duplicates are expressed in alternative ancestral expression domains; mitfa and mitfb, where the “a” and “b” copies use different promotors (Lister, Close, & Raible, 2001); hematopoietic growth factor genes (Pazhakh & Lieschke, 2018); tlr5a and tlr5b (Voogdt, Wagenaar, & van Putten, 2018); fatty acidbinding protein genes (Laprairie, Denovan-Wright, & Wright, 2017), and many others (Pasquier et al., 2017). For zebrafish genetics, the consequence of the TGD followed by subfunctionalization is that researchers must examine both duplicates to draw valid inferences regarding the functions of the ancestral preduplication gene, and thus, to connect zebrafish research to human biology. The consequence of neofunctionalization as a mechanism for duplicate gene preservation is that one or both of the zebrafish gene ohnologs may possess novel roles not already present in the ancestral preduplication gene, and hence in humans.

The Zebrafish Genome Sequence Assembly With the accumulation of mutated genes to clone and the growing utility of zebrafish as an experimental model for vertebrate gene function, sequencing the zebrafish genome became a priority (Howe et al., 2013). Efforts to sequence the zebrafish genome from the TU strain began at the Wellcome Trust Sanger Institute (UK) in 2001 and culminated in the formal genome article published in 2013 (Howe et al., 2013). Initial forays sequenced DNAs extracted from about a 1000 TU individuals, a strategy intended to distinguish SNPs that were preexisting in the population from mutations chemically induced in this strain. The published genome assembly combined sequences from high-quality finished clone sequences and wholegenome shotgun sequences to assemble contigs and scaffolds that were then anchored onto the Sanger AB-TU meiotic map (SATmap) to achieve a chromosome-level assembly (Howe et al., 2013). The zebrafish genome assembly, currently in version GRCz11, is updated by the Genome Reference Consortium (https://www.ncbi.nlm.nih.gov/grc/ zebrafish) and the Alliance for Genome Resources (https://www.alliancegenome.org/). The zebrafish genome assembly can be accessed through multiple web-based browsers, such as ZFIN (http://zfin.org/ action/gbrowse/), Ensembl (http://www.ensembl. org/Danio_rerio/Info/Index), NCBI (https://www. ncbi.nlm.nih.gov/genome/gdv/?org¼danio-rerio), or UCSC (https://genome.ucsc.edu).

Major features of the zebrafish genome include its size, burden of repetitive sequences, genetic similarity to human genes, high rate of heterozygosity, and frequency of evolutionarily fixed chromosome rearrangements. With 1.674 GB (giga base pairs) in the assembly version GRCz11, the zebrafish genome is substantially larger than that of many teleosts, four times the fugu pufferfish genome (0.391 GB, version FUGU5) (Elgar, 1996), more than twice as large as medaka (0.734 GB, ASM223467v1) (Kasahara et al., 2007), and it reaches about half the size of the human genome (3.609 GB, GRCh38.p12) (Freeman et al., 2007). The bloating of the zebrafish genome is due to zebrafish-specific amplification of certain transposable elements, which occupy about 55% of the zebrafish genome (Chalopin, Naville, Plard, Galiana, & Volff, 2015; Howe et al., 2013). This huge content of repetitive DNA frustrated early zebrafish genome sequencing and assembly efforts. The current zebrafish genome annotation contains 25,592 coding genes and an additional 6599 noncoding genes, including 1046 microRNA genes (Desvignes, Beam, Batzel, Sydes, & Postlethwait, 2014). The GRCh38.p12 version of the human genome sequence contains just 20,465 annotated protein-coding genes, 20% fewer than zebrafish (25,592). The larger gene catalog in zebrafish might reflect the legacy of the teleost genome duplication, because many other well-studied teleost fish also have more annotated protein-coding genes than human (e.g., Mexican tetra, 26,698 (McGaugh et al., 2014) and medaka, 23,622 (Kasahara et al., 2007)), but spotted gar, which diverged from the teleost lineage before the TGD, has just 18,341 genes (LepOcu1 (Braasch et al., 2016)). About 70% of human genes have at least one ortholog in zebrafish, and reciprocally, about 70% of zebrafish genes have at least one ortholog in the human genome (Howe et al., 2013). About 47% of the human/zebrafish orthologs are one-to-one pairs. The finding that 70%e80% of human genes that have been associated with a disease or specific human phenotypes have an ortholog in zebrafish enhances the value of zebrafish as a model for human biology. Zebrafish has an exceedingly high rate of genetic polymorphisms. The sequencing of the two individual founders of the SATmap, each of which was completely homozygous at all loci, revealed about seven million SNPs, which is far greater than that seen between any two humans (Genomes Project et al., 2012; Howe et al., 2013). The sequence of a single zebrafish taken from the wild in northeastern India also had about seven million single-nucleotide and structural polymorphisms with respect to the reference genome sequence (Patowary et al., 2013). Chromosome rearrangements also impact zebrafish genetics. Some analysts had suggested that the zebrafish

I. Introduction

Genetics of Zebrafish Sex Determination

genome experienced more intrachromosomal rearrangements, such as inversions or transpositions, after the TGD than other fish lineages (Semon & Wolfe, 2007). After building synteny blocks using only genes with orthologs among all species tested (zebrafish, stickleback, medaka, green spotted pufferfish, and chicken, which have gene orders that are well-conserved from the ancestral stem bony vertebrate), zebrafish appears to have about the same intrachromosomal rearrangement rate as other teleosts: about 40% of genes that are neighbors in chicken have orthologs that are also neighbors in zebrafish, stickleback, medaka, and pufferfish (Howe et al., 2013). In contrast to intrachromosomal rearrangements, interchromosomal rearrangements, such as translocations and chromosome fissions and fusions, are more frequent in the zebrafish lineage than in some other teleost lineages. If no interchromosomal rearrangements had occurred after the TGD, then ohnolog pairs would occupy the same total number of chromosomes as twice the preduplication chromosome count. Each translocation or chromosome fission after the TGD, however, would increase the number of chromosomes with ohnolog pairs. Chromosome fusions, on the other hand, might decrease the number of chromosomes with ohnolog pairs, and the comparator species have fewer chromosomes than zebrafish, whose haploid number is 25 (stickleback, 21; medaka, 24; green spotted pufferfish, 21). Nevertheless, it takes more chromosomes in zebrafish to reach 1% of the total number of ohnologs than it does for any of the other species, consistent with more translocations and/or chromosome fusions in the zebrafish lineage than in the three percomorph fish tested (Howe et al., 2013). Despite this finding, most of the interchromosomal rearrangements observed comparing zebrafish to human are shared among teleosts; for example, contrast breaks in syntenies comparing Hsa7 to the zebrafish genome with breaks comparing Hsa7 to the medaka genome (Fig. 3.5). This result shows that these synteny breaks occurred before the divergence of the zebrafish and medaka lineages over 250 million years ago (Near et al., 2012).

Genetics of Zebrafish Sex Determination Zebrafish geneticists “using standard lines of zebrafish that have long been maintained in laboratories are often plagued by severe sex ratio distortions” (Lawrence, Ebersole, & Kesseli, 2008). This situation produces problems for researchers trying to mate specific rare genotypes and poses the interesting question of what causes an individual zebrafish to develop ovaries or testes. Laboratory strain zebrafish pass through a “juvenile ovary” phase, in which gonads form meiotic oocytes that die in

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individuals that become males, and survive in fish that become females (Rodriguez-Mari et al., 2010; Takahashi, 1977; Uchida, Yamashita, Kitano, & Iguchi, 2002; Wang, Bartfai, Sleptsova-Freidrich, & Orban, 2007). Animals containing gonads developing without germ cells become males (Siegfried & Nusslein-Volhard, 2008; Slanchev, Stebler, de la Cueva-Mendez, & Raz, 2005), and specifically, meiotic oocytes are required to maintain ovary differentiation (Dranow, Tucker, & Draper, 2013; Draper, McCallum, & Moens, 2007; Rodriguez-Mari et al., 2010, 2011), leading to the hypothesis that meiotic oocytes signal the gonadal support cells to maintain expression of aromatase (Kossack & Draper, 2019; Rodrı´guez-Marı´ & Postlethwait, 2011), the enzyme that converts testosterone to estrogen. Candidate signaling mechanisms include the FGF, BMP, and WNT signaling pathways (Dranow et al., 2016; Kossack et al., 2019; Leerberg, Sano, & Draper, 2017). Environmental factors can strongly affect the sex ratio in zebrafish, with harmful factors like gamma rays, hypoxia, high density, high temperature, altered thermocycles, and poor nutrition favoring male development (Abozaid, Wessels, & Horstgen-Schwark, 2012; Delomas & Dabrowski, 2018a, 2018b; Hosseini, Brenig, Tetens, & Sharifi, 2019; Liew et al., 2012; Ribas, Liew, et al., 2017; Ribas, Valdivieso, Diaz, & Piferrer, 2017; Santos, Luzio, & Coimbra, 2017; Shang, Yu, & Wu, 2006; Valdivieso, Ribas, & Piferrer, 2019; Villamizar, Ribas, Piferrer, Vera, & Sanchez-Vazquez, 2012; WalkerDurchanek, 1980). However, it is unlikely that zebrafish has an established environmental sex determination mechanism (ESD) like some fish and some sauropsids (reptiles) (Charnier, 1966; Conover, 1984; Lang & Andrews, 1994; Merchant-Larios & Diaz-Hernandez, 2013; Mork, Czerwinski, & Capel, 2014; OspinaAlvarez & Piferrer, 2008) because male and female zebrafish develop in tanks maintained at constant temperatures, and single pair matings show that zebrafish sex determination has a strong genetic basis (Liew et al., 2012). Nevertheless, the offspring of a single fully homozygous doubled haploid AB fish mated to a completely homozygous doubled haploid TU fish made genetically identical males and females, showing that genetic differences are not essential to make the two sexes in zebrafish (Howe et al., 2013). Several studies have attempted to isolate genetic markers linked to sex determination in zebrafish strains adapted to laboratories, but each identified different regions of the genome, often several regions that appear to be important in the same study (Anderson et al., 2012; Bradley et al., 2011; Howe et al., 2013; Liew et al., 2012; Liew & Orban, 2014; Orban, Sreenivasan, & Olsson, 2009; Siegfried, 2010; Tong, Hsu, & Chung, 2010; Wilson et al., 2014), suggesting a polygenic sex-determination system.

I. Introduction

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3. Zebrafish Genetics

(A)

Dre Chromosomes

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 Hsa7

(B)

Ola Chromosomes

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 Hsa7 0Mb

20Mb

40Mb

60Mb 80Mb Hsa7 Orthologs

100Mb

120Mb

140Mb

FIGURE 3.5 Most chromosome rearrangements occurred before the diversification of the largest group of teleosts. (A) A dot plot comparing human chromosome 7 to the zebrafish genome, with gene orders as in Hsa7. (B) A dot plot comparing Hsa7 to the medaka genome. Most breaks in synteny mapped onto the human chromosome are in the same place for both teleosts, as expected if most chromosome rearrangements occurred after the TGD but before the divergence of the medaka and zebrafish lineages more than 250 million years ago.

Two studies included zebrafish strains that had not been adapted to laboratory life using tens of thousands of DNA polymorphisms identified by sequencing the ends of restriction enzyme fragments (RAD-tags). These investigations identified a Sex-Associated Region on Dre4 (sar4) about 2 MB long at the distal tip of the long arm of chromosome Dre4 (Anderson et al., 2012; Wilson et al., 2014). Recall from Figs. 3.1 and 3.2 that Dre4 is a special chromosome, the only one with nearly an entire arm appearing as heterochromatic and late replicating, properties shared by many sex chromosomes, for example, the human Y chromosome. Identified polymorphisms showed that female zebrafish are heterogametic and male zebrafish tended to be homogametic, as expected for a WZ female/ZZ male chromosomal sex determination mechanism. Chromosomally, WZ

individuals sometimes developed as phenotypic males, but ZZ individuals never developed as females. This finding suggests that (1) the W chromosome contains a gene or allele that is necessary but not sufficient to cause a gonad to develop into an ovary, or (2) that two doses of the Z chromosome actively extinguish ovary development. The molecular genetic sex determining factor in sar4 has not yet been identified, and so we do not yet know how it works. Identifying the factor is stymied by two factors: the nature of sex chromosomes and the animals used for the zebrafish reference genome sequence. The long right arm of chromosome Dre4 is late replicating and largely heterochromatic (Fig. 3.1) and has a plethora of repetitive elements (Howe et al., 2013). These repetitive regions have thwarted the assembly of the

I. Introduction

References

sequenced genome around sar4. In addition, the reference genome sequence comes from the AB and TU strains of fish (Howe et al., 2013), which appear to have lost sex-specific genetic polymorphisms. The AB strain was produced from multiple rounds of gynogenesis (Walker-Durchanek, 1980), where offspring retain chromosomes only from the mother, each individual is homozygous for all loci, and gynogens tend to develop as males, presumably due to the deleterious effects of inbreeding (Delomas & Dabrowski, 2018a). The TU strain was developed to be lethal free by multiple single-pair matings (Mullins, Hammerschmidt, Haffter, & Nusslein-Volhard, 1994). In both cases, the accidental use of WZ females mated to sex-reversed WZ males could produce WW individuals, which can develop into either females or males (Wilson et al., 2014). Thus, by chance, both AB and TU strains could have drifted to become WW and still produce both males and females. If the sequenced genome comes from chromosomally WW fish, then it will lack a polymorphism for the primary genetic sex-determining mechanism. Over the more than two decades of laboratory domestication, the AB and TU strains, due to “natural” selection imposed by zebrafish geneticists, apparently evolved alternative environmental or genetic factors that increase the likelihood that males can develop, and these alleles are likely the weak factors identified by some genetic studies (Anderson et al., 2012; Bradley et al., 2011; Howe et al., 2013; Liew et al., 2012). Identifying the molecular genetic nature of the sex determinant in sar4 should be a research priority of zebrafish geneticists.

Conclusions Zebrafish provides a remarkably helpful system to investigate the molecular, cellular, and genetic mechanisms underlying a wide variety of processes common to all vertebrates. To fully leverage the zebrafish system, we must understand its genome, what it contains, and how it got here. The legacy of the teleost genome duplication and the partitioning of ancestral vertebrate gene functions between retained ohnologs provides a natural experiment useful for dissecting how the genome functions. Coupled with modern ways to study chromatin modifications and gene expression in single cells, zebrafish genetics will continue to contribute to understanding how animals with backbones develop, function, and evolve.

Acknowledgments The authors thank NIH grant R01 OD011116 for support.

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I. Introduction

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Takahashi, H. (1977). Juvenile hermaphroditism in the zebrafish, Brachydanio rerio. Bulletin of the Faculty of Fisheries Hokkaido University, 28, 57e65. Taylor, J. S., Braasch, I., Frickey, T., Meyer, A., & Van de Peer, Y. (2003). Genome duplication, a trait shared by 22000 species of ray-finned fish. Genome Research, 13(3), 382e390. https://doi.org/10.1101/ gr.640303. Taylor, J. S., Van de Peer, Y., Braasch, I., & Meyer, A. (2001). Comparative genomics provides evidence for an ancient genome duplication event in fish. Philosophical Transactions of the Royal Society of London B Biological Sciences, 356(1414), 1661e1679. https://doi.org/10.1098/ rstb.2001.0975. Tong, S. K., Hsu, H. J., & Chung, B. C. (2010). Zebrafish monosex population reveals female dominance in sex determination and earliest events of gonad differentiation. Developmental Biology, 344(2), 849e856. https://doi.org/10.1016/j.ydbio.2010.05.515. Traut, W., & Winking, H. (2001). Meiotic chromosomes and stages of sex chromosome evolution in fish: Zebrafish, platyfish and guppy. Chromosome Research, 9(8), 659e672. Retrieved from https://www.ncbi.nlm.nih.gov/pubmed/11778689. Uchida, D., Yamashita, M., Kitano, T., & Iguchi, T. (2002). Oocyte apoptosis during the transition from ovary-like tissue to testes during sex differentiation of juvenile zebrafish. Journal of Experimental Biology, 205(Pt 6), 711e718. Retrieved from http://www.ncbi.nlm. nih.gov/entrez/query.fcgi? cmd¼Retrieve&db¼PubMed&dopt¼Citation&list_ uids¼11914381. Ungar, A. R., & Moon, R. T. (1996). Inhibition of protein kinase A phenocopies ectopic expression of hedgehog in the CNS of wild-type and cyclops mutant embryos. Developmental Biology, 178(1), 186e191. https://doi.org/10.1006/dbio.1996.0209. Valdivieso, A., Ribas, L., & Piferrer, F. (2019). Ovarian transcriptomic signatures of zebrafish females resistant to different environmental perturbations. Journal of Experimental Zoology Part B: Molecular and Developmental Evolution, 332(3e4), 55e68. https://doi.org/ 10.1002/jez.b.22848. Varga, M. (2018). The doctor of delayed publications: The remarkable life of George Streisinger (1927-1984). Zebrafish, 15(3), 314e319. https://doi.org/10.1089/zeb.2017.1531. Vialle, R. A., de Souza, J. E. S., Lopes, K. P., Teixeira, D. G., Alves Sobrinho, P. A., Ribeiro-Dos-Santos, A. M., et al. (2018). Whole genome sequencing of the pirarucu (Arapaima gigas) supports

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independent emergence of major teleost clades. Genome Biology and Evolution, 10(9), 2366e2379. https://doi.org/10.1093/gbe/evy130. Villamizar, N., Ribas, L., Piferrer, F., Vera, L. M., & SanchezVazquez, F. J. (2012). Impact of daily thermocycles on hatching rhythms, larval performance and sex differentiation of zebrafish. PLoS One, 7(12). https://doi.org/10.1371/journal.pone.0052153. e52153. Voogdt, C. G. P., Wagenaar, J. A., & van Putten, J. P. M. (2018). Duplicated TLR5 of zebrafish functions as a heterodimeric receptor. Proceedings of the National Academy of Sciences of the United States of America, 115(14), E3221eE3229. https://doi.org/10.1073/ pnas.1719245115. Walker-Durchanek, R. C. (1980). Induction of germ line mutations by gamma-irradiation of zebrafish embryos (MS). Eugene, Oregon, USA: University of Oregon. Walker, C., & Streisinger, G. (1983). Induction of mutations by gammarays in pregonial germ cells of zebrafish embryos. Genetics, 103(1), 125e136. Retrieved from https://www.ncbi.nlm.nih.gov/ pubmed/17246099. Wang, X., Bartfai, R., Sleptsova-Freidrich, I., & Orban, L. (2007). The timing and extent of ‘juvenile ovary’ phase are highly variable during zebrafish testis differentiation. Journal of Fish Biology, 70, 33e44. Wilson, C. A., High, S. K., McCluskey, B. M., Amores, A., Yan, Y. L., Titus, T. A., et al. (2014). Wild sex in zebrafish: Loss of the natural sex determinant in domesticated strains. Genetics, 198(3), 1291e1308. https://doi.org/10.1534/genetics.114.169284. Woltering, J. M., & Durston, A. J. (2006). The zebrafish hoxDb cluster has been reduced to a single microRNA. Nature Genetics, 38(6), 601e602. https://doi.org/10.1038/ng0606-601. Woods, I. G., Kelly, P. D., Chu, F., Ngo-Hazelett, P., Yan, Y. L., Huang, H., et al. (2000). A comparative map of the zebrafish genome. Genome Research, 10(12), 1903e1914. Retrieved from http://www.ncbi.nlm.nih.gov/entrez/query.fcgi? cmd¼Retrieve&db¼PubMed&dopt¼Citation&list_uids¼11116086. Woods, I. G., Wilson, C., Friedlander, B., Chang, P., Reyes, D. K., Nix, R., et al. (2005). The zebrafish gene map defines ancestral vertebrate chromosomes. Genome Research, 15(9), 1307e1314. doi: gr.4134305 [pii] 10.1101/gr.4134305. Yang, Y., Wandler, A. M., Postlethwait, J. H., & Guillemin, K. (2012). Dynamic evolution of the LPS-detoxifying enzyme intestinal alkaline phosphatase in zebrafish and other vertebrates. Frontiers in Immunology, 3, 314. https://doi.org/10.3389/fimmu.2012.00314.

I. Introduction

C H A P T E R

4 Geographic Range and Natural Distribution Carole J. Lee, Charles R. Tyler, Gregory C. Paull Department of Biosciences, University of Exeter, Exeter, Devon, United Kingdom range of freshwater fish species (Jackson, Peres-Neto & Olden, 2001). Biotic factors, such as predation and competition, are also influential (Gaston, 2003). Determining the geographic range of a species can be tricky. In fact, it is usually impractical, in terms of time and budget, for a single researcher to conduct a complete survey across a large geographical area, and so data from a variety of sources, such as biological surveys, scientific reports, and museum specimens, are often relied on. The accuracy of this information can vary and without careful scrutiny errors in identification or taxonomy may perpetuate over time (Kottelat & Freyhof, 2007).

Geographic range and natural distribution are two of the fundamental characteristics of a species (Gaston, 2003). This chapter provides a broad overview of the range and distribution of zebrafish. It describes the typical habitats of wild zebrafish, the biotic and abiotic factors that determine range limits, and the potential implications of environmental change and human interference. Knowledge on wild zebrafish populations and the environment in which they live is vital for understanding the natural genetic and phenotypic variation, interpreting observations on their morphology, physiology, and behavior in the laboratory, and in designing and refining optimum housing systems and husbandry practices.

Geographic Range of the Zebrafish Geographic Range

The geographic range of the zebrafish extends across much of India, Bangladesh, and Nepal, from the Pakistan border in the west to the Myanmar border in the east, and from the foothills of the Himalaya in the north to the paddy fields of Karnataka in the south (Fig. 4.1; Engeszer, Patterson, Rao & Parichy, 2007). Geographically, this area comprises three major regions: the northern mountains, the Indo-Gangetic Plain, and the Indian peninsula. The northern mountains comprise the Himalaya and Hindu Kush ranges, which run from east to west in an arc across the entire northern boundary of India. South of the mountains, the IndoGangetic Plain encompasses the vast, fertile floodplains of the Indus, Ganges, and Brahmaputra rivers. These floodplains comprise most of northern India and almost all of Bangladesh. To the west lies the desert of Rajasthan, while to the south is the Indian peninsula, an immense plateau flanked by the coastal mountains of the Western and Eastern Ghats, which projects into the Indian Ocean (Thapar, 2004). As with most species, the zebrafish is not evenly distributed throughout its range. Reliable first-hand reports document populations in the streams and

The geographic range of a species comprises the regions in which that species can be found. The geographic ranges of fish species vary in size enormously. At one end of the spectrum are those whose distribution extends the entire world across tropical or temperate waters, such as the swordfish (Xiphias gladius) and the basking shark (Cetorhinus maximus) (Gaither, Bowen, Rocha & Briggs, 2016). At the other end of the spectrum are endemic species confined to a restricted area, such as the gizani (Ladigesocypris ghigii), which lives only in streams on the Greek island of Rhodes (Stoumboudi, Barbieri, Mamuris, Corsini-Foka & Economou, 2002), and the Quitobaquito pupfish (Cyprinodon macularius eremus), which is found in a single spring outflow in the Arizona desert (Douglas, Douglas & Brunner, 2001). The factors that limit geographic ranges are so varied and the interactions among them so complex that they are not fully understood for any single species (Gaston, 2009). Climate (especially temperature) and water chemistry (including levels of dissolved oxygen and acidity) play an important role in determining the geographic The Zebrafish in Biomedical Research https://doi.org/10.1016/B978-0-12-812431-4.00004-X

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© 2020 Elsevier Inc. All rights reserved.

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4. Geographic Range and Natural Distribution

FIGURE 4.1 The geographic range of the zebrafish comprises the foothills of the northern mountains, the Indo-Gangetic Plain, and the Indian peninsula.

FIGURE 4.2 Wild zebrafish found in Bangladesh. Photo by Gregory Paull.

tributaries of rivers in the Himalaya drainage system of Nepal (Dhital & Jha, 2002), the floodplains of the Ganges and Brahmaputra rivers in India and Bangladesh (Fig. 4.2; Spence et al., 2006; Engeszer et al., 2007; Arunachalam, Raja, Vijayakumar, Malaiammal & Mayden,

2013), and along the Western Ghats (Dahanukar, Raghavan, Ali, Abraham & Shaji, 2011). Compilations of older records report specimens found in Rajasthan (Datta & Majumdar, 1970) and in the peninsula’s southeast region (Menon, 1999). No systematic field studies have mapped the occurrence of the zebrafish throughout its range, and it may be missed from species surveys due to its small size and lack of commercial value (Spence et al., 2008). As a result, zebrafish may be more widely distributed than previously thought. It should be recognized that the geographic range of any species is dynamic. It expands, contracts, or shifts over time in response to changes in climate or physical geography (Thomas, 2010). The ability of the zebrafish to expand or move its range boundaries in response to environmental change is constrained by the physical barriers of the northern mountain ranges to the north, east and west, and by the Indian Ocean to the south. Within its range boundaries, the dispersal of zebrafish to new areas in response to changing conditions is limited to the floodplains and drainage basins that it inhabits and is influenced by interactions with other species (predators and competitors) whose abundance

I. Introduction

43

Natural Distribution

and diversity are also affected by climate (Thomas, 2010). Human activities, such as farming, flooding arable land to create paddy fields, and connecting previously unconnected water bodies with drainage ditches, may also influence the dispersal of zebrafish.

Home Range Most animals live within a home rangedan area that contains the resources that they need to avoid predation, find food, and reproduce (Welsh, Goatley & Bellwood, 2013). Usually, an animal’s resource requirements change as it grows and matures, and therefore the size of its home range changes with ontogeny. With fish, the smaller the animal, the higher the risk of predation (Miller, Crowder, Rice & Marschall, 1988). A young fish may restrict its home range and avoid risky locations to avoid predators. As it grows, the young fish’s vulnerability to predation decreases and its food requirement increases, allowing it to expand its home range. Finally, as the fish matures, its home range must be big enough to allow it find a mate and reproduce (Welsh et al., 2013). For example, gold-spot mullet (Liza argentea) spawn in coastal waters and the lower reaches of estuaries in tropical and temperate regions (Kendall & Gray, 2008). Larval mullet migrate into the estuaries and seek protection in dense vegetation or in mangrove forests where a labyrinth of tangled roots hampers the movements of large predators (Laegdsgaard & Johnson, 2001). As they grow, juvenile mullet develop greater mobility and can more easily escape from predators. This enables them to move to adjacent habitats, such as seagrass beds or mudflats, and then to open estuarine habitats where they feed on larger prey items before joining adult populations in deep waters (Laegdsgaard & Johnson, 2001). Little is known about the average size of a zebrafish home range. Studying the home range of fish in the low visibility of vegetated habitats or turbid waters is challenging, especially with a diminutive, highly mobile species such as the zebrafish. Tagging individual fish to radiotrack their movements has been used successfully to study larger species (Cooke et al., 2013) but has not been attempted with wild zebrafish. However, recent advances in technology have resulted in smaller, more efficient tags and passive integrated transponders that increase the potential to track smaller fish species and juvenile life stages (Cooke et al., 2016). Innovations continue, and the home range of the zebrafish should soon be testable. As water levels rise at the start of the monsoon, zebrafish are thought to migrate laterally from rivers, streams, and irrigation channels where they spend the dry months to flooded areas such as paddy fields where they spawn in the nutrient-rich waters. When water levels recede at the end of the monsoon, adults and

young-of-the-year migrate out of the floodplains and into rivers, streams, and channels (Engeszer et al., 2007). Little is known about this lateral migration, and the distances covered have not been measured. Zebrafish home range size therefore varies seasonally and may differ in streams versus floodplains due to the correlation of home range size with food supply (Minns, 1995). The spatial behavior of zebrafish at all life stages and throughout the seasons needs to be assessed to understand the extent of their home-ranging behavior.

Natural Distribution The terms geographic range and natural distribution are sometimes used interchangeably, but here we define natural distribution as “the specific areas, within its geographic range, in which a species occurs.” For example, the geographic range of the zebrafish includes the Western Ghats of India. Within the Western Ghats, the natural distribution of the species includes the Thunga River in the state of Karnataka and the Kabini River in the state of Kerala where populations of zebrafish have been found (Arunachalam et al., 2013). A species’ natural distribution may change over time, regardless of whether its geographic range moves, as habitats become more or less favorable due to changes in the physical or biological environment (Lawton, 1996). Much of the zebrafish’s natural distribution lies within the extensive floodplains of the Ganges and Brahmaputra rivers. Indeed, the zebrafish has been described as a “floodplain species” (Spence et al., 2008) and seems well adapted to constantly changing environmental conditions in the floodplains. Typical habitats of zebrafish include streams, drainage ditches, and ponds, often adjacent to rice paddies (McClure, McIntyre & McCune, 2006; Spence et al., 2006). In addition, Engeszer et al. (2007) and Arunachalam et al. (2013) found zebrafish in secondary and tertiary channels of large alluvial rivers. Together, these field studies report a range of habitat features in the specific locations where zebrafish were found. Some areas have still waters while others have slow to medium water flow. Water clarity ranges from clear to muddy, and substrates are recorded from silt at one end of the spectrum, through sand, gravel, pebbles, and boulders, to bedrock at the other end of the spectrum. Vegetation ranges from none at all to an abundance of submerged plants, with or without overhanging canopy. Some of these habitats are permanent while others are transitory and form during the monsoon. In summary, the zebrafish can be found in a diverse array of habitats throughout its huge geographic range; however, the geographic, climatic, and temporal factors that influence the distribution of zebrafish and the endogenous and exogenous pressures that drive them to occupy certain habitats are not well understood.

I. Introduction

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4. Geographic Range and Natural Distribution

Floodplains Floodplains are areas that are periodically inundated by the lateral overflow of rivers or lakes and/or by direct precipitation (Junk, Bayley & Sparks, 1989). They contain a medley of habitats and support diverse biotic communities. The floodplains of northern India and Bangladesh flood seasonally from June to October in response to monsoon rains and snow melt (Craig, Halls, Barr & Bean, 2004). Most of this vast region has been converted to agricultural land for the production of rice and other crops while fishing, by professional and subsistence fishers, contributes to floodplain livelihoods (Hoggarth et al., 1999). During monsoon rains, the floodplains are inundated with nutrient-rich waters, causing rapid growth of macrophytes and periphyton, increased concentrations of invertebrates, and an accumulation of detritus from vegetation and phytoplankton. Fish follow the advancing water’s edge and flooded areas become important food-rich nurseries for larval and juvenile fish. When water levels drop, vegetation becomes stranded, the food supply decreases, and surviving adult and young-of-the-year fish of many species migrate from the floodplains to channels, rivers, and other permanent water bodies (Bayley, 1988). Zebrafish are believed to spawn in shallow waters during the monsoon, and their opportunistic life history strategy of small body size, rapid growth, early maturity, and high fecundity allows them to reproduce and disperse while conditions are favorable and food is plentiful. Floodplain fishes rarely survive for more than a year (De Graaf, 2003), and field observations of wild zebrafish support this assumption (Spence, Fatema, Ellis, Ahmed & Smith,2007). Within the floodplains are a mosaic of habitat types, including major rivers and their secondary channels, canals, permanent lakes (baors), floodplain depressions (beels), excavated fish pits (kuas), and household ponds (mathels) (Hoggarth et al., 1999). Field studies have reported populations of zebrafish in most of these habitats.

Rivers The two major floodplain rivers within the zebrafish’s natural distribution are the Ganges and the Brahmaputra. Bedload sediments of sand and silt roll along the bottom of the main channels of both rivers, forming a sequence of dunes up to 6 m high, which slowly move along with the current (Garzanti et al., 2010). These shifting substrates are unfavorable for aquatic plants and invertebrates. As a result, vegetation tends to fringe the main channel borders and side channels where substrates are more stable (De Graaf, 2003). Fish abundance is highest at the channel borders where plant beds create diverse habitats and food is more plentiful (Junk, Bayley & Sparks, 1989).

FIGURE 4.3 Zebrafish have been found in tributaries of major rivers such as the Brahmaputra but have not been reported in the main rivers themselves. Photo by Gregory Paull.

Zebrafish have been captured in the tributaries and drainages of the Ganges and Brahmaputra, but none have been reported in the main rivers themselves (Fig. 4.3). Elsewhere in India, Arunachalam et al. (2013) collected zebrafish in a side channel of the Dikrong River, a major tributary of the Brahmaputra, and in a secondary channel of the Thunga River in the Western Ghats. During a field trip to investigate the habitats of wild zebrafish, Engeszer et al. (2007) traveled to northeast India and found populations in the Ghotimari River and in a tributary of the Jorai River, both in West Bengal, and further populations in the Dukan River in the hills of Meghalaya. The absence of zebrafish in major rivers is likely due to the effects of predation and competition or a lack of suitable food resources or spawning areas. Predators in rivers and streams affect habitat choice of prey species such as zebrafish, which tend to move into shallow waters or complex habitats or leave the site to avoid predators (Jackson, Peres-Neto & Olden, 2001). The effects of interspecific competition on river fish communities may also play a role in the absence of zebrafish from major rivers, although this has not been tested in the field.

Streams Stream types range from perennial snow- or monsoon-fed hill streams that drain the northern mountains and the Western Ghats to seasonal nalas that develop during the monsoon. Zebrafish are absent from fast-flowing hill streams but have been found in slower streams with shallow waters (Fig. 4.4). They appear to have restricted ranges of current velocity and water depth but tolerate variety in other habitat

I. Introduction

Natural Distribution

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FIGURE 4.4 Zebrafish have been found in slow streams with shallow waters and abundant vegetation. Photo by Gregory Paull.

FIGURE 4.5 Sampling zebrafish in a shallow pond in Bangladesh. Photo by Gregory Paull.

features, such as substrate type and the presence or absence of submerged vegetation. A detailed account of the stream habitat preferences of zebrafish is given in a field report by Arunachalam et al. (2013) who collected individuals from 11 streams in the Western Ghats and northern and northeastern states of India. The team found zebrafish in stream areas with overhanging vegetation or undercut banks and in places where alcoves or shallow pools had been created by bedrock or large boulders. Substrates varied from silt and sand to cobbles and bedrock, but all the habitats had low water flow (3.5e13.9 cm/s) and shallow depth (300 L per minute). Bag vessels are equipped with pressure gauges to notify technicians that bags need to be cleaned or replaced. Pressurized bag filters located just prior to UV disinfection equipment will greatly increase the efficiency of UV disinfection as water clarity (absence of suspended “fine” solids) is one of the most important factors in efficient UV disinfection (Malone, 2013). Pressurized bag filters have small footprints and high solids removal rates but can be labor-intensive, require consumables, and have a high water pressure requirement (Table 30.1). For smaller applications with lower flow rates, cartridge filters can be used. Many of the standalone zebrafish racks have integrated cartridge filters prior to chemical filters, and UV disinfection equipment for reasons stated above. Cartridge filters are available in a wide variety of particle removal sizes from 20 to 50 mm and can be comprised of stainless steel mesh to reduce consumable costs. Cartridge filter canisters are also equipped with pressure gauges to notify technicians that cartridges need to be cleaned or changed. Cartridge filters are also very good at removing the smallest suspended “fine” solids from system water (Table 30.1) (Figs. 30.5 and 30.6).

Foam fractionators, also called protein skimmers, use air bubbles and organic surfactants to create stable foams that remove suspended “fine” solids from culture water (Malone, 2013; Timmons & Ebeling, 2013). Surfactants are chemicals that have a hydrophobic “waterhating” portion and a hydrophilic “water-loving” portion (Timmons & Ebeling, 2013). Surfactants form when proteins are degraded in the system water. Protein surfactants that come into contact with air bubbles orient the charged hydrophilic portion of the molecules outward where they attract oppositely charged molecules in the water (suspended “fine” solids in particular) and trap them, creating a stable foam that overflows into a collection cup for disposal. The amount of skimming accomplished depends on the amount of surfactant and the amount of surface tension in the water. While foam fractionation is very effective in marine systems due to the high surface tension of the water, it is more erratic in freshwater systems and will probably have only marginal significance for freshwater zebrafish culture systems (Malone, 2013). This author is not aware of any zebrafish facilities that currently use foam fractionation as part of the filtration process though it has been discussed.

Biological Filtration Biological filtration is defined as the process by which dissolved wastes are removed from aquaculture water by bacterial action (Malone, 2013). The term biological

III. Husbandry

Biological Filtration

filtration, in the literature, most frequently refers to the process of converting dissolved nitrogenous wastes, such as toxic ammonia (NH3) and nitrite (NO 2 ) to less toxic nitrate (NO 3 ). Often overlooked are the important processes of mineralization by the heterotrophic microbial communities also found in these filters (Malone, 2013; Blacheton et al. 2013). Heterotrophic microbial communities within biofilters rapidly and efficiently mineralize a wide variety of dissolved organic materials, including carbohydrates, fats, and proteins (Malone, 2013; Schreier, Mirzoyan, & Keiko, 2010; Blacheton et al. 2013). Biological filters house the greatest variety and number of microbial organisms in recirculating systems (Rurangwa & Verdegem, 2015). Schreier et al. (2010) provides a list of the microbial diversity found in the biofilters of recirculating systems. The management of these microbial communities is reviewed in Rurangwa and Verdegem (2015). The process of nitrification is most important to the discussion of biological filtration for zebrafish husbandry. Two different groups of aerobic nitrifying bacteria drive this process: (1) the ammonia-oxidizing bacteria (AOB) that convert ammonia (NH3) to nitrite (NO 2) and (2) the nitrite-oxidizing bacteria (NOB) that convert nitrite to nitrate (NO 3 ). Ammonia production comes ultimately from the feed either as the nitrogenous waste from protein metabolism or from the mineralization of solid organic wastes, such as feces or uneaten feed, by heterotrophic microbial communities (Blancheton et al. 2013; Malone, 2013; Rurangwa & Verdegem, 2015).

FIGURE 30.6 Two fluidized sand filters. McWane Science Center.

Picture courtesy of

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Ammonia comes in two forms that exist in equilibrium and that are collectively referred to as total ammonia nitrogen (TAN). The optimal water quality conditions for the growth of AOB and NOB are generally a pH between seven to eight, alkalinity of 50e100 ppm (mg/ L), dissolved oxygen (DO) levels near saturation (6e9 ppm), and low concentrations of ammonia and nitrite. For detailed information regarding TAN, nitrite, nitrate, and nitrification, please refer to chapter 29 entitled Water Quality for Zebrafish Culture in this book. There are two basic types of biological filters found in aquaculture. Suspended growth biofilters grow suspended bacteria in the same tank as the culture organisms, and fixed film biofilters circulate treatment water in close proximity to bacteria attached to media in a location separate from the culture tank (Malone, 2013). Both biofilter types rely on the diffusion of dissolved organic materials for either mineralization or nitrification to occur. For this reason, both aeration and water circulation within biofilters is of paramount importance to optimize processes. Fixed film biofilters are the overwhelming favorite for most aquaculture applications and are the only method of biological filtration currently used for the culture of zebrafish in the research environment. Characteristics that influence the effectiveness of biological filtration include: the size and volume of the biofilter, the circulation of the water through the media, the retention time of the water in the biofilter, the water quality conditions within the biofilter, and the type of media used (Malone, 2013; Rurangwa & Verdegem, 2015; Timmons & Ebeling, 2013). It is currently presumed that the ammonia movement into biofilms is the single most important factor affecting the rate of biofiltration (Malone, 2013). The aquaculture industry is placing a strong emphasis on filter types that maximize the surface area of biofilms available to oxidize ammonia (Malone, 2013). Fixed film biofilters use a wide variety of media types with high surface area to volume ratios (Table 30.2). Several characteristics are considered when media are selected for biofilters (Timmons & Ebeling, 2013): (1) The void volume of the media. Media with larger open spaces among the surface area for bacterial attachment encourage water circulation, oxygen transfer, and reduce potential clogging. (2) The specific surface area of the media per unit of volume (ft2 per ft3 or m2 per m3). The greater surface area provides greater space for bacterial growth and more TAN removal per unit of media volume. (3) The TAN conversion rate of the media type (area and volume) as grams of TAN converted per ft2 or m2 of media per day or grams of TAN converted per ft3 or m3 of media per day.

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30. Recirculating Aquaculture Systems (RAS) for Zebrafish Culture

(4) The cross-sectional area and hydraulic loading rate. These are the area of media directed at the water flow (ft2 or m2) and the volume of water moved through the biofilter per unit of cross-sectional area per unit of time (gallons per ft2 per day, or m3 per m2 per day) respectively. Different media types and the characteristics of each are shown in Table 30.2. A large number of different biofilter types are commercially available for use with centralized filter systems in zebrafish culture facilities. There is less variety found in the stand-alone rack systems provided by the commercial suppliers. The ideal biofilter would have a maximal surface area, remove 100% of toxic ammonia and nitrite, maximize DO transfer, have a small footprint, use inexpensive media, have low pumping requirements, require little maintenance, and not capture solids (Timmons & Ebeling, 2013). Such a filter does not exist. There are many types of filters that can be used, and this chapter will focus on those types best suited for the culture of zebrafish used in research.

Moving Bed Bioreactors The moving bed bioreactors (MBBR) use 6e13 mm plastic media with moderate void volumes and protected internal surface areas (Kaldnes K1 media) for

the attachment of bacteria (Malone, 2013; Timmons & Ebeling, 2013). Media typically occupies 50%e70% of the reactor volume and is contained within the reactor by the screened pipe at the water outlet (Timmons & Ebeling, 2013). The MBBR is continuously and vigorously aerated, creating maximal water circulation for maximal diffusion of dissolved oxygen (DO) and nitrogenous wastes (Malone, 2013). These filters have only about ¼ to ½ the nitrification capacity of fluidized sand biofilters but feature high levels of aeration and CO2 removal (Malone, 2013). Guerdat et al. (2010) researched an MBBR from an intensive warm-water tilapia aquaculture system and observed a TAN removal rate of 267 g TAN/m3/day and a modest increase in DO as water exited the MBBR. MBBRs have a small footprint, low maintenance requirements, no requirement for backwash, a low construction cost, and are conceptually simple (Malone, 2013; Timmons & Ebeling, 2013). These filters cause moderate shearing of biofilms as media collide with each other inside the reactor (Timmons & Ebeling, 2013). MBBR-like biofilters are found in the sumps of several commercially available zebrafish racks, such as the Aquarius System stand-alone rack by Aquatic Enterprises and the Z-Hab System by Pentair Aquatic Eco-Systems. The advantages and disadvantages of MBBR are summarized in Table 30.3.

TABLE 30.2 Biological filter media examples provided with a brief description, the approximate size of the media, specific surface areas m2/m3, relative void volume, and approximate TAN conversion rates for warm-water aquaculture systems. Media

Description

Size

Surface area m2 per m3

Void volume

TAN conversion rate

a

a

Bio-Balls

Plastic spheres

3.8 cm

322

High

1e2 g/m2/dayd

Bio-Ballsa

Plastic spheres

2.5 cm

526a

High

1e2 g/m2/dayd

Bio-Filla

PVC ribbon

Ribbon

822a

Medium

No reference

a

Bio-Strata

Plastic blocks

Block

223e361

Medium

1e2 g/m2/dayd

Bio-Barrelsa

Plastic cylinders

2.5 cm

210a

High

1e2 g/m2/dayd

Kaldnes K1d

Plastic cylinders with protected surface area

1  0.7 cm

500d

High

267 g/m3/daye

Standard beadb

Spherical plastic beads

3e5 mm

1150e1475b

Low

380e518 g/m3/dayf

Enhanced nitrificationb

Pressed plastic beads

8e10 mm

1150e1475b

Low

586 g/m3/daye

Sandc

Sand or silica glass of various sizes

0.1 e 0.3 mmc

4000-20,000 m2/m3c

Low

667 g/m3/daye

a

a

Values derived from the Pentair Aquatic Eco-Systems 2016 Master Catalog. Values from Malone, Chitta, and Drennan (1993). c Values from Summerfelt 2006. d Values from Timmons and Ebeling (2013) for Kaldnes K1 media surface area and TAN removal rates for trickle filter media with specific surface areas from 100 to 300 m2/m3 e Observed values from Guerdat, Losordo, Classen, Osborne, and Delong (2010) for tilapia grown under intensive conditions at average temperature of 29 C. f Observed values for TAN removal rates from Sastry, DeLosReyes, Rusch, and Malone (1999) for tilapia and bubble-wash bead filters that were performing both mechanical and biological filtration. b

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Biological Filtration

TABLE 30.3

Comparison of biological filter types (biofilters) used for the aquaculture of zebrafish for research with the advantages and disadvantages of each filter type.

Biofilter

Advantages

Disadvantages

Moving bed bioreactors (MBBR) Media: Bioballs, biobarrels, kaldnes K1

Aeration increases DO Good CO2 removal Small footprint Low maintenance No backwash needed Low-cost construction

Nitrification rates are ¼ to ½ of fluidized sand filters Surface area of media is about ½ that of fluidized sand filters Not good removal of excess biofilms Requires good water circulation

Fluidized sand filters (FS) Media: Sand of various sizes Silica glass beads

Highest surface area Highest TAN removal rate Small footprint Highest biological loads Excellent transfer of DO and TAN Inexpensive media No backwash or water loss

High DO consumption High CO2 production Biofouling of sand media and bed growth High pumping costs Backup pumps and power necessity Heavy media Sand beds go anoxic quickly with loss of water flow

Floating bead filters (FB) Media: Floating plastic beads Standard beads or enhanced nitrification “EN” beads

High TAN removal rates High surface area media Light-weight media Easy to install and operate Small footprint Backwash removes excessive biofouling Mechanical and biological filtration Low pumping requirement Low water loss on the backwash

DO is consumed CO2 produced Relatively high head loss that can be variable Require frequent backwashing to work optimally If used for both mechanical and biological filtration nitrification rates are lower

Trickling media filters (TM) Media: Bioballs, biobarrels, biofill, kaldness K1

Simple to construct and operate Excellent aeration Excellent CO2 removal No mechanical or electrical requirement

Low surface area media Lower nitrification and TAN removal rates Biofouling of media no biofilm removal Requires maintenance

Fluidized Sand Filters Fluidized sand (FS) filters are tubular filter casings containing a bed of sand that is expanded by the up-flow of water from the bottom of the filter casing (Malone, 2013) (Fig. 30.6). The up-flow of water greatly expands the distance between individual sand grains and fluidizes the sand, thus expanding the entire sand bed within the filter casing. Well-engineered FS filters have excellent transport of DO, ammonia, and nitrite to nitrifying bacteria (Timmons & Ebeling, 2013). FS filters feature the greatest specific surface areas and the highest TAN removal rates (Malone, 2013). FS filters can remediate TAN at 0.6e1.0 kg/m3/day and typically maintain very low ammonia and nitrite levels; it has been reported that as much as 50%e90% of the TAN is removed each time the water passes through the filter (Malone, 2013; Timmons & Ebeling, 2013). The continuous collisions of adjacent sand particles help to remove

excessive biofilm accumulations, and this increases nitrification efficiency (Malone, 2013). FS filters produce large amounts of carbon dioxide (CO2) and consume large amounts of oxygen (DO). Summerfelt and Sharrer (2004) reported that an FS filter attached to a salmonid production system produced an average of 4 ppm (mg/L) CO2 and consumed an average of 3.8 ppm (mg/L) of DO. These values represented 37% of the total CO2 produced and 35% of the total DO consumed by the entire system. These values are likely to be higher in warm water culture systems. The authors recommend that aeration/degassing would be most effective if located immediately after the biofilter (Summerfelt & Sharrer, 2004). Additional problems with FS filters include maintenance problems associated with biofouling (fine solids capture) and the high pumping costs associated with fluidizing sand (Timmons & Ebeling, 2013). Davidson, Helwig, and Summerfelt (2008)

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30. Recirculating Aquaculture Systems (RAS) for Zebrafish Culture

described an effective method for managing sand bed growth due to biofouling around sand grains. Sand with excessive biofouling has a low density and will float to the top of the FS bed. Authors used a submersible pump to move this low-density sand to the base of the FS filter. At the base of the filter, increased shearing removed the biofilm that then simply floated out of the filter. Managing FS filters in this way increased TAN and nitrite removal rates, decreased management labor, reduced bed growth, and reduced sand loss (Davidson et al. 2008). One pitfall of this FS filter management strategy is that it increases the amounts of fine solids that could irritate the gills of sensitive species (Summerfelt, 2006). FS filters can become compact and go anoxic quickly during a loss of pumping pressure or loss of power. For this reason, reliable back up pumps and power supplies are a necessity for FS filters. Different sizes and types of sand media can be used in FS filters. Davidson et al. (2008) studied FS filters using two different sizes of sand at 0.11 and 0.19 mm. They reported increased TAN and nitrite removal and increased DO consumption with the smaller sand size. Aquaneering Inc., for example, features an FS filter with a small spherical glass media (1000 m2/m3) and low void volume to support bacterial growth (Fig. 30.4). Water to be treated typically upflows through a static bed of floating beads that function as either a mechanical filter, a biological filter, or both simultaneously (Malone, 2013; Timmons & Ebeling, 2013). Aquaculture Systems Technologies (AST) has developed three different types of FB that suit a wide variety of culture applications. The bubble-wash bead filter, propeller-wash bead filter, and polygeyser bead filter, all have advantages and disadvantages that will not be discussed here. All FB filters have the advantage of high surface area and light-weight floating bead media that helps to facilitate good bacterial growth with effective wash capability to remove trapped solids. Beads are routinely washed by hydraulic, pneumatic, or mechanical means to remove accumulated solids and excessive biofilms (Malone, 2013; Timmons & Ebeling, 2013). FB filters have low pump pressure requirements, compact design, conserve water better than many other filters, and are easy to install, operate, and

backwash. The nitrification rates of FB filters approximate those of FS filters when used only for biological filtration (Malone, 2013). Nitrification rates for FB filters performing as both mechanical and biological filters are much lower at 380e518 g TAN/m3/day (Sastry et al. 1999). FB filters are not commonly used in zebrafish culture facilities at this time, although these filters have several desirable characteristics that could no doubt benefit such facilities. The advantages and disadvantages of FB filters are summarized in Table 30.3.

Trickling Media (TM) Filters Trickle filters (TM) use an open-top filter column packed with high void volume media to combine nitrification with aeration and degassing capability. The specific surface area and TAN conversion rates of the media used for TM filters is lower than that of other types of biofilters, but these issues are offset by the aeration and CO2 removal that these filters provide. Authors have reported a range of TAN removal rates from 0.1 to 0.9 g of TAN/m2/day for various TM filters operated at temperatures from 15 to 25 C (Eding et al. 2006; Timmons & Ebeling, 2013). The efficiency of these filters greatly depends on the water distribution at the top of the column and the efficiency of solids removal prior to water entering the TM filter (Eding et al. 2006; Timmons & Ebeling, 2013). Designs use flattened pore plates, spray nozzles, or other mechanisms to evenly distribute water to the top of the column. Water exiting TM filters is typically DO saturated and has very low CO2 concentrations. Solids not removed by efficient mechanical filtration upstream of TM filters will clog filter media, increase oxygen demand, and reduce TAN removal rates (Eding et al. 2006). TM filters are also easy to construct, relatively inexpensive, and have no mechanical or electrical requirements (Lawrence & Mason, 2012; Malone, 2013; Timmons & Ebeling, 2013). The advantages and disadvantages of TM filters are summarized in Table 30.3.

Chemical Filtration and Modification of Water Quality Chemical filtration is broadly interpreted herein to include any water treatment process where the water chemistry is altered by the addition or removal of chemicals from the water. This may occur by the processes of physical adsorption, water conditioning, or ion exchange.

Granular Activated Carbon Granular activated carbon (GAC) can be made from wood, charcoal, nutshells, fruit pits, bituminous coals,

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Chemical Filtration and Modification of Water Quality

lignite, peat, bone, and paper mill waste (Tchobanoglous, Burton, & Stensel, 2003; Yu, Yong, Han, & Ma, 2016). Material is heated at high temperature (700
C) in the absence of the oxygen necessary to sustain combustion. The material is then activated by oxidizing gases, such as steam and CO2, create a porous structure. The internal surface area created in the activated carbon has macropores >25 nm, mesopores from 1 to 25 nm, and micropores 100 mg/L

For 10 min after cessation of opercular movement

Lidocaine hydrochloride

>400 mg/L

For 10 min after cessation of opercular movement

2-Phenoxyethanol

800 mg/L

For 5 min

Hypothermia

0e4 C

For 5 min

>1800 mg/L

For larvae 3e8 dpf with exposure for over 1 h

>1800 mg/L

For larvae 3e8 dpf followed by an adjunctive method of euthanasia

Isoeugenol

>1500 mL/L

For larvae 14 dpf with exposure for at least 20 min after cessation of heartbeat

Hypothermia

0e4 C

Followed by an adjunctive method (bleach/decapitation/ maceration) after exposure for at least 20 min after cessation of heart beat for larvae 12 months), fish found in the system sumps with exposure to effluent water, or purposefully placed sentinel zebrafish. Each of these sample types has advantages and disadvantages and can be utilized to provide information about colony health.

Sentinel Fish Sentinel animals are purposefully placed into an animal colony to aid in the detection of disease. The use of sentinel fish in zebrafish facilities utilizing recirculating housing systems is analogous to the use of “dirty bedding” sentinels in rodent facilities. Sentinel

zebrafish are exposed to unfiltered, effluent wastewater from holding tanks from a specific rack, room, or facility, and therefore, have a much higher chance of being exposed to pathogens than colony fish in individual tanks on the system. As a result, fewer sentinel fish would be required for evaluation to detect disease at a given prevalence. The sensitivity for pathogen detection will vary based on the pathogen being evaluated, pathogen prevalence, age and strain of the fish tested, water quality, and other factors. For example, some pathogens, such as some Mycobacterium spp., are likely to be more prevalent in older fish, and thus may not lend itself to detection if using younger sentinel fish (Sasaki 2013; Marancik, 2019). Each independent holding system should have its own sentinel fish. The exposure of fish to effluent water can be accomplished by collecting fish housed in effluent “sumps” of the system, or by utilizing fish that are purposefully placed in tanks that are fed with effluent water from a specific sump, or a larger area of that system. The use of fish from the effluent sump(s) has both advantages and disadvantages. Many effluent sumps of large recirculating systems will become populated with fish originating from the holding system as a normal occurrence. System design and appropriate placement of baffles and handling of tanks will likely influence how many fry or fish are found in the sump. These fish are continuously exposed to large amounts of effluent water containing uneaten food, eggs, debris, and feces from all tanks draining to that sump. These fish are, thus, potentially exposed to high levels of any organisms present without using special equipment and/or fish produced or procured for this purpose. These fish are also being housed in an environment with sub-optimal water quality, which may make them more susceptible or more likely to develop clinical signs of disease. As a result, fish exposed to unfiltered effluent water are more likely to demonstrate histopathologic changes associated with decreased water quality, such as branchial hyperplasia (Marancik, 2019). Fish that reach the sump will no longer be available for research and should be removed routinely as standard practice to avoid the propagation of any diseases they may be carrying; and thus, are available for diagnostic testing. It has been documented that zebrafish of different genotypes may be more susceptible to certain pathogens (Whipps, Matthews, & Kent, 2008), and the genetic background of sump fish will be unknown. The genotype of fish present in a sump is likely heterogeneous, similar to that of the fish on the system; therefore, the population of sump fish may be more representative of the overall susceptibility of the represented colony fish to disease as opposed to using any one strain of wild type fish for testing. However, there may be an unequal distribution of genotypes if, for example, a

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Colony Fish

user of one line of fish more frequently has fish escaping into the sump. In addition, any previous experimental history, duration of exposure to effluent water, and age of the fish in the sump will be unknown. The advantages of using effluent sump fish should be balanced with the advantages of using sentinel fish that are purposefully placed in tanks fed with effluent water. Use of sentinel tanks with designated sentinel fish has several advantages to the use of colony fish, or sump fish, including sentinel fish that (1) are not actively being used for research, (2) can be exposed to effluent water from the larger overall holding system, albeit likely at a lower overall volume than fish housed in a sump, (3) are a known age and have had a known exposure period to effluent water, (4) can be easily observed on a daily basis and can be removed from the system quickly if there is evidence of illness, (5) are easy to remove from the system for testing, and (6) are of a known genotype. In addition, historical sentinel data comparisons can be made when sentinels come from the same source. Sentinel fish should be of a known pathogen status with susceptibility to the pathogens of interest and should be placed into the system in which they will be used as sentinels during the larval stage. This minimizes the risk of the introduction of new pathogens associated with the addition of adult fish from another source and allows environmental exposure to be evaluated for the maximum amount of time. A male sex bias toward susceptibility to P. neurophilia has been described (Chow, Xue, & Kent, 2016) and might be a consideration if excluding this pathogen; however, similar information is not yet available for other pathogens. The housing of all female fish in one tank is discouraged as this will result in the accumulation of eggs in the females, predisposing them to develop egg-associated inflammation. As mentioned previously, genotype may influence susceptibility to disease, for example, Tu¨bingen (TU) fish appear to be more susceptible to infection from M.chelonae (Murray et al., 2011a) and fish that have alterations to their immune system may be more sensitive to several infectious agents. Because the influence of genetic background and susceptibility to disease in zebrafish is still not well defined, the use of sentinel fish that are similar in genetic background to most of the fish used in the particular research colony (often a wild type strain, such as AB or TU) may be preferable. For most zebrafish pathogens, a minimum duration of system water exposure of 3e6 months is recommended to ensure the best chances of pathogen detection, although longer exposure often increases the chance of detecting certain diseases. Frequency of testing for each agent is discussed elsewhere, but will likely be determined based upon expected outcomes (i.e., transmission rate, availability of treatments, associated morbidity/mortality), and financial considerations.

Sentinels exposed to the same filtered system water as colony fish (postfiltration sentinels) can be used to evaluate the efficacy of the filtration and UV disinfection when results are compared with those from sentinel fish exposed to effluent water from that system. Similarly, sentinel fish placed on the quarantine system can be compared with results from both pre- and post filtration sentinels to evaluate the efficacy of quarantine procedures and to detect pathogens that have entered the quarantine system so that additional actions can be taken, if necessary, to prevent entry of the pathogen(s) into the conditioned colony (Martins et al., 2016). The use of contact sentinel fish, in which fish for sampling are purposefully cohoused in tanks also containing colony fish, has not been used extensively in zebrafish health surveillance programs. To utilize this strategy, zebrafish of a different phenotype can be cohoused with colony animals and subsequently sampled to evaluate the presence of disease in the conditioned colony animals. The number of fish required for evaluation is dependent upon the expected prevalence for each disease and the size of the colony, as these fish would not have greater potential pathogen exposure than the other fish within the colony, in contrast to fish exposed to system effluent water on a grander scale. Even so, this method allows testing without requiring the use of valuable research animals. It can be advantageous in this case to use one of the several phenotypically distinct zebrafish as contact sentinels, including those with hypopigmentation (Lister, Robertson, Lepage, Johnson, & Raible, 1999; White et al., 2008), variations in fin length (Iovine & Johnson, 2000), or altered pigmentation patterns, such as the leopard mutant (Watanabe et al., 2006). Current limitations of utilizing this strategy include limited vendor sources of specific pathogen-free fish and limited knowledge regarding the influence of genetic background on susceptibility to disease.

Colony Fish Direct sampling of clinically sick colony animals allows for the identification of pathogens which result in clinical disease and can be used to address concerns regarding morbidity and mortality within a colony. In these situations, a greater chance of disease detection may be achieved by sampling these animals, rather than a random sampling of colony animals (Collymore et al., 2016; Crim et al., 2017; Marancik, 2019). Clinical signs, location within the facility (room, holding system, rack, tank location, etc.), genotype, age, and experimental history should be recorded. Best results can be achieved when sick animals are submitted when they are still alive as tissues autolyze quickly following the

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35. Health Surveillance Programs

death of a fish. This is a reliable way to identify pathogens with a high prevalence without the need to purchase and sacrifice additional animals for screening; however, pathogens of a lower prevalence or that result in subclinical infections may be missed in this approach. The submission of aged (>12 months) colony zebrafish allows for evaluation of animals with prolonged duration of exposure to pathogens which may be present on a holding system. There are several pathogens, which are more easily detected with prolonged exposure, and which, often cause subclinical infections that would not be detected if only clinically ill fish are evaluated. For example, P. neurophilia is more prevalent in zebrafish over 1 year of age (Ramsay, Watral, Schreck, & Kent, 2009b). Older zebrafish are also more likely to be carriers of chronic subclinical or latent infections with Mycobacterium spp., and so may harbor greater numbers of bacterial organisms, making pathogen detection easier (Meritet et al., 2017). Removal of aged fish from a system is good practice from a biosecurity standpoint since they may be a reservoir for pathogens, and utilizing these fish as part of a health surveillance program provides a valuable use for them (Kent et al., 2009; Sasaki & Kishi, 2013).

Embryos Testing of embryos has been used as an adjunct method for detection of disease. The efficacy of this method of evaluation varies by the pathogen being evaluated, pathogen load, the spawning success of the adults utilized to generate the embryos, as well as the proportionate number of embryos from a clutch sampled. There are several flaws with the assumption that negative test results from embryos ensure that the spawning pair that produced those embryos is negative, or that all embryos in the same clutch are negative, as some pathogens may be spread intermittently, and therefore, may not be passed on to all offspring or even to some offspring at each spawning (Crim et al., 2017). For example, P. neurophilia is an obligate intracellular pathogen, and therefore, is only capable of growing and reproducing inside the cells of a host, although spores remain viable for long periods of time in the water. Therefore, P. neurophilia will be detected in embryos only if there is active external shedding of the organism during the spawning event allowing for extraovum transmission to embryos or rarely, true vertical (intraovum) transmission (Sanders et al. 2013). Other pathogens may be shed intermittently, or may be shed only if the animal is clinically ill. Less healthy animals are not as likely to participate in spawning or to produce sufficient embryos for evaluation, decreasing the sensitivity of this sampling source.

Environmental Samples Environmental samples can have diagnostic value for some zebrafish diseases. The sensitivity of environmental samples is likely to vary based on pathogen prevalence, exposure levels of the sample being evaluated, and the biology and life cycle of the agent. For example, the collection of tank detritus containing fish feces for double centrifugation with a saturated sugar solution is effective for the isolation and identification of P. tomentosa eggs (Murray & Peterson, 2015). However, the sensitivity of detection will likely vary based on the prevalence of the organism and the most prevalent life stage represented in the sample taken. In the case of P. tomentosa, for example, adult worms are found in the fish, and eggs in the environment, therefore, the life stage and numbers of the worms present will influence the numbers of eggs released into the environment that could be detected. The concentration of effluent water by filtration through a filter membrane followed by extraction of DNA for molecular diagnostic analysis allows a large volume of water to be evaluated as a single sample but may not be effective at detecting all pathogens. For example, real-time PCR of water concentrated by filtration has been shown to be useful for the detection of several Mycobacterium spp., but inconsistently detects other pathogens, such as Pseudoloma neurophilia and Pseudocapillaria tomentosa. Biofilms from tanks, gutters, sumps, or water lines on the system may be more useful for the evaluation of environmental bacteria, including Mycobacterium spp., or fungal pathogens having the potential to result in disease (Whipps et al., 2012). Areas exposed to high flows of effluent are likely to be of greater value, such as debris from the bottom of a specific tank or sump or from filter pads receiving effluent water. Evaluation of detritus has been shown to be sensitive for the detection of some Mycobacterium spp., and Pseudocapillaria tomentosa, with the evaluation of feces, also being sensitive for the detection of Pseudocapillaria tomentosa (Crim et al., 2017). This provides useful information which can be incorporated into the design of a health surveillance program. Interpretation of environmental results must be evaluated with knowledge and additional testing methodologies (e.g., histopathology) regarding the potential for the specific organism to cause disease and the particular risk to the colony based on the immune status of the fish and research goals. For example, the presence of environmental bacteria does not necessarily reflect infection of the fish on that system. In contrast, negative results may not be reflecting the presence of infection in fish for organisms that will only be present in the environment following the shedding of spores or eggs (i.e., Pseudoloma neurophilia, Pseudocapillaria tomentosa).

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Testing Methods

Testing Methods Testing methods and commercial diagnostic services for zebrafish have become more accessible in recent years, and both environmental and in vivo testing methodologies are available for some, but not all agents. Unfortunately, the most comprehensive method for detecting the presence of disease in zebrafish requires the inclusion of lethal testing methodologies. Several testing methods will be discussed briefly here as more comprehensive details regarding diagnostic procedures can be found elsewhere in this book. External parasites can be diagnosed in fish utilizing nonlethal methods, such as examination of external slide impressions or fin clip samples taken from anesthetized fish. A more comprehensive evaluation, however, can be conducted by utilizing fin clips, gill clips, or cutaneous wet mounts together when the entire body is available for diagnostic evaluation. When screening for Pseudoloma neurophilia exclusively, a simple brain prep can be utilized (Matthews et al., 2001). Gross necropsy incorporating external examination for external parasites, followed by internal necropsy with sterile culture and subsequent harvest of all organs for histopathology, is a method for a comprehensive evaluation of animal health and allows for detection of both infectious and noninfectious diseases. Noninfectious diseases that could be detected utilizing these testing methods include neoplasia, egg-associated inflammation, degenerative lesions, or other changes which may be indicative of poor water quality or the presence of toxins or nutritional deficiencies. Although complete histopathologic evaluation allows for a comprehensive evaluation, it may not always be the most sensitive method for all agents, and molecular diagnostic methods (i.e., polymerase chain reaction) may be more sensitive. Gross necropsy is best conducted immediately after euthanasia to avoid rapid autolysis of tissues. A dissecting microscope is often helpful for collection of samples and gross evaluation of organs. Histopathology results of animals found dead are often nonconclusive because of autolysis. Sterile samples may be cultured for identification of some pathogens as part of a necropsy. The collection of sterile culture samples requires advanced planning and technical skills to decrease sample contamination and to ensure diagnostic value. The potential bacterial species of interest should be considered before samples are collected to allow for the selection of the appropriate culture media and incubation temperature. Attempts to collect samples for bacterial culture in zebrafish should only occur immediately after euthanasia as the bacterial species present can change during postmortem autolysis of the fish. Before the fish is opened for culture, its

423

surface should be disinfected with 70% ethanol. Sterile instruments must be used to ensure sample integrity, and nonpowdered moistened gloves should be used to avoid contamination of samples with substances on the gloves. Organ cultures from the spleen and liver may allow isolation of Mycobacterium spp., and organ culture from the kidney (from either a dorsal or ventral approach) may be used for isolation of other organisms (Whipps et al., 2008; Zebrafish International Resource Center, 2016). Although culture may be diagnostic for known pathogenic organisms, positive results may be of questionable value unless evaluated in context with histopathologic findings or clinical disease. Molecular diagnostic methods, such as real-time polymerase chain reaction (PCR), conventional PCR, and matrix-assisted laser desorption ionization timeof-flight mass spectrometry (MALDI-TOF MS) are available for several known zebrafish pathogens. These testing methods can be used to evaluate live and dead animals, embryos, and environmental samples. PCR can be used to evaluate biofilms, detritus, food (both live and processed diets), feces, filter material, water, surface swabs, sperm, and microbial cultures. When used for the evaluation of live animals, there are several advantages to PCR. When compared with histopathology, PCR has greater sensitivity for small pathogenic burdens, as with some P. neurophilia infections, which may not be seen via histopathology (Kent et al., 2009; Murray et al., 2011b). Histopathology alone cannot be used to identify specific Mycobacterium spp., or the specific agent involved in other bacterial or fungal infections (Astrofsky, Schrenzel, Bullis, Smolowitz, & Fox, 2000). That is, a specific histological lesion may suggest an agent, but species-level identification requires molecular testing in most cases. Many fish can be pooled for molecular diagnostic evaluation, allowing for large numbers of fish to be sampled at once; and often allowing for a faster turnaround time compared to histopathology, standard microbial culture, or culturing of slow-growing bacteria (i.e., Mycobacterium spp.). Submission of a fresh-frozen whole fish is ideal to ensure representation of all organ systems for testing and to provide the most comprehensive information, especially for the evaluation of pathogens, such as Mycobacterium spp., which have a high limit of detection in PCR assays. However, fish that are found dead can be submitted for PCR analysis when histopathology would not provide any diagnostic value due to rapid tissue autolysis. Due to the small size of the zebrafish, it is often not practical to perform both histopathology and PCR on the same fish without decreasing the diagnostic sensitivity of both due to insufficient sample evaluation. However, a moderate level of sensitivity has been demonstrated when

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35. Health Surveillance Programs

evaluating formalin-fixed paraffin-embedded samples for some Mycobacterium spp. (Meritet et al., 2017) and may assist with speciation if acid-fast bacteria are noted during a histological evaluation. In summary, the testing methods utilized will often be based on the facility exclusion list and the best testing methodology available for the pathogens of greatest importance. A combination of gross necropsy with pathology and molecular diagnostic techniques, in addition to environmental testing, can likely provide the complete picture of the health status of a colony. The specific testing strategies used becomes increasingly more important if a facility wishes to exclude pathogens, which will not be eliminated with embryo surface disinfection.

