Báo cáo khoa học: Caspase-2 is resistant to inhibition by inhibitor of apoptosis proteins (IAPs) and can activate caspase-7

Caspase-2 is resistant to inhibition by inhibitor of apoptosis proteins (IAPs) and can activate caspase-7 Po-ki Ho1,2,3, Anissa M. Jabbour1,2,3, Paul G. Ekert1,4,5 and Christine J. Hawkins1,2,3

1 Murdoch Children’s Research Institute, Parkville, Australia 2 Children’s Cancer Centre, Royal Children’s Hospital, Parkville, Australia 3 Department of Paediatrics, University of Melbourne, Parkville, Australia 4 Department of Neonatology, Royal Children’s Hospital, Parkville, Australia 5 The Walter and Eliza Hall Institute, Royal Melbourne Hospital, Parkville, Australia

Keywords caspase-2; protease; caspase-7; S. cerevisiae; enzyme activity

Caspases are a family of cysteine proteases with roles in cytokine matur- ation or apoptosis. Caspase-2 was the first pro-apoptotic caspase identified, but its functions in apoptotic signal transduction are still being elucidated. This study examined the regulation of the activity of caspase-2 using recombinant proteins and a yeast-based system. Our data suggest that for human caspase-2 to be active its large and small subunits must be separ- ated. For maximal activity its prodomain must also be removed. Consistent with its proposed identity as an upstream caspase, caspase-2 could provoke the activation of caspase-7. Caspase-2 was not subject to inhibition by members of the IAP family of apoptosis inhibitors.

Correspondence C. Hawkins or P. Ekert, Murdoch Children’s Research Institute, Royal Children’s Hospital, Flemington Road, Parkville, VIC 3052 Australia Fax: +61 3 9345 4993 (CH); +61 3 9347 0852 (PE) Tel: +61 3 9345 5823 (CH); +61 3 9345 2548 (PE) E-mail: chris.hawkins@mcri.edu.au; paul.ekert@mcri.edu.au

(Received 10 November 2004, revised 7 January 2005, accepted 18 January 2005)

doi:10.1111/j.1742-4658.2005.04573.x

caspase family, the function of caspase-2 (Nedd-2 ⁄ Ich-1) remains somewhat elusive. Its substrate prefer- ence more closely aligns with that of the pro-apoptotic caspases than their cytokine processing relatives [5]. Of the mammalian caspases, caspase-2 is the most similar to the nematode apoptotic caspase, CED-3. This would also tend to imply that caspase-2 plays a pro- apoptotic role, yet caspase-2 deficient mice have an extremely subtle phenotype, arguing against a non- redundant role in programmed cell death [6,7].

has

recently

received

Caspase-2

The caspases are a family of cysteine proteases that typically cleave their substrates at aspartate residues [1]. Subclassification of family members has been based on various criteria including substrate specificity or structural features. For example, caspases-1, -4 and -5 are involved in the proteolytic maturation of cytokines including pro-interleukin-1b [2] and pro-interleukin-18 [3]. Caspases-8 and -9 are components of cell death signal transduction pathways and are classified as api- cal caspases. The primary role of these proteases, each of which has a long prodomain containing a protein is to proteolytically activate distal interaction motif, caspases (such as caspase-3 and caspase-7), which then catalyse the cleavage of numerous cellular substrates [4]. Despite being the second identified member of the

considerable attention, as several groups have sought to define its biological role in apoptosis signalling. Overexpressing caspase-2 provoked the release of pro-apoptotic mole- cules (including cytochrome c) from mitochondria [8],

Abbreviations AFC, 7-amino-4-trifluoromethyl coumarin; CARD, caspase activation and recruitment domain; GST, glutathione-S-transferease.

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found that

known caspase inhibitors and the ability of caspase-2 to activate effector caspases. In addition, we explored the relationship between proteolytic processing of caspase-2 and its enzymatic activity. Our data suggest that processing of human caspase-2 is required for maximal activity. Unlike other caspases, caspase-2 could not be inhibited by mammalian inhibitor of apoptosis proteins (IAPs). Caspase-2 was able to acti- vate caspase-7, suggesting that caspase-2 can function as an apical caspase.

Results

whilst diminished caspase-2 expression or a peptide caspase-2 inhibitor blocked etoposide-induced cyto- chrome c release from mitochondria [9]. This suggests that caspase-2 may function upstream of the mitoch- ondrial changes associated with stress-induced apopto- sis. This could be recapitulated in vitro [10] and has been proposed to occur via direct caspase-2-mediated permeabilization of mitochondrial membranes [11]. Lassus et al. suppression of caspase-2 expression provided equivalent protection to that con- ferred by Apaf-1 downregulation, against apoptosis induced by DNA damage [12]. The involvement of caspase-2 in TRAIL-induced apoptosis has also been reported recently, placing this enzyme upstream of Bid cleavage in the pathway [13].

High level expression of pro-caspase-2 is lethal in yeast

Like caspase-9, caspase-2 bears a caspase activation and recruitment domain (CARD) in its amino-terminal prodomain. The role of the CARD (in caspase-9 at least) is to permit binding to aggregated adaptor pro- teins, leading to autoactivation through ‘induced proxi- mity’ [14]. Consistent with this, forced dimerization of caspase-2 provoked its activation [15], and fusing the caspase-2 prodomain to caspase-3 resulted in caspase-3 autoactivation [16]. Recent findings by Baliga et al. indicated that dimerization is the key determinant for initial activation of murine pro-caspase-2 [17]. The phy- siological mechanism through which the prodomain might trigger activation of caspase-2 is still unclear. A molecular pathway has been proposed to link caspase- 2 to members of the tumour necrosis factor receptor family via an adaptor molecule (RAIDD ⁄ CRADD) and intermediaries (RIP, TRADD, FADD and TRAFs) [18,19]. However, this has not been directly demonstrated and death ligand-mediated apoptosis proceeds normally in caspase-2-deficient cells [7]. Other putative caspase-2 adaptors have been proposed [20,21], but verification of their relevance in physiologi- cal settings has not yet been published. Tinel and Tschopp recently reported a complex they designated the ‘PIDD-osome’ comprising caspase-2, RAIDD and PIDD, the formation of which promoted apoptosis following p53-dependent DNA damage [22]. Further, caspase-2 is recruited into a high molecular weight complex independent of the apoptosome components Apaf-1 and cytochrome c [23]. It has also been recently postulated that caspase-2 may influence apoptosis [24] and ⁄ or nuclear factor-jB activation [25] through mech- anisms unrelated to its enzymatic activity.

