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INTRODUCTION |
Cisplatin is used alone or in combination with other agents in the
treatment of many solid tumors and metastatic cancers (including ovarian, testicular, bladder, head and neck, lung, cervical, and breast
cancers) that are no longer amenable to local treatments such as
surgery and/or radiotherapy. The major disadvantage of this
antineoplastic agent is dose-dependent and cumulative
nephrotoxicity (1-3). A single therapeutic dose of cisplatin causes
kidney damage within 48 h (4, 5). Therefore, the dose used in
cancer chemotherapy is limited by the risk of acute or chronic renal
failure (1-3, 6-8). Renal proximal tubular cells
(RPTC)1 are the major target
for cisplatin toxicity within the kidney, as cisplatin nephrotoxicity
manifests primarily as proximal tubule dysfunction (6-8). Cisplatin
alters a variety of RPTC functions including DNA, mRNA, protein
syntheses, transport of organic and inorganic solutes, activities of
some ATPases, 
-glutamyl transpeptidase, and organization of
intracellular cytoskeleton (2-12).
In in vitro conditions, concentrations higher than 50 µM cisplatin result in necrosis whereas lower
concentrations induce apoptosis (13, 14). Multiple mechanisms have been
implicated in cisplatin-induced nephrotoxicity including DNA damage,
oxidative stress, mitochondrial dysfunction, and alteration in signal
transduction pathways involved in apoptosis. Cisplatin-induced
apoptosis in renal epithelial cells is generally considered to be the
result of DNA damage and is associated with caspase-3 activation (15, 16). This event is followed by cleavage of poly(ADP-ribose) polymerase
(PARP), chromatin condensation and DNA fragmentation, dissociation of
different proteins from cytoskeleton, and disruption of intermediate
filament organization of cytoskeleton (5, 11, 16). Oxidative stress has
been also implicated in cisplatin-induced nephrotoxicity. Cisplatin
treatment increases renal lipid peroxidation, blood urea nitrogen,
creatinine, and oxidized glutathione levels, but depletes renal
glutathione (17-19). Large bolus of intravenous glutathione blocks
these effects and cisplatin-induced nephrotoxicity (10).
Mitochondria are a major target of cisplatin in cancer cells. In
ovarian cancer, HeLa, and 3T3 cells, cisplatin treatment decreases
mitochondrial uptake of rhodamine 123 and induces morphological changes
in mitochondria concomitant with cytochrome c release to the
cytoplasm (20-22). Cisplatin-resistant cells exhibit inhibition of
cytochrome c release from mitochondria as a result of
overexpression of Bcl-xL (23, 24). Mitochondrial injury has been
proposed as an early event in cisplatin toxicity in RPTC (9). Higher concentrations of cisplatin that induce cell necrosis (0.1-2.0 mM) cause disruption of the mitochondrial respiratory chain
as a result of inhibition of complexes I-IV (2, 9, 17). The residual
electron flow through the respiratory chain is the source of reactive
oxygen species, but formation of reactive oxygen species is not a
direct cause of the renal cell toxicity induced by high concentrations
of cisplatin (17, 25). These changes are accompanied by the decrease in
state 3 respiration and intracellular ATP content (9). Cisplatin also
inhibits mitochondrial phosphate transport, possibly by a direct
interaction with the mitochondrial phosphate carrier (26). So far,
however, mitochondrial dysfunction has been reported only in renal
proximal tubules exposed to higher concentrations of cisplatin.
Therefore, the first goal of this study was to determine whether lower
concentrations of cisplatin cause alterations in mitochondrial function
that may contribute to toxicity of this drug.
Cisplatin activates multiple signal transduction pathways, which can
lead to different cellular responses (27). In various cancer cells,
cisplatin activates the MEK1 cascade and members of the
mitogen-activated protein kinase family including c-Jun N-terminal
kinase, p38, and extracellular signal-regulated kinases 1 and 2 (ERK1/2) (27-29). Activation of these kinases results in p53
phosphorylation, activation of caspase-3, generation of catalytic domains of different isozymes of protein kinase C (PKC), including PKC-
, PKC-
, and PKC-
, and is followed by cell death (30-32). PKC isozymes have been implicated in cisplatin toxicity in renal proximal tubules as well (4, 33). It has been shown that cisplatin
administration in vivo increases protein levels of PKC-
in renal proximal tubules and that pretreatment with general protein kinase inhibitors protects kidneys from cisplatin-induced injury (4).
On the other hand, there are reports showing that the exposure of renal
cortical slices to cisplatin decreases total PKC activity (33).
PKC- and ERK1/2 are present in RPTC and play an important role in
the responses of these cells to various types of injury. However, the
effect of cisplatin on PKC- and ERK1/2 in RPTC is not known, and it
is unknown whether these kinases are involved in cisplatin
nephrotoxicity. Therefore, the second goal of this study was to
determine whether PKC- and/or ERK1/2 play roles in cisplatin-induced
injury in RPTC and whether these kinases mediate cisplatin effects on
mitochondrial function.
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EXPERIMENTAL PROCEDURES |
Materials--
Female New Zealand White rabbits (2.0-2.5 kg)
were purchased from Myrtle's Rabbitry (Thompson Station, TN).
Cisplatin (cis-diamminedichloroplatinum(II)) was supplied by Aldrich.
5,5',6,6'-Tetrachloro-1,1',3,3'-tetraethylbenzimidazolylcarbocyanine iodide (JC-1) and 4',6-diamidino-2-phenylindole dihydrochloride (DAPI) were obtained from Molecular Probes (Eugene, OR).
L-Ascorbic acid-2-phosphate magnesium salt and cell culture
media were obtained from Wako BioProducts (Richmond, VA) and
Invitrogen, respectively. Caspase-3 and caspase-9 fluorometric
substrates (DEVD-7-amino-4-trifluoromethyl coumarin (AFC) and
LEHD-AFC, respectively), AFC, and buffers for caspase assays were
purchased from BioVision (Palo Alto, CA). PKC- inhibitor (Go6976)
and MEK inhibitors (PD98059 and UO126) were supplied by Calbiochem (La
Jolla, CA). zVAD-fmk was purchased from Biomol (Plymouth Meeting, PA).
Protease inhibitor and phosphatase inhibitor mixtures were supplied by
Oncogene (San Diego, CA) and Sigma, respectively. Phospho-p44/42 ERK,
phospho-PKC-, and active caspase-3 antibodies were obtained from
Cell Signaling Technology (Beverly, MA). Phospho-PKC- and
phospho-PKC- antibodies were purchased from Upstate Biotechnology
(Lake Placid, NY). PKC- and p44/42 ERK antibodies were obtained from
BD Transduction Laboratories (San Diego, CA) and cytochrome
c antibody from Santa Cruz (Santa Cruz, CA). Anti-rabbit and
anti-mouse IgG coupled to horseradish peroxidase were supplied by
Kirkegaard & Perry Laboratories (Gaithersburg, MD) and Supersignal
Chemiluminescent Substrate by Pierce. The sources of the other
reagents and cell culture hormones have been described previously
(34).
