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 Previous Article | Table of Contents | Next Article 
Blood, Vol. 92 No. 9 (November 1), 1998:
pp. 3042-3049
RAPID COMMUNICATION
Phosphorylated Forms of Activated Caspases Are Present in Cytosol From
HL-60 Cells During Etoposide-Induced Apoptosis
By
Luis M. Martins,
Timothy J. Kottke,
Scott H. Kaufmann, and
William C. Earnshaw
From the Institute of Cell & Molecular Biology, University of
Edinburgh, Edinburgh, UK; and the Division of Oncology Research, Mayo
Clinic, Rochester, MN.
 |
ABSTRACT |
Treatment of HL-60 human leukemia cells with etoposide induces
apoptotic cell death and activation of at least 18 electrophoretically distinct cysteine-dependent aspartate-directed protease (caspase) isoforms, several of which differ only in their isoelectric points. The
purpose of the present study was to determine whether activated caspases are phosphorylated. Phosphatase treatment of cytosolic extracts containing active caspases followed by affinity labeling with
N-(N -benzyloxycarbonylglutamyl-N -biotinyllysyl)aspartic
acid [(2,6-dimethylbenzoyl)oxy] methyl ketone (Z-EK(bio)D-aomk)
showed a mobility shift in several of the labeled species, suggesting
that phosphorylated forms of these enzymes are present in the extracts.
Metabolic labeling with 32P followed by etoposide treatment
and subsequent affinity purification of affinity-labeled caspases
confirmed that at least three caspase species were
phosphorylated. To detect effects of the phosphorylation on enzymatic
activity, caspase-mediated cleavage of
aspartylglutamylvalinylaspartyl-7-amino-4-trifluoromethylcoumarin (DEVD-AFC) and poly(ADP-ribose) polymerase (PARP) by phosphorylated and
dephosphorylated extracts was measured. No significant changes in
Km or vmax were detected using DEVD-AFC. In
contrast, a slight, but significant enhancement of PARP cleavage was
observed in dephosphorylated extracts, suggesting that phosphorylation
of active caspases could have an inhibitory effect on enzyme activity.
These observations, which provide the first evidence that caspases are
phosphoproteins, suggest that caspases may be targets for some of the
growing list of protein kinases that are involved in apoptotic events.
© 1998 by The American Society of Hematology.
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INTRODUCTION |
OVER THE PAST 5 years it has become clear
that the initiation and execution of apoptotic cell death involve a
complex network of cysteine-dependent proteases termed
caspases.1-3 Caspases are expressed in healthy cells as
inactive zymogens. Early in apoptosis, autocatalytic
activation of certain procaspases triggers a protease cascade that
leads to activation of downstream caspases and other enzymes that
mediate apoptotic biochemical changes.4-6 How this protease
cascade is regulated is the subject of intensive study. Localization of
regulatory procaspases to certain subcellular compartments appears to
be one key aspect of this regulation.3 It is not currently
known whether posttranslational modifications such as protein
phosphorylation might also play a role in modulating the activity of
the mature caspases and/or regulating the caspase cascade.
Several lines of evidence have previously suggested that apoptosis can
be regulated by protein kinases and phosphatases.7-9 It has
been shown, for example, that Bruton's tyrosine kinase (BTK) is
required for the apoptotic response to ionizing radiation in chicken
DT40 lymphoma cells.10 Two other kinases whose activity is
upregulated during apoptosis, PCK 11 and
ASK1,12 have been implicated in activation of the Jun
N-terminal kinase/stress-activated protein kinase (JNK/SAPK)
pathway. This pathway, which is stimulated during Fas- and
ceramide-mediated apoptosis,13,14 as well as apoptosis that
occurs after growth factor withdrawal15 or anticancer drug
treatment,16,17 results in c-Jun-dependent transcriptional
activation, although it is clear that apoptosis can occur in the
absence of transcription as well.18,19 Recent studies have
also implicated the closely related Zip20 and
DAP21 kinases in cell death, although the exact role of
these kinases during apoptosis remains to be determined.
