1 Interdisciplinary Center for Clinical Research (IZKF), Westphalian
Wilhelms-University, D-48149 Münster, Germany
2 Department of Pharmacology and Toxicology, Westphalian Wilhelms-University,
D-48149 Münster, Germany
* Author for correspondence (e-mail: prehn{at}uni-muenster.de)
Accepted 24 October 2002
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Summary |
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Key words: Apoptosis, Mitochondrial membrane potential, Plasma membrane potential, Confocal imaging, Mitochondrial respiration
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Introduction |
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Confocal time-lapse imaging experiments in cells expressing a
cyt-C-green-fluorescent-protein (GFP) fusion protein have suggested that the
majority of cyt-C is released rapidly and efficiently during apoptosis
(Heiskanen et al., 1999), a
process that occurs in one large step
(Goldstein et al., 2000
) or
two or more steps (Luetjens et al.,
2001
; Scorrano et al.,
2002
). A change in mitochondrial ultrastructure may be required to
guarantee a complete release of cyt-C, because large quantities may normally
reside in the inaccessible intracristal space
(Luetjens et al., 2001
;
Scorrano et al., 2002
).
Although many studies have focused on the role of cyt-C release in the
activation of the caspase cascade, loss of large quantities of cyt-C will also
directly affect mitochondrial respiration, ATP production and free radical
production, resulting in an organelle dysfunction program
(Adachi et al., 1997
;
Cai and Jones, 1998
;
Luetjens et al., 2000
;
Mootha et al., 2001
).
Single-cell imaging experiments have also shown that the mitochondrial
membrane potential (
M) depolarizes after or concomitant
with the release of a cyt-C-GFP fusion protein
(Heiskanen et al., 1999
;
Waterhouse et al., 2001
), a
process that has been described as caspase dependent
(Bossy-Wetzel et al., 1998
;
Waterhouse et al., 2001
).
However, we and others have previously demonstrated that mitochondria are able
to transiently maintain a membrane potential after the release of cyt-C
(Bossy-Wetzel et al., 1998
;
Deshmukh et al., 2000
;
Krohn et al., 1999
), even in
the absence of caspase inhibitors
(Bossy-Wetzel et al., 1998
;
Krohn et al., 1999
). However,
interpretations of fluorescence changes of voltage-sensitive probes used in
these type of studies are hampered by the fact that these probes are also
sensitive to changes in plasma membrane potential (
P)
(Ehrenberg et al., 1988
;
Ward et al., 2000
).
Mitochondrial depolarization, loss of ATP and increased free radical formation
could indeed lead to a disturbance of cellular ion homeostasis and subsequent
P depolarization. In the present study, we therefore
analyzed
M and
P changes at the
single-cell level during apoptosis employing breast carcinoma and HeLa cells
expressing a cyt-C-GFP fusion protein.
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Materials and Methods |
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Cell culture and transfection
Generation and characterization of human breast carcinoma MCF-7 cells
stably expressing caspase-3 (MCF-7/Casp-3) and a cyt-C-GFP fusion protein have
been described (Luetjens et al.,
2001). We have previously shown that cyt-C-GFP is imported into
mitochondria and co-released with endogenous cyt-C after selective outer
membrane permeabilization with digitonin. Subcellular fractionation
experiments confirmed the concomitant release of endogenous cyt-C and
cyt-C-GFP from mitochondria during apoptosis. Generation and characterization
of HeLa D98 cells expressing the cyt-C-GFP fusion protein was performed as
described for the MCF-7/Casp-3 cells. For detection of effector caspase
activation by fluorescence resonance energy transfer (FRET) analysis,
MCF-7/Casp-3 or HeLa D98 cells were transiently transfected with plasmid
pmyc-CFP-DEVD-YFP DNA (0.6 µg) (Rehm et
al., 2002
; Tyas et al.,
2000
) and 6 µl LipofectaminTM reagent (Life Technologies)
per ml serum-free RPMI medium. Cells were cultivated on 35 mm glass-bottom
culture dishes (Willco BV, Amsterdam, The Netherlands).
Time-lapse confocal fluorescence microscopy
Cyt-C-GFP, TMRM and Dibac4(3) fluorescence was monitored and
quantified confocally using an inverted Olympus IX70 microscope attached to a
confocal laser scanning unit equipped with a 488 nm argon laser and a
60x oil fluorescence objective (Fluoview; Olympus, Hamburg, Germany).
