103 Cardiac Development in the Zebrafish

Chapter 1.4

Cardiac Development in the Zebrafish

Ian C. Scott1 and Deborah Yelon2

1The Hospital for Sick Children, Program in Developmental and Stem Cell Biology, Department of Molecular Genetics, University of Toronto, Toronto, Canada2Skirball Institute of Biomolecular Medicine, Developmental Genetics Program, New York University School of Medicine, New York, USA

I. IntroductIon

Over the past decade, the zebrafish (Danio rerio) has become a popular model organism for the study of ver-tebrate development. Since the optically transparent embryo develops rapidly, the entirety of embryogenesis can be visualized in real time (Kimmel et al., 1995). The zebrafish embryo is also readily amenable to embryologi-cal approaches such as microinjection, lineage tracing and transplantation. The feasibility of conducting large-scale classical genetic screens in zebrafish has further increased interest in its use for discovering genes essential for ver-tebrate development (Patton and Zon, 2001). In addition to forward (phenotype-driven) genetic approaches, the zebrafish also offers opportunities for reverse (gene-driven) genetic approaches, via the use of antisense morpholino (MO) oligonucleotides to target specific genes (Ekker, 2000). Together, the combination of optical accessibil-ity, embryonic manipulability, and feasibility of genetic approaches makes the zebrafish a unique and exciting model organism for developmental biologists.

The attributes of the zebrafish have special appeal for the analysis of heart development. The embryonic zebrafish heart can be imaged live and develops quickly, with the heartbeat starting by 22 hours post-fertilization (hpf). This has allowed discovery of mutations affecting heart development via straightforward visual examination of mutant embryos under a dissecting microscope. To date, a large number of mutations affecting multiple aspects of heart development have been isolated from a variety of forward genetic screens (Chen et al., 1996; Stainier et al., 1996; Alexander et al., 1998; Warren et al., 2000; Beis et al., 2005). Studies of mutant phenotypes have provided significant new insights into the genetic mechanisms

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underlying cardiogenesis. The ease with which MOs can be used to examine the roles of specific genes has fur-ther extended the number of cardiac researchers turning to the zebrafish model to gauge gene function. Analysis of cardiac defects in mutant or MO-injected embryos is relatively convenient, since zebrafish embryos can survive up to seven days post-fertilization (dpf) in the absence of cardiovascular function via oxygen diffusion. In con-trast, mouse mutations that compromise aspects of heart or vascular development typically cause early embryonic lethality, necessitating the use of complex temporal and spatial regulation of gene function to assay roles in later developmental steps.

Here, we review the utility of zebrafish for the study of heart development. We begin by summarizing the experi-mental approaches that have been particularly useful for analysis of heart formation. We then outline the major phases of heart development in zebrafish, emphasizing our current understanding of the molecular and cellular regula-tion of each step. Finally, we reflect on the application of the zebrafish model to the study of human heart disease. Throughout, we highlight the important impact of the combinatorial use of imaging, embryology and genetics in zebrafish on our understanding of how genes regulate car-diac development.

II. ExpErImEntal approachEs for analysIs of hEart dEvElopmEnt In zEbrafIsh

While most often recognized as an opportune organism for forward genetic screens, zebrafish have numerous other experimental strengths. This is, in large part, due to the

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part | 1 Heart Evolution104

fact that fertilization and embryonic development occur entirely external to the mother. Embryos can be collected, examined and manipulated at the earliest stages of devel-opment. Mated pairs of fish yield several hundred embryos per week, providing an abundance of material. Since adult zebrafish are small in size (typically 1.5–2 inches in length) and relatively easy to maintain, an individual labo-ratory can keep a variety of wild-type, mutant and trans-genic lines inexpensively. Therefore, a growing number of researchers are exploiting the zebrafish model for the study of heart development. In this section, we summa-rize the experimental approaches routinely used in the zebrafish.

II.a. Genetics

The application of forward genetic screens in the fruit fly Drosophila and the nematode Caenorhabditis elegans has had a profound impact on our understanding of the genetic events that govern animal development (Brenner, 1974; Nusslein-Volhard and Wieschaus, 1980). These phenotype-driven screens are ideally suited for discovering new genes whose function is critical to a particular developmental process. Although the Drosophila heart equivalent (the dorsal vessel) is a relatively simple organ, forward genetic screens in this organism have uncovered several genes that are also crucial for vertebrate heart development. One strong example is the Drosophila gene tinman (see Chapter 1.2); mutation of tinman eliminates the dorsal vessel (Bodmer, 1993) and mutation of the mouse homolog Nkx2-5 results in profound perturbations in heart development (Lyons et al., 1995) (see Chapter 9.1). Furthermore, mutations in NKX2-5 are causative for congenital heart disease (CHD) in humans (Schott et al., 1998). Thus, it appears that many of the genetic pathways that regulate heart development are highly-conserved throughout much of the animal kingdom.

Of course, the Drosophila dorsal vessel lacks much of the complexity found in the vertebrate heart. Forward genetic screens in the chick and frog are not feasible, whereas this approach in the mouse embryo is labor- intensive, due to its in utero development, and expensive, due to the costs associated with housing large populations of mice. The zebrafish represents an ideal solution. Due to the small size, high fecundity and relatively low costs of maintenance of zebrafish, laboratories can easily establish zebrafish populations sufficient in size to carry out for-ward genetic screens (Patton and Zon, 2001). As the trans-lucent zebrafish embryo develops externally in a simple salt solution, defects in heart development can be assayed rapidly in live embryos under a dissecting microscope. More detailed phenotypic examination is facilitated by screening with molecular markers, especially transgenes that express fluorescent reporters in specific cell types (Jin et al., 2007).

Traditionally, forward genetic screens in zebrafish have taken advantage of the potent mutagen N-ethyl- N-nitrosourea (ENU), an alkylating agent that introduces random point mutations into the genome (Patton and Zon, 2001). Male founder (F0) adults are treated with ENU, resulting in the mosaic accumulation of point mutations in their sperm. Founders are repeatedly bred with wild-type female fish to establish multiple F1 progeny per F0 male. These F1 fish are then bred to derive F2 families, in which half of the siblings from a given cross will carry a particu-lar point mutation. F2 siblings are then intercrossed, and embryos are screened for defects in heart development. This standard diploid genetic screen is very effective for recovering recessive mutations, and more than 300 muta-tions affecting discrete steps in zebrafish heart develop-ment have been identified using this approach (Chen et al., 1996; Stainier et al., 1996; Warren et al., 2000; Beis et al., 2005). As will be discussed later in this chapter, the proc-esses disrupted by these mutations include specification of cardiac progenitors, assembly of the linear heart tube, chamber morphogenesis and valve formation.

While forward genetic screens are very powerful, they can be time-consuming and laborious, given the gen-eration time of zebrafish (around three months) and the number of F2 families that must be established for a large-scale screen. An interesting feature of zebrafish is that haploid and diploid embryos can be created gynogeneti-cally, such that their genome is derived solely from their mother (Corley-Smith et al., 1996). This technique has been exploited in screens analyzing cardiac defects in the hemizygous haploid progeny of F1 generation females, greatly reducing the time and number of fish necessary for an effective screen (Alexander et al., 1998). Although haploid embryos do not survive beyond embryogenesis, the early steps of heart development proceed relatively normally. This allows rapid identification of mutations affecting heart development that can then be further char-acterized and mapped in the diploid state.

