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Keywords:

  • apoptosis;
  • caspase signaling;
  • cell differentiation;
  • cell migration and shaping;
  • cell proliferation

Abstract

  1. Top of page
  2. Abstract
  3. Introduction to caspase signaling
  4. Caspases in cell proliferation
  5. Caspases in cell differentiation
  6. Caspases in subcellular compartments
  7. Caspases in cell migration
  8. Caspases in cell shaping
  9. Concluding remarks
  10. Acknowledgments
  11. References

The caspases are a family of cysteine proteases that function as central regulators of cell death. Recent investigations in Caenorhabditis elegans, Drosophila, and mice indicate that caspases are essential not only in controlling the number of cells involved in sculpting or deleting structures in developing animals, but also in dynamic cell processes such as cell-fate determination, compensatory proliferation of neighboring cells, and actin cytoskeleton reorganization, in a non-apoptotic context during development. This review focuses primarily on caspase functions involving their enzymatic activity.


Introduction to caspase signaling

  1. Top of page
  2. Abstract
  3. Introduction to caspase signaling
  4. Caspases in cell proliferation
  5. Caspases in cell differentiation
  6. Caspases in subcellular compartments
  7. Caspases in cell migration
  8. Caspases in cell shaping
  9. Concluding remarks
  10. Acknowledgments
  11. References

Caspases are a family of cysteine proteases that are highly conserved in multicellular organisms and that function as central regulators of cell death. Apoptosis, the programmed cell death mediated by caspase activation, is a fundamental cellular response that regulates tissue homeostasis by eliminating unwanted cells, and it plays a crucial role in shaping the body during development. Inhibiting cell death signals causes morphological defects such as abnormal heart formation, exencephaly, and syndactyly in mammals (Vaux & Korsmeyer 1999; Ranger et al. 2001).

Cell-death pathways are well conserved in all metazoans, from invertebrates to vertebrates. Caspase, the key enzyme associated with cell death, was first identified as Ced-3 in Caenorhabditis elegans (Ellis & Horvitz 1986; Yuan et al. 1993). The first caspase found and reported as a Ced-3 homologue in mammals was ICE (interleukin-1β-converting enzyme), a cysteine protease responsible for processing and secreting proIL-1β (Cerretti et al. 1992; Thornberry et al. 1992; Miura et al. 1993). Therefore, the function of the first discovered mammalian Ced-3 homologue involved both apoptotic and non-apoptotic processes. While CED-3 is the sole caspase required for programmed cell death in C. elegans, multiple caspases are required for apoptosis in more complex organisms, such as flies and mammals (Table 1). This evolutionary expansion of the caspase family may have arisen around the dual purposes of executing apoptotic cell death and of carrying out multiple cellular processes that do not necessarily involve cell death. Caspases are synthesized as zymogens, and their activation requires specific cleavage at select aspartate residues. The initial processing of inactive caspase separates the large (p20) and small (p10) subunits, after which the N-terminal domain is removed to form the catalytically active protease (Degterev et al. 2003).

Table 1. Drosophila and mammalian caspases
SubfamilyDrosophilaMammals
  1. While the Caenorhabditis elegans genome has four caspase genes (ced-3, csp-1, -2, -3), only ced-3 is required in programmed cell death. CARD, caspase recruitment domain; DED, death effector domain.

CARD containing caspase (initiator caspase)DroncCaspase-1, Caspase-2, Caspase-4, Caspase-5, Caspase-9, Caspase-11, Caspase-12
DED containing caspase (initiator caspase)DreddCaspase-8, Caspase-10
Short prodomain caspase (executioner caspase)Drice, Dcp-1Caspase-3, Caspase-6, Caspase-7
OthersStrica/Dream, Decay, DammCaspase-14

Caspases can be classified into two categories, initiator caspases and executioner caspases (Table 1). Initiator caspases have a long N-terminal prodomain, which mediates the formation of protein complexes that provide the molecular platform for caspase activation and inhibition. Initiator caspases cleave a few specific substrates, including the zymogens of executioner caspases, which are activated by the cleavage. The activated executioner caspases then cleave their respective substrates, which elicit apoptotic cell death, along with its characteristic morphological features: membrane blebbing, pyknotic nuclei, cell rounding, and apoptotic vesicle formation (Clarke 1990).

The caspase regulation following apoptotic stimulation has been studied in detail, and the core machineries of the caspase activation pathway are conserved throughout evolution (Fig. 1) (Schafer & Kornbluth 2006). Caspase activation is regulated through either extrinsic or intrinsic signaling pathways. In the extrinsic pathway, Fas and tumor necrosis factor receptor (TNFR) stimulation leads to Caspase-8 activation. The intrinsic pathway, which is used for most apoptotic caspase activation in mammals, triggers the mitochondrial release of cytochrome c. Cytochrome c, Apaf-1 and procaspase-9 oligomerize to form what is called the apoptosome complex. The activated caspase-9 in this complex triggers caspase-3 activation, thereby executing apoptosis.

