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(Received for publication, June 20, 1997, and in revised form, August 4, 1997)
From The Program in Apoptosis and Cell Death, The Burnham
Institute, La Jolla, California 92037
ABSTRACT The observation that the nematode cell death
effector gene product Ced-3 is homologous to human
interleukin-1 INTRODUCTION Apoptotic cell death is a process that enables metazoans to
eliminate cells that are damaged, mislocated, or have become
superfluous, and is characterized by controlled proteolysis of cellular
components resulting from activation of an in-built program (2, 3). The
signal for the execution of the cell may come from various stimuli:
specific death receptor ligation (4), ionizing radiation (5),
anti-neoplastic drugs (6), and growth factor withdrawal (7). However,
despite the variety of death signals, the key features of execution
appear to be quite similar; the death signal converges upon the
activation of a number of proteases, which in turn cleave protein
substrates (8, 9), thus giving rise to characteristic apoptotic
morphology.
Since the discovery that ced-3, a key effector gene of
programmed cell death in Caenorhabditis elegans, exhibited
homology with interleukin 1 EXPERIMENTAL PROCEDURES Active caspases-3, -6, -7, and -8 were expressed
in Escherichia coli and isolated as described previously
(22, 24, 27). The expression constructs for caspases-3, -6, and -7 contained a His6 tag at the C terminus of the full-length
protein, while caspase-8 was constructed to have a His6 tag
at the N terminus replacing residues 1-216 of the zymogen. The
concentrations of the purified enzymes were determined from the
absorbance at 280 nm based on the molar absorption coefficients for the
caspases calculated from the Edelhoch relationship (28): caspase-3
( The pH
dependence of the hydrolysis of the substrate Z-DEVD-AFC were evaluated
in the pH range 5.5-10. The enzymatic reaction was carried out at
37 °C in the following buffers: 20 mM MES (pH 5.5-6.5),
20 mM HEPES (pH 6.2-7.3), 20 mM PIPES (pH
6.9-8.1), 20 mM Bicine (pH 7.8-9.0) or 20 mM
CHES (pH 8.8-10.0), containing NaCl, DTT (fresh), EDTA, CHAPS, and
sucrose at optimized concentrations as described under "Results and
Discussion." For reasons discussed later, the optimal buffer used as
a basis for further studies was 20 mM PIPES, 100 mM NaCl, 10 mM DTT, 1 mM EDTA,
0.1% CHAPS, 10% sucrose, pH 7.2. The enzyme concentrations used were
1.2 nM (caspase-3), 18 nM (caspase-6), 5 nM (caspase-7), and 80 nM (caspase-8). The
initial rates of enzymatic hydrolysis were measured by release of AFC
from the substrate Z-DEVD-AFC (0.1 mM) as emission at 505 nm upon excitation at 400 nm using a Perkin-Elmer LS50B fluorimeter equipped with a thermostated plate reader. The pH dependences of the
initial rates of hydrolysis for all four caspases were fitted to a bell
shape described for two ionizing groups by the equation
v = (limit × log(pH The sensitivity toward
Zn2+ and Ca2+ was determined in optimal buffer
(without EDTA), containing varying concentrations of ZnCl2 or CaCl2 as described above. A concentration of 20 mM The sensitivity toward ionic strength was determined in
optimal buffer containing varying concentrations of NaCl as described above.
The
stability of the four caspases was determined by incubating the enzymes
in optimal buffer at 0 °C or 37 °C. At various time points, a
sample was withdrawn and the activity was determined as described
above.
RESULTS AND DISCUSSION Heterologous expression of the caspases is
required to obtain sufficient amounts of starting material for a
rigorous characterization. Although the mechanism is not understood,
when expressed in E. coli, these caspases spontaneously
undergo what appears to be autoprocessing to yield the appropriate
subunits characteristic of the active enzymes (Fig.
