Zebrafish as a powerful vertebrate model system for in vivo studies of cell death

Seminars in Cancer Biology 17 (2007) 154–165

Review

Zebrafish as a powerful vertebrate model system for in vivo studies of cell death Ujwal J. Pyati a , A. Thomas Look a , Matthias Hammerschmidt b,∗ a

b

Dana-Farber Cancer Institute, 44 Binney Street, Boston, MA 02115, USA Max-Planck Institute of Immunobiology, St¨ubeweg 51, 79108 Freiburg, Germany

Abstract Understanding and manipulating cell death pathways are critical to our ability to treat human degenerative diseases and cancer. The zebrafish Danio rerio, a common aquatic pet, has evolved as a powerful tool for the discovery of genes regulating cellular suicide both during normal vertebrate development and after genetic or environmental insult. In this review, we describe the techniques that can be applied to studying cell death in zebrafish as well as highlighting what has been discovered so far. Finally, we discuss future perspectives in the field and how they relate to human disease. © 2006 Elsevier Ltd. All rights reserved. Keywords: Zebrafish; Apoptosis; Development; Cancer; Degenerative diseases; Genetics

Contents 1. 2.

3. 4.

5. 6. 7.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Zebrafish as a powerful model system to study cell death . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1. Molecular techniques for in vivo examination of cell death . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2. Forward genetic screens for unbiased gene discovery . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3. “Reverse Genetics” via target-selected ENU mutagenesis (TILLING) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4. Transient gain and loss-of-function studies (RNA, DNA and antisense morpholino oligonucleotide injection) . . . . . . . . . . . . . . . 2.5. Transgenic technologies . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.6. Pharmacological screens . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Apoptosis pathways in zebrafish . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Developmental cell death in zebrafish . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1. Cell death during normal development (pro-apoptotic regulators) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2. Cell death caused by loss-of-function mutations (anti-apoptotic regulators) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . p53 and DNA-damage-induced cell death in zebrafish . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Future perspectives: zebrafish as a model for neurodegeneration and cancer . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Abbreviations: AO, acridine orange; BAC, bacterial artificial chromosome; Bak, Bcl-2-antagonist/killer; Bcl-2, B-cell lymphoma 2; BH, Bcl-2 homology domain; Cad, Caspase-activated deoxyribonuclease; Caspase, cysteine-aspartate protease; DISC, death-inducing signaling complex; ENU, N-ethyl-N-nitrosourea; FADD, fas-associated death domain-containing protein; FasL, fas ligand; GFP, green fluorescent protein; Icad, inhibitor of Caspase-activated deoxyribonuclease; Mcl1, myeloid cell leukemia 1; MPNSTs, malignant peripheral nerve sheath tumors; OTR, ovarian TNF receptor; rag2, recombination activating gene 2; RFP, red fluorescent protein; RT-PCR, reverse-transcriptase polymerase chain reaction; TILLING, targeting induced local lesions in genomes; TNF, tumor necrosis factor; TRAIL, TNF-related apoptosis-inducing ligand; TUNEL, terminal deoxynucleotidyl transferase-mediated dUTP nick end labeling ∗

Corresponding author. Tel.: +49 761 5108 495; fax: +49 761 5108 744. E-mail address: [email protected] (M. Hammerschmidt).

1044-579X/$ – see front matter © 2006 Elsevier Ltd. All rights reserved. doi:10.1016/j.semcancer.2006.11.007

U.J. Pyati et al. / Seminars in Cancer Biology 17 (2007) 154–165

1. Introduction Over the past decade, the zebrafish has come to the forefront of modern biological research and has allowed the elucidation of a plethora of fundamental developmental processes. For a vertebrate model system, zebrafish offers the unique combination of being simple to maintain and house while also being very tractable to genetic and cell biological studies. The zebrafish generation time is as in mice, with adults reaching sexual maturity within 3 months of age (Fig. 1). However, fecundity is much higher. Every week, one zebrafish pair can yield hundreds of embryos in a single clutch. Furthermore, embryos and larvae are optically transparent, allowing easy visualization of cellular morphology and movement. Additionally, large-scale forward genetic screens have proven to be extremely fruitful for discovering mutants that reveal novel aspects of developmental morphogenesis and signaling networks. Furthermore, the high degree of homology between the zebrafish genome and that of humans makes such discoveries especially pertinent to human disease and development [1]. In this review, we first give an overview of the methods that make the zebrafish a powerful model system to further dissect the genetic control and to analyze the cellular mechanisms of programmed cell death. We continue with a summary of the current knowledge of the cell death pathways in zebrafish, with special emphasis on its high degree of conservation with mammals. We then describe processes of normal zebrafish development involv-

