(Received for publication, November 26, 1996, and in revised form, April 7, 1997)
From the Cardiovascular Research Center,
Massachusetts General Hospital-East, Charlestown, Massachusetts 02129 and Department of Medicine, Harvard Medical School, Boston,
Massachusetts 02115, ¶ Pediatric Infectious Disease, Massachusetts
General Hospital, Charlestown, Massachusetts 02129, and the
Manitoba Institute of Cell Biology, Manitoba Cancer Treatment
and Research Foundation, University of Manitoba, Winnipeg,
Manitoba R3E OV9, Canada
Members of the CED-3/interleukin-1-converting
enzyme (ICE) protease (caspase) family are synthesized as proforms,
which are proteolytically cleaved and activated during apoptosis. We
report here that caspase-2 (ICH-1/NEDD-2), a member of the ICE family, is activated during apoptosis by another ICE member, a caspase-3 (CPP32)-like protease(s). When cells are induced to undergo apoptosis, endogenous caspase-2 is first cleaved into three fragments of 32-33
kDa and 14 kDa, which are then further processed into 18- and 12-kDa
active subunits. Up to 50 µM
N-acetyl-Asp-Glu-Val-Asp-aldehyde (DEVD-CHO), a
caspase-3-preferred peptide inhibitor, inhibits caspase-2 activation
and DNA fragmentation in vivo, but does not prevent loss of
mitochondrial function, while higher concentrations of DEVD-CHO (>50
µM) inhibit both. In comparison, although the activity of
caspase-3 is very sensitive to the inhibition of DEVD-CHO (<50
nM), inhibition of caspase-3 activation as marked by
processing of the proform requires more than 100 µM
DEVD-CHO. Our results suggest that the first cleavage of caspase-2 is
accomplished by a caspase-3-like activity, and other ICE-like proteases
less sensitive to DEVD-CHO may be responsible for activation of
caspase-3 and loss of mitochondrial function.
Interleukin-1-converting enzyme
(ICE)1 caspase-1 (1, 2) was
identified as the first mammalian homolog of the
Caenorhabditis elegans cell death gene product CED-3 (3,
4). Subsequently, a growing number of ICE-like cysteine proteases have
been isolated and characterized, including caspase-2 (NEDD-2/ICH-1) (5,
6), caspase-3 (CPP32/YAMA/Apopain) (7, 8, 39), caspase-6 (Mch-2) (9),
caspase-4 (TX/Ich-2/ICErelII) (10-12), caspase-5
(ICErelIII) (12), caspase-7 (Mch-3/CMH-1/ICE-LAP3)
(13-15), caspase-8 (FLICE/MACH/Mch-5) (16-18), caspase-10 (Mch-4)
(18), and caspase-9 (ICE-LAP6/Mch-6) (19, 20). Increasing evidence
suggests that caspases play critical roles in the control of programmed
cell death (for review, see Refs. 21-23). Microinjection of an
expression vector encoding CrmA, a serpin encoded by cowpox virus,
inhibits the death of dorsal root ganglia neurons induced by nerve
growth factor deprivation (24). Viral inhibitors of caspases, p35 and
CrmA, inhibit serum withdrawal-, tumor necrosis factor-, and
Fas-induced apoptosis, as well as cytotoxic T lymphocyte (CTL)-mediated
apoptosis (6, 25-29). Ice
/
thymocytes
undergo apoptosis normally when treated with dexamethasome and
-irradiation but are partially resistant to Fas-induced apoptosis (30). Peptide inhibitors of caspases prevent programmed cell death when
administered to tissue culture cells and animals (31). These results
indicate that the ICE family plays important roles in mammalian
apoptosis. The roles played by individual members of the caspase family
in controlling apoptosis are the subjects of intensive debates and
investigations.
Nedd-2, the murine caspase-2, was identified by Kumar et al. (32) as a mRNA expressed mostly during early embryonic brain development and down-regulated in adult brain. Overexpression of Nedd-2 in cultured fibroblast and neuroblastoma cells results in cell death by apoptosis, which is suppressed by the expression of the human bcl-2 gene (5). Previous work in our lab has shown that the human caspase-2, Ich-1 (Ice and ced-3 homolog), encodes a protein that shares sequence similarities with ICE and CED-3 proteins (6). Two different forms of mRNA species derived from alternative splicing encode two proteins, ICH-1L and ICH-1S, which have antagonistic effects on cell death. ICH-1L (435 amino acids) contains sequence homologous to both p20 and p10 subunits of ICE, while ICH-1S (312 amino acids) is a truncated version of ICH-1L, containing only the p20 region. Previous studies of Ich-1 in our laboratory revealed that overexpression of Ich-1L induces programmed cell death, while overexpression of Ich-1S suppresses Rat-1 cell death induced by serum deprivation. These results suggest that Ich-1 may play an important role in both positive and negative regulation of programmed cell death. Apoptosis induced by ICH-1 is suppressed by overexpression of bcl-2, but not by crmA. Northern blotting and reverse transcription-PCR results showed that Ich-1 is expressed in many tissues and cells with tissue and developmental stage specificities. Expression of Ich-1 is detected in HeLa, THP.1, U937, and Jurkat cells. The expression patterns of these two alternatively spliced forms of Ich-1 show tissue-specific differences; expression of both Ich-1L and Ich-1S can be detected in heart, kidney, and embryonic and adult brain with the expression of Ich-1S being highest in embryonic brain, and only Ich-1L is expressed in adult thymus.
