© 1999 American Heart Association, Inc.
Cellular Biology |
The Mitochondrial Apoptotic Pathway Is Activated by Serum and Glucose Deprivation in Cardiac Myocytes
From the Departments of Medicine (Cardiology) and Cell Biology (S.B., S.M., A.L.W., R.N.K.), Albert Einstein College of Medicine, Bronx, NY; Division of Endocrinology (V.L.C., A.D.), Northwestern University Medical Center, Chicago, Ill; and Idun Pharmaceuticals (A.S.), La Jolla, Calif. S.B.'s present address is Department of Molecular Genetics, Weizmann Institute of Science, Rehovot, Israel.
Correspondence to Richard N. Kitsis, Departments of Medicine (Cardiology) and Cell Biology, Albert Einstein College of Medicine, 1300 Morris Park Ave, Bronx, NY 10461. E-mail kitsis{at}aecom.yu.edu
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Abstract—Many cell types undergo apoptosis under conditions of ischemia. Little is known, however, about the molecular pathways that mediate this response. A cellular and biochemical approach to elucidate such signaling pathways was undertaken in primary cultures of cardiac myocytes, a cell type that is especially sensitive to ischemia-induced apoptosis. Deprivation of serum and glucose, components of ischemia in vivo, resulted in myocyte apoptosis, as determined by nuclear fragmentation, internucleosomal cleavage of DNA, and processing of caspase substrates. These manifestations of apoptosis were blocked by zVAD-fmk, a peptide caspase inhibitor, indicating that caspase activity is necessary for the progression of apoptosis in this model. In contrast to control cells, apoptotic myocytes exhibited cytoplasmic accumulation of cytochrome c, indicating release from the mitochondria. Furthermore, both caspase-9 and caspase-3 were processed to their active forms in serum-/glucose-deprived myocytes. Caspase processing, but not cytochrome c release, was inhibited by zVAD-fmk, placing the latter event upstream of caspase activation. This evidence demonstrates that components of ischemia activate the mitochondrial death pathway in cardiac myocytes.
Key Words: apoptosis • cysteine proteinase • cytochrome c • mitochondria • ischemia
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Introduction |
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Myocardial ischemia is a pathological process that results in extensive cell death, a significant portion of which can be attributed to apoptosis. Myocyte apoptosis has been demonstrated in necropsy samples of humans suffering myocardial infarction1 2 3 4 as well as in rabbit, rat, and mouse models of continuous ischemia or transient ischemia followed by reperfusion.5 6 7 8 9 Cultured cardiac myocytes also undergo apoptosis in response to component stimuli of ischemia, such as hypoxia, serum and nutrient deprivation, and metabolic inhibition, with and without restoration of control conditions.10 11 12 13 14 15 16 The identities of the molecular signaling pathways that mediate ischemia-induced apoptosis are largely unknown, however. Previous experiments have implicated signaling pathways involving p53 and its transcriptional targets,3 6 8 10 11 17 growth factor withdrawal,12 13 and TNF and ceramide signaling.18 19 These pathways are likely to act in a redundant, overlapping fashion, as evidenced by the observation that abrogation of individual pathways does not prevent myocyte apoptosis during myocardial infarction in vivo.9
The caspase family of cysteine proteases serves as the central
executioners of apoptosis.20 Caspases are
synthesized as proenzymes that are activated by proteolytic
processing to form the active large (p20) and small (p10)
subunits.20 Once activated, caspases initiate a
cascade of proteolysis that involves further processing/activation of
additional caspases, and ultimately, cleavage of specific cellular
proteins, such as poly(ADP-ribose) polymerase, lamin, DNA fragmentation
factor (DFF)45/inhibitor of caspase-activated DNase (ICAD),
protein kinase C-
(PKC), and focal adhesion kinase
(FAK).20 Breakdown of these and other substrates leads to
the orderly dismantling of the apoptotic cell. It has recently
been demonstrated that caspases are processed in cardiac myocytes
during continuous ischemia21 and
ischemia-reperfusion in vivo.22 Significantly,
inhibition of caspase activity with a caspase pseudosubstrate blocks
myocyte apoptosis in these models.21 23 Thus, it
appears that caspases are central mediators of myocyte
apoptosis during ischemia.
Apoptotic stimuli activate the caspase proteolytic
cascade by inducing oligomerization and subsequent autocatalytic
processing of a subset of caspases known as the signaling
caspases.20 Oligomerization occurs at distinct cell
locations in the context of multiprotein structures, such as the
Fas/FADD complex at the plasma membrane or the Apaf-1 complex at the
mitochondria.20 Apaf-1 is an adaptor molecule that
recruits and activates caspase-9, but only when
simultaneously bound to dATP/ATP and cytochrome
c.24 25 26 Therefore, translocation of
cytochrome c from the mitochondrial intermembrane space into
the cytoplasm, where it binds Apaf-1, serves as a trigger for
apoptosis.27 28 29 30 The mechanism by which this
process is initiated and executed is not understood, although it is
known that antiapoptotic members of the Bcl-2 family block
cytochrome c release, whereas the proapoptotic
member Bax promotes it.28 29 31 32 Furthermore,
cytochrome c release can result from changes in
mitochondrial membrane permeability after loss of membrane potential
(
m).33 34 Loss of
m, a common, early characteristic of
apoptosis, is thought to result from dissipation of the
H+ gradient after opening of the permeability
transition (PT) pore in the inner mitochondrial membrane. Although
cytochrome c release can precede and occur independently of
decreased m,28 29 35
dissipation of m is sufficient to
activate the apoptosis program.33 34 36
During ischemia in vivo, myocyte mitochondria exhibit decreased
m and reduced function.37
Thus, the mitochondria may serve as a death trigger during
ischemia by promoting the release of cytochrome
c.
