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Blood, Vol. 93 No. 12 (June 15), 1999:
pp. 4222-4231
Role of Caspase in a Subset of Human Platelet Activation Responses
By
Anna Shcherbina and
Eileen Remold-O'Donnell
From The Center for Blood Research and the Department of Pediatrics,
Harvard Medical School, Boston, MA.
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ABSTRACT |
Platelets function to protect the integrity of the vascular wall. A
subset of platelet activation responses that are especially important
for thrombus formation include exposure of phosphatidylserine and
release of microparticles, which generate procoagulant surfaces. The
resemblance of these platelet activation processes to events occurring
in nucleated cells undergoing apoptosis suggests a possible role for
caspases, which are major effector enzymes of nucleated cell apoptosis. We demonstrate here the presence of caspase-3 in human
platelets and its activation by physiological platelet agonists. Using
cell-permeable specific inhibitors, we demonstrate a role for a
caspase-3-like protease in the agonist-induced (collagen plus thrombin
or Ca2+ ionophore) platelet activation events of
phosphatidylserine exposure, microparticle release, and cleavage of
moesin, a cytoskeletal-membrane linker protein. The role of caspase-3
in platelet activation is restricted rather than global, because other
activation responses,  granule secretion, shape change, and
aggregation were unaffected by caspase-3 inhibitors. Experiments with
two classes of protease inhibitors show that caspase-3 function is
distinct from that of calpain, which is also involved in late platelet
activation events. These findings show novel functions of caspase and
provide new insights for understanding of platelet activation.
© 1999 by The American Society of Hematology.
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INTRODUCTION |
PLATELETS ARE THE major players of
hemostasis and thrombosis. Among other responses to physiological
agonists, activated platelets expose negatively charged
phosphotidylserine (PS) on the platelet exterior and release
PS-positive microparticles, both of which contribute to fibrin
deposition by providing competent surfaces for assembly of coagulation
factors and thrombus formation.1 Increased levels of
circulating microparticles are associated with various thrombotic
disorders, including activated coagulation, transient ischemia,
myocardial infarction, and postsurgery in cardiopulmonary bypass
patients.2-6 For these reasons, studies to delineate the
mechanisms of PS exposure and microparticle release are extremely
important for understanding platelet function.
Exposure of PS and release of microparticles occur only within the
larger process of platelet activation. Conversely, the activation
response of platelets is distinguished by a multiplicity of subevents
and regulatory mechanisms. Particularly noteworthy is the sequential
nature of the response and the temporal arrangement of individual
steps. Several parallels can be discerned between the programmed
activation response of platelets and programmed death of
nucleated cells the apoptotic process. Common characteristics include
the prominence of coordinated morphological changes and, to a large
extent, the quality of irreversibility. In platelet activation, as in
apoptosis, transbilayer migration of phosphatidylserine to the outer
membrane leaflet is an important event, facilitating interaction with
phagocytic cells and, in the case of platelets, generating, in
addition, the procoagulant surface. Also, the process of platelet
microparticle formation is morphologically similar to the membrane
blebbing phase of nucleated cell apoptosis. These shared features
prompted us to examine a possible role in platelet activation for
caspases, a protease family intimately associated with the programmed
death of nucleated cells.
Caspases, which are cysteinyl proteases that cleave after aspartic
acid, are key effectors of apoptosis. The family consists of at least
11 enzymes in humans.7-9 In response to apoptotic signals,
the proform of an initiator caspase containing an adapter domain, such
as caspase-8, -9, or -10, is recruited to a surface membrane complex or
a mitochondrial-derived complex, where it is autoproteolytically
processed to the two subunit active form. The initially activated
caspase processes/activates other procaspases that lack adapter
domains, including caspase-3 and -7, and the latter, the effector
caspases, incapacitate essential homeostatic pathways by limited
cleavage of specific targets.
Another hallmark of nucleated cell apoptosis is cleavage of select
cytoskeletal proteins by effector caspases.10,11 We focus
in this study on one cytoskeletal protein, moesin, the only member in
human platelets12,13 of the ERM
(ezrin-radixin-moesin) family proteins, which stabilize surface
projections by linking the underlying actin cytoskeleton with the
plasma membrane.14,15 In response to agonist, platelet
moesin undergoes rapid phosphorylation, localizes transiently to newly
formed filopodial/lamellipodial projections,12,16 and
subsequently is proteolytically cleaved,16a a reaction
expected to terminate moesin's linker function and facilitate late
platelet cytoskeletal changes required for clot retraction and
microparticle release. Moesin cleavage requires calpain
(Ca2+-activated protease), which is required also for
microparticle release17,18; however, pure calpain alone
does not cleave moesin,13 suggesting a requirement for a
second protease.
