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Blood, Vol. 94 No. 5 (September 1), 1999:
pp. 1683-1692
Calpain Functions in a Caspase-Independent Manner to Promote
Apoptosis-Like Events During Platelet Activation
By
Beni B. Wolf,
Joshua C. Goldstein,
Henning R. Stennicke,
Helen Beere,
Gustavo P. Amarante-Mendes,
Guy S. Salvesen, and
Douglas R. Green
From the Division of Cellular Immunology, La Jolla Institute for
Allergy and Immunology, San Diego, CA; the Department of Internal
Medicine, University of California, San Diego, CA; and The Program for
Apoptosis and Cell Death, The Burnham Institute, La Jolla, CA.
 |
ABSTRACT |
Apoptosis and platelet activation share common morphological and
biochemical features. Because caspases are essential mediators of
apoptosis, we examined whether platelets contain these proteinases and
use them during platelet activation. Human platelets contained caspase-9, caspase-3, and the caspase activators APAF-1 and cytochrome c as shown by sodium dodecyl sulfate-polyacrylamide gel electrophoresis and Western blotting. Upon treatment with cytochrome c and dATP, platelet cytoplasmic extracts recapitulated apoptotic events, including
sequential activation of procaspase-9 and procaspase-3 and subsequent
proteolysis of caspase substrates. Calcium ionophore-stimulated platelets also recapitulated apoptotic events, including cell shrinkage, plasma membrane microvesiculation, phosphatidyl serine externalization, and proteolysis of procaspase-9, procaspase-3, gelsolin, and protein kinase C- . Strikingly, however, these events occurred without caspase activation or release of mitochondrial cytochrome c, suggesting a role for a noncaspase proteinase. Supporting this, inhibition of the calcium-dependent proteinase, calpain, prevented caspase proteolysis, `apoptotic' substrate cleavage, and
platelet microvesiculation. In vitro, purified calpain cleaved recombinant procaspase-9 and procaspase-3 without activating either caspase, confirming the inhibitor studies. These data implicate calpain
as a potential regulator of caspases and suggest that calpain, not
caspases, promotes apoptosis-like events during platelet activation.
© 1999 by The American Society of Hematology.
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INTRODUCTION |
PROFOUND ALTERATIONS in cellular
architecture characterize apoptotic cell death. Regardless of the
initiating signal, the cell shrinks, the plasma membrane blebs and
vesiculates, and phosphatidylserine (PS) redistributes to the cell
surface. Meanwhile, the chromatin condenses, the nucleus breaks up, and
the DNA fragments into oligonucleosomal pieces. These events culminate
in removal of the apoptotic cell by neighboring phagocytes without
induction of an inflammatory response.1
Nuclear collapse and DNA fragmentation are dramatic and universal
features of apoptosis. However, the nucleus itself is not required for
apoptosis because apoptotic stimuli readily induce apoptotic
morphological features in anucleate cytoplasts.2-4 Furthermore, platelets, small anucleate cells derived from bone marrow
megakaryocytes, undergo events similar to apoptosis upon activation
with hemostatic agents like thrombin and collagen or stimulation with
pharmacological agents like calcium ionophore.5-8 Regardless of the stimulus, the platelet shrinks and extends filopodia, the plasma membrane blebs and microvesiculates, and PS redistributes to
the platelet surface.5-8 These events promote hemostasis by inducing platelet aggregation, platelet adhesion to injured blood vessels, and activation of the coagulation cascade.
Caspase proteinases are centrally involved in apoptotic signaling and
execution.1 Caspases are aspartate-specific cysteine proteinases that exist as latent zymogens, but once activated by
apoptotic signals, they promote apoptosis by specific limited proteolysis of key cellular substrates. Death stimuli typically facilitate autoactivation of initiator caspases such as caspase-8 and
caspase-9. The adapter molecule FADD initiates caspase-8 activation following ligation of death receptors such as Fas.9,10
Chemotherapeutic agents, UV-irradiation, and other apoptotic stimuli
cause release of mitochondrial cytochrome c into the
cytosol.11,12 Cytochrome c then binds to apoptotic
proteinase activating factor-1 (APAF-1) and this complex, along with
adenine nucleotides, promotes caspase-9 autoactivation.13
Active caspase-8 also causes release of mitochondrial cytochrome c,
thereby linking the activation pathways.14,15 Once
activated, both caspase-8 and caspase-9 activate caspase-3, which in
turn cleaves other caspases, fodrin, gelsolin, protein kinase C-
(PKC), and DNA fragmentation factor-45 (DFF45).1,16-22 Proteolysis of these and other caspase substrates induces the hallmarks
of apoptotic cell death.
Calpain, a calcium-dependent proteinase, may also function in apoptosis
by cleaving cytoskeletal proteins such as fodrin, actin, and the
pro-apoptotic protein Bax.18,23,24 It is unclear, however,
whether calpain functions upstream or downstream of caspases in
apoptosis.18,25-27 During platelet activation, calpain
promotes platelet microvesiculation, clot retraction, and proteolysis
of fodrin and other cytoskeletal components.28-31
Therefore, calpain functions in both apoptosis and platelet activation.