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ing programmed death. Some of them are conserved among the vertebrate kingdom, while others are more specific for anamniotes. In addition, we give an overview of apoptosis caused by mutations in particular genes or by DNA-damage. Finally, we provide a perspective on the future of this important field of research, with focus on its potential contributions to a better understanding of cancer and degenerative diseases. 2. Zebrafish as a powerful model system to study cell death 2.1. Molecular techniques for in vivo examination of cell death The transparency of the zebrafish embryo makes observing cell death quite easy even in live animals, since tissues become opaque upon dying [2]. Additionally, common molecular methods allow for precise identification of dying cells in vivo, such as TUNEL (terminal deoxnucleotidyl transferase-mediated dUTP nick end labeling) of fragmented DNA [3], staining with the vital dye acridine orange (AO) [4] that becomes fluorescent inside acidic lysosomes enriched in apoptotic, but not necrotic cells [5], immunostaining for activated Caspase-3 [2,6], and trypan blue exclusion stainings to assay cell membrane integrity [7] (see Fig. 2 for sample images). For a detailed description of apoptotic assays in zebrafish, see Negron and Lockshin [2]. Since many embryos or larvae can be processed in parallel, screening for

Fig. 1. The zebrafish life-cycle. Progression of an embryo from the one-cell stage (0 h post-fertilization) to the mid-somitogenesis stage (∼18 h post-fertilization), when the trunk musculature can be clearly seen in the chevron-shaped somites, to the 24 h stage, when most axial patterning (head, trunk, and tail) is complete. Embryos hatch during day 2 of development, from when on they are called larvae, and larvae start to feed at day 5. After 3 weeks of development, some metamorphosis takes place, involving thickening of the skin, development of scales, and sex differentiation, while sexual maturity is reached at 3 months post-fertilization. Then, one breeding pair can spawn hundreds of embryos in 1 week.

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Fig. 2. Examples of dying cells in zebrafish embryos. (A) Fluorescent TUNEL-positive cells in a 30 hpf wild-type embryo after ␥-irradiation (from [18]; copyright 2005, National Academy of Sciences U.S.A.). (B) DAB-stained TUNEL-positive cells in the olfactory organ of a wild-type embryo at 48 hpf (transverse section; from [38]; copyright 2001, with permission from Elsevier). (C) Acridine orange-positive cells in the adenohypophysis of a 28 hpf fgf3 mutant fish (from [96]; reproduced with permission of the Company of Biologists).

mutants in apoptotic pathways is quite feasible. Furthermore, TUNEL-positive cells can be counted to quantify the effects of mutations or other manipulations on cell death. Finally, cell death assays can be combined with other labeling procedures to look for tissue-specific apoptosis during normal development, or after genetic or environmental insult, all of which will be discussed below. 2.2. Forward genetic screens for unbiased gene discovery The transparency of the zebrafish embryo and its quick development (see Fig. 1) allows high-throughput and easy identification of tissue malformations. In the mid-1990s, large-scale three-generation screens for N-ethyl-N-nitrosourea (ENU)induced recessive mutations causing altered embryonic or larval morphology were carried out (Fig. 3), spawning a huge pool of mutants with developmental defects ranging from heart and vascular defects to tail defects to neural degeneration [8,9]. Also, insertional mutagenesis has been established, using modified retroviruses [10]. Although insertional mutagenesis rates are lower than for ENU, they have the advantage that the affected genes can be more easily isolated, as they are tagged by the inserted DNA. Recently, a collection of 315 insertional zebrafish mutants and their corresponding genes have been published [10]. In the case of ENU mutagenesis, which induces the exchange of single nucleotides, mutated genes have to be isolated using positional cloning via meiotic segregation linkage analysis [11]. With near completion of the zebrafish genome, positional cloning sped up considerably, and over the past years, approximately 100 genes affected in the different mutants have been cloned and further characterized (see http://zfin.org). Additionally, the use of transgenic technology (see below) has allowed for the development of screens that isolate mutants in very subtle and dynamic developmental processes such as lateral line development [12], where the tissue of interest is marked with GFP and observed as it develops over time. These unbiased approaches serve as a huge advantage in gene discovery compared to mice or Xenopus laevis, two other premier vertebrate model systems. In Xenopus laevis, they are impossible, due to a recent genome duplication that has occurred in this species. Therefore, people have started to establish the diploid and smaller frog Xenopus tropicalis as an alternative model system [13]. In mouse, large-scale mutagenesis screens