Properties of caspase-2 were assessed using a yeast- based system we have previously exploited to character- ize other caspases and apoptotic pathways [26–28]. This system capitalizes on the observation that some caspases kill yeast upon enforced high-level expression. In order for caspases to kill yeast, they must both be able to autoactivate and their proteolytic specificity must permit cleavage of essential yeast proteins. To assess the activity of caspase-2 in yeast, various con- structs encoding different forms of the protein (Fig. 1) were transformed into yeast (Fig. 2A). Expression of pro-caspase-2 using the Gal 1 ⁄ 10 promoter affected yeast growth only marginally (compare growth in lane 2 to that of an empty vector transformant in lane 1). Increasing the pro-caspase-2 expression level, by intro- ducing an additional expression construct under dif- ferent nutritional selection, elicited more substantial lethality (lane 3). A caspase-2 cleavage site mutant (D152A), from which the prodomain could not be removed, was also expressed at a high level using two plasmids with different nutritional selections. Com- pared with equivalent expression of wild-type pro- caspase-2 (lane 3), this mutant exhibited only marginal toxicity (lane 4) suggesting that removal of the prodo- main contributes to full enzymatic activity. Consistent with this observation, a truncation mutant lacking almost all of the caspase-2 prodomain (caspase-2D1)149) killed yeast more efficiently than full-length caspase-2 (compare lane 7 with lane 2). An artificially autoacti- vating version of caspase-2 (rev-caspase-2), in which the small subunit precedes the prodomain and large subunit [29], killed yeast readily (lane 6). The catalyti- cally inactive mutant pro-caspase-2C303A was unable to kill yeast (lane 5) implying that the lethality of wild- type caspase-2 in yeast was due to its enzymatic activity. The expression of the prodomain (caspase-2D150)435) had no effect on yeast viability (lane 8).

If caspase-2 functions as an apical caspase, it may process and activate downstream caspases. We sought to characterize the molecular events downstream of human caspase-2 activation. In particular we focused on the susceptibility of caspase-2 to suppression by

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Fig. 1. Schematic illustration of the caspase- 2 proteins used in this study. Mutated resi- dues are listed above wild-type caspase-2 and are depicted with black circles.

substrate to assess caspase-2 activity [5]. VDVADase activity was detected in lysates from yeast expressing all forms of caspase-2 that were capable of autoprocessing (Fig. 2C). The most lethal forms of caspase-2 had the highest VDVADase activity (lanes 3, 6 and 7), while ly- sates from yeast that survived (lanes 1, 5 and 8) did not cleave the peptide substrate. Yeast transformed with one wild-type caspase-2 plasmid or the D152A mutant were killed only inefficiently, however, their lysates exhibited significant VDVADase activity. This may indi- cate that the biochemical assay is a more sensitive meas- ure of caspase-2 activity than the yeast death assay.

Caspase-2 is not inhibited by mammalian IAP proteins

To investigate the auto-processing of pro-caspase-2 in yeast, we immunoblotted lysates obtained from yeast expressing these different forms of caspase-2 with an antibody recognizing an epitope in the large subunit. In lysates from yeast expressing wild-type pro-caspase-2, a partial cleavage product was detected, in addition to the fully processed large subunit (Fig. 2B). Like the wild-type enzyme, the cleavage site mutant pro-caspase-2D152A was processed efficiently between the large and small subunits, however, the mutation at D152 prevented it from being further processed to separate the prodomain from the large subunit. Caspase-2C303A remained intact as a result of the abolished catalytic activity. Rev-caspase-2, despite its ability to efficiently kill yeast, was only incompletely processed. A proportion of caspase-2D1)149 was cleaved to remove the small subunit, thereby permitting detec- tion of the dissociated large subunit.

The activities of these different forms of caspase-2 were also analysed biochemically using a fluorogenic caspase-2 substrate. In this assay, the activity of an enzyme is reflected by the efficiency with which it cleaves the substrate to release free 7-amino-4-trifluoro- methyl coumarin (AFC). The caspase-2-specific fluoro- genic synthetic peptide Z-VDVAD-AFC was used as a

Members of the mammalian IAP family contribute to the regulation of apoptotic pathways in part by their inhibition of caspases-3, -7 and -9 [30]. Other mamma- lian caspases (-1, -6, -8 and -10) are known to be resist- ant to inhibition by IAPs [30], but the susceptibility of caspase-2 to direct inhibition by IAPs has not been reported to date. To explore the sensitivity of caspase-2 to IAP inhibition, we tested whether coexpression of IAPs would suppress caspase-2-dependent yeast death.