The high efficiency of ENU mutagenesis is unfortu-nately counterbalanced with the challenge of identifying the affected genes. Positional cloning of ENU-induced mutations typically involves high resolution linkage analy-sis requiring the isolation of 500–3,000 mutant embryos (Bahary et al., 2004). While the nearly complete sequence of the zebrafish genome has facilitated positional cloning, identification of a mutated gene can still take many months of work. An alternative approach to ENU mutagenesis is insertional mutagenesis using retroviruses. While not as efficient as ENU in introducing mutations, retroviral mutagenesis has the advantage of marking mutated genes with a molecular tag, making their identification possible in a much shorter period of time (Amsterdam et al., 1999). A similar insertional approach employing transposons to disrupt genes is currently being developed (Sivasubbu et al., 2007; Nagayoshi et al., 2008).

chapter | 1.4 Cardiac Development in the Zebrafish 105

The ability to modify genes in mice precisely via homologous recombination in embryonic stem cells has revolutionized the study of gene function in vertebrates (Capecchi, 2005). The power of this technique has been further extended by the use of spatially- and temporally-regulated gene modifications. Unfortunately, homologous recombination approaches are not currently feasible in zebrafish, and it will be challenging to overcome the tech-nical hurdle of the early establishment of germ cell fate through inheritance of maternally provided cytoplasmic determinants (Raz, 2003). While the introduction of MOs has been extremely beneficial (see below), the availability of a loss-of-function mutant is often required to ensure that the MO-induced phenotype is specific. A promising devel-opment has been the use of targeting induced local lesions in genomes (TILLING) to isolate mutations in a given gene (Sood et al., 2006). In this reverse-genetic approach, PCR targeted against a known gene is used to screen DNA samples derived from a library of F1 fish following ENU mutagenesis. Mutations are detected by sequencing or by using a restriction enzyme that recognizes DNA base pair mismatches. In the coming years, the large-scale applica-tion of TILLING and insertional mutagenesis will create an invaluable resource of mutations in many (if not all) zebrafish genes. Commercial and academic consortia are currently scaling-up efforts toward this goal.

While they are not strictly gene-based, chemical genetic approaches have recently become popular in zebrafish (Peterson et al., 2000; Murphey and Zon, 2006). The aqueous habitat of the zebrafish embryo makes it particularly suitable for treatment with small molecules. Inhibitors of known specificity can be used to ascertain the role of select signaling pathways in heart development. An advantage is that chemicals can be added and removed at defined time points, allowing for the temporal resolution of requirements for specific signals. Complementing this targeted approach, the large clutch size of zebrafish makes it possible to screen through large libraries of several thou-sand small molecules in search of previously uncharacter-ized chemicals that cause specific developmental defects, or that mitigate or rescue specific mutant phenotypes. As will be discussed later in this chapter, this chemical genetic approach may prove to be a powerful method for analyzing zebrafish models of human heart disease, since the effects of small molecules may illuminate disease mechanisms and suggest possible treatments.

II.b. regulation of Gene activity

Besides standard genetic approaches, additional tech-niques are available to analyze gene function in zebrafish. These strategies take advantage of the ease of introduc-ing exogenous material into the zebrafish embryo at early stages. At the 1-cell stage, the zebrafish embryo is large,

roughly 0.7 mm in diameter. It is therefore straightforward to microinject RNA, DNA, or protein into the 1-cell stage embryo using a fine glass needle. Following fertilization, a stream of maternally deposited proteins and RNA flows from the yolk into the embryo proper; these gene products are instrumental in supporting the initial phases of embryo-genesis (Kimmel and Law, 1985). Thus, material microin-jected into the yolk at the 1-cell stage is passed into the cell, inherited by its descendants, and thereby distributed into every cell of the embryo. As embryonic cell cleav-ages proceed, a finer glass needle can similarly be used to transfer material to individual blastomeres, although this technique becomes impractical beyond the 1,000-cell stage without the use of more sophisticated iontophoresis equipment. Given the large clutch sizes and ease of injec-tion of zebrafish, researchers can routinely inject hundreds of embryos in a single session.

Because of the convenience of microinjection, the advent of chemically-modified antisense MO DNA oligo-nucleotides has allowed the selective knockdown of genes in the zebrafish embryo (Ekker, 2000). Injected at the 1-cell stage, MOs target either the translational start site or splice sites of a specific mRNA. Interaction of an MO and its target sterically blocks translation or splicing, effec-tively interfering with gene function. MOs are highly sta-ble, so they remain effective for at least the first five days of embryogenesis, providing ample time to analyze the major phases of heart development. While MOs provide powerful tools for analysis of gene function, care must be taken in the interpretation of MO results. The efficacy of individual MOs varies widely and must be determined empirically. Moreover, MOs may have off-target effects and sometimes appear to cause nonspecific toxicity (Robu et al., 2007). The amount of MO injected should be care-fully titrated to aid in gauging specific versus nonspecific effects. Importantly, multiple independent MOs, targeting different regions of a gene, should also be used to verify that an observed phenotype is gene-specific. In C. elegans, use of bacterial libraries encoding double-stranded RNA (dsRNA) has proven to be an effective means of knocking-down genes in a high-throughput manner (Simmer et al., 2003). In contrast to MOs, dsRNA does not currently seem to be effective for specific gene knockdown in zebrafish (Oates et al., 2000).

As a complement to loss-of-function approaches like MO knockdown, gain-of-function experiments, in which wild-type genes or their modified forms are overexpressed, are often informative. In vitro transcribed RNA is read-ily translated following injection. When available, RNA encoding dominant-negative or constitutively active forms of a protein can be used for functional and epistasis analy-sis. Typically, gain-of-function experiments in zebrafish overexpress a gene throughout the entire embryo from the 1-cell stage. Unlike the case in the frog Xenopus laevis, the fates of early blastomeres in the zebrafish embryo are not


reproducible (Kimmel et al., 1990; Ho and Kimmel, 1993), so injections cannot specifically target the heart lineage. To achieve this, transgenic approaches are required. DNA transgenes injected at the 1-cell stage result in mosaic expression in F0 embryos. The zebrafish cardiac myosin light chain 2 (cmlc2) promoter drives strong expression specifically in cardiomyocytes (Huang et al., 2003). In F0 injected embryos it is feasible to achieve expression in a large proportion of the myocardium. To attain stable, nonmosaic transgenic lines, F0 embryos are raised to adulthood and screened for germline transmission of the transgene. Use of transposons to derive transgenic ani-mals has been shown to increase the frequency of germline transmission greatly, and has become a favored method for transgenesis in zebrafish (Kawakami, 2005).

Gene overexpression can often cause a lethal pheno-type. Temporal control of transgene expression is therefore desirable. Additionally, temporally-regulated transgene expression can allow examination of when gene activity is required. The zebrafish hsp70 promoter provides a useful tool to conditionally overexpress gene products throughout the embryo (Halloran et al., 2000). This heat-shock pro-moter is quiescent at standard temperature (28.5°C), but is activated on short incubations at 37°C. Other methods, including the binary tetracycline-regulated and Gal4/UAS systems, provide additional means of temporally and spa-tially restricting transgene expression in zebrafish (Koster and Fraser, 2001; Huang et al., 2005). A further option is a recombination-based system, in which transgenic expres-sion of Cre can modify a second transgene (Langenau et al., 2005). The future derivation of multiple regionally restricted cardiac promoters will increase the usefulness of these transgenic approaches for gene misexpression.

II.c. Embryological manipulation

In order to analyze the cellular consequences of altering gene function in the zebrafish embryo, it is extremely ben-eficial to employ embryological techniques such as fate mapping and transplantation. The external development of the zebrafish embryo makes such embryonic manipulations possible. Embryonic cells can be individually labeled, tracked in real time, and transferred between embryos. When coupled with the availability of mutants, the use of MOs and the ability to misexpress genes, embryological approaches in zebrafish can reveal a wealth of informa-tion about the mechanisms regulating fate assignment and developmental potential.

