点击显示 收起
【摘要】 Previously, we showed that physiological functions of renal proximal tubular cells (RPTC) do not recover following S -(1,2-dichlorovinyl)- L -cysteine (DCVC)-induced injury. This study investigated the role of protein kinase C- (PKC- ) in the lack of repair of mitochondrial function in DCVC-injured RPTC. After DCVC exposure, basal oxygen consumption (Q O 2 ), uncoupled Q O 2, oligomycin-sensitive Q O 2, F 1 F 0 -ATPase activity, and ATP production decreased, respectively, to 59, 27, 27, 57, and 68% of controls. None of these functions recovered. Mitochondrial transmembrane potential decreased 53% after DCVC injury but recovered on day 4. PKC- was activated 4.3- and 2.5-fold on days 2 and 4, respectively, of the recovery period. Inhibition of PKC- activation (10 nM Go6976) did not block DCVC-induced decreases in mitochondrial functions but promoted the recovery of uncoupled Q O 2, oligomycin-sensitive Q O 2, F 1 F 0 -ATPase activity, and ATP production. Protein levels of the catalytic -subunit of F 1 F 0 -ATPase were not changed by DCVC or during the recovery period. Amino acid sequence analysis revealed that -, -, and -subunits of F 1 F 0 -ATPase have PKC consensus motifs. Recombinant PKC- phosphorylated the -subunit and decreased F 1 F 0 -ATPase activity in vitro. Serine but not threonine phosphorylation of the -subunit was increased during late recovery following DCVC injury, and inhibition of PKC- activation decreased this phosphorylation. We conclude that during RPTC recovery following DCVC injury, 1 ) PKC- activation decreases F 0 F 1 -ATPase activity, oxidative phosphorylation, and ATP production; 2 ) PKC- phosphorylates the -subunit of F 1 F 0 -ATPase on serine residue; and 3 ) PKC- does not mediate depolarization of RPTC mitochondria. This is the first report showing that PKC- phosphorylates the catalytic subunit of F 1 F 0 -ATPase and that PKC- plays an important role in regulating repair of mitochondrial function.
【关键词】 renal proximal tubular cells mitochondria F F ATPase
S -(1,2- DICHLOROVINYL )- L - CYSTEINE (DCVC), a nephrotoxic metabolite of the environmental contaminants trichloroethylene and dichloroacetylene, produces cell injury and death in renal proximal tubular cells (RPTC) ( 10, 14 - 18, 20, 26, 28, 29, 35, 36, 39 ). DCVC exposure induces mitochondrial dysfunction, depletion of cellular ATP, modification of thiols, and decreases in active Na + transport, Na + -K + -ATPase activity, and Na + -dependent glucose uptake ( 12, 14, 16, 26, 28, 29 ). Mitochondrial dysfunction is a hallmark of DCVC-induced toxicity. DCVC exposure decreases mitochondrial respiration, total adenine nucleotide pool, and mitochondrial glutathione content, alters the concentration of several citric acid cycle intermediates, reduces the activity of isocitrate dehydrogenase and succinate:cytochrome c oxidoreductase, and inhibits succinate-linked state 3 respiration ( 1, 15 ). Furthermore, DCVC impairs the ability of mitochondria to retain Ca 2+ and to generate the mitochondrial membrane potential ( m ), which leads to extensive mitochondrial swelling, release of cytochrome c, and activation of caspase-3 ( 6, 7, 37 ).
The return of RPTC physiological functions is critical for the restoration of normal renal function. We showed that RPTC fully recover physiological functions following an oxidant injury but not following DCVC-induced injury, which suggests that DCVC inhibits regenerative responses in RPTC ( 29 ). For example, the mitochondrial function remains suppressed in DCVC-injured RPTC for 6 days following the treatment ( 29 ). Our previous studies reported that epidermal growth factor (EGF), pharmacological concentrations of ascorbic acid, collagen IV, and longer exposures to phorbol ester promote recovery of mitochondrial function in DCVC-injured RPTC ( 23, 24, 26, 28, 29 ). However, the mechanisms underlying the lack of mitochondrial repair and the pathways by which EGF, ascorbic acid, collagen, and phorbol ester promote the return of RPTC functions in DCVC-injured RPTC remain unknown.
