From oogenesis through gastrulation: developmental regulation of apoptosis

Seminars in Cell & Developmental Biology 16 (2005) 215–224

Review

From oogenesis through gastrulation: developmental regulation of apoptosis Jessica Greenwood a,b , Jean Gautier b,∗ a

b

Integrated Program in Cellular, Molecular, and Biophysical Studies, Columbia University College of Physicians and Surgeons, 701 W. 168th Street, NY 10032, New York, USA Department of Genetics and Development, Hammer Health Sciences Center Room 1620, Columbia University College of Physicians and Surgeons, 701 W. 168th Street, New York, NY 10032, USA Available online 1 February 2005

Abstract Apoptosis is a mechanism employed by multicellular organisms throughout development as a means of eliminating damaged or otherwise unwanted cells. From oogenesis through fertilization and gastrulation, organisms use an array of cell- and tissue-specific mechanisms to regulate the apoptotic program in response to stress or developmental cues. Since cell death regulation is tightly interwoven with cell cycle and checkpoint controls, and embryos of the fly, fish and frog exhibit unique embryonic cell cycle regulation, it is of great interest to understand how early embryos coordinate these cellular functions. © 2004 Elsevier Ltd. All rights reserved. Keywords: Apoptosis; Programmed cell death; Cell cycle; Checkpoints; Development

Contents Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.1. Molecular components of apoptosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2. Developmental PCD . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1. Apoptosis in the germline. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1.1. C. elegans . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1.2. Drosophila . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1.3. MAPK pathway and germline apoptosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2. Developmental PCD following fertilization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3. Induced cell death. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1. Resistance of embryos to induced apoptosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1.1. Drosophila . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1.2. Vertebrates . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2. Connecting DNA damage, cell cycle checkpoints and apoptosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2.1. DNA damage response . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2.2. Cell cycle regulation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4. Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.

∗

Corresponding author. Tel.: +1 212 305 9573 (Lab)/305 9586 (O); fax: +1 212 923 2090. E-mail address: [email protected] (J. Gautier).

1084-9521/$ – see front matter © 2004 Elsevier Ltd. All rights reserved. doi:10.1016/j.semcdb.2004.12.002

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1. Introduction Throughout development, apoptosis is employed by multicellular organisms to eliminate damaged, unwanted or unneeded cells. While the effectors of cell death remain generally consistent from cell type to cell type and are evolutionarily conserved, the upstream activators of the cell death machinery may vary depending on many factors including, but not limited to, stage of differentiation, cell cycle phase, and the type of death-inducing stimulus. Here, we will discuss both developmental programmed cell death (PCD), defined as apoptosis observed during the development of a wildtype organism under normal conditions, and ectopic or stressinduced apoptosis. From oogenesis through fertilization and gastrulation, embryos use various mechanisms to regulate both the developmental and the induced apoptotic program. This review, while not comprehensive in its coverage, aims to provide examples of stage-specific mechanisms used by different organisms to regulate cell death. In addition, the field is just beginning to flesh out the molecular connections between cell death regulation and cell cycle control during embryogenesis. We will highlight some of the recent work in this exciting area. 1.1. Molecular components of apoptosis The evolutionarily conserved core components of the cell death machinery have been well characterized through molecular genetics and biochemical analyses. Pioneering studies in the nematode, Caenorhabditis elegans, took advantage of the highly reproducible pattern of PCD in the developing hermaphrodite to identify ced-3 and ced-4 as necessary effectors of somatic cell death [1,2]. CED-3, the C. elegans caspase, is present as an inactive zymogen prior to the initiation of the cell death program. Proteolytic activation of proCED-3 is mediated by its interaction with CED-4. CED-9, an essential negative regulator of the cell death pathway and homolog of anti-apoptotic Bcl-2 in vertebrates, is localized to the mitochondria where it sequesters CED-4 preventing its association with CED-3 [3]. CED-9 is itself inhibited by EGL-1, a pro-apoptotic BH3-domain-containing protein that is necessary for all somatic cell death in C. elegans [2]. While these core components are conserved in higher eukaryotes, additional regulatory controls have been added to the apoptotic pathway throughout evolution [4]. Two separate pathways, the intrinsic and the extrinsic, mediate the initiation of a caspase cascade, where activation of the apical caspase results in subsequent cleavage and activation of downstream pro-caspases. The extrinsic pathway is initiated in response to death receptor binding and results in the activation of apical caspase-8 and dredd in vertebrates and Drosophila, respectively. This pathway is of particular importance in the regulation of apoptosis in the immune system in mammals [5–7]. In contrast, the intrinsic pathway is activated in response to stress and developmental cues. Cytochrome c release from mitochondria, a characteristic feature of the in-

trinsic pathway in vertebrates, is essential for the formation of the apoptosome, a large multimeric complex consisting of apical caspase-9, Apaf-1, which is the mammalian CED-4 homolog, and cytochrome c itself. The release of cytochrome c from mitochondria is controlled by pro- and anti-apoptotic Bcl-2 family members [8]. In Drosophila, the apical caspase, Dronc, is activated by the Apaf-1 homolog, Dark, however, the role of cytochrome c in Drosophila cell death is unclear, as it does not seem to be released from the mitochondria, but undergoes a detectable conformational change in apoptotic cells and is capable of binding Dark [9]. Although initiation of the caspase cascade is an important regulatory step in the control of apoptosis, the regulation of caspase activity at a step downstream of cytochrome c release is also critical. In flies and vertebrates, this regulation is characterized by the antagonistic interactions between inhibitor of apoptosis proteins (IAPs) and pro-apoptotic proteins for which reaper (rpr) is the prototype [10,11]. IAP family members interact directly with and inhibit processed forms of cellular caspases. This inhibition is relieved through binding of IAPs to upstream pro-apoptotic activators, known collectively as IAP antagonists. In a screen for genes required for apoptosis in Drosophila, the H99 chromosomal region was found to be necessary for nearly all PCD in the developing fly [12]. Three genes, rpr, hid (head involution defective) and grim are located within this region and each encodes a protein that contains an N-terminal RHG motif, which mediates the IAP/antagonist interaction. In mammals, IAPs are commonly upregulated in tumors, thus promoting the enhanced survival of cancer cells [10]. IAP antagonists, Smac/DIABLO and Omi/HtrA2, have also been identified in mammals [13–16]. Under non-apoptotic conditions, these proteins localize to the intermembrane space of the mitochondria; in response to apoptotic stimuli, they are released into the cytosol, where they promote cell death, at least in part, through the inhibition of IAPs [13,14].

