* IDUN Pharmaceuticals, La Jolla, California 92037; and Novartis Pharma AG, CH-4002 Basel, Switzerland
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Abstract |
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The mechanism by which membrane-bound Bcl-2 inhibits the activation of cytoplasmic procaspases is unknown. Here we characterize an intracellular, membrane-associated form of procaspase-3 whose activation is controlled by Bcl-2. Heavy membranes isolated from control cells contained a spontaneously activatable caspase-3 zymogen. In contrast, in Bcl-2 overexpressing cells, although the caspase-3 zymogen was still associated with heavy membranes, its spontaneous activation was blocked. However, Bcl-2 expression had little effect on the levels of cytoplasmic caspase activity in unstimulated cells. Furthermore, the membrane-associated caspase-3 differed from cytosolic caspase-3 in its responsiveness to activation by exogenous cytochrome c. Our results demonstrate that intracellular membranes can generate active caspase-3 by a Bcl-2-inhibitable mechanism, and that control of caspase activation in membranes is distinct from that observed in the cytoplasm. These data suggest that Bcl-2 may control cytoplasmic events in part by blocking the activation of membrane-associated procaspases.
Key words: apoptosis; Bcl-2; caspase; cytochrome c; programmed cell death ![]() |
Introduction |
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THE caspases are a family of cysteine proteases that
are essential effectors of the apoptotic process
(Yuan et al., 1993; Alnemri et al., 1996
; Cohen,
1997
; Miller, 1997
; Salvesen and Dixit, 1997
). Caspases are
synthesized as inactive zymogens, which are activated by
proteolytic processing to yield large (~18 kD) and small
(~12 kD) subunits that associate to form active enzymes (Thornberry et al., 1992
; Nicholson et al., 1995
; Stennicke
and Salvesen, 1997
). Diverse apoptotic stimuli cause the
activation of specific caspases which then initiate a protease cascade by proteolytically processing additional caspases (Srinivasula et al., 1996
; Yu et al., 1998
). Once activated, these downstream caspases kill cells by cleaving
specific molecular targets that are essential for cell viability or by activating proapoptotic factors (Liu et al., 1997
;
Salvesen and Dixit, 1997
; Enari et al., 1998
). Although
caspases have been generally shown to be cytosolic proteins (Miller et al., 1993
; Nicholson et al., 1995
; Li et al.,
1997b
), immunochemical studies have suggested that in
some instances, caspases might also be associated with the nucleus or plasma membrane (Singer et al., 1995
; Krajewska et al., 1997
; Krajewski et al., 1997
; Posmantur et al.,
1997
). Recently published data has indicated an association of certain caspases with mitochondria and endoplasmic reticulum (Chandler et al., 1998
; Mancini et al., 1998
).
The Bcl-2 family constitutes another key set of regulators of the apoptotic pathway. These proteins can function
to inhibit or induce apoptosis in a wide variety of cell systems (Oltvai and Korsmeyer, 1994; Reed, 1997
). Bcl-2
family proteins contain one to four conserved domains,
designated BH1-BH4, and most family members contain a
COOH-terminal transmembrane anchor sequence which
allows them to be associated with cellular membranes including the outer membrane of the mitochondria, the nuclear envelope and the endoplasmic reticulum (Krajewski
et al., 1993
; Lithgow et al., 1994
; Yang et al., 1995
; Reed,
1997
). The over-expression of Bcl-2 has been shown to inhibit the activation of cytoplasmic caspases after apoptotic
stimuli in several cell systems (Armstrong et al., 1996
;
Boulakia et al., 1996
; Chinnaiyan et al., 1996
; Srinivasan et al.,
1996
). However, it remains unclear how the membrane-bound Bcl-2 exerts control over the soluble cytoplasmic caspases.
Recent experiments have suggested several possible
mechanisms for Bcl-2 family function. The Bcl-2 homologue Bcl-xL has been shown to be structurally similar to
the diphtheria toxin channel-forming protein (Muchmore
et al., 1996), and several Bcl-2 family members have been
shown to form ion channels in vitro using reconstituted systems (Minn et al., 1997
; Schendel et al., 1997
; Schlesinger et al., 1997
). These data have led to the hypothesis
that Bcl-2 family members might function in cells as transmembrane channels (Vander Heiden et al., 1997). Other
experiments demonstrated that Bcl-2 and Bcl-xL block the
release of cytochrome c from mitochondria, preventing
cytochrome c-mediated caspase activation (Kluck et al.,
1997
; Yang et al., 1997
). This work suggested that Bcl-2 and Bcl-xL might act directly at the level of cytochrome c
release. However, microinjection experiments have demonstrated that inhibition of apoptosis by Bcl-xL cannot be
explained only by effects on cytochrome c compartmentalization (Li et al., 1997a
). Yet other experiments have
shown that CED-9, a Bcl-2 family member from the
roundworm Caenorhabditis elegans (Horvitz et al., 1994
),
biochemically interacts with the adapter protein CED-4,
blocking the CED-4-dependent activation of the caspase
CED-3 (Chinnaiyan et al., 1997
; Ottilie et al., 1997
; Seshagiri and Miller, 1997
; Spector et al., 1997
; Wu et al., 1997
).
This work suggested that the mammalian Bcl-2 family
members may similarly control apoptosis by directly affecting caspase activation mechanisms. Indeed, recent data
indicates that Bcl-xL can bind to the mammalian CED-4
homologue Apaf-1, at least under some conditions (Hu et
al., 1998
; Pan et al., 1998
).
Previous work has demonstrated that Bcl-2 inhibits the
onset of apoptosis, but once apoptosis is initiated, Bcl-2
does not impede the process (McCarthy et al., 1997). This
suggested that if Bcl-2 exerted direct control over caspases, it did not directly block the downstream caspases
that effect cell killing, but rather, might affect regulatory
mechanisms that trigger the downstream events. This
prompted us to consider the existence of such triggering mechanisms in the Bcl-2-containing membrane compartments of the cell, and specifically, whether regulated
caspases might be present there. This report describes the
identification and characterization of membrane-derived
caspase-3, the activation of which is suppressed by expression of Bcl-2.