Protein kinase C (PKC) is a family of serine/threonine protein kinases, which play a central role in cell signaling and the regulation of a variety of cellular functions, including growth, differentiation, motility, contraction, ion and macromolecule secretion, electrolyte transport, synaptic transmission, axonal regeneration, tumor promotion, cell aging, apoptosis, and survival ( 8 ). Each PKC isozyme is involved in the regulation of different functions and has a unique role in the cell. PKC-, one of the classic PKC isozymes, has been implicated in cellular proliferation and differentiation ( 22, 33 ) as well as cell injury and death ( 25 ). A previous study by this laboratory showed that PKC- mediates mitochondrial dysfunction, decreases in active Na + transport, and apoptosis in cisplatin-injured RPTC, suggesting an important role of PKC- in the regulation of mitochondrial function during cell injury and death ( 25 ). Specifically, cisplatin-induced activation of PKC- inhibits oxidative phosphorylation by decreasing electron transport rate and F 1 F 0 -ATPase activity, suggesting an important role of PKC- in the regulation of F 1 F 0 -ATPase activity ( 25 ).
F 1 F 0 -ATPase is localized in the inner mitochondrial membrane and is responsible for ATP synthesis under physiological conditions ( 2, 3 ). Under some pathological conditions leading to the loss of m, F 1 F 0 -ATPase operates in a reverse mode and hydrolyzes ATP. The enzyme has multiple subunits forming two major complexes, F 0 and F 1 ( 3, 4 ). F 1 is a water-soluble catalytic complex made up of five subunits ( 3 3 ), with the catalytic site located on the -subunit. F 0 is made up of several integral membrane proteins that form a proton channel ( 3, 4, 32, 34 ). The activity of this enzyme is tightly regulated to synchronize ATP synthesis with cellular energy expenditure. Protein phosphorylation, a posttranslational modification producing changes in enzymatic conformation and activity, may represent one of the mechanisms responsible for regulation of F 1 F 0 -ATPase activity. It has been shown that platelet-derived growth factor induces tyrosine phosphorylation of the -subunit of F 1 F 0 -ATPase ( 42 ). Indeed, our search of the amino acid sequence of F 1 F 0 -ATPase revealed several possible PKC phosphorylation sites on subunits,, and as well as on coupling factor 6.
PKC- has also been implicated in cellular repair and regenerative processes. Renal regeneration following folic acid- or DCVC-induced injury in vivo is associated with the downregulation of PKC- protein levels in kidney homogenates ( 9, 43 ). Our previous study also suggested that PKC- is one of the PKC isozymes involved in the repair of mitochondrial function and active Na + transport following DCVC injury in RPTC, but the mitochondrial targets of this PKC isozyme have not been examined ( 26 ). Therefore, the aim of the present study was to determine the role of PKC- in the inhibition of the repair of mitochondrial function and to elucidate the mitochondrial targets of PKC- in DCVC-injured RPTC.
MATERIALS AND METHODS
Materials. Female New Zealand White rabbits (2.0-2.5 kg) were purchased from Myrtle's Rabbitry (Thompson Station, TN). DCVC was synthesized according to the method of Moore and Green ( 31 ). 5,5',6,6'-Tetrachloro-1,1',3,3'-tetraethylbenzimidazolylcarbocyanine iodide (JC-1) and the antibodies against F 1 F 0 -ATPase subunits were obtained from Molecular Probes (Eugene, OR). The cell culture media (a 50:50 mixture of DMEM and Ham's F-12 nutrient mix without phenol red, pyruvate, and glucose) were purchased from MediaTech Cellgro (Herndon, VA). PKC- inhibitor, Go6976, recombinant PKC-, -, and - were supplied by Calbiochem (La Jolla, CA). Protease inhibitors and an ATP Bioluminescence Assay Kit HS II were obtained from Roche (Mannheim, Germany), and phosphatase inhibitors cocktail was obtained from Sigma (St. Louis, MO). Tris-glycine gels were supplied by BioWhittaker Molecular Applications (Rockland, ME). A Silver Stain Plus kit and nitrocellulose membranes were obtained from Bio-Rad (Hercules, CA). Phospho-PKC- and PKC- antibodies were purchased from Upstate Biotechnology (Lake Placid, NY) and BD Transduction Laboratory (San Diego, CA), respectively. Phosphoserine-PKC substrate antibody and phosphothreonine antibody were obtained from Cell Signaling Technologies (Beverly, MA) and Biomol (Plymouth Meeting, PA), respectively. Anti-mouse IgG coupled to horseradish peroxidase was supplied by Kirkegaard & Perry Laboratory (Gaithersburg, MD) and Supersignal Chemiluminescent Substrate by Pierce (Rockford, IL). Redi vue [ - 32 P]ATP (3,000 Ci/mmol) was purchased from Amersham Biosciences (Piscataway, NJ). Protein A/G agarose was obtained from Santa Cruz Biotechnology (Santa Cruz, CA), an Amicon Diaflo apparatus with a PM-10 filter (10,000-molecular mass cutoff point) was obtained from Millipore (Bedford, MA), and phosphocellulose disks were from Whatman (Clifton, NJ). The sources of the other reagents and cell culture hormones have been described previously ( 25, 30 ).
