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Vol. 280, Issue 2, 638-649, 1997
Division of Toxicology, Leiden/Amsterdam Center for Drug Research, Leiden University, 2300 RA Leiden, The Netherlands
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Abstract |
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 Top
Abstract
Introduction
Materials & Methods
Results
Discussion
References
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Cisplatin-induced nephrotoxicity was studied in porcine proximal
tubular cells, focusing on the relationship between mitochondrial damage, reactive oxygen species (ROS) and cell death. Cisplatin specifically affected mitochondrial functions: complexes I to IV of the
respiratory chain were inhibited 15 to 55% after 20 min of incubation
with 50 to 500 µM, respectively. As a result, intracellular ATP was
decreased to 70%. The mitochondrial glutathione (reduced form)
(GSH)-regenerating enzyme GSH-reductase (GSH-Rd) activity was reduced
by 20%, which contributed to a 70% reduction of GSH levels and ROS
formation. The residual electron flow through the mitochondrial
respiratory chain was the source of ROS because additional inhibition
of the complexes I to IV reduced ROS formation. Because cisplatin
affects both GSH-Rd and complexes I to IV, cells were incubated with
N,N
-bis(2-chloroethyl)-N-nitrosourea (inhibitor of GSH-Rd) and
inhibitors of the different complexes. Only
N,N-bis(2-chloroethyl)-N-nitrosourea with rotenone (complex I
inhibitor) induced ROS formation, which indicates that inhibition of
complex I and inhibition of the GSH-Rd is probably the cause of ROS
formation. However, the resulting ROS is not the cause of cell death
because diphenyl-p-phenylene-diamine and deferoxamine,
which completely prevented ROS, could not prevent cell death.
Similarly, the antioxidants did not completely prevent the decrease in
activity of complexes I to IV, ATP or GSH levels. In conclusion, ROS
formation does occur during cisplatin-induced toxicity, but it is not
the direct cause of cell death.
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Introduction |
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Top
Abstract
Introduction
Materials & Methods
Results
Discussion
References
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The antitumor drug cisplatin is
used for the treatment of cancer of a wide range of tissues (Borch,
1987
). The major side effect, nephrotoxicity, is dose limiting and
occurs either acutely or after repeated treatments. In rats, cisplatin
exerts its effects mainly in the S3 segment of the proximal
tubule, which results in necrotic lesions (Doyban et al.,
1980); in man, additional damage is found in the distal part of the
tubules (Gonzalez-Vitale et al., 1977). A wealth of
histopathological data, mainly in rats and mice, is available about
cisplatin-induced nephrotoxicity; in addition, several biochemical
effects have been described in vitro as well as in
vivo, among which alterations in capacity of several active
transport systems (Miura et al., 1987), loss of
mitochondrial function (Gordon and Gattone, 1986),
Na+,K+-ATPase activity (Brady et
al., 1993; Kim et al., 1995) and lipid peroxidation
(Zhang et al., 1994; Hannemann and Baumann, 1988). However,
the sequence and relative importance of these effects are still
unclear.
Mitochondria seem to play an important role in cisplatin-induced
nephrotoxicity (Gordon and Gattone, 1986; Brady et al.,
1993;). Accordingly, we demonstrated that exposure of freshly isolated PPTC in suspension to cisplatin resulted in loss of mitochondrial membrane potential (
) (Kruidering et al., 1994). The
decrease in preceded cell death, implying that damage to the
mitochondria is an early event in the cascade of events leading to cell
death. However, how cisplatin damages the mitochondria remains unclear.
Oxidative damage has been proposed as a mechanism of cisplatin-induced
renal cell death in vitro as well as in vivo
(Kameyama and Gemba, 1991a; Zhang and Lindup, 1994; Sugihara et
al., 1987; Gemba et al., 1988). However, these studies
did not investigate the relationship between mitochondrial dysfunction
and oxidative damage, leaving the question of cause and consequence
unanswered. Mitochondria continuously convert 1 to 2% of the consumed
oxygen to superoxide (Richter et al., 1995); therefore, they
are an important source of ROS. Superoxide anion is produced in the
respiratory chain by reaction of oxygen with iron-sulfur centers in
complex I and by partially reduced ubiquinone and cytochrome b in
complex III. A simplified scheme of the respiratory chain is given in figure 1.
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Mitochondria are protected from oxidative damage in several ways.
Superoxide and hydrogen peroxide are metabolized by Mn-containing superoxide dismutase and the Se-containing glutathione peroxidase, respectively. Vitamins C and E, glutathione and ubiquinol-10 are important for additional scavenging of ROS. Clearly, mitochondria both
form and scavenge ROS; disturbances of the balance, caused by changes
in the electron flow or in the defense mechanisms, can lead to an
overproduction of ROS. ROS attack lipids, proteins and nucleic acids
nonspecifically, which results in (more mitochondrial) dysfunction
(Richter et al., 1995).
In this study, we investigated the roles of mitochondrial dysfunction and formation of ROS in cisplatin-induced cell death of PPTC in detail. Our results suggest that impaired respiratory activity rather than the formation of ROS is the key event in the pathway leading to renal cell death.