2. Developmental PCD 2.1. Apoptosis in the germline In this section we will discuss developmental cell death in the germline of invertebrates and amphibians. It is worth noting that extensive investigation into the regulation and role of oocyte apoptosis in mammals has been conducted. These have been reviewed elsewhere and will not be addressed here [17,18]. 2.1.1. C. elegans In the nematode, C. elegans, 1090 somatic cells are generated during the development of the hermaphrodite, of which, 131 undergo developmental programmed cell death in a highly reproducible manner [19–21]. In addition to somatic cell death, approximately half of the female germ cells un-

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dergo apoptosis. Interestingly, syncitial germ cell apoptosis occurs only during oogenesis, specifically as germ cells are about to exit the pachytene stage of meiotic prophase I. During this process, germ cell nuclei undergoing apoptosis are quickly cellularized away from the syncitium, most likely to sequester apoptotic factors from acting on the remaining nuclei [22]. Genetic analysis revealed that the machinery involved in the execution of germ cell death is similar to that in somatic cells; ced-3 or ced-4 loss-of-function(lf) mutations prevent germ cell death, and ced-9(lf) mutants exhibit increased levels of germ cell death. Genes involved in the phagocytosis of somatic cell corpses, such as ced-1,-2,5,-6,-7, and ced-10 are also implicated in germ cell corpse phagocytosis. However, in contrast to somatic cells, where a ced-9 gain-of-function(gf) mutation completely blocks cell death, a ced-9(gf) mutation does not prevent physiological germ cell death. In addition, egl-1, which is required for all somatic cell death [23], has no role in physiological germ cell apoptosis [22]. These findings provide a clear example of how cells in different stages of development within an organism may use different regulatory mechanisms to control programmed cell death. Furthermore, this raises the question of what makes meiotic and mitotic cells respond differently with respect to apoptosis. To address this issue specifically, (lf) mutants of the Ras/MAPK pathway were examined for a change in the levels or timing of germline apoptosis [22]. The germ cells of Ras/MAPK mutants fail to exit pachytene arrest and also do not undergo PCD. It is not yet clear whether the MAPK pathway plays a direct role in regulating apoptosis, or whether its role is indirect, by regulating the differentiation of oocytes. We will discuss in Section 2.1.3 the possible direct role of the MAPK pathway in meiosis-specific apoptosis. 2.1.2. Drosophila During Drosophila oogenesis, apoptosis is used for two distinct purposes: (1) to eliminate defective egg chambers at early and mid-oogenesis and (2) to resorb every nurse cell in late oogenesis [18,24]. Egg chambers consist of somatic follicle cells, 15 germline-derived nurse cells, and one oocyte. Nurse cells support the growth of the oocyte by synthesizing and depositing maternal factors into the oocyte through intercellular bridges called ring canals. At stage 11 of egg chamber development, nurse cells rapidly transfer the majority of their cellular contents into the oocyte through a process called ‘dumping’. As dumping occurs, each nurse cell undergoes apoptosis. Since apoptotic nurse cells remain connected to the oocyte, a mechanism must be present to protect the oocyte from the active cell death machinery and to prevent complete cytoskeletal destruction. Using an antibody that recognizes the activated form of an effector caspase, Drice, Peterson et al. observed that active Drice is present in a punctate pattern surrounding nurse cell nuclei [25]. The aggregation of Drice may help to keep its activity localized, thus protecting the cytoskeleton as well as the connected oocyte. Therefore, organisms have developed different strategies to prevent the spreading of a death signal throughout a syncitium by (1)

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physically isolating dying nuclei by cellularization in C. elegans or (2) concentrating the apoptotic signal via the local activation of caspases in Drosophila. Interestingly, unlike most somatic cell death, nurse cell death is not dependent on rpr, hid or grim, as an H99 deletion does not prevent nurse cell death [26]. In addition Diap1 overexpression does not inhibit nurse cell death, whereas Diap1 overexpression blocks apoptosis in different stages of oogenesis and somatic cells [25]. At an earlier stage in oogenesis (stages 7–8), defective egg chambers are eliminated in what is thought of as a midoogenesis checkpoint. Apoptosis of entire egg chambers has been observed due to defects such as nutrient deprivation, chemical treatment, or the inability to synthesize or respond to the steroid hormone ecdysone [24]. Genetic analyses have highlighted some of the molecular features of apoptosis in mid-oogenesis; while there are similarities to other phases of developmental apoptosis, there are some differences as well. First, a mutation in the effector caspase gene, dcp-1 prevents germline cell death during mid-oogeneis, but has no effect on nurse cell death during late oogenesis [27]. In fact, ectopic expression of Dcp-1 in mature egg chambers inhibits the cytoskeletal alterations and nurse cell dumping normally seen in dying nurse cells [25]. The caspase Drice is also active during mid-oogenesis in dying egg chambers, but whereas its activity is tightly localized in subcellular aggregates in late oogenesis, Drice actvity is more robust and found ubiquitously in dying stage 8 egg chambers. In further contrast to nurse cell apoptosis during late oogenesis, apoptosis at this stage is responsive to levels of Diap1, as diap1(lf) mutants exhibit increased egg chamber degeneration at midoogenesis [28]. Differences aside, apoptosis during both midand late-oogenesis, is rpr-, hid-, and grim-independent, thus demonstrating the unique regulatory characteristics of germ cell apoptosis in Drosophila. 2.1.3. MAPK pathway and germline apoptosis Meiotic-specific signaling pathways may play a role in the regulation of germ cell death and may explain, in part, its unique characteristics (Fig. 1). Drosophila Mos (dmos), an upstream component of the MAPK pathway and regulator of meiosis in vertebrates is not essential for meiosis or development in Drosophila but dmos mutant females have a lower fertility rate. Compared to wild-type, ovaries from dmos mutant females have more apoptotic egg chambers at stages 7–8, suggesting that Dmos normally promotes survival at mid-oogenesis [29]. In starfish oocytes, MAPK signaling also plays a role in regulating apoptosis. Following hormone treatment with 1-methyladenine (1-MA), starfish oocytes undergo germinal vesicle breakdown (GVBD) followed by apoptosis, 8–12 h later, if fertilization does not occur. In the current model, Mos/MEK/MAPK(ERK) signaling, which is active at GVBD, promotes survival, whereas p38 MAPK, which becomes active at the time of cell death, promotes apoptosis [30]. In addition, cell-free extracts derived from Xenopus

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it is unclear whether MAPK promotes survival via a similar mechanism in organisms other than fly.