Isolation and culture of RPTC. Renal proximal tubules were isolated from rabbit kidneys by the iron-oxide perfusion method and cultured in 35-mm culture dishes in improved conditions as previously described ( 30 ). The culture medium was a 50:50 mixture of DMEM and Ham's F-12 nutrient mix without phenol red, pyruvate, and glucose, supplemented with 15 mM NaHCO 3, 15 mM HEPES, and 6 mM lactate (pH 7.4, 295 mosmol/kgH 2 O). 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 media immediately before daily media change (2 ml/dish).
DCVC treatment of RPTC monolayer. RPTC monolayers reached confluence within 6 days and were treated with DCVC (200 µM, 90 min) on day 7 of culture. After DCVC exposure, the monolayer was washed with fresh warm (37°C) medium and cultured for an additional 4 days. In experiments using the PKC- inhibitor, RPTC were pretreated for 1 h with 10 nM Go6976 and then exposed to DCVC. Go6976 was added daily starting with the media change immediately following DCVC exposure. RPTC samples were taken at various time points after DCVC exposure for measurements of mitochondrial functions, biochemical analyses, immunoprecipitation, and immunoblotting.
Oxygen consumption. RPTC monolayers were gently detached from the dishes using a rubber policeman and transferred to the oxygen consumption (Q O 2 ) measurement chamber. Q O 2 was measured using a Clark-type electrode as described previously ( 25, 29, 30 ). Oligomycin-sensitive Q O 2 was used as a marker of oxidative phosphorylation and was calculated as a difference between basal Q O 2 and oligomycin-insensitive Q O 2, which was measured in the presence of oligomycin (0.6 µg/ml). Uncoupled Q O 2 was used as a marker of the electron transfer rate and integrity of complexes of the respiratory chain and was measured in the presence of FCCP (2 µM).
Measurement of intracellular ATP content. Intracellular ATP content in RPTC lysates was measured by the luciferase method using an ATP Bioluminescence Assay Kit HS II according to the manufacturer's protocol.
ATP production rate. The assessment of state 3 respiration (the maximum rate of ATP synthesis) was carried out by a modified method of Borkan et al. ( 2 ). In brief, the culture media were aspirated and replaced with 1 ml of a buffer solution resembling an intracellular electrolyte milieu (120 mM KCl, 5 mM KH 2 PO 4, 10 mM HEPES, 1 mM MgSO 4, and 2 mM EGTA, adjusted to pH 7.4 with KOH), containing digitonin (0.1 mg/ml) and 5 mM glutamate + 5 mM malate as the substrates. The reaction was initiated by adding excess ADP (2 mM final concentration) and carried out for 5 min at 37°C. Initial experiments determined that ATP production in these conditions was linear for 10 min. The reaction was terminated by adding an aliquot of ice-cold perchloric acid (3% final concentration), and the suspension was snap-frozen in liquid nitrogen. After thawing, the suspension was spun down at 15,000 g x 1 min at 4°C. The supernatant was neutralized to pH 7.5 and centrifuged again at 15,000 g x 10 min at 4°C. The final supernatant was analyzed for ATP content using an ATP Bioluminescence Assay Kit HS II as described above. The initial pellet was assayed for protein content following solubilization in a buffer containing 100 mM Tris·HCl (pH 7.5), 150 mM NaCl, and 0.05% Triton X-100.
m. m Was assessed as described previously ( 25 ) using JC-1, a cationic dye that exhibits potential-dependent accumulation and formation of red fluorescent J-aggregates in mitochondria, which is indicated by a fluorescence emission shift from green (525 nm) to red (590 nm). At different time points of the recovery period, RPTC monolayers were loaded with 10 µM JC-1 for 30 min at 37°C. After loading, media were aspirated and monolayers were put on ice, washed with ice-cold PBS, scraped off culture dishes, washed, and resuspended in PBS. Fluorescence was determined by flow cytometry (FACSCalibur, BD Biosciences) using excitation by a 488-nm argon-ion laser. The JC-1 monomer (green) and the J-aggregates (red) were detected separately in FL1 (525 nm) and FL2 (590 nm) channels, respectively. m Is presented as JC-aggregates/JC-1 monomer ratio.