A fate mapping or lineage tracing approach is used to determine the progeny of a labeled cell or group of cells retrospectively. This is especially useful when molecu-lar markers specific to a progenitor population do not exist, as is the case for myocardial progenitors in the early embryo prior to and during gastrulation (Fig. 1A). Putative

progenitor cells can be labeled at early stages, and the fate, migration and growth of these cells can be determined from the locations and numbers of labeled cells found at later stages (Fig. 2A–D). Individual cells can be labeled via direct injection of fluorophore-conjugated dextrans. However, beyond the 1,000-cell stage, when microinjec-tion of single cells is technically challenging, it is particu-larly useful to label cells with a caged fluorescein dextran conjugate; its fluorescence is inhibited by a chemical moiety until it is activated by UV light (Kozlowski et al., 1997) (Fig. 2A–D). Similarly, the photoconvertible pro-tein Kaede changes its fluorescence to emit red rather than green light following UV exposure (Hatta et al., 2006). By finely focusing UV light via microscopy, individual cells or groups of cells in embryos injected with caged fluores-cein or kaede RNA at the 1-cell stage can be labeled at any optically accessible stage and location and followed sub-sequently. These strategies have been employed to gener-ate fate maps of myocardial and endocardial progenitors in wild-type zebrafish embryos (Stainier et al., 1993; Lee et al., 1994; Keegan et al., 2004; Schoenebeck et al., 2007) (Fig. 1A,B). Comparison of the wild-type fate map to fate maps in embryos with cardiac defects has been instru-mental in elucidating whether and how particular genes or pathways impact cardiac fate assignment (Keegan et al., 2004, 2005; Schoenebeck et al., 2007).

Transplantation approaches provide a powerful means to assay the potential of a cell in various environments (Carmany-Rampey and Moens, 2006) (Fig. 2E–H). Using a fine glass needle and mild suction, cells are removed from a donor embryo and placed into a host embryo, typically at midblastula stages. The donor embryo is labeled with a lineage tracer, such that the fate of donor-derived cells in the host embryo can be followed over time. This provides an assay to gauge the autonomy, or cellular requirement, of gene function. By placing mutant or wild-type cells into a host of the contrary genotype, the influence of environ-ment on phenotype can be observed.

II.d. Imaging

Aside from providing aesthetic pleasure, the optical clar-ity and rapid development of the zebrafish embryo make it an ideal model organism for high-resolution imaging of the developing heart (Schoenebeck and Yelon, 2007). Internal organs are readily visualized via whole-mount in situ hybridization and immunohistochemistry. Online databases contain the results of high-throughput in situ analysis of thousands of genes, and numerous myocardial and endothelial markers exist for zebrafish (Thisse et al., 2004). Furthermore, the optical properties of the zebrafish embryo allow detailed cellular and subcellular examina-tion of the cell biological traits of the developing heart. Myocardial cell polarity, cytoskeletal organization and cell


(A) (B) (C) (D) (E)

40% epiboly 7 somites 21 somites 30 hpf 48 hpf

fIGurE 1 Major phases of heart formation in the zebrafish embryo. (A) Lateral view of the blastula at the 40% epiboly stage (5 hpf), depicting locations of ventricular (red) and atrial (yellow) progenitors as determined by fate mapping (Keegan et al., 2004). (B) Dorsal view, anterior up, at the 7-somite stage (12.5 hpf), depicting locations of ventricular and atrial progenitors in the lateral mesoderm as determined by fate mapping (Schoenebeck et al., 2007). (C) By the 21-somite stage (19.5 hpf), cardiac fusion has created a cardiac cone at the embryonic midline. Atrial (yellow) cardiomyocytes surround the ventricular (red) cardiomyocytes (Berdougo et al., 2003). (D) At 30 hpf, heart tube elongation is complete. (E) Frontal view, anterior up, at 48 hpf; the ventricle and atrium are morphologically distinct and asymmetrically looped. Artwork based on images from Schoenebeck and Yelon (2007).

(A) (B) (C) (D)

(E) (F) (G) (H)

fIGurE 2 Techniques for embryological manipulation in zebrafish. (A–D) Fate mapping using caged fluorescein dextran as a lineage tracer. (A) Caged fluorescein dextran is injected into the embryo at the 1-cell stage. The lineage tracer is inherited by all of the cells of the embryo, but does not fluoresce until it is photoactivated. (B) Photoactivation using UV light creates fluorescence in selected cells. In this example, five blastomeres (green) are selected at the margin of the embryo at 40% epiboly, as in the fate-mapping experiments of Keegan and colleagues (2004). (C) The location of photoactivated cells is recorded relative to anatomical landmarks or transgene expression patterns. In this view of the animal pole of the embryo at 50% epiboly, the longitudinal location (90°) of the photoactivated cells is recorded relative to the expression of Tg(gsc:gfp) at the dorsal midline. (D) The fate of photoactivated cells is determined at later developmental stages. Here, immunohistochemistry at 44 hpf detects fluorescein (blue) in two cardiomyocytes derived from the photoactivated cells. (E–H) Transplantation using fluorescein dextran as a lineage tracer. (E) Fluorescein dextran is injected into the donor embryo at the 1-cell stage. All of the cells of the donor embryo are therefore fluorescent. (F) Using mild suction, cells are removed from the donor embryo at the midblastula stage. (G) The removed cells are transferred to a host embryo. Placement near the margin of the host embryo facilitates contribution to cardiac lineages. (H) The fate of donor-derived cells is determined at later developmental stages. Only the donor-derived cells contain the lineage tracer. In this example, two fluorescent cardiomyocytes are detectable at 44 hpf.


shape can be analyzed in fixed samples via immunofluo-rescence and confocal microscopy (Trinh and Stainier, 2004; Trinh et al., 2005; Rohr et al., 2006; Auman et al., 2007). Transgenic zebrafish that express cytoplasmic, nuclear or membrane-localized fluorescent proteins allow dynamic analysis of cell movement, cell number and cell shape in the living embryo (Rottbauer et al., 2002; Auman et al., 2007; Holtzman et al., 2007). It is particularly excit-ing to use transgenic zebrafish for time-lapse analysis during stages of heart development when considerable morphogenesis is occurring, including the assembly of the heart tube, the expansion of the cardiac chambers and the formation of the atrioventricular valve (Beis et al., 2005; Rohr et al., 2006; Holtzman et al., 2007). High-resolu-tion imaging of the live heart is, of course, complicated by the heartbeat, which by 48 hpf occurs at a rate of roughly 180 beats per minute in the zebrafish embryo (Baker et al., 1997). Heartbeat can be temporarily inhibited by addition and removal of chemicals such as 2,3-butanedi-one monoxime and the anesthetic tricaine (Bartman et al., 2004).

Although it is often desirable to stop the heart in order to collect images, it is also of interest to use imag-ing techniques to assess cardiac function. By combining high-speed microscopy and custom computational algo-rithms with transgenic embryos expressing fluorescent proteins in the myocardium, endothelium and circulating red blood cells, it is possible to measure fractional short-ening, flow velocity, ejection fraction and additional functional parameters (Hove et al., 2003; Forouhar et al., 2006). Additionally, calcium indicator dyes and transgenic calcium sensor proteins can be used to assess cardiac con-duction in the live embryo (Sedmera et al., 2003; Milan et al., 2006; Arnaout et al., 2007). These techniques are fundamental for analysis of the large number of zebrafish mutations that affect cardiac function. The ongoing devel-opment of more rapid microscopic techniques will allow more detailed analysis of cell morphology in a functional heart. One promising technique is selective plane illumi-nation microscopy (SPIM), which is well-suited to imag-ing samples the size of the zebrafish heart (Huisken et al., 2004; Huisken and Stainier, 2007).

Altogether, the confluence of genetic, embryological and imaging techniques available for zebrafish facilitates what has been termed an “in vivo cell biology” approach (Beis and Stainier, 2006). A variety of cell behaviors – including fate assignment, differentiation, proliferation, adhesion, polarity, movement, shape change, contractility and conduction – can be examined in the context of the developing embryo. This presents an ideal model for the direct analysis of how genes regulate heart development at the level of the cell. Thus, work in zebrafish can reveal key mechanisms by which mutations are translated into devel-opmental defects and disease.

III. mEchanIsms rEGulatInG zEbrafIsh hEart dEvElopmEnt

In a relatively short period of time, our understanding of the mechanisms regulating zebrafish heart development has expanded dramatically. Large-scale genetic screens have yielded plentiful collections of mutations, and the characterization of mutant phenotypes and cloning of mutated genes have revealed key genetic pathways con-trolling cardiogenesis. In concert with these genetic stud-ies, high-resolution imaging of the developing heart has provided insight into the fundamental cellular mechanisms underlying cardiac morphogenesis. In this section, we out-line the major phases of the first two days of heart develop-ment in zebrafish, emphasizing our current understanding of the molecular and cellular regulation of each step.