2.2. Developmental PCD following fertilization

Fig. 1. Mos/MAPK pathway as a specific regulator of germ cell death. The Mos/MAPK pathway regulates meiosis completion. MAPK family signaling in different cellular contexts has been shown to promote survival or apoptosis. In oocytes of Drosophila, starfish, and Xenopus, the Mos/MAPK pathway seems to prevent apoptosis, whereas in starfish, p38 MAPK may act to promote cell death. MAPK signaling is one of the regulatory controls that may give germ cell apoptosis some of its unique characteristics.

eggs exhibit several hallmarks of apoptosis, including caspase activation, following prolonged incubation [31,32]. Extracts arrested in M-phase (CSF extracts) are more resistant to caspase activation than interphase extracts made from the same eggs, but both will eventually activate the apoptotic machinery [33]. The apoptotic resistance in M-phase extracts is mediated downstream of cytochrome c release and is thought to be dependent on MAPK signaling. Because transcription and protein synthesis are absent in these egg extracts, it is likely that post-translational modifications of target proteins are responsible for the apoptotic resistance observed. However, no protein target has been identified and it is not yet clear how MAPK signaling communicates with the cell death machinery. In Drosophila embryos, the Ras/MAPK pathway is involved in the transcriptional downregulation of the hid gene, and in the inhibitory phosphorylation of the Hid protein [34,35]. As no hid homolog has been identified in vertebrates,

Following fertilization, embryonic cells of the fly, frog and fish begin to divide rapidly. Cell cycles during this stage of development consist of only S- and M-phases and are driven by maternally stored products; zygotic transcription is absent in the early embryo. Transition from maternal to zygotic control is achieved at the maternal/zygotic transition (MZT). In frog, zygotic transcription is initiated at the mid-blastula transition (MBT) [36,37]. At this stage, gap phases are introduced into embryonic cell cycles, cell division becomes asynchronous, and cell motility is first observed (Fig. 2) [36,38]. By the onset of gastrulation, maternal cell cycle regulators have been removed and zygotic control over cell cycles begins [38,39]. The regulation of the onset of both the MBT and the EGT (early gastrulation transition) has been attributed to developmental timers [40]. The timing of the MBT is regulated by the nucleus-to-cytoplasm volume ratio [37], however, the mechanism controlling the EGT has not been identified. Interestingly, no developmental PCD has been reported in C. elegans, fly, zebrafish or frog in the period between fertilization and gastrulation (Fig. 2). As we will discuss in the next section, apoptosis cannot be induced prior to gastrulation in the frog or the fish, despite the fact that the apoptotic machinery is present. The reason apoptosis is suppressed during embryonic cleavage is unknown. This lack of apoptosis is counter-intuitive as the rapid, unchecked, embryonic cell cycles are more likely to yield damaged genomes, normally a potent apoptotic trigger. Two distinct features of these embryos could account for this situation. First, these embryos

Fig. 2. Acquisition of the ability to undergo stress-induced apoptosis is developmentally regulated. Early embryos of Drosophila and Xenopus are refractory to induced apoptosis. In Drosophila, several pathways could be involved in early apoptosis since the onset and the regulation of apoptosis varies with the type of stress used to trigger apoptosis. In Xenopus, all stress-induced apoptosis initiates at the early gastrulation transition (EGT), following the activation of zygotic transcription, the remodeling of the cell cycle and the acquisition of damage checkpoints. MBT: Midblastula transition.

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rely on maternally stockpiled proteins and mRNAs for all cellular functions. In the absence of transcription, there is no selective pressure to keep the genome (the template for transcription) intact or to eliminate damaged genomes. Second, the rapidly dividing cells of the early embryo lack checkpoints that normally monitor genomic integrity. With the introduction of checkpoints, we see the initiation of cell death, suggesting that there may be communication between the cell cycle machinery, the checkpoint machinery and the cell death machinery. These relationships will be discussed in Section 3.2.

3. Induced cell death 3.1. Resistance of embryos to induced apoptosis 3.1.1. Drosophila In addition to developmental cell death, organisms may use apoptosis as a mechanism to protect against irreparable environmental damage incurred during development or as an adult. In Drosophila, developmental cell death can be detected under normal conditions, at approximately 7 h after egglaying (ael), or stage 11, at which point, the embryo has already undergone gastrulation and has reached the fully extended germ band stage (Fig. 2) [41]. By irradiating 3–4 h old embryos with high doses of gamma-irradiation, apoptotic cells appear 2 h earlier than the onset of PCD in non-irradiated embryos. Molecular genetic characterization of the pathway responsible for the induction of cell death implicates the H99 chromosomal region, containing rpr, hid and grim, as necessary for the majority of irradiation-induced apoptosis [12]. In addition, induction of dark, the Drosophila Apaf-1 homolog, is necessary for irradiation-induced apoptosis in the early embryo [28]. Interestingly, UV-irradiation induces apoptosis in the Drosophila embryo earlier in development than gammairradiation and is developmentally regulated [42]. When exposed to UVB or UVC, but not UVA, cells in the early embryo (0–3 h ael) undergo apoptosis prior to gastrulation. dark expression is induced and this upregulation is necessary for the activation of apoptosis in early embryos. In contrast, rpr expression does not change in the early embryo following UV irradiation. Conversely, no induction of dark is observed in TUNEL-positive cells of older embryos (5–10 h ael) upon exposure to UVC, whereas rpr is induced in these cells following UVC but not UVB. Finally, later embryos (12 h + ael) exhibit resistance to UV-induced apoptosis, as no induction of dark, rpr or TUNEL staining was observed with high doses of UV. Ectopic cell death is also observed in embryos with a diap1(lf) mutation. An embryo with a homozygous null mutation in diap1 progresses normally through gastrulation, but at stage 7, development is arrested and embryonic cells undergo massive apoptosis 90 min later [43,44]. However, a