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A

B

C

Fig. 2. Caspase-2 kills yeast. (A) A semi- quantitative assay compares the effect of transgenes on yeast growth and viability. Yeast cells were transformed with the indi- cated plasmids. Suspensions of each trans- formant were prepared at standardized concentrations. Serial dilutions were made and spotted onto solid inducing minimal media vertically down the plate. Colony size indicates growth rate and colony number reflects cell viability. (In every experiment, each dilution was also spotted onto a repressing plate to verify that equivalent numbers of each transformant were spot- ted; data not shown). (B) Anti-caspase-2 immunoblotting of lysates from the indica- ted transformants. The presumed identities of each band are shown to the left (pro, pro- domain; L, large subunit; S, small subunit). (C) The ability of caspase-2 to cleave the flu- orogenic peptide substrate Z-VDVAD-AFC. Native lysates obtained from yeast were incubated with Z-VDVAD-AFC. Fluorescence was monitored over time and the maximal rate of increase in free AFC was calculated and graphed. Error bars indicate SD (n ¼ 4).

peptide substrate Z-VDVAD-AFC. Caspase-2 activity was not affected by the presence of XIAP (Fig. 3D), whereas XIAP significantly reduced the activity of caspase-3, as demonstrated previously [33]. The pres- ence of p35 led to a decrease in both caspase-2 and caspase-3 activities. Inactive mutants of p35 (p35D87A) and XIAP (XIAPD148A) were unable to inhibit either caspase.

Caspase-2 can promote caspase-7 catalytic activity

We had previously established that the inhibitors p35 and p49 could rescue yeast from caspase-2 mediated death [31], so these baculoviral proteins were used as positive controls. Caspase-3 effectively killed yeast and this could be blocked by XIAP (also known as hILP), MIHB (cIAP-1 ⁄ hIAP-2 ⁄ BIRC2) and MIHC (cIAP- 2 ⁄ hIAP-1), as well as p35 and p49 (Fig. 3A). In contrast, the mammalian IAPs could not inhibit yeast death induced by expression either of full-length pro-caspase-2 (Fig. 3B) or of truncated caspase-2 lacking the prodo- main (Fig. 3C). As expected, the baculoviral caspase inhibitors p35 and p49 protected caspase-2-expressing yeast (Fig. 3B,C).

To explore the potential for caspase-2 to functionally interact with other caspases, we exploited the dose- dependent caspase-2-mediated yeast toxicity illustrated in Fig. 2. Caspase-2 was coexpressed in yeast from a single plasmid either alone (yielding weak lethality) or

To confirm these observations using a biochemical approach, purified caspase-2 was mixed with recombin- ant XIAP or the inactive mutant XIAPD148A [32], then assayed for its ability to cleave the fluorogenic penta-

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D

B

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Fig. 3. IAPs do not inhibit caspase-2. The caspase expression plasmids used to kill yeast were (A) Caspase-3-lacZ (B) pGALL- (LEU2)-caspase-2 with pGALL-(URA)-cas- pase-2 or (C) pGALL-(URA)-caspase-2D1)149. Yeast transformed with the indicated plas- mids were spotted as described in the legend to Fig. 1. (D) The indicated combina- tions of caspase, fluorogenic substrate and inhibitor were mixed together and the fluor- escence resulting from the caspase-medi- ated substrate cleavage was monitored and calculated as described in the legend to Fig. 1. Error bars indicate SD (n ¼ 3).

[35]

together with the nonlethal caspases-3, -4, -6, -7 and -9 (Fig. 4A). Yeast death was used as an indicator of caspase activity. Co-expression of caspase-2 with caspase-7 led to a pronounced increase in yeast death, compared to that triggered by either caspase alone (compare lane 12 with lanes 2 and 11). Much weaker synergy was also reproducibly observed between caspase-2 and -3 (compare lane 6 with lanes 2 and 5). We then tested the ability of lysates from these yeast to cleave a fluorogenic caspase-3 substrate (Ac-DEVD- AFC) or a caspase-2 substrate (Z-VDVAD-AFC). Caspase-2 activity was not enhanced by coexpression of caspases-3 or -7. However, significantly more clea- vage of Ac-DEVD-AFC was observed when caspase-2 was coexpressed with caspase-7 (or, to a lesser extent with caspase-3) (Fig. 4B).

lanes 2 and 4 in Figs 5A–C). Full length caspase-7 was unprocessed and did not kill yeast (lane 9), whereas caspase-7 coexpressed with caspase-2 was activated and toxic to yeast (lane 5). The activation of caspase-7 by caspase-2 depended on caspase-2 catalytic activity since coexpression of catalytically inactive caspase-2 with caspase-7 did not yield enzymatic activity (neither VDVADase nor DEVDase) and did not kill yeast (Figs 5A–C, lane 6). However, caspase-2 activation was independent of caspase-7 as caspase-2 proteolytic activity was the same in the presence of active or enzy- matically inactive caspase-7 (compare Fig. 5B and C lanes 5 and 7). Two positive controls were used for caspase-7 activation. First, caspase-7D1)53, which lacks the prodomain region and is constitutively active in mammalian cells [34] and in yeast (lane 10). Second, as previously reported for caspase-3 [27], caspase-7 was activated by a constitutively active (lane 11). This autoacti- caspase-9 (rev-caspase-9) vating caspase-9 protein, which could activate caspase- 3 [27] or caspase-7 (Fig. 5A–C, lane 11), was not able to co-operate with caspase-2 to kill yeast (Fig. 4A, lane 14). Together, these data suggest that caspase-2 may lie upstream of caspase-7, and not downstream of caspase-9, in apoptotic pathways.

To further investigate the apparent synergy between caspase-2 and caspase-7, plasmids encoding different forms of these enzymes were transformed into yeast in various combinations and their effects on enzyme clea- vage, enzyme activity and yeast growth determined (Fig. 5). As before, high level expression of caspase-2 resulted in an active enzyme, able to efficiently kill yeast, whereas lower expression levels of caspase-2 had in vitro activity but weak killing activity (compare

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Fig. 4. Co-expression of caspases-2 and -7 enhances yeast lethality. (A) The indicated combinations of caspases were coexpre- ssed in yeast and their ability to promote yeast death was compared to the lethality arising from expression of single caspases. (B) The activities of caspases were assayed in lysates from yeast expressing individual caspases, or the indicated combinations of caspases. Substrate cleavage was calcula- ted from the maximal rate of free AFC released through cleavage by 30 lg of yeast lysate. Error bars indicate SD (n ¼ 3).