Health Report A health surveillance report should be available for all zebrafish colonies and should include a summary of recent tests, test results, as well as significant historical information. A housing and husbandry report may also be created to describe standard housing and husbandry procedures, including an overview of the quarantine and health surveillance programs and a description of acceptable and nonacceptable pathogens for that facility. Please see Table 35.3 for an example of a health surveillance report. An example of a housing and husbandry report has also been published (Collymore et al., 2016). Health reports should be made available to all researchers utilizing the facility. Pathogen status should be included when reporting experimental results in publications and should be critically evaluated by researchers to determine what influence, if any, pathogen status may have on their studies. In addition, health information should be disclosed whenever zebrafish are transferred between facilities. If not readily provided when arrangements for the transfer of fish are made, the receiving institution should request a health surveillance report prior to receiving the fish. It is imperative to disclose all positive test results because the presence of some diseases, especially those that have no treatments and are not prevented by embryo surface disinfection, can have a devastating effect on both animal health and research programs.

Disease Prevention An adequate health surveillance program, the provision of a stable aquatic environment, and adherence to appropriate quarantine and biosecurity practices are all important aspects of disease prevention. Adequate detection of disease is required in order to quickly identify and remove sick fish and minimize transmission of

disease. Similarly, there are other practices which can help minimize the transmission of new or endemic diseases. All efforts should be made to maintain breeding stock that is as free as possible from diseases. This can be achieved by replacing the breeding stock with fish of a known health status every 6e12 months. Ideally, fish used for spawning are only crossed with fish from one other tank or group of tanks. Because the rate of transmission for some pathogens, such as P. neurophilia, increases during spawning events, spawning tanks or mass spawning chambers should be disinfected using a validated method, after each spawning event (Peneyra et al., 2018). Older fish are more likely to be infected with pathogens, such as P. neurophilia or Mycobacterium spp., and are more likely to succumb to illness from disease and/or to potentially transmit disease as they age or are exposed to experimental stress (Keller & Murtha, 2004; Parikka et al., 2012; Spitsbergen & Kent, 2003). Colony culling of all fish older than 1e1.5 years of age has been recommended. Alternatively, these older fish may be moved to quarantine (Borges et al., 2016; Collymore et al., 2016) if they are valuable as aged fish for experimental reasons. If maintained in the colony for longer periods of time, these fish are likely to serve as reservoirs of infection. They may also eventually die in their tank and subsequently expose their tank mates to disease after oral ingestion of spores or other infectious particles from their carcass occurs (Murray et al., 2011b). As mentioned earlier, these culled fish may also be useful for health surveillance purposes. Routine monitoring of system water quality parameters ensures a stable environment for zebrafish and has a direct influence on zebrafish health. Alterations in water quality can result in stress and subsequent expression of or susceptibility to disease. Water quality monitoring should include confirmation of adequate water exchanges in recirculating systems, confirmation of adequate gross particle filtration, and frequent monitoring and verification of values for important parameters including pH, temperature, conductivity, hardness/alkalinity, levels of nitrogenous waste, and contaminants, such as chlorine. Stocking density can also impact water quality. Recirculating water systems should include adequately sized UV disinfection units to assist with the prevention of tank to tank transmission of pathogens. The level of UV irradiation required to eliminate aquatic pathogens has been published (Harper & Lawrence, 2011). The functioning of the UV lights should be verified on a frequent basis, and the UV light bulbs and associated quartz sleeves should be replaced as per the manufacturer’s recommendations. One of the most important aspects of biosecurity involves the prevention of pathogen entry into a facility.

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Disease Prevention

TABLE 35.3

Example of a health surveillance reporta.

Health Surveillance Report for Zebrafish Instuon:

Date:

Building

Re-circulang

Room

Flow through

System/Rack Idenficaon Contact person: (name, e-mail, phone)

Stac Other Recent tesng

Sampling Tested Subjet Locaon (pre(sennel, colony fish, sump fish, environmental)

Historical results Collected

over ___ filtraon, postmonths # filtraon, main colony, Age of Exposure Tesng Tesng Tesng Sampling posives/ Sampling # posives/ # tested # tested Date me Frequency Method Laboratory Date quaranne ) fish

Bacteria Aeromonas hydrophila Edwardsiella ictaluri Flavobacterium columnare Mycobacterium spp. Mycobacterium abscessus Mycobacterium chelonae Mycobacterium fortuitum Mycobacterium haemophilum Mycobacterium marinum Mycobacterium peregrinum

Microsporidia Pseudoloma neurophilia Pleistophora hyphessobryconis

Protozoa Ichthyophthirius mulfilis Piscinoodinum pillulare

Fungi Saprolegnia brachydanis

Parasites Pseudocapillaria tomentosa

Addional Agents Infecous spleen and kidney necrosis virus (ISKNV) Spring Viremia of Carp Virus (SVCV)

Pathology:

Additonal comments:

a

Adapted with modifications from Collymore et al., (2016).

Limitation of the entry of pathogens associated with live fish and embryos is managed with an appropriately designed import and quarantine program. This is discussed in detail in another chapter of this book. Limitation of the entry of pathogens by individuals handling and caring for the fish must also be

considered. Individuals that provide care to fish at home or at other locations (e.g., pet stores, schools) should be counseled regarding necessary disease prevention precautions, which may include not handling other aquatic species in the morning prior to entering the zebrafish facility. Personal Protective Equipment

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(PPE) requirements should be determined for all areas where zebrafish are manipulated, including laboratories, holding rooms, and quarantine. To avoid the transfer of water from room to room, disinfection mats or baths, shoe covers or dedicated shoes may be used. Due to the possibility of exposure to zoonotic pathogens, such as Mycobacterium spp., the use of gloves and thorough washing of hands and arms when leaving the aquatic facilities is recommended. Limiting the entry of pathogens in the feed provided to animals is another important aspect of biosecurity and also must be considered during disease evaluation. Most dry diets utilize a fish protein source and information regarding measures taken to ensure that the protein source does not contain pathogenic organisms. The quality assurance methods utilized by the vendor providing the feed should be critically evaluated. For example, does the vendor test for aquatic pathogens or autoclave or irradiate the diet? Irradiation is one method for preventing the transmission of disease in animal diets (Reuter, Livingston, & Leblanc, 2011; Watson, 2013). In the absence of defined quality assurance for feed, testing for the presence of pathogens, via PCR is an option. Similarly, the feeding of a live diet, such as paramecium, Artemia (brine shrimp) or rotifers (i.e., Brachionus spp.), carries an inherent risk. It has been demonstrated that paramecium, rotifers, and Artemia can be used as a vector for the transmission of Mycobacterium and other bacteria, including Aeromonas spp. (Cantas, Sørby, Alestro¨m, & Sørum, 2012; Mason et al., 2016; Peterson et al., 2013). Artemia are nonselective filter feeders and can also be infected with microsporidia and other parasites (Hoj, Bourne, & Hall, 2009; Me´ndez-Hermida, 2007). Crustaceans have been documented to pass microsporidian spores through their digestive tract, experimentally resulting in heavier infections than fish infected through direct IP injection (Solter, 2014) and have been suggested as a vector for cryptosporidiosis in other fish (Me´ndez-Hermida, Go´mez-Couso, & Parasitol Res, 2007). Artemia is commonly sourced from natural lakes with varying levels of lead, arsenic, chromium, and other metal contaminants or pollutants with limited quality control (Peterson & Gustin, 2008; Tye, 2018). This results in a food source with varying nutritional content that can potentially introduce pathogens or other nonbiological contaminants. Workflow for feeding and other procedures should begin with provision of care to embryos and larvae first since they are less likely to be infected with a disease, then to adult fish, then aged fish, and finally to quarantined fish. Because of the risk for water splashing to tanks below during removal of tanks or other manipulations, it has been recommended that younger fish be placed above older fish on racks when possible and

that the younger fish be fed first (Mason et al., 2016). Ideally, the staff providing care to quarantine areas are distinct from those providing care to the conditioned colony. If staff providing care to zebrafish also provide care to other animal species, an appropriate workflow and PPE procedures for these individuals must be determined based on the health status and transmissibility of pathogens of all animal species being cared for. Prevention of tank to tank transmission of disease within a facility is closely associated with the proper handling of tanks and the use of validated cleaning methods. Nets and other supplies should be used for only one tank and should be adequately disinfected and thoroughly dried, using a validated method, prior to being used in another tank. Similarly, feeding implements ideally should not come into contact with the lids of multiple tanks during a feeding session. Cleaning methods for tanks, baffles, lids, and enrichment devices should all be validated to be effective and should utilize methods that are either nontoxic to the fish or that incorporate appropriate rinsing procedures to avoid the possibility of chemical residues (Martins et al., 2016). Regular cleaning and sanitization of holding tanks will allow for adequate visualization of the fish during health monitoring and also will prevent the accumulation of bryozoan and biofilms. Sanitization methods may be evaluated for efficacy by utilizing ATP-based monitoring systems, cultures, or PCR (Collymore, Porelli, Lieggi, & Lipman, 2014; Garcia & Sanders, 2011) and for safety utilizing chemical detection kits or embryo toxicity studies. Tank biofilms colonized with Mycobacteriumspp. have been associated with transmission of this disease (Chang, 2019), emphasizing the need for regular sanitization of holding tanks, and any enrichment in the holding tanks, utilizing validated methods. The impact on fish health must be considered if adding plants or other structural enrichment devices into an aquatic environment. The release of chemical components (e.g., phthalates, Bisphenol A) may have direct or indirect biologic effects on physiology, pharmacokinetics, behavior, and neoplasia (Lawrence & Mason, 2012; Mathieu-Denoncourt et al., 2015). These structures also increase the surface area for microbial growth which can impact water chemistry and algal and bacterial growth. The structure may also provide a good location for the harboring of pathogens (i.e., parasites, Mycobacterium spp.). As such, all enrichment devices require adequate sanitization, by a validated method, prior to being moved into a different tank. These devices may also provide additional surface area for chemical binding of cleaning agents, so if a chemical method of disinfection is chosen, care should be taken to ensure no residual chemical remains. Other considerations include the physical presence of structural

III. Husbandry

References

devices and any associated potential to cause physical harm (i.e., skin damage leading to secondary infections, choking, etc.) and finally any behavioral impact (alteration in swimming patterns, territory aggression, etc.). Pest control in a zebrafish facility must take into account aquatic, terrestrial, and airborne parasites. Due to the sensitivity of zebrafish to chemicals in their environment, prevention by maintaining a clean facility is of paramount importance. This includes minimizing the amount of food and other organic material available to pests, by maintaining food in closed containers, maintaining lids on tanks and sumps, minimizing food on lids of tanks, avoiding standing water, and flushing floor and sink drains daily. The maintenance of snails, algae-eating fish, or other organisms within a zebrafish holding system should be avoided as these animals often have an unknown health status and have the potential to transmit disease either directly (i.e., P. hyphessobryconis, P.neurophilia) or indirectly (i.e., P. tomentosa, Transversotrem spp.) (Sanders et al. 2010, 2016; Womble, Cox-Gardiner, Cribb, & Bullard, 2015). As with other animal species, adequate cleanliness of the facility, verification of appropriate lighting (levels, cycling, wavelength), and development of an emergency response plan are all important for providing adequate care. In summary, health surveillance of laboratory zebrafish colonies is necessary to ensure animal welfare and responsible use of animals. Timely recognition of the presence of infectious agents will be of assistance in the appropriate management of these conditions and will allow for safe collaboration, including the sharing of different zebrafish lines, with other scientists. In addition, several zebrafish pathogens result in pathology that may influence research, therefore, reporting the presence of these pathogens in scientific publications is of paramount importance to ensure adequate transparency of research results.

References ´ ., Reuter, G., Sadeghi, M., Deng, X., Altan, E., Kubiski, S. V., Boros, A Creighton, E. K., Crim, M. J., & Delwart, E. A. (2019). Highly divergent picornavirus infecting the gut epithelia of zebrafish (Danio rerio) in research institutions worldwide. zebrafish, 16(3), 291e299. https://doi.org/10.1089/zeb.2018.1710. Epub 2019 Apr 2. Astrofsky, K. M., Schrenzel, M. D., Bullis, R. A., Smolowitz, R. M., & Fox, J. G. (2000). Diagnosis and Management of atypical Mycobacterium spp. infections in established laboratory zebrafish (Brachydanio rerio) facilities. Comparative Medicine, 50, 666e672. Baker, D. G. (1998). Natural pathogens of laboratory mice, rats, and rabbits and their effects on research. Clinical Microbiology Reviews, 11(2), 231e266. Balla, K. M., Lugo-Villarino, G., Spitsbergen, J. M., Stachura, D. L., Hu, Y., Ban˜uelos, K., et al. (2010). Eosinophils in the zebrafish: Prospective isolation, characterization, and eosinophilia induction by helminth determinants. Blood, 116(19), 3944e3954.

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Bermudez, R., Losada, A. P., de Azevedo, A. M., Guerra-Varela, J., Perez-Fernandez, D., Sanchez, L., et al. (2018). First description of a natural infection with spleen and kidney necrosis virus in zebrafish. Journal of Fish Diseases, 41, 1283e1294. https://doi.org/ 10.1111/jfd.12822. Binesh, C. P. (2013). Mortality due to viral nervous necrosis in zebrafish Danio rerio and goldfish Carassius auratus. Diseases of Aquatic Organisms, 104, 257e260. https://doi.org/10.3354/dao02605. Borges, A. C., Pereira, N., Franco, M., Vale, L., Pereira, M., Cunha, M. V., et al. (2016). Implementation of a zebrafish health program in a research facility: A 4-year retrospective study. Zebrafish, 13(Suppl. 1), S-115eS-126. Broussard, G. W., Norris, M. B., Schwindt, A. R., Fournie, J. W., Winn, R. N., Kent, M. L., et al. (2009). Chronic Mycobacterium marinum infection acts as a tumor promoter in Japanese medaka (Oryzias latipes). Comparative biochemistry and physiology. Toxicology & Pharmacology : CB, 149(2). Cantas, L., Sørby, J. R. T., Alestro¨m, P., & Sørum, H. (2012). Culturable gut microbiota diversity in zebrafish. Zebrafish, 9(1), 26e37. Chang, C. T., Lewi, J., & Whipps, C. M. (2019). Source or Sink: Examining the Role of Biofilms in Transmission of Mycobacterium spp. in Laboratory Zebrafish. Zebrafish, 16(2), 197e206. https:// doi.org/10.1089/zeb.2018.1689. Chow, F. W., Xue, L., & Kent, M. L. (2016). Retrospective study of the prevalence of Pseudoloma neurophilia shows male sex bias in zebrafish Danio rerio (Hamilton-Buchanan). Journal of Fish Diseases, 39(3), 367e370. Collymore, C., Crim, M. J., & Lieggi, C. (2016). Recommendations for health monitoring and reporting for zebrafish research facilities. Zebrafish, 13(Suppl. 1), S-138eS-148. Clark, T. S., Pandolfo, L. M., Marshall, C. M., Mitra, A. K., & Schech, J. M. (2018). Body condition scoring for adult zebrafish (Danio rerio). Journal of the American Association for Laboratory Animal Science, 57(6), 698e702. https://doi.org/10.30802/AALAS-JAALAS-18-000045. Collymore, C., Watral, V., White, J. R., Colvin, M. E., Rasmussen, S., Tolwani, R. J., & Kent, M. L. (2014b). Tolerance and efficacy of emamectin benzoate and ivermectin for the treatment of Pseudocapillaria tomentosa in laboratory zebrafish (Danio rerio). Zebrafish, 11(5), 490e497. https://doi.org/10.1089/zeb.2014.1021. Crim, M. J., Lawrence, C., Livingston, R. S., Rakitin, A., Hurley, S. J., & Riley, L. K. (2017). Comparison of antemortem and environmental samples for zebrafish health monitoring and quarantine. Journal of the American Association for Laboratory Animal Science, 56(4), 412e424. Cronan, M. R., & Tobin, D. M. (2014). Fit for consumption: Zebrafish as a model for tuberculosis. Disease Models & Mechanisms, 7(7), 777e784. Franklin, C. L. (2006). Microbial considerations in genetically engineered mouse research (Vol. 47, pp. 141e155). Institute for Laboratory Animal Research. Ferguson, J., Watral, V., Schwindt, A., & Kent, M. L. (2007). Spores of two fish Microsporidia (Pseudoloma neurophilia and Glugea anomola) are highly resistant to chlorine. Diseases of Aquatic Organisms, 76, 205e214. Garcia, R. L., & Sanders, G. E. (2011). Efficacy of cleaning and disinfection procedures in a zebrafish (Danio rerio) facility. Journal of the American Association for Laboratory Animal Science, 50(6), 895e900. Gaulke, C. A., Martins, M. L., Watral, V., Kent, M. L., & Sharpton, T. J. (2016). Parasitic infection by Pseudocapillaria tomentosa is associated with a longitudinal restructuring of the zebrafish gut microbiome. BioRxiv Beta. https://doi.org/10.1101/076596. Harper, C., & Lawrence, C. (2011). The laboratory zebrafish. Boca Raton, FL: Taylor and Rancis Group, LLC (Chapter 2).

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Hawke, J. P., Kent, M., Rogge, M., Baumgartner, W., Wiles, J., Shelley, J., et al. (2013). Edwardsiellosis caused by Edwardsiella ictaluri in laboratory populations of zebrafish Danio rerio. Journal of Aquatic Animal Health, 25(3), 171e183. Hoj, L., Bourne, D. G., & Hall, M. R. (2009). Localization, abundance and community structure of bacteria associated with Artemia: Effects of nauplii enrichment and antimicrobial treatment. Aquaculture, 293, 278e285. Institute for Laboratory Animal Research. (2011). Guidance for the description of animal research in scientific publications. Washington (DC): National Academies Press. Iovine, M. K., & Johnson, S. L. (2000). Genetic analysis of isometric growth control mechanisms in the zebrafish caudal Fin. Genetics, 155(3), 1321e1329. Keller, E. T., & Murtha, J. M. (2004). The use of mature zebrafish (Danio rerio) as a model for human aging and disease. Comparative Biochemistry and Physiology - C Toxicology and Pharmacology, 138(3), 335e341. Kent, M. L., & Bishop-Stewart, J. K. (2003). Transmission and tissue distribution of Pseudoloma neurophilia (Microsporidia) of zebrafish, Danio rerio (Hamilton). Journal of Fish Diseases, 26, 423e426. Kent, M. L., Bishop-Stewart, J. K., Matthews, J. L., & Spitsbergen, J. M. (2002). Pseudocapillaria tomentosa, a nematode pathogen, and associated neoplasms of zebrafish (Danio rerio) kept in research colonies. Comprative Medicine, 52, 654e658. Kent, M. L., Feist, S. W., Harper, C., Hoogstraten-Miller, S., Mac Law, J., Sa´nchez-Morgado, J. M., et al. (2009). Recommendations for control of pathogens and infectious diseases in fish research facilities. Comparative Biochemistry & Physiology, 149(2), 240e248. Kent, M. L., Harper, C., & Wolf, J. C. (2012). Documented and potential research impacts of subclinical diseases in zebrafish (Vol. 53, pp. 126e134). Institute for Laboratory Animal Research, 2. Kent, M. L., Matthews, J. L., Bishop-Stewart, J. K., Whipps, C. M., Watral, V., Poort, M., et al. (2004). Mycobacteriosis in zebrafish (Danio rerio) research facilities. Comparative Biochemistry and Physiology Part C: Toxicology & Pharmacology, 138, 383e390. Kent, M. L., Watral, V. G., Kirchoff, N. S., Spagnoli, S. T., & Sharpton, T. J. (2016). Effects of subclinical Mycobacterium chelonae infections on fecundity and embryo survival in zebrafish. Zebrafish, 13(Suppl. 1), S-88eS-95. Kilkenny, C., Browne, W. J., Cuthill, I. C., Emerson, M., & Altman, D. G. (2010). Improving bioscience research reporting: The ARRIVE guidelines for reporting animal research. PLoS Biology, 8(6), e1000412. Lawrence, C., & Mason, T. (2012). Zebrafish housing systems: a review of basic operating principles and considerations for design and functionality. ILAR Journal, 53(2), 179e191. Lawrence, C., Eisen, J. S., & Varga, Z. M. (2016). Husbandry and health program survey synopsis. Zebrafish, 13(Suppl. 1), S-5eS-7. Lister, J. A., Robertson, C. P., Lepage, T., Johnson, S. L., & Raible, D. W. (1999). Nacre encodes a zebrafish microphthalmia-related protein that regulates neural-crest-derived pigment cell fate. Development, 126, 3757e3767. Martins, S., Monteiro, J. F., Vito, M., Weintraub, D., Almeida, J., & Certal, A. C. (2016). Toward an integrated zebrafish health management program supporting cancer and neuroscience research. Zebrafish, 13(S1), S47eS55. Marancik, D., Collins, J., Afema, J., & Lawrence, C. (2019). Exploring the advantages and limitations of sampling methods commonly used in research facilities for zebrafish health inspections. Laboratory Animals. epub ahead of print. https://doi.org/10.1177/ 0023677219864616 Martins, M. L., Watral, V., Rodrigues-Soares, J. P., & Kent, M. L. (2017). A method for collecting eggs of Pseudocapillaria tomentosa (Nematoda: Capillariidae) from zebrafish Danio rerio and efficacy of heat

and chlorine for killing the nematode’s eggs. Journal of Fish Diseases, 40, 169e182. Mason, T., Snell, K., Mittge, E., Melancon, E., Montgomery, R., McFadden, M., et al. (2016). Strategies to mitigate a Mycobacterium marinum outbreak in a zebrafish research facility. Zebrafish, 13(Suppl. 1), S-77eS-87. Mathieu-Denoncourt, J., Wallace, S. J., de Solla, S. R., & Langlois, V. S. (2015). Plasticizer endocrine disruption: highlighting developmental and reproductive effects in mammals and non-mammalian aquatic species. General and Comparative Endocrinology, 219, 74e88. Matthews, J. L., Brown, A. M. V., Larison, K., Bishop-Stewart, J. K., Rogers, P., & Kent, M. L. (2001). Pseudoloma neurophilia n. g., n. sp., a new microsporidium from the central nervous system of the zebrafish (Danio rerio). The Journal of Eukaryotic Microbiology, 48, 227e233. Meijer, A. H., Verbeek, F. J., Salas-Vidal, E., Corredor-Adamez, M., Bussman, J., van der Sar, A. M., et al. (2005). Transcriptome profiling of adult zebrafish at the late stage of chronic tuberculosis due to Mycobacterium marinum infection. Molecular Immunology, 42, 1185e1203. Me´ndez-Hermida, F., Go´mez-Couso, H., & Parasitol Res, A.-M.,E. (2007). Possible involvement of Artemia as live diet in the transmission of cryptosporidiosis in cultured fish. Parasitology Research, 101(3), 823e827. Meritet, D. M., Mulrooney, D. M., Kent, M. L., & Lo¨hr, C. V. (2017). Development of quantitative real-time PCR assays for postmortem detection of Mycobacterium spp. common in zebrafish (Danio rerio) research colonies. Journal of the American Association for Laboratory Animal Science, 56(2), 131e141. Mocho, J.-P. (2016). Three-dimensional screen: A comprehensive approach to the health monitoring of zebrafish. Zebrafish, 13(Suppl. 1), S-132eS-137. Murray, K. N., Bauer, J., Tallen, A., Matthews, J. L., Westerfield, M., & Varga, Z. M. (2011). Characterization and management of asymptomatic Mycobacterium infections at the zebrafish international resource center. Journal of the American Association for Laboratory Animal Science: JAALAS, 50(5), 675e679. Murray, K. N., Dreska, M., Nasiadka, A., Rinne, M., Matthews, J. L., Carmichael, C., et al. (2011). Transmission, diagnosis, and recommendations for control of pseudoloma neurophilia infections in laboratory zebrafish (Danio rerio) facilities. Comparative Medicine, 61(4), 322e329. Murray, K. N., & Peterson, T. S. (2015). Pathology in practice. P tomentosa infection in zebrafish. Journal of the American Veterinary Medical Association, 246(2), 201e203. https://doi.org/10.2460/ javma.246.2.201. Nogueira, C. L., Whipps, C. M., Matsumoto, C. K., Chimara, E., Droz, S., Tortoli, E., et al. (2015). Mycobacterium saopaulense sp. nov., a rapidly growing mycobacterium closely related to members of the Mycobacterium chelonaeeMycobacterium abscessus group. International Journal of Systematic and Evolutionary Microbiology, 65(Pt 12), 4403e4409. Parikka, M., Hammare´n, M. M., Harjula, S.-K. E., Halfpenny, N. J. A., Oksanen, K. E., Lahtinen, M. J., et al. (2012). Mycobacterium marinum causes a latent infection that can be reactivated by gamma irradiation in adult zebrafish. PLoS Pathogens, 8(9), e1002944. Peneyra, S. M., Cardona-Costa, J., White, J., Whipps, C. M., Riedel, E. R., Lipman, N. S., & Lieggi, C. (2018). Transmission of Pseudoloma neurophilia in laboratory zebrafish (Danio rerio) when using mass spawning chambers and recommendations for chamber disinfection. Zebrafish, 15(1), 63e72. https://doi.org/10.1089/ zeb.2017.1493. Peterson, T. S., Ferguson, J. A., Watral, V. G., Mutoji, K. N., Ennis, D. G., & Kent, M. L. (2013). Paramecium caudatum enhances transmission and infectivity of Mycobacterium marinum and Mycobacterium

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Further reading

chelonae in zebrafish (Danio rerio). Diseases of Aquatic Organisms, 106(3), 229e239. Peterson, C., & Gustin, M. (2008). Mercury in the air, water and biota at the great salt lake (Utah, USA). The Science of the Total Environment, 405, 255e268. Pham, L. N., Kanther, M., Semova, I., & Rawis, J. F. (2008). Methods for generating and colonizing gnotobiotic zebrafish. Nature Protocols, 3, 1862e1875. Ramsay, J. M., Watral, V., Schreck, C. B., & Kent, M. L. (2009). Husbandry stress exacerbates mycobacterial infections in adult zebrafish, Danio rerio (Hamilton). Journal of Fish Diseases, 32(11), 931e941. Ramsay, J. M., Watral, V., Schreck, C. B., & Kent, M. L. (2009). Pseudoloma neurophilia infections in zebrafish Danio rerio: Effects of stress on survival, growth, and reproduction. Diseases of Aquatic Organisms, 88, 69e84. Reuter, J. D., Livingston, R., & Leblanc, M. (2011). Management strategies for controlling endemic and seasonal mouse parvovirus infection in a barrier facility. Lab Animal, 40, 145e152. Sanders, J. L., Lawrence, C., Nichols, D. K., Brubaker, J. F., Peterson, T. S., Murray, K. N., et al. (2010). Pleistophora hyphessobryconis (Microsporidia) infecting zebrafish (Danio rerio) in research facilities. Diseases of Aquatic Organisms, 91(1), 47e56. Sanders, J. L., Peterson, T. S., & Kent, M. L. (2014). Early development and tissue distribution of Pseudoloma neurophilia in the zebrafish, Danio rerio. The Journal of Eukaryotic Microbiology, 61(3), 238e246. Sanders, J. L., Watral, V., Clarkson, K., & Kent, M. L. (2013). Verification of intraovum transmission of a microsporidium of vertebrates: Pseudoloma neurophilia infecting the zebrafish, Danio rerio. PLoS One, 8(9), e76064. Sanders, J., Watral, V., & Kent, M. L. (2012). Microsporidiosis in zebrafish research facilities (Vol. 53, pp. 106e113). Institute for Laboratory Animal Research. Sanders, J. L., Watral, V., Stidworthy, M. F., & Kent, M. L. (2016). Expansion of the known host range of the microsporidium, pseudoloma neurophilia. Zebrafish, 13(Suppl. 1), S102eS106. Sasaki, T., & Kishi, S. (2013). Molecular and chemical genetic approaches to developmental origins of aging and disease in zebrafish. Biochimica et Biophysica Acta, 1832(9), 1362e1370. https://doi.org/10.1016/j.bbadis.2013.04.030. Simon, R. C., & Schtll, W. B. (1984). Tables of sample size requirements for detection of fish infected by pathogens: Three confidence levels for different infection prevalence and various population sizes. Journal of Fish Diseases, 7, 515e520. Solter, L. F. (2014). Epizootiology of microsporidiosis in invertebrate hosts. In L. M. Weiss, & J. J. Becnel (Eds.), Microsporidia: Pathogens of opportunity (pp. 165e194). Wiley-Blackwell. Spagnoli, S., Sanders, J., & Kent, M. L. (2017). The common neural parasite Pseudoloma neurophilia causes altered shoaling behaviour in adult laboratory zebrafish (Danio rerio) and its implications for neurobehavioural research. Journal of Fish Diseases, 40, 443e446. Spagnoli, S. T., Sanders, J. L., Watral, V., & Kent, M. L. (2016). Pseudoloma neurophilia infection combined with gamma irradiation causes increased mortality in adult zebrafish (Danio rerio) compared to infection or irradiation alone: New implications for studies involving immunosuppression. Zebrafish, 13(S1), S107eS114. Spagnoli, S., Xue, L., & Kent, M. L. (2015). The common neural parasite Pseudoloma neurophilia is associated with altered startle response habituation in adult zebrafish (Danio rerio): Implications for the zebrafish as a model organism. Behavioural Brain Research, 291, 351e360. Spagnoli, S. T., Xue, L., Murray, K. N., Chow, F., & Kent, M. L. (2015). Pseudoloma neurophilia: A retrospective and descriptive study of

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nervous system and muscle infections, with new implications for pathogenesis and behavioral phenotypes. Zebrafish, 12(2), 189e201. Spitsbergen, J. M., Buhler, D. R., & Peterson, T. S. (2012). Neoplasia and neoplasm associated lesions in laboratory colonies of zebrafish emphasizing key influences of diet and aquaculture system design. ILAR Journal/National Research Council, Institute for Laboratory Animal Resources, 53(2), 114e125. Spitsbergen, J. M., & Kent, M. L. (2003). The state of the art of the zebrafish model for toxicology and toxicologic pathology researchd advantages and current limitations. Toxicologic Pathology, 31(Suppl. l), 62e87. Tye, M. T., Montgomery, J. E., Hobbs, M. R., Vanpelt, K. T., & Masino, M. A. (2018). An adult zebrafish diet contaminated with chromium reduces the viability of progeny. Zebrafish, 15(2), 179e187. https://doi.org/10.1089/zeb.2017.1514. Van der Vaart, M., Spaink, H. P., & Meijer, A. H. (2012). Pathogen recognition and activation of the innate immune response in zebrafish. Advances in Hematology, 159807. Watanabe, M., Iwashita, M., Ishii, M., Kurachi, Y., Kawakami, A., Kondo, S., et al. (2006). Spot pattern of leopard Danio is caused by mutation in the zebrafish connexin41.8 gene. EMBO Reports, 7(9), 893e897. Watson, J. (2013). Unsterilized feed as the apparent cause of a mouse parvovirus outbreak. Journal of the American Association for Laboratory Animal Science: JAALAS, 52(1), 83e88. West, K., Miles, R., Kent, M. L., & Frazer, J. K. (2014). Unusual fluorescent granulomas and myonecrosis in Danio rerio infected by the microsporidian pathogen pseudoloma neurophilia. Zebrafish, 11(3), 283e290. Whipps, C. M., Dougan, S. T., & Kent, M. L. (2007). Mycobacterium haemophilum infections of zebrafish (Danio rerio) in research facilities. FEMS Microbiology Letters, 270(1), 21e26. Whipps, C. M., Lieggi, C., & Wagner, R. (2012). Mycobacteriosis in zebrafish colonies. ILAR Journal/National Research Council, Institute of Laboratory Animal Resources, 53(2), 95e105. Whipps, C. M., Matthews, J. L., & Kent, M. L. (2008). Distribution and genetic characterization of Mycobacterium chelonae in laboratory zebrafish Danio rerio. Diseases of Aquatic Organisms, 82(1), 45e54. Whipps, C. M., Moss, L. G., Sisk, D. M., Murray, K. N., Tobin, D. M., & Moss, J. B. (2014). Detection of autofluorescent Mycobacterium chelonae in living zebrafish. Zebrafish, 11(1), 76e82. White, R. M., Sessa, A., Burke, C., Bowman, T., LeBlanc, J., Ceol, C., et al. (2008). Transparent adult zebrafish as a tool for in vivo transplantation analysis. Cell Stem Cell, 2(2), 183e189. Womble, M. R., Cox-Gardiner, S. J., Cribb, T. H., & Bullard, S. A. (2015). First record of Transversotrema Witenberg, 1944 (Digenea) from the Americas, with comments on the taxonomy of Transversotrema patialense (Soparkar, 1924) Crusz and Sathananthan, 1960, and an updated list of its hosts and geographic distribution. The Journal of Parasitology, 101(6), 717e725. Zebrafish International Resource Center. (2016). General diagnostics and necropsy protocols. Retrieved from https://zebrafish.org/wiki/ health/disease_manual/general_diag_and_necropsy_protocols.

Further reading Collymore, C., Porelli, G., Lieggi, C., & Lipman, N. S. (2014a). Evaluation of 5 cleaning and disinfection methods for nets used to collect zebrafish (Danio rerio). Journal of the American Association for Laboratory Animal Science: JAALAS, 53(6), 657e660.

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C H A P T E R

36 Importation and Quarantine Christine Lieggi Center of Comparative Medicine and Pathology, Memorial Sloan Kettering Cancer Center and Weill Cornell Medicine, New York, NY, United States of America

Introduction The importation of new zebrafish increases the risk of colony animals being exposed to pathogens if the imported fish are not managed appropriately. In an existing facility, it is important to identify any pathogens already present and determine the ones that will be excluded from the conditioned colony. Determination of excluded pathogens should be based on the direct impact of the pathogens on animal health (i.e., associated morbidity, mortality) and research results, their zoonotic potential, and financial resources available. An importation and quarantine program should be designed to minimize risk to the existing colony, while still allowing for scientific collaboration, sharing of fish lines, and adherence to regulations that require tracking numbers of vertebrate animals used in research.

Importation Request and Approval Defined animal requisition and importation procedures should be agreed upon between the researchers, veterinarians, zebrafish facility staff, and the Institutional Animal Care and Use Committee (IACUC) to ensure that all aspects of importation and regulatory control are met. Currently, commercial vendors dedicated to the production of Specific Pathogen Free (SPF) zebrafish for research do not exist. As a result, most fish obtained from outside sources are obtained through a zebrafish repository, such as the Zebrafish International Resources Center (ZIRC; University of Oregon, Eugene, OR, United States of America) or the European Zebrafish Resource Center (Karlsruhe Institute of Technology, Institute of Toxicology and Genetics, Eggenstein-Leopoldshafen, Germany); other research labs around the world that can provide SPF fish (SARL

The Zebrafish in Biomedical Research https://doi.org/10.1016/B978-0-12-812431-4.00036-1

Facility, Oregon State University, Corvallis, OR, United States of America); or other laboratories that may not be able to provide health status. As a result, the importation of zebrafish may best be handled like that for rodents or other species from noncommercial sources. A centralized animal ordering program is preferable to achieve the greatest control over the process. Importation requests should be cross-checked, following the institution’s standard animal ordering procedures, to confirm that the number and genetic line of zebrafish requested are described in an approved IACUC protocol and that the number of animals (including embryos and larvae) imported are deducted accurately from that protocol. If the receiving institution wishes to exclude specific pathogens, the health status of the exporting colony should be evaluated, ideally by a veterinarian, to determine if the shipment should be accepted, if special accommodations are needed for pathogen control, or if it should be rejected. For example, fish requested from a colony infected, or potentially infected, with an excluded pathogen may be accepted with a requirement for the administration of a specific treatment prior to shipment or upon arrival (e.g., surface disinfection of embryos, anthelminthic treatment, etc.). Alternatively, such a shipment may require further testing upon arrival, or standard quarantine procedures may be sufficient to prevent transmission of that specific pathogen, or unknown pathogens, to the conditioned colony. Some sources may be designated as “approved vendors.” Granting this status to a vendor generally requires the exporting facility to conduct routine colony health monitoring and to accurately disclose those results, which, at the time of this writing, is not well standardized and is inconsistently done for zebrafish. Even if a health report is available, the information must be reviewed with the understanding that results are always

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retrospective and may not reflect the current health status of the colony and thus, an understanding of the biosecurity practices of the exporting facility is also required. If a detailed health history and testing results are not available, the importing institution may attempt to retrieve details regarding the import, quarantine, and health monitoring procedures of the exporting institution to better analyze risk, or may request additional animals to be shipped to allow for more stringent quarantine strategies, including diagnostic testing, to be implemented. When receiving additional animals for testing upon arrival, it should be recognized that these fish will only be representative of a small group of fish from the colony and will likely only be valuable for detecting disease conditions that are highly prevalent in the source colony. Therefore, diagnostic testing on these animals would best be used to evaluate the health status of offspring generated from these specific animals, rather than used to extrapolate the health status of the originating colony. Some of these strategies will be discussed more thoroughly in the quarantine section below. Once the responsible facility administrator has reviewed all information provided on colony health status and determined the most appropriate location or handling procedures to be used upon arrival of the imported fish, it should be verified that there is adequate space in the quarantine area to accommodate them. Even if embryos or larvae are being received, accommodations should be planned for adult fish in the conditioned colony for the eventual housing of larvae derived from surface disinfected embryos.

Receipt Zebrafish are typically exported as either embryos or adults, and there are advantages and disadvantages to each method. Embryos may be surface disinfected prior to shipping to decrease potential contaminants that can impact water quality during shipping and decrease the risk of transmission of some pathogens to the receiving facilities, and a relatively large number of embryos can be held in a small container. Embryos do not require an external source of nutrition and do not generate large amounts of waste during transport. When imported embryos/larvae are received they are typically maintained on the quarantine system until they are able to produce embryos on their own, which can then be surface disinfected for placement into the main conditioned colony; this process may take 2e3 months. Additional considerations are required when shipping adults to alleviate the adverse effects of nitrogenous waste production and decreased water

oxygenation. Shipments of adult fish must be limited to smaller numbers in a given volume of water, when compared to embryos, to allow for safe shipping. Please refer to the Exportation and Transport chapter for more details regarding regulations and methods for shipping embryos and adults. Receipt procedures will differ based on the life stage of animals; however, the first step for all life stages is to inspect the shipping container for damage before it is opened for removal of animals. Upon receipt of embryos, the container should first be evaluated for integrity and location of embryos; it will be easier to remove embryos from the container if they are suspended in water. Embryos should be removed from the shipping container as soon as possible upon receipt. The embryos should be transferred to a clean container filled with embryo media or autoclaved system water, and the shipping water should be discarded down the drain. The easiest way to do this is to pour the contents of the shipping container through a nonabrasive strainer to catch the embryos, which are then gently rinsed, while still in the strainer, with embryo media or autoclaved system water. No more than 50e100 embryos should then be transferred into a standard 90 mm Petri dish filled with embryo media or autoclaved system water. It is important to limit the density of embryos and to use a clean media to minimize fungal, protozoan, and bacterial contamination and to provide sufficient aeration (Nu¨sslein-Volhard and Dahm, 2002, chap. 1). The embryos should be evaluated and any nonviable embryos (see Fig. 36.1) or debris removed, as this will contribute to the deterioration of water quality within the Petri dish. The embryos are then placed in a dedicated quarantine incubator and maintained at 28.5
2 C. If embryos were not surface disinfected prior to shipping, facility quarantine policies would dictate if surface disinfection of embryos upon arrival is required. However, embryos must not be disinfected once they begin to emerge from the chorion as this may lead to deformities or death of embryos/larvae. Adult zebrafish should be inspected as soon as possible upon arrival to evaluate their overall condition and health status. Fish that appear to be stressed will require an accelerated acclimation process to get them into better water conditions more quickly. From the time adult fish are placed into shipping containers of static water, conditions of the water begin to change. Fish consume oxygen and release carbon dioxide, which combines with water forming a weak acid and decreasing pH. Because nitrogenous waste is also released by fish during transport, the decreased pH is somewhat beneficial because it decreases the ratio of toxic, un-ionized ammonia (NH3) to nontoxic, ionized

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FIGURE 36.2 Without opening the bag, the shipping bag containing imported fish is placed in a static tank or other container filled with water that is the same temperature as the holding system to allow the water in the bag to acclimate.

FIGURE 36.1 No magnification. The chorion of a viable developing embryo will be transparent (white arrow with black outline), whereas nonviable embryos will appear opaque (white arrow with red outline). This clutch demonstrated a high level of embryo death following exposure to 100 ppm sodium hypochlorite for 10 min.

ammonia (NHþ 4 ), also known as ammonium. Once the shipping bag is opened and exposed to the environment, the carbon dioxide is released into the environment, resulting in a quick increase in pH of the water, shifting the balance toward a higher level of toxic ammonia. Therefore, the shipping bag holding fish should not be opened until fish are ready to be transferred out of the bag (Harper and Lawrence, 2011). It is also important to remember that the water inside and on the surface of the bag maybe contaminated with unwanted organisms; transfer of these organisms to the receiving facility should be avoided. Prior to removing fish from the bag, they should be acclimated to the temperature on the holding system. To achieve this while minimizing contamination risk, a static tank can be filled with water at the same temperature as that of the holding system, and the shipping bag can be placed into this tank to allow the water in the bag to acclimate (Fig. 36.2). It is estimated that acclimation of small volumes of water requires about 5 min per degree of temperature difference.

When temperature acclimation is satisfactory, cut a hole in the bag and then pour the water and fish through this hole into a perforated strainer or breeding tank insert to separate the fish from the shipping water (Figs. 36.3 and 36.4). Then quickly place the fish into their new holding tank without placing the net inside the new tank, and discard shipping water down the drain (Fig. 36.5). Any equipment exposed to the shipping water should not contact the system water or facility equipment until after it has been adequately disinfected. Continue to evaluate the fish after the transfer, and allow fish to acclimate to their new environment for at least 24 h prior to feeding and 2e5 weeks prior to breeding. It is important to minimize the stress of animals during transportation and receipt and to provide a stable environment once animals are received into a new facility. Stress is known to result in several physiologic changes in fish, including cortisol release (Ramsay et al., 2006, 2009a; Schreck, 1996), which can lead to immunosuppression, allowing a previously subclinical infection to exert clinical effects and result in shedding of organisms into the environment (Dror et al., 2006; Ramsay, Watral, Schreck, & Kent, 2009b). Because of this, it is important to ensure that the receipt of animals is as stress free as possible, and handling of animals, once they have arrived, should be minimized to allow them to acclimate to their new environment. For this reason, it is recommended that fish are simply observed for overall health status for the first 2e 5 weeks after arrival before breeding or any other manipulations begin.

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FIGURE 36.3

Example of equipment preparation for the transfer of imported adult fish onto a quarantine holding system following water temperature acclimation. There is a tank and a net to be used for separating fish from the shipping water and a labeled tank filled with system water that will be used to house new fish in the quarantine holding system.

Quarantine Goals

FIGURE 36.4 Following water temperature acclimation, the fish are separated from the shipping water and moved into a holding tank. A hole is cut in the bag, and the contents of the shipping bag are poured through a net. The captured shipping water is then disposed of down the drain.

The establishment of defined quarantine procedures is the first step in ensuring biosecurity during the importation of fish from outside sources. Because there are currently limited commercial sources of zebrafish guaranteed to be free of known pathogens, quarantine of incoming fish is extremely important. In the case of zebrafish, the purposes of quarantine are to allow the import of fish from colonies that may have an undefined health status, provide the opportunity to evaluate these for the presence of disease, prevent disease from incoming fish to be transferred to the main colony, and allow for acclimation of incoming fish. When the main conditioned colony is shared by several different laboratories, the importance of quarantine increases as any break in biosecurity will have the potential to affect a large number of users. This risk is amplified further when the conditioned colony is maintained on a recirculating holding system.

General Quarantine Practices

FIGURE 36.5 Fish are quickly placed into their new holding tank without putting the net inside the new tank.

The most universally utilized quarantine concept used by zebrafish facilities is an “eggs only” policy, meaning that only larvae derived from surface disinfected embryos are allowed to be placed on the housing system used for the conditioned colony. Therefore, all imported embryos, larvae, or adults are initially placed into the quarantine system to undergo treatments, procedures, or evaluations. All embryos generated in quarantine are (at minimum) surface disinfected prior to being placed as larvae into the designated nursery area within the conditioned colony. Once the imported line is established on the main system, the imported fish on the quarantine system should be culled. Imported fish should not be allowed to remain on the quarantine

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Quarantine Strategies

system longer than necessary to decrease the amount of fish that are potentially exposed to the disease each time a new import is added, and to allow for adequate space when new imports are requested. Generally, the entire process for an import should be completed in 70% (Xu et al., 2008). Experimental infections with ISKNV in other fish species have also resulted in high morbidity and mortality. For example, in one experimental infection study, all (n ¼ 40) mandarinfish (S. chuatsi) maintained over 20 C and exposed by intracoelomic injection died in less than 25 days, with average time to death for each treatment group ranging from 9 to 15 days (He et al., 2002). Importantly, however, these experimental infections were achieved via intracoelomic injection, which is not the natural route of infection. RNA viruses: A greater diversity of RNA viruses of fish has been experimentally evaluated in the zebrafish model, including representatives of three families: Birnaviridae, Nodaviridae, and Rhabdoviridae. Zebrafish have been experimentally infected with IPNV (LaPatra, Barone, Jones, & Zon, 2000; Seeley, Perlmutter, & Seeley, 1977), and Betanodavirus spp., including Malabar grouper nervous necrosis virus (Lu et al., 2008), RGNNV (Furusawa et al., 2007), and SGNNV (Morick et al., 2015). Zebrafish are experimentally susceptible to infection with at least four viral species that infect fish from the family Rhabdoviridae: IHNV (Ludwig et al., 2011), snakehead rhabdovirus (Phelan et al., 2005), Spring viremia of carp virus (SVCV) (Lopez-Munoz, Roca, Sepulcre, Meseguer, & Mulero, 2010; Sanders, Batts, & Winton, 2003), and VHSV (Novoa et al., 2006). Experimental infection of zebrafish with IHNV demonstrated

the utility of the zebrafish model for temperature-shift experiments (Ludwig et al., 2011). As zebrafish are poikilothermic and can survive a wide range of temperatures, the temperature at which infected embryos or larvae are maintained can be shifted several degrees to stop viral replication at various time points to better characterize the course of infection (Ludwig et al., 2011).

Experimental Infection Studies with Mammalian Viruses Zebrafish have also been used as a model organism to study several viruses that infect humans and other mammals. Not all human or mammalian viruses are capable of infecting zebrafish, however. Several important factors include the breadth of each viral host range, the expression of zebrafish orthologs of known viral receptors, and the necessary viral incubation temperature (Goody, Sullivan, & Kim, 2014). Some authors have noted that because zebrafish are commonly housed at 28 C, the utility of the zebrafish model for human infections might be limited (Lieschke & Currie, 2007). However, not all human viruses need to be maintained at 37 C for viral replication. Moreover, zebrafish can be slowly acclimated to a wide range of temperatures and have been maintained successfully at 37 C to facilitate the study of mammalian pathogens (Sanders et al., 2015). DNA viruses: Herpes simplex virus type (HSV)-1 is a member of the family Herpesviridae and is closely related to HSV-2 and varicella zoster virus. HSV-1 was the first human virus used to experimentally infect zebrafish in a study that demonstrated the reduction of viral load in response to the antiviral acyclovir in contrast to increased viral loads and mortality when treated with cyclophosphamide (Burgos, Ripoll-Gomez, Alfaro, Sastre, & Valdivieso, 2008). RNA viruses: Mammalian RNA viruses that have been studied in zebrafish are chikungunya virus (CHIKV) (Palha et al., 2013), sindbis virus (SINV) (Passoni et al., 2017), influenza A virus (IAV) (Gabor et al., 2014), and vesicular stomatitis virus (VSV) (Guerra-Varela et al., 2018). The zebrafish model is attractive for research using these viruses because other animal models do not permit efficient visualization of hostepathogen interactions. SINV and CHIKV are arboviruses in the genus Alphavirus in the Togaviridae family. SINV is spread by Culex mosquitos, whereas CHIKV is spread by Aedes mosquitoes. IAV is in the Orthomyxoviridae family. Experimental infection using a fluorescent IAV demonstrated a pattern of vascular endothelial infection and visualization of a reduction in fluorescence in infected larvae treated with an antiviral compound, illustrating the utility of the zebrafish model for antiviral screens (Gabor et al., 2014). VSV rarely causes zoonotic

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infections and also infects insect vectors, but is primarily known as a virus causing economic losses among livestock. Zebrafish embryos were highly susceptible when infected by microinjection into the yolk, with rapid viral spread to the CNS followed by other tissues just before death (Guerra-Varela et al., 2018).

Comments on Detection, Diagnosis, Risk Assessment, and Decision-making Husbandry and health monitoring practices for zebrafish are underdeveloped relative to rodent biomedical research models. To date, three naturally occurring viral infections have been reported in zebrafish (Altan et al., 2019; Bermudez et al., 2018; Binesh, 2013), and no zebrafish suppliers currently offer stocks that are guaranteed to be virus-free. It is important for the biomedical research community to identify and characterize naturally occurring zebrafish viruses and move toward the use of zebrafish that are free of adventitious viruses. The identification of viral pathogens in zebrafish will improve the utility of zebrafish as a model organism by improving zebrafish welfare, reducing confounding experimental variability, and providing insight regarding the most effective biosecurity measures in aquatic research facilities.

Detection and Diagnosis As viral pathogens are identified in zebrafish, monitoring should be put into place for producers, resource centers, established research colonies, and zebrafish quarantine units. As has been suggested for rodent colonies, prospective sampling according to a prearranged schedule permits effective health monitoring, and sampling can be increased in response to suspicion or evidence of viral infection (Suckow, Weisbroth, & Franklin, 2005). Many of the biosecurity practices that are widely practiced in rodent research facilities are only beginning to become recognized as critically important for zebrafish research, such as the use of vendor lists, purpose-bred pathogen-free animals, entry quarantine, pathogen exclusion lists, routine sentinel health monitoring, and environmental monitoring; however, widespread application of these practices will facilitate the exclusion and containment of infectious agents from zebrafish colonies (Crim et al., 2017).

Risk Assessment and Decision-making Biosecurity considerations: When a previously unknown viral infection is detected in a zebrafish colony as the result of a break in biosecurity, the next steps include an assessment, followed by actions to contain

the outbreak, interrupt viral transmission to naı¨ve fish, reduce the viral burden of the system, and reduce the impact of infection on research objectives. There are a number of important factors to consider during the initial and subsequent assessments to evaluate the options for control and/or elimination. These include any threat to human or animal health, the impact of infection on research objectives, the potential for spread to other systems, rooms, or institutions, the efficacy of possible measures, the potential disruption of any corrective measures to research, and the time and expense required. Importantly, many of these considerations require some understanding of viral stability and mode of transmission, in addition to virulence and shedding in different age groups of immunocompetent and immunocompromized zebrafish.

Restrictions on Zebrafish Movement SVCV, a commercially important viral pathogen of common carp and other fishes, has been studied experimentally using the zebrafish as a model (Encinas et al., 2013; Lopez-Munoz et al., 2010; Sanders et al., 2003; Wang et al., 2017). Because laboratory zebrafish were shown to be experimentally susceptible to infection with SVCV under experimental conditions, zebrafish were controversially added to the list of SVCVsusceptible species in the World Organization for Animal Health (OIE) Aquatic Manual, which was issued to support the implementation of the OIE Aquatic Animal Health Code (Hanwell et al., 2016). Zebrafish have been experimentally infected by both intracoelomic injection and immersion challenges (Encinas et al., 2013; Lopez-Munoz et al., 2010; Ruyra et al., 2014; Sanders et al., 2003). As a result of SVCV experimental infection studies using zebrafish, the Canadian Food Inspection Agency placed restrictions on the importation of zebrafish models into Canada, increasing the difficulty of sharing genetically engineered zebrafish lines internationally, and therefore adversely impacting the Canadian biomedical research community (Hanwell et al., 2016).

Conclusions Zebrafish are susceptible to viral infections, as demonstrated by a naturally occurring outbreak of VER among zebrafish obtained from the ornamental fish trade for research purposes, an outbreak of clinical disease due to natural ISKNV infection in laboratory zebrafish (Bermudez et al., 2018), the recent discovery of a widely distributed and prevalent novel enteric picornavirus, ZfPV-1 (Altan et al., 2019), and experimental susceptibility to both fish and mammalian viruses. The relative lack of information available with

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respect to naturally occurring viral infections in zebrafish is surprising, considering the widespread use of zebrafish as both a model organism and an ornamental species. Many facilities now only allow the introduction of surface-disinfected embryos into main systems to exclude new or unknown pathogens (Kent et al., 2009). However, zebrafish were shared among investigators at various institutions for many years before this practice was common, and viruses that are vertically transmitted might not be eliminated by surface disinfection. The conditions of zebrafish culture in rack systems with tanks plumbed in parallel also reduce pathogen transmission among fish (Crim et al., 2017). The lack of information available about naturally occurring viral infections primarily reflects limited diagnostic data and scientific investigation in this area (Crim & Riley, 2012). Historically, only a small number of research institutions implemented health monitoring programs for zebrafish colonies, with minimal diagnostic investigation of colony morbidity and mortality at most institutions. Moreover, the existing programs primarily relied on histopathology, which lacks sensitivity for viral infections, as viral inclusion bodies or pathognomonic lesions are not evident for many viral infections. Importantly, as the increasing use of metagenomic approaches will likely result in the discovery of novel viruses in zebrafish, the experimental investigation of viral epizootiology, pathogenesis, and transmission in immunocompetent and immunocompromized zebrafish lines will be extremely important to inform biosecurity practices in zebrafish facilities. Husbandry and health monitoring practices for zebrafish are currently immature relative to those for rodent models, although that is beginning to change. As the zebrafish continues to grow in importance as a model organism, greater investigation into naturally occurring viral diseases and reliance on evidence-based biosecurity measures will be necessary and critical for protecting zebrafish health, welfare, and data integrity.

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Sanders, J. L., Watral, V., Clarkson, K., & Kent, M. L. (2013). Verification of intraovum transmission of a microsporidium of vertebrates: Pseudoloma neurophilia infecting the Zebrafish, Danio rerio. PLoS One, 8(9), e76064. https://doi.org/10.1371/journal.pone.0076064. Sanders, J. L., Zhou, Y., Moulton, H. M., Moulton, Z. X., McLeod, R., Dubey, J. P., et al. (2015). The zebrafish, Danio rerio, as a model for Toxoplasma gondii: An initial description of infection in fish. Journal of Fish Diseases, 38(7), 675e679. https://doi.org/10.1111/jfd.12393. Sano, T., Morita, N., Shima, N., & Akimoto, M. (1991). Herpesvirus cyprini: Lethality and oncogenicity. Journal of Fish Diseases, 14(5), 533e543. Saraceni, P. R., Romero, A., Figueras, A., & Novoa, B. (2016). Establishment of infection models in zebrafish larvae (Danio rerio) to study the pathogenesis of Aeromonas hydrophila. Frontiers in Microbiology, 7, 1219. https://doi.org/10.3389/fmicb.2016.01219. Seeley, R. J., Perlmutter, A., & Seeley, V. A. (1977). Inheritance and longevity of infectious pancreatic necrosis virus in the zebra fish, Brachydanio rerio (Hamilton-Buchanan). Applied and Environmental Microbiology, 34(1), 50e55. Shen, C. H., & Steiner, L. A. (2004). Genome structure and thymic expression of an endogenous retrovirus in zebrafish. Journal of Virology, 78(2), 899e911. Shi, J., Zhang, H., Gong, R., & Xiao, G. (2015). Characterization of the fusion core in zebrafish endogenous retroviral envelope protein. Biochemical and Biophysical Research Communications, 460(3), 633e638. https://doi.org/10.1016/j.bbrc.2015.03.081. Song, J. Y., Kitamura, S., Jung, S. J., Miyadai, T., Tanaka, S., Fukuda, Y., et al. (2008). Genetic variation and geographic distribution of megalocytiviruses. Journal of Microbiology, 46(1), 29e33. Suckow, M. A., Weisbroth, S. H., & Franklin, C. L. (2005). The laboratory rat. Academic Press. Sudthongkong, C., Miyata, M., & Miyazaki, T. (2002). Iridovirus disease in two ornamental tropical freshwater fishes: African lampeye and dwarf gourami. Diseases of Aquatic Organisms, 48(3), 163e173. https://doi.org/10.3354/dao048163. Su, Y., Xu, H., Ma, H., Feng, J., Wen, W., & Guo, Z. (2015). Dynamic distribution and tissue tropism of nervous necrosis virus in juvenile pompano (Trachinotus ovatus) during early stages of infection. Aquaculture, 440, 25e31. Thiery, R., Cozien, J., Cabon, J., Lamour, F., Baud, M., & Schneemann, A. (2006). Induction of a protective immune response against viral nervous necrosis in the European sea bass Dicentrarchus labrax by using betanodavirus virus-like particles. Journal of Virology, 80(20), 10201e10207. https://doi.org/10.1128/ JVI.01098-06.

Valero, Y., Arizcun, M., Esteban, M. A., Bandin, I., Olveira, J. G., Patel, S., et al. (2015). Nodavirus colonizes and replicates in the testis of Gilthead seabream and European sea bass modulating its immune and reproductive functions. PLoS One, 10(12), e0145131. https://doi.org/10.1371/journal.pone.0145131. Wagner, H., Simon, D., Werner, E., Gelderblom, H., Darai, C., & Flugel, R. M. (1985). Methylation pattern of fish lymphocystis disease virus DNA. Journal of Virology, 53(3), 1005e1007. Wang, Y., Zhang, H., Lu, Y., Wang, F., Liu, L., Liu, J., et al. (2017). Comparative transcriptome analysis of zebrafish (Danio rerio) brain and spleen infected with spring viremia of carp virus (SVCV). Fish and Shellfish Immunology, 69, 35e45. https://doi.org/10.1016/ j.fsi.2017.07.055. Wang, Y. Q., Lu, L., Weng, S. P., Huang, J. N., Chan, S. M., & He, J. G. (2007). Molecular epidemiology and phylogenetic analysis of a marine fish infectious spleen and kidney necrosis virus-like (ISKNVlike) virus. Archives of Virology, 152(4), 763e773. https://doi.org/ 10.1007/s00705-006-0870-4. Weber, E. S., 3rd, Waltzek, T. B., Young, D. A., Twitchell, E. L., Gates, A. E., Vagelli, A., et al. (2009). Systemic iridovirus infection in the Banggai cardinalfish (Pterapogon kauderni Koumans 1933). Journal of Veterinary Diagnostic Investigation, 21(3), 306e320. doi: 21/3/306 [pii]. Willis, D. B., & Granoff, A. (1980). Frog virus 3 DNA is heavily methylated at CpG sequences. Virology, 107(1), 250e257. Xiang, Z., Dong, C., Qi, L., Chen, W., Huang, L., Li, Z., et al. (2010). Characteristics of the interferon regulatory factor pairs zfIRF5/7 and their stimulation expression by ISKNV Infection in zebrafish (Danio rerio). Developmental and Comparative Immunology, 34(12), 1263e1273. https://doi.org/10.1016/j.dci.2010.07.003. Xiong, X. P., Dong, C. F., Xu, X., Weng, S. P., Liu, Z. Y., & He, J. G. (2010). Proteomic analysis of zebrafish (Danio rerio) infected with infectious spleen and kidney necrosis virus. Developmental and Comparative Immunology, 35(4), 431e440. doi:S0145-305X(10)00274-0 [pii] 10.1016/j.dci.2010.11.006. Xu, X., Zhang, L., Weng, S., Huang, Z., Lu, J., Lan, D., et al. (2008). A zebrafish (Danio rerio) model of infectious spleen and kidney necrosis virus (ISKNV) infection. Virology, 376(1), 1e12. doi:S00426822(07)00829-X [pii]10.1016/j.virol.2007.12.026. Yoshikoshi, K., & Inoue, K. (1990). Viral nervous necrosis in hatcheryreared larvae and juveniles of Japanese parrotfish, Oplegnathus fasciatus (Temminck & Schlegel). Journal of Fish Diseases, 13(1), 69e77.

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43 Nonexperimentally Induced Neoplastic and Proliferative Lesions in Laboratory Zebrafish Sean T. Spagnoli1, Katrina N. Murray2 1

Biomedical Sciences, Oregon State University College of Veterinary Medicine, Corvallis, OR, United States of America; 2 Zebrafish International Resource Center, University of Oregon, Eugene, OR, United States of America

Introduction When considering neoplasia in any species, it is best to begin by examining the cell populations most likely to become neoplastic. In mammals, we are broadly familiar with common neoplasms arising from rapidly dividing cell populations: Lymphosarcoma from replicating lymphocytes, squamous cell carcinomas from high-turnover epithelial cells, and intestinal adenocarcinomas from the labile intestinal epithelium. By applying these basic principles correlating neoplasia risk with rapidly replicating cell populations, it might be assumed that we can make reasonable predictions concerning the types of neoplasms we might observe in any species. However, because of the wide spectrum of factors including environmental influences, genetic predispositions, and the role of chance in the development of neoplasms, it is nearly impossible to predict the types and frequencies of neoplasms that may develop between different species. For example, mast cell tumors, which are common in dogs, are exceedingly rare in humans. While all species can develop teratomas and nephroblastomas, frequencies vary between species. Furthermore, embryonic neoplasias and choristomas (the growth of normal tissue in abnormal places) of certain types can be common in some species but unheard-of in others. In human medicine, we tend to think of the development of cancers secondary to exposure to certain environmental factors, such as toxins that can act as initiators and promoters. However, viruses like hepatitis and human papillomaviruses are responsible for many cases of neoplasia, particularly in the developing world. In the realm of veterinary medicine, viral infections are known to cause a wide range of neoplasms. Bovine

The Zebrafish in Biomedical Research https://doi.org/10.1016/B978-0-12-812431-4.00043-9

leukemia virus, avian lymphoid leucosis, equine sarcoid, and plasmacytoid leukemia are just a few examples of neoplasms associated with viral infections (Truyen & Lochelt, 2006). Nonviral infectious entities, such as gastric carcinomas in humans caused by Helicobacter pylori (Correa & Blanca Piazuelo, 2011), should also be considered. In the following chapter, we will discuss both common and uncommon zebrafish tumors observed in zebrafish research facilities, focusing on pathogenesis and comparative pathology. Since zebrafish are a relatively recent introduction to the lab animal stable, we are continually discovering new trends and disease entities. Therefore, this chapter is written with the support of peer-reviewed data where possible, but we also introduce newer, previously unpublished entities that the authors have observed during their tenures as researchers and diagnosticians. Diagnosis of neoplasia in zebrafish relies primarily on microscopic or histological techniques because grossly visible lesions and readily identifiable clinical signs are generally nonspecific. The most common clinical signs of neoplasia in zebrafish are emaciation with or without scoliosis, both of which are nonspecific. Similarly, a distended abdomen could indicate a coelomic tumor; however, this is the primary differential only in males. In females, it is more likely that this represents egg-binding (egg-associated inflammation with fibroplasia) or simple gravidity. In continuing with the trend, exophthalmia can be due to etiologies that include neoplasia, granulomatous inflammation, or gas bubble disease (See chapter on Water Quality and Idiopathic Diseases of Laboratory Zebrafish Murray et al.). Even mass-like lesions have multiple possible etiologies. White or pink dermal masses can be neoplasms but

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are just as likely to be granulomas due to mycobacterial infection, parasitic infections, or transdermal migration of egg-associated inflammatory debris. The only relatively reliable gross signs of neoplasia in zebrafish are dark-colored external masses, which tend to be diagnosed as melanomas, and ventral mandibular lesions appearing in multiple fish with thyroid lesions (either hyperplasia or neoplasia). Although these clinical presentations are more likely to be neoplastic than inflammatory, histology is still needed for confirmation. Unlike mammalian neoplasms, naturally occurring malignancies in zebrafish rarely metastasize. Intestinal adenocarcinomas can spread through the coelom by carcinomatosis, and thyroid neoplasia may occur as a solitary mass with additional coelomic foci of proliferative follicles, but it is unclear whether this represents metastasis or activation of ectopic tissue (Spitsbergen, Buhler, & Peterson, 2012; Fournie, Wolfe, Wolf, Courtney, Johnson, & Hawkins, 2005). Hematopoietic tumors, such as lymphosarcoma, can spread to multiple organs, but are commonly considered multicentric, rather than metastatic. In general, spontaneous tumors in zebrafish can result in severe organ compression and displacement and aggressive local invasion but for most tumors, vascular metastasis has not been confirmed. Because of the immense utility of zebrafish for studies of genetic manipulation, toxicology, and xenotransplantation, the number of potential neoplasms in these animals is effectively infinite (Langenau, 2016). Therefore, this chapter will only focus on neoplasms that we consider to be “naturally occurring” in cultivated zebrafish strains used for experimental work, that is, neoplasms that have arisen in multiple fish from multiple laboratories over time in the absence of any consistent experimental manipulation. Where information is available about potential infectious or environmental etiologies, we include a section on methods for control and treatment. However, as most tumors identified outside of carcinogenesis studies arise spontaneously, no methods of prevention or treatment in laboratory zebrafish are currently known. Although a complete experimental history is often not provided, it is important to consider the previous genetic manipulations that may give rise to unexpected lesions and presentations. The images in this chapter, as well as most of the information presented, have been gleaned from the authors’ careers along with the ZIRC zebrafish diagnostic service’s archives, representing 18 years’ worth of cases from around the world. We would like to recognize, in this chapter, the contributions of Jan Spitsbergen to the field with her seminal paper on zebrafish neoplasia (Spitsbergen et al., 2012). Please refer to Table 43.1 for details regarding the types and relative frequencies of tumors identified by the ZIRC diagnostic service.