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dark/diap1 double mutant does not arrest development and does not exhibit increased cell death at stage7, demonstrating that dark is necessary for the activation of caspases in the ectopic cell killing seen in diap1 mutants [28]. In mammals, deletion of individual IAP family members does not lead to increased cell death in the developing organism, suggesting that during evolution, even more restrictive regulatory requirements were imposed on the apoptotic pathway [45]. Finally, Drosophila Bcl-2 family members also play a role in both developmental and induced apoptosis [46]. Overexpression of a pro-survival Bcl-2 homolog, buffy, in early embryos inhibits developmental PCD [47]. debcl, a proapoptotic Bcl-2 family member is expressed in regions of cell death during development [48]. Overexpression of Debcl leads to increased PCD throughout development. Genetic interaction experiments have placed the Bcl-2 family members downstream or in a parallel pathway to RHG proteins and IAPs and upstream of the apical caspase, Dronc [47]. How the embryo integrates each of these pathways into a specific developmental context remains unclear. 3.1.2. Vertebrates In zebrafish and Xenopus, there exists a general resistance to apoptosis prior to gastrulation. The ability to undergo apoptosis coincides with the introduction of checkpoints in the early embryo (Fig. 2). A frog or fish embryo that has been treated with a threshold death-inducing insult prior to the MBT will undergo rapid and synchronous cell death at the onset of gastrulation (EGT). This response is triggered by gamma-irradiation [49,50], camptothecin [51], both of which induce DNA damage, nocodazole [52], which inhibits microtubule polymerization, hydroxyurea and aphidicolin [50,51,53], which block DNA replication, ␣amanitin [50,54], which inhibits transcription and cycloheximide [50,51,54], which blocks protein synthesis. Inhibitors of transcription or translation can induce apoptosis, suggesting that cell death in the early embryo is regulated by maternal protein components. Conserved effector caspases have been implicated in this cell death, as injection of peptide inhibitors of caspases significantly delay the onset of apoptosis [50,51]. In addition, regulators acting upstream must be conserved, as the overexpression of Bcl-2 delays apoptotic initiation [50]. Finally, overexpression of a constitutively active form of Akt, a protein that promotes survival in mammalian cells, in irradiated Xenopus embryos delays the onset of apoptosis [55]. The exact nature of apoptotic resistance in Xenopus embryos is unknown at this time. Interestingly, while overexpression or inhibition of some proteins can delay the onset of apoptosis (beyond the EGT), there are no reports of apoptosis induced prior to the EGT. Overexpression of downstream effectors, such as active caspase-3, or overexpression of upstream regulators is predicted to induce at least some of the hallmarks of apoptosis, however, no such reports have been made. Since cytochrome c release is necessary for the activation of caspase-9, it would be interesting to determine if cytochrome c release takes place prior to the EGT and if this

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step is rate limiting. It is likely that both cytochrome c release, mediated by pro-apoptotic Bcl-2 family members, and caspase inhibition relief, mediated by the rpr/IAP pathway, need to take place in parallel to in order to fully activate the cell death program at the EGT. To date, no rpr, Smac or Omi homologs have been identified in Xenopus. As mentioned above, despite the absence of apoptosis in early embryos, functional apoptotic machinery is present in Xenopus embryos. Indeed, cell-free systems derived from Xenopus unfertilized eggs have been instrumental in the study of the biochemical regulation of apoptosis [31,32]. These extracts can be fractionated into a heavy membrane fraction, containing mitochondria, and a cytoplasmic fraction. When the two fractions are mixed, cytochrome c is released and caspases are activated. Alternatively, prolonged incubation of non-apoptotic extracts will eventually lead to caspase activation. These extracts have helped to gain insight into the function of Reaper in vertebrates [56]. Addition of recombinant Rpr protein to Xenopus extracts induces caspase activity in a mitochondria-dependent manner, demonstrating that the pro-apoptotic function of Rpr is at least partially conserved. In a biochemical screen for Xenopus proteins that interact with Rpr, Scythe was identified. When Scythe is removed from extract, Rpr-induced caspase activity is prevented, suggesting that Scythe is a pro-apoptotic factor [57]. However, incubation of extracts with excess Scythe suppresses Rprinduced caspase activation. These seemingly contradictory results can be resolved with a model in which a pro-apoptotic factor, X, is sequestered by Scythe; Rpr binding then releases factor X [58]. Scythe homologs have been identified in both Drosophila and human; it will be interesting to see whether Scythe interacts with Rpr orthologs, Smac and Omi in mammalian cells. Other types of cell-free extracts can be made from Xenopus eggs and embryos to study the regulation of apoptosis. For example, apoptotic extract can be generated from gastrulation stage embryos treated with DNA-damaging agents prior to the MBT. As these extracts activate cell death in vivo, they are particularly useful for studying the downstream events of apoptosis [50] (Greenwood and Gautier, unpublished data). 3.2. Connecting DNA damage, cell cycle checkpoints and apoptosis 3.2.1. DNA damage response Before activating the apoptotic program following ectopic damage, a cell must first recognize the damage, and if it is irreparable, a signal must be sent to the apoptotic machinery. Following DNA damage, cells rely on checkpoints to arrest the cell cycle, allowing DNA repair or apoptosis to take place to avoid the propagation of a compromised genome. The ability of cells to respond to damage, as well as their decision to repair, arrest, or die changes throughout development and evolution. DNA damage-induced apoptosis does not take place in somatic cells of the developing C. elegans hermaphrodite.