Fluorescence transmitted the first dicroic mirror with more than 90%
transmission above 505 nm was divided with a second dichroic mirror at 550 nm
and detected after transmission of a 510-540 nm bandpass filter [GFP or
Dibac4(3)] or a 565 nm high pass emission filter (TMRM). There was
no TMRM fluorescence detectable in the GFP or Dibac4(3) channel.
The crosstalk between the average pixel intensity of GFP in the TMRM channel
was less than 10% of the average pixel intensity in the GFP channel. The
maximum change owing to GFP fluorescence in the TMRM channel occurred during
the release of the cyt-C-GFP fusion protein. The resulting change was within
the standard deviation of the average pixel intensity of single cells in the
TMRM channel (approx. 5%). Fluorescence was detected from a 0.7 µm thick
confocal section (full width half maximum).
The membrane-permeant, cationic probe TMRM distributes across cellular
membranes according to the Nernst equation and accumulates in the negatively
charged mitochondrial matrix (Ehrenberg et
al., 1988). TMRM has little effects on the respiratory chain
activity at the concentration used in the present study (30 nM), and has a
lower membrane-bound fraction than TMRE
(Scaduto et al., 1999
).
Dissipation of the inner mitochondrial membrane H+ gradient using
10 µM FCCP resulted in a transient peak in the TMRM fluorescence intensity
in less than 50% of control MCF-7/Casp-3 cells, suggesting that TMRM was used
below the quenching limit (Ward et al.,
2000
). Saturation of mitochondrial TMRM fluorescence was reached
at 250 nM extracellular probe concentration. For
P
measurements, we used the anionic probe Dibac4(3). The Nernstian
behaviour of this probe allows measurements of slow
P
changes by confocal microscopy (Dall'Asta
et al., 1997
). Dibac4(3) fluorescence was detected with
the 510-540 nm bandpass filter and showed a negligible overlap into the TMRM
channel.
For time-lapse imaging, culture dishes were mounted onto the microscope
stage that was equipped with a temperature-controlled inlay (HT200,
Minitüb, Tiefenbach, Germany). In control experiments constant
fluorescence values were monitored for 24 hours in case of cyt-C-GFP, 18 hours
in the case of TMRM and 12 hours in the case of Dibac4(3). For
induction of apoptosis, cells were incubated with 100 ng/ml TNF- plus 1
µg/ml CHX or 3 µM staurosporine (STS) directly on the stage after 1 hour
of equilibration with 30 nM TMRM and subsequent equilibration with 1 µM
Dibac4(3) for 1 hour when indicated. The medium was enriched with
10 mM HEPES (pH 7.4) and thoroughly mixed to ensure a proper distribution of
the drugs. To prevent evaporation the media was covered with embryo-tested
paraffin oil. Dibac4(3) measurements were carried out in serum-free
RPMI buffered with 10 mM HEPES because of the extensive binding of this probe
to proteins. Image data were obtained using Fluoview 2.0 software (Olympus)
and Kalman filtered from three scans for each image.
Image processing and remodeling of TMRM fluorescence kinetics
The quantitative analysis of the fluorescence images was performed using
the UTHSCSA ImageTool program (developed at the University of Texas Health
Science Center at San Antonio, Texas and available from the internet by
anonymous FTP from
maxrad6.uthscsa.edu).
For analysis of TMRM uptake in single cells, the fluorescent mitochondrial
regions were segmented from the non-fluorescent regions. After background
subtraction the average fluorescence intensity per pixel was calculated. The
release kinetics of cyt-C are shown as standard deviation (s.d.) from the
average pixel intensity of individual cells. Compartmentalized cyt-C-GFP
contributes to a high s.d. of pixel intensities, and homogeneously distributed
cyt-C-GFP is represented by a low s.d. Data are presented as a percentage of
the average of 10 values detected before release of cyt-C-GFP.
The TMRM kinetics from individual cells were fitted with the sigmoidal
Boltzmann equation:
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Remodeling of TMRM kinetics was performed according to Ward et al.
(Ward et al., 2000) under the
assumption of the Nernstian behavior of the dye. Briefly, the TMRM
concentration in mitochondria at a given extracellular concentration results
from both the
M and the
P.
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In these equations, T represents the temperature in °K,
F is the Faraday and R the Rydberg constant. The remodeling
takes the mitochondrial volume fraction (5%), the diffusion constant of TMRM
across cellular membranes (0.01 per second) and the quenching limit of TMRM
(700 µM) into account. Values were calculated as described previously
(Poppe et al., 2001).