However, whether caspases participate in platelet activation is unknown.
In this report, we examine whether platelets contain caspases and use
them during platelet activation. We found that human platelets contain
caspase-9, caspase-3, and the caspase activators APAF-1 and cytochrome
c. In vitro, these proteins recapitulate the cytoplasmic events of
apoptosis. Surprisingly, however, we found that calpain, not caspases,
promotes apoptosis-like events during platelet activation. Therefore,
platelet activation is not equivalent to apoptosis and certain aspects
of the apoptotic phenotype may occur without caspase activation.
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MATERIALS AND METHODS |
Materials.
Antibodies against the following proteins were purchased commercially:
caspase-3 (Pharmingen, La Jolla, CA), cytochrome c (Pharmingen), fodrin (Chemicon International, Temecula, CA), PKC (Santa Cruz Biotechnology, Santa Cruz, CA), gelsolin (Sigma, St Louis,
MO), CD3 (Sigma), and glycoprotein (gp) IIb (Southern Biotech, Birmingham, AL). Dr Xiadong Wang (University of Texas Southwestern Medical Center, Dallas, TX) provided anti-DFF45 polyclonal
antibodies.22 Antisera against APAF-1 and caspase-9 was
prepared by immunizing rabbits with an APAF-1 peptide
(S38EEEKVRNEPTQQQR52) or with recombinant
caspase-9. Horseradish peroxidase-conjugated secondary antibodies were
from Amersham (Arlington Heights, IL). N-acetyl-Asp-Glu-Val-Asp-p-nitroanilide (DEVD-pNA),
benzoyloxycarbonyl-Asp-Glu-Val-Asp-amino-4-trifluoro-methyl-coumarin (DEVD-AFC), N-acetyl-Tyr-Val-Ala-Asp-AFC (YVAD-AFC),
N-acetyl-Val-Glu-Ile-Asp-AFC (VEID-AFC),
N-acetyl-Leu-Glu-His-Asp-AFC (LEHD-AFC), succinyl-Leu-Leu-Val-Tyr-AFC (LLVY-AFC), and benzoyloxycarbonyl-Val-Ala-Asp(OMe)-fluoromethyl ketone
(ZVAD-fmk) were from Enzyme Systems Products (Livermore, CA).
Benzoyloxycarbonyl-Val-Ala-Asp-fluoromethyl ketone (ZVAD-fmk*) was from
Bachem. All other reagents were from Sigma.
Proteinases.
Proteolytically active caspases and caspase zymogens were expressed in
Escherichia coli and purified as described.16,32 Active caspase-3 was active site titrated with ZVAD-fmk*.16 35S-procaspase-9 was prepared using a procaspase-9 cDNA
(provided by Dr Emad Alnemri, Thomas Jefferson University,
Philadelphia, PA) and the TnT reticulocyte lysate system (Promega,
Madison, WI).33 Porcine µ-calpain was purchased from
Calbiochem (San Diego, CA) and active site titrated with calpeptin.
Hereafter we will refer to µ-calpain simply as calpain. Granzyme B
(13 U/µg) was from Enzyme Systems Products.
Platelets and cells.
Platelets were isolated from human platelet concentrates (San Diego
Blood Bank, San Diego, CA) or from whole blood obtained from normal
human donors using a protocol approved by the Review Board for Human
Studies at the La Jolla Institute for Allergy and
Immunology.34 A mixed population of lymphocytes and
granulocytes was obtained from the buffy coat after hypotonic lysis of
red blood cells.35 Jurkat cells were originally from
American Type Culture Collection (ATCC; Rockville, MD) and maintained
as described.36
Sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE)
and Western blotting.
Whole-cell lysates or cytosolic extracts were denatured under reducing
conditions and subjected to SDS-PAGE using the Laemmli buffer system.
Resolved components were electrotransferred to nitrocellulose membranes
and analyzed by Western blotting with the indicated
antibodies.36,37
Cell-free apoptosis.
Cytosolic extracts from Jurkat cells and buffy coat cells were prepared
as described.36,37 Extracts were prepared from platelets
using the same method except that platelets were subjected to 3 freeze/thaw cycles before homogenization. Extract protein concentrations were determined with the Bradford dye reagent (BioRad). Cell-free apoptosis was initiated with cytochrome c and dATP as indicated in the figure legends. At the indicated times, aliquots were
withdrawn for proteinase assay and Western blot analysis.37 To detect caspase-dependent DNase activity, extracts were incubated with  106 rat liver nuclei and buffer, caspase-3 (100 nmol/L), or cytochrome c (10 µmol/L) and dATP (1 mmol/L). DNA
fragmentation was visualized via agarose gel electrophoresis and
ethidium bromide staining.36
Platelet activation.
Before activation, freshly isolated platelets were suspended at a
concentration of 2 × 107/mL in buffer A (10 mmol/L
HEPES pH 7.4, 137 mmol/L NaCl, 2.7 mmol/L KCl, 1.7 mmol/L
MgCl2, 3 mmol/L CaCl2, 25 mmol/L glucose, 0.05% bovine serum albumin). Platelets were treated with buffer or
calcium ionophore A23187 (2 µmol/L) and incubated at 37°C for the
indicated times. Cell shrinkage (decreased forward scatter) and PS
externalization were monitored by flow cytometry as
described.4 In addition, whole-cell lysates were prepared
and analyzed for caspase activity, caspase processing, and apoptotic
substrate cleavage. For comparison, control and UV-treated (apoptotic)
Jurkat cells were also analyzed.38
Platelet subcellular fractionation.