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Materials and Methods |
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Cell Lines and Cell Production
697 human lymphoblastoid cells stably infected with a retroviral expression construct containing bcl-2 cDNA or a control neomycin resistance
gene (697-Bcl-2 and 697-neo cells1, respectively; obtained from Dr. John
Reed, Burnham Institute; Miyashita and Reed, 1993) were used in these
studies. The cells were maintained in mid-log phase growth in RPMI 1640 medium (Irvine Scientific) supplemented with 10% FBS (Hyclone), 0.2 mg/ml G-418 (GIBCO BRL) and 0.1 mg/ml penicillin/streptomycin (Irvine Scientific). Murine dopaminergic MN9D cells (obtained from Dr. A. Heller, University of Chicago) were grown in MEM (Irvine Scientific)
supplemented with 10% FBS, 2 mM glutamine and 0.1 mg/ml penicillin/
streptomycin. Mouse brain cortical cells were prepared at E15 of gestation
in Hank's buffered saline solution (Irvine Scientific) with 15 mM Hepes.
The tissue was briefly dissociated with 0.1% trypsin and washed thoroughly with MEM supplemented with 10% FBS and 0.4 mg/ml DNase I
(Sigma Chemical Co.), gently triturated and flash frozen. The human
breast carcinoma cell line T47D was obtained from American Type Culture Collection and cultured as suggested by the manufacturer. MCF-7
cells stably transfected with an expression plasmid coding for procaspase-3 was kindly supplied by Dr. C. Froelich (Northwestern Healthcare
Research Institute, Evanston, IL).
Subcellular Fractionation
Frozen cell pellets containing ~109 cells were thawed and resuspended in
cold hypotonic buffer (10 mM Na-Hepes, 5 mM MgCl2, 42 mM KCl, pH
7.4) supplemented with 1 mM PMSF, 1 µg/ml leupeptin, 1 µg/ml pepstatin
A, 5 µg/ml aprotinin, 0.1 mM EDTA, 0.1 mM EGTA, and 5 mM DTT
(Sigma Chemical Co.) to a density of ~1.5 × 108 cells/ml. The samples
were incubated on ice for 30 min at which time the cells were lysed using
30-40 strokes with a Dounce homogenizer. The sample was centrifuged
twice for 10 min at 500 g, 4°C to separate the nuclei. The nuclear pellets
were then washed twice in the same buffer supplemented with 1.6 M sucrose, yielding the nuclear fraction. The supernatant was then centrifuged
at 14,000 g for 30 min at 4°C to pellet the heavy membranes. The heavy
membranes were washed three times with 1.5 ml cold hypotonic buffer
containing protease inhibitors and DTT. The washed membranes were resuspended in hypotonic buffer so that the total protein concentration was ~2 mg/ml, yielding the heavy membrane fraction, that was either flash
frozen or used immediately for enzymatic measurements without freezing.
The 14,000 g supernatant was centrifuged at 100,000 g for 30 min at 4°C,
yielding a supernatant (cytoplasmic fraction) and a pellet (light membrane
fraction). Protein concentrations were measured using Protein Assay Kit
II (Bio-Rad Laboratories) with bovine serum albumin as the calibration
standard. In some experiments, cell pellets were lysed as above, but without a freezing step. To test effects of cytochrome c on caspase activity,
some samples were treated with 10 µg/ml bovine cytochrome c (Sigma Chemical Co.) throughout the entire isolation procedure. In some experiments, mitochondrial fractions were prepared from lysed 697-neo and
697-Bcl-2 cells by the rat liver mitochondrial methods of Mancini and collaborators (Mancini et al., 1998) and used without freezing.
Western Immunoblotting
Subcellular fractions (50 µg protein per lane) were resolved by SDS-PAGE on 12% or 16% gels (Novex) and transferred to Immobilon PVDF
membranes (Millipore). Membranes were blocked in PBS and 0.1% Tween
20 (PBST) + 0.4% casein (I-block, Tropix). Blots were incubated in 1 µg/ml
primary antibody diluted in PBST/casein for 1 h. After three washes in
PBST, blots were incubated for one hour in 1:15,000 dilutions of alkaline
phosphatase conjugated goat anti-rabbit IgG or goat anti-mouse IgG
(Tropix) in PBST/casein. Blots were then washed twice with PBST, twice
in assay buffer (10 mM diethanolamine, pH 10.0, 1 mM MgCl2), and then
incubated in 250 µM chemiluminescent substrate CSPD (Tropix) in assay
buffer and exposed to Biomax film (Kodak) overnight. In some cases, after the secondary antibody incubations, the blots were washed with 10 mM
Tris, pH 9.5, 1 mM MgCl2. The blots were then incubated for 30 min in
1.25 µg/ml DDAO phosphate (Amersham) dissolved in the Tris buffer.
The blots were scanned using the STORM fluorescence imager (Molecular Dynamics). The antibodies used were against Bcl-2 (clone 7; Transduction Labs), caspase-3 (Srinivasan et al., 1998), cytochrome c (clone
7H8.2C12; PharMingen), cytochrome oxidase, subunit IV (clone 1A12-A12; Molecular Probes), D4-GDP dissociation inhibitor (D4-GDI; a kind
gift of Dr. G. Bokoch, Scripps Research Institute, La Jolla, CA) and
poly(ADP-ribose) polymerase (PARP) (clone C2-10; Enzyme Systems).