TABLE 43.1 Tumors, hyperplastic, and dysplastic lesions diagnosed at the ZIRC zebrafish diagnostic service between 2006 and 2016. Total number of fish examined by histology: 16,169 (Includes fish from ZIRC quarantine and husbandry rooms, internal sick fish, and nonclinical sentinels). Intestinal epithelial hyperplasia, dysplasia, dysplasia, or neoplasia

322

Seminoma

182

Biliary or pancreatic duct hyperplasia

169

Ultimobranchial gland hyperplasia, adenoma, or adenocarcinoma

82

Thyroid hyperplasia, adenoma, or adenocarcinoma

52

Melanoma

5

Chordoma

20

Hepatocellular carcinoma

17

Peripheral nerve sheath tumor

52

Spindle cell tumor (nonspecific)

11

Lymphoma/lymphosarcoma

14

Nephroblastoma/ependymoma

10

Hemangioma

7

Seminomas Description The most common neoplasm reported in zebrafish is the seminoma (Spitsbergen et al., 2012). In mammals, seminomas are derived from primitive male germ cells and appear as disorganized sheets of spherical spermatogonia-like cells with marked variation in cell and nucleus size. Normal cellular architecture is generally obliterated, and the absence of mature spermatozoa in the masses likely indicates that these tumors are nonfunctionaldthat is, no fertile sperm is produced. Mitotic rates are generally high, and these neoplasms have the potential for local infiltration, as well as distant metastasis (Hayes & Sass, 2012). In contrast, in zebrafish, most of the putative seminomas that we have observed exhibit minimal deviation from normal testis tissue at a histological level, presenting a challenge for their diagnosis as a true neoplasm (Figs. 43.1e43.4).

Pathobiology and Clinical Signs Seminomas, when evident macroscopically, usually present as swollen abdomens in males. When opened, the coelom is filled by a pink, fleshy mass. Larger masses with more mature sperm appear less pink. Many seminomas in zebrafish, however, are incidental and are

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FIGURE 43.1 Normal testis H&E. (A) 200 magnification. (B) 400 magnification.

(B)

(A)

(C)

FIGURE 43.2 Common well-differentiated seminoma. (A and B) While the structure of the mass appears similar to that of the normal testis (Fig. 43.1), it can often take up the majority of the coelomic cavity. H&E. 20. (B) H&E. 200. (C) Seminomas not uncommonly contain female germ cells at various primordial stages of development (Arrow). Vitellogenic oocytes are rare to nonexistent in these tumors. H&E 400.

discovered on microscopic evaluation of clinically normal fish. It is unclear whether these tumors represent functional testicular tissue, and their effect on fecundity has not been measured. However, the masses are composed of all normal cell types, including mature sperm. At the Zebrafish International Resource Center (ZIRC) we occasionally see presumptive seminomas during testis dissection for sperm cryopreservation, and these testes yield greater than normal amounts of sperm. However, the sperm from these presumptive seminomas has not been individually assayed for activity and viability in in vitro fertilization as ZIRC pools sperm samples from several fishes to cryopreserving a line. A distinct etiology for spontaneously occurring 2tumors of this type has not been identified; however,

hormonal influences are likely contributors. The ZIRC diagnostic service typically diagnoses seminomas in 1%e2% of the submitted fish per year. To date, there is no documented zebrafish line predisposition. Although the masses can be quite large, filling the majority of the coelomic cavity, they are not typically associated with loss of body condition or signs of compromise to other organs.

Diagnosis Diagnosis is made based on postmortem histology. As many of the extremely enlarged testes appear essentially normal at a microscopic/tissue level, it raises the question of whether or not these are true neoplasms. A

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FIGURE 43.3 (AeD) Some well-differentiated seminoma variants have complete sperm maturation, but earlier stages of spermatogenesis make up the majority of the tumor. (A) H&E 40. (B) H&E 200. (C) H&E 400. (D and E) Other variants lack maturation beyond early germ cell stages, and no spermatozoa are present. (D) H&E 200. (E) H&E 400. (F) Ovarian dysgerminoma. H&E 200.

(B)

(A)

(C)

FIGURE 43.4 Cystic seminomas. Even within a single seminoma, different areas can have variable degrees of spermatogenic maturation. (A) H&E 20. (B) H&E 40x. (C) H&E 40.

neoplasm is defined as a pathological overgrowth of abnormal cells, while hyperplasia is defined as either a physiologic or pathologic overgrowth of normal cells. As stated previously, there is a fine line between benign neoplasia and hyperplasia, particularly because hyperplasia generally predisposes a cell line to the development of neoplasia. While we currently call this particular entity a seminoma, the presence of normal,

albeit enlarged, testicular tissue could indicate that, rather than a neoplastic process producing a benign tumor of testicular tissue, it is, in fact, diffuse hyperplasia of the testis in these animals. Conversely, evidence that these are neoplasms (i.e., well-differentiated benign neoplasia) is supported by the fact that they grow into large, ovoid, multinodular, expansive masses in which the overall architecture of

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Intestinal Carcinomas

FIGURE 43.5 Intestine. (A) Normal intestine is lined by a single layer of columnar epithelial cells. Unlike the mammalian intestine, the zebrafish intestine has an extremely limited submucosal layer and very few resident leukocytes H&E 200. (B) Enteritis with intestinal hyperplasia. As opposed to the smooth profile of intestinal folds in (A), the mucosal surface is thickened and irregular, and there is “stacking” of nuclei. The submucosa is expanded by moderate numbers of leukocytes. H&E 200. (C) This intestinal adenocarcinoma presents as a severely infiltrative mass composed of duct- and tubule-like structures lined by epithelial cells embedded in a scirrhous and inflamed stroma. H&E 100x.

the testis is markedly distorted (although there is spermatogenic progression and some tubule-like structures occasionally remain). In Japanese medaka, similar well-differentiated testicular lesions do metastasize (J. Wolf, EPL, pers. comm.), an indicator of malignancy. Finally, hyperplastic lesions theoretically regress following the removal of the stimulus for proliferation. To date, regression has not been documented, but this would admittedly be hindered by the fact that many of these masses are diagnosed by postmortem histology on clinically normal zebrafish. Less common than the “hyperplastic-type” seminomas are a group of seminomas that are composed primarily of earlier sperm developmental stages. These can either have a solid or a cystic appearance, and portions of these tumors may undergo differentiation into normal-appearing testicular tissue with the patchy appearance of mature spermatozoa. It is also not uncommon to find various stages of oocyte development in seminomas, which also contributes to the argument against a hyperplastic process. Dysgerminomas, primitive tumors of oocytes, have been observed in female zebrafish, although they are far less common than seminomas in males. These appear similar to mammalian dysgerminomas: They are infiltrative, aggressive tumors composed of pleomorphic spherical cells in contrast to more well-organized zebrafish seminomas.

Control and Treatment It is unknown whether or not well-differentiated testicular masses are responding to some type of hormonal stimulation. If multiple seminomas are diagnosed in a tank or location, exposure to an external stimulus

should be considered. Spitsbergen et al. noted that seminomas are one of the most frequently diagnosed tumors associated with carcinogen exposure (Spitsbergen et al., 2012).

Intestinal Carcinomas Description No neoplastic entity in these animals demonstrates the progression from inflammation to hyperplasia to neoplasia better than the intestinal carcinoma. While some intestinal carcinomas may arise spontaneously, a distinct linkage between chronic inflammation and the development of intestinal carcinomas has been observed in zebrafish. Furthermore, it has been shown that these tumors, or rather, their as-yet-unidentified tumorcausing agent is transmissible, and animals in the transmission study showed distinct stages of chronic enteritis, epithelial hyperplasia, dysplasia, and finally, neoplasia (Burns et al., 2018; Paquette et al., 2015). Burns et al. (2018) showed that these tumors are caused by a transmissible agent. They also found a single 16S rRNA 30 sequence, which was most similar to Mycoplasma penetrans, to be highly enriched in the donors and recipients compared to disease-free controls using general microbiome evaluations. Furthermore, the original donor fish and exposed fish populations were positive for the Mycoplasma sp. using a targeted PCR test, while corresponding unexposed control fish were negative. Whereas this Mycoplasma sp. was directly associated to the neoplasm, a direct cause and effect link has yet to be verified. Paquette et al. (2013) showed that the occurrence of these intestinal tumors was not

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directly related to diet or zebrafish strain, and they usually occur in fish older than 1 year. The intestinal nematode Pseudocapillaria tomentosa is another transmissible cause of intestinal cancer resulting from chronic inflammatory processes rather than direct change to the genome as with viruses (Kent, Bishop-Stewart, Matthews, & Spitsbergen, 2002; Spitsbergen et al., 2012). There does not appear to be any recognizable difference morphologically between Mycoplasma-associated, Pseudocapillaria-associated, and spontaneously arising intestinal neoplasms (Figs. 43.5e43.7). Intestinal carcinomas in zebrafish are associated with general malaise, including lethargy and emaciation (Burns et al., 2018), but the large retrospective survey conducted by Paquette et al. (2013) showed that the tumors also frequently occur in subclinical fish. There are no obvious macroscopic changes in affected fish, even during necropsy when the intestines are closely examined.

Diagnosis Diagnosis of intestinal carcinomas in zebrafish can be difficult as the most common type of intestinal carcinoma is considered the “small cell” carcinoma (Spitsbergen et al., 2012). These neoplasms tend to manifest subgrossly as thickenings in the wall of the gut with loss of intestinal folds. Upon close examination, one can observe small nests of one to three cells invading transmurally, often infiltrating the coelomic cavity and spreading along serosal surfaces (carcinomatosis). Paquette et al. (2015) provided evidence that the tumors are of epithelial origin based on immunohistochemistry using epithelial and neural mammalian antibodies. Frequently, these neoplasms are associated with intestinal perforation. The difficulty of identifying small cell intestinal carcinomas is threefold: neoplastic cells are usually severely atypical and bear little resemblance to normal intestinal epithelium, severe inflammation and chronic fibrosis can obscure neoplastic cells, and the distortion and thickening of the gut wall secondary to the desmoplastic

FIGURE 43.6 Transmural invasion of intestinal adenocarcinoma. (A) Intestinal folds have been replaced by an infiltrative neoplasm. Tubular structures lined by epithelium are observed within the tunica muscularis (arrows). H&E 100. (B) Intestinal folds are blunted and effaced by the infiltrative neoplasm lined by a layer of hyperplastic epithelium (star). Tubular structures extend through the tunica muscularis (arrow) and into the serosal surface (arrowhead). The separation of the overlying epithelium from the lamina propria in this image is likely artifactual. H&E 200.

FIGURE 43.7 Duct-like versus small cell adenocarcinomas. (A) Duct-like adenocarcinomas continue to form tubular/duct-like structures with distinct lumens (arrow). H&E 400x. (B) Small cell adenocarcinomas no longer form duct-like structures but form infiltrative clusters of polygonal cells (arrow) that could be mistaken for inflammatory cells. H&E 400.

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response can present a tangential face to the histologic section. That is, rather than receiving a full transverse section of gut, which is either circular or ovoid with a lumen present, the microtome “grazes” the edge of the thickened gut which shows portions of muscularis and lamina propria with no readily apparent lumen. In this way, it becomes difficult to identify the characteristic features of the tumor. It is therefore important to consider deeper recuts of samples that contain suspect carcinomas. There exist more adenomatous-type intestinal carcinomas, which appear similar to the intestinal epithelium and recapitulate intestinal fold structure. They are less common, but just as invasive as small cell carcinomas (Burns et al., 2018). Differentiation of hyperplastic lesions from neoplastic lesions is based on invasion of neoplastic cells into the tunica muscularis, as preneoplastic atypical hyperplastic cells generally remain within the lamina propria.

Control and Treatment Intestinal neoplasms can arise spontaneously, in which case no control measures exist. If enteritis is observed in multiple fish with occasional neoplasms, P. tomentosa should be ruled-out. Treatments with ivermectin, emamectin, and fenbendazole have been described for P. tomentosa infections (Collymore et al., 2014; Maley, Laird, Rinkwitz, & Becker, 2013). If multiple fish in a tank are diagnosed with preneoplastic and neoplastic lesions of the gut, transmissible tumors should be considered when P. tomentosa has been eliminated as a differential. This is an active area of research, and currently, no treatment is recommended. Although a particular16S rRNA sequence similar to M. penetrans has been associated with the tumors (Burns et al., 2018), it is unknown whether the identified Mycoplasma sp. is always pathogenic or if there are particular strains of Mycoplasma, some of which are pathogenic and some of which may be considered normal flora (Fig. 43.8).

Ultimobranchial Gland Description The ultimobranchial gland in cyprinids is located just ventral to the musculature of the pharynx. It produces calcitonin, and it functions in osmotic regulation and calcium homeostasis. Pathobiology and Linical Signs Ultimobranchial hyperplasia, adenomas, and adenocarcinomas occur in laboratory zebrafish. Presently we have not identified a specific line or age of zebrafish that is predisposed to the lesions. Zebrafish in the wild in India live in water with a wide range of conductivity

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levels, 10e271 mS (Engeszer, Patterson, & Parichy, 2007) compared to those routinely used in zebrafish facilities (Chapter 29). The main action of calcitonin, which is produced by the ultimobranchial organ, is to increase bone calcium content and decrease the blood calcium level when it rises above normal concentrations. Hence, while no distinct etiology has been proven, electrolyte and osmotic imbalances are possible stimulants for hyperplasia based on the organ’s function. Chronic hyperplasia is a risk factor for neoplasia. Hyperplastic and neoplastic changes to the ultimobranchial gland are not typically visible during postmortem examination and are not generally associated with clinical disease. Diagnosis Ultimobranchial gland lesions are typically diagnosed by evaluation of histological sections. In paramedian sections of bisected animals, the gland is frequently small enough that it is out of the plane of section and therefore cannot be evaluated. It can, however, become enlarged up to three times normal size (up to roughly 50 mm), revealing a number of ducts or acini with central lumens lined by columnar epithelium. These lesions are another important example of the fine line between hyperplasia and neoplasia: Often, an ultimobranchial gland adenoma has the appearance of a diffusely enlarged, well-differentiated gland. In other words, much like the common hyperplastic-type seminoma, there is rarely a distinct area, where one can differentiate normal from tumor tissue. What may be considered a neoplastic process may actually be a hyperplastic process in response to some external environmental stimulus. As with seminomas, it is not known if the lesions regress with the removal of environmental stimuli. Lesions that are obviously anaplastic with a high mitotic index are appropriately diagnosed as adenomas. A diagnosis of adenocarcinoma can be made if neoplastic cells invade surrounding tissues. Control and Treatment If ultimobranchial gland pathology is routinely diagnosed in a fish population, it may be appropriate to consider environmental stimuli that could initiate these changes. While it has not been definitively proven, it is possible that one cause of nephrocalcinosis could be aberrant calcium metabolism. It may be prudent to monitor for coincidences of ultimobranchial gland pathology and nephrocalcinosis. Ultimobranchial hypertrophy has been associated with migration and spawning in the Atlantic salmon (Deville & Lopez, 1970). We have not noticed a predilection for ultimobranchial changes associated with any particular line, sex, or the age of laboratory zebrafish. However, in the laboratory, conditions are kept relatively constant, and fish are spawned throughout the year.

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FIGURE 43.8

Ultimobranchial gland lesions. (A) Normal gland. H&E 200. (B) Hyperplastic ultimobranchial gland. Note the increased size compared to the gland in (A) along with the irregular margins. H&E 100. (C) Ultimobranchial gland adenoma. Note the extremely large size and irregular appearance of cells. H&E 200. (D) Ultimobranchial gland adenoma. Simple to pseudostratified columnar epithelial cells surround small tubular lumens. The columnar cells have basilar nuclei and slightly flocculent pale eosinophilic cytoplasm. H&E 400.

Whether ultimobranchial gland anatomy and physiology fluctuates in zebrafish in the wild has not been reported (Fig. 43.9).

Lymphosarcoma and Other Hematopoietic Tumors Description Lymphoid tumors have been reported in numerous wild and captive fishes. As with mammals and birds, at least some of these neoplasms are caused by oncogenic viruses (Coffee, Casey, & Bowser, 2013). Zebrafish have proven to be useful models of hematopoietic neoplasias (Langenau et al., 2003; Rasighaemi, Basheer, Liongue, & Ward, 2015; Zhuravleva et al., 2008). Infectious agents have yet to be associated with naturally occurring lymphoid neoplasms in zebrafish. Pathobiology and Clinical Signs Lymphosarcoma in zebrafish is a highly aggressive neoplasm derived from lymphocytes. T lymphocytes originate in the thymus, while B lymphocytes, along with other erythroid and myeloid cells, are produced in the kidney. Fish with lymphosarcoma most commonly present with emaciation and nonspecific

signs of illness. When lesions are grossly visible, they can present as discrete masses, or diffuse epidermal invasion can result in skin pallor. A common gross manifestation of dermal lymphosarcoma is infiltration of the tissues surrounding the head, resulting in a “lionhead” appearance. Epidermal infiltration can also have an appearance similar to scale pocket edema, which makes the fish present as if it had “dropsy.” Exophthalmia may result a from retro-orbital accumulation of malignant cells, although this is a nonspecific finding as any retro-orbital, space-occupying lesion, including granulomas, can result in exophthalmia. Diagnosis This neoplasm is composed of leukocytes that do not recapitulate any particular tissue structure. Therefore, the most common appearance is that of an infiltrative process wherein sheets of cells with the appearance of lymphocytes invade, expand, and destroy normal tissues. Diagnosis is usually confirmed with postmortem histology. Impression smears of externally presenting masses may reveal sheets of monomorphic round cells with a high nuclear to cytoplasmic ratio. These cells generally have medium-sized nuclei and very little cytoplasm, making H&E stained tissues appear dark blue when viewed

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(D)

(C)

(B)

(A)

(F)

(E)

FIGURE 43.9 Lymphosarcoma. (A) Lymphosarcoma in the renal interstitium. The only leukocytes present are neoplastic lymphocytes which separate renal tubules. H&E 100x. (B) Lymphosarcoma in the liver. Sheets of neoplastic lymphocytes fill sinusoids and compress adjacent hepatic cords. H&E 100x. (C) Oral cavity. Sheets of neoplastic lymphocytes infiltrate just under the mucosal epithelium. H&E 400. (D) Whole body and skin. Neoplastic cells fill the coelom (bottom of the frame) and infiltrate the musculature just under the scales (top). H&E 40x. (E) Epidermis and dermis. Neoplastic lymphocytes invade beneath the scales and cause them to lift. H&E 100x. (F) Neoplastic lymphocytes arranged in sheets. Giemsa 400.

subgrossly. Many of these neoplasms likely arise from the renal interstitium, where most of the zebrafish hematopoiesis occurs. Microscopically, the renal interstitium is very commonly affected. In these cases, most to all other hematopoietic precursors have been replaced by neoplastic lymphocytes that expand the interstitial space and separate renal tubules. It is possible that lymphosarcoma that infiltrates the gills and the head arises from the thymus, but many fish with these lesions also have kidney lesions, and so a specific origin is unclear. While neoplastic lymphocytes can invade any and all organs of the zebrafish’s body, many of them appear to have a particular tropism for the epithelium, infiltrating just beneath the basement membrane of the epidermis, oral mucosal epithelium, and even the gills. Unlike the mammalian counterpart of epitheliotrophic lymphoma, however, neoplastic cells do not generally infiltrate between epithelial cells, and so-called Pautrier’s abscesses have not been observed. Other hematopoietic tumors have been observed in zebrafish during the authors’ tenures as diagnosticians. These neoplasms frequently appear as sheets of hematopoietic round cells, morphologically unlike lymphocytes, that have a similar appearance to hematopoietic

precursor cells, usually of the myeloid lineage. They are generally localized to the kidney and spleen, although neoplastic cells can be observed in vascular profiles, as in leukemias. Unfortunately, in the absence of a the distinct identity of a known genetic knockout and/or appropriate immunohistochemical stains, it is difficult to identify the particular cell type responsible for these neoplasms. In general, it helps to think of the renal interstitium as bone marrow. In a bone marrow biopsy, if adipocytes and the various stages of hematopoiesis are replaced by a single monomorphic population of round cells, a hematopoietic tumor is likely to be present. The same is true for the renal interstitium with the added benefit that one can also identify expansion by separation of renal tubules. Caution is warranted, in these cases, however, since interstitial expansion or hyperplasia can occur physiologically as a response to inflammation, causing tubules to be separated by a mixed population of normal hematopoietic cells. Differentiation between a hyperplastic and neoplastic hematopoietic population is made based on the presence of all hematopoietic stages (hyperplasia) or the presence of a monomorphic population of cells in a single stage (neoplasia).

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FIGURE 43.10 Peripheral nerve sheath tumor. (A) These tumors are composed of short, intersecting streams and bundles of spindle-shaped cells. H&E 200. (B) Spindle cells are stacked in a storiform pattern similar to those observed in mammals. H&E 400.

Control and Treatment To date, infectious agents have not been associated with lymphoid neoplasms in laboratory zebrafish. Although, it should be considered as a possible etiology if multiple fish are affected since tumors have been associated with retroviruses in other fishes (Coffee et al., 2013). The genetic background should also be considered, as lines have been engineered to recapitulate hematopoietic neoplasias (Rasighaemi et al., 2015) (Fig. 43.10).

Soft Tissue Srcoma/Peripheral Nerve Sheath Tumors Soft tissue sarcomas are one of the more common neoplasms diagnosed in laboratory zebrafish. These tumors can occur in any part of the body, and clinical presentation varies with affected tissues.

zebrafish can develop other types of soft tissue sarcomas, those morphologically similar to mammalian peripheral nerve sheath tumors are, by far, the most common and best described. When other soft tissue sarcomas are identified that do not fit this morphological pattern, comparison to similar tumors in other cyprinids is warranted. Again, due to the lack of validated immunohistochemical stains in zebrafish, identification of specific cells of origin for soft tissue sarcomas, including peripheral nerve sheath tumors, is difficult and diagnoses at this point in time are made based solely on morphology in histological sections. It is entirely possible that soft tissue sarcomas in zebrafish represent a broad range of cell types beyond the common “peripheral nerve sheath tumor.” Overall, we tend to refer to nonspecific soft tissue sarcomas as “spindle cell sarcomas” of course, providing that the primary cell type is spindle-shaped.

Pathobiology and Clinical Signs

Control and Treatment

These tumors can present as single or multiple swellings externally. Tumors that arise in the retrobulbar region cause massive, unilateral exophthalmia. Clinical signs can be absent or nonspecific and include emaciation and general malaise. If tumors are internally invasive, they can appear as amorphous white masses that surround and adhere to organs upon dissection.

A distinct etiology for these tumors in zebrafish is unknown, and they have not correlated these tumors with a particular wild-type line or sex. However, we often diagnose soft tissue sarcomas in zebrafish with mutations in the p53 gene (Fig. 43.11).

Optic Pathway Tumor

Diagnosis In zebrafish, the most common soft tissue sarcoma is the peripheral nerve sheath tumor, which generally appears as a solid, highly infiltrative, densely cellular mass composed of short streams and bundles of spindle-shaped cells. It is very similar in appearance to mammalian peripheral nerve sheath tumors, and most tumors bear a distinct storiform pattern. These neoplasms can be highly aggressive, invading, and effacing large amounts of normal tissue both in the exterior and the interior of the animal (Spitsbergen et al., 2012). While

Description An interesting tumor that does not seem common in other laboratory animal species is a neural tumor with multiple areas of differentiation that resemble various layers of the retina. The closest description in the literature is found in Stolin et al. (2014), wherein they called the tumors simply “optic pathway tumors” and found that they were likely of glial origindmost likely the multipotent Muller gliadbased on molecular and cellular characteristics.

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FIGURE 43.11 (A) Normal retina 200x. (B) Ocular pathway tumor with multiple areas of varied retinal differentiation. H&E 20. (CeF) demonstrate these varying areas. (C) Ganglion cells and nerve fibers. H&E 200. (D) Photoreceptor layer. H&E 400. (E) Retinal pigmented epithelium. H&E 200. (F) Plexiform and nuclear layers. H&E 200

Pathobiology and Clinical Signs Exophthalmia is the most common lesion associated with these tumors. They can appear as pink, fleshy masses behind the eye although this is nonspecific since any mass effect in orbit would have a similar appearance.

known; however, culling of animals with this lesion may prevent transmission of tumor-associated genes to offspring (Fig. 43.12).

Diagnosis Diagnosis is made via postmortem histology. These tumors can be highly variable in appearance based on the proportions of retinal layers represented in each lesion. These are considered distinct from retinoblastomas because there is no identified abnormality in rb1, and morphologically, the degree of differentiation into cells representing different retinal layers is far more advanced than the simple rosettes observed in well-differentiated retinoblastomas. Also, to the authors’ knowledge, there have been no confirmed diagnoses of spontaneously arising retinoblastomas in zebrafish.

Description Zebrafish can develop spontaneous melanomas. Although not as well known for these lesions as their Xiphophorus counterparts (Meierjohann & Schartl, 2006), zebrafish are also utilized for studies of melanoma biology. Several lines of zebrafish have been established with mutations in conserved genes affecting specification, differentiation, and function of melanocytes (Ceol, Houvras, White, & Zon, 2008).

Control and Treatment In the Stolin study (2014), these tumors developed in heterozygous Tg(flk1:RFP)is18 transgenic adult fish. While this means these tumors are likely caused by genetic abnormalities, a distinct etiology for spontaneously occurring tumors of this type has not been identified. Therefore, no methods of control or prevention are

Melanoma

Pathobiology and Clinical Signs A distinct etiology for spontaneously occurring tumors of this type has not been identified. Ultraviolet light is a possible contributor, as in mammals, although this has not been definitively shown in zebrafish. Mitchell, Paniker, and Douki, (2009) showed that early life stages of a Xiphophorus backcross hybrid developed melanomas following UVB exposure. Zebrafish studies may elucidate some of the outstanding genetic questions in melanoma biology (Ceol et al., 2008).

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43. Nonexperimentally Induced Neoplastic and Proliferative Lesions in Laboratory Zebrafish

FIGURE 43.12 Melanoma. (A) Dermal melanomas present subgrossly as a variably pigmented subdermal mass. H&E. 20. (B) The neoplasm is moderately invasive into the adjacent musculature (arrow). H&E. 100. (C) The neoplasm is composed of intersecting streams of spindle-shaped to stellate cells containing variable amounts of intracytoplasmic brown pigment granules.

These tumors can present anywhere inside or outside the body, usually as pigmented masses. Amelanotic melanomas may also occur, but their diagnosis is difficult and will be discussed later. Disruption of internal organs could result in physiological alterations that present as nonspecific morbidity, e.g., edema, lethargy, reduced spawning activity, and inanition. While locally aggressive, metastasis has rarely been confirmed outside of experimental conditions. Diagnosis Diagnosis is made on postmortem histology. As in mammals, neoplastic melanocytes can take on a number of appearances, but generally, these tumors appear as sheets of irregular, spindle-shaped to stellate and occasionally rounded cells containing variable amounts of intracytoplasmic melanin pigment. These tumors are generally locally infiltrative and may occasionally show neuroendocrine-type “packeting.” It is important to differentiate melanomas from other melanistic lesions, such as those associated with chronic inflammation. These lesions are generally composed of clusters of melanomacrophages, which are not always easy to differentiate from neoplastic melanocytes. Generally, melanomacrophages contain intracytoplasmic vacuolesdpresumably phagolysosomes containing degenerate melanin. Because these are the products of phagocytosis and digestion, pigmented granules in macrophages are relatively variable in size and shape, while granules in melanocytes are small and uniform in size. Also, while melanomacrophages can have a moderate amount of pleomorphism based on their ameboid characteristics, more frequently, they tend to have round to polygonal appearance with less pleomorphism than is usually observed in melanomas. Furthermore, the margins of inflammatory melanistic lesions will be relatively well demarcated or at least bounded by resident anatomical structures. This differs from melanocytic neoplasms, which tend to infiltrate between

the cells of surrounding anatomic structures. Of course, due to the variable appearance of both lesions, there is likely to be some morphological overlap. FontanaMasson staining can help in highlighting the shape of melanocytic granules, which may aid in differentiation. Amelanotic melanomas presumably occur spontaneously in zebrafish, although without a robust array of immunohistochemical stains, it is difficult to confirm these diagnoses. A presumptive diagnosis may be made based on the irregular appearance of a mesenchymal neoplasm that does not quite conform to any particular cell type morphologically. Neuroendocrine packeting may aid in this presumption, and FontanaMasson staining may reveal rare melanin granules. Due to the notorious variability of melanoma morphologies, it is possible that a large number of amelanotic melanomas in zebrafish may not be confirmed (Fig. 43.13).

Extraspinal Chordoma Description Like most teleosts, zebrafish retain prominent notochord remnants within their vertebrae for their entire lives. In mammals, the notochord remnant is reduced to the nucleus pulposa of the intervertebral disks. Chordomas are tumors of notochord remnants that most commonly affect the axial skeleton in humans, and in terms of domestic animal species, the tails of ferrets. Zebrafish are unique in that chordomas most frequently occur in the intestinal tract, although axial skeleton chordomas have also been reported (Cooper, Murray, Spagnoli, & Spitsbergen, 2015). Pathobiology and Clinical Signs While extraskeletal soft tissue chordomas are exceedingly rare in humans and mammals, they are considered to be a background lesion in aged zebrafish. Although not exceedingly common, extraspinal soft tissue chordomas

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FIGURE 43.13 Chordomas. (A) Normal vertebral body with notochord remnant (star). H&E 200. (B) Intestinal chordoma (star). H&E 40. (C) The chordoma arising in the lamina propria is composed of physaliphorous cells and elevates the overlying mucosal epithelium (arrow). H&E. 200.

in the zebrafish occur most frequently in the intestine (Cooper & Spitsbergen, 2016). Extraspinal chordomas generally do not elicit clinical symptoms. They are usually too small to be observed on gross postmortem examination. Whether or not the masses are large enough to disrupt digestion and feeding and potentially contribute to weight loss or other nonspecific signs associated with aging, is not known. A distinct etiology for spontaneously occurring tumors of this type has not been identified. Spitsbergen et al. speculated that sonic hedgehog b mutations could be involved in tumor development in the notochord and intestine (Cooper et al., 2015). Diagnosis Diagnosis is made on postmortem histology. Chordomas are multilobulated and contain large, vacuolated polygonal cells with fine cytoplasmic processes (physaliphorous cells). They are similar in appearance to the notochord remnants in the vertebrae (Fig. 43.14).

Hepatocellular Tumors Description Hepatocellular tumors have been described from countless wild fishes, particularly associated with polluted waters. Whereas viruses are important causes of liver tumors in humans and other mammals, they have only been linked to anthropogenic toxicants (Myers, Landal, Krahn, & McCain, 1991) or specific natural hepatotoxins, such as aflatoxin (Sinnhuber, Hedricks, Walse, & Putnam, 1997) in fish. They have also been experimentally induced in numerous laboratory studies, including those with zebrafish (Spitsbergen et al., 2012). Pathobiology and Clinical Signs These tumors arise from hepatocytes and can be discrete, focal masses in the liver. The degree of demarcation is highly variable, and often low-magnification

evaluation of the liver as a whole is the best method of initial identification. On necropsy, these tumors may appear as soft tissue swellings on the liver, but they are generally too small to be identified grossly. Diagnosis Diagnosis is made based on postmortem histology. Hepatocellular tumors in zebrafish can have a number of appearances, but are generally, poorly demarcated and composed of sheets of disorganized polygonal cells similar in appearance to, but smaller or larger than, adjacent hepatocytes. They can sometimes be difficult to differentiate from normal adjacent liver tissue, particularly in emaciated animals with generalized reduced hepatocellular cytoplasm. They are frequently associated with hemorrhage and severely vacuolated hepatocytes, and it can be difficult to discern characteristic neoplastic hepatocytes in the midst of hemorrhage and degenerate cells. Small patches of shrunken hepatocytes or vacuolated hepatocytes have also been observed within nonneoplastic livers, and it is unclear whether these represent degenerate changes or preneoplastic lesions. As mentioned in the Water Quality and Idiopathic Disease of Laboratory Zebrafish chapter (Murray et al. This book.), the relationship between hepatocellular karyomegaly and neoplasia is unclear. In mammals, it is extremely difficult to differentiate between hepatocellular adenomas and hepatocellular carcinomas, since they can have similar microscopic morphologies. Differentiation between benign and malignant tumors generally depends on the size and clinical behavior: Malignant neoplasms, which tend to be larger, metastasize while benign neoplasms do not. Since metastases in zebrafish tumors are rare in all tumor types, we cannot rely on clinical behavior for a diagnosis. Examining for characteristics of malignancy may, therefore, be the best way to distinguish between hepatocellular adenomas and carcinomas. Tumors with severe cellular and nuclear pleomorphism, high mitotic

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43. Nonexperimentally Induced Neoplastic and Proliferative Lesions in Laboratory Zebrafish

FIGURE 43.14 (A) Hepatocellular adenoma (star). The mass is an irregular aggregate of vacuolated hepatocytes paler than the adjacent normal tissue. H&E. 100. (B) Hepatocellular adenoma. Hepatocytes have vacuolated cytoplasm with moderate nuclear pleomorphism (star). H&E. 400. (C) Hepatocellular carcinoma. The mass is locally destructive and contains large areas of hemorrhage. H&E. 20. (D) Hepatocellular carcinoma. Hepatocytes display marked nuclear and cellular atypia (star). There are frequently multifocal regions of lipid-laden hepatocytes throughout these tumors (arrow). H&E 400.

rates, necrosis, and infiltration of the surrounding tissue, along with multicentric intrahepatic lesions and/or carcinomatosis are best-termed carcinomas. Small, discrete masses with cells similar in appearance to normal hepatic tissue are best-termed adenomas. The identification of benign hepatic lesions may be further complicated by an entity, common in aged canines, known as nodular hyperplasia. This is usually a benign, focal, or multifocal overgrowth of normal hepatic tissue with loss of portal architecture. In a case where multiple nodules of normally appearing hepatocytes are present, this would best be termed nodular hyperplasia. Distinguishing focal nodular hyperplasia from a hepatocellular adenoma is difficult and should be made based on the altered appearance of cells in the mass. To the authors’ knowledge, spontaneously occurring nodular hyperplasia has not been specifically identified in zebrafish. As it a laboratory animal, it is worth discussing the appearance of preneoplastic “altered foci,” particularly in the light of carcinogenesis studies. While basophilic, eosinophilic, clear cell, and mixed cell foci, are well described in mice, they are uncommon in zebrafish. Naturally occurring altered foci have been observed

anecdotally by the authors, but they have not been reported in the literature, nor is it clear if these lesions are preneoplastic in zebrafish. Control and Treatment Hepatocellular neoplasms, including those of fishes, are often caused by anthropogenic toxicants. However, it is well recognized that natural toxins, such as aflatoxin, cause such tumors in fish as well (Myers et al., 1991; Sinnhuber et al., 1997). A distinct etiology for spontaneously occurring tumors of this type in zebrafish has not been identified. Carcinogens most frequently affect the liver in both wild-type and mutant zebrafish (Spitsbergen et al., 2012) (Fig. 43.15).

Biliary and Pancreatic Ductal Lesions Description Liver and pancreatic tumors arising from ductal structures occur in laboratory zebrafish. These tumors must be differentiated from ductal hyperplasia, particularly in the context of toxicant research with regard to carcinogenesis.

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FIGURE 43.15

(A) Bile duct adenoma. Focally within the liver, there is a mass composed of tortuous ducts embedded in a dense collagenous stroma. H&E 200x. (B) Bile duct adenocarcinoma. The liver is severely expanded and infiltrated by a mass composed of tortuous dysplastic bile ducts, giving the liver a bulging, rounded appearance. H&E 20. (C) Biliary adenocarcinoma. Incomplete bile ducts are lined by atypical epithelial cells, often forming ducts filled with proteinaceous material. H&E 400.

Pathobiology and Clinical Signs Biliary and pancreatic ducts normally course through the parenchyma of their respective organs to the intestinal tract. Therefore, tumors arising from these ducts can involve any combination of the liver, pancreas, and intestine. These can rarely appear as masses that protrude above the liver capsule. Carcinomatosis is also possible. Clinical signs are generally nonspecific for emaciation and general malaise. Diagnosis Diagnosis is made based on postmortem histology. As in mammals, it is extremely difficult to differentiate biliary from pancreatic ductal tumors based on appearance alone. It is even more difficult in zebrafish due to the anatomical proximity of the pancreas and the liver, and so unless it is very clear that the tumor “belongs” to one organ or the other, it may be practical to lump the two entities together into simply a “ductal” tumor. These neoplasms are generally composed of tortuous, variably sized ducts lined by squamous to columnar epithelium and embedded in variable amounts of loosely organized fibrous connective tissue. They can be highly invasive. To further complicate matters, there is an entity in zebrafish wherein there is diffuse, presumably benign, hyperplasia of the biliary tree in which most of the ducts observed throughout the liver are tortuous, reduced in diameter, and lined by squamous to low cuboidal epithelium. They are generally associated with a moderate amount of fibrous connective tissue. They are frequently associated with granulomas that contain pigmented acellular material, presumably bile, although the material does not stain with Hall’s bile stain and is distinct in color and character from melanin (Spagnoli, Xue, & Kent, 2015). Rather than forming distinct masses with infiltration into the surrounding tissue, as is typical of ductal tumors, bile duct hyperplasia occurs

throughout the entire biliary tree and atypical cellular features are generally absent. These hyperplastic populations may be preneoplastic, although this has not been demonstrated. Most lesions of diffuse biliary hyperplasia tend to occur in clinically normal fish submitted for sentinel screening. While biliary adenomas and carcinomas are focal to multicentric lesions that are either well demarcated or locally infiltrative, biliary hyperplasia occurs diffusely throughout the biliary tree without forming discrete nodules. Furthermore, bile ducts in biliary hyperplasia generally lack cellular atypia, mitotic figures, and necrosis. Control and Treatment A distinct etiology for spontaneously occurring tumors of this type has not been identified. However, TL line fish appear to be particularly predisposed to bile duct hyperplasia (Spitsbergen et al., 2012). This should be considered when choosing a line for toxicology and carcinogenesis studies where the liver is expected to be affected (Fig. 43.16).

Nephroblastoma/Ependymoma Description Nephroblastomas have been reported in a number of fish species and after lymphoma, are the second most common primary renal neoplasms in fish (Lombardini, Hard, & Harshbarger, 2014). These tumors have been associated with water or environmental factors in Japanese eel (Masahito, Ishikawa, Okamoto, & Sugano, 1992) and carcinogen exposure in trout (Bailey, Williams, & Hendricks, 1996). Given the described response in rainbow trout, Spitsbergen et al. noted the remarkable lack of nephroblastomas in zebrafish exposed to carcinogens (Spitsbergen et al., 2000).

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(A)

(B)

(C)

(D)

(E)

(F)

FIGURE 43.16 (A) Normal kidney tubules. H&E 400. (B) Normal spinal cord with central canal lined with ependymal cells. H&E 400. (C) Nephroblastoma. Note the primitive glomerular structures embedded in the mesenchymal cell population (arrow). H&E 200. (D) The majority of the tumor is composed of epithelium-lined tubules (arrow) embedded in dense primitive mesenchyme (blastema, star). (E) Ependymoma. A highly infiltrative mass effaces the spinal cord and invades surrounding tissue. H&E 40. (F) The neoplasm is composed of tubules recapitulating the central canal (arrow) embedded in a matrix that resembles neuropil. H&E 400.

Pathobiology and Clinical Signs These tumors can cause scoliosis, kyphosis, or lordosis. Otherwise, clinical signs are generally nonspecific for emaciation and general malaise. A distinct etiology for spontaneously occurring tumors of this type has not been identified. Presently, we have not recognized any predilection for these tumors related to sex or a particular line of zebrafish. Mutations in WT1, encoding Wilms tumor protein, results in pediatric renal cancer in humans. Two paralogs, wt1a and wt1b, have been identified in zebrafish and activation experiments indicate that they have different functions in renal development compared to their human counterparts (Perner, Englert, & Bollig, 2007).

Diagnosis Diagnosis is made based on postmortem histology. These tumors can exceedingly difficult to differentiate from each other, due to the proximity of the spinal cord and kidney as well as their similar appearances. It is for this reason that they are discussed in a combined category here. Generally, these tumors contain both mesenchymal and epithelial components that have the appearance of tubules lined by high cuboidal epithelium embedded in sheets of poorly differentiated spindleshaped cells. If glomerular structures are observed in the tumor, then it can definitively be called a nephroblastoma. However, these structures can be difficult to observe microscopically in normal zebrafish, and their

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FIGURE 43.17 (A) Normal periocular vascular rete. H&E 200. (B) Periocular hemangioma (star). H&E 40. (C) Intramuscular hemangioma (star). H&E 20. (D) Hemangioma composed of variably sized, blood-filled cavities separated by thin connective tissue trabeculae lined by flattened endothelial cells (arrow). H&E 400.

absence does not necessarily rule out nephroblastoma. Neoplastic tubules in nephroblastomas are also generally surrounded by primitive blastema which has a densely packed appearance compared to ependymomas, which may contain components similar to neuropil. Ependymomas may also form rosettes. Both of these neoplasms are highly aggressive and can invade and efface adjacent kidney, musculature, spinal cord, and vertebrae.

Hemangioma Description Hemangiomas are benign tumors of endothelial structures. Carcinogen-associated and spontaneous hemangiomas have been described in several different fishes, including zebrafish (Couch, 1995; Fournie, Herman, Couch, & Howse, 1998; O’Hagan & Raidal, 2006; Spitsbergen et al., 2000). Pathobiology and Clinical Signs Hemangiomas arise from endothelial cells. They have been described in the heart, gills, liver, and in the retrobulbar space (Couch, 1995; Grizzle & Thiyagarajah,

1988; O’Hagan & Raidal, 2006), but presumably can occur in any vascularized tissue. These tumors generally do not cause clinical signs. Rarely, they may appear as soft, red masses anywhere in the body, although most are too small to be observed grossly. Diagnosis Diagnosis is made based on postmortem histology. Generally, these tumors are quite well organized and composed of well structured, but irregularly sized and tortuous capillaries lined by flattened to slightly plump endothelial cells. Most are intramuscular but some can occur within the vascular rete caudal to the eye. Because of their relatively well-organized appearance, an argument can be made that some of these represent a more benign hamartomatous lesion, such as angiomatosis or vascular hamartoma. However, they can be locally aggressive with severe tissue destruction despite the benign appearance of the endothelium, which argues against a benign, hamartomatous lesion. The distinction between benign and malignant lesions should be made based largely on an evaluation of adjacent tissue destruction first, and if present, features of malignancy.

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FIGURE 43.18 Thyroid tumors/hyperplasia. (A) Anatomic location of “normal” thyroid tissue (arrow). H&E 20. (B) Normal thyroid follicle lined by flattened epithelial cells and filled with colloid (arrow). H&E 200. (C) Thyroid adenoma. The mass, arising ventral to the gills and visible externally, is pedunculated. H&E 20. (D) Thyroid adenoma. Frequently, epithelial cells fail to form follicles, producing clusters of polygonal cells with minimal cellular atypia (arrowhead). H&E 200. (E) Thyroid carcinoma. The mass appears highly infiltrative and destructive to the surrounding tissue. (H&E 20). (F) Follicles are frequently lined by more than one layer of high cuboidal epithelium (arrows). Mitotic figures are frequent. (arrowheads). H&E 200.

Control and Treatment Couch described a high incidence of hemangioendothelial neoplasms in Rivulus marmoratus (Couch, 1995). He postulated a number of potential etiologies, including avian sarcoma virus associated with a chicken liver diet, latent sarcoma virus activation, general activation of oncogenes and downregulation of tumor suppressor genes, and carcinogen exposure. Hemangiomas are not common in laboratory zebrafish, and to

date, we have not identified cohorts with a high prevalence of this tumor that might indicate any of these etiologies (Fig. 43.17).

Thyroid Tumors Description The thyroid gland in zebrafish is unencapsulated, and generally, follicles of various sizes run along the ventral

IV. Diseases

References

aorta, from the bulbous arteriosus to the first gill arch (Wendl et al., 2002). The occurrence of thyroid follicles in ectopic locations is common in some species, but only occasionally recognized in zebrafish. Single cases of thyroid tumors can occur, but the diagnosis of these tumors typically involves multiple fish in a facility or tank in response to extrinsic factors, especially iodine deficient conditions. As these conditions generally instigate hyperplasia of thyroid tissue, cases of multiple tumors should be carefully evaluated, considering environmental conditions and genetic backgrounds of the affected stocks. Specific criteria for the microscopic distinction between hyperplasia and neoplasia are provided. Pathobiology and Clinical Signs Thyroid tumors generally present as reddened masses on the ventral mandible. Masses have also been observed extending dorsally to the snout area and at the vent. Macroscopic masses may be the only clinical sign observed. However, perturbations in thyroid hormone levels could result in other abnormalities. Thyroid hormone is involved in masculinization of larval zebrafish, and hypothyroid females can give rise to female-skewed progeny (Mukhi, Torres, & Patino, 2007; Sharma, Tang, Mayer, & Patino, 2016). Diagnosis Diagnosis of thyroid tumors is made by histological examination. Fournie et al. described a range of thyroid proliferative lesions, highlighting the importance of adhering to specific criteria and standardized nomenclaons, Zimmermann and Galetti concluded that iodine deficiency can act as a tumor promoter and is a risk factor for thyroid cancer in people (Zimmermann & Galetti, 2015). Therefore, cases of multiple thyroid lesions should be carefully evaluated using histologic criteria to determine whether true neoplastic changes are present. Genetic background of the affected stocks should also be considered, as the effects of iodine deficiency in a mutant or transgenic line that is prone to tumors may increase the likelihood of follicular cell neoplastic changes. The diagnosis of multiple thyroid proliferative lesions should be followed by an assessment of food and water parameters and correction of any nutrient imbalances, particularly iodine. Significant regression and recovery of lesions following these adjustments may be evidence for observed masses being a result of hyperplasia. Conversely, these findings could indicate a high hormone or nutrient responsive neoplasm in which removal of the inciting stimulus causes a reduction in the size of neoplastic cells to the point, where they can no longer be identified in 5 mm sections (Fig. 43.18).

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References Bailey, G. S., Williams, D. E., & Hendricks, J. D. (1996). Fish models for environmental carcinogenesis: The rainbow trout. Environmental Health Perspectives, 104, 5e21. Burns, A. R., Watral, V., Sichel, S., Spagnoli, S., Banse, A. V., Mittge, E., et al. (2018). Transmission of a common intestinal neoplasm in zebrafish by cohabitation. Journal of Fish Diseases, 41, 569e579. Ceol, C. J., Houvras, Y., White, R. M., & Zon, L. I. (2008). Melanoma biology and the promise of zebrafish. Zebrafish, 5(4), 247e255. Coffee, L. L., Casey, J. W., & Bowser, P. R. (2013). Pathology of tumors in fish associated with retroviruses: A review. Veterinary Pathology, 50(3), 390e403. Collymore, C., Watral, V., White, J. R., Colvin, M. E., Rasmussen, S., Tolwani, R. J., et al. (2014). Tolerance and efficacy of emamectin benzoate and ivermectin for the treatment of Pseudocapillaria tomentosa in laboratory zebrafish (Danio rerio). Zebrafish, 11(5), 490e497. Cooper, T. K., Murray, K. N., Spagnoli, S., & Spitsbergen, J. M. (2015). Primary intestinal and vertebral chordomas in laboratory zebrafish (Danio rerio). Veterinary Pathology, 52(2), 388e392. Cooper, T. K., & Spitsbergen, J. M. (2016). Valvular and mural endocardiosis in aging zebrafish (Danio rerio). Veterinary Pathology, 53(2), 504e509. Correa, P., & Blanca Piazuelo, M. B. (2011). Helicobacter pylori infection and gastric adenocarcinoma. US Gastroenterology and Hepatology Reviews, 7(1), 59e64. Couch, J. A. (1995). Invading and metastasizing cardiac hemangioendothelial neoplasms in a cohort of the fish rivulus-marmoratus e unusually high prevalence, histopathology, and possible etiologies. Cancer Research, 55(11), 2438e2447. Deville, J., & Lopez, E. (1970). Histophysiology of ultimobranchial body at some phases (in fresh water) of life cycle of salmon (Salm salar-L). Archives d’Anatomie Microscopique et de Morphologie Experimentale, 59(4), 393e402. Engeszer, R. E., Patterson, L. B., & Parichy, D. M. (2007). Zebrafish in the wild: A review of natural history and new notes from the field. Zebrafish, 4(1), 21e40. Fournie, J. W., Herman, R. L., Couch, J. A., & Howse, H. (1998). Tumors of heart and endothelial cells. In C. J. Dawe, J. C. Harshbarger, S. R. Wellings, & J. D. Strandberg (Eds.), Pathobiology of spontaneous and induced neoplasms in fishes: Comparative characterization, nomenclature and literature. New York: Academic Press. Fournie, J. W., Wolfe, M. J., Wolf, J. C., Courtney, L. A., Johnson, R. D., & Hawkins, W. E. (2005). Diagnostic criteria for proliferative thyroid lesions in bony fishes. Toxicologic Pathology, 33(5), 540e551. Grizzle, J. M., & Thiyagarajah, A. (1988). Diethylnitrosamine-induced hepatic neoplasms in the fish Ribulus oscellatus marmoratus. Diseases of Aquatic Organisms, 5, 39e50. Hayes, H. M., & Sass, B. (2012). Testicular tumors: Species and strain variations. In H. E. Kaiser (Ed.), Comparative aspects of tumor development (Vol. 5, pp. 106e119). Springer Science and Business Media. Kent, M. L., Bishop-Stewart, J. K., Matthews, J. L., & Spitsbergen, J. M. (2002). Pseudocapillaria tomentosa, a nematode pathogen of zebrafish (Danio rerio) kept in research colonies and associated neoplasms. Comparative Medicine, 52, 362e367. Langenau, D. M. (Ed.). (2016). Cancer and zebrafish: Mechanisms, techniques, and models. Springer. Langenau, D. M., Traver, D., Ferrando, A. A., Kutok, J. L., Aster, J. C., Kanki, J. P., et al. (2003). Myc-induced T cell leukemia in transgenic zebrafish. Science, 299(5608), 887e890. https://doi.org/10.1126/ science.1080280. Lombardini, E. D., Hard, G. C., & Harshbarger, J. C. (2014). Neoplasms of the urinary tract in fish. Veterinary Pathology, 51(5), 1000e1012. https://doi.org/10.1177/0300985813511122.

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Maley, D., Laird, A. S., Rinkwitz, S., & Becker, T. S. (2013). A simple and efficient protocol for the treatment of zebrafish colonies infected with parasitic nematodes. Zebrafish, 10(3), 447e450. https:// doi.org/10.1089/zeb.2013.0868. Masahito, P., Ishikawa, T., Okamoto, N., & Sugano, H. (1992). Nephroblastomas in the Japanese eel, anguilla-japonica temminck and schlegel. Cancer Research, 52(9), 2575e2579. Meierjohann, S., & Schartl, M. (2006). From mendelian to molecular genetics: The Xiphophorus melanoma model. Trends in Genetics, 22(12), 654e661. Mitchell, D. L., Paniker, L., & Douki, T. (2009 NoveDec). DNA damage repair and photoadaptation in a Xipophorus fish hybrid. Photochemistry Photobiology, 85(6), 1384e1390. Mukhi, S., Torres, L., & Patino, R. (2007). Effects of larval-juvenile treatment with perchlorate and co-treatment with thyroxine on zebrafish sex ratios. General and Comparative Endocrinology, 150(3), 486e494. https://doi.org/10.1016/j.ygcen.2006.11.013 powerpoint. Myers, M. S., Landal, J. T., Krahn, M. M., & McCain, B. B. (1991). Relationships between hepatic neoplasms and related lesions and exposure to toxic chemicals in marine fish from the U.S. West Coast. Envinonmental Health Prespectives, 90, 7e15. O’Hagan, B. J., & Raidal, S. R. (2006). Surgical removal of retrobulbar hemangioma in a goldfish (Carassius auratus). Veterinary Clinics of North America: Exotic Animal Practice, 9(3), 729e733. Paquette, C. E., Kent, M. L., Buchner, C., Tanguay, R. L., Guilleman, K., Mason, T. J., & Peterson, T. S. (2013 Jun 10). A retrospective study of the prevalence and classification of intestinal neoplasia in zebrafish (Danio rerio). Zebrafish, (2), 228e236. Paquette, C. E., Kent, M. L., Peterson, T. S., Wang, R., Dashwood, R. H., & Lo¨hr, C. V. (2015). Immunohistochemical characterization of intestinal neoplasia in zebrafish (Danio rerio) indicates epithelial origin. Diseases of Aquatic Organisms, 116, 191e197. Perner, B., Englert, C., & Bollig, F. (2007). The Wilms tumor genes wt1a and wt1b control different steps during formation of the zebrafish pronephros. Developmental Biology, 309(1), 87e96. Rasighaemi, P., Basheer, F., Liongue, C., & Ward, A. C. (2015). Zebrafish as a model for leukemia and other hematopoietic disorders. Journal of Hematology and Oncology, 8.

Sharma, P., Tang, S., Mayer, G. D., & Patino, R. (2016). Effects of thyroid endocrine manipulation on sex-related gene expression and population sex ratios in Zebrafish. General and Comparative Endocrinology, 235, 38e47. Sinnhuber, R.0., Hedricks, J. D., Walse, J. H., & Putnam, G. B. (1997). Neoplasms in rainbow trout, a sensitive animal model for environmental carcinogensis. Spagnoli, S., Xue, L., & Kent, M. L. (2015). The common neural parasite Pseudoloma neurophilia is associated with altered startle response habituation in adul zebrafish (Danio rerio): Implications for the zebrafish as a model organism. Behavioural Brain Research, 291, 351e360. Spitsbergen, J. M., Buhler, D. R., & Peterson, T. S. (2012). Neoplasia and neoplasm-associated lesions in laboratory colonies of zebrafish emphasizing key influences of diet and aquaculture system design. ILAR Journal, 53(2), 114e125. Spitsbergen, J. M., Tsai, H. W., Reddy, A., Miller, T., Arbogast, D., Hendricks, J. D., et al. (2000). Neoplasia in zebrafish (Danio rerio) treated with N-methyl-N’-nitro-N-nitrosoguanidine by three exposure routes at different developmental stages. Toxicologic Pathology, 28(5), 716e725. Stolin, S. L., Wang, Y., Mauldin, J., Schultz, L. E., Lincow, D. E., Brodskiy, P. A., et al. (2014). Molecular and cellular characterization of a zebrafish optic pathway tumor line implicates glia-derived progenitors in tumorigenesis. PLoS One, 9(12). Truyen, U., & Lochelt, M. (2006). Relevant oncogenic viruses in veterinary medicine: Original pathogens and animal models for human disease. In T. Dittmar, K. S. Zaenker, & A. Schmidt (Eds.), Infection and inflammation: Impacts on oncogenesis (Vol. 13, pp. 101e117). Wendl, T., Lun, K., Mione, M., Favor, J., Brand, M., Wilson, S. W., et al. (2002). pax2.1 is required for the development of thyroid follicles in zebrafish. Development, 129(15), 3751e3760. Zhuravleva, J., Paggetti, J., Martin, L., Hammann, A., Solary, E., Bastie, J. N., et al. (2008). MOZ/TIF2-induced acute myeloid leukaemia in transgenic fish. British Journal of Haematology, 143(3), 378e382. Zimmermann, M. B., & Galetti, V. (2015). Iodine intake as a risk factor for thyroid cancer: A comprehensive review of animal and human studies. Thyroid Research, 8, 8.

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44 Special Procedures for Zebrafish Diagnostics Michael L. Kent1, Katrina N. Murray2, Kay Fischer3, Christiana Lo¨hr3,4, Donna Mulrooney3, Justin L. Sanders3,4 Departments of Microbiology and Biomedical Sciences, Oregon State University, Corvallis, OR, United States of America; 2Zebrafish International Resource Center, University of Oregon, Eugene, OR, United States of America; 3 Oregon Veterinary Diagnostic Laboratory, Oregon State University, Corvallis, OR, United States of America; 4 Department Biomedical Sciences, Oregon State University, Corvallis, OR, United States of America 1

Environmental Samples Water quality-related diseases are one of the major causes of morbidity in zebrafish (See Hammer (2020), Chapter 30): Water Quality Chapter; Murray, Lains, & Spagnoli, 2020, Chapter 45). Inappropriate water chemistry parameters or the presence of certain toxicants in the water are the main causes of acute mortality. Therefore, all disease investigation should include documentation of the most common water quality parameters (i.e., temperature, pH, ammonia, nitrate, conductivity, and alkalinity). Depending on the system, data on chlorine, copper, etc. should be obtained. All of these data are usually available from the facility records, but if not, should be obtained by the diagnostician using the appropriate water quality testing kits or meters. Testing of pathogens in the environment is discussed below under “Nonlethal Testing.”

Clinical examination Fish should initially be evaluated while in their home environment. Behavior within the tank may be useful for the indication of an emerging disease, and some behavioral changes are useful for presumptive diagnoses. For example, if fish are flashing (rubbing on the sides of tanks), this may indicate that they are infected with external parasites. Other indications of the disease include cessation of feeding, lethargy, and abnormal position in the aquarium. Fish with impaired gills or other respiratory problems will often have rapid opercular movements and be gathered near the water surface,

The Zebrafish in Biomedical Research https://doi.org/10.1016/B978-0-12-812431-4.00044-0

whereas fish suffering supersaturation will also demonstrate respiratory distress, but will congregate at the bottom of the tank. Macroscopic changes are seldom pathognomonic with zebrafish diseases, but they do provide useful information. Erythema is a common presentation with sick zebrafish but can be caused by both water quality problems and infectious agents. Distinct ulcers on the flanks can be caused by both infectious agents, such as Mycobacterium marinum or by a noninfectious inflammatory process, such as egg-associated inflammation (Murray et al., 2020, Chapter 45). Emaciation is associated with most of the common infectious diseases in zebrafish; Pseudoloma neurophilia, Pseudocapillaria tomentosa (see Kent and Sanders, 2020, Chapter 43) or mycobacteriosis (Whipps & Kent, 2020, Chapter 44).

Selecting Fish for Diagnostic Evaluation Examinations should be conducted on diseased fish that are collected while still alive (moribund) as dead fish left in water autolyze extremely rapidly, hence many histological changes are obliterated by postmortem autolysis. However, dead fish may be suitable for some parasitological examinations and for observing obvious macroscopic pathological changes. Additional information can be obtained from textbooks that provide general guidelines for conducting health and disease investigations and necropsies on fish (e.g., Noga, 2010), and some that are specifically directed to zebrafish (Westerfield, 2000; Harper & Lawrence, 2011; Kent et al., 2009 [ZIRC online manual]).

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External Examination and Gross Necropsy Although their small size presents some necropsy challenges, general procedures used with larger fishes can be employed with some modifications. With patience and practice necropsies can be efficiently accomplished with the aid of a stereo dissecting microscope and small, fine dissecting instruments (particularly high-quality fine tip forceps). Fish may be euthanized by two approved methods, either rapid hypothermia (icing) or by an overdose of tricaine methanesulfonate (MS-222). Collymore (2020) provides a review of euthanasia using these methods. Note surface abnormalities (e.g., frayed fins, cloudy eyes, ulcers, skin discolorations, parasites, and tumors). Prepare wet mounts of the skin mucus and a few scales by scraping the surface of the fish with a coverslip and placing the coverslip on a glass slide. Some water may be added to the preparation, so that the area between the slide and the coverslip is completely filled with liquid. Examine the wet mount with a compound microscope, starting with low power, for detecting external parasites and bacteria, such as gliding bacteria Flavobacterium spp. Reducing the light and lowering the condenser will produce higher contrast, which will make the microscopic parasites and other pathogens more visible. Remove the operculum (Fig. 44.1). Note the color of the gills (pale gills usually indicate anemia). Check for parasites, cysts, excessive mucus, and hemorrhages with a dissecting microscope. Prepare a wet mount by removing the entire gill arch with attached filaments with scissors, then place the arch in a large drop of freshwater on a glass slide. The cartilaginous arch is too thick for high magnification examination, and hence, should be separated from the filaments with a scalpel blade and discarded. Then carefully overlay the preparation with a coverslip. Examine for

small parasites, fungi, and bacteria using a compound microscope. Open the visceral cavity as described in Fig. 44.1, revealing the visceral organs. The fish should be opened with the right side down for better exposure of the liver (Fig. 44.2). Note if ascites, hemorrhages, enlarged spleen, or other abnormalities are present. Multiple, whitish cysts in the visceral organs are suggestive of granulomas (e.g., Mycobacterium spp. infections). Expose the kidney by removing the swim bladder and note any kidney abnormalities. Many diseases cause enlargement or discoloration of the kidney, but this may be difficult to see in zebrafish due to their small size. Examine the heart for any abnormalities. Examine squash preparations of organs with a compound microscope to detect encysted parasites, fungi or microscopic granulomas. Squash preparations are made by removing a small piece of tissue, and gently squashing it between a slide and coverslip so that a thin preparation suitable for examination with a compound microscope or dissecting microscope is made. Leishman’s Giemsa or Diff-Quik (Dade Behring AG, Newark, DE) stained imprints of the kidney or other affected organs are useful for detection of protozoa and bacteria. Remove a piece of tissue, blot it on a clean paper towel to remove most of the blood, and lightly touch the cut surface of the tissue on a clean glass slide. Several imprints from the same piece of tissue can be made on one slide. Air-dry the preparation for approximately 1/2 h. Fix the slide for 5e10 min in absolute methanol for Giemsa stains or the fixative provided with the Diff-Quik kit. The slide can then be stained with Giemsa or Diff-Quik, or shipped to a diagnostic laboratory. For Mycobacterium spp., imprints and tissue smears are stained with acid-fast stains. Pseudoloma neurophilia, the most common parasite of zebrafish, can be detected by carefully removing the cranial cap (Fig. 44.1 e Step 5), removing the hindbrain

FIGURE 44.1 Dissection of a zebrafish. White lines designated demark location of incisions. (1) remove operculum. (2) Make an incision starting at dorsal posterior of opercular cavity and extend toward anus. (3) Connect incision with a cut starting at the ventral posterior of opercular cavity. (4) Cut dorsally to connect with Cut 2 and remove abdominal flank. (5) Remove cranial cap to access neural tissues for wet mounts. Black line ¼ incision prior to preservation to assure proper fixation of visceral organs.

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Histopathology

Spores occur in aggregates and appear more opaque than neural tissue (Fig. 44.3).

Histopathology

FIGURE 44.2 Internal anatomy of zebrafish. (A) Female (B) Male. (C) Swim bladder lifted to show kidney, which is dorsal to swim bladder. Images courtesy of Dr. Jennifer Matthews, Zebrafish Internal Resource Center. Modified by Robert Pianavilla.

and pieces of the anterior spinal cord, and preparing a wet mount. Avoid bone fragments, as these will prevent optimal thin preparations for microscopy.

Histopathology is often the primary or first-line diagnostic test for zebrafish (Collymore, Crim, & Lieggi, 2016; Murray, Varga, & Kent, 2016). A major strength of histopathology is that it facilitates the documentation of changes in a variety of tissues and organs with no a priori assumptions relating to a specific disease. It allows for examination of all tissues for infectious pathogens, changes related to suboptimal water quality, neoplasia, and numerous other noninfectious pathologies. In fact, the initial diagnosis and descriptions of many of the major pathogens affecting zebrafish in research facilities, including P. neurophilia (Matthews, Brown, Larison, Bishop-Stewart, Rogers, & Kent, 2001), M. chelonae (Whipps, Matthews, & Kent, 2008), Pleistophora hyphessobryconis (Sanders et al., 2010), Edwardsiella ictaluri (Hawke et al., 2013), and Myxidium streisingeri (Whipps, Murray, & Kent, 2015) were made by screening histological sections of moribund zebrafish submitted to diagnostic services, such as the Zebrafish International Resource Center Diagnostic Pathology Service (https://zebrafish.org/health/index.php). If possible, withhold feed to fish for 2e3 days before they are euthanized. This reduces postmortem autolysis of the intestine and liver and reduces the likelihood of ingesta from entering the gill cavity and causing postmortem changes. Also, hard food items in the intestine cause difficulties (e.g., chatter) when preparing sections. It is critical to preserve fish for histology as soon as possible after fish are dead to avoid postmortem changes. If possible, do not use dead fish because significant autolytic changes may occur in 15e20 min after death. Fish are preserved in formalin-based fixatives. Davidson’s or Dietrich’s solutions are preferred because these fixatives also contain acidic acid and ethanol, and thus

FIGURE 44.3

Wet mount preparation of neural tissue revealing Pseudoloma neurophilia, showing progressively higher magnifications (AeC). At lower magnification, aggregates of spores appear more opaque than surrounding tissues. Higher magnifications (B and C) (e.g., X 400 or 1000) demonstrates the typical oval microsporidian spore morphology with a posterior vacuole. (A), Bar ¼ 100 mm, (B and C). Bar ¼ 10 mm. IV. Diseases

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provide better penetration and fixation of all organs, good cellular detail and some decalcification of bone. Tissue preserved in formalin-based fixatives are generally considered less suitable for PCR from tissue blocks, but we found that there was no difference in success with mycobacteria PCR tests on zebrafish preserved in 10% formalin compared to Dietrich’s (Peterson, Kent, Ferguson, Watral, & Whipps, 2013). Most zebrafish submitted for histopathology evaluation are euthanized, fixed, and the entire fish submitted for histological processing. The visceral cavity of the fish is opened by cutting an incision starting at the posterior part of the opercular cavity and extending it to about 2/3 the length of the coelomic cavity. Then the whole fish is placed in fixative at approximately 1:20 (v/v) tissue to fixative. Injection of fixative into the oral cavity and coelom is another effective method to assure proper fixation (G. Sanders, University of Washington, pers. comm.). Fixative is injected into the coelomic cavity until it is extended but not ruptured. This volume will vary greatly with the size of the fish but on average ranges from less than 0.5 mL to up to w1 mL for zebrafish. A 3 mL syringe and a 23 or 21 gauge needle can be used. Multiple needles will be required if multiple fish are being processed at once. When utilizing the oral injection procedure, account for at least 0.5 mL because fish are injected after needle placement until fixative backs up into the mouth and out the down side gill operculum. The fish is then place in fixative as described above. Fixation should be adequate as long as the fish are then placed in enough fixative (i.e., 20:1, fixative to fish carcass). The entire fish can be processed (usually cut in sagittal sections) and representative organs can be visualized in multiple sections on one or two slides (Fig. 44.4). Preparation of high-quality histological slides from whole, small fish requires some additional steps and attention to detail compared to processing homogenous pieces of organs. Some general considerations are (1) Decalcification: whereas zebrafish are small, the skeleton of adult fish is ossified, and hence, often requires

decalcification for optimal section. Cal-Ex II (formic acid) (Fisher Chemical) is recommended as more aggressive solutions, such as those containing HCl may inhibit acid-fast staining of Mycobacterium spp. in sections (Kent et al. 2006). (2) Whole-body sagittal sections: most often, the preserved fish is bisected, and both halves processed “cut side down.” The spinal cord should be included in sections for detection of Pseudoloma neurophilia. Obtaining good sections of the spinal cord can be challenging. Double-edged razor blades are cut into two pieces with scissors and used for bisecting the fish. A scalpel blade or single-edge razor blade is unacceptable due to the thickness of the blades. Grasp the fish with forceps with the head facing to the left and place the blade on the other side of the fish slightly off of the spinal cord and make a clean cut through the fish. By firmly grasping the fish with the forceps, it may be possible to straighten out the fish with spinal deformities prior to bisecting. Fish lacking pigment (e.g., Caspers) can be particularly difficult, as it is nearly impossible to visualize the centerline. Cassettes of fish that are brittle are placed in a jar of dedicated Cal-ex II for about 4e8 h before processing. (3) Facing the block to obtain spinal cord in the section requires careful embedding, and it is important to press the fish to the bottom of the mold starting with the half of the fish that does not contain the spinal cord. Clean the embedded cassettes of any extra paraffin, especially the part of the cassette that touches the back of the microtome. A dissecting microscope with a light source angled from the side is set up near the microtome to enhance visualization. The cassettes are faced by taking a few turns at 10 mm, stopping, and examining at the cut surface under the microscope and resuming until the spinal cord comes into view and the coelomic organs are exposed. The spinal cord may appear before the organs. In this case, cut sections and then face deeper to obtain the coelomic organs. Use charged adhesion slides and dry the slides thoroughly before staining. Fish within the blocks may be dry and should soak on a cold tray with water for at least 30 min after facing and prior to sectioning. A few squirts of a 5% solution of Downey brand fabric softener can be added to the cold tray water to enhance the softening but should be judiciously used, as it can interfere with forming a sectioning ribbon.