Although the mitochondrial/cytochrome c death pathway mediates apoptosis in response to many stimuli,38 39 40 41 its involvement in ischemia-induced apoptosis has not been demonstrated in any cell system. To address this issue, components of ischemia, namely serum and glucose deprivation, were examined in a cell culture model of cardiac myocyte apoptosis. We find that cytochrome c is released from the mitochondria of all apoptotic myocytes, coinciding with activation of both caspase-9 and caspase-3. Furthermore, cytochrome c release occurs independently of caspase activity. These data provide evidence implicating the caspase-9/Apaf-1 mitochondrial death pathway in myocyte apoptosis and suggest a mechanism by which ischemia leads to cardiac myocyte apoptosis.
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For all experiments, chemicals were purchased from Sigma, unless otherwise noted. Tissue culture reagents were obtained from Gibco-BRL unless stated otherwise. Pregnant female rats or 1-day-old neonatal rats were supplied by either Taconic Farms (Germantown, NY) or Charles River Laboratories (Wilmington, MA). All animal experimental protocols were approved by the review board of the Animal Institute of the Albert Einstein College of Medicine.
Primary Myocyte Cultures
Primary cultures of neonatal rat cardiac myocytes were prepared
as described.42 Cells were plated on coated dishes (Falcon
Primaria, Becton Dickinson Labware) at a density of 350
cells/mm2 in plating medium, which consisted of a
1:1 mixture of DMEM and Ham's F12 supplemented with 10% defined FBS
(Hyclone Laboratories, Inc), 100 U/mL penicillin, 0.1 mg/mL
streptomycin, 0.25 µg/mL amphotericin B, and 1 mg/mL BSA.
Approximately 24 hours after plating, cardiac myocytes were washed
twice with PBS, and treatment medium was added. For controls, treatment
medium consisted of RPMI 1640 (containing 11 mmol/L
D-glucose), supplemented with 10% FBS, whereas
deprivation medium consisted of glucose-free, serum-free
RPMI 1640, with or without addition of 1 mmol/L
2-deoxy-D-glucose. In certain experiments, 100
µmol/L zVAD-fmk (benzyloxycarbonyl-Val-Ala-Asp(OMe)-fluoromethyl
ketone) or vehicle, DMSO, was added to the medium.
Ladder Assays
Adherent and floating cells were collected and lysed in the
following (in mmol/L): Tris (pH 8.0) 10, NaCl 100, and EDTA 25, as
well as 0.5% SDS and 1.0 mg/mL proteinase K at either 37°C for 4
hours or overnight at room temperature. DNA was extracted from the
digested cells as previously described9 and subjected to
electrophoresis on 1.4% agarose gels.
Immunocytochemistry
Cells on coverslips were fixed in 3.7% formaldehyde, blocked
with 10% normal goat serum and 0.4% Triton X-100 in PBS, and
incubated with primary antibody diluted in 2% normal goat serum and
0.4% Triton X-100. Mouse monoclonal anti–cytochrome c
(Pharmingen) was used at a dilution of 1:500; rabbit polyclonal
anti-ventricular myosin light chain 2 antibody (MLC2v; a
kind gift of Dr Kenneth Chien, University of California, San Diego),
1:50; and rabbit polyclonal CM1, which recognizes only the processed
p20 subunit of activated caspase-3,43 1:500.
After washing coverslips with PBS/0.1% Tween-20, cells were incubated
with secondary antibody consisting of FITC-conjugated anti-rabbit IgG
(Jackson ImmunoResearch Laboratories, Inc, diluted 1:75) or Texas
Red–conjugated anti-mouse IgG (Molecular Probes, diluted 1:250).
Finally, cells were counterstained with the DNA binding dye DAPI. To
stain for mitochondria, unfixed cells were incubated with 100
µmol/L MitoTracker Red (Molecular Probes) for 30 minutes at
37°C. After washing and fixation in 3.7% formaldehyde, cells were
stained further with anti–cytochrome c antibody as
described above, using FITC-conjugated anti-mouse IgG (Molecular
Probes) as secondary antibody.
Western Blot Analysis
A polyclonal anti–caspase-9 antibody was generated by
immunizing rabbits with bacterially expressed, recombinant human
caspase-9 (amino acids 139 to 416). The affinity-purified antibody
recognized both the unprocessed pro–caspase-9 protein and the
processed 35-kDa intermediate (lacking the p10 subunit) on Western
blots. It did not cross-react with recombinant caspase-1, -2, -3, -6,
-7, -8, or -10 (data not shown).
Western blotting was performed on lysates prepared from
serum-/glucose-deprived or control myocytes. Adherent and floating
cells were collected and resuspended in 60 mmol/L Tris-Cl (pH
6.8), 1% SDS, 10% glycerol, and 0.7 mol/L ß-mercaptoethanol.
Samples were boiled and were subjected to electrophoresis on 8% or
13% polyacrylamide gels and then transferred to Immobilon-P
nylon membranes (Millipore Corp). After blocking in 5% nonfat milk and
0.05% Tween-20 in PBS, blots were incubated with antibodies to FAK
(1:1000, mouse monoclonal antibody, Transduction Laboratories), PKC
(1:2000, rabbit polyclonal antibody, Santa Cruz Biotechnology), or
caspase-9 (1:12 000). For analysis of mitochondrial proteins,
membranes were blocked in 10% nonfat milk for 2 hours and incubated
overnight at 4°C with mouse monoclonal antibodies to denatured
cytochrome c (1:500, Pharmingen), cytochrome oxidase IV (COX
IV; 0.1 µg/mL, Molecular Probes) or actin (1:500, Sigma). Secondary
antibody consisted of a 1:75 000 dilution of horseradish peroxidase
(HRP)–conjugated goat anti-rabbit IgG (for caspase-9 blots, Pierce) or
1:2000 dilution of HRP-conjugated goat anti-mouse IgG (for
mitochondrial analysis, Sigma) and was detected with Super
Signal Ultra Chemiluminescent Substrate (Pierce) on Kodak BioMax Light
film (Eastman Kodak). For other antibodies, secondary antibody
consisted of 1:2000 dilutions of HRP-conjugated anti-rabbit IgG or
anti-mouse IgG (Southern Biotechnology Associates, Inc), which was
detected by enhanced chemiluminescence (Amersham Life Sciences,
Inc).