The relationship between caspase-2 processing and enzymatic activity

determine

(Fig. 6B)

enzyme

to

Previous work had illustrated that human pro- caspase-2 can be processed at residue D152 to remove its prodomain, and at residues D316 and D330 to dissociate the large and small subunits and release a small linker peptide [36]. To investigate the impact of these cleavage events on the enzymatic activity of caspase-2, recombinant caspase-2 proteins harbouring one or a combination of mutated D152, D316 or D330 residues were generated and expressed in bacteria and the protein purified (Figs 1 and 6A).

We observed that full length recombinant caspase-2 has about a 10-fold lower activity than a commonly used amino-terminal truncation lacking most of the prodomain (D1–149) [37] (Fig. 6B). We therefore used this truncated caspase-2 to test the effects on activity of mutating the D152, D316 and D330 residues. We immunoblotted the purified caspase-2 enzymes with an antibody recognizing an epitope within the large subunit of caspase-2, to determine whether enzyme autoprocessing had occurred (Fig. 6A) and then tes- ted cleavage of a caspase-2 specific fluorogenic sub- strate activity. Retention of at least one cleavage site between the

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Fig. 5. Caspase-2 activates caspase-7 in yeast. (A) The indicated plasmids were transformed into yeast and transformants spotted onto inducing medium to visualize their impact on yeast growth. (B) Immuno- blotting was used to detect caspase pro- cessing. The presumed identities of each band are shown to the left (pro, prodomain; L, large subunit; S, small subunit). (C) The abilities of the yeast lysates to cleave the caspase-2 (Z-VDVAD-AFC) and caspase-7 (Ac-DEVD-AFC) substrates were assayed. Substrate cleavage was calculated from the maximal rate of free AFC released through cleavage by 30 lg of yeast lysate. Error bars indicate standard deviations (n ¼ 3).

it was evident

large and small subunits (D316 and ⁄ or D330) permit- ted autocatalytic separation of the subunits (Fig. 6A) and yielded active enzymes (Fig. 6B). Fusion of the linker to the small subunit (D330A) had a slightly greater deleterious effect on enzyme activity than (Fig. 6B). In fusion to the large subunit

(D316A)

contrast, mutation of both D316 and D330 sites abolished auto-processing (Fig. 6A) and dramatically reduced enzymatic activity (Fig. 6B). Using higher amounts of enzyme (100 nm), that mutation of both of these cleavage sites decreased activity by 840-fold (data not shown). The C303A

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Fig. 6. Caspase-2 processing is necessary for activation. (A) Caspase-2 enzymes with the indicated mutations were generated in bacteria and immunoblotted to determine the extent of auto-processing. The pre- sumed identities of each band are shown to the right (pro, prodomain; L, large subunit; S, small subunit). (B) The abilities of wild- type recombinant caspase-2 or the indicated mutants to cleave the fluorogenic substrate Z-VDVAD-AFC were monitored as described in previous legends. Two independent prep- arations of each enzyme were used. (C) The indicated plasmids encoding wild-type or cleavage site mutants of casapse-2 (or empty vectors) were transformed into yeast and transformants spotted onto inducing medium to visualize their impact on yeast growth.

caspase-2 correlates with the degree to which the large subunit is separated from the small subunit.

active site mutant was completely inactive (Fig. 6B), even at 100 nm (data not shown). Full length clea- vage site mutants were expressed from two plasmids in yeast and their impact on yeast viability assayed. The D316A and D330A single mutants were toxic to yeast, however, yeast expressing the double D316, 330A mutant survived (Fig. 6C). Together, these data activity of human suggest

the proteolytic

that

We also tested the abilities of the caspase-2 mutant enzymes to cleave protein substrates. Cellular sub- strates (Bid, PARP, catalytically inactive pro-caspase-2 and pro-caspase-7) were expressed as glutathione- S-transferease (GST)-fusion proteins, incubated with caspase-2 enzymes and subjected to the various

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Discussion

the

cleavage product

A unique merit of the yeast system used here is that it is free from the potential interference of other mamma- transduction pathway compo- lian apoptotic signal nents, allowing the expression of the gene of interest in a naive yet eukaryotic cell-based environment. We have previously used this system to reconstitute caspase-9 activation by Apaf-1 [27] and the core nematode pro- grammed cell death pathway [28]. Here, we harnessed this system to analyse the regulation of caspase-2 activ- ity, exploiting the observation that overexpressed caspase-2 kills yeast in a concentration dependent man- ner, requiring a catalytically active enzyme. Purified, recombinant proteins were also used to verify much of the data generated from the yeast system.

SDS ⁄ PAGE. Cleavage of the substrates was assessed by staining with Coomassie blue (Fig. 7A) and immu- noblotting (Fig. 7B). All caspase-2 proteins that were active in the fluorogenic assay (Fig. 6B) were also able to cleave Bid at aspartate 60 [10] and a catalytically inactive GST-tagged pro-caspase-2 (Fig. 7A). The size of implied that processing occurred between the large and small subunits. PARP was cleaved by caspases-3 and -7, but not by caspase- 2. GST-tagged pro-caspase-2C303A was not processed by caspases-3, -7 or -8; cleavage products were not detected by Coomassie blue staining (Fig. 7A) or by immunoblotting (Fig. 7B). (Processing of PARP or Bid by these enzymes confirmed that they were active). Having observed the activation of caspase-7 by caspase-2 in yeast, we examined the processing of GST-tagged pro-caspase-7C186A by caspase-2 in this system. The cleavage of GST-pro-caspase-7C186A by active caspase-2 (as well as by caspases-3 and -8) was detected by immunoblotting with an antibody that recognizes cleaved caspase-7 (Fig. 7B).