The present report breaks new ground by demonstrating the presence and
novel function of caspase in human platelets. We identify the zymogen
form of the effector caspase, caspase-3, as a component of human
platelets and demonstrate that procaspase-3 becomes activated when
isolated platelets are stimulated by physiological agonists. A specific
cell-permeant inhibitor of caspase-3 is shown to abrogate agonist-induced PS exposure and microparticle release. The caspase inhibitor also prevents cleavage of the structural protein moesin in
activated platelets. Comparative experiments with protease inhibitors
showed that both caspase and calpain function in agonist-induced late
events of platelet activation (PS exposure, microparticle release, and
moesin cleavage) and demonstrate that the two proteases have distinct roles.
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MATERIALS AND METHODS |
Platelet isolation.
Freshly drawn blood from normal healthy donors, who gave written
consent, was collected in acid-citrate-dextrose (ACD; NIH formula A) in
plastic and fractionated immediately at ambient temperature. Cells were
counted using a MAX-M Blood Cell Analyzer (Coulter Corp, Hialeah, FL).
The blood was centrifuged at 200g for 12 minutes to separate
platelet-rich plasma (PRP). Additional ACD was added (1 part ACD per 3 parts PRP) and platelets were pelleted at 800g for 15 minutes.
The platelets were resuspended in platelet buffer (10 mmol/L
Tris-hydroxymethyl-methyl-2-aminoethane sulphonic acid [TES], pH 7.2, 136 mmol/L NaCl, 2.6 mmol/L KCl, 0.5 mmol/L
NaH2PO4, 2 mmol/L MgCl2, 0.1%
glucose, and 0.1% bovine albumin) and, after the addition of ACD (20%
of final volume) and prostacyclin (1 µg/mL; Calbiochem, San Diego,
CA), were centrifuged at 800g for 10 minutes. The isolated
platelets contained no detectable erythrocytes and less than 1 leukocyte per 4,000 platelets. For activation experiments, platelets
were suspended in platelet buffer and allowed to recover for 90 minutes
at 37°C to ensure their resting state.
Peptidase assay.
Platelets (5 × 108) in 1 mL of platelet buffer with 2 mmol/L CaCl2 and 3 µmol/L A23187 were incubated with
stirring in flat-bottom polyethylene vials (14-mm diameter) at 37°C
for the indicated time and were lysed by adding 1/3 vol of 4% Triton
X-100, 8 mmol/L EGTA, 20 mmol/L dithiothreitol, 200 µg/mL aprotinin,
200 µg/mL benzamidine, and 200 µg/mL leupeptin. The Triton lysates
(200 µL) were combined with 0.1 mmol/L of
N-acetyl-Asp-Glu-Val-Asp-p-nitro anilide (DEVD-pNA; Enzyme Systems
Products, Livermore, CA) or N-acetyl-Tyr-Val-Ala-Asp-p-nitroanilide
(YVAD-pNA; Sigma, St Louis, MO). The reactions were
incubated for 2 hours in flat-bottom 96-well plates at 37°C with
monitoring of OD405 nm. The mean  OD per minute was
calculated from the linear range of the reaction.
Immunoblotting.
Platelets (5 × 108/mL) were lysed by adding an equal
volume of 2% sodium dodecyl sulfate (SDS), 120 mmol/L Tris-HCl, pH
6.8, 4% mercaptoethanol, 100 µg/mL leupeptin, 4 mmol/L EGTA, and 2 mmol/L diisopropyl fluorophosphate (DFP) and heating for more than 3 minutes at 100°C. The lysates were fractionated by
SDS-electrophoresis on 8 or 12% polyacrylamide gels (Novex, San Diego,
CA).19 Polypeptides were transferred to nitrocellulose at
constant 80 mA at approximately 22°C for 16 hours. The membranes
were blocked with 2% normal rabbit serum in phosphate-buffered saline
with 0.05% Tween-20 for 20 minutes, washed, and incubated for 2 hours
with clone 38 moesin monoclonal antibody (MoAb; 100 ng/mL) or C31720
caspase-3 MoAb (250 ng/mL; both from Transduction Laboratories,
Lexington, KY), or with caspase-3 MoAb CPP32/p20-E8 (1 µg/mL; Santa
Cruz, Santa Cruz, CA) or B27D8 µ-calpain MoAb20 (1 to
10,000 dilution of ascites), or, after blocking with 2% normal goat
serum, with rabbit antibodies to caspase-3 (1 to 1,500 dilution;
Stratagene, La Jolla, CA). The membranes were washed, incubated with
125I-labeled secondary antibody, and exposed to
Phosphor-screen. Bands were quantified using Phosphor-Imager Storm 860 and Image Quant v1.1 program (Molecular Dynamics, Sunnyvale, CA). As a
control for the detection of active caspase-3, isolated mononuclear
cells were lysed at 1.5 × 107/mL in 0.5% NP-40, 10 mmol/L Tris HCl, pH 7.4, 150 mmol/L NaCl, 2 mmol/L DFP, 2 mmol/L EGTA,
and 50 µg/mL leupeptin and, after clarification by centrifugation,
the lysate was incubated for 1 to 3 hours with 25 µg/mL of granzyme
B21 (kindly provided by Dr Zhinan Xia, Center for Blood
Research, Boston, MA).