Cellular Fractionation
Cardiac myocytes were washed twice with ice-cold PBS and
collected by centrifugation at 200g for 10
minutes at 4°C. The cell pellets were then resuspended in 400 µL of
extraction buffer containing (in mmol/L) mannitol 220, sucrose 68,
HEPES, (pH 7.4) 20, KCl 50, EGTA 5, MgCl2 2, EDTA
1, and DTT 1, as well as protease inhibitors. Cells were
homogenized for 40 strokes using a glass Dounce
homogenizer and a B pestle. Cell
homogenates were then subjected to
centrifugation at 14 000g for 30 seconds to
pellet nuclei, and supernatants were centrifuged once more at
200 000g for 30 minutes. Two micrograms of the resulting
supernatants (consisting of soluble proteins) and pellets (containing
mitochondria, microsomes, and insoluble proteins) were fractionated on
12.5% SDS-polyacrylamide gels as described above.
Measurement of Intracellular ATP Content
The intracellular ATP content of control and
serum-/glucose-deprived myocytes was measured with the ATP
bioluminescent assay kit (Sigma). Cells were lysed directly in
somatic-cell ATP-releasing agent, and the lysate was assayed according
to the manufacturer's instructions using a 1:25 dilution of the ATP
assay mix (a solution containing the firefly luciferase protein and
luciferin). Light emitted was measured using the Monolight luminometer
(model 2010, Analytical Luminescence Laboratory). ATP content was
calculated by comparison with a standard curve derived from known
concentrations of ATP, ranging from 0.01 to 10 pmol.
Mitochondrial Reductase Activity
Cellular reductase activity of live cultured myocytes was
determined by measuring the reduction of MTT.44 At each
end point, treatment medium was replaced with fresh, serum-free medium
(with glucose for controls, with 1 mmol/L 2-deoxyglucose for
serum-/glucose-deprived samples) containing 2.4 mmol/L MTT at pH
7.4. Cells were incubated with MTT medium for 1 hour at 37°C with
occasional mixing. After solubilization in N-propanol (FisherBiotech),
absorbance was measured at 560 nm using the UltraSpec III UV/visible
spectrophotometer (Pharmacia Biotech, Inc).
Measurement of m
Mitochondrial membrane potential was assessed using both
JC-145 and rhodamine 12346 staining. After
treatment in control or deprivation medium, cardiac
myocytes on coverslips were incubated in serum-free medium containing
10 µmol/L JC-1 (Molecular Probes) or 25 µmol/L rhodamine
123 at 37°C for 5 minutes (JC-1) or 10 minutes (rhodamine 123).
During this step, the glucose and deoxyglucose contents of the medium
were maintained as during the treatment period. After staining,
cultures were washed twice with the original treatment medium.
Coverslips were then removed from culture dishes and inverted onto a
slide. The live (unfixed) cardiac myocytes were viewed as follows: for
JC-1, excitation at 450 to 490 nm with sampling at 
520 nm, and for
rhodamine 123, excitation at 546 nm with sampling at 590 nm. As a
positive control for loss of m, cells were
treated with 50 mmol/L sodium cyanide and 62.5 µg/mL oligomycin
(mixture of oligomycin A, B, and C) at 37°C for 30 minutes.
Statistical Analysis
For ATP and MTT studies, triplicate samples were examined at
each of 6 time points. Results were subjected to ANOVA and Tukey post
hoc statistical analysis with differences considered
significant at P<0.05. Data are reported as the
mean±SD.
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Deprivation of Serum and Glucose Induces Apoptosis in Cultured Cardiac Myocytes
Ischemia is a complex physiological process, in which multiple changes contribute to cellular death. Among these are deprivation of nutrients, growth, and survival factors. To study the effects of these component stimuli on myocyte apoptosis, primary neonatal rat cardiac myocytes were cultured in the absence of serum and glucose. To further limit glucose metabolism, the nonmetabolizable glucose analogue 2-deoxy-D-glucose (1 mmol/L) was added to the medium in some experiments. Myocyte contraction, an ATP-dependent process, was not observed after 4 hours of treatment. By 6 to 15 hours, myocytes exhibited a shriveled, constricted shape. Many round, floating cells were observed, and by 24 hours, there were few adherent cells remaining (Figure 1B
). In contrast, myocytes
incubated in control medium containing glucose and serum were confluent
and well spread, with flattened morphology (Figure 1A), and beat
rapidly in synchrony. The morphological changes observed were
suggestive of apoptosis. In fact, internucleosomal DNA
fragmentation was observed in the deprived, but not control, plates, at
24 and 30 hours (Figure 2A). Serum and
glucose deprivation without addition of 2-deoxyglucose also
induced apoptosis, but the process was much slower;
apoptotic DNA ladders were first evident at 48 to 60 hours
(Figure 2B). Serum deprivation alone did not induce
apoptosis in the time course examined (data not shown),
although it has been reported to cause myocyte death at later time
points.12 13 Likewise, addition of 1 mmol/L
2-deoxyglucose to control medium or to serum-containing glucose-free
medium had no effect on myocyte morphology (data not shown). This may
be due to the presence of glucose or other energy sources in serum.
Alternatively, survival factors present in serum may actively
suppress the apoptotic signal.
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Apoptosis was confirmed on a cell-by-cell level by examining
cellular and nuclear morphology with an antibody to the sarcomeric
protein MLC2v and the DNA binding dye DAPI, respectively. Control
myocytes were flat and striated, with large, round nuclei (Figure 3A and 3B). This contrasted with the
serum-/glucose-deprived cells, which were shrunken and lacked
striations, suggestive of sarcomeric breakdown (Figure 3D).
Three to six percent of myocytes that were still adherent to the plate
exhibited fragmented nuclei with condensed chromatin, a clear
indication of apoptosis (Figure 3C). This quantity
underestimates the total extent of death, however, because the vast
majority of cells had detached from the culture plate (see below).
