J Biol Chem, Vol. 274, Issue 32, 22635-22645, August 6, 1999
Characterization of Caspase Processing and Activation in
HL-60 Cell Cytosol Under Cell-free Conditions
NUCLEOTIDE REQUIREMENT AND INHIBITOR PROFILE*
Peter W.
Mesner Jr.a,
Keith C.
Bibleb,
Luis M.
Martinscd,
Timothy J.
Kottkea,
Srinivasa M.
Srinivasulae,
Phyllis A.
Svingena,
Tamie J.
Chilcotef,
Guriq S.
Basif,
Jay S.
Tungf,
Stan
Krajewskig,
John C.
Reedg,
Emad S.
Alnemrie,
William C.
Earnshawch, and
Scott H.
Kaufmannai
From the Divisions of a Oncology Research and
b Medical Oncology, Mayo Clinic,
Rochester, Minnesota 55901, the c Institute of Cell & Molecular Biology, University of Edinburgh,
Edinburgh, EH9 3JR Scotland, United Kingdom, the
e Center for Apoptosis Research and Department of Microbiology
and Immunology, Kimmel Cancer Institute, Thomas Jefferson
University, Philadelphia, Pennsylvania 19107, f Elan
Pharmaceuticals, South San Francisco, California 94080, and
g Burnham Institute, La Jolla, California 92037
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ABSTRACT |
The present studies compared caspase activation
under cell-free conditions in vitro and in
etoposide-treated HL-60 leukemia cells in situ.
Immunoblotting revealed that incubation of HL-60 cytosol at 30 °C in
the presence of cytochrome c and ATP (or dATP) resulted in
activation of procaspases-3, -6, and -7 but not -2 and -8. Although
similar selectivity was observed in intact cells, affinity labeling
revealed that the active caspase species generated in vitro
and in situ differed in charge and abundance. ATP and dATP
levels in intact HL-60 cells were higher than required for caspase
activation in vitro and did not change before caspase activation in situ. Replacement of ATP with the poorly
hydrolyzable analogs 5'-adenylyl methylenediphosphate, 5'-adenylyl
imidodiphosphate, or
5'-adenylyl-O-(3-thiotriphos-phate) slowed caspase
activation in vitro, suggesting that ATP hydrolysis is
required. Caspase activation in vitro was insensitive to
phosphatase and kinase inhibitors (okadaic acid, staurosporine, and
genistein) but was inhibited by Zn2+, aurintricarboxylic
acid, and various protease inhibitors, including 3,4-dichloroisocoumarin,
N
-p-tosyl-L-phenylalanine
chloromethyl ketone,
N-p-tosyl-L-lysine
chloromethyl ketone, and
N-(N-benzyloxycarbonylphenylalanyl)alanine
fluoromethyl ketone, each of which inhibited recombinant caspases-3,
-6, -7, and -9. Experiments with anti-neoepitope antiserum confirmed
that these agents inhibited caspase-9 activation. Collectively, these
results suggest that caspase-9 activation requires nucleotide
hydrolysis and is inhibited by agents previously thought to affect
apoptosis by other means.
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INTRODUCTION |
Apoptosis is a morphologically distinct form of physiological cell
death that is widely observed in nature (1, 2). Studies performed over
the past 5 years have revealed that many of the changes observed in
apoptotic cells result from the action of a family of
cysteine-dependent aspartate-directed intracellular proteases (now termed caspases) on their substrates (3-7). As a
result, considerable attention is now focused on understanding the
factors that control caspase activation and activity.
Several pathways of caspase activation have been identified. One
involves ligation of certain receptors (e.g. CD95 or the type 1 tumor necrosis factor-
receptor), recruitment of adaptor proteins such as FADD/Mort1, binding of procaspases-8 and -10 to the
adaptor molecules, activation of these initiator caspases, and
subsequent proteolytic activation of the downstream caspases-3, -6, and
-7 (7-10). This canonical pathway appears to account, at least in
broadstroke, for the events initiated by ligation of a number of death receptors.
An alternative pathway of caspase activation (7, 11) appears to be
involved in other apoptotic deaths (12, 13). Many proapoptotic stimuli
cause mitochondria to release cytochrome c to the cytosol,
where it binds to a docking protein called Apaf-1, inducing a
conformational change in Apaf-1 that facilitates binding and activation
of procaspase-9 (14-19). Caspase-9 then proteolytically activates
caspases-3 and -7; the former activates caspase-6 (16, 18).
Previous studies have indicated that this cytochrome
c/Apaf-1/caspase-9 pathway can be reconstituted in
vitro by incubating cytosol from nonapoptotic cells with purified
cytochrome c and dATP (13, 14, 20, 21). Although this
pathway has been intensively studied, several aspects remain poorly
understood. First, the role of ATP or dATP is unclear. Because mutation
of the ATP-binding site abolishes the caspase activating activity of
the Apaf-1 homolog ced-4 (22, 23), it has generally been assumed that
the nucleoside triphosphate is hydrolyzed during caspase activation.
Consistent with this possibility, ATP depletion has been shown to
abrogate caspase activation and subsequent apoptotic events in damaged
cells (24, 25). Srinivasula et al. (18), however, observed
that a fragment of Apaf-1 can facilitate caspase-9 activation in a
nucleotide-independent fashion. More recently, Kuida et al.
(26) reported that caspase activation can occur upon incubation of
brain cytosol in the absence of exogenous dATP and cytochrome
c. These observations raise the possibility that nucleotide
hydrolysis might not be required for caspase activation. Second,
because the vast majority of studies have focused upon the activation
of caspase-3, it is unclear how faithfully the events produced in
vitro reflect the selective activation of a subset of the
available procaspases that is observed in situ. Finally, the
effects of various inhibitors on this caspase activation pathway remain
to be elucidated.
A number of inhibitors of the apoptotic process have been previously
identified. Early experiments established that Zn2+ (27)
and ATA1 (94) decrease
thymocyte apoptosis. Although these effects were initially attributed
to inhibition of apoptotic nucleases, subsequent observations have
indicated that Zn2+ inhibits active caspases (28-30) or
apoptotic events further upstream (31). A variety of additional
compounds have been observed to inhibit apoptosis in intact cells.
These include the caspase inhibitor ZVAD-fmk (32); agents such as TLCK,
TPCK, and DCI (33-43), which are often regarded as serine protease
inhibitors; and ZFA-fmk (39), which is considered a specific inhibitor
of sulfhydryl-dependent cathepsins. Although it is now
clear that ZVAD-fmk can inhibit caspase-9 activation (13, 20) and
activity (44), the possibility that the other agents inhibit caspase
activation has not been previously explored.
In the present study, we have compared caspase activation in HL-60
cells in situ and in HL-60 cytosol in vitro.
Earlier studies established that etoposide-induced apoptosis in HL-60
cells is accompanied by cleavage of multiple caspase substrates (34, 45, 46) and that this cleavage can be inhibited by treatment of cells
with TPCK, TLCK, or Zn2+ (34). Subsequent reports have
demonstrated that a subset of the available caspase precursors is
activated during the course of apoptosis in this model system (47) and
that cytochrome c release from mitochondria precedes caspase
activation (15, 48). In the present study, the cohort of caspases
activated by treatment of HL-60 cytosol with cytochrome c
and ATP in vitro has been compared with that activated
in situ during etoposide-induced apoptosis. In addition, the
nucleotide requirements in vitro have been compared with the
nucleotide pools available in intact cells. Finally, the cell-free
system was utilized to explore the possibility that small molecule
inhibitors of apoptosis, including ATA and protease inhibitors that are
usually thought to inhibit other types of proteases, might act by
inhibiting caspase activation.