Activation of platelet caspase.
Platelets (5 × 108) in 1 mL of platelet buffer with 2 mmol/L CaCl2 in flat-bottom polyethylene vials were
incubated while stirring with 3 µmol/L A23187, 1 U/mL of human
thrombin, 10 µg/mL of collagen (native collagen fibrils from equine
tendons; Collagen Reagent Horn; Nycomed Arzneimittel GmbH, Munich,
Germany), or the combination of thrombin and collagen at 37°C for
20 minutes with stirring. The reaction was terminated by solubilizing
with SDS in preparation for immunoblotting.
Treatment with caspase and calpain inhibitors.
For inhibition experiments,
carbobenzoxy-Asp(OMe)-Glu(OMe)-Val-Asp(OMe)-fluoromethylketone
(DEVD-fmk), carbobenzoxy-Tyr-Val-Ala-Asp(OMe)-fluoromethyl ketone
(YVAD-fmk), carbobenzoxy-Phe-Ala-fluoromethylketone (FA-fmk), or
diluent was added to platelets during the final 60 minutes of
preincubation, and calpeptin and E64d were added during the final 10 minutes. For moesin cleavage experiments, fluoromethylketone stocks
were prepared in dimethylsulfoxide, the recommended solvent, or
dimethylformamide. For PS exposure experiments, we found that preincubation of platelets with dimethylsulfoxide (0.01% to 1.0%), but not with dimethylformamide, significantly increased A23187-induced exposure of PS (data not shown). Therefore, fluoromethylketone stocks
for PS exposure and all other experiments were prepared in
dimethylformamide (0.2% final concentration). Using this diluent, incubation of platelets with fluoromethylketones for 1 hour did not
alter resting platelet values of any of the parameters studied. Calpeptin and E64d stocks were prepared in ethanol or dimethylformamide.
Platelet activation and flow cytometry.
For PS exposure and microparticle release experiments, 107
platelets in 200 µL platelet buffer with 2 mmol/L CaCl2
were placed in siliconized 7 × 45 mm glass cuvettes at 37°C
in an aggregation meter (DP-247E; Sienco, Morrison, CO). A23187 (1 µmol/L), thrombin (human; 1 U/mL; Sigma), or thrombin (1 U/mL) plus
collagen (20 µg/mL) was added from a diluted stock and, after an
initial mixing, incubation was continued without stirring at 37°C
for 1 to 20 minutes. The platelet suspensions were transferred to an
approximately 22°C water bath and, after 1 minute, 50 µL was
combined with fluorescein isothiocyanate-labeled annexin V (annexin
V-FITC; 1 µg/mL; PharMingen, San Diego, CA) and phycoerythrin
(PE)-labeled anti-CD41 (GPIIb) MoAb (150 ng/mL; Coulter/Immunotech,
Miami, FL) and incubated for 10 minutes at approximately 22°C.
Samples were diluted fivefold with platelet buffer with 2 mmol/L
CaCl2 for immediate analysis by flow cytometry.
Samples were acquired and analyzed using a FACS-Calibur flow cytometer
and CellQuest software (Becton Dickinson, Mountain View, CA). Particles
were gated for CD41+ to distinguish platelets and
platelet-derived microparticles from electronic noise. The lower limit
of the platelet gate was defined on the forward scatter profile of
resting platelets, and CD41+ particles smaller than that
were considered microparticles.
To measure  granule secretion, 107 platelets were
activated as described above, and the reaction was stopped after 0, 30, or 60 seconds by adding an equal volume of 2% paraformaldehyde. Aliquots of the fixed platelets were incubated for 10 minutes with 1 µg/mL PE-labeled anti-CD62P (clone AC1.2 MoAb; Becton Dickinson, San
Jose, CA) and examined by flow cytometry.
To measure aggregation in response to thrombin or A23187,
107 platelets were incubated as described above with
stirring, and light transmission was measured as a function of time
according to the manufacturer's instruction.
Moesin degradation assay.
After preincubation of 5 × 108 platelets in 1 mL
platelet buffer in flat-bottom polyethylene vials, CaCl2 (2 mmol/L) and A23187 (3 µmol/L) were added and incubation was continued
for 10 or 20 minutes at 37°C with stirring. The reaction was
terminated by solubilizing with SDS.
Statistical analysis.
The Student's paired t-test was used to calculate P values.
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RESULTS |
Detection of caspase-3 in platelets.
To test for the presence of caspase, isolated platelets were lysed with
Triton X-100 and peptidase activity was measured by chromogenic assay
with the p-nitroanilid derivative of Asp-Glu-Val-Asp as
substrate (DEVD-pNA). Substrates based on DEVD are specific for
effector caspases, including caspase-3.7,8 DEVD-pNA
cleaving activity was detected at low levels in lysates of resting
platelets and was significantly increased on stimulation of platelets
with the Ca2+ ionophore A23187 (P < .01),
reaching maximal levels at 5 minutes (Fig
1A). In parallel assays with YVAD-pNA, a caspase-1 substrate, no activity was detected (data not shown).