Isolation of RPTC mitochondria. Mitochondria were isolated from RPTC by the method of Lash and Sall ( 19 ). RPTC were homogenized in ice-cold isolation buffer [225 mM sucrose, 10 mM Tris·HCl, 10 mM potassium phosphate (pH 7.4), 5 mM MgCl 2, 20 mM KCl, 2 mM EGTA, protease and phosphatase inhibitors cocktail] using a Dounce homogenizer and centrifuged at 1,000 g for 5 min at 4°C. The supernatant was collected and centrifuged at 15,000 g for 10 min at 4°C. The pellet containing RPTC mitochondria was washed twice in the isolation buffer and spun down again 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, and 100 mM KCl and used for measurement of F 1 F 0 -ATPase activity. In some experiments, the final mitochondrial pellet was resuspended in 50 µl of modified radioimmune precipitation assay (RIPA) buffer (50 mM Tris·HCl, 150 mM NaCl, 1 mM EGTA, 1% Triton X-100, 1 mM Na 3 VO 4, 1 mM NaF, and the protease and phosphatase inhibitors cocktail; pH 7.4) and used for immunoblot analysis.
Measurement of F 1 F 0 -ATPase activity. F 1 F 0 -ATPase activity was determined in freshly isolated RPTC mitochondria by measuring the release of P i from ATP according to Law et al. ( 21 ) with modifications described previously ( 23 ).
Isolation of F 1 F 0 -ATPase. Freshly isolated rabbit renal cortical mitochondria were used for the isolation of F 1 F 0 -ATPase using the method described by Catterall and Pedersen ( 5 ). Briefly, mitochondria were resuspended and homogenized in a homogenization buffer (3.0 mM Tris·HCl and 5.0 mM EDTA; pH 7.5) at a protein concentration of 50 mg/ml. After homogenization, the sample was sonicated using a Vibra Cell sonicator (Sonics and Materials, Danbury, CT) at maximum speed for 2 min on ice and then centrifuged at 150,000 g for 45 min at 0°C. The pellet was resuspended in a washing buffer (3.0 mM Tris·HCl and 50.0 mM EDTA; pH 7.5) at a protein concentration of 20 mg/ml and centrifuged at 150,000 g for 45 min at 0°C. The pellet was then washed five times in the washing buffer and centrifuged at 150,000 g for 45 min. The final pellet was resuspended in a washing buffer containing 10% ethylene glycol and 4.0 mM ATP, incubated for 16 h at room temperature, and centrifuged at 150,000 g for 45 min at room temperature. The pellet was resuspended in a buffer containing 250 mM sucrose, 3.0 mM Tris·HCl, and 5.0 mM EDTA (pH 7.5) and sonicated for 30 min (three 10-min intervals) in a 22°C water bath followed by immediate centrifugation at 150,000 g for 90 min at room temperature. The clear supernatant was applied to a DEAE cellulose column (2.0 x 8.0 cm) preequilibrated with 20 mM potassium phosphate, 5.0 mM EDTA (pH 7.5), and the enzyme was eluted with a linear gradient of potassium phosphate (50-250 mM; pH 7.5) containing 5.0 mM EDTA at a flow rate of 2 ml/min. The fractions eluted by 150-200 mM potassium phosphate were pooled and concentrated using an Amicon Diaflo apparatus with a PM-10 filter (10,000 cutoff point), and the concentrated sample was snap-frozen in liquid nitrogen and stored at -70°C until used in assays.
