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Blood, Vol. 96 No. 3 (August 1), 2000:
pp. 1030-1038
IMMUNOBIOLOGY
Caspase-independent commitment phase to apoptosis in activated
blood T lymphocytes: reversibility at low apoptotic insult
Céline Dumont,
Antoine Dürrbach,
Nicolas Bidère,
Matthieu Rouleau,
Guido Kroemer,
Ghislaine Bernard,
François Hirsch,
Bernard Charpentier,
Santos A. Susin, and
Anna Senik
From the Laboratoire d'Immunologie Cellulaire et de
Transplantation, Laboratoire de l'Apoptose, Cancer et Immunité,
Villejuif, France; Unité INSERM 343, Hôpital de
l'Archet, Nice, France.
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Abstract |
Little is known about the mechanisms of programmed death triggered
in T lymphocytes by stimuli that can bypass caspase activation. Anti-CD2 monoclonal antibody and staurosporine are such apoptosis inducers because they operate in the presence of broad-spectrum caspase
inhibitors BOC-D.fmk and Z-VAD.fmk. A system was devised, based on the isolation according to density of activated blood T cells
progressively engaged in the apoptotic process. This allowed definition of a sequence of caspase-dependent and
caspase-independent apoptogenic events that are triggered by anti-CD2
and staurosporine. Thus, a commitment phase to apoptosis was defined
that is entirely caspase independent and that is characterized by cell
volume loss, partial chromatin condensation, and release into the
cytosol and the nucleus of mitochondrial "apoptosis-inducing factor
" (AIF). Committed cells were viable, displayed a high mitochondrial
inner transmembrane potential (  m), and lacked large-scale and
oligonucleosomal DNA fragmentation. Mitochondrial release of AIF was
selective because cytochrome c was retained in mitochondria of the very same cells. Mitochondrial release of cytochrome c occurred later, at
the onset of the execution phase of apoptosis, concurrently with
m collapse, poly (ADP-ribose) polymerase cleavage, and DNA
fragmentation. The apoptogenic events of this commitment phase are
reversible if the strength of the stimulus is low and of short duration.
(Blood. 2000;96:1030-1038)
© 2000 by The American Society of Hematology.
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Introduction |
Apoptosis or programmed cell death is a
self-destruction process characterized by stereotyped ultrastructural
changes including condensation of the nucleus and cytoplasm, membrane
blebbing,1 and external display of phosphatidylserine, a
signal for recognition and engulfment of apoptotic cells by adjacent
cells. In mammalian cells, the onset of apoptosis correlates with the
activation of a family of cysteine proteases called caspases, which are
constitutively expressed as inactive zymogens in the cytosol. Caspases
constitute a potent machinery that cleaves crucial proteins of the
nucleus and the cytoskeleton after Asp residues, and thus induce the
phenotypic changes of apoptosis, including advanced chromatin
condensation and internucleosomal DNA fragmentation (reviewed in Cryns
and Yuan2). Evidence from an increasing number of
experimental systems, using broad-spectrum caspase inhibitors, however,
supports the notion that programmed cell death can also proceed in a
caspase-independent manner, with stereotyped features 3-8
(reviewed in Borner and Monney9). For example, we have
previously shown that in activated T cells exposed to some apoptotic
inducers (anti-CD2 monoclonal antibody [mAb] or staurosporine) in the
presence of Z-VAD.fmk or BOC-D.fmk, cell death occurs with only partial
condensation of the chromatin and without internucleosomal DNA
fragmentation, but maintains many other features of apoptosis, such as
shrinkage of the cytoplasm and the nucleus,  m collapse, membrane
blebbing, and externalization of phosphatidylserine.6
Remarkably, the expression of the adenovirus protein E4orfA in rodent
fibroblasts induces apoptosis without even activating
caspase-3.7 The same is true for T lymphocytes subjected to
cross-linking of the human immunodeficiency virus-1 (HIV-1) coreceptor
CXCR4 by specific antibodies.10 Chicken erythrocytes
exposed to staurosporine die from apoptosis yet lack active proteases
that cleave Z-VAD.afc, a classical caspase substrate, suggesting that
caspase activation may not be involved at all in this system of cell
death.11
The cellular mechanisms that account for caspase-independent programmed
cell death are still elusive. In most cells subjected to an apoptotic
insult in the presence of Z-VAD.fmk or BOC-D.fmk, dissipation of the
mitochondrial inner transmembrane potential (m) is
detected.3,6,7 The loss of m is mediated by opening
of the mitochondrial permeability transition (PT) pores12 and is believed to be an early and irreversible event in the apoptotic process.13 Caspase-unrelated death triggers or effectors
might be released from mitochondria concomitantly with this
mitochondrial dysfunction, thus triggering a caspase-independent cell
death pathway. One critical effector might be apoptosis-inducing factor (AIF), a flavoprotein with striking homology with the reduced form of
nicotinamide adenine dinucleotide oxidoreductases.14 AIF
resides in the intermembrane space of mitochondria and is translocated
to the nucleus on opening of PT pores. When microinjected into cells in
the presence of Z-VAD.fmk, recombinant AIF induces the same cellular
changes as those observed during caspase-independent cell
death.14
In most cells undergoing apoptosis, AIF may exert its apoptogenic
effects in concert with mitochondrial factors involved in caspase
activation, in particular with cytochrome c, a factor that is
translocated from the intermembrane space of mitochondria to the
cytosol after a death signal, and which, together with Apaf-1, dATP and
caspase-9, activates pro-caspase-3 (reviewed in Cryns and
Yuan2 and Wolf and Green15). A number of other intermembrane space proteins are also released from mitochondria during
apoptosis, including pro-caspase-9,16
pro-caspase-3,17 and the Hsp10 and Hsp60 chaperons, which
assist pro-caspase-3 activation by upstream proteases.18,19
Hence, an important issue is to know the sequence of caspase-dependent
and caspase-independent cellular events that lead to the demise of
cells during apoptosis. In activated T cells exposed to anti-CD2 or
staurosporine, both caspase-dependent and a caspase-independent
pathways of cell death are triggered.6 In the present
study, we sought to characterize the ordering of cellular events that
define the commitment phase of apoptosis in these models. Progressive
cell shrinkage and concomitant increase in cell density occur early
after apoptotic induction.20 This allows us to separate
cells with increasing density, which represent successive stages of the
apoptotic process, on discontinuous Percoll gradients.21
Using this approach, we exposed large activated T cells to
apoptosis-inducing stimuli, and thereafter fractionated them according
to density. In each fraction, we monitored signs of apoptosis
associated with mitochondrial alterations such as m collapse,
release of cytochrome c and AIF, as well as signs of nuclear apoptosis
such as chromatin condensation and DNA fragmentation. We also
analyzed whether these events depended on caspase activation. For
comparison, the same parameters were investigated in cells exposed to
anti-CD95, a stimulus that directly triggers a caspase cascade in
peripheral T lymphocytes.22
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Materials and methods |
Reagents
The CD2 mAb were GT2, T111, and D66.6
Anti-CD95 mAb (CH-11) was from Immugenex Corp (Los Angeles, CA).
Anti-PARP mAb (C2-10) was purchased from Dr G. G. Poirier (Montreal
University, Montreal, Quebec, Canada). The mAbs recognizing the native
form of cytochrome c (6H2.B4) or its denatured form (7H8.2C12) were
from PharMingen (Becton Dickinson, Le Pont de Claix, France).
Anticytochrome oxidase subunit II mAb (12C4-F12) was from Molecular
Probes, (Interchim, Montluçon, France). Antiactin rabbit serum
was from Sigma (Saint Quentin Fallavier, France). Rabbit
anti-AIF was raised against a mixture of AIF peptides.14
Z-VAD.fmk (benzyloxycarbonyl-Val-Ala-Asp[Ome]-fluoromethyl ketone)
and BOC-D.fmk (Boc-Asp[Ome]-fluoromethyl ketone) were purchased from Enzyme Systems Products (Dublin, CA).
T lymphocyte isolation, culture conditions, and induction of
cell death
Peripheral blood leukocytes were isolated from blood bank
leukophoresis packs obtained from healthy volunteers (Blood Transfusion Center of Hôpital Saint Louis, Paris). After Ficoll-Isopaque density (d = 1.078) gradient centrifugation, adherent cells were removed by incubation on plastic dishes and passage over nylon wool
columns. T lymphocytes (6 × 106 in the wells of
6-well flat-bottomed plates; Nunc, Roskide, Denmark) were stimulated
for 4 to 5 days with the mitogenic GT2+T111 CD2 mAb pair (2 µg/mL) plus 100 U/mL interleukin-2 (IL-2) (Roussel Uclaf,
Romainville, France). The death signals were given by
T111+ D66 (2µg/mL), CH-11 (250 ng/mL), or staurosporine
(0.1-0.5 µmol/L; from Sigma).