A

for

B

We have shown that prodomain removal increases caspase-2 activity, when expressed in yeast or in bac- teria. For generation of active human caspase-2, pro- cessing is also required between the small and large subunits (at D316 and ⁄ or D330). Mutation of either site had little effect on enzyme activity or toxicity to yeast but mutation of both sites abolished both enzyme activity and yeast killing. These observations differ somewhat from previously reported analyses of the human and mouse murine caspase-2. Firstly, enzymes vary in their propensity for autoprocessing between the large subunit and the linker. We have shown that human caspase-2 almost completely auto- processes at this point (D316), as indicated by the effi- cient separation of large and small subunits of the D330A mutant. In contrast, mutation of the murine equivalent of the human residue D316 (D333) alone prevented autoprocessing of caspase-2 [17,38]. This species difference persisted when the mouse and human mutants were generated using the identical bac- terial expression systems (B. Baliga and S. Kumar, personal communication), ruling out any technical explanations the variation. Secondly, human caspase-2 which was prevented from autoprocessing between the large and small subunits was almost totally inactive, however, the D333G mutant of murine caspase-2 that could not autoprocess retained about one-fifth of wild-type enzyme activity [17,38]. Further investigations will hopefully clarify the mechanisms underlying these curious species differences.

Fig. 7. Substrate cleavage by wild-type caspase-2, its cleavage site mutants and other caspases. (A) GST-tagged, enzymatically inactive pro-caspase-2 or caspase substrates were incubated with the indi- cated purified recombinant caspases (as detailed in the experimen- tal section). The reactions were then subjected to SDS ⁄ PAGE and the gels stained with Coomassie blue to visualize cleavage. (B) The more sensitive technique of immunoblotting was used to detect cleavage of catalytically inactive pro-caspase-2 or pro-caspase-7.

Coexpression of caspase-2 with caspase-7 in yeast was significantly more toxic than expression of either protein alone. Although this result could reflect an additive effect of two mildly lethal stimuli, two pieces of evidence suggest that caspase-2 activation of caspase-7 accounts for the combined lethality. Firstly, caspase-2 cleaved caspase-7 in vitro. Secondly, lysates from yeast

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Experimental procedures

Plasmid construction

expressing caspases-2 and -7 had higher DEVDase activity (indicative of caspase-7 activity) than those from yeast only expressing caspase-7. However, the VDVADase activity (reflecting caspase-2 activity) of the lysates from double transformants was similar to that of lysates from yeast only expressing caspase-2.

pGALL-(LEU2)-caspase-4,

Previously published data demonstrated that caspa- ses-1, -2, -3, -4, -6, -7, -11 and CED-3 could all cleave caspase-2, to varying extents [36,38–40]. In contrast, our data indicates that concentrations of caspases-3, -7 and -8 capable of efficiently processing known physio- logical substrates (PARP or Bid) could not cleave an inactive mutant of pro-caspase-2. This discrepancy probably relates to differences in the relative concen- trations and ⁄ or purity of the enzymes and substrates used. The studies cited above used either unspecified amounts of unpurified enzyme or enzyme concentra- tions four times [40] or over 11 times [36] that used here. In the previous studies, reticulocyte lysates con- taining 35S-labelled wild-type caspase-2 were used as substrates. These lysates would contain endogenous reticulocyte proteins that may potentially influence the processing of caspase-2. To avoid any such indi- rect effects, we used purified, catalytically inactive caspase-2 as a substrate.

pGALL-(URA)-caspase-2C303A,

from apoptosis

cells

produce to

The IAP family of apoptosis inhibitors exert their pro-survival effect, at least in part, through suppres- sion of caspases-3, -7 and -9 [33,41,42]. The IAPs XIAP, MIHB and MIHC could not inhibit other casp- ases including -1, -6, -8 and -10 [30], but their ability to directly inhibit caspase-2 has not been previously published. Caspase-2-dependent yeast death was unaf- fected by coexpression of XIAP, MIHB and MIHC although, as we previously reported [31], p35 and p49 could inhibit caspase-2 in this system. Furthermore, XIAP, the most potent caspase inhibitor of the IAP family, did not impede the ability of recombinant caspase-2 to cleave a synthetic substrate. It was previ- ously observed that IAPs partially protected tissue cul- ture induced by caspase-2 overexpression [43]. In the light of our findings, this inefficient protection was probably due to IAP-medi- ated inhibition of caspase-7, which was likely activated by the overexpressed caspase-2.

fragment a SalI ⁄ BamHI

In summary, this study illustrated that, at least in the absence of an activating adaptor, generation of active human caspase-2 requires separation of its large and small subunits. In the context of autoactivation, removal of the prodomain also enhances proteolytic activity. Caspase-2 can act as an apical caspase, pro- moting the activation of caspase-7. Unlike caspases-3, -7 and -9, caspase-2 was resistant to inhibition by members of the IAP family.