III.a. overview of stages of zebrafish development

Following fertilization, the zebrafish embryo develops in a simple salt solution at 28.5°C (Kimmel et al., 1995). The embryo is found on top of a yolk, and is surrounded by a transparent chorion that provides protection. A series of rapid cell divisions converts the large single cell of the embryo into an unpatterned cap of over 1,000 small cells by 3 hpf. Maternally provided protein, RNA and other bio-molecules play key roles in supporting these early cleav-ages. At the midblastula transition (roughly 3 hpf), zygotic transcription commences and is required for the initia-tion of gastrulation and further embryonic development. Morphogenesis of the embryo begins at dome stage (just after 4 hpf) as the yolk pushes upwards into the embryo, moving cells away from the inner core and creating a monolayer, termed the epiblast, that covers the upper 30% of the yolk. This is followed by epiboly – the spreading of the epiblast vegetally over the yolk. At 40% epiboly (5 hpf) gastrulation commences, with cells at the margin (the leading edge of the embryo over the yolk) involuting underneath the epiblast to form a new cell layer, termed the hypoblast. This process is critical to embryogenesis, as cells of the hypoblast will form the mesodermal and endo-dermal germ layers. Gastrulation movements consist of a combination of convergence towards the embryonic midline and extension along the future anterio–posterior (A–P) axis. The combination of these convergent extension (CE) move-ments converts the blastula into an organized, multi-layered embryo with discernible axes.

Gastrulation and epiboly are completed in the zebrafish embryo at tailbud stage (10 hpf), by which time the yolk is completely engulfed. The embryo now has an obvious anteroposterior axis, with a presumptive head and tail, and the notochord is apparent at the embryonic midline. Between 10 and 24 hpf, a number of major developmental


events occur. Rudimentary organs become evident; for example, the eye is visible by 12 hpf, and the otic vesi-cle forms by 14 hpf. Somite formation begins, with new somites being added initially at a rate of 2–3 somites per hour. At the same time, the embryo lengthens in the antero-posterior dimension, primarily via lengthening of the tail.

Heartbeat in the zebrafish embryo commences at 22 hpf, with blood flow apparent by 24 hpf. Between 24 and 48 hpf, the embryo straightens, with the head mov-ing from a position curled over the yolk to be in line with the trunk axis. Spontaneous contractions of skeletal mus-cle become more frequent, and pigmentation arises in the eyes and body. The vascular network in the embryo becomes more elaborate over this time period, as angio-genesis results in a fine network of vessels branching from the primary vasculature in a stereotypic fashion. By 48 hpf, much of organogenesis is completed and the embryo is motile and responsive to external stimuli. Over the next 24 hours, the embryo hatches from its chorion and starts to inflate its swim bladder. By 120 hpf, the embryo has depleted the energy stores present in the yolk and begins eating. Sexual maturity is reached at 2.5–3 months of age. The strikingly rapid and external development of zebrafish makes it an excellent model for the study of cardiogenesis.

III.b. cardiac progenitor specification

Fate maps of several vertebrate species indicate that car-diac progenitor cells ingress early during gastrulation, migrating to reach their destination in the anterior lat-eral plate mesoderm (ALPM) (Schoenwolf and Garcia-Martinez, 1995; Tam et al., 1997). In the zebrafish embryo, extensive fate-mapping studies have determined the ori-gins of cardiac progenitors prior to and following gastrula-tion (Stainier et al., 1993; Lee et al., 1994; Keegan et al., 2004; Schoenebeck et al., 2007). At early blastula stages (256 to 512 cells), progenitors are found at the ventro-lateral margin, 90° to 270° from the dorsal midline of the embryo (Stainier et al., 1993). The multipotent myocardial progenitor cells are intermingled with cells fated to form other tissues, including muscle and blood. Endocardial progenitor cells are also found near the margin, at more ventral positions closer to 180° (Lee et al., 1994). In the early blastula, individual myocardial progenitor cells can contribute to both the atrium and ventricle, indicating that cardiac chamber lineages are not yet separated at this stage of development (Stainier et al., 1993). After an additional hour of development at midblastula stage (1,000–2,000 cells), individual atrial and ventricular myocardial progeni-tor cells can be identified via fate mapping, indicating line-age separation by this time.

At the onset of gastrulation, the distribution of myocar-dial progenitors has shifted to encompass bilateral regions 60°–140° from the dorsal midline (Keegan et al., 2004)

(Fig. 1A). Roughly 10–20% of blastomeres within the first three rows next to the embryonic margin in this area will give rise to myocardial progeny. Organization of the atrial and ventricular progenitors is evident at this stage of devel-opment. Progenitors of ventricular myocardium are found closer to the margin and the shield, whereas those of atrial fate are found further from the margin in more ventral positions (Fig. 1A). In contrast, endocardial progenitors do not demonstrate a chamber-specific organization, and individual cells can contribute to the endocardium of both chambers. The spatial organization of ventricular and atrial myocardial progenitors persists after gastrulation, when they occupy discrete regions of the ALPM (Schoenebeck et al., 2007) (Fig. 1B). Fatemapping experiments indicate that ventricular progenitors occupy a more medial portion of the ALPM than atrial progenitors (Fig. 1B), and this mediolateral organization correlates well with the subse-quent complementary mediolateral expression patterns of the chamber-specific genes ventricular myosin heavy chain (vmhc) and atrial myosin heavy chain (amhc) (Yelon et al., 1999; Berdougo et al., 2003).

The regions of the early embryo containing myocar-dial progenitors qualify as heart fields, areas of the embryo possessing cardiac developmental potential (Jacobson and Sater, 1988). Transplantation of donor cells from noncar-diogenic regions of the blastula into the heart fields of host embryos can confer myocardial fate, although not with 100% efficiency (Lee et al., 1994; Scott et al., 2007). Thus, the signals available to cells within the heart fields are clearly important for cardiac specification. However, as other types of mesendodermal progenitor cells also arise from these regions, additional signals must be critical for commitment to cardiac lineages. These signals may be delivered at later stages, as the progenitor cells migrate to and reside within the ALPM.

Studies of zebrafish mutations have implicated multi-ple signaling pathways in the regulation of cardiac speci-fication within the heart field. For example, mutations in components of the Bmp and Fgf signaling pathways indi-cate that these pathways are essential for promoting myo-cardial fate assignment. In swirl (bmp2b) and acerebellar (fgf8) mutants, the number of cardiomyocytes is severely decreased compared to their wild-type siblings; this myo-cardial deficiency is preceded by gene expression defects, including reduced expression of the transcription factor gene nkx2.5, thought to be an early marker of myocar-dial progenitors (Reifers et al., 2000; Reiter et al., 2001). A similar phenotype is exhibited by one-eyed pinhead mutants, which lack an essential coreceptor in the Nodal signaling pathway (Reiter et al., 2001). Furthermore, in acerebellar (fgf8) and one-eyed pinhead mutants, the effects on ventricular cardiomyocytes are more dramatic than the effects on atrial cardiomyocytes, suggesting that Fgf8 and Nodal signaling may be involved in chamber fate assignment. Indeed, fate-map analysis in embryos


with reduced levels of Nodal signaling supports a model in which differential exposure to Nodal ligands, secreted from the embryonic margin, promotes the assignment of ventricular identity in the myocardial progenitors closest to the margin (Keegan et al., 2004).

Integration of Bmp, Fgf and Nodal signaling is likely to regulate expression of pivotal transcription factor genes that drive cardiomyocyte differentiation. Mutation of the transcription factor gene faust (gata5) results in reduced expression of nkx2.5 and severe cardiomyocyte deficien-cies, especially for ventricular cardiomyocytes (Reiter et al., 1999). Epistasis experiments suggest that Gata5 functions downstream of Bmp2b and Nodal signaling (Reiter et al., 2001); although Fgf8 is also necessary for normal expression of nkx2.5, it does not seem to be essen-tial for gata5 expression. Mutants lacking the transcription factor gene hands off (hand2) also have too few cardiomy-ocytes but do not exhibit defects in nkx2.5 or gata5 expres-sion, suggesting a downstream or parallel role for Hand2 (Yelon et al., 2000). Fate mapping in hands off (hand2) mutants reinforces this notion: the dimensions of the heart fields appear unchanged in hands off (hand2) mutants, yet production of differentiated cardiomyocytes from these fields is inefficient (Schoenebeck et al., 2007).