1). Processing at interdomain Asp residues was confirmed for all of the recombinant proteases by sequencing of the N termini of the two subunits (see Refs. 22 and 27
for further details). Note that both the large and small subunits
migrate as homogeneous bands in SDS-PAGE, with the exception of the
large subunit of caspase-6, which, based on N-terminal sequencing,
represents alternative cleavages at the C terminus of the large
subunit. The N-terminal peptides of caspases-3, -6, and -7 were also
removed during processing, as demonstrated to occur during Fas-mediated
apoptosis in vivo (22). Caspase-8 could not be expressed as
a full-length protein and thus was engineered with a 21-residue linker
(24, 27) that replaces the 216-residue N-terminal segment removed
during its activation in vivo after Fas ligation (25, 26).
Consequently, with the exception of the terminal purification tags, the
proteins are essentially identical to the forms identified or expected
in vivo in apoptotic cells. On the basis of titration with
the active site-directed caspase inhibitor Z-DEVD-fluoromethyl ketone,
all proteases were 100% active, based on absorbance at 280 nm, with
the exception of caspase-8, which was 50% active.
Common to all the caspases
is a distinct preference for aspartic acid in the P1
position.2 In the present
study, we have used Z-DEVD-AFC as the test substrate even though the
four caspases investigated exhibit some degree of P4
preference. Use of this substrate is suitable, inasmuch as we are
interested in the properties of the caspases and not in the possible
effects that the change of various parameters may have on the protein
substrates.
Initially, the requirements for various components were investigated in
a buffer based on that used by Thornberry et al. for caspase-1 (11): 20 mM HEPES, 100 mM NaCl, 10 mM DTT, 0.1% CHAPS, 10% sucrose, pH 7.4. Table
I demonstrates the effects of removal of
some of these component from the assay buffer. All four caspases lose
over 40% of their activity upon removal of CHAPS from the buffer, and
the effect is more dramatic with caspase-6 than with caspases-3, -7, and -8. Only minor beneficial effects are found with sucrose and NaCl;
in the case of caspase-6, there is a significant reduction in activity
in the presence of NaCl, which will be discussed in detail below.
However, 100 mM NaCl is required in the assay buffer to
maintain a consistent ionic strength when varying pH. A relatively high
concentration of DTT (10 mM) is required for full activity
of the recombinant enzymes. They may be preactivated by DTT and
the DTT removed by gel filtration; however, if neither reducing agent
nor EDTA is present in the exchange buffer, the activity declines
rapidly, presumably due to oxidation of the catalytic cysteine (data
not shown). EDTA (1 mM) is incorporated into the assay
buffer to avoid inactivation by trace metals.
Table I.
Effect of removing various compounds from the assay buffer
Volume 272, Number 41,
Issue of October 10, 1997
pp. 25719-25723
©1997 by The American Society for Biochemistry and Molecular Biology, Inc.
and
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS AND DISCUSSION
FOOTNOTES
ACKNOWLEDGEMENTS
REFERENCES
-converting enzyme (caspase-1) has led to the discovery
of at least nine other human caspases, many of which are implicated as
mediators of apoptosis. Significant interest has been given to aspects
of the cell biology and substrate specificity of this family of
proteases; however, quantitative descriptions of their biochemical
characteristics have lagged behind. We describe the influence of a
number of environmental parameters, including pH, ionic strength,
detergent, and specific ion concentrations, on the activity and
stability of four caspases involved in death receptor-mediated
apoptosis. Based on these observations, we recommend the following
buffer as optimal for investigation of their characteristics in
vitro: 20 mM
piperazine-N,N
-bis(2-ethanesulfonic acid) (PIPES),
100 mM NaCl, 10 mM dithiothreitol, 1 mM EDTA, 0.1% 3-[(3-cholamidopropyl)dimethylammonio]-2-hydroxy-1-propanesulfonic acid (CHAPS), 10% sucrose, pH 7.2. Caspase activity is not affected by
concentrations of Ca2+ below 100 mM, but is
abolished by Zn2+ in the submicromolar range, a common
characteristic of cysteine proteases. Optimal pH values vary from 6.8 for caspase-8 to 7.4 for caspase-3, and activity of all is relatively
stable between 0 and 150 mM NaCl. Consequently, changes in
the physiologic pH and ionic strength would not significantly alter the
activity of the enzymes, inasmuch as all four caspases are optimally
active within the range of these parameters found in the cytosol of
living and dying human cells.