Immunocytochemistry
T47D human breast carcinoma cells, MCF7 human breast carcinoma cells
transduced with a control vector or caspase-3 expression vector (Yang et al.,
1998; MCF7/cont and MCF7/casp-3, respectively) were cultured on
8-chamber permanox slides (Nalge Nunc International Corporation). The
MCF7/cont and MCF7/casp-3 cells were cultured in separate wells on the
same 8-chamber slide. When the cells reached 40-50% confluence, they
were fixed in ice-cold 10% formalin for 20 min, washed twice with PBS
and immunostained immediately. For immunostaining, fixed cells were incubated for one hour at room temperature in blocking buffer (2% normal
goat serum, 2% BSA, 0.2% nonfat milk powder, 0.4% Triton X-100 in
PBS). Cells were then incubated with affinity-purified anti-caspase-3 rabbit polyclonal antibody CSP3 (Srinivasan et al., 1998
) or purified rabbit
IgG (PharMingen; 0.3-1.2 µg/ml), plus anti-cytochrome c mouse monoclonal antibody (clone 6H2.B4; PharMingen, 0.25 µg/ml) diluted in blocking buffer, for 1 h at room temperature. After three 5-min washes in wash
buffer (PBS/0.1% Tween 20), cells were incubated for 1 h at room temperature with 0.8 µg/ml each of goat anti-rabbit IgG Alexa 488 conjugate and
goat anti-mouse Alexa 594 conjugate (Molecular Probes). Finally, cells
were washed three times, 5 min each, in wash buffer. The chamber divisions were removed and the cells were coverslipped under Citiflor mounting fluid (Ted Pella, Inc.). Immunstained cells were visualized by laser
scanning confocal microscopy and conventional fluorescence microscopy;
procaspase-3 and cytochrome c immunostaining were visualized with
FITC and Texas red filters, respectively. The confocal images are single
0.4-µm optical sections.
Enzyme Activity and Inhibition Studies
Caspase activity was measured by mixing 50 µl of an enzyme-containing fraction and 200 µl of 25 µM acetyl-Asp-Glu-Val-Asp-aminomethylcoumarin (acDEVD-amc) substrate in ICE buffer (20 mM Hepes, 1 mM EDTA, 0.1% CHAPS, 10% sucrose, 5 mM DTT, pH 7.5) in duplicate 96-well Cytoplate wells (Perseptive Biosystems). Product formation was monitored by the increase in fluorescence (ex = 360 nm, em = 460 nm) over 1-2 h at 30°C using the CytoFluor 4000 plate reader (Perseptive Biosystems). For kinetic studies, the substrate concentration was varied in the range 1-100 µM. For inhibition studies the enzyme was pretreated with 150 µl inhibitor for 30 min at room temperature before the addition of 50 µl of 50 µM substrate solution. Inhibitor IC50 values were determined using the equation:
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where FL/
t is the observed initial rate of fluorescence change at inhibitor concentration [I] and (
FL/
t)o is the initial rate fluorescence change
for the uninhibited enzyme.
Caspase Activation
Heavy membrane samples were diluted to 1 mg/ml in hypotonic buffer or
in 0.25 M sucrose, 10 mM MOPS, 2 mM EDTA, pH 7.4 (Mancini et al.,
1998) containing 5 mM DTT with or without 1% NP-40. Caspase activation was induced by adding either 60-160 ng/ml recombinant murine
caspase-1 (in bacterial lysate), 2 µg/ml of purified human granzyme B
(Enzyme Systems Products) or buffer, and incubating the samples for 60 min at 30°C or 37°C. After the activation period, the heavy membrane pellet was removed from the sample by centrifugation for 10 min at 14,000 g
at 4°C. The acDEVD-amc cleaving activities in the resulting supernatants
were corrected for the activity of the exogenous enzymes. To examine the
time course of spontaneous activation of caspase activity from membranes, 50 µl of heavy membrane slurry containing 50-100 µg total protein was mixed with 200 µl hypotonic buffer containing 25 µM acDEVD-amc substrate and 6 mM DTT in 96-well Cytoplates and fluorescence was
measured over time. At selected time points, aliquots were removed from some wells, centrifuged for 10 min at 14,000 g to remove the heavy membranes, and then the supernatant was added back into the 96-well plate to
measure the soluble acDEVD-amc cleavage activity. In some experiments, subcellular fractions were treated with 1 µg/ml bovine cytochrome
c (Sigma Chemical Co.) and 50 µM dATP (New England Biolabs) for 40 min at 30°C before measurement of caspase activity.
Production of Recombinant Caspase-1 and Caspase-3 Proteins
BL21 (DE3) cells harboring a plasmid containing the cloned human
caspase-3 cDNA (Fernandes-Alnemri et al., 1994; provided by Dr. E. Alnemri, Thomas Jefferson University, Philadelphia, PA) ligated into the
BamHI/XhoI sites of pET21b (Novagen) were grown in one liter LB medium containing 0.1 mg/ml ampicillin at 37°C. When the culture density
reached A600 = 1, IPTG (Sigma Chemical Co.) was added to a concentration of 1 mM and the culture was incubated at 25°C for 3 h. The cells
were harvested by centrifugation at 2,000 g for 15 min at 4°C. The cells
were lysed using one freeze-thaw cycle in 100 ml binding buffer (20 mM
Tris-HCl, 500 mM NaCl, 5 mM imidazole, 0.1% Triton X-100) with 0.1 mg/ml lysozyme. Cell debris was removed from the sample by centrifugation at 20,000 g, for 30 min at 4°C. The lysed cells were treated just before centrifugation with 0.5 mM MgCl2 and 2 µg/ml DNase I (Sigma Chemical Co.) to reduce viscosity. The supernatant was filtered through a 0.45-µm
syringe filter and loaded onto a 1 ml Ni2+-charged HiTrap chelating column (Amersham Pharmacia) at a 1 ml/min flow rate. The column was
washed at 1 ml/min with 10 ml binding buffer followed by 10 ml binding
buffer containing 60 mM imidazole. The caspase-3 protein was eluted
from the column using a 30-ml linear gradient of imidazole (60-500 mM).