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| Fig 1.
Detection of caspase-3 in resting and activated
platelets. (A) Caspase-3-like peptidase activity (cleavage of
DEVD-pNA) of Triton X-100 lysates of resting platelets (0 time
point) and platelets activated by A23187. Data shown are the mean ± SEM (n = 5). Peptidase activity was significantly increased relative
to resting platelets after 5, 10, and 20 minutes with A23187
(*P < .01). In the sample 20 min + DEVD, the inhibitor
DEVD-fmk was present during platelet activation and peptidase assay.
(B) Caspase-3 antigen detected by immunoblot. Resting platelets and
platelets activated with A23187 for 10, or 20 minutes were stained with
caspase-3-specific MoAb C31720. Molecular weight marker positions are
shown on the left and an arrow indicates 32-kD procaspase-3. The lanes
MNC (mononuclear cell lysates treated without and with granzyme B)
control for the ability to detect active caspase-3, the large subunit
of which (p17) is indicated by the arrow on the right. (C).
Procaspase-3 antigen quantified by immunoblots. Data shown are the mean ± SEM (n = 4). Platelet content of procaspase-3 antigen was
significantly decreased relative to resting platelets after 10 (*P < .02) and 20 minutes (*P <.003) with
A23187.
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Because caspase enzymes have overlapping substrate specificities,
platelets were also tested for caspase-3 antigen by immunoblotting. The
specific anti-caspase-3 MoAb C31720, which recognizes an epitope in
the large subunit, detected a 32-kD band in resting platelets, procaspase-3,22,23 the single-chain zymogen form of the
molecule (Fig 1B, left lane). On treatment with A23187, the platelet
content of procaspase-3 decreased significantly over time (Fig 1B and C; P < .02 at 10 minutes and P < .003 at 20 minutes). However, p17, the large subunit of active caspase-3, was not
detected. A positive control for the ability to detect p17 was provided by mononuclear cell lysates treated with granzyme B (Fig 1B, right lane). Similar platelet immunoblot results were obtained also with
caspase-3 antibodies from rabbit and with another MoAb (CPP32/p20-E8; data not shown). These findings demonstrate the presence of caspase-3 zymogen in resting platelets and its activation in A23187-treated platelets. The decrease of zymogen combined with the absence of sufficient active caspase-3 for immunoblot detection strongly suggests
that the active protease is short lived in platelets.
Physiological agonists activate platelet caspase-3.
Because A23187 is a potent but nonphysiological stimulus, we asked
whether processing of platelet caspase-3 zymogen is induced also by
physiological agonists. Thrombin, collagen, and the combination thrombin plus collagen were each found to induce processing of procaspase-3 (Table 1). The extent of
procaspase-3 processing varied; the order of agonist efficiency
was A23187 > thrombin + collagen > either collagen or
thrombin.
Caspase inhibitor abrogates agonist-induced phosphatidylserine
exposure.
We next examined whether caspase is involved in the movement of
negatively charged PS from the inner to the outer platelet membrane
leaflet, an activation reaction synonymous with generation of the
procoagulant surface. Platelets were treated with DEVD-fmk, a
cell-permeant inhibitor of caspase-3-like proteases, and then stimulated with agonist. Exposed PS was measured at fixed time points
by binding of annexin V-FITC.18 In response to the potent stimulant A23187, exposure of PS was rapid and extensive; 80% ± 4% (n = 4) of platelets became PS positive in 5 minutes (eg, Fig 2A). Incubation with DEVD-fmk
substantially inhibited/delayed A23187-induced PS exposure (Fig 2B). At
100 µmol/L, DEVD-fmk caused 74% ± 2% inhibition of PS exposure
at 3 minutes and 40% ± 3% at 5 minutes, and FA-fmk, a chemically
similar compound lacking caspase inhibitory activity, failed to inhibit
PS exposure (Fig 2B). Lower concentrations of DEVD-fmk were also
inhibitory, eg, 25 µmol/L inhibited A23187-induced PS exposure by
35% ± 5% at 3 minutes and 52% ± 2% at 5 minutes
(n = 3).
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| Fig 2.
Inhibition by the caspase inhibitor DEVD-fmk of PS
exposure in platelets activated by A23187 or thrombin plus collagen.
Platelets were preincubated for 1 hour with no inhibitor (A), or
FA-fmk, or DEVD-fmk (100 µmol/L in [B] and 25 µmol/L in [C])
and were treated for the indicated time with A23187 (1 µmol/L) or
thrombin (1 U/mL) plus collagen (20 µg/mL). (A) Platelets without
inhibitor treatment. Histograms of annexin V-FITC binding show
time-dependent A23187-induced conversion of platelets from PS-negative
to PS-positive. (B) Inhibition by DEVD-fmk of A23187-induced PS
exposure. The data are the mean percentages ± SEM (n = 4) of
annexin V-positive platelets as a function of time with A23187. Note
that pretreatment with DEVD-fmk, but not with FA-fmk, significantly
inhibited PS exposure (*P < .005 at 3 minutes and
P < .009 at 5 minutes). (C) Inhibition by DEVD-fmk
of thrombin plus collagen-induced PS exposure (n = 4). Pretreatment
with DEVD-fmk significantly inhibited PS exposure (*P < .001 at 5 minutes, P < .008 at 10 minutes, and P < .01 at 20 minutes). The unstimulated baseline values in (B) and (C) (dashed
lines) are not significantly different for platelets preincubated
without (shown) or with inhibitors.