However, meiotic germ cells activate the apoptotic program following gamma irradiation [59]. There is an approximate 10-fold increase in the number of germ cell corpses following treatment of L4 hermaphrodites with ionizing radiation. Irradiation-induced apoptosis in germ cells is restricted to the pachytene stage of oogenesis and is dependent on ced3 and ced-4. However, in contrast to physiological egl-1independent germ cell death, irradiation-induced apoptosis is severely compromised in an egl-1(lf) mutant. Also, a ced9(gf) mutation does not block physiological germ cell death, but does prevent irradiation-induced apoptosis, suggesting that even within the same cell type, different mechanisms are used to control the decision to die. In addition to apoptosis, ionizing radiation induces cell cycle arrest in C. elegans mitotic germ cells and sensitizes progeny to lethality [59]. As these cellular mechanisms can generally be attributed to checkpoint proteins, it is reasonable to investigate the role of conserved checkpoint factors in irradiation-induced germ cell apoptosis. In mammalian somatic cells, the tumor suppressor, p53, plays a critical role in maintaining genomic integrity by arresting cell cycle progression and/or activating apoptosis in the presence of DNA damage (Fig. 3). p53 directly participates in the decision to undergo cell cycle arrest, through transcriptional activation of genes of cell cycle regulators, such as p21Cip1 , or to undergo apoptosis by induction of death genes, such as Bax. This is in contrast to invertebrates where p53 functions downstream of the branching point of this decision. The C. elegans p53 homolog, cep-1, is essential for irradiation-induced apoptosis in the germline, whereas a cep-1(lf) mutant undergoes normal physiological germ cell death and somatic cell death [60,61]. Interestingly, as opposed to mammalian p53, cep-1 is not essential for irradiation-induced cell cycle arrest. Overexpression of wild-type cep-1 in somatic cells leads to an increased number of corpses, but cell cycle arrest is not detected. These results are consistent with studies conducted in Drosophila, where dmp53 was found to be important for damage-induced apoptosis, but not damage-induced cell cycle arrest (Fig. 3) [62,63]. Interestingly, a direct link between the p53 checkpoint and apoptosis has been demonstrated, as p53 can activate transcription of the rpr gene [64]. Some conserved checkpoint genes, however, have been identified that are essential for both cell cycle arrest and apoptosis in the invertebrate germline following irradiation. Specifically, hus-1, rad-5 and mrt2 mutants exhibit normal developmental cell death, but are defective in radiationinduced germ cell death and proliferation arrest [59,65,66]. In Schizosaccharomyces pombe, Rad-1 (mrt-2 homolog) and Hus-1 have been implicated in the DNA damage response and are involved in telomere length maintenance. Genetic experiments in C. elegans have placed hus-1 and mrt-2 together in a pathway parallel to rad-5. A clue to understanding the link between the checkpoint machinery and apoptosis came with the discovery that ionizing radiation induces egl-1 transcription in a hus-1 and cep-1 dependent manner [65]. Since cep-1 is only involved in the activation of apoptosis in re-

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Fig. 3. Coordination of cell cycle arrest and apoptosis. Following DNA damage, cells or embryos activate a coordinated response leading to cell cycle arrest or apoptosis. p53 is at the branching point of this decision in mammalian cells and plays an important regulatory role in both cell death and apoptosis. In contrast, p53 in invertebrates regulates only cell death and lies downstream of the decision between cell cycle arrest and cell death.

sponse to irradiation, it is likely that hus-1 and mrt-2 communicate with a separate pathway to induce proliferation arrest in C. elegans. Finally, an upstream negative regulator of the apoptotic pathway has been identified: abl-1 [67]. Mutations in abl-1 lead to higher basal and radiation-induced germline apoptosis. This phenotype can be rescued by mutating checkpoint genes, including hus-1, mrt-2, rad-5 and cep-1. Further clarification of the relationship between checkpoint genes and apoptotic genes begins with the continued identification of cellular factors that play a regulatory role in germ cell death. In a genome-wide study, anti-apoptotic genes were identified using RNAi-mediated knockdown of 86% of the protein-coding genes [68]. Of the 21 gene inactivations that led to increased germ cell death, 16 act through the cep1checkpoint. The remaining 5, including ced-9 and pmk-3, a p38 MAPK homolog, are hus-1 and cep1-independent. While the role of ced-9 has already been established, it will be interesting to learn more about how pmk-3 regulates apoptosis in the worm, as MAPK family members have been shown to be important for the regulation of germ cell apoptosis in other systems (Fig. 1) [69]. 3.2.2. Cell cycle regulation One critical target of the DNA damage response is the cell cycle engine driven by cyclin-dependent kinases (Cdks). In vertebrates, the upstream kinases ATM and ATR, phosphorylate and activate checkpoint kinases Chk2(Cds1) and Chk1, respectively [70,71]. Upon checkpoint initiation in G2, Chk1 and Cds1 phosphorylate and inhibit Cdc25C, a Cdc2activating phosphatase. When damage is detected prior to or during S phase, Cdc25A, an activator of CyclinE/Cdk2 is phosphorylated and inhibited by Cds1. The role of checkpoint kinases, Xenopus Chk1 (XChk1) and Xenopus Cds1 (XCds1) has been addressed by overexpressing dominant negative forms of these kinases in early Xenopus embryos, where checkpoints are not normally present [72,73]. Inter-

estingly, overexpression of a kinase-dead, dominant negative Chk1, but not Cds1in the early Xenopus embryo results in apoptosis at the EGT, suggesting that XChk1 activity is necessary for the prevention of apoptosis. Upon further inspection, it was observed that embryos expressing dominant negative Chk1 underwent normal replication and cell division prior to the MBT, but at the MBT cells underwent two additional rounds of rapid division, whereas in control embryos, cell cycles slow down at the MBT with the introduction of gap phases. Inhibition of another gene implicated in DNA replication, geminin, induces apoptosis at the EGT as well [74]. In mammalian somatic cells, inhibition of checkpoint proteins does not generally lead to cell death. The complete penetrance of cell death in the checkpoint-inhibited embryos demonstrates a developmentally specific dependence on checkpoint function, and may indicate direct communication between cell cycle components and the apoptotic machinery. What are the specific links between the cell cycle and cell death machinery? The cell cycle is driven by cyclins and cyclin-dependent kinases. Cdc2 in a complex with cyclin A or cyclin B is necessary for M-phase progression, whereas CycA/Cdk2 and CycE/Cdk2 are needed in S-phase. In order to progress from one cell cycle phase to the next, Cdk activity oscillates; if Cdc2 activity remains high, the cell cannot exit M-phase [75]. The initiation of a cell cycle checkpoint ultimately leads to regulation of Cdk activity to block progression of the cell cycle. In Xenopus, cyclin A1 is present in unfertilized eggs through early embyrogenesis, but is rapidly degraded at the MBT. Cyclin A2, on the other hand, is present at low levels in early embryos, but begins to accumulate after the MBT [38,76]. Some interesting links between the cell cycle and cell death have been observed in Xenopus. When embryos are irradiated prior to the MBT, CycA1/Cdk2 activity is prolonged compared to unirradiated embryos [49]. Furthermore, cyclin A2 is a substrate for caspases [53,77]. Caspase-mediated