Platelets were preincubated for 30 minutes at room temperature in the
absence or presence of ZVAD-fmk (25 µmol/L) or calpeptin (25 µmol/L) and then stimulated with ionophore A23187. After 15 minutes,
subcellular fractions were prepared by differential centrifugation as
decribed.28,29 This procedure yields a 15,000g
pellet (P15) and supernatant (S15) as well as a 100,000g pellet
(P100), which represent the mitochondrial, cytosolic, and
microvesicular fractions, respectively. The P15 and S15 fractions were
analyzed by SDS-PAGE and Western blotting with anti-cytochrome c
antibodies to detect release of mitochondrial cytochrome
c.11 The P100 fractions were suspended in 50 µL of
caspase buffer (20 mmol/L PIPES, 100 mmol/L NaCl, 10 mmol/L
dithiothreitol, 1 mmol/L EDTA, 0.1%
3-[(3-cholamidopropyl)-dimethylammonio]-1-propanesulfonic acid
(CHAPS), 10% sucrose, pH 7.2). Aliquots (10 µL) of each sample were
subjected to SDS-PAGE and Western blotting with anti-gp IIb antibodies
to assess microvesiculation.28,29
Proteinase assays.
To detect caspase-3-like activity, samples (75 µg protein) from
cell-free reactions were incubated at 37°C for 30 minutes with
DEVD-pNA (100 µmol/L in caspase buffer). Substrate hydrolysis was
measured by sample absorbance at 406 nm. A molar extinction coefficient
of 10,000 M 1cm1 was used to
determine the concentration of pNA generated from substrate hydrolysis.
For studies involving platelet activation and apoptosis, we determined
caspase activity using more sensitive AFC-based fluorogenic substrates.
LEHD-AFC and DEVD-AFC were used to detect caspase-9-like and
caspase-3-like activity, respectively.39 Whole-cell
lysates (25 µg protein) were incubated with 100 µmol/L LEHD-AFC or
DEVD-AFC in caspase buffer at 37°C. Initial rates of substrate
hydrolysis were determined using a Tecan SpectroFluor fluorimeter in
the kinetic mode. Excitation was at 400 nm and emission was at 500 nm
(slit widths 10 nm). A standard curve of AFC concentration versus
fluorescence was used to determine the concentration of AFC generated
from substrate hydrolysis.
Inhibitor studies with purified proteinases.
Purified calpain or caspase-3, each at a final concentration of 100 nmol/L, were incubated with ZVAD-fmk (0 to 10 µmol/L) or calpeptin (0 to 10 µmol/L) for 20 minutes at room temperature. Residual enzyme
activity was then measured fluorimetrically using LLVY-AFC (100 µmol/L) for calpain or DEVD-AFC (100 µmol/L) for caspase-3.39,40 Calpain reactions were conducted in calpain buffer (caspase buffer lacking EDTA and supplemented with 1 mmol/L CaCl2) and caspase-3 reactions were conducted in caspase
buffer. Initial rates of substrate hydrolysis were determined at each inhibitor concentration. Results were plotted as percent maximal enzyme
activity (determined in the absence of inhibitor) versus the logarithm
of the inhibitor concentration.
Reaction of calpain with procaspase-9 and procaspase-3 in vitro.
Calpain was reacted with 35S-procaspase-9 or
recombinant procaspase-3 for various times at 37°C in calpain
buffer as detailed in the legend for Fig 6. Reaction
products were analyzed by SDS-PAGE followed by autoradiography
(35S-procaspase-9) or staining with Coomassie brilliant
blue (procaspase-3).
To determine if calpain activates procaspase-3, procaspase-3 (0.5 µmol/L) was reacted with calpain (100 nmol/L) for 15 minutes at
37°C and then caspase-3 activity was measured by monitoring DEVD-AFC hydrolysis as described above. As a positive control, procaspase-3 was activated with granzyme B (1 U) under the same conditions. We also examined whether granzyme B could activate calpain-processed caspase-3 because preliminary studies showed that
calpain-cleaved procaspase-3 was not active. To accomplish this, we
treated procaspase-3 with calpain for 15 minutes and then with granzyme
B (1 U) for an additional 15 minutes before DEVDase assay.
Identification of calpain cleavage sites in procaspase-3 and
procaspase-9.
Fifty micrograms of each caspase zymogen was digested with 100 nmol/L
calpain in calpain buffer for 15 minutes at 37°C. The active site
mutant of caspase-9 was used because this allows purification of large
amounts of the zymogen from E coli. Reaction products were
separated by 8% to 18% SDS-PAGE, blotted to polyvinylidene fluoride membranes and the major fragments sequenced by
Edman degradation using an Applied Biosystems 476A Protein Sequencer.
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RESULTS |
Identification of platelet caspases and caspase activators.