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Platelets also expose PS when stimulated by thrombin plus collagen, in
which case the response is slower and less extensive; 25% ± 4% of
platelets became PS-positive by 20 minutes (Fig 2C, no inhibitor).
Pretreatment of platelets with DEVD-fmk, but not with FA-fmk (not
shown), significantly inhibited platelet PS exposure induced by
thrombin plus collagen (Fig 2C). DEVD-fmk at 25 µmol/L caused 85% ± 6% inhibition at 5 minutes, 45% ± 8% at 10 minutes, and
55% ± 10% inhibition at 20 minutes (Fig 2C). The finding of specific inhibition by DEVD-fmk of PS exposure in response to both
A23187 and to thrombin plus collagen strongly indicates that a
caspase-3-like enzyme is involved in agonist-induced translocation of
platelet PS to the outer membrane leaflet.
Caspase inhibitor abrogates agonist-induced microparticle release.
PS exposure in activated platelets is closely linked with the release
of microparticles (Zwaal and Schroit1 and Discussion). The
effect of DEVD-fmk on microparticle release was examined using a flow
cytometric assay to quantify microparticles (Materials and Methods).
Platelet pretreatment with DEVD-fmk, but not with FA-fmk, was found to
substantially inhibit/delay microparticle release in response to A23187
(Fig 3A). The extent of inhibition by 100 µmol/L DEVD-fmk was 42% ± 1% at 3 minutes and 32% ± 2% at
5 minutes (Fig 3A). Lower DEVD-fmk concentrations were also inhibitory;
25 µmol/L caused 25% ± 2% inhibition at 3 minutes and 8% ± 3% inhibition at 5 minutes (n = 3). For thrombin plus collagen-treated
platelets, microparticle release was also inhibited by DEVD-fmk.
DEVD-fmk at 25 µmol/L caused 57% ± 3% inhibition at 5 minutes,
47% ± 4% at 10 minutes, and 58% ± 7% at 20 minutes (Fig
3B); FA-fmk had no inhibitory effect (data not shown).
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| Fig 3.
Inhibition by DEVD-fmk, but not by FA-fmk, of
microparticle release from platelets stimulated by (A) A23187 or (B)
thrombin plus collagen. Platelets were preincubated as indicated for 1 hour with no inhibitor, FA-fmk, or DEVD-fmk (100 µmol/L in [A] and
25 µmol/L in [B]) and were then treated for the indicated time with
A23187 (1 µmol/L) or thrombin (1 U/mL) plus collagen (20 µg/mL).
Shown are the mean number of released microparticles (MP) ± SEM (n
= 4). The unstimulated values (dashed lines) are not significantly
different for platelets preincubated without (shown) and with
inhibitors. Microparticle release was significantly inhibited in
A23187-stimulated platelets at 3 (*P < .001) and 5 minutes
(*P < .01) and in thrombin plus collagen-stimulated platelets
at 5 (*P < .008), 10 (*P < .01), and 20 minutes
(*P < .01).
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Caspase inhibitor fails to prevent agonist-induced platelet
aggregation and secretion of granules.
Studies were performed to determine whether DEVD-fmk inhibits other
platelet activation responses, including the early responses of
aggregation and granule secretion. Aggregometer tracings showed
that pretreatment with DEVD (100 µmol/L) did not alter the time
course or amplitude of the aggregation response to A23187 (Fig 4A) or to thrombin (Fig 4B). To
monitor granule secretion, we measured surface expression of CD62P
(P-selectin), an granule membrane protein.24
Pretreatment with DEVD-fmk failed to alter thrombin-induced
upregulation of CD62P. Thirty seconds after thrombin addition, CD62P
expression, which was negative on resting platelets, was partially
upregulated in both DEVD fmk-pretreated and control platelets and was
maximally upregulated in both after 60 seconds (Fig 4C). DEVD also
failed to prevent agonist-induced platelet shape change, which was
detected in flow cytometry by the alteration of forward and side
scatter profiles (data not shown). These findings strongly suggest that
caspase-3 is not required for the early platelet activation responses
of granule secretion, shape change, and aggregation.
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| Fig 4.