Recombinant murine caspase-1 was expressed using BL21 (DE3) pLys S cells harboring pET3ap30mICEFLAG plasmid (a generous gift of Drs. H.R. Horvitz and Ding Xue, Massachusetts Institute of Technology) which contains the p30 caspase-1 cDNA inserted into the NdeI/BamHI sites of the pET3a expression vector (Novagen). A 3-liter culture was grown at 37°C in induction medium (20 g/liter tryptone, 10 g/liter yeast extract, 6 g/liter NaCl, 3 g/liter Na2HPO4, 1 g/liter KH2PO4, 1 mM MgCl2, 0.1 mM CaCl2, pH 7.4) containing 0.1 mg/ml ampicillin and 0.025 mg/ml chloramphenicol. When the culture reached a density of A600 = 1.0, IPTG was added to 1 mM and the culture was shaken at 25°C for 3 h. The cells were collected by centrifugation at 2,000 g for 15 min at 4°C and resuspended in 100 ml cold buffer containing 25 mM Tris-HCl, pH 8.0, 25 mM KCl, 0.1% Triton X-100, and 0.1 mg/ml lysozyme (InovaTech). The cells were lysed using one freeze/thaw cycle and the lysate was clarified by treating the sample with 2 µg/ml DNase I, 0.5 mM MgCl2 for 60 min and then centrifuging at 20,000 g for 30 min at 4°C to remove cell debris.
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Results |
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Characterization of Subcellular Fractions from 697 Cells
Subcellular fractions were prepared from 697 cells stably
infected with retroviral constructs expressing either bcl-2
cDNA or a neomycin resistance gene (697-Bcl-2 and 697-neo cells, respectively; Miyashita and Reed, 1993). Nuclear, heavy membrane, light membrane, and cytosolic
fractions were isolated from these cells, and were characterized by Western blot analysis with antibodies specific for proteins with distinct known subcellular distributions.
Antibodies used were directed against cytochrome oxidase, specific for mitochondrial inner membrane (Tzagoloff, 1982
), PARP, specific for nuclei (Berger, 1985
),
D4-GDP dissociation inhibitor (D4-GDI), specific for cytoplasm (Na et al., 1996
) and Bcl-2. As shown in Fig. 1,
the mitochondrial marker was found almost exclusively in
the heavy membrane fraction, the nuclear marker only
in the nuclear fraction, and the cytoplasmic marker only in
the cytoplasmic fraction. Thus, the fractionation methods
used generated fractions with the expected subcellular distribution of marker proteins. Importantly, we could not
detect cytoplasmic contamination of the nuclear and membrane fractions, and detected only minimal mitochondrial
contamination of nuclear fractions (the diffuse D4-GDI
reactive band in the nuclear fraction shown in Fig. 1 is
nonspecific). Western analysis of fractions from 697-neo
cells with an antibody to human Bcl-2 (Fig. 1) demonstrated strong reactivity in nuclear and heavy membrane
fractions, weaker reactivity in the light membrane fraction, and undetectable signal in cytoplasm, in accord with
previous results (Krajewski et al., 1993
; Yang et al., 1995
;
Lithgow et al., 1994
). Similar analysis of fractions from
697-Bcl-2 cells showed significant overexpression.
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Subcellular Distribution of acDEVD-amc Cleavage Activity in 697 Cells
Preliminary experiments indicated that caspase activity
was associated with membranes derived from unstimulated cells. To determine the subcellular distribution of
such caspases, we quantitated the caspase activity in the
subcellular fractions from 697-neo cells by incubating
them with the substrate acDEVD-amc, and measuring the increase in fluorescence over the subsequent 2 h.
acDEVD-amc is a useful substrate for all caspases characterized to date, with the exception of caspase-2 (Talanian et al., 1997; data not shown). While most of the
acDEVD-amc cleavage activity (~75%) was in the cytoplasmic fraction, a substantial amount of the cleavage activity was found in the nuclear, heavy membrane and
light membrane fractions (Fig. 2, a and c). The major
acDEVD-amc cleaving activity in each fraction was indeed caspase activity since it was potently blocked by
specific caspase inhibitors (Table I, column 1, and data
not shown).
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Bcl-2 Suppresses Membrane-derived Caspase Activity
Next, we examined the effect of Bcl-2 on the caspase activities in the various subcellular fractions. When subcellular fractions derived from 697-Bcl-2 cells were
prepared and incubated with acDEVD-amc substrate,
substantially reduced caspase activity was observed in the
nuclear and heavy membrane fractions compared with
697-neo cells (Fig. 2 b). This Bcl-2 effect was evident when the caspase activity was measured on a per cell basis or
per mg protein and resulted in an 80-90% reduction in
caspase activity in these fractions (Fig. 2, b and d). The effect of Bcl-2 expression on caspase activity in these fractions was specific, since little if any suppression was seen in
the activities observed in the cytoplasmic or light membrane fractions (Fig. 2, a-d). These observations suggested
that the membrane-associated caspase activity was not simply derived from a small percentage of apoptotic cells
in the 697-neo cultures whose numbers were suppressed in
the 697-Bcl-2 cultures. If that were the case, we would also
have expected to see major differences in caspase activities between cytoplasmic fractions derived from 697-neo
vs. 697-Bcl-2 cells. Indeed, control experiments demonstrated that when 697-neo cells were induced to undergo
apoptosis by staurosporine treatment, the major increase in caspase activity was found in the cytoplasm (data not
shown). The ability of Bcl-2 to suppress membrane-associated caspase activity was not limited to the 697 lymphoblastoid cells, since similar effects were observed in Jurkat
T cells and FL5.12 cells (data not shown). Since our data,
as well as other published studies, have demonstrated that
Bcl-2 protein is found predominantly in nuclear envelope
and heavy membrane fractions (Fig. 1; Krajewski et al.,
1993; Yang et al., 1995
), our results were compatible with
the possibility that Bcl-2 might act locally to regulate this
membrane-derived caspase activity. In an effort to begin analyzing such mechanisms, we further characterized this
membrane-derived, Bcl-2-suppressible caspase activity
and focused our efforts on the heavy membrane fraction.