Immunohistochemistry and Diagnostic Antibodies

FIGURE 44.4 Sagittal histological section of a male zebrafish showing major organs. K ¼ kidney, G ¼ Gills, B ¼ Brain, SC ¼ spinal cord, H ¼ heart, In ¼ intestine, G ¼ granulomas due to mycobacteriosis.

The whole-body sagittal section provides the ability to have the important organs on one slide, and therefore, provides an excellent format for positive and negative controls within the same section. Antibodies directed specifically to zebrafish antigens are becoming more

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available. For example, as of 2017, Genetex list some 450 antibodies that cross-react with zebrafish. Antibodies developed against epitopes in human and mouse proteins often have useful cross-reactivity with conserved proteins in zebrafish. Hence, tissue or cell type-specific antibodies can be used to characterize -neoplasms to elucidate tissue or cell of origin (Paquette et al., 2015). Nevertheless, antibodies not specifically developed against zebrafish epitopes must be carefully validated to avoid overinterpretation of findings (Ramos-Vara, 2005; Bordeaux et al., 2010). Some companies, such as AbCam (Eugene, Oregon) also offers trials where an antibody is purchased, evaluated, and when results are reported back to the company (e.g., results with zebrafish), the researcher is credited the cost of the antibody. This strategy allows for the zebrafish community to contribute to a growing list of mammalian antibodies with known cross-reactivity (do or do not work) in zebrafish. Quality of immunohistochemistry stains can be negatively impacted by improper fixation. Inappropriate fixation times can result in poor epitope preservation and antibody binding.

Bacteriology Bacteriology can be performed as part of the initial diagnostic evaluation, or when bacteria are observed via histopathology in conjunction with pathologic lesions and clinical morbidity or mortality is present in other fish within the population. Some bacterial infections, such as surface gliding bacteria and Mycobacterium spp., can be identified to the genus or family level using simple Gram or acid-fast stains (e.g., Ziehl Neelsen) or distinct histological presentations. However, a more precise diagnosis of other bacterial infections often requires isolation of the bacteria in culture. Only freshly euthanized fish should be used for bacteriological examinations as dead fish in the water are essentially worthless due to invasion of tissues by non-pathogenic bacteria. The optimal organs for isolating systemic bacteria are the kidney and spleen. Aerocystitis is commonly associated with bacterial

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infections, and hence, the culture of the swim bladder is often useful. The surface of the fish should be disinfected with 70% ethanol, and dissecting instruments should be sterilized. Two methods are used for obtaining inocula from the kidney or swim bladder, the dorsal approach, or through the coelomic cavity. Dorsal: Make an incision anterior to the dorsal fin, extending through the muscle and spinal cord, into the kidney or even more ventral into the swim bladder (Fig. 44.5A and B). Small inoculating loops are used to obtain samples, which are streaked on blood agar for Gram-negative bacteria, and specialized media for mycobacteria (See Whipps and Kent (2020), Chapter 44). Plates are then delivered to the appropriate diagnostic laboratory for further diagnosis. Placing inocula in liquid transport media is not recommended because it is difficult to avoid all extraneous, nonpathogenic bacteria. And these can quickly overgrow slower growing bacteria, such as Mycobacterium spp. and Edwardsiella ictaluri. Mycobacteria are often slow-growers and require special media. However, they have characteristics (i.e., thick, waxy cell wall) that provide selective advantages for culture and isolations. Tissues are often decontaminated overnight in 1% cetyl pyridinium chloride (Whipps, Dougan, & Kent, 2007). Cultures are usually grown on Middlebrook (MB) 7H10 and Lowensteine Jensen (LJ), and M. haemophilum requires 7H10 agar supplemented with OADC and 60 mM hemin (Whipps et al. 2007; Whipps & Kent, 2020, Chapter 44). For gliding bacteria (Cytophaga, Flexibacter and Flavobacterium spp.) are usually cultured from skin or gills, and Cytophaga Medium (Anacker & Ordal, 1959) is recommended. Although the lesions may exhibit massive numbers of gliding bacteria, other bacteria (e.g., Aeromonas spp.) may overgrow the former. Therefore, the best approach to obtain pure cultures of gliding bacteria is to homogenize the infected tissue (e.g., gills, skin, and muscle) in sterile water, and inoculate plates in serial log dilutions. Gram-stained preparations may reveal bacteria when they are numerous in infected tissue. Smear or imprint suspect tissues thinly on a glass slide, air dry and fix the slide by gently heating the slide over an open flame

FIGURE 44.5

Culture for bacteria from kidney, dorsal approach. (A) Arrow ¼ location of the incision to access the kidney. (B) Histological section showing the path of incision, through dorsal muscle, spinal cord (SC) and vertebrae to access the kidney (arrow). Bar ¼ 50 mm.

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for 3e5 s. Gram stain kits are available from scientific supply houses and include instructions for their use. Likewise, mycobacteria can be visualized in imprints stained with acid fast stains.

Bacteriology With Histopathology Bacterial culture and histopathology can be performed on the same fish. This is particularly useful because of the risk of bacterial contamination of internal structures, and histology provides useful information to discern if the bacteria actually represent an infection. False-negative culture results may occur with an inaccurate sampling of focal infections or with slow-growing and difficult to culture bacteria (such as Mycobacterium haemophilum). False-positive cultures may occur with skin or gut contamination of samples, which is common with small fish, and in mixed bacterial infections, where the easily cultured bacteria are not responsible for the observed fish morbidity and mortality. For Gram negative septicemia and aerocystis, we usually use the dorsal approach to perform aseptic cultures of the kidney and swim bladder (Fig. 44.5). For mycobacteria, we recommend removing a large portion of the spleen and part of the liver. These can then be examined by culture, acid-fast imprints or preserved in ethanol or frozen for PCR analysis. The remaining carcasses are fixed in Dietrich’s fixative and processed for histological evaluation. We have found that both of the methods to obtain samples for bacteriology do not significantly compromise histological evaluations on the same fish.

Virology Zebrafish are susceptible to numerous fish viruses, and even some human viruses. However, we are aware of two documented natural outbreaks of viral diseases in zebrafish (See Crim (2020), Chapter 42). As with bacterial diseases, isolation of viruses in culture may be required to diagnose a viral disease, and culture is best conducted on tissues collected from freshly killed fish. If this is not practical, the fish should be refrigerated for no longer than 24 h before examination. As a last resort, fish for virus examination can be frozen. The specimens are then transported to a qualified fish virology laboratory. As has been the case with other relatively new forms of fish culture, as fish health researchers thoroughly investigate outbreaks and incorporate tissue culture in their examinations, it is very likely that “new” viruses and viral diseases will be recognized in zebrafish.

Molecular Diagnositic Tests Traditional or qPCR tests have been developed for several zebrafish pathogens for examining fish tissues, which include P. neurophilia (Sanders & Kent, 2013; Whipps & Kent, 2006), Mycobacterium marinum, M. chelonae and M. haemophilum (Meritet, Mulrooney, Kent, & Loehr, 2017), and Edwardsiella ictaluri (Griffin et al., 2016). Crim, Lawrence, Livingston, Rakitin, Hurley, & Riley (2017) described PCR tests for Mycobacterium spp., Pseudoloma neurophilia, and Pseudocapillaria tomentosa, and evaluated them for testing fish tissues versus environmental samples (see Nonlethal Testing). There are countless species of Mycobacterium that occur in the aquatic environment, and many are nonpathogenic or only potential opportunists. Therefore, molecular tests require the use of well-designed specific primers utilizing a variety of genes. With zebrafish mycobacteria, these have most often been the hsp 70 gene (Meritet et al., 2017; Whipps et al., 2007). DNA fingerprint methods can also be employed to further characterize variations in strains within an outbreak (Ostland et al., 2008; Whipps et al., 2008). For bacteria, molecular identification can be performed directly on fish tissues (Whipps et al., 2007; 2008) or on isolated colonies from cultures. The former has certain advantages as culture, as stated above, may be misleading. For example, a population of wild mollies and swordtails suffered an epidemic of mycobacteriosis based on the presence of granulomas. Here M. chelonae was cultured from a few fish, but the actual etiologic agent was an M. triplex-like organism that was never cultured (Poort, Whipps, Watral, Font, & Kent, 2006). In recent years methods for retrival of DNA sequences from formalin preserved tissues, including those in paraffin blocks have significantly improved. Usually, it is more difficult to retrieve useful DNA from tissues that have been stored in formalin for extended periods of time. However, Peterson et al. (2013) were able to obtain diagnostic sequences of mycobacteria from about 50% of zebrafish in paraffin blocks in which tissues were stored in either Dietrich’s or 10% formalin for up to 45 days before processing. Alternatively, DNA can be extracted from cores made in tissue blocks, allowing for precise targeting of bacteria within lesions (Meritet et al., 2017) (Fig. 44.6). This approach is particularly useful with mycobacteria as nonpathogenic Mycobacterium spp. often colonize the intestine, and thus, may confused diagnoses with whole-body sections. Conversely, it may be more difficult to extract DNA from dense, solid cores that thin tissue sections. Although about half of the paraffin blocks samples do not yield mycobacterial sequences, most cases involve submitting multiple fish. And this approach was useful

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FIGURE 44.6 Coring blocks to target granulomas with acid fast bacteria for PCR analysis. (A) Granulomas with bacteria are matched to blocks. (B and C) are whole in blocks, with arrows demarking location of cores.

for conducting a retrospective study on the first occurrence of M. marinum in a large facility (Mason et al., 2016).

Nonlethal Testing Some mutants or transgenic lines are available in only very small numbers, and therefore euthanasia of enough fish to obtain sufficient confidence that negative results are correct for the presence of a particular pathogen is often not practical. However, the small, confined tanks and frequent spawning of zebrafish provide advantages for developing such tests based on water, biofilms, feces, gametes, or spawn water. This is consistent with the concept of screening for the presence of pathogens in mouse colonies using feces or bedding (Bauer et al., 2016; Livingston & Riley, 2003). Mycobacterial pathogens zebrafish occur both in the feces and water, and as biofilms on solid surfaces within tanks (Whipps et al., 2007; 2008; Mason et al., 2016). Accordingly, Crim et al. (2017) found that mycobacteria could readily be detected in feces, detritus, and water. Detritus and feces were useful for detecting P. tomentosa, which is consistent with the life cycle involving releasing of eggs in feces. Sugar centrifugation tests to detect parasite eggs in feces are fundamental in veterinary diagnostics, and can be employed to detect eggs of Pseudocapillaria tomentosa in feces collected from zebrafish tanks (Martins, Watral, RodriguesSoares, & Kent, 2017; Murray & Peterson, 2015). This is particularly useful, as the eggs of this nematode have the distinctive capillarid morphology with bipolar plugs, and hence can be easily differentiated from other nematode eggs and other organisms found in tank detritus.

Crim et al. (2017) found that environmental samples, including water, feces, and detritus, were not useful for detecting the microsporidium Pseudoloma neurophilia. In contrast, water collected following a spawning (e.g., from spawning tanks) is useful for PCR testing for P. neurophilia as the parasite is common in eggs and other material in the ovary (Sanders & Kent, 2011). This study found that very few eggs are positive by PCR, which agrees with Crim et al. (2017). PCR methods have recently been developed that have improved the detection of DNA from environmental samples (eDNA). One of these is droplet digital PCR (ddPCR) (Rothrock et al., 2013). We are using this format to detect zebrafish pathogens in water. This approach partitions the sample into hundreds of millions of water-in-oil droplets before thermal cycling; essentially running thousands of PCR reactions, and it thus greatly reduces inhibition with nontarget DNA and other environmental inhibitors. At this point, we have improved detection of P. neurophilia in tank water and can detect P. tomentosa DNA in water long before eggs are released in feces from adult female worms. Most diagnostic laboratories do not use the droplet digital system, but with its accuracy and sensitivity, it provides a very useful foundation for developing other PCR water tests. Blood can be used for nonlethal testing for pathogens from larger fish (Whipps et al., 2005). Although challenging due to their small size, blood can be nonlethally collected from blood using special techniques (Zang, Shimada, Nishimura, Tanaka, & Nishimura, 2015). Presently there are no reported blood tests for zebrafish pathogens.

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Treatment Details for treating specific infectious and noninfectious diseases are provided in other chapters (Kent & Sanders, 2020, Chapter 43; Whipps & Kent, 2020, Chapter 44; Murray et al., 2020, Chapter 45). Here we provide a general overview of common methods used for administering antibiotics and antiparasitic compounds. There are many texts that review treating infectious diseases in food and ornamental fishes (TrevesBrown, 2000; Noga, 2010). These approaches and methods can generally be applied to zebrafish, but laboratory zebrafish provide some unique challenges. First, concerns about nonprotocol induced variation should be considered when administering therapuetants to fish being used in experiments. Second, many therapeutants are administered by adding them directly to water, particularly for external parasites (Kent & Sanders, 2020, Chapter 43), and some of these may be detrimental to beneficial bacteria in biological filters (Pedersen, 2009). Last, some drugs are administered by injection, and this is often impractical for many zebrafish laboratories.

Treatments Delivered in Water Application of chemotherpeutants directly to water to control external parasites and bacterial infections is a common practice in aquaculture. Bath treatments, most often formalin, for about 1 hour or extended periods are effective for treating external parasites (See Kent & Sanders, 2020, Chapter 43). Formalin reduces dissolved oxygen levels, so it is important to maintain active aeration of the treated water. Sodium chloride baths are routinely used for reducing parasite burdens (Noga, 2010), and they offer an additional benefit in that fish with skin and gill infections often have reduced osmoregulatory capabilities, and hence adding salt at the appropriate level will reduce hypoosmotic stress. Emamectin benzoate, an analog of ivermectin, is a macrocyclic lactone. This class of drugs is well recognized for its efficacy for treating intestinal nematodes, including by external delivery. A commercial product of emamectin benzoate (Lice-Solve) is available for treating external copepods by adding the product to water. We recently showed that this product is very effective for treating Pseudocapillaria tomentosa infections using bath treatments (Kent, Watral, Gaulke, & Sharpton, 2019).

Oral Treatments Treatment of feed is now the most common approach in large aquaculture facilities, and more recently, public

aquariums are employing this approach. With aquaculture, large quantities of drugs are incorporated into the diet as a premix by feed manufactures, or the diet is top coated with drug mixtures after pellets are made (Burridge, Weiss, Cabeelos, Pizzarro, & Bostick, 2010; Noga, 2010). This approach has been used for treating P. tomentosa infection with emamectin benzoate and ivermectin in zebrafish (Collymore et al., 2014; Kent et al., 2019). Gelatin-based diets containing antibiotics are often used in public aquariums, and this approach has been used to deliver antibiotics for treating mycobacteriosis in zebrafish (Chang, Doerr, & Whipps, 2017).

References Anacker, R. L., & Ordal, E. J. (1959). Studies on the myxobacterium Chondrococcus columnaris. I Serological typing. Journal of Bacteriology, 78, 25e32. Bauer, B. A., Besch-Williford, C., Livingston, R. S., Crim, M. J., Riley, L. K., & Myles, M. H. (2016). Evaluation of rack design and disease prevalence on detection of rodent pathogens in exhaust debris samples from individually ventilated caging systems. Journal of the American Association for Laboratory Animal Science, 55, 782e788. Bordeaux, J., Welsh, A. W., Agarwal, S., Killiam, E., Baquero, M. T., Hanna, J. A., et al. (2010). Antibody validation. Biotechniques, 48, 197e209. Burridge, L., Weiss, J. S., Cabeelos, F., Pizzarro, J., & Bostick, K. (2010). Chemical use in salmon aquaculture: A review of current practices and possible environmental effects. Aquaculture, 306, 7e23. Chang, C. T., Doerr, K. M., & Whipps, C. M. (2017). Antibiotic treatment of zebrafish mycobacteriosis: Tolerance and efficacy of treatments with tigecycline and clarithromycin. Journal of Fish Diseases, 40, 1473e1485. Collymore, C. (2020). Analgesia, anesthesia, and anesthisia. Chapter 39. In S. C. Cartner, J. S. Eisen, S. C. Farmer, K. J. Guillemen, M. L. Kent, & G. F. Sanders (Eds.), The zebrafish in biomedical research: Biology, husbandry, diseases, and research application. Elsevier. Collymore, C., Crim, M. J., & Lieggi, C. (2016). Recommendations for health monitoring and reporting for zebrafish research facilities zebrafish. Zerbrafish, 13(S1), S138eS148. Collymore, C., Watral, V., White, J. R., Colvin, M. E., Rasmussen, S., Tolwani, R. J., et al. (2014). Tolerance and efficacy of emamectin benzoate and ivermectin for the treatment of Pseudocapillaria tomentosa in laboratory zebrafish (Danio rerio). Zebrafish, 11, 490e497. Crim, M. J., Lawrence, C., Livingston, R. S., Rakitin, A., Hurley, S. J., & Riley, L. K. (2017). Comparison of antemortem and environmental samples for zebrafish health monitoring and quarantine. Journal of the American Association for Laboratory Animal Science, 56, 412e424. Griffin, M. J., Reichley, S. R., Greenway, T. E., Quiniou, S. M., Ware, C., Gao, D. X., et al. (2016). Comparison of Edwardsiella ictaluri isolates from different hosts and geographic origins. Journal of Fish Diseases, 39, 947e969. Crim, M. J. (2020). Viral Diseases e Chapter 42. Hammer, H. S. (2020). Water quality. Chapter 30. In S. C. Cartner, J. S. Eisen, S. C. Farmer, K. J. Guillemen, M. L. Kent, & G. F. Sanders (Eds.), The zebrafish in biomedical research: Biology, husbandry, diseases, and research application. Elsevier. Harper, C., & Lawrence, C. (2011). Boca Raton: The Laboratory Zebrafish CRC Press. Hawke, J. P., Kent, M. L., Rogge, M., Baumgartner, W., Wiles, J., Shelly, J., et al. (2013). Edwardsiellosis caused by Edwardsiella

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ictaluri in laboratory populations of zebrafish Danio rerio. Journal of Aquatic Animal Health, 25, 171e183. Kent, M. L., Feist, S. W., Harper, C., Hoogstraten-Miller, S., Law, J. M., Sa´nchez-Morgado, J. M., et al. (2009). Recommendations for control of pathogens and infectious diseases in fish research facilities. Comparative Biochemistry and Physiology ePart C (Toxicology and Pharmacology), 149, 240e248. Kent, M. L., Heidel, J. R., Watral, V. G., Bishop-Stewart, J. K., Matthews, J. L., & Ostland, V. (2006). Some decalcification procedures inhibit acid fast staining of Mycobacterium spp. in tissue sections. American Fisheries Society/Fish Health Section. Newsletter, 34(1), 15e17. Kent, M. L., & Sanders, J. L. (2020). Parasites of zebrafish. In The zebrafish in biomedical research: Biology, husbandry, diseases, and research applications. Editors THIS BOOK. Kent, M. L., Watral, V., Gaulke, C. A., & Sharpton, T. J. (2019). Further evaluation of the efficacy of emamectin for treating Pseudocapillaria tomentosa (Dujardin 1843) in zebrafish Danio rerio (Hamilton 1822). Journal of Fish Diseases (in press). Livingston, R. S., & Riley, L. K. (2003). Diagnostic testing of mouse and rat colonies for infectious agents. Lab Animal, 32, 44e51. Martins, M. L., Watral, V., Rodrigues-Soares, J. P., & Kent, M. L. (2017). A method for collecting eggs of Pseudocapillaria tomentosa (Capillariidae) from zebrafish Danio rerio and efficacy of heat and chlorine for killing the nematode’s eggs. Journal of Fish Diseases, 40, 169e182. Mason, Snell, K., Mittge, E., Melancon, E., Montgomery, R., McFadden, M., et al. (2016). Strategies to mitigate a Mycobacterium marinum outbreak in a zebrafish research facility. Zebrafish, 13(S1). S-77-S-87. Matthews, J. L., Brown, A. M. V., Larison, K., Bishop-Stewart, J. K., Rogers, P., & Kent, M. L. (2001). Pseudoloma neurophilia n.g., n.sp., a new genus and species of Microsporidia from the central nervous system of the zebrafish (Danio rerio). The Journal of Eukaryotic Microbiology, 48, 229e235. Meritet, D. M., Mulrooney, D. M., Kent, M. L., & Loehr, C. V. (2017). Development of a development of quantitative real-time PCR assays for postmortem detection of Mycobacterium spp. common in zebrafish (Danio rerio) research colonies. Journal of the American Association for Laboratory Animal Science, 56, 1e11. Murray, K., Lains, D., & Spagnoli, S. (2020). Water quality and idiopathic diseases. Chapter 45. In S. C. Cartner, J. S. Eisen, S. C. Farmer, K. J. Guillemen, M. L. Kent, & G. F. Sanders (Eds.), The Zebrafish in Biomedical Research: Biology, Husbandry, Diseases, and Research Application. Elsevier. Murray, K. N., & Peterson, T. S. (2015). Pathology in practice. P. tomentosa infection in zebrafish. Journal of the American Veterinary Medical Association, 246, 201e203. Murray, K. N., Varga, Z. M., & Kent, M. L. (2016). Biosecurity and health monitoring at the zebrafish international resource center. Zebrafish, 13(S1), S30eS38. Noga, E. J. (2010). Fish disease: Diagnosis and treatment (2nd ed.). St. Louis, MO: Mosby-Year Book, Inc., 367 pp. Ostland, V. E., Watral, V., Whipps, C. M., Austin, F. W., St-Hilaire, S., Westerman, M. E., et al. (2008). Biochemical, molecular, and virulence characteristics of select Mycobacterium marinum isolates in hybrid striped bass (Morone chrysops x Morone saxatilis) and zebrafish (Danio rerio). Diseases of Aquatic Organisms, 79, 107e118. Paquette, C. E., Kent, M. L., Peterson, T. S., Wang, R., Dashwood, R. H., & Lo¨hr, C. V. (2015). Immunohistochemical characterization of intestinal neoplasia in zebrafish (Danio rerio) indicates epithelial origin. Diseases of Aquatic Organisms, 116, 191e197.

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Pedersen, L. F. (2009). Fate of water borne therapeutic agents and associated effectson nitrifying biofilters in recirculating aquaculture systems. Aalborg Universitet: Institut for Kemi, Miljø og Bioteknologi, Aalborg Universitet. Peterson, T. S., Kent, M. L., Ferguson, J. A., Watral, V. G., & Whipps, C. M. (2013). Comparison of fixatives and fixation time on PCR detection of Mycobacterium in zebrafish Danio rerio. Diseases of Aquatic Organisms, 104, 113e120. Poort, M. J., Whipps, C. M., Watral, V. G., Font, W. F., & Kent, M. L. (2006). Molecular characterization of a Mycobacterium species in non-native poeciliids in Hawaii using DNA sequences. Journal of Fish Diseases, 29, 181e185. Ramos-Vara, J. A. (2005). Technical aspects of immunohistochemistry. Veterinary Pathology, 42, 405e426. Rothrock, M. J., Hiett, K. L., Kiepper, B. H., Ingram, K., & Hinton, A. (2013). Quantification of zoonotic bacterial pathogens within commercial poultry processing water samples using droplet digital PCR. Advances in Microbiology, 3, 403e411. Sanders, J. L., & Kent, M. L. (2011). Development of a sensitive assay for the detection of Pseudoloma neurophilia in laboratory populations of the zebrafish Danio rerio. Diseases of Aquatic Organisms, 96, 145e156. Sanders, J. L., & Kent, M. L. (2013). Verification of intraovum transmission of a microsporidium of vertebrates: Pseudoloma neurophilia infecting the zebrafish, Danio rerio. PLoS One, 8(9), e76064. https://doi.org/10.1371/journal.pone.0076064. PMC3781086. Sanders, J. L., Lawrence, C., Nichols, D. K., Brubaker, J. F., Peterson, T. S., Murray, K. N., & Kent, M. L. (2010). Pleistophora hyphessobryconis (Microsporidia) infecting zebrafish (Danio rerio) in research facilities. Diseases of Aquatic Organisms, 91, 47e56. Treves-Brown, K. M. (2000). Vol 3. Methods of drug administration. In K. M. Treves-Brown (Ed.), Applied fish pharmacology springer (pp. 1e15). Westerfield, M. (2000). The zebrafish book. A guide for the Laboratory.Use of zebrafish (Danio rerio) (4th ed.). Eugene: University of Oregon Press, 2000. Whipps, C. M., Burton, T., Watral, V. G., St-Hilaire, S., & Kent, M. L. (2005). Assessing the accuracy of a polymerase chain reaction test for Ichthyophonus hoferi in Yukon River Chinook salmon (Oncorhynchus tshawytscha). Diseases of Aquatic Organisms, 8, 141e147. Whipps, C. M., Dougan, S. T., & Kent, M. L. (2007). Mycobacterium haemophilum infections of zebrafish (Danio rerio) in research facilities. FEMS Microbiology Letters, 270, 21e26. Whipps, C. M., & Kent, M. L. (2006). Polymerase chain reaction detection of Pseudoloma neurophilia, a common microsporidian of zebrafish (Danio rerio) reared in research laboratories. Contemporary Topics in Laboratory Animal Science, 45, 13e16. Whipps, C. M., & Kent, M. L. (2020). Bacterial and fungal diseases of zebrafish. Chapter 44. In S. C. Cartner, J. S. Eisen, S. C. Farmer, K. J. Guillemen, M. L. Kent, & G. F. Sanders (Eds.), The zebrafish in biomedical research: Biology, husbandry, diseases, and research application. Elsevier. Whipps, C. M., Murray, K. N., & Kent, M. L. (2015). Occurrence of a myxozoan parasite Myxidium streisingeri n. sp. in laboratory zebrafish Danio rerio. The Journal of Parasitology, 10, 86e90. Whipps, C. M., Matthews, J. L., & Kent, M. L. (2008). Distribution and genetic characterization of Mycobacterium chelonae in laboratory zebrafish (Danio rerio). Diseases of Aquatic Organisms, 82, 45e54. Zang, L., Shimada, Y., Nishimura, Y., Tanaka, T., & Nishimura, N. (2015). Repeated blood collection for blood tests in adult zebrafish. Journal of Visualized Experiments, (102), e53272. https:// doi.org/10.3791/53272. See comment in PubMed Commons below.

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Further reading Barton, C. L., Johnson, E. W., & Tanguay, R. L. (2016). Faciility design and health management program at the Sinnhuber aquatic research laboratory. Zebrafish, 13(Suppl. 1), 39e43. Borges, A. C., Pereira, N., Franco, L. V., Pereira, M., Cunha, M. V., Amaro, A., et al. (2016). Implementation of a zebrafish health program in a research facility: A 4-year retrospective study. Zebrafish, 13(S1). S-115-S-126. Chang, C. T., Amack, J. D., & Whipps, C. M. (June 2016). Zebrafish embryo disinfection with povidoneeiodine: Evaluating an alternative to chlorine bleach. Zebrafish, 13(S1). S-96-S-101. Chang, C. T., Colicino, E. G., DiPaola, E. J., Al-Hasnawi, H.,J., & Whipps, C. M. (2015). Evaluating the effectiveness of common disinfectants at preventing the propagation of Mycobacterium spp. isolated from zebrafish. Comparative Biochemistry and Physiology ePart C (Toxicology and Pharmacology), 178, 45e50. Ferguson, J., Watral, V., Schwindt, A., & Kent, M. L. (2007). Spores of two fish Microsporidia (Pseudoloma neurophilia and Glugea anomola) are highly resistant to chlorine. Diseases of Aquatic Organisms, 76, 205e214. Hobbs, M. R., Shankaran, S. S., & James, W. L. (2016). Controlling endemic pathogensdchallenges and opportunities zebrafish. Zebrafish, 13(S1). S-66-71. Kent, M. L., Buchner, C., & Tanguay, R. L. (2014). Toxicity of chlorine to zebrafish embryos. Diseases of Aquatic Organisms, 107, 235e240. Kent, M. L., Buchner, C., Watral, V. G., Sanders, J. L., LaDu, J., Peterson, T. S., et al. (2011). Development and maintenance of a specific pathogen free (SPF) zebrafish research facility for Pseudoloma neurophilia. Diseases of Aquatic Organisms, 95, 73e79. Kent, M. L., Harper, C., & Wolf, J. C. (2012). Documented and potential research impacts of subclinical diseases in zebrafish. ILAR Journal, 53, 126e134. Kent, M. L., & Kieser, D. (2003). Avoidance of introduction of exotic pathogens with Atlantic salmon reared in British Columbia. In C.S. Lee, & P. J. O’Bryen (Eds.), Biosecurity in aquaculture production

systems: Exclusion of pathogens and other undesirables. World aqualt. Soc., Baton Rouge, Louisiana (pp. 43e50). Lawrence, C., Ennis, D. G., Harper, C., Kent, M. L., Murray, K., & Sanders, G. E. (2012). The challenges of implementing pathogen control strategies for fishes used in biomedical research. Comparative Biochemistry and Physiology ePart C (Toxicology and Pharmacolony), 155, 160e166. Legendre, L., Guillet, B., Leguay, E., Meunier, E., Labrut, S., Keck, N., et al. (2016). A network for monitoring health and husbandry practices in aquatic research facilities. Zebrafish, 13(S1). S-56-S-65. Martins, S., Monteiro, J. F., Vito, M., Weintraub, D., Almeida, J., & Certal., A. C. (2016). Toward an integrated zebrafish health management program supporting cancer and neuroscience research. Zebrafish, 13(S1), S47eS55. Murray, K. N., Bauer, J., Tallen, A., Matthews, J. L., Westerfield, M., & Varga, Z. (2011). Characterization and management of asymptomatic Mycobacterium infections at the zebrafish international resource center. Comparative Medicine, 50, 675e679. Murray, K. N., Dreska, M., Nasiadka, A., Rinne, M., Matthews, J. L., Carmichael, C., et al. (2011). Transmission, diagnosis, and recommendations for control of Pseudoloma neurophilia infections in laboratory zebrafish (Danio rerio) facilities. Comparative Medicine, 61, 322e329. Paquette, C. E., Kent, M. L., Murray, M., Guillemin, K., Mason, T. J., Buchner, C., et al. (2013). Intestinal neoplasia in zebrafish. Zebrafish, 10, 228e236. Paquette, C. E., Kent, M. L., Peterson, T. S., Wang, R., Dashwood, R. H., & Lo¨hr, C. V. (2015). Immunohistochemical characterization of intestinal neoplasia in zebrafish (Danio rerio) indicates epithelial origin. Diseases of Aquatic Organisms, 116, 191e197. Peterson, T. S., Spitsbergen, J. M., Feist, S. W., & Kent, M. L. (2011). The Luna stain, an improved selective stain for detection of microsporidian spores in histologic sections. Diseases of Aquatic Organisms, 95, 175e180.

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45 Zebrafish as a Model to Understand Vertebrate Development Narendra H. Pathak, Michael J.F. Barresi Smith College, Northampton, MA, United States of America

Introduction to Embryology Developmental Stages and Conservation Chordates, more colloquially referred to as vertebrates, cleave the zygote into thousands of cells with each daughter cell half the size of its parent. These cleavages continue until cells start to move, grow, and specialize as different types of cells during gastrulation. A seemingly chaotic amount of tissue morphogenesis occurs over the course of gastrulation that precisely positions the three earliest germ tissue types: ectoderm, endoderm, and mesoderm. Ectodermal cells give rise to the nervous system located toward the back or dorsal-most region of the embryo, as well as contribute to the epidermis that covers the entire embryo. A primary outcome of these gastrulation movements is the proper positioning of both the endodermal and mesodermal derivatives within the embryo along all the basic axes (See also reviews on early development Hasley, Chavez, Danilchik, Wu¨hr, & Pelegri, 2017; O’Farrell, 2015; Stower & Bertocchini, 2017). Importantly, the unifying feature of chordates is the notochord, which is formed at the midline of the gastrula during the initiation of mesoderm development (Annona, Holland, & D’Aniello, 2015). The central nervous system, which is the first major organ system to develop following gastrulation, forms from the creation of a rudimentary tube sitting atop the notochord. This neural tube ultimately creates the brain and spinal cord along the full length of the anterior to the posterior axis (O’Connell, 2013). Soon after the onset of neural tube formation the entire vertebrate trunk elongates, portions of which become sequentially segmented into similarly sized blocks of cells known as

The Zebrafish in Biomedical Research https://doi.org/10.1016/B978-0-12-812431-4.00045-2

somites (Be´naze´raf & Pourquie´, 2013; Hubaud & Pourquie´, 2014; Maroto, Bone, & Dale, 2012; Rashid et al., 2014; Resende, Andrade, & Palmeirim, 2014). Differentiation of somites will contribute to the development of skeletal muscle, portions of the circulatory system, and most notably the bones and cartilage of the vertebrae. The embryonic period culminates with organogenesis, which marks the time when an array of different organ types likes the heart, gut, kidney, liver, pancreas, and others are established throughout the embryo (Miquerol & Kelly, 2013; Zorn & Wells, 2009). The last moments of embryogenesis are known as the “pharyngula” stage, a stage when all vertebrates physically look the most similar (Collazo, 2000). Generally speaking, each embryonic part from the regions of the brain and eyes to the heart and segmented trunk are all similarly located and structurally resemble each other to such a degree that the conservation of their development is visibly undeniable. It is this biological conservation that enables researchers to use different vertebrate species as proxies or “model systems” to gain valuable insight into how humans may even develop (Dietrich, Ankeny, & Chen, 2014).

Genes and Development Cleavage, gastrulation, neurulation, somitogenesis, and organogenesis are the progressive stages of embryonic development that are made possible through the precisely timed regulation of gene expression. With the sole exception of the gametes (sex cells), all the zygotederived cells that make up an individual organism possess identical genomes (for instance, the DNA sequences in one’s gut epithelial cell is identical to the

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DNA sequences found within the photoreceptor cell of their eye). This fact presents a profound realization of one of the key mechanisms of embryonic development, that of differential gene expression. Cell differentiation is based on the different ways cells use the genome to turn on (express) or off (repress) specific genes. The resulting composition of genes expressed will lead to the repertoire of proteins for a given cell that will provide its unique structural and functional properties. Therefore, embryonic development is foundationally governed by controlling which genes are expressed at different times and places. Most recently, zebrafish researchers have advanced the use of single-cell RNA sequencing to characterize all the transcripts (mRNAs) present in all the cells of embryos at different embryonic stages (reviewed in Harland, 2018). Analyses of this enormous amount of data with nearest-neighbor computational approaches and cross-referencing with known gene functions has enabled the visualization of “developmental trees”d3-Dimensional representations of the transcriptomic identity of each cell as it matures over the course of embryogenesis (Farrell et al. 2018; Wagner et al. 2018). See “movie S1” from Farrell et al., 2018 to see the trajectories of cell specification over early embryogenesis. Due to the importance that a cell’s transcriptome plays in defining its identity, the field of developmental biology has been ever captivated with determining how cells acquire these patterns of differential gene expression. Over the course of the blastula and gastrula stages, cells appear visibly similar and will, however, gradually become spatially distinct across the embryo. The environment cells experience in different regions of the embryo can profoundly influence the genes those cells ultimately express and consequently, the cell fates they adopt. Much research in developmental biology has been devoted to identifying and characterizing the myriad of regionally restricted signals present throughout embryogenesis and how such signals influence gene expression, cell differentiation, morphogenesis, and other developmental outcomes (Perrimon, Pitsouli, & Shilo, 2012; Wilcockson, Sutcliffe, & Ashe, 2017). Several families of signaling proteins have been found to be essential for embryonic development, these include, but are not limited to, the superfamily of Transforming Growth Factors (TGFs, like bone morphogenic proteins or BMPs), and families of Wnts, Fibroblast Growth Factors (Fgf), and Hedgehogs (like Sonic hedgehog, Shh). Secretion of these proteins into the extracellular environment or their active transport can be received and interpreted by receptors in a cell’s outer (plasma) membrane, which triggers progressive changes inside the cell resulting in a diversity of responses like alterations in gene expression, remodeling of the cytoskeleton to influence cell shape and movement, or

promotion of cell division. Much attention has been placed on deciphering the array of downstream transcription factor proteins that these signaling systems effect, efforts that are providing insights into the whole gene regulatory networks responsible for specific tissue type development (Ettensohn, 2013; Ferg et al., 2014; Nowick & Stubbs, 2010; Peter, 2017; Peter and Davidson, 2009, 2017). Use of the zebrafish model system has greatly advanced our understanding of the genetic, molecular, and cellular mechanisms governing each stage of vertebrate embryogenesis.

The Emergence of Zebrafish as a Versatile Vertebrate Model System to Understand Embryology From the fruit fly to the mouse, different model systems have provided unique advantages to dissecting the developmental mechanisms of embryogenesis. Among the many diverse advantages the zebrafish model system has to offer, it was initially adopted for research primarily because of its strong applicability to the study of embryonic development (Kimmel, Ballard, Kimmel, Ullmann, & Schilling, 1995). For practical reasons the small size of the adult zebrafish enables many thousands of fish to be housed in a relatively small laboratory space making the zebrafish significantly more cost-effective than other vertebrate model systems. Most importantly, a single female and male zebrafish can produce hundreds of embryos a week. Being able to house thousands of adult fish and pairs of which producing hundreds of embryos all together makes genetic analyses feasible. At this time, it is clear that the numerous forward genetic screens conducted on zebrafish have yielded the field’s greatest contributions to our understanding of embryogenesis. Moreover, the last 2 decades of zebrafish research has also amassed an ever-growing battery of powerful genetic and molecular approaches to both see and manipulate specific genes, cells, tissues, and whole physiological systems. Namely the creation of robust transgenic tools for cell-type specific reporter fish lines to visualize phenotypes even in live embryos, and for gene inducible techniques to answer both gain- or loss-of-function questions (as exemplified and reviewed in Halpern et al., 2008; Higashijima, Hotta, & Okamoto, 2000; Moro et al., 2013; Weber & Ko¨ster, 2013). Lastly, the ever-evolving approaches to capture and interrogate the zebrafish genome are providing novel insights from the specification of cell lineages to the comparative evolution of fish and humans (Gehrke et al., 2015; Harland, 2018; McCluskey & Postlethwait, 2015; McKenna et al., 2016; Pan et al., 2013; Pasquier et al., 2017).

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Zebrafish Research Advances Our Understanding of the Mechanisms Governing Development

Critical to making any gene or embryological manipulation effective is the ability to see or assess changes in embryonic morphology and progression. The accessibility of the zebrafish’s external development, paired with the embryo’s near transparency permits easy and direct microscopic visualization of each major embryonic structure. Additionally, this unique visual accessibility, combined with its rapid embryogenesis over a 24-hour period, allows researchers to quite simply watch development over time (Karlstrom & Kane, 1996). Today this advantage is being exploited to even higher degrees of resolution with the advent of more sophisticated live embryo imaging techniques, like laserscanning confocal, two-photon, and light-sheet microscopy (Keller, 2013; Keller, Schmidt, Wittbrodt, & Stelzer, 2008; Royer et al., 2016). Researchers have also taken advantage of the visible accessibility of zebrafish by perfecting the use of physical manipulatives, such as, but not limited to, the bathing of embryos with specific compounds, chemicals, and other environmental alterations, the microinjection of dyes, DNA and RNA, or even the transplantation of cells from one embryo to another. Thus, the zebrafish model system offers many advantages that have dramatically helped to reveal the developmental mechanisms that are driving the vertebrate embryogenesis.

Zebrafish Research Advances Our Understanding of the Mechanisms Governing Development Early Development In a “Fish-Shell” Cleavages and Maternal Contributions: Externally developing vertebrates like zebrafish have provided a remarkable advantage to studying the mechanisms of early development from conception through gastrulation (See Fig. 45.1). Whether glaring though the outer protective chorion shell or experimentally observing a dechorionated embryo, simply being able to watch each cell division during blastula development has demonstrated some of the defining features of embryonic cleavages in vertebrates. Zebrafish cleavage is incomplete or meroblastic such that a blastoderm (cells of the embryo proper) is built with each cell division in the animal pole, while a single yolk cell is retained in the vegetal hemisphere and an outermost enveloping layer (EVL) develops over the embryo. These animal pole restricted cleavages occur as near synchronous waves of mitoses that pass across the entire blastoderm, each time resulting in an equal halving of the cell size. As the amount of cytoplasm to DNA approaches more equal proportions, cell movement becomes visible which marks the moment when a blastula begins gastrulation (Kane, Warga, & Kimmel, 1992; Kane & Kimmel, 1993).

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Prior to conception critical preparatory events occur during oogenesis, namely the storing of maternal mRNA (transcripts) and protein contributions in the oocyte. These maternal contributions are themselves sufficient to drive all the mechanics of cell division during cleavage stages. For instance, futile cycle mutants fail to undergo pronuclear fusion leading to only 2 cells of the blastula actually possessing DNA containing nuclei, and despite this lack of DNA all futile cycle cells still undergo normal cleavages (Dekens, Pelegri, Maischein, & Nu¨sslein-Volhard, 2003). However, maternal factors alone are not sufficient for continued development past the blastula stage, and active transcription from the “zygotic genome” is required (Lee et al., 2013). This moment of shifting from maternal to zygotic control is known as the “midblastula transition” or “maternal-zygotic transition” (MZT) (Lee, Bonneau, & Giraldez, 2014; Miccoli, Dalla Valle, & Carnevali, 2017; Wragg & Mu¨ller, 2016). Investigations into the zebrafish MZT have provided important insights into the role that microRNAs (miRNAs) play in degrading the maternal RNA stores to facilitate the rapid transition of developmental control to the zygotic genome (Lee et al., 2014). miRNAs do not code for a protein but rather can function in the RNA-induced silencing complex (RISC) as a sequencespecific guide to target the complex’s ability to degrade RNA transcripts. More specifically, miR-430 has been demonstrated to be essential for the MZT as it is responsible for both translational repression and the targeted decay of hundreds of maternally provided transcripts (Bazzini, Lee, & Giraldez, 2012; Giraldez et al., 2006). As these maternal mRNAs diminish over the course of cleavages, the expression of zygotic mRNAs ramps up to regulate the rest of embryogenesis. Gastrulation and Axis Determination: The zebrafish blastula culminates into a perfect sphere with nearly all of the animal pole composed of embryonic cells and the vegetal pole occupied by the yolk (See Fig. 45.1). Gastrulation is first visibly evident by the asymmetric collection of cells at the margin, which resembles a small bulge at the surface equidistant between the poles - as if this very early embryo were holding a shield. As a result of this physical feature, the start of zebrafish gastrulation is commonly referred to as the “shield” stage. Important gastrulating cell behaviors occur prior to the visible shield, namely the movements of radial intercalation that drive epiboly. Deeper blastula cells of the epiblast move radially outward to join more superficially positioned layers of the epiblast. Such cell intercalation causes the entire population of blastula cells to flatten and consequently spread outward toward the vegetal pole. This is the gastrulation movement called epiboly, and it will assist in powering the complete coverage of the yolk with

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FIGURE 45.1 An Illustrative overview of zebrafish embryonic development. This developmental series progresses left to right and from top to bottom. The first four illustrations are representative of the cleavage period during which cells get smaller upon every division. Primordial germplasm is evident at the two-cell stage along the division plane (pink), while four discrete primordial germ cells (PGC) can be seen at the 64 cell stage that establishs four multicell clusters by the sphere stage (two shown). At the sphere stage, the enveloping layer covers the animal hemisphere stretching down to the yolk syncytial layer (YSL) where yolk syncytial nuclei and microtubules extend toward the vegetal pole. Next, gastrulation commences, as many mechanisms like contraction along these microtubules draws down the embryonic cells from 50% epiboly to 80% and finally full closure of the blastopore at the tailbud stage. The first indication of dorsal and ventral axes is visible at the shield stage, where the concentrated convergence of cells upon the midline produces a physical budge of cells. The most dorsal cells are destined to contribute to the development of the nervous system (green). Dorsal is to the right of each embryo shown from shield to tailbud. Individual representative cells are mapped at the shield stage in locations and colors indicative of their future fates (see structure color coding key). The pointed shape of these illustrated cells also denotes their directional migration/movement during gastrulation. Kupffer’s vesicle is an early sign of posterior growth and left-right patterning (white circular structure in the tail from tailbud to 18 hpf). Mesoderm induction during gastrulation will give rise to the axial mesodermal structure of the notochord (light mauve color delineated with dotted lines), and as the tail extends the paraxial mesodermal structures of the somites (light blue block/chevron-shaped structures transparently illustrated) will sequentially form from anterior to posterior over time. The PGCs complete their long migratory journey around 24 hpf and develop into the gonads. Placodal structures in the ectoderm (neural and epidermal tissues) build the eye (yellow), ear (honey color) and nose (orange) organs. Development of the heart and circulatory system begins following the tailbud with bilaterally positioned cells that migrate over to the midline and converge to form the heart (only one population of cells shown, red), after which an elaborate vasculature system is created (only the dorsal aorta and vena cava with posterior blood island shown; high to low red gradient represents flow directionality). Development of the gut, kidney, liver, and pancreas are all illustrated in the approximate positions (see key). Vascularization of the glomerulus (kidney) is shown at 48 hpf. Key for staging embryos between 24 hpf and day 2 is the position of the lateral line primordium, which migrates from the brain to the tail atop of the middle of somites depositing neuromasts along the way (see key). 48 hpf is known as the long pec stage due to the presence of the extended pectoral fin tissue (show above the yolk, gray pattern). Although pigment can be first seen at 24.5 hpf in the dorsal region of the retina (eye, brown), body pigment increasingly appears between days 1 and 2 (only the dorsal most melanocytes are shown, 48 hpf). Illustration by Michael J.F. Barresi ©2018.

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Zebrafish Research Advances Our Understanding of the Mechanisms Governing Development

ectodermal cells by 10 h postfertilization (hpf) or the tailbud stage (Bruce, 2016). Differential expression of the types and amounts of cell surface adhesion receptors, such as from the cadherin family, have been shown to be responsible for these radial intercalation movements. Loss of E-cadherin in the half-baked zebrafish mutant causes it to fail to progress beyond 70% epiboly and ultimately mutants arrest in development (Kane, McFarland, & Warga, 2005; McFarland, Warga, & Kane, 2005). In addition, epiboly progression in zebrafish is also influenced by the yolk syncytial layer that forms at the vegetalmost edge of the blastoderm. Of most significant relevance is the elaborate cytoskeletal networks affiliated with the external yolk syncytial nuclei (eYSN) (See Fig. 45.1 Bruce, 2016). The eYSN (and their centrosomes) project an expansive network of longitudinal microtubules toward the vegetal pole, as well as organize an actomyosin contractile ring at the blastoderm edge. By exposing zebrafish gastrulae to cytoskeletal blocking drugs applied to their growth media or using targeted laser ablation methods, researchers have shown that both the microtubules and actomyosin elements are in fact required for completion of blastopore closure and the full enwrapping of the embryo by the external enveloping layer (Behrndt et al., 2012; Solnica-Krezel & Driever, 1994; Stra¨hle & Jesuthasan, 1993). As epiboly pulls the blastoderm toward the vegetal pole, novel cell behaviors are simultaneously occurring at the location of the “shield.” The midline positioned shield is akin to the dorsal blastopore lip (DBL) in frogs and salamanders or the node in mouse and chick (Thisse & Thisse, 2015). The shield represents the presumptive dorsal side of the embryo, as well as the location of mesendodermal induction (Heisenberg & Tada, 2002; Rohde & Heisenberg, 2007). Recently, researchers using light-sheet microscopy have recorded the position, movements and division patterns of every individual cell through the entire process of zebrafish gastrulation. This imaging has revealed a slowing in cell division specifically in the presumptive shield prior to any visible thickening, as well as quantifying the cell dynamics driving the primary internalization of the hypoblast by involution (Keller, 2013; Keller et al., 2008). This level of quantification from a single cell to the whole organism is unparalleled for the analysis of vertebrate embryogenesis. The tissue thickening that happens at the shield is due to the opposed mediolateral intercalation of cells on both sides of the midline. This midline convergence of cell movement produces a buildup of cells at the presumptive shield location that provides the cellular infrastructure to support early neural keel development (beginnings of the brain and spinal cord) development, as well as powering the involuting movement of presumptive hypoblast cells. Involution is the inward

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folding of the blastoderm that is first initiated at the shield and will create a second layer of cells underlying the epiblast called the hypoblast. As the hypoblast forms, it is rapidly patterned into dorsal and ventral endodermal and mesodermal precursor cells. Importantly, chordamesoderm, the predecessor of the notochord, is also derived from the cells of the involuting shield (Concha & Adams, 1998; Cooper & D’amico, 1996; D’Amico & Cooper, 2001; Warga & Kimmel, 1990). The notochord is the defining feature of chordates and possesses important structural and regulatory roles throughout embryonic development (Annona et al., 2015; Lawson & Harfe, 2017). The notochord is formed at the quintessential midline through the bilateral convergence of hypoblast cells upon this location (See Fig. 45.1 Shindo, 2018). Chordamesoderm cells uniquely form a fluid-filled vacuole and a thick extracellular matrix that together build its characteristic rod-shaped morphology, which helps to provide skeletal-like support for the elongation of the embryonic axis (Ellis, Bagwell, & Bagnat, 2013a; 2013b). Moreover, the notochord will function as a central source of patterning signals to all surrounding tissues, such as the overlying neural keel, the bilaterally positioned paraxial mesoderm, as well as ventral mesendoderm derivatives like the gut and vasculature (Anderson & Stern, 2016). Interestingly, forward genetic screens using zebrafish identified many key transcription factors regulating notochord development (exemplified in Odenthal et al., 1996). Loss of floating-head (Xnot) and no tail (brachyury) homeodomain transcription factors result in a complete absence of the notochord (Fig. 45.2AeD). However, due to differences in the resulting cell fates that occupy the now vacant midline in these mutants (muscle or mesenchyme for floating-head and no tail respectively), researchers were able to decipher the cell lineages that floating-head and no tail were normally preventing by the promotion of notochord development (Amacher, Draper, Summers, & Kimmel, 2002; Halpern, Ho, Walker, & Kimmel, 1993; Melby, Kimelman, & Kimmel, 1997; Schulte-Merker, van Eeden, Halpern, Kimmel, & Nu¨sslein-Volhard, 1994; Talbot et al., 1995). Characterization of these genes illustrated the fundamental regulatory roles that transcription factors play on whole gene networks to control cell fate decisions during development. Lastly, both forward and reverse genetic approaches have identified many essential genes for the correct patterning of germ layer cell fates across all axes of the developing gastrula. This work has demonstrated a prominent role for morphogenetic signaling proteins in regulating differential gene expression and cell fate induction during early development (For more extensive reviews see Thisse & Thisse, 2015; Zinski, Tajer, & Mullins, 2017a). In fact, complex spatial and temporal models

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FIGURE 45.2 Early development. (A-D) no tail (brachyury) is required for the development of the notochord (no) as the notochord is completely lost in ntl mutants seen from lateral (A,B) and cross-sectional (C,D) views (Amacher et al., 2002). (EeH) Lateral views of 30 hpf embryos showing the ventralizing effects of bmp2b misexpression and the dorsalizing effects of a loss of bmp2b or misexpression of chordin (Pomreinke et al., 2017). (IeN) nanos-1 expressing primordial germ cells overlap specifically with sdf-1 expression patterns during PGC migration unless sdf1a or its receptor cxcr4 are knocked down by morpholino (MO) (Doitsidou et al., 2002).

of signaling systems involving the Bmp, Nodal, Wnts, and Fgf pathways have all been implicated in early axis determination (as demonstrated in Zinski et al., 2017). For example, a loss of the signals encoded by bmp4/7 or their downstream responsive transcription factors smad1/5 result in a failure to develop more ventral structures like the tail, whereas loss of the Bmp antagonist, Chordin, causes the reciprocal effect of a reduction in head and neural tissues that are representative of more dorsal fates (Fig. 45.2EeH Pomreinke et al., 2017). Primordial Germ Cells: Independent of the three germ layers, there is yet another important cell type developing during these early stages of embryogenesis, the germ cells. The germ cells referred to here will go on to form the gametes or sex cells within the gonads (ovaries or testes). Amazingly, known germ cell-specific transcripts (mRNA) can be detected in the two-cell stage embryo, and over the next few cleavages, they form four asymmetrically positioned clumps localized to plasma membrane borders (See Fig. 45.1 Braat, Zandbergen, van de Water, Goos, & Zivkovic, 1999). These clumps contain not only RNAs but proteins and mitochondria that together are known as the germplasm. Prior to the 1000-cell stage, these four clumps of germplasm are

inherited into four separate cells at which point are considered the primordial germ cells (PGCs). The first four PGCs are relatively equally dispersed across the blastula, which is followed by an elaborate process of cell migration, ultimately to the site of gonadal differentiation (Paksa & Raz, 2015). PGC migration is arguably one of the most remarkable developmental feats. Over the course of several divisions, the PGC population reaches near 50 cells in total. These cells actively migrate from their four different starting points to the presumptive gonads, which is a journey through the remaining cleavages and all of the chaotic movements of gastrulation and segmentation until they finally reside in two bilaterally positioned cell clusters just ventral to somites 8e10 (See Fig. 45.1 Raz, 2003). Analysis of PGC development in zebrafish has provided great insight into the mechanisms regulating their guidance to the presumptive gonad. Specifically, being able to watch PGCs in the live embryos with specific transgenic reporters has led to a “tumble and run” model of PGC migratory behaviors (Blaser et al., 2005; Reichman-Fried, Minina, & Raz, 2004). In this model, PGCs migrate briefly in one direction and then pause to reevaluate their new

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Zebrafish Research Advances Our Understanding of the Mechanisms Governing Development

location before migrating again. PGCs are following precisely positioned guidance cues in the embryonic environment, which PGC specific transmembrane receptors recognize as attractive or repulsive (Paksa & Raz, 2015). One guidance system that is evolutionarily conserved is the secreted protein Stromal Derived Factor 1 (Sdf1a/1b) and its PGC expressed receptor CXCR4 (Fig. 45.2IeN). Based on both the expression pattern of Sdf1a/1b and loss of function testing for both sdf and cxcr4, the Sdf1 ligands function as attractive, migratory promoting cues for which the Cxcr4 receptor interprets (Doitsidou et al., 2002). In this way, PGCs will “run” toward the greatest concentrations of Sdf1a/1b, which is displayed in a gradient of expression leading directly to the gonadal precursor cell domain. Kupffer’s vesicledDeterminant of organ laterality: Unpaired internal organs of zebrafish, such as the liver, gut, pancreas, and heart display a characteristic asymmetry in their position either to the left or right side of the midline. This asymmetry has been determined to be regulated by the transient, spherical cyst-like structure termed Kupffer’s vesicle (KV), which encloses a fluid-filled lumen. Functionally, KV is equivalent to the embryonic node of mammals (Nonaka et al., 1998) and can be observed in zebrafish embryos between 4 and 15 somite stages underneath the notochord and posterior to the yolk tube extension (See Fig. 45.1 Essner et al., 2002). The KV is formed of epithelial cells, each projecting a single cilium from its apical surface. Higher columnar shape and density of ciliated cells in the more anterior-dorsal section of the KV creates an overall leftward flow within the lumen fluid (Wang, Manning, & Amack, 2012). This induces preferential leftward expression of components in the Nodal-Lefty signaling pathway that instructs unpaired organs to form in an asymmetric manner, which can be visualized by uneven symmetry in the expression of genes, such as southpaw and pitx (reviewed in Joseph Yost, 1999). Gene mutations that affect the motility of KV cilia have also been shown to perturb cardiac laterality during zebrafish development (Choksi, Babu, Lau, Yu, & Roy, 2014; KramerZucker, 2005 ). The progenitors of KV originate from a specialized cluster of approximately 25 dorsal forerunner cells (DFC) that emerge around the shield stage (Melby, Warga, & Kimmel, 1996). The DFCs do not involute during gastrulation, but instead, migrate toward the leading edge of the blastoderm margin, move deeper and then undergo mesenchyme to epithelial transformations (Cooper & D’amico, 1996). KV architecture defects seen upon chemical inhibition of Nodal signaling and in mutants of nodal interacting proteins, such as gdf3 suggests that the nodal pathway is also necessary for proper patterning of the KV during its early development (Pelliccia, Jindal, & Burdine, 2017). The development

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of KV also depends upon the T-box transcription factor encoding genes no-tail (brachyury) and tbx16, which regulate KV’s mesenchymal to epithelial cellular transition (MET) (Amack, Wang, & Yost, 2007). Although the specific signal that induces leftward patterns of gene expression remains unclear and debated, many mutations that perturb left-right (L-R) asymmetry during zebrafish development have been traced to cilia genes. Further investigation of these cilia genes, as well as the observed correlations of asymmetric patterns in membrane potential, are promising areas for development.

Development of the Central Nervous System Neurulation: The nervous system, which is the first organ system to develop, is centrally located at the dorsal midline. The earliest morphogenesis and specification of neural ectoderm occurs at the beginning of gastrulation, and this tissue will develop into the presumptive brain and spinal cord. Prior to the visible elaboration of the brain into its fore-, mid- and hindbrain regions and the spinal cord with its extended motor and sensory nerves, a rudimentary tube first needs to be constructed. From Sea squirt to primates all chordates develop an embryonic neural tube through a process called neurulation. Evolutionarily related to the invertebrate “nerve chord,” the overall mechanisms for nervous system development across species are highly conserved. Although careful comparison of cell and tissue dynamics underlying neurulation between anamniotes (fish and frogs) and amniotes (birds and mammals) reveals some important differences in the ways by which such similar tubes are formed (Harrington, Hong, & Brewster, 2009; Korzh, 2014; Lowery & Sive, 2004). All neurulation begins with the pseudoepithelialization of the neural ectoderm into a plate. Interestingly, how teleosts like zebrafish change this plate into a tube is the primary difference from some other vertebrate species (Cearns, Escuin, Alexandre, Greene, & Copp, 2016). For instance, the neural plates of mouse and chick physically bend at discrete points parallel to the rostral-caudal axis, which serves to bring the folded edges in direct opposition to each other leading to their fusion and consequent separation from the presumptive surface epithelium (reviewed in Massarwa, Ray, & Niswander, 2014). The zebrafish neural plate does not undergo this type of folding morphogenesis. Rather, the bilateral convergentextension movements during late gastrulation continue to force the stacking of pseudostratified epithelial cells upon the midline directly above the developing notochord. This mounting compaction of centrally positioned cells forms a precursor structure called the neural keel, which will continue to compress into a similarly epithelialized neural rod.

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The functioning adult nervous system requires a central ventricle or lumen in which cerebral spinal fluid can flow. Amniotes maintain the continuity of apical to basal polarity throughout folding and fusion of the neural plate, which establishes the central ventricle upon closure. In contrast, many of the cells of the zebrafish neural rod cross the imaginary plane of the midline with some cells even maintaining contact to both lateral edges of the rod (Cearns et al., 2016). For creating a lumen through the full extent of the neural rod (formally referred to as the neurocoel), the midline needs to gain apical polarity. This is the step during zebrafish neurulation known as cavitation, during which the subcellular recruitment of apical junctions is targeted to the absolute midline of the neural rod. Polarized cell divisions connected to these new apical junctions establish an overall apical to basal polarity within each half of the developing neural tube, which creates the neurocoel and the completion of neurulation (Buckley et al., 2013). Not surprisingly, adhesion proteins like N-Cadherin and E-Cadherin are required for the proper morphogenesis of the neural tube in all vertebrate species tested (Biswas, Emond, & Jontes, 2010; Detrick, Dickey, & Kintner, 1990; Hong & Brewster, 2006; Lele et al., 2002; Morita et al., 2010; Radice et al., 1997). More recently, researchers have used the zebrafish as a model to understand better the mechanisms governing the establishment of apical to the basal polarity that is suggested to be necessary for neurocoel formation. By both visualizing and directly testing the requirements of key apically restricted proteins like Partitioning defective 3 (Pard3), we now know that these apical proteins are first shuttled along microtubules within a cell to a position aligned with the neural rod’s midline center (Buckley et al., 2013; Tawk et al., 2007). Remarkably, such positioning of these apical-defining proteins will occur at any asymmetric location as long as it is aligned at the midline, which ultimately coordinates cell division upon this plane, creating two symmetrical daughter cells. Loss of function of Pard3 or other related apical proteins causes a failure of an organized apical-basal axis, as well as a lack of lumen development (Buckley et al., 2013). Neural tube patterning: The adult central nervous system (CNS) functions through the precise organization of an enormous diversity of different neuronal networks from the forebrain to the most posterior portion of the spinal cord. Neurons extend long, cell processes called axons to a target cell where a synapse is formed, and the electrical to the chemical transfer of neuronal signaling takes place. Moreover, the CNS is equally occupied by different types of glial cells that function to provide the insulating myelin sheath around axons (oligodendrocytes), support synapse signaling, provide structural support, establish the blood-brain barrier

(astrocytes or astroglia), and offer immune responses (microglia). Lastly, neural stem cells reside in both the developing and adult brain. Radial glial cells (considered a type of astroglia) serve as the neural stem cell throughout embryogenesis and fetal development in all vertebrates; however, while radial glia maintain this role in the adult zebrafish brain they are transformed into a different neural stem cell morphology called B-cells in the adult mammalian brain (Johnson et al., 2016; Paridaen & Huttner, 2014; Taverna, Go¨tz, & Huttner, 2014). How is the rich diversity of cell types and patterns of circuits established in the developing CNS? The greater simplicity of the zebrafish embryonic brain paired with its transparency for microscopic observation and amenable experimental tools have made it a favorite model system to explore the regulatory mechanisms governing nervous system development and disease (Schmidt, Stra¨hle, & Scholpp, 2013). Similarly, across all vertebrates, organization of the zebrafish CNS can be defined as a multisegmented structure along both the anterior to posterior and dorsal to ventral axes. Most obvious is the clear segmentation of the brain into its forebrain, midbrain and hindbrain portions, of which the hindbrain is further subdivided into seven successive regions called rhombomeres (Gahtan & Baier, 2004; Schilling & Knight, 2001; Wilson, Brand, & Eisen, 2002). These rhombomere segments house specific sets of neuronal lineages many constituting the branchiomotor circuitry of the cranial nerves, among others (Chandrasekhar, 2004). As with organisms across phyla, differential expression of the Hox family of homeodomain-containing transcription factors serve to set in motion the specification of the cell types from head to tail, including cell differentiation within the CNS. Prior to the visible morphological characteristics of rhombomere segmentation, different combinations of hox gene expression prefigure these presumptive rhombomere regions, and such segmented hox expression in the hindbrain has been shown to be conserved even to the phylogenetic base of vertebrate species like the lamprey (Krumlauf, 2016; Parker & Krumlauf, 2017). It should be referenced that the lineage leading to teleosts experienced an additional genomic duplication as compared to other vertebrate lineages, and as such, for instance, zebrafish do possess substantially more hox genes than mouse (Kuraku & Meyer, 2009; Postlethwait, 2006, 2007). This kind of genetic overlap in hox genes has demonstrated low penetrance to single gene loss experiments. When the entire paralog group 1 (hoxb1a and hoxb1b) was knocked out in zebrafish; however, specific deficits in the neuronal identities in the hindbrain were revealed, like the loss of the large Mauthner neurons found in rhombomere 4 (Fig. 45.3AeF Weicksel, Gupta, Zannino, Wolfe, & Sagerstro¨m, 2014).