Pretreatment of platelets with DEVD-fmk (100 µmol/L for
1 hour) does not alter agonist-induced platelet aggregation or
granule secretion. (A and B) Absence of effect of DEVD-fmk on platelet
aggregation induced by (A) A23187 (1 µmol/L) or (B) thrombin (1 U/mL). (C) Absence of effect of DEVD-fmk on thrombin-induced
granule secretion. The histograms show surface expression of the
granule membrane marker CD62P, which is absent on resting platelets
(left) and upregulated on treatment with thrombin for 30 seconds
(middle) or 60 seconds (right). Note that DEVD-fmk pretreatment (dashed
lines) did not alter thrombin-induced upregulation of CD62P.
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Caspase inhibitor abrogates agonist-induced cleavage of platelet
moesin.
The effect of DEVD-fmk was also examined on another late platelet
activation response, cleavage of the cytoskeletal linker protein
moesin. In the absence of inhibitor, A23187 induced cleavage of 60% ± 5% (n = 4) of moesin molecules in 20 minutes
(Fig 5A, lanes 1 and 2). Cleavage of moesin
was substantially inhibited in platelets pretreated with DEVD-fmk (lane
3). The extent of inhibition of A23187-induced moesin cleavage
increased over the DEVD-fmk range of 5, 25, and 50 µmol/L and was
complete at 50 and 100 µmol/L (Fig 5B). A23187-induced moesin
cleavage was not inhibited in platelets pretreated with FA-fmk and
minimally affected in platelets pretreated with YVAD-fmk, a related
inhibitor with primary specificity for caspase-1-like protease. These
findings strongly suggest that a caspase-3-like enzyme is required for the platelet activation event of moesin cleavage.
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| Fig 5.
Inhibition by DEVD-fmk of A23187-induced cleavage of
platelet moesin. (A) Platelets were preincubated for 1 hour without
inhibitor or with DEVD-fmk (50 µmol/L) and treated for 20 minutes
with A23187. The platelet suspensions were lysed by the addition of SDS
(with protease inhibitors). Shown is an immunoblot stained with clone
38 antimoesin MoAb. (B) Quantitation of moesin immunoblots. The methods
used are the same as in (A), except that platelets were preincubated
with varying concentrations of FA-fmk, YVAD-fmk, and DEVD-fmk. Shown
are the mean percentages of inhibition (±SEM, n = 4) of
moesin cleavage relative to platelets preincubated without inhibitor.
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Caspase activation and calpain activation are independent events.
Previous studies demonstrated an important role for another protease,
Ca2+-activated neutral protease (calpain), in a subset of
platelet activation events. Specific cell permeant reagents, including E64d and calpeptin, which prevent conversion of procalpain to calpain,
inhibit agonist-induced cleavage of select platelet cytoskeletal proteins, release of microparticles, and generation of prothrombinase activity.17,18 To determine whether DEVD-fmk alters
platelet functions by acting on calpain, the latter protein was
measured in DEVD-fmk-pretreated platelets. Reciprocal experiments were performed on calpeptin-pretreated platelets. Neither calpeptin nor
DEVD-fmk detectably altered the resting platelet content of µ-procalpain (Fig 6, top panel, first 3 lanes) or procaspase-3 (lower panel, first 3 lanes). Moreover, A23187
induced the conversion of procalpain to calpain in platelets
preincubated with DEVD-fmk, but not in calpeptin-pretreated platelets
(Fig 6, top panel, last 4 lanes). Similarly, A23187 induced the
processing of procaspase-3 in platelets preincubated with calpeptin,
but not in DEVD-fmk-pretreated platelets (lower panel, last 4 lanes).
These findings indicate that the alteration of platelet function by
DEVD-fmk does not rely on acting through calpain; similarly, the
effects of calpeptin do not entail acting through caspase-3.
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| Fig 6.
Effects of pretreatment with calpeptin or DEVD-fmk on the
content of µ-procalpain/µ-calpain (top panel) and procaspase-3
(lower panel) in resting and A23187-treated platelets. Platelets were
preincubated with no inhibitor or calpeptin (50 µg/mL) or DEVD-fmk
(25 µg/mL) and were lysed by the addition of SDS, immediately or
after stimulation with A23187 for 10 or 20 minutes. Shown are
immunoblots of 1.5 × 107 platelets stained with B27D8
anti-µ-calpain (top panel) or C31730 anti-caspase-3 (lower panel)
MoAb.
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Effects of calpain inhibitor and caspase inhibitor on PS exposure and
microparticle release.
To compare the roles of calpain and caspase, platelets were pretreated
with calpeptin and examined for agonist-induced PS exposure and
microparticle release. Similar to previous reports,17,18 calpeptin pretreatment significantly inhibited microparticle release in
response to thrombin plus collagen (Fig
7B). The extent of inhibition of microparticle release was similar for
calpeptin (69% ± 6% inhibition at 20 minutes; Fig 7B) and
DEVD-fmk (58% ± 7% inhibition at 20 minutes; Fig 3B), and the
inclusion of DEVD-fmk together with calpeptin did not further inhibit
residual microparticle release (Fig 7D).
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| Fig 7.