To determine whether human platelets contain caspases and their
activators, we subjected platelet lysates to Western blotting with
antibodies against these proteins. We analyzed equivalent amounts of
Jurkat cell lysates for comparison. As shown in
Fig 1, comparable amounts of APAF-1,
cytochrome c, caspase-9, and caspase-3 were present in platelets and
Jurkat cells. Both caspase-9 and caspase-3 were detected solely as the
zymogen forms in both extracts. No APAF-1 or caspase-9 immunoreactivity
was detected when preimmune rabbit serum was substituted for immune
antiserum (not shown). Because platelets contain abundant amounts of
adenine nucleotides,41 platelets contain all components
necessary for mitochondria-dependent caspase activation.
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| Fig 1.
Human platelets contain caspases and caspase activators.
Whole-cell lysates (25 µg protein) were subjected to SDS-PAGE and
Western blotting with antibodies against apoptotic proteins (APAF-1,
cytochrome c, caspase-9, caspase-3, DFF45), a platelet marker (gpIIb),
a T-cell marker (CD3), and actin as a loading control. The presented
blots were performed with lysates from platelet concentrates.
Comparable results were obtained with freshly isolated platelets.
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To assess the purity of our platelet preparations, we subjected
platelet lysates to Western blotting with antibodies against gpIIb, a
platelet marker, and antibodies against CD3, a component of the T-cell
receptor. Although the preparation showed immunoreactivity to gpIIb,
the platelets contained no detectable CD3 immunoreactivity (Fig 1).
Additionally, platelet extracts (50 µg protein) contained no
detectable caspase-activated DNase activity in a cell-free assay and
lacked DFF-45, a caspase substrate that must be cleaved before DNA
fragmentation in nucleated cells (Fig 1 and data not shown).21,22 Since caspase-dependent DNA fragmentation was detectable in Jurkat cell extracts or buffy coat extracts with as
little as 0.1 µg of extract (not shown), we conclude that our platelet preparations were not significantly contaminated with nucleated cells. Comparable results were obtained with freshly isolated
platelets and platelets isolated from platelet concentrates.
Cytochrome c and dATP-dependent caspase activation.
We next determined if cytochrome c and dATP would activate platelet
caspase-9 and caspase-3 in vitro. To accomplish this, we incubated
platelet cytosolic extracts with and without cytochrome c and dATP and
then analyzed procaspase processing by SDS-PAGE and Western blotting.
As shown in Fig 2, cytochrome c and dATP initiated rapid sequential processing of procaspase-9 and procaspase-3 to their proteolytically active fragments. Procaspase processing coincided with detection of caspase-3-like DEVDase activity in the
cytochrome c and dATP-treated platelet extracts (Fig 2C).39 No caspase activation occurred in the absence of cytochrome c and dATP.
Thus, cytochrome c and dATP initiate sequential activation of caspase-9
and caspase-3 in platelet cytosolic extracts as in extracts from
nucleated cells.33,37
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| Fig 2.
Cytochrome c and dATP initiate sequential caspase-9 and
caspase-3 activation in platelet cytosolic extracts. Platelet cytosolic
extracts derived from hospital concentrates (final concentration, 5 mg
protein/mL) were treated with buffer or cytochrome c (10 µmol/L) and
dATP (1 mmol/L) and then analyzed by SDS-PAGE and Western blotting (35 µg protein/lane). (A) Procaspase-9 processing in response to
cytochrome c and dATP. (B) The blot from (A) was stripped and reprobed
with anti-caspase-3 antibodies. At the indicated times after
incubation with buffer ( ) or cytochrome c and dATP ( ), aliquots
(75 µg protein) of cell-free reactions were analyzed for activity
against DEVD-pNA (C). (A) and (B) are representative of 3 independent
experiments. In (C), error bars represent the SEM, N = 3.
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Cytochrome c and dATP-dependent caspase activation promotes
proteolysis of apoptotic substrates.
As shown in Fig 3, cytochrome c and
dATP-dependent caspase activation in platelet extracts leads to
proteolysis of gelsolin, PKC, and fodrin, proteins cleaved during
apoptosis.17-20 Cleavage of these substrates was evident
within 30 minutes and nearly complete by 3 hours. In the absence of
cytochrome c and dATP, no substrate cleavage was observed, even at the
latter time points. Thus, cytochrome c and dATP initiate caspase
activation and subsequent cleavage of caspase substrates in platelet
extracts. Therefore, platelet extracts recapitulate cytosolic events
that occur in apoptotic cells and in cytosolic extracts from nucleated
cells.
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| Fig 3.
Cytochrome c and dATP-dependent caspase activation
promotes proteolysis of apoptotic substrates. Cell-free reactions were
prepared as described in the legend to Fig 2. At the indicated times,
portions (50 µg protein) of the extract were removed for Western
blotting with antibodies against PKC (A), gelsolin (B), or fodrin
(C). Molecular weights (kD) are listed for the endogenous and cleaved
forms.
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A comparison of platelet activation and apoptosis.