have been initiated recently [14], however, they are extremely space-demanding and costly, and usually unaffordable for a “regular” university institute. 2.3. “Reverse Genetics” via target-selected ENU mutagenesis (TILLING) Recombinant gene knockout techniques have not been developed in zebrafish as yet. However, as an alternative reverse genetics strategy, the targeting induced local lesions in genomes (TILLING) technique has been established. For this purpose, a library of male F1 offspring of ENU-mutagenized fish is generated (individuals are heterozygous for induced mutations). Sperm of these F1 fish is frozen for later establishment of stable lines, while genomic DNA of the fish soma is screened for mutations in given genes via PCR amplification with gene-specific primers, followed by digestion with the mismatchcutting enzyme CelI and/or sequencing [15,16]. So far, this procedure has been successfully employed to discover mutations in over 100 zebrafish genes (Edwin Cuppen, Utrecht, personal communication), two of which have recently been published, one in the recombination activating gene rag1 [17], the other in the tumor suppressor p53 (see more below) [18]. 2.4. Transient gain and loss-of-function studies (RNA, DNA and antisense morpholino oligonucleotide injection) As with Xenopus embryos, zebrafish embryos are very amenable to microinjection, making gain and loss-of-function studies quite easy. Synthetic mRNAs of interest can be microinjected at the one-cell stage, and phenotypes can be monitored subsequently to discover the effects of overexpressing the genes of interest. Additionally, molecular techniques such as in situ hybridization and RT-PCR can be used to examine effects at the gene expression level. Injected mRNA usually starts to be translated shortly after injection. As an alternative to mRNA, plasmid DNA can be injected. Depending on the promoter used, this can lead to later or tissue-specific expression of the inserted cDNA. However, in contrast to RNA, injected DNA does not distribute uniformly throughout the embryo, leading to mosaic situations. While this can be useful to address cell autonomy issues, it usually does not give the full gain-of-function phenotype. For

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Fig. 3. A three-generation recessive screen to identify homozygous mutants in cell survival pathways. Wild-type adult males or isolated spermatids are mutagenized using ENU or retroviral suspensions, and used to fertilize eggs from non-mutagenized females, yielding an F1 generation with individual carriers (heterozygotes) of randomly induced mutations. These F1 fish are crossed to wild-type adults, creating F2 families in which each mutation inherited from the F1 parent is carried by 50% of the siblings. Next, F2 siblings are intercrossed to generate F3 offspring that are used for screening. Statistically, every 4th F2 × F2 pair consists of two carriers of a particular mutation, yielding 25% homozygous fish in the F3 offspring. In this theoretical screen, acridine orange (AO) is used to quickly identify increased or reduced levels of cell death in F3 embryos. Once a mutant is identified, it can be crossed to a polymorphic strain of wild-type fish, followed by the identification of F1 hybrid carriers to generate mutant and wild-type F2 offspring for meiotic mapping. Symbol (+) indicates a wild-type allele, asterisk (*) indicates a mutant allele, males are depicted in brown, and females are depicted in yellow. Green staining in the F3 homozygous mutant embryo depicts an increase in acridine orange-positive cells. For simplicity, only one mutation is shown throughout the figure.

loss-of-function studies, morpholino antisense oligonucleotides have emerged as an efficient tool to knock down given gene products in vivo [19]. Sequences of such chemically stabilized oligonucleotides are selected so that they specifically hybridize with the 5 un-translated region, the translational start site, or splice sites of the targeted mRNA, thereby preventing proper translation or splicing and causing greatly reduced protein levels. By analogy with “mutants”, such morpholino-injected embryos are called “morphants”. Morpholinos have been used in a variety of zebrafish studies to rapidly ascertain the effect of gene lossof-function within a few days, compared with the months that are required to make gene knockouts in mice. Researchers can also co-inject morpholinos targeted to different gene products or inject morpholinos into mutant embryos for combinatorial lossof-function experiments. This is especially useful for apoptotic studies, where multiple factors often intersect to influence cell survival in different contexts. The drawback to these gain-and loss-of-function approaches is that injected RNAs are only stable for less than 1 day, and injected morpholinos for 3 days, exclud-

ing studies of cell death in older larvae, juveniles, or adults. For these later stages, it is essential to have mutant or stable transgenic lines. 2.5. Transgenic technologies Transgenesis in zebrafish has progressed to the point where one can easily create a transgenic fish line within the span of 6 months [20]. Notably, for in vivo marking of particular cell types, green fluorescent protein (GFP) or red fluorescent protein (RFP) have been placed downstream of zebrafish regulatory sequences to drive fluorescent reporter expression in a wide variety of different tissues [21–23]. Furthermore, for temporally controlled overexpression studies, transgenic lines have been generated in which particular cDNAs are under the control of heatshock-sensitive promoter elements. Transgene expression in such transgenic fish can be efficiently activated at any desired stage of development simply by increasing the water temperature (from 28 to 39 ◦ C) (e.g. [24]). To generate trans-