Isolation of Proximal Tubules and Culture Conditions--
Rabbit
renal proximal tubules were isolated by the iron oxide perfusion method
and grown in 35-mm culture dishes in optimized conditions as described
previously (34). The purity of the renal proximal tubular segments
isolated by this method is ~96%. The culture medium was a 50:50
mixture of Dulbecco's modified Eagle's medium and Ham's F-12
nutrient mix without phenol red, pyruvate, and glucose, supplemented
with 15 mM NaHCO3, 15 mM Hepes, and 6 mM lactate (pH 7.4, 295 mosmol/kg). Human transferrin (5 µg/ml), selenium (5 ng/ml), hydrocortisone (50 nM),
bovine insulin (10 nM), and L-ascorbic
acid-2-phosphate (50 µM) were added to the medium
immediately before daily media change (2 ml/dish).
Cisplatin Treatment of RPTC Monolayers--
RPTC monolayers
reached confluence within 6 days and were treated with cisplatin on day
6 of culture. Samples of RPTC were taken at various time points
(between 0.5 and 24 h) of cisplatin exposure for immunoblotting,
measurements of mitochondrial functions, and caspase activities.
Various inhibitors (Go6976, PD98059, UO126, zVAD-fmk) were added
0.5 h prior to cisplatin treatment. Control RPTC were treated with
diluent (dimethyl sulfoxide; 0.1% final concentration)
Mitochondrial Function--
Mitochondrial function was assessed
by measurements of oxygen consumption, intracellular ATP content, and
mitochondrial transmembrane potential.
Oxygen Consumption (QO2)--
RPTC monolayers were
gently detached from the dishes with a rubber policeman, suspended in
37 °C culture medium, and transferred to the QO2
measurement chamber. QO2 was measured polarographically using Clark type electrode as described previously (34). Basal QO2 was measured in each run prior to any additions.
Oligomycin-sensitive QO2 was used as a marker of oxidative
phosphorylation, measured in the presence of oligomycin (0.6 µg/ml),
and calculated as a difference between basal and oligomycin-insensitive
QO2. Uncoupled QO2 was used as a marker of the
activity of electron transport through the respiratory chain and was
measured after addition of carbonyl cyanide
p-(trifluoromethoxy)phenylhydrazone (FCCP; 2 µM).
Measurement of Intracellular ATP Content--
ATP content in
RPTC was measured by the luciferase method in freshly prepared cellular
lysates using the ATP bioluminescence assay kit supplied by Roche
Molecular Biochemicals and instructions provided by the manufacturer.
Mitochondrial Membrane Potential
(
m)--
m was assessed using
JC-1, a cationic dye that exhibits potential-dependent
accumulation and formation of red fluorescent J-aggregates in
mitochondria. In contrast, changes in plasma membrane potential do not
affect the JC-1 status. The JC-1 monomer accumulates in the cytoplasm,
where it produces green fluorescence. Formation of J-aggregates in the
mitochondria is indicated by a fluorescence emission shift from green
(525 nm) to red (590 nm). Mitochondrial depolarization is indicated by
a decrease in the red/green fluorescence intensity ratio. At different
time points of cisplatin exposure, RPTC monolayers were loaded with 10 µM JC-1 for 30 min at 37 °C. After loading, media were
aspirated, monolayers kept on ice, washed twice with ice-cold
phosphate-buffered saline (PBS), scraped off culture dishes, washed,
and resuspended in PBS. Fluorescence was analyzed by flow cytometry
(FACSCalibur, BD Biosciences) using excitation by a 488-nm argon-ion
laser. The JC-1 monomer (green) and the J-aggregate (red) were detected
separately in FL1 (emission, 525 nm) and FL2 (emission, 590 nm)
channels, respectively.
The JC-1 accumulation in mitochondria of control and cisplatin-treated
RPTC was visualized by fluorescence microscopy. RPTC were loaded with
JC-1 as described above, washed twice, and overlaid with ice-cold PBS.
Live RPTC monolayers were examined under a Zeiss fluorescent microscope
(Axioskop) using water immersion objective.
Isolation of RPTC Mitochondria and Cytosol--
Mitochondria
were isolated from RPTC as described by Lash and Sall (35). RPTC were
homogenized in the ice-cold isolation buffer (225 mM
sucrose, 10 mM Tris-HCl, 10 mM potassium
phosphate (pH 7.0), 5 mM MgCl2, 20 mM KCl, 0.1 mM phenylmethylsulfonyl fluoride, 2 mM EGTA, 1 mM dithiothreitol, and protease
inhibitor mixture) using Dounce homogenizer and centrifuged at
1,000 × g for 10 min at 4 °C. Supernatant was
collected and centrifuged at 15,000 × g for 10 min at
4 °C. The pellet was washed twice by re-suspending in the isolation
buffer followed by centrifugation at 15,000 × g for 10 min at 4 °C. The final mitochondrial pellet was resuspended in 10 mM Tris-HCl buffer (pH 7.4) containing 25 mM
sucrose, 75 mM mannitol, 100 mM KCl, 0.05 mM K2EDTA, 5 mM
H3PO4, and used for measurement of
F0F1-ATPase activity and immunoblotting. The supernatant collected after the first centrifugation at 15,000 × g for 10 min (mitochondrial pellet) was further spun down at 100,000 × g for 30 min at 4 °C, and the supernatant
resulting from this centrifugation was used as the cytosolic fraction.
Measurement of F0F1-ATPase
Activity--
F0F1-ATPase activity was
determined in freshly isolated RPTC mitochondria by measuring the
release of Pi from ATP as described by Law et
al. (36).
Cytochrome c Release from Mitochondria--
Cytosolic and
mitochondrial fractions were isolated from RPTC at various time points
of cisplatin exposure. Mitochondria were isolated from RPTC as
described above. The supernatant resulting from the mitochondrial spin
was further centrifuged at 100,000 × g for 30 min at
4 °C to obtain the cytosolic fraction, which was used to examine
protein levels of cytochrome c by means of immunoblotting.
Active Na+ Transport--
Ouabain-sensitive
QO2 was used as a marker of active Na+
transport in RPTC as described previously (34). Ouabain-sensitive QO2 was measured in the presence of 1.0 mM
ouabain and calculated as a difference between basal and
ouabain-insensitive QO2.
Measurement of Na+/K+-ATPase
Activity--
RPTC lysates were prepared as described previously (37).
Briefly, 1.0 mg of RPTC protein was added to 0.1 ml of 25 mM imidazole buffer (pH 7.0) containing 0.065% SDS, 1%
bovine serum albumin, and a phosphatase inhibitor mixture. Following
incubation for 10 min at 22 °C, 0.6 ml of 0.3% bovine serum albumin
in 25 mM imidazole buffer was added to lower the SDS
concentration and aliquots were used for measurement of
Na+/K+-ATPase activity.