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cleavage of CyclinA2 removes the destruction box, thereby stabilizing the protein. The enhanced stability of CyclinA2 keeps Cdk2 active, effectively arresting the cell cycle. The cleavage also prevents p27Xic1 from binding and inhibiting the CycA2/Cdk2 complex. Interestingly, it was shown that incubation of recombinant cleaved CycA2/Cdk2 in extracts induced fragmentation of DNA in the absence of detectable caspase activity, suggesting that the cleaved CycA2/Cdk2 complex plays a direct role in regulating one of the hallmarks of apoptosis. Furthermore, overexpression of cyclin E in the early Xenopus embryo results in rapid uniform apoptosis at the EGT [78]. The reason embryos activate the apoptotic program following Cyclin E overexpression may be due to the fact that Cyclin E overexpression blocks DNA synthesis. As the embryos continue to divide, DNA is damaged with each cell cycle. When cell cycle checkpoints are activated at the EGT, the irreparable damage is recognized and the apoptotic program is initiated. Finally, the cell cycle regulator Wee1 has been implicated in caspase activation in Xenopus cellfree extracts [79]. Wee1 was identified as a Crk-interacting protein; Crk is an adaptor protein in the phosphotyrosine signaling pathway essential for caspase activation in the in vitro Xenopus system. Wee1 is an inhibitor of Cdc2, however, its Cdc2-inhibiting activity is not sufficient to induce apoptosis in the cell-free system, as Myt1, another Cdc2-inhibiting protein does not accelerate apoptosis. The dual function of Wee1 as a potent inhibitor of Cdc2 and as an activator of cell death suggests that it may play a role in the decision to arrest the cell cycle or to induce apoptosis.

4. Conclusion Recent studies of the regulation of apoptosis during early development have highlighted the ability of the embryo to integrate cell cycle, checkpoint and cell death controls. Meiotic cell cycles at the end of oogenesis and early mitotic cell cycles at the onset of development display characteristic regulatory features with respect to apoptosis. Genetic studies in C. elegans and Drosophila have been instrumental in the identification and ordering of components of the cell death pathway. In addition, these organisms regulate apoptosis in a developmental stage-specific manner. Early embryos of vertebrates also exhibit developmentally regulated apoptosis. Zebrafish and Xenopus embryos acquire the ability to activate the cell death program only at gastrulation. This timing correlates with the introduction of cell cycle checkpoints, and as we have discussed, it is likely that the cell cycle machinery and the apoptotic machinery communicate directly. The specific mechanisms involved will likely be elucidated through the examination of genes affecting early embryonic development. Interestingly, the majority of mutations identified in a recent maternal-effect mutant screen in zebrafish affect embryo viability, however, apoptosis was not detectable until early gastrulation [80,81]. A screen for maternal-effect mutants that exhibit precocious or delayed

apoptosis onset following DNA damage would likely uncover maternal cell cycle, checkpoint and apoptotic factors involved in the regulation of the acquistion of the cell death program in the early embryo. Coordination of cell cycle checkpoints and apoptosis is an essential feature of early embryos. This coordination is also critical for somatic cells where it plays a crucial role in preventing the occurrence of cancer. By understanding how embryos first establish the relationships between apoptosis and checkpoints, and by studying how this coordination has been modified through evolution, we will further our understanding of how the loss of these regulatory mechanisms contributes to disease.

Acknowledgements We thank members of the laboratory for helpful discussion and critical reading of the manuscript.

References [1] Ellis HM, Horvitz HR. Genetic control of programmed cell death in the nematode C. elegans. Cell 1986;44:817–29. [2] Metzstein MM, Stanfield GM, Horvitz HR. Genetics of programmed cell death in C. elegans: past, present and future. Trends Genet 1998;14:410–6. [3] Chen F, Hersh BM, Conradt B, Zhou Z, Riemer D, Gruenbaum Y, et al. Translocation of C. elegans CED-4 to nuclear membranes during programmed cell death. Science 2000;287:1485–9. [4] Danial NN, Korsmeyer SJ. Cell death: critical control points. Cell 2004;116:205–19. [5] Nagata S, Golstein P. The Fas death factor. Science 1995;267:1449–56. [6] Mizuno T, Zhong X, Rothstein TL. Fas-induced apoptosis in B cells. Apoptosis 2003;8:451–60. [7] Green DR, Droin N, Pinkoski M. Activation-induced cell death in T cells. Immunol Rev 2003;193:70–81. [8] Scorrano L, Korsmeyer SJ. Mechanisms of cytochrome c release by proapoptotic BCL-2 family members. Biochem Biophys Res Commun 2003;304:437–44. [9] Varkey J, Chen P, Jemmerson R, Abrams JM. Altered cytochrome c display precedes apoptotic cell death in Drosophila. J Cell Biol 1999;144:701–10. [10] Salvesen GS, Duckett CS. IAP proteins: blocking the road to death’s door. Nat Rev Mol Cell Biol 2002;3:401–10. [11] Bergmann A, Yang AY, Srivastava M. Regulators of IAP function: coming to grips with the grim reaper. Curr Opin Cell Biol 2003;15:717–24. [12] White K, Grether ME, Abrams JM, Young L, Farrell K, Steller H. Genetic control of programmed cell death in Drosophila. Science 1994;264:677–83. [13] Du C, Fang M, Li Y, Li L, Wang X. Smac, a mitochondrial protein that promotes cytochrome c-dependent caspase activation by eliminating IAP inhibition. Cell 2000;102:33–42. [14] Verhagen AM, Ekert PG, Pakusch M, Silke J, Connolly LM, Reid GE, et al. Identification of DIABLO, a mammalian protein that promotes apoptosis by binding to and antagonizing IAP proteins. Cell 2000;102:43–53. [15] Suzuki Y, Imai Y, Nakayama H, Takahashi K, Takio K, Takahashi R. A serine protease, HtrA2, is released from the mitochondria and interacts with XIAP, inducing cell death. Mol Cell 2001;8:613–21.