Phosphorylation of F 1 F 0 -ATPase by PKC-. In vitro phosphorylation of isolated F 1 F 0 -ATPase by recombinant PKC- was performed using a modification of the assay described by Yasuda et al. ( 41 ). Briefly, experiments were carried out at 30°C in an incubation mixture (25 µl) containing 20 mM Tris, pH 7.5, 0.35 mM EDTA, 0.35 mM EGTA, 1 mM CaCl 2, 20 mM MgCl 2, 6 mM -mercaptoethanol, 10 µM PMA, 280 µg/ml L - -phosphatidyl- L -serine, 2.4 µg F 1 F 0 -ATPase, and 30 nM PKC-. Histone type III phosphorylation by PKC- was used as a positive control. Reaction was started by the addition of ATP (10 µM, final concentration) with trace amounts of [ - 32 P]ATP (60,000 cpm/µl). Under these assay conditions, phosphotransferase activity of PKC- was linear for at least 5 min. The reaction was stopped after 5 min by adding Laemmli sample buffer (60 mM Tris·HCl, pH 6.8, containing 2% SDS, 10% glycerol, 100 mM -mercaptoethanol, and 0.01% bromophenol blue) and boiling samples for 5 min ( 13 ). Proteins were resolved by SDS-PAGE, gels were dried, and 32 P incorporation was determined by autoradiography. In some experiments, the reaction was stopped by spotting the samples on P81 phosphocellulose disks, the disks were washed twice with 1% phosphoric acid and twice with deionized water, and 32 P incorporation was quantified by liquid scintillation spectrometry.
To determine the identities of phosphorylated proteins, immunoblot analysis of the isolated F 1 F 0 -ATPase (2 µg) was performed using antibodies against various subunits of F 1 F 0 -ATPase. To visualize proteins resolved on gels, some gels were fixed and stained with silver stain using the Silver Stain Kit and the protocol provided by the manufacturer (Bio-Rad).
To examine the effect of phosphorylation by PKC- on F 1 F 0 -ATPase catalytic activity, the phosphorylation assay was performed as described above using nonradioactive ATP and the enzymatic activity of F 1 F 0 -ATPase was determined in the reaction mixture by measuring the release of P i from ATP ( 25 ).
Immunoblotting. Immunoblot analysis was used to assess protein levels of PKC-, PKC-, PKC-µ, phospho-PKC-, phospho-PKC-, phospho-PKC-µ, PKC-phosphorylated serine, phosphothreonine, and F 1 F 0 -ATPase subunits in total RPTC homogenates, mitochondria, and F 1 F 0 -ATPase preparations. Samples were lysed and boiled for 5 min in Laemmli sample buffer, and proteins were separated by SDS-PAGE and transferred electrophoretically to a nitrocellulose membrane. Blots were blocked in Tris-buffered saline (TBS) 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. After being washed in TBS containing 0.05% Tween 20 (TBS-T), the membranes were incubated for 1 h in the presence of secondary IgGs coupled to horseradish peroxidase and washed again in TBS-T. The supersignal chemiluminescent system was used for protein detection. Quantification of the results obtained from immunoblotting and autoradiography was performed using scanning densitometry.
Immunoprecipitation. RPTC monolayers were washed twice with ice-cold PBS, scraped off the dishes, lysed in ice-cold modified RIPA buffer for 15 min at 4°C, and spun down at 14,000 rpm for 10 min at 4°C. The supernatants (500 µg protein) were precleared with a protein A/G agarose bead slurry for 1 h and incubated overnight (both at 4°C) with the monoclonal antibody specific to the -subunit of the F 1 F 0 -ATPase (5 µg). Nonimmune mouse IgG was used as a negative control. The protein A/G agarose bead slurry was added to capture the immunocomplexes, and the incubation was continued for an additional 1 h at 4°C. Protein A/G agarose-attached immunocomplexes were harvested by centrifugation at 14,000 rpm at 4°C for 10 min and washed four times with ice-cold RIPA buffer. The final pellet was resuspended in Laemmli buffer, boiled for 5 min to dissociate immunocomplexes from protein A/G agarose beads, spun down at 14,000 rpm for 2 min at 4°C, and the supernatant was used for immunoblotting with antibodies specific to 1 ) PKC-phosphorylated serine, 2 ) phosphothreonine, and 3 ) the -subunit of the F 1 F 0 -ATPase. Antibody against PKC-phosphorylated serine detects phosphorylated serine residues when they are within a PKC consensus motif.
DNA content was determined using a CyQUANT Cell Proliferation Assay Kit (Molecular Probes) and the manufacturer's protocol. RPTC samples were prepared for DNA assay as described previously ( 32 ). 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 ± SE and were analyzed for significance by ANOVA. Multiple means were compared using Fisher's protected least significance difference test with a level of significance of P < 0.05. RPTC isolated from an individual rabbit represented one experiment ( n = 1) consisting of data obtained from 2 to 10 culture plates.