Na+/K+-ATPase activity was determined in
cellular lysates by measuring the difference between total ATPase
activity and ouabain-insensitive ATPase activity as described
previously (37).
Measurement of Caspase-3 Activity--
Caspase-3 activity was
quantified by fluorometric detection of free AFC after cleavage from
DEVD-AFC. In brief, the media were aspirated from the culture dishes,
then the RPTC monolayers were washed with ice-cold PBS, scraped off
culture dishes, and spun down at 1,000 × g for 2 min.
The pellet was resuspended in cell lysis buffer (BioVision, Palo Alto,
CA), incubated on ice for 10 min, and centrifuged at 15,000 × g for 10 min at 4 °C. The pellets were discarded and the
supernatants used for caspase assays. Cell lysates were incubated for
1 h at 37 °C in the presence of reaction buffer optimized for
caspase activity assays (BioVision), 1 mM dithiothreitol,
and 50 µM DEVD-AFC. The fluorescence was read at 380/500
nm (excitation/emission), and the amount of product cleaved was
determined from the AFC standard curve.
Immunoblotting--
Immunoblot analysis was used for the
measurement of protein levels of both total and phosphorylated forms of
ERK1/2, PKC-, PKC-, and PKC- in the total RPTC homogenates and
RPTC mitochondria and also for the assessment of protein levels of
cytochrome c and active caspase 3 in RPTC cytosol. RPTC
homogenates and mitochondrial and cytosolic fractions of RPTC were
lysed and boiled for 10 min in Laemmli sample buffer (60 mM
Tris-HCl, pH 6.8 containing 2% SDS, 10% glycerol, 100 mM
-mercaptoethanol, and 0.01% bromphenol blue) (38). Proteins were
separated using SDS-PAGE. Following electroblotting of the proteins to
a nitrocellulose membrane, blots were blocked for 1 h in
Tris-buffered saline buffer containing 0.5% casein and 0.1% Tween 20 (blocking buffer), and incubated overnight at 4 °C in the presence
of primary antibodies diluted in the blocking buffer. Following
washing, the membranes were incubated for 1 h with anti-rabbit or
anti-mouse IgG coupled to horseradish peroxidase and washed again. The
supersignal chemiluminescent system was used for protein detection and
scanning densitometry for the quantification of results.
Assessment of Apoptosis--
RPTC nuclei were visualized by DAPI
staining. The monolayers were fixed in 3.7% formaldehyde for 15 min,
rinsed with PBS, and incubated in the presence of 8 µM
DAPI for 2 h at room temperature. Following staining, RPTC
monolayers were rinsed with PBS, coverslips mounted, and the nuclei
evaluated under a Zeiss Fluorescent Microscope. Total and apoptotic
nuclei were counted in 6-8 different areas of each monolayer using two
plates per each experimental group.
Protein Assay--
Protein concentration in all samples was
determined using bicinchoninic acid assay with bovine serum albumin as
the standard.
Statistical Analysis--
Data are presented as means ± S.E. and were analyzed for significance by analysis of variance.
Multiple means were compared using Student-Newman-Keuls test.
Statements of significance were based on p < 0.05. Renal proximal tubules isolated from an individual rabbit represented a
separate experiment (n = 1) consisting of data obtained
from two to four plates.
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RESULTS |
Oxygen Consumption (QO2)--
Basal QO2 in
control confluent quiescent RPTC monolayers was 21.1 ± 1.2 nmol
of O2/mg of protein/min. Exposure of RPTC to 5 and 10 µM cisplatin for 48 h had no effect on basal
QO2 (data not shown). Two-hour exposure to 50 µM cisplatin decreased basal QO2 by 23%, but
this decrease was transient and basal QO2 returned to
control level at 4 h of the treatment (Fig.
1A). However, at 24 h of
cisplatin exposure, basal QO2 in RPTC decreased by 49% and
was accompanied by cell death (Figs. 1A and
12B).
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Fig. 1.
The effect of cisplatin on basal oxygen
consumption (QO2) and oligomycin-sensitive QO2
(A) and uncoupled QO2
(B). RPTC were treated with 50 µM
cisplatin, and samples were taken at 1, 2, 4, 8, 12, and 24 h of
cisplatin exposure for measurements of QO2. Control RPTC
were treated with vehicle (Me2SO) alone. Basal
QO2 in controls did not change over the course of 24 h. Basal QO2 ( ) represents the total amount of oxygen
consumed by RPTC and was measured as described under "Experimental
Procedures." Oligomycin-sensitive QO2 ( ) was measured
in the presence of oligomycin (0.6 µg/ml) and calculated as a
difference between basal and oligomycin-insensitive QO2.
Uncoupled QO2 ( ) was measured after addition of FCCP (2 µM). Results are the average ± S.E. of six
experiments (RPTC isolations).
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Oligomycin-sensitive QO2 (a marker of oxidative
phosphorylation) in control confluent quiescent RPTC monolayers was
15.3 ± 1.0 nmol of O2/mg of protein/min. Exposure of
RPTC to 5 and 10 µM cisplatin for 48 h did not have
a significant effect on oligomycin-sensitive QO2 (data not
shown). Exposure to 50 µM cisplatin had effects on
oligomycin-sensitive QO2 similar to the effects it had on
basal QO2, but the decreases in oligomycin-sensitive
QO2 (42 and 66% at 2 and 24 h, respectively) were
more pronounced than the reduction in basal QO2 (Fig.
1A). Oligomycin-insensitive QO2 in RPTC was not
changed by cisplatin exposure.
Uncoupled QO2 (a marker of electron transport through the
respiratory chain) in confluent quiescent control RPTC monolayers was
51.6 ± 5.0 nmol of O2/mg of protein/min. Exposure of
RPTC to 5 and 10 µM cisplatin for 48 h had no effect
on uncoupled QO2 (data not shown). Exposure of RPTC to 50 µM cisplatin decreased uncoupled QO2 by 25 and 53% at 2 and 24 h of treatment, respectively (Fig.
1B). These data show that: 1) cisplatin induces a transient reduction in RPTC respiration during early exposure, followed by a
sustained decrease in RPTC respiration during later time points of
exposure; and 2) oxidative phosphorylation and the electron transport
chain are the targets of cisplatin in RPTC mitochondria.
F0F1-ATPase Activity and Intracellular ATP
Content--
As shown in Fig.
2A, exposure of RPTC for
1 h to 50 µM cisplatin decreased activity of
mitochondrial F0F1-ATPase by 43%. This decrease preceded the reductions in basal, oligomycin-sensitive, and
uncoupled QO2s (Figs. 1 and 2A). The activity of
F0F1-ATPase remained decreased until the end of
cisplatin exposure in RPTC (24 h).
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Fig. 2.
The effect of cisplatin on
F0F1-ATPase activity (A) and
intracellular ATP content (B). A, RPTC
were treated with 50 µM cisplatin and mitochondria were
isolated at 1, 2, 4, 8, 12, 18, and 24 h of cisplatin exposure.