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Materials and Methods |
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Top
Abstract
Introduction
Materials & Methods
Results
Discussion
References
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Chemicals
BSA, glycine, collagenase (EC3.4.24.3) (from Clostridium
histolyticum), cis-diamine-dichloroplatinum (II)(Pt),
R123, propidium iodide, ethyleneglycol-bis-(
-aminoethylether)
N,N,N,N-tetraacetic acid, ubiquinone, cytochrome cIII,
N-D-maltoside, Oxa, rot, TTFA, anti, firefly lantern
extract (crude luciferin/luciferase), GSH and GSSG, GSH-Rd, TBHP and
cumene hydroperoxide were from Sigma Chemical Co. (St. Louis, MO).
DPPD was purchased from Aldrich (Brussels, Belgium). Potassium cyanide
(KCN) was from Janssen Chimica (Tilburg, the Netherlands). HEPES and
sodium pyruvate were obtained from Boehringer Mannheim (Mannheim,
Germany). Nycodenz (Iohexol) was from Nycomed AS (Oslo, Norway).
DesferalR (deferoxamine) was from Ciba-Geigy AG (Basle,
Switzerland). Diethylmaleate was from Merck (Darmstadt, Germany).
(R,S)-3-Hydroxy-4-pentenoic acid was synthesized
as described in Shan et al. (1993). Dih123 was from
Molecular Probes (Eugene, OR). BCNU was kindly provided by Prof. E.W.
Vogel, Leiden University.
Cell Isolation
Cell isolation was performed as described before (Kruidering
et al., 1993). The kidneys with intact capsule were taken
from pigs in the slaughterhouse approximately 15 to 20 min after
electrocution; the renal artery and vein were flushed with ice-cold
Eurocollins, pH 7.4, consisting of 177 mM glucose, 10 mM
NaHCO3, 15 mM KCl, 42 mM K2HPO4 and
15 mM KH2PO4, supplemented with 2 mM glycine. After transport to the laboratory on ice, the capsule was removed and
the cortex was dissected on ice. The minced cortex was washed three
times with Ca++-and Mg++-free Hanks-HEPES
buffer, pH 7.4, containing 25 mM HEPES and 2 mM glycine (buffer A).
Thirty grams of cortex were incubated at 37°C with 25 ml of 0.07%
(w/v) collagenase in buffer A, containing 1 mM deferoxamine and 4 mM
CaCl2. After 55 min, 125 ml cold buffer A, including 4 mM
CaCl2 and 1.5% BSA (w/v) (buffer C), was added to stop
degradation. The cell suspension was filtered through a nylon gauze
with a pore size of 80 µm and washed twice by centrifugation (5 min,
80 × g 4°C). The suspension was mixed and proximal
tubular cells were purified from this suspension by centrifugation,
with use of a discontinuous density gradient consisting of layers of 17%, 11.3% and 8.5% (w/v) Nycodenz, which was prepared as follows. A
34% (w/v) stock of Nycodenz was prepared in a solution containing 6.7 mM KCl, 1.22 mM CaCl2, 10 mM HEPES, pH 7.4. This was mixed 1:1 and 1:3 with buffer C to obtain 17% and 8.5% (w/v), respectively. The cells were resuspended in 20 ml buffer and mixed with 10 ml 34%
Nycodenz to obtain 11.3%. The gradient was layered carefully in two
50-ml glass tubes. First, 5 ml of 17% Nycodenz was put in the tube; on
top of this, 10 ml of cells in 11.3% Nycodenz was layered carefully
without mixing the layers. Finally, this was topped with 5 ml 8.5%
Nycodenz solution. The tubes were centrifuged for 6 min at 2300 × g 4°C. The cells which accumulated in bands at the
interface between the upper two layers were removed with a capillary
pipette and pooled by centrifugation in buffer C (90 × g for 5 min at 4°C). Subsequently, cells were washed twice
with buffer C (80 × g for 6 min at 4°C). The maximum
period between the death of the animals and the end of the isolation
was 2.5 h. Viability was determined by trypan blue exclusion.
Purity of the resulting cell suspension was determined by counting the
percentage 
-glutamyl transpeptidase positive cells (Rutenberg
et al., 1969) and by determination of the -glutamyl
transpeptidase activity of the cells before and after purification. The
final cell preparation, lacking distal tubular and endothelial cells,
had a viability of more than 90% and purity of at least 80% and
routinely 90% [i.e., -glutamyl transpeptidase positive
cells (Kruidering et al., 1993)].
Incubations
Freshly isolated cells were incubated in buffer C, at a density of 0.5·106 cells/ml at 37°C in a humidified atmosphere (95% air/5% CO2) to allow the cells to recover from isolation stress. After 20 min, cells were centrifuged and resuspended in buffer C without BSA (buffer D). If necessary, cells were preincubated for 20 min as indicated in the legends.