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Zebrafish Research Advances Our Understanding of the Mechanisms Governing Development

Because of the importance that hox genes have in regulating the activity of whole-cell type determining gene regulatory networks, much research has focused on addressing how the correct pattern of hox gene expression is first established across the embryo. Recall that cell position within the embryo matters. Morphogenetic signaling is one critical mechanism known to influence the differential expression of hox genes along both the anterior to posterior and dorsoventral axes. One essential inducer of hox genes is retinoic acid, which is dispersed in a gradient with its highest expression in the anterior embryo. Interestingly retinoic acid can enter a cell and function directly as a transcription factor to regulate the expression of a variety of genes that include those that encode many Hox transcription factors. Loss of retinoic acid signaling in pathway-specific zebrafish mutants, gene-targeted morpholinos, or pharmacological inhibitors all cause changes in the pattern of hox gene expression in the hindbrain with predictable losses in the differentiation of specific neuronal types and rhombomere identity (Alexandre et al., 1996; Emoto, Wada, Okamoto, Kudo, & Imai, 2005; Hernandez, Putzke, Myers, Margaretha, & Moens, 2007; Lee & Skromne, 2014; Maves & Kimmel, 2005; Moens & Prince, 2002; Parker, Bronner, & Krumlauf, 2014; Shimizu, Bae, & Hibi, 2006). Some of the greatest advances in our understanding of the mechanisms governing neural tube patterning have come from an analysis of cell fate determination along the dorsal to the ventral axis of the developing spinal cord. As found in all vertebrates, opposing dorsal and ventral gradients of morphogenetic signals establish the pattern of cell fates along this axis (Sagner & Briscoe, 2017). In the zebrafish spinal cord, sensory neurons occupy the most dorsal locations, interneurons and later forming oligodendroglia within the middle third of the spinal cord, and motor neurons sit just above the ventral-most structure, the floorplate (Lewis & Eisen, 2003). The floorplate is an important location, where many commissural neurons send their axons across the midline to connect the two hemispheres of the central nervous system (See floorplate in Fig. 45.1), while the sensory and motor neurons project their axons out of the spinal cord to innervate their nonneural target cells. This vertically stepped organization of the spinal cord is first patterned by the opposing gradients of Bmps and Wnts from the dorsal neural tube and overlaying surface ectoderm and Sonic hedgehog (Shh) secreted from ventrally situated structures of the floorplate and underlying notochord (Le Dre´au & Martı´, 2012). Cells of the neural tube that experience higher levels of Shh will adopt more ventral fates like motorneurons, whereas those cells exposed to higher concentrations of Bmp/ Wnts will mature into more dorsally positioned neurons. Importantly, it is the combined balance of these

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overlapping morphogens that establish the correct pattern of cell fates along the dorsoventral axis of the neural tube (Sagner & Briscoe, 2017). Shh and Bmp/ Wnt signaling function to antagonize one another by regulating specific downstream transcription factors that promote a given cell identity while also crossrepressing the transcription factors required for alternative fates. Many mutants of the Hedgehog pathway were discovered in the first forward genetic screens using zebrafish, like shh (sonic-you), smoothened (slowmuscle-omitted and smo), you (scube2), gli2 (you-too), iguana (Dzip1), and hedgehog interacting protein (hip1) to name some (Barresi, Stickney, & Devoto, 2000; Chen, Burgess, & Hopkins, 2001; Karlstrom, Talbot, & Schier, 1999; Koudijs et al., 2005; Schauerte et al., 1998; Sekimizu et al., 2004; Varga et al., 2001; Woods & Talbot, 2005). Predictably, loss of Hedgehog signaling leads to an expansion of dorsal cell types at the expense of more ventral cell types, while its misexpression replaces dorsal domains with more ventral cell type identities (as exemplified by Guner & Karlstrom, 2007; Park, Shin, & Appel, 2004; Ravanelli & Appel, 2015). Thus, morphogens along both the anterior to posterior and dorsal to ventral axes play major roles in determining the cell fates across the entire developing neural tube. Neural crest development: The last hallmark of neural tube formation is the development of a transient, multipotent stem cell population known as neural crest cells. Neuroepithelial cells located at the dorsal-most region of the forming neural tube undergo a characteristic epithelial to mesenchymal transition (EMT). During this EMT the adherent epithelial cells atop the neural tube reduce their cell junctions and actively migrate away into peripheral tissues (Klymkowsky, Rossi, & Artinger, 2010; Kwak et al., 2013; Theveneau & Mayor, 2012). Neural crest cells exiting the neural tube along the spinal cord (known as trunk neural crest) will follow guidance cues outside the CNS expressed on the surface of cells or laden within the extracellular matrix. Through a ligandreceptor recognition system, cues like ephrin-ephs, semaphorin-neuropilins, and homodimerizing cadherin receptors serve to influence the trajectory of neural crest cell migration through attraction, repulsion, or adherence (reviewed in Shellard & Mayor, 2016 and exemplified by Berndt & Halloran, 2006; Yu & Moens, 2005). Aside from their migratory behavior, one of the most uniquely important features of neural crest cells is that they are multipotent stem cells, and are thus able to contribute to several disparate types of lineages. The cell type generated by a neural crest cell progenitor depends on their specification within the neural tube, as well as the signals they experience along their journey and at their final destination (Kalcheim & Kumar, 2017; Kelsh & Raible, 2002; Martik & Bronner, 2017). Trunk neural crest has been shown to give rise to sensory

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neurons of the dorsal root ganglion, the Schwann glial cells that myelinate peripheral nerves, and most noticeably pigment cells like melanocytes. The neural crest cells that emanate from more cephalic regions (known as the cranial neural crest) contribute to pharyngeal arch development, the cartilage and bone of the skull and jaw, the cranial nerves, and neurons of the enteric nervous system. Lastly, those cephalic crest cells that contribute to portions of the heart are called cardiac neural crest. The generation of neural crest labeling transgenic lines (sox10 being a common driver) has enabled remarkable live-cell imaging in zebrafish that is continually revealing new insights into the mechanisms of neural crest migration and differentiation (Antonellis et al., 2008; Dougherty et al., 2012; Gfrerer, Dougherty, & Liao, 2013; Rodrigues, Doughton, Yang, & Kelsh, 2012). For instance, trunk neural crest demonstrates a “follow-the-leader” sort of migratory streaming that laser ablation analyses have shown require cell-to-cell interactions. In contrast, cranial neural crest cells move

as a whole population through a process known as “collective migration” (Carmona-Fontaine et al., 2011; Richardson et al., 2016). The types of neural crest and their associated mechanisms of migration behaviors appear to be largely conserved across vertebrate species (Kee, Hwang, Sternberg, & Bronner-Fraser, 2007). The processes regulating neural crest progenitor specification toward different lineages remains a major focus of the investigation, for which the zebrafish is proving to be a worthy model. The proneural transcription factor Neurogenin1 has been demonstrated to promote differentiation into dorsal root ganglion cells as opposed to the glia that wraps the dorsal root ganglion axons (Fig. 45.3GeH McGraw, Nechiporuk, & Raible, 2008). Interestingly, loss of sox10 expression in the colorless mutant is necessary for these same glial progenitor cells to develop into chromatophores and contribute to the melanocyte lineage (Fig. 45.4A,B Dutton et al., 2001; Kelsh & Eisen, 2000; Kelsh et al., 1996). Work has even progressed to a fine charting of the precise combinations

FIGURE 45.3 Building a Brain. (AeF) Early segmentation of the hindbrain is seen by the discrete banding expression of hoxb1a, krox20, and fgf3. Rhombomere four identity requires both hoxb1a and hoxb1b transcription factors as seen by the loss of segment boundaries (B,D) and loss of Mauthner neurons (3A10 antibody labeling); (F) in the double mutants (Weicksel et al., 2014). (G,H) Dorsal root ganglion precursor cells require neurogenin1 for neural development as opposed to the glial, Schwann cell identity. The green transgenic reporter labeling of DRG precursors show their expression of glial markers in neurogenin1 mutants (McGraw et al., 2008). (I,J) Lateral view of axon labeling in and out of the spinal cord with the Znp1 antibody at 48 hpf. Loss of the receptor plexin A3 causes significant pathfinding errors of motor axons as they leave the spinal cord (Palaisa & Granato, 2007).

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FIGURE 45.4 Building with Ectoderm. (A,B) sox10 is a master regulatory transcription factor required for melanocyte (pigment) cell fate throughout the body but not pigmented epithelium of the retina as evident in the colorless (sox10) mutant (B) Data generously provided by Ellie Melancon and Judith Eisen; see also (Dutton et al., 2001). (CeH) Radial progression of cell development in the retina over time (Almeida et al., 2014). (IeN) The transcription factors dlx3b and dlx4b are required for the whole generation of hair cells (Tg(pou4f3:GAP-GFP)þ) and otoliths during inner ear development (Schwarzer et al., 2017). (OeQ) The lateral line primordium (P) demonstrates collective migration upon the horizontal myoseptum of the somites as it progresses from the hindbrain to the tip of the tail depositing neuromasts along the way (Q). Data generously provided by Damian Dalle Nogare and Ajay Chitnis.

of gene expression changes that occur over the course of neural crest cell differentiation from progenitor cell to enteric neuron (Taylor, Montagne, Eisen, & Ganz, 2016). The delaminating cell behaviors displayed by neural crest cells during their emergence from the dorsal neural tube are routinely compared to the behavioral characteristics of metastasizing cancers cells in the adult (Gallik et al., 2017). Recently, the well-known neural crest marker crestin was found to be induced in melanomas in the adult zebrafish, and forced overexpression of sox10 in melanocytes was capable of triggering melanoma formation (Kaufman et al., 2016). Therefore, the study of neural crest development in zebrafish may very well be directly preparing us to understand better, cancer in those tissues whose cells originated from neural crest in the embryo.

Connecting up the nervous system- Axon guidance: The neuronal “wiring” of an adult organism is organized in a stereotypical pattern for which a scaffold of pioneering axons first laid down during embryogenesis (Lewis & Eisen, 2003; Wilson, Ross, Parrett, & Easter, 1990). Whether it is a dorsal root ganglion neuron adjacent to the somite, a motor neuron in the spinal cord or even a retinal ganglion cell neuron in the eye, once a neuron is born it must establish a physical connection or synapse with its target cell that may be located great distances away. A newly born neuron will extend a long, membranous process called an axon that is led by a highly motile and dynamic growth cone tip (Pacheco & Gallo, 2016). This growth cone houses a specific repertoire of chemical-sensing receptors capable of relaying the guidance information of environmental cues to the

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cytoskeletal motors within the growth cone. Such cues like netrins, slits, semaphorins, and ephrins that we described previously in the guidance of migrating neural crest cells also function to attract or repel pathfinding axons (Morales & Kania, 2017; Seiradake, Jones, & Klein, 2016). In zebrafish, axon guidance mechanisms have and continue to be defined for many different pathways, such as those for the spinal motor nerves, optic nerves, commissural pathways, and sensory efferent nerves to name a few. For instance, the first commissural axons to cross the midline of the central nervous system (CNS) occurs in the diencephalon of the forebrain by postoptic commissural axons at about 24 hpf. This commissure is soon followed by the crossing of anterior commissural axons in the telencephalon and then by retinal ganglion cell axons that form the optic chiasm also at the diencephalic midline. Prior to and during commissure formation, a midline spanning population of astroglial cells appears to provide a supportive substrate for pathfinding across the midline. In addition, the surrounding expressions of Slit2 and Slit3 function to repel all three of these axon types by serving to channel axons across the midline and prevent inappropriate axon wandering (Barresi, Hutson, Chien, & Karlstrom, 2005). Loss of the slit receptor roundabout2 causes optic nerves to be led “astray,” crossing the midline multiple times in aberrant positions (Hutson & Chien, 2002). Many more guidance cues and pathfinding behaviors have been identified and first characterized in the zebrafish embryo. For instance, the semaphorin receptor Plexin A3 was shown to be required for the proper exit positioning of motor nerves out of the spinal cord (Fig. 45.3I,J; Palaisa & Granato, 2007). Moreover, it was by examining the live pathfinding of peripheral sensory axons in the zebrafish epidermis that identified how random repulsive interactions between axons can create a perfectly elaborated pattern of sensory arbors (Sagasti, Guido, Raible, & Schier, 2005).

Ectodermal Derivatives Like all vertebrates, zebrafish sense their external environment through their eyes, ears, nose, and skin, as well as with a unique set of mechanosensory lateral line organs specialized to detect the flow of water (See Fig. 45.1). Interestingly, zebrafish larvae can respond to diverse sensory stimuli even before hatching, and thus, must form functional response circuits while their sensory organs and nervous system are still developing. The rapid, ex utero development and transparency of zebrafish has been instrumental in revealing close developmental links between the nervous system and sensory organs by precisely revealing the location, timing, and

processes involved in the origins of specific cell types and the formation of their functional connections. Primordia of specialized sensory organs known as cranial placodes first appear in the early vertebrate embryo as focal thickenings of the ectoderm in the vicinity of the developing brain (Schlosser, 2014). Specific cranial placodes can generate all cell types within an individual sensory organ through a series of differentiation events or inductive interactions with cells or secreted factors of the neural ectoderm, neural crest, endoderm, and mesoderm. Studies in mouse, chick, and frogs have defined specific gene members of the Six, Pax, Eya, Id, Dlx, Iro and Fox transcription factor families as the identifying molecular markers for individual placodes (reviewed in Streit, 2004; 2018). Expression analysis of these markers in developing zebrafish has helped to trace the origins of all placode precursors to an arc-shaped preplacodal region (PPR), which surrounds anteriorlateral aspects of the developing neural tube and brackets cells of the prospective neural crest (Kozlowski, Murakami, Ho, & Weinberg, 1997; Solomon & Fritz, 2002; Whitlock & Westerfield, 2000). Early lineage tracing studies demonstrated a close spatial relationship between progenitors of the epidermis, cranial placodes, neural crest, and nervous system. During gastrulation, these progenitors differentiate from the ectoderm in response to graded concentrations of BMP, Wnt, and FGF morphogens found along the dorsoventral embryonic axis (Kudoh, Wilson, & Dawid, 2002; Reichert, Randall, & Hill, 2013). Beginning with the epidermis, we describe the development of individual zebrafish sensory structures highlighting major transitions and morphogenic interactions that shape the ontogeny of diverse cell types within each organ. Epidermal development: The natural habitats of zebrafish are water bodies ranging from very low salinity to brackish ones (Harper & Lawrence, 2016). Zebrafish can thrive in such diverse habitats in part due to adaptations of their epidermal cells, which can regulate skin permeability by directly sensing the presence of distinct organic and inorganic ions, and in response to acute changes in their concentrations. Zebrafish do not development dermal scales until 30 days postfertilization (dpf), and therefore, protective adaptations of the epidermis are particularly essential to prejuvenile zebrafish survival (Sire & Akimenko, 2003). The epidermis is an expansive sensory organ comprised of a loosely organized but diverse array of sensory ionocytes. Unlike terrestrial vertebrates, the zebrafish epidermis exclusively consists of living cells and is not covered by moisture conserving keratinized layers. The epidermis is a pseudostratified epithelium, in which cells from adjacent layers connect to each other via desmosomes and to the basement membrane via hemidesmosomes

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Zebrafish Research Advances Our Understanding of the Mechanisms Governing Development

(Le Guellec, Morvan-Dubois, & Sire, 2003). Although the adult epidermis has three layers, it forms as a bilayer in the embryo, which first develops an outer or enveloping layer (EVL) from the surface blastoderm and later forms the inner epidermal blastoderm layer (EBL) during gastrulation. The EBL covers the entire surface of embryos at 90% epiboly in a single layer and begins to fuse at the embryonic midline at the 14 somite stage (Schmitz, Papan, & Campos-Ortega, 1993). Cell types in the mature epidermis of teleosts include keratinocytes, mucus-secreting cells, four types of ionocytes that regulate entry of specific cations, sensory cells, and alarm substance-secreting club cells (Henrikson & Matoltsy, 1967a; 1967b; 1967c). These diverse cell types do not randomly occur across the surface and width of the epidermis but localize on the basis of their ontogeny and local cellular interactions. Keratinocytes predominantly occur in the outermost layer and have mechanical resistance due to actinbased surface microridges. Ionocytes, secretory cells, and undifferentiated cells occur in the middle or basal epidermal layers (Chang & Hwang, 2011; Hawkes, 1974; Le Guellec et al., 2003). Lineage tracing experiments in zebrafish show that all epidermal cell types originate from non-neural ectodermal cells located near the ventral axis of the early embryo (Kimmel, Warga, & Schilling, 1990; Sagerstro¨m, Gammill, Veale, & Sive, 2005). Consistent with high levels of BMP in this region, epidermal progenitors distinctly express the BMP-induced gene DNp63 (Lee & Kimelman, 2002). Both keratinocytes and ionocytes differentiate from common progenitors under the influence of Delta-Notch and Foxi3a/b signaling proteins (Esaki et al., 2009; Hsiao et al., 2007; Ja¨nicke, Carney, & Hammerschmidt, 2007). Distinct skin ionocytes resemble kidney cells both in morphology and in the expression of ion transporter proteins, and interestingly, their proliferation, differentiation, and turnover are regulated by environmental factors, such as acidity and cold temperatures (Chou et al., 2008; Horng, Lin, & Hwang, 2009; Hwang, 2009). While the characterization of epidermal defects in the psoriasis mutant indicates that secreted factors could influence the differentiation of keratinocytes (Webb, Driever, & Kimelman, 2008), little is known about the development of mucus or alarm substance-secreting cells in zebrafish skin. It has recently been shown that all mature epidermal cell types exhibit phagocytic behaviors, and this function may serve to trim axon termini or remove debris from apoptotic neurons that are densely interspersed between the epidermal layers (Rasmussen, Sack, Martin, & Sagasti, 2015). Epidermis specifically refers to the outermost skin layer but is also a loose term for the entire skin. Thus, it is pertinent to consider how the zebrafish skin

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increases in mechanical strength and develops its characteristic pigment stripes. Zebrafish biologists are well aware that it is more difficult to penetrate larvae than embryos either with a microinjection needle, antibodies or riboprobes. This is mainly due to the presence of a hard cuticle over the skin, the bulk of which is composed of epidermal cell secretions and debris. The epidermis begins to secrete mucus after 24 hpf but does not develop a collagenous envelope until 72 hpf as it is secreted by the later developing hypodermis. Although the thickness of skin increases with time and higher degrees of collagen crosslinking, it takes about 30 days to get covered by dermal scales, which arise from epidermal placodes through epidermishypodermis interactions (Le Guellec et al., 2003). Interestingly, the defining pattern of horizontal blueblack lines interspersed on zebrafish skin is generated by xanthophore, iridophore and melanophore pigments derived from neural crest cells (reviewed in Kelsh, Harris, Colanesi, & Erickson, 2009). In contrast to other vertebrates, zebrafish pigment cells typically do not invade the epidermis, but rather migrate along specific paths namely between somites and the neural tube (medial path) or between somites and non-neural ectoderm (lateral path). Different chromatophores will migrate along different paths; such that, melanoblasts move along both medial and lateral paths, whereas iridophores are restricted to the medial path and xanthophores to the lateral path. Melanophores begin to appear after 24 hpf first in the cephalic region and progressively over the trunk where they occur in four stripes, dorsal, lateral, ventral and yolk syncytial (Hirata, Nakamura, & Kondo, 2005). The characterization of the zebrafish pigment cell specification, colorless (sox10) mutant, indicated that the identity of particular pigment cells is not defined by their migration paths but is rather determined at the time of specification itself (see Fig. 45.4A,B Dutton et al., 2001; Kelsh & Eisen, 2000; Kelsh et al., 1996). Intriguingly, pigment pattern defects in the choker mutant indicate that germ cell migration and axonal guidance cues like the Sdf-1 factor also appear to play a role in melanophore migration (Svetic et al., 2007). With its ready accessibility, the zebrafish epithelium represents a powerful alternative to tissue culture models to understand the development, physiology, and regulation of epithelial cells. Interestingly, a zebrafish enhancer trap screen has identified stable but mosaic patterns in the epidermal expression of Gal4 transgenes indicating that epidermal cells from specific locations may have distinct identities based upon local tissue interactions (Eisenhoffer et al., 2017). PlacodesdThe eye with its lens: The zebrafish eye is the first peripheral sensory organ to emerge toward the end of gastrulation and appears prominently large relative to the developing embryo. Zebrafish are tetrachromats

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that can perceive light in the visible and ultraviolet spectrum; however, the overall structure of their eye is quite similar to that of mammals. It is a spherical chamber with a transparent lens in the center of its exposed surface surrounded by a protective, light-proof sclera that encloses a laminar sensory retina. The outermost retinal layer is comprised of pigmented epithelial cells that support rod and cone photoreceptors, as well as the inner layers of bipolar, horizontal and amacrine neurons, and Mu¨ller glia. Lastly, the essential retinal ganglion cells transmit light stimuli from the eye to the optic tectum within the forebrain (Gestri, Link, & Neuhauss, 2012). During early embryogenesis, the eyes begin to develop from a central eye field or anlage within the most anterior and central section of the preplacodal region specified by the expression of transcription factors Six3, Rx3, and Pax6. This central eye anlage is split into bilateral retinal primordia by the anterior migration of cells from the adenohypophyseal placode triggered by factors secreted by the axial mesoderm that includes TGFb, FGF, and Shh. Thus, zebrafish deficient in these factors either due to genetic mutations or exposure to chemical inhibitors form one central fused eye instead of two bilateral ones (reviewed in Sinn & Wittbrodt, 2013). The first step during eye formation involves evagination of the epithelium in the forebrain region driven by migration of retinal precursor cells (see Fig. 45.1). This results in the formation of an optic vesicle whose molecular signaling interactions with the overlying ectoderm lead to the development of the lens placode around 16 hpf. Subsequently, the bilayered optic vesicle invaginates to form an optic cup that gives rise to the laminar neural retina surrounded by pigmented epithelium. Within the retina, the ganglion cells form as the eye’s first neurons around 32 hpf followed by cells of the inner nuclear layer. Rods and cones do not form until 55 hpf. All cells of the retina form circumferentially in the germinal zone near the ciliary margin. Thus, the oldest retinal cells are found toward the center, whereas, the newest form around the periphery (Fig. 45.4CeH Almeida et al., 2014; Sinn & Wittbrodt, 2013). PlacodesdOlfactory: Zebrafish rely on the sense of smell to forage, find mates, and detect alarm signals released from injured individuals in a group. The odors are transduced to specific olfactory sensory neurons via nares that appear as pits situated above the mouth and underneath the eyes. Interestingly, zebrafish have evolved unique olfactory adaptations to rapidly detect trace amounts of odorant molecules diffused in surrounding water (Hansen & Eckart, 1998). Focusing on the early development of the olfactory organ, we outline salient sensory and epithelial cell specializations that occur within the olfactory placode. Olfactory placodes are formed by the convergence of precursor cells from the anterolateral edges of the

preplacodal region, which first appear around the 17e18 somite stage as transient but distinctive ectodermal thickenings expressing the olfactory marker dlx3 (see Fig. 45.1 Whitlock & Westerfield, 2000). Around 24 hpf the periphery of olfactory placodes can be identified by a dense whorl of motile multicilia extending from columnar epithelial cells (Pathak, Obara, Mangos, Liu, & Drummond, 2007). The central portion of the olfactory placodes contains a rosette-like arrangement of neuroepithelial progenitor cells that differentiate into olfactory sensory neurons, support cells, glial cells, and neuroendocrine cells expressing Gonadotropinreleasing hormone (GnRH). At this stage, transient adendritic pioneer neurons can be observed connecting the olfactory epithelium with the olfactory bulb. Following programmed cell death of the pioneer neurons, three mature olfactory cell types emerge, ciliated, microvillus, and crypt neurons (reviewed in Whitfield, 2013). Olfactory neurons are distinguishable by the expression of olfactory marker protein and the expression of olfactory receptors for distinct odorant classes (Yoshihara, 2009). Photoconversion experiments in zebrafish have distinguished that the microvillar olfactory neurons actually originate from neural crest (Saxena, Peng, & Bronner, 2013). PlacodesdEar and Lateral line organs: The ear enables vertebrates to hear and maintain positional equilibrium by sensing linear acceleration and rotational movement. Although the zebrafish ear is not prominently visible externally like our outer ear structures, its organization does resemble that of the semicircular canals in the mammalian inner ear. It comprises three orthogonally arranged vestibular structures known as the utricle, saccule, and lagena, which end in sensory epithelial patches or maculae formed by specialized hair cells. The apical surface of these hair cells projects mechanosensory kinocilia that also tether large crystalline structures termed otoliths, which amplify and transmit mechanical stimuli to the brain via bipolar neurons that form the statoacoustic ganglion (reviewed in Whitfield, Riley, Chiang, & Phillips, 2002). Zebrafish mutants that exhibit otolith or inner ear structural defects, vestibular dysfunction and cilia abnormalities associated with sensory hair cells are unable to maintain an upright balance, and as a result such ear mutants swim abnormally either on their sides, upside down or in circles (Granato et al., 1996; Schibler & Malicki, 2007; Stooke-Vaughan, Huang, Hammond, Schier, & Whitfield, 2012; Whitfield et al., 1996). All cell types within the ear including the sensory epithelium, neurons and supporting cells originate from the otic placode, which prominently appears in a 16 hpf embryo as a thickening in the ectoderm posterior to the eyes and underneath the hindbrain rhombomeres 6 and 7 (See Fig. 45.1 Alsina & Whitfield, 2017). The otic

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Zebrafish Research Advances Our Understanding of the Mechanisms Governing Development

and epibranchial (cranial sensory nerves) placodes originate commonly from a posterior section of the preplacodal region showing enriched expression of the transcription factor Pax2 (McCarroll et al., 2012). Migrating cells from the neural crest later separate the otic placode into its distinct position as observed by the expression of transcription factors dlx3b and dlx4b. By 24 hpf the otic placode invaginates into an otic vesicle containing a large anterior otolith associated with the utricle and a smaller posterior otolith associated with the saccule. Zebrafish double mutants of the dlx3b and dlx4b genes result in a dramatically smaller otic vesicle with reduced sensory hair cell development and lack otoliths (Fig. 45.4IeN; Schwarzer, Spieß, Brand, & Hans, 2017). The hair cells differentiate from the columnar epithelial cells along the periphery of the otic vesicle at distinct times; the first ones emerge prior to otoliths and anchor them. The otoliths begin to form around 18e20 hpf as crystals of calcium carbonate that nucleate around a matrix of glycoproteins, both secreted by supporting epithelial cells within the otic placode (Wu, Freund, Fraser, & Vermot, 2011). Zebrafish sense movement patterns in surrounding water via a series of special lateral line organs, which like the ear are mechanosensory. The lateral line organs are formed by a series of individual neuromasts distributed all along the trunk and around the eyes and ears (See Fig. 45.1). Each neuromast, in turn, consists of a central cluster of sensory epithelium formed by hair cells surrounded by a rosette of supporting epithelium and encircled by mantle cells (reviewed in Dalle Nogare & Chitnis, 2017). Neuromasts can be visualized in live, unstained zebrafish using microscopes equipped with Nomarski Differential Interference Contrast (DIC) microscopy or with fluorescent styryl dyes, such as DASPEI or FM 1e43 which are preferentially endocytosed by hair cells (Ou, Simon, Rubel, & Raible, 2012). The early development of lateral line organs is understood mostly with regard to the posterior lateral line primordia (pLLp), which originates from the neural crest around 22 hpf as a placode formed by 140 cells that migrates caudally under the skin (reviewed in Dalle Nogare & Chitnis, 2017). Prior to beginning its migration, the pLLp differentiates bipolar sensory neurons, which extend an anterior axon into the hindbrain and a posterior axon into the migrating primordium (Fig. 45.4OeQ Hava et al., 2009; Knutsdottir et al., 2017; Nogare et al., 2017; Sarrazin et al., 2010; Dalle Nogare et al., 2014). The path of these migrating cells is guided by affinity-based interactions between Sdf-1 (also called Cxcl12a), a chemokine secreted by cells of the horizontal myoseptum and Cxcr receptors expressed by the migrating pLLp cells. As the cluster migrates,

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cells at the leading edge display a mesenchymal character and express Cxcr4b whereas those at the trailing edge begin to acquire apicobasal polarity typical of epithelial cells and the express Cxcr7b receptor (Dambly-Chaudie`re, Cubedo, & Ghysen, 2007; Haas & Gilmour, 2006; Sape`de, Rossel, Dambly-Chaudie`re, & Ghysen, 2005; Valentin, Haas, & Gilmour, 2007). This epithelial transition occurs within a cluster of w20 cells which stop migrating and begin to constrict at their apical ends to form rosettes described as protoneuromasts. The central cell within each protoneuromast differentiates into a progenitor of sensory hair cells. pLLp cells that stop migrating but are left out of the protoneuromasts instead deposit as interneuromasts, which can later proliferate to produce more neuromasts. Neuromasts continue to be sequentially deposited up to 48 hpf. The somite position reached by the leading edge of the pLLp is used to accurately stage the development of embryos after the first 24 h. Neuromast formation is regulated by a limiting balance between Wnt and FGF signaling pathways (Ma & Raible, 2009). Initially, the entire pLLp expresses high levels of Wnts, which activates expression of both Wnt, as well as FGF ligands. However, FGF producing cells do not respond to Wnt activators and Wnt expressing cells produce inhibitors of FGF signaling which results in establishment of an FGF signaling center at the trailing edge (Aman, Nguyen, & Piotrowski, 2011; Aman & Piotrowski, 2008; Head, Gacioch, Pennisi, & Meyers, 2013; Kozlovskaja-Gumbriene_ et al., 2017; Nikaido, Navajas Acedo, Hatta, & Piotrowski, 2017; Tang, Lin, He, Chai, & Li, 2016).

Germ Layer Integration to Build the Craniofacial Skeleton The head skeleton is structurally essential for both its protection of the brain and its functional roles in eating and hearing, and in the case of a fish, even breathing. Therefore, it is not surprising that malformations in the morphogenesis of the craniofacial skeleton during embryonic development are associated with a large range of disorders in humans, such as cleft palate, DiGeorge syndrome, or Treacher-Collins syndrome to name a few (Machado & Eames, 2017). Although the boney fish’s head has over two times as many bone elements as the mammalian head skeleton, much conservation in overall structure and function exists between the two classes; additionally, the larval zebrafish craniofacial skeleton represents a simple organization of discrete parts making it a conserved and tractable model for embryological and disease relevance (Medeiros & Crump, 2012; Mork & Crump, 2015).

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For instance, bones of the fish jaw and viscerocranium are evolutionarily homologous to the bones making up the mammalian middle ear. By the early larval stages, the head cartilage elements are forming and will eventually ossify into bone (Kimmel et al., 1998). As mentioned earlier, most of the cartilage and bone in the head is directly descendant from cranial neural crest cells (Knight & Schilling, 2006). From single-cell microinjection to cell transplantation and laser light-induced transgenic procedures, lineage tracing approaches have demonstrated that the location of cranial neural crest cells along the neural tube and the timing of their exit reproducibly contributes to cartilage of particular regions of specific elements (Mork & Crump, 2015). One of the most fascinating aspects of craniofacial development however, is the requirement to first form pharyngeal arches, which are transient embryonic structures bilaterally repeated along the rostral to caudal axis of the ventral half of the head that contribute to the gills, jaw and head skeleton among other tissues (Frisdal & Trainor, 2014). Although cranial neural crest cells primarily make up the pharyngeal arches, the entire morphogenesis of craniofacial development involves the direct interactions of cells from the mesoderm, endoderm, and ectoderm. Beginning at 20 hpf and ventral to the otic placode (ear), seven- bilateral, pharyngeal arches extend further ventral to the locations of element-specific chondrogenesis on each side of the developing jaw. Imagine the pharyngeal segments as pairs of oars being extended into the water from seven rowers seated in line within a common canoe. As mentioned above, the pharyngeal arch is made up of several types of cells, and in extending this rowing analogy, the central core of the oar would be made of mesodermal cells surrounded by successive rings of endodermal cells, cranial neural crest cells, and a final circumferential ring of ectodermal cells. Exciting recent research into zebrafish pharyngeal development is starting to reveal how these multilayered arches are formed. Cells of the endodermal layer, called the pharyngeal pouches, play an important role in dividing the streams of cranial neural crest cells into the seven separate arches, whereas, the initially positioned ventral mesoderm functions to provide a guidance cue for the directionality of arch elongation (Mork & Crump, 2015). The Tbx1 transcription factor is expressed by the mesodermal cells, and it has been shown to regulate Wnt11r and Fgf8a in subpopulations of the mesoderm. Wnt11r and Fgf8, in turn, function to properly coordinate endodermal pouch morphogenesis and attract the collective migration of the cranial neural crest cells, respectively (Fig. 45.5 Choe & Crump, 2014). This is substantially relevant to human health as mutations in the Tbx1 gene are known to be the cause of DiGeorge syndrome (Chieffo et al., 1997).

FIGURE 45.5 Building a Face. (A, B) Ventral view of Alcian Blue staining of cartilage elements in the jaw and branchial regions of 5-day old wild type (A) and tbx1 mutant (B) larvae. Loss of tbx1 leads to reduced and disorganized pharyngeal pouches at 34 hpf (C, D; AntiAlcama labeling, green), which causes the ultimate loss of craniofacial elements later in development (B). Data generously provided by Gage Crump. See also (Choe & Crump, 2014).

Segmenting the Trunk for Muscle, Bone and Much More The reach of hox genes: The stretch of hox gene influence on patterning anterior to posterior tissue identity does not stop at the hindbrain as described earlier, but is a robust mechanism for positional patterning from the olfactory placodes (nose) to the tips of caudal fin rays (tail) and all the parts in between. For instance, positioning of the pectoral fin is under the delineation of hox gene expression and its regulation homologous to the mechanisms of tetrapod forelimb development. Furthermore, patterning of the developing pectoral fin along its own anterior to the posterior axis is governed by highly conserved mechanisms involving hox genes among many others (Ahn & Ho, 2008). In particular, Sonic hedgehog morphogenetic signaling from the posterior fin bud and the antagonistic signals of Fgf8 and retinoic acid along the distal to proximal axes lead to the outgrowth of the fin, which is accomplished in part through the upregulation of progressively more 50 hox genes over time that induce progressively more distal cell derivatives of the fin (Akimenko & Ekker, 1995; Hoffman, Miles, Avaron, Laforest, & Akimenko, 2002; Laforest et al., 1998; Noordermeer et al., 2014; Prykhozhij & Neumann, 2008; Sakamoto et al., 2009; Zhao et al., 2009). One of the most obvious axial structures in the embryonic zebrafish is the repeated, chevron-shaped blocks of paraxial mesoderm running down the length of the trunk and tail. In the embryo, these segments are called somites, portions of which give rise to the myotome or skeletal muscle, to the sclerotome or cartilage and bone of the vertebrae, and to some vascular derivatives among others (Stickney, Barresi, & Devoto,

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2000). As with all segmented organisms, the identity type of each segment is highly regulated by hox genes; such that, the anterior border of hox gene expression strongly influences the cell types that will develop in each segment (Iimura & Pourquie´, 2007). As an example, hoxc6 correlates with the cervical to thoracic transition in mouse, chick, and zebrafish; however, while the overall hox-mediated mechanisms of identity are conserved, there is a discord in the expression of certain hox genes at more posterior transition sites between boney fish and amniotes (Burke, Nelson, Morgan, & Tabin, 1995; Morin-Kensicki, Melancon, & Eisen, 2002). Importantly, the regulation of hox gene expression by the opposing gradients of and antagonistic signaling by retinoic acid from the anterior mesoderm and Fgf8 from the posterior mesoderm are highly conserved across vertebrate species, including zebrafish. Somitogenesis and tail elongation: Although establishing different cell identities across the entire anterior to the posterior axis is critical, the boundaries of paraxial mesoderm segmentation represent a particularly unique physical feature that prepares this embryonic tissue for later differentiation. Somite formation or somitogenesis is the cyclical cutting of progressively more posterior paraxial mesoderm into roughly equal-sized blocks. A somite constitutes the mesoderm occupied between

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segment boundaries. Understanding the developmental underpinnings of somitogenesis have been part of intense research for many decades due to its common occurrence in all segmented creatures, its medical relevance to human disorders like scoliosis, as well as to biologist’s pure phenomenological fascination (Boswell & Ciruna, 2017; Eckalbar, Fisher, Rawls, & Kusumi, 2012; Grimes et al., 2016). The rapid external development of zebrafish paired with its amenability to timelapse microscopy and cell tracking techniques have all garnered zebrafish as a superb model to study somite formation. The zebrafish paraxial mesoderm is divided up into roughly 35 bilaterally positioned somites. Somite formation and maturation occur in remarkably regular order in a rostral to caudal direction over the trunk between 10 and 24 hpf (Fig. 45.6C and E). So standardized is somite development that a bilateral pair of somites is reliably formed every 30 min (at 28.5 C), and researchers use the total number of somites as a metric of embryonic age (e.g., 18 somites at 18 hpf, 22 somites at 20 hpf, 30 somites at 24 hpf) (See Fig. 45.1 Kimmel et al., 1995). Forming somites is a complex process involving at least four moving parts: one- the convergence of mesoderm toward the midline to both populate the presumptive anterior presegmental plate

FIGURE 45.6 Slicing up the segments. (AeF) Dorsal (A, B) and lateral (CeF) views of 12- and 30-somite wild type (AeC,E) and txb6 mutant (D,F) embryos. Tbx6 transcript (A) and protein (B) are expressed within presomitic mesoderm that disappears as a somite matures. MF20 is a marker for skeletal muscle fiber differentiation. Somites fail to form in fused somites (tbx6) mutants (D, F). (GeJ) Loss of proper segmentation in the tbx6 mutant embryo leads to disorganization of the adult vertebral elements with many instances of vertebral fusions (I,J). Data shown in AeJ was generously provided by Stephen H. Devoto.

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and power the caudal extension of the trunk, two- the generation of new, migratory cells from the caudal progenitor domain at the tailbud to further drive growth of the posterior presegmental paraxial mesoderm, threethe inflation of chordamesoderm cells to elongate and harden the notochord further supporting posterior extension, and four- the sequential and targeted epithelialization of paraxial mesoderm cells at presumptive boundary locations during trunk extension. Together these different cell behaviors lead to the periodic segmentation of the paraxial mesoderm in an anterior to the posterior direction over the course of tail elongation (Yabe & Takada, 2016). Despite the complexity of somitogenesis, the molecular components coordinating all of its moving parts are gaining clarity and found to be remarkably conserved across vertebrate species. Much insight has been garnered from mutagenesis screens that have identified many essential genes, which include but are not limited to T-box transcription factors, genes within the Notch and Fgf pathways, and cell adhesion receptors as in Eph-Ephrin signaling. One of the most dramatic phenotypes is seen when the tbx6 transcription factor is lost in the fused-somites mutant, which results in a complete loss of any somitic furrows (Fig. 45.6AeF Nikaido et al., 2002; Windner, Bird, Patterson, Doris, & Devoto, 2012). It is hypothesized that Tbx6 functions to specify the anterior half of presumptive somites leading to its Eph-ephrin mediated epithelialization (Barrios et al., 2003). However, the timing of a somite boundary forming is dictated by the cycling expression (every 30 min) of Notch-Delta signaling factors from the tailbud through the paraxial mesoderm and culminating at the next segment to be created (Holley, Ju¨lich, Rauch, Geisler, & Nu¨sslein-Volhard, 2002). Notch and Delta interact at the plasma membranes between presomitic paraxial mesoderm cells, which enables the transference of pathway activation like a wave from the tail to the last somite boundary. Simultaneously, the location of epithelialization is determined in part by the opposing antagonistic actions of Fgf8 from the tailbud and retinoic acid from more anterior, already segmented paraxial mesoderm. A caudally emanating gradient of Fgf8 morphogenetic signaling functions to repress epithelialization of the paraxial mesoderm, and as the tailbud continues to grow caudally, this Fgf8 gradient slowly releases cells from its repression (Reifers et al., 1998; Shimozono, Iimura, Kitaguchi, Higashijima, & Miyawaki, 2013). Paraxial mesoderm cells experiencing the right balance of retinoic acid stimulation, lack of Fgf8 repression, and the cresting wave of Notch-Delta activation will undergo a MET transition and form a somitic boundary (reviewed in Aulehla & Pourquie´, 2010). Severe congenital disorders of the vertebral elements like spondylocostal dysostosis that cause many

vertebrae to be fused and malformed are directly associated with mutations in genes of the Notch-Delta pathway (Penton, Leonard, & Spinner, 2012). The zebrafish provide a powerful model to better understand the developmental processes behind somitogenesis and potentially the causes of related human disorders. Somite patterning to make skeletal muscle and vertebra: Failure to properly form somites in humans causes spine defects because the cells that generate the bones of vertebrae originate from somitic cells (Fig. 45.6GeJ). The ordered segmentation of somites helps to guide the proper organization of the developing skeleton in the trunk, as well as guide the position of invading spinal motor nerves and the organized pattern of skeletal muscle fibers (Bernhardt, Goerlinger, Roos, & Schachner, 1998; Ward, Evans, & Stern, 2017; Zeller et al., 2002). Soon after a somite has formed it will begin to differentiate. The cartilage and bone precursors, called sclerotome, develop in the most ventral region of the somite, where they lose their epithelial constraints and migrate toward and around the notochord and neural tube. In an unintuitive manner, the sclerotome from the anterior half of one somite will fuse with the sclerotome of the posterior half of its neighboring somite to establish one vertebral elementda process known as resegmentation (Ward et al., 2017). As the revered salmon steak would reveal, a vast majority of the remaining somite will develop into muscle. In an adult fish steak, the small, dark triangle of muscle at the outer edge represents a focus of slow-twitch muscle fibers, while the remainder of the body wall musculature is all fast-twitch muscle fibers (Devoto, Melanc¸on, Eisen, & Westerfield, 1996). This is relevant because the slow-twitch muscle precursor cells are morphologically one of the first cell types to differentiate in the somite. These embryonic slow muscle precursor cells, named adaxial cells due to their direct adjacency to the notochord, will be the first to express myogenic genes-like myoD. Shortly after somite formation, these adaxial cells transform into fiber-like mesenchymal cells and complete a remarkable movement out to the superficial most edge of the somite, where they will differentiate into slow muscle fibers positioned in a near-perfect parallel monolayer. Fast-twitch muscle differentiation is initiated following the adaxial migration (Corte´s et al., 2003; Daggett, Domingo, Currie, & Amacher, 2007; Devoto et al., 1996; Henry, McNulty, Durst, Munchel, & Amacher, 2005; Stellabotte, Dobbs-McAuliffe, Ferna´ndez, Feng, & Devoto, 2007). Interestingly, axon guidance cues are differentially expressed by slow muscle and fast muscle precursor cells to properly guide spinal motor axons out of the spinal cord and into the developing musculature (Rodino-Klapac & Beattie, 2004; Zhang, Lefebvre, Zhao, & Granato, 2004).

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It has been well known that the amniote somite differentiates into a similar ventral sclerotomal region and a more dorsal myotome; still the myotome is derived from an outermost epithelial structure known as the dermomyotome. Only recently has the teleost dermomyotome been described (Devoto et al., 2006; Stellabotte & Devoto, 2007). The zebrafish dermomyotome is derived from the most anterior border cells (ABCs) of the newly formed somite, which will gradually become positioned to the outermost portion of the myotome, “external” even to the slow muscle monolayer. Similar to the amniote dermomyotome, these external cells function to support myotome growth (Feng, Adiarte, & Devoto, 2006; Stellabotte et al., 2007). Research across vertebrate species has helped to identify the array of patterning signals derived from adjacent tissues, namely the notochord, neural tube, surface epidermis, and lateral plate mesoderm. Although characterization of these signals in zebrafish is an ongoing area of investigation, a huge wealth of important factors has been identified by noticing changes in the shape of the somite following gene specific loses. Mutants in genes of the hedgehog pathway were originally called you-class mutants due to the nonchevron, blocky, or “U”-shaped somite they exhibited (Amsterdam et al., 2004; Barresi et al., 2000; Chen et al., 2001; Schauerte et al., 1998; van Eeden et al., 1996); reviewed in (Stickney et al., 2000). As mentioned earlier, Sonic hedgehog is secreted from the notochord, which is required for the induction of adaxial cells into slow-twitch muscle fibers (Barresi et al., 2000, 2001; Du, Devoto, Westerfield, & Moon, 1997; Lewis et al., 1999). It is hypothesized that failure to form the superficial monolayer of slow muscle fibers contributes to the resulting U-shape of the somite. Lastly, an emerging “area” where zebrafish are turning out to be an important model is occurring between the somites, at the myotendinous junction. Tendon development involves the sculpting of extracellular matrixes and elaborate connective machinery to muscle and bone, and in the case of the zebrafish myotendinous junction helping to connect the muscle fibers of one somite to another (Snow & Henry, 2009). A critical feature of somite boundary formation is the early deposition of extracellular matrix proteins like Fibronectin and Collagen. Of particular relevance is how defects in the composition or function of matrix factors in the zebrafish myotendinous junction resemble human muscular dystrophies (Charvet et al., 2013; Telfer, Busta, Bonnemann, Feldman, & Dowling, 2010). In fact, the sheer mass and direct accessibility of zebrafish muscle has made significant contributions to the modeling of human muscular dystrophies (Goody, Carter, Kilroy, Maves, & Henry, 2017; Li, Hromowyk, Amacher, & Currie, 2017).

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Development of the Circulatory System Zebrafish is a simple yet elegant vertebrate model that has yielded great insights into early cardiovascular system development. Using a stereomicroscope alone, one can view how the heart, vessels, and blood cells form in the externally developing zebrafish larva. Although the zebrafish heart has only two chambers, it has many structural features found in mammals. The fact that gas exchange through skin alone can suffice for zebrafish larvae to survive up to 5 days has allowed circulation defective mutants to reveal intermediate steps during cardiovascular development. Details regarding early morphogenesis of individual zebrafish cardiovascular organs have emerged mostly from analyses of cardiovascular cell type-specific gene expression and live tracing of fluorescent reporter transgenes (Fig. 45.7AeC; reviewed in Staudt & Stainier, 2012). In the discussion below, we highlight how and where distinct cells within individual circulatory organs originate, differentiate, and migrate in the embryo and how cellular processes and morphogenetic factors influence their assembly. Heart development: The heart is the first circulatory organ that prominently appears as a contractile structure over the anteromedial aspect of yolk in a 24 h old zebrafish embryo (See Fig. 45.1). The early heart is a single, straight tube connected to vessel precursors of the cardinal vein and dorsal aorta. Heart assembly occurs in two discrete steps, first of which forms a tubular structure and the next separates it into ventricular and atrial compartments while adding cells for their growth. The ontogeny of myocardial progenitors during distinct stages of heart assembly has been determined through the expression pattern of evolutionarily conserved transcription factors like nkx2.5. These studies have revealed that myocardial progenitors originate from bilateral fields near the blastula margins. In addition, expression analysis of the atrial cardiomyocyte-specific myosin gene amhc and the ventricular myocyte specific myosin gene vmhc show that progenitors of distinct heart chambers are spatially segregated in the blastula and occupy distinct positions in the Anterior Lateral Plate Mesoderm (ALPM) (reviewed in Bakkers, 2011; Staudt & Stainier, 2012). Initial heart assembly occurs in the primary heart field area that is medially located behind the eyes. Around 20 somite stage, myocardial and endocardial progenitors from bilateral areas within the ALPM migrate into the primary heart field and fuse to form a cone-shaped cluster termed the cardiac disk. Later, cells within the cardiac disk involute to form the linear heart tube. Medial migration of myocardial progenitors is promoted by secreted Extracellular Matrix (ECM)

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FIGURE 45.7 Organogenesis. (AeC) Development of the circulatory system has been observed with microangiography using fluorescent molecules (A,C) and endothelial cell-specific transgenic reporters (B, tg(fli1a:GFP)). A basic yet functional vasculature is present at 24 hpf that serves as a stereotyped foundation for the elaboration of a highly sophisticated system of capillaries and veins to develop throughout the entire zebrafish from 48 hpf onward (A). The intersegmental blood vessels and cerebral vasculature are visible in lateral (B) and dorsal (C) views, respectively. Data generously provided by Brant Weinstein. (DeG) Bilaterally positioned heart progenitor cells actively migrate to the midline to form the primary heart field that then becomes asymmetrically localized. Fibronectin is an extracellular matrix protein that is essential for successful heart progenitor cell migration and development of the primary heart field as seen by the pericardial edema (F, arrowhead) and sustained bilaterality with scattered progenitor cells in natter (fibronectin) mutants (F, G) (Trinh & Stainier, 2004). (HeK) Expression of glucagon, insulin, and somatostatin by the pancreatic alpha, beta, and delta cells, respectively (H,J). mnx1 is required for insulin-secreting beta cells (I,K). Data generously provided by Gokhan Dalgin and Victoria Prince. See also (Dalgin et al., 2011). (LeO) The embryonic kidney is represented by the glomerulus (marked by podocin and nephrin expression) and the pronephric duct (also expressing nephrin) (L,N). Loss of wt1 results in the specific loss of glomerulus formation (M,O) (Tomar, Mudumana, Pathak, Hukriede, & Drummond, 2014).

proteins, such as Fibronectin, whose deficiency in the natter mutant induces a cardia bifida-like phenotype (Fig. 45.7DeG Trinh & Stainier, 2004). As the heart matures around 48 hpf it loops to form a ventricle on the right side and atrium to the left with distinct and conspicuous outer and inner curvatures. From a dorsal view, the linear heart tube can be seen oriented toward the left eye of wildtype embryos around 27 hpf. This normal asymmetry in the embryonic heart position is regulated by expression of endoderm associated nodal

related genes southpaw (spaw) and pitx2 that begins around the Kupffer’s vesicle (KV) and becomes progressively restricted to the anterior, left side. Heart laterality is reversed in the spaw gene mutant straightforward (Noe¨l et al., 2013), as well as in many cilia gene mutants due to perturbation of KV signal sidedness (reviewed in Bakkers, Verhoeven, & Abdelilah-Seyfried, 2009). Besides the outer myocardial layer, the early heart consists of an inner endocardium, which originates from endothelial progenitors. Endocardial progenitors

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also originate in the ALPM but are located relatively anterior to the myocardial progenitors and migrate medially in response to Vegf (Reviewed in Haack & Abdelilah-Seyfried, 2016). The endocardium, in conjunction with hemodynamic forces, plays important roles in coordinating radial movements of myocardial cells and in regulating heart size (de Pater et al., 2009). The loss of endocardium and endothelial cells in cloche mutant embryos greatly increases the heart volume but reduces the thickness of its walls (Holtzman, Schoenebeck, Tsai, & Yelon, 2007). The AV valve separating the two chambers is also derived from the endocardium. The heart grows substantially after 48 hpf and forms distinct internal divisions across the atrial and ventricular chambers. However, later growth of the heart does not occur by the proliferation of existing myocardial cells within the initial heart tube but by addition of differentiating mesenchymal cells from the secondary heart field (SHF) area. The SHF located in the pharyngeal mesoderm is identifiable by the expression of the latent TGFb binding protein 3 (ltbp3) (Zhou et al., 2011). Late differentiating cardiomyocytes from the SHF add cells to the ventricle, atrium, and the outflow tract (de Pater et al., 2009). The outermost protective heart layer or the pericardium is the last to form around 72 hpf and includes heterogeneous cells attracted from outside the heart field areas after these undergo an epithelial to mesenchymal transition (Peralta, Gonza´lez-Rosa, Marques, & Mercader, 2014; Serluca, 2008). Pericardial edema is a frequent phenotype associated with poor heart function and circulation caused by the expansion of space between the myocardium and pericardium due to fluid buildup. Vascular development: Zebrafish larvae exhibit a stereotypic closed-loop pattern of vessels that circulate blood from the heart to peripheral tissues and back (See Figs. 45.1 and 45.7AeC). As the pattern of vessel network is reproducible across different zebrafish larvae, it is easy to detect perturbations induced by genetic or chemical manipulation of cellular processes. Significantly, the cellular origins of zebrafish arteries, veins, and capillaries (including those of lymph) are also similar to that of other vertebrates (Gore, Monzo, Cha, Pan, & Weinstein, 2012). The cells that form blood vessels belong to a special category of endothelial cells known as angioblasts, whose identity can be detected by their expression of genes, such as scl or flk-1(Kabrun et al., 1997). In zebrafish, the earliest precursors of vascular endothelial cells have been traced to the ventral mesoderm in shield stage embryos. Around the 14e16 somite stage of development, clusters of endothelial cells specified to become angioblasts migrate toward the midline and in rostral and caudal directions. The rostral stream of migrating

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angioblasts reaches pharyngeal mesoderm and ALPM, whereas the caudal stream reaches the posterior lateral plate mesoderm (PLPM) and spreads along the trunk between the endoderm and hypochord (Bussmann, Bakkers, & Schulte-Merker, 2007). Around 24 hpf, the dorsal aorta (DA) and cardinal vein (CV) are the first major undivided vessels that emerge in the trunk. In contrast to other vertebrates, the DA and CV do not fuse together in zebrafish; however, steps in their formation have revealed significant insights into conserved cellular mechanisms that guide blood vessel formation in vertebrates (Isogai, Horiguchi, & Weinstein, 2001). Blood vessels form de novo via the process of vasculogenesis involving coalescence of adjacent angioblasts (Fouquet, Weinstein, Serluca, & Fishman, 1997). Interestingly, it has been demonstrated in zebrafish that the arterial and venous fates of major vessels are predetermined: regulated by Notch signaling pathway angioblasts committed to form arteries express genes encoding the membrane ligand EphrinB2 and components of Notch signaling pathway; whereas venous angioblasts express the ephrinB4 gene encoding the receptor that recognizes EphrinB2 (Lawson et al., 2001). In response to somite secreted VegF only the DA begins to sprout a set of uniformly spaced, perpendicular Intersegmental Vessels (ISV) from its dorsal aspect. ISV assembly represents a distinct process of angiogenesis that sprouts new vessels from preexisting ones. Studies with zebrafish fli1a transgenics show that during angiogenesis, the endothelial cells at specific locations within the walls of the DA divide and differentiate to sprout a distinct “tip cell” that extends filopodia in all directions. To begin migrating, the “tip cell” first divides asymmetrically to create itself and a new daughter “stalk cell” near its connection to the parent vessel. Extending dorsally, the ISVs stay confined between the somites in response to secreted chemorepellent cues. Upon reaching the level of the dorsal neural tube, the ISVs give out rostral and caudal branches that merge to form the dorsal lateral anastomotic vessel (DLAV) (Isogai et al., 2001). Initially the ISVs do not show arterial or venous characteristics but do so later in response to blood flow (Isogai, Lawson, Torrealday, Horiguchi, & Weinstein, 2003). Lastly, newly formed vessels do have a lumen which forms later as endothelial cells form vacuoles that fuse inside the vessel (Kamei et al., 2006). Live imaging in zebrafish has shown that lymph vessels and capillaries form after cardiovascular development (Yaniv et al., 2006). Although lymph vessels are closely associated with those of blood, the two are not directly connected. Lymph endothelial cells are derived from venous endothelial cells but distinguishable via unique expression of the vegf-c or prox-1 genes (Ku¨chler et al., 2006). The trunk lymphatic vessels emerge from parachordal vessels, which are derived

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from secondary sprouts of the posterior cardinal vein and migrate along arterial but not venous ISVs (Mulligan & Weinstein, 2014). Hematopoesis: In developing zebrafish embryos, circulating blood cells first appear shortly after 24 hpf when the heart starts beating. Cells of blood like those of vessels originate from endothelial cells, presumably derive from a common precursor termed hemangioblasts. Zebrafish blood shows the presence of all major cell types of the erythroid and myeloid lineage, including granulocytes, macrophages, neutrophils, and platelets (reviewed in Carroll & North, 2014). Zebrafish blood cells form in two waves, and these differ with respect to their origin in the embryos. Between 12 and 24 hpf an initial or primitive wave of hematopoiesis generates cells of the erythroid lineage from the inner mass of cells of the PLPM (Detrich et al., 1995) and macrophages from a rostral location of the ALM (Herbomel, Thisse, & Thisse, 1999; Lieschke et al., 2002). However, these cells are limited in their ability to meet the oxygen needs of growing embryos. Thus, similar to mammalian blood development, a second or definitive wave of hematopoiesis around 30 hpf generates multipotent hematopoietic stem cells (HSCs) from hemogenic endothelium in the ventral wall of the dorsal aorta designated as the aorta-gonadal-mesonephros (AGM) (Bertrand et al., 2010; Kissa & Herbomel, 2010). Zebrafish mutants of the runx1 gene that is expressed in the AGM show severe loss of definitive blood cell lineages at 5 dpf (Kalev-Zylinska et al., 2002; Lam, Hall, Crosier, Crosier, & Flores, 2010). Around 36 hpf, most HSCs circulate in the blood and settle into supportive niches of the kidney or migrate to a posterior region in the tail termed Caudal Hematopoietic Tissue (CHT), which represent sites of continued hematopoiesis from 3e4 dpf through adulthood (Ma, Zhang, Lin, Italiano, & Handin, 2011).

Organogenesisdthe Kidney The pronephros: The kidney in zebrafish embryos or the pronephros is an anatomically simple structure comprising a pair of tubular nephrons (See Fig. 45.1). The zebrafish nephron is functionally similar to that of higher vertebrates, comprised of three functional segments, which, from the anterior to posterior, are the glomerulus, proximal tubule, and distal duct. The pronephros develops from intermediate mesoderm progenitors adjacent to those of the blood and distinguishable by the expression of transcription factors encoding genes pax2, pax8, lhx1 and hnf1ba (reviewed in Drummond & Davidson, 2010). During somitogenesis, distinct pronephric segments form as cells of the intermediate mesoderm undergo mesenchyme to epithelial transition (MET) and differentiate under the influence

of retinoic acid from the anterior paraxial mesoderm (Wingert et al., 2007). The most anterior section of the pronephros or the glomerulus occupies a medial position in the embryo, posterior to the fin buds near the third somite. The glomerulus is essentially a network of leaky blood capillaries covered by a unique type of epithelial cells termed podocytes characterized by multiple apical foot processes. The interdigitating podocyte foot processes form a barrier around the capillary endothelial cells and filter blood by excluding cells and solutes on the basis of molecular size and charge. Pronephric cells within the glomerulus originate from a cluster of progenitors expressing paralogs of the Wt1 transcription factor (Majumdar & Drummond, 1999). The segments immediately posterior to the glomerulus include the pronephric neck and proximal tubule, which are highly endocytic and serve to reabsorb the bulk of leaked macromolecules (Anzenberger et al., 2006). These segments are distinguished by the expression of the transcription factor encoding gene pax2a. In zebrafish mutants of the wt1, pax2, and hnf1 transcription factors, one kidney segment expands at the expense of the other (Fig. 45.7LeO Majumdar, Lun, Brand, & Drummond, 2000; Naylor, Przepiorski, Ren, Yu, & Davidson, 2013). Notch-dependent signaling regulates differentiation of pronephric epithelial cells into transporting or multiciliated cell types after 24 hpf (Liu, Pathak, Kramer-Zucker, & Drummond, 2007). Around 24 hpf, the anterior pronephros develops a characteristic lateral bend due to extension of the neck segment, which progressively contracts beyond 48 hpf due to fluid flow driven collective migration of proliferating cells (Vasilyev et al., 2009). The very posterior pronephric segment, including the exterior tubular opening, is a highly proliferative zone distinguished by its expression of the oncogene c-ret (Vasilyev et al., 2009). The distal tubule of the kidney is closely associated with the corpuscle of Stannius. This is a cluster of cells marked by the T-box transcription factors Tbx2a/b that regulates calcium and phosphate levels (Drummond, Li, Marra, Cheng, & Wingert, 2017). The zebrafish kidney begins to function around 48 hpf as the glomerular podocytes differentiate to filter blood. Physiological adaptations of kidney enable zebrafish to thrive in freshwater by wasting a high volume of water while reabsorbing essential solutes. The zebrafish kidney, unlike mammals, lacks smooth muscle and instead relies on highly motile multicilia that act as fluid pumps to continually expel urine. Embryonic kidney defects in zebrafish mutants prominently manifest phenotypes, such as glomerular cysts and/or severe pericardial or general edema. A majority of cystic kidney phenotypes have revealed the importance that cilia genes play in the cellular mechanisms associated

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Zebrafish Research Advances Our Understanding of the Mechanisms Governing Development

with complex human disorders of not only the kidney but across multiple organs. In the adult zebrafish, the kidney appears as a dense bilateral network of, branched, pigmented tubules under the dorsal body wall between the gills and caudal fins. The growth of pronephric into adult mesonephric kidney occurs by the addition of more nephron units rather than proliferation within the initial nephron (Zhou, Boucher, Bollig, Englert, & Hildebrandt, 2010).

OrganogenesisdEndoderm Derived Organs The digestive system of zebrafish, which includes the intestinal tract, liver, and pancreas originates mainly from the embryonic endoderm. Its early development could potentially reveal what defects may underlie metabolic disorders, such as diabetes, and has thus, spurred excitement in the analysis of zebrafish endoderm mutants and transgenics (Field, Ober, Roeser, & Stainier, 2003). These efforts are defining a trajectory of specification and assembly of distinct digestive structures beginning as early as the blastula stage, although the digestive tract does not open from the mouth to anus until 5 days postfertilization. Severe defects in all endoderm derived structures in the zebrafish cyclops and squint mutants have revealed an essential role for the secreted morphogen Nodal and members of the FoxA transcription factor family in specifying distinct regions of the digestive tract (Dougan, Warga, Kane, Schier, & Talbot, 2003; Niu, Shi, & Peng, 2010). Consistently, nodal is expressed at high levels near the origins of endoderm within a few layers of circumferential cells near the blastula margin between the yolk and overlying cells of the embryo (Fauny, Thisse, & Thisse, 2009). With the onset of gastrulation these endodermal cells involute forming large, flat cells with bipolar filopodial extensions that eventually converge dorsally and distribute along the anteroposterior embryonic axis (Pe´zeron et al., 2008). With the onset of somitogenesis, these cells distribute sparsely along the most ventral aspect of the embryo and coalesce medially to form a rod-like structure along the midline of a 20 hpf embryo (Miles, Mizoguchi, Kikuchi, & Verkade, 2017). This endodermal rod specializes into anterior pharyngeal endoderm at the level of the first somite, and a posterior alimentary tract. In the following section, we summarize how the developing gut specializes into three functionally distinct segments; an anterior intestinal bulb, a midintestine, and posterior-intestine and gives rise to the liver and pancreas as outgrowths of the intestinal bulb. Gut development: The distinct segments of zebrafish gut are lined by epithelial cells that in essence, become transformed from the endoderm into gut between 26 and 52 hpf (See Fig. 45.1). During this process, the

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rod-like endoderm grows and generates a bilayer of cells. Subsequently, this bilayered structure begins to form interspersed cavities as the cells form apical membranes. The cavities eventually join to form a continuous lumen, which by 76 hpf extends across its entire length (Ng et al., 2005). While no cell death occurs during the formation of intestinal cavities, the development of distinct membrane polarity needs to happen during a precise window of development. For instance, the delayed membrane polarization in the heart and soul (has) mutant produces multiple lumens in the intestinal bulb (Horne-Badovinac et al., 2001). The intestinal bulb, which forms the most anterior segment of the alimentary canal, has only one lumen but multiple folds directly connects the esophagus to the intestine. The intestinal bulb serves a similar role as the stomach, having highly proliferative secretory cells restricted to the basal layers. However, the intestinal bulb lacks the stomach’s characteristic acid-secreting cells. The mid-intestine is comprised of enterocytes, as well as enteroendocrine cells interspersed with mucus-secreting goblet cells. Interestingly the same mid-intestinal enteroendocrine cells can secrete both glucagon and somatostatin, unlike cells of the pancreas, where these are secreted by distinct cell types (Ng et al., 2005). The posterior intestine is mainly comprised of cuboidal epithelium. While there are no folds in the middle and posterior intestine, the intestinal bulb develops multiple folds around 4 dpf, when the anus also opens to the outside. By 5 dpf, the entire intestine becomes enveloped by a thickened layer of mesenchyme, the yolk gets depleted, and the embryo begins to actively eat. Liver development: The liver plays important roles in metabolism, detoxification, and fat digestion, and can be first distinguished by expression of differentiated liver markers, such as Ceruloplasmin (Cp) (Korzh, Emelyanov, & Korzh, 2001). cp expression at the 16 somite stage demonstrates the early left sidedness of the liver, located in the embryo near the junction of the intestinal bulb at about the first somite axial position (See Fig. 45.1). Cell fate analysis with gut-specific reporter lines shows that liver progenitors reside asymmetrically within late blastula stage embryos, in ventrolateral and dorsolateral positions along the left side and right sides respectively. Beside fluorescent transgenes, liver cells can be easily visualized by fluorescently labeled modified phospholipids. During the 24e28 hpf, the hepatocyte progenitors aggregate as a distinct projection within the intestinal primordium, near the spot where the prospective intestinal bulb also loops. Around 50 hpf as the liver buds and grows further, the tissue connecting it to the intestinal bulb becomes restricted to form the bile duct. The liver becomes vascularized by endothelial cells, which encapsulate it partially around 60 hpf with complete invasion by 72 hpf (Field et al., 2003).

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45. Zebrafish as a Model to Understand Vertebrate Development

Pancreas development: The zebrafish pancreas, like other vertebrates, is an important exocrine and endocrine gland associated with the intestine. In the developing zebrafish embryo, pancreas precursor cells can be distinguished by the expression of the pdx1 (pancreatic and duodenal homeobox1) homolog as a bilateral cluster adjacent to the midline around the 10 somite stage (Biemar et al., 2001). The initial pancreas appears to contain a single endocrine islet surrounded by exocrine tissue. These coalesce by the 18 somite stage to form a single domain located on the right side of the midline (See Fig. 45.1). The quintessential function of the pancreas is its production of hormones, including insulin. The pancreas is best known for its alpha, beta, and delta cell types that function to secrete glucagon, insulin, and somatostatin, respectively (Fig. 45.7H,J). While insulin expression can be observed within the pancreas as early as 12 hpf, somatostatin and glucagon expression appears later at 16 and 24 hpf respectively. The mnx1 transcription factor plays a required role in promoting insulin-secreting beta-cell development at the expense of alpha cell fates while having no influence over delta cell development (Fig. 45.7I,K Dalgin et al., 2011). Moreover, the Hedgehog pathway plays an essential role in the development of the endocrine pancreas, which is lost in mutants like sonic you (shh) and smoothened (DiIorio, Moss, Sbrogna, Karlstrom, & Moss, 2002; Roy, Qiao, Wolff, & Ingham, 2001; Tehrani & Lin, 2011).

Summary The multitude of experimental advantages from its size, accessibility, development speed, and visible tractability to its genetic, molecular, and cellular approaches have all made the zebrafish a powerful model system to uniquely piece together the complexities underlying vertebrate development. The conservation of these developmental mechanisms across phyla has provided great and often direct relevance of the investigations in zebrafish to human development and disease. An emergent theme in development is how only a limited array of signaling molecules is reiteratively used to induce distinct tissues by establishing different patterns of conserved, cell-type determining genes that predominantly vary only in their spatiotemporal expression within a given tissue. As new experimental tools and systems-level approaches improve, it will be fascinating to watch how our understanding of the complex interactions between cells, tissues, and even between the embryo and its environment function together to shape the vertebrate embryodthrough the perspective of the zebrafish.