Examination of death receptor signaling pathways has provided even
stronger evidence linking kinases with the induction of apoptosis. The
serine/threonine kinase RIP22 appears to play an important
role in the initiation of tumor necrosis factor- (TNF-)-induced
cell death by mediating interactions between the type I TNF-
receptor (TNFR1)23 and an adapter protein RAIDD,24 thereby facilitating recruitment of procaspase-2
to the TNFR1 signaling complex. More recently,
RIP2/ CARDIAK,25,26 a RIP-related kinase that can bind to
the intracellular domains of Fas/CD95 and TNFR1,26 was
found to bind to and promote activation of procaspase-1 (but not
caspases  2, 4, 9) in vitro,25
providing the first example of a direct interaction between a kinase
and a procaspase. However, it is not clear whether RIP2/CARDIAK
phosphorylates procaspase-1, as the binding between the two molecules
requires the so-called caspase recruitment domain27 rather
than the kinase domain of RIP2.25 Forced overexpression of
RIP2 was shown to induce apoptosis,26 but it remains to be
determined whether procaspase-1 (or any other procaspase) is a
pertinent substrate when RIP2 kinase initiates apoptotic signaling.
In addition to acting as positive regulators of apoptosis, kinases can
also be anti-apoptotic. The Bcr/Abl kinase, for example, delays the onset of apoptosis in granulocytes from patients with chronic myelogenous leukemia,28 possibly by delaying
engagement of the apoptotic machinery.29-31 Likewise,
activation of a kinase cascade involving phosphatadylinositol-3 kinase
and Akt (protein kinase B) has been observed to inhibit apoptosis that
ordinarily occurs after withdrawal of interleukin-2 (IL-2), IL-3 or
integrin-mediated signals.32-34
The biochemical basis for the effects of many of these kinases on
apoptotic events is incompletely understood. Several members of the
Bcl-2 family of apoptotic regulators are apparently regulated at least
in part by phosphorylation.35 Serine phosphorylation of
Bcl-2 has been reported to be either anti-apoptotic36,37 or
proapoptotic,38 depending on the site that is
phosphorylated. For example, paclitaxel-induced phosphorylation of
Bcl-2 results in the disruption of Bcl-2/Bax interactions, the release
of Bax to the cytosol, and activation of the apoptotic
pathway.39 Similarly, signaling through the
phosphatidylinositol 3-kinase/Akt pathway leads to phosphorylation of
the proapoptotic Bcl-2 family member BAD, disrupting its interactions
with anti-apoptotic Bcl-2 family members and altering its ability to
induce apoptosis.40-43 However, these observations do not
rule out the possibility that other components of the apoptotic
machinery might also be regulated by phosphorylation.
Phosphatases have also been implicated in control of the apoptotic
pathway. Association of the tyrosine phosphatase FAP-1 with the
carboxyl terminal domain of the death receptor CD95/Fas has been
reported to inhibit Fas-mediated apoptosis,44 raising the
possibility that phosphorylation of Fas or some other component may be
required for activation of this pathway. Conversely, drug-induced apoptosis in human leukemia cells is accompanied by dephosphorylation of the retinoblastoma susceptibility protein (Rb).45,46 In at least one cell line, serine/threonine phosphatase inhibitors can
prevent both the drug-induced dephosphorylation of Rb and apoptosis,47 suggesting that dephosphorylation of Rb or
some other cellular component is required in this system for engagement of the apoptotic machinery.
Recent studies using affinity labeling and two-dimensional gel
electrophoresis to identify the caspase species activated during apoptosis showed that a number of labeled polypeptides differed only in
their isoelectric points.48,49 This observation raised the
possibility that caspases might be regulated by posttranslational modifications. In view of the growing evidence that kinases and phosphatases are involved in the regulation of apoptosis, the objective
of the present study was to determine whether active caspases are
phosphorylated in vivo and to perform an initial assessment of the
consequences of this modification on enzymatic activity.
| |
MATERIALS AND METHODS |
Materials.
Reagents were obtained from the following suppliers: sodium
orthovanadate, dialyzed fetal bovine serum (FBS), and d-biotin from
Sigma (St Louis, MO);  phosphatase from New England Biolabs (Beverly, MA); ortho[32P]phosphate from Amersham
(Arlington Heights, IL); okadaic acid and DEVD-fmk from
Calbiochem (Cambridge, MA); Nonidet P40 from BioRad (Richmond, CA);
sodium deoxycholate from BDH (Poole, UK); immunopure immobilized avidin
from Pierce (Rockford, IL); Sephadex G-25 medium and Immobiline
drystrips (pH 4-7L, 11 cm) from Pharmacia (Uppsala, Sweden); and other
reagents as previously specified.48
Cell culture.