-converting enzyme
(ICE1 or caspase-1), the
involvement of proteolytic enzymes in apoptosis has been an issue of
significant interest (10-12). This has resulted in the cloning of
several mammalian genes encoding ICE/Ced-3 homologues, known commonly
as caspases (13), several of which are important for promotion of the
death pathway in mammals (reviewed in Ref. 9). However, with the
notable exception of caspase-1 (14-16), little attention has been
given to the key biochemical properties of these enzymes, which is
important for understanding the effect of the intracellular environment
on their activity. For example, changes in pH, redox potential, and
Zn2+ concentration all have effects on apoptosis (17-21).
In the present article, we present a characterization of some of the
basic biochemical properties of four of the caspases. We have chosen to
focus on those that play a central role in the apoptotic pathway
initiated by ligation of the death receptors Fas and tumor necrosis
receptor 1: caspase-3 (Yama/CPP32/apopain), caspase-6 (Mch2), caspase-7 (Lap3/Mch3/CMH1), and caspase-8 (FLICE/MACH) (22-26).
280 = 26000 M
1
cm1), caspase-6 (280 = 26000 M1 cm1), caspase-7
(280 = 24510 M1
cm1), and caspase-8 (280 = 27390 M1 cm1).
Carbobenzoxy-Asp-Glu-Val-Asp-7-amino-4-trifluoromethyl coumarin (Z-DEVD-AFC) was purchased from Enzyme System Products. DTT was from
Diagnostic Chemicals Limited. Sucrose was from Mallinckrodt. All other
chemicals were from Sigma. Z-DEVD-fluoromethyl ketone was the kind
gift of Joe Krebbs, IDUN Pharmaceuticals.
pKa1))/(log(2 × pH pKa1 pKa2) + log(pH pKa1) + 1)) using Grafit 3.01 (29).
-mercaptoethanol was used to replace DTT because it
does not chelate zinc to the same extent as DTT. The influence of the
concentration of the reductant on the zinc sensitivity was exploited
for caspase-3 using varying concentrations of -mercaptoethanol. The
inhibition of the caspases by ZnCl2 was fitted to an
equation describing simple competitive inhibition, v = Vmax/(1 + ((Ks/[S]) × (1 + ([Zn2+]/KZn)))) using Grafit 3.01 (29).
Fig. 1.
SDS-PAGE analysis of the purified recombinant
caspases. Approximately 5 µg of each caspase was electrophoresed
in a linear 5-15% SDS-PAGE gel (40), followed by staining with
Coomassie Blue. The gel demonstrates the presence of the large and
small subunits characteristic of the activated enzymes, as well as the purity of the preparations used in this study.
[View Larger Version of this Image (62K GIF file)]
Caspase-3
Caspase-6
Caspase-7
Caspase-8
CHAPS
0.59
0.03
0.39
0.30
Sucrose
0.95
0.82
0.81
0.92
NaCl
0.97
1.69
0.78
0.97
Only minor differences were observed in the pH
profiles of the four caspases. The bell-shaped pH dependence signifies
the existence of one active form of the enzyme with the increase in activity most likely due to the de-protonation of the catalytic Cys
residue. In this respect, the caspases closely resemble other unrelated
cysteine proteases in their activity pH profiles (30). Caspase-3 was
found to be active over a broader pH range with an optimum slightly
higher than the other three (see Fig. 2). Although we have analyzed the pH dependence of all four enzymes as a
simple bell-shaped curve, there is a faster than expected drop-off in
activity at low pH, most clearly observed with caspases-3 and -6. This
indicates that more than one group is protonating, possibly another
group on the enzyme, or the substrate carboxylate(s), inasmuch as the
three-dimensional structure of caspases-1 and -3 demonstrates binding
of unprotonated side-chains in its specificity pockets (31-33). The pH
dependence of these caspases also indicates that they all are fully
active within the pH range found in normal as well as apoptotic cells,
designated by the shaded background in Fig. 2 (18, 21). It
is possible that changes in pH during apoptosis may affect caspase
activity indirectly by altering the structure of a particular set of
natural substrates. However, this hypothetical event would change only
the susceptibility of the substrate, not the activity of the
caspases.
