Cardiac Regeneration in the Zebrafish Model System

Chapter 12.2

Cardiac Regeneration in the Zebrafish Model System Kenneth Poss Department of Cell Biology, Howard Hughes Medical Institute, Duke University Medical Center, Durham, NC, USA

I.  Introduction Acute myocardial infarction, typically caused by coronary artery occlusion and ischemia, is a leading cause of death worldwide. For those fortunate enough to survive myocardial infarction, necrotic myocardium is replaced in the next few weeks by collagen-rich scar tissue. The scar provides a rapid solution; however, it is not contractile. Indeed, cardiac scarring, thought to be irreversible, weakens the heart while also increasing susceptibility to compensatory pathology, additional infarction events and organ failure. Many scientists and clinicians believe that myocardial infarction patients represent the greatest potential beneficiaries from regenerative medicine, which aims to restore functional tissue through stem or progenitor cell manipulation. Indeed, the development of therapies that can facilitate survival or regenerative replacement of destroyed myocardium would be of enormous social and economic impact. One promising approach to effecting cardiac regeneration in humans is to establish means to purify and transplant progenitor cells. This new and exciting pursuit is the subject of other chapters within this textbook (see Chapters 13.2, 14.1 and 14.2). A complementary method is to identify successful examples of cardiac regeneration among nonmammalian vertebrates, dissect how this success is achieved, and then attempt to apply this information to mammals via provision of the appropriate regenerative stimuli. Indeed, the general capacity for organ regeneration is remarkably elevated in certain lower vertebrates such as urodele amphibians and teleost fish (see Chapter 12.1). This disparity in regenerative capacity is perhaps most prominent in the heart, where little or no regeneration occurs in mammals. By contrast, a relatively new laboratory model system, the Heart Development and Regeneration Copyright © 2010 Elsevier Inc. All rights of reproduction in any form reserved.

zebrafish, displays a surprising degree of regeneration after cardiac injury. Already, we have learned information from studies of zebrafish heart regeneration that brings a new perspective on human heart disease and potential regenerative therapies. This chapter will review what is known about mechanisms of heart regeneration in zebrafish. In addition to addressing “how” regeneration occurs, it will touch on questions of “why”. That is, why is the capacity for natural heart regeneration limited to nonmammalian species?

II.  Regeneration Stem and progenitor cells have been identified from most mammalian organs, including brain, skeletal muscle, liver, lung, skin, kidney, blood, bone, reproductive tissues, intestine and heart. These cells ostensibly function to maintain homeostasis, the replacement of cells damaged and lost during day-to-day performance of organ function. Some progenitor cells also have the potential to renew tissue after minor injury, or even after major damage or removal of structures. Robust examples of mammalian organ regeneration are demonstrated by blood and liver. Transplanted hematopoietic stem cells have a tremendous capacity to repopulate the hematopoietic system of an irradiated mouse, and the liver is able to renew its mass within two weeks after removal of two-thirds of all hepatic tissue (Wagers et al., 2002; Taub, 2004; Shizuru et al., 2005). On the other hand, not all mammalian organs are equally capable of regeneration. The heart and central nervous system are particularly resistant to regeneration after injury, despite the presence of one or more progenitor cell types. In the brain, the inability to regenerate is thought to contribute to morbidity caused by neurodegenerative disease 839

840

or stroke. The regenerative deficiency of the mammalian heart is believed to have a huge impact on human disease, as it provides little defense against myocardial infarction. For hundreds of years, biologists have been fascinated and inspired by the spectacular regenerative abilities of many nonmammalian species (see also Chapter 12.1). The invertebrate hydra and planarian models have a seemingly-unlimited capacity for regeneration. These species are capable of regenerating a full head and tail from an incredibly small piece of tissue, and hydra can even form from cells coaxed together after dissociation of an intact animal (Fujisawa, 2003; Reddien and Sanchez Alvarado, 2004). Indeed, their mere propagation requires major regenerative capacity, since asexual reproduction in these animals occurs by budding or fission. Lizards, frogs and fish have served as vertebrate models for spectacular regeneration; yet salamander research has traditionally dominated the field. The work of Spallanzani reported in 1768 was the first to illuminate the ability of salamanders to regenerate an entire amputated limb (Spallanzani, 1768). Since then, newts or axolotls have been found to regenerate tail, spinal cord, retina, lens, jaws, portions of intestine, and brain tissue (Tsonis, 1996). In particular, the regenerating limb has been a subject of fascination for regeneration researchers. Following amputation of the newt limb, a thin layer of epithelial cells called the wound epidermis quickly migrates to cover the stump. In the ensuing weeks, interactions between the wound epidermis and the underlying stump mesenchyme stimulates formation of the regeneration blastema, a mass of undifferentiated, proliferative progenitor cells. The blastema, a distinguishing feature of many additional examples of regeneration, ultimately gives rise through patterning and differentiation events to the new cartilage, bone and muscle of the restored limb. Growth factors such as fibroblast growth factors (Fgfs) or bone morphogenetic proteins released from the wound epidermis, and/or from the nerves that penetrate the newly-formed blastema, influence blastemal formation and function (Brockes and Kumar, 2005; Kumar et al., 2007). A major outstanding issue in developmental biology is just how an amputated structure is capable of such effective mobilization and organization of progenitor cells. There are multiple lines of evidence that demonstrate the ability of differentiated newt cells to reduce their differentiation status, or dedifferentiate, to create a more plastic or progenitor cell state. For instance, when multinucleate skeletal myotubes were labeled with a membraneimpermeable dye and injected into a limb blastema, they were observed to contribute mononuclear cell progeny, including chondrocytes, to the restored structure (Lo et al., 1993). In order for this to occur, myotubes ostensibly downregulate contactile genes like myosin heavy chain and undergo fragmentation, phenomena that have been observed in newt myotubes cultured in vitro (Kumar et al., 2004). For many years, dedifferentiation has been trumpeted as a

PART  |  12  The Regenerative Heart

seminal aspect of appendage regeneration. Still, it remains unclear to what extent dedifferentiation, as opposed to employment of a separate reserve of stem or progenitor cells, contributes to blastema formation. Notably, a different study showed evidence that cells which are positive for the skeletal muscle satellite marker Pax7 donate cells to the newt blastema and restored limb (Morrison et al., 2006). In planarians, a reserve population of stem cells called neoblasts is recruited to the injury site to create blastemal tissue (Reddien and Sanchez Alvarado, 2004). While there are interesting mechanistic differences displayed by nature’s examples of regeneration, there is typically a shared formula for regenerative success; the effective activation and cultivation of progenitor cells.

III.  Capacity for heart regeneration III.A.  The Mammalian Heart Slight improvements in the survival or regeneration of human cardiac muscle after ischemia are predicted to have major impacts on quality of life. Even with a relaxed definition of regeneration, involving any replacement of lost myocardium through the production of new cardiomyocytes, significant natural cardiac regeneration of any form has not been confirmed in mammals. These observations represent multiple models of injury, including ischemic infarction, burning, freezing, mechanical injury, chemical injury, etc. (Rumyantsev, 1977). The liver regenerates by activating proliferation of its major structural cells, hepatocytes, after injury. By contrast, it is generally-accepted that mature mammalian cardiomyocytes have little or no capacity for cell division (Pasumarthi and Field, 2002). Studies of rodent cardiac growth have suggested that cardiomyocyte hypertrophy, an increase in cell size, is responsible for cardiac growth after the first few days of life, as opposed to cardiomyocyte hyperplasia, an increase in cell number (Li et al., 1996). Further supporting the idea that in vivo division of mature mammalian cardiomyocytes is rare is their inability to survive for long periods or to divide in cell culture. Interestingly, recent studies have indicated that cultured adult cardiomyocytes can be stimulated to divide at low frequencies in vitro by adding fibroblast growth factors (Fgfs) and/or inhibiting p38 MAP kinase (Engel et al., 2005). It seems logical that large, contractile cardiomyocytes are not highly-proliferative; yet existing cardiomyocytes need not divide in order for regeneration potentially to take place. Notably, several groups have identified multiple types of undifferentiated or poorly-differentiated cardiac progenitor cells in the postnatal heart that have the potential to mature into contractile cells. At least three rare cell types with the potential to create cardiomyocytes postnatally have been described, expressing either the pan-stem cell marker c-kit (Beltrami et al., 2003), the transcription factor islet1

Chapter  |  12.2  Cardiac Regeneration in the Zebrafish Model System

(Laugwitz et al., 2005), or the cell surface marker stem cell antigen 1 (Laugwitz et al., 2005). Given findings over the years that most mammalian organs are equipped with one or more progenitor cell types, these discoveries have been exciting, if not surprising. The mystery, however, is that the injured mammalian heart fails to harness the potential of progenitor cells to create new cardiomyocytes after injury.

