Zebrafish as a Model to Understand Vertebrate Development

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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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Zebrafish Research Advances Our Understanding of the Mechanisms Governing Development

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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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.

References diIorio, P. J., Moss, J. B., Sbrogna, J. L., Karlstrom, R. O., & Moss, L. G. (2002). Sonic hedgehog is required early in pancreatic islet development. Developmental Biology, 244, 75e84. Ahn, D., & Ho, R. K. (2008). Tri-phasic expression of posterior hox genes during development of pectoral fins in zebrafish: Implications for the evolution of vertebrate paired appendages. Developmental Biology, 322, 220e233. Akimenko, M. A., & Ekker, M. (1995). Anterior duplication of the Sonic hedgehog expression pattern in the pectoral fin buds of zebrafish treated with retinoic acid. Developmental Biology, 170, 243e247. Alexandre, D., Clarke, J. D., Oxtoby, E., Yan, Y. L., Jowett, T., & Holder, N. (1996). Ectopic expression of Hoxa-1 in the zebrafish alters the fate of the mandibular arch neural crest and phenocopies a retinoic acid-induced phenotype. Development (Cambridge, England), 122, 735e746. Almeida, A. D., Boije, H., Chow, R. W., He, J., Tham, J., Suzuki, S. C., et al. (2014). Spectrum of fates: A new approach to the study of the developing zebrafish retina. Development (Cambridge, England), 141, 1971e1980. Alsina, B., & Whitfield, T. T. (2017). Sculpting the labyrinth: Morphogenesis of the developing inner ear. Seminars in Cell and Developmental Biology, 65, 47e59. Amacher, S. L., Draper, B. W., Summers, B. R., & Kimmel, C. B. (2002). The zebrafish T-box genes no tail and spadetail are required for development of trunk and tail mesoderm and medial floor plate. Development (Cambridge, England), 129, 3311e3323. Amack, J. D., Wang, X., & Yost, H. J. (2007). Two T-box genes play independent and cooperative roles to regulate morphogenesis of ciliated Kupffer’s vesicle in zebrafish. Developmental Biology, 310, 196e210. Aman, A., Nguyen, M., & Piotrowski, T. (2011). Wnt/b-catenin dependent cell proliferation underlies segmented lateral line morphogenesis. Developmental Biology, 349, 470e482. Aman, A., & Piotrowski, T. (2008). Wnt/beta-catenin and Fgf signaling control collective cell migration by restricting chemokine receptor expression. Developmental Cell, 15, 749e761. Amsterdam, A., Nissen, R. M., Sun, Z., Swindell, E. C., Farrington, S., & Hopkins, N. (2004). Identification of 315 genes essential for early zebrafish development. Proceedings of the National Academy of Sciences of the United States of America, 101, 12792e12797. Anderson, K. V., & Ingham, P. W. (2003). The transformation of the model organism: A decade of developmental genetics. Nature Genetics, 33(Suppl. l), 285e293. Anderson, C., & Stern, C. D. (2016). Organizers in development. Current Topics in Developmental Biology, 117, 435e454. Annona, G., Holland, N. D., & D’Aniello, S. (2015). Evolution of the notochord. EvoDevo, 6, 30. Antonellis, A., Huynh, J. L., Lee-Lin, S.-Q., Vinton, R. M., Renaud, G., Loftus, S. K., et al. (2008). Identification of neural crest and glial enhancers at the mouse Sox10 locus through transgenesis in zebrafish. PLoS Genetics, 4, e1000174. Anzenberger, U., Bit-Avragim, N., Rohr, S., Rudolph, F., Dehmel, B., Willnow, T. E., et al. (2006). Elucidation of megalin/LRP2dependent endocytic transport processes in the larval zebrafish pronephros. Journal of Cell Science, 119, 2127e2137. Aulehla, A., & Pourquie´, O. (2010). Signaling gradients during paraxial mesoderm development. Cold Spring Harbor Perspectives in Biology, 2, a000869. Bakkers, J. (2011). Zebrafish as a model to study cardiac development and human cardiac disease. Cardiovascular Research, 91, 279e288. Bakkers, J., Verhoeven, M. C., & Abdelilah-Seyfried, S. (2009). Shaping the zebrafish heart: From left-right axis specification to epithelial tissue morphogenesis. Developmental Biology, 330, 213e220.

V. Scientific Research

References

Barresi, M. J., D’Angelo, J. A., Herna´ndez, L. P., & Devoto, S. H. (2001). Distinct mechanisms regulate slow-muscle development. Current Biology, 11, 1432e1438. Barresi, M. J. F., Hutson, L. D., Chien, C.-B., & Karlstrom, R. O. (2005). Hedgehog regulated Slit expression determines commissure and glial cell position in the zebrafish forebrain. Development (Cambridge, England), 132, 3643e3656. Barresi, M. J., Stickney, H. L., & Devoto, S. H. (2000). The zebrafish slow-muscle-omitted gene product is required for Hedgehog signal transduction and the development of slow muscle identity. Development (Cambridge, England), 127, 2189e2199. Barrios, A., Poole, R. J., Durbin, L., Brennan, C., Holder, N., & Wilson, S. W. (2003). Eph/Ephrin signaling regulates the mesenchymal-to-epithelial transition of the paraxial mesoderm during somite morphogenesis. Current Biology, 13, 1571e1582. Bazzini, A. A., Lee, M. T., & Giraldez, A. J. (2012). Ribosome profiling shows that miR-430 reduces translation before causing mRNA decay in zebrafish. Science, 336, 233e237. Behrndt, M., Salbreux, G., Campinho, P., Hauschild, R., Oswald, F., Roensch, J., et al. (2012). Forces driving epithelial spreading in zebrafish gastrulation. Science, 338, 257e260. Be´naze´raf, B., & Pourquie´, O. (2013). Formation and segmentation of the vertebrate body axis. Annual Review of Cell and Developmental Biology, 29, 1e26. Berndt, J. D., & Halloran, M. C. (2006). Semaphorin 3d promotes cell proliferation and neural crest cell development downstream of TCF in the zebrafish hindbrain. Development (Cambridge, England), 133, 3983e3992. Bernhardt, R. R., Goerlinger, S., Roos, M., & Schachner, M. (1998). Anterior-posterior subdivision of the somite in embryonic zebrafish: Implications for motor axon guidance. Developmental Dynamics: An Official Publication of the American Association of Anatomists, 213, 334e347. Bertrand, J. Y., Chi, N. C., Santoso, B., Teng, S., Stainier, D. Y. R., & Traver, D. (2010). Haematopoietic stem cells derive directly from aortic endothelium during development. Nature, 464, 108e111. Biemar, F., Argenton, F., Schmidtke, R., Epperlein, S., Peers, B., & Driever, W. (2001). Pancreas development in zebrafish: Early dispersed appearance of endocrine hormone expressing cells and their convergence to form the definitive islet. Developmental Biology, 230, 189e203. Biswas, S., Emond, M. R., & Jontes, J. D. (2010). Protocadherin-19 and N-cadherin interact to control cell movements during anterior neurulation. The Journal of Cell Biology, 191, 1029e1041. Blaser, H., Eisenbeiss, S., Neumann, M., Reichman-Fried, M., Thisse, B., Thisse, C., et al. (2005). Transition from non-motile behaviour to directed migration during early PGC development in zebrafish. Journal of Cell Science, 118, 4027e4038. Boswell, C. W., & Ciruna, B. (2017). Understanding idiopathic scoliosis: A new zebrafish school of thought. Trends in Genetics, 33, 183e196. Braat, A. K., Zandbergen, T., van de Water, S., Goos, H. J., & Zivkovic, D. (1999). Characterization of zebrafish primordial germ cells: Morphology and early distribution of vasa RNA. Developmental Dynamics: An Official Publication of the American Association of Anatomists, 216, 153e167. Bruce, A. E. E. (2016). Zebrafish epiboly: Spreading thin over the yolk. Developmental Dynamics: An Official Publication of the American Association of Anatomists, 245, 244e258. Buckley, C. E., Ren, X., Ward, L. C., Girdler, G. C., Araya, C., Green, M. J., et al. (2013). Mirror-symmetric microtubule assembly and cell interactions drive lumen formation in the zebrafish neural rod. The EMBO Journal, 32, 30e44. Burke, A. C., Nelson, C. E., Morgan, B. A., & Tabin, C. (1995). Hox genes and the evolution of vertebrate axial morphology. Development (Cambridge, England), 121, 333e346.