Percoll gradients
The T cells were fractionated on discontinuous Percoll gradients
(Pharmacia Biotech, Saclay, France). Six different concentrations of
Percoll in phosphate-buffered saline (PBS) 10% fetal calf serum (FCS)
were used (37.5%, 40%, 42.5%, 45%, 47.5%, and 60%). Fractions recovered at each interface from the top to the bottom of the gradients
were numbered F1 to F5. The low buoyant-density F1 fraction, primarily
constituted of dead cells, was discarded.
Flow cytometric analyses of m and of phosphatidylserine
externalization
To evaluate changes in inner mitochondrial transmembrane potential
m, cells were stained for 15 minutes at 37° with 40 nmol/L of
the potential sensitive fluorescent dye DiOC6
(3.3'-diethyloxacarbocyanine) from Molecular Probes.
Phosphatidylserine externalization was detected by staining the cells
with annexin V-FITC from Euromedex (Souffelweyersheim, France).
Hypodiploid cell assessment and microscopic detection of
chromatin condensation
Cells (5 × 105) were washed in PBS with 5.5 mmol/L glucose and fixed overnight in ethanol (70% in water, at
4°C). Cells were then resuspended in 0.5 mL PBS containing 50 µg/mL propidium, 100 U/mL RNAse A (Sigma) and incubated for 1 hour at
room temperature. The DNA content of 104 cells was
monitored by cytofluorometry using a Coulter Epics profile II analyzer.
To assess chromatin condensation, cells were fixed with 3%
paraformaldehyde and incubated for 10 minutes at room temperature with
5 µmol/L 4',6-diamino-2-phenylindole, dihydrochloride (DAPI,
from Molecular Probes). Cells were rinsed, resuspended in Mowiol
mounting medium, and analyzed by conventional (Leica Microsystèmes, Rueil-Malmaison, France) or confocal fluorescence microscopy.
Subcellular fractionation and immunoblotting
T cells (5-10 × 106) were washed twice in PBS
and resuspended in 500 µL of ice-cold extraction buffer23
containing 220 mmol/L mannitol, 68 mmol/L sucrose, 50 mmol/L PIPES-KOH,
pH 7.4, 50 mmol/L KCl, 5 mmol/L sodium EGTA, 2 mmol/L
MgCl2, 1 mmol/L dithiothreitol (DTT), and a cocktail of
protease inhibitors (from Boehringer Mannheim, Meylan, France). After
incubation on ice for 30 minutes, cells were homogenized with 60 pestle
strokes of a glass homogenizer (Polylabo, Strasbourg, France). Unbroken
cells and nuclei were pelleted by centrifugation at 760g for 10 minutes at 4°C. The resulting supernatants were centrifuged at
10 000g for 15 minutes to yield S10 supernatants and pellets
enriched in mitochondria. The latter was resuspended in extraction
buffer supplemented with 1% Triton X-100. The S10 supernatants were
further centrifuged at 100 000g for 1 hour and the final S100
supernatants were either used as such or concentrated for 2 hours at
4°C on a 3 mw cut-off Centricon apparatus (Amicon Bioseparation,
Millipore, ST Quentin en Yvelines, France). S100 and mitochondrial
fractions were stored at  80°C.
Immunoblot analyses
Five to 20 µg of S100 cytosolic extracts and 5 µg of
mitochondrial extracts were separated by sodium dodecyl
sulfate-polyacrylamide gel electrophoresis (SDS-PAGE; 12%
polyacrylamide) and transferred onto a polyvinylidene difluoride (PVDF)
membrane (Immobilon-P, Millipore). After overnight incubation in PBS
containing 3% bovine serum albumin (BSA) and 0.1% Tween 20, the
membranes were incubated for 2 hours at room temperature with rabbit
anti-AIF serum (1:2000 diluted), with anticytochrome c (1:2000) or with
anticytochrome c oxidase (1:500). The blots were stripped by using a
Western blot recycling kit (Euromedex) and reprobed with antiactin
(1:500). The immunoreactive proteins were visualized by using
horseradish peroxidase-coupled (HRP) goat antimouse IgG (Amersham, Les
Ulis, France) or HRP-conjugated sheep antirabbit IgG (Biotest, BUC, France) and enhanced chemiluminescence detection system (ECL kit, Amersham). To evaluate PARP cleavage, 3 × 105 cells
were washed and solubilized in 10 µL Laemmli buffer. Cell lysates were then subjected to 8% SDS-PAGE and electroblotted onto a
PVDF membrane. Blots were stained with anti-PARP (1:10 000) and
revealed as above.
Confocal laser scanning microscopy analysis of cytochrome c and AIF
localization
Cells were fixed with 3% paraformaldehyde in PBS for 30 minutes at
4°C, washed with PBS, and incubated with ClNH4 (50 mmol/L) before being permeabilized with 0.05% Triton X-100 for 5 minutes at room temperature. After 3 washings, the cells were incubated for 1 hour at 4°C with rabbit anti-AIF and with mouse
anticytochrome c, respectively, diluted at 1:200 and 1:100 in PBS
supplemented with 0.5% BSA and 2% FCS. The cells were stained with
fluorescein isothiocyanate (FITC)-conjugated AffiniPure goat antirabbit
IgG and tetrarhodamine isothiocyanate (TRITC)-conjugated AffiniPure donkey antimouse IgG from Jackson Immunoresearch Laboratories (Immunotech, Luminy-Marseille, France). For localization of
mitochondria, the cells were loaded with the mitotracker CMXRos red dye
(Molecular Probes) before fixation, permeabilization, and labeling with
the desired FITC-conjugated AffiniPure antibodies. Cells were examined using a CLSM confocal microscope (Leica).
Field inversed gel electrophoresis (FIGE)
One percent agarose plugs containing 1 × 106
cells were digested with proteinase K (0.5 mg/mL) at 37°C for 48 hours in the presence of 0.5 mol/L EDTA and 10 mg/mL lauroyl sarcosine.
This step was followed by electrophoresis for 24 hours in 1% agarose gels using a FIGE mapper equipment from Bio-Rad Laboratories (Ivry Sur
Seine). The gels were run for 24 hours in 0.5× TBE buffer (pH
8.3) at 170 V with a ramping rate changing from 5 to 15 seconds. Molecular weight standards (midrange PFG marker I) were purchased from
Biolabs (Ozyme, Montigny le Bretonneux).
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Results |
Fractionation according to density of activated T-cell populations
exposed to CD2 apoptotic signaling: definition of a commitment
phase of apoptosis and comparison with anti-CD95-treated T cells
Within 2 hours following the receipt of CD2 apoptotic signal, in
vitro activated T cells begin to shrink without losing membrane integrity and without exposing phosphatidylserine. Previous studies have determined that full-blown apoptosis occurs later, yielding after
4 hours approximately 30% to 40% apoptotic cells (with low m),
which rapidly become permeable to trypan blue.6 These observations led us to postulate that the first 2 hours of CD2 stimulation corresponded to a commitment phase and that initiation of
cell shrinkage was one of the events characterizing commitment to
apoptosis. A starting population of large activated T cells, homogeneous in terms of cell size, was isolated as F2-I cells in the
low buoyant density fraction of a discontinuous density Percoll
gradient. This F2-I cell population was then exposed for 2 hours to CD2
apoptotic signaling and, after extensive washing, fractionated on a
second Percoll gradient into 5 fractions (F2-II to F5) displaying
progressive increase in density and progressive decrease in cell volume
(Figure 1A) and forward side scatter
(Figure 1D). Control F2-I cells that had not been exposed to apoptotic insult remained at the low buoyant density level of the secondary gradients. Immediately after fractionation, most cells were viable as
assessed by trypan blue exclusion and morphologic examination. When
replaced for 16 hours in medium containing IL-2, large cells of the
F2-II fraction remained viable, whereas in the subsequent fractions,
increasing proportions of cells underwent apoptosis, yielding
35 ± 11% dead cells in F3, 58 ± 9% in F4,
and81 ± 7% in F5. The carry-over of residual anti-CD2 mAb did
not account for cell death because subjecting F2-I cells to a short
acid pulse (pH 3, 1 minute on ice), at the end of the signalization
period, removed all residual membrane-bound CD2 antibodies but did not modify the number of cells recovered in each fraction, nor the rate of
cell death occurring thereafter (not shown). This indicated that the
apoptotic process was launched despite interruption of the apoptotic
insult. Kinetic experiments, carried out to examine the distribution of
anti-CD2-treated cells in the gradient fractions over time (Figure 1B),
strongly supported the notion that F3 and F4 cells were intermediary
between F2 and F5 cells. The presence of BOC-D.fmk or Z-VAD.fmk (50 µmol/L, to target most caspases) during the entire experimental
procedure did not alter the distribution of condensed cells along the
density gradients nor the percentages of cells committed to death in
each fraction. Thus, the successive cell populations recovered from
Percoll gradients after a short period of CD2 apoptosis induction were
progressively enriched in cells committed to caspase-independent death.