The F0F1-ATPase activity assay was performed at
31 °C in 10 mM Tris-HCl, pH 8.2, containing 200 mM KCl, 3 mM MgCl2, and RPTC
mitochondria. The reaction was initiated by the addition of ATP (5 mM) and terminated after 5 min by adding 3 M
trichloroacetic acid to precipitate protein. The inorganic phosphate
concentration in the supernatant was determined using Sumner reagent.
Each mitochondrial sample was run in the absence and presence of
oligomycin (10 µg/ml), and the F0F1-ATPase
activity was expressed as the oligomycin-sensitive phosphate
production. Results are the average ± S.E. of three experiments
(RPTC isolations). B, RPTC were treated with 50 µM cisplatin and samples were taken at 1, 2, 4, 8, 18, and 24 h of cisplatin exposure for measurements of intracellular
ATP content. Results are the average ± S.E. of five experiments
(RPTC isolations).
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The intracellular ATP concentration was examined to determine whether
cisplatin-induced toxicity in RPTC was caused by decreases in
intracellular ATP levels. The ATP content in cisplatin-treated RPTC
decreased by 32 and 25% at 6 and 8 h of exposure, respectively, but was not different from control levels at 18 and 24 h of
exposure (Fig. 2B). The decreases in ATP content were
preceded by transient reductions in the basal, oligomycin-sensitive,
and uncoupled QO2s and the activity of
F0F1-ATPase. However, sustained decrease in oxidative phosphorylation at 24 h of cisplatin treatment did not affect ATP content in these cells. These data suggest that cisplatin reduces ATP-consuming processes in RPTC.
Mitochondrial Membrane Potential
(m)--
Mitochondrial respiration results in the
generation of a proton and pH gradients across the inner mitochondrial
membrane and produces the membrane potential (m),
which represents most of the energy of the proton gradient. Lipophilic
cations such as JC-1 accumulate in the mitochondrial matrix driven by
the electrochemical gradient (negative inside the mitochondrion). The
higher the m, the more polarized is the mitochondrial
membrane, and more JC-1 is taken up into the mitochondrial matrix. Once
taken up into the mitochondrial matrix, JC-1 forms aggregates that
fluoresce red (emission, 590 nm), whereas JC-1 in the cytosol exists in a monomeric form that fluoresces green (emission, 525 nm). Thus, an
increase in intensity of red fluorescence of JC-1 indicates higher
m and mitochondrial membrane hyperpolarization,
whereas loss of red and increased green fluorescence indicates
decreased m and mitochondrial membrane
depolarization. The ratio of red to green fluorescence is dependent
only on the mitochondrial membrane potential and not on the other
factors such as plasma membrane potential, mitochondrial size, shape,
and density that might affect a single component fluorescence signal
such as red fluorescence.
Using fluorescent microscopy, we determined that the red fluorescence
(JC-1 aggregates) was localized in the mitochondria (Fig.
3). Control RPTC had elongated
yellow/orange and red fluorescing mitochondria, against a diffused
greenish background corresponding to monomeric JC-1 in the cytoplasm
(Fig. 3A). The green (monomer) fluorescence corresponds to
values of m higher than 
140 mV (39). The orange
fluorescence in mitochondria reflects m of approximately 150 mV (40). Exposure of RPTC to cisplatin for 18 and
24 h resulted in an increase in red fluorescence in the mitochondria and a decrease in green fluorescence in the cytoplasm (Fig. 3, B and C). The increase in overall red
fluorescence was a result of both the increase in the intensity of
fluorescence in the mitochondria and the increase in the number of
red-fluorescing mitochondria. Exposure of control RPTC to mitochondrial
uncoupler (FCCP) resulted in the loss of red staining in RPTC,
indicating mitochondrial membrane depolarization (Fig.
3D).
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Fig. 3.
Mitochondrial hyperpolarization induced by
cisplatin exposure in RPTC. RPTC were treated with 50 µM cisplatin and loaded with 10 µM JC-1 for
30 min at 37 °C, washed twice with ice-cold PBS, and overlaid with
ice-cold PBS. The live RPTC monolayers were examined under Zeiss
fluorescent microscope (Axioskop) using water-immersion objective.
Original magnification, ×400. A, controls. B,
RPTC treated with 50 µM cisplatin for 18 h.
C, RPTC treated with 50 µM cisplatin for
24 h. Inset, a single cell (original magnification,
×800). D, mitochondrial depolarization in RPTC induced by
FCCP (2 µM). These images are representative of three
independent experiments (cell isolations).
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Flow cytometry was used for the quantification of m
in RPTC. m was assessed by the measurement of both
red fluorescence (JC-1 aggregates present in mitochondria) and
red/green fluorescence ratio (JC-1 aggregate/JC-1 monomer ratio). The
JC-1 aggregate/monomer ratio increased by 43% at 2 h of cisplatin
exposure (Fig. 5B). This increase was transient, and
m returned to control levels at 4 h of cisplatin
exposure. However, at 12, 18, and 24 h of cisplatin exposure,
m increased 1.4-, 1.9-, and 2.4-fold, respectively (Figs.
4A and
5, A and B).
Previously, it has been shown that a 12% change in fluorescence signal
of JC-1 reflects a 10-mV change in the membrane potential (41).
Therefore, the increases in JC-1 fluorescence at 12, 18, and 24 h
of cisplatin exposure in RPTC correspond to m of
180, 230, and 270 mV, respectively. The early (2 h) increase in
m followed the decrease in
F0F1-ATPase activity and was accompanied by
decreases in basal, oligomycin-sensitive, and uncoupled respiration of
RPTC (Figs. 1, 2A, and 5, A and B). These results show that cisplatin treatment in RPTC results in a
transient hyperpolarization of the mitochondrial membrane that occurs
early during the exposure and is followed by sustained hyperpolarization of the mitochondrial membrane that precedes RPTC
apoptosis.
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Fig. 4.
Quantification of JC-1 accumulation in
mitochondria (red fluorescence) and cytoplasm (green fluorescence) in
cisplatin-treated RPTC. RPTC monolayers were exposed to 50 µM cisplatin for 24 h and loaded with 10 µM JC-1 for 30 min at 37 °C. After loading, media were
aspirated and monolayers kept on ice, washed twice with ice-cold PBS,
scraped off culture dishes, washed, and resuspended in PBS.
Fluorescence was analyzed by flow cytometry (BD Biosciences
FACSCalibur) using excitation by 488 nm argon-ion laser. The JC-1
monomer (green fluorescence) and J-aggregate (red fluorescence) were
detected in FL1 (emission, 525 nm) and FL2 (emission, 590 nm) channels,
respectively. A, the effect of cisplatin on JC-1
accumulation in RPTC mitochondria. B, the effect of
inhibition of ERK1/2 (50 µM PD98059) on cisplatin-induced
accumulation of JC-1 in mitochondria. C, the effect of
inhibition of PKC- (10 nM Go6976) on cisplatin-induced
accumulation of JC-1 in mitochondria. Experiments were performed five
times with comparable results.