583

Bussmann, J., Bakkers, J., & Schulte-Merker, S. (2007). Early endocardial morphogenesis requires Scl/Tal1. PLoS Genetics, 3, e140. Carmona-Fontaine, C., Theveneau, E., Tzekou, A., Tada, M., Woods, M., Page, K. M., et al. (2011). Complement fragment C3a controls mutual cell attraction during collective cell migration. Developmental Cell, 21, 1026e1037. Carroll, K. J., & North, T. E. (2014). Oceans of opportunity: Exploring vertebrate hematopoiesis in zebrafish. Experimental Hematology, 42, 684e696. Cearns, M. D., Escuin, S., Alexandre, P., Greene, N. D. E., & Copp, A. J. (2016). Microtubules, polarity and vertebrate neural tube morphogenesis. Journal of Anatomy, 229, 63e74. Chandrasekhar, A. (2004). Turning heads: Development of vertebrate branchiomotor neurons. Developmental Dynamics: An Official Publication of the American Association of Anatomists, 229, 143e161. Chang, W.-J., & Hwang, P.-P. (2011). Development of zebrafish epidermis. Birth Defects Research Part C: Embryo TodayeReviews, 93, 205e214. Charvet, B., Guiraud, A., Malbouyres, M., Zwolanek, D., Guillon, E., Bretaud, S., et al. (2013). Knockdown of col22a1 gene in zebrafish induces a muscular dystrophy by disruption of the myotendinous junction. Development (Cambridge, England), 140, 4602e4613. Chen, W., Burgess, S., & Hopkins, N. (2001). Analysis of the zebrafish smoothened mutant reveals conserved and divergent functions of hedgehog activity. Development (Cambridge, England), 128, 2385e2396. Chieffo, C., Garvey, N., Gong, W., Roe, B., Zhang, G., Silver, L., et al. (1997). Isolation and characterization of a gene from the DiGeorge chromosomal region homologous to the mouse Tbx1 gene. Genomics, 43, 267e277. Choe, C. P., & Crump, J. G. (2014). Tbx1 controls the morphogenesis of pharyngeal pouch epithelia through mesodermal Wnt11r and Fgf8a. Development (Cambridge, England), 141, 3583e3593. Choksi, S. P., Babu, D., Lau, D., Yu, X., & Roy, S. (2014). Systematic discovery of novel ciliary genes through functional genomics in the zebrafish. Development (Cambridge, England), 141, 3410e3419. Chou, M.-Y., Hsiao, C.-D., Chen, S.-C., Chen, I.-W., Liu, S.-T., & Hwang, P.-P. (2008). Effects of hypothermia on gene expression in zebrafish gills: Upregulation in differentiation and function of ionocytes as compensatory responses. Journal of Experimental Biology, 211, 3077e3084. Collazo, A. (2000). Developmental variation, homology, and the pharyngula stage. Systematic Biology, 49, 3e18. Concha, M. L., & Adams, R. J. (1998). Oriented cell divisions and cellular morphogenesis in the zebrafish gastrula and neurula: A time-lapse analysis. Development (Cambridge, England), 125, 983e994. Cooper, M. S., & D’amico, L. A. (1996). A cluster of noninvoluting endocytic cells at the margin of the zebrafish blastoderm marks the site of embryonic shield formation. Developmental Biology, 180, 184e198. Corte´s, F., Daggett, D., Bryson-Richardson, R. J., Neyt, C., Maule, J., Gautier, P., et al. (2003). Cadherin-mediated differential cell adhesion controls slow muscle cell migration in the developing zebrafish myotome. Developmental Cell, 5, 865e876. Daggett, D. F., Domingo, C. R., Currie, P. D., & Amacher, S. L. (2007). Control of morphogenetic cell movements in the early zebrafish myotome. Developmental Biology, 309, 169e179. Dalgin, G., Ward, A. B., Hao, L. T., Beattie, C. E., Nechiporuk, A., & Prince, V. E. (2011). Zebrafish mnx1 controls cell fate choice in the developing endocrine pancreas. Development (Cambridge, England), 138, 4597e4608. Dalle Nogare, D., & Chitnis, A. B. (2017). A framework for understanding morphogenesis and migration of the zebrafish posterior lateral line primordium. Mechanisms of Development, 148, 69e78.

V. Scientific Research

584

45. Zebrafish as a Model to Understand Vertebrate Development

Dalle Nogare, D., Somers, K., Rao, S., Matsuda, M., ReichmanFried, M., Raz, E., et al. (2014). Leading and trailing cells cooperate in collective migration of the zebrafish posterior lateral line primordium. Development (Cambridge, England), 141, 3188e3196. Dambly-Chaudie`re, C., Cubedo, N., & Ghysen, A. (2007). Control of cell migration in the development of the posterior lateral line: Antagonistic interactions between the chemokine receptors CXCR4 and CXCR7/RDC1. BMC Developmental Biology, 7, 23. Dekens, M. P. S., Pelegri, F. J., Maischein, H.-M., & Nu¨sslein-Volhard, C. (2003). The maternal-effect gene futile cycle is essential for pronuclear congression and mitotic spindle assembly in the zebrafish zygote. Development (Cambridge, England), 130, 3907e3916. Detrich, H. W., Kieran, M. W., Chan, F. Y., Barone, L. M., Yee, K., Rundstadler, J. A., et al. (1995). Intraembryonic hematopoietic cell migration during vertebrate development. Proceedings of the National Academy of Sciences of the United States of America, 92, 10713e10717. Detrick, R. J., Dickey, D., & Kintner, C. R. (1990). The effects of Ncadherin misexpression on morphogenesis in Xenopus embryos. Neuron, 4, 493e506. Devoto, S. H., Melanc¸on, E., Eisen, J. S., & Westerfield, M. (1996). Identification of separate slow and fast muscle precursor cells in vivo, prior to somite formation. Development (Cambridge, England), 122, 3371e3380. Devoto, S. H., Stoiber, W., Hammond, C. L., Steinbacher, P., Haslett, J. R., Barresi, M. J. F., et al. (2006). Generality of vertebrate developmental patterns: Evidence for a dermomyotome in fish. Evolution and Development, 8, 101e110. Dietrich, M. R., Ankeny, R. A., & Chen, P. M. (2014). Publication trends in model organism research. Genetics, 198, 787e794. Doitsidou, M., Reichman-Fried, M., Stebler, J., Ko¨prunner, M., Do¨rries, J., Meyer, D., et al. (2002). Guidance of primordial germ cell migration by the chemokine SDF-1. Cell, 111, 647e659. Dougan, S. T., Warga, R. M., Kane, D. A., Schier, A. F., & Talbot, W. S. (2003). The role of the zebrafish nodal-related genes squint and cyclops in patterning of mesendoderm. Development (Cambridge, England), 130, 1837e1851. Dougherty, M., Kamel, G., Shubinets, V., Hickey, G., Grimaldi, M., & Liao, E. C. (2012). Embryonic fate map of first pharyngeal arch structures in the sox10: Kaede zebrafish transgenic model. Journal of Craniofacial Surgery, 23, 1333e1337. Drummond, I. A., & Davidson, A. J. (2010). Zebrafish kidney development. Methods in Cell Biology, 100, 233e260. Drummond, B. E., Li, Y., Marra, A. N., Cheng, C. N., & Wingert, R. A. (2017). The tbx2a/b transcription factors direct pronephros segmentation and corpuscle of Stannius formation in zebrafish. Developmental Biology, 421, 52e66. Du, S. J., Devoto, S. H., Westerfield, M., & Moon, R. T. (1997). Positive and negative regulation of muscle cell identity by members of the hedgehog and TGF-beta gene families. The Journal of Cell Biology, 139, 145e156. Dutton, K. A., Pauliny, A., Lopes, S. S., Elworthy, S., Carney, T. J., Rauch, J., et al. (2001). Zebrafish colourless encodes sox10 and specifies non-ectomesenchymal neural crest fates. Development (Cambridge, England), 128, 4113e4125. D’Amico, L. A., & Cooper, M. S. (2001). Morphogenetic domains in the yolk syncytial layer of axiating zebrafish embryos. Developmental Dynamics: An Official Publication of the American Association of Anatomists, 222, 611e624. Eckalbar, W. L., Fisher, R. E., Rawls, A., & Kusumi, K. (2012). Scoliosis and segmentation defects of the vertebrae. Wiley Interdisciplinary Reviews. Developmental Biology, 1, 401e423. van Eeden, F. J., Granato, M., Schach, U., Brand, M., Furutani-Seiki, M., Haffter, P., et al. (1996). Mutations affecting somite formation and

patterning in the zebrafish, Danio rerio. Development (Cambridge, England), 123, 153e164. Eisenhoffer, G. T., Slattum, G., Ruiz, O. E., Otsuna, H., Bryan, C. D., Lopez, J., et al. (2017). A toolbox to study epidermal cell types in zebrafish. Journal of Cell Science, 130, 269e277. Ellis, K., Bagwell, J., & Bagnat, M. (2013a). Notochord vacuoles are lysosome-related organelles that function in axis and spine morphogenesis. The Journal of Cell Biology, 200, 667e679. Ellis, K., Hoffman, B. D., & Bagnat, M. (2013b). The vacuole within: How cellular organization dictates notochord function. BioArchitecture, 3, 64e68. Emoto, Y., Wada, H., Okamoto, H., Kudo, A., & Imai, Y. (2005). Retinoic acid-metabolizing enzyme Cyp26a1 is essential for determining territories of hindbrain and spinal cord in zebrafish. Developmental Biology, 278, 415e427. Esaki, M., Hoshijima, K., Nakamura, N., Munakata, K., Tanaka, M., Ookata, K., et al. (2009). Mechanism of development of ionocytes rich in vacuolar-type H(þ)-ATPase in the skin of zebrafish larvae. Developmental Biology, 329, 116e129. Essner, J. J., Vogan, K. J., Wagner, M. K., Tabin, C. J., Yost, H. J., & Brueckner, M. (2002). Lefteright development: Conserved function for embryonic nodal cilia. Nature, 418, 37e38. Ettensohn, C. A. (2013). Encoding anatomy: Developmental gene regulatory networks and morphogenesis. Genesis (New York, N.Y.: 2000), 51, 383e409. Farrell, J. A., Wang, Y., Riesenfeld, S. J., Shekhar, K., Regev, A., & Schier, A. F. (2018). Single-cell reconstruction of developmental trajectories during zebrafish embryogenesis. Science, (6392), 360. Fauny, J.-D., Thisse, B., & Thisse, C. (2009). The entire zebrafish blastula-gastrula margin acts as an organizer dependent on the ratio of Nodal to BMP activity. Development (Cambridge, England), 136, 3811e3819. Feng, X., Adiarte, E. G., & Devoto, S. H. (2006). Hedgehog acts directly on the zebrafish dermomyotome to promote myogenic differentiation. Developmental Biology, 300, 736e746. Ferg, M., Armant, O., Yang, L., Dickmeis, T., Rastegar, S., & Stra¨hle, U. (2014). Gene transcription in the zebrafish embryo: Regulators and networks. Briefings in Functional Genomics, 13, 131e143. Field, H. A., Ober, E. A., Roeser, T., & Stainier, D. Y. R. (2003). Formation of the digestive system in zebrafish. I. Liver morphogenesis. Developmental Biology, 253, 279e290. Fouquet, B., Weinstein, B. M., Serluca, F. C., & Fishman, M. C. (1997). Vessel patterning in the embryo of the zebrafish: Guidance by notochord. Developmental Biology, 183, 37e48. Frisdal, A., & Trainor, P. A. (2014). Development and evolution of the pharyngeal apparatus. Wiley Interdisciplinary Reviews. Developmental Biology, 3, 403e418. Gahtan, E., & Baier, H. (2004). Of lasers, mutants, and see-through brains: Functional neuroanatomy in zebrafish. Journal of Neurobiology, 59, 147e161. Gallik, K. L., Treffy, R. W., Nacke, L. M., Ahsan, K., Rocha, M., GreenSaxena, A., et al. (2017). Neural crest and cancer: Divergent travelers on similar paths. Mechanisms of Development, 148, 89e99. Gehrke, A. R., Schneider, I., de la Calle-Mustienes, E., Tena, J. J., Gomez-Marin, C., Chandran, M., et al. (2015). Deep conservation of wrist and digit enhancers in fish. Proceedings of the National Academy of Sciences of the United States of America, 112, 803e808. Gestri, G., Link, B. A., & Neuhauss, S. C. F. (2012). The visual system of zebrafish and its use to model human ocular diseases. Developmental Neurobiology, 72, 302e327. Gfrerer, L., Dougherty, M., & Liao, E. C. (2013). Visualization of craniofacial development in the sox10: Kaede transgenic zebrafish line using time-lapse confocal microscopy. Journal of Visualized Experiments, e50525.