In the F2-I population subjected to CD95 ligation for 2 hours, the
fractionation procedure gave rise to cells mostly committed to death,
irrespective of their density. As expected, anti-CD95-induced cell
death was strongly inhibited by the caspase inhibitor BOC-D.fmk (Figure
1C).
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| Fig 1.
Fractionation of T cells after apoptotic induction.
Activated T-cell populations exposed for 2 hours to CD2 or
CD95 apoptotic induction and fractionated according to density on
discontinuous Percoll gradients give rise to fractions progressively
enriched in cells committed to apoptosis. (A) Homogeneous populations
of large activated T cells (F2-I) obtained from the low buoyant density
fractions of a first primary Percoll gradient were exposed for 2 hours
to CD2 mAb (D66 +T111, 2 µg/mL) and were then
fractionated again on secondary Percoll gradients to yield fractions
F2-II, F3, F4, and F5 with decreasing cell volumes. Columns are mean
cell volumes ± SD of determinations performed in 3 different
experiments, as measured by using a Coulter counter and a channelyser.
 --- indicates percent dead cells in the fractionated cell
populations 2 hours after apoptotic induction, just after their
recovery from the gradients, as assessed by trypan blue uptake and cell
morphology (n = 40);  ---, percent dead cells in fractionated
cell populations replaced after washing in culture for 18 hours in IL-2
containing medium (n = 10);  ---, percent dead cells in cells
exposed to 50 µmol/L BOC-D.fmk during apoptotic induction and during
subsequent culture in IL-2 containing medium (n = 3);  ---,
percent dead cells similarly exposed to 50 µmol/L Z-VAD.fmk. Values
are means ± SD. (B) Activated (F2-I) T cells exposed for various
periods of time to anti-CD2 mAb. At each time point, the percentages of
cells recovered in the F2-II ( ---), F3 (---), F4
(---), and F5 (---) Percoll fractions were estimated. It is
seen that the percentages of F2-II cells progressively decreased within
2 hours, whereas the percentages of F3, and to a lesser extent, those
of F4 and F5 cells concomitantly increased. After 3 hours, the
percentages of F2-II cells were stabilized, but a further rise in F5
cells occurred at the expense of F3 and F4 cells. (C) Fractionation of
F2-I cells treated for 2 hours with an anti-CD95 at 250 ng/mL
(n = 10). Symbols are the same as in panel A. Cells treated in
parallel with 50 µmol/L BOC-D.fmk did not shrink and were therefore
recovered in majority in the low buoyant fraction (F2-II) of the
secondary Percoll gradients (). (D) Forward scatter (FSC) analysis
showing the reduction of cell size from F2-II to F5. Numbers refer to
the mean forward scatter. SSC, side scatter.
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Partial chromatin condensation precedes m collapse, PARP
cleavage activity, and large scale DNA fragmentation in anti-CD2
treated cells
Using the lipophilic cationic fluorochrome
DiOC6(3), which localizes to
mitochondria as a consequence of m, we observed that the vast
majority of cells in fractions F2-II to F4 exhibited, 2 hours after CD2
apoptotic induction, a high mitochondrial transmembrane potential,
similar to that of the starting F2-I population. F5 was the sole
fraction displaying m collapse (with only about 45%
DiOC63high cells versus about 88% in control
F2-I, Figure 2A). As previously reported,6 the loss of m in this fraction was
caspase-independent (not prevented by BOC-D.fmk, Figure 2B). From F2-II
to F4, there was no significant generation of hypodiploid cells and in
cells exposing phosphatidylserine on their surface. A small increase in
the percentages of hypodiploid cells and annexin V+ cells
was, however, observed in F5 (Figure 2A,C). Because no proteolytic
cleavage of the nuclear PARP into its 85-kd signature fragment (mainly
performed by caspase-3 and caspase-7)24 was detected from
F2-II to F4, whereas it was detected in F5 (Figure 2D), one can
conclude that the caspase cascade was not activated until F5. By
comparison, all fractionated cell populations recovered 2 hours after
CD95 cross-linking displayed PARP cleavage and most of them had lost
their mitochondrial transmembrane potential. In activated T
lymphocytes, caspase-8 is, in fact, very rapidly recruited to the
death-inducing signaling complex (DISC) on CD95 ligation.22
This indirectly causes mitochondrial damage and cytochrome c release
through the cleavage and activation of the death agonist
BID.25,26
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| Fig 2.
Apoptotic alterations among the fractionated cell
populations of activated T lymphocytes recovered 2 hours after their
exposure to CD2 and CD95 apoptotic stimuli.
(A)  --- indicates percent cells with high m as measured in
each fraction with DIOC6 3;  ---, percent
cells with condensed nuclei as visualized by DAPI staining; ---,
percent hypodiploid cells; ---, percent annexin-V-positive cells.
Values are means ± SD of 20 determinations in the CD2 apoptotic
system and of 10 determinations in the CD95 system. B)
Effect of 50 µmol/L BOC-D.fmk on the occurrence of m disruption
in F5 cells of the CD2 system and in F2-II cells of the CD95 system.
Data shown are representative of 3 different experiments. (C) Propidium
iodide staining of DNA from ethanol-permeabilized cells. Values
indicate the percentages of hypodiploid cells (linear scales are
represented). (D) Total lysates of 5 × 105 cells
were analyzed by Western blot using an anti-PARP mAb. Experiments are
representative of 6 independent determinations.
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One striking nuclear change observed in the fractionated cell
populations generated 2 hours after CD2 apoptotic signaling was
progressive chromatin condensation, starting from F3. Thus, 36 ± 15% cells with condensed chromatin were scored in F3 by
DAPI staining, 51 ± 12% in F4, and 95 ± 9% in F5 (versus
about 5% in F2-I, n = 3). Ultrastructural examination of the
fractionated cell populations (Figure 3)
gave a more precise insight into nuclear morphology, indicating that
45% F3 cells and approximately 64% F4 cells had their
heterochromatin condensed in discrete clumps abutting the nuclear
membrane, whereas more condensed chromatin was seen in F5 cells. The
presence of 50 µmol/L BOC-D.fmk during apoptosis signaling did not
affect the partial chromatin condensation detected in F3 and F4 cells.
This caspase inhibitor, however, prevented the advanced chromatin
condensation detected in F5. Moreover, BOC-D.fmk inhibited all signs of
nuclear apoptosis in anti-CD95-treated cells. We wondered whether
large-scale chromatin fragmentation, which precedes internucleosomal
fragmentation,27 would coincide with the chromatin
condensation seen in the CD2 system. However, no cleavage of DNA could
be detected by FIGE from F2-II to F4 fractions of anti-CD2-treated
cells, whereas cleavage to large kilobase pair sized fragments (50 kbp)
was generated in F5 (Figure 4A). BOC-D.fmk
inhibited the formation of these fragments (Figure 4B), corroborating
the notion that large-scale DNA fragmentation may, in certain models,
strictly depend on upstream caspase activity.28,29
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| Fig 3.
Ultrastructural changes occurring in the fractionated
cell populations recovered 2 hours after apoptotic induction.