Because activated platelet extracts demonstrated caspase activity in
vitro, we wished to determine whether caspases function during platelet
activation. To accomplish this, we stimulated freshly isolated
platelets with calcium ionophore A23187 and then examined caspase
processing and activity. Naive and UV-irradiated Jurkat cells were also
analyzed to compare apoptotic events with similar events during
ionophore stimulation. We observed cell shrinkage (decreased forward
scatter) and PS externalization by fluorescence-activated cell sorting
(FACS) analysis in both ionophore-stimulated platelets and
UV-treated (apoptotic) Jurkat cells (not shown). However, these events
occurred within 15 minutes in ionophore-treated platelets, but required
6 hours to occur in apoptotic cells. In the studies presented in
Fig 4, greater than 90% of A23187-treated platelets were PS positive, compared with 1.5% of untreated platelets. With Jurkat cells, 68% of UV-treated cells were PS positive, compared with 3% of untreated cells.
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| Fig 4.
Caspase activation accompanies apoptosis but not platelet
activation. Platelets were activated with calcium ionophore A23187 (2 µmol/L) for 15 minutes at 37°C. For comparison, control and
apoptotic (6 hours post UVB-treatment) Jurkat cells were examined in
parallel with platelets. Platelet activation and apoptosis were
confirmed by monitoring cell shrinkage (decreased forward scatter) and
PS externalization by FACS analysis as detailed in the text. Cell
lysates were analyzed by SDS-PAGE and Western blotting (50 µg
protein/lane) with antibodies against caspase-9 (A) and caspase-3 (B).
Fifty micrograms of lysate were also assayed against LEHD-AFC and
DEVD-AFC to detect caspase-9-like (A, bottom panel) and
caspase-3-like activity (B, bottom panel). In (C), control and
A2387-treated platelets were subjected to subcellular fractionation.
The cytosolic (S15) and mitochondrial (P15) fractions (40 µg protein)
were then subjected to SDS-PAGE and Western blotting with
anti-cytochrome c antibodies. The presented data are representative of
3 independent experiments.
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As shown in Fig 4, apoptosis correlated with processing of procaspase-9
and procaspase-3 and the onset of caspase activity (Fig 4A and B).
After 6 hours, caspase-9 immunoreactivity was no longer detectable in
the apoptotic Jurkat cells; however, a low level of caspase-9-like
activity was detected using the substrate LEHD-AFC.39 The
apoptotic Jurkat cells contained only the active form of caspase-3 and
showed activity against the caspase-3 substrate DEVD-AFC.39
Procaspase processing also occurred in the ionophore-stimulated platelets; however, no caspase activity was detected (Fig 4A and B).
Procaspase-9 was cleaved to multiple fragments, primarily to an
30-kD form. This contrasts to the 35-kD active form of caspase-9 observed in response to cytochrome c and dATP (Fig 2). A
slight increase in LEHDase activity was observed in the
ionophore-stimulated platelets; however, this result was not
statistically significant as assessed by the paired Student's
t-test. Similarly, procaspase-3 was processed to an 30-kD
form during platelet activation, which is distinct from the active p17
caspase-3 fragment observed during apoptosis and in response to
cytochrome c and dATP (Figs 2 and 4B). No caspase-3-like DEVDase
activity was detected after ionophore stimulation. Thus, procaspase
processing occurs without detectable caspase activation during
ionophore stimulation. Furthermore, cytochrome c did not translocate to
the platelet cytosol (S15) during platelet activation (Fig 4C),
eliminating the possibility that caspase processing requires cytochrome
c release or APAF-1.
To confirm that the preceding findings were not limited to ionophore
stimulation, we examined caspase processing and activity after
treatment of platelets with physiological platelet agonists. We
observed procaspase-3 processing to an 30-kD form after treatment of
platelets with thromboxane agonist U46619, thrombin, or thrombin and
collagen; however, no caspase-3-like DEVDase activity was detected
(not shown). Procaspase-9 processing was not examined; however, none of
the preceding treatments induced activity against the caspase-9
substrate LEHD-AFC. Furthermore, anti-Fas antibodies did not activate
platelets or induce platelet caspase activation (not shown). Therefore,
platelet activation apparently occurs independently of mitochondria-
and death-receptor-dependent caspase pathways.
ZVAD-fmk and calpeptin inhibit microvesiculation, caspase processing,
and substrate cleavage during platelet activation.
We next sought to determine whether calpain might promote
apoptosis-like events during platelet activation or whether platelet activation proceeds with a very low level of caspase activity (below
the assay detection level). To accomplish this, we examined the effect
of ZVAD-fmk and calpeptin, 2 cell-permeable proteinase inhibitors, on
platelet activation. ZVAD-fmk inhibits most caspases and calpeptin
potently inhibits calpain.42,43 Calpeptin and other calpain
inhibitors like MDL 28,170 therefore inhibit calpain-dependent platelet
activation events.28 We incubated platelets with buffer, ZVAD-fmk, or calpeptin for 30 minutes and then stimulated them with
ionophore A23187. After 15 minutes of activation, we assessed how the
inhibitors affected various platelet activation parameters.
Ionophore A23187 induced platelet microvesiculation, PS
externalization, caspase processing, and cleavage of the `apoptotic' substrates, gelsolin and PKC (Fig 5).