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Fig. 5.
The effect of cisplatin exposure on
m in RPTC. RPTC
monolayers were treated and analyzed as described in the legend to Fig.
4. A, the effect of cisplatin on the average red
fluorescence of JC-1 in RPTC. B, the effect of cisplatin on
the red/green fluorescence ratio of JC-1 in RPTC. Results are the
average ± S.E. of seven experiments (RPTC isolations).
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Cytochrome c Release--
Cisplatin exposure in RPTC induced the
release of cytochrome c from the mitochondria to the
cytosol. The protein levels of cytochrome c in the cytosolic
fraction of RPTC increased 5-fold at 6 h of cisplatin exposure and
continued to increase until 18 h of the treatment (Fig.
6). Cytochrome c translocation
to the cytosol followed the initial transient disruption of
mitochondrial function and preceded sustained decreases in oxidative
phosphorylation electron transport rate, and hyperpolarization of the
mitochondrial membrane (Figs. 1, 5, and 6). These data show that: 1)
cisplatin exposure in RPTC induces the release of cytochrome
c from mitochondria to the cytosol, and 2) cytochrome
c release from mitochondria occurs following initial
hyperpolarization of the mitochondrial membrane and the decrease in
respiration. These data provide additional evidence for the disruption
of mitochondrial function caused by cisplatin in RPTC.
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Fig. 6.
The effect of cisplatin exposure on the
release of cytochrome c to RPTC cytosol and activation
of caspase-3. At different time points of cisplatin exposure, RPTC
cytosol was isolated as described under "Experimental Procedures"
and protein levels of cytochrome c and active caspase 3 (17-19-kDa cleaved fragment) determined by immunoblotting.
A, the effect of cisplatin exposure on the release of
cytochrome c to the RPTC cytosol. B, the effect
of cisplatin on caspase-3 cleavage in RPTC. C, the effect of
ERK1/2 inhibition (50 µM PD98059) on caspase-3 cleavage
during cisplatin exposure in RPTC. D, the effect of PKC-
inhibition (10 nM Go6976) on caspase-3 cleavage during
cisplatin exposure in RPTC. Experiments were performed three times with
comparable results.
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Active Na+ Transport--
Active Na+
transport is an ATP-consuming process, as it is driven by
Na+/K+-ATPase. The consumption of oxygen
associated with production of ATP required for maintaining the
Na+/K+-ATPase activity and active
Na+ transport (ouabain-sensitive QO2) accounts
for ~50% of basal oxygen consumption in RPTC. Ouabain-sensitive
QO2 (used as a marker of active Na+ transport)
in control RPTC was 9.5 ± 1.1 nmol of
O2/mg of protein/min. One- and 2-h exposures to cisplatin
decreased ouabain-sensitive QO2 by 44 and 36%,
respectively. Ouabain-sensitive QO2 remained decreased
until 4 h of cisplatin exposure and transiently returned to
control levels at 8 and 12 h of treatment (Fig.
7A). This transient increase
in ouabain-sensitive QO2 between 8 and 12 h was
accompanied by a decrease in intracellular ATP content (Figs. 2 and
7A). At 24 h of cisplatin exposure, ouabain-sensitive
QO2 was reduced by 67% in comparison with controls (Fig.
7A). The decline in ouabain-sensitive QO2
occurred prior to any significant decreases in basal,
oligomycin-sensitive, or uncoupled QO2 (Figs. 1 and 6).
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Fig. 7.
The effect of cisplatin on active
Na+ transport. A, the effect of cisplatin
on ouabain-sensitive QO2. RPTC were treated with cisplatin,
and samples were taken at 1, 2, 4, 8, 12, and 24 h of exposure for
measurements of QO2. Ouabain-sensitive QO2 in
controls did not change over the course of 24 h. Ouabain-sensitive
QO2 was measured in the presence of 1.0 mM
ouabain and calculated as a difference between basal and
ouabain-insensitive QO2. B, the effect of
cisplatin on the activity of Na+/K+-ATPase in
RPTC. RPTC were treated with 50 µM cisplatin, and samples
were taken at 2 and 24 h of exposure for measurements of
Na+/K+-ATPase activity as described under
"Experimental Procedures."  , control RPTC treated with vehicle
(Me2SO); , RPTC treated with 50 µM
cisplatin. Results are the average ± S.E. of six independent
experiments (RPTC isolations).
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The activity of Na+/K+-ATPase was decreased 25 and 69% at 2 and 24 h of cisplatin exposure, respectively (Fig.
7B). These data show that the cisplatin-induced decrease in
active Na+ transport in RPTC: 1) is an early event and
occurs prior to any alterations in mitochondrial function, and 2) is
not caused by reduced levels of intracellular ATP. These data also
suggest that one of the major ATP-consuming processes (active
Na+ transport) is significantly reduced early during
cisplatin exposure.
Activation of PKC and ERK by Cisplatin--
Fig.
8A shows that exposure of RPTC
to 50 µM cisplatin was associated with the activation of
PKC-. The levels of phosphorylated PKC- in cell homogenate
increased 1.8- and 2.5-fold at 0.5 and 1 h of cisplatin exposure,
respectively, and remained increased within the first 8 h of
treatment (Fig. 8A). Go6976 (10 nM), an inhibitor of PKC-, abolished the phosphorylation of PKC- in cisplatin-treated RPTC (Fig. 8C). In contrast, cisplatin
exposure had no effect on PKC- or PKC- (Fig. 8, E and
F). Phosphorylated PKC- was also present in RPTC
mitochondria (Fig. 8G). However, cisplatin did not affect
the phosphorylation of mitochondrial PKC- (Fig. 8G).
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Fig. 8.
The effect of cisplatin on
PKC-, PKC-, and
PKC- in RPTC. RPTC were treated with 50 µM cisplatin and samples were taken at 0, 0.5, 1, 2, 4, 6, 8, 12, 18, and 24 h for measurements of protein levels of
phosphorylated (active) and total PKC-, PKC-, and PKC- using
immunoblotting. Samples were processed as described under
"Experimental Procedures" and proteins separated using 10%
SDS-PAGE. Following electroblotting of the proteins to a nitrocellulose
membrane, blots were blocked for 1 h in Tris-buffered saline
containing 0.5% casein and 0.1% Tween 20, and incubated overnight at
4 °C in the presence of anti-phospho-PKC-, anti-phospho-PKC-,
anti-phospho-PKC- antibodies, or anti-PKC- antibody diluted in
the blocking buffer. Following washing, the membranes were incubated
for 1 h with anti-rabbit or anti-mouse IgG coupled to horseradish
peroxidase and washed again. The supersignal chemiluminescent system
was used for protein detection and scanning densitometry for
quantification of results. A-F, protein levels of
phospho-PKC-, phospho-PKC-, and phospho-PKC- in RPTC
homogenates. G-J, protein levels of phospho-PKC- in RPTC
mitochondria. Go6976 (10 nM) was added 1 h prior to
cisplatin exposure. Presented data are representative of three
independent experiments (cell isolations).