V. Scientific Research

References

Giraldez, A. J., Mishima, Y., Rihel, J., Grocock, R. J., Van Dongen, S., Inoue, K., et al. (2006). Zebrafish MiR-430 promotes deadenylation and clearance of maternal mRNAs. Science, 312, 75e79. Goody, M. F., Carter, E. V., Kilroy, E. A., Maves, L., & Henry, C. A. (2017). “Muscling” throughout life: Integrating studies of muscle development, homeostasis, and disease in zebrafish. Current Topics in Developmental Biology, 124, 197e234. Gore, A. V., Monzo, K., Cha, Y. R., Pan, W., & Weinstein, B. M. (2012). Vascular development in the zebrafish. Cold Spring Harbor Perspectives in Medicine, 2, a006684. Granato, M., van Eeden, F. J., Schach, U., Trowe, T., Brand, M., Furutani-Seiki, M., et al. (1996). Genes controlling and mediating locomotion behavior of the zebrafish embryo and larva. Development (Cambridge, England), 123, 399e413. Grimes, D. T., Boswell, C. W., Morante, N. F. C., Henkelman, R. M., Burdine, R. D., & Ciruna, B. (2016). Zebrafish models of idiopathic scoliosis link cerebrospinal fluid flow defects to spine curvature. Science, 352, 1341e1344. Grunwald, D. J., & Eisen, J. S. (2002). Headwaters of the zebrafish d emergence of a new model vertebrate. Nature Reviews Genetics, 3, 717e724. Guner, B., & Karlstrom, R. O. (2007). Cloning of zebrafish nkx6.2 and a comprehensive analysis of the conserved transcriptional response to Hedgehog/Gli signaling in the zebrafish neural tube. Gene Expression Patterns, 7, 596e605. Haack, T., & Abdelilah-Seyfried, S. (2016). The force within: Endocardial development, mechanotransduction and signalling during cardiac morphogenesis. Development (Cambridge, England), 143, 373e386. Haas, P., & Gilmour, D. (2006). Chemokine signaling mediates selforganizing tissue migration in the zebrafish lateral line. Developmental Cell, 10, 673e680. Halpern, M. E., Ho, R. K., Walker, C., & Kimmel, C. B. (1993). Induction of muscle pioneers and floor plate is distinguished by the zebrafish no tail mutation. Cell, 75, 99e111. Halpern, M. E., Rhee, J., Goll, M. G., Akitake, C. M., Parsons, M., & Leach, S. D. (2008). Gal4/UAS transgenic tools and their application to zebrafish. Zebrafish, 5, 97e110. Hansen, A., & Eckart, Z. (1998). The peripheral olfactory organ of the zebrafish, Danio rerio: An ultrastructural study. Chemical Senses, 23, 39e48. Harland, R. M. (2018). A new view of embryo development and regeneration. Science, 360(6392), 967e968. Harper, C., & Lawrence, C. (2016). The laboratory zebrafish. CRC Press. Harrington, M. J., Hong, E., & Brewster, R. (2009). Comparative analysis of neurulation: First impressions do not count. Molecular Reproduction and Development, 76, 954e965. Hasley, A., Chavez, S., Danilchik, M., Wu¨hr, M., & Pelegri, F. (2017). Vertebrate embryonic cleavage pattern determination. Advances in Experimental Medicine and Biology, 953, 117e171. Hava, D., Forster, U., Matsuda, M., Cui, S., Link, B. A., Eichhorst, J., et al. (2009). Apical membrane maturation and cellular rosette formation during morphogenesis of the zebrafish lateral line. Journal of Cell Science, 122, 687e695. Hawkes, J. W. (1974). The structure of fish skin. I. General organization. Cell and Tissue Research, 149, 147e158. Head, J. R., Gacioch, L., Pennisi, M., & Meyers, J. R. (2013). Activation of canonical Wnt/b-catenin signaling stimulates proliferation in neuromasts in the zebrafish posterior lateral line. Developmental Dynamics: An Official Publication of the American Association of Anatomists, 242, 832e846. Heisenberg, C. P., & Tada, M. (2002). Zebrafish gastrulation movements: Bridging cell and developmental biology. Seminars in Cell and Developmental Biology, 13, 471e479.

585

Henrikson, R. C., & Matoltsy, A. G. (1967a). The fine structure of teleost epidermis. 1. Introduction and filament-containing cells. Journal of Ultrastructure Research, 21, 194e212. Henrikson, R. C., & Matoltsy, A. G. (1967b). The fine structure of teleost epidermis. II. Mucous cells. Journal of Ultrastructure Research, 21, 213e221. Henrikson, R. C., & Matoltsy, A. G. (1967c). The fine structure of teleost epidermis. 3. Club cells and other cell types. Journal of Ultrastructure Research, 21, 222e232. Henry, C. A., McNulty, I. M., Durst, W. A., Munchel, S. E., & Amacher, S. L. (2005). Interactions between muscle fibers and segment boundaries in zebrafish. Developmental Biology, 287, 346e360. Herbomel, P., Thisse, B., & Thisse, C. (1999). Ontogeny and behaviour of early macrophages in the zebrafish embryo. Development (Cambridge, England), 126, 3735e3745. Hernandez, R. E., Putzke, A. P., Myers, J. P., Margaretha, L., & Moens, C. B. (2007). Cyp26 enzymes generate the retinoic acid response pattern necessary for hindbrain development. Development (Cambridge, England), 134, 177e187. Higashijima, S., Hotta, Y., & Okamoto, H. (2000). Visualization of cranial motor neurons in live transgenic zebrafish expressing green fluorescent protein under the control of the islet-1 promoter/ enhancer. The Journal of Neuroscience: The Official Journal of the Society for Neuroscience, 20, 206e218. Hirata, M., Nakamura, K.-I., & Kondo, S. (2005). Pigment cell distributions in different tissues of the zebrafish, with special reference to the striped pigment pattern. Developmental Dynamics: An Official Publication of the American Association of Anatomists, 234, 293e300. Hoffman, L., Miles, J., Avaron, F., Laforest, L., & Akimenko, M.-A. (2002). Exogenous retinoic acid induces a stage-specific, transient and progressive extension of Sonic hedgehog expression across the pectoral fin bud of zebrafish. International Journal of Developmental Biology, 46, 949e956. Holley, S. A., Ju¨lich, D., Rauch, G.-J., Geisler, R., & Nu¨sslein-Volhard, C. (2002). her1 and the notch pathway function within the oscillator mechanism that regulates zebrafish somitogenesis. Development (Cambridge, England), 129, 1175e1183. Holtzman, N. G., Schoenebeck, J. J., Tsai, H.-J., & Yelon, D. (2007). Endocardium is necessary for cardiomyocyte movement during heart tube assembly. Development (Cambridge, England), 134, 2379e2386. Hong, E., & Brewster, R. (2006). N-cadherin is required for the polarized cell behaviors that drive neurulation in the zebrafish. Development (Cambridge, England), 133, 3895e3905. Horne-Badovinac, S., Lin, D., Waldron, S., Schwarz, M., Mbamalu, G., Pawson, T., et al. (2001). Positional cloning of heart and soul reveals multiple roles for PKC lambda in zebrafish organogenesis. Current Biology, 11, 1492e1502. Horng, J.-L., Lin, L.-Y., & Hwang, P.-P. (2009). Functional regulation of Hþ-ATPase-rich cells in zebrafish embryos acclimated to an acidic environment. American Journal of PhysiologyeCell Physiology, 296, C682eC692. Hsiao, C.-D., You, M.-S., Guh, Y.-J., Ma, M., Jiang, Y.-J., & Hwang, P.-P. (2007). A positive regulatory loop between foxi3a and foxi3b is essential for specification and differentiation of zebrafish epidermal ionocytes. PLoS One, 2, e302. Hubaud, A., & Pourquie´, O. (2014). Signalling dynamics in vertebrate segmentation. Nature Reviews Molecular Cell Biology, 15, 709e721. Hutson, L. D., & Chien, C. B. (2002). Pathfinding and error correction by retinal axons: The role of astray/robo2. Neuron, 33, 205e217. Hwang, P.-P. (2009). Ion uptake and acid secretion in zebrafish (Danio rerio). Journal of Experimental Biology, 212, 1745e1752.