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Figure 1.  Conservation of caspase-dependent cell death machinery. Functional homologues across species are represented by the same color and shape. Caspases and the proapoptotic member of the Bcl-2 family, Apaf-1, are evolutionarily well conserved. The negative regulation of CED-4 by CED-9 can be cancelled by EGL-1, a BH3-only member of the Bcl-2 superfamily, and CED-4 consequently promotes activation of the caspase CED-3 in Caenorhabditis elegans. EGL-1 binding to CED-9 also activates CED-9’s killing function. In Drosophila, Dark promotes activation of the initiator caspase Dronc, and multi-domain Bcl-2 family members Debcl and Buffy may regulate this activation. Drosophila inhibitor of apoptosis protein 1 (DIAP1) inhibits Dronc and the effector caspases Drice and Dcp-1. RHG family proteins such as Rpr, Hid, Grim, Sickle, and Jafrac-2 promote cell death, in part by disrupting DIAP1’s ability to inhibit caspase activity. In mammals, Apaf-1-dependent caspase-9 activation is regulated by proapoptotic Bcl-2 family proteins such as Bax and Bak, which are inhibited by the antiapoptotic Bcl-2 family. BH3-domain-only proteins facilitate the Bax- and Bak-dependent release of mitochondrially localized proteins, including cytochrome c (cyt c). HtrA2/Omi and Smac/Diablo, functional homologues of RHG proteins, are also released from mitochondria in apoptotic cells. The binding of a death ligand to its corresponding receptor activates caspase-8. In some cell types, caspase-8 directly activates executioner caspases, and in others, Caspase-8 processes the BH3-only protein Bid to promote Bax activation. Dotted arrows indicate possible interactions between components of the apoptotic pathway. BH3, Bcl-2 homology 3; CED, cell-death abnormal; EGL, egg-laying defective; Rpr, Reaper; Hid, head involution defective.

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In Drosophila, the initiator caspase dronc and the executioner caspase drice are responsible for most of the embryonic apoptosis, and null mutant alleles of these genes prevent most embryonic apoptosis (Chew et al. 2004; Daish et al. 2004; Waldhuber et al. 2005; Xu et al. 2005, 2006; Kondo et al. 2006; Muro et al. 2006). A similar reduction in embryonic apoptosis occurs in the Drosophila apaf-1 (dark/hac-1/dapaf-1) mutant, suggesting that the Apaf-1/caspase activation pathway is essential for apoptosis in the Drosophila embryo (Akdemir et al. 2006; Srivastava et al. 2007). While cells from knockouts of non-redundant apoptosome components (apaf-1 or caspase-9) are resistant to various apoptotic stimuli, apaf-1/caspase-9-independent cell death is still observed during development (Schafer & Kornbluth 2006). Dark/dronc-independent cell death is observed in some Drosophila post-embryonic tissues, including the midgut, suggesting that apoptosome-independent cell death is associated with tissue development (Denton et al. 2009).

While caspases are prominently associated with apoptosis, recent research findings indicate that they also play dynamic roles in compensatory proliferation of neighboring cells and cell-fate determination, as well as in actin cytoskeleton reorganization, which impacts a cell’s shape and migration (Kuranaga & Miura 2007; Lamkanfi et al. 2007; Feinstein-Rotkopf & Arama 2009; Yi & Yuan 2009). Caspases can also signal through protein–protein interactions independently of enzymatic activity, as in the caspase activation of nuclear factor-κB (NF-κB) (Lamkanfi et al. 2007). However, this review will focus mainly on the enzymatic activity and functions of caspases.

Caspases in cell proliferation

  1. Top of page
  2. Abstract
  3. Introduction to caspase signaling
  4. Caspases in cell proliferation
  5. Caspases in cell differentiation
  6. Caspases in subcellular compartments
  7. Caspases in cell migration
  8. Caspases in cell shaping
  9. Concluding remarks
  10. Acknowledgments
  11. References

Cell proliferation has been a primary focus of study in developmental biology, and caspases are closely involved in several features of cell proliferation. Caspase-8 can be both a positive and negative regulator of lymphocyte proliferation. Patients with homozygous caspase-8 mutations show defective T lymphocyte, B lymphocyte, and natural killer cell activation (Chun et al. 2002). Mice with a conditional caspase-8 knockout in the peripheral T cells also show defective proliferation of these cells following T-cell receptor (TcR) activation (Salmena et al. 2003). However, the long-term effect of the murine T-cell-specific caspase-8 knockout is a lymphoproliferative phenotype, not an immunodeficiency (Salmena & Hakem 2005).