At t = 0 cisplatin (from a freshly prepared 100 mM
stock in dimethyl sulfoxide) was added to the cells; the cells were
gently shaken every 10 min. At the indicated time points, samples were taken and analyzed as described below. The samples for flow cytometry were taken directly from the incubation tubes, whereas the samples for
determination of enzymatic activities, or ATP and GSH content were
washed twice with buffer, snap-frozen in liquid nitrogen and stored at
80°C until use. Incubations for determination of ROS were performed
as described below.
Flow Cytometry
and viability.
and viability were determined
by analyzing the R123 and propidium iodide fluorescence intensity with
a FACScan flow cytometer (Becton Dickinson, San Jose, CA) equipped with
an argon laser, with the Lysis software program (Becton Dickinson).
R123 is a cationic dye that accumulates in the negatively charged inner side of the mitochondria. When the potential drops, less R123 accumulates in the mitochondria, which results in a lower fluorescence signal. The potential was measured as follows: at the indicated times,
a 500-µl sample of the cell suspension was taken and transferred to
an Eppendorf minivial. To this sample, 100 µl of 6 µM R123 in
buffer D was added. After incubation for 10 min at 37°C, the cell
suspension was centrifuged for 5 min at 80 × g. The
cell pellet was resuspended in 200 µl of buffer D, containing 0.2 µM R123 and 10 µM propidium iodide, to prevent loss of R123 and to stain nonviable cells, respectively. The samples were transferred to
FACScan tubes and analyzed immediately. Analysis was performed at a
flow rate of 60 µl/min. R123 fluorescence was detected by the FL1
detector with an emission detection limit below 560 nm. Propidium
iodide fluorescence was detected by the FL3 detector, with emission
detection above 620 nm. Per sample 3,000 to 5,000 cells were counted
(Van de Water et al., 1993).
ROS.
For determination of ROS, samples taken at the
indicated time points were directly transferred to FACScan tubes.
Dih123 (10 µM, final concentration) was added and cells were
incubated at 37°C in a humidified atmosphere (95% air/5%
CO2) for 10 min. At t = 9, propidium iodide
(10 µM, final concentration) was added, and cells were analyzed by
flow cytometry at 60 µl/min. Nonfluorescent Dih123 is cleaved by ROS
to fluorescent R123 and detected by the FL1 detector as described above
for (Van de Water 1995).
Atomic Absorption Spectroscopy
For determination of cisplatin content, 500-µl samples were taken from the incubations, centrifuged and washed twice with buffer D. The pellet was dried at 65°C for 30 to 60 min, resuspended in 100 µl phosphate-buffered saline and dissolved by addition of 200 µl of 2 N NaOH. After 2 h at 55°C, the samples were neutralized by addition of 100 µl of 4 M HCl and kept at 4°C until use. Cisplatin content was measured by a Perkin Elmer spectrophotometer (model 4000) at 265.9 nm, after electrothermic atomization (2,600°C) in a graphite furnace. K2PtCl6 was used as standard for calibration.
Determination of Enzymatic Activity
The frozen cells (80°C) were thawed on ice and mixed with
Triton X-100 to a final triton concentration of 0.3% (w/v).
Lactate dehydrogenase was determined spectrophotometrically by
following the decrease of NADH at 340 nm according to Bergmeyer et al. (1965).
Glucose-6-phosphatase activity was determined by the release of
phosphate from 0.03 M glucose 6-phosphate in 0.1 M citrate buffer, pH
6.5, in a volume of 0.5 ml. The reaction was started by addition of the
lysed cell homogenate. After precipitation of the protein with 2 ml
ice-cold 5% (w/v) TCA, phosphate was determined in the supernatant
with the molybdate assay (Harper, 1965).
Acid phosphatase was determined by release of phosphate from 50 mM
-glycerophosphate at 37°C in a 0.1 M citrate buffer, pH 4.6, in a
final volume 0.6 ml. The reaction was started by addition of the lysed
cell homogenate. After precipitation of the protein with 2 ml ice-cold
5% (w/v) TCA, phosphate was determined in the supernatant by the
molybdate assay (Harper, 1965)
Catalase was determined spectrophotometrically by following the
decomposition of H2O2 in 50 mM potassium
phosphate buffer, pH 7.0, at 240 nm as described by Aebi (1974).
GSH-Rd activity was determined spectrophotometrically as described in
Bilzer et al. (1984). To 600 µl of 100 mM potassium phosphate buffer, pH 7.0, containing 200 mM KCl and 1 mM EDTA, 10 µl
of 6 mM NADPH and 50-µl sample were added. After stabilization of the
signal, 10 µl of 60 mM GSSG (in H2O) were added and the decrease in absorbency at 340 nm was followed at 25°C.
GSH-Px activity was determined by a modification of the assay described
in Lawrence and Burk (1976). To 600 µl of 100 mM potassium phosphate
buffer, pH 7.0, containing 200 mM KCl, 10 µl 6 mM NADPH, 60 µl 20 mM GSH, 50 µl GSH-Rd (6 U/ml) and 50 µl sample were added. After
stabilization of the signal, 50 µl of 4 mM cumene hydroperoxide was
added and the decrease in absorbency at 340 nm was followed at 25°C.