Fig. 2. The pH dependence of the four caspases for the hydrolysis of the synthetic peptide substrate Z-DEVD-AFC. The dependence was fitted to a bell shape characterized by the following pKa values: pKa1 = 6.4 and pKa2 = 8.6 for caspase-3, pKa1 = 6.9 and pKa2 = 7.2 for caspase-6, pKa1 = 6.5 and pKa2 = 7.7 for caspase-7, and pKa1 = 6.0 and pKa2 = 7.7 for caspase-8. The shaded area illustrates the pH range found in normal and apoptotic cells, with the latter favoring lower pH (18, 21).
[View Larger Version of this Image (26K GIF file)]
Effects of Ionic Strength
We used NaCl in the range 0-1
M in assay buffer to address the dependence of ionic
strength on caspase activity. Differential effects were found depending
on the enzyme, with caspases-3 and -8 having fairly flat profiles,
whereas caspases-6 and -7 demonstrated maximal activity at 0.03 M and 0.25 M (Fig.
3). Although the activity of caspase-6
declined faster than the others as ionic strength increased, none of
the enzymes demonstrated substantial adverse effects on their activity
in the physiologic range of ionic strength (34), designated by the
shaded background in Fig. 3. The apparent stability to
substantial changes in ionic strength indicates that this would not be
limiting during commitment to apoptosis.
Fig. 3. NaCl dependence of caspases. Caspases-3 (
), -6 (
), -7 (
), and -8 (
) were incubated under optimal
buffer conditions, with the indicated concentration of NaCl, and
initial rates of substrate hydrolysis determined. The rates of
hydrolysis have been normalized to the rate of hydrolysis in the
absence of NaCl. The shaded area illustrates the range of
ionic strength normally found in the cytosol (34).
[View Larger Version of this Image (27K GIF file)]
Effects of Zn2+and Ca2+
Several
studies have reported that Zn2+ inhibits apoptosis.
Originally, this effect was believed to be due to the inhibition of
nucleases; however, caspase-6 (17) and, more recently, caspase-3 (20)
have been found to be inhibited completely by 2 mM
Zn2+. The influence of transition metal ions on the
activity of cysteine proteases has been well established for a long
time; for instance, members of the papain family are sensitive to
Zn2+, mercury, and various organomecurials (35, 36).
Because DTT chelates Zn2+, we compared caspases for
sensitivity to this ion in the presence of 20 mM
-mercaptoethanol, which we determined to be the concentration of
this reductant required for optimal activity of the recombinant enzymes
(data not shown). Due to the inherent tendency of Zn2+ to
react with thiols, we can only obtain an apparent binding constant and,
under these conditions, all the caspases are inhibited by small amounts
of Zn2+, although there are significant differences in the
affinity (Fig. 4). Caspase-6 is most
readily inhibited by Zn2+, completely inactivated by 0.1 mM, and caspase-3 is the least sensitive, requiring more
than 1 mM for complete inactivation. To estimate the real
binding affinity, we probed the influence of reductant on the
inhibition of caspase-3 by Zn2+. Not surprisingly, there
was a significant influence of the concentration of -mercaptoethanol
on the KZn,app giving rise to values converging on an approximate value of 0.15 µM (Fig.