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EXPERIMENTAL PROCEDURES |
Materials--
Reagents were obtained from the following
suppliers: horse heart cytochrome c, Tween 20, DCI, TLCK,
TPCK, okadaic acid, Na3VO4, staurosporine,
genistein, pNA, nucleotides and nucleotide analogs from
Sigma; adjuvants from Ribi (Hamilton, MT); recombinant human caspases-3, -6, and -7 from PharMingen (San Diego, CA); recombinant human caspase-9 from Chemicon (Temecula, CA); DEVD-AFC from Biomol (Plymouth Meeting, PA); ZVAD-fmk from Bachem Bioscience (King of
Prussia, PA); and VEID-AFC, LEHD-AFC, DEVD-pNA,
ZVAD(OMe)-fmk, and ZFA-fmk from Enzyme Systems Products (Dublin, CA).
All other reagents were obtained as previously indicated (47, 49).
Antibodies--
Rabbit antisera that recognize caspase-3 or
caspase-7 were raised by injecting female New Zealand White rabbits
with the large subunit of recombinant human caspase-3 or -7. In brief,
the indicated large subunit was expressed as a fusion protein with
glutathione S-transferase using the pGEX-2T plasmid
(Amersham Pharmacia Biotech), purified by affinity chromatography on
glutathione-agarose, released from the fusion protein with thrombin,
subjected to SDS-PAGE, and excised from the gel. Rabbits were
inoculated at monthly intervals with polyacrylamide gel chips
containing 100-200 µg of purified protein mixed with Ribi adjuvant.
Positive sera were affinity purified by adsorption to a column
containing the indicated large subunit (purified as described above and
electroeluted from the SDS-polyacrylamide gel) followed by elution with
50 mM glycine hydrochloride (pH 2). Affinity purified
antibodies were concentrated by vacuum dialysis and stored at
70 °C. Control experiments revealed that these antibodies
recognized their corresponding antigens but not caspase-6 or caspase-8.
Rabbit antisera that recognize human caspases-6 and -8 were raised
using purified recombinant His6-tagged proteins (17).
Rabbit antisera that recognize neoepitopes generated upon activation of
caspases-3 and -9 were generated using synthetic peptides (CIETD and
CPEPD, respectively) coupled to keyhole limpet hemocyanin. Mouse
monoclonal anti-caspase-2 was purchased from Transduction Laboratories
(Lexington, KY). Peroxidase-coupled affinity purified secondary
antibodies were from Kirkegaard & Perry Laboratories (Gaithersburg, MD).
Tissue Culture--
HL-60 acute myelogenous leukemia cells
(provided by Dr. Robert Abraham, Duke University, Durham, NC) were
cultured in RPMI 1640 medium containing 10% heat-inactivated fetal
bovine serum, 100 units/ml penicillin G, 100 µg/ml streptomycin, and
2 mM glutamine. Cultures were incubated at 37 °C in
humidified air with 5% (v/v) CO2 and maintained in
exponential growth phase. Cell size was determined on a Coulter counter
equipped with a channel analyzer and calibrated using polystyrene beads
of defined diameter (Becton-Dickinson, San Jose, CA). To induce
apoptosis, cells were treated with 68 µM etoposide for
the indicated length of time (47).
Cytosol Preparation--
All steps were performed at 4 °C.
Cells were sedimented at 200 × g for 10 min, washed
twice with RPMI 1640 containing 10 mM HEPES (pH 7.4), and
resuspended at 3 × 108 cells/ml in buffer A (25 mM HEPES (pH 7.5 at 4 °C), 5 mM
MgCl2, 1 mM EGTA, supplemented immediately
before use with 1 mM DTT and the following protease
inhibitors: 1 mM PMSF, 10 µg/ml pepstatin A, and 10 µg/ml leupeptin). After a 20-min incubation, cells were disrupted by
20-50 strokes in a pre-chilled Dounce homogenizer with a tight-fitting
pestle and sedimented at 800 × g for 10 min. The
postnuclear supernatant was sedimented at 100,000 × gmax for 60 min in a Beckman TL-100
ultracentrifuge. The resulting cytosol (supernatant) was placed in
Spectra/Por 12-14-kDa cut-off dialysis tubing (Spectrum, Laguna Hills,
CA); dialyzed against several changes of Buffer A for 12-18 h to
deplete endogenous nucleotides; supplemented with 1 mM DTT,
1 mM PMSF, 10 µg/ml pepstatin A, 10 µg/ml leupeptin;
and frozen in 100-µl aliquots at 70 °C. Protein concentrations
in cytosol preparations (assayed by the method of Smith et
al. (50)) were 2.5-7 mg/ml. Cytosol prepared and stored in this
manner was capable of undergoing caspase activation in vitro
for at least 14 months.
Caspase Activation--
Cell-free caspase activation reactions
(40 µl final volumes) were assembled on ice in wells of 96-well
microtiter plates by sequential addition of ice-cold buffer A,
additives (e.g. nucleotides, cytochrome c, and
inhibitors prepared in buffer A and adjusted to pH 7.5), and freshly
thawed cytosol containing 50 µg of protein. Reactions were initiated
by placing the microtiter plate in a humidified 30 °C incubator.
After incubation for the indicated times, reaction mixtures were
harvested for immunoblotting, affinity labeling, or caspase activity assays.
Immunoblotting--
At the completion of the incubation, samples
were diluted with 10 µl of 4× sample loading buffer (8% SDS, 250 mM Tris-HCl (pH 6.8 at 20 °C), 40% (v/v) glycerol, 20%
(v/v) 
-mercaptoethanol) and heated to 65 °C for 10 min. Samples
containing 50 µg of protein were subjected to electrophoresis for
50-60 min at 200 V on SDS-polyacrylamide minigels containing 15%
(w/v) acrylamide. Polypeptides were electrophoretically transferred to
polyvinylidene fluoride membranes for 50 min at 4 °C. Membranes were
stained with fast green FCF to confirm equal loading and transfer of
samples. Blots were then washed twice in TS buffer (150 mM
NaCl containing 10 mM Tris-HCl, pH 7.4 at 20 °C),
incubated for 
30 min in TS buffer containing 10% (w/v) powdered
nonfat milk (TSM) to block unoccupied protein-binding sites, reacted
with primary antibodies diluted in TSM buffer for 14-16 h, washed
three times (15 min each) with calcium- and magnesium-free phosphate-buffered saline (PBS) containing 0.05% (w/v) Tween 20, reacted for 1 h with peroxidase-coupled secondary antibodies
diluted in PBS containing 3% (w/v) powdered nonfat milk, washed three times in PBS, 0.05% Tween 20 (15 min each) and once in PBS, and subjected to enhanced chemiluminescent detection.
Affinity Labeling--
After activation as described above,
aliquots containing 50 µg of cytosolic protein were incubated for 15 min at 37 °C with 1 µM ZEK(bio)D-aomk.
Previous experiments have revealed that p20 subunits of all caspases
tested (caspases-1, -2, -3, -4, -6, -7, and -8) can be labeled with
this reagent (47, 51). At the completion of the incubation, samples
were diluted with 1/2 volume of 3× SDS sample buffer (9% (w/v)
SDS, 0.15 M Tris-HCl (pH 6.8 at 20 °C), 6 mM
EDTA, 45% (w/v) sucrose, 0.03% (w/v) bromophenol blue, 10% (v/v)
-mercaptoethanol), heated to 95 °C for 3 min, subjected to
SDS-PAGE on 16% (w/v) acrylamide gels, transferred to nitrocellulose,
probed with peroxidase-labeled streptavidin, and visualized using
ECL-enhanced chemiluminescence reagents.
Two-dimensional analysis was performed using isoelectric focusing for
the first dimension and SDS-PAGE for the second dimension as described
(49). Labeled polypeptides were visualized using peroxidase-coupled
streptavidin followed by SuperSignalTM ULTRA chemiluminescence reagent
(Pierce). Caspases expressed in Sf9 cells (47) were subjected to
this analysis in parallel to permit identification of the labeled
polypeptide species.