(A) Sizeable proportions of activated T cells sedimenting in the F3 and
F4 fractions of Percoll gradients after CD2 signaling display partial
chromatin condensation. F2-I cells were exposed for 2 hours to
anti-CD2, fractionated on density Percoll gradients, and examined by
electron microscopy. Values inside the brackets are the percentages of
cells displaying the same morphology as that represented in the
pictures. (Bi) In the CD2 system, the presence of BOC-D.fmk (50 µmol/L) during apoptotic induction does not prevent the appearance of
F5 cells with partial chromatin condensation, contrasting with the
strong condensation seen in F5 cells generated in the absence of
caspase inhibitor. (Bii) Cells treated with anti-CD95 in the presence
of BOC-D.fmk retain a normal nuclear morphology and are still recovered
in the low buoyant density fraction of Percoll gradients (at the level
of F2-II).
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| Fig 4.
DNA cleavage.
(A) Large-scale DNA fragmentation does not occur in F3 and
F4 cells committed to apoptosis after CD2 apoptotic induction but does
occur in a BOC-D.fmk inhibitable manner. (B) F5 cells entered the
execution phase. Genomic DNA from 2 × 106 cells of
each Percoll fraction was analyzed by pulse-field gel
electrophoresis.
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On the whole, it appears that in the CD2 apoptotic system, the
commitment phase to apoptosis, represented by F3 and F4 cells, is
characterized by a reduction in cell size and by partial chromatin condensation, both being caspase independent. The subsequent execution phase of apoptosis is represented by frankly shrunken F5 cells, an
important proportion of which displays caspase-independent m loss
and caspase-dependent nuclear events such as advanced chromatin
condensation, large-scale DNA fragmentation, and PARP cleavage. In our
experimental conditions, the execution phase was already at work in
F2-II cells generated after CD95 signaling, as evidenced by the early
m loss affecting the majority of cells, and by PARP degradation.
These events preceded cell shrinkage and advanced chromatin
condensation, because only about 27% of F2-II cells displayed
chromatin condensation, and this at a rather low extent (Figure 3).
Subcellular redistribution of AIF but not cytochrome c during the
commitment phase of CD2-induced apoptosis
Subcellular fractionation of F3 to F4 cells (pooled to gain
sufficient material), followed by immunoblot detection, did not allow
detection of significant mitochondrial release of cytochrome c (Figure
5A), even when the S100 cytosolic extracts
were concentrated up to 4-fold (data not shown), demonstrating the
exclusive mitochondrial localization of cytochrome c in those cells.
Adequacy of the subcellular fractionation procedure was assessed by
using an anticytochrome oxidase mAb. In anti-CD95-treated cells,
therelease of cytochrome c into the cytosol already occurred in F2-II.
We also examined by this technique whether mitochondrial AIF was
released into the cytosol, using a rabbit antiserum to AIF. The
specificity of anti-AIF detection was assessed in whole cell lysates by
comparison with the preimmune serum obtained from the same rabbit
(Figure 5B). Two forms of AIF were apparently recognized by the
anti-AIF serum in SDS-PAGE analysis, one with an apparent 67-kd
molecular mass, which corresponded to the migration of recombinant AIF
(Figure 5C), and the other with an apparent 57-kd molecular mass.
Detection of both forms was strongly attenuated by preincubating the
anti-AIF serum with a cocktail of AIF-derived peptides (Figure 5B). The molecular mass of recombinant AIF, deduced from the cDNA incoding AIF14 and confirmed by mass spectrometry (not shown), is 57 kd. Its apparent higher molecular weight detected in SDS-PAGE may be
due to ionic interaction with the gel matrix, as shown for another
ferredoxin oxidoreductase.30 Variations in the isoelectric points of AIF molecules might account for the differential migration patterns in SDS-PAGE. After CD2 apoptotic signaling, AIF was detected in the cytosol of pooled F3 to F4 cells but not in the cytosol of
control F2-I cells or of F2-II cells. Data not shown indicated that the
mitochondrial extracts from F3 to F4 cells still contained AIF. These
data suggested that at least part of AIF was released from mitochondria
during the commitment phase of apoptosis in anti-CD2-treated cells,
whereas cytochrome c was retained in these organelles. In F2-II cells
from anti-CD95-treated cells, which served as positive controls, both
cytochrome c and AIF were released into the cytosol.
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| Fig 5.
Mitochondrial AIF is released into the cytosol before
cytochrome c in activated T cells subjected to CD2 apoptotic induction.
(A) Cytosolic and mitochondrial extracts (5 µg) were
prepared from cell populations that have been fractionated on Percoll
density gradients after a 2-hour apoptotic induction with anti-CD2 or
with anti-CD95. F3 and F4 cells were pooled to gain sufficient
material. Total cell lysates were obtained from control activated F2-I
cells. Immunoblots were probed with anticytochrome c (Cyt.c), with
anticytochrome c oxidase subunit II (Cyt.c.ox.II) to detect the
presence of mitochondrial material, or with antiactin to monitor
protein loading in cytosolic extracts. (B) Specificity of AIF detection
by immunoblot analysis. Total lysates of 5 × 105
activated T cells (F2-I) were probed with a preimmune rabbit serum
(lane 1), or with a rabbit AIF antiserum (lane 2), or with the same AIF
antiserum preincubated for 2 hours at 4°C under agitation with 1 mmol/L of a mixture of immunogenic AIF-derived peptides (amino acids
151-170, 166-185, 181-200). (C): Immunoblot analysis of cytosolic
extracts from anti-CD2 and anti-CD95 treated cells by using AIF
antiserum. Recombinant AIF (5 ng) and total cell lysates (5 µg) were
used as positive controls. Data are representative of 5 experiments.
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The respective cellular localizations of cytochrome c and AIF were
further examined by laser scanner confocal microscopy in anti-CD2-treated cells that were simultaneously labeled with
anti-cytochrome c and anti-AIF. The immunostaining of untreated F2-I
cells demonstrated a punctate staining pattern for cytochrome c that
matched the staining of AIF, consistent with the localization of these
molecules in mitochondria (Figure 6A, B).
This was confirmed by loading the cells with Mitotracker Red CMXRos,
which localizes to mitochondria, and by staining cytochrome c or AIF in
green. In this case, the punctate red staining pattern of CMXRos
perfectly matched that of cytochrome c or AIF (not shown). After CD2
apoptotic signaling, cytochrome c remained in the mitochondria of all
fractionated populations except F5 in which a diffuse distribution
pattern throughout the cytosol was observed, even when apoptotic
induction was performed in the presence of BOC-D.fmk or of Z-VAD.fmk
(to ensure that most caspases would be inhibited; Figure 6C). In
contrast, AIF was translocated earlier, as deduced from the diffuse
anti-AIF staining already exhibited by the large majority ( 95%) of
F3 cells but not by F2-II cells, not committed to death. In some F3
cells, AIF was detected in the cytosol and to a lesser extent in the
nucleus, whereas in F4 and F5 cells, it was detected equally well in
the cytosol and the nucleus, consistent with the translocation of AIF
through the cytosol toward the nucleus.14 The increased intensity of anti-AIF staining after release from mitochondria may
reflect a better accessibility of the antigens recognized by the AIF
antiserum because no increase in the total amount of AIF could be
detected by Western blot analysis in any cell lysate, when compared to
the lysates of control F2-I cells (not shown). As observed for
cytochrome c, BOC-D.fmk and Z-VAD.fmk had no effect on AIF
redistribution (Figure 6C), indicating that this was a caspase-independent phenomenon. On the whole, the release of AIF from
mitochondria is an early and apparently selective event because it
occurs before the release of cytochrome c.
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| Fig 6.
Differential subcellular localization of cytochrome c and
AIF during the commitment phase of anti-CD2-induced apoptosis as
visualized by confocal immunofluorescence microscopy.
(A) In F3 and F4 cells, the punctate cytosolic distribution pattern of
cytochrome c differs from the diffuse staining pattern of AIF
throughout the cytosol and the nucleus. (B) A microscopic field
displays several F3 cells to show that most of them exhibit a
differential distribution pattern for AIF and for cytochrome. (C,D) The
presence of BOC-D.fmk or Z-VAD.fmk (50 µmol/L) during apoptotic
induction and Percoll fractionation does not prevent the mitochondrial
release of cytochrome c and AIF in F5 cells.
|
|
Volume cell loss, partial chromatin condensation, and release of AIF
may be reversible at an early stage of caspase-independent cell death
commitment
When added to large F2-I cells, 250 nmol/L staurosporine provoked
within 2 hours the shrinkage of 70% or more of the cells in a
caspase-independent manner (not prevented by BOC-D.fmk, not shown).