Determination of Enzymatic Activity of Complexes I to IV of the Respiratory Chain
Enzymatic activities of the complexes I to IV were determined by dual wavelength spectrophotometry with an Aminco Dual Wavelength 2 ATM UV-VIS spectrophotometer (Silver Spring, MD). All concentrations below are final concentrations.
Complex I (NADH:ubiquinone oxidoreductase) activity was determined at
340 nm with 380 nm as reference wavelength, with a slit width of 3.0 nm
according to Estornell et al. (1993). The assay was
performed with 10 to 30 µg protein in a final volume of 1 ml of
buffer, pH 7.4, containing 10 mM Tris-HCl, 50 mM KCl, 1 mM EDTA and 2 mM KCN. After addition of 75 µl of 1 mM NADH and stabilization of the
signal, the reaction was started by addition of 100 µl of 1 mM
ubiquinone-10. The activity was calculated from the rate of decrease of
NADH (e = 5.5 mM
1 cm1) per µg
protein.
Complex II (succinate dehydrogenase) activity was determined by the
difference in absorbency between 270 and 330 nm according to Estornell
et al. (1993). The assay was performed with 10 to 30 µg
protein in a final volume of 1 ml of 50 mM potassium phosphate buffer,
pH 7.4, containing 100 µM EDTA, 1 mM KCN and 0.1% (w/v) BSA. After
addition of 80 µl of 1 mM ubiquinone-0 and stabilization of the
signal, the reaction was started by addition of 100 µl of 0.1 M
sodium succinate. The activity was calculated from the rate of decrease
in ubiquinone (e = 9.6 mM1 cm1).
Complex III (Ubiquinol-cytochrome c reductase) activity was determined
by the difference in absorbency between 550 and 580 nm according to
Birch-Machin et al. (1993b). The assay was performed with 10 to 30 µg protein in a final volume of 1 ml of 25 mM potassium phosphate buffer, pH 7.2, containing 5 mM MgCl2, 2 mM KCN,
2.5 mg/ml BSA, 2 µg/ml rotenone and 0.5 mM N-D-maltoside.
After addition of 10 µl of 3.5 mM ubiquinol and stabilization of the
signal, the reaction was started by the addition of 10 µl of 1.5 mM
cytochrome cIII. The activity was calculated from the rate of reduction
of cytochrome cIII (e = 19 mM1 cm1).
Complex IV (cytochrome c oxidase) activity was determined by the
difference in absorbency between 550 and 580 nm according to
Birch-Machin et al. (1993a). The assay was performed with 10 to 30 µg protein in a final volume of 1 ml of 25 mM potassium phosphate buffer, pH 7.0, containing 0.5 mM N-D-maltoside.
After addition of 10 µl of 1.5 mM cytochrome cII and stabilization of the signal, the reaction was started by the addition of 10 to 30 µg
cells. The activity was calculated from the rate of increase in
absorbency caused by oxidation of cytochrome cII to cytochrome cIII
(e = 19 mM1 cm1).
All activities were expressed per microgram of protein, which was
determined according to Lowry et al. (1951).
Determination of ATP Content
ATP content was determined in 1-ml samples taken from the
incubations. The samples were washed once with buffer D, resuspended in
400 µl of 10% (v/v) HClO4, frozen in liquid nitrogen and
stored at 80°C until use. After thawing and addition of 50 µl of
0.5 M potassium phosphate buffer, solutions were further neutralized by
addition of 90 µl of 10 N KOH and centrifuged at maximal speed in an
eppendorf centrifuge. ATP was assayed by the luciferin/luciferase method optimized according to Kimmich et al. (1975) in
buffer consisting of 21 mM glycylglycine, containing 6.6 mM
Na2HAsO4 and 4.2 mM MgCl2, adjusted
to pH 8.05 with NaOH. Two milliliters of buffer was mixed with 50 µl
of 10 mg/ml firefly extract in H2O. After addition of 50 µl sample, luminescence was measured. Values were compared with a
calibration curve with ATP as standard.
Determination of GSH
GSH content was determined according to Saville (1958). PPTC
were centrifuged at 80 × g for 3 min. The pellet was
extracted with 300 µl of 20% w/v TCA. After centrifugation at
14,000 × g for 10 min, GSH was determined in the
supernatant.
Statistical Analysis
All values are expressed as mean ± S.E.M. The statistical evaluation was performed with analysis of variance. Results were considered significant if P < .05.
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Results |
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Top
Abstract
Introduction
Materials & Methods
Results
Discussion
References
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Effects of Cisplatin on Mitochondrial Activities
Respiratory activity.
Incubation of freshly isolated PPTC in
suspension with cisplatin in concentrations varying from 5 to 100 µM
resulted in a concentration-dependent decrease in and viability.
The decrease in preceded the loss of viability (fig.
2).
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Effects of cisplatin on intracellular ATP content.