Measurement of Caspase Activity--
After incubation under
cell-free conditions as described above, the ability of cytosol
preparations to cleave DEVD-pNA was measured as described by
Datta et al. (52). Reactions were assembled in microtiter
plate wells by adding 160 µl of buffer B (100 mM HEPES
(pH 7.5), 20% (v/v) glycerol, 5 mM DTT, and 0.5 mM EDTA) containing 125 µM
DEVD-pNA to wells containing 50 µg of cytosolic protein in
40 µl of buffer A. Plates were incubated at 37 °C in a Molecular
Dynamics (Sunnyvale, CA) Thermomax plate reader. Release of free
pNA, which absorbs at 405 nm, was monitored continuously. Absorbance values were converted to picomoles using a standard curve
based on free pNA.
The effect of various inhibitors on activated caspases was assessed as
described previously (47). In brief, recombinant caspases-3, -6, -7, or
-9 in 50 µl of buffer A were incubated for 5 min at 20-22 °C with
the indicated concentration of the inhibitor and then diluted with 225 µl of freshly prepared buffer C (25 mM HEPES (pH 7.5),
0.1% (w/v) CHAPS, 10 mM DTT, 100 units/ml aprotinin, 1 mM PMSF) containing 100 µM substrate and
incubated for 2 h at 37 °C. The following substrates were
utilized: DEVD-AFC for caspases-3 and -7; VEID-AFC for caspase-6; and
LEHD-AFC for caspase-9. Reactions were terminated by addition of 1.225 ml of ice-cold buffer C. Fluorescence was measured using an excitation wavelength of 360 nm and emission wavelength of 475 nm. Reagent blanks
containing 50 µl of buffer A and 225 µl of buffer C were incubated
at 37 °C for 2 h and then diluted with 1.225 ml of ice-cold buffer C. Standards containing 0-1500 pmol of AFC were utilized to
determine the amount of fluorochrome released. Results are the mean of
3-5 determinations at each inhibitor concentration.
Nucleotide Measurement--
After treatment with 68 µM etoposide for the indicated lengths of time, aliquots
containing 1.3 × 108 HL-60 cells were washed twice
with ice-cold PBS and lysed by incubation for 15 min at 4 °C in 0.5 M perchloric acid. All additional steps were performed at
4 °C. Following removal of insoluble macromolecules by sedimentation
at 1600 × g for 15 min, the perchloric acid extract was neutralized with 1 M KOH containing 0.33 M
potassium phosphate (pH 7.4 at neutralization) and sedimented at
1600 × g for 15 min to remove precipitated potassium
perchlorate. Aliquots of the neutralized supernatant were subjected to
high performance liquid chromatography within 60 min of the final
sedimentation step using conditions that permitted separation and
quantitation of ATP, dATP, ADP, and dADP as well as a number of
additional nucleotide species. In brief, 60 µl of the resulting
supernatant was subjected to HPLC analysis using a Beckman 125 dual
pump gradient system equipped with 507e autosampler, 168 diode array
detector, and IBM personal computer 350 with Beckman Gold software. A
Brownlee MPLC Newguard anion-exchange precolumn (3.2 × 15 mm × 7 µm) and a Whatman Partisil-10 SAX column (4.6 × 250 mm × 5 µm) preequilibrated for at least 30 min with 99% mobile
phase A (7 mM NH4H2PO4,
pH 3.8) and 1% mobile phase B (250 mM
NH4H2PO4, pH 4.5, containing 500 mM KCl) were used for all analyses. Separation was
accomplished using a flow rate of 2 ml/min and the following elution
gradient: 0-10 min, 99% mobile phase A, 1% mobile phase B; 10-30
min, linear gradient to 95% mobile phase A, 5% mobile phase B; 30-75
min, linear gradient to 100% mobile phase B. Absorbance was measured at 259 nm. Peaks were identified by coelution with authentic
nucleotides in the described solvent system as well as a separate
solvent system consisting of 1% aqueous triethylamine and methanol
using a C18 HPLC column (Beckman Ultrasphere ODS 4.6 × 250 mm).
Known amounts of ATP, dATP, ADP, and dADP were utilized to construct standard curves, which were then employed to determine the amounts of
various nucleotides in the perchloric acid extracts.
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RESULTS |
Caspase Activation in a Cell-free System Prepared from HL-60 Cell
Cytosol--
Previous studies have demonstrated that caspases can be
activated when cytosol from nonapoptotic cells is incubated with
cytochrome c and ATP at 30 °C (13, 14, 20, 21, 48, 53).
Our initial experiments (Fig. 1) were
performed to characterize this process in cytosol prepared from HL-60
human leukemia cells. Incubation of dialyzed HL-60 cytosol with 100
µM ATP and 1 µM cytochrome c
led to caspase-3 activation. Processing of endogenous procaspase-3 to
the active enzyme was demonstrated by immunoblotting, which showed loss
of the 32-kDa precursor and appearance of a new 17-kDa polypeptide that
reacts with affinity purified antibodies raised against the recombinant
caspase-3 17-kDa subunit (Fig. 1A, top panel); by appearance
of an activity that cleaves DEVD-pNA (Fig. 1B), a
preferred substrate of caspase-3; and by the appearance of polypeptides
that comigrate with recombinant caspase-3 species after reaction with
the affinity labeling reagent ZEK(bio)D-aomk (see
below).
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Fig. 1.
Caspase activation in HL-60 cytosol under
cell-free conditions. A, specific activation of a
subset of caspases in vitro. Dialyzed HL-60 cytosol was
treated with 500 µg/ml cytochrome c and 1 mM
dATP for the indicated length of time and then harvested for
immunoblotting with antibodies that recognize the large subunits of the
indicated active caspases. Locations of the procaspase species
(open arrows) and large subunits of active species
(closed arrows) are indicated. Note that procaspases-3, -6, and -7 are cleaved, whereas procaspases-2 and -8 are not. B,
detection of caspase activity after incubation of cytosol with
cytochrome c and dATP in vitro. After incubation
for 30 min with dATP in the absence () or presence (+) of cytochrome
c, aliquots of cytosol were assayed for ability to cleave
DEVD-pNA. C, HL-60 cells treated with 68 µM etoposide for 0 h (lane 1) or 4 h
(lane 2) were probed as described in A.
Procaspase-6 (not shown) was not reliably detected in the whole cell
lysates with the available antibody. Note that a subset of caspase
precursors are cleaved at this time point in situ.
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Previous results have established that etoposide-induced apoptosis in
intact HL-60 cells is accompanied by selective activation of a subset
of the procaspase species present in these cells (47, 48). In
particular, caspases-3 and -7 are activated by 4 h, whereas
caspases-2 and -8 are not (Fig. 1C) (37, 47,
48).2 To determine whether
similar selectivity is observed in vitro, cytosol incubated
with ATP and cytochrome c was probed with antibodies that
recognize several different caspases (Fig. 1A). Processing of procaspases-3 and -7 was detected within 30-60 min of the start of
the incubation. Processing of the caspase-6 zymogen was also observed,
albeit with slightly slower kinetics. In contrast, cleavage of
procaspases-2 and -8 was not observed (Fig. 1A, lower
panels), confirming that this in vitro system of
caspase activation recapitulates the selectivity observed in intact cells.
In further experiments, affinity labeling with
ZEK(bio)D-aomk was utilized to compare the cohort of
caspases activated in vitro with those detected in cytosol
of etoposide-treated HL-60 cells (Fig.
2). Although analysis by unidimensional
(Fig. 2A) and two-dimensional gel electrophoresis (Fig.
2B) confirmed that active caspase species were detected
after incubation of cytosol with cytochrome c and dATP
in vitro, the labeling patterns were significantly different
from those observed when cytosol was analyzed after caspases were
activated in intact cells. Earlier analysis using unidimensional gels
indicated that the pattern of active caspases in cytosol of
etoposide-treated cells is relatively simple early in the course of
caspase activation and becomes more complicated over time (47). In
contrast, during caspase activation in vitro, the labeling
pattern was most complex early in the course of the incubation and
became simpler over time (cf. lanes 3 and
7 in Fig. 2A). Comparison of the species detected
in cytosol after activation in vivo and in vitro
(cf. lanes 1 and 3 in Fig.