When recovered in the high buoyant density fraction of Percoll
gradients, those cells were similar in many aspects to the F4
cells generated after CD2 apoptotic insult: about half of
them were committed to apoptotic death as determined by subsequent culture in IL-2-containing medium, but most displayed high m, moderate chromatin condensation, and diffuse immunostaining of AIF
spread into the cytoplasm and the nucleus, contrasting with the
punctate immunostaining of cytochrome c. Noteworthy, in both the CD2
and the staurosporine systems, the release of AIF from mitochondria was
not necessarily correlated with cell death. For example, although at
least 95% of F3 and F4 cells generated during CD2 apoptotic induction
exhibited a diffuse immunostaining of AIF, only part of them
(respectively, about 37% and about 50%) died in subsequent culture,
suggesting that the apoptogenic factor AIF was degraded or held in
check in the surviving cells. To study this aspect more in detail, we
sought to obtain an homogenous population of cells just engaged in the
commitment phase of apoptosis, assuming that a weak and brief apoptotic
insult would induce AIF release, without causing cell death. This
possibility was examined in staurosporine-treated cells, which allows
control of the strength of the apoptotic stimulus. When activated T
lymphocytes (F2-I cells) were exposed to only 100 nmol/L staurosporine
for 2 hours, they shrunk and displayed partial condensation of their
chromatin, events that were not inhibited by BOC-D.fmk (Figure
7A). The shrunken cells sedimented in the
F4 and F5 fractions of Percoll density gradients, displayed a high
mitochondrial transmembrane potential (82 ± 5%
DiOC6(3)high cells for F4 and 70 ± 5%
DiOC6(3)high cells for F5, n = 13), had
intact DNA as assessed by FIGE (Figure 7B), and lacked PARP cleaving
activity (not shown). Confocal laser scanning microscopy revealed that
AIF had lost its punctate mitochondrial distribution and had been
translocated to the cytosol and the nucleus, whereas cytochrome c was
still retained in the mitochondria (Figure 7C). When washed and
replaced in IL-2-containing medium, F4 and F5 cells did not undergo
accelerated apoptosis as compared to control F2-I cells (a result of 6 different experiments). On the contrary, they dilated and recovered
within 24 hours a normal size, and they continued to incorporate
tritiated thymidine (70 800 ± 7308 cpm in F4 and
47 634 ± 7798 cpm in F5 versus 66 711 ± 10 726 in F2-I;
mean ± SD of 3 experiments). The chromatin was no longer condensed
(Figure 8A), and AIF immunostaining tended to regain a punctate distribution pattern, indicative of mitochondrial localization (Figure 8B). It therefore appears that the initial apoptogenic events triggered during the caspase-independent commitment phase to apoptosis in activated T lymphocytes may be reversible, provided the strength of the apoptotic stimulus is weak. Rescue mechanisms may be operative, in particular those that degrade AIF and
repair the DNA alterations responsible for chromatin
condensation.
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| Fig 7.
Characteristics of activated T cells subjected for 2 hours to 100 nmol/L staurosporine.
(A) The ultrastructural changes seen in pooled F4 and F5 cells treated
with staurosporine (STS), that is, cell shrinkage and partial chromatin
condensation, persist in the presence of BOC-D.fmk. (B) Absence of
large-scale DNA fragmentation in F4 and F5 cells contrasting with the
approximate 50-kbp sized fragments seen in the F5 fraction of the CD2
system. 1, F2-I; 2, F4; 3, F5; 4, F5 generated after a 2-hour exposure
to anti-CD2 (positive control). (C) Confocal immunofluorescence
analysis of the localization of cytochrome c and AIF demonstrating that
only mitochondrial AIF has been translocated to the cytosol and the
nucleus.
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| Fig 8.
Reversibility of the morphologic alterations and of
mitochondrial AIF release induced at low apoptotic insult.
(A) F2-I cells that have been treated with 100 nmol/L staurosporine
(STS) for 2 hours were washed and cultured for 24 hours in
IL-2-containing medium. The electron micrographs show that the shrunken
cells with partially condensed chromatin generated after 2 hours of
apoptotic induction recover within 24 hours a normal aspect provided
staurosporine is removed. (B) Confocal microscopy analysis showing that
AIF tends to recover a punctated mitochondrial localization 24 hours
after the removal of staurosporine (compare with Figure 7C).
|
|
| |
Discussion |
This study was aimed at deciphering the ordering of
caspase-dependent and caspase-independent apoptogenic events that are triggered in activated T lymphocytes, using as apoptotic inducers anti-CD2 mAb and staurosporine. The latter stimulus was in particular suitable for analyzing the relationship between the strength of apoptotic induction and the ensuing events. We have set up a system based on the isolation, according to density, of T cells progressively engaged in the apoptotic process. This allowed us to demonstrate that
the commitment phase to apoptosis in the models investigated was
caspase independent and that it was characterized by cell shrinkage,
partial chromatin condensation, and translocation of AIF to the cytosol
and the nucleus. The notion of commitment as such was deduced from the
propensity of the cells to undergo accelerated cell death in culture,
after removal of the apoptotic stimulus. Release of cytochrome c from
mitochondria occurred at a more advanced stage of apoptosis,
concurrently with m collapse and PARP cleavage activity,
indicating that the cells had entered the execution phase of apoptosis.
We have also demonstrated that the initial events of the commitment
phase were reversible at low apoptotic insult.
Most T lymphocytes located in the commitment phase of apoptosis
displayed a high mitochondrial inner membrane potential. Significant m loss was only detected after 1 to 2 hours of further culture (not shown). Yet a mitochondrial response was detected during commitment to apoptosis, which consisted in the caspase-independent translocation of AIF from the intermembrane space of mitochondria to
the cytosol and the nucleus. This release was selective because cytochrome c was retained in the mitochondria of the very same cells as
examined by laser scanning confocal microscopy and Western blot
analysis. Therefore, the initial mitochondrial release of AIF appears
not to be dependent on the overall release of intermembrane proteins,
as is the case for apoptotic cells with swollen mitochondria, the outer
membranes of which can be physically disrupted.31 Instead,
the lymphocytes that have entered the apoptotic pathway possessed
mitochondria that were structurally intact as viewed by electron
microscopy. That AIF was released without occurrence of the major
mitochondrial perturbation represented by m collapse is at odds
with previous studies.14,32 However, mitochondrial dysfunction, due to subtle changes in m (not detectable by
standard methods), may account for the mitochondrial release of AIF in T cells committed to apoptosis. Such subtle changes have been detected
in isolated mitochondria incubated with Bax, a proapoptotic member of
the Bcl-2 family.33 Usually, insertion of Bax at µmol/L concentrations into isolated mitochondria results in m collapse and in cytochrome c release,34,35 events that are
associated with the interaction of Bax with components of the
mitochondrial PT complex.35-37 However, Pastorino and
coworkers33 have shown that at nmol/L concentrations, Bax
does not induce detectable mitochondrial depolarization. Yet, several
mitochondrial intermembrane space proteins are released (although not
completely), including cytochrome c and adenylate kinase.33
This release is due to transient and nonsynchronous opening and closing
of the PT pores of a fraction of mitochondria, hardly detected by
standard methods, but readily detected by using the cyclosporin
A-sensitive release of preloaded calcein. Such a scenario may well
occur in the mitochondria of intact cells because Bax moves from
cytosol to mitochondria after a death signal.38 Not only
would this scenario help in reconciling the opposite points of view
existing in the literature about the temporal relationship between
cytochrome c release and m loss (see the review by Bernardi et
al39), but it could also explain why AIF is released from
mitochondria of cells with high m. However, if such were the
case, the question remains of the reason why AIF and cytochrome c were
not translocated simultaneously in the lymphocytes committed to
apoptosis. This important question awaits elucidation.