RESULTS
DCVC-induced injury in RPTC. Exposure of confluent RPTC monolayers to 200 µM DCVC for 90 min resulted in 30 and 43% cell loss at 4 and 24 h following the treatment, as indicated by the monolayer DNA content (26.3 ± 7.9 and 21.6 ± 3.0 µg/plate at 4 and 24 h, respectively, following DCVC treatment vs. 37.7 ± 3.0 µg/plate in controls). Monolayer DNA content in DCVC-injured RPTC did not recover during the 4 days following injury (23.3 ± 3.8 µg/plate in DCVC-injured RPTC vs. 40.8 ± 3.3 µg/plate in controls). However, visual inspection under a microscope showed that monolayer confluence recovered by day 4 following DCVC treatment (data not shown). PKC- inhibitor Go6976 had no effect on DNA content in control or DCVC-injured RPTC (21 and 46% cell loss at 4 and 24 h, respectively).
Activation of PKC- during RPTC repair following DCVC injury. Figure 1 shows that the recovery of RPTC following DCVC exposure was associated with the activation of PKC-. The ratio of phosphorylated PKC- to total PKC- in RPTC homogenates increased 4- and 2.5-fold on days 2 and 4, respectively, of the recovery period ( Fig. 1 D ). Go6976 abolished the activation of PKC- in DCVC-treated RPTC at all time points studied ( Fig. 1 A ). DCVC-induced injury had no effect on phosphorylation of PKC- ( Fig. 1 C ) and PKC-µ (data not shown).
Fig. 1. Protein levels of phosphorylated PKC- ( A ), total PKC- ( B ), phosphorylated PKC- ( C ), and PKC- activation (the ratio of phosphorylated PKC- to total PKC- levels; D ) in renal proximal tubular cells (RPTC) during recovery following S -(1,2-dichlorovinyl)- L -cysteine (DCVC)-induced injury. The results (quantified by densitometry) are the average ± SE of 3 independent experiments (RPTC isolations). Blots are representative of 3 independent experiments.
Activation of mitochondrial PKC- during RPTC repair following DCVC-induced injury. Our previous report demonstrated that PKC- is present in RPTC mitochondria ( 25 ). The present data show that the lack of recovery of mitochondrial function following DCVC-induced injury is associated with increased levels of active PKC- in RPTC mitochondria ( Fig. 2 A ). The ratio of phosphorylated PKC- to total PKC- in the mitochondria isolated from DCVC-injured RPTC was not changed during the early recovery period (1-4 h) following the injury ( Fig. 2 ). However, on day 4 of the recovery period, the ratio of phosphorylated PKC- to total PKC- was threefold higher in mitochondria of DCVC-injured RPTC than in controls ( Fig. 2, B and C ). Treatment of RPTC with Go6976 had no effect on the ratio of phosphorylated PKC- to total PKC- at 4 h following DCVC exposure but it prevented the activation of PKC- on day 4 ( Fig. 2, B and C ). These results show that mitochondrial PKC- is activated during the late recovery period following DCVC injury and that PKC- inhibitor Go6976 prevents the activation of mitochondrial PKC-.
Fig. 2. A : PKC- activation in RPTC mitochondria following DCVC-induced injury. B : effect of Go6976 on PKC- activation in RPTC mitochondria following DCVC-induced injury. C : ratio of phosphorylated PKC- to total PKC- levels in RPTC mitochondria following DCVC-induced injury. The results (quantified by densitometry) are the average ± SE of 5 independent experiments (RPTC isolations). Blots are representative of 5 independent experiments.
Mitochondrial respiration. Basal and uncoupled Q O 2 s were used as markers of mitochondrial respiration. Specifically, basal Q O 2 reflects the overall function of the mitochondria. The basal Q O 2 was decreased by 50% at 4 h following DCVC treatment and did not recover over time ( Fig. 3 A ). The inhibition of PKC- activation by Go6976 did not affect DCVC-induced decreases in basal Q O 2 but promoted the recovery of basal Q O 2 in DCVC-injured RPTC on day 4 ( Fig. 3 A ).
Fig. 3. Oxygen consumption (Q O 2 ) in RPTC during recovery following DCVC-induced injury. Results are the average ± SE of 3 independent experiments (RPTC isolations). Values with dissimilar superscripts ( a-c ) on a given day are significantly different ( P < 0.05) from each other.
Uncoupled Q O 2 was used as a marker of the mitochondrial electron transfer rate and the integrity of complexes of the respiratory chain. Uncoupled Q O 2 declined by 60% in DCVC-treated RPTC at 4 h following the exposure and remained decreased on day 4 of the recovery period ( Fig. 3 B ). The inhibition of PKC- activation by Go6976 promoted the recovery of uncoupled Q O 2 in DCVC-injured RPTC ( Fig. 3 B ).