Membrane-derived Caspase Activity Reflects Spontaneous Activation and Membrane Release
It was possible that the membrane associated caspase activity was due either to an active membrane-bound enzyme, or alternatively, to the spontaneous activation and release of a soluble active enzyme. We therefore designed a set of experiments to distinguish between these two possibilities. First, to freshly prepared heavy membranes derived from 697-neo cells (neo-membranes), we immediately added hypotonic buffer and acDEVD-amc substrate at room temperature, and measured the emergence of amc fluorescence over a 90-min period (Fig. 3 a). The data demonstrate that there is little detectable fluorescence change over the first 15 min of incubation, but after this lag period, the rate of amc production increases markedly (Fig. 3 a). These results indicated that the freshly prepared membranes did not contain active caspase, but that activation occurred spontaneously during the incubation period. To assess whether this newly activated caspase was soluble or membrane bound, membranes were incubated for different periods of time, after which the samples were centrifuged and the resulting supernatants were assayed for caspase activity with acDEVD-amc substrate. These data demonstrated that very little caspase activity was present in the supernatant initially, but that soluble caspase activity appeared thereafter (Fig. 3 b). Quantitative analysis of these data demonstrated that for each supernatant, fluorescence increased linearly, indicating that once released from the membranes, no further activation occurred. Furthermore, the slopes of these curves (Fig. 3 b) approximate the instantaneous slopes of the corresponding time points in the progress curve for the heavy membrane slurry (Fig. 3 a). Therefore, all of the caspase-3 activity can be accounted for in the supernatant fraction, indicating that all active enzyme had been released from the membranes. In contrast to the neo-membranes, membranes derived from the 697-Bcl-2 cells (Bcl-2-membranes) failed to generate significant acDEVD-amc cleaving activity (Fig. 3 a).
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Procaspase-3 Is Present in Heavy Membranes from Both 697-neo and 697-Bcl-2 Cells
The lack of acDEVD-amc cleaving activity in the Bcl-2-membranes could be due either to the absence of activatable procaspase or suppression of procaspase activation. To distinguish between these alternatives, we first
performed Western blot analysis on the membrane fractions with antibodies specific for caspase-3, since the measured acDEVD-amc cleavage activity is in fact due to
caspase-3 (see below). The results (Fig. 4 a) demonstrate
the presence of a caspase-3 reactive band that is of similar
intensity in both the neo-membranes and Bcl-2-membranes, and that is approximately the size expected for
the procaspase zymogen. Interestingly, the electrophoretic
mobility of the membrane-derived bands was slightly slower than that of cytoplasmic procaspase-3. To further
demonstrate the presence of procaspase-3 in both neo-
and Bcl-2-membranes, we attempted to activate these
fractions by treatment with exogenous caspase-1, since
procaspases can be activated by proteolytic cleavage at aspartic acid residues between their large and small subunits (Srinivasula et al., 1996; Stennicke and Salvesen, 1997
;
Salvesen and Dixit, 1997
). As we have shown above, membranes derived from Bcl-2 cells showed almost no caspase
activity when measured under our standard conditions.
However, treatment of the Bcl-2-membranes with caspase-1 caused a robust induction of enzymatic activity (Fig. 4 b). The neo-membranes were also activated by exogenous caspase-1. But importantly, after activation, the
resulting caspase activities from the Bcl-2- and neo-membranes were always similar, within a factor of two (Fig. 4
b). Together with the procaspase-3 immunoblot data, this
supports the conclusion that comparable levels of procaspase-3 are present in neo- and Bcl-2-membranes.
|
Caspase-1-treatment of membranes not only activated the endogenous caspase activity, but also released it from the membranes, since the activity remained in the supernatant when the membranes were removed by centrifugation (Fig. 4 b). This induction and release were due to the proteolytic activity of caspase-1, since the caspase-1 activation could be completely blocked by 200 nM acYVAD-aldehyde which inhibits caspase-1, but not the membrane caspase, at this concentration (data not shown). Our results indicate that both neo- and Bcl-2-expressing cells contain similar amounts of a membrane-associated inactive procaspase that can be activated by caspase-1. However, without exogenous caspase treatment, only membranes derived from the neo-expressing cells demonstrated spontaneous caspase activation.
To further document the presence of procaspase-3 in
heavy membrane fractions we performed immunocytochemical studies, using adherent cell lines for ease of experimentation. First, we demonstrated that our affinity-purified antibody CSP3, generated against recombinant caspase-3 and used in our Western blots (Fig. 4 a; Srinivasan et al., 1998), was specific when used as an immunocytochemical reagent. Our staining results showed that the
antibody did not react with MCF7/cont breast carcinoma
cells (Fig. 5 f) which lack procaspase-3 due to a genetic
deletion (Li et al., 1997a
; Janicke et al., 1998
). However,
the antibody showed intense staining when reacted with
MCF7/casp-3 cells overexpressing recombinant procaspase-3 (Fig. 5 d). To analyze the distribution of endogenous procaspase-3 in an untransfected cell line we used
T47D breast carcinoma cells. The caspase-3 antibody demonstrated both diffuse and punctate staining (Fig. 5 a).
Much of the punctate staining colocalized with mitochondria, as visualized by anti-cytochrome c antibody (Fig. 5, b
and c). Nonspecific purified rabbit IgG did not stain these
cells (Fig. 5 h). These results confirm that procaspase-3 immunoreactivity associates with heavy membrane elements
in cells, as was also shown using other cell types (Mancini
et al., 1998
).
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Enzymological Characterization of the Induced and Spontaneous Caspase Activities
We further characterized the membrane-derived caspase
activities by measuring the inhibition of acDEVD-amc
cleavage by several peptide aldehyde inhibitors (Table I).