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genic fish, researchers can simply microinject their transgene construct into one-cell stage embryos and grow these injectants to adulthood. Subsequently, the fish can be crossed and – via their offspring – examined for stable genomic integration of the transgene and transmission to the germline. Once again, these studies are greatly facilitated by the transparency of the zebrafish embryo. Transgenic technologies have improved over the years to allow for up to 50% transgenesis rates using the Isce-1 meganuclease [25] or Tol2 transposable element [26] systems. Also, the advent of BAC recombination techniques has permitted researchers to efficiently place reporter genes downstream of uncharacterized promoters so that they can save the laborious steps of isolating regulatory sequences and subcloning reporter constructs prior to transgenesis [27]. The streamlining of these techniques continues, and the use of transgenic zebrafish lines for apoptotic studies should accelerate both the discovery and analysis of tissue-specific apoptotic events. For example, transgenic zebrafish overexpressing Caspase-3 were recently generated, allowing researchers to monitor Caspase3-dependent apoptotic events during both embryogenesis and adult stages [28]. 2.6. Pharmacological screens To develop drug treatments that modulate cellular suicide in humans, it will be essential to have the ability to screen through multiple pharmacological compounds very quickly and at low costs with a clear readout of the compounds’ effectiveness. The zebrafish provides a perfect model for such screens, since many embryos or larvae can be screened and analyzed at once using very small amounts of material [29,30]. Taking this approach, researchers have already begun to discover novel compounds that can rescue faulty vascular network formation [31] or cell cycle defects [32]. In the latter case, 16,320 small molecules (an entire chemical library) were screened within 16 weeks to discover a single compound that specifically reduced the elevated mitosis rate in mutant embryos while leaving that of wild-type embryos unaltered. This elegant screen combined the power of zebrafish genetics with the ease of chemical uptake and phenotypic analysis to find a potential anticancer agent that warrants further testing in mammalian models. By using the fish as a starting point in such a screen, researchers can sift through many compounds without having any bias about which molecules or pathways the chemicals may target. This is analogous to forward genetic screens, where the readout is phenotypic alteration without knowledge of the molecular changes involved. Since the ultimate goal of any therapy is treatment of the phenotype, this strategy is especially valuable. Extrapolating such a technique to cell death models, one can easily imagine the possibilities for both uncovering new cell death pathways and for finding potential treatments of aberrant death programs at the same time [33]. 3. Apoptosis pathways in zebrafish Two recent articles delineated the intrinsic and extrinsic apoptosis pathways in zebrafish [6,34] (see Fig. 4). The

intrinsic pathway, characterized by mitochondrial alterations, cytochrome C release, and Caspase activation, includes both proand anti-apoptotic members of the Bcl-2 family. Pro-apoptotic Bax and Bak directly elicit cytochrome C release into the cytoplasm. In resting cells, the activity of Bax and Bak is blocked by anti-apoptotic members of the Bcl-2 family, such as Bcl-2 and Mcl-1, which contain the BH1-4 domains characteristic of this family. Activation of the intrinsic apoptosis pathway, for instance by stress-induced up-regulation of p53 protein levels (see chapter 4), leads to the activation of BH3-only (containing just the BH3 domain) pro-apoptotic proteins like p53 Upregulated Mediator of Apoptosis (Puma). In the case of Puma, irradiation-induced activation occurs via direct transcriptional activation of the puma gene by p53, as shown both in zebrafish embryos and in cell culture. As a result, the anti-apoptotic Bcl-2 proteins are neutralized, which allows Bak and Bax to oligomerize and to release cytochrome C from the mitochondria. This in turn leads to the activation of Caspases, which proteolyze downstream apoptotic inhibitors such as inhibitor of Caspase-activated deoxyribonuclease (ICAD), allowing others (such as CAD, Caspase-activated deoxyribonuclease) to execute the death program. Many zebrafish members of the pro- and anti-apoptotic intrinsic pathway have been identified and cloned through EST and synteny comparisons with the human and mouse homologs [6,35]. Furthermore, using morpholino antisense technology, zebrafish Mcl1a and Mcl1b (anti-apoptotic Bcl-2 members) were shown to be essential for proper cell viability during zebrafish embryogenesis, while zebrafish Bax1 and Puma (pro-apoptotic Bcl-2 members) are required for ␥irradiation-induced apoptosis [6]. These data indicate substantial functional similarity between zebrafish and mammalian Bcl-2 family members, establishing the zebrafish as a relevant model for studying the intrinsic apoptotic pathway. The mammalian extrinsic apoptosis pathway is triggered by extracellular ligands, such as the Fas ligand (FasL) or the Apo2 ligand/tumor necrosis factor-related apoptosis-inducing ligand (Apo2L/TRAIL), which signal through the Fas receptor or other death receptors to activate a death-inducing signaling complex (DISC), eliciting Caspase-mediated cell death [36]. Zebrafish homologs of FasL and Apo2L/TRAIL, tumor necrosis factor (TNF), and numerous death-domain receptors were recently cloned and shown to cause diverse cell death responses within the embryo [34]. Over expression of three of the Apo2L/TRAIL homologs (termed Death Ligands 1a, 1b, and 2) individually causes shortening of the anteroposterior (head-tail) axis and extensive cell death in the notochord, suggesting that specific cells are competent to respond to these death signals. Such a model is supported by the restricted expression pattern of individual death ligand, receptor, and effector mRNAs in tissues of the early embryo. For example, zebrafish OTR (the closest homolog to a human Apo2/TRAIL receptor) is strongly expressed in the early notochord, as is FADD (Fas-associated death domain-containing protein), a critical adaptor protein that relays the extrinsic death signal to the inside of the cell, while other receptors show specific expression in other embryonic tissues such as pronephric ducts or somitic muscle [34]. Overexpression of death receptors themselves, the FADD adaptor,