To investigate the mechanism and function of caspase-2 (NEDD-2/ICH-1) in apoptosis, we examined the processing and activation of caspase-2 when cells undergo apoptosis. We demonstrate here that caspase-2 is processed and activated in a specific temporal sequence when cells are induced to undergo apoptosis by diverse stimuli. Our results show that caspase-2 is activated by a caspase-3 (CPP32)-like protease when cells are induced to undergo apoptosis. Moreover, caspase-2 activation can be distinguished from activation of caspase-3 and loss of mitochondrial function by their sensitivity to inhibitors of the ICE family.
Staurosporine, 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT), and other molecular biology grade reagents were purchased from Sigma. N-Acetyl-Asp-Glu-Val-Asp-aldehyde (DEVD-CHO), N-acetyl-Tyr-Val-Ala-Asp-chloromethylketone (YVAD-CMK), and N-acetyl-Tyr-Val-Ala-Asp-aldehyde (YVAD-CHO) were obtained from Bachem Bioscience, Inc. (King of Prussia, PA).
Cell CulturesJurkat cells were grown in RPMI 1640 medium (Life Technologies, Inc.) with 10% fetal calf serum. HeLa cells were grown in Dulbecco's modified Eagle's medium (Life Technologies, Inc.) supplemented with 10% fetal calf serum.
Constructions of Expression Plasmids for Caspase-1, -2, and -3 in Bacteria and Site-directed Mutagenesis of Caspase-2BamHI sites were introduced at the 5 and 3
ends of the p30 domain of caspase-1 and full-length caspase-3 by PCR
amplification. For caspase-1 p30, oligonucleotide primers
5
-CGCGGATCCTGGCACATTTCCAGGAC-3
(5
primer) and
5
-CGCGGATCCTAAGGAAGTATTGGC-3
(3
primer) were used. For caspase-3,
two primers, 5
-CGCGGATCCGGAGAACACTGAAAACTC-3
(5
primer) and
5
-CGCGGGATCCTACCATCTTCTCACTTGG-3
(3
primer), were used. For
caspase-3 p30, XhoI sites were introduced at the both ends
by PCR using two primers: 5
-GCGCTCGAGGGTCCTGTCTGCCT-3
(5
primer) and
5
-CGGCTCGAGGTGACATCATGTGGG-3
(3
primer). The PCR products were
cloned into pBluescript (Promega, Madison, WI), and their sequences
were confirmed by DNA sequencing (U. S. Biochemical Corp.). Each
fragment was inserted into the BamHI or
XhoI site of pET-15b (Novagen, Madison, WI). The
resulting plasmids were transformed into Escherichia coli
strain BL21(DE3).
Site-directed mutagenesis of caspase-2 was carried out by PCR. Two
primers, D316E primer (5-GGGGATCCTGCGTGGTTCTTTCCCTCTTGTTGGTC-3
) and
D330E primer (5
-GCAGGATCCCCTGGGTGCGAGGAGAGTGATGCCGGTAAAG-3
) were used
to mutate both Asp-316 and Asp-330 to Glu. To generate caspase-2
(D316E) mutant, PCR was performed using caspase-2 5
primer
(5
-GCGCTCGAGCTGATGGCCGCTG-3
) and D316E primer and wild type caspase-2
cDNA as a template. The PCR fragment was cloned into pBluescript.
The BamHI-digested PCR fragment was used to replace the
corresponding wild type fragment in caspase-2. D330E primer and
caspase-2 3
primer (5
-CGGCTCGAGA CATCATGTGGG-3
) were used in a
similar procedure to generate caspase-2 (D330E) mutant.
E. coli BL21(DE3) transformed with plasmids
expressing caspase-1, -2, and -3 genes
were grown in LB media to exponential phases, and induced with 0.4 mM isopropyl-1-thio--D-galactopyranoside for
2 h. Cells were pelleted, resuspended in lysis buffer (30 mM Tris-HCl, pH 7.5, 0.1 mM NaCl, 1 mM DTT, 0.1 mM EDTA, 1% Nonidet P-40, and 20 µg/ml PMSF), and sonicated. The supernatant after centrifugation at
14,000 × g for 15 min was used in enzymatic cleavage
assays. The protein concentration was determined by BCA assay (Pierce),
and aliquots were stored at
80 °C.