Cell-cycle regulators are substrates for caspase-3. Caspase-3 cleavage of the cyclin-dependent kinase (CDK) inhibitor p27 promotes lymphoid cell proliferation (Frost et al. 2001). In other situations, however, Caspase-3 acts to prevent proliferation. Binding of the CDK inhibitor p21 to PCNA, a cofactor for DNA polymerase δ, promotes cell proliferation. Caspase-3 can cleave p21 at its C-terminal PCNA-binding sites, causing the PCNA to be released. Similarly, caspase-3 knockout mice exhibit B-cell hyperproliferation (Woo et al. 2003).

Caspases can also integrate the processes of cell death and cell proliferation to shape tissue maintenance. Animal tissues such as the Drosophila larval imaginal disc have a striking regenerative capacity not only during development, but also in adulthood. A large number of cells undergo cell death when such tissues are accidentally injured. The final shape and size of the healed tissue is often the same as the uninjured tissue, owing to additional cell proliferation that compensates for the cell loss. This phenomenon is called compensatory proliferation. The local induction of apoptosis by ectopic toxin expression results in elevated cell proliferation around the apoptosis site, suggesting that cells can perceive apoptosis in their vicinity and undergo cell division until the original cell number is restored (Milan et al. 1997).

The expression of the Drosophila cell death-inducing genes reaper (rpr) or head involution defective (hid) in the imaginal disc, together with p35, a potent inhibitor of effector caspases such as Drice and Dcp-1, activates the signaling cascade as far as activation of the initiator caspase Dronc, and the cells fail to undergo apoptosis. The resultant “undead” cells ectopically express Wingless (WG) and Decapentaplegic (DPP), which induce uncontrolled cell proliferation (Huh et al. 2004a; Perez-Garijo et al. 2004, 2005; Ryoo et al. 2004). The Drosophila caspase-9 orthologue Dronc was found to be required for the compensatory proliferation in this experimental system, suggesting that it is involved in inducing mitogen expression (Huh et al. 2004a; Kondo et al. 2006; Wells et al. 2006).

In contrast, in the case of differentiating eye tissues in Drosophila, the undead apoptotic cells induce compensatory proliferation by upregulating Hedgehog (Hh), not Wnt or Dpp (Fan & Bergmann 2008). Therefore, caspases may coordinate both cell death and compensatory proliferation during development and regeneration. This compensatory proliferation signal activated by apoptotic cells has been also found in another animal. In Hydra, during basal head regeneration after midgastric bisection, cells from the interstitial lineage immediately undergo cell death. These apoptotic cells provide an immediate but transient source of Wnt3 that activates the Wnt-β-catenin pathway in the surrounding cycling cells, which rapidly divide (Chera et al. 2009). In the cases of both Drosophila and Hydra, caspase and the intensity of the apoptotic process play a role in tissue regeneration, but the caspase substrates that induce compensatory proliferation remain to be investigated.

Caspases in cell differentiation

  1. Top of page
  2. Abstract
  3. Introduction to caspase signaling
  4. Caspases in cell proliferation
  5. Caspases in cell differentiation
  6. Caspases in subcellular compartments
  7. Caspases in cell migration
  8. Caspases in cell shaping
  9. Concluding remarks
  10. Acknowledgments
  11. References

Caspase activation during apoptosis ultimately disrupts the nuclear structure. Some cells lose their nuclei during differentiation, and this can be considered a specialized form of apoptosis. Such cells include keratinocytes, megakaryocytes, erythrocytes, and lens cells, and it is not surprising that the terminal differentiation of these cells involves caspases (Lamkanfi et al. 2007). Caspases can mediate irreversible signal transduction, a type of signal regulation that may be suitable for determining cell fate. Activated caspases are found in human peripheral blood monocytes undergoing macrophage colony-stimulating factor (M-CSF)-induced differentiation into macrophages, and inhibiting caspases inhibits this differentiation (Sordet et al. 2002). A caspase-8 conditional knockout in bone-marrow cells prevents myelomonocytic-lineage cells from differentiating into macrophages (Kang et al. 2004). Caspase-8 cleaves receptor-interacting protein (RIP), a death-domain-containing kinase that regulates NF-κB, thus downregulating NF-κB during macrophage differentiation (Rebe et al. 2007). Caspase-3 is required for skeletal muscle differentiation in vitro, as seen by defective myotube and myofiber formation in the primary myoblasts isolated from caspase-3 knockout mice (Fernando et al. 2002). Sterile Twenty-like kinase (MST1), which is required for muscle differentiation, is activated when it is cleaved by Caspase-3 (Fernando et al. 2002). Caspase-8 is required for the syncytial fusion of cytotrophoblasts, which forms the syncytiotrophoblasts, during differentiation of the human placental villous trophoblasts (Black et al. 2004). Caspase-3-deficient mice show decreased osteogenic differentiation of the bone marrow stromal stem cells, resulting in delayed ossification and decreased bone mineral density (Miura et al. 2004). Notch1 is activated during the commitment stage of embryonic keratinocyte differentiation, and Notch1 signaling induces caspase-3. The activation and increase in caspase-3 are not sufficient to induce apoptosis; instead, caspase-3 cleaves and activates PKC-δ, which is a positive regulator of keratinocyte differentiation. In capsase-3-deficient mice, interfollicular embryonic keratinocyte proliferation is increased, but differentiation is decreased. Therefore, caspase-3 is required to commit embryonic keratinocytes to terminal differentiation (Okuyama et al. 2004). In vitro experiments showed that caspase-3 is required for the differentiation of Bergmann glia in neural cells (Oomman et al. 2006). A transient increase in caspase-3 activity is observed during differentiation from neurospheres or PC12 cells (Rohn et al. 2004; Fernando et al. 2005), and administering a caspase-3 inhibitor prevents neural differentiation. Caspase-1 activity also increases in, and is essential to, PC12 cell differentiation (Vaisid et al. 2005).