Epifluorescence microscopy
CFP-DEVD-YFP-expressing cells equilibrated with 30 nM TMRM were treated
with STS or STS plus z-VAD-fmk and then placed in a heated (37°C) chamber
(Minitüb) mounted on a Nikon TE 300 microscope stage (Nikon,
Düsseldorf, Germany). Fluorescence was observed using a 20x S-Fluor
objective. The microscope was equipped with a polychroic mirror and
filterwheels in the excitation and emission light path containing the
appropriate filter sets (polychroic mirror with more than 90% reflexion from
411 to 438 nm, between 491 and 506 nm, and between 593 and 627 nm; CFP:
excitation 436±10 nm, emission 480±20 nm; YFP: excitation
500±20 nm, emission 535±30 nm; FRET: excitation 436±10
nm, emission 535±30 nm; TMRM: excitation 500±20 nm, emission 570
nm long pass filter; AHF Analysentechnik, Tübingen, Germany). Images were
recorded using a CCD camera (Visicam, Visitron Systems, Puchheim, Germany).
The imaging setup was controlled by MetaMorph software (Universal Imaging,
West Chester, PA). During control experiments bleaching of the probes was
negligible.
Kinetics of FRET disruption
Images were processed using MetaMorph software (Universal Imaging, West
Chester, PA). CFP/YFP emission ratios were obtained by dividing the integrated
fluorescence intensity values of single cells
(Rehm et al., 2002). To
compare individual cells, time courses of the emission ratios were scaled by
defining the baseline ratio before the onset of FRET disruption as one.
Changes in TMRM uptake of individual cells were monitored by total intensity
in the TMRM-sensitive channel after subtraction of the YFP overlap. The
baseline of total intensity was defined as 100% in cells 1 hour after the
equilibration with 30 nM TMRM.
Preparation of whole cell extracts and western blotting
Cells were collected at 200 g for 5 minutes and washed with
phosphate-buffered saline (PBS). The cell pellet was resuspended in lysis
buffer [62.5 mM Tris HCl pH 6.8, 10% (v/v) glycerin, 2% (w/v) sodium dodecyl
sulfate (SDS), 1 mM phenylmethylsulfonyl fluoride (PMSF), 1 µg/ml pepstatin
A, 1 µg/ml leupeptin and 5 µg/ml aprotinin]. Cell homogenates were
centrifuged at 15,000 g and 4°C for 15 minutes. Protein
content was determined with the Pierce Micro-BCA Protein Assay (KMF, Cologne,
Germany). An equal amount of protein (20 or 40 µg) was loaded onto
SDS-polyacrylamide gels. Proteins were separated at 120 V for 1.5 hours and
then blotted to nitrocellulose membranes (Protean BA 83; 2 µm; Schleicher
& Schuell, Dassel, Germany) in transfer buffer [25 mM Tris, 192 mM
glycine, 20% methanol (v/v) and 0.01% SDS]. The blots were blocked with 5%
non-fat dry milk in TBST (15 mM Tris-HCl pH 7.5, 200 mM NaCl and 0.1%
Tween-20) at room temperature for 2 hours. Membranes were incubated with a
mouse monoclonal anti-Na+/K+ ATPase -1 subunit
antibody (clone C464.6, 1:2000, Upstate Biotechnology, Lake Placid, NY), a
mouse monoclonal anti-Na+/K+ ATPase ß-1 subunit
antibody (clone C464.8, 1:2000, Upstate Biotechnology), a rabbit polyclonal
anti-caspase-3 antibody (H-277, 1:1000, Santa Cruz Biotechnology, CA), a mouse
monoclonal anti-poly(ADP ribose) polymerase (PARP) antibody (clone C2-10,
1:2000, Pharmingen Becton Dickinson, Hamburg, Germany), or a mouse monoclonal
anti-
-tubulin antibody (clone DM 1A; 1:5000, Sigma, Missouri).
Membranes were washed with TBST six times for 10 minutes and incubated with
anti-mouse or anti-rabbit peroxidase-conjugated secondary antibodies (Promega,
Madison, WI) for one hour. Blots were washed and developed using the ECL
chemiluminescence detection reagent (Amersham Pharmacia, Buckinghamshire, UK).
Membranes were stripped in standard stripping buffer (2% SDS, 62.5 mM
Tris-HCl, 100 mM 2-mercaptoethanol, pH 6.8) at 60°C for 20 minutes, washed
twice in TBST for 10 minutes and reprobed.