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Fig. 4. Intrinsic (left side of cell) and extrinsic (right side of cell) apoptosis pathways in zebrafish. Upon cellular stress, the intrinsic apoptosis pathway can be triggered, beginning with activation of p53, which transcribes genes encoding BH3-only pro-apoptotic proteins such as Noxa and Puma. In resting cells, BH3-only proteins are bound and kept inactive by anti-apoptotic Bcl-2 family members. This blockage is overcome by high levels of BH3-only proteins after cellular stress. The BH3-only proteins induce oligomerization of the pro-apoptotic Bcl-2 family members Bak and Bax (potentially by direct binding), inducing mitochondrial outer membrane permeabilization (MOMP) and subsequent Cytochrome C release. Cytoplasmic Cytochrome C activates the Apaf-1 complex, which converts activator pro-caspases (such as Caspase 9) into their mature form. Activator Caspases cleave at certain sites including aspartate residues in executioner pro-Caspases (such as Caspases-3 and 7), inducing proteolysis of Icad, the inhibitor of Cad. Free Cad moves into the nucleus, where it cleaves DNA. In the extrinsic cell death pathway, Fas ligand (for example) binds its receptor Fas, inducing activation of FaDD and the death-inducing signaling complex. This, in turn, converts the activator pro-Caspase (Caspase 8) to its mature form, which cleaves and activates executioner pro-Caspases. In both types of cell death, the chromatin condenses and DNA fragments, giving two hallmarks of apoptosis. Shown in blue are the assays described in chapter 1.1 that can be used in zebrafish to assess cell death at distinct levels.

or Caspases in early embryos cause widespread death and early lethality, likely because a large excess of any of these transducers of extrinsic death signals can activate the death response in any cell. 4. Developmental cell death in zebrafish 4.1. Cell death during normal development (pro-apoptotic regulators) In contrast to mammals, cell death during zebrafish development does not occur before the mid-blastula transition (MBT), when zygotic transcription begins [2]. At the start of gastrulation (approximately 6 h post-fertilization), the first apoptotic cells become apparent in electron micrographs [37]. During and after this stage, dying cells are engulfed by neighboring cells in a process that depends upon expression of the phosphatidylserine receptor (PSR) [37]. A series of TUNEL experiments performed

on zebrafish embryos between 12 and 96 h post fertilization (early segmentation to larval stages) provided a map of developmental apoptosis (24 hpf map shown in Fig. 5A) [38]. High levels of developmental apoptosis were observed throughout the brain and nervous system as well as the eye, the nose (olfactory placode), the urogenital (excretory) system, and the tail bud. Detailed analyses have revealed that cell death in the ganglion cell layer and inner cell layer of the zebrafish retina resembles that in mammals [39]. Within the excretory system, selected apoptosis of a small number of cells of the proctodeum seems to be required for proper development of the cloaca, the common opening of the gut and kidney for waste excretion [24]. Here, sustained signaling by Bone morphogenetic proteins (Bmp) seems to be essential for the induction of apoptosis. Thus, proctodeal cells remain alive and the cloaca fails to open upon temporally controlled blockage of Bmp signaling, raising the possibility that defective Bmp signaling might also underlie similar human anorectal malformations [24].

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Fig. 5. Zones of cell death in 24 hpf zebrafish embryos. (A) Regions of normal cell death during development. Reference articles for the individual tissues are indicated in superscript. (B) Regions in which developmental cell death is altered in zebrafish mutant, morphant (morpholino-injected), or transgenic embryos at 24 hpf. Reference articles for the individual tissues are indicated in superscript. Note that cell death within the developing skin has also been reported [55], but this was left off the figure for simplicity. (C) Regions of cell death in zebrafish upon toxic insults (e.g. ␥- or UV-irradiation). The most extensive death consistently occurs in the brain and spinal cord, likely because these are the tissues with highest cell death rates during normal development.

Other processes of zebrafish development involving regulated cell death are most likely less conserved between fish and mammals, but might nevertheless serve as similarly good models for human pathologies. Thus, compared to mammals, developing zebrafish remain much longer in a hermaphrodite state (until age of 3 weeks; sexual maturity at 12 weeks). Still, during these first 3 weeks, thousands of early diplotene oocytes are generated that subsequently undergo programmed cell death, particularly during male sex differentiation. Such oocyte death might serve as a model for normal or premature reproductive senescence in mammalian females [40]. Another example is the Rohon-Beard neurons, a population of transient trunk sensory cells. Their function is later taken over by sensory neurons from the dorsal root ganglia, while Rohon-Beard neurons themselves become

apoptotic in a Caspase-9-dependent manner, driven by a combination of Neurotrophin signaling and electric neural activity [41,42]. While Rohon-Beard neurons might specifically exist in teleost and amphibian larvae, the principle of transient neuronal populations is not unique to anamniote vertebrates. In mammals, it is represented by Cajal-Retzius cells, which have an important impact on cerebral cortex development [43]. A third example is mechanosensory neuromast cells of the fish and amphibian lateral line system. Central hair cells of the neuromasts undergo both regular and insult-induced cell death, in which case they are replaced by proliferating mantle supporting cells [44]. Similar mechanisms seem to underlie the regeneration of mechanosensory hair cells in the mammalian inner ear, and might therefore be relevant for a better understanding of hearing loss in humans.