5). From these results, it is quite
evident that Zn2+ is a good inhibitor of the caspases,
albeit very dependent on the thiol content, and therefore presumably
the redox potential of the cell. The influence of Ca2+ was
investigated in a similar manner and was found to have no effect on the
activity of any of the caspases at concentrations up to 100 mM (data not shown). Thus, the reported role of
Ca2+ in apoptosis (see, for example, Ref. 37) is unlikely
to be due to any effect on the caspases.
Fig. 4. Sensitivity of the four caspases to the presence of Zn2+. Caspases were incubated under optimal buffer conditions, with DTT replaced by
-mercaptoethanol, at
the indicated concentration of Zn2+, and initial rates of
substrate hydrolysis determined. The apparent binding constants for
Zn2+ to the individual caspases
(KZn,app) are 8.8 µM for
caspase-3, 0.3 µM for caspase-6, 1.7 µM for
caspase-7, and 1.9 µM for caspase-8.
[View Larger Version of this Image (23K GIF file)]
Fig. 5. Influence of the concentration of
-mercaptoethanol on the apparent binding constant for
Zn2+ to caspase-3. The influence of
-mercaptoethanol on KZn,app were investigated
using the concentrations 0.25 mM (
), 0.5 mM (
), 1 mM (
), 2 mM (
), 4 mM
(), 8 mM (), 16 mM (), and 32 mM ().
[View Larger Version of this Image (25K GIF file)]
In Vitro Stability
On the basis of the foregoing results, the
optimal general caspase buffer was designated as 20 mM
PIPES, 100 mM NaCl, 10 mM DTT, 1 mM
EDTA, 0.1% CHAPS, 10% sucrose, pH 7.2. The stability of the four
caspases was tested by incubating the enzymes at 0 °C or 37 °C in
the optimal assay buffer and determining the activity at various times
(Fig. 6). None of the caspases showed any
decrease in activity at 0 °C over the 150-min period. Caspases-3 and
-6 retained full activity for 150 min at 37 °C, whereas caspases-7 and -8 showed an appreciable decrease in activity. To verify that the
decrease in activity observed at 37 °C with caspases-7 and -8 was
not due to sample variation, the experiment was performed with two
different preparations of these enzymes giving rise to almost identical
results, reducing the probability that the decrease in activity is
associated with sample variations. The reason for the decrease in
activity is not clear; however, SDS-PAGE analysis of caspases incubated
at 0 °C and 37 °C for 150 min does not reveal any indications of
degradation (data not shown). Based on these observations, the most
probable explanation is a conformational change, possibly due to slow
dissociation of the subunits after dilution into assay buffer, as
originally described for caspase-1 (11). This assumption is supported
because the decrease in activity observed with caspase-8 appears to
approach a level of approximately 60%, and remains there for an
extended period of time. Whether such dissociation occurs in a cell
under physiologic conditions remains an open question, but it is
evident that none of the investigated caspases undergo autolysis that
will significantly affect their role in apoptosis. This is in contrast
to caspase-1, which has been shown to inactivate spontaneously by
autolytic degradation of its small subunit (38).
Fig. 6. In vitro stability of the caspases. The stability of the caspases in optimal buffer at 0 °C (
) and 37 °C () were determined by measuring the
residual activity as a function of time.
[View Larger Version of this Image (21K GIF file)]
Biologic Perspective
The results demonstrate that all four
caspases are optimally active under normal physiologic conditions. We
have to activate the recombinant enzymes by adding thiols, presumably
because of reversible modification of the catalytic cysteine during
expression and purification. In vivo, however, the
glutathione balance would favor the reduced form, with the result that,
once processed from their single chain zymogens, the caspases would be
fully active. We do not rule out the possibility that natural caspase
substrates are affected by changes in environmental parameters that
would alter their susceptibility to specific proteolysis in
vivo. In this context, caspase-1 was shown to exhibit a marked
salt dependence due to effects of NaCl on the substrate
pro-interleukin-1, but not on a synthetic peptidyl substrate (39).