ZVAD-fmk and calpeptin attenuated microvesiculation, caspase
processing, and substrate cleavage; however, calpeptin was a more
potent inhibitor. By contrast, these inhibitors did not significantly
alter PS externalization or cell shrinkage (Fig 5B and not shown). We
observed no activity against the caspase substrates LEHD-AFC, DEVD-AFC,
VEID-AFC, or YVAD-AFC in the absence or presence of inhibitors (Fig 4
and not shown), suggesting that ZVAD-fmk inhibits a noncaspase
proteinase(s) that promotes proteolysis of caspases, gelsolin and
PKC.39 Because ZVAD-fmk inhibited the same events as
calpeptin, the data suggest that ZVAD-fmk inhibits calpain and that
calpain cleaves caspases and `apoptotic' substrates.
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| Fig 5.
Calpain inhibition prevents platelet microvesiculation,
caspase processing, and apoptotic substrate cleavage, but not PS
externalization. Platelets were preincubated with vehicle (lanes 1 and
2), ZVAD-fmk (25 µmol/L; lane 3) or calpeptin (25 µmol/L; lane 4)
for 30 minutes at room temperature and then treated with buffer (lane
1) or A23187 (lanes 2 through 4) for 15 minutes at 37°C.
Microvesicles were then isolated from platelet supernatants as
described in Materials and Methods and analyzed by Western blotting
(10-µL aliquots/lane) with anti-gpIIb antibodies (A). The graph in
(A) shows the extent of microvesiculation as assessed by gpIIb
densitometry (arbitrary units). In (B), plasma membrane PS
externalization was determined by annexin V-FITC binding and FACS
analysis. (C) Whole-cell lysates analyzed by Western blotting
with antibodies against caspase-9, caspase-3, gelsolin, and
PKC (50 µg protein/lane). (D) Inhibition of 100 nmol/L purified
calpain and caspase-3 by ZVAD-fmk ( ) and calpeptin (). Error bars
represent the SEM, N = 4.
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To determine the relative potency of ZVAD-fmk and calpeptin for calpain
and caspase-3, we incubated the purified proteinases with various
concentrations of the inhibitors and then determined residual
proteinase activity against fluorogenic substrates. Both ZVAD-fmk and
calpeptin effectively inhibited calpain, with both inhibitors
completely abolishing calpain activity by 1 µmol/L inhibitor (Fig
5D). ZVAD-fmk potently inhibited caspase-3; however, calpeptin was a
poor caspase-3 inhibitor (Fig 5D). Together with the preceding data,
these results suggest that ZVAD-fmk and calpeptin inhibit calpain
during platelet activation and this inhibition prevents caspase
processing and `apoptotic' substrate cleavage.
Reaction of purified calpain with procaspase-9 and procaspase-3 in
vitro.
We next incubated purified calpain with 35S-procaspase-9 to
determine if the proteinase directly processed the zymogen. As shown in
Fig 6A, calpain rapidly processed the 45-kD
procaspase-9 to several fragments. Major fragments of 30 to 34 kD
and 11 kD were observed. Nearly complete procaspase-9 processing was
observed within 10 minutes and no further proteolysis was observed
after 1 hour of incubation. ZVAD-fmk and calpeptin abolished
procaspase-9 processing (Fig 6A, lanes 8 and 9).
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| Fig 6.
Purified calpain removes the prodomain of caspase-9 and
caspase-3 via specific limited proteolysis. Calpain (100 nmol/L) was
reacted with 35S-procaspase-9 (5 µL; A) or recombinant
procaspase-3 (10 µmol/L; B) in the absence and presence of calpeptin
(1 µmol/L) or ZVAD-fmk (1 µmol/L) and the products were analyzed by
SDS-PAGE and autoradiography (A) or staining with Coomassie Blue (B).
To determine whether calpain activates procaspase-3, we treated the
procaspase with calpain and then measured caspase-3 activity by
monitoring DEVD-AFC hydrolysis as shown in (C). The graph shows
DEVD-AFC hydrolysis by procaspase-3 (1), procaspase-3 reacted with
calpain (2), granzyme B (3), or calpain then granzyme B (4) (error bars
represent the SEM, N = 3). Columns 5 and 6 show the DEVDase activity
of calpain and granzyme B, respectively. In (D), purified procaspase-9
and procaspase-3 were digested with calpain and the products sequenced
by Edman degradation. The insets show the calpain cleavage sites. C287
and C163 denote the caspase-9 and caspase-3 active site cysteines. The
shaded areas between the caspase large and small subunits represent the
inter-domain linker that is removed upon caspase activation.
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Calpain also showed activity against purified procaspase-3 (Fig 6B).
Calpain processed the zymogen to an 30-kD fragment within 15 minutes
at 37°C. ZVAD-fmk and calpeptin both attenuated procaspase-3 processing (lanes 5 and 6). The 16-kD band present in all lanes represents a contaminant, which does not promote procaspase-3 activation, as the preparation showed no activity against DEVD-AFC (Fig
6C). Calpain-treated procaspase-3 showed no activity against DEVD-AFC,
indicating that calpain cleavage does not activate procaspase-3 (Fig
6C). By contrast, granzyme B, a potent caspase activator, activated
procaspase-3 within 15 minutes at 37°C, as shown by the appearance
of DEVDase activity.16 Granzyme B readily activated calpain-treated procaspase-3, indicating that calpain cleavage does not
inactivate procaspase-3.