The partial condensation of the chromatin observed in the nuclei of the
lymphocytes committed to apoptosis was caspase independent, allowing us
to discard the intervention of caspase-activated factors involved in
chromatin condensation, namely caspase-activated DNAse (CAD),40 caspase-6,41 and the nuclear factor
acinus.42 Recombinant AIF directly causes chromatin
condensation in isolated nuclei, and microinjected AIF antiserum
prevents the nuclear effects (peripheral chromatin condensation and
digestion of chromatin into 50-kbp fragments) induced by staurosporine
in intact cells.14 Released endogenous AIF might therefore
account for chromatin condensation in the lymphocytes committed to
death of our system. Note that cleavage of DNA into 50-kbp fragments
was not detected in these lymphocytes, unless they had entered the
execution phase of apoptosis cells. But in this case, caspase activity
was clearly upstream of large scale DNA fragmentation, in line with
other cell models.28,29 It is possible that in situ the
effects of released AIF might be subjected to cell type-specific
regulation mechanisms with regard to chromatin condensation and DNA
fragmentation, and that caspases may be differentially involved in
these effector events. The causal relationship between the
mitochondrial release of AIF and the early nuclear manifestations of
apoptosis in activated T lymphocytes remains to be established. As to
volume loss induced by apoptotic inducers, it is believed to be tightly
linked to enhanced K+ efflux.43 Shrunken
lymphocytes display both decreased intracellular [K+] and caspase-dependent or independent m
loss.44,45 However, in the committed cells of our study,
the overall mitochondrial transmembrane potential was normal. It would
be interesting to measure [K+] in these populations and
establish the temporal relationship between the expected K+
efflux and the release of AIF, the sole indication thus far of mitochondrial perturbation in our determinations.
A most striking result observed in this study was the reversibility
of cell shrinkage, of chromatin condensation, and of mitochondrial release of AIF, provided the strength of the stimulus was low and its
application of short duration. After cessation of apoptotic insult,
cells pretreated with 100 nmol/L staurosporine for 2 hours recovered a
normal morphology within 18 hours in proliferated as well as control
cells. AIF was no longer present in the cytosol and the nucleus, but
instead was primarily localized in mitochondria, as visualized by
confocal microscopy. When higher concentrations of staurosporine were
used (250-500 nmol/L for 2 hours), the vast majority of shrunken cells
displayed a diffuse distribution pattern of AIF throughout the cytosol
and the nucleus, but only half of them were committed to death.
Similarly, in most F3 and F4 cells of the CD2 system, AIF exhibited a
uniform cellular distribution but important proportions of those cells
survived after removal of anti-CD2 mAb. This suggests that rescue
mechanisms were counteracting the death process and that at low
apoptotic insult they were able to save a majority of cells, whereas at
higher (or prolonged) apoptotic insult they became overwhelmed. The
fact that released AIF had disappeared from the cytosol of saved cells
also suggests that AIF was rapidly degradated. Some recent studies have
demonstrated that the cytochrome c release46,47 and the
initial collapse of m occurring in certain cell death-inducing
conditions48,49 may be reversible. In our setting, the
reversal of chromatin condensation and of cell shrinkage further
emphasize the diversity of repair mechanisms (still to be investigated)
within the different compartments of cells committed to apoptosis.
In the lymphocytes of our study, m loss and mitochondrial
cytochrome c release were coincident with the entry in the execution phase of apoptosis as evidenced by PARP cleavage activity. Both apoptogenic events still occurred in the presence of BOC-D.fmk or
Z-VAD.fmk. They could be followed, in these conditions, by an entirely
caspase-independent cell death process.6 It is now admitted
that when caspase activity is not inhibited, multiple interconnected
amplification loops, either caspase-dependent or caspase-independent,
are involved in most apoptotic models, including CD95-induced
apoptosis.50,51 In our attempt to define the ordering of
caspase-dependent and caspase-independents apoptogenic events that are
triggered in T lymphocytes by signals that do not rely on the "death
receptors" of the tumor necrosis factor-receptor family, we have
characterized a caspase-independent commitment phase to apoptosis.
These data might apply to the caspase-independent cell death triggered
in CD4+ T lymphocytes via the coreceptor of HIV-1, namely
CXCR4,10 and by HIV-1 infection.52 In addition,
the notion of reversible commitment to apoptosis might help developing
strategies to prevent undesired cell death.
| |
Acknowledgments |
We thank Dr N. Zamzami for helpful discussions and A. Oudin for
skillful technical assistance.
| |
Footnotes |
Submitted December 2, 1999; accepted March 30, 2000.
Supported by grants from the CNRS, from the Etablissement
Français des Greffes, the Association pour la Recherche sur le Cancer, and the Hôpital Universitaire de Bicêtre,
Faculté de Médecine de Paris Sud.
Reprints: Anna Senik, Equipe d'Immunologie Cellulaire et de
Transplantation, ERS 1984 du CNRS, 19 rue Guy Moquet, 94801 Villejuif,
France; e-mail: asenik{at}infobiogen.fr.
The publication costs of this
article were defrayed in part by
page charge payment. Therefore,
and solely to indicate this fact,
this article is hereby marked
"advertisement"
in accordance with 18 U.S.C.
section 1734.
| |
References |
1.
Wyllie AH, Kerr JFR, Currie AR.
Cell death: the significance of apoptosis.
Int Rev Cytol.
1980;68:251-306[Medline]
[Order article via Infotrieve].
2.
Cryns V, Yuan J.
Proteases to die for.
Genes Dev.
1998;12:1551-1570[Free Full Text].
3.
Xiang J, Chao DT, Korsmeyer SJ.
BAX-induced cell death may not require interleukin 1 beta-converting enzyme-like proteases.
Proc Nat Acad Sci U S A.
1996;93:14559-14563[Abstract/Free Full Text].
4.
Sarin A, Williams MS, Alexander-Miller MA, Berzofsky JA, Zacharchuk CM, Henckart PA.
Target cell lysis by CTL granule exocytosis is independent of ICE/CED-3 family protease.
Immunity.
1997;6:209-215[Medline]
[Order article via Infotrieve].
5.
McCarthy NJ, Whyte MKB, Gilbert CS, Evan GI.
Inhibition of Ced-3-related proteases does not prevent cell death induced by oncogenes, DNA damage, or the Bcl-2 homolog Bak.
J Cell Biol.
1997;136:215-227[Abstract/Free Full Text].
6.
Déas O, Dumont C, MacFarlane M, et al.
Caspase-independent cell death induced by anti-CD2 and staurosporine in activated human peripheral T lymphocytes.
J Immunol.
1998;161:3375-3383[Abstract/Free Full Text].
7.
Lavoie JN, Nguyen M, Marcellus RC, Branton PE, Shore GC.
E4orf4, a novel adenovirus death factor that induces p53-independent apoptosis by a pathway that is not inhibited by zVAD-fmk.
J Cell Biol.
1998;140:637-645[Abstract/Free Full Text].
8.
Monney L, Otter I, Olivier R, et al.
Defects in the ubiquitin pathway induce caspase-independent apoptosis blocked by Bcl-2.
J Biol Chem.
1998;11:6121-6131.
9.
Borner C, Monney L.
Apoptosis without caspases: an inefficient molecular guillotine?
Cell Death and Differentiation.
1999;6:497-507[Medline]
[Order article via Infotrieve].
10.
Berndt C, Möpps B, Angermüller S, Gierschik P, Krammer PH.
CXCR4 and CD4 mediate a rapid CD95-independent cell death in CD4+ T cells.
Proc Natl Acad Sci U S A.
1998;95:12556-12561[Abstract/Free Full Text].
11.
Weil M, Jacobson MD, Raff MC.
Are caspases involved in the death of cells with a transcriptionally inactive nucleus? Sperm and chicken erythrocytes.
J Cell Sci.
1998;111:2707-2715[Abstract].
12.
Bernardi P.
Mitochondrial transport of cations: channels, exchangers and permeability transition.
Physiol Rev.
1999;79:1127-1155[Abstract/Free Full Text].
13.
Zamzami N, Marchetti P, Castedo M, et al.
Reduction in mitochondrial potential constitutes an early irreversible step of programmed lymphocyte death in vivo.
J Exp Med.
1995;181:1661-1672[Abstract/Free Full Text].
14.
Susin SA, Lorenzo HK, Zamzami N, et al.
Molecular characterization of mitochondrial apoptosis-inducing factor.
Nature.
1999;397:441-446[Medline]
[Order article via Infotrieve].
15.
Wolf BB, Green DR.
Suicidal tendencies: apoptotic cell death by caspase family proteinases.
J Biol Chem.
1999;274:20049-20052[Free Full Text].
16.
Susin SA, Lorenzo HK, Zamzami N, et al.
Mitochondrial release of caspase-2 and -9 during the apoptosic process.
J Exp Med.
1999;189:381-394[Abstract/Free Full Text].
17.
Mancini M, Nicholson DW, Roy S, et al.
The caspase-3 precursor has a cytosolic and mitochondrial distribution: implications for apoptotic signaling.
J Cell Biol.
1998;140:1485-1495[Abstract/Free Full Text].
18.
Samali A, Cai J, Zhivotovski B, Jones DP, Orrenius S.