III.B.  The Amphibian Heart The possibility of natural cardiac regeneration in amphibians has been examined in frogs, newts and axolotls, with much of the original literature reporting work of Soviet scientists in the 1960s (Rumyantsev, 1977). This body of work indicated that amphibians survive massive mechanical injury to the ventricle, including removal of as much as one-quarter of the chamber. This resilience is a feat in itself, and is likely to reflect a lesser reliance on vigorous circulation than mammalian species. While there is some variability in reports and reviews on the extent of regeneration after injury, the prevailing view is that some cardiac regeneration occurs after resection (Becker et al., 1974; Oberpriller and Oberpriller, 1974; Neff et al., 1996). There is definitive evidence for proliferative activity, including the presence of mitotic figures in cardiomyocyte nuclei as visualized by standard histology and transmission electron microscopy. Furthermore, a high frequency of 3H-thymidine incorporation, and more recently, incorporation of the nucleotide analog bromodeoxyuridine (BrdU), was reported in cardiomyocytes of the injured ventricle (Flink, 2002). Perhaps most strikingly, a recent study found that cardiomyocytes labeled with a dye and transplanted into the limb blastema lost expression of contractile genes, and appeared in some cases to convert to skeletal muscle tissue, a process that would be considered “transdifferentiation” (Laube et al., 2006). The idea that cardiomyocyte cytokinesis occurs in newts after injury was bolstered by studies of cardiomyocytes isolated from the adult ventricle and cultured in vitro, in which cardiomyocytes readily divided (Oberpriller et al., 1995). It is not clear which molecules stimulate cardiomyocyte division; however, newts are reported to have a heterogeneous population of cardiomyocytes, some of which may be better-suited to division (Bettencourt-Dias et al., 2003). Despite evidence for in vitro cytokinesis and in vivo cell-cycle entry and mitosis in the newt, the predominant response to resection injury is scarring (Oberpriller and Oberpriller, 1974). That is, while it appears that adult newt cardiomyocytes have the potential to regenerate and do so to some degree, there is only minor replacement of cardiac muscle after resection injury. This paradoxical result may have something to do with the injury model itself. For instance, cardiomyocytes within minced myocardium that has been grafted to an injured ventricle will show proliferation, and can form a contiguous, contractile mass (Bader and Oberpriller, 1979). Another idea is that growth of scar tissue in the injured newt ventricle

841

simply overwhelms that of new myocardium. This idea of competition between regeneration and scarring has recently re-emerged and been addressed through use of zebrafish (Poss et al., 2002b), as described later in this chapter.

III.C.  The Zebrafish Heart The zebrafish Danio rerio has become a popular model system for understanding the development of the vertebrate embryo. A large community of zebrafish embryologists has focused on early patterning and organogenesis, with particularly strong inroads achieved in gastrulation and cardiovascular, blood and brain development. The zebrafish provides embryologists with a transparent specimen that develops quickly outside the mother, and a model system that is highly-amenable to forward genetic approaches (Grunwald and Eisen, 2002). The recent popularity of the zebrafish model system has encouraged the exploration of its regenerative capacity. It has been known for many years that other teleost species can regenerate amputated fins. In fact, the great geneticist Thomas Hunt Morgan was a leader in the field of regeneration before his famous change in direction to questions of inheritance in Drosophila, publishing multiple manuscripts that investigated teleost fin regeneration (Morgan, 1906). Zebrafish have five fin types, two of which are paired, all comprised of several segmented, bony fin rays. Each ray contains two concave, facing hemirays that surround nerves, blood vessels and connective tissue. All seven fins can regenerate after as much as 95% of the tissue is amputated. The caudal fin is most often used in regeneration experiments, and completes regeneration in 10–14 days. The process of fin regeneration is epimorphic, involving the formation and proliferation of an undifferentiated and presumably multipotent blastema (Akimenko et al., 2003; Poss et al., 2003). The simple structure of the fin, its external accessibility and the fact that its amputation causes no significant lifestyle changes are key reasons for its success as a model organ. Many groups now study zebrafish fin regeneration, with the goal of exploiting the tools and techniques developed by embryologists to apply them to fundamental questions of regulation during regeneration. For instance, how does injury stimulate the formation of a blastema? How does an injury stump remember the appropriate structures to restore? Additional regenerating structures of zebrafish may provide particular relevance to disease. Adult teleost retinae also regenerate, producing new rods and cones after an acute injury or after complete ablation by phototoxicity (Vihtelic and Hyde, 2000). Retinal regeneration appears to involve a progenitor cell-based mechanism; here, stem cells in the inner nuclear layer give rise to new photoreceptor neurons after injury (Otteson and Hitchcock, 2003). Amazingly, spinal cord tissue can regenerate after a complete transection. In a process that takes about six weeks, ~80% of animals given a posterior injury achieve

842

PART  |  12  The Regenerative Heart

functional recovery (Becker et al., 1997). This phenomenon is based on axonal regeneration, the striking ability of central nervous system neurons to recover, traverse the lesion and re-establish functional connections. Thus, adult zebrafish, like newts and axolotls, are capable of regenerating many structures that mammals cannot. As discussed later, zebrafish also bring certain unique practical advantages to the study of regeneration. Because adult zebrafish effectively regenerate multiple structures, it is logical to be curious about their natural capacity for heart regeneration. Ischemic myocardial injury to the zebrafish ventricle would be highly-representative of human disease, and myocardial infarction models are routinely used in small mammals such as mouse and rat. However, the zebrafish ventricle is diminutive (~1 mm3), as is the coronary vascular network, making ischemic injury models of myocardial infarction a difficult task. Research groups have recently examined the effects of removing approximately 20% of the ventricle by surgical resection with iridectomy scissors (Fig. 1A) (Poss et al., 2002b; Raya et al., 2003). By mechanical removal of ventricular myocardium, these studies examined what would occur after the strongest possible stimulus for regeneration. Resection injury penetrates the ventricular lumen, releasing a large amount of blood. As in the amphibian heart, the injury clots quickly, and the organ sustains sufficient contractile force to continue to drive circulation. (A)

Over the next month, a remarkable series of events occurs in response to ventricular resection. First, the clot that seals the apex matures within several days into fibrin, a complex milieu containing serum factors and degenerated erythrocytes. In the infarcted mammalian ventricle, fibrin deposition attracts fibroblasts and inflammatory cells, and is a precursor to scarring. Surprisingly, fibrin is not replaced by scar tissue during cardiac repair in zebrafish; in fact, little or no collagen is retained by 1–2 months after injury (Poss et al., 2002b). Instead, the fibrin is supplanted by cardiac muscle, restoring a contiguous ventricular wall (Fig. 1B). While the perfect pyramidal shape of the uninjured ventricle is typically not renewed, and the restored muscle is predominantly of the compact type that comprises the ventricular wall, the process restores approximately the amount of muscle that was lost (Poss et al., 2002b).

IV.  Myocardial progenitor   cells and zebrafish heart regeneration IV.A.  New Cardiomyocytes are Born   during Heart Regeneration As described in other chapters of this text, cardiac hypertrophy is one method that the mammalian heart uses to

outflow tract valve

atrium ventricle

0 dpa

(B)

fibrin

9 dpa

14 dpa

fibrin

30 dpa

Figure 1  Cardiac regeneration in zebrafish. (A) Hematoxylin and eosin stain of the adult zebrafish heart before and after 20% ventricular resection. (B) Ventricular sections stained for Myosin heavy chain to identify cardiac muscle (brown) and aniline blue to identify fibrin (blue). Mature fibrin seals the wound by 7 dpa, and is gradually replaced by cardiac muscle. The wall is typically restored by 30 dpa. Figure adapted with permission from Poss et al. (2002b).

Chapter  |  12.2  Cardiac Regeneration in the Zebrafish Model System

increase cardiac mass, either in the exercised heart or in the damaged or diseased heart. By contrast, restoration of the wall of the zebrafish ventricle reflects hyperplasia, the creation of new cardiomyocytes. Labeling studies with BrdU, designed to detect cells that have entered S-phase, found that cardiomyocyte cell-cycle entry is stimulated by injury. In the uninjured ventricle, little proliferation is observed. However, by one week after injury, BrdU-labeled cardiomyocytes were seen adjacent to the wound (Poss et al., 2002b; Raya et al., 2003). The number of labeled cardiomyocytes peaked at 14 days post-amputation (dpa), and BrdU labeling was also observed at 30 dpa in an outer rim of peripheral cardiomyocytes in the restored ventricular wall (Fig. 2). Condensed chromatin and mitotic figures were observed at low frequencies in cardiomyocytes near the injury site. Thus, adult zebrafish cardiomyocytes, like newt cardiomyocytes, are capable of hyperplasia in vivo, and they are somehow stimulated by injury to proliferate.