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Treatment of RPTC with cisplatin also resulted in the activation of
ERK1/2 (Fig. 9A).
Phosphorylation of ERK1/2 in the cell homogenate was increased by 50%
at 0.5 h and 4-fold at 1 h of cisplatin exposure (Fig. 9,
A and I). At 24 h of cisplatin treatment, the phosphorylation of ERK1/2 was increased 17-fold in comparison with
controls (Fig. 9, A and I). Cisplatin exposure
also induced ERK1/2 activation in RPTC mitochondria (Fig.
9E). Protein levels of active ERK1/2 were very low in the
mitochondria of control RPTC but increased 4-,and 40-fold in
mitochondria of cisplatin-treated RPTC at 2 and 24 h of the
exposure (Fig. 9E). Total ERK1/2 protein was abundant in the
mitochondria of control RPTC and was not altered by cisplatin exposure
(Fig. 9G), which suggested that the increase in
phosphorylated ERK1/2 levels in the mitochondria of cisplatin-treated RPTC was not a result of ERK1/2 translocation to mitochondria. Pretreatment of RPTC with 50 µM PD98059 prevented the
phosphorylation of ERK1/2 in cell homogenates and mitochondria isolated
from cisplatin-treated RPTC (Fig. 9, B and
F).
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Fig. 9.
Activation of ERK1/2 during cisplatin
exposure in RPTC. RPTC were treated with 50 µM
cisplatin, and samples were taken at 0, 0.5, 1, 2, 4, 6, 8, 12, 18, and
24 h for measurements of protein levels of phosphorylated and
total ERK1/2 using immunoblotting. Immunoblotting was performed as
described in the legend to Fig. 8. A-D, protein levels of
phospho- and total ERK1/2 in RPTC homogenates. E-H, protein
levels of phospho- and total ERK1/2 in RPTC mitochondria. PD98059 (50 µM) and Go6976 (10 nM) were added 1 h
prior to cisplatin treatment. Presented data are representative of
three independent experiments (cell isolations). I,
cisplatin-induced ERK1/2 activation quantified by densitometry. Results
are the average ± S.E. of three independent experiments (RPTC
isolations).
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These data show that cisplatin activates PKC- and ERK1/2 in RPTC but
has no effect on PKC- or PKC-. Our results also demonstrate that
cisplatin exposure results in ERK1/2 activation in RPTC mitochondria.
Effect of Inhibition of PKC- Activation on Cisplatin-induced
Changes in Mitochondrial Function and Active Na+
Transport--
We have examined whether PKC- activation is involved
in mediating cisplatin-induced decreases in mitochondrial function and active Na+ transport. Treatment of RPTC with Go6976 alone
for 24 h had no effect on oligomycin-sensitive QO2
(12.0 ± 0.9 versus 12.7 ± 1.4 nmol of
O2/min/mg of protein in control and Go6976-treated cells, respectively), m (aggregate/monomer fluorescence
ratio: 1.6 ± 0.5 versus 1.5 ± 0.1 in control and
Go6976-treated cells, respectively), ouabain-sensitive QO2
(8.5 ± 0.8 versus 9.1 ± 0.8 nmol of
O2/min/mg of protein in control and Go6976-treated cells,
respectively), and Na+/K+-ATPase activity
(290 ± 6 versus 254 ± 64 milliunits/mg of
protein in control and Go6976-treated cells, respectively).
Pretreatment of RPTC with Go6978 prior to cisplatin exposure had no
effect on transient decreases in oligomycin-sensitive QO2
at 2 h but prevented sustained decreases in this function at
24 h of treatment (Fig.
10A). Likewise, inhibition
of PKC- activation by cisplatin had no effect on the early
hyperpolarization of the mitochondrial membrane but prevented the
sustained increase in m at 24 h of the exposure
(Figs. 4C and 10C). Consistent with the lack of
effect on the early and transient mitochondrial dysfunction, inhibition
of PKC- activation did not prevent translocation of cytochrome
c from mitochondria to the cytosol (data not shown). Furthermore, pretreatment of RPTC with Go6976 prevented reduction in
Na+/K+-ATPase activity and ouabain-sensitive
QO2 in cells treated with cisplatin for 24 h (Fig.
11, A and B).
However, the initial transient decreases in
Na+/K+-ATPase activity and ouabain-sensitive
QO2 were independent of PKC- activation (Fig. 11,
A and B). In addition, PKC- inhibitor did not
prevent cisplatin-induced decreases in uncoupled QO2 (Fig. 10C).
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Fig. 10.
The effect of inhibition of PKC- and
ERK1/2 activation on transient (2 h) and sustained (24 h) changes in
oxidative phosphorylation (A), electron transport rate
(B), and the mitochondrial membrane potential
(C) induced by cisplatin (50 µM) in RPTC.
The monolayers were treated and RPTC functions analyzed as
described under "Experimental Procedures." White
columns, controls; black columns, 50 µM cisplatin; light gray
columns, 50 µM PD98059 + 50 µM
cisplatin; dark gray columns, 10 nM Go6976 + 50 µM cisplatin;
hatched columns, 3 µM UO126 + 50 µM cisplatin; striped columns, 50 µM zVAD-fmk + 50 µM cisplatin. Results are
the average ± S.E. of three to five independent experiments (RPTC
isolations).
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Fig. 11.
The effect of inhibition of PKC- and
ERK1/2 activation on transient (2 h) and sustained (24 h) changes in
ouabain-sensitive oxygen consumption (A) and
Na+/K+-ATPase activity (B) induced
by cisplatin (50 µM) in RPTC. The monolayers were
treated and ouabain-sensitive oxygen consumption and
Na+/K+-ATPase activity analyzed as described
under "Experimental Procedures." White
columns, controls; black columns, 50 µM cisplatin; light gray
columns, 50 µM PD98059 + 50 µM
cisplatin; dark gray columns, 10 nM Go6976 + 50 µM cisplatin. Results are the
average ± S.E. of three independent experiments (RPTC
isolations).
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These data show that the inhibition of cisplatin-induced activation of
PKC-: 1) prevents late and sustained (24 h) but not transient (2 h)
decreases in oxidative phosphorylation and active Na+
transport, 2) prevents sustained (24 h) but not transient (2 h)
increases in m, 3) has no effects on
cisplatin-induced alterations in electron transport chain, and 4) does
not block the release of cytochrome c from mitochondria.
Therefore, our results show that PKC- mediates sustained decreases
in hyperpolarization of the mitochondrial membrane and oxidative
phosphorylation but not the decreases in electron transfer rate.
Moreover, the sustained decrease in active Na+ transport in
cisplatin-treated RPTC is also mediated by PKC-. Our data also
suggest that PKC- mediates events that occur after the release of
cytochrome c from mitochondria.