Jurkat cells were induced to undergo apoptosis by a
variety of agents including staurosporine and anti-Fas monoclonal
antibody CH-11 (Kamiya Biomedical Co., Thousand Oaks, CA), whereas HeLa cells were treated with a combination of TNF (R&D Systems,
Minneapolis, MN) and cycloheximide. The percentage of cell death was
measured either by trypan blue exclusion or MTT assays. For trypan blue exclusion assay, Jurkat cells or trypsinized HeLa cells were incubated with 0.4% trypan blue solution (Sigma) for 10 min, and more than 200 cells were scored on a hemocytometer. Alternatively, MTT assays were
performed as described (33). Briefly, 5 × 104 cells
(50 µl) were subcultured in RPMI 1640 (phenol red-free) supplemented
with 10% fetal calf serum in a 96-well plate, and treated with
apoptosis-inducing agents for various time periods. For MTT assay, 5 µl of MTT agent (5 mg/ml in RPMI 1640 (phenol red-free)) was added
and further incubated for 2 h. Equal volumes of 0.05 N
HCl in isopropanol were then added, and cells were disrupted by
pipetting up and down. Cell viabilities were determined
colorimetrically by using an automated 96-well plate reader (Molecular
Devices, Sunnyvale, CA) and SOFTmax software to measure absorbance at
570-650 nm.
Detection of DNA fragmentation was performed as described by Eastman (34). Briefly, a 2% agarose gel was prepared by pouring 350 ml of 2% agarose in TAE buffer in a large (20 × 34 cm) horizontal gel support. Once the gel solidified, the top section of gel immediately above the comb was removed, and filled with 1% agarose, 2% SDS, 64 µg/ml proteinase K. After treated with 200 ng/ml anti-Fas monoclonal antibody in the presence of different amounts of DEVD-CHO for 20 h, Jurkat cells were harvested by centrifugation at 1000 rpm, and excess medium was removed. The cell pellets were resuspended in 15 µl of sample buffer (5% glycerol, 5 mM Tris, pH 8.0, 0.05% bromphenol blue, and 5 mg/ml RNase A), and directly loaded into the wells. After electrophoresis for 14 h at 60 V at room temperature, the gel was stained with 0.5 µg/ml ethidium bromide in water for 1 h, and destained in water overnight. The picture was taken using the Gel Doc 1000 system (Bio-Rad).
Western BlottingThe protein samples were subjected to
SDS-PAGE, and then transferred to Immobilon-P membranes (Millipore,
Bedford, MA) using a semi-dry transfer apparatus (Pharmacia Biotech
Inc.). The membranes were blocked in TBST buffer (20 mM
Tris-HCl, pH 7.5, 150 mM NaCl, 0.2% Tween 20) containing
5% nonfat dried milk overnight at 4 °C. Membranes were then blotted
with various primary antibodies with different dilutions for 2 h
at room temperature. After washing three times in TBST, membranes were
subsequently incubated with horseradish peroxidase-conjugated secondary
antibodies (either goat anti-mouse or goat anti-rabbit) (Southern
Biotechnology, Birmingham, AL) for 45 min. After washing in TBST,
proteins were detected by ECL (Amersham) according to the
manufacturer's instructions. Primary antibodies were diluted as
follow: polyclonal antibody for caspase-2 with a dilution of 1:3000,
polyclonal antibody C-20 (Santa Cruz) for caspase-2 C terminus
(416-435 residues; SEYCSTLCRHLYLFPGHPPT) with a dilution of 1:200,
monoclonal antibody for caspase-3 (Transduction Laboratories) with a
dilution of 1:2000, polyclonal antibody for PARP with a dilution of
1:1000, and monoclonal antibody for -tubulin (Sigma) with a dilution
of 1:5000.
Jurkat cells
(1 × 108) were treated with 1 µM
staurosporine for various time periods, and cytosolic lysates were
prepared as described with minor modification (35). Briefly, cells were washed twice with cold RPMI 1640, and resuspended in 400 µl of extraction buffer (10 mM HEPES, pH 7.0, 40 mM
glycerophosphate, 50 mM NaCl, 2 mM
MgCl2, 5 mM EGTA, and 1 mM DTT)
containing protease inhibitors (1 mM PMSF, 1 µg/ml
leupeptin, 0.5 µg/ml aprotinin). After four cycles of freezing and
thawing, crude extracts were obtained by centrifugation at 12,000 × g for 15 min at 4 °C. The cell lysates were further
centrifuged at 100,000 × g for 60 min, and the
resulting supernatant was used as the cytosolic fraction. The protein
concentration was determined by BCA protein assay (Pierce), and
aliquots were stored at 80 °C.