Caspase is a critical enzyme in stem cell differentiation. Non-apoptotic caspase-3 activation has been observed in several stem cell lineages, including embryonic stem cells (ESCs), hematopoietic stem cells (HSCs), and neural stem cells (Miura et al. 2004). Caspase-3-deficient mice exhibit significant bone defects during early development; the alteration of the transforming growth factor-β (TGF-β)/Smad2 pathway and of cell-cycle progression decreases the osteogenic differentiation of bone marrow stromal stem cells (BMSSCs) (Miura et al. 2004). Based on the premise that stem-cell differentiation corresponds with a loss of capacity for self-renewal, two recent studies explored the role of Caspase-3 as a probable gatekeeper of stem-cell function (Fujita et al. 2008; Janzen et al. 2008). Stem cells lacking caspase-3 had marked defects in differentiation (Fujita et al. 2008). Fujita et al. reasoned that caspase-3 mediates its effects through the targeted cleavage of a pluripotent factor, and identified Nanog as a critical target substrate, since the expression of a caspase-3-resistant Nanog promoted ESC self-renewal while inhibiting differentiation (Fujita et al. 2008). Janzen et al. (2008) reported that the loss of caspase-3 resulted in an accumulation of phenotypically defined long-term repopulating HSCs, with a corresponding reduction in circulating mature hematopoietic cells. Cytokine-mediated signals were also elevated in caspase-3-deficient HSCs.

Macrochaetes, typical structures of the Drosophila peripheral nervous system, are external sensory organs located on the notum. Four large macrochaetes are observed on the wild-type scutellum. However, one extra macrochaete often appears on each side of the scutellum in dark-mutant flies (Fig. 2) (Kanuka et al. 1999; Rodriguez et al. 1999). Both dark mutants and dominant-negative Dronc-expressing flies have an extra sensory organ precursor (SOP) cell on each side, showing that caspase activation is involved in controlling SOP cell formation in the scutellum area (Kanuka et al. 2005). Furthermore, the increased number of SOP cells seen in caspase-inhibited wing discs is not a result of the artificial survival of cells that otherwise would have undergone apoptotic cell death, but rather is due to inhibited apoptosis-independent caspase activity (Kanuka et al. 2005). Genetic screening identified a novel substrate cleaved by the Dark-dependent caspase drICE, the Shaggy46 protein. This is an isoform encoded by the shaggy gene, a Drosophila orthologue of gsk-3β, that is essential for the negative regulation of Wingless signaling. Cleaving Shaggy46 converts it to the active isoform Shaggy10, which contributes to SOP cell formation (Fig. 3) (Kanuka et al. 2005).

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Figure 2.  Non-apoptotic caspase functions in Drosophila. Caspase activation mediates non-apoptotic functions, including sensory organ precursor (SOP) development, dendrite pruning, aristae shaping, sperm individualization, and border cell migration.

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Figure 3.  Non-apoptotic caspase regulation mechanisms and physiological functions. Caspases have dynamic roles not only in controlling apoptosis, but also in non-apoptotic processes such as cell-fate determination and actin cytoskeleton reorganization (cell shaping and migration). Mechanisms controlling caspase activation are essential for maintaining cell integrity, and these are carried out by the inhibitor of apoptosis proteins (IAPs). DIAP1 suppresses caspase activation by binding directly to caspases and promoting their degradation (Dronc) or non-degradative inactivation (executioner caspases). DIAP1 degradation is promoted by its DmIKKε-induced phosphorylation. DmIKKε maintains caspase activity at the threshold required for non-apoptotic functions, as a determinant of the DIAP1 protein level. Shaggy46 protein has been identified as an executioner caspase substrate during sensory organ precursor (SOP) differentiation. DmIKKε acts as a negative regulator of F-actin polymerization. DIAP1 positively regulates border cell migration and cell shaping by downregulating Dronc. DIAP2 controls caspase in non-apoptotic contexts by interacting with activated drICE to prevent cell death. In invertebrates, Drosophila sperm differentiation is the only known system requiring cytochrome c to activate caspase. The Drosophila genome contains two cytochrome c genes, cyt-c-d and cyt-c-p; only cyt-c-d is required for caspase activation, which is mediated by Dark-dependent Dronc activation during sperm individualization. Subcellular compartmentalization of activated caspase during spermatid differentiation is regulated through Soti and dBruce. Soti competes with an inhibitor of the apoptosis protein dBruce, which is a target for the Cullin-3-based complex necessary for caspase activation during spermatid terminal differentiation. Soti is expressed in a subcellular gradient within spermatids, and in turn promotes a similar dBruce gradient, resulting in an inverse gradient pattern of caspase activation.