HL-60 cells were cultured in RPMI 1640 medium containing 10% (vol/vol)
heat-inactivated FBS and gentamicin (50 µg/mL) at concentrations of
less than 106 cells/mL to insure logarithmic growth. For
metabolic labeling, cells (5 × 105 cells/mL) were
centrifuged and resuspended in phosphate-free RPMI 1640 medium
containing 10% (vol/vol) dialyzed FBS and 10 µCi/mL
ortho[32P]phosphate. Cells were labeled for 24 hours at
37°C before induction of apoptosis.
Induction of apoptosis and preparation of cytosolic extracts.
Etoposide was added to HL-60 cells at a concentration of 68 µmol/L, a
concentration shown to induce apoptotic cell death in previous
studies.29,48 After 5 hours, cytosol was prepared from
washed cells as previously described,48 with the exception that okadaic acid was included at a final concentration of 20 nmol/L.
Preparation of concentrated extracts from HL-60 cells.
After drug treatment, all steps were performed at 4°C (modified
from Lazebnik et al50). Cells were sedimented
at 300g for 5 minutes, and washed with incomplete MIG buffer
(50 mmol/L piperazine-N,N -bis[2-ethanesulfonic acid] [PIPES,
pH 7.0], 50 mmol/L KCl, 10 mmol/L EDTA, 10 mmol/L sodium
orthovanadate). Cells were washed again in complete MIG buffer,
consisting of incomplete buffer supplemented just before use with 20 nmol/L okadaic acid, 20 µmol/L cytochalasin B, 1 mmol/L dithiothreitol (DTT), 100 µmol/L -phenylmethylsulfonyl fluoride, and 1 µg/mL chymostatin, leupeptin, antipain, and pepstatin
A. The cell pellet was resuspended in 1/3 vol of complete
MIG buffer and transferred to a 2-mL microcentrifuge tube. Cells were
disrupted by sonication at 4°C for 30 seconds at 100 W using a
Misonix Model XL2010 sonicator equipped with a 1/8"
microtip probe (Branson Ultrasonics, Danbury, CT). The cell lysate was
centrifuged at 250,000gmax for 2 hours, yielding a
clear extract. Protein concentration of the extract, measured by the
Bradford assay,51 was estimated to be 40 mg/mL.
Phosphatase treatment of HL-60 cytosolic extracts.
Aliquots containing 300 µg of cytosolic protein were supplemented
with 8 U/mL of phosphatase in the presence or absence of 10 mmol/L
sodium orthovanadate and incubated at 30°C for 30 minutes. At the
end of the incubation, the reaction was terminated by addition of
sodium orthovanadate to a final concentration of 10 mmol/L to each
tube. Samples were frozen at 80°C until analyzed by affinity
labeling with Z-EK(bio)D-aomk followed by immunoblotting (see below) or
assayed for DEVD-AFC cleavage activity as previously described.29,48
Affinity labeling and inhibitor competition experiments.
Aliquots containing 5.2 mg of cytosolic protein from etoposide-treated
HL-60 cells were incubated for 5 minutes at 37°C with the
tetrapeptide inhibitor DEVD-fmk or the same volume of dilulent (DMSO).
Z-EK(bio)D-aomk was then added to a final concentration of 1 µmol/L and the incubation was continued for 15 minutes at 37°C.
Purification of caspases from radiolabeled extracts.