843

IV.B.  Participation of Progenitor Cells What is the origin of proliferative cardiomyocytes during regeneration? Some details were revealed by BrdUlabeling studies. The authors found that proliferation events were enhanced at the apical edge of the regenerate throughout the regenerative process. Extensive pulse– chase studies, in which BrdU was “pulsed” for seven days during the peak proliferative period, but then “chased” for 16–46 days without label, were also helpful. These experiments pointed to a mechanism in which muscle is restored by new cardiomyocyte production at this gradually shifting proliferative edge, with earlier proliferating cardiomyocytes displaced inward (Poss et al., 2002b). This finding implied that heart regeneration may share some mechanisms with a process like fin regeneration, where a proliferative domain is maintained in a specific region of the blastema as regenerative growth proceeds distally (Nechiporuk and Keating, 2002; Poss et al., 2002a).

(A)

(B)

(C)

(D)

Figure 2  Cardiomyocyte cell-cycle entry during zebrafish heart regeneration. (A) Confocal image of a heart section of an unamputated fish labeled for 7 days with BrdU, stained for myosin heavy chain to identify cardiomyocytes (red), BrdU to detect cycling cells (green) and DAPI to detect nuclei (blue). (B, C) Cardiomyocyte BrdU incorporation (arrowheads) is activated at the amputation plane by 7 dpa (0–7 dpa BrdU labeling) (B) and 14 dpa (7–14 dpa BrdU labeling) (C). (D) 30 dpa (23–30 dpa BrdU labeling). Myocyte BrdU incorporation continues as the ventricular wall expands. Labeling is usually limited to the most epicardial myocytes of the restored wall (arrowheads). Figure adapted with permission from Poss et al. (2002b).

844

How cardiomyocyte proliferation is initiated and maintained at the apical edge of the regenerate is a critical issue. When considering possible mechanisms, it is important to keep in mind the recent identification of postnatal cardiac progenitor cells in mammalian cardiac tissue (see Chapters 13.1 and 14.3). Indeed, there are several likely mechanisms for the origin of new muscle during regeneration: (1) differentiated, contracting cardiomyocytes in existing myofibers could be stimulated to enter the cell-cycle, divide, and reform the apex; (2) regeneration could proceed through the recruitment of undifferentiated progenitor cells that form new, proliferative cardiomyocytes; or (3) dedifferentiation of cardiomyocytes, including downregulation of contractile genes like cardiac myosin light chain 2 (cmlc2) and ventricular myosin heavy chain, could produce more plastic cardiomyocytes, or even progenitor cells, that divide and redifferentiate. Certainly, there could be multiple mechanisms taking place simultaneously. These possibilities were addressed in a set of experiments designed to detect whether an undifferentiated intermediate structure like a blastema exists during heart regeneration (Lepilina et al., 2006). In that study, transgenic strains were used that expressed either enhanced green fluorescent protein (EGFP), or nuclear-­localized DsRed2, a red fluorescent protein, behind the zebrafish cmlc2 promoter (Mably et al., 2003; Burns et al., 2005). Cmlc2 is the earliest known marker for differentiated embryonic cardiomyocytes, and its expression is maintained in adult cardiac muscle (Yelon et al., 1999). Employment of the cmlc2;nuc-DsRed2 transgenic strain allowed visualization of many individual cardiomyocytes within the injured apex undergoing transitions in contractile gene expression during regeneration (Fig. 3A). To determine whether a definitive undifferentiated progenitor cell exists, developmental timing assays using a double transgenic cmlc2:nuc-DsRed2;cmlc2:EGFP strain were initiated. Because EGFP and DsRed2 have very different maturation and stability characteristics, this strain could temporally and spatially characterize activity at the cmlc2 promoter during regeneration (Baird et al., 2000; Verkhusha et al., 2001; Bevis and Glick, 2002; Verkhusha et al., 2003). For example, the emergence of tissue positive for EGFP (fast-fluorescing) and negative for DsRed (slowfluorescing) at the apical edge of the regenerate indicated that aforementioned transitions in Cmlc2 expression represent increases in differentiation from an undifferentiated state. By contrast, these assays revealed no evidence that reductions in Cmlc2-expression occur during regeneration. Coincident with these differentiation events was the emergence of cells at the resection plane with enhanced expression of multiple markers, including nkx2.5, tbx20 and hand2, that are also expressed by cardiac progenitors during embryonic heart development in zebrafish (Fig. 3B) (Chen and Fishman, 1996; Griffin et al., 2000; Yelon et al., 2000). BrdU-labeling experiments at 7 dpa indicated that

PART  |  12  The Regenerative Heart

(A)

(B)

Figure 3  Creation of a heart field during regeneration. (A) Sections through uninjured and injured cmlc2:nuc-DsRed2 ventricles. Cardiomyocyte nuclear fluorescence is uniform in uninjured and 3 dpa ventricles. A subpopulation of weakly-fluorescing nuclei manifest by 14 dpa (arrowheads) indicating cardiomyocytes differentiating from undifferentiated progenitor cells. Magenta line in high magnification insets delineates high-fluorescing cardiomyocytes (above line) from low-fluorescing cardiomyocytes. By 22 dpa, many low-fluorescing cardiomyocyte nuclei are detectable at apex. (B) The embryonic heart field markers hand2, nkx2.5 and tbx20 are each detectable by in situ hybridization at 7 dpa in a domain of enhanced expression at the apical edge of the regenerate (brackets), a region of the most recent myocardial differentiation events. Expression of cmlc2 indicates differentiated muscle. Figure adapted with permission from Lepilina et al. (2006).

the cardiomyocytes newly-differentiating from an undifferentiated state (as suggested by cmlc2 reporter transgenic strains) preferentially incorporated BrdU. These and other findings from the study argued that undifferentiated cardiac progenitor cells similar to those that make up the embryonic heart field are responsible for generating new, proliferative cardiomyocytes during regeneration. That is, the adult zebrafish heart somehow organizes a new heart field after injury, a field that can associate with existing cardiac muscle and build on it with new cardiomyocytes (Fig. 4). In this way, the injured heart employs a structure analogous to a blastema to regenerate new muscle, just like an amputated limb or fin.

Chapter  |  12.2  Cardiac Regeneration in the Zebrafish Model System

Day 0

Day 3 Outflow tract

Ventricular wall

Atrium Amputation site

Day 7

Myocardial progenitor

Day 14

clot

Days 30 to 60

New muscle Fibrin

Proliferating cardiomyocyte

Figure 4  Model for cardiac progenitor cell participation during heart regeneration. Myocardial progenitor cells first originate in the wound by 3–4 dpa, express embryonic heart field markers, associate with existing muscle, and differentiate. For the next few weeks, regenerative morphogenesis occurs in a shifting wave of progenitor cell seeding, cardiomyocyte differentiation and proliferation events, to contribute new layers of muscle.

IV.C.  Possible Origins of Progenitor Cells The source of progenitor cells that could establish a heart field during regeneration is unknown. That is, what is the nature and location of these cells prior to their acquisition of cardiac fate? Possible sources include reserve populations similar to adult mammalian cardiac progenitors, positive for Islet1, c-Kit, or Sca1. In this way, the injured heart would access progenitor cells by activating an existing pool that is similar to skeletal muscle satellite cells. Alternatively, myocardial progenitor cells may be generated by transdifferentiation of other cell types in the heart like the endocardium or epicardium, or even through dedifferentiation of existing muscle. Although no evidence for dedifferentiation was observed during zebrafish heart regeneration, a recent study of the injured newt heart suggested that a rapid dedifferentiation program is stimulated after injury that can create progenitor cells (Laube et al., 2006). In that study, which used a ventricular clamping injury model that does not remove tissue, the authors reported that a rapid reduction of contractile gene expression occurs within one day of injury. They also found that newt cardiomyocytes isolated from the heart and transplanted into limb blastemata show a similar downregulatory response. While it is possible that these regulatory changes reflect some aspect of a stress response and not a true dedifferentiation program, it suggests a provocative method by which to create cardiac progenitor cells. Lineage tracing technologies should be helpful to resolve the question experimentally of how progenitor cells arise during heart regeneration. That is, one would label

845

certain cardiac or extracardiac cell types, injure the animal and allow regeneration, and then assess whether labeled descendants include cardiomyocytes of the completed regenerate. Labeling techniques might involve introducing fluorescent dyes to directly tag cells, but this could prove technically difficult in the contracting adult heart. In fact, genetic tracing methods that involve Cre-recombinase have been immensely successful in mice. Cre excises genomic DNA positioned between small regions of DNA sequence called loxP sites. In genetic lineage tracing experiments, Cre-recombinase expression is driven in a specific cell type, while a second transgene allows strong expression of a reporter gene only after a loxp-flanked transcriptional stop sequence is excised. This approach succeeded in identifying islet1-positive cells as progenitors to differentiated cardiomyocytes (Cai et al., 2003; Laugwitz et al., 2005). It was also recently employed in an inducible manner to determine the extent to which undifferentiated progenitor cells might contribute new cardiomyocytes in either the aging or infarcted adult mouse heart. Here, the authors’ irreversibly-labeled cardiomyocytes using Cre-based technology and found that, while no new cardiomyocyte maturation appears to occur during normal aging, there is evidence for progenitor cell-based cardiomyocyte derivation after injury (Hsieh et al., 2007). Although it is critical to determine how progenitors of new muscle first originate in the injured zebrafish heart, the evidence that zebrafish effectively influence progenitor cells to achieve cardiac regeneration is a provocative finding, lending optimism to the idea that mammalian cardiac stem cells may someday be coaxed toward therapeutic regeneration.