Identification of the calpain cleavage sites in procaspase-9 and
procaspase-3.
To identify the calpain cleavage sites in procaspase-9 and
procaspase-3, we digested each zymogen with calpain and sequenced the
products by Edman degradation. Calpain cleaved procaspase-9 primarily
at Val120 and to a lesser extent at Ile115 and
Glu143 (Fig 6D). These cleavage sites are all located
between the procaspase-9 prodomain and the large subunit.44
Therefore, processing at any of these sites would not activate the
caspase. Similarly, calpain removed a fragment of the procaspase-3
prodomain by cleaving after Ser7.45 This
cleavage would not promote procaspase-3 activation, consistent with the
preceding activation studies.
| |
DISCUSSION |
In this study, we found that human platelets contain functional
pro-apoptotic caspases. Surprisingly, however, we found that calpain,
not caspases, promotes events resembling apoptosis during platelet
activation. This suggests that platelet activation is not equivalent to
apoptosis and that certain aspects of the apoptotic phenotype may occur
without caspase activation.
Platelets contain caspase-3, caspase-9, and the caspase activators
APAF-1 and cytochrome c, at comparable levels to nucleated cells (Fig
1). Furthermore, the extent and rate of caspase-3 activation and
apoptotic substrate cleavage observed in platelet extracts is similar
to that observed in extracts from nucleated cells, suggesting that the
2 extracts are functionally equivalent (Figs 2 and
3).33,36,37 However, platelets lack DFF45 (Fig 1) and consequently contain no detectable caspase-dependent DNase (DFF40) activity. The platelet cell-free system should therefore be useful to
dissect the role of DFF45, DFF40, and other molecules in nuclear apoptosis and DNA fragmentation. Furthermore, because megakaryocytes undergo apoptosis, study of megakaryocyte differentiation may provide
insight into developmental regulation of caspases and DNases.46
During apoptosis, caspases mediate cell shrinkage, plasma membrane
microvesiculation, PS externalization, and apoptotic substrate cleavage.1 We observed comparable events during platelet
activation; surprisingly, however, we found that caspases do not
participate in this process. Three lines of evidence support this
finding. First, apoptosis, but not platelet activation, correlated with procaspase processing and activation (Fig 4). Second, calpain inhibition, not caspase inhibition, prevented apoptosis-like events during A23187 stimulation (Fig 5). Third, there was no release of
mitochondrial cytochrome c during platelet activation, an event critical for caspase activation in many forms of apoptosis (Fig 4).
Because we also failed to detect caspase activation after treatment of
platelets with thrombin, thrombin and collagen, or a thromboxane
analog, we conclude that platelet activation occurs independently of
caspase activation.
Because platelet activation occurs irreversibly, the process results in
platelet death. In this context, platelet activation might represent a
nonapoptotic form of cell death that involves neither caspases nor
other pro-apoptotic molecules. Supporting this, many platelet
activation events including adhesion to endothelial cells, extension of
filopodia, self-aggregation, and release of intracellular granules
distinguish this process from apoptosis.5-7 G-protein-coupled receptors, adhesion receptors, and second messenger systems involving protein phosphorylation, calcium influx, and metabolism of inositol polyphosphates and diacyl glycerol mediate these
events; however, definitive roles for these signaling molecules in
apoptosis have not been established.5-7 Furthermore,
platelet activation proceeds much more rapidly than apoptosis and does not involve nuclear events. Thus, certain aspects of platelet activation are distinct from apoptosis and do not require caspases or
other apoptotic signaling molecules.
Alternatively, platelet activation pathways may use pro-apoptotic
molecules other than caspases. For example, Vanags et al8 recently detected upregulation of Bax, a pro-apoptotic Bcl-2 family member, in activated platelets. Bax can induce caspase-independent cell
death and this may contribute to platelet demise during
activation.47,48 Additionally, noncaspase proteinases such
as calpain might cleave caspase substrates to recapitulate apoptotic
events during platelet activation. Cleavage of caspase substrates like
gelsolin (Fig 6), PKC (Fig 6), fodrin, and pp125-focal adhesion
kinase by calpain supports this hypothesis.17,49-51
Two surprising findings in this study concern the calcium-dependent
proteinase, calpain. First, we found that the broad specificity caspase
inhibitor, ZVAD-fmk, potently inhibits calpain (Fig 5). Although
calpain prefers Tyr, Met, or Arg in the P1 position of peptide substrates, it cleaves procaspase-9 after a Glu residue (Fig 6)
and glucagon after an Asp residue.40 Therefore, our finding
that ZVAD-fmk inhibits calpain is not entirely unexpected, especially
since the fluoromethyl ketone (fmk) group efficiently modifies the
catalytic thiol of cysteine proteinases. Although this study focused on
µ-calpain, m-calpain demonstrates similar substrate specificity and
thus may also be susceptible to ZVAD-fmk.40 Because
ZVAD-fmk inhibits caspase-3 and calpain, inhibitor studies alone cannot
distinguish the biological activities of these proteinases. Caution
should therefore be used when using ZVAD-fmk to study apoptotic events.