After radiolabeled extracts were reacted with Z-EK(bio)D-aomk,
unreacted affinity label was removed by passing the lysates through a
Sephadex G25 gel filtration column (1.5 mL of packed resin). The flow
through was diluted with an equal volume of denaturation buffer (0.5%
wt/vol sodium dodecyl sulfate [SDS], 20 mmol/L PIPES-KOH, pH 7.0, 20% [vol/vol]  -mercaptoethanol) and boiled for 5 minutes. Four
volumes of correction buffer (1% [vol/vol] Nonidet P40, 1% [wt/vol] sodium deoxycholate, 2 mmol/L EDTA, and 10 mmol/L PIPES-KOH, pH 7.0) was added. The corrected lysate (900 µL) was incubated with
60 µL of immunopure immobilized avidin for 1 hour with agitation at
4°C. The resin was washed three times with 1 mL of wash buffer (0.05% [wt/vol] SDS, 10 mmol/L PIPES, 4% -mercaptoethanol, 0.8% [vol/vol] Nonidet P40, 0.8% [wt/vol] sodium deoxycholate, and 1.6 mmol/L EDTA), and once with 1 mL of final wash buffer (10 mmol/L PIPES,
10% [vol/vol] -mercaptoethanol, 2 mmol/L EDTA). The bound
proteins were eluted by boiling the resin for 5 minutes in elution
buffer (50 mmol/L Tris-HCl [pH 8.6], 15% [wt/vol] sucrose, 2 mmol/L EDTA, 3% [wt/vol] SDS, and 2 mmol/L d-biotin).
Western blot analysis.
Samples containing purified radiolabeled caspases were subjected to
SDS-polyacrylamide gel electrophoresis (SDS-PAGE) on 16% (wt/vol)
acrylamide gels and transferred to nitrocellulose. Radioactivity was
quantitated using a Storm 840 Phosphorimager (Molecular Dynamics, Sunnyvale, CA), and digital data were processed using the
ImageQuant software package. The same nitrocellulose membrane was later
probed with peroxidase-labeled streptavidin and visualized by enhanced chemiluminescence.29,48
Alternatively, samples treated with phosphatase and labeled with
Z-EK(bio)D-aomk were precipitated using methanol/chloroform and
resuspended in 100 µL of urea solubilization buffer (8 mol/L urea,
4% [vol/vol] Nonidet P-40, 2% [vol/vol] -mercaptoethanol, 20%
[vol/vol] BioLyte 3-10 carrier ampholytes). Two-dimensional gel
electrophoresis was performed in a Multiphor II Unit using immobilized
pH gradients as previously described.52 Samples were
focused on an immobilized linear pH gradient isoelectric focusing gel
(110 mm, pH 4 to 7) for 30,500 volt-hours, transferred to
the top of a 16% (wt/vol) SDS-polyacrylamide gel, resolved at 15 mA
for 6 hours, transferred to nitrocellulose, and probed with
streptavidin as described above.
Analysis of poly(ADP-ribose) polymerase (PARP) cleavage activity.
Extracts treated with phosphatase were incubated with 125 ng of
purified bovine PARP at 37°C for various amounts of time. Proteins
were resolved by SDS-PAGE on 7.5% (wt/vol) acrylamide gels and
transferred to nitrocellulose. PARP was detected using C-II-10
monoclonal antibody and visualized by enhanced fluorescence (ECF).
Quantitation on the phosphorimager was performed as described above.
| |
RESULTS |
Detection of active caspases in extracts from apoptotic HL-60 cells.
Our previous studies showed that at least eight electrophoretically
distinct caspase species, including multiple isoforms of caspase-3 and
caspase-6, were detectable in cytosol prepared from apoptotic HL-60
cells.48 To more completely characterize the caspases
activated during HL-60 cell apoptosis, whole-cell extracts were
prepared from sonicated cells and reacted with Z-EK(bio)D-aomk, an
affinity labeling reagent that covalently labels the large subunits of
all caspases previously tested.48 After labeling, the
extracts were subjected to affinity chromatography on avidin agarose.
Polypeptides eluted from the column with free biotin were resolved by
two-dimensional gel electrophoresis using isoelectric focusing with an
immobilized pH gradient in the first dimension and SDS-PAGE in the
second. The region of the gel containing polypeptides in the 15- to
25-kD molecular-weight range (the known molecular weights
of the caspase large subunits53) is shown in
Fig 1. Using this methodology, 18 distinct
Z-EK(bio)D-aomk-reactive species were observed (Fig 1).
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| Fig 1.