V.  Nonmyocardial cells and   heart regeneration How are zebrafish cardiac cells successfully propelled to regenerate? Determining nonmyocardial influences, or describing the niche, is of equal importance to characterizing the progenitors themselves. Prominent nonmyocardial tissues in the adult zebrafish heart are the ­epicardium, a thin epithelial layer enveloping the chambers, the endocardium, a layer of endothelial cells that lines the inner trabecular myofibers, and the coronary vasculature that provides nutrition to the myocardial wall. Multiple other cell types undoubtedly exist in adult zebrafish hearts which have not been well-characterized, such as a cardiac fibroblast population similar to fibroblasts well-represented in mammalian hearts. A recent study drew attention to the epicardium, a tissue that in its adult form does not receive much research attention. During embryonic cardiac development, epicardial cells migrate out and over the naked myocardium from a structure called the proepicardial organ, undergo epithelial-to-mesenchymal transition (EMT), and invade

846

PART  |  12  The Regenerative Heart

Uninjured

3 dpa

14 dpa

tbx18

Figure 5  Whole-mount (top) and section (bottom) in situ hybridization of uninjured and 3 and 14 dpa adult zebrafish hearts for tbx18, a marker of the embryonic epicardium. Expression is low or undetectable in the uninjured heart, but induced at high levels in epicardial tissue surrounding the atrium (a) and ventricle (v) for tbx18. By 14 dpa, tbx18 expression is localized to the apical wound (arrowhead). Figure adapted with permission from Lepilina et al. (2006).

the underlying subepicardial space and myocardial wall (see Chapters 5.1 and 5.2). After doing so, they differentiate into endothelial and smooth muscle cells of the coronary vasculature (Dettman et al., 1998; Olivey et al., 2004), as well as into cardiac fibroblasts. The epicardium is the primary and possibly sole source of blood vessels for the embryonic heart, and thus is a critical structure for cardiac growth. Recent studies identified interesting and important roles for the epicardium during myocardial regeneration in zebrafish. Epicardial cells exhibit a rapid and robust response to partial ventricular resection injury, proliferating and expressing genes that are also expressed by the embryonic epicardium, tbx18 and raldh2, within 1–2 days of resection (Lepilina et al., 2006). Strikingly, epicardial cells covering both cardiac chambers, not only that tissue adjacent to the injured apex, become developmentally-activated in this manner. Presumably through migration of this activated epicardium, the injured apex is covered with a new epithelial layer of cells within 7–14 dpa (Fig. 5). Then, in a process highly-reminiscent of epicardial cell activity during embryonic heart development, a subpopulation of cells from the overlying epicardium invades the regenerating tissue and settles several cell layers deep into new muscle. The epicardial-derived cells in the regenerate are likely to possess a similar role as they do in the embryo, as progenitors for vascular tissue. Indeed, the regenerate undergoes heavy vascularization by 14 dpa, coincident with epicardial cell invasion (Lepilina et al., 2006). Therefore, it

appears that a primary function of the dramatic response of the epicardium is to create an abundance of tissue with the capacity to identify and vascularize the regenerating muscle as it accumulates (Fig. 6). This response of the zebrafish epicardium provokes several questions. For instance, how is the entire epicardium, including that surrounding the atrium, activated by resection of the ventricular apex? Also, how does the epicardium target the injury, and to what extent is this response required for myocardial regeneration? Lepilina et al. addressed this second question by examining the participation of the Fgf signaling pathway during regeneration (Lepilina et al., 2006). Fgfs, a large family of secreted ligands, have been shown to promote epithelial-to-mesenchymal transition in cultured epicardial cells, and thus present a candidate for recruiting epicardial cells to the site of regeneration (Morabita et al., 2001). In support of this idea, the investigators found that an Fgf ligand, fgf17b, was induced in injured and regenerating myocardium, while Fgf receptors fgfr2 and fgfr4 were expressed in epicardially-localized cells invading the regenerate. To test the idea that Fgfs target the epicardial cells to the regenerate, they used an inducible transgenic line that allowed heat-activated expression of a dominant-negative Fgf receptor (Lee et al., 2005). Myocardial regeneration arrested at ~14 dpa in animals undergoing Fgfr inhibition. Associated with the defect were failures in both epicardial epithelial-to-mesenchymal transition and neovascularization of the regenerate (Lepilina et al., 2006). These results

847

Chapter  |  12.2  Cardiac Regeneration in the Zebrafish Model System

Day 0

Day 3 Activated epicardium

Outflow tract

Ventricular wall

Atrium Amputation site

Day 7

New muscle Fibrin

Clot

Day 14

New coronary vessels

Days 30 to 60

Vascularized regenerate

Figure 6  Model for epicardial cell activation and participation in heart regeneration. Resection of the ventricular apex stimulates rapid expansion of the entire epicardium by 3 dpa (black dots). By 7 dpa, raldh2/tbx18-positive epicardial cells begin to surround and invade the wound. At 14 dpa, epicardial-derived cells undergo epithelial-to-mesenchymal transition in response to apparent signals from the injury site like Fgfs, and vascularize the regenerate (green dots). Presence of new coronary vasculature by 14 dpa extends progenitor cell activity and facilitates regenerative restoration and expansion of the ventricular wall.

suggest that the epicardial response and formation of coronary vessels are required to maintain regeneration. The authors proposed that Fgf signaling acts to support regeneration by progenitor cells by directing developmentallyactivated epicardial cells to vascularize the injury site. Indeed, there are likely to be many other molecular interactions between epicardium and myocardium that facilitate regeneration. Furthermore, it will be very interesting to understand the roles of other cardiac cell types, as well as critical signals for mediating their influences, in additional studies that dissect zebrafish heart regeneration.

VI.  Molecular genetic approaches to zebrafish heart regeneration The ability to observe cardiomyocyte division in vitro, as well as the potential for grafting and lineage studies successfully performed in studies of appendage regeneration (Bader and Oberpriller, 1979; Lo et al., 1993; Echeverri and Tanaka, 2002), represent strengths of amphibian model systems for studying heart regeneration. In fact, these approaches would be more difficult to apply in smaller, fully-aquatic adult zebrafish. The zebrafish has a unique position, however, in that it is currently the only highly-regenerative vertebrate model system that is amenable to molecular genetic approaches. Thus, the molecular mechanisms of zebrafish heart regeneration are beginning to receive attention. While there are now functional data that demonstrate a requirement for Fgf signaling during heart regeneration (Lepilina et al., 2006), other signaling pathways have also

been implicated. An earlier study found an induction of notch1b receptor mRNA and deltaC ligand mRNA in the endocardial cell layer of the injured heart, suggesting roles for Notch signaling that require future functional exploration (Raya et al., 2003). Lien et al. recently examined mRNA expression profiles during heart regeneration at multiple timepoints after injury, using Affymetrix microarrays that facilitate genome-wide assays of gene expression (Lien et al., 2006). Several differentially-expressed genes were identified, predicted to be involved in wound healing, tissue remodeling, or cardiomyocyte hyperplasia. It is fascinating that nonmammalian vertebrates typically show successful regeneration in many organs that are nonregenerative in mammals (e.g. retina, appendage, heart …), not just one organ system. This suggests that elevated regenerative capacity is a trait represented by a common set of regenerative mechanisms. In fact, by comparing heart regeneration gene expression profiles with those obtained from regenerating fin tissue, the authors suggested the existence of a set of “regeneration core molecules”. Attention was focused on two members of the Platelet derived growth factor (Pdgf) family, Pdgf-a and Pdgf-b, which showed increased expression after injury by microarray and/or RT-PCR and was shown by others to increase DNA synthesis in cultured newt cardiomyocytes (Soonpaa et al., 1992). The authors found a similar effect of stimulating BrdU incorporation in cultured zebrafish cardiomyocytes, supporting the idea that Pdgf is an injury-activated cardiomyocyte mitogen, and future studies are thus likely to focus on in vivo functions of Pdgf. This study illustrates how microarray and/or in situ hybridization screens can yield many candidate genes during heart regeneration.