Data analysis and statistics
For quantification of changes in TMRM and Dibac4(3) uptake, all
cells within a given field were analyzed that (a) demonstrated cyt-C-GFP
release or TMRM fluorescence loss and (b) could be monitored for at least
60-90 minutes afterward. Data are given as means±s.e.m. For statistical
comparison, ANOVA and subsequent Tukey test were employed (SPSS 10.0, SPSS
Inc., Chicago, USA). P values smaller than 0.05 were considered to be
statistically significant.
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Results |
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The decrease in mitochondrial TMRM uptake during apoptosis is not
sensitive to a broad-spectrum caspase inhibitor
We subsequently addressed the question of whether the decrease in TMRM
uptake required the activation of caspases. MCF-7/Casp-3 cells expressing
cyt-C-GFP were treated simultaneously with 3 µM STS and 200 µM of the
broad-spectrum caspase inhibitor z-VAD-fmk. Although the onset of the
cyt-C-GFP release was delayed in cells treated with STS plus z-VAD-fmk (STS:
206±11 minutes, STS+zVAD-fmk: 311±32 minutes,
n=16 cells each in eight separate experiments, P<0.05),
the individual release kinetics were not altered by caspase inhibition
(Fig. 2A,B) (Goldstein et al., 2000;
Luetjens et al., 2001
). We
could also detect a decrease in mitochondrial TMRM uptake in z-VAD-fmk-treated
cells after the release of cyt-C-GFP. Similar to the results obtained in cells
treated with STS alone, the depolarization reached a steady-state level after
60 minutes (Fig. 2B,C).
However, the average TMRM intensity at this steady state was significantly
higher than in cells treated with STS alone (see also
Fig. 5D). We could not detect
any recovery of TMRM uptake in the presence of z-VAD-fmk up to 90 minutes
after the release of the cyt-C-GFP fusion protein
(Fig. 2B,C). In select
experiments, cells were monitored up to 6 hours after cyt-C-GFP release and
also revealed no recovery of
M-cyt-C in the
presence of z-VAD-fmk. By contrast, z-VAD-fmk potently protected cells that
had released cyt-C-GFP from apoptotic morphological changes induced by STS,
including cell shrinkage and blebbing (Fig.
2D,E). Treatment with z-VAD-fmk also completely inhibited the
processing of caspase-9 as well as caspase-3 and 7 into active subunits as
detected by western blot analysis (data not shown). These findings were not
confined to MCF-7/Casp-3 cells, since a z-VAD-fmk-independent decrease in
mitochondrial TMRM uptake was also observed in HeLa D98 cells stably
transfected with cyt-C-GFP (Fig.
2F).
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Effect of FCCP or oligomycin on the steady-state level of
M after the release of cyt-C-GFP
STS-treated cells that had released cyt-C-GFP showed a significantly
reduced, but stable, TMRM uptake, suggesting that cells had established a new
M (
M-cyt-C).
Treatment with the protonophore FCCP was able to further dissipate
mitochondrial TMRM uptake, both in the absence
(Fig. 3A) or presence (data not
shown) of 200 µM z-VAD-fmk. The ability of mitochondria to maintain
M-cyt-C after outer mitochondrial membrane
permeabilization could be due to a reversal of the
FOF1-ATP-synthase, consuming ATP and thereby generating
M-cyt-C. We treated cells with the
FOF1-ATP-synthase inhibitor oligomycin (5 µM) 60
minutes after the addition of STS but prior to the release of the cyt-C-GFP
fusion protein. In these cells, the release of cyt-C-GFP was always followed
by a decrease in mitochondrial TMRM uptake to background levels within a 60
minute time period (Fig. 3B-D).
Interestingly, cellular necrosis frequently followed the total decrease in
TMRM fluorescence, as indicated by a sudden decrease in the GFP fluorescence
intensity (Fig. 3B, indicated
by arrows). The average time to cellular necrosis was 83±14 minutes
after the cyt-C-GFP release. Addition of oligomycin to cells that already had
released cyt-C-GFP and maintained a stable
M-cyt-C led to a rapid and complete
dissipation of TMRM uptake (Fig.
3E).