However, changes in the pH and ionic strength of the cytosol would not
significantly alter the activity of the enzymes themselves, inasmuch as
all four caspases are optimally active within the range of these
parameters found within cell cytosols, irrespective of their metabolic
status.
FOOTNOTES
Supported by Danish Natural Science Foundation Grant 9600412.
§ To whom correspondence should be addressed: The Burnham Institute, 10901 N. Torrey Pines Rd., La Jolla, CA 92037. Tel.: 619-646-3114; Fax: 619-646-3189; E-mail: gsalvesen{at}ljcrf.edu.
1 The abbreviations used are: ICE, interleukin-1
-converting enzyme; Z, carbobenzoxy; DTT,
dithiothreitol; MES, 2-(N-morpholino)ethanesulfonic acid;
CHES, 2-(N-cyclohexylamino)ethanesulfonic acid; PIPES,
piperazine-N,N-bis(2-ethanesulfonic acid); CHAPS,
3-[(3-cholamidopropyl)dimethylammonio]-2-hydroxy-1-propanesulfonic acid; Bicine, N,N-bis(2-hydroxyethyl)glycine; AFC,
7-amino-4-trifluoromethyl coumarin; PAGE, polyacrylamide gel
electrophoresis.
2 Binding site nomenclature is in accordance with the nomenclature of Schechter and Berger (1).
ACKNOWLEDGEMENTS
We thank Yuri Lazebnik for helpful discussion of this manuscript, Qiao Zhou for performing the active site titration of the caspases, and Annamarie Price and Scott Snipas for technical assistance. We thank Joe Krebbs for providing us with Z-DEVD-fluoromethyl ketone.
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P. W. Mesner Jr., K. C. Bible, L. M. Martins, T. J. Kottke, S. M. Srinivasula, P. A. Svingen, T. J. Chilcote, G. S. Basi, J. S. Tung, S. Krajewski, et al. Characterization of Caspase Processing and Activation in HL-60 Cell Cytosol Under Cell-free Conditions. NUCLEOTIDE REQUIREMENT AND INHIBITOR PROFILE J. Biol. Chem., August 6, 1999; 274(32): 22635 - 22645. [Abstract] [Full Text] [PDF]
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B. Zech, M. Wilm, R. van Eldik, and B. Brune Mass Spectrometric Analysis of Nitric Oxide-modified Caspase-3 J. Biol. Chem., July 23, 1999; 274(30): 20931 - 20936. [Abstract] [Full Text] [PDF]
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S. Krajewski, M. Krajewska, L. M. Ellerby, K. Welsh, Z. Xie, Q. L. Deveraux, G. S. Salvesen, D. E. Bredesen, R. E. Rosenthal, G. Fiskum, et al. Release of caspase-9 from mitochondria during neuronal apoptosis and cerebral ischemia PNAS, May 11, 1999; 96(10): 5752 - 5757. [Abstract] [Full Text] [PDF]
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H. R. Stennicke, Q. L. Deveraux, E. W. Humke, J. C. Reed, V. M. Dixit, and G. S. Salvesen Caspase-9 Can Be Activated without Proteolytic Processing J. Biol. Chem., March 26, 1999; 274(13): 8359 - 8362. [Abstract] [Full Text] [PDF]
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A. Weidemann, K. Paliga, U. Durrwang, F. B. M. Reinhard, O. Schuckert, G. Evin, and C. L. Masters Proteolytic Processing of the Alzheimer's Disease Amyloid Precursor Protein within Its Cytoplasmic Domain by Caspase-like Proteases J. Biol. Chem., February 26, 1999; 274(9): 5823 - 5829. [Abstract] [Full Text] [PDF]
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Q. L. Deveraux and J. C. Reed IAP family proteins---suppressors of apoptosis Genes & Dev., February 1, 1999; 13(3): 239 - 252. [Full Text]