The IC50 values for the inhibition of acDEVD-amc activity
derived from activated Bcl-2 membranes are quite similar
to those for the inhibition of the activity derived from neo-membranes, suggesting that caspase-1 activates the same
procaspase in both membrane preparations. Furthermore, these IC50 values are similar to those for the spontaneously
activated acDEVD-amc activity derived from neo-membranes, suggesting that the spontaneous and caspase-1-induced activities derive from the same caspase. In all
cases, the inhibition data fit well to a simple competitive
inhibition curve as described in the Materials and Methods, suggesting that each acDEVD-amc activity arose
from a single caspase rather than a mixture of enzymes.
The observed IC50 values for the membrane associated
caspases are very similar to those for purified fully processed recombinant human caspase-3. Kinetic measurements also indicate that Km values for hydrolysis of
acDEVD-amc by the membrane-derived caspases (10 µM)
are similar to that observed with fully processed caspase-3
(Nicholson et al., 1995). NH2-terminal microsequence
analysis of activated, affinity-purified heavy membrane
caspase confirms that this enzyme is indeed human caspase-3 (manuscript in preparation).
To determine if the presence of membrane-associated
caspase activity is a general property of mammalian cells,
we measured the acDEVD-amc cleavage activity in heavy
membranes from two other cell sources: mouse E15 primary brain cortical cells and the mouse dopaminergic
MN9D cell line (Choi et al., 1992). Heavy membrane fractions were prepared using identical procedures to those
used for the 697 cells and were activated with caspase-1.
These fractions contained a membrane-associated caspase
activity with similar cleavage activities per mg protein as
observed in 697 cells (data not shown) and that was
blocked by caspase inhibitors with a similar potency to
that observed with fractions derived from 697 cells or with
recombinant caspase-3 (Table I). We conclude that the existence of membrane-derived caspase activity is not specific to 697 cells, but appears to be a more general phenomenon.
Addition of Exogenous Cytochrome c Does Not Activate Membrane-associated Procaspase-3
Several recent reports have shown that the release of cytochrome c from mitochondria can cause the activation of
cytoplasmic caspase-3 (Liu et al., 1996; Li et al., 1997a
).
Other reports have demonstrated that cytochrome c is released from mitochondria after apoptotic insults and that
Bcl-2 can inhibit that release (Kluck et al., 1997
; Yang et al.,
1997
). Thus it was possible that the difference we observed
between caspase activities in heavy membranes from Bcl-2-
and neo-expressing cells simply reflected inhibition by
Bcl-2 of cytochrome c release during preparation of the
heavy membrane fractions or during subsequent incubation of these fractions. To investigate this possibility, we
performed cell fractionation in the presence of exogenous
cytochrome c and measured whether this influenced caspase activation. If the Bcl-2-membranes had low caspase
activity because of a Bcl-2 effect on cytochrome c sequestration, then the addition of exogenous cytochrome c during membrane fractionation should increase the caspase
activity derived from those membranes to the levels seen
in membranes from neo-cells. Accordingly, during the
fractionation procedure for heavy membranes from neo-
and Bcl-2-expressing cells, we added 10 µg/ml cytochrome c to the cell lysate immediately after homogenization, and
10 µg/ml to the buffers used to suspend and wash the
heavy membranes. This concentration of cytochrome c
was chosen since it represents the estimated total amount
of cytochrome c present in the starting cell pellets (Li et al.,
1997a
). Finally, these membranes were resuspended in 1 µg/ml cytochrome c plus 50 µM dATP, incubated, and
then assayed for acDEVD-amc cleaving activity (Fig. 6 a). This activity was compared with that from our usual membrane preparations prepared without cytochrome c, and
incubated without cytochrome c or dATP. The data demonstrate that inclusion of cytochrome c during membrane
fractionation and incubation has no effect on membrane-derived caspase activity; the activity in the membranes derived from Bcl-2-expressing cells remained low compared
with the activity in the neo-membranes, and furthermore,
there was also no effect of cytochrome c on the caspase activity derived from the neo-membranes (Fig. 6 a). Although the cytochrome c treatments did not activate the
membrane-associated caspase, the enzyme could still be
activated by subsequent treatment with exogenous caspase-1 (data not shown). The lack of a cytochrome c effect on the activation of the membrane caspase was not
due to an inactive preparation of cytochrome c, since the
acDEVD-amc cleavage activity of the cytoplasmic fractions from both neo and Bcl-2 cells were strongly activated
by inclusion of cytochrome c during fractionation and assay (Fig. 6 b). We conclude that Bcl-2 expression suppresses the activation of the membrane-associated procaspase-3, but that this effect is not overcome by addition
of exogenous cytochrome c. Furthermore, Bcl-2 overexpression did not affect the ability of cytochrome c to activate caspase-3 in cytoplasmic fractions.
|
Release of Membrane-associated Caspase Activity Is Not Due to Simple Leakage from Organelles
A recent report described the presence of procaspase-3 in
the intermembrane space within mitochondria (Mancini
et al., 1998). Thus, it was possible that this material could
account for the activatable caspase activity that we measured in our mitochondria-containing heavy membrane
fractions. Furthermore, it was possible that the spontaneous activity that we measured in membrane fractions from 697-neo cells was due to leakage of active caspase from
mitochondria, and that mitochondria isolated from 697-Bcl-2 cells were simply less leaky (Yang et al., 1997
). However, several experiments suggested that the activity we
measured was not due to leakage from mitochondria, and
that the activity is distinct from that described by Mancini et al. (1998)
.