Presence of a preapoptotic complex of pro-caspase-3, Hsp60 and Hsp10 in the mitochondrial fraction of Jurkat cells.
EMBO J.
1999;18:2040-2048[Medline]
[Order article via Infotrieve].
19.
Xanthoudakis S, Roy S, Rasper D, et al.
Hsp60 accelerates the maturation of procaspase-3 by upstream activator proteases during apoptosis.
EMBO J.
1999;18:2049-2056[Medline]
[Order article via Infotrieve].
20.
Wyllie AH, Kerr JFR, Currie AR.
Cell death: the significance of apoptosis.
Int Rev Cytol.
1980;68:251-306.
21.
Cohen GM, Sun XM, Fearnhead H, et al.
Formation of large molecular weight fragments of DNA is a key committed step of apoptosis in thymocytes.
J Immunol.
1994;153:507-516[Abstract].
22.
Scaffidi C, Kirchhoff S, Krammer PH, Peter ME.
Apoptosis signaling in lymphocytes.
Curr Opin Immunol.
1999;11:277-285[Medline]
[Order article via Infotrieve].
23.
Bossy-Wetzel E, Newmeyer DD, Green DR.
Mitochondrial cytochrome c release in apoptosis occurs upstream of DEVD-specific caspase activation and independently of mitochondrial transmembrane depolarization.
EMBO J.
1998;17:37-49[Medline]
[Order article via Infotrieve].
24.
Tewari M, Quan LT, O'Rourke K, et al.
Yama/CPP32 , a mammalian homolog of CED-3, is a crmA-inhibitable protease that cleaves the death substrate poly (ADP-ribose) polymerase.
Cell.
1995;81:801-809[Medline]
[Order article via Infotrieve].
25.
Li H, Zhu H, Xu CJ, Yuan J.
Cleavage of BID by caspase 8 mediates the mitochondrial damage in the Fas death pathway of apoptosis.
Cell.
1998;94:491-501[Medline]
[Order article via Infotrieve].
26.
Luo X, Budiharjo I, Zou H, Slaughter C, Wang X.
Bid, a Bcl2 interacting protein, mediates cytochrome c release from mitochondria in response to activation of surface death receptors.
Cell.
1998;94:481-490[Medline]
[Order article via Infotrieve].
27.
Oberhammer FA, Wilson JW, Dive C, Morris ID, Hickman JA.
Apoptotic death in epithelial cells: cleavage of DNA to 300 and /or 50 kb fragments prior to or in the absence of internucleosomal fragmentation.
EMBO J.
1993;12:3679-3684[Medline]
[Order article via Infotrieve].
28.
Samejima K, Toné S, Kottke TJ, et al.
Transition from caspase-dependent to caspase-independent mechanisms at the onset of apoptosis execution.
J Cell Biol.
1998;143:225-239[Abstract/Free Full Text].
29.
Sakahira H, Enari M, Ohsawa Y, Uchiyama Y, Nagata S.
Apoptotic nuclear morphological change without DNA fragmentation.
Curr Biol.
1999;9:543-546[Medline]
[Order article via Infotrieve].
30.
Böhme H, Schrautemeier B.
Comparative characterization of ferredoxins from heterocysts and vegetative cells of Anabaena variabilis.
BBA.
1987;891:1-7.
31.
Vander Heiden MG, Chandel NS, Williamson EK, Schumacker PT, Thompson CB.
Bcl-XL regulates the membrane potential and volume homeostasis of mitochondria.
Cell.
1997;91:627-637[Medline]
[Order article via Infotrieve].
32.
Susin SA, Zamzani N, Castedo M, et al.
Bcl-2 inhibits the mitochondrial release of an apoptogenic protease.
J Exp Med.
1996;184:1331-1342[Abstract/Free Full Text].
33.
Pastorino JG, Tafani M, Rothman RJ, Marcineviciute A, Hoek JB, Farber JL.
Functional consequences of the sustained or transient activation of Bax on the mitochondrial permeability transition pore.
J Biol Chem.
1999;274:31734-31739[Abstract/Free Full Text].
34.
Jürgensmeier JM, Xie Z, Devereaux Q, Ellerby L, Bredesen D, Reed JC.
Bax directly induces release of cytochrome c from isolated mitochondria.
Proc Natl Acad Sci U S A.
1998;95:4997-5002[Abstract/Free Full Text].
35.
Narita M, Shimizu S, Ito T, et al.
Bax interact with the permeability transition pore to induce permeability transition and cytochrome c release in isolated mitochondria.
Proc Natl Acad Sci U S A.
1998;95:14681-14686[Abstract/Free Full Text].
36.
Marzo I, Brenner C, Zamzami N, et al.
Bax and adenine nucleotide translocator cooperate in the mitochondrial control of apoptosis.
Science.
1998;281:2027-2031[Abstract/Free Full Text].
37.
Shimizu S, Narita M, Tsujimoto Y.
Bcl-2 family proteins regulate the release of apoptogenic cytochrome c by the mitochondrial channel VDAC.
Nature.
1999;399:483-487[Medline]
[Order article via Infotrieve].
38.
Wolter KG, Hsu YT, Smith CL, Nechustan A, Xi XG, Youle RJ.
Movement of Bax from the cytosol to mitochondria during apoptosis.
J Cell Biol.
1997;139:1281-1292[Abstract/Free Full Text].
39.
Bernardi P, Scorrano L, Colonna R, Petronilli V, Di Lisa F.
Mitochondria and cell death: mechanistic aspects and methodological issues.
Eur J Biochem.
1999;264:687-701[Medline]
[Order article via Infotrieve].
40.
Liu X, Pi P, Widlak P, et al.
The 40-kDa subunit of DNA fragmentation factor induces DNA fragmentation and chromatin condensation during apoptosis.
Proc Natl Acad Sci U S A.
1998;95:8461-8466[Abstract/Free Full Text].
41.
Takahashi A, Alnemri ES, Lazebnik YA, et al.
Cleavage of lamin A by Mch2 but not CPP32: multiple interleukin 1-converting enzyme-related proteases with distinct substrate recognition properties are active in apoptosis.
Proc Natl Acad Sci U S A.
1996;93:8395-8400[Abstract/Free Full Text].
42.
Sahara S, Aoto M, Eguchi Y, Imamoto N, Yoneda Y, Tsujimoto Y.
Acinus is a caspase-3 activated protein required for apoptotic chromatin condensation.
Nature.
1999;401:168-172[Medline]
[Order article via Infotrieve].
43.
Yu SP, Yeh CH, Sensi SL, et al.
Mediation of neuronal apoptosis by enhancement of outward potassium current.
Science.
1997;278:114-117[Abstract/Free Full Text].
44.
Dallaporta B, Hirsch T, Susin SA, et al.
Potassium leakage during the apoptotic degradation phase.
J Immunol.
1998;160:5605-5615[Abstract/Free Full Text].
45.
Bortner CD, Cidlowski JA.
Caspase independent/dependent regulation of K+, cell shrinkage, and mitochondrial membrane potential during lymphocyte apoptosis.
J Biol Chem.
1999;274:21953-21962[Abstract/Free Full Text].
46.
Chen Q, Takeyama N, Brady G, Watson AJM, Vive C.
Blood cells with reduced mitochondrial membrane potential and cytosolic cytochrome c can survive and maintain clonogenicity given appropriate signals to suppress apoptosis.
Blood.
1998;92:4545-4553[Abstract/Free Full Text].
47.
Martinou I, Desagher S, Eskes R, et al.
The release of cytochrome c from mitochondria during apoptosis of NGF-deprived sympathetic neurons is a reversible event.
J Cell Biol.
1999;144:883-889[Abstract/Free Full Text].
48.
Kerr KM, Suleiman MS, Halestrap AP.
Reversal of permeability transition during recovery of hearts from ischemia and its enhancement by pyruvate.
Am J Physiol.
1999;276:496-502.
49.
Minamikawa T, Williams DA, Bowser DN, Nagley P.
Mitochondrial permeability transition and swelling can occur reversibly without inducing cell death in intact human cells.
Exp Cell Res.
1999;246:26-37[Medline]
[Order article via Infotrieve].
50.
De Maria R, Lenti L, Malisan F, et al.
Requirement for GD3 ganglioside in CD95- and ceramide-induced apoptosis.
Science.
1997;277:1652-1655[Abstract/Free Full Text].
51.
Kawahara A, Ohsawa Y, Matsumura H, Uchiyama Y, Nagata S.