The inductive signals that promote cardiac specification are counterbalanced by repressive signals that restrict the formation of cardiac progenitors. In contrast to zebrafish mutations causing a shortage of cardiomyocytes, muta-tion of neckless (retinaldehyde dehydrogenase 2) causes

a significant cardiomyocyte surplus (Keegan et al., 2005). Retinaldehyde dehydrogenase 2 controls the rate-limit-ing step of retinoic acid (RA) synthesis; therefore, the neckless mutant phenotype, together with the phenotype of embryos treated with inhibitors of RA signaling (Fig. 3A–D), suggests that RA signaling is critical for restricting cardiomyocte production. Furthermore, fatemap analysis in embryos with reduced RA signaling has shown that RA signaling during gastrulation stages is essential for setting limits on the number of myocardial progenitors. Canonical Wnt signaling also appears to inhibit cardiac specification, although its influence varies at different developmental stages (Ueno et al., 2007). Use of heat-shock induc-ible transgenes that activate or inhibit Wnt/-catenin sig-naling has demonstrated that signaling during gastrulation blocks expression of nkx2.5, whereas signaling at earlier stages promotes nkx2.5 expression. In addition to the RA and canonical Wnt pathways, the pathways that control endothelial and hematopoietic specification contribute to the restriction of cardiac specification (Schoenebeck et al., 2007). Zebrafish mutants deficient in vessel and blood lin-eages, such as cloche, exhibit ectopic cardiomyocytes and enlarged hearts. Fate mapping indicates that this pheno-type is caused by fate transformation of a territory located adjacent to the heart field; normally, this region produces vessel and blood progenitors but not cardiac progenitors. It is not yet known how the multiple repressive influences on cardiac specification work in opposition to the inductive influences on this process. Future studies will illuminate

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fIGurE 3 Examples of signaling pathways regulating cardiac progenitor specification in zebrafish. (A–D) RA signaling restricts the number of car-diomyocytes in the zebrafish embryo. (A, B) Dorsal views, anterior to the top, comparing cmlc2 expression at the 16-somite stage (17 hpf) in wild-type (wt) embryos and siblings treated with the RA receptor antagonist BMS189453. Inhibition of RA signaling creates a significant surplus of cardiomyo-cytes residing in bilateral populations in the ALPM. Images adapted from Keegan et al. (2005). (C, D) Frontal views of hearts at 48 hpf; both chambers express the transgene Tg(cmlc2:DsRed2-nuc) (red; Mably et al., 2003) and atria are labeled with the anti-atrial myosin heavy chain (Amhc) antibody S46 (green). Inhibition of RA signaling causes the formation of enlarged hearts containing an excess number of cardiomyocytes. Images courtesy of J. S. Waxman and D. Yelon (unpublished data). (E–H) Agtrl1b signaling promotes cardiomyocyte formation in the zebrafish embryo. (E, F) Dorsal views, anterior to the top, comparing nkx2.5 expression at the 10-somite stage (14 hpf) in wt and grinch (agtrl1b) mutant embryos. Loss of Agtrl1b signaling inhibits formation of nkx2.5-expressing myocardial progenitors. (G, H) Lateral views, anterior to the left, of cmlc2 expression at 48 hpf. Inhibition of Agtrl1b signaling results in a severe deficiency of cardiomyocytes. Images adapted from Scott et al. (2007).


how the network of inductive and repressive pathways is integrated before, during and after gastrulation stages to establish an appropriate number of cardiac progenitors.

It is unclear at present if myocardial fate assignment is simply a matter of migration to the proper region in the ALPM. The path traveled by mesendodermal progeni-tor cells during gastrulation may be important to ensure their exposure to the appropriate mixture of inductive and repressive signals over a suitable length of time. The com-plementary organization of the myocardial fate map in the blastula and the myocardial fate map in the ALPM may reflect organized patterns of cell movements during gas-trulation. Several signals are known to be required for effi-cient gastrulation (Rohde and Heisenberg, 2007). However, the regulation of migration patterns of specific progenitor populations during gastrulation is not yet understood. In the grinch mutant, myocardial progenitors are lost, as evi-denced by an absence of nkx2.5 expression (Scott et al., 2007) (Fig. 3E–H). The grinch locus encodes the G protein-coupled receptor (GPCR) Agtrl1b, a relative of the chem-okine GPCRs. Transplantation experiments demonstrate that Agtrl1b signaling is required autonomously in myo-cardial progenitors during the initial phases of gastrulation (Scott et al., 2007). MO knockdown of the Agtrl1b ligand Apelin inhibits directed cell migrations during gastrula-tion (Zeng et al., 2007). Agtrl1b signaling may therefore represent a key pathway that regulates migration of cardiac progenitors to the correct position in the ALPM for the initiation of cardiogenesis.

III.c. heart tube assembly

By 16 hpf (14-somite stage), myocardial differentiation is underway within the bilateral heart fields of the zebrafish ALPM. The differentiating cardiomyocytes, while not yet contractile, begin expressing genes encoding sarcom-ere components, including cardiac myosin light chain 2 (cmlc2) in all cardiomyocytes, vmhc in ventricular car-diomyocytes, and amhc in atrial cardiomyocytes (Yelon et al., 1999; Berdugo et al., 2003). Between 16 and 20 hpf, these bilateral sheets of cells, sandwiched between the extraembryonic yolk syncytial layer (YSL) on their ven-tral side and the anterior endoderm on their dorsal side, migrate toward the embryonic midline, where they will begin to assemble the heart tube (Fig. 1C,D). Time-lapse studies utilizing a myocardial reporter transgene have revealed that cardiomyocytes migrate medially in an orderly and predictable manner (Holtzmann et al., 2007) (Fig. 4A–C). The coherent pattern of cell behavior is con-sistent with the finding that the migrating cardiomyocytes begin to form a polarized epithelium as they approach the embryonic midline (Trinh and Stainier, 2004) (Fig. 4D,E). As the bilateral cardiomyocyte populations approach each other, they create a ring of cells referred to as the cardiac

cone, through a process termed cardiac fusion (Fig. 1C). Cardiac fusion begins with contacts between posterior sub-sets of the contralateral cardiomyocyte populations; then, connections between anterior subsets of contralateral cells complete formation of the cone (Fig. 4A–C). Time-lapse analysis has indicated that particular angular movements of the most anterior and posterior cardiomyocytes create the specific circumference of the myocardial cone, engulf-ing the centrally located endocardial precursors (Bussmann et al., 2007; Holtzman et al., 2007).

At 20 hpf (22-somite stage), the cardiac cone is a lumenized structure projecting in a dorsoventral direction (Glickman and Yelon, 2002). Atrial cardiomyocytes reside in its broad base, and ventricular cardiomyocytes are found more dorsally and medially in the apex of the cone (Fig. 1C). The cone then tilts posteriorly, placing the ventricle posterior to the atrium. In this orientation, the cone gradu-ally elongates, generating a primitive myocardial tube

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fIGurE 4 Medial migration of cardiomyocytes is an orderly, coher-ent process. (A–C) Images from a time-lapse study of cardiac fusion in a wild-type embryo expressing the myocardial reporter transgene Tg(cmlc2:egfp). Dorsal views, anterior to the top, at the (A) 16-somite; (B) 18-somite; and (C) 20-somite stages. Cardiac fusion begins with con-tacts between posterior cardiomyocytes (B) and concludes with contacts between anterior cardiomyocytes (C). Images adapted from Holtzman et al. (2007). (D–G) The migrating myocardium is a polarized epithelium, and its integrity requires Fibronectin function. (D, F) Transverse sections, dorsal to the top, of wt and natter (fibronectin) mutant embryos at the 20-somite (19 hpf) stage. (E, G) Magnified views of the right side of each section. Expression of Tg(cmlc2:egfp) (pseudocolored blue) indicates bilateral locations of cardiomyocytes, and immunohistochemistry for -catenin (red) and PRKCi (green) indicates protein localization. (D, E) The myocardium exhibits apicobasal polarity, with -catenin local-ized basolaterally and PRKCi localized apicolaterally. (F, G) Loss of Fibronectin disrupts myocardial polarity and causes cardia bifida. Images adapted from Trinh and Stainier (2004).