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Myocytes with nuclei in earlier stages of fragmentation, including
margination of DNA at the nuclear periphery, were observed in medium
containing 
0.1 mmol/L 2-deoxyglucose (Figure 3C
, inset).
Such nuclei were seen only rarely in the presence of 1 mmol/L
2-deoxyglucose. This is consistent with the kinetics of cell
death being more rapid in the presence of high concentrations of the
metabolic inhibitor, such that intermediate
stages of death were less likely to be observed.
Caspases Mediate Cardiac Myocyte Apoptosis During
Serum/Glucose Deprivation
Proteolysis of caspase substrates provides a marker for
apoptosis in general and caspase activity in particular. To
determine whether caspases were activated in
serum-/glucose-deprived myocytes, Western blot analysis of
caspase substrates PKC and FAK was performed. In fact, both
substrates were proteolyzed in a time-dependent manner after
deprivation (Figure 4A). By 13 hours, both FAK (120 to
125 kDa) and PKC (80 kDa) were processed to their predicted caspase
cleavage products of 77 to 85 kDa and 40 kDa,
respectively.47 48 By 24 hours, the majority of the
precursor proteins was proteolyzed. In control cells, a low level of
processing was observed; this may reflect basal caspase activity or
nonspecific proteolysis at hypersensitive sites during preparation of
the cell lysates.
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To determine whether caspase activity was necessary for the progression
of myocyte apoptosis during ischemia, cardiac myocytes
were subjected to serum/glucose deprivation in the presence
of either the caspase pseudosubstrate inhibitor zVAD-fmk or
vehicle alone. The caspase inhibitor completely blocked
internucleosomal DNA fragmentation in serum-/glucose-deprived cells
(Figure 2C). Consistent with the absence of DNA ladders,
zVAD-fmk prevented nuclear fragmentation as well (see Figure 9D)
and preserved sarcomeric staining (data not shown). Inhibition of
caspase activity also blocked proteolysis of caspase substrates FAK and
PKC (Figure 4B). Thus, caspases are necessary for the
execution of myocyte apoptosis during serum/glucose
deprivation. Despite the prevention of the terminal
features of apoptosis, however, cellular morphology and
contractile capability of the remaining adherent cells did not improve
to normal (data not shown), unless glucose and serum were restored.
This suggests that energy depletion or other caspase-independent events
may also contribute to cellular damage in this model.
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The Mitochondrial Death Pathway Is Activated in
Apoptotic Myocytes
Treatment of myocytes with serum-free, glucose-free medium
containing 2-deoxyglucose resulted in a decrease in cellular ATP levels
within the first 2 hours, which ultimately declined to levels that were
30% of control cells (Figure 5A). This reflects the absence of
the glycolytic production of ATP, as well as the limited redox
potential of the mitochondria that results from the reduced supply of
substrates (eg, pyruvate) for the oxidative respiratory pathway. In
fact, an impairment in mitochondrial redox activity was apparent soon
after initiation of the stimulus, as assessed by the conversion of the
tetrazolium dye MTT to its reduced form, a reaction mediated by
mitochondrial reductases.44 MTT reductase activity fell
precipitously to 45% of control levels within 4 hours of treatment
(Figure 5B). It remained steady at 40% for the next 5 hours.
These changes occurred before any signs of cell death were observed,
indicating that the decrease in metabolism and the
consequent fall in energy production was not a result of the
loss of live cells, but was rather due to a metabolic
impairment within a population that was still viable.
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In light of these changes in mitochondrial function, we assessed
whether there was any decrease in mitochondrial membrane potential
using the potential-sensitive dye JC-1. Myocytes exhibited loss of
m after a 6-hour incubation in medium
lacking glucose and serum and containing 2-deoxyglucose (Figure 5C).
Loss of membrane potential persisted at 18 hours (data not
shown). In contrast, no change in m was
detectable after 2 hours of deprivation conditions (data
not shown). Similar results were obtained with rhodamine 123 staining
(data not shown). Thus, loss of m lagged
slightly behind changes in cellular ATP content and MTT reductase
activity.
Because dissipation of the mitochondrial potential can be sufficient to
activate the apoptotic pathway,33 34 36 we
hypothesized that the mitochondrial impairments observed here might
lead to the induction of apoptosis through the release of
cytochrome c. To test this hypothesis, the subcellular
localization of cytochrome c was determined by Western blot
analysis of fractionated cell lysates (Figure 6). Cytochrome c was
present exclusively in the insoluble, membrane fraction of control
myocytes (compare lanes 1 and 3). In contrast, after 18 hours of serum
and glucose deprivation, cytochrome c was
observed in both the membrane (lane 2) and cytosolic (lane 4)
fractions. The appearance of cytochrome c in the cytosol was
not due to contamination of the cytosolic fraction with mitochondria,
as the mitochondrial membrane protein COX IV was found exclusively in
the membrane fraction (compare lanes 1 and 2 with 3 and 4). To confirm
translocation of cytochrome c to the cytosol in individual
apoptotic cells, immunostaining was performed
(Figure 7). Control cells with normal
nuclear morphology (Figure 7A) exhibited a wispy, punctate,
subcytoplasmic pattern of cytochrome c
immunostaining (Figure 7C), which coincided with a
marker for mitochondria, MitoTracker Red (Figure 7G through 7I). In
contrast, serum-/glucose-deprived myocytes with fragmented,
apoptotic nuclei (Figure 7D) always exhibited a diffuse,
cytoplasmic pattern of cytochrome c
immunostaining (Figure 7F). Notably, those deprived
cells in which the nuclei remained intact (ie, nonapoptotic)
retained the punctate cytochrome c
immunostaining (data not shown). Thus, in response to
serum/glucose deprivation, cytochrome c
translocated from the mitochondria into the cytoplasm specifically in
apoptotic cells.