High-resolution, high-sensitivity detection of active
caspases in apoptotic HL-60 cell extracts by two-dimensional gel
electrophoresis. (A) Mobilities of Z-EK(bio)D-aomk reactive caspases on
two-dimensional PAGE. Caspases were partly purified from extracts of
apoptotic HL-60 cells (see Materials and Methods), labeled with
Z-EK(bio)D-aomk, resolved by 2-dimensional PAGE using an immobilized pH
gradient, transferred to nitrocellulose membrane, reacted with
peroxidase-conjugated streptavidin, and detected by ECL. (B) Indexing
of active caspases purified from HL-60 cytosol. Shaded circles
correspond to species previously shown to comigrate with known
caspases. Spots A1, C1, C3 comigrate with Caspase-3, while spot B1
comigrates with Caspase 6.48 The migration of relevant
molecular-weight markers is shown at the left of (A) and (B). (C)
Isoelectric points of the detected caspases. (D) Control experiments
performed with ortho[32P]phosphate labeled extracts
showed the presence of Z-EK(bio)D-aomk reactive species in extracts
prepared from etoposide treated cells (lanes 2, 2), but not in
extracts from control healthy cells (lanes 1, 1).
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A number of observations indicate that these Z-EK(bio)D-aomk-labeled
species are active caspases: (1) Z-EK(bio)D-aomk efficiently labels
cloned caspases.48 (2) Extracts made by an identical method
from nonapoptotic control cells, which lack active
caspases,48 lack 15- to 25-kD polypeptides that react with
Z-EK(bio)D-aomk (Fig 1D). (3) Treatment with the irreversible caspase
inhibitors YVAD-cmk, DEVD-fmk, or Z-VAD-fmk inhibits labeling with
Z-EK(bio)D-aomk (eg, see Fig 3B).48,54 (4) Several of the
labeled species shown in Fig 1 comigrate in two-dimensional gels with
cloned human caspases expressed in Sf9 cells. Spots A1, C1, and C3
comigrate with caspase-3; and spot B1 comigrates with
caspase-6.48 (5) Direct sequencing of two-dimensional gel
spots obtained after a similar purification yielded sequences
corresponding to active caspase-3 and caspase-6.49 Collectively, these observations indicate that the
Z-EK(bio)D-aomk-reactive species detected in Fig 1A are active
caspases. Morover, the results of this experiment indicate that the
pattern of caspase activation in this cell line is substantially more
complex than previously suspected based on conventional two-dimensional
gel analysis of cytosol alone.
Effect of phosphatase treatment on the pattern of affinity labeled
caspases.
Several of the labeled caspase species detected in Fig 1 differed from
one another only in their isoelectric points. To investigate the
possibility that some of these species might represent phosphorylated forms, cytosol was prepared from etoposide-treated HL-60 cells in the
presence of the phosphatase inhibitor okadaic acid, reacted with
Z-EK(bio)D-aomk, and treated with phosphatase (8 U/mL, 30 minutes
at 30°C) in the presence or absence of sodium orthovanadate as a
phosphatase inhibitor. Cytosol was used for this analysis to decrease
the number of reactive species, thereby simplifying the analysis. In
the absence of inhibitor, phosphatase induced a substantial
alteration of the gel pattern (Fig 2),
suggesting that at least some of the species are phosphorylated forms
of the large subunits of active caspases. Quantitation of the active caspases detected by enhanced chemiluminescence (ECL)
showed that the five species whose mobility shifted upon phosphatase
treatment corresponded to approximately 30% of the covalently bound
affinity label or, to a first approximation because labeling is
stoichiometric, 30% of the active caspases detected by this protocol.
In additional experiments (not shown), similar results were obtained
when potato acid phosphatase was substituted for phosphatase.
Moreover, similar changes were observed when the order of treatment
with Z-EK(bio)D-aomk and phosphatase was reversed.
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| Fig 2.
Effect of phosphatase treatment on the two-dimensional
gel pattern of active caspases. Z-EK(bio)D-aomk treated cytosol from
etoposide-treated HL-60 cells was incubated with 400 U phosphatase
in the presence (A) or absence (B) of the inhibitor sodium
orthovanadate, then subjected to analysis by two-dimensional PAGE.
Black arrows point to caspases that disappear upon phosphatase
treatment. (C) Indexing of the active caspases present in (A). Filled
circles correspond to caspases that disappear upon phosphatase
treatment. (D) Bar chart illustrating the relative abundance of the
various species shown in (A) (average of three independent experiments,
with the standard deviation indicated). Also shown is the nomenclature
of the various active caspases detected in this experiment together
with their corresponding isoelectric points. Species indicated by an
asterisk (*) correspond to caspases that disappear upon
dephosphorylation.