(Fig. 1D). At the same time, the endocardial precursors migrate from their central position to generate an endothe-lial lining for the tube (Bussmann et al., 2007). The geom-etry of the developing heart is tightly confined by its position above the yolk and beneath the head. With resorp-tion of the yolk, the atrium, attached to the venous inflow, shifts to a more posterior position, leaving the heart in its familiar anteroposterior orientation, with the ventricle positioned more anteriorly (Stainier and Fishman, 1992).

Genetic analysis has revealed that the endoderm, YSL and endocardium all contribute to regulation of the dynamic patterns of cell behavior that drive cardiomyo-cyte migration and cardiac fusion. Inhibition of cardio-myocyte migration results in the formation of two separate hearts in lateral positions, a phenotype known as cardia bifida (Glickman and Yelon, 2002). Remarkably, bifid hearts can each contain a ventricle and an atrium (Yelon et al., 1999); cardiac fusion therefore appears dispensable for myocardial differentiation, even though it is essential for proper alignment with the vascular system. Mutations in casanova (sox32), bonnie and clyde, faust (gata5) and one-eyed pinhead all disrupt endoderm specification and cause cardia bifida, suggesting that the endoderm provides an important signal or substrate utilized by migrating car-diomyocytes (Peyrieras et al., 1998; Reiter et al., 1999; Kikuchi et al., 2000, 2001). The molecular underpinnings of endodermal-myocardial interactions remain mysteri-ous, although these may somehow involve sphingolipid sig-naling, since transplantation experiments indicate that the sphingosine-1-phosphate receptor gene miles apart (edg5) plays a cell nonautonomous role in promoting myocardial migration (Kupperman et al., 2000). In addition to putative interactions with the endoderm, migrating cardiomyocytes require interactions with components of the extracellular matrix, particularly Fibronectin, which is deposited by the YSL. This requirement is demonstrated by the cardia bifida phenotype of natter (fibronectin) mutants (Trinh and Stainier, 2004). Mutation of natter (fibronectin) also dis-rupts apicobasal polarity in cardiomyocytes (Fig. 4F,G), suggesting that formation of a polarized epithelium is an important prerequisite for the coordination of myocardial migration (Trinh and Stainier, 2004; Trinh et al., 2005).

Once interactions with the endoderm and YSL have recruited cardiomyocytes toward the embryonic midline, interactions between the myocardium and the endocardium regulate the cell behaviors that create the specific shape of the cardiac cone (Holtzman et al., 2007). Mutation of cloche, which eliminates the endocardium, alters the pat-tern of cardiomyocyte movements during cardiac fusion. Although cloche mutant embryos do not display cardia bifida, the cloche mutant cardiac cone has a significantly distorted morphology. Although the molecular nature of the relevant myocardial-endocardial interactions is not yet clear, it seems that communication between myocardium and endocardium controls the induction, direction and

duration of angular cardiomyocyte movements during the final phase of cardiac fusion. Thus, the endocardium helps to organize the myocardium into a configuration appropri-ate for the assembly of the heart tube.

Much less is known about the mechanisms that drive the tilting and elongation of the cardiac cone to create the heart tube. One emerging theme is that the regulation of apicobasal polarity is critical during heart tube elongation, just as it is during cardiomyocyte migration. Mutations in the genes heart and soul (prcki), snakehead (atp1a1a. 1) and nagie oko (mpp5) inhibit heart tube elongation; mutant hearts appear as arrested cardiac cones or stunted heart tubes (Yelon et al., 1999; Horne-Badovinac et al., 2001; Peterson et al., 2001; Shu et al., 2003; Yuan and Joseph, 2004; Rohr et al., 2006). PRKCi and Mpp5 are compo-nents of apically-localized protein complexes, and analy-sis of prkci and mpp5 mutants demonstrates that both genes are required cell-autonomously for normal apico-basal polarity of the myocardium and heart tube elonga-tion (Rohr et al., 2006) (Fig. 5). The function of Atp1a1a.1 also relates to the maintenance of myocardial apicobasal polarity, since its activity as an ion pump is required for the maintenance of junctional belts that connect the myo-cardial epithelium (Cibrian-Uhalte et al., 2007). Together, these data point to the importance of organization and coherence of the myocardium during the transformation of the cardiac cone into the heart tube.

III.d. morphogenesis of the cardiac chambers and atrioventricular cushions

Between 24 and 48 hpf, the simple heart tube gradually transforms into a two-chambered organ, with an atrium and a ventricle separated by the constriction of the atrio-ventricular canal (Fig. 1D,E). The zebrafish heart tube is initially positioned with its ventricular end pointing toward the right side of the embryo and its atrial end pointing toward the left side of the embryo (Fig. 1D). Left–right asymmetry of the heart is maintained as the tube loops to create an S-shaped structure (Fig. 1E). By 36 hpf the ven-tricle is clearly displaced to the right of the atrium and an obvious pinching appears at the atrioventricular (AV) canal. While cardiac looping is underway, chamber curva-tures emerge through a process called ballooning, in which localized bulges deform the cylindrical wall of the heart tube (see Chapter 3.2, Vol. I). By 48 hpf each expanded chamber exhibits two characteristic curvatures: a bulging curvature called the outer curvature (OC); and a recessed curvature called the inner curvature (IC).

Recent studies have started to uncover the cellular mechanisms responsible for zebrafish chamber morphogen-esis. In particular, analysis of cardiomyocyte morphology and organization in the zebrafish heart has demonstrated that regional changes in cell size and shape are associated


with curvature formation in the ventricle (Auman et al., 2007). Within the linear heart tube, all ventricular cardio-myocytes are similarly small and round (Fig. 6A). As cham-ber curvatures emerge, both OC and IC cells increase their surface area (Auman et al., 2007). Notably, however, only OC cells significantly change their shape, becoming flat-tened and elongated (Fig. 6B), whereas IC cells maintain a cuboidal morphology (Fig. 6C). Thus, regionally-confined cell shape changes underlie the acquisition of chamber morphology; the elongation and orientation of OC cells, coupled with the cuboidal shape of IC cells, create the characteristic curvatures of the expanded ventricle.

The dynamic cellular remodeling of the heart tube is essential for creating chambers of appropriate functional

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fIGurE 5 Heart tube elongation requires normal apicobasal polarity in the myocardium. Dorsal views, anterior to the top, at the (A, C, E) 20-somite and (B, D, F) 28-somite (23 hpf) stages, comparing myocardial morphology of (A, B) wt embryos with that of embryos injected with (C, D) anti-prkci MO or (E, F) anti-mpp5 MO. Expression of Tg(cmlc2:egfp) (green) indicates arrangement of cardiomyocytes, and immunohistochem-istry for PRKCi (red) and ZO-1 (blue) facilitates visualization of the mid-line. In wt embryos, tilting and elongation of the cardiac cone (A) creates a primitive heart tube (B) by the 28-somite stage. (C, D) Loss of PRKCi blocks both tilting and elongation, and (E, F) loss of Mpp5 delays cardiac fusion and blocks further steps of heart tube assembly. Images adapted from Rohr et al. (2006).

capacity, yet this entire process must occur while the heart is beating. Interestingly, studies of zebrafish mutants with functional deficiencies have demonstrated that cardiac function has a potent influence on chamber curvature for-mation. For example, mutation of the weak atrium (amhc) locus, which encodes an atrium-specific myosin heavy chain, disrupts atrial contractility and, consequently, hin-ders blood flow (Berdougo et al., 2003). Additionally, weak atrium (amhc) mutants have significant ventricular defects; their ventricular cardiomyocytes fail to enlarge or elongate normally (Fig. 6D,E), resulting in an abnormally small and round ventricle (Auman et al., 2007). Since amhc is not expressed in the ventricle, this ventricular