The
is the driving force of ATP production. Therefore, the decrease in
could affect the ATP content. Indeed, exposure of PPTC to 50 µM cisplatin for 20 min resulted in a significant decrease from
9 ± 1 to 5.7 ± 0.8 nmol ATP/mg protein (mean ± S.E.M., n 
3, P 
.05). Antioxidants could only
slightly prevent the decrease in ATP content induced by 50 µM
cisplatin. PPTC treated with antioxidants and 50 µM cisplatin
contained 6.4 ± 0.6 nmol ATP/mg protein, whereas PPTC exposed to
cisplatin only contained 5.5 ± 0.5 nmol ATP/mg protein.
Unfortunately, pretreatment of PPTC with DPPD or deferoxamine decreased
the ATP levels. However, all PPTC exposed to 100 and 500 µM cisplatin
for 20 min contained 5.6 ± 0.8 and 5.2 ± 0.8 nmol ATP/mg
protein, irrespective of if cells were treated with 20 µM DPPD or 1 mM DES before and during exposure to cisplatin (fig. 3).
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Effect of cisplatin on nonmitochondrial enzymes. To evaluate whether cisplatin also affected enzymes located in other parts of the cell, several enzymatic activities were assessed.
Cytosolic lactate dehydrogenase was only slightly inhibited by cisplatin (7 ± 0.8%); glucose-6-phosphatase, located in the endoplasmatic reticulum, catalase (peroxisomes) and acid phosphatase (lysosomes) was not altered significantly (n 4, P .05).
Control values of the enzymatic activities (mean ± S.E.M.) were:
lactate dehydrogenase, 17 ± 2 mmol NADH/min/mg protein;
glucose-6-phosphatase, 8 ± 0.9 µmol Pi/min/mg protein; acid
phosphatase, 55 ± 8 µmol Pi/min/mg protein; and catalase,
0.7 ± 0.1 mmol/min/mg protein.
Formation of ROS
Time course of decrease in versus ROS
formation.
Incubation of the PPTC with cisplatin caused a time-
and concentration-dependent generation of ROS, as determined by the
fluorescent probe Dih123 with flow cytometry (fig. 4A).
The decrease in occurred before ROS formation (fig. 4B): 10 min
after addition of 100 and 500 µM cisplatin, a significant decrease of
to 84 and 65% of control was found; whereas, at this time
point, no significant amounts of ROS had been formed (fig. 4B). Only
after 20 min, ROS formation became evident at cisplatin concentrations greater than 100 µM; at cisplatin concentrations of 50 µM and lower, ROS formation was not detectable until after 30 to 40 min (fig.
4A).
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and viability (not shown).
Effect of cisplatin on GSH-Rd and GSH-Px.
One of the
mechanisms that may underlie cisplatin-induced oxidative damage is
inhibition of enzymes that protect against oxidative stress. Therefore,
we determined the effect of cisplatin on GSH-Rd and GSH-Px, responsible
for GSH regeneration from GSSG and hydrogen peroxide metabolism,
respectively. Incubation of PPTC with cisplatin resulted in inhibition
of both enzymes. Within 20 min of exposure to 100 µM cisplatin,
GSH-Rd decreased to 80% of control, whereas GSH-Px activity, which was
inhibited less, decreased to 90% of control. The inhibition of GSH-Rd
was concentration and time dependent (fig. 5).
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Effect of cisplatin on GSH content. GSH-Rd regenerates GSH from GSSG; therefore, inhibition of GSH-Rd may lead to decreased GSH levels. Indeed, within 20 min the GSH content of PPTC exposed to 500 µM cisplatin decreased from 4.8 ± 0.8 nmol GSH/mg cell protein to 1.2 ± 0.4 nmol GSH/mg cell protein.
Role of mitochondria in formation of ROS.
Mitochondria are
often the source of ROS (Dawson et al., 1993). Therefore, we
studied the effect of inhibitors of the complexes I to IV of the
respiratory chain on cisplatin-induced ROS formation. However, because
the probe we used for ROS, Dih123, is cleaved into R123, the resulting
fluorescent signal possibly not only depends on ROS formation but also
on the . To assess to what extent the observed effects on ROS
production were caused by a lowering of , we determined the
effect of the uncoupler CCCP on the detection of ROS induced by TBHP.
Incubation of the PPTC with 500 µM TBHP decreased the by 15%
and caused formation of ROS within 15 min. Incubation of the PPTC with
the uncoupler 60 µM CCCP alone caused a decrease in , but no
ROS. However, coincubation of the PPTC with 500 µM TBHP and 60 µM
CCCP showed that CCCP further decreased the of the PPTC exposed
to TBHP, but this decrease did not hamper the detection of ROS by
Dih123. The Dih123 fluorescent signal in PPTC exposed to THBP and CCCP was 172 ± 18% of control, whereas the Dih123 fluorescent signal was 151 ± 13 in PPTC exposed to TBHP alone, which indicated that an effect on does not interfere with determination of ROS
formation measured with Dih123.
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-tocopherol (Van
de Water et al., 1995). TTFA inhibits the Q cycle activity of complex II, while Oxa, which competes for succinate, inhibits complex II at another site, which is involved in its antioxidant function. To evaluate at which site cisplatin inhibited complex II, we
studied the effect of TTFA and Oxa on cisplatin-induced ROS.