V. Scientific Research

586

45. Zebrafish as a Model to Understand Vertebrate Development

Iimura, T., & Pourquie´, O. (2007). Hox genes in time and space during vertebrate body formation. Development Growth and Differentiation, 49, 265e275. Isogai, S., Horiguchi, M., & Weinstein, B. M. (2001). The vascular anatomy of the developing zebrafish: An atlas of embryonic and early larval development. Developmental Biology, 230, 278e301. Isogai, S., Lawson, N. D., Torrealday, S., Horiguchi, M., & Weinstein, B. M. (2003). Angiogenic network formation in the developing vertebrate trunk. Development (Cambridge, England), 130, 5281e5290. Ja¨nicke, M., Carney, T. J., & Hammerschmidt, M. (2007). Foxi3 transcription factors and Notch signaling control the formation of skin ionocytes from epidermal precursors of the zebrafish embryo. Developmental Biology, 307, 258e271. Johnson, K., Barragan, J., Bashiruddin, S., Smith, C. J., Tyrrell, C., Parsons, M. J., et al. (2016). Gfap-positive radial glial cells are an essential progenitor population for later-born neurons and glia in the zebrafish spinal cord. Glia, 64, 1170e1189. Joseph Yost, H. (1999). Diverse initiation in a conserved left-right pathway? Current Opinion in Genetics and Development, 9, 422e426. Kabrun, N., Bu¨hring, H. J., Choi, K., Ullrich, A., Risau, W., & Keller, G. (1997). Flk-1 expression defines a population of early embryonic hematopoietic precursors. Development (Cambridge, England), 124, 2039e2048. Kalcheim, C., & Kumar, D. (2017). Cell fate decisions during neural crest ontogeny. International Journal of Developmental Biology, 61, 195e203. Kalev-Zylinska, M. L., Horsfield, J. A., Flores, M. V. C., Postlethwait, J. H., Vitas, M. R., Baas, A. M., et al. (2002). Runx1 is required for zebrafish blood and vessel development and expression of a human RUNX1-CBF2T1 transgene advances a model for studies of leukemogenesis. Development (Cambridge, England), 129, 2015e2030. Kamei, M., Saunders, W. B., Bayless, K. J., Dye, L., Davis, G. E., & Weinstein, B. M. (2006). Endothelial tubes assemble from intracellular vacuoles in vivo. Nature, 442, 453e456. Kane, D. A., & Kimmel, C. B. (1993). The zebrafish midblastula transition. Development (Cambridge, England), 119, 447e456. Kane, D. A., McFarland, K. N., & Warga, R. M. (2005). Mutations in half baked/E-cadherin block cell behaviors that are necessary for teleost epiboly. Development (Cambridge, England), 132, 1105e1116. Kane, D. A., Warga, R. M., & Kimmel, C. B. (1992). Mitotic domains in the early embryo of the zebrafish. Nature, 360, 735e737. Karlstrom, R. O., & Kane, D. A. (1996). A flipbook of zebrafish embryogenesis. Development (Cambridge, England), 123, 461. Karlstrom, R. O., Talbot, W. S., & Schier, A. F. (1999). Comparative synteny cloning of zebrafish you-too: Mutations in the hedgehog target gli2 affect ventral forebrain patterning. Genes and Development, 13, 388e393. Kaufman, C. K., Mosimann, C., Fan, Z. P., Yang, S., Thomas, A. J., Ablain, J., et al. (2016). A zebrafish melanoma model reveals emergence of neural crest identity during melanoma initiation. Science, 351, aad2197. Kee, Y., Hwang, B. J., Sternberg, P. W., & Bronner-Fraser, M. (2007). Evolutionary conservation of cell migration genes: From nematode neurons to vertebrate neural crest. Genes and Development, 21, 391e396. Keller, P. J. (2013). In vivo imaging of zebrafish embryogenesis. Methods (San Diego, Calif.), 62, 268e278. Keller, P. J., Schmidt, A. D., Wittbrodt, J., & Stelzer, E. H. K. (2008). Reconstruction of zebrafish early embryonic development by scanned light sheet microscopy. Science, 322, 1065e1069. Kelsh, R. N., Brand, M., Jiang, Y. J., Heisenberg, C. P., Lin, S., Haffter, P., et al. (1996). Zebrafish pigmentation mutations and the processes of neural crest development. Development (Cambridge, England), 123, 369e389.

Kelsh, R. N., & Eisen, J. S. (2000). The zebrafish colourless gene regulates development of non-ectomesenchymal neural crest derivatives. Development (Cambridge, England), 127, 515e525. Kelsh, R. N., Harris, M. L., Colanesi, S., & Erickson, C. A. (2009). Stripes and belly-spots – a review of pigment cell morphogenesis in vertebrates. Seminars in Cell and Developmental Biology, 20, 90e104. Kelsh, R. N., & Raible, D. W. (2002). Specification of zebrafish neural crest. Results and Problems in Cell Differentiation, 40, 216e236. Kimmel, C. B., Ballard, W. W., Kimmel, S. R., Ullmann, B., & Schilling, T. F. (1995). Stages of embryonic development of the zebrafish. Developmental Dynamics: An Official Publication of the American Association of Anatomists, 203, 253e310. Kimmel, C. B., Miller, C. T., Kruze, G., Ullmann, B., BreMiller, R. A., Larison, K. D., et al. (1998). The shaping of pharyngeal cartilages during early development of the zebrafish. Developmental Biology, 203, 245e263. Kimmel, C. B., Warga, R. M., & Schilling, T. F. (1990). Origin and organization of the zebrafish fate map. Development, 108, 581e594. Kissa, K., & Herbomel, P. (2010). Blood stem cells emerge from aortic endothelium by a novel type of cell transition. Nature, 464, 112e115. Klymkowsky, M. W., Rossi, C. C., & Artinger, K. B. (2010). Mechanisms driving neural crest induction and migration in the zebrafish and Xenopus laevis. Cell Adhesion and Migration, 4, 595e608. Knight, R. D., & Schilling, T. F. (2006). Cranial neural crest and development of the head skeleton. Advances in Experimental Medicine and Biology, 589, 120e133. Knutsdottir, H., Zmurchok, C., Bhaskar, D., Palsson, E., Dalle Nogare, D., Chitnis, A. B., et al. (2017). Polarization and migration in the zebrafish posterior lateral line system. PLoS Computational Biology, 13, e1005451. Korzh, V. (2014). Stretching cell morphogenesis during late neurulation and mild neural tube defects. Development Growth and Differentiation, 56, 425e433. Korzh, S., Emelyanov, A., & Korzh, V. (2001). Developmental analysis of ceruloplasmin gene and liver formation in zebrafish. Mechanisms of Development, 103, 137e139. Koudijs, M. J., den Broeder, M. J., Keijser, A., Wienholds, E., Houwing, S., van Rooijen, E. M. H. C., et al. (2005). The zebrafish mutants dre, uki, and lep encode negative regulators of the hedgehog signaling pathway. PLoS Genetics, 1, e19. Kozlovskaja-Gumbrien_e, A., Yi, R., Alexander, R., Aman, A., Jiskra, R., Nagelberg, D., et al. (2017). Proliferation-independent regulation of organ size by Fgf/Notch signaling. ELife, 6. Kozlowski, D. J., Murakami, T., Ho, R. K., & Weinberg, E. S. (1997). Regional cell movement and tissue patterning in the zebrafish embryo revealed by fate mapping with caged fluorescein. Biochemistry and Cell Biology, 75, 551e562. Kramer-Zucker, A. G. (2005). Cilia-driven fluid flow in the zebrafish pronephros, brain and Kupffer’s vesicle is required for normal organogenesis. Development, 132, 1907e1921. Krumlauf, R. (2016). Hox genes and the hindbrain: A study in segments. Current Topics in Developmental Biology, 116, 581e596. Ku¨chler, A. M., Gjini, E., Peterson-Maduro, J., Cancilla, B., Wolburg, H., & Schulte-Merker, S. (2006). Development of the zebrafish lymphatic system requires VEGFC signaling. Current Biology, 16, 1244e1248. Kudoh, T., Wilson, S. W., & Dawid, I. B. (2002). Distinct roles for Fgf, Wnt and retinoic acid in posteriorizing the neural ectoderm. Development (Cambridge, England), 129, 4335e4346. Kuraku, S., & Meyer, A. (2009). The evolution and maintenance of Hox gene clusters in vertebrates and the teleost-specific genome duplication. International Journal of Developmental Biology, 53, 765e773. Kwak, J., Park, O. K., Jung, Y. J., Hwang, B. J., Kwon, S.-H., & Kee, Y. (2013). Live image profiling of neural crest lineages in zebrafish transgenic lines. Molecules and Cells, 35, 255e260.