Effect of Inhibition of ERK Activation on Cisplatin-induced Changes
in Mitochondrial Function and Active Na+ Transport--
We
next addressed whether activation of ERK1/2 plays a role in the
cisplatin-induced changes in mitochondrial function and active
Na+ transport. RPTC were pretreated with 50 µM PD98059, a specific MEK1 inhibitor, to inhibit
cisplatin-induced ERK activation. Oxidative phosphorylation,
electron transport rate, m, and active
Na+ transport were assessed at 2 and 24 h of cisplatin
exposure in PD98059-pretreated RPTC. Treatment of RPTC for 24 h
with PD98059 alone had no effect on oligomycin-sensitive
QO2 (12.0 ± 0.9 versus 11.5 ± 1.5 nmol of O2/min/mg of protein in control and PD98059-treated cells, respectively), m (aggregate/monomer
fluorescence ratio: 1.7 ± 0.4 versus 2.1 ± 0.4 in control and PD98059-treated cells, respectively), ouabain-sensitive
QO2 (8.5 ± 0.8 versus 8.7 ± 1.1 nmol
of O2/min/mg of protein in control and PD98059-treated
cells, respectively), and Na+/K+-ATPase
activity (290 ± 6 versus 291 ± 22 milliunits/mg
of protein in control and PD98059-treated cells, respectively).
Treatment of RPTC with PD98059 prior to cisplatin exposure had no
effect on the transient decrease in oligomycin-sensitive QO2 at 2 h but reduced sustained decreases in
oligomycin-sensitive QO2 at 24 h of the exposure (Fig.
10A). Likewise, pretreatment of RPTC with ERK1/2 inhibitor
prior to cisplatin exposure did not abolish the transient
hyperpolarization of the mitochondrial membrane at 2 h but
prevented the sustained increases in m (Figs. 4B and 10C). Another MEK1 inhibitor, UO126 (3 µM), also abolished the sustained increases in
m in cisplatin-treated RPTC (Fig. 10C).
Consistent with the lack of effect on transient mitochondrial dysfunction, inhibition of ERK1/2 activation did not prevent
translocation of cytochrome c from mitochondria to the
cytosol (data not shown). In contrast, inhibition of the ERK1/2
activation had no effect on cisplatin-induced decreases in the electron
transport rate (Fig. 10B).
Inhibition of the ERK activation did not affect the transient decrease
in ouabain-sensitive QO2 and
Na+/K+-ATPase activity in cisplatin-treated
RPTC but fully protected against sustained decreases in these functions
(Fig. 11, A and B).
Interestingly, inhibition of PKC- activation in cisplatin-treated
RPTC did not abolish ERK1/2 phosphorylation (Fig. 9, D and
H), which suggests that ERK1/2 activation by cisplatin does not occur through a PKC--mediated pathway.
These data demonstrate that inhibition of cisplatin-induced ERK1/2
activation: 1) reduces, in part, sustained but not transient decreases
in oxidative phosphorylation; 2) prevents sustained increases in
m in cisplatin-treated RPTC; and 3) prevents
sustained decreases in active Na+ transport. Thus, our
results show that ERK1/2 mediates cisplatin effects on sustained
changes in mitochondrial functions of RPTC such as hyperpolarization of
the mitochondrial membrane and decreases in oxidative phosphorylation
but not on the decreases in the electron transport rate. Furthermore,
the sustained decrease in the active Na+ transport during
cisplatin exposure is also mediated by ERK1/2. Finally, our data
suggest that ERK1/2 and PKC- mediate cisplatin-induced mitochondrial
dysfunction and decreases in active Na+ transport through
independent signaling pathways.
Caspase Activation--
Caspase-3 cleavage and activity were
evaluated to determine whether the alterations in mitochondrial
function during cisplatin exposure are associated with caspase
activation. No cleavage of caspase-3 was observed during the first
6 h of cisplatin exposure. The first evidence of the cleaved form
of caspase-3 was found at 12 h, and the protein levels of the
cleaved caspase-3 further increased at 18 and 24 h of cisplatin
exposure (Fig. 6B). At 12 and 24 h of cisplatin
exposure, caspase-3 activity was increased 3.9-fold (2.49 ± 0.14 nmol/h/mg of protein) and 14.3-fold (9.12 ± 1.93 nmol/h/mg of
protein), respectively, in comparison with controls (0.64 ± 0.27 nmol/h/mg of protein). Pretreatment of RPTC with 50 µM
PD98059 prior to cisplatin exposure abolished caspase-3 cleavage and
decreased caspase-3 activity by 44% at 24 h of cisplatin treatment (Figs. 6C and
12A). Similarly,
pretreatment with another specific MEK inhibitor, UO126, decreased
cisplatin-induced caspase-3 activity by 36% at 24 h of the
exposure (Fig. 12A). Likewise, the pretreatment of cells
with Go6978 (PKC- inhibitor) abolished cisplatin-induced caspase-3
cleavage and decreased caspase-3 activity by 38% (Figs. 6D
and 12B). We also addressed whether caspase activation plays
a role in cisplatin-induced hyperpolarization of the mitochondrial membrane. Pretreatment of RPTC with caspase inhibitor, 50 µM zVAD-fmk, prior to cisplatin treatment inhibited
caspase-3 activation (Fig. 12B) but did not prevent
increases in m (Fig. 10C).
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Fig. 12.
The effect of cisplatin on caspase-3.
Caspase-3 activity was quantified by fluorometric detection of free AFC
after cleavage from 50 µM DEVD-AFC, and the amount of
product cleaved was determined from the AFC standard curve.
A, the effect of inhibitors of ERK1/2 activation (50 µM PD98059 and 3 µM UO126) on cisplatin (50 µM)-induced caspase-3 activation. B, the
effect of inhibition of PKC- activation (10 nM Go6976)
or treatment with caspase inhibitor (50 µM Z-VAD) on
cisplatin (50 µM)-induced caspase-3 activation. Results
are the average ± S.E. of four to nine independent experiments
(RPTC isolations).
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These data show that: 1) cisplatin exposure activates caspase-3, 2)
inhibition of ERK1/2 and PKC- activation decreases cisplatin-induced caspase-3 activation, and 3) inhibition of caspase activation does not
block mitochondrial membrane hyperpolarization.
Assessment of Cell Death--
Cisplatin exposure induced changes
in the RPTC nuclear morphology consistent with apoptosis.