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or and For expression in yeast, coding regions of human genes were cloned into the pGALL yeast vectors under the regu- the inducible Gal 1 ⁄ 10 promoter [26]. Yeast lation of vectors pGALL-(HIS3), pGALL-(LEU2) and pGALL- (URA) have been described previously [27,35]. Plasmids pGALL-(LEU2)-caspase-2, pGALL-(URA)-caspase-2, casp- pGALL-(URA)- ase-3-LacZ, caspase-7D1)53, pGALL-(HIS3)-p35 and pGALL-(HIS3)- p49 have been reported [27,31,35]. Other plasmids were constructed as follows: Pro-caspase-2 PCR product, gener- ated with primers 1 and 2, was cut with BglII ⁄ XbaI and ligated into BamHI ⁄ XbaI cut vectors to produce pGALL- (HIS3)-caspase-2 and pGALL-(HIS3)-FLAG-caspase-2. To make pGALL-(LEU2)-rev-caspase-2, the carboxyl terminal fragment was amplified with primers 3 and 4, digested with BglII ⁄ XbaI and ligated into pGALL-(LEU2) to give pGALL-(LEU2)-rev-caspase-2-C. The amino-terminal frag- ment was generated with primers 5 and 6, cut with XhoI ⁄ XbaI, and ligated into pGALL-(LEU2)-rev-caspase-2-C to generate the final construct. pGALL-(LEU2)-caspase- 2C303A, pGALL-(HIS3)- caspase-2C303A and pGALL-(HIS3)-FLAG-caspase-2C303A were produced by replacing a SpeI ⁄ BamHI cut fragment with a PCR product generated with primers 1 and 7. pGALL-(HIS3)-FLAG-caspase-2D1)149 was cloned by ligat- ing a NdeI-digested and blunt-ended then BamHI cut frag- ment from pET23a-caspase-2D1)149 into SpeI-digested and blunt-ended then BamHI cut pGALL-(HIS3)-FLAG-ca- spase-2. A PCR product generated with primers 1 and 8 was cut with BglII ⁄ XbaI and ligated into BamHI ⁄ XbaI cut pGALL-(LEU2)-caspase-2D153)435. vector pGALL-(HIS3)-FLAG-caspase-2D152A and pGALL-(HIS3)- FLAG-caspase-2D316A were made by replacing a SalI ⁄ BamHI cut fragment with PCR products generated with primer pairs 9, 10 and 11, 12, respectively. pGALL-(HIS3)- FLAG-caspase-2D330A was produced by replacing a Bam- HI ⁄ XbaI cut fragment with a PCR product generated with primers 13 and 14. To make pGALL-(HIS3)-FLAG- caspase-2C303A; D152,316A, in pGALL-(HIS3)-FLAG-caspase-2C303A was replaced with a PCR product generated using primers 9 and 12. It was sub- sequently used to produce pGALL-(HIS3)-FLAG-cas- pase-2 C303A; D152, 316, 330Aby replacing aSalI ⁄ BamHI fragment into pGALL-(HIS3)-FLAG-caspase-2D330A. SpeI ⁄ BamHI fragments isolated from pGALL-(HIS3)-FLAG-Caspase- 2D1)149 or pGALL-(HIS3)-FLAG-Caspase-2D152A were used to replace part of the coding region in pGALL-(HIS3)- Caspase-2, pGALL-(URA)-Caspase-2 and pGALL-(LEU2)- Caspase-2 to make pGALL-(HIS3)-Caspase-2D1)149 and pGALL-(URA)-Caspase-2D1)149 pGALL-(HIS3)- pGALL-(LEU2)-Caspase-2D152A, Caspase-2D152A

P.-k. Ho et al.

Caspase-2 can activate caspase-7 and is resistant to IAPs

pGALL-(HIS3)-Caspase-2D316A,

pGALL-(URA)-Caspase-2D330A,

generated

and into SpeI ⁄ XbaI

primers and 15

and pGALL-(HIS3)-p35D87A as a template [31]. Using a template kindly provided by John Silke that contained XIAPD148A, the coding region of XIAP was amplified with primers 28 and 29, cut with BamHI ⁄ EcoRI and inserted into pGEX-6P3 to give pGEX6P3-XIAPD148A. To construct pGEX6P3-BidD60A, first a PstI-digested PCR product amplified with primers 30 and 31 was used to replace an internal fragment in pBluescriptII(SK+)-Bid [46] to gener- ate pBluescriptII(SK+)-BidD60A. The coding region was then amplified with primers 32 and 33, digested with Bam- HI ⁄ EcoRI and cloned into pGEX-6P3 to give the final construct. A SpeI ⁄ XbaI fragment excised from pGALL- (LEU2)-caspase-2C303A was ligated into pBluescriptII(SK+) to produce pBluescriptII(SK+)-caspase-2C303A. A blunt- ended SpeI-cut then NotI-digested fragment isolated from the above construct was ligated into a blunt-ended EcoRI- cut then NotI-digested vector to yield pGEX6P3-caspase- 2C303A. The coding region of truncated PARP was released with EcoRI ⁄ NotI from pADH-(TRP1)-mycPARPD338)1013 [47] and ligated into pGEX-6P3. The construct was then cut with BamHI, blunt-ended and re-ligated to produce pGEX6P3-mycPARPD338)1013. A BamHI ⁄ XbaI fragment was excised from pGALL-(URA)-Caspase-7C186A and cloned into pBluescriptII(SK+) to give pBluescriptII(SK+)- Caspase-7C186A, from which a BamHI ⁄ NotI fragment was isolated and ligated into pGEX-6P3 to yield pGEX6P3- Caspase-7C186A.

pGALL- respectively. (HIS3)-Caspase-2D330A, pGALL-(HIS3)-Caspase-2D316,330A, pGALL-(HIS3)-Caspase-2D152,316,330A, pGALL-(URA)-Cas- pase-2D316A, pGALL- (URA)-Caspase-2D316,330A and pGALL-(URA)-Caspase- 2D152,316,330A were ligating SpeI ⁄ XbaI by fragments released from pGALL-(HIS3)-FLAG-caspase- 2D316A, pGALL-(HIS3)-FLAG-caspase-2D330A, pGALL- (HIS3)-FLAG-caspase-2D316,330A pGALL-(HIS3)- FLAG-caspase-2D152,316,330A digested pGALL-(HIS3)-Caspase-2 and pGALL-(URA)-Caspase-2, respectively. The coding regions of pro-caspases-3, -6 and -7 were excised with BamHI ⁄ XbaI from pGALL-(URA)-cas- pase-3 [27], pEF-Mch2 (made from a vector kindly provided by E. Alnemri [44]) and pCUP1-(LEU2)-caspase-7 [35] and ligated into pGALL-(LEU2) to generate pGALL-(LEU2)- caspase-3, pGALL-(LEU2)-caspase-6 and pGALL-(LEU2)- caspase-7, respectively. pGALL-(URA)-caspase-7 was made by ligating the coding region of caspase-7 cut with Bam- HI ⁄ XbaI into pGALL-(URA). To make pGALL-(URA)-cas- pase-7C186A, the carboxyl-terminal fragment was amplified digested with BamHI ⁄ with 14, XbaI and ligated into pGALL-(URA) to give pGALL- (URA)-caspase-7C186A-C. The amino terminal fragment was generated with primers 16 and 17, cut with BamHI ⁄ XhoI, and ligated into pGALL-(URA)-caspase-7C186A-C to give the final construct. pGALL-(HIS3)-XIAP and pGALL-(HIS3)- MIHB were constructed by ligating EcoRI ⁄ NotI cut PCR products amplified with primer pairs 18, 19 and 20, 21, into pGALL-(HIS3). The coding region of respectively, MIHC was isolated from pADH-(TRP1)-MIHC [27] with EcoRI ⁄ NotI and cloned into pGALL-(HIS3) to give pGALL-(HIS3)-MIHC.