V. Scientific Research

References

Laforest, L., Brown, C. W., Poleo, G., Ge´raudie, J., Tada, M., Ekker, M., et al. (1998). Involvement of the sonic hedgehog, patched 1 and bmp2 genes in patterning of the zebrafish dermal fin rays. Development (Cambridge, England), 125, 4175e4184. Lam, E. Y. N., Hall, C. J., Crosier, P. S., Crosier, K. E., & Flores, M. V. (2010). Live imaging of Runx1 expression in the dorsal aorta tracks the emergence of blood progenitors from endothelial cells. Blood, 116, 909e914. Lawson, L. Y., & Harfe, B. D. (2017). Developmental mechanisms of intervertebral disc and vertebral column formation. Wiley Interdisciplinary Reviews. Developmental Biology, 6. Lawson, N. D., Scheer, N., Pham, V. N., Kim, C. H., Chitnis, A. B., Campos-Ortega, J. A., et al. (2001). Notch signaling is required for arterial-venous differentiation during embryonic vascular development. Development (Cambridge, England), 128, 3675e3683. Le Dre´au, G., & Martı´, E. (2012). Dorsal-ventral patterning of the neural tube: A tale of three signals. Developmental Neurobiology, 72, 1471e1481. Le Guellec, D., Morvan-Dubois, G., & Sire, J.-Y. (2003). Skin development in bony fish with particular emphasis on collagen deposition in the dermis of the zebrafish (Danio rerio). International Journal of Developmental Biology, 48, 217e231. Lee, M. T., Bonneau, A. R., & Giraldez, A. J. (2014). Zygotic genome activation during the maternal-to-zygotic transition. Annual Review of Cell and Developmental Biology, 30, 581e613. Lee, M. T., Bonneau, A. R., Takacs, C. M., Bazzini, A. A., DiVito, K. R., Fleming, E. S., et al. (2013). Nanog, Pou5f1 and SoxB1 activate zygotic gene expression during the maternal-to-zygotic transition. Nature, 503, 360e364. Lee, H., & Kimelman, D. (2002). A dominant-negative form of p63 is required for epidermal proliferation in zebrafish. Developmental Cell, 2, 607e616. Lee, K., & Skromne, I. (2014). Retinoic acid regulates size, pattern and alignment of tissues at the head-trunk transition. Development (Cambridge, England), 141, 4375e4384. Lele, Z., Folchert, A., Concha, M., Rauch, G.-J., Geisler, R., Rosa, F., et al. (2002). parachute/n-cadherin is required for morphogenesis and maintained integrity of the zebrafish neural tube. Development (Cambridge, England), 129, 3281e3294. Lewis, K. E., Currie, P. D., Roy, S., Schauerte, H., Haffter, P., & Ingham, P. W. (1999). Control of muscle cell-type specification in the zebrafish embryo by Hedgehog signalling. Developmental Biology, 216, 469e480. Lewis, K. E., & Eisen, J. S. (2003). From cells to circuits: Development of the zebrafish spinal cord. Progress in Neurobiology, 69, 419e449. Lieschke, G. J., Oates, A. C., Paw, B. H., Thompson, M. A., Hall, N. E., Ward, A. C., et al. (2002). Zebrafish SPI-1 (PU.1) marks a site of myeloid development independent of primitive erythropoiesis: Implications for axial patterning. Developmental Biology, 246, 274e295. Li, M., Hromowyk, K. J., Amacher, S. L., & Currie, P. D. (2017). Muscular dystrophy modeling in zebrafish. Methods in Cell Biology, 138, 347e380. Li, H.-H., Huang, P., Dong, W., Zhu, Z.-Y., & Liu, D. (2013). [A brief history of zebrafish research–toward biomedicine]. Yi Chuan, 35 (4), 410e420, Review. Chinese. PubMed PMID: 23659931. Liu, Y., Pathak, N., Kramer-Zucker, A., & Drummond, I. A. (2007). Notch signaling controls the differentiation of transporting epithelia and multiciliated cells in the zebrafish pronephros. Development (Cambridge, England), 134, 1111e1122. Lowery, L. A., & Sive, H. (2004). Strategies of vertebrate neurulation and a re-evaluation of teleost neural tube formation. Mechanisms of Development, 121, 1189e1197. Machado, R. G., & Eames, B. F. (2017). Using zebrafish to test the genetic basis of human craniofacial diseases. Journal of Dental Research, 96, 1192e1199.

587

Majumdar, A., & Drummond, I. A. (1999). Podocyte differentiation in the absence of endothelial cells as revealed in the zebrafish avascular mutant, cloche. Developmental Genetics, 24, 220e229. Majumdar, A., Lun, K., Brand, M., & Drummond, I. A. (2000). Zebrafish no isthmus reveals a role for pax2.1 in tubule differentiation and patterning events in the pronephric primordia. Development (Cambridge, England), 127, 2089e2098. Ma, E. Y., & Raible, D. W. (2009). Signaling pathways regulating zebrafish lateral line development. Current Biology, 19, R381eR386. Maroto, M., Bone, R. A., & Dale, J. K. (2012). Somitogenesis. Development (Cambridge, England), 139, 2453e2456. Martik, M. L., & Bronner, M. E. (2017). Regulatory logic underlying diversification of the neural crest. Trends in Genetics, 33, 715e727. Massarwa, R., Ray, H. J., & Niswander, L. (2014). Morphogenetic movements in the neural plate and neural tube: Mouse. Wiley Interdisciplinary Reviews. Developmental Biology, 3, 59e68. Maves, L., & Kimmel, C. B. (2005). Dynamic and sequential patterning of the zebrafish posterior hindbrain by retinoic acid. Developmental Biology, 285, 593e605. Ma, D., Zhang, J., Lin, H., Italiano, J., & Handin, R. I. (2011). The identification and characterization of zebrafish hematopoietic stem cells. Blood, 118, 289e297. McCarroll, M. N., Lewis, Z. R., Culbertson, M. D., Martin, B. L., Kimelman, D., & Nechiporuk, A. V. (2012). Graded levels of Pax2a and Pax8 regulate cell differentiation during sensory placode formation. Development (Cambridge, England), 139, 2740e2750. McCluskey, B. M., & Postlethwait, J. H. (2015). Phylogeny of zebrafish, a “model species,” within Danio, a “model genus. Molecular Biology and Evolution, 32, 635e652. McFarland, K. N., Warga, R. M., & Kane, D. A. (2005). Genetic locus half baked is necessary for morphogenesis of the ectoderm. Developmental Dynamics: An Official Publication of the American Association of Anatomists, 233, 390e406. McGraw, H. F., Nechiporuk, A., & Raible, D. W. (2008). Zebrafish dorsal root ganglia neural precursor cells adopt a glial fate in the absence of neurogenin1. The Journal of Neuroscience: The Official Journal of the Society for Neuroscience, 28, 12558e12569. McKenna, A., Findlay, G. M., Gagnon, J. A., Horwitz, M. S., Schier, A. F., & Shendure, J. (2016). Whole-organism lineage tracing by combinatorial and cumulative genome editing. Science, 353, aaf7907. Medeiros, D. M., & Crump, J. G. (2012). New perspectives on pharyngeal dorsoventral patterning in development and evolution of the vertebrate jaw. Developmental Biology, 371, 121e135. Melby, A. E., Kimelman, D., & Kimmel, C. B. (1997). Spatial regulation of floating head expression in the developing notochord. Developmental Dynamics: An Official Publication of the American Association of Anatomists, 209, 156e165. Melby, A. E., Warga, R. M., & Kimmel, C. B. (1996). Specification of cell fates at the dorsal margin of the zebrafish gastrula. Development, 122, 2225e2237. Meunier, R. (2012). Stages in the development of a model organism as a platform for mechanistic models in developmental biology: Zebrafish, 1970e2000. Studies in History and Philosophy of Biological and Biomedical Sciences, 43, 522e531. Miccoli, A., Dalla Valle, L., & Carnevali, O. (2017). The maternal control in the embryonic development of zebrafish. General and Comparative Endocrinology, 245, 55e68. Miles, L. B., Mizoguchi, T., Kikuchi, Y., & Verkade, H. (2017). A role for planar cell polarity during early endoderm morphogenesis. Biology Open, 6, 531e539. Miquerol, L., & Kelly, R. G. (2013). Organogenesis of the vertebrate heart. Wiley Interdisciplinary Reviews. Developmental Biology, 2, 17e29.