848

PART  |  12  The Regenerative Heart

How does one definitively test the function of candidate genes during heart regeneration? The use of heat-shock inducible transgenes to block Fgfr function was effective in identifying requirements for Fgf signaling (Lepilina et al., 2006). Continued use and improvement of inducible transgenic approaches will facilitate examination of other molecular signaling pathways during regeneration (Stoick-Cooper et al., 2007). In addition, adenoviruses may provide a useful means to deliver expression cassettes locally to the heart, as used recently to dissect fin regeneration (Kawakami et al., 2006). To supplement inducible transgenesis-targeting candidate genes, heart regeneration can potentially be dissected through antisense technology and chemical genetics. For antisense approaches, most zebrafish embryologists employ morpholinos, which are stable, modified oligonucleotides complementary to start- or splice-sites that block translation or splicing (Nasevicius and Ekker, 2000). Morpholinos have also been delivered to the regenerating adult spinal cord or fin with some effects (Becker et al., 2004; Thummel et al., 2006). Whether morpholinos can be used to knockdown adult cardiac gene expression is unknown. A more commonly-used antisense technology for other model systems is RNA interference, which takes advantage of the cellular machinery that generates microRNAs (Pasquinelli and Ruvkun, 2002; Chapter 10.3). However, RNA interference was not immediately effective in zebrafish (Oates et al., 2000), and has yet to be adopted by zebrafish researchers. Interfering RNAs have an advantage over morpholinos in that they could potentially be expressed behind a tissuespecific or inducible promoter as part of a transgenic construct to knockdown expression in adult fish. With respect to chemical genetics, approaches relevant to regeneration have already been performed. Myoseverin ENU-mutagenized male

was identified from a library of purine compounds for its ability to initiate cleavage of cultured, multinucleate myotubes into proliferative, mononuclear cells (Rosania et al., 2000). Relevant to cardiovascular biology in zebrafish, chemical modulators of heart rate (Burns et al., 2005) and vascular development (Peterson et al., 2004) have been identified. While chemical genetics may seem a tedious way to dissect cardiac regeneration, it is possible that its combination with transgenic reporter strains would be fruitful. For instance, an injected compound, analogous to a mutation, may block expression of a regeneration marker in an injured fish, or even induce expression of a regeneration marker in an uninjured animal. An entire library could be screened in such a way to find those compounds that enhance or inhibit some aspect of regeneration. Forward genetics, which starts with a phenotype and works toward identification of the disrupted gene, has long been the tool of choice for researchers who work with zebrafish. Several recent studies have employed forward genetics to identify genes essential for regeneration of the caudal fin, using ethylnitrosourea (ENU), an alkylating point mutagen, to mutagenize the adult male germline (Chapter 11.2). In 1995, Johnson and Weston pioneered this approach, showing that screens for temperature-sensitive (ts) mutations can be fruitful in zebrafish, and that these screens can identify mutants with conditional defects in appendage regeneration (Johnson and Weston, 1995). One assumption of this type of screen is that many genes important for regeneration also have earlier roles in ontogenetic development; hence, the use of temperature changes to identify conditional alleles (Fig. 7). Importantly, this screen is also designed to identify strong, non-ts mutations in hypothetical regeneration-specific genes, or non-ts alleles that affect

F1

F2 2N oocytes

X

Wildtype female

Early pressure parthenogenesis Amputate, place at 33°C conditions

Potential regeneration mutant Screen after 5–10 days Figure 7  Mutagenesis and genetic screening strategy to detect temperature-sensitive fin regeneration mutants. N-ethyl-N-nitrosourea (ENU) is an alkylating mutagen commonly used in zebrafish and mice. Early pressure parthenogenesis involves fertilization of oocytes with UV-irradiated sperm, followed by treatment with hydrostatic pressure. Such a step generates diploid progeny, some homozygous for ENU-generated mutations, in a manner that saves facility space, as well as one generation of mating crosses (Streisinger et al., 1981; Johnson et al., 1995). At 2–3 months of age, caudal fins are amputated, and animals are moved from 25°C to 33°C for one week. Regeneration mutants can easily be detected after 5–10 days incubation at 33°C.

Chapter  |  12.2  Cardiac Regeneration in the Zebrafish Model System

regeneration specifically in genes that function in both embryogenesis and regeneration. A later mutagenesis screen found additional mutants and, with the establishment of genetic markers and progress of genome sequencing efforts, the disrupted genes were mapped to chromosomal loci (Poss et al., 2002b). Positional cloning revealed the affected genes for four of these mutants: (1) mps1, a cell-cycle regulator essential for the mitotic checkpoint (Poss et al., 2002b); (2) sly1, involved in intracellular vesicular trafficking (Nechiporuk et al., 2003); (3) hsp60, a chaperone essential for the response to stress (Makino et al., 2005); and (4) fgf20a, one of many zebrafish Fgfs (Whitehead et al., 2005). Will forward genetics be applied to discover new genes essential for heart regeneration? The best genetic screens identify mutations disrupting a consistent and easilyobserved trait. To fit this description, several modifications to the current analysis of zebrafish heart regeneration could be implemented. First, the current resection injury model will likely need to be altered to provide a more precise injury, changes that may involve chemical lesions or promoter-driven expression of cytotoxic proteins. There have been recent findings that demonstrate the ability of a bacterial nitroreductase system to cause inducible, cell type-specific lesions in zebrafish embryos. Here, nitroreductase is expressed in transgenic zebrafish using a tissuespecific promoter, and converts an exogenously supplied prodrug metronidazole to a cytotoxic product (Curado et al., 2007; Pisharath et al., 2007). Ideally, this modified injury model would normally be repaired quickly, since the current resection model takes a lengthy 1–2 months or more to reach approximate completion. Second, assessment of regeneration is currently tedious, requiring histological processing of the entire ventricle. A higherthroughput method of assessing successful regeneration would be greatly advantageous. It is interesting that two of the four genes shown to be essential for fin regeneration were also found to be critical for heart regeneration (Poss et al., 2002b; Makino et al., 2005). In particular, the gene mps1 is upregulated in a small number of cells of both the regenerating fin and heart, and its function is required for each regenerative event (Poss et al., 2002a,b). Therefore, a subset of heart regeneration mutants can be obtained through secondary screening of fin regeneration mutants. Additionally, the use of transgenic reporter strains should facilitate directed forward genetic screens for heart regeneration mutants. For instance, fli1:EGFP marks endothelial cells and has been used to identify mutants in vascular development (Lawson and Weinstein, 2002; Lawson et al., 2003). Similarly, a “gut” GFP strain that marks the liver, pancreas and intestine has been employed to reveal new mutations that disrupt development of these normallyconcealed structures (Field et al., 2003). The appropriate fluorescent reporter strains could potentially enhance identification and/or characterization of mutations that disrupt heart regeneration.

849

VII.  Why does the zebrafish   heart regenerate? Why have zebrafish retained the ability to regenerate? It is thought that the capacity for organ regeneration is an ancestral condition that has occasionally been diminished in the course of vertebrate evolution (Scadding, 1977; Goss, 1992; Wagner and Misof, 1992). Thus, most biologists suspect that the molecular machinery to optimize regeneration is present but poorly-utilized in mammals. By this line of reasoning, zebrafish heart regeneration may represent an optimal utilization of universal cardiac machinery. Indeed, the findings that progenitor cells of some form exist both in the regenerative zebrafish heart, and in the hearts of less-regenerative mammals supports this idea. Zebrafish have ostensibly found some method to optimize the activity of progenitor cells, perhaps either by maintaining more cells, or by harboring a more cultivating environment for regeneration. Also, both mammalian and nonmammalian hearts contain an epicardial cell layer, yet zebrafish have found some way to activate the epicardium after injury, a process linked with essential neovascularization of regenerating muscle (Lepilina et al., 2006). This result points to the adult mammalian epicardium as a potential cellular source to assist myocardial regeneration or survival. Indeed, mammalian myocardial infarcts typically show poor or insufficient neovascularization, a response that many are trying to improve experimentally. Recent findings have indicated that the G-actin sequestering protein, Thymosin-ß4, may influence the mammalian epicardium. Treatment of adult cardiac explants with Thymosin-ß4 induced the migration of fibroblasts, endothelial and smooth muscle cells as assessed by gene expression and cellular morphology (Smart et al., 2007). In addition, in vivo Thymosin-ß4 treatment could partially restore cardiac survival and function following coronary ligation (Bock-Marquette et al., 2004). Notably, Thymosinß4 expression is induced in the injured zebrafish heart, suggesting that fish naturally release this epicardial stimulant on injury (Lien et al., 2006). It was also shown recently that delivery of Fgfs by release from peptide nanofibers, a gradual local delivery system, can increase neovascularization and reduce infarct size in the ischemic rodent heart (Engel et al., 2006). Related to this, zebrafish have a natural ability to synthesize Fgfs after myocardial injury, a signal that appears to recruit Fgf receptor-expressing epicardial-derived cells toward regenerating muscle (Lepilina et al., 2006). Thus, what has been and what will be discovered about zebrafish heart regeneration is quite likely to illuminate possible strategies for enhancing regeneration in the mammalian heart (see Chapter 14.4). Interestingly, there are numerous differences in the biology of teleosts and mammals, as well as specific differences in cardiomyocyte cellular structure and anatomy,