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The existing literature implicates caspases and their regulators in non-apoptotic functions, raising further compelling questions of how caspase activity is controlled to prevent apoptosis. Evidence exists for at least two mechanisms by which a cell safely activates caspase for non-apoptotic functions. Mechanisms to control caspase activation are essential for maintaining cell integrity, and much of this function is carried out by the inhibitor of apoptosis proteins (IAPs), originally found in baculoviruses (Fig. 3). DIAP1, which contains a carboxy-terminal RING finger domain and functions as an E3 ubiquitin ligase, suppresses caspase activation by binding directly to caspases and promoting their degradation (Wilson et al. 2002) or non-degradative inactivation (Ditzel et al. 2008). The proapoptotic proteins Rpr, Hid, and Grim promote this DIAP1 degradation during periods of programmed cell death (Hays et al. 2002; Ryoo et al. 2002; Wilson et al. 2002; Yoo et al. 2002). Programmed cell death is initiated when caspases are released from DIAP1 inhibition. Therefore, the balance between the DIAP1 protein level and caspase activation determines whether cells will survive, or die by apoptosis.

A genetic modifier screen for genes that regulate caspase activation identified a novel regulator of caspase activation that determines the threshold of caspase activation by regulating DIAP1 turnover (Kuranaga et al. 2006). Drosophila IKK-related kinase (DmIKKε) is a homologue of the non-canonical members of IκB kinase (IKKε/IKKι or NAK/T2K/TBK1), which regulate the activation of NF-κB or interferon regulatory factor (IRF)-3 and -7 in mammals (Kawai & Akira 2006). DmIKKε determines the level of DIAP1 by modifying its phosphorylation. Ectopic DmIKKε expression causes DIAP1 phosphorylation and degradation. NAK/TBK1/T2K, the mammalian homologue of DmIKKε, phosphorylates mammalian XIAP, a potent caspase inhibitor. NAK/TBK1/T2K expression promotes XIAP phosphorylation and degradation under low-serum culture conditions, thus suggesting that the IKK-related kinase function in IAP phosphorylation and degradation is conserved in Drosophila and mammalian cells (Kuranaga et al. 2006). DmIKKε may maintain caspase activity at the threshold required for non-apoptotic functions as a determinant of the DIAP1 protein level (Fig. 3). This is consistent with the observation that knocking down DmIKKε leads to an increase in SOP cells, which phenocopies the dark mutant (Kuranaga et al. 2006).

Another mechanism that restrains or shuts down caspase activity under such non-apoptotic conditions is the Drosophila inhibitor of apoptosis protein 2 (DIAP2), which controls the level of caspase activity in living cells (Fig. 3). Although diap2-deficient cells remain viable in vivo, they have increased drICE activity and are sensitized to cell death following treatment with sublethal X-irradiation doses. DIAP2 forms a covalent adduct with the catalytic portion of drICE, and it requires a functional RING finger domain to target drICE for ubiquitination and block cell death. These data suggest that DIAP2 controls caspase in a non-apoptotic context, by efficiently interacting with activated drICE to prevent cell death (Ribeiro et al. 2007).

Caspase-3 also plays a dynamic role in neural activation. In the zebra finch auditory forebrain, birdsong exposure increased the concentration of activated caspase-3 in the caudomedial nidopallium (NCM), and inhibiting caspase-3 activity in the NCM during song training disrupted habituation memory development (Huesmann & Clayton 2006). In this case, activated caspase-3 was observed in both the NCM and in unstimulated regions of the brain, but was normally bound to the inhibitor of apoptosis protein XIAP to prevent unwanted cell death (Huesmann & Clayton 2006). These results support a mechanism in which active caspase-3 is always present in living cells, but is sequestered by its inhibitors and released only transiently for essential non-apoptotic functions.