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Apoptotic myocytes that contained fragmented nuclei (Figure 7D) and cytoplasmic cytochrome c (Figure 7F) also
stained positively with the CM1 antibody, which recognizes only the
processed, activated form of caspase-3 (Figure 7E). In
contrast, nonapoptotic deprived cells with intact nuclei and
mitochondria-localized cytochrome c were CM1 negative (data
not shown), as were control cells (Figure 7A through 7C). The
concordance among these 3 parameters (cytochrome
c translocation, caspase-3 activation, and nuclear
fragmentation) was absolute in >3000 myocytes examined in 8
independent preparations of cells.
Interestingly, in the early phases of apoptosis, active
caspase-3 was compartmentalized to particular subcellular locations.
Figures 7J and 7K show an example of a myocyte cultured in the absence
of serum and glucose but without addition of 2-deoxyglucose. The DNA in
the nucleus of this cell is marginated at the nuclear membrane (Figure 7J), and CM1 staining is predominantly in the nucleus and at the cell
periphery (Figure 7K). Although the exact identities of these
subcellular compartments have not been determined in these experiments,
it is notable that this phenomenon has also been observed in cardiac
myocytes subjected to other apoptotic stimuli such as
staurosporine (A.S., unpublished data, 1998).
The link between cytochrome c release and activation of
caspase-3 is caspase-9, the first caspase activated in the
mitochondrial pathway.25 26 We therefore generated a
monospecific polyclonal antibody to caspase-9 and performed Western
blot analysis to determine whether the caspase was
proteolytically processed (Figure 8A). In
control lanes, the full-length caspase-9 precursor was observed at
46 kDa. Only minimal processing was observed, which may reflect
basal activation of the caspase, or more likely, autocatalysis of
aggregated protein after cell lysis. No differences were observed after
13 hours of serum/glucose deprivation. We cannot exclude
the possibility, however, that undetectable levels of processed
caspase-9 enzyme are generated that are sufficient to activate
the proteolytic cascade. In fact, caspase substrates are already
proteolyzed at this time (Figure 4A), which suggests that some
upstream signaling caspase is active at 13 hours. By 18 hours, however,
the 46-kDa caspase-9 precursor was converted to a 35-kDa fragment,
which represents the first processing intermediate (removal of
p10) during caspase-9 activation.26 An additional band of
30 kDa was also observed; this most likely reflects removal of the
prodomain.49 The abundance of the caspase-9 precursor and
the intermediates substantially decreased by 24 hours, presumably
because of further processing to the final p10 and p20 subunits, which
were not detectable with this antibody. Thus, caspase-9 is
activated by serum/glucose deprivation in cardiac
myocytes.
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Cytochrome C Release Occurs Independently of
Caspase Activity
Cytochrome c release from the mitochondria can occur
through caspase-dependent or independent
pathways.28 30 35 50 51 We therefore investigated the
effects of inhibiting caspase activity on the mitochondrial pathway. As
stated previously, treatment of serum-/glucose-deprived cells with the
caspase inhibitor zVAD-fmk completely blocked nuclear
fragmentation (Figure 9D). Furthermore,
no CM1 staining was observed (Figure 9E), denoting inhibition of
caspase-3 processing. In addition, proteolysis of caspase-9 was
inhibited (Figure 8B), although this was not complete. Perhaps
zVAD-fmk is not capable of completely blocking caspase-9 autocatalysis
on cytochrome c release, even though it is an effective
inhibitor of further caspase activity. Similar results have
been reported previously with the viral caspase inhibitor
CrmA, which blocked Fas-induced apoptosis and caspase-8
activity, but not caspase-8 processing.52 Most
notably, however, cytochrome c release from the mitochondria
was not blocked (Figure 9F, cell on right). In fact, 40% of
myocytes cultured in medium lacking serum and glucose but containing
2-deoxyglucose and zVAD-fmk exhibited diffuse, cytoplasmic cytochrome
c immunostaining in the absence of caspase-3
activation or nuclear fragmentation. This suggests that at least 40%
of the population had initiated the apoptotic program, which
would have advanced to death had caspases been activated. Thus,
cytochrome c translocation in this model occurs through a
caspase-independent mechanism.
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Serum and glucose deprivation, in combination with the addition of 2-deoxyglucose, induces time-dependent apoptosis in cultured neonatal rat cardiac myocytes, as evidenced by changes in nuclear morphology, internucleosomal DNA fragmentation, caspase activation, and cleavage of caspase substrates. Caspase activation in this model occurs, at least in part, via the mitochondrial death pathway, as evidenced by the translocation of cytochrome c from the mitochondria to the cytosol and the activation of both caspase-9 and caspase-3. Moreover, the release of cytochrome c occurs independently of caspase activity. The apoptotic stimulus used in this model involves several components of ischemia in vivo, including deficiency of survival and growth factors and some nutrients. Other important aspects of ischemia, such as hypoxia, were not examined. However, the sufficiency of the component stimuli to activate the caspase-9/Apaf-1 mitochondrial death pathway in vitro suggests that this pathway mediates ischemia-induced apoptosis in vivo as well.
When cells were cultured in serum-/glucose-free medium containing 2-deoxyglucose, activated caspase-3 was localized throughout the cytosol. In contrast, when 2-deoxyglucose was omitted, processed caspase-3 was observed at specific subcellular locations, namely the nucleus and periphery of the cell. One interpretation of these observations is that the less severe inhibition of glycolysis resulting from omission of 2-deoxyglucose might involve only a subset of those caspases activated by the more severe stimulus. Alternatively, the absence of 2-deoxyglucose may slow the kinetics of apoptosis, allowing the identification of intermediate stages, such as the initial activation of specific subcellular pools of caspases. A previous study using confocal microscopy and immunoelectron microscopy concluded that pro–caspase-3 is located in both the cytoplasm and the mitochondrial intermembrane space, and apoptotic stimuli led to the activation of the mitochondrial pool.53 The disparity between the pattern of activation observed in the aforementioned study and the current report may reflect differences in cell type or death stimulus. More generally, however, the observations in both studies suggest that there are distinct sites at which pools of inactive procaspases reside and become activated when the cell receives an apoptotic stimulus. These sites might represent locations where effector caspases are targeted to ensure contact with either upstream signaling caspases or downstream substrates. Studies that identify these compartments as well as proteins that colocalize with activated caspase-3 should be informative in this regard.