850

PART  |  12  The Regenerative Heart

all of which might contribute to regenerative variability. Unlike mammals, zebrafish can grow throughout most of adulthood, a phenomenon called “indeterminate growth” (Jordan, 1905). In fact, their growth can be affected markedly by changes in nutrition and population density (Goldsmith et al., 2006). It is thus possible that the capacity to replace cardiac tissue rapidly in teleosts has been retained in evolution as a function of the need for robust animal and cardiac growth. Indeed, a recent study has found that experimentally-induced adult cardiac growth in zebrafish is hyperplastic, and appears to rely on the same signals present or required during cardiac regeneration (Wills et al., 2008). Some differences in cardiac anatomy exist between mammals and teleosts. The zebrafish ventricle has a thin wall of compact muscle surrounding a much larger compartment of myofibers organized into elaborate trabeculae. It is intriguing that this structure is very similar to that of the embryonic mammalian ventricle prior to its septation and fusion of trabeculer myofibers into a thick, vascularized wall (Sedmera et al., 2000). That the mammalian heart has a more differentiated, contractile anatomy is apparent not only in gross cardiac structure, but also in cellular features. Teleost cardiomyocytes are 2–10 times smaller, mononucleated, have a greatly-reduced sarcoplasmic reticulum and lack the T-tubule system found in skeletal muscle and mammalian cardiac muscle (Farrell, 1992). One might speculate that the teleost heart is better designed for growth and regeneration, while the mammalian heart is better designed for sheer contractile force. Nevertheless, none of the mentioned differences between

lower and higher vertebrate hearts preclude the idea that the mammalian heart could be stimulated to regenerate, especially if that regeneration is due to mobilization of a progenitor cell population. An interesting concept that has emerged from initial findings is that regeneration and fibrosis are competing events in the vertebrate heart. That is, if there is a capacity for injurystimulated cardiomyocyte hyperplasia beyond a certain threshold, regenerative mechanisms will overcome scarring. Results consistent with this idea came from experiments with zebrafish possessing a ts mutation in the cell-cycle checkpoint kinase Mps1 (Poss et al., 2002b). As mentioned earlier, mps1 mutants were initially identified based on their defects in caudal fin regeneration. Serendipitously, mps1 mutants also showed defects in cardiac regeneration at a temperature restrictive for the mutation (Poss et al., 2002b). Instead of regenerating muscle in response to ventricular resection injury, mps1 mutants repaired wounds by forming large, collagen-rich scars. Inhibition of Fgf signaling also stunts cardiac regeneration and causes scarring (Lepilina et al., 2006). These results indicate that even vertebrates with high cardiac regenerative capacity have a default scarring mechanism; normally, regeneration somehow restricts this pathway (Fig. 8). The implication is exciting; perhaps by stimulating regeneration in a poorly-regenerative system like the mammalian heart, scarring events characteristic of myocardial infarction would be restricted by new muscle formation. Similarly, deterring cardiac scarring mechanisms would perhaps favor regeneration in mammals.

Normal regeneration

Outflow tract

Little or no scarring Ventricular wall

Atrium Amputation site

Fgf receptor inhibition or mps1 mutation Scar tissue Figure 8  Evidence that regeneration and scarring are complementary pathways in the injured heart. Zebrafish normally regenerate a resected ventricular wall, with little or no scarring (top). However, if Fgf signaling or the function of the cell-cycle regulator Mps1 are disrupted after injury, regeneration is blocked or arrests prematurely. In the absence of optimal regeneration, collagen-rich scar tissue completes repair of the injury. This result suggests a default mechanism of scarring that is similar to mammalian cardiac repair and is enabled by a lack of regeneration.

Chapter  |  12.2  Cardiac Regeneration in the Zebrafish Model System

VIII.  Summary and future directions Unlike mammals, zebrafish have the capacity to regenerate cardiac muscle removed by mechanical injury. They do so by stimulating the creation of new cardiomyocytes at the injury site, and by restricting scar formation. Research has essentially just begun to dissect the underlying mechanisms. Initial findings indicate that the regenerating heart establishes a field of cardiac progenitors, much like the embryonic heart field, as a source of new, proliferative cardiomyocytes. Regenerative cardiogenesis then proceeds in a wave of progenitor cell seeding, cardiomyocyte differentiation and proliferation. Concomitantly, the epicardial cell layer that surrounds the myocardium proper responds to injury by expressing embryonic epicardial markers and proliferating. Then, in a process of epithelial-to-mesenchymal transition reminiscent of embryonic epicardial cells, the wound and regenerating muscle become populated by invading epicardial-derived cells. These epithelial-tomesenchymal transition events, and the neovascularization that accompany them, require the Fgf signaling pathway, ostensibly with Fgf ligands synthesized in injured/ regenerating muscle recruiting epicardial-derived cells that express Fgf receptors. These events point to an intricate orchestration of different cardiac cell types to carry out regeneration in the injured zebrafish heart. In future experiments, multiple injury models should be examined in attempts to parallel human myocardial infarction; i.e., models that focally destroy myocardium without its removal. These experiments would directly examine whether and how infarction injuries can be regenerated. Second, optimal culture conditions for adult zebrafish cardiac cells should be established, so that in vitro progenitor cell differentiation or other regeneration paradigms can be dissected, or chemical genetic strategies explored. Third, genetic lineage tracing tools that have been so successful for elucidating progenitor cell biology in mammalian systems must be introduced to the zebrafish system. Combined with the identification of additional molecular markers, these experiments would conclusively define the cellular origin of regenerated myocardium. Fourth, there must be new tools that facilitate an appropriate genetic screen to reveal informative mutations detrimental to cardiac regeneration. Such mutations would hold the key to discovery of new factors that carry out regenerative events. When regenerative mechanisms in zebrafish achieve cellular and molecular definition, detailed comparisons with nonregenerative mammalian myocardium will be possible, comparisons that should provide insight into increasing the regenerative ability of the human heart.

References Akimenko, M.A., Mari-Beffa, M., Becerra, J., Geraudie, J., 2003. Old questions, new tools, and some answers to the mystery of fin regeneration. Dev. Dyn. 226, 190–201.