These results demonstrate that basal respiration and the electron transfer rate in mitochondria do not recover in DCVC-injured RPTC and that inhibition of PKC- activation promotes the return of these functions.
m. Mitochondrial respiration results in the generation of proton and pH gradients across the inner mitochondrial membrane and produces the transmembrane potential ( m ), which represents most of the energy of the proton gradient. m In RPTC was assessed by the measurement of changes in the amount of JC-1 aggregates (red fluorescence) and J-aggregate/JC-1 monomer ratio. The amount of JC-1 aggregates decreased in DCVC-injured RPTC at 24 h following the treatment ( Fig. 4 A ). The J-aggregate/JC-1 monomer ratio in control RPTC was 1.29 ± 0.20, decreased by 52.8% in DCVC-injured RPTC at 24 h following the treatment, and returned to control levels by day 4 ( Fig. 4 B ). Go6976 had no effects on m in control or DCVC-injured RPTC ( Fig. 4 B ). These results show that DCVC-induced decreases in m recover over time and that the decrease and recovery of m following DCVC injury are not mediated by PKC-.
Fig. 4. Mitochondrial membrane potential ( m ) in RPTC during recovery following DCVC-induced injury. A : aggregate form of JC-1 in control and DCVC-injured RPTC on day 1 following exposure. Histogram is representative of 3 independent experiments. B : ratio of aggregate/monomeric form of JC-1. The results are expressed as a percentage of controls and are the average ± SE of 3 independent experiments (RPTC isolations). Values with dissimilar superscripts ( a and b ) on a given day are significantly different ( P < 0.05) from each other.
ATP production. The rate of ATP synthesis was measured during state 3 (the maximal capacity of mitochondria to generate ATP) in digitonin-permeabilized RPTC incubated in the presence of ADP and metabolic substrates linked to the respiratory complex I (glutamate + malate). The rate of ATP production in control RPTC was 67.4 ± 10.6 nmol/mg protein. Treatment with Go6976 for 4 h and 4 days had no effect on the ATP production rate ( Fig. 5 A ). ATP production was decreased 32% at 4 h following DCVC exposure and did not recover on day 4 ( Fig. 5 A ). However, treatment with Go6976 during the recovery period promoted the return of the ATP synthesis rate ( Fig. 5 A ). These results demonstrate that PKC- activation following DCVC-induced injury in RPTC mediates decreases in ATP production.
Fig. 5. ATP production rate ( A ) and intracellular content of ATP ( B ) in RPTC during recovery following DCVC-induced injury. Results are the average ± SE of 3 independent experiments (RPTC isolations). Values with dissimilar superscripts ( a and b ) on a given day are significantly different ( P < 0.05) from each other.
Intracellular ATP. As shown in Fig. 5 B, intracellular ATP content decreased by 32 and 40%, respectively, at 4 and 24 h and did not recover on day 4 following DCVC treatment. Treatment with Go6976 during the recovery period did not promote the return of intracellular ATP in DCVC-injured RPTC ( Fig. 5 B ). These results show that despite the return of the ATP synthesis rate, inhibition of PKC- activation following DCVC-induced injury does not result in the recovery of intracellular ATP content.
Oxidative phosphorylation. Oxidative phosphorylation and ATP synthesis are the fundamental functions of mitochondria. Oligomycin-sensitive Q O 2 was used as a marker of oxidative phosphorylation in RPTC. Oligomycin-sensitive Q O 2 declined 69% at 4 h following DCVC treatment and did not return throughout the recovery period ( Fig. 6 ). Inhibition of PKC- activation had no effect on the decreases in oligomycin-sensitive Q O 2 at 4 h but promoted, in part, the return of oligomycin-sensitive Q O 2 on day 4 following DCVC-induced injury ( Fig. 6 ). These results show that oxidative phosphorylation does not recover in DCVC-injured RPTC and that PKC- activation mediates the inhibition of the recovery of oxidative phosphorylation in this model.
Fig. 6. Oligomycin-sensitive Q O 2 in RPTC during recovery following DCVC-induced injury. Oligomycin-sensitive Q O 2 was measured in the presence of oligomycin (6 µg/ml) and was calculated as the difference between basal and oligomycin-insensitive Q O 2. Results are the average ± SE of 3 independent experiments (RPTC isolations). Values with dissimilar superscripts ( a and b ) on a given day are significantly different ( P < 0.05) from each other.