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Purification and detection of active caspases in etoposide-treated
ortho[32P]phosphate-labeled HL-60 cells.
To confirm that the caspases whose mobility shifted after treatment of
extracts with phosphatase were in fact phosphorylated, HL-60 cells
were labeled in vivo with ortho[32P]phosphate and treated
with etoposide to induce apoptosis (Fig 3A). The active caspases present in extracts prepared from these metabolically labeled cells were derivatized with Z-EK(bio)D-aomk and
purified using avidin agarose. To distinguish purified
[32P]-labeled caspases from other labeled cellular
proteins that also bound to the avidin-agarose, control extracts were
pre-incubated with the covalent caspase inhibitor DEVD-fmk before
derivatization with Z-EK(bio)D-aomk. This had no effect on background
binding of extract proteins to the avidin-agarose, but completely
abolished the Z-EK(bio)D-aomk-dependent binding of
[32P]-labeled activated caspases (Fig 3B, lanes 2,
6). Purified [32P]-labeled proteins were resolved
by SDS-PAGE, transferred to nitrocellulose, and exposed for 7 days to a
phosphorimager plate. The same membrane was later incubated with
peroxidase-coupled streptavidin and visualized by enhanced
chemiluminescence. The mobility of proteins detected by ECL, previously
shown to correspond to active caspases,48,49,54 and the
labeled species detected using the phosphorimager were then compared
(Fig 3B). Quantitation of the phosphorimager counts corresponding to
the rectangular areas A5, A5, and A6 shown in Fig 3B enabled us
to construct the profiles shown in Fig 3C. This analysis showed that
phosphorylated species 5 and 8 comigrated on one-dimensional gels with
active caspases, previously designated IRP1 and IRP4.54
Furthermore, recovery of species 5 and 8 was significantly diminished
after pretreatment of extracts with DEVD-fmk, consistent with the
notion that both species are indeed active caspases. One additional
labeled species (no. 2) was also substantially decreased after DEVD-fmk pretreatment (see Fig 3C, merged). Although this species did not comigrate with one of the known biotinylated caspases labeled with
Z-EK(biotin)D-aomk, it may correspond to a minor caspase species that
is labeled to relatively high specific activity with 32P
under this protocol. When combined with the
two-dimensional analysis of phosphatase-treated caspases,
these data suggest that several active caspases are indeed
phosphorylated in HL-60 cells.
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| Fig 3.
Detection of 32P-labeled active caspases in
extracts from apoptotic HL-60 cells. (A) Experimental protocol. HL-60
cells labeled with ortho[32P]phosphate were exposed to
etoposide to induce apoptosis. Concentrated extracts from these cells
were reacted with Z-EK(bio)D-aomk (±pretreatment with DEVD-fmk) and
bound to monomeric avidin-agarose. The input, flow-through, and eluted
fraction (with 0.1% SDS) were resolved on a 16% SDS-polyacrylamide
gel and then analyzed (B) by autoradiography of the 32P
(lanes 1 through 6) or by ECL (lanes 1 through 6). Four
percent of the total extract was loaded for lanes 1, 2, 3, 4, 1,
2, 3, 4. Eighty percent of the purified protein
fraction was loaded for lanes 5, 6, 5, 6. Lanes 2, 4, 6, 2, 4, 6 correspond to cytosol preincubated with
DEVD-fmk before Z-EK(bio)D-aomk labeling. The rectangular areas, A5,
A5, A6 were subsequently subjected to quantitative analysis
using the phosphorimager. (C) Quantitation of the phosphorimager data
(counts per minute) of areas A5, A6, and densitometry (arbitrary units)
for area A5. The box designated "MERGED" corresponds to a
superimposition of A5 and A6, showing the relative position of the ECL
bands in A5 as black rectangles on the x-axis.
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Effects of dephosphorylation of active caspases on cleavage of
peptide and native protein substrates.