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4.2. Cell death caused by loss-of-function mutations (anti-apoptotic regulators) Myriad genetic networks specify and pattern distinct tissue types in the developing zebrafish embryo. Perturbation of these networks can result in aberrant formation or maintenance of tissues in the developing body, often resulting in cell death. In the various screens over 100 mutants with higher apoptosis rates specifically in the developing central nervous system (CNS; brain, spinal cord and eyes) have been discovered [45–52]. The CNS contains numerous proliferative zones where new neurons are born, and a proper balance between neuronal birth and death is essential for development and functionality of the nervous system [38]. Perturbations of the genetic system regulating this balance might underlie the excessive cell death observed in the different mutants. In addition to brain and spinal cord, there have been reports of loss-of-function experiments leading to altered cell death in such diverse tissues as neural crest [53], pituitary gland [54], notochord [55], lateral line [56], heart [57–59], muscle [60], blood [61–64], excretory system [24,65], germ cells [66–68], and skin [55] (see Fig. 5B). We will briefly review some of this work to give a flavor for cell death studies underway in the zebrafish that might also be relevant for a better understanding of the cellular and molecular basis of a wide spectrum of human diseases, such as craniofacial or cardiovascular malformations, pituitary hypoplasia, blood disorders, muscle degeneration, or infertility. Recently, a zebrafish mutant in the transcription factor Foxd3 was shown to have abnormal neural crest development, resulting in severe malformation of the jaw [53]. It was revealed that much of this defect results from aberrant cell death within the migrating neural crest cells, lessening the number of jaw chondrocyte precursors. A similar death within a population of migrating cells was observed in the small heart mutant, defective in Na+ /K+ ATPase, which develops abnormal cardiac tissue due to excessive death within the migrating cardiac precursor cells [57]. Other studies have focused on the connection between cell differentiation and cell survival. In the developing adenohypophysis, a component of the pituitary gland that generates different hormones regulating basic body functions, loss of the basic helix-loop-helix transcription factor Ascl1a leads to failed cell differentiation coupled with subsequent apoptosis in some cell lineages, while other lineages survive in an undifferentiated state [54]. This is similar to human pituitary syndromes, in which pituitary dysfunction is or is not linked to pituitary hypoplasia. A similar connection of compromised differentiation and enhanced apoptosis is seen in erythrocytes of zebrafish mutants in the Transcriptional Intermediary factor 1␥, leading to dyserythropoiesis similar to human congenital dyserythropoietic anemia [64]. Comparable dyserythropoietic phenotypes are also obtained upon cell-specific mitotic defects associated with erythroid band 3 deficiency [63], while loss of the chaperone Hspa9b leads to compromised mitochondrial function, enhanced oxidative stress, and apoptosis distinctly in blood

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cells, recapitulating the ineffective hematopoesis of the human myelodysplastic syndrome [62]. Another class of ENU-induced zebrafish mutants displays degeneration of somitic muscle cells [69]. One of these mutations disrupts the zebrafish ortholog of the X-linked human Duchenne muscular dystrophy (DMD) gene, which encodes a structural protein required for proper muscle attachment [60]. This demonstrates the suitability of the zebrafish to identify and characterize genes involved in human muscle degeneration, and it will be interesting to identify the molecular nature of the other zebrafish mutations causing similar phenotypes [69]. In addition to transcription factors, structural proteins, and signaling networks, RNA-binding proteins have been shown to be essential for cell survival in specific developmental contexts. In zebrafish, three separate RNA-binding proteins were shown to be important for germ cell survival [66–68]. Mutant and morphant primordial germ cells that lack functional Nanos, Dead End, or Staufen proteins do not move to the genital ridges and fail to differentiate into mature germ cells, thus abrogating their survival, and causing infertility. One interesting variant of apoptosis is anoikis, the death of cells upon detachment from neighboring cells or from the extracellular matrix [70]. Studies in cell culture have suggested that anoikis can result from activation of either the intrinsic or extrinsic cell death pathways [71–75], and that integrin signaling at sites of cell attachment plays a major role in maintaining cell survival [76]. While anoikis has not been specifically analyzed in the zebrafish, it is likely that cell death or developmental defects described in some zebrafish studies are actually a result of anoikis, when tissue integrity and/or integrin signaling is lost. Thus, antisense-mediated loss of Integrin-linked kinase (Ilk) causes widespread patterning defects of the larval zebrafish vasculature, consistent with the requirement of Ilk for cell survival of cultured mouse endothelial cells [77]. Another likely zebrafish model for anoikis are perp morphants, which display severe cell death of skin keratinocytes [55]. The phenotype suggests that the transmembrane protein Perp, although required as a pro-apoptotic factor downstream of p53 for DNA-damageinduced cell death [55], is an essential anti-apoptotic factor to promote survival of keratinocytes during normal development. Normally, basal keratinocytes are firmly attached to the underlying basement membrane, a state in which Integrin signaling might prevent cell death. However, during keratinocyte proliferation or epithelial remodeling, cells have to transiently detach. During this time, Integrin signaling should be diminished, and it is tempting to speculate that Perp might serve to block the anoikis program normally activated in the absence of Integrin signaling. Similarly, metastasizing basal or squamous cancer cells must overcome anoikis to survive and proliferate after their epithelial-mesenchymal transitions and during their migration to new tissue locations. 5. p53 and DNA-damage-induced cell death in zebrafish Upon DNA-damage-inducing insults such as UV- or ␥irradiation, cells utilize a p53-dependent pathway to kill themselves through apoptosis [78] (see Figs. 4 and 5C). The