Caspase-independent cell killing by Fas-associated protein with death domain.
J Cell Biol.
1998;143:1353-1360[Abstract/Free Full Text].
52.
Gandhi RT, Chen BK, Straus SE, Dale JK, Lenardo MJ, Baltimore D.
HIV-1 directly kills by a Fas-independent mechanism.
J Exp Med.
1998;187:1113-1122[Abstract/Free Full Text].
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Commitment to Apoptosis in CD4+ T Lymphocytes Productively Infected with Human Immunodeficiency Virus Type 1 Is Initiated by Lysosomal Membrane Permeabilization, Itself Induced by the Isolated Expression of the Viral Protein Nef
J. Virol.,
October 15, 2007;
81(20):
11426 - 11440.
[Abstract]
[Full Text]
[PDF]
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C. Munoz-Pinedo, A. Guio-Carrion, J. C. Goldstein, P. Fitzgerald, D. D. Newmeyer, and D. R. Green
Different mitochondrial intermembrane space proteins are released during apoptosis in a manner that is coordinately initiated but can vary in duration
PNAS,
August 1, 2006;
103(31):
11573 - 11578.
[Abstract]
[Full Text]
[PDF]
|
|
|
|
|
|
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M. Laforge, N. Bidere, S. Carmona, A. Devocelle, B. Charpentier, and A. Senik
Apoptotic Death Concurrent with CD3 Stimulation in Primary Human CD8+ T Lymphocytes: A Role for Endogenous Granzyme B
J. Immunol.,
April 1, 2006;
176(7):
3966 - 3977.
[Abstract]
[Full Text]
[PDF]
|
|
|
|
|
|
|
S.-C. Chen, C.-C. Huang, C.-L. Chien, C.-J. Jeng, H.-T. Su, E. Chiang, M.-R. Liu, C. H. H. Wu, C.-N. Chang, and R.-H. Lin
Cross-linking of P-selectin glycoprotein ligand-1 induces death of activated T cells
Blood,
November 15, 2004;
104(10):
3233 - 3242.
[Abstract]
[Full Text]
[PDF]
|
|
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J. Fombonne, S. Reix, R. Rasolonjanahary, E. Danty, S. Thirion, G. Laforge-Anglade, O. Bosler, P. Mehlen, A. Enjalbert, and S. Krantic
Epidermal Growth Factor Triggers an Original, Caspase-independent Pituitary Cell Death with Heterogeneous Phenotype
Mol. Biol. Cell,
November 1, 2004;
15(11):
4938 - 4948.
[Abstract]
[Full Text]
[PDF]
|
|
|
|
|
|
|
T. Decker, M. Oelsner, R. J. Kreitman, G. Salvatore, Q.-c. Wang, I. Pastan, C. Peschel, and T. Licht
Induction of caspase-dependent programmed cell death in B-cell chronic lymphocytic leukemia by anti-CD22 immunotoxins
Blood,
April 1, 2004;
103(7):
2718 - 2726.
[Abstract]
[Full Text]
[PDF]
|
|
|
|
|
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M. Okada, S. Adachi, T. Imai, K.-i. Watanabe, S.-y. Toyokuni, M. Ueno, A. S. Zervos, G. Kroemer, and T. Nakahata
A novel mechanism for imatinib mesylate-induced cell death of BCR-ABL-positive human leukemic cells: caspase-independent, necrosis-like programmed cell death mediated by serine protease activity
Blood,
March 15, 2004;
103(6):
2299 - 2307.
[Abstract]
[Full Text]
[PDF]
|
|
|
|
|
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S. Nikiforow, K. Bottomly, G. Miller, and C. Munz
Cytolytic CD4+-T-Cell Clones Reactive to EBNA1 Inhibit Epstein-Barr Virus-Induced B-Cell Proliferation
J. Virol.,
November 15, 2003;
77(22):
12088 - 12104.
[Abstract]
[Full Text]
[PDF]
|
|
|
|
|
|
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N. Bidere, H. K. Lorenzo, S. Carmona, M. Laforge, F. Harper, C. Dumont, and A. Senik
Cathepsin D Triggers Bax Activation, Resulting in Selective Apoptosis-inducing Factor (AIF) Relocation in T Lymphocytes Entering the Early Commitment Phase to Apoptosis
J. Biol. Chem.,
August 15, 2003;
278(33):
31401 - 31411.
[Abstract]
[Full Text]
[PDF]
|
|
|
|
|
|
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C. Zhang, J. Baffi, S. W. Cousins, and K. G. Csaky
Oxidant-induced cell death in retinal pigment epithelium cells mediated through the release of apoptosis-inducing factor
J. Cell Sci.,
May 15, 2003;
116(10):
1915 - 1923.
[Abstract]
[Full Text]
[PDF]
|
|
|
|
|
|
|
G Thuret, C Chiquet, S Herrag, J-M Dumollard, D Boudard, J Bednarz, L Campos, and P Gain
Mechanisms of staurosporine induced apoptosis in a human corneal endothelial cell line
Br. J. Ophthalmol.,
March 1, 2003;
87(3):
346 - 352.
[Abstract]
[Full Text]
[PDF]
|
|
|
|
|
|
|
C.-Y. Liu, A. Takemasa, W. C. Liles, R. B. Goodman, M. Jonas, H. Rosen, E. Chi, R. K. Winn, J. M. Harlan, and P. I. Chuang
Broad-spectrum caspase inhibition paradoxically augments cell death in TNF-alpha -stimulated neutrophils
Blood,
January 1, 2003;
101(1):
295 - 304.
[Abstract]
[Full Text]
[PDF]
|
|
|
|
|
|
|
V. Mateo, E. J. Brown, G. Biron, M. Rubio, A. Fischer, F. L. Deist, and M. Sarfati
Mechanisms of CD47-induced caspase-independent cell death in normal and leukemic cells: link between phosphatidylserine exposure and cytoskeleton organization
Blood,
September 26, 2002;
100(8):
2882 - 2890.
[Abstract]
[Full Text]
[PDF]
|
|
|
|
|
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F. Li, H. P. Mao, K. L. Ruchalski, Y. H. Wang, W. Choy, J. H. Schwartz, and S. C. Borkan
Heat stress prevents mitochondrial injury in ATP-depleted renal epithelial cells
Am J Physiol Cell Physiol,
September 1, 2002;
283(3):
C917 - C926.
[Abstract]
[Full Text]
[PDF]
|
|
|
|
|
|
|
N. Bidere, M. Briet, A. Durrbach, C. Dumont, J. Feldmann, B. Charpentier, G. de Saint-Basile, and A. Senik
Selective Inhibition of Dipeptidyl Peptidase I, Not Caspases, Prevents the Partial Processing of Procaspase-3 in CD3-activated Human CD8+ T Lymphocytes
J. Biol. Chem.,
August 23, 2002;
277(35):
32339 - 32347.
[Abstract]
[Full Text]
[PDF]
|
|
|
|
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|
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H. Ferry-Dumazet, O. Garnier, M. Mamani-Matsuda, J. Vercauteren, F. Belloc, C. Billiard, M. Dupouy, D. Thiolat, J. P. Kolb, G. Marit, et al.
Resveratrol inhibits the growth and induces the apoptosis of both normal and leukemic hematopoietic cells
Carcinogenesis,
August 1, 2002;
23(8):
1327 - 1333.
[Abstract]
[Full Text]
[PDF]
|
|
|
|
|
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J.-H. Lai, L.-J. Ho, K.-C. Lu, D.-M. Chang, M.-F. Shaio, and S.-H. Han
Western and Chinese Antirheumatic Drug-Induced T Cell Apoptotic DNA Damage Uses Different Caspase Cascades and Is Independent of Fas/Fas Ligand Interaction
J. Immunol.,
June 1, 2001;
166(11):
6914 - 6924.
[Abstract]
[Full Text]
[PDF]
|
|
|
|
|
|
|
F. Petit, D. Arnoult, J.-D. Lelievre, L. M.-d. Parseval, A. J. Hance, P. Schneider, J. Corbeil, J. C. Ameisen, and J. Estaquier
Productive HIV-1 Infection of Primary CD4+ T Cells Induces Mitochondrial Membrane Permeabilization Leading to a Caspase-independent Cell Death
J. Biol. Chem.,
January 4, 2002;
277(2):
1477 - 1487.
[Abstract]
[Full Text]
[PDF]
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Top
Abstract
Introduction