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C H A P T E R

46 Zebrafish as a Model for Revealing the Neuronal Basis of Behavior* Kimberly L. McArthur1,2,a, Dawnis M. Chow1,a, Joseph R. Fetcho1 1

Dept. of Neurobiology and Behavior, Cornell University, Ithaca, NY, United States of America; 2Dept. of Biology, Southwestern University, Georgetown, TX, United States of America

Introduction Wh ile much of the original work using the zebrafish model in neuroscience was focused on developmental questions (Eaton, Farley et al., 1977; Eisen, 1991; Eisen, Pike, & Debu, 1989; Grunwald, Kimmel, Westerfield, Walker, & Streisinger, 1988; Kimmel, 1982; Kimmel, Sessions, & Kimmel, 1981; Myers, Eisen, & Westerfield, 1986; Streisinger, Coale, Taggart, Walker, & Grunwald, 1989), zebrafish offer major advantages for revealing how vertebrate brains produce behavior (Fetcho & Liu, 1998; Kimmel, Eaton, & Powell, 1980). This role for fish might not seem so obvious, but the advantages of small size, transparency, and genetic tools that lie at the heart of the power of the zebrafish model also catalyze its role in studies of brains and behavior. These studies are typically not directed toward understanding fish per se, but rather have as their goal the discovery of principles underlying brain function that apply to vertebrates broadly. This is possible because, to a first approximation, all vertebrate brains are the same (Butler & Hodos, 1996).

Vertebrate Brains Have Much in Common Across Species Nearly every region in the nervous system, with the notable exception of the multilayered cerebral cortex in mammals, is present in all vertebrates, including such important regions as the olfactory bulbs, retina,

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telencephalon, hippocampus, amygdala, thalamus, hypothalamus, optic tectum, basal ganglia, hindbrain, cerebellum, spinal cord and the major dopaminergic, serotonergic, and noradrenergic neuromodulatory systems (Butler & Hodos, 1996). This is not too surprising because vertebrates evolved to do many of the same behaviors. They all use their senses (vision, hearing, smell, proprioception) to move about in coordinated ways to find food, water, shelter, and mates, and avoid becoming food. They learn about the external world through experience and store that information to produce adaptive behavior. While the execution (swimming vs. walking for example) might be different across animals, the brain regions and neural computations that process internal and external sensory information and produce appropriate outputs have much in common across species, allowing us to use zebrafish to inform us about how brains and spinal cords work across vertebrates generally.

The Challenge of Understanding How Brains Generate Behavior Our understanding of the generation of behavior by vertebrate brains is still in its infancy; it is one of the greatest remaining biological puzzles. To set the stage for the power of zebrafish and its place along the path to revealing brain function, a short account of what is needed to explain how any particular behavior is

Authors contributed equally.

* Supported in part by grants from the NIH (NINDS: F32-NS083099, F32-NS084654 and R01-NS026539).

The Zebrafish in Biomedical Research https://doi.org/10.1016/B978-0-12-812431-4.00046-4

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Copyright © 2020 Joseph Fetcho. Published by Elsevier Inc. All rights reserved.

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produced by a brain will provide a useful context for discussion.

Behavior The challenge begins with a proper definition of the behavior of interest. It could be a relatively simple motor behavior, such as the escape behavior that fish use to avoid predators or a seemingly simple decision to turn to the left or right. On the other hand, the behavior might be much more complex, such as learned avoidance of dangerous places, navigation through a complex environment (3D for fish), or chasing down a prey item. In either case, the behavior must be studied rigorously to define how it is shaped by sensory information from the environment and prior experience, and the details of the movements that underlie it. These transformations from sensation to decision and action, and the influence of behavioral history (learning and memory) and internal state (hunger, fear, sexual, or general arousal, etc.), are what must be explained at the level of the neurons and circuits in the brain and spinal cord.

Neurons The next critical step is to reveal what neurons are involved by somehow monitoring their activity in the brain. Given that most brain regions are heterogeneous, with neurons of different function and connectivity, the ability to resolve the activity of individual cells is critical. Electrophysiological or imaging approaches are methods of choice. Electrophysiology, such as wholecell patch recording from individual cells, can resolve the exact firing pattern of the neurons and even the synaptic inputs from other cell types. It also allows the detailed study of the electrical/ion channel properties of the cells, which influence how they respond to synaptic input from other neurons. The study of how these properties and circuit function can be modified by neuromodulators that can reshape activity without altering physical connections between neurons also demands electrophysiology (Lovett-Barron, Andalman et al., 2017). The limitations of electrophysiology are its invasiveness (surgical exposure and paralysis are typically needed) and an inability to record from more than a few neurons at once. Both of these limitations are overcome by using imaging, done most commonly with fluorescent calcium indicators, which change intensity when calcium levels in neurons rise (or fall) during activity. Here, the benefit of the transparency and small size of the larval fish is enormous, as large numbers of neurons, even deep in the brain, can be imaged with single-cell resolution to tie the activity of both individual neurons and neuronal populations to

behavior. The benefit, however, comes at the cost of temporal resolution, as the millisecond and submillisecond resolutions of electrophysiology are impossible to achieve with the slow kinetics of calcium sensors and their indirect ties to the actual electrical activity. Fortunately, zebrafish are accessible both to electrophysiological and imaging approaches (Fetcho & O’Malley, 1995; Legendre & Korn, 1995), so one can obtain both high temporal resolution and population-level information about the ties between active neurons and the behavior of interest.

Wiring The patterns of neuronal activity during a behavior are not sufficient to reveal how that behavior is produced by a brain because the brain uses connections/ wiring between neurons to compute and transform external and internal information into appropriate action. The precise wiring is critical, although it sometimes does not get the attention it deserves. It is what defines the circuit! This is often missing from studies of zebrafish because it is one of the hardest bits of information to obtain. The most direct path to it is to record from pairs of neurons, then activate one and monitor the electrophysiological response in the other, to reveal the type (chemical, electrical, excitatory, inhibitory, neuromodulatory) of synapse, its strength, and the plasticity of the connection in response to different patterns of activation of the presynaptic cell. This pairwise (and even triple) recording can be done in zebrafish with optical targeting of electrodes to particular neurons (Koyama et al., 2016). It is hard, but feasible. Newer approaches to reveal connectivity combine optical activation of neurons with optical (or better, electrophysiological) monitoring of responses in potentially connected cells (Forster, Dal Maschio et al., 2017). Another approach to connectivity that is just now reaching fruition in zebrafish is electron microscopic reconstruction of large regions of the brain, which reveals the pattern of connections of the neurons because both neuronal processes and their input and output synapses are visible at EM resolution. This does not provide direct functional information about connection strength or synaptic properties, but it is still critical information as it forms a ground plan for circuit wiring. Remarkably, there are still very few cases of any brain, in which, we know even how many output connections a single neuron has and how they are distributed because it is so difficult to obtain them with physiological methods. Still, this is critical for understanding circuit function and computations. EM provides this and can do so even for entire regions of the brain. In the small larval zebrafish, this opens access

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A History of Zebrafish as a Model for Understanding the Generation of Behavior

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to the baseline structural connectivity anywhere in the brain via whole-brain EM sectioning (Hildebrand et al., 2017; Wanner, Genoud, Masudi, Siksou, & Friedrich, 2016). The combination of light level imaging of structure, even at high resolution with expansion microscopy (Freifeld et al., 2017), and activity with EM level connectivity anywhere in the brain makes zebrafish unique among the vertebrates as a model for revealing how brains produce behavior.

and can be added to expand its predictive power. The goal is the simplest model that explains what is known, has large predictive value, and hopefully reveals features of the computational algorithms used to transform internal and external sensation and stored information into adaptive actions.

Models

The Beginning

The complexity of the wiring in brains, the varying electrical properties of different neurons and the multitude of interactions among cells makes understanding how the brain generates behavior a challenge, even with activity and wiring in hand. We typically cannot look at the data and intuit how computation works. This demands models, grounded in the data, that simulate the neuronal interactions, so that we can see whether the information we have captures what is needed to produce observed patterns of activity appropriate for driving behavior (Koyama et al., 2016; Naumann et al., 2016). Importantly, such models should have predictive power, since understanding and explanation in science is synonymous with predictive power. Indeed, even brains themselves are mostly concerned with predicting what to do next. Tests of the predictions assess model quality. Zebrafish are especially amenable for testing predictions because the model predictions often take the form of what happens to the behavioral output (and activity of other cells in the circuit) when particular neurons are removed or activated or inactivated. The transparency of the zebrafish allows for very specific optical and genetic perturbations, including application of light-activated proteins for activating or silencing neurons, discussed in later sections (Douglass, Kraves, Deisseroth, Schier, & Engert, 2008; Kimura, Satou et al., 2013).

All of the steps toward revealing the neuronal basis of behavior summarized in the previous sections become easier if one can simply look to the brain of an animal and see everything that is happening all at once. Larval zebrafish are now close to offering that, but reaching the required level of sophistication took several decades of effort starting with the genetic studies of George Streisinger in the late 1970s and early 1980s. Working nearly alone, so as not to risk the careers of his students and postdocs, Streisinger pioneered methods for the induction and study of mutations in the zebrafish genome using gamma rays (Chakrabarti, Streisinger, Singer, & Walker, 1983; Stahl, 1994). He also developed methods of inducing haploidy in the progeny of mutants, which facilitated the identification of recessive lethal mutations, as the haploid zygotes can survive for several days after fertilization (Kimmel, 1989; Patton & Zon, 2001; Streisinger et al., 1989; Streisinger, Walker, Dower, Knauber, & Singer, 1981). These pioneering genetic methods, along with in vivo imaging of zebrafish during development (Eisen, Myers, & Westerfield, 1986; Liu & Westerfield, 1988), initial studies of motor innervation and function by the Oregon group (Liu & Westerfield, 1988), and their generosity in helping others with the model, catalyzed large-scale screens for mutations affecting embryogenesis in zebrafish in Boston (USA) and Tubingen (Germany), which identified roughly 2000 genes necessary for normal development (Driever, Solnica-Krezel et al., 1996; Haffter, Granato et al., 1996).

The Goal In the end, we can claim to understand the behavioral features under study when we have a model built upon known information about cell types, their electrical properties and activity, and their wiring that both produces the output seen during the behavior and that generates predictions that have been confirmed by empirical biological methods. The model need not include everything, and likely will not, as biological systems are complex at many levels. Predictions will eventually falter, but further biological studies will reveal critical features that are missing from the model

A History of Zebrafish as a Model for Understanding the Generation of Behavior

Mutagenesis and Behavior Studies of genetic effects on behavior took a leap forward in zebrafish work because the big screens included behavior-based assays in the larvae that took advantage of the early development of motor behavior in order to identify mutations producing defects in motility and optomotor processing. In the screening by Granato et al (Granato, Van Eeden et al., 1996), mutagenized progeny were examined for motility defects in response to touch (which develops in normal embryos at around 24 h postfertilization). This screening

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identified 166 mutations that resulted in motility defects and included mutations that affected muscle development, incapacitated proteins necessary for synaptic function at the neuromuscular junction or centrally, and genes necessary for correct wiring of the nervous system. Another interesting screen exploited the tendency of larval zebrafish to swim along and visually track moving gratings (termed the optomotor response). In an optomotor-based assay (Neuhauss, Biehlmaier et al., 1999), mutants from the Tubingen screen (450 lines) were exposed to a moving grating displayed on a monitor, and those that failed to accumulate at one end of a rectangular arena were examined for defects in visual anatomy and sensitivity. This screen identified 25 mutants, 13 of which exhibited retinal dystrophy (loss or degeneration of cell layers in the retina) and others that exhibited defects, such as lens degeneration, deficiency in the pigment melanin, miswiring of axons, and absence of retinal ganglion cells, among other defects.

The Quantitative Study of Behavior in Larval Zebrafish Investigations into the neuronal basis of behavior in normal and mutant zebrafish depend on a critical foundation in the careful quantitative analysis of the behaviors. Early on, scientists were concerned with the relationship between activity in the giant Mauthner cell and the escape behavior it was thought to initiate in teleost fish (including the zebrafish), and sought to relate the activity of this cell to the behavior (Eaton & Farley, 1975). Because the escape response (C-start) elicited by this cell is so fast, there were early technological limitations in being able to record the movement of the animal with sufficient temporal resolution. In a Cstart, fish make a characteristic fast bend to one side (so that the animal resembles a C) before a weaker bend to the other side followed by lower amplitude bends as the animal swims away (Kimmel, Patterson, & Kimmel, 1974). Kimmel et al. (Kimmel et al., 1974) were able to reveal the latency of this rapid response by using a photoresistor to detect breaks in a beam of light. This method determined that a fast C-start response develops after 4 days postfertilization (dpf) in zebrafish, although slower responses were present after 3 dpf. The C-start has a latency of 15 ms in the larvae, with similar latencies (and general movement patterns) in adult animals. Advances in video technology allowed higher rates of image acquisition. Eaton et al. (Eaton, Farley et al., 1977) filmed larval escapes to a probe touch at 133 frames per second and were able to obtain more detailed information about the series of movements larvae produce during a C-start escape.

At 44e48 hpf and 88e95 hpf, analysis of these images revealed a response latency of 10.2 and 12.3 ms respectively. Video imaging of larvae at 1000 Hz with custom-written automated computer analysis (Liu & Fetcho, 1999), and more recently with software to analyze multiple animals simultaneously (Burgess & Granato, 2007), has refined this number even further; in response to an acoustic tone, 6 day old larvae produced either a short latency C-Start with a mean of 5.3 ms or a longer-latency C-start with similar movement kinematics as the fast one, but with a latency averaging 28.2 ms. Although early behavioral studies focused on C-start escapes, the combination of high-speed imaging and detailed quantification they pioneered have provided the foundation for inquiries into circuits controlling other aspects of zebrafish behavior. Classification of slow and fast swimming behaviors, routine turns, and prey-capture behaviors by careful quantification of tail bend angles and frequencies, swim velocities, and other fin and eye movements, provided a valuable reference point for many studies of zebrafish neural circuits (Budick & O’Malley, 2000; Burgess & Granato, 2007).

The Advent of in vivo Functional Imaging in Zebrafish Taking advantage of the transparency of zebrafish, researchers took a key step toward understanding the behavioral function of neurons by performing the first imaging of neuronal activity of single cells in the brain and spinal cord of an intact vertebrate (Fetcho & O’Malley, 1995; O’Malley, Kao, & Fetcho, 1996) (Figure 46.1A). Neurons were filled by injection of dextran tagged synthetic calcium indicators that are picked up by neuronal processes into the muscle or spinal cord and the labeled neurons were imaged with confocal microscopy in restrained fish embedded in agar. These indicators increase their fluorescence as calcium levels rise in electrically active neurons. The early experiments revealed the potential to simply watch the activity of cells even deep in a vertebrate brain. However, it was important to develop methods for filling neurons with indicators that did not depend on damaging their processes to facilitate indicator uptake. This was done initially by injecting calcium indicator into single-cell embryos and raising them to larval stages, at which time, cells throughout the body, including brain and spinal cord, were filled with an indicator that could be used to detect neuronal activity (Cox & Fetcho, 1996). A better way was on the horizon, however, with the demonstration of the ability to make transgenic fish with the beautiful expression of fluorescent proteins (Higashijima, Hotta, & Okamoto, 2000; Higashijima, Okamoto, Ueno,

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FIGURE 46.1 (A) Calcium green-dextran backfilled Mauthner neuron exhibits a dramatic increase of fluorescence during an escape response (hotter colors represent higher image intensities, see color bar lower left). Each image is 400 ms apart, and the scale bar is 10 microns. Inset in bottom right indicates the level of fluorescence in each image. Adapted from (O’Malley et al., 1996). (B) Cameleon expressed throughout the nervous system, and in particular, Rohon-Beard (RB) sensory neurons in the spinal cord (left, arrows). The RB neurons exhibit an increase in the YFP/CFP ratio in response to an electrical stimulus applied at 5 s. Adapted from (Higashijima et al., 2003). (C) Whole-brain light-sheet imaging during visual adaptation to moving gratings. Larval fish was suspended over a video display (top left) while the motor responses were monitored from spinal ventral roots. Tuning of individual neurons was assessed by their response to gratings moving in a particular direction (traces one to five, see arrows on bottom) and each neuron was color coded by its preferred direction (see bottom arrows). Each box on the left is a magnified region from the image on the right, corresponding to habenula, tectum, and hindbrain. The preferred tuning of all imaged neurons during the course of the experiment is shown by the color map on the left. Adapted from (Freeman, Vladimirov et al., 2014). (D) A triple patch recording showing the activity of the ipsi and contralateral Mauthner neuron after depolarization of a feedforward interneuron. Adapted from (Koyama et al., 2016). (E) Chr2-mCherry (green) expressed in spinal cSF neurons and GFP expressed in a sensory-related CoPA interneuron (top). Targeting the cSF neurons with blue light (bottom) causes current to flow into the CoPA neuron due to synaptic contact between the cells (arrows top). Adapted from (Hubbard et al., 2016).

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Hotta, & Eguchi, 1997; Long et al., 1997). In parallel, the first genetically encoded fluorescent calcium sensor was engineered by adding a calcium-binding domain to a fluorescent protein (Miyawaki, Llopis et al., 1997). Subsequently, zebrafish were the first vertebrates with a transgenic fluorescent indicator of neuronal activity expressed in all neurons (Higashijima, Masino, Mandel, & Fetcho, 2003) (Figure 46.1B). The early genetically encoded calcium sensors worked in fish but had relatively small signals, so there was a lag in their adoption after an initial successful demonstration of their functionality (Higashijima et al., 2003). Most early works applied various synthetic indicators, although there was some application of the early genetic indicators by the Friedrich lab (Li, Mack et al., 2005), which also did important early imaging work of exposed portions of adult zebrafish brains (Friedrich & Korsching, 1997). This has changed recently with the optimization of indicators that can produce robust calcium transients, in the best situations detecting calcium transients from even single action potentials (Chen, Wardill et al., 2013; Nagai, Yamada, Tominaga, Ichikawa, & Miyawaki, 2004; Tian, Hires et al., 2009). Many of these are based on an approach, in which, a calcium-sensitive protein linker (calmodulin) is placed in a split fluorescent protein such that the conformation changes upon calcium binding in a way that partially reconstitutes the normal structure of the fluorescent protein to produce a fluorescence increase (Nagai, Sawano, Park, & Miyawaki, 2001). Optimization of these so-called GCaMPs over the last decade has generated a variety of indicators with varying time-constants, sensitivity to changes in calcium concentrations, and improved brightness (Chen, Wardill et al., 2013; Nagai et al., 2004; Tian, Hires et al., 2009). Improved indicators expressed under the original pan-neuronal promoter produce robust signals in neurons throughout the brain in larval fish (Ahrens, Li et al., 2012; Ahrens, Orger et al., 2013). These new indicators, along with other technological developments including very sensitive, fast cameras and fast laser-based imaging with sheets of light swept through the brain, has led to the recent imaging of activity of nearly every neuron in the larval brain at about two brains a second as a fish restrained in agar attempted to move (Ahrens, Orger et al., 2013) (Figure 46.1C). In this case, the movements themselves can be observed by freeing portions of the body, as done initially in early functional imaging experiments (Ritter, Bhatt, & Fetcho, 2001), or motor activity monitored by recording from motor nerves in paralyzed fish (Masino & Fetcho, 2005). Active neurons can then be mapped onto a zebrafish brain atlas (Randlett, Wee et al., 2015), or they can be monitored in fish with known neuronal subtypes

labeled in a different color than the calcium indicator to reveal more about the active neurons. Zebrafish is the only vertebrate in which simultaneous whole-brain imaging of neuronal activity with simultaneous behavioral monitoring is possible, after a history of innovations in the model. Revealing circuits, however, depends on knowing more about the neurons than their activity. Fortunately, transgenic approaches have produced beautiful lines that label neurons with different neurotransmitter phenotypes and transcription factor identities (Bae, Kani et al., 2009; Higashijima, 2008; Kinkhabwala, Riley et al., 2011; Satou et al., 2013). Labeling fluorophores targeted to the membrane can also reveal the detailed projections of the cells (Forster, Arnold-Ammer et al., 2017; Pan, Freundlich et al., 2013). The combination of calcium imaging in one color with imaging of morphology or cell type markers in a different color provides the link between activity and some of the other key features that determine a neuron’s role in a behavioral circuit. This combination has been used in many ways, but one example is the combination of structural and functional imaging (Farrar, Kolkman, & Fetcho, 2018; Kimura, Satou et al., 2013; Kinkhabwala, Riley et al., 2011; Koyama, Kinkhabwala, Satou, Higashijima, & Fetcho, 2011) that revealed a striking pattern of alternating glycinergic and glutamatergic columns in the hindbrain, an organization that is less evident in adult vertebrates as neurons migrate (Higashijima, Schaefer, & Fetcho, 2004). This columnar patterning reflects a ground plan for circuit formation during hindbrain development. Mapping this ground plan in zebrafish revealed the transcription factors that define these columns, the morphologically different cell types localized to particular columns, the disposition of neurons for particular circuits within the columnar pattern, and the orderly recruitment of neurons during swimming by location/birth order in columnsdpatterns also evident in spinal cord (McLean, Fan, Higashijima, Hale, & Fetcho, 2007; McLean & Fetcho, 2009). While much of the work with functional imaging has focused on calcium sensing, zebrafish offer the opportunity to look at fluorescently tagged synaptic markers in vivo (Chow, Zuchowski, & Fetcho, 2017; Niell, Meyer, & Smith, 2004), which has allowed studies of synapse formation during circuit construction in development (Niell et al., 2004), as well as the first in vivo imaging of the activity-dependent translocation of a kinase implicated in changes in synaptic strength to synaptic sites in vivo (Gleason et al., 2003). The possibilities for tying molecular dynamics to synaptic and circuit dynamics are enormous in zebrafish, but still largely untapped.

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A History of Zebrafish as a Model for Understanding the Generation of Behavior

Electrophysiology in Zebrafish Electrophysiology provides high temporal resolution information about neuronal activity that is missed by the slower calcium imaging. The giant Mauthner cell and its role in the initiation of escape from predators in fishes and amphibians discussed earlier was the focus of early physiological studies in fish (Diamond, 1971; Faber & Korn, 1978). The escape response can be elicited experimentally by mechanical or acoustic stimuli, such as touch or water pressure waves (Eaton, Bombardieri et al., 1977; Eaton & Farley, 1975; Eaton, Nissanov, & Wieland, 1984; Kimmel et al., 1974). The M-cell is much larger than other neurons, and as such, produces a noticeable change in the electric field in and around the M-cell and, possibly, even outside of the fish when an action potential occurs. These can be recorded utilizing extracellular electrodes. Early on, such recordings, along with a high-speed video of escaping zebrafish larvae, revealed that the response latency in larvae was much the same as in adult animals (Eaton & Farley, 1975). The M-cell could be detected responding to stimuli as early as 40 hpf, but with increases in response amplitude and decreases in escape duration up to 100 hpf (Eaton, Farley et al., 1977). While extracellular recordings are useful, they often do not allow identification of the neuron being monitored, can suffer from low signal-to-noise, and may not permit satisfactory discrimination between action potentials generated by nearby neurons. Other studies, initially in goldfish, used sharp electrodes inserted into individual cells (Faber, Fetcho, & Korn, 1989; Korn & Faber, 2005) to reveal circuit connections of identified neurons. Single-cell electrophysiology from identified neurons in larval zebrafish, however, reached fruition with the application of patch electrophysiology, which was already being widely used in other animals, especially mammals. This approach was critical because larval neurons are smaller than mammalian cells (many are 5e10 um in diameter) and difficult to record with sharp electrodes, but amenable to patch recording, in which the recording electrode (usually a hollow glass pipette tapered to a polished tip filled with a physiological solution, a few microns in diameter) seals onto a patch of membrane that is then ruptured to give electrical access to the inside of the neuron. The neuron can then also be filled with dye via the recording electrode to reveal its structure. The earliest patch physiology in zebrafish was performed by Legendre and Korn (1994, 1995) and was used to study the quantal nature of the synaptic release of glycine (an inhibitory neurotransmitter) onto M-cell, and the voltage-dependence of the glycine receptor channel conductance. Patch recording from spinal neurons followed later (Drapeau, Ali, Buss, & Saint-Amant, 1999). Extending physiological

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work on the M-cell in zebrafish beyond studies of inhibitory conductances in larvae, Hatta and Korn used whole-cell patch recording to characterize the electrophysiological properties of the M-cell in adult zebrafish and compare its properties to the known ones of adult goldfish (Hatta & Korn, 1998). Physiological properties between the two species were very similar. This early work in zebrafish larvae and adults, along with anatomical studies suggested that a class of interneurons called cranial relay neurons played a major role in Mauthner cell circuitry in zebrafish, as previously shown for goldfish (Faber et al., 1989; Korn & Faber, 2005). However, it would take simultaneous paired and triple neuron patch recordings a decade later to definitively show the circuit connectivity of cranial relay neurons and the M-cell in zebrafish larvae and to begin to reveal synaptic connectivity of neurons in the larval brain and spinal cord (Bhatt, McLean, Hale, & Fetcho, 2007; Koyama et al., 2011; Koyama et al., 2016; Satou, Kimura et al., 2009) (Figure 46.1D). Whole-cell recordings allowed scientists to narrow down the defects present in some of the zebrafish mutants generated in the Tubingen screen. For example, a careful electrophysiological analysis of the twitch once mutant, whose body movements fatigue easily, revealed a previously undiscovered consequence of a mutation in the protein rapsyn, which was known to localize acetylcholine receptors on the surface of muscle fibers (Ono, Shcherbatko, Higashijima, Mandel, & Brehm, 2002). Such a mutation causes muscles to become less responsive to high-frequency stimulation, leading to quick fatigue in response to stimulidan observation that presaged the identification of human patients with similar deficits. Similar physiology led to critical discoveries about synaptic transmission and the functional organization of nerve-muscle connections (Wang & Brehm, 2017; Wen et al., 2013; Wen, McGinley, Mandel, & Brehm, 2016). Patch electrophysiology was also key to revealing the circuit properties of interneuron-motoneuron connections and network behavior in the spinal cord. Early work recording from single spinal motoneurons, red/ white muscle fibers, and interneurons with characteristic morphology gave indications of the spinal motoneurons and interneurons active during locomotion in larvae (Drapeau et al., 1999; Saint-Amant & Drapeau, 2000) that complemented prior studies of motoneurons in adults (Liu & Westerfield, 1988). Paired recordings of a specific interneuron class (Chx10 CiDs) in the spinal cord and large primary motoneurons that innervate massive numbers of muscle fibers in a body segment revealed that the motoneurons receive excitatory input from this class (Bhatt et al., 2007). This circuit is active during fast swimming and escapes. However, during slow swimming, paired recordings (and functional

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imaging of populations) revealed that a different class of commissural interneurons, the MCoDs, provide drive to the smaller secondary motoneurons that innervate fewer muscle fibers (McLean et al., 2007; McLean, Masino, Koh, Lindquist, & Fetcho, 2008; Ritter et al., 2001). Paired recordings between the CiDs and MCoDs reveal that when the CiD network is active, the MCoD network is inhibited (they are mutually exclusive in the activity), which revealed, as a general principle, that different interneuron networks are used in the spinal cord depending on locomotor speed and strength. The findings from these studies in larval zebrafish were later confirmed to operate across vertebrates moving at different speeds. Mutations of interneuron networks in both mice and horses walking, galloping or bounding revealed defects at certain speeds depending on which, interneurons were disrupted (Andersson, Larhammar et al., 2012; Crone, Zhong, Harris-Warrick, & Sharma, 2009; Kullander, Butt et al., 2003). Today, electrophysiological approaches continue to reveal with high temporal resolution the physiological activity of cells and the connectivity that underlies behavior.

Circuit Perturbations Zebrafish offer powerful ways to perturb neuronal activity to test ideas and formal models of the role of those neurons in behavior. Early perturbations in larvae used laser ablation of specific neurons, followed by behavioral testing. Calcium imaging of activity within the reticulospinal neurons in hindbrain suggested that several large reticulospinal neurons (a class of neurons in the hindbrain, which projects into the spinal cord), along with the Mauthner cells, contributed to the fast escape response (O’Malley et al., 1996). To demonstrate their respective contributions, Liu & Fetcho (Liu & Fetcho, 1999) used a focused excitation laser to kill calcium-dye loaded neurons via a phototoxicity effect. These experiments revealed that two non-Mauthner reticulospinal neurons were important, along with the Mauther cell, in head-touch triggered escapes, whereas the Mauthner cell acted independently of them in tail-triggered escapes. The development of highpower long-wavelength lasers for multiphoton fluorescence imaging allowed for direct, specific ablation of neurons via heating effects, with the care necessary to avoid damaging the surrounding tissue. Even higher power lasers and multiphoton excitation can be used for ablation and cutting of processes in vivo with minimal heating (Koyama et al., 2016). Another method for cell ablations relies on introducing a nitroreductase enzyme into zebrafish neurons, which metabolizes the prodrug metronidazole into a cytotoxic compound (Curado et al., 2007). This method has the advantage

of being genetically targeted to specific cell types and can be used to ablate larger populations of neurons that would be more time-consuming to achieve using laser ablations. For example, nitroreductase mediated ablation of serotonergic dorsal raphe neurons was used to demonstrate their role in visual sensitivity (Yokogawa, Hannan, & Burgess, 2012). Until relatively recently, stimulation during singlecell electrophysiology was the only method available for affecting the activity of a specific neuron without ablation or chemical reagents. This changed with the advent of the field of optogenetics, in which, lightsensitive ion channels (e.g., channelrhodopsin from the alga Chlamydomonas reinhardtii), genetically expressed by neurons, may be used to excite or inhibit cells when they are exposed to specific wavelengths of light (Boyden, Zhang, Bamberg, Nagel, & Deisseroth, 2005). This approach was easy to implement and rapidly adopted in neuroscience in many models. In the earliest published use of optogenetics in zebrafish, mechanosensory neurons in the trigeminal nucleus and spinal cord were transgenically modified to express channelrhodopsin, and exposure to blue light was sufficient to trigger an escape response in 24 hpf embryos (Douglass et al., 2008)dwith optogenetic activation of mechanosensory neurons simulating a noxious touch stimulus. Understandably, optogenetics has revolutionized the field of neuroscience, with impact on zebrafish neuroscience as well. Since the earliest days, a wide variety of more effective optogenetic constructs have been engineered from the original ion channels, and new tools specifically for zebrafish are available (Forster, Dal Maschio et al., 2017). Optogenetics may be used to reveal the behavioral consequences of circuit activation. For example, expression of ChR2 and halorhodopsin specifically in Chx10 (a transcription factor) positive hindbrain neurons was used to conclusively link their activity with the initiation of (and stopping, in the case of halorhodopsin) swimming in the animal (Kimura, Satou et al., 2013). In other work, activating subsets of a group of spinally projecting midbrain cells (the nucleus of the medial longitudinal fasciculus) elicited smooth tail bending in a directionspecific manner, implicating these neurons in the generation of body posture and steering (Thiele, Donovan, & Baier, 2014). Another application of optogenetics in zebrafish is to fill or supplement a role traditionally occupied by paired-patch electrophysiologyddemonstrating the connectedness of neurons in a neural circuit. A recent example comes from studies of the cerebrospinal fluid contacting neurons (CSF-cNs), which are now known to be a type of proprioceptive sensory neuron in the spinal cord that modulate bending movements

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Advantages and Disadvantages of Zebrafish Compared to Other Model Organisms

(Hubbard, Bo¨hm, Prendergast, Tseng, Newman, Stokes, & et al., 2016). Hubbard and colleagues demonstrated that these neurons connect both to a specific type of primary motoneuron (the CaP, an individually identifiable cell present in each repeating segment of the tail) and a sensory interneuron by recording the motoneurons or interneurons with traditional electrophysiology while using patterned light to stimulate a subset of CSF-cNs expressing channelrhodopsin (Figure 46.1E). A definitive advantage of optogenetics over electrophysiology for neuronal activation studies is the ability to stimulate multiple individual neurons in a class at once, or in any sequence, in a non-invasive manner. This allowed the authors of this study to examine the convergence of input from the CSF-cNs to motoneurons. Functional imaging and optogenetics applied in zebrafish have also included studies of eye movements, where imaging of activity revealed how a neuronal integrator was implemented at the population level (Miri et al., 2011)da discovery that informed work on primates (Joshua & Lisberger, 2015). Subsequent work applied optogenetics to push the network dynamics of the brain from one regime to another in order to test mathematical models of network function (Gonc¸alves, Arrenberg, Hablitzel, Baier, & Machens, 2014). In that work, none of the tested models fully predicted the results, and the experiments led to the creation of a new model that had not been anticipated.

Imaging and Perturbations in Freely Swimming Zebrafish The vast majority of modern imaging and perturbation techniques only work well in restrained fish. However, there are a handful of methods developed for recording and perturbing neuronal activity in freely swimming zebrafish larvae. One system for recording the activity of unrestrained animals relies on bioluminescence using the jellyfish protein aequorin. Aequorin emits light upon a rise in calcium during neuronal activity in the absence of light stimulation, unlike fluorescent proteins (Naumann, Kampff, Prober, Schier, & Engert, 2010). Another approach conditionally converts a green fluorophore (CaMPARI) to red based on the activity level of neurons, when and only when they are in the presence of UV light. This allows scientists to reveal populations of neurons that were active specifically during a particular interval of a behavioral task at the time the UV light was applied (Fosque, Sun et al., 2015). Calcium responses can also be imaged in unrestrained fish at times when they are not moving (Muto & Kawakami, 2016), but imaging of cells in larvae while they are moving is a much greater challenge only now being attacked

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(Kim et al., 2017). Finally, a number of perturbation approaches have been developed and used in animal models to genetically render neurons sensitive to chemical compounds to which they are normally insensitive. The compounds can then be applied to activate (or silence) specific sets of neurons. This methodology relies on introducing chemically activated ion channels under genetic control. For example, zebrafish neurons are not normally sensitive to capsaicin or to menthol but can be made sensitive by the introduction of ligand-gated ion channels from other species (Chen, Chiu, McArthur, Fetcho, & Prober, 2016). These approaches, along with recent technological developments to perform highresolution fluorescence imaging in moving animals, provide a path to studies of neuronal function in freely behaving animals (Kim et al., 2017; Symvoulidis, Lauri et al., 2017).

Advantages and Disadvantages of Zebrafish Compared to Other Model Organisms for the Study of the Neural Basis of Behavior What is a Model Organism in the Context of Systems Neuroscience? A model organism is one of a few species that have been chosen for extensive study by large communities of biologists. These few species typically: share a relatively short generation time, facilitating genetic approaches; share a wide range of available tools and techniques; and are amenable to genetic manipulation. The implicit assumption of studying model organisms is that the neurobiological principles gleaned from a small number of species will ultimately be widely applicable to many species (including humans) because of shared organizational features and shared behavioral problems that all species must solve. These are coupled with similar principles underlying the circuit level solutions of shared behavioral problems, such as finding food or mates, determining what is good or bad in the world, or making adaptive behavioral choices. Of course, much is also to be gleaned from studies of non-models with specialized behavioral abilities, ala Krogh’s principle, which states “For a large number of problems there will be some animal of choice or a few such animals on which it can be most conveniently studied,” especially as genetic engineering technology moves toward powerful methods like CRISPR, which allows access to genetic manipulation and labeling in species more broadly (Albadri, Del Bene, & Revenu, 2017; Kimura, Hisano, Kawahara, & Higashijima, 2014; Krogh, 1929; Liu & Westerfield, 1988).

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Nematodes, Flies, and Mice Versus Zebrafish The relative merits of zebrafish for behavioral work can best be appreciated through comparison with other model organisms used in neuroscience today. Here, we focus on nematode worms, fruit flies, zebrafish, and mice, due to their wide use and genetic accessibility. The nematode roundworm C. elegans is quite small (about 1 mm in length) and exists in a male form and a hermaphrodite form. The development of the animal is highly stereotypical across individuals. As there are only 302 neurons in the adult animal, it was possible for developmental biologists to track the lineage of each cell from the single-cell stage (Sulston & Horvitz, 1977; Sulston, Schierenberg, White, & Thomson, 1983) and to identify specific cells across different individuals. Not only is every neuron individually identifiable, but the entire network of synaptic and electrical connectivity between neurons has been mapped with serial electron microscopy (White, Southgate, Thomson, & Brenner, 1986). C. elegans exhibits a number of interesting behaviors, such as movement toward sources of food, movement away from noxious stimuli, and mating behaviors (Bargmann, 1993). They are amenable to genetic manipulation, and all of the genetically expressed molecular tools known to work in zebrafish also work in C. elegans. Like the larval zebrafish, nematode worms are completely transparent, so it is possible to perform fluorescence microscopy anywhere in their bodies. However, the small scale of the animal and the internal pressurization needed to maintain its body integrity make electrophysiological recordings substantially more challenging than in fish (Goodman, Hall, Avery, & Lockery, 1998). Additionally, the neurons of the nematode worm typically function using graded potentials, unlike the action potentials utilized by most neurons in other animals (Lockery & Goodman, 2009). The main difference from other models, however, is that C. elegans manages to accomplish the challenges of survival and reproduction within its natural environment with many fewer neurons than more complex invertebrates and vertebrates. The distinction between nervous systems that solve behavioral problems with fewer neurons (as in C. elegans) versus many more neurons (as in zebrafish) gets to the heart of how neurons generate behavior. In terms of nervous system complexity and behavioral repertoire, the fruit fly Drosophila melanogaster seems not wildly different from zebrafish. Larval zebrafish may exhibit less behavioral complexity than adult fruit flies because of the added requirements of flight and mating in the fly. Adult zebrafish, however, have additional behavioral complexity and cognitive abilities. Importantly, both free-swimming larval fish and adults must produce critical behaviors well enough for survival to

live to reproduce; in that sense, understanding the behavior in a larval animal is just as important as in adults, something that is also true for larval and adult flies. The adult fruit fly brain has roughly 135,000 neurons (Alivisatos et al., 2012), similar in amount to that of a larval zebrafish. Also, like zebrafish, fruit flies have sensory organs for vision, olfaction, taste, and touch/proprioception, although sensitivity to auditory stimuli may be more limited (Albert & Go¨pfert, 2015; Gillespie & Walker, 2001; Vosshall & Stocker, 2007). The wealth of genetic and molecular tools available to label these systems in Drosophila is better than those available for zebrafish. Differences in the organization between vertebrate and invertebrate brains provide some unique advantages (and disadvantages) for Drosophila. Invertebrate neurons may rely less on using large numbers of neurons to drive behavior (via what is called population coding) than vertebrate ones (Pearson, 1993), and as a result, smaller subsets of neurons may have a similar genetic and functional identity in Drosophila. It has been possible to isolate small populations of interneuronsdin some cases, single interneuronsdgenetically in the fly (Fischbach & Dittrich, 1989; Jenett, Rubin, Ngo, Shepherd, Murphy, Dionne, & et al., 2012). Single neurons with big behavioral effects are more common in insects, such as Drosophila, than in vertebrates, with the notable exception of the Mauthner cell. However, differences between the fly brain and the brain of vertebrates also lead to disadvantages in flies. The small size of the neurons and axonal and dendritic processes distant from the cell body make electrophysiological recording of the relevant activity of the neurons (which may occur in fine processes of tiny cells) more challenging in flies, although this is also an issue in many larval zebrafish neurons. Even calcium imaging data may have some unresolvable ambiguity, as active regeneration of voltage changes happening in neuronal processes may not be reflected in the calcium levels measured at the cell body. Furthermore, insights gained from studying neural circuits in zebrafish are more likely to apply to the more closely related humans than work in flies, if the goal is simply to understand humans rather than reveal principles of the biological organization more generally. The fruit fly is not as optically transparent as larval fish. Fruit flies also pupate between maggot stage and adults, so it is difficult to follow the development of neural circuits in the same individuals. In contrast, the development and formation of neural circuits are accessible at any stage in larval zebrafish because of its transparency. Mice are the most popular model organism for systems neuroscience because of their close phylogenetic relationship with humans. Mice, unlike zebrafish, possess a neocortex, which is the subject of many studies in systems neuroscience because of its role in complex

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Major Avenues of Investigation in Zebrafish With a Couple of Case Studies

behavioral tasks. Mice do exhibit cognitively sophisticated behaviors that do not exist in zebrafish, such as acoustic communication, social bonding, and caring for young (although there are fish that have some of these) (Bass & Baker, 1990). Despite relatively long generation times compared to flies and worms, there are sophisticated molecular tools available to mouse researchers that do not have close analogs in other model systems. For example, it is possible to introduce transgenic constructs into the brains of mice by applying viruses that infect particular types of neurons (Warden, Selimbeyoglu et al., 2012). Other synaptic crossing viruses, such as rabies virus, have been engineered to jump from neuron to neuron, tracing out the circuit of connected cells involved in a particular behavior (Callaway & Luo, 2015). Furthermore, mice can be trained to perform complex tasks and conditioned to fear certain stimuli, and in this way, have been fundamental in developing theories of memory formation and learning at a network level (Betley, Xu et al., 2015; Harvey, Collman, Dombeck, & Tank, 2009; Redondo et al., 2014). Because of their young age, larval zebrafish seem to have a lesser capacity for learning in some, probably less biologically relevant tasks. These differences may be mitigated by using adult zebrafish for studies of learning and memory, where appropriate. The large scale of the mouse brain also provides some advantages for optogenetics. Because it is so large, it is easier to excite neurons using light in freely moving mice because minimicroscopes can be mounted on their heads. For the most part, however, the large, opaque brain and the skull are a disadvantage for optical imaging. In sum, each model, as one might expect, has strengths and weaknesses. The special advantages for studying the neuronal basis of behavior in zebrafish are optical and electrophysiological access to neurons anywhere (and even everywhere at once, using optical whole-brain imaging) in an intact vertebrate brain in an animal model with established tools to genetically target neurons.

Major Avenues of Investigation in Zebrafish With a Couple of Case Studies Because of its unique advantages, the zebrafish serves as a model system for the study of many neural circuits and behaviors. Zebrafish have been used to study a variety of visual behaviors (Helmbrecht, Dal Maschio, Donovan, Koutsouli, & Baier, 2018; Portugues & Engert, 2009), including the optokinetic (Beck, Gilland, Tank, & Baker, 2004; Chen, Bockisch et al., 2014; Emran et al., 2007; Kubo et al., 2014; Portugues, Feierstein, Engert, & Orger, 2014; Schoonheim, Arrenberg, Del Bene, & Baier, 2010) and optomotor (Ahrens, Huang et al., 2013;

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Maaswinkel & Li, 2003; Naumann et al., 2016; Portugues, Haesemeyer, Blum, & Engert, 2015; Quirin et al., 2016) responses to visual motion. Indeed, these visual responses have been used as behavioral assays in mutagenesis screens to identify genes involved in visual system development and function (Brockerhoff et al., 1995; Muto, Orger et al., 2005; Neuhauss, 2003). Zebrafish also serve as a model of gaze (Beck et al., 2004; Bianco, Ma et al., 2012; Easter & Nicola, 1997; Greaney, Privorotskiy, D’Elia, & Schoppik, 2017; Mo, Chen et al., 2010) and postural stabilization (Ehrlich & Schoppik, 2017; Hubbard et al., 2016; Migault, van der Plas et al., 2018; Roberts, Elsner, & Bagnall, 2017; Semmelhack et al., 2014), as well as spatial navigation strategies, such as phototaxis (Ahrens, Huang et al., 2013; Burgess, Schoch, & Granato, 2010; Guggiana-Nilo & Engert, 2016; Horstick, Bayleyen, Sinclair, & Burgess, 2017; Lee, Ferrari, Vallortigara, & Sovrano, 2015). The olfactory system also plays critical behavioral roles in both larval and adult fish (Friedrich & Korsching, 1997; Li, Mack et al., 2005; Yaksi, von Saint Paul, Niessing, Bundschuh, & Friedrich, 2009). Considerable work has focused on motor behaviors and their control by the brain, including swimming (Ahrens, Li et al., 2012; Bagnall & McLean, 2014; Bhatt et al., 2007; Bianco, Kampff, & Engert, 2011; Borla, Palecek, Budick, & O’Malley, 2002; Budick & O’Malley, 2000; Burgess & Granato, 2007; Fidelin, Djenoune et al., 2015; Gahtan, Tanger, & Baier, 2005; Granato, Van Eeden et al., 1996; Hubbard et al., 2016; Kimura, Satou et al., 2013; Kinkhabwala, Riley et al., 2011; Liu & Westerfield, 1988; McLean & Fetcho, 2009; McLean et al., 2007; McLean et al., 2008; Menelaou, VanDunk, & McLean, 2014; Montgomery, Wiggin, Rivera-Perez, Lillesaar, & Masino, 2016; Mu, Li, Zhang, & Du, 2012; Patterson, Abraham, MacIver, & McLean, 2013; Portugues et al., 2015; Ritter et al., 2001; Sankrithi & O’Malley, 2010; Satou, Kimura et al., 2009; Thiele et al., 2014; Trivedi & Bollmann, 2013; Warp, Agarwal et al., 2012; Wiggin, Peck, & Masino, 2014; Wyart, Del Bene et al., 2009), escape movements (Burgess & Granato, 2007; Dunn, Gebhardt et al., 2016; Eaton & Emberley, 1991; Eaton & Farley, 1975; Eaton et al., 1984; Eaton, Bombardieri et al., 1977; Eaton, Lavender, & Wieland, 1982; Fetcho & O’Malley, 1995; Kimmel et al., 1980; Kinkhabwala, Riley et al., 2011; Korn & Faber, 2005; Koyama et al., 2011; Koyama et al., 2016; Lacoste, Schoppik et al., 2015; Lambert, Bonkowsky, & Masino, 2012; Liu & Fetcho, 1999; Liu, Bailey, & Hale, 2012; McLean et al., 2007; Mu et al., 2012; O’Malley et al., 1996; Prugh, Kimmel, & Metcalfe, 1982; Pujala & Koyama 2019; Ritter et al., 2001; Satou, Kimura et al., 2009; Takahashi, Narushima, & Oda, 2002; Temizer, Donovan, Baier, & Semmelhack, 2015; Thorsen & Hale, 2005; Thorsen & Hale, 2007; Thorsen, Cassidy, & Hale, 2004; Trivedi &

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Bollmann, 2013; Yao, Li et al., 2016), fin movements (Green & Hale, 2012; Green, Ho, & Hale, 2011; Hale, 2014; Hale, Katz, Peek, & Fremont, 2016), opercular movements(McArthur & Fetcho, 2017) and rheotaxis (Haehnel-Taguchi, Akanyeti, & Liao, 2014; Levi, Akanyeti, Ballo, & Liao, 2015; Liao & Haehnel, 2012; Olszewski, Haehnel, Taguchi, & Liao, 2012; Oteiza, Odstrcil, Lauder, Portugues, & Engert, 2017). In the realm of more complex behaviors, zebrafish exhibit fear responses and learned fear conditioning (Agetsuma, Aizawa et al., 2010; Amo, Fredes et al., 2014; Duboue, Hong, Eldred, & Halpern, 2017; Okamoto, Agetsuma, & Aizawa, 2012), which can be used to study neural circuitry related to anxiety and associative learning. Zebrafish also show typical circadian activity patterns and have been used to study circuit mechanismsdand consequencesdof vertebrate sleep (Gandhi, Mosser, Oikonomou, & Prober, 2015; Kaslin, Nystedt, Ostergard, Peitsaro, & Panula, 2004; Oikonomou & Prober, 2017; Prober, Rihel, Onah, Sung, & Schier, 2006; Yokogawa, Marin et al., 2007; Zhdanova, Wang, Leclair, & Danilova, 2001). Zebrafish have complex social behaviors like shoaling (Buske & Gerlai, 2012; Canzian, Fontana, Quadros, & Rosemberg, 2017; Hinz & de Polavieja, 2017; Saverino & Gerlai, 2008), which have been used as behavioral assays to study the developmental impact of early ethanol and nicotine exposure (Buske & Gerlai, 2011; Fernandes, Rampersad, & Gerlai, 2015; Miller, Greene, Dydinski, & Gerlai, 2013). These are a just sample of behavioral abilities - efforts are underway to categorize the many behaviors in this model (Cachat, Stewart et al., 2011; Kalueff, Gebhardt et al., 2013; Mu et al., 2019). To illustrate how zebrafish and associated technologies have been used to investigate the neural basis of behavior, we present two case studies in which this animal model’s unique advantages have been used to advance our understanding of sensorimotor processing. In each case, the scientific progression parallels (and depends upon) the course of methodological innovation. Furthermore, these two cases follow similar scientific trajectories: initial characterization of stereotyped behavior, followed by efforts to reveal the identity and activity patterns of the neurons involved, with subsequent utilization of that basic understanding to ask even more diverse and sophisticated questions. Regarding each type of behavior, we outline major findings about its neural underpinnings and suggest fundamental principles derived from these discoveries.

Short-Latency Escapes Behavior in goldfish: Like many animals, fish respond to sudden aversive stimuli with a startle responseda

quick, decisive maneuver that orients the animal away from a potentially threatening stimulus (Eaton & Emberley, 1991; Hale et al., 2016). An aversive visual, auditory, or tactile stimulus evokes a short-latency escape maneuver, typically comprised of a sharp Cshaped turn followed by a rapid acceleration away from the threat (Eaton & Emberley, 1991). This behavior was initially well-characterized in adult goldfish, and early work in goldfish identified one of the neurons responsible for driving it: the Mauthner cell (M-cell). Identifying premotor neurons in goldfish: In normal fish, there is one Mauthner cell on each side of the hindbrain. This specialized reticulospinal neuron receives multisensory input onto its prominent ventral and lateral dendrites, and projects directly to primary motor neurons in the contralateral spinal cord (Korn & Faber, 2005; Zottoli & Faber, 2000). Electrophysiological recordings in goldfish demonstrated that the M-cell fires a single action potential in response to an aversive stimulus, which drives contraction of the body and tail muscles on the contralateral side (Fetcho & Faber, 1988; Prugh et al., 1982)devoking the short-latency turn away from the threat. However, following M-cell ablation, goldfish could still generate some shortlatency escapes (Eaton et al., 1982), indicating that other neurons must also be involved. Further, anatomical studies suggested additional candidates: two M-cell segmental homologs, MiD2 and MiD3 (Lee & Eaton, 1991; Lee, Eaton, & Zottoli, 1993). Identifying premotor neurons in zebrafish: Like goldfish, larval zebrafish execute short-latency escapes away from threatening stimuli (Figure 46.2A). However, larval zebrafish provided better in-vivo accessibility than goldfish (in terms of size and optical transparency) to facilitate further progress in our understanding of escape circuitry. Using backfills with dextran-conjugated calcium indicators to monitor neuronal activity in larval zebrafish, O’Malley et al. (1996) demonstrated that the M-cell and its homologs (MiD2 and MiD3) are active during escapes in response to tactile stimuli to the head, whereas the M-cell alone is active during escapes in response to tail stimulation. This supported a prediction made by Foreman and Eaton (1993) that additional reticulospinal neurons participate along with the M-cell in driving stronger head-evoked escapes (vs. weaker, M-cell-dependent tail-evoked escapes). Further, by refining a non-invasive method for laser ablation of single neurons, Liu and Fetcho (1999) observed that short-latency escapes to head stimuli were robust to M-cell loss alone but were abolished by ablation of the M-cell along with MiD2 and MiD3 reticulospinal neuronsdconsistent with the original hypothesis. Interneuronsdaction selection: The M-cell and its homologs provide the shortest synaptic pathway from an aversive sensory input to an escape behavior, but

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FIGURE 46.2 Early advances in the neural basis of short-latency escapes in larval zebrafish. (A) Example of an escape response to a unilateral water pulse delivered to the head. The single-asterisk marks the onset of the response; the double-asterisk marks the frame of the maximal C-bend. Images were collected at 1000 frames/s, and every third frame is shown. Modified from Liu and Fetcho (1999). (B) (i) Top left: Dorsal view of a larval zebrafish (4 dpf) and the kinematics of escape responses to head and tail stimuli, each compiled from frames captured at 1000 frames/s. Note the larger bend in response to a head stimulus. Scale bar ¼ 1 mm. Top right: Schema showing experimental preparation used for whole-cell recordings from the spinal cord, with simultaneous extracellular recordings of tail motor activity. Bottom: DIC (left) and fluorescent (middle) views of a MiP motor neuron in the spinal cord, targeted for whole-cell recording and filled with dye. White arrows mark the MiP axon. Scale bars ¼ 20 mm (right) DIC view of extracellular ventral root recording. (ii) Whole-cell recordings from a CaP motor neuron during electrical stimuli of increasing intensity, applied to the head or tail. Simultaneous recordings from the ventral root (VR) are also shown. Black arrows mark truncated stimulation artifacts. Modified from Bhatt et al. (2007). (C) Calcium activity from an array of CiD interneurons in the spinal cord during escapes elicited by head versus tail stimuli. The y axis is the ratio of calcium green to Alexa Fluor 647 divided by the mean ratio from 10 frames collected prestimulus. The escape movement occurs during the gap in the plot, when the cells move briefly out of the focal plane. Note that head and tail stimuli evoke responses in largely the same set of CiDs, consistent with changes in response amplitude being mediated by changes in the magnitude of interneuron activitydrather than the addition of more active neurons. Modified from Bhatt et al. (2007). (D) Summary of shortlatency escape circuitry in the zebrafish hindbrain. Modified from Hale et al. (2016).

researchers had reason to suspect that other neuronal populations optimize the functionality of the escape and further shape motor output. For example, the M-cell fires a single action potential, an all-or-none response (Nissanov, Eaton, & Didomenico, 1990)dbut the details of the motor output (though relatively stereotyped) vary based on the parameters of the stimulus. Further, early calcium imaging experiments revealed the influence of inhibitory networks in shaping M-cell

activation (Takahashi et al., 2002). In addition, calcium imaging in zebrafish spinal cord indicated that both primary and secondary motor neurons in the tail respond strongly during escapes (Fetcho & O’Malley, 1995)dalthough the M-cell only provides direct input to the primary motoneurons. Thus, interneurons must be involved, at a minimum, in modulating motor output and distributing motor commands. Once again, neuronal candidates were identified based on

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morphology (Hale, Ritter, & Fetcho, 2001) and backfilled with calcium indicator and imaged during behavior (Ritter et al., 2001). Two spinal interneuron subtypes (the ipsilateral descending CiD and the contralateral descending MCoD) were investigated, and Ritter et al. (2001) discovered that their activity is behaviorspecific: CiDs are active during escapes but not spontaneous swims, and MCoDs are active during spontaneous swims but not escapes. This study provided evidence for an additional participant in the escape circuit, but it also demonstrated that different interneuronal populations can be deployed to generate distinct behaviors. Following the development of a fictive swimming preparation in paralyzed zebrafish (Masino & Fetcho, 2005), researchers used whole-cell patch-clamp recordings (shown for motor neurons in Figure 46.2B) to show that individual CiDs (active during escapes) increase their firing rate during stronger escape maneuvers (Bhatt et al., 2007). In the same study, simultaneous calcium imaging from multiple CiDs showed that stronger escapes did not recruit additional CiDs from the population (Figure 46.2C). These experiments, taken together, are consistent with increased motor output strength carried out by modulation of active interneurons’ firing rates, rather than changes in the size of the active population. Interneuronsdvisual stimulus selectivity: Since these early studies, the widespread adoption of classical approaches (such as electrophysiology) and the advent of transgenic methodologies for use in larval zebrafish (Higashijima et al., 2000)dwith the subsequent explosion in the number and variety of available transgenic linesdhas spurred further progress in our understanding of the neural circuitry underlying short-latency escapes. For example, the use of genetically expressed calcium indicators enabled deeper investigations of the sensory side of the escape behaviordincluding the responses of tactile sensory neurons (Higashijima et al., 2003) and of the visual circuits involved in generating an escape response to looming stimuli (Dunn, Gebhardt et al., 2016). Looming stimuli elicit C-bend escapes that are abolished in mutants lacking retinal ganglion cells and impaired in larvae with unilateral ablations of tectal neuropil (Temizer et al., 2015). Twophoton calcium imaging in restrained larvae identified looming-specific regions of tectum that might participate in these escapes (Dunn, Gebhardt et al., 2016; Temizer et al., 2015). A study from Yao, Li et al. (2016) used an impressive variety of approachesdincluding electrophysiology, pharmacology, and optogeneticsd to reveal how dopaminergic inputs can act (via modulation of inhibitory glycinergic interneurons) to increase the specificity of visually evoked escapes by facilitating M-cell responses to looming stimuli and suppressing escape responses to other visual stimuli.

Interneuronsdauditory/vestibular and tactile stimulus selectivity: Other studies have investigated the role of hindbrain interneurons in ensuring optimal function of the escape circuit. For example, spiral fiber neurons receive contralateral sensory inputs and wrap their axons around the axon hillock of the contralateral Mcell (Lacoste, Schoppik et al., 2015), where they form excitatory electrical and chemical synapses (Koyama et al., 2011). Based on studies of mutant fish lacking the hindbrain commissure formed by their axons (Lorent, Liu, Fetcho, & Granato, 2001), spiral fiber neurons have long been thought to play a role in short-latency escape circuitry. Lacoste, Schoppik et al. (2015) proposed that this indirect interneuron pathway ensures that brief, weak stimuli do not evoke strong short-latency escapesdconsistent with their optogenetic experiments demonstrating that coincident activation of spiral fiber neurons enhances M-cell responses (and short-latency escape behavior) to weak stimuli or sensory noise. Thus, under ethological conditions, weak dendritic inputs will only elicit an M-cell response if they also excite the spiral fiber neurons (and persist long enough for the direct and indirect excitation to overlap in time). Interneuronsdlaterality: Koyama et al. (2016) investigated another example of an interneuron motif that optimizes escape circuit function, focusing on feedforward inhibitory neurons located near the M-cell. These neurons receive ipsilateral sensory input and project to both the ipsilateral and contralateral M-cells (Korn & Faber, 1975)dthough, importantly, each inhibits the contralateral M-cell more strongly (Koyama et al., 2016). In a study combining electrophysiology, calcium imaging, modeling, and behavior, Koyama et al. (2016) showed how these feedforward inhibitory neurons enhance short-latency escapes by ensuring a quick, lateralized response. Reciprocal inhibition of M-cells (and other feedforward inhibitory neurons) ensures that left and right sensory inputs compete to ultimately produce a strong movement away from a threat, even in the presence of ambiguous stimuli. This work provides a circuit-level account of how a simple, left/right decision is implemented in the brain (see also (Shimazaki, Tanimoto, Oda, & Higashijima, 2018) for a recent careful look at another inhibitory control mechanism in the Mcell network). Circuit development: Because we understand something about the various interneurons participating in escapes, this circuit was used as a test case to determine if hindbrain interneurons are recruited into specific circuits in an orderly way, based on their time of differentiation and positioning during early development. In larval zebrafish, hindbrain interneurons are organized into stripes according to their expression of specific transcription factors (Kinkhabwala, Riley et al., 2011). The neurons in each stripe share the same neurotransmitter

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Major Avenues of Investigation in Zebrafish With a Couple of Case Studies

identity and gross axonal morphology. Further, neurons are organized within a stripe according to relative age, with older neurons typically located in the most ventral portion of the stripe. Importantly, the dorsoventral position also correlates with neuronal excitability, suggesting that a neuron’s position in a transcription factor stripe might predict its functional role in a motor circuit. Conversely, if you understand the functional role of a specific interneuron, you should be able to predict its locationdwhich stripe it inhabits, as well as its dorsoventral position. Indeed, Koyama et al. (2011) used patch-clamp recordings and cell morphology to confirm this hypothesis for the interneuronal components of the M-cell escape circuit, including the spiral fiber neurons and feedforward inhibitory neurons. Conclusion: Short-latency escape behavior in larval zebrafish has been used to establish basic principles of sensorimotor control, to reveal interneuron circuit motifs that support optimal behavioral output, including one used to implement two-alternative decisions, and to facilitate our understanding of how circuits recruit specific neurons during development (circuit summary in Figure 46.2D). Further, the pursuit of these objectives propelled many of the methodological innovations that make larval zebrafish such an attractive model for neurobiological study.

Prey Capture Behavior: Soon after they acquire the ability to swim, larval zebrafish begin foraging for food and pursuing small prey, such as paramecia (Muto & Kawakami, 2013). Like short-latency escapes, prey capture maneuvers involve both sensory and motor processingdto select and transform appropriate sensory input into patterned muscle activity. An episode of prey capture behavior includes a series of slow swims and smallangle turns by which the larva approaches and re-orients itself relative to the prey, followed by a strike to engulf the prey (Budick & O’Malley, 2000; Borla et al., 2002; Patterson et al., 2013). Using a closed-loop virtual reality system to simulate prey capture in restrained larvae, researchers showed that the sequence of orienting turns (J-turns) is always preceded by binocular convergence of the eyesdnot to direct the fish’s gaze toward the prey, but to create an area of binocular overlap in front of the fish that facilitates prey tracking and capture (Bianco et al., 2011; Trivedi & Bollmann, 2013). In addition, studies in juvenile and adult zebrafish indicate that prey capture behaviors become more flexible and effective during development, associated with a transition from separate J-turns and approach swims to a single complex maneuver merging orientation and approach (Westphal & O’Malley, 2013).

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Identifying premotor neurons: Early studies found that M-cell ablations did not affect prey capture (Borla et al., 2002), so researchers began investigating other reticulospinal neurons that might be involved in generating prey capture maneuvers. Because the behavior relies heavily on visiondand laser ablation of the retinotectal neuropil causes a prey capture deficit (Gahtan et al., 2005)dcandidate reticulospinal neurons would need to receive visual input (directly or indirectly) from the tectum. In zebrafish, the nucleus of the medial longitudinal fasciculus (nMLF) includes two neurons (per side: MeLr and MeLc) that both extend dendrites into the deep output layers of the optic tectum and project to the spinal cord (Gahtan et al., 2005; Sankrithi & O’Malley, 2010). As assessed by a feeding assay, laser ablation of MeLr and MeLc decreases the number of successful prey captures without disrupting other motor behaviors. Further, a combined unilateral ablation strategy provided evidence that the tectum and the nMLF are part of the same pathway in the prey capture circuit (Gahtan et al., 2005). Thus, detection of visual prey stimuli by the optic tectum likely activates neurons in the nMLF to initiate or sustain prey capture maneuvers. Indeed, optogenetic activation of the anterior tectum can initiate J-turns (Fajardo, Zhu, & Friedrich, 2013). Consistent with these results, calcium imaging in freely swimming larvae indicates that anterior tectal activity precedes prey capture maneuvers (Muto, Ohkura, Abe, Nakai, & Kawakami, 2013), and quantitative analyses of tectal activity in restrained fish indicate that some neurons contain information about both sensory input and motor output (Bianco & Engert, 2015). Sensory processingdsize selectivity: In executing prey capture behaviors, it is important that zebrafish correctly identify potential prey. Larvae must discriminate between a small moving object (which might be suitable prey and should be approached) and a large moving object (which might be a predator and should be avoided). Thus, there should be some mechanism in the sensory circuitry for discriminating between small and large visual stimuli. Using genetically expressed calcium indicators, researchers discovered a region in the deep layers of optic tectum containing projection neurons that are more responsive to small stimuli than to large stimuli (Del Bene, Wyart et al., 2010). Using pharmacology, laser ablations, and transgenic strategies for blocking synaptic transmission in specific neuronal populations, researchers concluded that tectal inhibitory interneurons are necessary for establishing size filtering in deep layers of tectumdand that interfering with these microcircuits eliminates those projection neurons’ size tuning (Barker & Baier, 2013). However, it is worth noting that other studies using calcium imaging have found evidence for prey stimulus selectivity in the axons of some retinal ganglion cells (Semmelhack et al., 2014),

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46. Zebrafish as a Model for Revealing the Neuronal Basis of Behavior

FIGURE 46.3 Advances in the neural basis of prey capture behavior in larval zebrafish. (A) Example of pursuit and capture of a paramecium by a zebrafish larva. Time projection is shown in the upper left panel, illustrating the travel paths of individual paramecia during 3.3s of imaging (200 frames total, imaged at 60Hz). Remaining panels show a single prey capture sequence (3s total), with frames chosen to highlight sequence components: approach swims (0e0.83s), pursuit swims (1.44e2.56s), and the final capture swim (2.56e3.0s). Dashed arrows indicate the heading of the paramecium, and solid arrows indicate the heading of the larva. Note that as the zebrafish pursues its prey, it makes orienting turns that bring its heading into alignment with the path of the paramecium. Adapted from Gahtan et al. (2005). (B) Prey capture relies on visual processing in the optic tectum. (i) Laser ablations were used to eliminate the retinotectal visual pathway. (ii) Prey capture behavior was evaluated by quantifying the fraction of available paramecia remaining over the course of 5h total feeding time. Note that darkness severely disrupts prey capture, and tectal lesions also cause dramatic impairment. (iii) Blind lakritz mutants normally cannot hunt, providing further evidence for the importance of visual input. Adapted from Gahtan et al. (2005). (C) Deep layers of the optic tectum neuropil exhibit selectivity for small, prey-like stimuli. (i) Regions of interest highlight superficial (orange) and deep (green) layers of tectal neuropil. (ii-iv) Calcium responses of tectal neuropil to three types of visual stimuli. (v) Ratios of maximum responses in deep and superficial tectal neuropil layers to bars of increasing width. (vi-viii) Average maximum responses of deep and superficial neuropil layers to three types of visual stimuli. Note that deep layers respond less than superficial layers to full-screen flashes and large bars, consistent with selectivity for smaller stimuli. Adapted from Del Bene et al. (2010). D) Feeding state affects visual processing in the tectum. (i) Schematic diagram of the zebrafish retinotectal pathway. (ii) Calcium indicator expression in the optic tectum was used to observe neuronal activity to prey-like stimuli. (iii) Examples of tectal neuron calcium signals in response to visual stimuli of different sizes. (iv) Comparison of the cumulative percentages of weighted mean response (WMR) angles for tectal neurons in starved and fed larvae. Note that starved larvae have more tectal neurons tuned for small, prey-like stimuli, consistent with a state-dependent modulation of visual processing favoring prey capture behaviors in starved larvae. Modified from Filosa et al. (2016).

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The Future for Zebrafish in Studies of the Neuronal Basis of Behavior

raising the possibility that some stimulus filtering may occur before processing in the tectum. Stimulus selectivity and behavioral state: Filosa, Barker, Dal Maschio, and Baier (2016) conducted an important study of how behavioral state might influence sensory processing (Figure 46.3D). They first observed that an ambiguous visual stimulus (intermediate in size) can evoke either prey capture or avoidance maneuvers, but starving the larvae increases the likelihood that they treat the stimulus as prey (and not a threat). Starved fish also have lower cortisol levels, suggesting the involvement of the hypothalamic-pituitary-interrenal (HPI) axisdpreviously shown to be involved in the regulation of food intake in fish (Bernier & Peter, 2001). Because the HPI axis can modulate serotonin levels (Fox & Lowry, 2013), they pursued a role for serotonin and found that the activity of serotonergic neurons in the raphe nucleus is elevated in starved fish (Filosa et al., 2016). Moreover, the increase in avoidance (in response to ambiguous stimuli) caused by prefeeding is abolished by selective serotonin reuptake inhibitors (SSRIs), and starved fish lacking serotonergic neurons (killed via nitroreductase ablation) behave like prefed fish. These results, taken together, are consistent with a model, wherein satiety activates the HPI axis, thereby decreasing serotonin release from neurons in the raphe nucleus. But what mediates the effect on behavior? The same study also used calcium imaging to observe the effect of starvation on size selectivity in the optic tectum, revealing that starvation induces a serotonin-mediated shift in the population-level tuning of interneurons and periventricular neuronsdwhich may bias motor output in favor of prey capture in starved larvae. Thus, behavioral state modulates how sensory stimuli are represented in the brain and how they are transformed into motor output. ‘Conclusion: Prey capture behavior has provided a window into the neural basis of ethologically relevant stimulus selection, motor sequence generation, and state-dependent sensorimotor processing. These studies capitalized on many of the methodological approaches first pioneered to study the escape circuitry, including electrophysiology, laser ablations, and imaging with genetically encoded calcium indicators. Further, because prey capture (unlike escapes) comprises a sequence of behaviors modified by the stimulus in real time, these studies have innovated in their use of closed-loop virtual reality arenas for calcium imaging in restrained zebrafish. Researchers investigating this behavior can now move to take advantage of increasingly sophisticated tools for the dissection of neuronal connectivity in larval zebrafish that subserve it.