and pET23a-caspase2D1)149

and

pET23a-caspase-2D1)149 fragment internal in

pET23a-caspase2D1)149;D316A,

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1411

For expression in bacteria, coding regions of genes were cloned into pET23a(+) (Novagen, Madison, WI, USA) or pGEX6P-3 (Amersham Biosciences, Uppsala, Sweden). pGEX6P3-XIAP [45], pGEX6P3-Bid [46] and pET23a-p35 [28] have been previously described. The coding region of pro-caspase-2 was amplified with primers 21 and 22, cut with NdeI ⁄ XhoI and ligated into pET23a to give pET23a- pET23a-casp- caspase2. ase2D1)149; D152A were constructed using NdeI ⁄ XhoI cut PCR products amplified with primer pairs 24, 23 and 25, 23, respectively. HindIII ⁄ EcoRI fragments released from pGALL-(LEU2)-caspase-2C303A, pGALL-(HIS3)-FLAG- pGALL-(HIS3)-FLAG-caspase-2D330A, caspase-2D316A, pGALL-(HIS3)-FLAG-caspase-2D316,330A pGALL- (HIS3)-FLAG-caspase-2D152,316,330A were used to replace or an pET23a-caspase-2D1)149;D152A to generate pET23a-caspase- 2D1)149;C303A, pET23a- caspase2D1)149;D330A, pET23a-caspase2D1)149;D316,330A and respectively. pET23a-caspase2D1–149; D152 316 330 A, pET23a-p35D87A was constructed by ligating a NdeI ⁄ HindIII cut PCR product generated with primers 26 and 14 The primers used were: 1, 5¢-GGAAGATCTACTAG TATGGCCGCTGACAGGGGACGC-3¢; 2, 5¢-GCTCTAG ACTATGTGGGAGGGTGCCTTGGG-3¢; 3, 5¢-GCAGAT CTATGGACCAACAAGATGGAAAG-3¢; 4, 5¢-CGTCT AGACTCGAGTCCATCTTGTTGGTCTGTGGGAGGGT GTCCTGG-3¢; 5, 5¢-GGCTCGAGATGGCCGCTGACAG GGGACGC-3¢; 6, 5¢-GCTCTAGACTAATCTTGTTGGT CAACCC-3¢; 7, 5¢-GGGGATCCTGCGTGGTTCTTTCC ATCTTGTTGGTCAACCCCACGATCAGTCTCATCTCC ACGGGCGGCCTG-3¢; 8, 5¢-GCTCTAGATTAATCTTT ATTGTCTAGGGAGTGTTCC-3¢; 9, 5¢-GGCGTCGACA GATACTGTGGAACACTCCCTAGACAATAAAGCTGG TCCTGTCTGC-3¢; 10, 5¢-GCGGATCCTGCGTGGTTCT TTCCATC-3¢; 11, 5¢-GGCGTCGACAGATACTGTGGAA CACTCCC-3¢; 12, 5¢-GCGGATCCTGCGTGGTTCTTTC CAGCTTGTTGGTCAACCC-3¢; 13, 5¢-GCGGATCCCCC GGGTGCGAGGAGACTGCTGCCGG-3¢; 14, 5¢-CTTTA TTATTTTTATTTTATTGAGAGGGTGG-3¢; 15, 5¢-GCG GATCCCTCGAGAAACCCAAACTCTTCTTCATTCAG GCTGCCCGAGGGACCGAGCTTG-3¢; 16, 5¢-CCACTTT AACTAATACTTTCAACATTTTCGG-3¢; 17–5¢-GGCCTC GAGAAGGGTTTTGCATC-3¢; 18, 5¢-GGATTCATGACT TTTAACAGTTTTGAAGG3¢; 19, 5¢-CCCCCGCGGCCG CTTAAGACATAAAAATTTTTTGCTTG-3¢; 20, 5¢-GGA ATTCATGCACAAAACTGCCTCCC-3¢; 21, 5¢-CCCCCG 22, CGGCCGCTTAAGAGAGAAATGTACGAAC-3¢; 5¢-GGCAGATCTCATATGGCCGCTGACAGGGGACGC- 3¢; 23, 5¢-CCCTCGAGTGTGGGAGGGTGTCCTGGG-3¢;

P.-k. Ho et al.

Caspase-2 can activate caspase-7 and is resistant to IAPs

25,

Yeast transformation and death assays

24, 5¢-GAGATCTCATATGAATAAAGATGGTCCTGTC TGC-3¢; 5¢-GGCAGATCTCATATGAATAAAGCT GGTCCTGTCTGC-3¢; 26, 5¢-GGAATTCCATATGTGTG TAATTTTTCCGGTAG-3¢; 27, 5¢-CCCTCGAGTTTAAT TGTGTTTAATATTAC-3¢; 28, 5¢-GCGGATCCATGACT TTTAACAGTTTTGAAGG-3¢; 29, 5¢-GAGAATTCTTAA GACATAAAAATTTTTTGCTTG-3¢; 30, 5¢-GCCTGCAG ACTGCTGGCAACCGCAGCAGCCACTCGAGG-3¢; 31, 5¢-GCCTGCAGCAGCTGCTCCAGGGCAGTGGCCAGG TCCCTGTTCCGGTCCTCCTCCGACCGGCTGGTGTTC CTGAGTTG-3¢; 32, 5¢-GCGGATCCATGGACTGTGAG GTCAACAACGG-3¢; 33, 5¢-GAGAATTCTCAGTCCAT CCCATTTCTGGC-3¢.