851

Bader, D., Oberpriller, J., 1979. Autoradiographic and electron microscopic studies of minced cardiac muscle regeneration in the adult newt, notophthalmus viridescens. J. Exp. Zool. 208, 177–193. Baird, G.S., Zacharias, D.A., Tsien, R.Y., 2000. Biochemistry, mutagenesis, and oligomerization of DsRed, a red fluorescent protein from coral. Proc. Natl. Acad. Sci. USA 97, 11984–11989. Becker, C.G., Lieberoth, B.C., Morellini, F., Feldner, J., Becker, T., Schachner, M., 2004. L1.1 is involved in spinal cord regeneration in adult zebrafish. J. Neurosci. 24, 7837–7842. Becker, R.O., Chapin, S., Sherry, R., 1974. Regeneration of the ventricular myocardium in amphibians. Nature 248, 145–147. Becker, T., Wullimann, M.F., Becker, C.G., Bernhardt, R.R., Schachner, M., 1997. Axonal regrowth after spinal cord transection in adult zebrafish. J. Comp. Neurol. 377, 577–595. Beltrami, A.P., Barlucchi, L., Torella, D., Baker, M., Limana, F., Chimenti, S., Kasahara, H., Rota, M., Musso, E., Urbanek, K., Leri, A., Kajstura, J., Nadal-Ginard, B., Anversa, P., 2003. Adult cardiac stem cells are multipotent and support myocardial regeneration. Cell 114, 763–776. Bettencourt-Dias, M., Mittnacht, S., Brockes, J.P., 2003. Heterogeneous proliferative potential in regenerative adult newt cardiomyocytes. J. Cell Sci. 116, 4001–4009. Bevis, B.J., Glick, B.S., 2002. Rapidly maturing variants of the Discosoma red fluorescent protein (DsRed). Nat. Biotechnol. 20, 83–87. Bock-Marquette, I., Saxena, A., White, M.D., Dimaio, J.M., Srivastava, D., 2004. Thymosin beta4 activates integrin-linked kinase and promotes cardiac cell migration, survival and cardiac repair. Nature 432, 466–472. Brockes, J.P., Kumar, A., 2005. Appendage regeneration in adult vertebrates and implications for regenerative medicine. Science 310, 1919–1923. Burns, C.G., Milan, D.J., Grande, E.J., Rottbauer, W., MacRae, C.A., Fishman, M.C., 2005. High-throughput assay for small molecules that modulate zebrafish embryonic heart rate. Nat. Chem. Biol. 1, 263–264. Cai, C.L., Liang, X., Shi, Y., Chu, P.H., Pfaff, S.L., Chen, J., Evans, S., 2003. Isl1 identifies a cardiac progenitor population that proliferates prior to differentiation and contributes a majority of cells to the heart. Dev. Cell 5, 877–889. Chen, J.N., Fishman, M.C., 1996. Zebrafish tinman homolog demarcates the heart field and initiates myocardial differentiation. Development 122, 3809–3816. Curado, S., Anderson, R.M., Jungblut, B., Mumm, J., Schroeter, E., Stainier, D.Y., 2007. Conditional targeted cell ablation in zebrafish: a new tool for regeneration studies. Dev. Dyn. 236, 1025–1035. Dettman Jr, R.W., Denetclaw, W., Ordahl, C.P., Bristow, J., 1998. Common epicardial origin of coronary vascular smooth muscle, perivascular fibroblasts, and intermyocardial fibroblasts in the avian heart. Dev. Biol. 193, 169–181. Echeverri, K., Tanaka, E.M., 2002. Ectoderm to mesoderm lineage switching during axolotl tail regeneration. Science 298, 1993–1996. Engel, F.B., Schebesta, M., Duong, M.T., Lu, G., Ren, S., Madwed, J.B., Jiang, H., Wang, Y., Keating, M.T., 2005. P38 MAP kinase inhibition enables proliferation of adult mammalian cardiomyocytes. Genes Dev. 19, 1175–1187. Engel, F.B., Hsieh, P.C., Lee, R.T., Keating, M.T., 2006. FGF1/p38 MAP kinase inhibitor therapy induces cardiomyocyte mitosis, reduces scarring, and rescues function after myocardial infarction. Proc. Natl. Acad. Sci. USA 103, 15546–15551. Farrell, A., Jones, D.R., 1992. Fish Physiology: The Heart. Academic Press, San Diego, CA.

852

Field, H.A., Dong, P.D., Beis, D., Stainier, D.Y., 2003. Formation of the digestive system in zebrafish. II. Pancreas morphogenesis. Dev. Biol. 261, 197–208. Flink, I.L., 2002. Cell cycle reentry of ventricular and atrial cardiomyocytes and cells within the epicardium following amputation of the ventricular apex in the axolotl, Amblystoma mexicanum: confocal microscopic immunofluorescent image analysis of ­ bromodeoxyuridine-labeled nuclei. Anat Embryol. (Berlin) 205, 235–244. Fujisawa, T., 2003. Hydra regeneration and epitheliopeptides. Dev. Dyn. 226, 182–189. Goldsmith, M.I., Iovine, M.K., O’Reilly-Pol, T., Johnson, S.L., 2006. A developmental transition in growth control during zebrafish caudal fin development. Dev. Biol. 259, 450–457. Goss, R.J., 1992. The evolution of regeneration: adaptive or inherent? J. Theor. Biol. 159, 241–260. Griffin, K.J., Stoller, J., Gibson, M., Chen, S., Yelon, D., Stainier, D.Y., Kimelman, D., 2000. A conserved role for H15-related T-box transcription factors in zebrafish and Drosophila heart formation. Dev. Biol. 218, 235–247. Grunwald, D.J., Eisen, J.S., 2002. Headwaters of the zebrafish – emergence of a new model vertebrate. Nat. Rev. Genet. 3, 717–724. Hsieh, P.C., Segers, V.F., Davis, M.E., Macgillivray, C., Gannon, J., Molkentin, J.D., Robbins, J., Lee, R.T., 2007. Evidence from a genetic fate-mapping study that stem cells refresh adult mammalian cardiomyocytes after injury. Nat. Med. 13, 970–974. Johnson, S.L., Weston, J.A., 1995. Temperature-sensitive mutations that cause stage-specific defects in Zebrafish fin regeneration. Genetics 141, 1583–1595. Johnson, S.L., Africa, D., Horne, S., Postlethwait, J.H., 1995. Half-tetrad analysis in zebrafish: mapping the ros mutation and the centromere of linkage group I. Genetics 139, 1727–1735. Jordan, D.S., 1905. A Guide to the Study of Fishes. Henry Holt and Co, NY, NewYork. Kawakami, Y., Rodriguez Esteban, C., Raya, M., Kawakami, H., Marti, M., Dubova, I., Izpisua Belmonte, J.C., 2006. Wnt/beta-catenin signaling regulates vertebrate limb regeneration. Genes Dev. 20, 3232–3237. Kumar, A., Velloso, C.P., Imokawa, Y., Brockes, J.P., 2004. The regenerative plasticity of isolated urodele myofibers and its dependence on MSX1. PLoS Biol. 2, E218. Kumar, A., Godwin, J.W., Gates, P.B., Garza-Garcia, A.A., Brockes, J.P., 2007. Molecular basis for the nerve dependence of limb regeneration in an adult vertebrate. Science 318, 772–777. Laube, F., Heister, M., Scholz, C., Borchardt, T., Braun, T., 2006. Re-programming of newt cardiomyocytes is induced by tissue regeneration. J. Cell Sci. 119, 4719–4729. Laugwitz, K.L., Moretti, A., Lam, J., Gruber, P., Chen, Y., Woodard, S., Lin, L.Z., Cai, C.L., Lu, M.M., Reth, M., Platoshyn, O., Yuan, J.X., Evans, S., Chien, K.R., 2005. Postnatal isl1 cardioblasts enter fully differentiated cardiomyocyte lineages. Nature 433, 647–653. Lawson, N.D., Weinstein, B.M., 2002. In vivo imaging of embryonic vascular development using transgenic zebrafish. Dev. Biol. 248, 307–318. Lawson, N.D., Mugford, J.W., Diamond, B.A., Weinstein, B.M., 2003. phospholipase C gamma-1 is required downstream of vascular endothelial growth factor during arterial development. Genes Dev. 17, 1346–1351. Lee, Y., Grill, S., Sanchez, A., Murphy-Ryan, M., Poss, K.D., 2005. Fgf signaling instructs position-dependent growth rate during zebrafish fin regeneration. Development 132, 5173–5183.