Effects of calpeptin and DEVD-fmk on PS exposure (A and
C) and microparticle release (B and D). Platelets were pretreated as
indicated with no inhibitor or calpeptin (50 µg/mL) or calpeptin plus
DEVD-fmk (25 µg/mL) and stimulated for the indicated times with
thrombin plus collagen (1 U/mL; 20 µg/mL). (A) PS exposure induced by
thrombin plus collagen was significantly increased by calpeptin at 5, 10, and 20 minutes (*P < .01, P < .001, and
P < .002, respectively; n = 4). (C) PS exposure induced by
agonist and enhanced by calpeptin was significantly inhibited by
DEVD-fmk at 5, 10, and 20 minutes (P < .01, P < .007, and P < .001, respectively). (B) Thrombin plus
collagen-induced microparticle release was significantly inhibited by
calpeptin at 5, 10, and 20 minutes (P < .02, P < .005, and P < .0003, respectively) and (D) was not further
inhibited by DEVD-fmk and calpeptin. Unstimulated values (dashed
lines), with the exception of PS exposure after calpeptin pretreatment
(shown in [A]), were not significantly different for platelets
preincubated without and with inhibitors.
|
|
On the other hand, calpeptin pretreatment did not inhibit PS exposure
in response to thrombin plus collagen; rather, PS exposure was
significantly increased (140% ± 8% increase at 20 minutes of
stimulation; Fig 7A). Calpeptin pretreatment also slightly increased
spontaneous PS exposure (Fig 7A, 0-minute value). Enhancement of
thrombin plus collagen-induced PS exposure was observed also when
platelets were pretreated with the chemically unrelated cell permeant
calpain inhibitor E64d (data not shown), and enhancement by E64d was
previously reported for A23187-induced PS exposure.18 Inclusion of DEVD-fmk together with calpeptin in the platelet pretreatment significantly and substantially inhibited PS exposure in
response to thrombin plus collagen, including abrogation of the
enhancing effect of calpeptin (Fig 7C). Together, these findings show
that caspase and calpain both function in specific late events of
platelet activation and that the two proteases have distinct functional roles.
| |
DISCUSSION |
Our studies with specific reagents demonstrate the presence of at least
one caspase family protease in platelets. Peptidase activity
corresponding to the effector protease caspase-3 was detected in
lysates of resting platelets and was significantly increased in
A23187-activated platelets. Specific antibodies identified the zymogen
procaspase-3 in resting platelets, and stimulation with agonist induced
processing of procaspase-3. Active platelet caspase-3 appears to be
short-lived, because it did not accumulate at levels adequate for
detection by immunoblot.
Processing/activation of platelet procaspase-3 was induced by several
characterized platelet agonists. The order of agonist effectiveness in
activating procaspase-3, A23187 > thrombin + collagen > either
thrombin or collagen (Table 1), is the same as their order of
effectiveness in inducing PS exposure,25,26 microparticle
release,25,26 and moesin cleavage.16a
Three agonist-induced platelet activation responses were found to be
significantly inhibited by the specific caspase inhibitor DEVD-fmk,
which has the greatest specificity for the effector proteases caspase-3
and caspase-7.7,8 DEVD-fmk inhibited A23187-induced
cleavage of the cytoskeletal protein moesin in a dose-dependent manner
and inhibition was complete at 50 µmol/L. DEVD-fmk inhibited exposure
of platelet PS and release of microparticles in response to A23187 and
to thrombin plus collagen. The extent of DEVD-fmk inhibition of PS
exposure was dose-dependent. The finding of inhibition of these three
platelet responses by DEVD-fmk strongly indicates that caspase-3, or a
caspase with specificity similar to caspase-3, is required for a subset
of late-occurring platelet activation events: moesin cleavage, PS
exposure, and microparticle release. In contrast, treatment with
DEVD-fmk had no effect on other (earlier) events of platelet
activation, including granule secretion, shape change, and
aggregation. Thus, the caspase-3 inhibitor is not a global inhibitor of
platelet responses, but rather is directed to a subset of events,
including those that are important for the generation of the
procoagulant platelet and microparticle surfaces.
Another protease, calpain, was previously shown to be involved in a
subset of platelet activation events, not including aggregation, shape
change, or secretion,27 and we, therefore, performed
parallel testing of calpain inhibitors and caspase inhibitors. These
experiments showed that the calpain inhibitor calpeptin and the
caspase-3 inhibitor DEVD-fmk are comparably effective in preventing
microparticle release in thrombin plus collagen-treated platelets. On
the other hand, calpeptin pretreatment substantially increases PS
exposure in response to thrombin plus collagen, whereas DEVD-fmk
inhibits agonist-induced PS exposure, including abrogating the increase effected by calpeptin.