M depolarization during apoptosis is
synchronized to a z-VAD-fmk-sensitive
P
depolarization
P depolarization has been reported to occur during
apoptosis (Bortner and Cidlowski,
1999
; Bortner et al.,
2001
), but its relation to outer mitochondrial membrane
permeabilization or
M depolarization during apoptosis
is unclear. We analyzed changes in
M and
P during apoptosis by simultaneous confocal imaging of
TMRM and Dibac4(3) uptake in MCF-7/Casp-3 cells. Uptake of the
negatively charged Dibac4(3) and subsequent reversible binding to
cytoplasmic proteins can be detected in response to depolarization of
P (Dall'Asta et al.,
1997
). Control experiments revealed that
M
depolarization with 10 nM valinomycin
(Poppe et al., 2001
;
Rottenberg and Wu, 1998
)
caused a decrease in TMRM uptake in MCF-7/Casp-3 cells, whereas
Dibac4(3) fluorescence remained stable
(Fig. 4A). Subsequent
P depolarization with the
Na+/K+-ATPase inhibitor ouabain (100 µg/ml) increased
cellular Dibac4(3) uptake (Fig.
4A). Owing to the overlap of Dibac4(3) and GFP
fluorescence emission spectra, the temporal relationship between cyt-C-GFP
release and
P depolarization could not be directly
monitored. However, this relationship could be indirectly monitored by the
kinetics of TMRM uptake. Cells were treated with 3 µM STS after
equilibration with TMRM and Dibac4(3). At different time after
addition of STS, individual cells showed a decrease in the TMRM fluorescence
signal and a simultaneous increase in the Dibac4(3) fluorescence
(Fig. 4B). These changes were
always coupled. To calculate the average kinetics of the TMRM and
Dibac4(3) fluorescence changes, the onset of TMRM fluorescence loss
was set to time zero (Fig. 4C).
The increase in Dibac4(3) fluorescence was completed 50 minutes
after the onset of the TMRM fluorescence changes.
|
We subsequently examined the effect of caspase inhibition on
P depolarization. After equilibration with
Dibac4(3) and TMRM, MCF-7/Casp-3 cells were treated with 3 µM
STS plus 200 µM z-VAD-fmk. In agreement with the data above
(Fig. 2), we also observed a
decrease in TMRM fluorescence in the presence of z-VAD-fmk, indicating the
onset of
M depolarization in the individual cells
(Fig. 4D,E). However, a
concomitant
P depolarization could not be detected. In
intact cells,
P is largely maintained by the activity
of the Na+/K+-ATPase. Immunoblot analysis of whole cell
extracts of MCF-7/Casp-3 cells treated with 3 µM STS showed a significant
degradation of the regulatory ß-subunit of
Na+/K+-ATPase. The degradation correlated with the
accumulation of the caspase-derived 85 kDa cleavage product of PARP
(Fig. 4F), and the processing
of procaspase-3 into active subunits (Fig.
4G). By contrast, a degradation of the
-subunit of
Na+/K+-ATPase could not be detected. The degradation of
the ß-subunit of Na+/K+-ATPase and the activation
of procaspase-3 were potently blocked in cells treated with STS plus z-VAD-fmk
(Fig. 4G).
Simulation of TMRM fluorescence changes due to outer mitochondrial
membrane permeabilization in a virtual cell
To evaluate quantitatively the contribution of M and
P to the changes in TMRM fluorescence after outer
mitochondrial membrane permeabilization, we employed a method introduced by
Ward et al. (Ward et al.,
2000
). In this approach, TMRM fluorescence changes are calculated
on the basis of Nernst calculations of fluorescence in the extracellular,
cytoplasmic and mitochondrial compartments, taking the mitochondrial volume
fraction, diffusion constant of TMRM across cellular membranes and the
quenching limit of TMRM into account (see Materials and Methods). Similar to
our previously reported findings in STS-treated hippocampal neurons and
medulloblastoma cells (Poppe et al.,
2001
), we observed that
M hyperpolarized
from -150 mV to -160 mV in the MCF-7/Casp-3 cells within 2 hours of addition
of STS (data not shown). The remodeling therefore starts with
M=-160 mV and
P=-70 mV. The
increase in the Dibac4(3) intensity from 100±3% to
182±20% indicated a depolarization of
P to
-58±4 mV calculated with the Nernstian equation
(Dall'Asta et al., 1997
). In
STS-treated cells, we obtained a good fit to our experimentally determined
TMRM kinetics, remodeling a
P depolarization of 10 mV
(t1/2=30 minutes) and a
M depolarization of
32 mV (t1/2=30 minutes) (Fig.