Coincubation of the cells with cisplatin and Oxa increased cisplatin-induced ROS formation from 371 ± 30 to 664 ± 40 (fig. 6B). This potentiation of cisplatin-induced ROS was dependent on
respiratory activity, since further addition of TTFA, antimycin, rotenone or KCN, reduced the ROS levels from 664% to almost the control level (fig. 6B).
Contribution of cisplatin-induced effects to ROS content.
The
above results showed that cisplatin induced an impaired antioxidant
status of the PPTC: the GSH content was reduced and regeneration of GSH
was inhibited. In addition, the respiratory activity was decreased. We
studied whether decreased GSH-Rd activity alone or in combination with
decreased respiratory activity would be sufficient for formation of
ROS. PPTC were incubated with BCNU (inhibitor of GSH-Rd) in combination
with the inhibitors of complexes I to IV; BCNU alone caused little ROS
(fig. 7A).
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and viability; after 3 h of exposure decreased to
40 ± 7% of control and viability to 45 ± 9% (fig. 7, B
and C).
Coincubation of the PPTC with BCNU and TTFA (II), Oxa (II) or anti
(III) did not result in formation of ROS (fig. 7A). Moreover, the
effects on and viability were less pronounced, than those of
BCNU and cisplatin. After 3 h of exposure and viability were 60 ± 7% of control, compared with 45 ± 9% obtained
with BCNU/cisplatin (fig. 7, B and C).
KCN is an inhibitor of complex IV activity (Lehninger, 1972). However,
it is also known to be a potent displacer of bound cisplatin (Naser
et al., 1988). To check whether the observed effects by KCN
were caused by inhibition of complex IV or by displacement of
cisplatin, we determined cisplatin content of the PPTC in presence and
absence of 1 mM KCN.
PPTC exposed to 100 µM cisplatin for 2 h contained 4 nmol
Pt/µg protein, which was reduced by simultaneous addition of 1 mM KCN
to 0.5 nmol Pt/µg. Likewise, the Pt content of PPTC exposed to 500 µM was reduced from 9 to 2 nmol Pt/µg protein. Values are mean ± S.E.M., n 3, and were significantly different
from cells incubated with cisplatin alone (P < .05). This
indicated that prevention of ROS formation by KCN, as shown in fig. 6A,
may be merely caused by displacement. Therefore, the exact role of
complex IV activity in cisplatin-induced ROS formation can not be
studied with KCN.
Role of ROS formation in cisplatin-induced cell death. To evaluate the role of ROS formation in cisplatin-induced cell death, we studied the effects the antioxidant DPPD and the iron chelator deferoxamine.
The ROS formation induced by cisplatin could be completely prevented by DPPD and deferoxamine. However, these compounds could not prevent the decrease in, nor prevent cell death (fig. 8, A-C). Identical results were obtained with rotenone and BCNU; i.e., although formation of ROS was prevented, DPPD and
deferoxamine could not prevent the effects on and viability. As
a control, PPTC were incubated with 500 µM TBHP, which is known to
cause cell death by peroxidation. Prevention of the TBHP-induced ROS with DPPD and deferoxamine, significantly reduced the loss of
and completely prevented the PPTC from cell death (fig.
9, A-C). This indicated that if ROS formation is the
main cause of cell death, its prevention protects PPTC from cell death.
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Discussion |
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Top
Abstract
Introduction
Materials & Methods
Results
Discussion
References
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In a previous paper, freshly isolated PPTC were validated
as an in vitro model to detect nephrotoxicity by studying
the effect of mercuric chloride, cisplatin, para-aminophenol
and halogenated alkenes on viability and mitochondrial membrane
potential. The cells responded, time- and dose-dependently, to the
nephrotoxic compounds with a decrease in and loss of viability.
The sensitivity of the porcine cells to detect toxic effects
corresponded favorably with in vitro systems derived from
other animals (Kruidering et al., 1994).
This study provides evidence that cisplatin-induced mitochondrial
dysfunction is caused by inhibition of complexes I to IV of the
respiratory chain, which results in decreased intracellular levels of
ATP. This selectivity for mitochondria is probably caused by
accumulation of cisplatin in the negatively charged inner space of the
mitochondria because of the positive charge of aquated complexes of
cisplatin. Indeed cisplatin has been reported to accumulate in
mitochondria of kidney cells in vitro (Gemba and Fukuishi,
1991; Kameyama and Gemba, 1991b) as well as in mitochondria of kidney
and liver cells in vivo (Singh, 1989; Gemba et
al., 1987; Rosen et al., 1992).
Role of mitochondria in the formation of ROS.
Because
oxidative damage has been suggested as the main cause of
cisplatin-induced renal cell death and several antioxidants and radical
scavengers alleviate cisplatin-induced nephrotoxicity in
vitro (Zhang et al., 1992; Sadzuka et al.,
1992b; Kameyama and Gemba, 1991a) as well as in vivo (Gemba
et al., 1988), we studied the role of ROS formation.