In vitro translations of 35S-labeled proteins were done by using the TNT-coupled transcription/translation kit (Promega) in the presence of [35S]methionine. 35S-Labeled proteins were incubated with either bacterial lysates or staurosporine-treated Jurkat cytosolic lysates in a reaction buffer (20 mM Tris-HCl, pH 7.5, 10 mM DTT, 0.1 mM EDTA) for 1-2 h at 30 °C, in the presence of protease inhibitors (1 mM PMSF, 0.5 µg/ml aprotinin). The reactions were terminated by addition of equal volume of 2 × protein lysis buffer, and analyzed by SDS-PAGE. In Granzyme B (GB) cleavage assay, 35S-labeled caspase-2 or caspase-3 was incubated with 20 of ng GB in a reaction buffer (20 mM Tris, pH 8.0, 100 mM NaCl, and 1 mM DTT) at 30 °C for 1 h.
Analysis of Target Cell Proteins following Cytotoxic T Lymphocyte (CTL)-mediated CytolysisAlloreactive murine CTL (F3B4,
anti-H-2b, or FC4 and G4, anti-H-2d) were
harvested 4-6 days after stimulation and purified by Ficoll-Hypaque density gradient centrifugation. Target cells (EL-4, H-2b;
P815, H-2d) were washed in fresh supplemented RPMI medium.
Target cells (~6 × 105) were mixed with CTL at an
effector:target ratio of 1-2.5 in a final volume of 200 µl of medium
in microcentrifuge tubes. Control samples were prepared by adding the
cells directly to 1 ml of PBS wash buffer containing the protease
inhibitors diisopropyl fluorophosphate (4 mM) and
para-hydroxymercurobenzoate (2 mM) and
immediately harvested. The remaining samples were centrifuged briefly
at 500 rpm, and then incubated at 37 °C for 45-90 min. The
incubated samples were diluted with 1 ml of washing buffer and pelleted
at 2000 rpm for 2 min in a microcentrifuge. The supernatants were
aspired, and the cell pellets were dissolved in 200 µl of PBS
solubilization buffer containing 1% Nonidet P-40, 0.1% SDS, diisopropyl fluorophosphate, and
para-hydroxymercurobenzoate) for 15 min on ice. The samples
were centrifuged at 3000 rpm for 3 min in a microcentrifuge. The
supernatants were transferred to fresh microcentrifuge tubes, and
precipitated in 1.2 ml of cold acetone. After overnight storage at
20 °C, the extracted proteins were recovered by centrifugation,
dried by vacuum centrifugation, and analyzed by SDS-PAGE.
Members
of the caspase family are synthesized as precursors of approximately
45-50 kDa. Activation of the caspases involves proteolytic cleavages
of the precursors at specific Asp residues into a large subunit of
approximately 20 kDa and a small subunit of approximately 10 kDa. To
determine whether caspase-2 is cleaved and activated when cells undergo
apoptosis, a rabbit polyclonal antibody was generated against purified
His-tagged caspase-2 protein expressed in E. coli. On
Western blots, this antibody recognizes a 48-kDa polypeptide, the
molecular mass predicted for caspase-2 precursor protein, in Jurkat and
HeLa cells as well as non-human cell lines including Rat-1 and COS
cells (Fig. 1 and data not shown). This
48-kDa protein is specifically absent from tissues of
caspase-2/
mutant mice generated by gene
targeting technique, which further confirms the identity of this 48-kDa
protein as the product of caspase-2 locus (data not shown). In several
human cell lines, as well as in mice, this polyclonal anti-caspase-2
antibody also detects a 37-kDa polypeptide, which is not altered in
caspase-2
/
mutant mice, and thus is not from caspase-2
locus (data not shown). No cross-reactivity of this antibody to
caspase-1, caspase-3, and caspase-4 was observed using Western blot
analysis (data not shown).
To examine whether caspase-2 is activated during apoptosis, we induced
apoptosis in Jurkat cells by treatment with anti-Fas antibody in the
presence or absence of cycloheximide, which inhibits protein synthesis
and potentiates apoptosis, or staurosporine, a broad spectrum protein
kinase inhibitor that induces apoptosis in a variety of cells (36).
Total cell lysates were collected at different time points and
subjected to Western blot analysis using the polyclonal anti-caspase-2
antibody. In these experiments, processing of pro-caspase-2 was first
detected as the appearance of a 32-33-kDa doublet at 1-h time point
and an 18-kDa peptide at 4-h time point (Fig. 1A). The
degree of caspase-2 processing correlates very well with the extent of
cell death. A similar processing pattern of caspase-2 was observed with
anti-Fas antibody alone, but with a delayed time course of cell death
and caspase-2 processing (data not shown). Processing of caspase-2 was
also detected in HeLa cells that were induced to die by TNF and
cycloheximide (27) (Fig. 1B). These observations suggest
that caspase-2 is activated in apoptosis and its processing may be an
important regulatory step for caspase-2.