Whereas DEVD-fmk completely inhibited agonist-induced moesin cleavage,
inhibition of PS exposure was only partial. Possibly, low levels of
caspase that escape inhibition may suffice to support partial PS
exposure, but not moesin cleavage. Alternatively, caspase may be
absolutely required for moesin cleavage, but platelets may have
caspase-dependent and caspase-independent pathways for activating PS
exposure. Support for the latter possibility is provided by the finding
that PS exposure in nucleated cells undergoing apoptosis is also
abrogated by inhibitors of caspase-3, including DEVD-fmk.28,29 However, in Jurkat T cells, PS exposure is
caspase-dependent when apoptosis is induced by Fas ligand, but
caspase-independent when the inducing agent is cytolytic
granules.30
For rapid exposure of PS, platelets rely on a
Ca2+-dependent mechanism, likely involving the recently
cloned lipid scramblase, a transmembrane protein with a
Ca2+ binding motif that rapidly moves phospholipids between
membrane leaflets.31,32 Other requirements for activation
of this mechanism are not known. Induction of platelet PS exposure and
microparticle release generally occur as linked processes. For example,
thrombin, a potent stimulator of aggregation, is a weak inducer of both PS exposure and microparticle release, which are best stimulated by
A23187 or thrombin + collagen or collagen in that order of effectiveness.25,26 In Scott syndrome, an inherited
bleeding disorder characterized by impaired lipid scramblase activity, ie, failure of agonist-induced PS exposure, patient platelets also fail
to release microparticles.33 By strictly regulating intracellular Ca2+ levels34 or through the use
of calpain inhibitors, PS exposure can be induced without microparticle
release (eg, Fig 7), but agonist-induced microparticle release has not
been observed in the absence of PS exposure. Such instances of apparent
linkage have led several investigators to hypothesize that PS exposure (loss of lipid asymmetry) is required for microparticle
release.1,35
Thus, of the agonist-induced activation events examined in this study,
aggregation, shape change, and granule secretion, previously found to
be unaffected by calpain inhibition,27 are also unaffected by caspase-3 inhibition (Table 2). PS
exposure was inhibited by caspase-3 inhibitor and not by calpain
inhibitor; rather, (agonist-induced) PS exposure was substantially
enhanced in calpeptin-pretreated platelets. In a recent study, another
platelet activation event, filopod extension in response to A23187, was
also enhanced in calpeptin-pretreated platelets.36 Finally,
moesin cleavage and microparticle release are prevented by either
calpain inhibition or caspase-3 inhibition (Table 2).
Together, these findings strongly indicate that caspase-3 functions in
agonist-induced activation of the PS exposing mechanism (Table 3), possibly by cleavage/activation
of a component of the PS exposing enzyme complex or by
cleavage/destruction of an inhibitory/regulatory protein. Based on the
findings that PS exposure is potentiated by calpeptin and E64d, we
hypothesize that calpain normally terminates the action of the PS
exposing system (Table 3). Calpain may act also to terminate filopod
extension.36 Whether caspase-3 acts directly at the level
of micro particle release cannot be concluded from the present data,
because inhibition of PS exposure by DEVD-fmk may suffice to prevent
microparticle release. Likewise, the apparent requirement for both
calpain and caspase-3 to cleave moesin requires further study. These
cumulative findings confirm a direct role for calpain in microparticle
release17,18 and demonstrate a distinct role for caspase-3
in activating PS exposure in response to agonist.
Although best known as effectors of apoptosis, caspases function also
in maturation of the cytokines, interleukin-1 by caspase 137,38 and interleukin-16 by caspase 3.39 Also,
caspase-3 activity was found in nonapoptotic mitogen-activated T
cells,40-42 and a caspase-3-like protease functions in
terminal differentiation of lens epithelial cell.43 The
present platelet findings represent an additional example of caspase
function in a nonapoptotic setting.
Recent studies have identified naturally occurring human caspase
inhibitors, the IAP (inhibitors-of-apoptosis) family of protease inhibitors44 and the related tumor cell protein
survivin.45 These discoveries suggest pharmacological
therapies to induce apoptosis, eg, of tumor cells, or to prevent
apoptosis, eg, in neurodegenerative disease. These efforts have
relevance to platelet function in coagulation and thrombosis in that
agents that inhibit caspases may be useful to prevent pathological
thrombotic events. From another perspective, it would be prudent that
clinical trials to alter apoptosis by targeting caspases be designed
bearing in mind the possible effect of the therapy on platelet function
in coagulation and thrombosis.
Altogether, these findings provide a basis for in-depth studies of the
molecular mechanisms of caspase family protease action in platelet
activation and programmed generation of procoagulant platelets.
| |
ACKNOWLEDGMENT |
The authors thank Drs Zhinan Xia and Judy Lieberman for providing
granzyme B, Dr Anthony Bretscher for advice, Dr Dianne Kenney for
advice and the use of equipment, and Drs John Hartwig and Andrey
Prodeus for critical reading of the manuscript. We are grateful to the
blood donors for their cooperation.
| |
FOOTNOTES |
Submitted September 9, 1998; accepted January 25, 1999.
Supported by National Institutes of Health Grant No. AI39574.
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 Eileen Remold-O'Donnell, PhD, The Center
for Blood Research, 800 Huntington Ave, Boston, MA, 02115; e-mail:
remold{at}cbr.med.harvard.edu.
| |
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TOP
ABSTRACT
INTRODUCTION