Consistent with the caspase-dependent memory consolidation in zebra finches, a recent study showed that synaptic depression and AMPA receptor internalization in mouse hippocampal neurons requires caspase-9 and caspase-3/7 activity, and can be blocked by the overexpression of the anti-apoptotic proteins Bcl-xL and XIAP (Li et al. 2010). Li et al. demonstrated that stimulating N-methyl-D-aspartate (NMDA) receptors transiently activates caspase-3 without causing cell death, and long-term synaptic depression cannot be induced by stimulating NMDA receptors in hippocampal slices from caspace-3 knockout mice (Li et al. 2010).

Caspases in subcellular compartments

  1. Top of page
  2. Abstract
  3. Introduction to caspase signaling
  4. Caspases in cell proliferation
  5. Caspases in cell differentiation
  6. Caspases in subcellular compartments
  7. Caspases in cell migration
  8. Caspases in cell shaping
  9. Concluding remarks
  10. Acknowledgments
  11. References

Some cells appear to have mechanisms that resist caspase-mediated cell death even in the face of high levels of caspase activation. One mechanism involves sequestering the caspase activity in specific subcellular regions (Arama et al. 2003; Huh et al. 2004b; Kuo et al. 2006; Williams et al. 2006). Eliminating excessive dendrites and axonal projections is important for shaping the neuronal architecture and neural circuits. During insect metamorphosis, larval neurons undergo massive loss and reconstruction of dendrites and axons without cell death. This dendritic pruning requires the ubiquitin-proteasome system and ubcD1, an E2 ubiquitin-conjugating enzyme involved in the degradation of the caspase-inhibitory molecule DIAP1 (Fig. 2) (Ryoo et al. 2002; Kuo et al. 2006). DIAP1 prevents activation of the initiator caspase Dronc, and UbcD1 activation is likely to lead to the degradation of DIAP1 and the subsequent activation of Dronc (Fig. 3). A loss-of-function dronc mutant prevents dendritic pruning, and its dendrites are labeled by antibodies that detect activated Caspase-3-like activity (Fig. 2) (Kuo et al. 2006; Williams et al. 2006). The dendritic pruning process resembles apoptosis in that it includes cytoskeleton disruption, fragmentation, and the cleanup of cellular fragments by phagocytic cells. The localized caspase activation in dendrites may induce apoptotic changes that affect only parts of cells, without executing cell death.

The mechanisms allowing localized caspase activation for dendritic pruning while preventing cell death have recently been elucidated. One study suggested that the molecular subcellular destruction mechanisms involved in developmental neurite pruning and injury-induced neurite degeneration (Wallerian degeneration) are similar (Nikolaev et al. 2009; Schoenmann et al. 2010). Nikolaev et al. (2009) indicated that caspase-6 was required for the axonal pruning in cultured dissociated mouse dorsal root ganglion (DRG) neurons after trophic-factor deprivation, while caspase-3 was not. Schoenmann et al. (2010) demonstrated that the combination of NAD+ supplementation and the inhibition of either caspase-3 or caspase-6 in axons efficiently blocked both caspase-3 or caspase-6 expression and axonal degeneration. Moreover, Schoenmann et al. (2010) demonstrated that the expression of mouse Wld (Wallerian degeneration slow) protein, which mainly consists of the full-length sequence of the NAD+ biosynthetic enzyme Nmnat1, in Drosophila could suppress dendritic pruning in sensory neurons, and caspase activation was still observed even in dendrites that escaped degeneration. Therefore, caspases and the NAD+-sensitive pathway, which operate in parallel to execute the degeneration process, may cooperate in the evolutionarily conserved neurite destruction systems during development and after injury.

Spermatid individualization is another example of localized caspase activation (Fig. 2). During sperm differentiation in Drosophila, 64 haploid spermatids of each cyst are connected by cytoplasmic bridges; these bridges are then removed, and most of the cytoplasm is expelled to form the individual sperm. This process of spermatogenesis is termed individualization. Immunoreactivity for activated caspase-3 is detected in the individualization complex (IC), a cytoskeletal membrane complex that moves along the length of the cyst to the sperm tail (Arama et al. 2003; Huh et al. 2004b). Arama et al. demonstrated that active-caspase (CM1 antibody) (Arama et al. 2003) staining occurs throughout the spermatids, although in a gradient and not only around the IC, and that dark and dronc mutants fail to complete individualization (Arama et al. 2006). Mutants of another initiator caspase, dredd, and of its adaptor molecule dfadd, which can activate Dredd, have partial individualization defects (Huh et al. 2004b), as does a mutant of the executioner caspase drice (Muro et al. 2006).

The Drosophila genome contains two cytochrome c genes, cyt-c-d and cyt-c-p. A mutation in cyt-c-d also causes defects in IC caspase activation and in spermatid individualization (Arama et al. 2003, 2006). Arama et al. indicated that only cyt-c-d is required to activate caspase during spermatid differentiation, whereas cyt-c-p is required for respiration in the soma (Arama et al. 2006). While cytochrome c plays crucial roles in apoptosome-mediated caspase activation in mammals, the existence of a comparable cytochrome c function in Drosophila is still controversial. Cytochrome c is not required to activate caspase in stress-induced apoptosis in Drosophila (Dorstyn et al. 2004); however, specifically cyt-c-d, and not the somatic cyt-c-p, was shown to activate caspase in caspase-dependent SOP development and retinal cell death (Fig. 3) (Arama et al. 2006; Mendes et al. 2006).