The results presented in Fig 2 show that the intensity of one
polypeptide previously shown to comigrate in two-dimensional gels with
caspase-3 (spot C1)48 increased upon phosphatase treatment of extracts. This suggests that a portion of the activated caspase-3 is
phosphorylated in HL-60 cells undergoing apoptosis. Analysis of the
three-dimensional structure of caspase-3, based on the reported atomic
coordinates,55 showed several putative phosphorylation sites, including a protein kinase C (PKC) consensus phosphorylation site, on the p20 subunit in close proximity to the substrate binding pocket. PKC and PKC have been reported to be cleaved and
activated during apoptosis in a caspase-dependent
manner.11,56,57 More recently, activation of PKC has
also been reported to occur before58 or after59
caspase activation in HL-60 cells. Although the precise role of PKC
isoforms during apoptosis clearly remains to be clarified, these
observations led us to ask whether activity of caspase-3-like enzymes
was altered in phosphatase-treated samples. A priori, we expected any
such effects to be small, because the great bulk of the active
caspase-3 in these extracts (spots A1 and C1) is apparently not
phosphorylated and shows no mobility shift following phosphatase
treatment (Fig 2). In fact, when caspase activity was measured using
the tetrapeptide substrate DEVD-AFC, phosphatase treatment failed to
alter either the vmax or Km ( 20 µmol/L)
for this substrate (Fig 4A). However,
recent experiments have suggested that tetrapeptide substrates might
not accurately reflect all aspects of caspase-mediated
cleavage.60 To investigate whether phosphorylation might
influence the interaction of active caspases with physiological
protein substrates, we compared the ability of control and
dephosphorylated extracts to cleave the canonical caspase substrate
PARP (Fig 4B). These experiments showed that full-length PARP is
cleaved more effectively in the dephosphorylated extracts than in
control extracts. Quantitation performed on results obtained from
three independent experiments indicated that PARP cleaving activity of
dephosphorylated extracts is approximately 60% higher than that of
control extracts (Fig 4B, bottom).
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| Fig 4.
DEVD-AFC and PARP cleavage activity assays using
dephosphorylated and control extracts. (A) Measurements using a
fluorogenic assay showed no differences between the ability of
dephosphorylated (open symbols) and control extracts (closed symbols)
to cleave the synthetic substrate DEVD-AFC. Inset, data replotted by
the method of Lineweaver and Burke. (B) Dephosphorylated
extracts exhibit enhanced ability to cleave PARP. (Top) PARP cleavage
was examined by enhanced chemofluorescence. Lanes 1 and 2 correspond to
PARP and extract incubated alone, respectively. (This extract contains
the cleaved PARP fragment.) Lanes 3 and 4, cleavage of PARP by
dephosphorylated extract. Lane 5, PARP was incubated in the presence of
control extract following pretreatment with DEVD-fmk. (There is no
significant decrease in intact PARP the fragment is from the extract
as in lane 2.) Lanes 6 and 7, cleavage of PARP by control extract.
(Bottom) Quantitation of the PARP cleavage results (average of three
independent experiments, with the standard deviations indicated).
Downward black bars correspond to the relative decrease (relative to
the starting level) in the amount of full-length PARP remaining after
incubation for 20 minutes in phosphatase-treated and control extracts.
Both the decrease in intact PARP and corresponding increase in the
cleaved fragment are enhanced in phosphatase-treated extracts.
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| |
DISCUSSION |
In the present study, we used two complementary techniques to evaluate
the possibility that caspases are phosphoproteins. Results of these
experiments not only provide the first evidence that some activated
caspase species contain covalently bound phosphate, but also provide an
indication that phosphorylation might inhibit the activity of caspase
species against certain substrates. These observations have potentially
important implications for current understanding of the manner in which
apoptosis is regulated.
Previous studies have shown that multiple caspase species differing
only in isoelectric point are detectable in the cytosol of apoptotic
leukemia cells.48,49 We have extended those studies by
demonstrating that phosphatase treatment of cytosol results in
disappearance of some species and increases in others (Fig 2). To
confirm this result using independent techniques, caspases were
affinity purified from 32P-labeled cells after induction of
apoptosis (Fig 3). At least two caspase species were found to contain
covalently bound 32P. Coupled with the previous reports of
charge heterogeneity among activated caspases,48,49 these
results provide strong evidence that activated caspases are
phosphoproteins.