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The Future for Zebrafish in Studies of the Neuronal Basis of Behavior In spite of the remarkable advances that make zebrafish such a powerful vertebrate model for revealing the neuronal basis of behavior, the glimpses we have provided are just the beginning. The number of labs using fish is growing rapidly, as are the tools that set zebrafish apart from other models. This last section looks to things on the horizon that ensure that the fish model is not going away any time soon as a path to revealing how vertebrate brains work.

EM Connectivity on Order Several recent reports make strong use of electron microscopy (EM) to produce serial EM sectioning of the larval zebrafish brain, to reconstruct the connectome of the olfactory bulb, and to reconstruct the connectivity of neurons in the oculomotor system after a functional study by calcium imaging. These point to a fruitful interface between synaptic level EM morphology and functional studies of circuits in zebrafish (Hildebrand, Cicconet et al., 2017; Vishwanathan et al., 2017; Wanner et al., 2016). While a connectome of the entire larval nervous system remains a challenge, mostly because of limitations in automated tracing, it is less wildly out of reach than in other, larger vertebrate brains. The ability to reveal connectivity within regions is critical for constraining circuit models, so we can expect many regional reconstructions in zebrafish to become available soon. Even more fruitful in the short term, however, will be the ability to quickly reconstruct the projections of specific functionally identified neurons, to tie function to exact structural connectivity. As this becomes routine, we can expect that the future will offer the ability to compare the connectivity of identified neurons between animals with different experiences, such as exposure to a specific learning paradigm. This will provide an unprecedented view of how experience reshapes connectivity in neural circuits. If the EM processing and reconstruction becomes more automated and commercialized, it may be possible to simply send tissue and obtain connectivity data from individual animals, much like genomic data.

Transynaptic Mapping Viruses that cross synaptic connections offer a potential path to revealing the set of inputs to a class of neurons without the need for an EM reconstruction. Rabies virus-based approaches modified to jump only one synapse already provide information in mammals

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about the set of neurons connected directly to a target population (Kim, Jacobs, Ito-Cole, & Callaway, 2016). There remain limitations due to toxicity of the viruses, as well as lingering concerns about whether there is leakage of the labeling to adjacent but unconnected neurons, but the tools are improving substantially in mammals. Similar efforts using vesicular stomatitis virus (rVSV) are underway in zebrafish as part of an effort to extend the range of species available for viral tracing, but these efforts are still in progress (Beier, Mundell, Pan, & Cepko, 2016). As these reach fruition, they will provide a direct path to in vivo tracing in zebrafish. An ability to transynaptically express activity indicators or optogenetic constructs, when combined with the fish’s transparency, will provide functional and behavioral information for specific components with known connectivity.

The Physiological Holy GraildGenetically Encoded Voltage Imaging The calcium indicators used to monitor neuronal activity in vivo are only indirectly coupled to the transmembrane electrical activity that actually contributes to behavior. Efforts to optically monitor membrane potential have a long history, with challenges centering around small signals and phototoxicity. Both issues are a result of the need for the indicators to be localized in the cell membrane, a small and vulnerable region of the neuron, unlike the cytosol where calcium indicators typically reside. Still, there are many efforts underway to improve genetically encoded indicators of voltage, with entire families of them now in existence (Ouzounov, Wang et al., 2017). Improvements are coming quickly, and single-cell resolution in vivo has been achieved in some species (Xu, Zou, & Cohen, 2017). While efforts in zebrafish have not yet been resoundingly successful (Kibat, Krishnan, Ramaswamy, Baker, & Jesuthasan, 2016), there is little doubt that the optical access of the model will lead to a burst of voltage imaging if reliable, easy-to-use voltage indicators become available, which seems imminent (Piatkevich et al., 2018; Adam et al., 2019; Abdelfattah et al., 2019). Still, the opposition of membranes of adjacent neurons will make separation of signals from adjacent cells difficult, and deeper imaging with the superior optical sectioning of multiphoton microscopy may not work for voltage indicatorsd because of the limited number of photons available from restricted regions of the membrane imaged at the kilohertz rates needed to detect action potentials that last a millisecond or so. The voltage indicators will likely prove most useful for sparsely labeled animals.

Direct to Circuits in Zebrafish One promise of zebrafish that builds upon prior optical approaches to circuits involves a tool that is still under development but could be revolutionary when combined with the optical access to the larval brain. There are now several new approaches to activate genes using light with cellular specificity in culture, and hints of potential for in-vivo application (e.g., (Polstein & Gersbach, 2015)). This ability could prove very powerful when combined with whole-brain calcium imaging, as in a recent combination of whole-brain imaging and laser ablation (Vladimirov, Wang et al., 2018). One could imagine utilizing whole-brain calcium imaging to identify “interesting” candidate neurons, which participate in a particular behavior, and then use light-induced gene expression to cause those candidate neurons to express a genetically encoded label or control element (e.g., optogenetic constructs to turn them on or off, designer ligands for experiments in freely swimming animals, electron-dense markers for EM, fluorescent proteins, or voltage indicators). These candidate neurons may be ones that become active during the behavior or could even be neurons that change their activity after learning. The cells’ structure could be revealed by the light-activated expression of membrane-targeted proteins. Inducing expression of optogenetic constructs would allow perturbations in neuronal activity, to test hypotheses about those neurons’ specific role in behavior. Light induction of EM markers would even allow later reconstruction of those neurons’ connectivity throughout the brain. This would offer the possibility of moving more directly from a specific behavior to formulating and testing predictions about the neural basis of that behavior(Dal Maschio, Donovan, Helmbrecht, & Baier, 2017).

Circuits and Behavior From Embryo to Adult Most of the studies of brain and behavior in zebrafish focus on larval fish, typically less than 3 weeks of age. This is a practical choice, as the larvae are small and translucent, so optical tools and targeted recording are easy. The fish must survive on their own after hatchingdmeaning that they execute a wide variety of behaviors and simple forms of learning - so the larvae have a major place in understanding brains and behavior. A strong understanding of even simple behaviors, such as movement, visual-motor orientation, feeding, and simple forms of learning still elude us on the scale of the whole nervous system. However, while larval behavior is just as consequential as any other behavior (dead larvae have no

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TABLE 46.1

Advantages and disadvantages of zebrafish by comparison to other model systems for the study of the neural basis of behavior. C. elegans

D. melanogaster

D. rerio (larvae)

M. musculus

Number of neurons

302

w135,000

w130,000

w75 million

Body length

1 mm

3 mm

4 mm

7.5e10 cm

Time to sexual maturity

4 days

1 week

12 weeks

10 weeks

Example behaviors studied at the level of neural circuits

Locomotion, Taxis, foraging, mating

Flight, crawling (larvae), foraging, mating, Aggression, visual-motor reflexes

Swimming, escape, visual-motor reflexes, prey capture, phototaxis,

Aggression, mating, fear conditioning, spatial learning, contextual memory, foraging, Maternal care, communication

Optical accessibility

Transparent

Surgical window

Transparent

Surgical window

Electrophysiology

Difficult

Difficult

Feasible

OK

Unique advantages

Full connectome

Defined genetic access to small groups of individually identifiable neurons

Best vertebrate model for optical microscopy.

Most similar to humans. Synaptic tracing viruses. Has a cortex.

Disadvantages

Less behavioral complexity; graded neurons

Opaque cuticle; neuronal computations take place in dendrites

Limited capacity for learning and memory in the larval stage.

Has a cortex. Large scale and high complexity of brain.

offspring!), the fish brain and behavior does change as the fish grows to reach sexual maturity at about 3 months of age. For example, a very small cerebellum in larvae grows to a large and recognizably vertebrate cerebellum in adults. With growth and adulthood, comes more complex social interactions, seemingly more sophisticated motor control and learning, and new behaviors, such as mating. The advantages of the transparency are lost, however, which would seem to undermine the utility of the model later in life. Recent innovations will likely circumvent that. The development of longer wavelength three-photon microscopy and the attendant ability of long wavelengths to more easily penetrate opaque tissue allows deeper imaging into previously inaccessible regions of adult brains (Ouzounov, Wang et al., 2017). The depth of imaging possible with three-photon microscopy (on the order of 1.5 mm) matches the thickness of an adult zebrafish brain. The combination of three-photon microscopy and some adaptive optical tools with the pigmentless Casper fish lines will likely allow optical access anywhere in the intact living adult zebrafish brain. This would make zebrafish the first vertebrate where brain function in behavior can be studied in the same animal from embryo to adult, opening up many questions of how brain circuits and behavior change with maturity and experience, and eventually degenerate with age. Table 46.1.

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C H A P T E R

47 Zebrafish as a Model to Understand Human Genetic Diseases Jennifer B. Phillips, Monte Westerfield Institute of Neuroscience, University of Oregon, Eugene, OR, United States of America

Zebrafish Versus Human: How Similar are They? The completion of the human genome project and, more recently, the shift to Whole Exome Sequencing (WES) and Whole Genome Sequencing (WGS) has further accelerated the number of identified human disease genes. At the time of this writing, the Online Mendelian Inheritance in Man (OMIM) database lists over 5000 diseases for which the clinical phenotypes have a known molecular basis, with more confirmed disease genes being identified regularly. This wealth of new data accessible to clinicians, though, would not have been possible without the contributions of model organisms, such as the zebrafish in identifying and validating disease loci. Since their original use as a vertebrate model (Streisinger, Walker, Dower, Knauber, & Singer, 1981), zebrafish have been illuminating and informing our knowledge of human development, genetics, and disease. Throughout the late 20th century, the previously segregated fields of evolution, development, and genetics increasingly converged, united by the common thread of molecular biology. Despite the divergent evolutionary processes that have occurred in the teleost and human lineages over the past 450 million years, the ancestral vertebrate “operations manual” can be clearly identified in both organisms. Although the specifics of how a fertilized egg develops into a fish versus a human are obviously and importantly different, the patterns of cell movement, tissue differentiation, and organogenesis are strikingly conserved. The retained similarities extend even to the structure and function of their genomes. Analysis of the arrangements of genes on fish and human chromosomes reveals many regions where linear organization of the genes is preserved

The Zebrafish in Biomedical Research https://doi.org/10.1016/B978-0-12-812431-4.00047-6

(Catchen et al., 2011), whereas analysis of the genes themselves highlight that zebrafish orthologs for 70% of all human genes, and a striking 84% of human disease genes, have been identified (Howe et al., 2013). This conservation indicates a close adherence to the development, structure, and function of our common vertebrate ancestor and strengthens the rationale for characterizing zebrafish, despite their divergent freshwater adaptations, as a legitimate tool for understanding the human genetic disease. Moreover, even significant anatomical differences are often surmountable in terms of modeling human genetic disorders. For example, in humans, hematopoietic stem cells in the bone marrow give rise to the different classes of blood cells, several of which can be affected by genetic mutations leading to various diseases. Zebrafish lack bone marrow, but instead have a population of hematopoietic stem cells in the kidney, which gives rise to all of the analogous blood cells, and thus, have still contributed valuable insights into the mechanisms underlying blood cancers and other disorders (reviewed in Choudhuri, Fast, & Zon, 2017; Rissone & Burgess, 2018). Similarly, although the zebrafish telencephalon is rudimentary compared to the mammalian cerebrum, zebrafish neuronal connections and neurological functions, including learning, memory, social behavior, and anxiety, are genetically and physiologically influenced in ways similar to that of humans (Kalueff, Stewart, & Gerlai, 2014; Newman, Ebrahimie, & Lardelli, 2014; Matsui, 2017; Velkey, Boles, Betts, Kay, Henenlotter, Wiens, 2019). Another potentially influential difference in disease pathology between zebrafish and humans is the regenerative capacity of many zebrafish tissues, which in principle could compensate for or mask defects precipitated by genetically regulated cell loss. This concern is

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© 2020 Elsevier Inc. All rights reserved.

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47. Zebrafish as a Model to Understand Human Genetic Diseases

largely resolved by the observation that numerous zebrafish models of degenerative disease have been characterized (Matsui & Takahashi, 2017; Sher, 2017; Sa´nchez, Azcona, & Paisa´n-Ruiz, 2018; Dona et al., 2018; Mishra et al., 2019) and further allayed by the availability of mutant backgrounds inhibiting the normal regeneration process (Pei et al., 2016) that could be used to unmask more completely the degenerative potential of a human disease gene under study.

An Expanding Toolkit for Generating Zebrafish Models of Human Genetic Disease Mutagenesis Screens As with many other genetic model organisms, parallel and often reciprocal progress in genetic analysis and genomics of the zebrafish has increasingly led to more tractability in the ways in which gene function can be analyzed and manipulated. Initial large-scale forward genetics screens were conducted by inducing random mutagenesis via chemical exposure or viral insertion, (Mullins, Hammerschmidt, Haffter, & Nu¨ssleinVolhard, 1994; Haffter et al., 1996; Amsterdam, Nissen, Sun, Swindell, Farrington, Hopkins, 2004), screening the progeny at embryonic or larval stages for phenotypes of interest, and characterizing the developmental or functional anomaly while working to map the genetic cause of the defect to a precise locus within the genome. Concomitant efforts to sequence the zebrafish genome provided an increasing catalog of genetic markers on each of the 25 zebrafish chromosomes to assist in the mapping efforts (Ransom & Zon, 1999; Shimoda et al., 1999; Kelly et al., 2000). The efforts of the global zebrafish community over the past 40 years have isolated and characterized the recessive loss of function mutations in scores of genes linked to inherited disease in humans. Broadly, forward genetics screens center on observing phenotypic abnormalities in morphology and/or behavior. Evaluation of large pools of larvae from mutagenized parents for defects in the development of craniofacial skeletal elements, for example, led to the discovery of several alleles of the zebrafish mutation jellyfish. These animals exhibit profound and ultimately lethal malformations in the skeletal elements of the head and trunk. Further research to identify the causative gene revealed mutations in sox9a, a transcription factor with a varied and important function in cell-fate specification during embryonic development (Yan et al., 2002). The zebrafish phenotype of sox9a mutants bears a striking similarity to the clinical symptoms observed in Campomelic dysplasia, which in

humans is caused by mutations in SOX9 (OMIM #114290). The data gleaned from characterizing these mutants led not only to new insights on how cartilage forming cells organize during skeletal development, but further provided important information about a severe human genetic disorder. Behavioral screens can detect anomalous phenotypes in embryos or larvae with mild or undetectable changes to their morphology. A screen for mutants with diminished visual function, assessed by the ability to track moving stripes known as the optokinetic response, netted a large number of distinct lines that shared this common behavioral defect despite a wide range of underlying genetic causes. One such mutant initially termed “no optokinetic response a” (noa) (Brockerhoff, Hurley, Janssen-Bienhold, Neuhauss, Driever, & Dowling, 1995), exhibited impaired visual function despite a histologically normal retina. Additionally noted phenotypes included reduced physical activity, poor feeding behavior, and early larval death. Using positional cloning methods, Brockerhoff and colleagues identified dlat as the affected gene in these mutants (Taylor, Hurley, Van Epps, & Brockerhoff, 2004). Dihydrolipoamide S-acetyltransferase (Dlat) is a subunit in the Pyruvate dehydrogenase (PDH) complex, a crucial component in the metabolic process that produces energy from carbohydrates to power a number of neurological functions. Defects in the PDH complex in humans cause neurological defects, failure to thrive, and premature death. The advent of an animal model in which to study defects in the pyruvate dehydrogenase pathway not only provided new information on the molecular basis of the disorder, but also an opportunity to explore ways to treat it. When zebrafish dlat mutants were provided with a ketogenic diet, their visual and neurological symptoms improved along with their survival. Up to this point, some PDH patients had been advised to consume a ketogenic diet with low carbohydrate, high fatty acid content, but the clinical evidence for this recommendation was limited. Analyzing the connection between diet modifications and metabolic function in the dlat mutants provided strong validation for this approach in the clinic, and more rigorous clinical trials have since demonstrated positive outcomes for PDH patients following this protocol (Sofou, Dahlin, Hallbo¨o¨k, Lindefeldt, Viggedal, Darin, 2017). Far from obsolete, mutagenesis screens are still used to good effect, particular in the discovery of genes important for postdevelopmental or more specific behavioral functions (Pelegri & Mullins, 2016; Henke et al., 2017; Gerlai et al., 2017; Kegel et al., 2019; and see Table 47.1).

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An Expanding Toolkit for Generating Zebrafish Models of Human Genetic Disease

TABLE 47.1

Data samples provided by the Zebrafish Information Network (ZFIN), taken at three intervals between September 2017 and April 2019 reveal the growing number of published genetic disease models and the changing frequencies with which various genetic models are being used.

Total zebrafish genetic disease models in published studies

2017

2018

2019

‘17-’19 increase

1334

1447

1582

þ19%

Forward mutagenesis screen approaches N-ethyl-N-nitrosourea

260

267

333

þ 28%

Viral insertion

102

119

124

þ 22%

Reverse Genetic approaches Morpholino

1217

1356

1396

þ 15%

TILLING

46

64

80

þ 74%

ZFN

29

30

40

þ 38%

TALENS

65

115

136

þ109%

CRISPR

55

187

247

þ349%

Morpholino Knockdown Although undeniably powerful, the genetic screens described above are perennially limited by the “luck of the draw,” especially because the nature of mutagenesis can be limited by areas within the genome that are more or less susceptible to disruption by chemical or insertional mutagenesis. As human genetic studies advanced through the late 20th century and more molecular identities were conferred on known hereditary diseases, the need for targeted disruption of disease genes in the zebrafish model was increasingly pressing. The methodology of using an injection of double-stranded RNA to knockdown candidate gene function that was tremendously powerful in fly and worm models proved to be ineffective in zebrafish (Zhao, Cao, Li, & Meng, 2001). Thus, the introduction of morpholino oligonucleotides (short, single-stranded base sequences arranged on a modified morpholine backbone to impart stability) as a means of temporarily targeting a chosen gene for disruption by blocking mRNA translation or pre-mRNA splicing (Nasevicius & Ekker, 2000; Draper et al., 2001) was heralded with justifiable enthusiasm by the zebrafish community. With this new tool in hand, “morphant” models of human disease burgeoned over the next decade. One significant benefit to human health was the ability to validate candidate disease genes quickly. Advances in the sensitivity and cost-effectiveness of

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genomic analysis in the clinical setting provided new potential answers to unsolved genetic disease riddles, but the high number of sequence variations in every human genome can reduce confidence in attributing symptoms to defects in a gene not previously identified as causative. The ability to deplete the function of the orthologous gene(s) in zebrafish and quickly validate or rule out candidates derived from sequence analysis, is a significant asset to genetic diagnosis. Morpholino knockdown of candidate genes has been used in numerous collaborations between basic researchers and clinicians to vet novel genes identified in sequence analysis of patients. Hanno Bolz and colleagues (Beck et al., 2014) detected lesions in the POC1 gene of individuals with severe symptoms indicative of defective primary cilia. Primary cilia are integral to the functions of numerous tissues throughout the body, including the retina, kidney, and brain. These cilia emanate from centrosome-like microtubule-organizing centers made up of dozens of proteins. Although many inherited ciliopathies are known to be caused by defects in the genes that encode these centrosomal proteins, there is also considerable redundancy in this system, to the extent that uncharacterized factors in this category cannot be automatically assumed to cause disease when mutated. Morpholino knockdown of poc1b in zebrafish produced defects in retinal and renal cilia, consistent with the patients’ symptoms. These data combined with the family genotyping and in silico modeling of the predicted changes to the POC1B protein were sufficient to diagnose the families genetically and categorize other truncating mutations in POC1B as likely pathogenic. In other cases, zebrafish morpholino models have been used to exclude certain alleles from pathogenic categorization. In another case investigated by Bolz & colleagues (Elsayed et al., 2015), siblings with genetically undiagnosed deafness were found to have a homozygous mutation in the ciliopathy gene, AHI1. Not only did these affected individuals show no sign of ciliopathy themselves, but other family members with normal hearing also carried the same homozygous mutation. The allele in question was predicted to produce a C-terminal truncation of the protein, but a known truncating mutation mapping downstream of the newly identified one had previously been linked to severe ciliopathy. Use of two morpholinos to 50 and 30 regions of the transcript showed distinct differences, with the earlier disruption causing widespread cilia dysfunction consistent with early truncating human alleles, whereas the downstream disruption showed no symptoms. This analysis underscores the importance of being able to validate pathogenicity so that accurate information

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47. Zebrafish as a Model to Understand Human Genetic Diseases

about disease-causing alleles can be provided to human geneticists and genetic counselors worldwide. Although the discoveries netted with morpholino knockdown were undeniably valuable in the pursuit of understanding human disease genes, one limitation is the relatively short-lived efficacy period of morpholinos, which, in most cases, restricted analysis of the morphant phenotype to the first week of development or less. Off-target effects, giving strong and deleterious phenotypes unrelated to the targeted specific gene can confound interpretation of the results. In some cases, differences have been noted between morphants and genetic mutants (Kok et al., 2015; Novodvorsky et al., 2015), although even this initially dispiriting finding is leading to a deeper understanding of gene regulation in zebrafish and how transcript-mediated genetic buffering can compensate for loss of function mutations (Rossi et al., 2015; Tuladhar et al., 2019). Overall, morpholinos presented the zebrafish community with early and fruitful opportunities to leverage the power of candidate gene approaches to modeling human disease. Morpholinos remain a valuable tool in analyzing human disease genes in zebrafish, particularly in clinical collaborations due to the speed with which loss of function results can be obtained (Table 47.1).

Targeting Induced Local Lesions in Genomes (TILLING) As morpholino knockdown efforts gained prominence in the zebrafish research community, a concerted effort to capitalize on the permanence of genetic mutation and to increase the library of zebrafish mutants was mounted, first in smaller collaborations (Moens, Donn, Wolf-Saxon, & Ma, 2008) and later by the Zebrafish Mutation Project undertaken by the Sanger Institute (Zebrafish Mutation Project (https://www.sanger.ac. uk/resources/zebrafish/zmp/). Which sought to create a loss of function alleles for every protein-coding gene in the zebrafish genome. As in the classic forward genetic screens, random mutagenesis was initiated by chemical mutagen exposure. But this time, instead of screening offspring for phenotypes of interest, collections of mutagenized genomes were screened for deleterious mutations in genes of interest by DNA sequencing. These TILLING mutants were subsequently cataloged and made available for study through the Zebrafish International Resource Center (ZIRC) in Eugene, Oregon, USA. At the time of this writing, the Zebrafish Mutation Project has isolated 37,624 alleles with putative loss of function point mutations, many of which are available for distribution from both the ZIRC and the European Zebrafish Resource

Center (EZRC) in Karlsruhe, Germany. Considering the overlap of disease genes in the respective genomes of human and zebrafish, this library of zebrafish mutants represents an important resource for modeling human disease. For example, careful analysis of several different Sanger alleles of the chromatin-modifying factor kdm2aa has illuminated a new genetic contributor to melanoma formation (Scahill et al., 2017), adding a valuable tool to the global research efforts addressing this challenging and often fatal form of skin cancer.

Gene Editing Zebrafish studies of human disease genes gained increasing international attention through the early 21st century, and optimization of the methods described above has reduced the inherent limitations on each to an impressive degree. The limits of modeling human genetic disease in zebrafish have been attenuated even further by the introduction of targeted gene editing that uses sequence-specific guided nucleases to find and create disruptions in a chosen genetic location. The first success at this targeted mutagenesis in zebrafish was obtained by generating customized zincfinger nucleases (ZFNs; Doyon et al., 2008; Foley et al., 2009). ZFNs are synthesized to recognize a specific DNA sequence and, via nuclease action, create breaks in the double helix. When DNA repair mechanisms are activated to re-join the broken ends of the DNA strand, small stretches of sequence can be randomly deleted or inserted, creating a shift in the reading frame of the genetic code. When introduced into zebrafish zygotes, these constructs create mutations that can be propagated throughout multiple cells of the developing embryo, including the germline cells, thus creating heritable changes in the genes of interest. The exciting potential of this new tool was tempered only by the labor-intensive nature of generating custom ZFNs, but parallel efforts to discover more effective and efficacious ways of altering selected regions of DNA soon provided alternatives. Shortly after the advent of ZFNs, a breakthrough harnessed Transcription Activator Like Effector Nucleases (TALENs), derived from a bacterial pathogen of plants, to edit zebrafish genes (Huang, Xiao, Zhou, Zhu, Lin, Zhang, 2011; Sander et al., 2011). This system consists of customizable DNA binding subunits (TALEs), each recognizing a specific nucleotide base that can be assembled in any order to recognize specific sequences. When these custom protein sequences are attached to a nuclease, such as Fok1, the whole assembly can create doublestranded breaks in the DNA at the binding site. Just on the heels of this methodological development, research to characterize Clustered Regularly

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Disease Models: What Genes do we Choose to Target?

Interspersed Short Palindromic Repeats (CRISPR), originally found in bacterial DNA and associated with CRISPR-associated (Cas) DNA cutting enzymes, led to the discovery that these short palindromic repeats could be used along with gene-specific guide RNA to bring a DNA-cutting enzyme to a selected position on the chromosome (Qi et al., 2013; Cong et al., 2013). Zebrafish researchers quickly jumped on this new technique (Chang et al., 2013; Hwang et al., 2013; Jao, Wente, & Chen, 2013), and currently, CRISPR/Cas9 system is the most widely used of the three gene-editing methods, largely due to the ease and low cost with which the constructs can be generated (Table 43.1). A rapidly expanding list of zebrafish mutants derived from CRISPR/Cas-generated disruption includes new models of atrial fibrillation (Ahlberg et al., 2018), congenital neutropenia (Pazhakh, Clark, Keightley, & Lieschke, 2017), osteoporosis (Zhang et al., 2017), structural heart and renal defects (Ta-Shma et al., 2017), scoliosis (Gao et al., 2017), thrombocytopenia (Marconi et al., 2019), and Wolf-Hirshhorn Syndrome (Yu et al., 2017), to name only a few. The ability to create frameshift mutations at precise locations within the genome conferred unprecedented power to the ongoing work to generate and learn from zebrafish models of human diseases. Along with the ability to “break” a gene, thereby disrupting function, these methods provide the opportunity to change the DNA code in more specific ways, using the event of a double-stranded DNA break to overwrite the native code. By providing a template strand of DNA containing both homologous sequences from the targeted region and customized new code to be incorporated, researchers now have the potential to change the gene sequence in even more refined ways, for example introducing an in-frame missense change to emulate a specific human allele with known or unknown disease-causing potential (Gopal et al., 2015; Armstrong, Liao, You, Lissouba, Chen, Drapeau, 2016; Boel et al., 2018; Prykhozhij et al., 2018;), or generating a reporter sequence (Hoshijima, Jurynec, & Grunwald, 2016; Kesavan, Chekuru, Machate, & Brand, 2017). With these technical advances, the further goal of using knock-in strategies to repair mutations without disrupting the reading frame, thus restoring the ability of the organism to make a wild-type protein product, is not far behind. The potential for exploring genetic variants of unknown significance (VUS) and for research into repairing genetic defects by overwriting them with better code has barely been tapped as of this writing, but as with the advent of every new tool described herein, the more broadly used the technique within the zebrafish community, the more potential there is for innovation and variation that will carry research forward.

623

Disease Models: What Genes do we Choose to Target? The gene-targeting tools described above, combined with a superior array of transgenic lines, vital dyes, and other methods of visualizing molecular physiological processes in action, have set up the zebrafish as an undeniably powerful system in which to explore any known or suspected human disease gene. Zebrafish models have been used both to study and to screen potential therapies for common genetic diseases, such as cancers (Anelli et al., 2017; Lu et al., 2017; Oppel et al., 2019; Tulotta, Stefanescu, Chen, Torraca, Meijer, Snaar-Jagalska, 2019), cardiovascular and kidney disorders (Minchin & Rawls, 2017; Chang et al., 2017; Schenk, Mu¨ller-Deile, Kinast, & Schiffer, 2017; van Rooijen et al., 2018; Zhao, Zhang, Sips, & MacRae, 2019), and cystic fibrosis (Zhang, Liu, & Chen, 2018). But although defects in human genes are responsible collectively for many millions of clinical cases, some genetic disorders are vanishingly rare. A disease affecting fewer than 200,000 people at any given time is defined as “rare” in the United States, but a growing number of genetic disorders affecting far fewer than that number, sometimes as few as a single patient, are reported. The struggle for equitable research funding for rare and extremely rare diseases far exceeds the scope of this chapter, but it is fair to say that clinicians, researchers, advocacy groups, and affected individuals alike are hopeful that the advancements that have made zebrafish a more tractable and versatile disease model for human afflictions will benefit the rare, as well as the common among them. To this end, collaborative organizations, such as the Undiagnosed Diseases Network (Chao, Liu, & Bellen, 2017; Wangler et al., 2017) have placed special emphasis not only on enhanced clinical workups and genomic analysis for patients with rare genetic disorders, but on the development of a high-throughput animal model core to explore the function of novel sequence variants implicated in unusual human genetic disorders. CRISPR/Cas9 zebrafish models of candidate genes implicated in these rare presentations are providing crucial validation of the genotypephenotype correlation required to make a genetic diagnosis (Ferreira et al., 2018; Burrage et al., 2019; Lahrouchi et al., 2019). Modeling human disease “on demand” is one way in which zebrafish is being harnessed at the interface of basic and clinical research, with data from human patients directly influencing the creation and analysis of their zebrafish proxies. With increasing frequency, the current flows in the other direction as well, as research in zebrafish disease models informs new clinical directions. The generation and ongoing analysis of

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zebrafish mutants has provided new therapeutic targets for the treatment of cancers (Guan et al., 2019) craniofacial and other disorders (Seda et al., 2019), hearing loss (He, Bao, & Li, 2017), Niemann-Pick disease (Tseng et al., 2018), and a range of other genetic disorders. The ease with which zebrafish can be subjected to panels of chemicals that might enhance or suppress an underlying genetic condition also feeds directly into the foundational research required before seeking approval for clinical trials. Beyond the broad array of published work, the widespread uptake of gene editing techniques, including work on knock-out and knock-in endpoints, has provided vital data on the advantages and challenges of tissue-specific methods for altering DNA sequences in individuals with genetic disorders. The reciprocal influence of zebrafish models on the human health issues they seek to illuminate and ameliorate is only beginning, and the potential impact that our small but powerful model organism can have on the vast field of human genetic disease is realized with each new insight that moves the entire community forward.

Acknowledgments The authors are grateful to Sabrina Toro and the ZFIN development team for providing statistics on curated zebrafish disease models. This work was supported by NIH Grants U54NS093793 and P01 HD22486.

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C H A P T E R

48 Zebrafish as a Model for Investigating AnimaleMicrobe Interactions Travis J. Wiles1, Karen J. Guillemin1,2 1

Institute of Molecular Biology, University of Oregon, Eugene, OR, United States of America;2Humans and the Microbiome Program, CIFAR, Toronto, ON, Canada

Introduction Over the past decade, a boom in DNA sequencing studies has revealed that the bodies of humans and other animals are inhabited by an amazing diversity of microscopic wildlife, including bacteria, archaea, viruses, and eukaryotic microorganisms, such as fungi and protozoa (Blaser et al., 2016; McFall-Ngai et al., 2013). These microbial consortia are now widely recognized to have far-reaching impacts on animal biology, from promoting normal growth and development to inciting infection and disease, making them the focus of intense biomedical research. However, unraveling the mechanisms by which resident microbes live and interact with each other, and their animal hosts remains incredibly challenging due to their intricate and multifarious nature. Animalemicrobe interactions play out over a range of temporal and spatial scales at various concealed locations across the host’s body, for example, deep within the intestinal tract, and involve numerous cell and tissue types. For understanding these complex interactions, multiple complementary experimental models are needed. The laboratory mouse is often employed because of its anatomic and genetic similarity to humans, and thus, it has yielded many important insights (Kostic, Howitt, & Garrett, 2013). However, the mouse’s relatively slow development, inaccessible internal tissues, and expensive husbandry make certain experimental approaches unfeasible with this model. Alternatively, highly amenable in vitro technologies have been developed that use cultured animal cells (typically human-derived) to generate so-called organson-chips, but the ability of these distilled experimental approaches to recapitulate higher-order interactions involving multiple organ systems, developmental

The Zebrafish in Biomedical Research https://doi.org/10.1016/B978-0-12-812431-4.00048-8

processes, or behavior is severely limited (Benam et al., 2015). Offering a balance between these approaches is the zebrafish, Danio rerio (Burns & Guillemin, 2017). In this chapter, we describe the attributes that make this popular aquarium fish an attractive model organism for studying animalemicrobe interactions and illustrate through several vignettes of cutting-edge research how it is propelling the field forward.

The Zebrafish: An Adaptable and Multifaceted Model Organism In this section, we overview five attributes that make the zebrafish an ideal model organism for studying animalemicrobe interactions. These attributes include a conserved vertebrate immune system, the ability to be associated with a wide range of microbes, high fecundity and small size, rapid ex-utero development, and amenability to live imaging. Our descriptions of these attributes are not exhaustive; rather, they are meant to serve as introductions to each topic. In addition to the utility of the zebrafish to model human infectious diseases caused by single pathogens, we also highlight the potential of the system to inform our understanding of biomedically relevant interactions between humans and their complex resident microbial communities, known as microbiota.

Vertebrate Immune System Understanding how microbes influence human health and disease motivates a great deal of research on animalemicrobe interactions. Because of this, model

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systems capable of capturing aspects of human biology have immediate biomedical relevance. The zebrafish meets this need through its conserved vertebrate physiology, which shares considerable overlap with humans despite the 450 million years of evolution that separates them (Rauta, Nayak, & Das, 2012; Renshaw & Trede, 2012; Sunyer, 2013; Trede, Langenau, Traver, Look, & Zon, 2004). What makes the zebrafish particularly useful in studying animalemicrobe interactions is its immune system. The immune system plays a frontline role in mediating tolerance to a large number of normally benign and beneficial members of resident microbiota while allowing rapid detection and defense against those that are harmful. Importantly, as in humans, zebrafish have two interconnected arms of their immune system, referred to as innate and adaptive immunity. For the first 3e4 weeks of development zebrafish possess only innate immunity, which provides a unique opportunity to study the contributions of this system to animalemicrobe interactions in isolation (Lam, Chua, Gong, Lam, & Sin, 2004). The zebrafish innate immune system comprises an array of protein receptors and signaling networks that are expressed by cells throughout a variety of tissues (Rauta et al., 2012; Trede et al., 2004). Innate immunity proteins that are widely studied in zebrafish, for which there are human homologs, include various pattern recognition receptors (e.g., Toll-like receptors) that recognize microbederived molecules, the cytokines TNFa and IL1b, the signaling protein MyD88, and the transcription factor NF-kB (Jault, Pichon, & Chluba, 2004; Trede et al., 2004). Additional conserved features of the zebrafish innate immune system include antimicrobial proteins (e.g., lysozyme, C-reactive protein, and complement) and phagocytic myeloid cells (e.g., neutrophils and macrophages) that serve as first responders to sites of infection and inflammation (Harvie & Huttenlocher, 2015; Li et al., 2007; Lieschke, Oates, Crowhurst, Ward, & Layton, 2001; Rauta et al., 2012). Characterization of the adaptive immune system in zebrafish is more limited, but adult fish are clearly capable of elaborating an adaptive immune response that is highly similar to that of humans and other mammals (Rauta et al., 2012; Sunyer, 2013; Trede et al., 2004). Generally, the adaptive immune system of teleost fish is made up of T and B lymphocyte lineages that express the T cell receptor (TCR) and a repertoire of immunoglobulins, respectively. Other fundamental components of teleost adaptive immunity include the major histocompatibility complex (MHC) and the recombination-activating genes (RAG) 1 and 2. MHC molecules are expressed on the surface of many different cell types and present self and nonself antigens to T cells, whereas RAG1 and 2 are critical for generating TCR and immunoglobulin diversity and ultimately, T and B lymphocyte function.

Natural and Surrogate Microbial Associations Zebrafish naturally associate with a multitude of commensal, beneficial, and pathogenic microorganisms (Roeselers et al., 2011; Stephens et al., 2016). An exciting and rapidly growing area of research aims to understand the diverse community of microbes that thrive within the zebrafish intestine, particularly the bacteria. The intestinal microbiota of zebrafish is an appealing system to study for several reasons (Burns & Guillemin, 2017). Foremost, it contains hundreds of bacterial species that can interact with each other and influence vital aspects of zebrafish biology (Bates et al., 2006; Hill, Franzosa, Huttenhower, & Guillemin, 2016; Stephens et al., 2016). The complexity of the zebrafish microbiota falls in between that of the simple gut communities of invertebrate models, such as the fruit fly, which comprise tens of bacterial species, and that of mammalian models, such as mouse and human, which comprise hundreds to thousands of species (Kostic et al., 2013). The intermediate complexity of the zebrafish microbiota aids tractability while at the same time recapitulating the diversity found within microbial communities of high biomedical interest, such as those within the human intestine. In addition, as with the intestinal microbiota of humans, the zebrafish microbiota undergoes characteristic changes in the types of bacterial species it contains as the host develops and ages. The codevelopment of both zebrafish and humans with their resident gut microbes, is therefore dynamic, but in ways that appear to conform to a general ontogenetic pattern (Palmer, Bik, DiGiulio, Relman, & Brown, 2007; Stephens et al., 2016; Vangay, Ward, Gerber, & Knights, 2015). Another notable feature of the zebrafish intestinal microbiota is that many of its bacterial members are related to lineages known to associate with and influence the health of humans (Roeselers et al., 2011). Bacteria belonging to the phylum Proteobacteriad particularly the genera Vibrio, Aeromonas, Shewanella, Plesiomonas, Acinetobacter, Pseudomonas, Enterobacter, and Escherichiadare frequent residents within the zebrafish gut (Rolig et al., 2017). Intriguingly, proteobacterial lineages are highly abundant in the human intestinal microbiota over the first 3 years of life and their presence during this time has been linked to decreased incidence of allergy and chronic disease (Palmer et al., 2007; Vangay et al., 2015; Vatanen et al., 2016). In a similar vein, some lineages of Escherichia can promote intestinal homeostasis, a phenomenon that has been observed in humans, mice, and zebrafish (Behnsen, Deriu, SassoneCorsi, & Raffatellu, 2013; Deriu et al., 2013; Rolig et al., 2017). However, in stark contrast to their beneficial activities, proteobacterial lineages are widely recognized as some of the most notorious human and animal

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The Zebrafish: An Adaptable and Multifaceted Model Organism

pathogens. Moreover, increased Proteobacteria abundance has been shown to play a role in inflammatory bowel disease and is correlated with elevated mucosal inflammation during pregnancy (Gevers et al., 2014; Koren et al., 2012). Therefore, understanding how zebrafish interact with this interesting and highly relevant group of bacteria promises to help illuminate many animalemicrobe relationships, especially those involving humans. The zebrafish also allows microbial associations to be experimentally controlled, which is a particularly powerful feature of this model organism (Melancon et al., 2017). Zebrafish embryos can be derived germ-free fairly easily and raised in the presence of defined microbial communities. This is achieved by surface sterilizing embryos with a series of washes in embryo medium containing low amounts of bleach and iodine. After sterilization, animals can be kept in standard tissue culture flasks containing filtered embryo medium as larvae when they can use endogenous yoke stores for nutrients. Rearing older germ-free zebrafish requires more labor-intensive husbandry and thus has been employed less often in research (Phelps et al., 2017; Rendueles et al., 2012). These sterilized animals can be maintained germ-free or in association with specific microbes via inoculation into the water column. Microbial associations with zebrafish can be further controlled with genetically engineered microbes. Many of the bacterial lineages native to zebrafish are generally amenable to established genetic techniques, with new approaches continuing to be developed that further improve their genetic tractability (Wiles et al., 2018). A major benefit of using genetically engineered bacteria in animalemicrobe interaction research is that it provides an opportunity to track bacteria, for example, via the expression fluorescent proteins, as they colonize host tissues (Schlomann, Wiles, Wall, Guillemin, & Parthasarathy, 2018; Wiles et al., 2016). There is also exciting potential to control bacterial behavior and the manner, in which, they interact with the host using inducible genetic switches that are triggered by small diffusible molecules added to the water. Aside from native microbial associations, the zebrafish has been incredibly useful as a surrogate host for modeling a wide range of humanemicrobe interactions. Microbiota transplantation experiments have been performed with fecal samples from both mice and humans, demonstrating the possibility of reconstituting communities of mammalian gut-derived microbes in zebrafish (Rawls, Mahowald, Ley, & Gordon, 2006). Zebrafish are also used to study pathogenic interactions with individual human-derived bacterial isolates, including Escherichia coli, Staphylococcus aureus, Salmonella typhimurium, and the fungus Candida albicans (Gratacap, Rawls, & Wheeler, 2013; Prajsnar, Cunliffe,

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Foster, & Renshaw, 2008; van der Sar et al., 2003; Wiles, Bower, Redd, & Mulvey, 2009). Larval zebrafish, which defend against infection solely through their innate immune system, are especially well-suited for dissecting the early stages of pathogenic encounters. Illustrating this point, a larval zebrafish infection model was used to test whether human isolates of extraintestinal pathogenic E. coli (ExPEC) use the pore-forming toxin, a-hemolysin, to evade killing by neutrophils and macrophages (Wiles et al., 2009). Such activity was hypothesized over a decade earlier, but an adequate model system to test this idea was not available (Wiles & Mulvey, 2013). An alternative hypothesis put forth at the time was that a-hemolysin is required for liberating nutrients critical for bacterial growth through the lysis of host cells. In zebrafish hosts, in which, neutrophils and macrophages were depleted via morpholino-mediated knockdown of the transcription factor PU.1, it was shown that attenuated ExPEC mutants lacking a-hemolysin expression were capable of causing a lethal infection. This result simultaneously refuted the hypothesis that a-hemolysin is required for ExPEC nutrient liberation and demonstrated that its function is required to overcome early innate immune defenses mediated by neutrophils and macrophages.

High Fecundity and Small Size The high fecundity and relatively small size of zebrafish make them exceptionally amenable to highthroughput experimental schemes. In a single cross, a mating pair can produce hundreds of offspring that grow to be 3.5e4.5 mm in length as larvae and 20e25 mm as adults. These attributes make it possible to generate and maintain tens to thousands of animals for any given experiment, and thus, are advantageous in animalemicrobe interaction research for three main reasons. First, it is feasible to carry out forward genetic screens for host genes involved in mediating animale microbe interactions, which is an endeavor rarely undertaken in vertebrate organisms (Tobin et al., 2010; Trede et al., 2004). For example, and as further detailed below, chemical mutagenesis of zebrafish has been used to identify genetic loci important for combating Mycobacterium marinum infection (Tobin et al., 2010). Second, many different experimental conditions can be rapidly tested and analyzed in parallel. Notably, the small size of embryonic and larval zebrafish has led to the innovation of robotic injection systems capable of delivering bacteria into thousands of embryos per hour in combination with morpholino-based gene knockdown or drug treatments (Carvalho et al., 2011; Ordas et al., 2015; Pardo-Martin et al., 2010; Spaink et al., 2013; Takaki, Cosma, Troll, & Ramakrishnan, 2012). Lastly, zebrafish boost statistical power in experiments by allowing large sample sizes to

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be used, which is crucial for addressing the highly variable and complex interactions that occur between animals and associated microbes (Rolig et al., 2017, 2015).

Rapid Ex-utero Development It has become apparent that how humans and other animals grow and develop is intimately connected to the microbes they associate with. This expanded view of what governs animal development challenges the very concept of individuality. Consequently, the emerging idea is that to understand the fundamental biology of animals, the composite system of the animal and microbial cells and their functionsdknown as the “holobiont”dmust be taken into account (Gilbert, Sapp, & Tauber, 2012; McFall-Ngai et al., 2013). Indeed, the links between animal development and resident microbes are being studied in a variety of organisms, including humans; however, much is still to be learned about the extent of these interactions and their mechanisms. The rapid ex-utero development of the zebrafishd which can go from a fertilized egg to a multicellular animal with a beating heart within 1 day and a functional digestive system within four daysdmakes this model organism an incredibly sensitive system for studying animalemicrobe interactions during vertebrate development (Kanther & Rawls, 2010). Several studies have revealed that bacteria profoundly influence gene expression, proliferation, and differentiation within the intestinal tract of developing zebrafish (Bates et al., 2006, 2007; Camp, Jazwa, Trent, & Rawls, 2012, 2014; Cheesman, Neal, Mittge, Seredick, & Guillemin, 2011; Davison et al., 2017; Kanther et al., 2011; Lickwar et al., 2017; Marjoram et al., 2015; Semova et al., 2012). It was found that bacteria trigger gene expression patterns involving nutrient metabolism and innate immunity within the zebrafish intestine that mirror patterns seen in mice (Lickwar et al., 2017). Intriguingly, it was recently discovered that zebrafish HNF4A, which is a member of an ancient metazoan family of hepatocyte nuclear factor 4 (HNF4) transcription factors, plays a central role in mediating microbiota-dependent gene expression changes within the intestinal epithelium (Davison et al., 2017). Intriguingly, sequence variants of human HNF4A have been linked to the incidence of inflammatory bowel diseases and colon cancer (Barrett et al., 2009; Chellappa et al., 2016). Therefore, the microbiotaeHNF4A axis characterized in zebrafish may represent an evolutionarily conserved signaling conduit between animals and their resident bacteria (Davison et al., 2017). Another key player in the dialogue between microbiota and the developing zebrafish intestine is the innate immunity protein MyD88 that functions as an adaptor

for multiple innate immune receptors, including both Toll-like receptors and Interleukin-1 receptors. For example, MyD88 was found to be required for the induction intestinal alkaline phosphatase (Iap) by the bacterial cell wall component lipopolysaccharide (LPS), also known as endotoxin (Bates, Akerlund, Mittge, & Guillemin, 2007). LPS can elicit severe and even fatal inflammatory responses in animals. However, through upregulation of the LPS-detoxifying Iap enzyme, moderate amounts of LPS promote mucosal tolerance to resident bacteria within the intestine. MyD88 was also found to play a role in mediating host responses to resident microbiota that shape intestinal development and homeostasis. For example, MyD88, in combination with canonical Wnt signaling, is required for normal intestinal epithelial cell proliferation in response to the presence of secreted bacterial products (Cheesman et al., 2011). Similarly, MyD88 was found to mediate microbiota-derived cues that modulate Notch signaling and influence the balance of absorptive versus secretory cells in the intestine (Troll et al., 2018). Bacteria can also influence developmental processes at sites outside of the intestine. It has been discovered that some bacterial members of the zebrafish intestinal microbiota encode a secreted protein capable of stimulating the expansion of insulin-producing pancreatic b cells that are crucial for controlling metabolic homeostasis (Hill et al., 2016). Intriguingly, this protein, which has been dubbed b cell expansion factor A (BefA), can be detected in the genomes of various bacteria that live within the human intestine. In addition, the zebrafish is emerging as a tractable model for studying the role of microbiota in neurobehavioral development, which because of the strong legacy of zebrafish in neurobiological research, promises to help uncover novel mechanisms that govern the so-called gutebrain axis (Phelps et al., 2017).

Optical Transparency and Amenability to Live Imaging Conventional approaches for studying animale microbe interactions largely neglect their spatial and temporal organization, yet these elements play defining roles in biological systems throughout nature. Offering a solution to this problem is the zebrafish, which has an optically transparent body during its larval and juvenile stages. This unique characteristic of the zebrafish, combined with its amenability to a variety of live imaging techniques and the expression of fluorescent proteins for tracking the activity of host and microbial cells, provides a literal window into the spatiotemporal organization of animalemicrobe interactions within a living host (Jemielita et al., 2014; Parthasarathy, 2018).

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The Zebrafish: Forging and Overturning Paradigms in AnimaleMicrobe Interaction Research

For example, researchers used state-of-the-art live imaging to track zebrafish intestinal colonization patterns of two fluorescently marked bacterial symbionts, Vibrio and Aeromonas (Wiles et al., 2016). On their own in mono-association, each symbiont forms a distinct population architecture within a different region of the intestine from the other. The consequences of these colonization patterns became evident after Vibrio and Aeromonas were monitored together within the same host. Vibrio induces sudden and dramatic collapses in Aeromonas abundance that occasionally result in complete extinction events while Vibrio grows just as it does during mono-association. Closer inspection of the underlying mechanism driving this apparent competitive interaction revealed that Aeromonas lives as rigid multicellular aggregates that are highly sensitive to peristaltic flow. By contrast, Vibrio exists mostly as highly motile single cells that readily adapt to the expanding and contracting environment of the intestine. Remarkably, in hosts with a dysfunctional enteric nervous system and thus, altered peristaltic activity, the competitive interaction between Vibrio and Aeromonas is neutralized, and these two bacteria coexist. This study shows that the host can exert physical control over the microbes that inhabit the intestine and highlights that changing the physical mechanics of the intestine may be a strategy for microbiota engineering. The capacity of the zebrafish to reveal the state of animalemicrobe interactions in real time also greatly enhances the possibilities for screening and characterization of specific phenotypes. For example, highthroughput imaging of zebrafish using flow cell sorting systems, coupled with robotic manipulation technologies like those mentioned earlier, represent a powerful screening platform (Carvalho et al., 2011; Pardo-Martin et al., 2010; Spaink et al., 2013). However, “low-tech” approaches based on fluorescence stereomicroscopy and manual analysis have also proven robust and fruitful. For example, using transgenic zebrafish that have fluorescently marked neutrophils, numerous bacterial members of the zebrafish intestinal microbiota were screened for their ability to induce neutrophil recruitment, which is a readout of inflammation (Rolig, Parthasarathy, Burns, Bohannan, & Guillemin, 2015). This work led to two important observations. First, several different bacterial lineages exhibit antiinflammatory activity, suggesting that this trait may be common among intestinal symbionts and that resident microbiota are active manipulators of their environment. And second, minor community members can disproportionately modulate immune responses. In other words, symbiotic bacteria that are present at a relatively low abundance can still have a large effect on the immune system.

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The Zebrafish: Forging and Overturning Paradigms in AnimaleMicrobe Interaction Research As outlined in the section above, the zebrafish is a multifaceted and highly amenable model organism for studying animalemicrobe interactions. In this section, we summarize two sets of studies that showcase the utility of the zebrafish and its ability to reveal novel aspects of animalemicrobe systems.

Immunity and Interhost Dispersal of Intestinal Microbiota Humans and other animals are associated with diverse communities of microbes throughout their lives that have the capacity to dramatically influence their health and development. Although these symbiotic relationships appear to follow general patterns, in terms of the way they assemble and the types of microbes they involve, there is still a great deal of variation that exists between individuals that cannot yet be explained. A pair of recent studies using zebrafish have now shown that dispersal of microbes between hosts appears to play a major role (Burns et al., 2017; Stagaman, Burns, Guillemin, & Bohannan, 2017). The zebrafish is an excellent animal model for addressing potential sources of microbiota variation, exemplifying three of the features discussed above: first, zebrafish innate and adaptive immunity can be experimentally distinguished and are known to act as microbial filters in mammals; second, the small size of zebrafish makes them amenable to multiple experimental housing conditions that, for example, differ in the extent of microbial dispersal, and third, the animals’ fecundity allows many replicate zebrafish hosts to be used in a single experiment to adequately capture and quantify variation. In these two studies, intestinal microbiomes were profiled from wild-type, and immunocompromised zebrafish deficient for either innate or adaptive immunity (myd88 / and rag1 / respectively), and the hosts’ immune genotypes were found to contribute to the composition of their intestinal microbiota, in agreement with previous observations made in mice. Unexpectedly, the cohousing conditions had a profound impact on the microbiota, with conditions that facilitated interhost dispersal of microbes almost entirely negating the filtering effects of the immune system. These studies indicate that the factors governing microbiota composition are not necessarily inherent to an individual host, but also shaped by the metacommunity of hosts and microorganisms with which each individual coexists. Similar patterns of interhost transmission of microbiomes are beginning to emerge from studies of human populations (Brito et al., 2019). Zebrafish promises to be a powerful system

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for modeling the epidemiology and mechanisms of interhost transmission of the microbiota.

Fighting Mycobacterium Infection is an Inflammatory Balancing Act Tuberculosis (TB) is a disease in humans that is caused by the bacterium M. tuberculosis. According to the World Health Organization, in 2017, 10 million people worldwide developed symptomatic TB, and 1.6 million died from the disease (World Health Organization, 2018). Despite over a 100 years of research on M. tuberculosis pathogenesis, it is still unclear what determines a person’s susceptibility to TB and of those who fall ill, why some respond to treatment while others do not (Ramakrishnan, 2013). Shockingly, new therapeutic strategies for TB have not been implemented for several decadesdthe current vaccine is 90 years old, and the current treatment regimen is over 50 years old (Ramakrishnan, 2013). New insights into this devastating human disease come from research on a related disease in zebrafish caused by M. marinum. A hallmark of both zebrafish and human Mycobacterium infection is the formation of granulomas, which are immune cell complexes mostly made up macrophages that are typically thought to protect the host by walling off the pathogen and controlling its growth (Ramakrishnan, 2013). However, live imaging of granuloma formation and dynamics in zebrafish has called this deeply rooted paradigm into question. It was found that granulomas actually promote the growth of M. marinum and remarkably, M. marinum attracts new and permissive macrophages to established granulomas to spur further growth and dissemination (Davis & Ramakrishnan, 2009). Not only has the zebrafish model challenged conventional wisdom about human TB pathogenesis, but it has also uncovered new leads into host genetic susceptibility to this pathogen. A forward genetic screen of hundreds of chemically mutagenized larval zebrafish hosts infected with fluorescently marked M. marinum, which allowed disease progression to be monitored in real time, identified the gene encoding leukotriene A4 hydrolase (LTA4H) as a critical player in the host’s defense against M. marinum (Tobin et al., 2010). Interestingly, follow up studies revealed that both LTA4H deficiency, as well as excess, led to increased disease susceptibility (Roca & Ramakrishnan, 2013; Tobin et al., 2012). Ultimately, it was discovered that LTA4H is part of a delicate balancing act in the inflammatory response to M. marinumdnot enough results in a lack of pathogen containment, whereas too much, results in collateral damage that cripples host defenses. Remarkably, these observations were found to mirror similar

pathogenic interactions in humans infected with M. tuberculosis. Humans who are genetically predisposed to have either low or high levels of LTA4H are more susceptible to TB compared to those with intermediate levels (Tobin et al., 2012, 2010). This represents an exciting breakthrough in TB research because it uncovered the underlying intricacies of the disease while at the same time highlighting that new TB treatments may need to involve personalized regimens that take into account host genetics.

Outlook Advancements in high-throughput DNA sequencing and omics technologies continue to expose new facets of animalemicrobe interactions. A growing awareness that microbes play a more diverse and far-reaching role in animal biology than previously thought has transformed the field into a melting pot of collaborative and cross-disciplinary research. A major goal moving forward is to dissect the mechanisms that underlie the overwhelming number of newly discovered animale microbe relationships. As described in this chapter, the laboratory zebrafish represents an ideal animal model for addressing this challenge. Not only is the zebrafish useful in uncovering the cellular and molecular details of animalemicrobe interactions, it is also well-suited to tackle many emerging areas of interest. In particular, zebrafish facilitate the experimental interrogation of ecological and population-level interactions, as well as microbial impacts on animal development. In addition, the amenability of zebrafish to large-scale comparative studies will help identify unifying principles of animale microbe interactions. With the continued innovation of genome editing tools and gnotobiotic husbandry techniques, the zebrafish is poised to make major contributions in the exploration of animals and their microbial associations.

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C H A P T E R

49 Targeted Editing of Zebrafish Genes to Understand Gene Function and Human Disease Pathology Alberto Rissone1, Johan Ledin2, Shawn M. Burgess1 Translational and Functional Genomics Branch, National Human Genome Research Institute, National Institutes of Health, Bethesda, MD, United States of America; 2Department of Organismal Biology, Science for Life Laboratory, Uppsala University, Uppsala, Sweden

1

Background Why Modify Gene Expression in Zebrafish? The zebrafish model system is a particularly powerful tool to investigate gene function, and for more than 3 decades (Detrich, Westerfield, & Zon, 1999), researchers have used various techniques to modify the zebrafish genome, studying the genetic basis of organism development and homeostasis. In addition, zebrafish genome editing can be performed to generate zebrafish models recapitulating pathological processes displayed in human diseases (Langheinrich, 2003; Santoriello & Zon, 2012). A common strategy consists of the inactivation of the gene/genes homologous to the human genes associated with the disease. A “sick” zebrafish that recapitulates the aspects of the human disease pathology may aid the understanding of disease mechanisms and the development of new clinical treatments.

Gain Versus Loss of Gene Function Genome editing procedures often involve the interference of gene expression to generate two types of gene changes: “loss-of-function” and “gain-of-function.” In loss-of-function experiments, the target gene is necessary for a specific biological process if the complete removal of a functional protein encoded by that gene disrupts the process. Generating loss-of-function alleles in genes of interest is the most common type of genome modification experiment in zebrafish. Usually, a “nonsense” or “frame-shift” mutation introduced in The Zebrafish in Biomedical Research https://doi.org/10.1016/B978-0-12-812431-4.00049-X

the coding sequence presents a high probability of resulting in a “null allele,” meaning the complete abolishment of protein function. Nonsense alleles have a premature signal to end translation of the protein (stop codons). Frame-shift alleles have a small insertion or deletion of bases that shifts the normal three base pair reading frame used by the cell. Insertions or deletions that are not multiples of three (e.g. a five base pair deletion or a two base pair insertion) will change the reading frame of the RNA, causing the protein to be “mistranslated,” resulting in an incorrect and usually truncated protein. Mutated alleles retaining some functionality are considered “hypomorphic,” and they can be the result of nonsynonymous mutations inducing amino acid substitutions in the protein code or small in-frame deletions that do not completely inactivate gene function. In contrast, a gain-of-function experiment adds (usually unwanted) functionality to the genome, by typically expressing genes outside of their natural spatiotemporal window or at higher levels than wanted in the cell. When ectopic expression of a gene induces new biological outcomes in strange, often unwanted locations, it indicates a regulatory role for the gene in that process. In addition, gain-of-function strategies have been developed to modify the zebrafish genome solely for experimental observation of biological processes, typically by the addition of controlled expression of detectable markers, such as green fluorescent protein (GFP) that can be observed in a live animal. The ability to share these “transgenic” lines between zebrafish laboratories has significantly contributed to the scientific popularity

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49. Understanding Human Diseases Using Zebrafish Genetics

of the zebrafish model system and enables researchers to compare the outcome of new experiments with previously published experiments without having to construct reporter lines over again in his or her laboratory (Udvadia & Linney, 2003). Until recently, gain-of-function experiments to edit the zebrafish genome typically required the synthesis of a DNA construct that was randomly integrated in the zebrafish genome. They used a gene “promoter” sequence that drove gene expression tissuedspecifically followed by the gene you want to be misexpressed. Notably, expression from randomly inserted constructs is strongly influenced by the region of insertion, and it is crucial to carefully evaluate the resulting expression pattern. Recently developed technology has opened up the exciting possibility that rather than relying on random insertion, it is now possible to guide insertion of DNA constructs to a specific location in the genome using programmable nucleases, which allows for more precise control of gene expression (Kimura, Hisano, Kawahara, & Higashijima, 2014) (discussed below in “Sequence integration” section).

Forward and Reverse Genetics (Historical and Conceptual Differences in Research Approach) The experimental process for linking genotype to phenotype follows one of two major procedures: “forward” (classical) or “reverse” genetics. Forward genetics procedures involve random mutagenesis of the zebrafish genome by mutagenic chemicals or injected viruses, followed by isolation of mutant phenotypes (Lawson & Wolfe, 2011). In zebrafish, these techniques were introduced in the early 1990s, and they were spectacularly successful in identifying major genetic components of embryo development. However, the huge number of mutated fish required and the efforts necessary for the identification of the affected gene using positional cloning limited the use of this approach in smaller zebrafish labs. In contrast, a reverse genetic experiment is designed to answer questions about the role/function of a specific protein by the introduction of targeted mutations followed by phenotypic characterization. Historically, in zebrafish, the first tool to be used for reverse genetic screening was “morpholinos” (MOs) (Nasevicius & Ekker, 2000). These were used to transiently block protein translation from the mRNA. Soon after, the TILLING (or Targeting Induced Local Lesions in Genome) system was developed in the early 2000s (Wienholds, Schulte-Merker, Walderich, & Plasterk, 2002). Although TILLING still required random mutagenesis, it allowed the screening of mutations in specific genes of interest bypassing the laborious positional cloning step, and for that reason, it quickly became popular until the advent of the targeted mutagenesis approaches in the late 2000s.

The first targeted mutagenesis tools for reverse genetics, based on genome-editing nucleases, such as Zinc Finger Nucleases (ZFNs) (Doyon et al., 2008; Meng, Noyes, Zhu, Lawson, & Wolfe, 2008) and Transcription Activator-Like Effector Nucleases (TALENs) (Bedell et al., 2012; Huang et al., 2011) developed quite recently (in 2008 and in 2011, respectively) in the zebrafish model system and were for a long time expensive and/or laborious. More recently (2013) the development of the Clustered Regularly Interspaced Short Palindromic Repeat/CRISPR associated protein 9 (CRISPR/Cas9) system (discussed below) has made targeted genome editing simple and affordable for most if not all zebrafish laboratories (Hwang, Fu, Reyon, Maeder, Kaini, et al., 2013; Hwang, Fu, Reyon, Maeder, Tsai, et al., 2013; Jao, Wente, & Chen, 2013; Varshney, Pei, et al., 2015).

Transient Methods for Altering Gene Expression The intrinsic features of the zebrafish model system make it useful in the manipulation of single or multiple target genes using transient gene knockdown and transgenic technology approaches.

Morpholinos One of the most rapid, inexpensive, and consequently, popular tools for performing reverse genetic analysis in zebrafish is morpholinos (MOs). MOs are modified antisense oligonucleotides designed to bind target RNAs, creating a transient gene knockdown effect (Bill, Petzold, Clark, Schimmenti, & Ekker, 2009; Eisen & Smith, 2008; Timme-Laragy, Karchner, & Hahn, 2012). The binding of morpholinos to the matching mRNA targets at the translational start-site directly inhibits the protein synthesis by sterically blocking the translation. The splice blocking MOs, targeting specific exon-intron boundaries of the pre-mRNA, instead interfere with the correct splicing mechanism. More recently, MOs have also been used to block microRNA (miRNA) activity directly targeting the guide strand of the microRNA, or indirectly targeting the miRNA binding regions on mRNA targets (Flynt, Rao, & Patton, 2017) (see Table 49.1 for a comparison between gene knockdown and knockout approaches in zebrafish). In zebrafish, MOs are delivered by direct injection into the fertilized embryos during the first stages of development (one- to the four-cell stage maximum). It is important to note that MO action is always dose dependent, and usually, it is limited to the first 3e5 days postfertilization (Timme-Laragy et al., 2012). In order to try to overcome this limitation, lightactivated MOs containing a photosensitive subunit (photo-MOs) have been developed (Tallafuss et al., 2012). The major advantage of photo-MOs is represented

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TABLE 49.1

Comparison between gene knock-down (left) and knock-out (right) approaches in zebrafish.

Targets

Morpholino

ZFN/TALEN/CRISPR Mutants

pre-mRNA, mRNA or miRNA

genomic DNA

Delivery

Direct injection in the embryo (1e4 cell stage)

Validation of activity on target sequences

1e2 weeks (RT-PCR or Western blot)

1 week (genotyping/sequencing)

Off-target effects

High probability (often mediated by activation of tp53 pathway)

Relatively low probability (reduced with outcrossing)

Effects

Partial to full knock-down (the effects are dose dependent and temporally limited to 120 hpf max)

Partial to full knock-out depending on the selected mutation

Proving specificity

At least 2 unique MOs required plus several controls

Genotype-phenotype correlation

Time required

Rapid (3e4 weeks)

Slow (4e7 month)

Cost

Relatively low

Low to moderate (based on the system used)

by the spatiotemporal control of the target gene expression, although with certain embryonic target regions or stages, the correct photo-activation using confocal microscope could sometimes be challenging. Since the early 2000s, MOs have been extensively used in studies of zebrafish development, to confirm the function of candidate genes and even to model human diseases. Nonetheless, lately, MO use has been criticized because of the induction of off-target effects, mostly mediated by p53 (Robu et al., 2007); as well as, in several cases, because they failed to replicate mutant phenotypes (Kok et al., 2015). However, in 2015, Rossi and colleagues (Rossi et al., 2015) showed that some MO-specific phenotypes observed in morphants are not present in the corresponding mutant animals because of a genetic compensation, reopening the long-standing controversial debate about the knockout versus knockdown approaches (Blum, De Robertis, Wallingford, & Niehrs, 2015; Schulte-Merker & Stainier, 2014;Stainier, Kontarakis, & Rossi, 2015). Although the debate is likely to continue, a general consensus has emerged that studies involving experiments with a new MO design should include a direct comparison with the corresponding genetic mutant phenotype (Stainier et al., 2017).

mRNA Injection and Pharmacological Approaches The overexpression of a target gene (Prelich, 2012) to study its function represents another very popular tool among the zebrafish research community. Specific variants of the gene can be introduced in the embryos by microinjection of capped mRNAs or DNA plasmids. The use of DNA plasmids will result in a variable

expression of the protein through the cells of the embryo generating a “mosaic” embryo, while the injection of capped mRNA usually produces a more homogenous but more temporally limited distribution. In this approach, the effects of the normal and the mutated variants of a transcript can be compared to understand their functions better during embryo development. This approach proved to be very helpful in confirming the effects of MO knockdowns and gene knockouts, as well as in testing the effects of specific mutated forms found in human patients. In some cases, the transcripts can be tagged with specific epitopes in order to visualize the protein localization during development. However, the limits of this approach are represented by the fact that the effect is transient, and often it results in a non-physiological ectopic expression of the protein. From this point of view, a pharmacological approach targeting elements of a defined pathway can directly or indirectly affect the gene expression of specific targets, and thus, could represent a reasonable alternative. Moreover, because chemicals can be applied and removed in a very precise time-dependent manner, a pharmacological agent may also represent a more physiologically relevant option (Skromne & Prince, 2008). A pharmacological compound typically affects entire families of gene products, which makes the phenotypic effects sometimes hard to link to the function of single genes. However, with knowledge of the spatiotemporal expression of individual members of gene families and partial biochemical characterization of the pharmacological reagent, it may be possible to attribute at least the phenotypic effects of the administration of pharmacological compounds to the subset of genes expressed in the area of the observed phenotype. This approach can be helpful to show, for example, a stage-specific effect of the target gene expression or to

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reduce toxicity or potential off-target effects due to ectopic expression of the target protein. However, when using a pharmacological approach, it is critical to be aware of potential pleiotropic effects due to a concentration-dependent specificity, and then the concentration in embryos should be carefully titrated. Finally, the genetic and pharmacological approaches can be used together to sustain and confirm each other (Lieschke & Currie, 2007; Skromne & Prince, 2008).