Inhibition of caspase activity by administration of zVAD-fmk blocked processing of caspases and their substrates, nuclear fragmentation, and DNA cleavage. In contrast, translocation of cytochrome c from the mitochondria occurred independently of caspase activation during serum/glucose deprivation, consistent with a direct link between the apoptotic signals and the mitochondria. In this respect, serum/glucose deprivation resembles apoptotic stimuli such as UV irradiation, staurosporine, and chemotherapeutic drugs, in which cytochrome c release occurs independently and upstream of caspase activation.28 35 It contrasts, however, with the recently described caspase-dependent pathway leading to cytochrome c release that occurs during Fas- and TNF-mediated apoptosis, which involves caspase-8–mediated cleavage of Bid and subsequent translocation of truncated Bid to the mitochondria.50 51
The mechanisms by which serum/glucose deprivation leads to cytochrome c release are not known. Possibilities include withdrawal of survival factors, which have been shown in other cell types to provide antiapoptotic signals through Akt-mediated phosphorylation of the proapoptotic Bcl-2 family member Bad, a Bcl-2/XL dimerization partner.54 When phosphorylated, Bad is sequestered away from Bcl-2/XL, enabling the antiapoptotic activity of the latter, which includes blocking release of cytochrome c from the mitochondria.28 29 Furthermore, caspase-9 is also phosphorylated by Akt, rendering it catalytically inactive.55 Thus, serum withdrawal in the cardiac myocyte model may activate a death signal through elimination of such survival signals.
Whatever upstream pathways transduce the serum/glucose
deprivation signal to the mitochondria, changes intrinsic
to the mitochondria are likely to ultimately mediate cytochrome
c release. We observed a loss of
m beginning 6 hours after the onset of
serum/glucose deprivation and continuing to at least 18
hours. Although loss of m is not necessary
for cytochrome c release in some
systems,28 29 35 it is often sufficient for
cytochrome c release to occur.36
Therefore, it is possible that loss of m
contributes to cytochrome c release in cardiac myocytes.
Loss of m is indicative of the opening of
large, nonselective channels in the inner membrane known as the PT
pore.34 It has been previously proposed that opening
of the PT pore would allow the entry of solutes and water into the
mitochondrial matrix, thereby causing swelling of the
matrix.34 Because of the greater surface area of the inner
membrane, this swelling would, in turn, cause rupture of the outer
mitochondrial membrane, liberating proteins that reside in the
intermembrane space, including cytochrome c. Additional
experiments are needed to determine whether loss of
m causes the initial release of cytochrome
c from cardiac myocytes or merely amplifies the release once
initiated by other mechanisms.
The data presented here strongly indicate that apoptosis occurs after serum/glucose deprivation. This does not exclude the possibility, however, that necrosis also occurs. In fact, in our model, it is difficult to distinguish between necrosis and apoptosis, because most of the apoptotic deaths are late events in which cellular membrane integrity is eventually lost in the absence of phagocytosis. Nevertheless, the presence of apoptotic myocytes in our model is in distinct contrast to a previous in vitro model of ischemia in which the majority of myocyte death that ensued was attributed to necrosis.14 The ischemic stimulus used in that study involved deprivation of glucose, serum, and oxygen, conditions that inhibit both glycolysis and oxidative respiration. As a result, ATP would be more severely exhausted compared with our model in which oxygen was maintained. Under the more stringent conditions, apoptosis, an ATP-dependent process,25 27 would not be favored. In fact, apoptosis is blocked in many cell types by severe depletion of ATP56 57 and is even converted to necrosis in some circumstances.57 58 59 Paradoxically, although ATP is required for apoptosis, mild ATP depletion, such as that occurring after blockade of only one component of the ATP production scheme, actually induces or enhances apoptosis.60 61 62 This may contribute to the activation of apoptosis in our model, as well as in a second cultured myocyte model of ischemia, in which oxidative respiration, but not glycolysis, was inhibited.16 It remains to be determined whether mild ATP depletion itself is a primary direct activator of the cytochrome c pathway.
Because apoptotic and necrotic stimuli both lead to mitochondrial damage, this organelle appears to be a point of convergence of the pathways that mediate these morphologically distinct forms of cell death. It is possible, therefore, that necrotic stimuli also lead to the release of cytochrome c from damaged mitochondria. Whether such release can successfully activate the remainder of the pathway leading to caspase activation under conditions of severe ATP depletion is unclear, however. Even if the Apaf-1/caspase-9 pathway were activated, it is unknown whether it mediates downstream necrotic events or merely exists as an "accidental" consequence that plays no role in the pathogenesis of necrotic death. The sorting out of these issues will shed light on the mechanism of necrosis and, in so doing, may call into question the notion that necrosis and apoptosis are completely distinct processes.
In conclusion, the experiments presented here provide evidence for the involvement of the mitochondrial signaling pathway in mediating apoptosis in response to components of ischemia in cardiac myocytes. Elucidation of the details by which this pathway is activated and regulated in cardiac myocytes may suggest novel strategies for the treatment of ischemic heart disease.
Note Added in Proof
Malhotra and Brosius63 recently showed that hypoxia
and glucose deprivation induce the release of cytochrome
c from mitochondria of neonatal cardiac myocytes.
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Acknowledgments |
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This work was supported by grants to R.N.K. from the National Institutes of Health (RO1 HL60665 and RO1 HL61550) and the American Heart Association, New York City affiliate. S.B. was supported by a predoctoral fellowship from the Howard Hughes Medical Institute. V.L.C. was supported by a Mentored Clinical Scientist Development Award from the National Institutes of Health (K08 CA01752) and by institutional research grants to Northwestern University from the Howard Hughes Medical Institute and the American Cancer Society. R.N.K is the Charles and Tamara Krasne Faculty Scholar in Cardiovascular Research of the Albert Einstein College of Medicine. Data in this paper are from a thesis submitted by S.B. in partial fulfillment of the requirements for the degree of Doctor of Philosophy in the Sue Golding Graduate Division of Medical Sciences, Albert Einstein College of Medicine, Yeshiva University. We thank Roberta Gottlieb for advice regarding mitochondrial preparations and Lan Bo Chen and Stine Kraeft for advice regarding measurement of mitochondrial membrane potential.