J. Greenwood, J. Gautier / Seminars in Cell & Developmental Biology 16 (2005) 215–224 [16] Vaux DL, Silke J. Mammalian mitochondrial IAP binding proteins. Biochem Biophys Res Commun 2003;304:499–504. [17] Tilly JL. Commuting the death sentence: how oocytes strive to survive. Nat Rev Mol Cell Biol 2001;2:838–48. [18] Baum JS, St. George J, McCall K. Programmed cell death in the germline. Semin Cell Dev Biol 2005;16:245–58. [19] Kimble J, Hirsh D. The postembryonic cell lineages of the hermaphrodite and male gonads in Caenorhabditis elegans. Dev Biol 1979;70:396–417. [20] Sulston JE, Horvitz HR. Post-embryonic cell lineages of the nematode, Caenorhabditis elegans. Dev Biol 1977;56:110–56. [21] Sulston JE, Schierenberg E, White JG, Thomson JN. The embryonic cell lineage of the nematode Caenorhabditis elegans. Dev Biol 1983;100:64–119. [22] Gumienny TL, Lambie E, Hartwieg E, Horvitz HR, Hengartner MO. Genetic control of programmed cell death in the Caenorhabditis elegans hermaphrodite germline. Development 1999;126:1011–22. [23] Conradt B, Horvitz HR. The C. elegans protein EGL-1 is required for programmed cell death and interacts with the Bcl-2-like protein CED-9. Cell 1998;93:519–29. [24] McCall K. Eggs over easy: cell death in the Drosophila ovary. Dev Biol 2004;274:3–14. [25] Peterson JS, Barkett M, McCall K. Stage-specific regulation of caspase activity in Drosophila oogenesis. Dev Biol 2003;260:113–23. [26] Foley K, Cooley L. Apoptosis in late stage Drosophila nurse cells does not require genes within the H99 deficiency. Development 1998;125:1075–82. [27] Laundrie B, Peterson JS, Baum JS, Chang JC, Fileppo D, Thompson SR, et al. Germline cell death is inhibited by P-element insertions disrupting the dcp-1/pita nested gene pair in Drosophila. Genetics 2003;165:1881–8. [28] Rodriguez A, Chen P, Oliver H, Abrams JM. Unrestrained caspasedependent cell death caused by loss of Diap1 function requires the Drosophila Apaf-1 homolog, Dark. EMBO J 2002;21:2189–97. [29] Ivanovska I, Lee E, Kwan KM, Fenger DD, Orr-Weaver TL. The Drosophila MOS ortholog is not essential for meiosis. Curr Biol 2004;14:75–80. [30] Sasaki K, Chiba K. Induction of apoptosis in starfish eggs requires spontaneous inactivation of MAPK (extracellular signalregulated kinase) followed by activation of p38MAPK. Mol Biol Cell 2004;15:1387–96. [31] Newmeyer DD, Farschon DM, Reed JC. Cell-free apoptosis in Xenopus egg extracts: inhibition by Bcl-2 and requirement for an organelle fraction enriched in mitochondria. Cell 1994;79:353–64. [32] Kornbluth S. Apoptosis in Xenopus egg extracts. Methods Enzymol 1997;283:600–14. [33] Tashker JS, Olson M, Kornbluth S. Post-cytochrome C protection from apoptosis conferred by a MAPK pathway in Xenopus egg extracts. Mol Biol Cell 2002;13:393–401. [34] Bergmann A, Agapite J, McCall K, Steller H. The Drosophila gene hid is a direct molecular target of Ras-dependent survival signaling. Cell 1998;95:331–41. [35] Kurada P, White K. Ras promotes cell survival in Drosophila by downregulating hid expression. Cell 1998;95:319–29. [36] Newport J, Kirschner M. A major developmental transition in early Xenopus embryos: I. characterization and timing of cellular changes at the midblastula stage. Cell 1982;30:675–86. [37] Newport J, Kirschner M. A major developmental transition in early Xenopus embryos: II. Control of the onset of transcription. Cell 1982;30:687–96. [38] Howe JA, Howell M, Hunt T, Newport JW. Identification of a developmental timer regulating the stability of embryonic cyclin A and a new somatic A-type cyclin at gastrulation. Genes Dev 1995;9:1164–76. [39] Howe JA, Newport JW. A developmental timer regulates degradation of cyclin E1 at the midblastula transition during Xenopus embryogenesis. Proc Natl Acad Sci USA 1996;93:2060–4.