A second interesting finding was that calpain cleaves procaspases
without activating them, both in vitro and during platelet activation
(Figs 5 and 6). Proteolysis of procaspase-3 removed a portion of the
caspase's prodomain. However, proteolysis did not inactivate caspase-3
because granzyme B activated the calpain-processed zymogen, in
agreement with studies we recently published concerning activation of
caspase-3 mutants.16 Full or partial deletion of the
procaspase-3 prodomain did not significantly alter procaspase-3 activation by caspase-8, caspase-10, or granzyme B. Furthermore, once
activated, the mutant proteins displayed equal reactivity toward
substrates and the baculovirus p35 caspase inhibitor. Together, these
results suggest that limited proteolysis of procaspase-3 does not alter
its interaction with other proteinases or inhibitors.
Calpain also removed the procaspase-9 prodomain during platelet
activation and in vitro in a purified system. Because binding of
caspase-9 to APAF-1 is necessary for its activity and because the
caspase-9 prodomain mediates this interaction,13,32,33 calpain-dependent removal of the prodomain could significantly abrogate
caspase-9 activation. Since calpain also did not activate procaspase-3
(Fig 5), this finding suggests that calpain must function downstream or
parallel to caspases during apoptosis. In fact, 2 recent reports
support this hypothesis.26,27 Alternatively, if calpain
acted on caspase-9 before an apoptotic stimulus, then removal of the
prodomain might promote cell survival by preventing caspase-9
activation and apoptosis.
In summary, human platelets contain functional pro-apoptotic caspases;
however, caspases do not function during platelet activation, even
though this process resembles apoptosis. What role might caspases play
in platelet biology? Because megakaryoctes can undergo apoptosis,
caspases might simply have been transferred to the platelet during
development. However, platelet production in vitro correlates with the
onset of megakaryocyte apoptosis, suggesting a potential role for
apoptosis and caspases in platelet production.46 By
contrast, because caspases and apoptosis often mediate cell senescence,
caspases might function during platelet senescence. Further study of
platelets and megakaryocytes will provide answers to these questions
and enhance our knowledge of the caspase system.
| |
ACKNOWLEDGMENT |
The authors thank Drs Xiadong Wang and Emad Alnemri for reagents.
| |
FOOTNOTES |
Submitted January 5, 1999; accepted May 4, 1999.
B.B.W. was supported by a Mentored Clinical Scientist Development Award
(CA75268-01). H.R.S. was supported by a grant from the Danish Natural
Science Foundation (9600412). G.S.S. and D.R.G. received funding from
the National Institutes of Health (NS37878 to G.S.S. and CA69831 and
AI40646 to D.R.G.). This is publication no. 300 from the La Jolla
Institute for Allergy and Immunology.
The publication costs of this
article were defrayed in part by
page charge payment. This article
must therefore be hereby marked
"advertisement"
in accordance with 18 U.S.C. section
1734 solely to indicate this fact.
Address reprint requests to Beni B. Wolf, MD, PhD, La Jolla Institute
for Allergy and Immunology, 10355 Science Center Dr, San Diego, CA
92121; e-mail: 102251.1444{at}compuserve.com.
| |
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S. Lankiewicz, C. Marc Luetjens, N. Truc Bui, A. J. Krohn, M. Poppe, G. M. Cole, T. C. Saido, and J. H. M. Prehn
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B. B. Wolf, M. Schuler, F. Echeverri, and D. R. Green
Caspase-3 Is the Primary Activator of Apoptotic DNA Fragmentation via DNA Fragmentation Factor-45/Inhibitor of Caspase-activated DNase Inactivation
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K. Blomgren, C. Zhu, X. Wang, J.-O. Karlsson, A.-L. Leverin, B. A. Bahr, C. Mallard, and H. Hagberg
Synergistic Activation of Caspase-3 by m-Calpain after Neonatal Hypoxia-Ischemia. A MECHANISM OF "PATHOLOGICAL APOPTOSIS"?
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V. Stoka, B. Turk, S. L. Schendel, T.-H. Kim, T. Cirman, S. J. Snipas, L. M. Ellerby, D. Bredesen, H. Freeze, M. Abrahamson, et al.
Lysosomal Protease Pathways to Apoptosis. CLEAVAGE OF Bid, NOT PRO-CASPASES, IS THE MOST LIKELY ROUTE
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B. B. Wolf, M. Schuler, W. Li, B. Eggers-Sedlet, W. Lee, P. Tailor, P. Fitzgerald, G. B. Mills, and D. R. Green
Defective Cytochrome c-dependent Caspase Activation in Ovarian Cancer Cell Lines due to Diminished or Absent Apoptotic Protease Activating Factor-1 Activity
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C. C. Q. Vu, C. D. Bortner, and J. A. Cidlowski
Differential Involvement of Initiator Caspases in Apoptotic Volume Decrease and Potassium Efflux during Fas- and UV-induced Cell Death
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TOP
ABSTRACT
INTRODUCTION