Techniques for Genome Editing Site-specific Nucleases The access to complete genomic sequence for a growing number of species in association with the advent of better technologies for genome modification

facilitated the gene function studies and opened the genome-editing era. Compared to older technologies, such as TILLING and random mutagenesis, the genome editing using ZFNs, TALENs, or CRISPR/Cas9 allowed direct targeting of specific genomic sequences and the easy selection of mutations, increasing the rate of identifying development and disease-related phenotypes (Varshney, Sood, & Burgess, 2015) (see Fig. 49.1). Moreover, the use of ZFNs, TALENs, and in particular the CRISPR/Cas9 system, reduces the problem of nonspecific mutations in the genome and repeated mutagenesis of the same gene, both common side effects of using random mutagenesis techniques. Genome editing consists of the use of proteins able to induce modification of genome sequences in specific locations of the genomic DNA. The sequence specificity is typically obtained using different kinds of DNA-binding

(A) FokI FokI = ZFN domains

(B) FokI FokI = TALE repeats

(C)

FIGURE 49.1 Schematic representation of ZFNs, TALENs, and CRISPR/Cas9 genome editing systems. Zinc Finger Nucleases (ZFNs) (A) and Transcription Activator-Like Effector Nucleases (TALENs) (B) technologies are based on fusing modified FokI nucleases able to cut the DNA only in the dimeric form to specific DNA binding motifs (ZFN domains or TALE domains, respectively). Different colors used for the DNA binding domains indicate different sequence specificities. (C) The Clustered Regularly Interspaced Short Palindromic Repeat/CRISPR associated protein 9 (CRISPR/Cas9) system requires the base pairing of a guide RNA (gRNA) downstream of a protospacer adjacent motif (PAM) in order to locate and induce double-strand breaks.

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TABLE 49.2

Comparison among different nuclease systems for genome editing. ZINC-FINGER nucleases

TALENs

CRISPR/Cas9

Recognition type

Protein-DNA

Protein-DNA

RNA-DNA

Cutting activity

Work as dimers

Work as dimers

Works as a monomer

Recognition site

9e12 nt/monomer

14-20 nt/monomer

w20 nt upstream a PAM site

Spacer

Required (5e7 nt)

Required (w14 nt)

No spacer required

Design

Limited (restrictions in triplets design)

Relatively flexible

Extremely flexible

Assembly

Laborious and time consuming

Moderately easy

Easy

Specificity

Usually high (but off target effect reported)

Usually high

Higher potential for off-target effects

Enzime

Coupling with non-specific nuclease FokI

Mechanism of action

Cas9

Induction of double strand brake in gDNA repaired by HDR or NHEJ mechanisms

Induced Mutations

Insertions and/or deletions

Mostly deletions

Insertions and/or deletions

Mutagenic efficiency

Low

Moderate

High

Multiplexing

No

No

Yes

Cost

Moderate

Moderate

Low

domains or using guide RNA molecules (see Table 49.2 for comparison among different nuclease systems). In the most straightforward approach, the use of engineered enzymes (nucleases) induces a double-stranded break in the DNA of specific regions of interest. Following the DNA double-strand break (DSB), two different cellular DNA repair mechanisms are induced (Symington & Gautier, 2011): (a) Homology Dependent Repair (HDR), which is rare, and it uses a template DNA supplied by the researcher to repair the DNA break (see below), or (b) nonhomologous end joining (NHEJ), which is more frequent in vivo, but highly error-prone (see Fig. 49.2). As a consequence, NHEJ repair mechanism is often accompanied by loss or gain of small fragments of DNA (usually ranging from 1 bp to w40 bps), which may induce frame-shift mutations in the coding sequence of the target gene impairing protein sequence and functionality. More recently, the use of different effector domains to perturb genome structure and function (such as recombinases, epigenetic modifiers, transcriptional repressors or activators, etc.) created a further level of versatility for the system (Hsu, Lander, & Zhang, 2014). In zebrafish, one of the major problems of using it as a genetic model was the teleost-specific genome duplication event that occurred w400 million years ago (Meyer & Van de Peer, 2005). Because of this ancient duplication event, zebrafish and other teleost species have a higher number of protein-coding genes (w26,000) compared to human, mouse, or chicken (Howe et al., 2013).

Approximately 20% of all zebrafish genes have two copies per haploid genome instead of the usual one copy. Thus, for one in five genes, knocking out one copy is insufficient for seeing the effects of a null allele as the second copy of the gene could compensate some or all functions in the animal, making the genetic analysis more difficult. In general, after duplication, the gene duplicates tend to diverge using one of the following models: (a) one copy is lost via acquired genomic mutations and only one copy survives (nonfunctionalization); (b) subfunctionalization occurs when each duplicated gene retains a subset of the original ancestral function; (c) neofunctionalization is the process in which one or both of the duplicates gain new functions; finally, (d) subfunctionalization can be followed by neofunctionalization events increasing the overall functional divergence (Ohno, 1970; Postlethwait, Amores, Cresko, Singer, & Yan, 2004; Rastogi & Liberles, 2005). Although the duplication event is thought to be at the base of the evolutionary success of teleost fish, obviously, it represents a problem for genome editing analyses. In most of the cases, there is a risk of compensatory effects by the untargeted gene copy. Fortunately, the multiplexing approach available with most of the genome editing technologies in zebrafish helps to overcome this limitation, allowing, for example, the direct targeting of both the duplicates simultaneously or even different members of the same gene family.

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NHEJ

FIGURE 49.2 Schematic representation of DNA repair mechanisms after DNA double-strand breaks. DNA double-strand breaks induced by nuclease activity can be repaired by nonhomologous end joining (NHEJ) (A) or Homology Dependent Repair (HDR) (B) mechanisms. NHEJ is the most frequent mechanism, but it often introduces an insertion or a deletion of bases (indel) at the genomic target site. Instead, the HDR pathway requires the presence of a template DNA sequence (donor DNA) and consequently is more rare and precise. It can be used to introduce exogenous DNA into the zebrafish genome. PAM ¼ protospacer adjacent motif.

specific sequence of three nucleotides (nt) in the major groove of DNA, and they developed a library of specific zinc-fingers. Combining several fingers in tandem allowed the targeting of unique 9e12 base pair sequences within the genome. Two different ZFN “arms” are created to recognize a 10e18 base pair target with a spacer of 5e7 nt between each half, which is where the FokI-induced double-strand break will occur. Researchers increased the specificity of the ZFNs system by the use of recombinant versions of FokI enzymes able to cut the DNA only when in dimeric form. The two different halves needed to be brought together (one on each side of the spacer) in order to cut at the locus. ZFNs first became popular in cell culture systems, and eventually, in 2008, they became the first genomeediting tool to be used in zebrafish (Doyon et al., 2008; Meng et al., 2008). However, ZFNs adoption in zebrafish was limited by low efficiency and flexibility, the complex assembly of their arms, significant off-target activity, and lastly, by the advent of TALENs (2011) and CRISPR/Cas9 (2013) that represented more efficient, faster systems with higher mutagenic throughput. After several years of ZFN use, TALENs emerged as a simpler-to-use alternative (Joung & Sander, 2013). TALEN technology is conceptually identical to ZFNs: the coupling of FokI nucleases to DNA binding motifs working as dimers to increase specificity (see Fig. 49.1B). In TALENs, the DNA binding motif is from the bacterial TALE repeats, which consist of 34 amino acids with the 12th and 13th residues consisting of a variable region (RVD) containing a simple code for binding to specific nucleotides. In contrast to ZFN domains, which bind to specific triplets of bases, each TALE repeat recognizes a single nucleotide. By changing only two amino acids in each RVD, researchers can generate a combination of motifs specific for any DNA sequence. While ZFN cutting induces insertion and deletion with the same frequency, TALENs rarely introduce insertions (Kim et al., 2013; Sood et al., 2013; Varshney, Pei, et al., 2015). Compared to ZFNs, TALEN technology represented a huge step forward in flexibility, efficiency, and design with similar, or potentially lower, off-target activity. However, compared to the CRISPR/Cas9 system, TALENs still present a reduced mutagenic throughput because of a time-consuming cloning process.

ZFNs and TALENs

CRISPR

Zinc Finger Nucleases are chimeric proteins, including DNA-binding domains typical of zinc finger-containing transcription factors and a bacterial FokI endonuclease domain (Urnov, Rebar, Holmes, Zhang, & Gregory, 2010) (see Fig. 49.1A). Researchers showed that each individual “zinc finger” was able to recognize and bind to a

The development of the CRISPR/Cas9 as a genomeediting tool was the consequence of discovering Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR) and CRISPR-associated (Cas) systems in prokaryotes (bacteria) (Wang, La Russa, & Qi, 2016). These unicellular organisms use a combination of short RNAs

(A)

PAM

INDEL

(B)

HDR

PAM

Left arm

Donor DNA

Right arm

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and proteins to target invading DNA sequences for cleavage. In essence, CRISPR/Cas provides a primitive adaptive immune system against plasmids and bacteriophage infections. In contrast to ZFNs and TALENs where the targeting of a specific genomic site is obtained using DNA-binding motifs, the CRISPR/Cas9 technology is based on base pairing using a guide RNA (gRNA) (see Fig. 49.1C). Once the guide finds the matching DNA sequence in the genome, the Cas9 endonuclease induces a double-strand break that can be repaired by the cell using HDR or NHEJ. The CRISPR/Cas9 tool proved to be a very efficient, versatile, and easy-to-use system, which has been used in several different biological contexts, from human cells to different animal models, zebrafish included (Varshney, Sood, et al., 2015). One of the major advantages of this system compared to ZFNs and TALENs is the broad genomic target coverage: the only limitation is the requirement of a PAM (or protospacer adjacent motif) site sequence directly upstream, the target region bound by the gRNA (20 base pairs long). For the most commonly used Cas9 protein, the PAM is the nucleotide sequence NGG, which occurs on average approximately once in every eight base pairs in the genome (Varshney, Sood, et al., 2015). Changing the 20 nt protospacer of the gRNA changes the target site and can be easily accomplished by subcloning the nucleotide sequence into a gRNA plasmid backbone or into a DNA fragment used directly for in-vitro transcription (Varshney, Pei, et al., 2015). In zebrafish, the Cas9 protein or mRNA encoding Cas9 and the gRNA (as a DNA cassette with a specific promoter or as an in-vitro synthetized gRNA of w100 nt) are introduced by coinjection during the earliest stages of embryo development (one- to the four-cell stage). The injection of multiple gRNAs is usually well tolerated in zebrafish embryos allowing the targeting in parallel of multiple genes simultaneously (multiplexing). Multiplexing can be utilized for several different purposes. It can be used, for example, to engineer the deletion of large genomic regions (two guides, one targeting each side of the desired genomic deletion) or to target duplicated genes or multiple members of the same gene family simultaneously. Other examples could be the targeting of multiple genes involved in the same signaling pathway or biological activity or unrelated genes to obtain compound heterozygous fish to reduce the space and the number of fish needed in the animal facility. Although most of the efforts so far have been focused on the targeting of gene coding sequences, recently some new studies are focusing on noncoding genomic regions, such as promoters, enhancers, and miRNA. In particular, Narayanan and colleagues (Narayanan et al., 2016) developed a multiplex approach to target miRNA gene families in vivo using zebrafish as the

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animal model. They generated inheritable mutations in multiple miRNA loci impairing the expression and activity of a miRNA family with negligible off-target effects (Narayanan et al., 2016). Finally, another interesting use of CRISPR/Cas9 has been recently published by Ablain, Durand, Yang, Zhou, and Zon (2015); (Ablain & Zon, 2016). They took advantage of the Tol2 transposon (transgenic) technology (Kawakami, 2007) to induce random integrations in the zebrafish genome of a CRISPR-based vector system for tissue-specific knockout of genes. Inside the vector, a U6 (a small nuclear RNA gene) promoter induces the ubiquitous and continuous expression of the gRNA, while the spatiotemporal expression of Cas9 is determined by a tissue-specific promoter. In this way, the effects of CRISPR/Cas9 system are limited to specific territories or tissues, potentially avoiding the embryonic lethality of some gene knockouts and the resulting animal potentially represents a more informative disease model. Notably, adding a second or multiple U6:gRNA cassettes in the vector backbone can easily allow a multiplexing approach. Compared to ZFNs and TALENs, one of the biggest concerns about the use of CRISPR/Cas9 is the increased potential for off-target effects induced using just 20 nt of sequence. So far, the data about nonspecific cleavage activity of CRISPR/Cas9 are limited. Moreover, the adoption of multiplexing approaches increases the concerns about the overall potential off-target effects of each injected gRNA. Varshney, Pei, et al. (2015) were able to detect off-target activity with germline transmission of the mutations in 1 of 25 predicted off-target-sites in exons, suggesting that the nonspecific activity of the system could be low but detectable. However, the potential offtarget activity is always sequence-dependent and could vary based on the gRNA used for targeting the genome. Several approaches were developed (Sander & Joung, 2014) to reduce the rate of off-target cleavages. One approach requires the use of an engineered version of Cas9 unable to cut the DNA (dead Cas9), fused with the same Fok1 endonuclease enzyme used in ZFN and TALEN systems. Similar to ZFNs and TALENs the increased specificity is obtained through targeting two different sequences to direct the Cas9-FokI fusion proteins to cut the same genomic site (Tsai et al., 2014) effectively doubling the length of the genomic targeting sequence. Another very efficient strategy used a mutant version of Cas9 (Cas9n or Cas9 nickase) able to induce only single-strand breaks in DNA and again using a pair of gRNAs targeting opposite DNA strands of the selected site. When the Cas9n cuts in off-target regions, the introduction of single-strand nicks can be efficiently repaired by the cells; however, the double nicks induced at the correct site create a double-stranded break that will be repaired by the more error-prone NHEJ system

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49. Understanding Human Diseases Using Zebrafish Genetics

(Ran et al., 2013). Although these methods can significantly reduce the off-target mutagenesis effects of CRISPR/Cas9 system, at the same time, they reduce its overall versatility, flexibility, and its multiplexing potential. However, in zebrafish, “out-crossing” the injected fish with wild-type strains, followed by identification of mutant “carriers” in the desired gene targets in most cases should be sufficient to segregate away any off-target activity.

Sequence Integration Random Insertion of Exogenous DNA In zebrafish, researchers have typically designed gainof-function experiments by random insertion of promoter and coding sequences in the genome using DNA transposons (Tol2, Sleeping Beauty or Ac/Ds systems) or restriction enzyme-based methods (meganuclease) (Grabher & Wittbrodt, 2008). While efficient, the expression of a randomly inserted sequence might be strongly influenced by copy number variation or epigenetic regulation in the region of the insertion making it crucial to carefully evaluate each resulting expression pattern by the inserted construct. Moreover, the added complexity of the positional effects on the function of randomly integrated constructs makes the comparisons between experiments difficult. In addition, correctly identifying promoters that express in the desired tissue-specific expression pattern is often difficult, with some regulatory DNA sequences tens or hundreds of kilobases away from the regulated gene. Recently, some reports indicated that the high efficiency of the CRISPR/Cas9 system can be a very powerful tool to induce a locus-specific integration of expression constructs (Auer, Duroure, De Cian, Concordet, & Del Bene, 2014; Kesavan, Chekuru, Machate, & Brand, 2017; Kimura et al., 2014; Ota et al., 2016) which, consequently, are expressed under the control of endogenous native regulatory context in the zebrafish genome. Although the integration can impair the expression of the endogenous gene (Ota et al., 2016) or it can be outside the target site inducing the expression of the reporter in unrelated cells (Kimura et al., 2014), this new generation of zebrafish transgenic lines represents a big step forward in the real-time analysis of the gene expression, avoiding the aforementioned limits of the random integration systems.

Introduction of Exogenous DNA by Nucleases Recent developments in editing technologies have inspired researchers using the zebrafish model to develop tools that induce site-specific insertion of DNA at the site of a double-stranded break in the

zebrafish genome (Hisano et al., 2015). DNA integration directed by DSBs has been achieved by multiple groups by leveraging three different cellular mechanisms: Homology Dependent Repair (HDR), Microhomology Mediated End Joining (MMEJ) and Nonhomologous End Joining (NHEJ) (Symington & Gautier, 2011).

Knock-in by HDR and MMEJ Mechanisms A cell can repair a DSB in a chromosome using homology dependent recombination where a DNA molecule with sequences matching both sides of the region of the DSB is used as a template (Sung & Klein, 2006) (see Fig. 49.2B). In zebrafish, exogenous DNA can be inserted by homology dependent recombination at the site of a DSB if the exogenous DNA is synthesized with long sequences identical to both sides of the DSB and the desired DNA insertion between the two sequencematching “arms.” Such identical sequences are commonly referred to as the homology arms, and typically they can be 1 Kb long or more (Hoshijima, Jurynec, & Grunwald, 2016; Shin, Chen, & Solnica-Krezel, 2014; Zhang, Huang, Zhang, & Lin, 2016; Zu et al., 2013). The recombination mechanism is typically error-free, and thus, well suited for integration into coding genes. However, the construction of donor DNA with long homology arms is relatively time consuming, which can limit the usefulness in zebrafish research projects where testing of several constructs is desired. In contrast, the MMEJ mechanism repairs DSBs by aligning microhomologous sequences internal to both ends of each side of a DSB. Several studies in zebrafish have successfully incorporated exogenous DNA at the site of a DSB using short homology arms ( T, C- > A or C- > G) in specific genomic locations with relatively high efficiency and a reduced occurrence of indel mutations. Moreover, using a catalytically dead Cas9 (able to

bind, but not to cut the DNA) instead of the Cas9 nickase, they were able to reduce the amount indels further, although with a lower base conversion efficiency (Zhang et al., 2017).

Conclusions Once efficient, precise, and robust methods for knockout, and more importantly, site-specific knock-in of exogenous DNA are developed, the zebrafish animal model could emerge as the preferred vertebrate system for many functional genomic studies. Several features make zebrafish particularly attractive in this regard. First, external fertilization and nearly transparent early larvae allow for direct observation of phenotypes or pathologies in a live animal. Second, the small size and relatively simple husbandry allow for larger-scale approaches that would be much more challenging in mammalian disease models, such as rodents. Finally, zebrafish represent an opportunity to reduce the use of rodent models when possible, an effort supported by most regulatory bodies that govern animal use. Zebrafish models represents a valid and economical alternative to reproduce mutations found in human patients (personalized medicine and gene therapy), to model lethal disease caused by dominant mutations, and finally, to study the effects of specific amino acid alterations in protein sequence/structure in vivo, in order to develop new and better drug treatments.

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Bill, B. R., Petzold, A. M., Clark, K. J., Schimmenti, L. A., & Ekker, S. C. (2009). A primer for morpholino use in zebrafish. Zebrafish, 6(1), 69e77. https://doi.org/10.1089/zeb.2008.0555. Blum, M., De Robertis, E. M., Wallingford, J. B., & Niehrs, C. (2015). Morpholinos: Antisense and sensibility. Developmental Cell, 35(2), 145e149. https://doi.org/10.1016/j.devcel.2015.09.017. Detrich, H. W., 3rd, Westerfield, M., & Zon, L. I. (1999). Overview of the zebrafish system. Methods in Cell Biology, 59, 3e10. Doyon, Y., McCammon, J. M., Miller, J. C., Faraji, F., Ngo, C., Katibah, G. E., et al. (2008). Heritable targeted gene disruption in zebrafish using designed zinc-finger nucleases. Nature Biotechnology, 26(6), 702e708. https://doi.org/10.1038/nbt1409. Eisen, J. S., & Smith, J. C. (2008). Controlling morpholino experiments: don’t stop making antisense. Development, 135(10), 1735e1743. https://doi.org/10.1242/dev.001115. Flynt, A. S., Rao, M., & Patton, J. G. (2017). Blocking zebrafish MicroRNAs with morpholinos. Methods in Molecular Biology, 1565, 59e78. https://doi.org/10.1007/978-1-4939-6817-6_6. Grabher, C., & Wittbrodt, J. (2008). Recent advances in meganucleaseand transposon-mediated transgenesis of medaka and zebrafish. Methods in Molecular Biology, 461, 521e539. https://doi.org/ 10.1007/978-1-60327-483-8_36. He, M. D., Zhang, F. H., Wang, H. L., Wang, H. P., Zhu, Z. Y., & Sun, Y. H. (2015). Efficient ligase 3-dependent microhomologymediated end joining repair of DNA double-strand breaks in zebrafish embryos. Mutation Research, 780, 86e96. https://doi.org/ 10.1016/j.mrfmmm.2015.08.004. Hisano, Y., Sakuma, T., Nakade, S., Ohga, R., Ota, S., Okamoto, H., et al. (2015). Precise in-frame integration of exogenous DNA mediated by CRISPR/Cas9 system in zebrafish. Scientific Reports, 5, 8841. https://doi.org/10.1038/srep08841. Hoshijima, K., Jurynec, M. J., & Grunwald, D. J. (2016). Precise genome editing by homologous recombination. Methods in Cell Biology, 135, 121e147. https://doi.org/10.1016/bs.mcb.2016.04.008. Howe, K., Clark, M. D., Torroja, C. F., Torrance, J., Berthelot, C., Muffato, M., et al. (2013). The zebrafish reference genome sequence and its relationship to the human genome. Nature, 496(7446), 498e503. https://doi.org/10.1038/nature12111. Hruscha, A., Krawitz, P., Rechenberg, A., Heinrich, V., Hecht, J., Haass, C., et al. (2013). Efficient CRISPR/Cas9 genome editing with low off-target effects in zebrafish. Development, 140(24), 4982e4987. https://doi.org/10.1242/dev.099085. Hsu, P. D., Lander, E. S., & Zhang, F. (2014). Development and applications of CRISPR-Cas9 for genome engineering. Cell, 157(6), 1262e1278. https://doi.org/10.1016/j.cell.2014.05.010. Huang, P., Xiao, A., Zhou, M., Zhu, Z., Lin, S., & Zhang, B. (2011). Heritable gene targeting in zebrafish using customized TALENs. Nature Biotechnology, 29(8), 699e700. https://doi.org/10.1038/nbt.1939. Hwang, W. Y., Fu, Y., Reyon, D., Maeder, M. L., Kaini, P., Sander, J. D., et al. (2013). Heritable and precise zebrafish genome editing using a CRISPR-Cas system. PLoS One, 8(7), e68708. https://doi.org/ 10.1371/journal.pone.0068708. Hwang, W. Y., Fu, Y., Reyon, D., Maeder, M. L., Tsai, S. Q., Sander, J. D., et al. (2013). Efficient genome editing in zebrafish using a CRISPRCas system. Nature Biotechnology, 31(3), 227e229. https://doi.org/ 10.1038/nbt.2501. Jao, L. E., Wente, S. R., & Chen, W. (2013). Efficient multiplex biallelic zebrafish genome editing using a CRISPR nuclease system. Proceedings of the National Academy of Sciences of the United States of America, 110(34), 13904e13909. https://doi.org/10.1073/pnas.1308335110. Joung, J. K., & Sander, J. D. (2013). TALENs: A widely applicable technology for targeted genome editing. Nature Reviews Molecular Cell Biology, 14(1), 49e55. https://doi.org/10.1038/nrm3486. Kawakami, K. (2007). Tol2: A versatile gene transfer vector in vertebrates. Genome Biology, 8(Suppl. 1), S7. https://doi.org/ 10.1186/gb-2007-8-s1-s7.

Kesavan, G., Chekuru, A., Machate, A., & Brand, M. (2017). CRISPR/ Cas9-Mediated zebrafish knock-in as a novel strategy to study midbrain-hindbrain boundary development. Frontiers in Neuroanatomy, 11, 52. https://doi.org/10.3389/fnana.2017.00052. Kim, Y., Kweon, J., Kim, A., Chon, J. K., Yoo, J. Y., Kim, H. J., et al. (2013). A library of TAL effector nucleases spanning the human genome. Nature Biotechnology, 31(3), 251e258. https://doi.org/ 10.1038/nbt.2517. Kimura, Y., Hisano, Y., Kawahara, A., & Higashijima, S. (2014). Efficient generation of knock-in transgenic zebrafish carrying reporter/ driver genes by CRISPR/Cas9-mediated genome engineering. Scientific Reports, 4, 6545. https://doi.org/10.1038/srep06545. Kok, F. O., Shin, M., Ni, C. W., Gupta, A., Grosse, A. S., van Impel, A., et al. (2015). Reverse genetic screening reveals poor correlation between morpholino-induced and mutant phenotypes in zebrafish. Developmental Cell, 32(1), 97e108. https://doi.org/10.1016/ j.devcel.2014.11.018. Langheinrich, U. (2003). Zebrafish: A new model on the pharmaceutical catwalk. BioEssays, 25(9), 904e912. https://doi.org/10.1002/ bies.10326. Lawson, N. D., & Wolfe, S. A. (2011). Forward and reverse genetic approaches for the analysis of vertebrate development in the zebrafish. Developmental Cell, 21(1), 48e64. https://doi.org/ 10.1016/j.devcel.2011.06.007. Lieschke, G. J., & Currie, P. D. (2007). Animal models of human disease: Zebrafish swim into view. Nature Reviews Genetics, 8(5), 353e367. https://doi.org/10.1038/nrg2091. Meng, X., Noyes, M. B., Zhu, L. J., Lawson, N. D., & Wolfe, S. A. (2008). Targeted gene inactivation in zebrafish using engineered zincfinger nucleases. Nature Biotechnology, 26(6), 695e701. https:// doi.org/10.1038/nbt1398. Meyer, A., & Van de Peer, Y. (2005). From 2R to 3R: Evidence for a fishspecific genome duplication (FSGD). BioEssays, 27(9), 937e945. https://doi.org/10.1002/bies.20293. Narayanan, A., Hill-Teran, G., Moro, A., Ristori, E., Kasper, D. M., C, A. R., et al. (2016). In vivo mutagenesis of miRNA gene families using a scalable multiplexed CRISPR/Cas9 nuclease system. Scientific Reports, 6, 32386. https://doi.org/10.1038/srep32386. Nasevicius, A., & Ekker, S. C. (2000). Effective targeted gene ‘knockdown’ in zebrafish. Nature Genetics, 26(2), 216e220. https:// doi.org/10.1038/79951. Ohno, S. (1970). Evolution by gene duplication. New York, NY: Springer. Ota, S., Taimatsu, K., Yanagi, K., Namiki, T., Ohga, R., Higashijima, S. I., et al. (2016). Functional visualization and disruption of targeted genes using CRISPR/Cas9-mediated eGFP reporter integration in zebrafish. Scientific Reports, 6, 34991. https://doi.org/10.1038/ srep34991. Postlethwait, J., Amores, A., Cresko, W., Singer, A., & Yan, Y. L. (2004). Subfunction partitioning, the teleost radiation and the annotation of the human genome. Trends in Genetics, 20(10), 481e490. https:// doi.org/10.1016/j.tig.2004.08.001. Prelich, G. (2012). Gene overexpression: Uses, mechanisms, and interpretation. Genetics, 190(3), 841e854. https://doi.org/10.1534/ genetics.111.136911. Ran, F. A., Hsu, P. D., Lin, C. Y., Gootenberg, J. S., Konermann, S., Trevino, A. E., et al. (2013). Double nicking by RNA-guided CRISPR Cas9 for enhanced genome editing specificity. Cell, 154(6), 1380e1389. https://doi.org/10.1016/j.cell.2013.08.021. Rastogi, S., & Liberles, D. A. (2005). Subfunctionalization of duplicated genes as a transition state to neofunctionalization. BMC Evolutionary Biology, 5, 28. https://doi.org/10.1186/1471-2148-5-28. Robu, M. E., Larson, J. D., Nasevicius, A., Beiraghi, S., Brenner, C., Farber, S. A., et al. (2007). p53 activation by knockdown technologies. PLoS Genetics, 3(5), e78. https://doi.org/10.1371/ journal.pgen.0030078.

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C H A P T E R

50 Zebrafish as a Platform for Genetic Screening James T. Nichols Department of Craniofacial Biology, University of Colorado Anschutz Medical Campus, Aurora, CO, United States of America

Introduction and History The mutant and its phenotype, caused by the lack of a single gene product, is a cleaner experiment than any transplantation, constriction or centrifugation could ever be. Christiane Nu¨ssleinVolhard (2012)

What is a genetic screen? Broadly defined, a genetic screen is a research technique aimed at uncovering the characteristics of an organism, its phenotype, which arises from the DNA sequences contained within that organism, its genotype. By observing phenotypes in organisms which differ when compared with “normal” or wild-type organisms, and then determining what DNA change caused the new phenotype, the function of the mutated DNA can be deduced. Most often, these changes to the DNA occur in genes. Thus, a genetic screen is a way to learn the function of genes. For example, if a mutant does not develop eyes, then one can infer that the mutated gene is required for eye formation. There are two major categories of genetic screens, known as forward and reverse. In a classical forward screen, the researcher starts with a phenotype that differs from the wild-type condition and then determines the gene that was changed. In contrast, in a reverse screen, the researcher starts with genes of interest and engineers the DNA changes to determine the phenotype that, if any, arises when the candidate genes are mutated. In zebrafish, reverse genetic approaches are enabled by recent advances in genome editing like CRISPR/Cas9, TALEN, zinc finger nucleases, and TILLING (Doyon et al., 2008; Huang et al., 2011; Hwang et al., 2013; Meng, Noyes, Zhu, Lawson, & Wolfe, 2008; Moens, Donn, Wolf-Saxon, & Ma, 2008; Rissone, Ledin, & Burgess, 2018). Moreover, transient methods like chemical treatments and gene knockdown are sometimes grouped into the reverse genetic category, although these nonheritable methods do not address how DNA relates to phenotype, and therefore, by definition, they The Zebrafish in Biomedical Research https://doi.org/10.1016/B978-0-12-812431-4.00050-6

are not genetic. While reverse genetic approaches are useful for understanding gene function, they are welldescribed in this volume and elsewhere (Lawson, 2016; Rissone et al., 2018). Here, I only consider further unbiased forward genetic screens, which remain a valuable and relevant technique even in the current era of powerful genome editing technologies. Why are zebrafish useful for screens? The similarities between humans and zebrafish may not be immediately apparent. But in fact, zebrafish development, anatomy, and genomes are all strikingly similar to other vertebrates including humans. Like all vertebrates, zebrafish develop from a single cell to a complex, integrated, multicellular animal and the cellular and molecular mechanisms of development that build these complex animals are largely similar, or conserved, across all vertebrates. During development, vertebrate organisms pass through several developmental stages and landmarks, which are shared with other vertebrates. One of these developmental stages, in particular, the pharyngula stage, is the so-called phylotypic period proposed to be the developmental stage when vertebrate morphology is most conserved between species (Duboule, 1994; Irie & Kuratani, 2011; Von Baer 1828). The outcome of development is the different anatomical features, which are also largely shared among fish and mammals. It is more unusual to find a derived character that is not shared across vertebrates than the vast number of anatomical features that are shared (Hyman, 1942). The “recipe book” guiding the development of the single cell to complex morphology is encoded in the full complement of DNA within the organism, its genome. The similarities between zebrafish and human genomes are evidenced by the discovery that approximately 70% of human genes have at least one clear zebrafish matching gene, or ortholog (Howe et al., 2013). There are also species-specific advantages for genetic screens that zebrafish enjoy compared to other animal

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model systems. Zebrafish, originally a popular aquarium pet, are very easy to keep because they are resistant to variable water quality, temperature, salinity, and pH. Zebrafish are straightforward to breed, and it is common to obtain hundreds of offspring per breeding pair each week, an asset for genetic studies. Zebrafish are translucent when young, so developing animals can be placed under the microscope to directly image tissues, cells, and molecules. Furthermore, they are externally fertilized so that all developmental stages are readily observed. These valuable characteristics allow screening for interesting changes in phenotype when fish genomes are mutated. How did zebrafish become a premier platform for genetic screening? The ability to start with an interesting phenotype and discover the causative genotype in zebrafish was made possible by pioneering scientists. In the 1960s, George Streisinger was using viruses that infect bacteria to address questions about gene function but wished to develop a vertebrate model system capable of mutational analysis of genes (Grunwald & Eisen, 2002). To this end, he began working to develop genetic methods in the zebrafish culminating with his generation of homozygous clones for the first time in a vertebrate (Streisinger, Walker, Dower, Knauber, & Singer, 1981). The now-famous, spontaneously occurring mutant “golden,” which alters pigmentation in the zebrafish was instrumental in this study and demonstrated that zebrafish could be used for mutational analyses like the forward screens. To understand the phenotype of a mutant, one must first understand the phenotype of the wild type. While Streisinger was developing genetic techniques, his colleague Charles Kimmel developed techniques for studying zebrafish development. Capitalizing on the advantages of the zebrafish, Kimmel used microscopy to observe live animals as they develop wild-type phenotypes. First developing techniques for tracking the development of a single neuron in live animals (Kimmel, 1972; Kimmel et al., 1978, 1981), followed by cell lineage analyses (Kimmel, 1989; Kimmel, Warga, & Schilling, 1990; Kimmel and Warga, 1987, 1988), and then a comprehensive description of embryonic

development (Kimmel, Ballard, Kimmel, Ullmann, & Schilling, 1995). With the tools for genetic and developmental analyses in place, the first genetic screens were carried out, uncovering induced mutations that altered the development of varied tissues (Felsenfeld, Walker, Westerfield, Kimmel, & Streisinger, 1990; Grunwald, Kimmel, Westerfield, Walker, & Streisinger, 1988; Walker & Streisinger, 1983). Meanwhile, large-scale genetic screens were revealing fantastic insight into the development of the Drosophila embryo (Nu¨sslein-Volhard & Wieschaus, 1980), motivating a similar big-screen approach in the burgeoning vertebrate model (Mullins, Hammerschmidt, Haffter, & Nu¨sslein-Volhard, 1994), thrusting the once humble aquarium pet into the limelight as a premier organism for genetic screens. What are the basic steps of a genetic screen? A genetic screen can be distilled into six steps which I will describe in detail in the remaining sections of this chapter. The simplified outline of a forward genetic screen is (1) mutagenesis, (2) recovery, (3) phenotyping, (4) genotyping, (5) validation, and (6) characterization.

Mutagenesis The goal of mutagenesis is to generate a permanent change in genomic DNA. There are several different mutagenesis options, and I will highlight the strengths and weaknesses of some commonly used mutagens below. Some variables to consider when choosing a mutagen are the type of lesion, the hazards to the researcher, and the target number of DNA lesions per genome. The number of lesions per genome is particularly important to consider because too few will result in an inefficient recovery of mutants, while too many will cause general sickness and increased chances of complex, difficult to recover mutations. N-ethyl-N-nitrosourea (ENU) is the most commonly used mutagen for the forward screens in zebrafish. This potent chemical mutagen has a high mutation rate and is known to primarily induce point mutations on a single strand (Kile & Hilton, 2005) (Fig. 50.1). Therefore, single genes are destroyed in ENU-induced mutant

FIGURE 50.1 Zebrafish mutagenesis. Chemical (ENU), radiation, and insertional mutagenesis are commonly used methods of inducing genomic DNA changes, or mutations. Each mutagen is expected to induce different types of DNA changes as indicated. V. Scientific Research

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lines, increasing the likelihood that a single candidate gene segregates with a phenotype. The primary negative to ENU mutagenesis is its toxicity; adult fish are known to jump out of tanks containing ENU to avoid exposure. Adding anesthetic to the mutagenesis solution helps to keep fish calm. At some doses, ENU is lethal to adult fish, and therefore, the concentrations must be kept low. However, the toxicity to adult fish can be overcome by repeated low-dose exposures, increasing mutagenesis while maintaining survival. Some of the dangers to humans from using ENU can be overcome by inactivating the mutagen with high pH and sodium thiosulfate (Westerfield, 1993). A second useful mutagen is an ionizing radiation which induces double-stranded DNA breaks. Doublestranded breaks promote many different types of DNA changes, such as small or large deletions, as well as chromosome rearrangements. These various changes can create useful versions of genes, known as alleles. For example, a deletion can completely remove an entire gene or several entire genes for that matter. If an entire gene is deleted, it is known as a null allele, or a version, in which, the gene is completely dead. Null alleles can lead to a complete understanding of a gene compared to a point mutation in which residual function can remain, known as a hypomorph. A complete description of different classes of alleles is presented in the characterization section below. The primary negative of irradiation as a mutagen is that, often, only a few lethal mutants are isolated; perhaps, because they are not transmitted through the germ line effectively (Mullins et al., 1994; Nusslein-Volhard & Dahm, 2002). Insertional mutagenesis is an effective method of introducing DNA changes into the zebrafish genome. Importantly, integrating foreign DNA sequences to disrupt genes fosters the rapid isolation of the DNA that was changed because such sequences act as “molecular flag” which can be easily located in the genome to determine the gene that was interrupted by the integrated foreign DNA. Moreover, experimentally useful sequences can be inserted, such as fluorescent tags to label cells or gene products or loxP sites which allow the removal of the causative DNA change (Clark et al., 2011). These sorts of reversion experiments address temporal questions about when a gene is required during development (Talbot et al., 2016). The downside of insertional mutagenesis is that there may be some sequence specificity, such that, the hotspots for integration may have increased insertion rates as opposed to uniform mutagenesis across the genome. In addition, the rate of insertional mutagenesis is lower than ENU mutagenesis. A less-efficient way of obtaining mutants is by relying on spontaneous mutation. Indeed, the first described zebrafish mutant, golden, was a spontaneous mutation propagated in the pet industry as a favorite of fish

hobbyists. Other spontaneous mutants from the pet trade include the leopard mutant, as well as longfin that has spotted and longfin phenotypes, respectively. While an inefficient way to generate mutants, interesting spontaneous mutant phenotypes can yield insight into the biology in zebrafish, just as in mouse, where the many “fancy mice” strains have been kept for centuries because of their appealing phenotype to hobbyists. Spontaneous mutations continue to occur in laboratory strains, and through careful inbreeding and husbandry techniques (Brooks & Nichols, 2017), these “fancy fish” mutants can easily be isolated when they appear. The final method that I wish to discuss does not concern a specific mutagen, but rather the background to which mutagenesis is applied. Secondary screens are a method in which the mutagen is applied to a zebrafish line already harboring a known mutation, further altering the phenotype. Some second mutations will enhance, while some will suppress the original mutant phenotype, revealing information about gene logic and circuitry. Successful secondary screens in invertebrate models demonstrate their utility in gleaning information about genetic pathways. But, thus far, there have been few secondary screens in zebrafish. This is likely because most of the characterized zebrafish mutants are lethal as homozygotes and display no phenotypes as heterozygotes. One way to circumvent this restriction is to mutagenize a homozygous viable mutant, or a heterozygote that is sensitive to further mutation as done for a chordin mutant (Kramer et al., 2002). A second clever approach to perform a secondary screen is to generate a stable transgenic rescue line allowing the uncovering of the suppressors of an otherwise lethal mutant (Bai et al., 2010).

Recovery After mutagenesis, the fish are screened to recover those with genetic mutations producing altered phenotypes. There are several different breeding strategies with advantages and disadvantages. Often, recovery strategy depends on the type of mutagenesis and what kinds of phenotypes the researcher is searching for. All recovery methods capitalize on the ability to readily breed zebrafish and propagate their genomes transgenerationally, either through natural crosses or laboratory manipulations. The traditional method for recovering mutant families is through inbreeding. In the familial inbreeding method, the first step is to cross mutagenized animals, which are mosaic for mutations in their germline, to wild-type animals, a process referred to as outcrossing. These mosaic adults are the parental generation or P0. The offspring of the P0 will produce nonmosaic carriers

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FIGURE 50.2 Mutant recovery. Careful breeding strategies are used to recover zebrafish mutants in a genetic screen. (A) Familial inbreeding requires more time and space than other methods because several generations of breeding are required, but it is not technically demanding and can identify phenotypes that manifest at all stages. (B) Haploid screening requires fewer generations, but it is only useful for screening early developmental stages and can be confounded by phenotypes that develop due to hemizygosity rather than a genetic lesion. (C) Gynogenetic diploid screening requires less time and space than familial inbreeding but is technically demanding.

of the mutation and are called the first filial generation or F1. Randomly intercrossing animals from the F1 produces the F2 generation, which is then intercrossed again and their offspring, the F3 generation, are screened for phenotypes. One-quarter of the F2 intercrosses will be between animals heterozygous for the same newly created mutation, and one-quarter of the F3 offspring from these crosses will be homozygous for the new mutations (Fig. 50.2A) (Haffter et al., 1996; NussleinVolhard & Dahm, 2002).

The family inbreeding method described above is time-consuming because of the requirement for several generations. Moreover, familial inbreeding requires substantial housing, heavily taxing facility space. Therefore, methods were developed to allow researchers to skip generations and screen families more rapidly. The simplest method to fast-forward a screen is to analyze haploid progeny. In this method, eggs from F1 females are fertilized with “dead” sperm that has had its genome destroyed by UV irradiation. The pseudo-fertilized egg

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will undergo somewhat normal development for several days, and half of these eggs will carry the new mutation and can thus present mutant phenotypes (Fig. 50.2B) (Westerfield, 1993). Haploid animals do not develop completely normally, so haploid offspring to be screened must be thoroughly scrutinized to determine if the phenotype is due to a mutation or the nonspecific problems that come from developing with only one copy of the genome instead of the usual two copies. Moreover, because the embryos will not develop past a few days, any phenotypes that arise beyond the early larval stage cannot be screened in haploid animals. To circumvent some of the problems associated with analyzing haploid individuals, the pseudo-fertilized eggs can be treated with either hydrostatic pressure or heat shock to block the second meiotic division, or the first mitotic division respectively (Fig. 50.2C), making them diploid because they now carry two copies of the genome; however, both copies are identical and come from the mother. These “gynogenetic diploids,” as well as haploid offspring are called parthenogenetic, meaning that they derive from an unfertilized egg from a single female parent. While parthenogenetic screens save time and space, they are technically more challenging. Sometimes a special screening strategy is required, dictated by the type of phenotype that is hunted. For example, screening for maternal-effect mutants, that is, phenotypes in offspring that result from a mutation in the mother requires an extra generation because the phenotype manifests in the offspring of the homozygous mutant animals. That is, the F3 homozygotes must be grown to adulthood to screen their F4 offspring for phenotypes (Dosch et al., 2004; Wagner, Dosch, Mintzer, Wiemelt, & Mullins, 2004).

Phenotyping Determining how the mutant phenotype differs from the wild type is a critical aspect of a genetic screen. A logical way to categorize phenotypes is by the developmental stage at which they manifest. Major lifestyle transitions like the onset of feeding and reproduction are rational ways to divide the zebrafish lifecycle and useful for dividing and categorizing mutant phenotypes based on when they appear (Fig. 50.3). By far, the largest class of characterized mutants are those that develop their phenotype during the embryonic and larval stage, appearing before the onset of feeding. Some mutants develop normally past the feeding stage, but manifest phenotypes in the late larva, juvenile or adult; these include many pigmentations and fin phenotypes and those that alter the larval to juvenile metamorphosis. The third class of mutants does not have obvious

phenotypes, but are sterile because of defects in their germline (Fig. 50.3). Finding a clean mutant phenotype requires an effective way to visualize and document the change in appearance. There are many ways of observing and phenotyping zebrafish mutants. The simplest method is observing the live, unstained organism with the naked eye for obvious phenotypes like those in the fancy fish trade (Fig. 50.3). Live unstained organisms can also be observed with microscopy-based methods, which reveal much more detail allowing a more comprehensive assessment of any mutant phenotypes. It is worth noting that the developing embryos are simply beautiful under the microscope, and phenotypes can be striking. Some phenotypes cannot be observed without first staining the animal. There are various methods for labeling cells with histological stains revealing anatomical features and tissues that cannot otherwise be seen. Molecular changes can also be observed when searching for phenotypes. For example, changes in gene expression patterns can be visualized with the RNA in situ hybridization technique, and proteins can be labeled with antibodies for visual and biochemical analysis. The advent of transgenes allows a dynamic look at phenotypes where traits like cell behavior and temporal changes in gene expression through development can be recorded from live animals. Sensory mutants are unresponsive to stimulation like touch and sound, and behavioral tests can assay cognitive ability. Careful phenotyping is important, and sometimes very subtle phenotypes can be found that will later turn out to be very interesting. Subtle phenotypes can later become more striking through selective breeding to make the phenotype more severe, or by recovering another more severe allele (Brooks & Nichols, 2017; Miller et al., 2007a). To harness the power of a genetic screen in order to determine gene function, the researcher must first start with a good clean phenotype. A robust phenotype is the biggest advantage of a forward screen, and great care should be dedicated to phenotyping.

Genotyping Genotyping is the determination of what DNA alteration is responsible for the altered phenotype. Often when screens are focused on specific tissues, many mutants will have similar phenotypes. The first step to genotyping these mutants is to determine if any of the mutants are disrupted in the same gene. A simple “complementation test” achieves this. Two mutants are said to complement each other if their hybrid offspring do not have the phenotype that is present within each

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FIGURE 50.3 Phenotyping. Genetic screens can identify phenotypes that manifest at different life stages. Wild-type phenotypes and examples of mutant phenotypes observable in live unstained animals are illustrated. Reproductive phenotypes images reproduced from Dosch, R., et al. (2004). Maternal control of vertebrate development before the midblastula transition: Mutants from the zebrafish I. Developmental Cell 6, 771e780 and embryonic and larval phenotypes images from Nichols, J. T., Pan, L., Moens, C. B., Kimmel, C. B. (2013). barx1 represses joints and promotes cartilage in the craniofacial skeleton. Development 140, 2765e2775 doi:10.1242/dev.090639.

mutant line. Therefore, simply outcrossing mutant lines to each other and observing the mutant phenotype informs the researcher that the same gene is disrupted in the two lines. This is known as failure to complement and allows the researcher to place mutants into complementation groups that likely, all are mutated in the same gene (Fig. 50.4). While, generally, very useful, this rule is not infallible as there can be nonallelic failure to complement when two different mutagenized genes can present a phenotype as “trans heterozygotes” and technically fail to complement. However, these cases are rare. To determine the precise DNA change in mutants, researchers take advantage of the fact that the DNA that the fish pass to the offspring are different than their own because their germ cells “mix-up” their genomes

when they divide during meiosis. When this mix-up, called recombination, occurs, any given two parts of the genome are more or less likely to stick together depending on how close together they are on the DNA strands. If they stick together, they are said to be linked. Genes that are very close together are rarely broken up and are said to be tightly linked. Because we know the location of many genomic landmarks, linkage analyses can give rough information about where in the genome the mutation is located. That is, by looking for genomic landmarks that are inherited in the next generation alongside the mutation, the researcher can narrow down, or map, the location of the mutation. Once a region or interval of the genome that contains the mutation is known, candidates in the area can be sequenced and, hopefully, a lesion can be discovered that

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Characterization

FIGURE 50.4

Complementation. A complementation test is an easy way to determine if two mutant lines with similar phenotypes harbor mutations in the same gene. If crossing two mutant lines to each other yield phenotypically mutant offspring, the mutants “fail to complement” and are likely caused by mutations in the same gene. If crossing the two mutant lines yields only wild-type offspring, the mutants “complement” each other, suggesting different genes are mutated in each line.

segregates with the mutant phenotype. Sophisticated methods of tracking genomic landmarks continue to be developed including sequencing all the genes in an animal or even the entire genome (Bowen, Henke, Siegfried, Warman, & Harris, 2012; Miller et al., 2007b, 2013; Obholzer et al., 2012; Stickney et al., 2002).

Validation Once a candidate mutation has isolated the causality of the mutation needs to be demonstrated. There are several methods of validation that are often used in concert. Multiple noncomplementing alleles with mutations in the same gene are perhaps the most convincing evidence the mutations are causative, but complementation is only useful for homozygous recessive alleles.

A common feature of many mutants is loss of the mRNA that the candidate gene encodes because mutated gene products that are not translated into protein properly are targeted for destruction through a cell-safety mechanism to remove mutated gene products called nonsense-mediated decay. If a candidate mutated gene has lower levels of mRNA in mutants, this can be evidence that the mutation is causative. Similarly, if the protein product of the candidate gene is somehow changed, either quantity or size or localization, further evidence has been obtained. However, this protein work requires a way to detect the protein, like an antibody, which unfortunately is not available for many gene products. Another convincing validation is to rescue the mutation by adding back either wild-type DNA sequences or mRNA from the suspected mutated gene. Finally, injecting an antisense morpholino (Stainier et al., 2017) which blocks either translation or splicing of mRNA gene products can phenocopy, or produce the same phenotype as the mutant, bolstering the case that a genetic lesion is causative. It is worth mentioning that morpholino phenotypes in isolation, that is, without an accompanying mutant, must be carefully considered as the field has come to realize that morpholino induced phenotypes often do not match those of the genetic mutant and can be misleading (Stainier et al., 2017). Nevertheless, they can provide supplementary evidence that a mutation is causative when mutant and morpholino phenotypes match. Researchers, to convincingly validate that a mutation is causative, should aim to satisfy several of the above validation methods in order to build a complete case that can be trusted.

Characterization The specific version of the gene that is mutated is called the allele. Alleles are assigned a letter code to designate the institution that generated the genomic feature, followed by a unique identifying number. For example, alleles generated the University of Colorado are denoted with co followed by a series of digits that are only assigned to a singular specific DNA sequence. For more nomenclature information, see The Zebrafish Information Network (ZFIN) nomenclature page: https://wiki.zfin.org/display/general/ZFINþZebrafish þNomenclatureþGuidelines. The traditional way to characterize mutant alleles is by the rules of Muller’s Morphs (Muller, 1932). According to this system, mutants come in four types or morphs. (1) Amorphs that are a complete loss of function or null. (2) Hypomorphs are a partial or weak loss of function. (3) Hypermorphs increase normal function and (3) Antimorphs oppose normal gene activity. Antimorphs are sometimes called dominant negatives, but antimorphs can also be recessive

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(Nichols et al., 2016; Sijacic, Wang, & Liu, 2011). There are established rules for interpreting what type of morph a mutant is, but they often rely on comparing multiple alleles within a single complementation group, which is not always possible. Up until this point, we have focused on mutations that occur within genes, coding mutations. But vast swaths of the genome that do not code for mRNA or protein are also important, and it is likely many unidentified mutants are in noncoding regions that act as regulatory regions. There are few noncoding mutants described in zebrafish (Schauerte et al., 1998; Sepich, Wegner, O’Shea, & Westerfield, 1998) because it can be difficult to assign noncoding genomic changes to a particular gene. As we learn more about gene regulation and noncoding DNA, it is likely more mutants in noncoding DNA will materialize in the zebrafish literature.

Concluding Remarks One outstanding question regarding zebrafish genetic screens is, are zebrafish screens saturated? A screen is said to be saturated if all the genes involved in a biological process are found. If no new mutations come from the genetic screen, only different alleles of the same genes, the screen is approaching saturation. Interestingly, with the rapid generation of CRISPR/Cas9 based reverse genetic mutants, we are learning that many single-gene mutants have no overt phenotype but when multiple family members are mutagenized a phenotype becomes apparent (Askary et al., 2017; Barske et al., 2016). These results suggest that even as screens approach saturation, we will still not know all the genes that are involved in a process because of genetic redundancy, or multiple genes doing the same job. These results motivate designing secondary screens in CRISPR/Cas9 reverse genetic backgrounds lacking overt phenotypes. This approach is predicted to uncover new complementation groups that might not be found by mutagenizing wild types. Amazing advancements in genome sequencing technologies have provided the full “ingredients list” for the DNA recipe book that guides development in many organisms, including zebrafish. But much work remains, the list of ingredients is not enough, and the challenge now is to understand how the ingredients work in concert to build a complex organism. That is, we need to learn about gene function. Staring with a clean mutant phenotype and working to find what genetic change causes that phenotype remains one of the most powerful ways to determine gene function. Clearly, there is still an important place in zebrafish biomedical research for unbiased forward genetic screens.

References Askary, A., et al. (2017). Genome-wide analysis of facial skeletal regionalization in zebrafish. Development, 144, 2994e3005. Bai, X., et al. (2010). TIF1g controls erythroid cell fate by regulating transcription elongation. Cell, 142, 133e143. Barske, L., et al. (2016). Competition between Jagged-Notch and Endothelin1 signaling selectively restricts cartilage formation in the zebrafish upper face. PLoS Genetics, 12, e1005967. Bowen, M. E., Henke, K., Siegfried, K. R., Warman, M. L., & Harris, M. P. (2012). Efficient mapping and cloning of mutations in zebrafish by low-coverage whole-genome sequencing. Genetics, 190, 1017e1024. Brooks, E., & Nichols, J. (2017). Shifting zebrafish lethal skeletal mutant penetrance by progeny testing. JoVE (Journal of Visualized Experiments), 127, e56200. Clark, K. J., et al. (2011). In vivo protein trapping produces a functional expression codex of the vertebrate proteome. Nature Methods, 8, 506e512. Dosch, R., et al. (2004). Maternal control of vertebrate development before the midblastula transition: Mutants from the zebrafish I. Developmental Cell, 6, 771e780. Doyon, Y., et al. (2008). Heritable targeted gene disruption in zebrafish using designed zinc-finger nucleases. Nature Biotechnology, 26, 702e708. Duboule, D. (1994). Temporal colinearity and the phylotypic progression: A basis for the stability of a vertebrate Bauplan and the evolution of morphologies through heterochrony. Development, 135e142. Felsenfeld, A. L., Walker, C., Westerfield, M., Kimmel, C., & Streisinger, G. (1990). Mutations affecting skeletal muscle myofibril structure in the zebrafish. Development, 108, 443e459. Grunwald, D. J., & Eisen, J. S. (2002). Headwaters of the zebrafishd emergence of a new model vertebrate. Nature Reviews Genetics, 3, 717e724. Grunwald, D. J., Kimmel, C. B., Westerfield, M., Walker, C., & Streisinger, G. (1988). A neural degeneration mutation that spares primary neurons in the zebrafish. Developmental Biology, 126, 115e128, 0012-1606(88)90245-X. Haffter, P., et al. (1996). The identification of genes with unique and essential functions in the development of the zebrafish, Danio rerio. Development, 123, 1e36. Howe, K., et al. (2013). The zebrafish reference genome sequence and its relationship to the human genome. Nature, 496, 498e503. Huang, P., et al. (2011). Heritable gene targeting in zebrafish using customized TALENs. Nature Biotechnology, 29, 699e700. Hwang, W. Y., et al. (2013). Efficient genome editing in zebrafish using a CRISPR-Cas system. Nature Biotechnology, 31, 227e229. Hyman, L. (1942). Comparative vertebrate anatomy. Universtiy of Chicago Press. Irie, N., & Kuratani, S. (2011). Comparative transcriptome analysis reveals vertebrate phylotypic period during organogenesis. Nature Communications, 2, 248. Kile, B. T., & Hilton, D. J. (2005). The art and design of genetic screens: Mouse. Nature Reviews Genetics, 6, 557e567. Kimmel, C. B. (1972). Mauthner axons in living fish larvae. Developmental Biology, 27, 272e275. Kimmel, C. B. (1989). Genetics and early development of zebrafish. Trends in Genetics, 5, 283e288. Kimmel, C. B., Ballard, W. W., Kimmel, S. R., Ullmann, B., & Schilling, T. F. (1995). Stages of embryonic development of the zebrafish. Developmental Dynamics, 203, 253e310. https://doi.org/ 10.1002/aja.1002030302. Kimmel, C. B., Sessions, S. K., & Kimmel, R. J. (1978). Radiosensitivity and time of origin of Mauthner neuron in the zebra fish. Developmental Biology, 62, 526e529.

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51 Zebrafish as a Platform for Drug Screening Tejia Zhang, Randall T. Peterson College of Pharmacy, University of Utah, Salt Lake City, UT, United States of America

Introduction Since the first demonstrated use of live zebrafish for chemical screening in 96-well format in 2000 (Peterson, Link, Dowling, & Schreiber, 2000), zebrafish chemical screens have expanded significantly to encompass a wide range of fish models, targeted pathways and phenotypic readouts (Rennekamp & Peterson, 2015). In the context of chemical screening, the zebrafish offers unique advantages, including rapid development, high fecundity, and small size, and optical transparency at the larval stage. These attributes allow highthroughput screening with larvae in 96- or 384-well plate format, and imaging-based readouts (Rennekamp & Peterson, 2015). In this chapter, we review chemical screening in the zebrafish model organism; we begin with a statistical overview of zebrafish chemical screens conducted since 2000, followed by more detailed discussions that address specific screens based on their areas of focus. We conclude by highlighting a few screens with significant clinical implications and bring to attention some of the current limitations and novel approaches toward addressing these limitations for future studies.

Zebrafish Chemical Screening, a Statistical Overview A survey of the existing literature between 2000 and June 2017 (Ncbi Resource Coordinators, 2017) uncovered 114 studies involving zebrafish chemical screening; while we highlight a number of these studies in this chapter, we do not claim this list to be exhaustive and apologize for any omissions made. Of the 114 studies identified, 56 focus on specific tissues to identify modulators of organogenesis or rescuers of tissue-specific pathologies (Fig. 51.1A and D) (Alvarez et al., 2009; Buckley et al., 2010; Chiu, Cunningham, Raible, Rubel, & Ou, 2008; Crawford et al., 2011; de Vrieze et al., 2015; The Zebrafish in Biomedical Research https://doi.org/10.1016/B978-0-12-812431-4.00051-8

Fleming, Sato, & Goldsmith, 2005; Hirose, Simon, & Ou, 2011; Huang, Lindgren, Wu, Liu, & Lin, 2012; Kitambi, McCulloch, Peterson, & Malicki, 2009; Lam et al., 2012; Lake, Tusheva, Graham, & Heuckeroth, 2013; Liang et al., 2015; Oppedal & Goldsmith, 2010; Papakyriakou et al., 2014; Sun, Dong, Khodabakhsh, Chatterjee, & Guo, 2012; Tran et al., 2007; Wang et al., 2010) (Asimaki et al., 2014; Choi et al., 2013; Coffin, Williamson, Mamiya, Raible, & Rubel, 2013; Gallardo et al., 2015; Kannan & Vincent, 2012; Leet et al., 2014; Liu et al., 2014; Milan, Peterson, Ruskin, Peterson, & MacRae, 2003; Namdaran, Reinhart, Owens, Raible, & Rubel, 2012; Ni et al., 2011; North et al., 2007; Ou et al., 2009; Owens et al., 2008; Peal et al., 2011; Shimizu et al., 2015; Tang, Xie, & Feng, 2015; Thomas et al., 2015; Vlasits, Simon, Raible, Rubel, & Owens, 2012; Yozzo, Isales, Raftery, & Volz, 2013; Yang et al., 2015), (Arulmozhivarman et al., 2016; Astin et al., 2014; Cao et al., 2009; Colanesi et al., 2012; de Groh et al., 2010; Garnaas et al., 2012; Hultman, Scott, & Johnson, 2008; Kawahara et al., 2011, 2014; Ko et al., 2016; Shafizadeh, Peterson, & Lin, 2004; Tamplin et al., 2015; Waugh et al., 2014; Westhoff et al., 2013; White et al., 2011; Yeh et al., 2009; Yeh & Munson, 2010; Yin, Evason, Maher, & Stainier, 2012; Zhen et al., 2013). Twelve studies use zebrafish in metabolic screens (Fig. 51.1A,B). The emergence of behaviors, such as eye movement, light response, and food seeking within the first week of development has enabled high-throughput behavioral screening in zebrafish larvae (Kalueff et al., 2013), and 10 such studies were identified by our search (Fig. 51.1B). Several screens have also been designed to target the inflammatory response (Hall et al., 2014; Liu et al., 2013; Robertson et al., 2014; Wang et al., 2014; Wittmann et al., 2015; Ye et al., 2017), toxin metabolism (Dimri et al., 2015; Jin et al., 2013; Legler et al., 2011; Nath et al., 2013; North et al., 2010; Padilla et al., 2012), and a diverse set of signaling pathways (Fig. 51.1A and C) (Gebruers et al., 2013; Hao et al., 2013; Le et al., 2013; Molina et al., 2009; Molina, Watkins, & Tsang, 2007;

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FIGURE 51.1 17 years of zebrafish chemical screening. (A) Zebrafish chemical screens by category, 2000e2017. (B) Metabolism-based zebrafish chemical screens. (C) Zebrafish chemical screens focusing on signaling pathways. (D) Tissue-specific zebrafish chemical screens.

Robertson et al., 2014; Torregroza, Evans, & Das, 2009; Weger, Weger, Nusser, Brenner-Weiss, & Dickmeis, 2012; Williams et al., 2015; Yu et al, 2008a, 2008b). Fifty five studies rely on wildtype zebrafish (Astin et al., 2014; Bouwmeester et al., 2016; Chiu et al., 2008; Coffin et al., 2013; Colanesi et al., 2012; Challal et al., 2014; Das, McCartin, Liu, Peterson, & Evans, 2010; de Groh et al., 2010; Fleming et al., 2005; Garnaas et al., 2012; Hirose et al., 2011; Kannan & Vincent, 2012; Kokel et al., 2010; Laggner et al., 2012; Lake et al., 2013; Li, Huang, Huang, Du, & Huang, 2012; Long et al., 2014; Lu et al., 2013; Maximino et al., 2014; Mendelsohn et al., 2006; Milan et al., 2003; Morash et al., 2011; Nath et al., 2015; North et al., 2007; Oppedal & Goldsmith, 2010; Ou et al., 2009; Owens et al., 2008; Peterson et al., 2000; Rahn et al., 2014; Rennekamp et al., 2016; Rihel et al., 2010; Sandoval et al., 2013; Sun et al., 2012; Tamplin et al., 2015; Tang et al., 2015; Thomas et al., 2015; Thorsteinson et al., 2009; Vlasits et al., 2012; Wang et al.,

2014; White et al., 2011; Zhen et al., 2013), 50 use transgenic lines (Alvarez et al., 2009; Andersson et al., 2012; Arulmozhivarman et al., 2016; Asimaki et al., 2014; Buckley et al., 2010; Choi et al., 2013; Crawford et al., 2011; de Vrieze et al., 2015; Gallardo et al., 2015; Gut et al., 2013; Gutierrez et al., 2014; Hall et al., 2014; Huang et al., 2012; Jimenez et al., 2016; Kitambi et al., 2009; Ko et al., 2016; Lam et al., 2012; Le et al., 2013; Leet et al., 2014; Li et al., 2015; Li et al., 2016; Li, Page-McCaw, & Chen, 2016; Liang et al., 2015; Liu et al., 2013; Liu et al., 2014; Molina et al., 2007; Molina et al., 2009; Namdaran et al., 2012; Ni et al., 2011; North et al., 2010; Papakyriakou et al., 2014; Raftery, Isales, Yozzo, & Volz, 2014; Robertson et al., 2014; Ridges et al., 2012; Rovira et al., 2011; Shafizadeh et al., 2004; Tran et al., 2007; Tsuji et al., 2014; Wang et al., 2010; Wang et al., 2014; Wang et al., 2015; Weger et al., 2012; Westhoff et al., 2013; Wittmann et al., 2015; Ye et al., 2017; Yeh & Munson, 2010; Yeh et al., 2009; Yin et al., 2012; Yozzo et al., 2013 ), and nine (Baraban, Dinday,

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Zebrafish Chemical Screening, a Statistical Overview

& Hortopan, 2013; Cao et al., 2009; Hultman et al., 2008; Kawahara et al., 2011; Kawahara et al., 2014; Paik, de Jong, Pugach, Opara, & Zon, 2010; Peal et al., 2011; Shimizu et al., 2015; Waugh et al., 2014) use mutant models (Fig. 51.2A), with the majority of studies focusing on larvae before eight days-post-fertilization (dpf), and four studies on adult zebrafish (Fig. 51.2B). (de Vrieze et al., 2015; Li et al., 2015; Maximino et al., 2014; Oppedal & Goldsmith, 2010). The majority of transgenic zebrafish encode fluorescent proteins (with GFP and its variants being the most common) for cell type-specific visualization: nine out of the nine angiogenesis screens use fli1or flik1-driven reporter lines that fluorescently label the entire vasculature (Alvarez et al., 2009; Crawford et al., 2011; Huang et al., 2012; Kitambi et al., 2009; Lam et al., 2012; Liang et al., 2015; Papakyriakou et al., 2014; Tran et al., 2007; Wang et al., 2010); and five out of the six

FIGURE 51.2 Zebrafish in chemical screening. (A) Types of zebrafish models for chemical screening, 2000e2017. (B) Age distribution of zebrafish screened. Studies with unclear or undesignated fish age were excluded; a few studies included screens in multiple age groups. t ¼ age of zebrafish.

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inflammation screens use neutrophil-specific reporter lines with lyz- or mpx-driven fluorescence (Hall et al., 2014; Liu et al., 2013; Robertson et al., 2014; Ye et al., 2017; Wang et al., 2014). Three studies rely on luminescence (luciferase)-based reporters (Asimaki et al., 2014; de Vrieze et al., 2015; Weger et al., 2012). Five studies use zebrafish carrying human oncogenes or additional human mutations of interest (Asimaki et al., 2014; Gutierrez et al., 2014; Le et al., 2013; Yeh & Munson, 2010; Yeh et al., 2009), and two studies use nitroreductase-expressing systems for tissue-specific ablation (Andersson et al., 2012; Ko et al., 2016), with or without additional fluorescent or luminescent reporters to facilitate tissue visualization. All nine mutant zebrafish lines used in screening were identified from forward genetic screens (Baraban et al., 2013; Cao et al., 2009; Hultman et al., 2008; Kawahara et al., 2011; Kawahara et al., 2014; Paik et al., 2010; Peal et al., 2011; Shimizu et al., 2015; Waugh et al., 2014). While forward genetics has contributed significantly toward the availability of zebrafish mutants for chemical screening, recent advancements in genome editing technologies, such as TALENS (Hwang, Peterson, & Yeh, 2014; Zu et al., 2013) and CRISPR-Cas9 (Gagnon et al., 2014; Liu et al., 2017) in zebrafish point toward a likely rise in targeted model generation, and pave the way for future chemical screens in new models. Approximately half of the 114 screens use commercially available libraries, while another 20% rely on academic and government sources (Fig. 51.3A). Approximately 6% of screens are performed with extracts from plants, fungi or soil rather than purified compounds (Fig. 51.3A). 1000e2000 is the most widely used compound range, with the largest screens falling within the 10,000e30,000 range (Fig. 51.3B). A number of studies were conducted with a relatively small number of compounds (