. Because the inhibitors of the respiratory
chain complexes decrease the , the lowered ROS production may be
the consequence of the effect on only. Therefore, we determined
to what extent the Dih123 signal was dependent on the decrease in
. Exposure of the PPTC to TBHP and the uncoupler CCCP clearly
showed that a decrease in does not directly result in a
decreased Dih123 signal and demonstrated that ROS can be detected in
spite of a decrease in .
The detectable amount of ROS is determined by the imbalance between
formation and scavenging of ROS. Cisplatin-induced inhibition of
complexes I to III will be expected to reduce the amount of ROS by
reducing the electron flow. However, exposure of PPTC to cisplatin
resulted in an increased ROS content in spite of the inhibition of
complexes I to IV, which suggests that the scavenging of ROS is
probably even more impaired by cisplatin. The mechanism of decreased
oxidant protection in cisplatin-induced ROS formation has been studied
extensively. Cisplatin affects many enzymes that protect the cells from
oxidative damage, among which Cu,Zn-superoxide dismutase, Mn-superoxide
dismutase and catalase (Sadzuka et al., 1992a,b, 1994). In
addition, cisplatin decreases intracellular GSH levels (Zhang and
Lindup, 1993; Somani et al., 1995), which we also observed.
This study shows that, also in PPTC, cisplatin-induced ROS formation
could be partially aggravated by the impaired antioxidant status of the
cell, because incubation of the PPTC with cisplatin resulted in a time-
and concentration-dependent inhibition of the enzymes GSH-Rd and
GSH-Px, in addition to the severe reduction of intracellular GSH
levels. GSH-Px is responsible for the detoxification of hydrogen
peroxide, resulting in the formation of GSSG. GSH-Rd subsequently
regenerates GSH from GSSG.
A decreased GSH content, or inhibition of GSH-Rd alone, is not
sufficient to induce ROS formation, as is clear from our observations that maximal depletion of GSH in the cytosol or in the mitochondria of
PPTC did not result in ROS formation, nor did incubation of the PPTC
with BCNU, which specifically inhibits GSH-Rd (Babson and Reed, 1978).
However, coincubation of the PPTC with inhibitors of complexes I to IV
clearly showed that only inhibition of complex I in combination with
BCNU resulted in ROS production. Because we found that cisplatin
inhibits both GSH-Rd and complex I enzymatic activity, our data suggest
that this may be the mechanism by which cisplatin-induced ROS formation
occurs. This is confirmed by the observations that incubation of the
PPTC with the combinations BCNU/cisplatin or BCNU/rotenone had
identical toxicity (fig. 6, B-C), whereas BCNU in combination with
TTFA, Oxa, anti or KCN had less pronounced effects on and
viability. Inhibition of complex I with rotenone or MPTP
(1-methyl-u-phenyl-1,2,3,6-tetrahydropyeidine) has been implicated in
formation of ROS before (Cortopassi and Wang, 1995). Inhibition of the
electron flow through complex I favors electron flow through complex
II. Electrons channeled via complex II produce about four
times more superoxide than those channeled via complex I
(Pryor, 1982); therefore, it is conceivable that inhibition of complex
I results in a higher production of superoxide anions, which in
combination with depletion of GSH will lead to oxidative damage.
Apart from GSH regeneration, other antioxidant functions exist in PTC.
Complex II of the respiratory chain is involved in regeneration of
-tocopherol. Also in PPTC exposed to cisplatin complex II exerts a
protective function, because cisplatin-induced ROS formation could be
potentiated by the complex II inhibitor Oxa. However, Oxa is also a
substrate for the citric acid cycle. Therefore, increased NADH
production and subsequent increase in electron flow through the
respiratory chain may also cause increased ROS. Our results do not
exclude this possibility.
In summary, these results show that only inhibition of complex I in
combination with reduced GSH-Rd activity resulted in ROS formation.
Cisplatin exerts both effects; therefore, cisplatin-induced ROS
formation can be mediated via simultaneous inhibition of
complex I and GSH-Rd.
Relevance of ROS for cisplatin-induced cell death.
In spite of
the observations that several agents such as deferoxamine (Kameyama and
Gemba, 1991a), DPPD (Sugihara et al., 1987), -tocopherol
(Hannemann and Baumann, 1988), procaine (Zhang et al., 1992)
and dithiothreitol (Zhang et al., 1994) prevented formation
of peroxides and cell damage both in vitro and in
vivo (Zhang and Lindup, 1994; Sugihara et al., 1987;
Gemba et al., 1988) we postulate that peroxide damage is not
the cause of cell death.
) and in vitro in mouse PTC
(Lieberthal et al., 1996).
Possible mechanism of cisplatin-induced cell death and clinical
relevance.