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fIGurE 6 Cardiac function influences morphogenesis of the cardiac chambers and atrioventricular cushions. (A–C) Regionally-confined cell shape changes underlie the emergence of chamber curvatures in the ven-tricle. Live wt hearts expressing Tg(cmlc2:egfp) and exhibiting mosaic expression of Tg(cmlc2:dsredt4). Arrows point to representative cells expressing both dsredt4 and egfp. Ventricular cells in the linear heart tube (LHT) at 28 hpf (A) and in the IC at 52 hpf (C) are relatively cuboi-dal, whereas cells in the OC at 52 hpf (B) are flattened and elongated. Images adapted from Auman et al. (2007). (D–F) Cardiomyocyte cell shape changes are regulated by cardiac function. Live hearts expressing Tg(cmlc2:egfp) and exhibiting mosaic expression of Tg(cmlc2:dsredt4) at 52 hpf. Arrows point to representative ventricular OC cells express-ing both dsredt4 and egfp. In contrast to the enlargement and elongation of OC cells in wt embryos (D), when blood flow is reduced, as in amhc mutant embryos (E), ventricular cardiomyocytes, including the OC cells, remain small and fail to elongate normally. (F) In vmhc mutant embryos, ventricular cardiomyocytes, including the OC cells, are excessively enlarged and elongated. Images adapted from Auman et al. (2007). (G–I) AV cushion formation is regulated by cardiac function. Lateral views of live embryos, anterior to the left, expressing the endothelial reporter transgene Tg(tie2:gfp) at 48 hpf. Red arrows point to the AV canal. (G) In wt embryos, enhanced expression of Tg(tie2:gfp) is visible in the AV endocardial cushions. In silent heart (sih; tnnt2) mutant (H) and cardio-funk (cfk; actc1) mutant embryos, cardiac contractility is disrupted and AV cushions do not form. Images adapted from Bartman et al. (2004).


phenotype is presumed to reflect a potent secondary effect of diminished blood flow on the cell shape changes that normally create chamber curvatures.

The precise mechanism by which blood flow promotes myocardial cell shape change remains unclear. One pos-sibility is that the shear forces produced by blood flow result in mechanosensation by the endocardium which then transmits another set of signals to the myocardium. Several zebrafish mutant phenotypes provide evidence for the importance of endocardial–myocardial signaling dur-ing chamber formation. The cloche mutation, which blocks endocardium formation, also causes dysmorphic, dilated chambers (Stainier et al., 1995; Liao et al., 1997). The heart of glass, santa (krit1) and valentine (ccm2) genes are also critical for regulating chamber morphology; muta-tion of any of these genes causes severe chamber dilation (Mably et al., 2003, 2006). Since all three of these genes are expressed in endothelial cells, these mutant phenotypes again implicate endocardial–myocardial signaling in the control of chamber shape.

Whereas blood flow encourages cells to enlarge and elongate, the inherent contractility of a cardiomyocyte seems to limit its degree of shape change. Mutation of the pickwick locus, encoding the giant scaffold protein Titin and mutation of the half-hearted locus, encoding the ventricle-specific myosin heavy chain Vmhc, both result in distortion of cardiomyocyte morphology (Xu et al., 2002; Auman et al., 2007) (Fig. 6F). Transplant experi-ments have demonstrated that these proteins are required cell-autonomously for maintenance of cardiomyocyte cell shape, indicating that the integrity of the contractile appa-ratus plays an important role in chamber morphogenesis. Altogether, the phenotypes of zebrafish mutants with both chamber shape defects and functional defects suggest that the acquisition of normal cardiomyocyte morphology requires a balance between external physical forces, such as blood flow, and internal physical forces, such as con-tractility. Future studies are likely to focus on the mech-anisms for the mechanotransduction of hemodynamic forces and on the cytoskeletal attributes that are linked to sarcomere integrity; potential interplay between the rel-evant pathways may provide the basis for establishing and maintaining cardiomyocyte cell shape during chamber cur-vature formation.

At the same time as chamber curvatures are emerging, AV valve morphogenesis is underway (Beis et al., 2005) (see Chapters 6.1 and 6.2). Valve formation initiates within the AV canal, a constriction of the heart tube found at the boundary between the atrium and the ventricle. Gene expression pat-terns also distinguish the developing AV canal; expression of bmp4 and versican becomes restricted to the AV myocardium and AV endocardial cells exhibit upregulation of endothelial reporter transgenes (Fig. 6G), restricted expression of notch1b and characteristic lateral localization of the adhe-sion molecule Dm-grasp (Walsh and Stainier, 2001; Beis

et al., 2005). Additionally, myocardial and endocardial cells in the AV canal begin to exhibit cell morphologies distinct from those in the flanking chambers. Notably, the AV endo-cardial cells transition from a squamous to a more cuboidal appearance and aggregate to establish endocardial cushions (Beis et al., 2005). Subsequent remodeling events convert these cushions into the leaflets of the AV valve.

A variety of zebrafish mutations provide insight into the mechanisms responsible for patterning the AV canal and executing AV valve morphogenesis. For example, jekyll (udp-glucose dehydrogenase) mutants have defects in AV canal patterning; they fail to restrict expression of bmp4, versican and notch1b, and do not form endocardial cushions (Walsh and Stainier, 2001). The enzymatic activ-ity of UDP-glucose dehydrogenase (Ugdh) is required for the modification of extracellular matrix proteins that facili-tate Wnt and Fgf signaling (Lander and Selleck, 2000), suggesting a mechanism by which Ugdh might regulate the signal transduction that is responsible for AV pattern-ing. In particular, the jekyll mutant phenotype might reflect a defect in Wnt signaling; activation of Wnt signaling via mutation of the tumor suppressor gene apc leads to exces-sive endocardial cushion formation beyond the boundaries of the AV canal, and inhibition of Wnt signaling blocks endocardial cushion formation (Hurlstone et al., 2003). In addition to requiring proper AV canal patterning, AV valve morphogenesis, like chamber morphogenesis, requires input generated by biomechanical forces. Mutation of genes encoding contractile proteins, such as the cardiac troponin T gene silent heart (tnnt2) and the cardiac actin gene cardiofunk (actc1), blocks endocardial cushion for-mation, suggesting that either shear forces, resulting from blood flow, or a stretch response, resulting from contrac-tility, trigger valve morphogenesis (Bartman et al., 2004; Beis et al., 2005) (Fig. 6G–I). The role of shear forces in promoting valve development has also been demonstrated by experiments implanting large beads into embryos; when the implanted bead obstructs blood flow, endocardial cushions do not form (Hove et al., 2003). However, not all aspects of AV canal development are dependent on cardiac function. Restriction of bmp4 expression to the AV canal proceeds normally, even when contractility is defective (Bartman et al., 2004). Also, formation of specialized slow conductive tissue in the AV myocardium, which is induced by endocardial-myocardial signaling, does not require car-diac contractility (Milan et al., 2006).

Together, studies of chamber and valve formation in zebrafish highlight the important interrelationship of form and function, chamber and valve morphogenesis are both required for, and dependent on, normal cardiac function. This suggests numerous etiologies for congenital heart defects, since small alterations in cardiac function could easily influence cardiac morphology, and vice versa. The 24 hpf heart tube has been described as a peristaltic pump, allowing blood flow via unidirectional propagation of


contractions from the atrial pole to the ventricular pole. However, recent examination of contractions and blood flow using high-speed confocal microscopy has suggested that the early heart tube acts instead as a suction pump (Forouhar et al., 2006). In this case, the precise engineer-ing required to allow the heart tube to pump blood prop-erly implies that even mild perturbations in morphogenesis will have dramatic effects on cardiac performance, lead-ing to potentially devastating consequences for cardiac development.