Following induction, bacteria were washed once in STE (10 mm Tris ⁄ HCl, pH 8.0, 150 mm NaCl, 1 mm EDTA), pelleted and frozen. His6 tagged caspase-2 and p35 were puri- fied using Ni2+–NTA agarose (Qiagen, Hilden, Germany) following manufacturer’s instructions. Quantification was performed by obtaining Western blot ECL signals and com- parison with standards of known concentration using a Gel-Doc system and quantity one analysis software (Biorad, Hercules, CA, USA). GST-fusion proteins were purified using glutathione sepharose (Amersham Biosciences) following the manufacturer’s instructions. XIAP and XIAPD148A were released from the GST-fusion partner by enzymatic cleavage using PreScission protease (Amersham fusion proteins Biosciences). For GST-fusion substrates, were left bound on the solid support. Quantification was per- formed by comparing bands against BSA standards using SDS ⁄ PAGE and staining with Coomassie blue.

Fluorogenic substrate cleavage assays

Preparation of yeast lysates

Saccharomyces cerevisiae strain W303a was used in all yeast transformation and death assays, as described previously [26,31].

peroxidase-conjugated horseradish

GST-fusion substrate cleavage assays

For immunoblotting, yeast were grown and induced for 7 h and lysates extracted as described previously [47]. Samples were resolved by SDS ⁄ PAGE on 12% gels, transferred to Hybond-P membrane (Amersham Biosciences), and probed with antibodies against caspase-2 [7] or caspase-7 (Cell Signaling, Beverly, MA, USA). Blots were washed with phos- phate-buffered saline ⁄ 0.5% Tween-20, and subsequently probed with goat anti-rat or donkey anti-mouse secondary Igs (Amersham Biosciences). Signals were developed using ECL reagents (Pierce, Rockford, IL, USA). Recombinant caspases were preactivated at 37 (cid:1)C for 30 min prior to the addition of fluorogenic substrates. Fluo- rogenic substrate cleavage assays were performed in an assay buffer containing 100 mm Hepes (pH 7.5), 10% sucrose, 0.1% CHAPS, 10 mm dithiothreitol, 25 mm Tris ⁄ HCl (pH 7.0), 75 mm NaCl, 0.5 mm EDTA and 0.2 mgÆmL)1 BSA with 50 lm of either Z-VDVAD-AFC or Ac-DEVD- AFC (Calbiochem, San Diego, CA, USA). For assays using yeast lysates, 30 lg of yeast lysates were used per 100 lL assay reaction. Assays using recombinant proteins contained 2 nm of preactivated caspase. Inhibition assays contained 2 nm of caspase-2 or 0.36 nm of caspase-3 (Biomol, Plymouth Meeting, PA, USA), with 0.5 lm of either XIAP, p35 or their inactive mutants per assay reaction.

Expression and purification of recombinant proteins

Native yeast lysates were prepared by resuspension in ice-cold lysis buffer (50 mm Tris ⁄ HCl, pH 7.0, 150 mm NaCl, 1 mm EDTA) followed by sonication for 15 s on ice. Lysates were cleared by centrifugation at 2300 g for 1 min and protein concentration was measured using the bicincho- ninic acid kit (Sigma, St. Louis, MO, USA) against a BSA standard curve.

Purified GST-fusion substrates bound on sepharose were used as substrates in the cleavage assays. Each 20-lL reac- tion contained 0.25 lm to 1.5 lm of GST-fusion protein on sepharose and 30 nm recombinant caspase (caspases-3, -7 and -8 were purchased from Biomol) in the same assay buf- fer used in fluorogenic substrate cleavage assays described above. The reactions were incubated at 37 (cid:1)C for 90 min and stopped by the addition of SDS ⁄ PAGE sample buffer, then boiled and subjected to SDS ⁄ PAGE. Gels were stained with Coomassie blue or immunoblotted with antibodies against caspase-2 or cleaved caspase-7 (Cell Signaling).

Acknowledgements

We thank B. Baliga, S. Read and S. Kumar for helpful comments on the manuscript and for permission to cite their unpublished data. We also thank E. Alnemri

FEBS Journal 272 (2005) 1401–1414 ª 2005 FEBS

1412

Constructs containing the genes of interest were transformed into Escherichia coli strain BL21(DE3)pLysS. Overnight cul- tures were grown until D600 reached 0.6–0.8. For His6- tagged caspase-2 and p35, inductions were carried out at 30 (cid:1)C for 3 h with 0.2 mm IPTG and for 4 h with 1 mm respectively. For GST-XIAP, GST-XIAPD148A, IPTG, GST-fusion substrates (Bid and BidD60A) and GST-fusion substrates (caspase-2C303A, myc-PARPD337)1013), inductions were carried out at 25 (cid:1)C for 3 h with 0.2 mm IPTG, for 4 h with 1 mm IPTG and for 8 h with 1 mm IPTG, respectively.

P.-k. Ho et al.

Caspase-2 can activate caspase-7 and is resistant to IAPs

for the Mch-2 plasmid. This work was supported by the Australian Research Council and a University of Melbourne Postgraduate Scholarship (to P.-K. H).

12 Lassus P, Opitz-Araya X & Lazebnik Y (2002) Require- ment for caspase-2 in stress-induced apoptosis before mitochondrial permeabilization. Science 297, 1352–1354.

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