V. Scientific Research

588

45. Zebrafish as a Model to Understand Vertebrate Development

Moens, C. B., & Prince, V. E. (2002). Constructing the hindbrain: Insights from the zebrafish. Developmental Dynamics: An Official Publication of the American Association of Anatomists, 224, 1e17. Morales, D., & Kania, A. (2017). Cooperation and crosstalk in axon guidance cue integration: Additivity, synergy, and fine-tuning in combinatorial signaling. Developmental Neurobiology, 77, 891e904. Morin-Kensicki, E. M., Melancon, E., & Eisen, J. S. (2002). Segmental relationship between somites and vertebral column in zebrafish. Development (Cambridge, England), 129, 3851e3860. Morita, H., Nandadasa, S., Yamamoto, T. S., Terasaka-Iioka, C., Wylie, C., & Ueno, N. (2010). Nectin-2 and N-cadherin interact through extracellular domains and induce apical accumulation of F-actin in apical constriction of Xenopus neural tube morphogenesis. Development (Cambridge, England), 137, 1315e1325. Mork, L., & Crump, G. (2015). Zebrafish craniofacial development: A window into early patterning. Current Topics in Developmental Biology, 115, 235e269. Moro, E., Vettori, A., Porazzi, P., Schiavone, M., Rampazzo, E., Casari, A., et al. (2013). Generation and application of signaling pathway reporter lines in zebrafish. Molecular Genetics and Genomics, 288, 231e242. Mulligan, T. S., & Weinstein, B. M. (2014). Emerging from the PAC: Studying zebrafish lymphatic development. Microvascular Research, 96, 23e30. Naylor, R. W., Przepiorski, A., Ren, Q., Yu, J., & Davidson, A. J. (2013). HNF1b is essential for nephron segmentation during nephrogenesis. Journal of the American Society of Nephrology, 24, 77e87. Ng, A. N. Y., de Jong-Curtain, T. A., Mawdsley, D. J., White, S. J., Shin, J., Appel, B., et al. (2005). formation of the digestive system in zebrafish: III. Intestinal epithelium morphogenesis. Developmental Biology, 286, 114e135. Nikaido, M., Kawakami, A., Sawada, A., Furutani-Seiki, M., Takeda, H., & Araki, K. (2002). Tbx24, encoding a T-box protein, is mutated in the zebrafish somite-segmentation mutant fused somites. Nature Genetics, 31, 195e199. Nikaido, M., Navajas Acedo, J., Hatta, K., & Piotrowski, T. (2017). Retinoic acid is required and Fgf, Wnt, and Bmp signaling inhibit posterior lateral line placode induction in zebrafish. Developmental Biology, 431, 215e225. Niu, X., Shi, H., & Peng, J. (2010). The role of mesodermal signals during liver organogenesis in zebrafish. Science China Life Sciences, 53, 455e461. Noe¨l, E. S., Verhoeven, M., Lagendijk, A. K., Tessadori, F., Smith, K., Choorapoikayil, S., et al. (2013). A Nodal-independent and tissueintrinsic mechanism controls heart-looping chirality. Nature Communications, 4, 2754. Nogare, D. D., Nikaido, M., Somers, K., Head, J., Piotrowski, T., & Chitnis, A. B. (2017). In toto imaging of the migrating Zebrafish lateral line primordium at single cell resolution. Developmental Biology, 422, 14e23. Nonaka, S., Tanaka, Y., Okada, Y., Takeda, S., Harada, A., Kanai, Y., et al. (1998). Randomization of lefteright asymmetry due to loss of nodal cilia generating leftward flow of extraembryonic fluid in mice lacking KIF3B motor protein. Cell, 95, 829e837. Noordermeer, D., Leleu, M., Schorderet, P., Joye, E., Chabaud, F., & Duboule, D. (2014). Temporal dynamics and developmental memory of 3D chromatin architecture at Hox gene loci. ELife, 3, e02557. Nowick, K., & Stubbs, L. (2010). Lineage-specific transcription factors and the evolution of gene regulatory networks. Briefings in Functional Genomics, 9, 65e78. Odenthal, J., Haffter, P., Vogelsang, E., Brand, M., van Eeden, F. J., Furutani-Seiki, M., et al. (1996). Mutations affecting the formation of the notochord in the zebrafish, Danio rerio. Development (Cambridge, England), 123, 103e115.

Ou, H., Simon, J. A., Rubel, E. W., & Raible, D. W. (2012). Screening for chemicals that affect hair cell death and survival in the zebrafish lateral line. Hearing Research, 288, 58e66. O’Connell, L. A. (2013). Evolutionary development of neural systems in vertebrates and beyond. Journal of Neurogenetics, 27, 69e85. O’Farrell, P. H. (2015). Growing an embryo from a single cell: A hurdle in animal life. Cold Spring Harbor Perspectives in Biology, 7. Pacheco, A., & Gallo, G. (2016). Actin filament-microtubule interactions in axon initiation and branching. Brain Research Bulletin, 126, 300e310. Paksa, A., & Raz, E. (2015). Zebrafish germ cells: Motility and guided migration. Current Opinion in Cell Biology, 36, 80e85. Palaisa, K. A., & Granato, M. (2007). Analysis of zebrafish sidetracked mutants reveals a novel role for Plexin A3 in intraspinal motor axon guidance. Development (Cambridge, England), 134, 3251e3257. Pan, Y. A., Freundlich, T., Weissman, T. A., Schoppik, D., Wang, X. C., Zimmerman, S., et al. (2013). Zebrabow: Multispectral cell labeling for cell tracing and lineage analysis in zebrafish. Development (Cambridge, England), 140, 2835e2846. Paridaen, J. T., & Huttner, W. B. (2014). Neurogenesis during development of the vertebrate central nervous system. EMBO Reports, 15, 351e364. Parker, H. J., Bronner, M. E., & Krumlauf, R. (2014). A Hox regulatory network of hindbrain segmentation is conserved to the base of vertebrates. Nature, 514, 490e493. Parker, H. J., & Krumlauf, R. (2017). Segmental arithmetic: Summing up the hox gene regulatory network for hindbrain development in chordates. Wiley Interdisciplinary Reviews. Developmental Biology, 6. Park, H.-C., Shin, J., & Appel, B. (2004). Spatial and temporal regulation of ventral spinal cord precursor specification by Hedgehog signaling. Development (Cambridge, England), 131, 5959e5969. Pasquier, J., Braasch, I., Batzel, P., Cabau, C., Montfort, J., Nguyen, T., et al. (2017). Evolution of gene expression after whole-genome duplication: New insights from the spotted gar genome. Journal of Experimental Zoology. Part B, Molecular and Developmental Evolution, 328, 709e721. de Pater, E., Clijsters, L., Marques, S. R., Lin, Y.-F., GaravitoAguilar, Z. V., Yelon, D., et al. (2009). Distinct phases of cardiomyocyte differentiation regulate growth of the zebrafish heart. Development (Cambridge, England), 136, 1633e1641. Pathak, N., Obara, T., Mangos, S., Liu, Y., & Drummond, I. A. (2007). The zebrafish fleer gene encodes an essential regulator of cilia tubulin polyglutamylation. Molecular Biology of the Cell, 18, 4353e4364. Pelliccia, J. L., Jindal, G. A., & Burdine, R. D. (2017). Gdf3 is required for robust Nodal signaling during germ layer formation and left-right patterning. ELife, 6, e28635. Penton, A. L., Leonard, L. D., & Spinner, N. B. (2012). Notch signaling in human development and disease. Seminars in Cell and Developmental Biology, 23, 450e457. Peralta, M., Gonza´lez-Rosa, J. M., Marques, I. J., & Mercader, N. (2014). The epicardium in the embryonic and adult zebrafish. Journal of Developmental Biology, 2, 101e116. Perrimon, N., Pitsouli, C., & Shilo, B.-Z. (2012). Signaling mechanisms controlling cell fate and embryonic patterning. Cold Spring Harbor Perspectives in Biology, 4, a005975. Peter, I. S. (2017). Regulatory states in the developmental control of gene expression. Briefings in Functional Genomic, 16, 281e287. Peter, I. S., & Davidson, E. H. (2009). Genomic control of patterning. International Journal of Developmental Biology, 53, 707e716. Peter, I. S., & Davidson, E. H. (2017). Assessing regulatory information in developmental gene regulatory networks. Proceedings of the National Academy of Sciences of the United States of America, 114, 5862e5869.

V. Scientific Research

References

Pe´zeron, G., Mourrain, P., Courty, S., Ghislain, J., Becker, T. S., Rosa, F. M., et al. (2008). Live analysis of endodermal layer formation identifies random walk as a novel gastrulation movement. Current Biology, 18, 276e281. Pomreinke, A. P., Soh, G. H., Rogers, K. W., Bergmann, J. K., Bla¨ßle, A. J., & Mu¨ller, P. (2017). Dynamics of BMP signaling and distribution during zebrafish dorsal-ventral patterning. ELife, 6. Postlethwait, J. H. (2006). The zebrafish genome: A review and msx gene case study. Genome Dynamics, 2, 183e197. Postlethwait, J. H. (2007). The zebrafish genome in context: Ohnologs gone missing. Journal of Experimental Zoology. Part B, Molecular and Developmental Evolution, 308, 563e577. Prykhozhij, S. V., & Neumann, C. J. (2008). Distinct roles of Shh and Fgf signaling in regulating cell proliferation during zebrafish pectoral fin development. BMC Developmental Biology, 8, 91. Radice, G. L., Rayburn, H., Matsunami, H., Knudsen, K. A., Takeichi, M., & Hynes, R. O. (1997). Developmental defects in mouse embryos lacking N-cadherin. Developmental Biology, 181, 64e78. Rashid, D. J., Chapman, S. C., Larsson, H. C., Organ, C. L., Bebin, A.-G., Merzdorf, C. S., et al. (2014). From dinosaurs to birds: A tail of evolution. EvoDevo, 5, 25. Rasmussen, J. P., Sack, G. S., Martin, S. M., & Sagasti, A. (2015). Vertebrate epidermal cells are broad-specificity phagocytes that clear sensory axon debris. The Journal of Neuroscience: The Official Journal of the Society for Neuroscience, 35, 559e570. Ravanelli, A. M., & Appel, B. (2015). Motor neurons and oligodendrocytes arise from distinct cell lineages by progenitor recruitment. Genes and Development, 29, 2504e2515. Raz, E. (2003). Primordial germ-cell development: The zebrafish perspective. Nature Reviews Genetics, 4, 690e700. Reichert, S., Randall, R. A., & Hill, C. S. (2013). A BMP regulatory network controls ectodermal cell fate decisions at the neural plate border. Development (Cambridge, England), 140, 4435e4444. Reichman-Fried, M., Minina, S., & Raz, E. (2004). Autonomous modes of behavior in primordial germ cell migration. Developmental Cell, 6, 589e596. Reifers, F., Bo¨hli, H., Walsh, E. C., Crossley, P. H., Stainier, D. Y., & Brand, M. (1998). Fgf8 is mutated in zebrafish acerebellar (ace) mutants and is required for maintenance of midbrain-hindbrain boundary development and somitogenesis. Development (Cambridge, England), 125, 2381e2395. Resende, T. P., Andrade, R. P., & Palmeirim, I. (2014). Timing embryo segmentation: Dynamics and regulatory mechanisms of the vertebrate segmentation clock. BioMed Research International, 2014, 718683. Richardson, J., Gauert, A., Briones Montecinos, L., Fanlo, L., Alhashem, Z. M., Assar, R., et al. (2016). Leader cells define directionality of trunk, but not cranial, neural crest cell migration. Cell Reports, 15, 2076e2088. Rodino-Klapac, L. R., & Beattie, C. E. (2004). Zebrafish topped is required for ventral motor axon guidance. Developmental Biology, 273, 308e320. Rodrigues, F. S. L. M., Doughton, G., Yang, B., & Kelsh, R. N. (2012). A novel transgenic line using the Cre-lox system to allow permanent lineage-labeling of the zebrafish neural crest. Genesis (New York, N.Y. : 2000), 50, 750e757. Rohde, L. A., & Heisenberg, C.-P. (2007). Zebrafish gastrulation: Cell movements, signals, and mechanisms. International Review of Cytology, 261, 159e192. Royer, L. A., Lemon, W. C., Chhetri, R. K., Wan, Y., Coleman, M., Myers, E. W., et al. (2016). Adaptive light-sheet microscopy for long-term, high-resolution imaging in living organisms. Nature Biotechnology, 34, 1267e1278.