As
described above, a polyclonal anti-caspase-2 antibody first detects the
appearance of 32-33-kDa doublets and then detects an 18-kDa
polypeptide during the course of apoptosis. The 32-33-kDa doublets may
be intermediate processing products, which may consist of either the
large subunit plus pro-domain or the large subunit plus the small
subunit. Since this polyclonal antibody against caspase-2 recognizes
the purified full-length but not the small subunit of caspase-2
expressed in E. coli (data not shown), the 32-33-kDa
products are likely to be the pro-domain plus the large subunit. To
verify this, we used a polyclonal peptide antibody that recognizes the
C terminus of caspase-2 to immunoblot the same lysate samples (Fig.
2, A and B). This
anti-C-terminal caspase-2 antibody recognizes the full-length caspase-2
and three additional polypeptides with estimated molecular masses of
44, 14, and 12 kDa, but not the 32-33-kDa doublets, confirming that
the 32-33-kDa products do not contain the C-terminal sequence. The
14-kDa product appeared at the same time point of apoptosis as the
32-33-kDa doublets did, suggesting that it is the
C-terminal-containing small subunit of caspase-2. The 12-kDa product
was detected much later than the 14-kDa, suggesting that the 12-kDa
peptide may be a further cleavage product of the 14-kDa product. There
was no change for the 44-kDa peptide in apoptosis, suggesting that it
is a protein not related to caspase-2 but is cross-recognized by this
anti-C-terminal antibody. This observation showed that caspase-2 is
activated by several distinct cleavage events, in which cleavage
between the large subunit and small subunit occurs first, followed by
cleavage(s) between the large subunit and the pro-domain and within the
14-kDa C-terminal domain. The possible cleavage sites are summarized in
Fig. 2C.
Activation of Caspase-2 Occurs Later than the Activation of Caspase-3-like Proteases when Cells Undergo Apoptosis
To determine possible interactions between caspase-2 and other family members, we compared the temporal activation profiles of caspase-2 and caspase-3. Jurkat cells and HeLa cells were induced to undergo apoptosis by incubation with anti-Fas antibody or staurosporine. The apoptotic cell lysates were immunoblotted with anti-caspase-3 monoclonal antibody as well as antibody specific for poly(ADP-ribose) polymerase (PARP), a substrate of caspase-3. Activation of caspase-3, as indicated by the disappearance of full-length caspase-3 and the appearance of the 89-kDa PARP cleavage product (PARP*), was observed shortly after treatment with apoptotic stimuli at a time point indistinguishable with the first appearance of the caspase-2 32-33-kDa doublets (Fig. 1A). Since the pro-domains of the caspase family often have inhibitory activity (12), and our in vitro data suggest that removal of the pro-domain is an essential event for activation of caspase-2 (see below), activation of caspase-2 as marked by the appearance of the 18-kDa cleavage product occurred at a much later time point than that of caspase-3 as marked by the cleavage of PARP (Fig. 1A). Thus, although the first cleavage of caspase-2 occurs at approximately the same time as the activation of caspase-3, activation of caspase-2 did not occur until 2-3 h later.
Caspase-2 Is a Substrate of Caspase-3 in VitroWhen cells
undergo apoptosis, caspase-2 may be activated by another caspase(s)
and/or by its self-catalytic activity. To address this issue, we tested
whether active caspase-2 and other ICE-like proteases are capable of
cleaving pro-caspase-2. When we expressed full-length caspase-2
cDNA in E. coli, we could not obtain active protease
activity even though 50% of full-length caspase-2 was processed into
pro-domain-large subunit and small
subunit.2 These findings
suggest that the N-terminal pro-domain has an inhibitory effect on
caspase-2 self-processing, especially on the processing between the
pro-domain and large subunit, and that the combination of
pro-domain-large subunit and the small subunit is inactive. To obtain
active caspase-2 protease, we expressed a fragment (named p30) in
E. coli containing a deletion of the N-terminal pro-domain
152 amino acid residues. As a control, we created a mutant which
contains Cys to Ser mutation in the coding region of the active site
pentapeptide QACRG (p30C-S). Using anti-caspase-2 antibodies in Western blot analysis of bacterial lysates, we found that
overexpressed wild type p30 was self-processed into 18-kDa and 14-kDa
polypeptides, while the p30C-S mutant remained intact, indicating that processing of wild type p30 is due to its own catalytic
activity. To determine whether such bacterially expressed caspase-2 p30
was active, we examined its ability to cleave full-length 35S-labeled in vitro translated pro-caspase-2.
As shown in Fig. 3A, caspase-2
p30 was capable of cleaving full-length caspase-2 into two polypeptides
of 34 kDa and 14 kDa, a pattern similar to the in vivo
results (Fig. 1). To explore the possibility that caspase-2 may be
cleaved by another member of the ICE family, we investigated whether
active caspase-1 and caspase-3 cleaved pro-caspase-2 in
vitro. Caspase-1 and caspase-3 cDNA were expressed in E. coli, and such caspase-1 and -3-expressing bacterial lysates were
found to efficiently cleave pro-IL-1 and PARP in vitro, respectively (data not shown). As shown in Fig. 3 A,
caspase-3 cleaved 35S-labeled pro-caspase-2 into two
polypeptides of 34 kDa and 14 kDa, while caspase-1 cleaved both
caspase-2 and caspase-3 very poorly. In contrast, neither caspase-3 nor
p30 of caspase-2 cleaved pro-caspase-1. These results suggest that
caspase-3 or a caspase-3-like member of the caspase family may act as
an activator of caspase-2.