2A) indicated that species a, b, d/e, and g are detected in
both samples, although the relative ratios appeared to be different. In
contrast, species c and f were detected only after caspase activation
in vitro.
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Fig. 2.
Affinity labeling of active caspase species
after activation in etoposide-treated cells or in vitro.
A, analysis by unidimensional SDS-PAGE. Cytosol from HL-60
cells treated with etoposide for 3 h (lane 1) or
cytosol from control cells treated with 500 µg/ml cytochrome
c and 1 mM dATP at 30 °C for the indicated
length of time (lanes 2-7) was incubated with 1 µM ZEK(bio)D-aomk and subjected to
unidimensional SDS-PAGE followed by blotting with peroxidase-coupled
streptavidin. a-g refer to discrete caspase species
detected by varying exposures of this blot. Results are representative
of seven separate experiments. B, after affinity labeling as
described for A, cytosol from HL-60 cells treated with
etoposide for 5 h (upper panel) or cytosol from control
cells treated with 500 µg/ml cytochrome c and 1 mM dATP at 30 °C for 5 h (middle panel)
was subjected to isoelectric focusing (from left to
right) followed by SDS-PAGE (from top to
bottom). C3 and C6 in upper two
panels refer to major species of caspases-3 and -6, respectively,
previously identified by this technique (47). White
arrowheads denote the mobility of recombinant caspases expressed
in Sf9 cells as determined in two-dimensional gels run in
parallel (see lower panel). Results are representative of
four separate experiments.
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These differences between the cohort of caspases activated in
vivo and in vitro were even more pronounced when active
caspase species were analyzed by two-dimensional isoelectric
focusing/SDS-PAGE (Fig. 2B). In particular, a major species
of active caspase-6 and multiple species of active caspase-3 were
detected after caspase activation in vivo (Fig. 2B,
upper panel) (47). Similar labeling patterns have been observed in
several cell lines after treatment of intact cells with various
proapoptotic stimuli (47, 48, 54). Previous studies have demonstrated
that the size and charge heterogeneity of these species reflect, at
least in part, the retention or removal of the caspase prodomain and
different degrees of caspase phosphorylation, respectively (49, 55,
56). Although the caspase-3 species were detectable after activation
in vitro (Fig. 2B, middle panel), the
major caspase-6 species was
not.3 Instead, several acidic
caspase species that were present in small amounts after caspase
activation in vivo were prominently labeled after caspase
activation in vitro. The identity of these novel species is
at present unknown.
To determine the influence of reaction conditions on caspase activation
in vitro, we systematically varied several of the experimental parameters. During the preparation of some batches of
cytosol, the dialysis step was omitted. Under these conditions, we
observed batch-to-batch variation in the requirement for exogenous cytochrome c and/or ATP. Some undialyzed HL-60 cytosol
preparations behaved as illustrated in Figs. 1 and 2, whereas others
facilitated caspase activation in the absence of exogenous cytochrome
c and ATP (data not shown) in a manner similar to extracts
recently described by Kuida et al. (26). In contrast, when
dialyzed cytosol was utilized, exogenous cytochrome c and
ATP were required. Unless otherwise indicated, all further experiments
utilized dialyzed cytosol.
Additional experiments (not shown) examined other parameters. Titration
experiments revealed that quantitative cleavage of procaspase-3 in
HL-60 cytosol required 1 µM cytochrome c, in agreement with observations made using cytosol from Xenopus
oocytes (53). NaCl or KCl concentrations above 60 mM were
observed to inhibit caspase-3 activation in HL-60 cytosol, a finding
consistent with results obtained using cytosol from rat thymocytes
(57). Finally, incubation of HL-60 cytosol with ATP and cytochrome
c resulted in more rapid and complete procaspase cleavage at
30 °C than at 37 °C. All subsequent activation studies were
performed at 30 °C in the absence of exogenous monovalent cations.
Nucleotide Requirements for Caspase Activation in HL-60
Cytosol--
Further experiments examined the nucleotide requirements
of the in vitro activation system. When 1 µM
cytochrome c was present, titration experiments revealed
complete processing of procaspase-3 in the presence of 100
µM ATP (data not shown). When other nucleotides were
tested at 1 mM, dATP or dADP could substitute for ATP,
whereas AMP, ADP-ribose, CMP, CTP, or UTP could not (Fig.
3A and data not shown).
Further examination revealed that dATP was effective at concentrations
below 10 µM (Fig. 3B, lane 5), although it
inhibited procaspase-3 activation at extremely high concentrations
(Fig. 3B, lane
8).4
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Fig. 3.
Effect of poorly hydrolyzable ATP analogs on
caspase activation. A, effect of selected nucleotide
analogs on caspase activation in vitro. Lane 1,
input cytosol. Lane 2-9, cytosol incubated for 5 h at
30 °C in absence (lane 2) or presence (lanes
3-9) of 500 µg/ml cytochrome c and absence
(lane 3) or presence (lanes 4-9) of the
indicated nucleotide at a final concentration of 1 mM. At
the completion of the incubation, cytosol was subjected to SDS-PAGE
followed by immunoblotting with affinity purified anti-caspase-3
antibody. Locations of procaspase-3 (open arrow) and its
enzymatically active cleavage product (closed arrow) are
indicated. B, effects of dATP, AMPPCP, and AMPPNP on caspase
activation in vitro. Lanes 1-3, dialyzed cytosol
incubated for 5 h at 30 °C in the absence (lane 1)
or presence (lanes 2 and 3) of 1 mM
ATP (lane 2) or 500 µg/ml cytochrome c
(lane 3). Lanes 4-17, dialyzed cytosol
supplemented with 500 µg/ml cytochrome c and the following
nucleotides: ATP at 1 mM (lane 4); dATP at 0.01, 0.1, 1, and 10 mM (lanes 5-8, respectively);
AMPPCP at 0.01, 0.1, 1, and 10 mM (lanes 9-12,
respectively); and AMPPNP at 0.01, 0.1, 1, 10, and 100 mM
(lanes 13-17, respectively). All samples were treated in
one experiment but applied to two separate minigels. Additional samples
confirmed that addition of cytochrome c + ATP resulted in
procaspase-3 activation with kinetics indistinguishable from those
illustrated in Fig. 1A. C, effect of ATP S on
caspase activation in vitro. Lane 1, input
cytosol. Lanes 2-5, cytosol incubated for 5 h at
30 °C in the absence or presence of exogenous ATP (1 mM)
and/or cytochrome c (500 µg/ml) as indicated. Lanes
6-9, cytosol incubated for 5 h at 30 °C in the presence
of 500 µg/ml cytochrome c and 0.1, 1, 5, or 10 mM
ATPS, respectively. For this experiment, undialyzed cytosol was
utilized. Activation of caspases upon addition of cytochrome
c (lane 4) presumably reflects the high levels of
endogenous adenine nucleotides (see Fig. 4).
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To determine whether the ATP/dATP requirement reflected the need to
hydrolyze a high energy phosphate bond, ATP was replaced with the
poorly hydrolyzable analogs AMPPCP, AMPPNP, or ATPS. As shown in
Fig. 3B, replacement of dATP with 1 mM AMPPCP or
1-10 mM AMPPNP resulted in partial cleavage of
procaspase-3 after 5 h. When compared with the quantitative
cleavage of procaspase-3 in <1 h (Fig. 1A; see also legend
to Fig. 3B), these results indicate that AMPPCP and AMPPNP
support caspase activation relatively poorly. In further experiments,
ATPS at a wide range of concentrations did not support caspase-3
processing in the in vitro system (Fig. 3C). On
the contrary, ATPS prevented activation by endogenous nucleotides in
the undialyzed cytosol used for this particular experiment
(cf. lanes 4 and 6, Fig.