First we tested whether the addition of 1% NP-40 to
neo-membranes affected the level of either spontaneous
activity or the activity induced by caspase-1 or granzyme
B. We reasoned that if procaspase and/or active caspase
was sequestered within organelles, then enhanced activity
would be measured in the presence of NP-40. Treatment
with 1% NP-40 was sufficient to release almost all of the
cytochrome c present in heavy membrane preparations
(data not shown). Furthermore, it was shown by Mancini
and colleagues that treatment of their mitochondrial preparations with 1% NP-40 allowed granzyme B to cleave
procaspase-3 whereas no cleavage was observed in the absence of detergent (Mancini et al., 1998). However, our results demonstrate that 1% NP-40 had little effect either on
spontaneous activity or the activity induced by treatment with caspase-1 or granzyme B (Fig. 7 a). Next, to analyze
whether membrane preparations from 697-Bcl-2 cells may
have low spontaneous activity due to enhanced sequestration of a caspase, we added acDEVD-amc to Bcl-2- and
neo-membrane preparations, incubated them in buffer
alone or buffer plus 1% NP-40, and measured the appearance of fluorescence (Fig. 7 b). The results indicate that 1% NP-40 had only a minor effect on the magnitude or
rate of fluorescence increase. Preparations derived from
697-Bcl-2 cells had low activity regardless of whether 1%
NP-40 was present, demonstrating that this low level of activity was not due to sequestration of an active caspase. Finally, we prepared mitochondrial fractions from 697-neo
and 697-Bcl-2 cells using the methods described by Mancini et al. (1998)
to more directly assess the relationship
between our results and their published data. As shown in
Fig. 7 c, fractions from both 697-neo and 697-Bcl-2 made
by these methods have granzyme B-activatable caspase activity in the absence of NP-40. However, in the presence
of 1% NP-40, granzyme B treatment yielded enhanced caspase activity (Fig. 7 c). Thus, under these conditions,
granzyme B generates caspase activity in both NP-40-independent and -dependent manners.
|
![]() |
Discussion |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
The present work was motivated by genetic and biochemical studies that suggested that the Bcl-2 homologue CED-9
functions by regulating the activity of the caspase CED-3
through protein-protein interactions (Horvitz et al., 1994;
Chinnaiyan et al., 1997
; Ottilie et al., 1997
; Seshagiri and
Miller, 1997
; Spector et al., 1997
; Wu et al., 1997
). Given
that Bcl-2 and the related death-inhibiting protein Bcl-xL
are both localized to intracellular membranes (Krajewski et al., 1993
; González-Garcia et al., 1994
), we reasoned
that these molecules may act locally to regulate a membrane-associated caspase. As an initial step in investigating
this hypothesis, we first demonstrated that intracellular
membrane fractions do in fact contain an activatable procaspase, which when characterized, was shown to be caspase-3. Immunocytochemical evidence further supported
the conclusion that heavy membrane components contain procaspase-3 (Fig. 5). Quantitatively, the amount of membrane-derived caspase activity is relatively small compared
with that in the cytoplasmic fraction; the total acDEVD-amc cleavage activity in the heavy membrane fraction was
generally ~5-10% of the total cytoplasmic activity in unstimulated cells (Fig. 2). In cells stimulated to undergo
apoptosis, there are increases in caspase activity in both
membrane and cytoplasmic fractions, but the percentage that is membrane-associated remains low (data not
shown). However, our results suggest that the pool of
membrane-associated caspase is uniquely regulated by
Bcl-2. Heavy membrane and nuclear fractions derived
from 697-Bcl-2 cells demonstrated only low levels of spontaneous activation of caspase activity compared with similar fractions derived from control 697-neo cells (Figs. 2, 3,
and 7 b). The heavy membranes from the Bcl-2-expressing
cells did, however, contain appreciable amounts of procaspase-3, measured in two ways: directly by Western
analysis (Fig. 4 a), and indirectly, by measurement of
acDEVD-amc cleaving activity after activation by exogenous caspase-1 (Fig. 4 b). This demonstrated that Bcl-2
was not affecting the ability of procaspase-3 to associate
with membranes, but rather, it exerted specific control
over enzyme activation. The Western blot analysis demonstrated that the membrane-associated procaspase-3 had an
electrophoretic mobility distinct from that of cytoplasmic procaspase-3. However, we do not yet know the biochemical basis for this observation.
There are many possible mechanisms by which procaspase-3 could become associated with heavy membranes.
It has recently been shown that soluble procaspase-3 is
present within the mitochondrial intermembrane space
(Mancini et al., 1998), and thus it was possible that this
material represented the Bcl-2-regulated activity that we
observed in our heavy membrane fractions. However,
whereas the procaspase sequestered in the intermitochondrial space is protected from activation by exogenous
granzyme B (Mancini et al., 1998
), we observed that heavy
membrane caspase activity in our preparations is readily
activated by exogenous caspase-1 (Fig. 4 b) or granzyme B
(Fig. 7). In addition, during the course of membrane-associated procaspase activation, active enzyme is continuously released into the medium (Fig. 3) in accord with the
idea that a bound, inactive procaspase is converted to a
soluble, active enzyme. Addition of the membrane-permeabilizing detergent NP-40 to our standard heavy membrane fractions had only a minimal effect on either spontaneous or caspase-1 activated caspase activity (Fig. 7, a
and b), suggesting that these activities were not sequestered within organelles. Thus, we favor the hypothesis that
the Bcl-2-regulated procaspase-3 is physically associated
with membranes and not simply soluble within a sequestered compartment, mitochondrial or other. However, we
did observe that addition of NP-40 enhanced the granzyme B-activated caspase activity when fractions were
prepared using methods designed to isolate intact mitochondria (Fig. 7 c). Thus, it is possible that mitochondria
may contain two pools of procaspase, one that is accessible
to activators only with detergent, and one accessible without detergent.