We also determined the abilities of three peptide inhibitors of the caspase family to inhibit ICH-1 protease activity in vitro. YVAD-CHO and DEVD-CHO are relatively specific inhibitors of caspase-1-like and caspase-3-like proteases, respectively. DEVD-CHO inhibits caspase-3 with Ki = 0.52 nM (37), whereas YVAD-CHO is a very potent inhibitor of caspase-1 (Ki = 0.76 nM) (2). Addition of YVAD-CMK (5 µM), an irreversible inhibitor of caspase-1-like proteases, inhibited the cleavage of pro-caspase-2 by p30 (data not shown). The caspase-2 activity, however, cannot be inhibited by YVAD-CHO (up to 400 µM, data not shown) and is insensitive to DEVD-CHO; only 50% inhibited at 10 µM DEVD-CHO with preincubation (Fig. 3B). In contrast, cleavage of caspase-2 by caspase-3 is much more sensitive to DEVD-CHO than that by caspase-2 itself: 50 nM DEVD-CHO inhibited the cleavage completely (Fig. 3C).
Processing of Caspase-2 and DNA Fragmentation, but Not Loss of Mitochondrial Function, Is Inhibited by Up to 50 µM DEVD-CHOOur in vitro cleavage results suggest that
caspase-3 or a caspase-3-like protease may act as an activator of
caspase-2. To elucidate the mechanism of caspase-2 activation during
apoptosis, we examined whether DEVD-CHO inhibited caspase-2 activation
and apoptosis in vivo. Previous studies have shown that
DEVD-CHO can inhibit apoptosis in cultured cells as well as in animals,
although the concentrations required are much higher than what is
needed to inhibit individual caspases in purified forms (38-42).
Jurkat cells were treated with anti-Fas antibody in the presence of
different concentrations of DEVD-CHO. Percentages of viable cells were
assessed by MTT assay (34), which measures mitochondrial function, and processing of caspase-2 was examined by immunoblotting using
anti-caspase-2 polyclonal antibody. As shown in Fig.
4A, approximately 50% of the
caspase-2 processing was inhibited by 10 µM DEVD-CHO, and 90% of caspase-2 processing was inhibited by 50 µM
DEVD-CHO, a concentration that completely inhibited PARP cleavage. In
contrast, caspase-3 activation as marked by the disappearance of
full-length caspase-3 was not affected by 10 µM DEVD-CHO,
and modestly affected by up to 100 µM DEVD-CHO (Fig.
4A). These results suggest that although the activity of
caspase-3 indicated by PARP cleavage is sensitive to DEVD-CHO,
caspase-3 itself is activated by a caspase less sensitive to DEVD-CHO.
DNA fragmentation was nearly half inhibited by 10 µM
DEVD-CHO, and almost completely inhibited by 50 µM
DEVD-CHO (Fig. 4B). In contrast, up to 50 µM
DEVD-CHO had no effect on cell viability as measured by MTT assay (Fig.
4). These results showed that loss of mitochondria function, activation of caspases-2 and -3, DNA fragmentation, and cleavage of PARP can be
distinguished by their differential sensitivities to the inhibition by
DEVD-CHO.
Activation of ICH-1 in a Cell-free System
To further explore
the identity of the upstream activator of caspase-2, we established a
cell-free system using staurosporine-induced apoptotic Jurkat cytosolic
lysate. Jurkat cells were induced to undergo apoptosis in the presence
of 1 µM staurosporine. Cytosolic extracts at different
time points of staurosporine treatment were isolated and incubated with
35S-labeled in vitro translated caspase-2 and
PARP for 2 h (Fig. 5A).
Cleavage of caspase-2 into 34 and 14 kDa and cleavage of PARP into 89 and 27 kDa in apoptosis induced by staurosporine occurred in a similar
time course as to that induced by anti-Fas antibody. Furthermore,
cleavage of both PARP and caspase-2 in this cell-free system was
sensitive to DEVD-CHO (50 nM) but insensitive to YVAD-CHO
(50 µM) (Fig. 5, B and C). These
results again suggest that caspase-2 is activated by caspase-3 or
caspase-3-like proteases during apoptosis.
Determination of Cleavage Sites of Caspase-2 Processing
Proteolytic activation of caspase proteases involves
cleavage of specific Asp residues in the precursor peptides. Based upon the homology between caspase-2 and 1 and the consensus sequence of
caspase-3 cleavage, several Asp residues in caspase-2 are candidates for processing sites (Asp-83, Asp-99, Asp-118, Asp-120, Asp-152, Asp-316, and Asp-330; only DNKD152G153 and
DQQD316G317 has caspase-3 cleavage consensus
sequence). To determine the processing sites of caspase-2 in
vitro, we mutated Asp residues at positions 316 and 330 to Glu
(D316E and D330E), and in vitro cleavage assays were
performed using these two mutants. As shown in Fig.