To elucidate the mechanisms of cyt-c-d action in IC caspase activation, male sterile mutants were screened against the phenotype for suppressed effector caspase activation at the onset of spermatid individualization (Arama et al. 2007). This screen revealed that a testis-specific Cullin-3-based E3 ubiquitin ligase complex was required for caspase activation in spermatids (Arama et al. 2007). In a similar experiment, Kaplan et al. (2010) identified Soti, which inhibits the Cullin-3-based E3 ubiquitin ligase complex required for caspase activation during spermatid terminal differentiation. They found that Soti competes with an inhibitor of the apoptosis protein dBruce (a target of the E3 complex) to bind Klhl10, the E3 substrate recruitment subunit (Fig. 3). Interestingly, Soti is expressed in a subcellular gradient within spermatids, and it in turn promotes the formation of a similar dBruce gradient, with the result that caspase activation occurs in an inverse graded pattern (Kaplan et al. 2010). These findings provide insight into how specific caspase regulation processes promote caspase-dependent differentiation while preventing cell death.

Apoptosis is defined by its unique morphological changes. Executioner caspase cleaves and activates ROCK-1 kinase, which in turn regulates cytoskeletal rearrangement to create the apoptotic-specific cell shape (Chang et al. 2006). It is therefore not surprising that apoptosis executioner caspase has the potential to regulate other cellular processes with dramatic cytoskeletal changes, such as apoptosis, dendrite pruning and spermatid individualization. However, initiator caspases may regulate the Rac-mediated cytoskeletal changes that are required for cell shaping and migration.

Caspases in cell migration

  1. Top of page
  2. Abstract
  3. Introduction to caspase signaling
  4. Caspases in cell proliferation
  5. Caspases in cell differentiation
  6. Caspases in subcellular compartments
  7. Caspases in cell migration
  8. Caspases in cell shaping
  9. Concluding remarks
  10. Acknowledgments
  11. References

In in vitro cell motility assays, caspase-8-null mouse embryonic fibroblasts (MEFs) are motility-deficient, while caspase-3-null MEFs are not. Cell motility requires calpain activity, and caspase-8-null cells have less calpain activity than wild-type cells. Calpain mediates Rac activation, and caspase-8-null cells activate Rac inefficiently (Helfer et al. 2006). Caspase-8-null MEFs are defective in generating lamellipodia, which is also initiated by Rac activation. Therefore, caspase-8 affects the calpain-mediated cellular migration processes (Helfer et al. 2006). Although caspase-8 promotes motility in MEFs, a loss of caspase-8 expression occurs in metastatic neuroblastoma in mice, and the expression of caspase-8 suppresses metastases (Stupack et al. 2006). Therefore, the caspase-8 loss seen in certain tumors may facilitate tumor invasion.

During Drosophila oogenesis, a group of follicle cells known as border cells migrates to the center of the developing egg chamber (Fig. 2). This phenomenon provides an excellent model for studying cell migration in vivo. The expression of dominant-negative Rac inhibits border cell migration, and the overexpression of DIAP1, but not of p35, rescues the migration defect. While p35 can suppress the executioner caspases Drice and Dcp-1, it does not inhibit the initiator caspase Dronc. Dominant-negative Dronc expression suppresses the migration defects caused by dominant-negative Rac. These results suggest that Dronc inhibits border cell migration downstream of Rac (Geisbrecht & Montell 2004). The expression of DmIKKε, a kinase that promotes DIAP1 degradation, inhibits border cell migration (Fig. 3) (Oshima et al. 2006). This is consistent with the observation that DIAP1 positively regulates border cell migration by downregulating Dronc.

Caspases in cell shaping

  1. Top of page
  2. Abstract
  3. Introduction to caspase signaling
  4. Caspases in cell proliferation
  5. Caspases in cell differentiation
  6. Caspases in subcellular compartments
  7. Caspases in cell migration
  8. Caspases in cell shaping
  9. Concluding remarks
  10. Acknowledgments
  11. References

The shape of the Drosophila antenna arista, a terminal segment of the antenna, is determined by a particular pattern of cells with specific morphologies. Mutant alleles of either diap1 or hid show an abnormal arista pattern (Fig. 2). The thread1 mutant (loss of function mutation of diap1) lacks aristal branching. The hidWR+X1 mutant (loss of function mutation of hid) has large and hairy arista (Cullen & Mccall 2004). As Hid antagonizes DIAP1′s caspase-inhibitory function, caspase activity appears to control arista morphology. DmIKKε, a DIAP1-degrading kinase, is a negative regulator of F-actin polymerization (Fig. 3) (Oshima et al. 2006). Dominant-negative DmIKKε expression causes an excess-branching phenotype that is suppressed by DIAP1 knockdown and enhanced by DIAP1 overexpression. A mild reduction in the expression of dronc and its activator dark by RNAi enhances the lateral branching phenotype caused by dominant-negative DmIKKε. This phenotype is not affected by p35 expression, suggesting that the shape of the arista is regulated by the initiator caspase Dronc, and not by the executioner caspases Drice or Dcp1 (Fig. 2) (Oshima et al. 2006).