5A). The mitochondrial TMRM fluorescence in a virtual cell treated
with STS and z-VAD-fmk was remodeled with a
M
depolarization of 32 mV (t1/2=17 minutes) in the absence of
P depolarization
(Fig. 5B). The remodeled TMRM
signal after the depolarization of
M equilibrates to a
value of 30% in a virtual cell treated with STS plus z-VAD-fmk, compared to
the value of 33.9±5.5% that we determined experimentally in our
confocal imaging experiments (Fig.
5D). The remodeled equilibrated TMRM signal of a virtual cell
treated with STS alone reached a value of 21.2% of the initial fluorescence,
compared to the experimentally determined value of 20.9±2.9%
(Fig. 5D). Finally, the effect
of oligomycin was successfully remodeled assuming a
M
depolarization of 65 mV (t1/2=30 minutes) and a
P depolarization of 10 mV (t1/2=30 minutes)
as detected with Dibac4(3) (data not shown)
(Fig. 5C,D).
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Discussion |
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M depolarization during apoptosis
Cyt-C transports electrons between complex III and cytochrome C oxidase.
Outer mitochondrial membrane permeabilization and subsequent release of large
quantities of cyt-C will slow down this electron flow, leading to an
impairment of mitochondrial respiratory chain activity and a reduced
H+ export from the mitochondrial matrix. Indeed,
M depolarization is considered a hallmark of apoptosis,
and ATP levels are known to decrease considerably during this process
(Adachi et al., 1997
;
Cai and Jones, 1998
;
Lemasters et al., 1999
;
Mootha et al., 2001
). In
isolated mitochondria, re-addition of small amounts of cyt-C restored
M and ATP production after outer mitochondrial membrane
permeabilization (Mootha et al.,
2001
), suggesting a direct role for the loss of cyt-C in these
bioenergetic alterations.
M depolarization can be
observed very shortly after the quantitative release of a cyt-C-GFP fusion
protein (Fig. 1)
(Heiskanen et al., 1999
;
Waterhouse et al., 2001
). The
majority of cyt-C (85-90%) is believed to reside in the intracristal space and
may not be directly accessible for release
(Ott et al., 2002
;
Scorrano et al., 2002
). It has
been suggested that the complete release of cyt-C requires a reorganization of
cristae and that small amounts of cyt-C (but presumably also cyt-C-GFP) are
released prior to this (Luetjens et al.,
2001
; Scorrano et al.,
2002
). Of note,
M depolarization could only
be detected after the complete release of cyt-C-GFP, suggesting that an
earlier release of smaller quantities of cyt-C was not sufficient to trigger
M depolarization. It is possible that cyt-C-GFP is less
effectively associated with the inner membrane and is thus lost more readily
upon outer membrane permeabilization. However,
M
clearly depolarized rapidly after the release of the fusion protein. It is
also unlikely that cyt-C-GFP substitutes for the electron transport function
of endogenous cyt-C, since previous experiments have estimated that the
cyt-C-GFP fusion protein represents less than 0.5% of total cellular cyt-C
(Goldstein et al., 2000
;
Luetjens et al., 2001
).
Cyt-C-GFP can therefore be considered a reliable indicator of the functional
consequences of a complete outer mitochondrial membrane permeabilization and
the accompanying loss of endogenous cyt-C.
We could demonstrate that M stabilizes to a new
potential,
M-cyt-C, after the release of
cyt-C-GFP, even in the absence of caspase inhibitors. These results are
consistent with our previous observations in cultured rat hippocampal neurons,
which demonstrated that mitochondria could still be depolarized with FCCP at a
time point when nuclear condensation and fragmentation was clearly evident
(Krohn et al., 1999
). Previous
studies using isolated mitochondria and permeabilized hepatocytes have
demonstrated that the stabilization of
M after the
release of cyt-C is caused by FOF1-ATP-synthase
reversal, since treatment with the FOF1-ATP-synthase
inhibitor oligomycin induced a rapid
M depolarization
(Madesh et al., 2002
;
Polster et al., 2001
).
However, in both studies cytosolic cyt-C may have been washed out or diluted
in the extracellular buffer solution to an extent that the concentration of
cyt-C was too low to maintain mitochondrial respiration. Clearly, the
energetics in cells with an intact plasma membrane may differ considerably
from isolated mitochondria or permeabilized cells. Using an approach similar
to our study, Waterhouse and co-workers
(Waterhouse et al., 2001
)
proposed that diffusion of released cyt-C back into the mitochondrial
respiratory chain maintained a
M after outer
mitochondrial membrane permeabilization. Our study provides experimental and
theoretical evidence for the concept that
FOF1-ATP-synthase reversal maintains
M-cyt-C also in intact cells. Treatment with
oligomycin either prior to or after the release of the cyt-C-GFP fusion
protein resulted in a rapid and complete depolarization of
M, and the experimentally determined TMRM fluorescence
changes could be remodeled in a virtual cell assuming
FOF1-ATP-synthase reversal. Likewise, Rego et al.