3C). The slowing or abrogation of the caspase activation
process by these nonhydrolyzable analogs suggests that hydrolysis of
the terminal phosphodiester bond in ATP plays an important role in the
caspase activation mechanism.
To compare the nucleotide requirements of this reaction in
vitro with the potential nucleotides available during apoptosis in situ, ATP and dATP were quantitated by HPLC (Fig.
4A) in HL-60 cells undergoing
etoposide-induced apoptosis. This analysis revealed a base-line ATP
concentration of 0.92 mM, a value that is within the range
previously reported for this cell line (58, 59). The same analysis
revealed a base-line dATP concentration of 0.17 mM. During
the first 2 h after addition of etoposide, which is the period of
time when caspase activation begins to occur in this cell line (47),
concentrations of ATP and dATP did not change appreciably (Fig.
4B). At later time points, ATP and dATP began to decline.
ADP and dADP levels decreased in a similar fashion (data not
shown).
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Fig. 4.
Effect of etoposide treatment on selected
HL-60 cell nucleotide pools. A, separation of
nucleotides under conditions of the HPLC assay. Perchloric acid
extracts from untreated HL-60 cells were subjected to HPLC as described
under "Experimental Procedures." The identity of the indicated
peaks was determined by their coelution with the indicated nucleotides
under two different chromatographic conditions and by the identity of
their ultraviolet absorption spectra with authentic samples.
B, levels of ATP and dATP in HL-60 cells treated with
etoposide for the indicated lengths of time. For each nucleotide,
levels were quantitated from data similar to that shown in A
and expressed relative to levels observed in untreated HL-60 cells
harvested at the beginning of this experiment. Base-line levels of ATP
and dATP in these cells were 0.92 and 0.17 mM,
respectively. ADP and dADP showed a similar pattern of changes. In the
same experiment, aliquots of cells were examined for nuclear
fragmentation after staining with Hoechst 33258 (% apoptotic
cells). Results are representative of four separate
experiments.
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Effects of Inhibitors on Caspase Activation in Vitro--
As
indicated in the Introduction, a variety of small molecule inhibitors
inhibit apoptosis in various model systems. To examine the possibility
that some of these might be acting by inhibiting caspase activation,
the effects of these compounds on caspase activation in the cell-free
system were assessed.
Zn2+ and ATA, two classical inhibitors of programmed cell
death (27, 94), inhibited activation of procaspase-3 when cytosol was
incubated with cytochrome c and ATP in vitro. As
illustrated in Fig. 5A, 0.5 mM Zn2+ inhibited procaspase activation.
Likewise, 40 µM ATA inhibited processing of
procaspase-3 under these conditions.
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Fig. 5.
Effect of selected inhibitors on caspase
activation in vitro. A, effects of
Zn2+ and ATA on caspase activation in vitro.
Lane 1, unsupplemented cytosol incubated at 30 °C for
5 h. Lanes 2-10, cytosol supplemented with 1 mM ATP, 500 µg/ml cytochrome c, and the
indicated concentration of ATA or ZnSO4 was incubated for
5 h at 30 °C and then subjected to immunoblotting with an
antiserum that recognizes procaspase-3 (arrowhead).
B, effects of phosphatase inhibitors on caspase activation
in vitro. Lanes 1 and 2, cytosol
incubated for 5 h at 30 °C in the absence (lane 1)
or presence (lane 2) of exogenous ATP (1 mM) and
cytochrome c (500 µg/ml). Lanes 3-9, cytosol
containing ATP and cytochrome c was supplemented with 1 mM staurosporine (lane 3), 100 µM
genistein (lane 4), 1 mM okadaic acid
(lane 5), or 0.1, 0.5, 1, or 10 mM
Na3VO4 (lanes 6-9, respectively),
prior to incubation at 30 °C for 5 h. Arrowhead,
procaspase-3. C, effects of various protease inhibitors on
caspase activation. Lanes 1' and 2', cytosol
incubated at 30 °C for 5 h in the absence (lane 1')
or presence (lane 2') of exogenous ATP (1 mM)
and cytochrome c (500 µg/ml). Lanes 1-3,
cytosol containing ATP and cytochrome c was supplemented
with increasing concentrations of iodoacetamide (IAA) (10, 100, and 500 µM), DEVD-fmk (1, 10, and 100 µM), VEID-fmk (1, 10, and 100 µM), ZVAD-fmk
(10, 100, and 1000 nM), ZVAD(OMe)-fmk (10, 100, and 1000 nM), TPCK (0.1, 0.5, and 1 mM), TLCK (0.1, 1, and 10 mM), or DCI (0.1, 0.25, and 0.5 mM),
respectively. D, effect of various concentrations of ZFA-fmk
on caspase-3 activation. Cytosol was incubated at 30 °C for 5 h
in the presence of 0.1, 0.5, 1, or 5 mM ZFA-fmk,
respectively.
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Recent studies have indicated that active caspases in etoposide-treated
HL-60 cells are phosphoproteins (49). Additional studies have
demonstrated that caspase-9 is also phosphorylated under some
conditions (60). Based on these results, the effects of inhibitors of
protein kinases (staurosporine and genistein) and protein phosphatases
(okadaic acid and Na3VO4) were investigated. Staurosporine, which inhibits calmodulin-dependent kinase,
cyclic AMP-dependent kinase, protein kinase G, and myosin
light chain kinase in the nanomolar range and protein kinase C in the
picomolar range (61), had no discernible effect on procaspase-3
activation at millimolar concentrations (Fig. 5B, lane 3).
Genistein, which inhibits tyrosine kinases at low micromolar
concentrations (62), was similarly unable to block procaspase-3
processing at millimolar concentrations (Fig. 5B, lane
4).
In additional experiments, okadaic acid, which inhibits protein
phosphatases 1, 2A, and 2B at concentrations below 1 µM
(63), had no discernible effect on procaspase-3 activation at
concentrations of up to 100 µM (Fig. 5B, lane
5). Na3VO4, which broadly inhibits protein
tyrosine phosphatases as well as alkaline phosphatase in the micromolar
range (64), was similarly ineffective in blocking procaspase-3
processing at submillimolar concentrations (Fig. 5B, lanes 6 and 7). These data suggest that alterations of caspase phosphorylation are not required for activation of effector caspases such as caspase-3 in HL-60 cytosol. On the other hand, higher concentrations of Na3VO4, which are known to
inhibit other enzymes involved in phosphate metabolism, including
ATPases (65, 66), did inhibit caspase activation (Fig. 5B, lane
9), consistent with data presented above suggesting that
nucleotide hydrolysis might be required for this process.
To examine the effects of various protease inhibitors on caspase
activation, cytosolic extracts were incubated at 4 °C for 5-10 min
with each inhibitor, then warmed to 30 °C for 5 h, and examined
for procaspase-3 cleavage. As indicated in Fig. 5C, a variety of protease inhibitors prevented procaspase-3 activation in a
dose-dependent manner. Iodoacetamide, an irreversible
sulfhydryl-blocking reagent, completely prevented procaspase-3
activation at concentrations in excess of 500 µM. This
reaction was also inhibited by several different caspase inhibitors,
including DEVD-fmk, VEID-fmk, and ZVAD-fmk. Interestingly, the methyl
ester ZVAD(OMe)-fmk was at least 20-50-fold less potent than the
parent compound in this reaction (cf. 4th and
5th panels, Fig. 5C).