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p53 pathway has become intensely studied over recent years because mutations in p53 are frequently associated with human cancers, pointing to a tumor-suppressing effect of p53 function, while treatments that restore or elevate p53 function in tumor cells appear as a reasonable approach to block malignant growth in vivo. The tumor-suppressing effect of p53 seems to be two-fold, blocking cell proliferation, while also promoting cell death. While analyses of the role of p53 in mediating cell death have been primarily performed in cell culture systems, the zebrafish offers the possibility to study this cell death pathway in vivo. By using the TILLING reverse genetic approach, Berghmans et al. were able to isolate p53 mutant zebrafish [18]. As in mouse, p53 is not required for normal early zebrafish development. However, gain of p53 function either by p53 mRNA injection [55], or by morpholino-based knock-down of its main negative regulator, the p53-specific ubiquitin ligase Mdm2 [33], leads to widespread embryonic apoptosis. Also, p53 is absolutely required for developmental cell death that occurs upon genetic or environmental insult, such as irradiation-induced apoptosis in developing zebrafish [18,33,55]. Here, p53 most likely acts by activating components of the intrinsic cell death pathway like Puma and Bax1 [6] (see above and Fig. 4). Similarly, zebrafish p53 is essential for the elimination of cells with stalled or compromised DNA replication during S-phase [79], and for the elimination of cells with increased DNA-damage due to the loss of the fancd2 gene, which is mutated in human patients suffering from Fanconia anemia [80]. In contrast to early development, loss of p53 function does affect normal cell behavior during adulthood. Thus, adult homozygous p53 zebrafish mutants display an increased predisposition for malignant peripheral nerve sheath tumor (MPNST) formation, revealing an essential tumor suppressor function of p53 in zebrafish, similar to its role in humans. The MPNSTs of p53 mutant zebrafish likely result from aberrant regulation of cell proliferation in the peripheral nervous system, modeling similar human pathologies. Since human tumors often display coupled mutations in p53 and other genes [81], it will be interesting to use the zebrafish system to systematically identify important p53 partners in genetic enhancer screens, searching for mutations that elevate the incidence or aggressiveness of MPNSTs in a p53 mutant background. A similar enhancement of malignancy has been observed when p53 deficiency in mouse lymphoid cells was combined with overexpression of the proto-oncogene Myc [82,83], leading to much higher proliferation and greatly reduced apoptosis rates. Along the same line, transgenic zebrafish with elevated Myc expression in given cell types could be used to screen for mutations that enhance or suppress particular malignancies such as T-cell leukemia [84] (see below) or neuroblastoma. 6. Future perspectives: zebrafish as a model for neurodegeneration and cancer In addition to fostering a basic understanding of the biology behind cell death, it will be important to utilize model systems such as zebrafish for finding new approaches to either reversing