Among the affinity-labeled caspase species that disappear from the
two-dimensional gels with phosphatase treatment are B1, which
comigrates with recombinant caspase-6, and four currently unidentified
species. These changes were accompanied by increased amounts of C1,
which comigrates with caspase-3. It is unlikely that any of the
unidentified species corresponds to either caspase-2 or caspase-1, the
only two caspases shown thus far to associate with protein kinases (the
former with RIP indirectly via the adapter RAIDD24 and the
latter directly with RIP2/CARDIAK25). Immunoblotting and
activity measurements previously failed to detect any significant maturation of procaspase-1 or -2 in HL-60 cells undergoing
etoposide-induced apoptosis.48
Despite the fact that only about 30% of the active caspase species
appear to be phosphorylated in these extracts (Fig 2), a reproducible
increase in PARP cleavage was observed after dephosphorylation of
cytosol (Fig 4B). This observation could mean that phosphorylation of
certain caspase species has a significant inhibitory effect on their
ability to cleave protein substrates. Because our detection of active
caspases depends on covalent labeling with the substrate analog
Z-EK(bio)D-aomk, we cannot at present rule out the possibility that
phosphorylated caspases are labeled less efficiently by this reagent.
If this were the case, then the data in Fig 2 might have underestimated
the level of caspase phosphorylation in apoptotic HL-60 cells. On the
other hand, although it is clear that the phosphatase acts directly
on certain caspases, as judged by the changes in their mobility in
two-dimensional gels, it is also difficult to rule out the possibility
that the enhanced PARP cleavage observed after phosphatase treatment is
due to dephosphorylation of another factor(s) that somehow modulates
PARP cleavage in these extracts. It will be important in the future to
map the phosphorylation sites on the caspases and identify the relevant
kinases. These experiments await the availability of high affinity
antibodies that can cleanly immunoprecipitate defined caspases from
apoptotic cell extracts.
In view of the evidence that phosphorylation plays an important role in
the apoptotic process (see Introduction), it is possible that
phosphorylation also affects other aspects of caspase function. For
example, phosphorylation has been shown to be an important factor
regulating the nuclear transport of certain
polypeptides.61-63 We have recently shown that C2 and C4,
two of the species whose mobility in two-dimensional gels is sensitive
to phosphatase treatment, are present in cytosol but absent from nuclei
of apoptotic HL-60 cells,29 raising the possibility that
phosphorylation might affect caspase targeting within apoptotic cells.
In MDA-MB-468 breast carcinoma cells, on the other hand, three of the
caspase species shown here to be phosphorylated (B1, C4, and C5) were found in the nuclei (T.J.K. and L.M.M., unpublished observations, January 1998). Thus, if phosphorylation is involved in
the regulation of caspase nuclear import, the phenomenon might vary
depending on the cell type.
It is also possible that phosphorylation is important in regulating
procaspases. Although the methods used in the present study detect only
the active caspases, preliminary experiments have shown the existence
of multiple charge isoforms of procaspase-2 and procaspase-3 in
untreated cells (T.J.K. and S.H.K., unpublished observations, May
1998). These observations raise the possibility that
changes in phosphorylation state might be involved in caspase activation and/or caspase interactions with regulatory
molecules.
Although the role(s) of caspase phosphorylation remain to be more fully
elucidated, the present report opens the way to the investigation of a
new level of caspase regulation in vivo. It will be important in future
experiments to identify both the phosphorylated species and the
relevant kinases in order to begin to understand the role of caspase
phosphorylation in the apoptotic response.
| |
FOOTNOTES |
Submitted June 17, 1998;
accepted August 12, 1998.
Supported by a Principal Research Fellowship from the Wellcome Trust
(W.C.E.) and Public Health Service Grant No. CA69008 (S.H.K. and
W.C.E.). L.M.M. was supported by a studentship from the Programa
Gulbenkian de Doutoramento em Biologia e Medicina. S.H.K. was a Scholar
of the Leukemia Society of America. W.C.E. is a Principal Research
Fellow of the Wellcome Trust.
Address reprint requests to William C. Earnshaw, PhD, Institute of Cell
& Molecular Biology, University of Edinburgh, Edinburgh, EH9 3JR,
Scotland, UK; e-mail: bill.earnshaw{at}ed.ac.uk.
The publication costs of this article were defrayed in part by page charge payment. This article must therefore be hereby marked "advertisement" is accordance with 18 U.S.C. section 1734 solely to indicate this fact.
| |
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Abstract
Introduction
Abstract