223

[40] Kirschner M, Newport J, Gerhart J. The timing of early developmental events in Xenopus. Trends Genet 1985;1:41–7. [41] Abrams JM, White K, Fessler LI, Steller H. Programmed cell death during Drosophila embryogenesis. Development 1993;117:29–43. [42] Zhou L, Steller H. Distinct pathways mediate UV-induced apoptosis in Drosophila embryos. Dev Cell 2003;4:599–605. [43] Wang SL, Hawkins CJ, Yoo SJ, Muller HA, Hay BA. The Drosophila caspase inhibitor DIAP1 is essential for cell survival and is negatively regulated by HID. Cell 1999;98:453–63. [44] Goyal L, McCall K, Agapite J, Hartwieg E, Steller H. Induction of apoptosis by Drosophila reaper, hid and grim through inhibition of IAP function. EMBO J 2000;19:589–97. [45] Harlin H, Reffey SB, Duckett CS, Lindsten T, Thompson CB. Characterization of XIAP-deficient mice. Mol Cell Biol 2001;21:3604–8. [46] Igaki T, Miura M. Role of Bcl-2 family members in invertebrates. Biochim Biophys Acta 2004;1644:73–81. [47] Quinn L, Coombe M, Mills K, Daish T, Colussi P, Kumar S, et al. Buffy, a Drosophila Bcl-2 protein, has anti-apoptotic and cell cycle inhibitory functions. EMBO J 2003;22:3568–79. [48] Colussi PA, Quinn LM, Huang DC, Coombe M, Read SH, Richardson H, et al. Debcl, a proapoptotic Bcl-2 homologue, is a component of the Drosophila melanogaster cell death machinery. J Cell Biol 2000;148:703–14. [49] Anderson JA, Lewellyn AL, Maller JL. Ionizing radiation induces apoptosis and elevates cyclin A1-Cdk2 activity before but not after the midblastula transition in Xenopus. Mol Biol Cell 1997;8:1195–206. [50] Hensey C, Gautier J. A developmental timer that regulates apoptosis at the onset of gastrulation. Mech Dev 1997;69:183–95. [51] Ikegami R, Hunter P, Yager TD. Developmental activation of the capability to undergo checkpoint-induced apoptosis in the early zebrafish embryo. Dev Biol 1999;209:409–33. [52] Ikegami R, Zhang J, Rivera-Bennetts AK, Yager TD. Activation of the metaphase checkpoint and an apoptosis programme in the early zebrafish embryo, by treatment with the spindle-destabilising agent nocodazole. Zygote 1997;5:329–50. [53] Stack JH, Newport JW. Developmentally regulated activation of apoptosis early in Xenopus gastrulation results in cyclin A degradation during interphase of the cell cycle. Development 1997;124:3185–95. [54] Sible JC, Anderson JA, Lewellyn AL, Maller JL. Zygotic transcription is required to block a maternal program of apoptosis in Xenopus embryos. Dev Biol 1997;189:335–46. [55] Finkielstein CV, Lewellyn AL, Maller JL. The midblastula transition in Xenopus embryos activates multiple pathways to prevent apoptosis in response to DNA damage. Proc Natl Acad Sci USA 2001;98:1006–11. [56] Evans EK, Kuwana T, Strum SL, Smith JJ, Newmeyer DD, Kornbluth S. Reaper-induced apoptosis in a vertebrate system. EMBO J 1997;16:7372–81. [57] Thress K, Henzel W, Shillinglaw W, Kornbluth S. Scythe: a novel reaper-binding apoptotic regulator. EMBO J 1998;17:6135–43. [58] Thress K, Evans EK, Kornbluth S. Reaper-induced dissociation of a Scythe-sequestered cytochrome c-releasing activity. EMBO J 1999;18:5486–93. [59] Gartner A, Milstein S, Ahmed S, Hodgkin J, Hengartner MO. A conserved checkpoint pathway mediates DNA damage–induced apoptosis and cell cycle arrest in C. elegans. Mol Cell 2000;5:435–43. [60] Derry WB, Putzke AP, Rothman JH. Caenorhabditis elegans p53: role in apoptosis, meiosis, and stress resistance. Science 2001;294:591–5. [61] Schumacher B, Hofmann K, Boulton S, Gartner A. The C. elegans homolog of the p53 tumor suppressor is required for DNA damageinduced apoptosis. Curr Biol 2001;11:1722–7. [62] Ollmann M, Young LM, Di Como CJ, Karim F, Belvin M, Robertson S, et al. Drosophila p53 is a structural and functional homolog of the tumor suppressor p53. Cell 2000;101:91–101.

224

J. Greenwood, J. Gautier / Seminars in Cell & Developmental Biology 16 (2005) 215–224

[63] Sogame N, Kim M, Abrams JM. Drosophila p53 preserves genomic stability by regulating cell death. Proc Natl Acad Sci USA 2003;100:4696–701. [64] Brodsky MH, Nordstrom W, Tsang G, Kwan E, Rubin GM, Abrams JM. Drosophila p53 binds a damage response element at the reaper locus. Cell 2000;101:103–13. [65] Hofmann ER, Milstein S, Boulton SJ, Ye M, Hofmann JJ, Stergiou L, et al. Caenorhabditis elegans HUS-1 is a DNA damage checkpoint protein required for genome stability and EGL-1-mediated apoptosis. Curr Biol 2002;12:1908–18. [66] Ahmed S, Alpi A, Hengartner MO, Gartner A. C. elegans RAD5/CLK-2 defines a new DNA damage checkpoint protein. Curr Biol 2001;11:1934–44. [67] Deng X, Hofmann ER, Villanueva A, Hobert O, Capodieci P, Veach DR, et al. Caenorhabditis elegans ABL-1 antagonizes p53mediated germline apoptosis after ionizing irradiation. Nat Genet 2004;36:906–12. [68] Lettre G, Kritikou EA, Jaeggi M, Calixto A, Fraser AG, Kamath RS, et al. Genome-wide RNAi identifies p53-dependent and -independent regulators of germ cell apoptosis in C. elegans. Cell Death Differ 2004. [69] Wada T, Penninger JM. Mitogen-activated protein kinases in apoptosis regulation. Oncogene 2004;23:2838–49. [70] Rouse J, Jackson SP. Interfaces between the detection, signaling, and repair of DNA damage. Science 2002;297:547–51. [71] Zhou BB, Elledge SJ. The DNA damage response: putting checkpoints in perspective. Nature 2000;408:433–9. [72] Carter AD, Sible JC. Loss of XChk1 function triggers apoptosis after the midblastula transition in Xenopus laevis embryos. Mech Dev 2003;120:315–23.

[73] Shimuta K, Nakajo N, Uto K, Hayano Y, Okazaki K, Sagata N. Chk1 is activated transiently and targets Cdc25A for degradation at the Xenopus midblastula transition. EMBO J 2002;21:3694– 703. [74] McGarry TJ. Geminin deficiency causes a Chk1-dependent G2 arrest in Xenopus. Mol Biol Cell 2002;13:3662–71. [75] Morgan DO. Cyclin-dependent kinases: engines, clocks, and microprocessors. Annu Rev Cell Dev Biol 1997;13:261–91. [76] Hartley RS, Rempel RE, Maller JL. In vivo regulation of the early embryonic cell cycle in Xenopus. Dev Biol 1996;173:408– 19. [77] Finkielstein CV, Chen LG, Maller JL. A role for G1/S cyclindependent protein kinases in the apoptotic response to ionizing radiation. J Biol Chem 2002;277:38476–85. [78] Richard-Parpaillon L, Cosgrove RA, Devine C, Vernon AE, Philpott A. G1/S phase cyclin-dependent kinase overexpression perturbs early development and delays tissue-specific differentiation in Xenopus. Development 2004;131:2577–86. [79] Smith JJ, Evans EK, Murakami M, Moyer MB, Moseley MA, Woude GV, et al. Wee1-regulated apoptosis mediated by the crk adaptor protein in Xenopus egg extracts. J Cell Biol 2000;151:1391– 400. [80] Dosch R, Wagner DS, Mintzer KA, Runke G, Wiemelt AP, Mullins MC. Maternal control of vertebrate development before the midblastula transition: mutants from the zebrafish I. Dev Cell 2004;6:771–80. [81] Wagner DS, Dosch R, Mintzer KA, Wiemelt AP, Mullins MC. Maternal control of development at the midblastula transition and beyond: mutants from the zebrafish II. Dev Cell 2004;6:781– 90.