Caspase2 is resistant to inhibition by inhibitor of apoptosis proteins (IAPs) and can activate caspase-7: Caspase2 can activate caspase-7 and is resistant to IAPs

Caspase-2 is resistant to inhibition by inhibitor ofapoptosis proteins (IAPs) and can activate caspase-7Po-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

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

interaction motif, is to proteolytically activate distal

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

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].

Caspase-2 has recently received 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],

Keywords

caspase-2; protease; caspase-7;

S. cerevisiae; enzyme activity

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: [email protected];

[email protected]

(Received 10 November 2004, revised 7

January 2005, accepted 18 January 2005)

doi:10.1111/j.1742-4658.2005.04573.x

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.

Abbreviations

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

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

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. found that 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].

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.

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

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

High level expression of pro-caspase-2 is lethal

in yeast

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).

Caspase-2 can activate caspase-7 and is resistant to IAPs P.-k. Ho et al.

1402 FEBS Journal 272 (2005) 1401–1414 ª 2005 FEBS

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

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

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.

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.

P.-k. Ho et al. Caspase-2 can activate caspase-7 and is resistant to IAPs


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 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-

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

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

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).



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).

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

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 [35] (lane 10).

Second, as previously reported for caspase-3 [27],

caspase-7 was activated by a constitutively active

caspase-9 (rev-caspase-9) (lane 11). This autoacti-

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.

A

B

C

D

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).



The relationship between caspase-2 processing

and enzymatic activity

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 (Fig. 6B) to determine enzyme activity.

Retention of at least one cleavage site between the

A

B

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).



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

fusion to the large subunit (D316A) (Fig. 6B). In

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), it was evident that

mutation of both of these cleavage sites decreased

activity by 840-fold (data not shown). The C303A

A

B

C

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).



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

suggest that the proteolytic activity of human

caspase-2 correlates with the degree to which the

large subunit is separated from the small subunit.

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

the various caspase-2 enzymes and subjected to

A

B

C

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.



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 the cleavage product 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).

Discussion

A unique merit of the yeast system used here is that it

is free from the potential interference of other mamma-

lian apoptotic signal transduction pathway compo-

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.

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

murine caspase-2. Firstly, the human and mouse

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 for 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.

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

A

B

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.



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.

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.

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 cells from apoptosis 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.

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.

Experimental procedures

Plasmid construction

For expression in yeast, coding regions of human genes

were cloned into the pGALL yeast vectors under the regu-

lation of the inducible Gal 1 ⁄ 10 promoter [26]. Yeast

vectors pGALL-(HIS3), pGALL-(LEU2) and pGALL-

(URA) have been described previously [27,35]. Plasmids

pGALL-(LEU2)-caspase-2, pGALL-(URA)-caspase-2, casp-

ase-3-LacZ, pGALL-(LEU2)-caspase-4, pGALL-(URA)-

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-(URA)-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

vector to produce pGALL-(LEU2)-caspase-2D153)435.

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, a SalI ⁄BamHI fragment 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-2C303A;D152, 316, 330AbyreplacingaSalI ⁄BamHIfragment

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 or pGALL-(HIS3)-

Caspase-2D152A and pGALL-(LEU2)-Caspase-2D152A,



respectively. pGALL-(HIS3)-Caspase-2D316A, pGALL-

(HIS3)-Caspase-2D330A, pGALL-(HIS3)-Caspase-2D316,330A,

pGALL-(HIS3)-Caspase-2D152,316,330A, pGALL-(URA)-Cas-

pase-2D316A, pGALL-(URA)-Caspase-2D330A, pGALL-

(URA)-Caspase-2D316,330A and pGALL-(URA)-Caspase-

2D152,316,330A were generated by ligating SpeI ⁄XbaIfragments released from pGALL-(HIS3)-FLAG-caspase-

2D316A, pGALL-(HIS3)-FLAG-caspase-2D330A, pGALL-

(HIS3)-FLAG-caspase-2D316,330A and pGALL-(HIS3)-

FLAG-caspase-2D152,316,330A into SpeI ⁄XbaI 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

with primers 15 and 14, digested with BamHI ⁄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,

respectively, into pGALL-(HIS3). The coding region of

MIHC was isolated from pADH-(TRP1)-MIHC [27] with

EcoRI ⁄NotI and cloned into pGALL-(HIS3) to give

pGALL-(HIS3)-MIHC.

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-

caspase2. pET23a-caspase2D1)149 and pET23a-casp-

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-

caspase-2D316A, pGALL-(HIS3)-FLAG-caspase-2D330A,

pGALL-(HIS3)-FLAG-caspase-2D316,330A and pGALL-

(HIS3)-FLAG-caspase-2D152,316,330A were used to replace

an internal fragment in pET23a-caspase-2D1)149 or

pET23a-caspase-2D1)149;D152A to generate pET23a-caspase-

2D1)149;C303A, pET23a-caspase2D1)149;D316A, pET23a-

caspase2D1)149;D330A, pET23a-caspase2D1)149;D316,330A and

pET23a-caspase2D1–149; D152 316 330 A, respectively.

pET23a-p35D87A was constructed by ligating a NdeI ⁄HindIII cut PCR product generated with primers 26 and 14

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.

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¢-CTTTATTATTTTTATTTTATTGAGAGGGTGG-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¢-CCCCCGCGGCCGCTTAAGAGAGAAATGTACGAAC-3¢; 22,

5¢-GGCAGATCTCATATGGCCGCTGACAGGGGACGC-

3¢; 23, 5¢-CCCTCGAGTGTGGGAGGGTGTCCTGGG-3¢;



24, 5¢-GAGATCTCATATGAATAAAGATGGTCCTGTC

TGC-3¢; 25, 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¢.

Yeast transformation and death assays

Saccharomyces cerevisiae strain W303a was used in all yeast

transformation and death assays, as described previously

[26,31].

Preparation of yeast lysates

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 horseradish peroxidase-conjugated goat

anti-rat or donkey anti-mouse secondary Igs (Amersham

Biosciences). Signals were developed using ECL reagents

(Pierce, Rockford, IL, USA).

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.

Expression and purification of recombinant

proteins

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 �C for 3 h with 0.2 mm IPTG and for 4 h with 1 mm

IPTG, respectively. For GST-XIAP, GST-XIAPD148A,

GST-fusion substrates (Bid and BidD60A) and GST-fusion

substrates (caspase-2C303A, myc-PARPD337)1013), inductions

were carried out at 25 �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.

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

Biosciences). For GST-fusion substrates, fusion proteins

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

Recombinant caspases were preactivated at 37 �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 lLassay 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.

GST-fusion substrate cleavage assays

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 �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



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).

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Caspase2 is resistant to inhibition by inhibitor of apoptosis proteins (IAPs) and can activate caspase-7: Caspase2 can activate caspase-7 and is resistant to IAPs

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