Treatment of RPTC with cisplatin for 24 h resulted in chromatin
condensation and nuclear fragmentation in 44.7 ± 3.6% cells
present in the monolayers (versus 3.0 ± 0.6% in Me2SO-treated controls)
(Fig. 13, A and
B; Table I). These changes
were preceded by caspase-3 activation (12 h) and sustained increases
m (starting at 12 h). The inhibition of ERK
activation (using PD98059) prior to cisplatin exposure decreased the
number of apoptotic cells to 12.0 ± 0.6% (Fig. 13D;
Table I). The inhibition of PKC- activation (using Go6976) prior to
cisplatin treatment resulted in a decrease in RPTC apoptosis to
18.0 ± 4.6% (Fig. 13F; Table I). The exposure of RPTC
to PD98059 or Go6976 alone for 24 h had no effects on nuclear
morphology and cell viability (Fig. 13, C and E;
Table I). Pretreatment with a general caspase inhibitor, 50 µM zVAD-FMK, did not protect against cisplatin-induced apoptosis in RPTC (data not shown).
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Fig. 13.
Cisplatin-induced alterations in nuclear
morphology in RPTC. At 24 h of cisplatin exposure, RPTC
monolayers were fixed in 3.7% formaldehyde, incubated in the presence
of 8 µM DAPI for 2 h at room temperature, and
evaluated under Zeiss fluorescent microscope (Axioskop). Pictures were
taken using Hamamatsu color chilled 3CCD digital camera. A,
control; B, 50 µM cisplatin; C, 50 µM PD98059; D, 50 µM PD98059 + 50 µM cisplatin; E, 10 nM Go6976;
F, 10 nM Go6976 + 50 µM cisplatin.
Presented data are representative of three independent experiments
(cell isolations). Original magnification, ×400.
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Table I
Percentage of nuclear condensation and fragmentation at 24 h of
cisplatin exposure in RPTC
Values with different letters (a, b,
c) are significantly different (P <0.05)
from each other.
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DISCUSSION |
The mechanisms and pathway(s) leading to cisplatin-induced RPTC
apoptosis, including the effects of cisplatin on RPTC mitochondria and
the role of mitochondrial dysfunction in cisplatin nephrotoxicity, have
not been elucidated. Therefore, we examined whether cisplatin, used at
pharmacological concentrations, induces mitochondrial dysfunction and
activates the mitochondrial stress pathway that leads to apoptosis. Our
results demonstrated that oxidative phosphorylation and the electron
transport chain were the mitochondrial targets of cisplatin
and suggested that the decrease in oxidative phosphorylation was a
result of the inhibition of mitochondrial
F0F1-ATPase activity. The decreases in RPTC
oxidative phosphorylation and electron transport rate were accompanied
by hyperpolarization of the mitochondrial membrane. Our observation is
in contrast with reports showing that apoptosis is associated with a
decrease in m caused by unrestrained opening of the
mitochondrial permeability transition pore (MPTP), which results in the
release of proapoptotic proteins from the mitochondrial intermembrane
space to the cytoplasm and activation of caspases (42-44). It has also
been suggested that cisplatin causes a decline in m
and opening of MPTP in RPTC as assessed by rhodamine 123 uptake (33,
45). However, these studies used a high concentration of cisplatin (2 mM) that causes oncosis and mitochondrial depolarization
rather than apoptosis. The concentration of cisplatin used in our
experiments (50 µM) resulted in RPTC apoptosis, and this
event was preceded by sustained hyperpolarization of the
mitochondrial membrane as demonstrated by both flow cytometry and
fluorescent microscopy.
Recent studies suggest that the initiation of apoptotic events is not
always associated with MPTP but, in some cases, with mitochondrial
membrane hyperpolarization and mitochondrial shrinkage (46, 47).
Apoptosis caused by the withdrawal of growth factors or interlukin-3
from cultured cells is associated with cytosolic acidification and
hyperpolarization of the inner mitochondrial membrane, which are
dependent on ATP synthase activity (48, 49). Furthermore,
staurosporine-induced apoptosis in neuronal cells is associated
with hyperpolarization of the mitochondrial inner membrane, which
precedes the release of cytochrome c (46, 50).
Dissipation of mitochondrial K+ and H+
gradients inhibits staurosporine-induced cytochrome c
release and attenuates apoptosis (46). In most eukaryotic cells,
m is generated by the respiratory chain or through
ATP-dependent reversal of mitochondrial ATP synthase. The
energy released by the electron transport pumps protons across the
mitochondrial inner membrane, generating an electrochemical and pH
gradient. Our results suggest that the increase in proton gradient was
not generated by an accelerated rate of electron transport but was caused by decreased activity of H+ pump of
F0F1-ATPase. This decrease preceded the
increase in m and the decrease in electron flow rate
through the respiratory chain. Hyperpolarization of the inner
mitochondrial membrane could be explained by the inability of
F0F1-ATPase to efficiently pump protons back
into the mitochondrial matrix through the proton channel of ATP
synthase. This would result in the accumulation of protons in the
cytoplasm and an increase in the proton gradient between the
mitochondrial matrix and the intermembrane space.
An alternative mitochondrial mechanism that leads to an increase in
m is the closure of the voltage-dependent
anion channel (VDAC) localized on the outer mitochondrial membrane
(51). The closure of VDAC results in a loss of permeability of the
outer mitochondrial membrane to various ions, limits metabolite flux across the outer membrane, inhibits mitochondrial respiration and
oxidative phosphorylation, and causes mitochondrial hyperpolarization (52, 53). Persistent loss of the outer membrane permeability leads to a
disruption of the mitochondrial membrane ion homeostasis, loss of the
membrane integrity, cytochrome c redistribution to the
cytosol, and apoptosis (52). VDAC closure causes hyperpolarization of the mitochondrial membrane, release of cytochrome c, and
apoptosis in fibroblasts following growth factor withdrawal (48).
Bcl-xL prevents apoptosis through promoting the open configuration of VDAC, which suggests that maintenance of the outer mitochondrial membrane permeability is necessary for cell survival (53). An inhibitory effect of cisplatin on VDAC could explain both the decreased
oxidative phosphorylation and the elevated m in
cisplatin-treated RPTC.
Another possible explanation for the increase in m is
the disruption of RPTC mitochondrial ion and volume homeostasis by cisplatin. It is possible that alterations in cytosolic K+,
Na+, and/or H+ concentrations contributed to
the hyperpolarization of the mitochondrial membrane. Our results
demonstrate that a marked decrease in active Na+ transport
(and Na+/K+-ATPase activity) paralleled the
decrease in F0F1-ATPase activity and preceded
increases in m in cisplatin-treated RPTC. Such a
large decrease in Na+/K+-ATPase activity would
cause a rapid decrease in cytosolic K+ and an increase in
cytosolic Na+ concentrations, and lead to increases in
cytosolic H+ concentration. Because ouabain-sensitive
QO2 in cisplatin-treated RPTC was decreased prior to
mitochondrial dysfunction and the increase in m, it
is likely that the changes in cytosolic ion levels preceded and drove
the subsequent changes in the mitochondrial membrane potential.
However, it is unlikely that these changes were the sole mechanism of
incr
and ERK1/2 Mediate Mitochondrial
Dysfunction, Decreases in Active Na+ Transport, and
Cisplatin-induced Apoptosis in Renal Cells*
yna
Nowak
TOP
ABSTRACT
INTRODUCTION