Received December 10, 1998; accepted June 14, 1999.
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 J. M. Hammel, C. A. Caldarone, T. L. Van Natta, L. X. Wang, K. F. Welke, W. Li, S. Niles, E. Barner, T. D. Scholz, D. M. Behrendt, et al. Myocardial apoptosis after cardioplegic arrest in the neonatal lamb J. Thorac. Cardiovasc. Surg., June 1, 2003; 125(6): 1268 - 1275. [Abstract] [Full Text] [PDF]
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 I. Korichneva, J. Waka, and U. Hammerling Regulation of the Cardiac Mitochondrial Membrane Potential by Retinoids J. Pharmacol. Exp. Ther., May 1, 2003; 305(2): 426 - 433. [Abstract] [Full Text] [PDF]
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P. Lee, M. Sata, D. J. Lefer, S. M. Factor, K. Walsh, and R. N. Kitsis Fas pathway is a critical mediator of cardiac myocyte death and MI during ischemia-reperfusion in vivo Am J Physiol Heart Circ Physiol, February 1, 2003; 284(2): H456 - H463. [Abstract] [Full Text] [PDF]
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T. L. Vanden Hoek, Y. Qin, K. Wojcik, C.-Q. Li, Z.-H. Shao, T. Anderson, L. B. Becker, and K. J. Hamann Reperfusion, not simulated ischemia, initiates intrinsic apoptosis injury in chick cardiomyocytes Am J Physiol Heart Circ Physiol, January 1, 2003; 284(1): H141 - H150. [Abstract] [Full Text] [PDF]
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 P A J Krijnen, R Nijmeijer, C J L M Meijer, C A Visser, C E Hack, and H W M Niessen Apoptosis in myocardial ischaemia and infarction J. Clin. Pathol., November 1, 2002; 55(11): 801 - 811. [Abstract] [Full Text] [PDF]
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W. Chao, Y. Shen, L. Li, and A. Rosenzweig Importance of FADD Signaling in Serum Deprivation- and Hypoxia-induced Cardiomyocyte Apoptosis J. Biol. Chem., August 23, 2002; 277(35): 31639 - 31645. [Abstract] [Full Text] [PDF]
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K. M. Regula, K. Ens, and L. A. Kirshenbaum Inducible Expression of BNIP3 Provokes Mitochondrial Defects and Hypoxia-Mediated Cell Death of Ventricular Myocytes Circ. Res., August 9, 2002; 91(3): 226 - 231. [Abstract] [Full Text] [PDF]
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R. A. Zoeller, T. J. Grazia, P. LaCamera, J. Park, D. P. Gaposchkin, and H. W. Farber Increasing plasmalogen levels protects human endothelial cells during hypoxia Am J Physiol Heart Circ Physiol, August 1, 2002; 283(2): H671 - H679. [Abstract] [Full Text] [PDF]
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T. Date, A. J Belanger, S. Mochizuki, J. A Sullivan, L. X Liu, A. Scaria, S. H Cheng, R. J Gregory, and C. Jiang Adenovirus-mediated expression of p35 prevents hypoxia/reoxygenation injury by reducing reactive oxygen species and caspase activity Cardiovasc Res, August 1, 2002; 55(2): 309 - 319. [Abstract] [Full Text] [PDF]
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J.-M. Li, A. M. Mullen, S. Yun, F. Wientjes, G. Y. Brouns, A. J. Thrasher, and A. M. Shah Essential Role of the NADPH Oxidase Subunit p47phox in Endothelial Cell Superoxide Production in Response to Phorbol Ester and Tumor Necrosis Factor-{alpha} Circ. Res., February 8, 2002; 90(2): 143 - 150. [Abstract] [Full Text] [PDF]
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 C. GILL, R. MESTRIL, and A. SAMALI Losing heart: the role of apoptosis in heart disease--a novel therapeutic target? FASEB J, February 1, 2002; 16(2): 135 - 146. [Abstract] [Full Text] [PDF]
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 S.-Y. Yu, S. J. Yoo, L. Yang, C. Zapata, A. Srinivasan, B. A. Hay, and N. E. Baker A pathway of signals regulating effector and initiator caspases in the developing Drosophila eye Development, January 7, 2002; 129(13): 3269 - 3278. [Abstract] [Full Text] [PDF]
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 K. C. Wollert, B. Fiedler, S. Gambaryan, A. Smolenski, J. Heineke, E. Butt, C. Trautwein, S. M. Lohmann, and H. Drexler Gene Transfer of cGMP-Dependent Protein Kinase I Enhances the Antihypertrophic Effects of Nitric Oxide in Cardiomyocytes Hypertension, January 1, 2002; 39(1): 87 - 92. [Abstract] [Full Text] [PDF]
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M. Gao, S. Fan, I. D. Goldberg, J. Laterra, R. N. Kitsis, and E. M. Rosen Hepatocyte Growth Factor/Scatter Factor Blocks the Mitochondrial Pathway of Apoptosis Signaling in Breast Cancer Cells J. Biol. Chem., December 7, 2001; 276(50): 47257 - 47265. [Abstract] [Full Text] [PDF]
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J. Shiraishi, T. Tatsumi, N. Keira, K. Akashi, A. Mano, S. Yamanaka, S. Matoba, J. Asayama, T. Yaoi, S. Fushiki, et al. Important role of energy-dependent mitochondrial pathways in cultured rat cardiac myocyte apoptosis Am J Physiol Heart Circ Physiol, October 1, 2001; 281(4): H1637 - H1647. [Abstract] [Full Text] [PDF]
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 D. Dyntar, M. Eppenberger-Eberhardt, K. Maedler, M. Pruschy, H. M. Eppenberger, G. A. Spinas, and M. Y. Donath Glucose and Palmitic Acid Induce Degeneration of Myofibrils and Modulate Apoptosis in Rat Adult Cardiomyocytes Diabetes, September 1, 2001; 50(9): 2105 - 2113. [Abstract] [Full Text] [PDF]