Results in Fig. 5, C and D, also illustrate
results obtained with other classes of protease inhibitors. TPCK and
TLCK, two chloromethyl ketones that inhibit
sulfhydryl-dependent proteases (67-69) as well as certain
serine proteases (70), abolished procaspase-3 activation at all
concentrations above 1 and 5 mM, respectively. DCI, which
is usually considered an inhibitor of serine proteases (71), completely
inhibited procaspase-3 activation at concentrations above 500 µM. In addition, ZFA-fmk, which is usually
considered a selective inhibitor of sulfhydryl-dependent
cathepsins (72), had two different effects on caspase-3 activation
(Fig. 5D). At low concentrations (Fig. 5D, lanes
1-3), ZFA-fmk inhibited removal of the prodomain from
active caspase-3 (manifested as cleavage from the 20-kDa to the 17-kDa
species). At higher concentrations, ZFA-fmk also inhibited cleavage of
procaspase-3 to the 20-kDa species (Fig. 5D, lane 4).
Effect of Inhibitors on Recombinant Caspase Activity--
The
preceding experiments raised the possibility that Zn2+,
ATA, DEVD-fmk, VEID-fmk, ZVAD-fmk, DCI, TPCK, TLCK, and ZFA-fmk might
be exerting their effects by inhibiting caspase activation. If so, one
logical target might be caspase-9, the apical caspase in the cytochrome
c/Apaf-1 pathway. To assess this possibility, recombinant
human caspase-9 was incubated with these agents for 5 min, diluted, and
assayed for ability to cleave its preferred substrate, LEHD-AFC.
Results of this analysis revealed that the fluoromethyl ketones
DEVD-fmk, VEID-fmk, ZVAD-fmk, ZVAD(OMe)-fmk, and ZFA-fmk all inhibited
caspase-9 activity (Fig. 6A).
In agreement with the in vitro activation data,
ZVAD(OMe)-fmk was at least 10-fold less potent than ZVAD-fmk. In
addition, ATA and Zn2+, which are commonly considered to be
nuclease inhibitors, inhibited caspase-9 (Fig. 6B). Finally,
TPCK, TLCK, and DCI, three agents that are frequently classified as
serine esterase inhibitors, also inhibited caspase-9 activity (Fig.
6B).
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Fig. 6.
Effects of various inhibitors on recombinant
caspase-9. Caspase-9 was incubated for 5 min on ice with the
indicated concentration of each inhibitor and then diluted 5.5-fold
with buffer A containing LEHD-AFC. After a 2-h incubation with
substrate at 37 °C, the release of fluorescent product was
determined as described under "Experimental Procedures." Results
were expressed relative to purified caspase-9 treated with diluent
alone and were plotted as a function of the initial inhibitor
concentration before addition of the substrate.
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Because the effects of ATA, DCI, ZFA-fmk, TPCK, and TLCK were somewhat
unanticipated, the effects of these compounds on the catalytic
activities of other purified recombinant human caspases were evaluated.
As indicated in Fig. 7, all of these
agents also inhibited purified caspases-3, -6, and -7 at submillimolar
or millimolar concentrations. In particular, DCI (Fig. 7, closed squares) inhibited caspase activity by 50% at concentrations that ranged from 500 µM (caspase-6) to 1 mM
(caspases-3 and -7). TPCK (Fig. 7, closed circles) inhibited
all three caspases at 200-1000 µM. Likewise, ZFA-fmk
(Fig. 7, open triangles) also inhibited the purified
caspases, with an IC50 in the 60 (caspase-7) to 1000 µM (caspase-3) range.
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Fig. 7.
Effects of various protease inhibitors on
purified recombinant caspases. Caspases-3, -6, and -7 were
incubated for 5 min on ice with the indicated concentration of each
inhibitor and then diluted 5.5-fold with buffer A containing DEVD-AFC
(caspases-3 and -7) or VEID-AFC (caspase-6). After a 2-h incubation
with substrate at 37 °C, the release of fluorescent product was
determined as described under "Experimental Procedures." Results
were expressed relative to purified caspases treated with diluent alone
and were plotted as a function of the initial inhibitor concentration
before addition of the substrate.
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Inhibition of Caspase-9 Activation--
To determine whether these
agents diminish caspase-9 activation, we utilized an antiserum raised
against a tetrapeptide epitope that is masked in the intact caspase-9
zymogen but is generated at the C terminus of the large subunit during
caspase maturation (7, 18). As indicated in Fig.
8A, this caspase-9 neoepitope antiserum recognized multiple bands in a preparation of purified caspase-9, including prominent species at 30-35 kDa (the previously reported size of the large subunit) (13, 18, 20) and another species at
Mr ~18,000 (thought to be a processed form of
the large subunit). When applied to multiple purified caspases, this
serum recognized active caspase-9 but not caspases-3, -6, or -7 (Fig. 8B, upper panel). In contrast, a corresponding caspase-3
neoepitope antiserum recognized caspase-3 but not caspases-6, -7, or -9 (Fig. 8B, lower panel).
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Fig. 8.
Effects of various protease inhibitors on
caspase-9 activation in HL-60 cytosol. A, reaction
between neoepitope antiserum and various caspase-9 species. Recombinant
His6-tagged human caspase-9 expressed in bacteria was
purified on nickel-agarose, subjected to SDS-PAGE, and probed with
anti-caspase-9 neoepitope antiserum. This antiserum recognized the
major Mr ~30-35,000 species present in this
preparation, as well as a less abundant Mr
~18,000 species. B, reaction of anti-neoepitope sera with
purified caspases. Caspases-3, -6, -7, and -9 were subjected to
SDS-PAGE followed by blotting with the indicated caspase neoepitope
antisera. C, activation of caspase-9 in HL-60 cytosol
in vitro. Dialyzed HL-60 cytosol was treated with 500 µg/ml cytochrome c and 1 mM dATP for the
indicated length of time and then harvested for immunoblotting with the
caspase-9 neoepitope antiserum. Open arrow
indicates a Mr ~65,000 cross-reactive
cytosolic polypeptide that serves as a loading control. D,
activation of caspase-9 in intact HL-60 cells. HL-60 cells treated with
diluent () or 68 µM etoposide for 1 h (+) were
washed, incubated for 14 h, and harvested for immunoblotting with
the caspase-9 neoepitope antiserum. The intense
Mr ~65,000 band (open arrow) served
as a loading control. Closed arrows, species detected in
apoptotic HL-60 cells. **, location of procaspase-9 when the blot was
reprobed with anti-procaspase-9 antibody. E, effects of
various inhibitors on etoposide-induced caspase activation in HL-60
cytosol. Lanes 1' and 2', cytosol incubated at
30 °C for 5 h in the absence (lane 1') or presence
(lane 2') of exogenous ATP (1 mM) and cytochrome
c (500 µg/ml). Lanes 1-3, cytosol containing
ATP and cytochrome c was supplemented with iodoacetamide
(IAA) (10, 100, and 500 µM), DEVD-fmk (1, 10, and 100 µM), VEID-fmk (1, 10, and 100 µM),
ZVAD-fmk (10, 100, and 1000 nM), ZVAD(OMe)-fmk (10, 100, and 1000 nM), TPCK (0.1, 0.5, and 1 mM), TLCK
(0.1, 1, and 10 mM), or DCI (0.1, 0.25, and 0.5 mM), respectively. At the completion of a 5-h incubation at
37 °C, samples were subjected to blotting with the caspase-9
neoepitope antiserum. Only the region between 15- and 25-kDa is
shown.
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When applied to HL-60 cytosol that had been incubated with cytochrome
c and dATP for varying lengths of time (Fig. 8C),
the caspase-9 serum recognized a Mr ~18,000
species that appeared within 30 min. A band of the same molecular
weight was also the major species detected when etoposide-treated HL-60
cells were examined (Fig. 8D). Binding to this species was
prevented by the immunizing
peptide.5 In both cases, a
Mr ~65,000 cross-reactive band that did not change during the course of the incubation served to confirm the equivalent loading of the various lanes (Fig. 8, C and
D, open arrowhead). These results suggest that
the Mr ~18,000 species of caspase-9 is the
predominant species generated during caspase activation in HL-60
cytosol in vitro and in vivo.