DIAP1 protein metabolism is critical for the temporal and quantitative control of caspases, because DIAP1 has a RING finger domain and functions as an E3 ubiquitin ligase. Koto et al. focused on how the balance of caspase roles in cell death and non-apoptotic functions is precisely maintained. They examined the protein turnover of the endogenous caspase inhibitor, DIAP1, which they monitored in the external SOP lineage of living Drosophila with a fluorescent probe PRAP (PRe-Apoptosis signal detecting probe based on DIAP1 degradation) (Fig. 4A). The SOP divides asymmetrically to make the shaft, socket, sheath cells, and then the neurons that innervate each sensory organ (Fig. 4B, C). They found that the DIAP1 level changes dramatically depending on the cell type and maturity (Fig. 4D). The physiological significance of the DIAP1 dynamics during sensory organ development was then studied by manipulating the DIAP1 level in SOP linage cells. Knocking down DmIKKε delayed DIAP1 degradation and caused the shorter, thicker bristle phenotype in the adult notum. On the other hand, either DIAP1 knockdown or Rpr overexpression resulted in bristle loss. These results suggest that the temporal regulation of DIAP1 turnover determines whether caspases function non-apoptotically in cellular shaping, or cause cell death (Koto et al. 2009).

image

Figure 4.  PRAP, a fluorescent probe for monitoring Drosophila inhibitor of apoptosis protein 1 (DIAP1) degradation and DIAP1 dynamics in the sensory organ precursor (SOP) lineage. (A) A schematic representation of PRAP. PRAP has mutations at the caspase-binding site, which prevent DIAP1 from inhibiting caspase activation. (B) External sensory organ. Sensory organs are formed by two external cells, the shaft (sf) and the socket (so) cells, and two internal cells, the internal neuron (n) and the sheath (sh) cell. (C) The SOP lineage. The progenitor cells pI, pIIa, pIIb, and pIIIb (white) divide to form the five cells of the external sensory organ shown in (B). The socket and shaft cells arise from the pIIa cell. The neuron, sheath, and glial (g) cells arise from the pIIb lineage. The glial cell soon dies by apoptosis. (D) PRAP protein dynamics in the SOP lineage at progressive stages of development. In vivo live imaging analysis of PRAP (green) began about 16 h after paparium formation (APF). The time-course of the imaging analysis is shown at the right bottom of the merged panel. Glial cell death is indicated by white arrowheads. Note that in the shaft cell (yellow arrowheads), PRAP vanishes immediately before bristle elongation. Each nucleus is marked by Histone2B-enhanced cyan fluorescent protein (ECFP) (magenta) and is outlined by a dashed line. Scale bar, 5 μm.

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Concluding remarks

  1. Top of page
  2. Abstract
  3. Introduction to caspase signaling
  4. Caspases in cell proliferation
  5. Caspases in cell differentiation
  6. Caspases in subcellular compartments
  7. Caspases in cell migration
  8. Caspases in cell shaping
  9. Concluding remarks
  10. Acknowledgments
  11. References

Caspases transduce irreversible or long-lasting signals through substrate cleavage. Strong caspase activation leads to the cell fate of apoptosis, but localized caspase activation, regulated activation levels, and substrate specificities appear to be involved in providing immunity to apoptosis and determining other cell fates. Caspases therefore have acquired multiple evolutionary activation mechanisms to control their unique signal-transducing roles in both apoptotic and non-apoptotic processes.

Acknowledgments

  1. Top of page
  2. Abstract
  3. Introduction to caspase signaling
  4. Caspases in cell proliferation
  5. Caspases in cell differentiation
  6. Caspases in subcellular compartments
  7. Caspases in cell migration
  8. Caspases in cell shaping
  9. Concluding remarks
  10. Acknowledgments
  11. References

I apologize to colleagues whose work could not be cited due to space limitations. I thank M. Miura and A. Koto for their help in preparing the figures. Studies by our group were supported in part by grants from the Takeda Science Foundation, the Uehara Memorial Foundation, the Kanae Foundation for the Promotion of Medical Science, and the Japanese Ministry of Education, Science, Sports, Culture and Technology.

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  8. Caspases in cell shaping
  9. Concluding remarks
  10. Acknowledgments
  11. References
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