6, the mutation at Asp316 (D316E)
completely blocked the pro-caspase-2 cleavage event by either caspase-2
and caspase-3 expressing bacterial lysate, or staurosporine-treated
Jurkat apoptotic cytosolic lysate (data not shown), whereas the D330E
mutation appears to alter the cleavage site, suggesting that Asp-316
was the primary cleavage site of caspase-2 by its activator. The
caspase-2 p30 double mutant bearing D316E/D330E was incapable of
self-processing and cleaving pro-caspase-2, indicating that processing
at Asp-316 is essential for caspase-2 proteolytic activity (data not
shown).
Caspase-2 Is Activated in CTL-mediated Apoptosis
CTL-mediated
cytotoxicity, the major cellular defense against virus-infected and
tumorigenic cells, is executed through two mechanisms:
perforin-granzyme B pathway (Ca2+-dependent)
and Fas signaling pathway (Ca2+-independent) (43, 44).
Previous studies have shown that CrmA, a specific inhibitor of
caspase-1, can inhibit CTL-mediated apoptosis, primarily by blocking
the Fas pathway (29). Granzyme B can cleave and directly activate
caspase-3 (45). It is of particular interest to examine whether
caspase-2 is activated in CTL-mediated apoptosis and is activated by
granzyme B directly. CTL-resistant (P815) and CTL-sensitive target
cells were incubated with CTL clone F3B4 in a ratio of 1:1. Caspase-2
is barely expressed in CTL, but highly expressed in target cells (Fig.
7A). Caspase-2 was fully
processed within 45 min in positive target cells EL4, whereas it
remained intact in negative control cells P815, indicating that
caspase-2 may also play a role in CTL-mediated apoptosis. We could not
observe processing products of caspase-2 since our anti-caspase-2
polyclonal antibody was generated against human caspase-2 and does not
recognize processed mouse caspase-2. To determine if granzyme B can
directly activate caspase-2, we determined if purified granzyme B (54) may cleave in vitro translated 35S-labeled
caspase-2. Such analysis showed that although granzyme B cleaves
caspase-3 efficiently, it cannot cleave caspase-2 (Fig. 7B).
Thus, activation of caspase-2 by CTL is most likely to be mediated
through the Fas pathway or indirectly by another caspase(s) activated
by granzyme B rather than granzyme B itself.
We have demonstrated that caspase-2 (NEDD-2/ICH-1), a member of
the ICE family, is activated when cells are induced to undergo apoptosis by diverse stimuli such as anti-Fas antibody, TNF, and
staurosporine. When cells are induced to undergo apoptosis, endogenous
caspase-2 is first cleaved into three fragments of 32-33 and 14 kDa,
which are then processed further into 18-kDa and 12-kDa active subunit.
When overexpressed in bacteria, the fragment of caspase-2 without its
N-terminal pro-domain was cleaved into two peptides of 18 and 12 kDa,
which are enzymatically active, similar to what has been reported (46).
The 18-kDa polypeptide detected by anti-caspase-2 antibody in apoptotic
cells is likely to be the large subunit of active caspase-2. Taken
together, our in vitro and in vivo observations
strongly suggest that caspase-2 is indeed activated when cells undergo
apoptosis.
The mechanism of activation of ICE/CED-3 cysteine proteases remains unclear so far. Two possible mechanisms, which are not mutually exclusive, may be involved. The first mechanism is that each member of the caspase family is activated through self-catalytic cleavage upon dissociation with a putative inhibitor(s). The evidence supporting this notion is that several members, when overexpressed in vitro, are capable of undergoing self-cleavage to generate active enzymes (10, 12, 47). The second possible mechanism is cross-activation whereby one caspase activates another one(s). We found that while caspase-2 activity in vitro is much less sensitive to the inhibition by DEVD-CHO than that of caspase-3, the activation of caspase-2 in cells, as indicated by the cleavage of pro-caspase-2, is as sensitive to the inhibition by DEVD-CHO as that of cleavage of PARP, an indicator of caspase-3-like activity. Our results suggest that caspase-2 is most likely to be activated by a caspase-3-like activity rather than by a self-activation mechanism. It has been shown that in in vitro assay systems, caspase-4 (TX/ICH-2) can process both pro-caspase-4 and pro-caspase-1 (10), and caspase-1 can process and activate pro-caspase-1 and caspase-3 (8). It is not clear, however, whether such cross-activation indeed occurs in cells undergoing apoptosis. Our study demonstrated that in vivo one member of the caspase family, caspase-2, is activated by another member of the caspase family, a caspase-3-like protease(s), when cells are induced to undergo apoptosis by staurosporine and anti-Fas antibody. Dr. Shige Nagata's laboratory has shown that when cells are induced to undergo apoptosis by anti-Fas antibody, there is a sequential activation of caspase-1-like and caspase-3-like proteases (48). Our results extended their observation by revealing downstream targets of the caspase-3-like proteases. The observation that caspase-2 was processed in CTL-mediated apoptosis, but granzyme B cannot cleave caspase-2 directly, also suggests that other factors mediate caspase-2 activation in perforin-granzyme B killing. Taken together, we propose a model of sequential activation involving three subfamilies of ICE/CED-3 proteases in the execution of programmed cell death. In this model, when cells are stimulated with a death signal such as anti-Fas antibody, a caspase-1-like protease(s) is activated first, followed by activation of a caspase-3-like protease(s) that may be mediated by the caspase-1-like activity, and then a caspase-3-like protease(s) activates caspase-2. We do not know, however, the exact identities of the upstream caspase-1- and caspase-3-like activity. Further studies using mutant mice that are defective in one or more members of the caspase family proteases are needed to clarify these questions.