589

Roy, S., Qiao, T., Wolff, C., & Ingham, P. W. (2001). Hedgehog signaling pathway is essential for pancreas specification in the zebrafish embryo. Current Biology, 11, 1358e1363. Sagasti, A., Guido, M. R., Raible, D. W., & Schier, A. F. (2005). Repulsive interactions shape the morphologies and functional arrangement of zebrafish peripheral sensory arbors. Current Biology, 15, 804e814. Sagerstro¨m, C. G., Gammill, L. S., Veale, R., & Sive, H. (2005). Specification of the enveloping layer and lack of autoneuralization in zebrafish embryonic explants. Developmental Dynamics, 232, 85e97. Sagner, A., & Briscoe, J. (2017). Morphogen interpretation: Concentration, time, competence, and signaling dynamics. Wiley Interdisciplinary Reviews. Developmental Biology, 6. Sakamoto, K., Onimaru, K., Munakata, K., Suda, N., Tamura, M., Ochi, H., et al. (2009). Heterochronic shift in Hox-mediated activation of sonic hedgehog leads to morphological changes during fin development. PLoS One, 4, e5121. Sape`de, D., Rossel, M., Dambly-Chaudie`re, C., & Ghysen, A. (2005). Role of SDF1 chemokine in the development of lateral line efferent and facial motor neurons. Proceedings of the National Academy of Sciences of the United States of America, 102, 1714e1718. Sarrazin, A. F., Nun˜ez, V. A., Sape`de, D., Tassin, V., DamblyChaudie`re, C., & Ghysen, A. (2010). Origin and early development of the posterior lateral line system of zebrafish. The Journal of Neuroscience: The Official Journal of the Society for Neuroscience, 30, 8234e8244. Saxena, A., Peng, B. N., & Bronner, M. E. (2013). Sox10-dependent neural crest origin of olfactory microvillous neurons in zebrafish. ELife, 2, e00336. Schauerte, H. E., van Eeden, F. J., Fricke, C., Odenthal, J., Stra¨hle, U., & Haffter, P. (1998). Sonic hedgehog is not required for the induction of medial floor plate cells in the zebrafish. Development (Cambridge, England), 125, 2983e2993. Schibler, A., & Malicki, J. (2007). A screen for genetic defects of the zebrafish ear. Mechanisms of Development, 124, 592e604. Schilling, T. F., & Knight, R. D. (2001). Origins of anteroposterior patterning and Hox gene regulation during chordate evolution. Philosophical Transactions of the Royal Society of London B Biological Sciences, 356, 1599e1613. Schlosser, G. (2014). Early embryonic specification of vertebrate cranial placodes. Wiley Interdisciplinary Reviews. Developmental Biology, 3, 349e363. Schmidt, R., Stra¨hle, U., & Scholpp, S. (2013). Neurogenesis in zebrafish - from embryo to adult. Neural Development, 8, 3. Schmitz, B., Papan, C., & Campos-Ortega, J. A. (1993). Neurulation in the anterior trunk region of the zebrafish Brachydanio rerio. Roux’s Archives of Developmental Biology: The Official Organ of the EDBO, 202, 250e259. Schulte-Merker, S., van Eeden, F. J., Halpern, M. E., Kimmel, C. B., & Nu¨sslein-Volhard, C. (1994). No tail (ntl) is the zebrafish homologue of the mouse T (Brachyury) gene. Development (Cambridge, England), 120, 1009e1015. Schwarzer, S., Spieß, S., Brand, M., & Hans, S. (2017). Dlx3b/4b is required for early-born but not later-forming sensory hair cells during zebrafish inner ear development. Biology Open, 6, 1270e1278. Seiradake, E., Jones, E. Y., & Klein, R. (2016). Structural perspectives on axon guidance. Annual Review of Cell and Developmental Biology, 32, 577e608. Sekimizu, K., Nishioka, N., Sasaki, H., Takeda, H., Karlstrom, R. O., & Kawakami, A. (2004). The zebrafish iguana locus encodes Dzip1, a novel zinc-finger protein required for proper regulation of Hedgehog signaling. Development (Cambridge, England), 131, 2521e2532. Serluca, F. C. (2008). Development of the proepicardial organ in the zebrafish. Developmental Biology, 315, 18e27.

V. Scientific Research

590

45. Zebrafish as a Model to Understand Vertebrate Development

Shellard, A., & Mayor, R. (2016). Chemotaxis during neural crest migration. Seminars in Cell and Developmental Biology, 55, 111e118. Shimizu, T., Bae, Y.-K., & Hibi, M. (2006). Cdx-Hox code controls competence for responding to Fgfs and retinoic acid in zebrafish neural tissue. Development (Cambridge, England), 133, 4709e4719. Shimozono, S., Iimura, T., Kitaguchi, T., Higashijima, S.-I., & Miyawaki, A. (2013). Visualization of an endogenous retinoic acid gradient across embryonic development. Nature, 496, 363e366. Shindo, A. (2018). Models of convergent extension during morphogenesis. Wiley Interdisciplinary Reviews. Developmental Biology, 7. Sinn, R., & Wittbrodt, J. (2013). An eye on eye development. Mechanisms of Development, 130, 347e358. Sire, J.-Y., & Akimenko, M.-A. (2003). Scale development in fish: A review, with description of sonic hedgehog (shh) expression in the zebrafish (Danio rerio). International Journal of Developmental Biology, 48, 233e247. Snow, C. J., & Henry, C. A. (2009). Dynamic formation of microenvironments at the myotendinous junction correlates with muscle fiber morphogenesis in zebrafish. Gene Expression Patterns, 9, 37e42. Solnica-Krezel, L., & Driever, W. (1994). Microtubule arrays of the zebrafish yolk cell: Organization and function during epiboly. Development (Cambridge, England), 120, 2443e2455. Solomon, K. S., & Fritz, A. (2002). Concerted action of two dlx paralogs in sensory placode formation. Development (Cambridge, England), 129, 3127e3136. Staudt, D., & Stainier, D. (2012). Uncovering the molecular and cellular mechanisms of heart development using the zebrafish. Annual Review of Genetics, 46, 397e418. Stellabotte, F., & Devoto, S. H. (2007). The teleost dermomyotome. Developmental Dynamics: An Official Publication of the American Association of Anatomists, 236, 2432e2443. Stellabotte, F., Dobbs-McAuliffe, B., Ferna´ndez, D. A., Feng, X., & Devoto, S. H. (2007). Dynamic somite cell rearrangements lead to distinct waves of myotome growth. Development (Cambridge, England), 134, 1253e1257. Stickney, H. L., Barresi, M. J., & Devoto, S. H. (2000). Somite development in zebrafish. Developmental Dynamics: An Official Publication of the American Association of Anatomists, 219, 287e303. Stooke-Vaughan, G. A., Huang, P., Hammond, K. L., Schier, A. F., & Whitfield, T. T. (2012). The role of hair cells, cilia and ciliary motility in otolith formation in the zebrafish otic vesicle. Development, 139, 1777e1787. Stower, M. J., & Bertocchini, F. (2017). The evolution of amniote gastrulation: The blastopore-primitive streak transition. Wiley Interdisciplinary Reviews. Developmental Biology, 6. Stra¨hle, U., & Jesuthasan, S. (1993). Ultraviolet irradiation impairs epiboly in zebrafish embryos: Evidence for a microtubule-dependent mechanism of epiboly. Development (Cambridge, England), 119, 909e919. Streit, A. (2004). Early development of the cranial sensory nervous system: From a common field to individual placodes. Developmental Biology, 276, 1e15. Streit, A. (2018). Specification of sensory placode progenitors: Signals and transcription factor networks. International Journal of Developmental Biology, 62(1e2-3), 195e205. Svetic, V., Hollway, G. E., Elworthy, S., Chipperfield, T. R., Davison, C., Adams, R. J., et al. (2007). Sdf1a patterns zebrafish melanophores and links the somite and melanophore pattern defects in choker mutants. Development (Cambridge, England), 134, 1011e1022. Talbot, W. S., Trevarrow, B., Halpern, M. E., Melby, A. E., Farr, G., Postlethwait, J. H., et al. (1995). A homeobox gene essential for zebrafish notochord development. Nature, 378, 150e157. Tang, D., Lin, Q., He, Y., Chai, R., & Li, H. (2016). Inhibition of H3K9me2 reduces hair cell regeneration after hair cell loss in the