To examine the effects of various inhibitors on caspase-9 activation,
the same samples shown in Fig. 5D were probed with the antiserum raised against the caspase-9 neoepitope. Results of this
analysis revealed that iodoacetamide, DEVD-fmk, VEID-fmk, ZVAD-fmk,
ZVAD(OMe)-fmk, TPCK, TLCK, DCI, Zn2+, and ATA inhibited the
generation of the Mr ~18,000 caspase-9 species
(Fig. 8E and data not shown). Given the fact that each active caspase-9 molecule can activate multiple procaspase-3 molecules, we anticipated that almost complete inhibition of caspase-9 activation and activity would be required to prevent procaspase-3 processing. Consistent with this prediction, higher concentrations of each agent
were required to inhibit caspase-3 activation as compared with
caspase-9 activation (cf. Figs. 5C and
8E). Collectively, these results suggest that the effects of
the indicated inhibitors involve inhibition of caspase-9 activation as
well as inhibition of activated caspases.
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DISCUSSION |
Recent studies from several laboratories have established that
addition of cytochrome c and adenine nucleoside
triphosphates to cytosol from control cells results in activation of
caspase-3 through a pathway that involves Apaf-1-mediated activation of procaspase-9 (13, 16, 18, 20). In the present study, we have extended
these observations by contrasting the cohorts of active caspases that
are generated in vitro and in situ, comparing the
nucleotide requirements in vitro to the adenosine nucleotide triphosphate levels available in intact cells, assessing the effects of
poorly hydrolyzable ATP analogs, and determining the effects of
selected inhibitors in the caspase activation process. These experiments address a number of unresolved questions about the activation of caspases under cell-free conditions.
Before embarking on these studies, we confirmed that the caspase
activation process in HL-60 cytosol in vitro was similar to
that observed in other model systems. Our studies demonstrated that the
process generated active forms of caspases-3, -6, and -7 in
vitro (Fig. 1A) but did not generate active forms of
caspases-2 or -8 (Fig. 1A and Fig. 2B). This
selectivity mirrors the activation process in intact HL-60 cells (Fig.
1C). Failure to activate caspase-2 (73, 74) and caspase-8
(74) has been reported in other cell-free systems, although it appears
that this selectivity might be cell type-dependent (21).
Our studies demonstrated that the activation process in dialyzed HL-60
cytosol was dependent upon addition of exogenous cytochrome
c and dATP or ATP (Fig. 3), consistent with results observed
in other laboratories (13, 14, 20, 21). A number of other nucleotides,
including AMP, ADP-ribose, CMP, CTP, or UTP, could not substitute for
ATP or dATP (Fig. 3A), in agreement with the results of Liu
et al. (14). Finally, our studies demonstrated that the
activation process was inhibited by 60-80 mM NaCl or KCl,
consistent with the results obtained in rat thymocyte cytosol (57).
Based on these similarities, it appears that the caspase activation
process in HL-60 cytosol is similar to that observed in other cell-free systems.
Some of the results obtained using the HL-60 cell-free system raise
questions about the relationship between conditions that result in
caspase activation in vitro and in situ. First,
as noted above, caspase activation in vitro is strongly
inhibited by buffers of physiological ionic strength. Second, the
activation process is more efficient in vitro at 30 °C
than at 37 °C. Third, results in Fig. 2B indicate that
the cohort of active caspase species detected by affinity labeling
after activation in vitro differs significantly from that
detected after activation in situ, possibly reflecting
differences in reactions that remove the caspase prodomains and alter
caspase phosphorylation. These observations raise the possibility that
one or more factors that modulate caspase activation in situ
at physiological ionic strength and temperature might have been lost
during the cell fractionation procedure. The similarities between the
results observed in HL-60 cytosol and other model systems (see above)
suggest that similar concerns might apply to many of the recently
described cell-free systems for caspase activation. Further studies are
required to evaluate the cause of the differences between caspase
activation in vitro and in vivo.
With these limitations in mind, the HL-60 system was utilized to study
nucleotide requirements and inhibitor sensitivity of caspase
activation. Studies employing cytosol from HeLa cells had previously
suggested that dATP might be specifically required for activation of
caspases (14). In particular, ATP was initially reported to be inactive
in vitro, whereas dATP was reportedly active, leading to the
speculation that elevations in dATP levels might occur after drug
treatment and contribute to the caspase activation process (14).
Consistent with this hypothesis, Wakade et al. (75) observed
a 40-fold increase in dATP levels when chick embryo sympathetic neurons
were induced to undergo apoptosis by treatment with 2-deoxyadenosine.
In the present study, nucleotide levels required for caspase activation
in cytosol under cell-free conditions were directly compared with
nucleotide pools in the same cell line as it underwent apoptosis.
Results of these studies indicated that caspase activation in HL-60
cytosol in vitro required as little as 10 µM
dATP or 100 µM ATP. In intact HL-60 cells, neither dATP
nor ATP increased during the induction of apoptosis by etoposide (Fig.
4B). Instead, base-line levels of both nucleotides were far
above those required for caspase activation in vitro. In
fact, levels of dATP observed in HL-60 cells are substantially higher
than previously reported in another leukemia cell line (76), raising
the interesting possibility that the exquisite sensitivity of HL-60
cells to a variety of proapoptotic agents might be related to the high
base-line dATP levels.
In subsequent experiments, ATP was replaced with AMPPCP, AMPPNP, or
ATPS. Results of these experiments help clarify the role of the
nucleotide in the caspase activation process. In ATP-requiring processes that depend on ATP binding rather than hydrolysis
(e.g. allosteric activation of pertussis toxin (77), the
release of eukaryotic initiation factor-2 from its ternary complex
(78), the binding of polyoma virus T antigen to its DNA-binding site (79), the activation of ubiquitin protein ligase (80), or release of
xylose reductase from its protein chaperone (81)), one or more of these
poorly hydrolyzable analogs has been observed to substitute fully for
ATP. In contrast, our experiments revealed that these analogs were poor
substitutes for ATP over a wide range of concentrations. In particular,
AMPPCP and AMPPNP permitted only slow caspase activation (Fig.
3B), and ATPS did not facilitate caspase activation at
all (Fig. 3C). Although these compounds are frequently
considered "nonhydrolyzable," previous studies have demonstrated
that various ATPases can hydrolyze these analogs at rates that range
from 0.0007 to 0.25 times the corresponding rates of ATP hydrolysis
(72, 82-85). The slowing of caspase activation in the presence of
these poorly hydrolyzable analogs (Fig. 3, B and
C) is consistent with a process that requires hydrolysis of
the terminal phosphodiester bond. Likewise, the inhibition of caspase
activation by millimolar concentrations of
Na3VO4 (Fig. 5B), which have
previously been shown to inhibit ATPases (65, 66), is consistent with
the suggestion that ATP or dATP hydrolysis is involved in this process.
In further experiments, the effects of a variety of inhibitors were
analyzed. A major advantage of the cell-free system is the lack of
permeability barriers that have been postulated to inhibit the action
of certain caspase inhibitors (86). Results obtained with the cell-free
system were compared with effects observed when recombinant caspases
were treated with the same inhibitors. For these experiments, we
utilized antisera that recognized caspase large subunits as well as an
antiserum raised against a neoepitope present in mature caspase-9 but
not the zymogen. This latter antiserum recognized caspase-9 but not
other caspases (Fig. 8B). In addition, this serum recognized
multiple species in a preparation of purified recombinant caspase-9,
including several 30-35-kDa species that correspond in molecular
weight to the reported sizes of active caspase-9 species (18) as well as a unique Mr ~18,000 species that appears to
represent further processing of the large subunit (Fig. 8A).
Interestingly, the major species detected by this antiserum after
caspase activation in vitro (Fig. 8C) or in
intact HL-60 cells (Fig. 8D) was the Mr ~18,000 species. A similar
Mr ~18,000 product of procaspase-9 activation
was recently observed by Susin et al. (87) but not by others
(13, 18, 20, 21). Further studies are required to understand the nature
and significance of the processing event that gives rise to this
species. Nonetheless, this reagent allowed us to examine caspase-9
activation under cell-free conditions.