(Rego et al., 2001
) could
convincingly remodel the kinetics of TMRM fluorescence changes induced by
oligomycin in STS-exposed neural cells, assuming that
FOF1-ATP-synthase reversal maintained
M-cyt-C, demonstrating the power of this
approach. We have also observed that the complex III inhibitor antimycin A
rapidly depolarized
M in intact cells to a level
similar to
M-cyt-C, but had no effect on
M-cyt-C in cells that had released cyt-C-GFP
(data not shown). There is also evidence that cyt-C may be released out of
apoptotic cells, hence limiting the availability of cyt-C for mitochondrial
respiration (Luetjens et al.,
2000
; Renz et al.,
2001
). It is therefore unlikely that the
M
of cells that have released cyt-C is maintained for significant time periods
by diffusion of cyt-C back to the mitochondrial inner membrane.
According to our simulation, the 80% decrease in average TMRM fluorescence
intensity after the release of cyt-C-GFP reflects a depolarization of 32 mV
assuming a M of -160 mV prior to the release. These
data suggest that FOF1-ATP-synthase reversal contributes
to the stabilization of
M-cyt-C at a
potential of -128 mV. Indeed, it has previously been shown that a
M depolarization of 10 mV with the complex III
inhibitor antimycin A was sufficient to trigger
FOF1-ATP-synthase reversal
(Ward et al., 2000
). However,
TMRM may also behave in a non-Nernstian manner owing to membrane and matrix
protein binding (Scaduto et al.,
1999
). Hence, quantification of
M
depolarization on the basis of TMRM uptake may lead to an overestimation of
M-cyt-C. The synthase activity is also
reduced with a decrease in
pH over the inner mitochondrial membrane
(Dimroth et al., 2000
;
Matsuyama et al., 2000
).
Increased lactate production and cytosolic acidification is likely to occur in
cells in which
M-cyt-C is maintained by
glycolytic ATP production. Both events have been shown to occur during
apoptosis (Matsuyama et al.,
2000
; Tiefenthaler et al.,
2001
). Interestingly, pH changes during apoptosis are sensitive to
Bcl-2 overexpression but independent of caspase activation
(Matsuyama et al., 2000
) (see
Discussion).
The switch from mitochondrial to glycolytic ATP production may be
facilitated in energized cells such as neurons or in tumor cells with higher
basal level of glycolysis. The energetic state of cells that have released
cyt-C resembles the state of cells depleted of mitochondrial DNA
(0 cells), which also lack mitochondrial respiration and
maintain their
M via
FOF1-ATP-synthase reversal. Of note, apoptosis in
0 cells is not significantly inhibited
(Jacobson et al., 1993
),
suggesting that glycolysis may provide sufficient amounts of ATP for the
execution of apoptosis. Of note, pretreatment with oligomycin not only leads
to a total breakdown of
M-cyt-C but also to
subsequent necrosis. It has previously been demonstrated that oligomycin
treatment is able to switch the cell death mode from apoptosis to necrosis
(Eguchi et al., 1997
;
Leist et al., 1997
). Our
results now demonstrate that cyt-C release is also important for the
occurrence and timing of necrosis in energetically deprived cells.
Role of caspases in M and
P depolarization
Simultaneous monitoring of (i) cyt-C-GFP and TMRM fluorescence changes,
(ii) DEVDase activity and TMRM uptake, and (iii) Dibac4(3) and TMRM
uptake in MCF-7/Casp-3 and HeLa D98 cells demonstrated that the depolarization
of M during apoptosis was insensitive to z-VAD-fmk.