Caspase-3 protease is activated by cleavage events at Asp-28/Ser-29 (between N-terminal pro-domain) and Asp-175/Ser-176 (between the large and the small subunits) to generate a large subunit of 17 kDa and a small subunit of 12 kDa (7), whereas pro-caspase-1 is activated through four cleavage events: two cleavages between the N-terminal prodomain (Asp-103/Ser-104 and Asp-119/Asn-120) and two between the large and small subunits (Asp-297/Ser-298 and Asp-316/Ala-317) (2). The temporal sequences of proteolytic cleavages during caspase-1 and -3 activation are not clear. We showed here that activation of caspase-2 occurs in distinct cleavage steps. The timing of the first cleavage between the large subunit and the small subunit coincides with the activation of caspase-3 and cleavage of PARP. This cleavage is inhibitable by DEVD-CHO in vivo and in vitro, although the active caspase-2 itself is much less sensitive to this inhibitor than that of caspase-3. These two observations suggest strongly that this first cleavage of caspase-2 is carried out by caspase-3 or a caspase-3-like protease. Our in vitro data indicate that a single cleavage between the large subunit and the small subunit of caspsae-3, however, is insufficient to activate caspase-2. The second cleavage of caspase-2, between the pro-domain and the large subunit, occurs much later at 4 h, when 25% of cells are dead as estimated by MTT assay. Neither caspase-3 nor active caspase-2 can carry out this second cleavage in vitro, suggesting that this cleavage is executed by an uncharacterized protease.
Apoptosis is usually measured by MTT assay, DNA fragmentation, or trypan blue exclusion (49). Each of these procedures measures a different parameter of cell viability. Trypan blue exclusion measures the integrity of cell membrane or permeability change. Disruption of the cytoplasmic membrane occurs relatively late in apoptosis. DNA fragmentation, representing an alteration in nuclei, occurs much earlier than changes in cell membrane permeability (our unpublished observation). The MTT assay is a quantitative colorimetric assay based on reduction of a tetrazolium salt, MTT. MTT is reduced within the active mitochondria of living cells by the enzyme succinate dehydrogenase (50). The salt is reduced to an insoluble blue formazan product in living cells but not in the mitochondria or cellular debris of dead cells. 70-80% of mitochondrial MTT reduction occurs subsequent to transfer of electrons from cytochrome c to cytochrome oxidase, but prior to the point of azide inhibition (51). Loss of mitochondrial function, a process beginning with a decrease in mitochondrial transmembrane potential, followed by mitochondrial uncoupling and generation of reactive oxygen species, precedes nuclear alteration (52). Recently, release of cytochrome c from mitochondria has been shown to be an early and essential step of apoptosis in a cell-free system induced by dATP (53). Our data showed here that there is a concentration of DEVD-CHO (50 µM), which inhibits the cleavage and activation of caspase-2 by a caspase-3-like activity and DNA fragmentation but does not alter viability as measured by MTT, suggesting that DEVD-CHO at that dose can block activation of the caspase family members such as caspase-2 but cannot block loss of mitochondrial function in apoptosis induced by anti-Fas antibody. These results indicate that activation of caspase-2 by a caspase-3-like activity is separable from the loss of mitochondrial function. Higher doses of DEVD-CHO, however, can inhibit loss of mitochondrial function as measured by MTT. Since the subfamily of caspase-1-like proteases that are mostly closely related to caspase-1 requires higher concentrations of DEVD-CHO for inhibition, this result suggests that there is an caspase-1-like activity further upstream from loss of mitochondrial function. This result is consistent with the report by Enari et al. (48), who showed that activation of an caspase-1-like activity precedes the activation of caspase-3-like activities in apoptosis induced by anti-Fas activity. It is not clear, however, in lieu of the recent report of caspase-8 (FLICE/MACH), an caspase-3-like protease containing MORT domain that allows direct coupling to the Fas receptor upon activation, the exact identity of this caspase-1-like activity.