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G. Lizard, S. Gueldry, O. Sordet, S. Monier, A. Athias, C. Miguet, G. Bessede, S. Lemaire, E. Solary, and P. Gambert Glutathione is implied in the control of 7-ketocholesterol-induced apoptosis, which is associated with radical oxygen species production FASEB J, December 1, 1998; 12(15): 1651 - 1663. [Abstract] [Full Text]
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H. R. Stennicke, J. M. Jurgensmeier, H. Shin, Q. Deveraux, B. B. Wolf, X. Yang, Q. Zhou, H. M. Ellerby, L. M. Ellerby, D. Bredesen, et al. Pro-caspase-3 Is a Major Physiologic Target of Caspase-8 J. Biol. Chem., October 16, 1998; 273(42): 27084 - 27090. [Abstract] [Full Text] [PDF]
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C. M. Carthy, D. J. Granville, K. A. Watson, D. R. Anderson, J. E. Wilson, D. Yang, D. W. C. Hunt, and B. M. McManus Caspase Activation and Specific Cleavage of Substrates after Coxsackievirus B3-Induced Cytopathic Effect in HeLa Cells J. Virol., September 1, 1998; 72(9): 7669 - 7675. [Abstract] [Full Text] [PDF]
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J. M. Zapata, R. Takahashi, G. S. Salvesen, and J. C. Reed Granzyme Release and Caspase Activation in Activated Human T-Lymphocytes J. Biol. Chem., March 20, 1998; 273(12): 6916 - 6920. [Abstract] [Full Text] [PDF]
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H.-k. Huang, C. A. P. Joazeiro, E. Bonfoco, S. Kamada, J. D. Leverson, and T. Hunter The Inhibitor of Apoptosis, cIAP2, Functions as a Ubiquitin-Protein Ligase and Promotes in Vitro Monoubiquitination of Caspases 3 and 7 J. Biol. Chem., August 25, 2000; 275(35): 26661 - 26664. [Abstract] [Full Text] [PDF]
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S. Luschen, S. Ussat, G. Scherer, D. Kabelitz, and S. Adam-Klages Sensitization to Death Receptor Cytotoxicity by Inhibition of Fas-associated Death Domain Protein (FADD)/Caspase Signaling. REQUIREMENT OF CELL CYCLE PROGRESSION J. Biol. Chem., August 4, 2000; 275(32): 24670 - 24678. [Abstract] [Full Text] [PDF]
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E. Leo, Q. L. Deveraux, C. Buchholtz, K. Welsh, S.-i. Matsuzawa, H. R. Stennicke, G. S. Salvesen, and J. C. Reed TRAF1 Is a Substrate of Caspases Activated during Tumor Necrosis Factor Receptor-alpha -induced Apoptosis J. Biol. Chem., March 9, 2001; 276(11): 8087 - 8093. [Abstract] [Full Text] [PDF]
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J. M. Angelastro, N. Y. Moon, D. X. Liu, A.-S. Yang, L. A. Greene, and T. F. Franke Characterization of a Novel Isoform of Caspase-9 That Inhibits Apoptosis J. Biol. Chem., April 6, 2001; 276(15): 12190 - 12200. [Abstract] [Full Text] [PDF]
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Y. Suzuki, Y. Nakabayashi, K. Nakata, J. C. Reed, and R. Takahashi X-linked Inhibitor of Apoptosis Protein (XIAP) Inhibits Caspase-3 and -7 in Distinct Modes J. Biol. Chem., July 13, 2001; 276(29): 27058 - 27063. [Abstract] [Full Text] [PDF]
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W. P. dela Cruz, P. D. Friesen, and A. J. Fisher Crystal Structure of Baculovirus P35 Reveals a Novel Conformational Change in the Reactive Site Loop after Caspase Cleavage J. Biol. Chem., August 24, 2001; 276(35): 32933 - 32939. [Abstract] [Full Text] [PDF]
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