Supporting the concept of FOF1-ATP-synthase reversal in
maintaining
M-cyt-C, we did not detect any
M recovery, even if the activation of the major
effector caspases was potently blocked with a broad spectrum caspase
inhibitor. Waterhouse and co-workers
(Waterhouse et al., 2001
)
previously reported that HeLa cells treated with z-VAD-fmk showed a rapid
recovery of
M depolarization after the release of
cyt-C-GFP. It is unlikely that these contrasting results are a consequence of
cell type differences, since we were also not able to detect a recovery of
M in z-VAD-fmk-treated HeLa cells, despite a complete
inhibition of DEVDase activity (see Results). In the present study, we could
demonstrate that
P depolarizes simultaneously to
M, but in a z-VAD-fmk-sensitive manner. We could also
demonstrate that
P depolarization contributed to the
resulting TMRM signal. The influence of
P
depolarization on the uptake of potential-sensitive cationic dyes critically
depends on the concentration and type of dye and is particularly pronounced
when concentrations are used that saturate mitochondrial uptake (`quenching
mode') (Ward et al., 2000
). In
the study of Waterhouse and co-workers
(Waterhouse et al., 2001
),
potential effects of
P depolarization on the uptake of
voltage-sensitive probes into mitochondria were not taken into account, and
experiments were performed with tetramethyl rhodamine ethyl-ester, which has a
significantly higher partition coefficient in mitochondrial membranes than
TMRM (Scaduto et al., 1999
).
Of note, our simulation of
M and
P changes in a virtual cell taking the
depolarization of
P detected with Dibac4(3)
into account confirmed our results detected with TMRM quantitatively.
Interestingly, z-VAD-fmk-treated cells that had released cyt-C-GFP showed a
stable
M-cyt-C did not exhibit a
depolarization of
P and were viable for several hours.
However, these cells may also eventually die, a process that may involve the
complete removal of mitochondria by autophagy or a final
M depolarization after the activation of the
permeability transition pore (Chang and
Johnson, 2002
; Xue et al.,
2001
).
Ion gradients across the plasma membrane are not stable during apoptosis
(Bortner and Cidlowski, 1999;
Bortner et al., 2001
;
Maeno et al., 2000
). Our
single-cell experiments demonstrate that similar to the release of
cyt-C-GFP the time point of
P depolarization
was set individually for each cell during apoptosis. The broad-spectrum
caspase inhibitor z-VAD-fmk not only blocked
P
depolarization but also inhibited apoptotic cell shrinkage and the degradation
of the regulatory ß-subunit of Na+/K+-ATPase,
suggesting that these were caspase-dependent events. Previous studies have
shown that the intracellular Na+ concentration increases prior to a
loss of plasma membrane integrity (Bortner
et al., 2001
) and that cell shrinkage in apoptotic Jurkat cells is
accompanied by a net efflux of ions due to an inactivation of
Na+/K+-ATPase (Mann
et al., 2001
). The latter authors also observed that
Na+/K+-ATPase subunits were degraded in populations with
reduced volume during apoptosis. Interestingly, it has previously been shown
that apoptosis is associated with a Bcl-2-sensitive activation of outward
K+ currents (Ekhterae et al.,
2001
; Yu et al.,
1997
). Because active cellular volume regulation requires
Na+/K+-ATPase activity, both events may act
synergistically in the induction of cell shrinkage. Cleavage products of the
Na+/K+-ATPase ß-subunit could not be detected in
the present study, but it should be noted that the N-terminal cytoplasmic
domains of the known ß-subunits do not contain canonical caspase cleavage
sites. This suggests that caspases may not directly cleave these subunits. The
ß-subunit of the Na+/K+-ATPase, however, regulates
the K+ affinity of the enzyme at the extracellular site
(Eakle et al., 1994
;
Shainskaya and Karlish, 1996
).
A truncation of the N-terminal cytoplasmic domain of the ß-subunit has
been shown to severely impair K+ affinity and to increase
ouabain-sensitive leak currents (Abriel et
al., 1999
; Hasler et al.,
1998
). Apart from a role in cell shrinkage during apoptosis,
caspase-mediated degradation of Na+/K+-ATPase and other
energy-consuming enzymes such as PARP (Ha
and Snyder, 1999
) may also help to transiently maintain cytosolic
ATP levels sufficient for the execution of apoptosis in cells that have
released cyt-C. Indeed, remodeling of TMRM fluorescence changes in the absence
or presence of z-VAD-fmk revealed a slower
M
depolarization when caspase activation was not blocked.
Our data demonstrate that outer mitochondrial membrane permeabilization
during apoptosis triggers a z-VAD-fmk-sensitive P
depolarization, as well as a z-VAD-fmk-independent
M
depolarization, and hence coordinates the activation of distinct, apoptotic
and necrotic cell death pathways.
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Acknowledgments |
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References |
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