The observation that the decrease in preceded
cell death induced by cisplatin (fig. 2) suggests an important role for
mitochondria as primary targets of cisplatin, which is in good
agreement with other reports in the mouse and rat in vivo
(Singh, 1989; Gordon and Gattone, 1986) as well as in vitro
(Zhang and Lindup, 1994; McGuiness and Ryan, 1994). We showed that
cisplatin-induced mitochondrial dysfunction was the result of
inhibition of the enzymatic complexes of the respiratory chain by
cisplatin, with decreased ATP levels as a consequence. Antioxidants
could not fully prevent the inhibition of the mitochondrial enzymes,
the decreases in ATP content nor cell death. Therefore, inhibition of
respiration probably is an important factor in cisplatin-induced cell
death. This hypothesis is supported by the observation that exposure of
PPTC to a combination low concentrations of inhibitors of complexes I
to IV of the MRC caused reduction of and cell death similar to
cisplatin (fig. 10).
|
) present in the active site of many enzymes,
cisplatin may exert its effect via inhibition of these
enzymes. Studies of the site and type of interaction of cisplatin at
the different enzymatic complexes will give more clues about the
mechanism of inhibition and the role of this inhibition in the pathway
leading to acute cell death. Because cisplatin is known to affect DNA,
RNA and protein synthesis, one could argue that the observed inhibition
of enzymatic activity is mediated this way. However, toxic effects
mediated in this way probably need more time to cause overt toxicity;
10 to 100 µM cisplatin did not have an affect on protein synthesis
within 6 h in human kidney cortical slices. Only after 12 h
could cisplatin-induced effects on protein synthesis be observed
(Fisher et al., 1994), making it less likely, although not
excluding, that cisplatin-induced inhibition of respiration, already
observed within 20 min, is caused by effects on protein synthesis.
In patients treated with cisplatin, plasma levels reach approximately
10 µM after a single dose. The concentrations we used in this study
are higher, but not unrealistic. Cisplatin usually is given to patients
in repeated doses with intervals of 1 to 3 weeks (Gandara et
al., 1989). However, 21 days after the last dose 13 to 40% of the
cisplatin is still present in the body (Fish et al., 1994),
which suggests that prolonged treatment may result in accumulation of
cisplatin. Indeed accumulation of cisplatin has been observed in
kidneys of patients treated with cisplatin (Stewart et al.,
1985; Tothill et al., 1992), which implies that studies of
the mechanisms underlying cisplatin-induced toxicity should include
concentrations higher than 10 µM.
Although this study suggests that ROS formation is not the direct cause
of cisplatin-induced toxicity, ROS formation may have significance for
cisplatin-induced nephrotoxicity in vivo, because administration of antioxidants can alleviate the nephrotoxicity substantially because of reduction of inflammatory reactions of cell-mediated toxicity and by reduction of the amount of ROS that is
released by a damaged cell. Yet, our results demonstrate that the
actual cause of cisplatin-induced cellular toxicity is not prevented by
antioxidants; therefore, cisplatin will still be nephrotoxic in spite
of the use of antioxidants.
| |
Acknowledgments |
|---|
The enthusiastic support of Jaap van Hellemond (Utrecht University, Utrecht, The Netherlands) with the enzymatic assays of complex I to IV activity is greatly appreciated. We thank Prof. Vogel for the generous gift of BCNU, and Dr. A.M. Fichtinger, TNO, Delft, The Netherlands, for her help with the atomic absorption spectroscopy.
| |
Footnotes |
|---|
Accepted for publication October 21, 1996.
Received for publication July 25, 1996.
1 This project was financially supported by the Dutch Foundation Platform Alternatives to Animal Experiments.
2 W.A. Jones Cell Science Center, Old Barn Road, P.O. Box 49, Lake Placid, NY 12946.
3 Dept. of Pathology, Leiden University, P.O. Box 9603, 2300 RC Leiden, The Netherlands.
Send reprint requests to: Ms Marieke Kruidering, Division of Toxicology, Leiden/Amsterdam Center for Drug Research, Leiden University, P.O. Box 9503, 2300 RA Leiden, The Netherlands.
| |
Abbreviations |
|---|
anti, antimycin;
BCNU, N,N-bis(2-chloroethyl)-N-nitrosourea;
BSA, bovine serum albumin;
CCCP, cyanide m-chlorophenylhydrazone;
DES, DesferalR
(deferoxamine);
Dih123, dihydrorhodamine-123;
DPPD, diphenyl-p-phenylene-diammine;
, mitochondrial membrane
potential;
EDTA, ethylenediaminetetraacetate;
GSH, glutathione (reduced
form);
GSH-Px, GSH peroxidase;
GSH-Rd, GSH reductase;
GSSG, glutathione
(oxidized form);
HEPES, N-2-hydroxyethylpiperazine-N-2-ethanesulfonic
acid;
MRC, mitochondrial respiratory chain;
Oxa, oxaloacetic acid;
PPTC, porcine proximal tubular cells;
Pt, cis-diamine-dichloroplatinum
(II);
R123, rhodamine-123;
ROS, reactive oxygen species;
rot, rotenone;
TBHP, tertiary butyl hydroperoxide;
TCA, trichloroacetic acid;
TTFA, thenoyltrifluoroacetone.
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References |
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Top
Abstract
Introduction
Materials & Methods
Results
Discussion
References
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