Caspase activity rises dramatically in the cytoplasm of
cells induced to undergo apoptosis and the generation of
this activity is blocked by various agents, including Bcl-2,
that inhibit apoptosis (Armstrong et al., 1996, 1997
; Chinnaiyan et al., 1996
; Srinivasan et al., 1996
). Thus, we considered whether the inhibition of caspase activation by
Bcl-2 in heavy membranes was an indirect consequence of
general apoptotic inhibition. For example, the caspase activity in neo-membranes could simply reflect the existence
of more apoptotic cells in unstimulated 697-neo cultures
vs. 697-Bcl-2 cultures. However, two observations argue
against this interpretation. First, the inhibition of caspase
activity by Bcl-2 in unstimulated cultures was only seen in
heavy membrane and nuclear fractions. However, in 697 cells induced to undergo apoptosis by staurosporine treatment, or in Jurkat cells stimulated with anti-Fas antibody,
the largest increase in caspase activity is seen in the cytoplasmic fraction, and this is blocked by Bcl-2 (data not
shown). The absence of an effect of Bcl-2 on the cytoplasmic caspase activity in unstimulated cells suggests that the
difference between activity in the neo-membranes and
Bcl-2-membranes is not simply a passive consequence of
more apoptotic cells in the 697-neo cultures. Note, however, that although the number of apoptotic cells in resting
697-neo cultures is low (<5%), this represents generally
two to three times more apoptotic cells than in 697-Bcl-2
cultures, as measured by Hoechst dye or annexin V staining (data not shown). Second, the activity that we measured in neo-membranes does not reflect procaspases that were activated during prior apoptotic events since these
membranes do not contain pre-existing active caspase;
rather, the activity is generated during subsequent incubation (Fig. 3). However, it is possible that the ability to autoactivate heavy membrane caspase is a specific property
of a subpopulation of pre-apoptotic cells. A recent report
suggests that caspases activated in the cytoplasm during
apoptosis can subsequently become membrane bound
(Chandler et al., 1998
) but this phenomenon does not appear to account for our observation of an activatable procaspase in membranes derived from unstimulated cells.
Another striking difference between membrane bound
and cytoplasmic caspase-3 relates to activation by cytochrome c. The membrane-associated procaspase showed
no activation in the presence of exogenous cytochrome c,
even when the cytochrome c was present throughout the
membrane isolation procedure and in the caspase assay
buffers. It is unlikely that the failure of cytochrome c to activate this caspase was due to an inability of the cytochrome c to obtain access to the procaspase, since activation was readily effected by exogenous granzyme B. It was
possible that cytochrome c failed to enhance caspase activation in neo-membranes because endogenous cytochrome c
was saturating the activation mechanism. However, this is
also unlikely because NP-40 treatment released endogenous cytochrome c but did not inhibit caspase activation
(Fig. 7, a and b). In contrast, cytoplasmic procaspase-3
could be robustly activated by cytochrome c when added
either during fractionation, during assay, or both (Fig. 6
and data not shown). Note also that cytochrome c effectively activated cytoplasmic caspases regardless of whether the cytoplasm was derived from neo- or Bcl-2-expressing
cells, in accord with published data indicating that Bcl-2
functions upstream of cytochrome c-induced caspase activation (Kluck et al., 1997; Duckett et al., 1998
). Thus, our
results imply the existence of two distinct procaspase-3 activation mechanisms: a Bcl-2-regulated pathway specific
to membranes and insensitive to exogenous cytochrome c;
and a cytoplasmic activation pathway, directly activated by
cytochrome c, but which is not directly Bcl-2-regulated.
However, the activation of the cytoplasmic procaspase-3 is
indirectly controlled by Bcl-2, since Bcl-2 blocks all downstream caspase events associated with apoptotic stimuli
(Armstrong et al., 1996
; Chinnaiyan et al., 1996
; Boulakia
et al., 1996
).
The fact that the heavy membrane procaspase-3 from
control cells becomes active spontaneously implies that it
undergoes proteolytic cleavage by some protease also
present in the membranes. It is possible that this cleavage
occurs through autoactivation without the intervention of
a separate activating protease, but this may be unlikely;
procaspase-3 in the cytoplasm does not self-activate, but
requires a first cleavage by caspase-9 before a second autocatalytic step (Martin et al., 1996; Li et al., 1997b
). Thus,
analogous to the caspase-9/procaspase-3 activating mechanism described in the cytoplasm, we speculate that the
heavy membrane and nuclear fractions also contain an activating caspase capable of cleaving the membrane-associated procaspase-3. This membrane-associated activating
caspase could be a form of caspase-9, or perhaps another caspase, that is regulated by Bcl-2.
The specific effect of Bcl-2 on the activation of the membrane-associated caspase suggests that this caspase may play an important role in controlling apoptosis. We speculate that the membrane-associated caspase might function as a specific trigger to promote downstream apoptotic activity leading ultimately to cytoplasmic caspase activation. If so, this would place Bcl-2 at a critical control point, regulating the trigger, and thereby inhibiting diverse apoptotic events in cells.
![]() |
Footnotes |
---|
Received for publication 27 March 1998 and in revised form 26 January 1999.
Address correspondence to Lawrence C. Fritz, IDUN Pharmaceuticals,
11085 N. Torrey Pines Road, Suite 300, La Jolla, CA 92037. Tel.: (619)
623-1330. Fax: (619) 625-2677. E-mail: lfritz{at}idun.com
We would like to thank Drs. E. Alnemri, C. Froelich, H.R. Horvitz, J. Reed, and D. Xue for cDNA clones and cell lines, E. Monosov for help with confocal microscopy, and Lisa Trout and Chris Knowles for invaluable assistance in the preparation of this manuscript.
![]() |
Abbreviations used in this paper |
---|
697-neo cells, 697 cells stably infected with a retrovirus expressing the neomycin resistance gene; 697-Bcl-2 cells, 697 cells stably infected with a retrovirus expressing human bcl-2 cDNA; acDEVD-amc, acetyl-Asp-Glu-Val-Asp-aminomethylcoumarin; acYVAD-ald, acetyl-Tyr-Val-Ala-Asp-aldehyde; Bcl-2-membranes, heavy membranes prepared from 697-Bcl-2 cells; neo-membranes, heavy membranes prepared from 697-neo cells; PARP, poly(ADP-ribose) polymerase.
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