zebrafish lateral line by down-regulating the Wnt and Fgf signaling pathways. Frontiers in Molecular Neuroscience, 9, 39. Taverna, E., Go¨tz, M., & Huttner, W. B. (2014). The cell biology of neurogenesis: Toward an understanding of the development and evolution of the neocortex. Annual Review of Cell and Developmental Biology, 30, 465e502. Tawk, M., Araya, C., Lyons, D. A., Reugels, A. M., Girdler, G. C., Bayley, P. R., et al. (2007). A mirror-symmetric cell division that orchestrates neuroepithelial morphogenesis. Nature, 446, 797e800. Taylor, C. R., Montagne, W. A., Eisen, J. S., & Ganz, J. (2016). Molecular fingerprinting delineates progenitor populations in the developing zebrafish enteric nervous system. Developmental Dynamics: An Official Publication of the American Association of Anatomists, 245, 1081e1096. Tehrani, Z., & Lin, S. (2011). Antagonistic interactions of hedgehog, Bmp and retinoic acid signals control zebrafish endocrine pancreas development. Development (Cambridge, England), 138, 631e640. Telfer, W. R., Busta, A. S., Bonnemann, C. G., Feldman, E. L., & Dowling, J. J. (2010). Zebrafish models of collagen VI-related myopathies. Human Molecular Genetics, 19, 2433e2444. Theveneau, E., & Mayor, R. (2012). Neural crest delamination and migration: From epithelium-to-mesenchyme transition to collective cell migration. Developmental Biology, 366, 34e54. Thisse, B., & Thisse, C. (2015). Formation of the vertebrate embryo: Moving beyond the Spemann organizer. Seminars in Cell and Developmental Biology, 42, 94e102. Tomar, R., Mudumana, S. P., Pathak, N., Hukriede, N. A., & Drummond, I. A. (2014). osr1 is required for podocyte development downstream of wt1a. Journal of the American Society of Nephrology, 25, 2539e2545. Trinh, L. A., & Stainier, D. Y. R. (2004). Fibronectin regulates epithelial organization during myocardial migration in zebrafish. Developmental Cell, 6, 371e382. Valentin, G., Haas, P., & Gilmour, D. (2007). The chemokine SDF1a coordinates tissue migration through the spatially restricted activation of Cxcr7 and Cxcr4b. Current Biology, 17, 1026e1031. Varga, Z. M., Amores, A., Lewis, K. E., Yan, Y. L., Postlethwait, J. H., Eisen, J. S., et al. (2001). Zebrafish smoothened functions in ventral neural tube specification and axon tract formation. Development (Cambridge, England), 128, 3497e3509. Vasilyev, A., Liu, Y., Mudumana, S., Mangos, S., Lam, P.-Y., Majumdar, A., et al. (2009). Collective cell migration drives morphogenesis of the kidney nephron. PLoS Biology, 7, e9. Wagner, D. E., Weinreb, C., Collins, Z. M., Briggs, J. A., Megason, S. G., & Klein, A. M. (2018). Single-cell mapping of gene expression landscapes and lineage in the zebrafish embryo. Science, 360(6392), 981e987. Wang, G., Manning, M. L., & Amack, J. D. (2012). Regional cell shape changes control form and function of Kupffer’s vesicle in the zebrafish embryo. Developmental Biology, 370, 52e62. Ward, L., Evans, S. E., & Stern, C. D. (2017). A resegmentation-shift model for vertebral patterning. Journal of Anatomy, 230, 290e296. Warga, R. M., & Kimmel, C. B. (1990). Cell movements during epiboly and gastrulation in zebrafish. Development (Cambridge, England), 108, 569e580. Webb, A. E., Driever, W., & Kimelman, D. (2008). Psoriasis regulates epidermal development in zebrafish. Developmental Dynamics: An Official Publication of the American Association of Anatomists, 237, 1153e1164. Weber, T., & Ko¨ster, R. (2013). Genetic tools for multicolor imaging in zebrafish larvae. Methods San Diego Calif, 62, 279e291. Weicksel, S. E., Gupta, A., Zannino, D. A., Wolfe, S. A., & Sagerstro¨m, C. G. (2014). Targeted germ line disruptions reveal general and species-specific roles for paralog group 1 hox genes in zebrafish. BMC Developmental Biology, 14, 25.

V. Scientific Research

References

Whitfield, T. T. (2013). Shedding new light on the origins of olfactory neurons. ELife, 2, e00648. Whitfield, T. T., Granato, M., van Eeden, F. J., Schach, U., Brand, M., Furutani-Seiki, M., et al. (1996). Mutations affecting development of the zebrafish inner ear and lateral line. Development (Cambridge, England), 123, 241e254. Whitfield, T. T., Riley, B. B., Chiang, M.-Y., & Phillips, B. (2002). Development of the zebrafish inner ear. Developmental Dynamics: An Official Publication of the American Association of Anatomists, 223, 427e458. Whitlock, K. E., & Westerfield, M. (2000). The olfactory placodes of the zebrafish form by convergence of cellular fields at the edge of the neural plate. Development (Cambridge, England), 127, 3645e3653. Wilcockson, S. G., Sutcliffe, C., & Ashe, H. L. (2017). Control of signaling molecule range during developmental patterning. Cellular and Molecular Life Sciences., 74, 1937e1956. Wilson, S. W., Brand, M., & Eisen, J. S. (2002). Patterning the zebrafish central nervous system. Results and Problems in Cell Differentiation, 40, 181e215. Wilson, S. W., Ross, L. S., Parrett, T., & Easter, S. S. (1990). The development of a simple scaffold of axon tracts in the brain of the embryonic zebrafish, Brachydanio rerio. Development (Cambridge, England), 108, 121e145. Windner, S. E., Bird, N. C., Patterson, S. E., Doris, R. A., & Devoto, S. H. (2012). Fss/Tbx6 is required for central dermomyotome cell fate in zebrafish. Biology Open, 1, 806e814. Wingert, R. A., Selleck, R., Yu, J., Song, H.-D., Chen, Z., Song, A., et al. (2007). The cdx genes and retinoic acid control the positioning and segmentation of the zebrafish pronephros. PLoS Genetics, 3, 1922e1938. Woods, I. G., & Talbot, W. S. (2005). The you gene encodes an EGF-CUB protein essential for Hedgehog signaling in zebrafish. PLoS Biology, 3, e66. Wragg, J., & Mu¨ller, F. (2016). Transcriptional regulation during zygotic genome activation in zebrafish and other anamniote embryos. Advances in Genetics, 95, 161e194. Wu, D., Freund, J. B., Fraser, S. E., & Vermot, J. (2011). Mechanistic basis of otolith formation during teleost inner ear development. Developmental Cell, 20, 271e278.

591

Yabe, T., & Takada, S. (2016). Molecular mechanism for cyclic generation of somites: Lessons from mice and zebrafish. Development Growth and Differentiation, 58, 31e42. Yaniv, K., Isogai, S., Castranova, D., Dye, L., Hitomi, J., & Weinstein, B. M. (2006). Live imaging of lymphatic development in the zebrafish. Nature Medicine, 12, 711e716. Yoshihara, Y. (2009). Molecular genetic dissection of the zebrafish olfactory system. Results and Problems in Cell Differentiation, 47, 97e120. Yu, H.-H., & Moens, C. B. (2005). Semaphorin signaling guides cranial neural crest cell migration in zebrafish. Developmental Biology, 280, 373e385. Zeller, J., Schneider, V., Malayaman, S., Higashijima, S., Okamoto, H., Gui, J., et al. (2002). Migration of zebrafish spinal motor nerves into the periphery requires multiple myotome-derived cues. Developmental Biology, 252, 241e256. Zhang, J., Lefebvre, J. L., Zhao, S., & Granato, M. (2004). Zebrafish unplugged reveals a role for muscle-specific kinase homologs in axonal pathway choice. Nature Neuroscience, 7, 1303e1309. Zhao, X., Sirbu, I. O., Mic, F. A., Molotkova, N., Molotkov, A., Kumar, S., et al. (2009). Retinoic acid promotes limb induction through effects on body axis extension but is unnecessary for limb patterning. Current Biology CB, 19, 1050e1057. Zhou, W., Boucher, R. C., Bollig, F., Englert, C., & Hildebrandt, F. (2010). Characterization of mesonephric development and regeneration using transgenic zebrafish. American Journal of PhysiologyeRenal Physiology, 299, F1040eF1047. Zhou, Y., Cashman, T. J., Nevis, K. R., Obregon, P., Carney, S. A., Liu, Y., et al. (2011). Latent TGF-b binding protein 3 identifies a second heart field in zebrafish. Nature, 474, 645e648. Zinski, J., Bu, Y., Wang, X., Dou, W., Umulis, D., & Mullins, M. C. (2017). Systems biology derived source-sink mechanism of BMP gradient formation. ELife, 6. Zinski, J., Tajer, B., & Mullins, M. C. (2018). TGF-b family signaling in early vertebrate development. Cold Spring Harb Perspect Biol. 10(6). a. Zorn, A. M., & Wells, J. M. (2009). Vertebrate endoderm development and organ formation. Annual Review of Cell and Developmental Biology, 25, 221e251.

V. Scientific Research