PART  |  12  The Regenerative Heart

Lepilina, A., Coon, A.N., Kikuchi, K., Holdway, J.E., Roberts, R.W., Burns, C.G., Poss, K.D., 2006. A dynamic epicardial injury response supports progenitor cell activity during zebrafish heart regeneration. Cell 127, 607–619. Lien, C.L., Schebesta, M., Makino, S., Weber, G.J., Keating, M.T., 2006. Gene expression analysis of zebrafish heart regeneration. PLoS Biol. 4, e260. Lo, D.C., Allen, F., Brockes, J.P., 1993. Reversal of muscle differentiation during urodele limb regeneration. Proc. Natl. Acad. Sci. USA 90, 7230–7234. Mably, J.D., Mohideen, M.A., Burns, C.G., Chen, J.N., Fishman, M.C., 2003. Heart of glass regulates the concentric growth of the heart in zebrafish. Curr. Biol. 13, 2138–2147. Makino, S., Whitehead, G.G., Lien, C.L., Kim, S., Jhawar, P., Kono, A., Kawata, Y., Keating, M.T., 2005. Heat-shock protein 60 is required for blastema formation and maintenance during regeneration. Proc. Natl. Acad. Sci. USA 102, 14599–14604. Morabito, C.J., Dettman, R.W., Kattan, J., Collier, J.M., Bristow, J., 2001. Positive and negative regulation of epicardial–mesenchymal transformation during avian heart development. Dev. Biol. 234, 204–215. Morgan, T.H., 1906. The physiology of regeneration. J. Exp. Zool. 3, 457–503. Morrison, J.I., Loof, S., He, P., Simon, A., 2006. Salamander limb regeneration involves the activation of a multipotent skeletal muscle satellite cell population. J. Cell Biol. 172, 433–440. Nasevicius, A., Ekker, S.C., 2000. Effective targeted gene “knockdown” in zebrafish. Nat. Genet. 26, 216–220. Nechiporuk, A., Keating, M.T., 2002. A proliferation gradient between proximal and msxb-expressing distal blastema directs zebrafish fin regeneration. Development 129, 2607–2617. Nechiporuk, A., Poss, K.D., Johnson, S.L., Keating, M.T., 2003. Positional cloning of a temperature-sensitive mutant emmental reveals a role for sly1 during cell proliferation in zebrafish fin regeneration. Dev. Biol. 258, 291–306. Neff, A.W., Dent, A.E., Armstrong, J.B., 1996. Heart development and regeneration in urodeles. Int. J. Dev. Biol. 40, 719–725. Oates, A.C., Bruce, A.E., Ho, R.K., 2000. Too much interference: injection of double-stranded RNA has nonspecific effects in the zebrafish embryo. Dev. Biol. 224, 20–28. Oberpriller, J.O., Oberpriller, J.C., 1974. Response of the adult newt ventricle to injury. J. Exp. Zool. 187, 249–253. Oberpriller, J.O., Oberpriller, J.C., Matz, D.G., Soonpaa, M.H., 1995. Stimulation of proliferative events in the adult amphibian cardiac myocyte. Ann. NY Acad. Sci. 752, 30–46. Oh, H., Bradfute, S.B., Gallardo, T.D., Nakamura, T., Gaussin, V., Mishina, Y., Pocius, J., Michael, L.H., Behringer, R.R., Garry, D.J., Entman, M.L., Schneider, M.D., 2003. Cardiac progenitor cells from adult myocardium: homing, differentiation, and fusion after infarction. Proc. Natl. Acad. Sci. USA 100, 12313–12318. Olivey, H.E., Compton, L.A., Barnett, J.V., 2004. Coronary vessel development: the epicardium delivers. Trends Cardiovasc Med. 14, 247–251. Otteson, D.C., Hitchcock, P.F., 2003. Stem cells in the teleost retina: persistent neurogenesis and injury-induced regeneration. Vision Res. 43, 927–936. Pasquinelli, A.E., Ruvkun, G., 2002. Control of developmental timing by micrornas and their targets. Annu. Rev. Cell. Dev. Biol. 18, 495–513. Pasumarthi, K.B., Field, L.J., 2002. Cardiomyocyte cell cycle regulation. Circ. Res. 90, 1044–1054. Peterson, R.T., Shaw, S.Y., Peterson, T.A., Milan, D.J., Zhong, T.P., Schreiber, S.L., MacRae, C.A., Fishman, M.C., 2004. Chemical

Chapter  |  12.2  Cardiac Regeneration in the Zebrafish Model System

suppression of a genetic mutation in a zebrafish model of aortic coarctation. Nat. Biotechnol. 22, 595–599. Pisharath, H., Rhee, J.M., Swanson, M.A., Leach, S.D., Parsons, M.J., 2007. Targeted ablation of beta cells in the embryonic zebrafish pancreas using E. coli nitroreductase. Mech. Dev. 124, 218–229. Poss, K.D., Nechiporuk, A., Hillam, A.M., Johnson, S.L., Keating, M.T., 2002a. Mps1 defines a proximal blastemal proliferative compartment essential for zebrafish fin regeneration. Development 129, 5141–5149. Poss, K.D., Wilson, L.G., Keating, M.T., 2002b. Heart regeneration in zebrafish. Science 298, 2188–2190. Poss, K.D., Keating, M.T., Nechiporuk, A., 2003. Tales of regeneration in zebrafish. Dev. Dyn. 226, 202–210. Raya, A., Koth, C.M., Buscher, D., Kawakami, Y., Itoh, T., Raya, R.M., Sternik, G., Tsai, H.J., Rodriguez-Esteban, C., Izpisua-Belmonte, J.C., 2003. Activation of Notch signaling pathway precedes heart regeneration in zebrafish. Proc. Natl. Acad. Sci. USA 100 (Suppl 1), 11889–11895. Reddien, P.W., Sanchez Alvarado, A., 2004. Fundamentals of planarian regeneration. Annu. Rev. Cell Dev. Biol. 20, 725–757. Rosania, G.R., Chang, Y.T., Perez, O., Sutherlin, D., Dong, H., Lockhart, D.J., Schultz, P.G., 2000. Myoseverin, a microtubule-binding molecule with novel cellular effects. Nat. Biotechnol. 18, 304–308. Rumyantsev, P.P., 1977. Interrelations of the proliferation and differentiation processes during cardiac myogenesis and regeneration. Int. Rev. Cytol. 51, 187–273. Scadding, S.R., 1977. Phylogenic distribution of limb regeneration capacity in adult Amphibia. J. Exp. Zool. 202, 57–68. Sedmera, D., Pexieder, T., Vuillemin, M., Thompson, R.P., Anderson, R.H., 2000. Developmental patterning of the myocardium. Anat. Rec. 258, 319–337. Shizuru, J.A., Negrin, R.S., Weissman, I.L., 2005. Hematopoietic stem and progenitor cells: clinical and preclinical regeneration of the hematolymphoid system. Annu. Rev. Med. 56, 509–538. Smart, N., Risebro, C.A., Melville, A.A., Moses, K., Schwartz, R.J., Chien, K.R., Riley, P.R., 2007. Thymosin beta4 induces adult epicardial progenitor mobilization and neovascularization. Nature 445, 177–182. Soonpaa, M.H., Oberpriller, J.O., Oberpriller, J.C., 1992. Stimulation of DNA synthesis by PDGF in the newt cardiac myocyte. J. Mol. Cell. Cardiol. 24, 1039–1046. Spallanzani, L., 1768. “An Essay on Animal Reproductions. T Becket, London, UK. Stoick-Cooper, C.L., Weidinger, G., Riehle, K.J., Hubbert, C., Major, M. B., Fausto, N., Moon, R.T., 2007. Distinct Wnt signaling pathways

853

have opposing roles in appendage regeneration. Development 134, 479–489. Streisinger, G., Walker, C., Dower, N., Knauber, D., Singer, F., 1981. Production of clones of homozygous diploid zebra fish (Brachydanio rerio). Nature 291, 293–296. Taub, R., 2004. Liver regeneration: from myth to mechanism. Nat. Rev. Mol. Cell. Biol. 5, 836–847. Thummel Jr, R., Bai, S., Sarras, M.P., Song, P., McDermott, J., Brewer, J., Perry, M., Zhang, X., Hyde, D.R., Godwin, A.R., 2006. Inhibition of zebrafish fin regeneration using in vivo electroporation of morpholinos against fgfr1 and msxb. Dev. Dyn. 235, 336–346. Tsonis, P.A., 1996. Limb Regeneration. Cambridge University Press, Cambridge, UK. Verkhusha, V.V., Otsuna, H., Awasaki, T., Oda, H., Tsukita, S., Ito, K., 2001. An enhanced mutant of red fluorescent protein DsRed for double labeling and developmental timer of neural fiber bundle formation. J. Biol. Chem. 276, 29621–29624. Verkhusha, V.V., Kuznetsova, I.M., Stepanenko, O.V., Zaraisky, A.G., Shavlovsky, M.M., Turoverov, K.K., Uversky, V.N., 2003. High stability of Discosoma DsRed as compared to Aequorea EGFP. Biochemistry 42, 7879–7884. Vihtelic, T.S., Hyde, D.R., 2000. Light-induced rod and cone cell death and regeneration in the adult albino zebrafish (Danio rerio) retina. J. Neurobiol. 44, 289–307. Wagers, A.J., Sherwood, R.I., Christensen, J.L., Weissman, I.L., 2002. Little evidence for developmental plasticity of adult hematopoietic stem cells. Science 297, 2256–2259. Wagner, G.P., Misof, B.Y., 1992. Evolutionary modification of regenerative capability in vertebrates: a comparative study on teleost pectoral fin regeneration. J. Exp. Zool. 261, 62–78. Whitehead, G.G., Makino, S., Lien, C.L., Keating, M.T., 2005. fgf20 is essential for initiating zebrafish fin regeneration. Science 310, 1957–1960. Wills, A.A., Holdway, J.E., Major, R.J., Poss, K.D., 2008. Regulated addition of new myocardial and epicardial cells fosters homeostatic cardiac growth and maintenance in adult zebrafish. Development 135, 183–192. Yelon, D., Horne, S.A., Stainier, D.Y., 1999. Restricted expression of cardiac myosin genes reveals regulated aspects of heart tube assembly in zebrafish. Dev. Biol. 214, 23–37. Yelon, D., Ticho, B., Halpern, M.E., Ruvinsky, I., Ho, R.K., Silver, L.M., Stainier, D.Y., 2000. The bHLH transcription factor hand2 plays parallel roles in zebrafish heart and pectoral fin development. Development 127, 2573–2582.