cell death in the case of degenerative diseases such as muscle dystrophies or neurodegeneration, or inducing cell death in the case of cancer. In the aging nervous system, excessive cell death is a major cause of human illnesses such as Parkinson’s disease and Alzheimer’s disease. In the latter disease, cell death is mediated by the gamma-secretase complex, which catalyzes the final cleavage of amyloid precursor protein to generate the toxic amyloid protein [85,86]. Mutations in the Presenilin enhancer-2 (Pen-2) gene are commonly associated with neurodegeneration in human Alzheimer’s disease patients, so understanding the functions of wild-type Pen-2 will be important for analyzing the biology of neuronal cell death. In two separate studies, zebrafish Pen-2 was shown to be critical for normal nervous system development, modeling neurodegeneration in humans [87,88]. Pen-2 morphant fish have excessive neuronal cell death, which is a p53-dependent phenotype, modeling symptoms of Alzheimer’s disease patients. People have also started to use the zebrafish to study Parkinson’s disease, where degeneration of given dopaminergic neurons leads to deregulated trembling. Treatment of zebrafish with dopaminergic neurotoxins such as MPTP, rotenone, and paraquat, in addition to causing developmental defects in treated larvae, induces movement alterations in both larval and adult fish, modeling the symptoms of human patients [89,90]. Utilizing such models, genetic screens can be designed to uncover suppressors of neurodegeneration, identifying promising targets for therapeutics to treat human neurodegenerative diseases. Furthermore, pharmacological screens as described above (see Section 2.6) could be applied for drug discovery. Elevated cell death as during degenerative diseases is usually caused by loss-of-function mutations in anti-apoptotic genes or gain-of-function mutations in pro-apoptotic genes. In contrast, human malignancies are due to loss-of-function mutations in genes that in addition to blocking cell proliferation normally promote apoptosis (tumor suppressors), or due to gain-of-function mutations in genes that in addition to promoting cell proliferation impair the cell death pathways (oncogenes) [91–94]. In these cases, cancerous cells are often rendered largely insensitive to death-inducing stimuli such as irradiation or chemotherapy. The ease and quickness of creating compound loss-of-function or gain-of-function scenarios in the zebrafish should make it a good system for manipulating tumor tissue and examining the efficacy of potential therapies. An example of tumor formation upon oncogene overexpression in zebrafish comes from a Myc transgenic zebrafish line, in which the Myc oncogene is highly and specifically expressed in T-lymphocytes under the control of the rag2 promoter [84]. Transgenic fish display massive T-cell over-proliferation and metastasis into other tissues, thus creating a valuable model for human T-cell leukemia. At present, the switches between proliferation and apoptosis downstream of Myc, and the exact mechanisms of the anti-apoptotic effects of Myc, remain largely mysterious. Thus, it will be crucial to find ways of enhancing apoptosis through by-passing those branches of the cell death pathways that are compromised by Myc overexpression. The Myc-overexpressing transgenic zebrafish might turn out as very helpful for this purpose. An initial study to prove the value of this system utilized a rag2-bcl-2

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transgenic line to over-express the anti-apoptotic Bcl-2 protein in T-lymphocytes, which led to a complete block of the intrinsic apoptotic pathway in T-cells after ␥-irradiation [61]. When this apoptosis-defective line was crossed to the Myc-overexpressing line, T-cell tumorigenesis was greatly enhanced, reinforcing the notion that pro-apoptotic mechanisms are at play even in Mycinduced tumors to attenuate cancer-causing potentials. In this light, systematic genetic screens as described above (see Section 2.2), but now in the Myc transgenic line, should enable researchers to identify novel suppressor or enhancers of the Myc phenotype. Similarly, small compound screens (see Section 2.6) should lead to the discovery of drugs mimicking or healing particular cancer types. 7. Conclusions As we have tried to emphasize in this review, the zebrafish is just being fully recognized as a good in vivo system to study cell death during development and after DNA-damage or malignant transformations. Remarkably, the zebrafish truly evolved as a powerful developmental model system only about 15 years ago, meaning that we have witnessed an exponential growth in the strength of the zebrafish system. According to the Zebrafish Information Network, there are now 472 zebrafish groups worldwide [95], with more and more medical doctors discovering the zebrafish as a complementary system for basic research on questions related to human diseases. Thus, expertise within the field is constantly growing beyond the initial purely academic interest in developmental biology, and opportunities for collaboration and development of new technologies are plentiful. There is no reason why the next decade should not similarly witness the increased use of the zebrafish for discovering, analyzing, and manipulating novel molecules that are critical for cell death pathways in all vertebrate organisms, including humans. Fishing out new cancer therapies should not be far behind. Acknowledgements U.J.P. is supported by the National Institutes of Health Program Training Grant in Molecular Hematology (5T32 HL07623-20), A.T.L. is supported in part by National Institutes of Health Grant CA119066-01, and M.H. in part by National Institutes of Health Grant 2R01-GM063904 and by the MaxPlanck Society. References [1] Postlethwait JH, Woods IG, Ngo-Hazelett P, Yan YL, Kelly PD, Chu F, et al. Zebrafish comparative genomics and the origins of vertebrate chromosomes. Genome Res 2000;10(12):1890–902. [2] Negron JF, Lockshin RA. Activation of apoptosis and caspase-3 in zebrafish early gastrulae. Dev Dyn 2004;231(1):161–70. [3] Yamashita M. Apoptosis in zebrafish development. Comp Biochem Physiol B Biochem Mol Biol 2003;136(4):731–42. [4] Ton C, Lin Y, Willett C. Zebrafish as a model for developmental neurotoxicity testing. Birth Defects Res A Clin Mol Teratol 2006;76(7):553–67. [5] Abrams JM, White K, Fessler LI, Steller H. Programmed cell death during Drosophila embryogenesis. Development 1993;117(1):29–43.

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