Iv. usE of zEbrafIsh as modEls of hEart dIsEasE

Despite the great progress made in identifying mutations associated with heart disease, heart dysfunction remains the leading cause of death in the Western world. Congenital heart disease is found in 1–2% of live births, with complex fetal or postnatal surgery in some cases being required for survival (Ransom and Srivastava, 2007). Additionally, cardiomyopathies and arrhythmias in adults often have an underlying genetic origin (Keating and Sanguinetti, 2001; Towbin and Bowles, 2002). Identification of mutations that cause or increase the likelihood of heart disease provides a means to pinpoint individuals at risk via genetic analysis. This offers an opportunity for the initiation of measures to prevent or mitigate the severity of disease. However, curative approaches are not currently available. It is diffi-cult at present to envisage gene therapy as an alternative to repair genetic defects in all cells of the heart. Therefore, a greater mechanistic understanding of how heart disease arises from genetic mutations is the current priority. This approach may indicate pathways to target for pharmaco-logical or dietary intervention.

Congenital heart disease primarily involves morpho-genetic defects. These include incomplete septation of the chambers, defects in valve development, hypoplastic chambers and anomalies in outflow tract septation/pattern-ing (Gruber and Epstein, 2004). The zebrafish does not require pulmonary circulation, and therefore the zebrafish heart is much simpler in design than the mammalian heart. In zebrafish, the single ventricle and atrium lack septa, and the outflow tract consists of a simple bulbus arterio-sus. However, the zebrafish bulbus is invested with smooth muscle (Grimes et al., 2006), as is observed in the mam-malian arterial pole. Additionally, valve formation in zebrafish appears to occur via a similar mechanism as in mammals (Chang et al., 2004; Beis et al., 2005), and the zebrafish ventricular myocardium forms a compact layer and trabeculae similar to those seen in mammals (Hu et al., 2000, 2001).

Despite the relative simplicity of the zebrafish heart, the genes that regulate heart development appear con-served. There are multiple examples in which mutations

of genes associated with human congenital heart disease cause cardiac developmental defects in zebrafish. The zebrafish heartstrings mutation disrupts tbx5, a gene asso-ciated with Holt-Oram syndrome (Basson et al., 1997; Li et al., 1997; Garrity et al., 2002). Similarly, mutation or MO knockdown of congenital heart disease genes such as tbx1, gata4, fog1 and raf1 cause early developmental heart defects in zebrafish (Piotrowski et al., 2003; Walton et al., 2006; Peterkin et al., 2007; Razzaque et al., 2007). It there-fore seems likely that the same molecular pathways enact common morphogenetic mechanisms in both the human and zebrafish heart. Since zebrafish cardiac morphogen-esis can be observed in real time, this model presents an excellent tool to study how congenital heart disease genes regulate heart development and to test the roles of candidate disease genes.

Forward genetic screens and the use of MOs in zebrafish have identified a number of phenotypes featuring physi-ological defects in which cardiac contractility or rhythm is compromised (Rottbauer et al., 2001; Sehnert et al., 2002; Xu et al., 2002; Berdougo et al., 2003; Langheinrich et al., 2003; Bartman et al., 2004; Ebert et al., 2005; Langenbacher et al., 2005; Rottbauer et al., 2006; Arnaout et al., 2007). Interestingly, many of these mutations affect human familial cardiomyopathy or arrhythmia genes, such as those encoding Titin or Kcnh2 (Xu et al., 2002; Langheinrich et al., 2003; Arnaout et al., 2007). While car-diomyopathies and arrhythmias are not strictly defined as developmental disorders, mutation of disease-associated genes causes profound defects in the zebrafish embryo. Given the experimental tractability of the zebrafish, these mutations provide excellent models to study the signals that lead to heart failure in humans. From the viewpoint of con-sidering mechanisms for cardiac repair, it is also important to note that the zebrafish heart has tremendous regenera-tive capacity (Poss et al., 2002). While the precise cellular and molecular basis for regeneration is not yet known, it appears that novel cardiomyocytes are generated to repair the injured zebrafish heart (Lepilina et al., 2006). A greater understanding of how this process is regulated in zebrafish may have significant implications for the repair of damage caused by heart failure in humans.

Aside from providing a means to study the mechanisms leading to disease, animal models are also useful for the evaluation of possible therapeutic interventions. The great success of a folic acid-rich diet in preventing the complex genetic spectrum of neural tube closure defects suggests that other congenital anomalies may also benefit from chemical intervention. Chemical genetics, in which large libraries of compounds are screened for specific biologi-cal effects, is one promising approach toward identifying useful small molecules. The zebrafish embryo provides an excellent in vivo model for this technique (Murphey and Zon, 2006). Successful chemical genetic screens in zebrafish have identified compounds that regulate


processes including pigmentation, cell-cycle progression, angiogenesis and heart rate (Milan et al., 2003; Peterson et al., 2004; Murphey et al., 2006; Ni-Komatsu and Orlow, 2007). In an example utilizing an automated method to measure heart rate, drugs known to cause long QT syn-drome in humans almost uniformly produced bradycardia and AV block in zebrafish embryos (Milan et al., 2003). Therefore, as with genetic pathways, drug effects may be conserved from zebrafish to human. Chemical sup-pression of a mutant cardiovascular phenotype has been demonstrated in the case of the zebrafish gridlock mutant (Peterson et al., 2004). Mutation of gridlock (hey2) causes aortic coarctation, resulting in the absence of blood flow to the trunk. Screening of several thousand chemicals revealed two modulators of VEGF signaling that restored proper blood flow to gridlock mutants. While it is tempt-ing to speculate that compounds able to rescue zebrafish mutant phenotypes could be clinically applicable, this may be a naïve hope. Zebrafish null mutants are not equiva-lent to patients heterozygous for a particular mutation. Furthermore, many compounds are likely to have undesir-able off-target effects. Even so, future pursuit of this strat-egy will provide molecular inroads to elaborate promising pathways with curative potential, further strengthening the utility of the zebrafish as a model of heart disease.

v. conclusIons

The past decade has been a highly productive period for the study of zebrafish heart development. By combining multiple genetic and embryological experimental appro-aches, investigators have taken considerable strides toward elucidating the mechanisms that drive heart patterning and morphogenesis, including and in addition to the work encapsulated here. Notably, the ability to study develop-ment at the cellular level in the zebrafish embryo has been especially useful for linking essential genes to the specific cell behaviors that they regulate. As the number of investi-gators using zebrafish increases and the sophistication of zebrafish technology advances, the pace of discovery will undoubtedly continue to accelerate. In particular, emerging methods for targeted gene inactivation, enhanced strategies for spatial and temporal manipulation of gene function, and new techniques for automated image analysis are likely to facilitate high-throughput analysis of phenotypes generated by forward, reverse and chemical genetics.

Importantly, future work is also likely to provide a clearer resolution of the parallels and contrasts between cardiogenic mechanisms in zebrafish and other vertebrates. It is certain that some aspects of heart development have changed over the course of vertebrate evolution; there are clear differences in the structure of fish and mammalian genomes and in the complexity of fish and mammalian hearts. It will be particularly exciting to elucidate models

of how the complex terrestrial heart evolved from the simpler aquatic organ. Studies in this regard are likely to focus on how the multiple heart fields found in amniotes (Buckingham et al., 2005) relate to the zebrafish heart fields; it is not yet clear whether zebrafish possess a second heart field equivalent (see Chapter 2.2). It will also be fas-cinating to examine whether advanced morphogenetic processes such as chamber septation are regulated by mech-anisms similar to those driving zebrafish heart tube assem-bly and chamber formation. Overall, it is probable that the commonalities between vertebrate hearts will ultimately outweigh their differences, at least in terms of the funda-mental mechanisms regulating many cardiac cell behaviors that are highly relevant to the etiology of human disease.

acknowlEdGmEnts

We thank S. Abdelilah-Seyfried, T. Bartman, D. Stainier, S. Rohr, L. Trinh and J. Waxman for providing images for the figures. We regret having to limit or omit discussion of many interesting studies due to space constraints. Work in the Scott laboratory is supported by funding from the Canadian Institutes of Health Research, the Natural Sciences and Engineering Research Council of Canada, and the Canada Foundation for Innovation; work in the Yelon laboratory is supported by funding from the National Institutes of Health, the American Heart Association and the March of Dimes.

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103 Cardiac Development in the Zebrafish