Based on recent observations that some active caspases are
phosphoproteins (49), we examined the effect of broad spectrum kinase
and phosphatase inhibitors on caspase activation. At concentrations that selectively inhibit kinases or phosphatases, these reagents had
little if any effect on caspase activation (Fig. 5B),
suggesting that phosphorylation or dephosphorylation of the effector
caspases does not play a major role in this process in
vitro.
In contrast, a variety of other agents that have previously been
reported to diminish apoptosis in intact cells inhibited caspase
activation in vitro. For example, Zn2+, which
was originally introduced into the apoptosis literature as a nuclease
inhibitor, inhibited caspase-9 activity (Fig. 6B) and
caspase activation (Fig. 5A). Although previous studies have identified Zn2+ as a potential caspase inhibitor (28-30),
the present study provides the first evidence that caspase-9 is a
pertinent target of this cation. Consistent with this hypothesis,
additional experiments have demonstrated that activation of caspase-9
and its downstream target caspase-3 is inhibited by Zn2+ in
intact cells.6
Similarly, our data raise the possibility that ATA might inhibit
apoptosis by acting as a caspase inhibitor. Although ATA is known to
inhibit endonucleases (88), studies performed over the past 20 years
have indicated that ATA can also inhibit proteases, including serine
proteases (89), the proteosome (90), calpains (91), and certain
caspases (92). Our results extend these previous studies by
demonstrating that ATA not only inhibits recombinant caspases-3, -6, -7, and -9 in vitro (Figs. 6 and 7) but also abrogates caspase activation under cell-free conditions (Fig. 5A).
The same analysis revealed that various caspase inhibitors prevented
caspase activation (Figs. 5C and 8E). Consistent
with results that were published as the present studies were nearing completion (13, 44), we observed that ZVAD-fmk inhibited caspase-9 activity (Fig. 6) and activation (Figs. 5C and
8E). ZVAD(OMe)-fmk exhibited lower potency as an inhibitor
in vitro, presumably because esterases that activate this
agent in intact cells were not available to deesterify it after
preparation of cytosol in the presence of the serine esterase inhibitor
PMSF (93). Interestingly, DEVD-fmk was only 10-fold less potent than
ZVAD-fmk as an inhibitor of caspase activation (Figs. 5C and
8E) and caspase-9 activity (Fig. 6). Although it has often
been argued that the anti-apoptotic effects of DEVD-fmk result from the
inhibition of caspase-3, these results raise the possibility
DEVD-fmk might also be acting by inhibiting caspase-9 at the 10-300
µM concentrations used in many experiments.
Caspase activation was also inhibited by protease inhibitors that are
not traditionally viewed as caspase inhibitors. The chloromethyl
ketones TPCK and TLCK, which have been reported to inhibit caspases-3
and -7 in crude bacterial lysates (69), inhibited caspase activation
(Figs. 5C and 8E). DCI and ZFA-fmk also prevented caspase activation under cell-free conditions (Figs. 5, C
and D, and 8E). Further analysis revealed that
all of these reagents could inhibit purified caspases-3, -6, -7, and -9 as well (Figs. 6 and 7). Collectively, the results in Figs. 5-8 raise
the possibility that these agents might be affecting apoptosis through
effects on caspase activation and activity rather than effects on
noncaspase proteases. In view of these results, previous studies that
utilized these inhibitors to support the view that noncaspase proteases play a role in apoptosis might need to be reinterpreted.
| |
ACKNOWLEDGEMENTS |
We gratefully acknowledge Gregory Gores for
advice and discussions, Guy Salvesen for the caspase-6 and caspase-8
used to raise antisera, and Deb Strauss for secretarial assistance.
| |
FOOTNOTES |
*
This work was supported in part by Public Health Service
Grant CA69008 (to S. H. K. and W. C. E.) and a studentship from
Programa Gulbenkian de Doutoramento em Biologia e Medicina (to
L. M. M.).The costs of publication of this
article were defrayed in part by the
payment of page charges. The article
must therefore be hereby marked
"advertisement" in
accordance with 18 U.S.C. Section
1734 solely to indicate this fact.
d
Present address: Signal Transduction Laboratory, London, UK.
h
Principal Research Fellow of the Wellcome Trust.
i
Leukemia Society of America Scholar during the performance
of this research. To whom correspondence should be addressed: 1301 Guggenheim, Mayo Clinic, 200 First St. S.W., Rochester, MN 55905. Tel:
(507) 284-8950; Fax: (507) 284-3906; E-mail:
Kaufmann.Scott@Mayo.edu.
2
When HL-60 cells are incubated with etoposide
for longer periods, procaspase-8 also decreases (S. H. K.,
unpublished observations), suggesting that its activation under these
circumstances is downstream of caspase-9 and caspase-3 as recently
suggested (13, 21).
3
Previous results, which have demonstrated that a
major active species of caspase-6 does not focus in this gel system
(47), appear to account for the lack of a major caspase-6 species on the two-dimensional gels (Fig. 2B, middle panel) despite the
cleavage of procaspase-6 in vitro (Fig.
1A).
4
Equal loading of protein in Fig. 3 and
subsequent figures was confirmed by staining blots with fast green and,
in many cases, by blotting with anti-procaspase-2. These controls are
omitted to simplify the figures.
5
P. A. Svingen and S. H. Kaufmann,
unpublished observations.
6
T. J. Kottke and S. H. Kaufmann,
unpublished observations.
| |
ABBREVIATIONS |
The abbreviations used are:
AMPPCP, 5'-adenylyl methylenediphosphate;
AMPPNP, 5'-adenylyl imidodiphosphate;
ATA, aurintricarboxylic acid;
ATPS, adenylyl-5'-O-(3-thiotriphosphate);
DCI, 3,4-dichloroisocoumarin;
DEVD-AFC, N-(N-acetylaspartylglutamylvalinyl)aspartate
7-amino-4-trifluoromethylcoumarin;
DEVD-pNA, N-(N-acetylaspartylglutamylvalinyl)aspartate
p-nitroanilide;
DTT, dithiothreitol;
LEHD-AFC, N-(N-acetylleucinylglutamylhistidyl)aspartate
7-amino-4-trifluoromethylcoumarin;
PMSF, -phenylmethylsulfonyl fluoride;
pNA, p-nitroaniline;
TLCK, N-p-tosyl-L-lysine
chloromethyl ketone;
TPCK, N-p-tosyl-L-phenylalanine
chloromethyl ketone;
VEID-AFC, N-(N-acetylvalinylglutamylisoleucyl)aspartate
7-amino-4-trifluoromethylcoumarin;
ZEK(bio)D-aomk, N-(N-benzyloxycarbonylglutamyl-N
-biotinyllysyl)
aspartic acid [(2, 6-dimethylbenzoyl)oxy]methyl ketone;
ZVAD-fmk, N-(N-benzyloxycarbonylvalinylalanyl)aspartate
fluoromethyl ketone;
PAGE, polyacrylamide gel electrophoresis;
HPLC, high pressure liquid chromatography;
CHAPS, 3-[(3cholamidopropyl)dimethylammonio]-1-propanesulfonic acid;
PBS, phosphate-buffered saline.
| |
REFERENCES |
| 1.
|
Wyllie, A. H.,
Kerr, J. F. R.,
and Currie, A. R.
(1980)
Int. Rev. Cytol.
68,
251-306[Medline]
[Order article via Infotrieve]
|
| 2.
|
Arends, M. J.,
and Wyllie, A. H.
(1991)
Int. Rev. Exp. Pathol.
32,
223-254[Medline]
[Order article via Infotrieve]
|
| 3.
|
Hannun, Y. A.
(1997)
Blood
89,
1845-1853 |
TOP
ABSTRACT
INTRODUCTION