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M. Xu, Y. Wang, A. Ayub, and M. Ashraf Mitochondrial KATP channel activation reduces anoxic injury by restoring mitochondrial membrane potential Am J Physiol Heart Circ Physiol, September 1, 2001; 281(3): H1295 - H1303. [Abstract] [Full Text] [PDF]
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R. M. Gurevich, K. M. Regula, and L. A. Kirshenbaum Serpin Protein CrmA Suppresses Hypoxia-Mediated Apoptosis of Ventricular Myocytes Circulation, April 17, 2001; 103(15): 1984 - 1991. [Abstract] [Full Text] [PDF]
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 R. NEVIÈRE, H. FAUVEL, C. CHOPIN, P. FORMSTECHER, and P. MARCHETTI Caspase Inhibition Prevents Cardiac Dysfunction and Heart Apoptosis in a Rat Model of Sepsis Am. J. Respir. Crit. Care Med., January 1, 2001; 163(1): 218 - 225. [Abstract] [Full Text]
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J. W. Adams, A. L. Pagel, C. K. Means, D. Oksenberg, R. C. Armstrong, and J. H. Brown Cardiomyocyte Apoptosis Induced by G{alpha}q Signaling Is Mediated by Permeability Transition Pore Formation and Activation of the Mitochondrial Death Pathway Circ. Res., December 8, 2000; 87(12): 1180 - 1187. [Abstract] [Full Text] [PDF]
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 T. Nakagawa and J. Yuan Cross-Talk between Two Cysteine Protease Families: Activation of Caspase-12 by Calpain in Apoptosis J. Cell Biol., August 21, 2000; 150(4): 887 - 894. [Abstract] [Full Text] [PDF]
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P. M. Kang, A. Haunstetter, H. Aoki, A. Usheva, and S. Izumo Morphological and Molecular Characterization of Adult Cardiomyocyte Apoptosis During Hypoxia and Reoxygenation Circ. Res., July 21, 2000; 87(2): 118 - 125. [Abstract] [Full Text] [PDF]
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P. M. Kang and S. Izumo Apoptosis and Heart Failure : A Critical Review of the Literature Circ. Res., June 9, 2000; 86(11): 1107 - 1113. [Full Text] [PDF]
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 Z. Lin, J. M. Weinberg, R. Malhotra, S. E. Merritt, L. B. Holzman, and F. C. Brosius III GLUT-1 reduces hypoxia-induced apoptosis and JNK pathway activation Am J Physiol Endocrinol Metab, May 1, 2000; 278(5): E958 - E966. [Abstract] [Full Text] [PDF]
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D. Ekhterae, Z. Lin, M. S. Lundberg, M. T. Crow, F. C. Brosius III, and G. Nunez ARC Inhibits Cytochrome c Release From Mitochondria and Protects Against Hypoxia-Induced Apoptosis in Heart-Derived H9c2 Cells Circ. Res., December 3, 1999; 85 (12): e70 - e77. [Abstract] [Full Text] [PDF]
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W. Wu, W.-L. Lee, Y. Y. Wu, D. Chen, T.-J. Liu, A. Jang, P. M. Sharma, and P. H. Wang Expression of Constitutively Active Phosphatidylinositol 3-Kinase Inhibits Activation of Caspase 3 and Apoptosis of Cardiac Muscle Cells J. Biol. Chem., December 15, 2000; 275(51): 40113 - 40119. [Abstract] [Full Text] [PDF]
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M. M-Y Chi, J. Pingsterhaus, M. Carayannopoulos, and K. H. Moley Decreased Glucose Transporter Expression Triggers BAX-dependent Apoptosis in the Murine Blastocyst J. Biol. Chem., December 15, 2000; 275(51): 40252 - 40257. [Abstract] [Full Text] [PDF]
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Y. I. Lee, S. Kang-Park, S.-I. Do, and Y. I. Lee The Hepatitis B Virus-X Protein Activates a Phosphatidylinositol 3-Kinase-dependent Survival Signaling Cascade J. Biol. Chem., May 11, 2001; 276(20): 16969 - 16977. [Abstract] [Full Text] [PDF]
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M. Chen, H. He, S. Zhan, S. Krajewski, J. C. Reed, and R. A. Gottlieb Bid Is Cleaved by Calpain to an Active Fragment in Vitro and during Myocardial Ischemia/Reperfusion J. Biol. Chem., August 10, 2001; 276(33): 30724 - 30728. [Abstract] [Full Text] [PDF]
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J.-M. Li, A. M. Mullen, S. Yun, F. Wientjes, G. Y. Brouns, A. J. Thrasher, and A. M. Shah Essential Role of the NADPH Oxidase Subunit p47phox in Endothelial Cell Superoxide Production in Response to Phorbol Ester and Tumor Necrosis Factor-{alpha} Circ. Res., February 8, 2002; 90(2): 143 - 150. [Abstract] [Full Text] [PDF]
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M. Akao, A. Ohler, B. O'Rourke, and E. Marban Mitochondrial ATP-Sensitive Potassium Channels Inhibit Apoptosis Induced by Oxidative Stress in Cardiac Cells Circ. Res., June 22, 2001; 88(12): 1267 - 1275. [Abstract] [Full Text] [PDF]
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D. Cesselli, I. Jakoniuk, L. Barlucchi, A. P. Beltrami, T. H. Hintze, B. Nadal-Ginard, J. Kajstura, A. Leri, and P. Anversa Oxidative Stress-Mediated Cardiac Cell Death Is a Major Determinant of Ventricular Dysfunction and Failure in Dog Dilated Cardiomyopathy Circ. Res., August 3, 2001; 89(3): 279 - 286. [Abstract] [Full Text] [PDF]
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