F 1 F 0 -ATPase activity. F 1 F 0 -ATPase catalyzes ATP synthesis using the energy of the proton gradient across the inner mitochondrial membrane. Exposure of RPTC to DCVC decreased activity of F 1 F 0 -ATPase by 41% at 4 h following the treatment ( Fig. 7 A ). F 1 F 0 -ATPase activity remained decreased over the 4-day recovery period ( Fig. 7 A ). Inhibition of PKC- activation by Go6976 had no effect on the F 1 F 0 -ATPase activity at 4 h following DCVC exposure but promoted full recovery of the F 1 F 0 -ATPase activity on day 4 ( Fig. 7 A ). These data demonstrate that the lack of recovery of F 1 F 0 -ATPase activity in DCVC-injured RPTC is mediated by PKC-.
Fig. 7. F 1 F 0 -ATPase activity and protein levels during recovery following DCVC-induced injury. A : F 1 F 0 -ATPase activity in RPTC mitochondria at 4 h and on day 4 of the recovery period following DCVC exposure. Results are the average ± SE of 3 independent experiments (RPTC isolations). Values with dissimilar superscripts ( a and b ) on a given day are significantly different ( P < 0.05) from each other. B : protein levels of the -subunit of F 1 F 0 -ATPase in RPTC homogenates during the recovery following DCVC exposure. C : protein levels of the -subunit of F 1 F 0 -ATPase in RPTC mitochondria during the recovery following DCVC exposure. Blots are representative of 3 independent experiments.
F 1 F 0 -ATPase protein levels. Decreases in F 1 F 0 -ATPase activity can be due to reduced enzymatic activity or decreased protein levels of the F 1 F 0 -ATPase. F 1 F 0 -ATPase is composed of multiple subunits and several regulatory peptides associated with these subunits. Because the -subunit is the catalytic subunit of F 1 F 0 -ATPase, we determined protein levels of the -subunit in cell homogenates and mitochondria isolated from DCVC-injured RPTC during the recovery period. As demonstrated in Fig. 7, B and C, protein levels of the -subunit of F 1 F 0 -ATPase in cell homogenates and mitochondria were not changed after DCVC-induced injury or throughout the recovery period. The inhibition of PKC- by Go6976 had no effect on protein levels of the -subunit of F 1 F 0 -ATPase in mitochondria isolated from control and DCVC-injured RPTC ( Fig. 7 C ). These results demonstrate that DCVC does not decrease protein levels of the catalytic subunit of F 1 F 0 -ATPase and suggest that the decreases in F 1 F 0 -ATPase activity in DCVC-injured RPTC are due to the reduced catalytic activity of F 1 F 0 -ATPase and not the amount of the enzyme.
Phosphorylation of F 1 F 0 -ATPase by PKC- in vitro. PKC, a serine/threonine kinase, phosphorylates a wide range of intracellular substrates. Several F 1 F 0 -ATPase subunits, including the -subunit ( Fig. 8 ), have PKC consensus amino acid motifs. An in vitro assay was performed to determine whether F 1 F 0 -ATPase subunits are phosphorylated by major PKC isozymes present in RPTC. Phosphorylation of F 1 F 0 -ATPase by recombinant PKC isozymes was determined using an in vitro kinase assay followed by liquid scintillation spectrometry. Among the three isozymes tested in this study, PKC-, PKC-, and PKC-, only PKC- phosphorylated F 1 F 0 -ATPase in vitro ( Fig. 9 A ).
Fig. 8. PKC consensus motifs on the -subunit of F 1 F 0 -ATPase. The sequence of the -subunit of F 1 F 0 -ATPase from Chaetosphaeridium globosum was obtained from National Center for Biotechnology Information (accession number Q8SLY2 ) and PKC consensus motifs were searched using ScanProsite.
Fig. 9. In vitro phosphorylation of F 1 F 0 -ATPase by PKC isozymes. A : phosphorylation of F 1 F 0 -ATPase by PKC-, -, and -. B : silver staining of subunits of the isolated F 1 F 0 -ATPase resolved by SDS-PAGE. C : representative autoradiogram of in vitro phosphorylation of F 1 F 0 -ATPase by PKC-. Lane 1 : recombinant PKC-. Lane 2 : isolated F 1 F 0 -ATPase. Lanes 3 and 4 : PKC- + F 1 F 0 -ATPase. Lane 5 : PKC- + histone type III (positive control). D : representative immunoblot of the isolated F 1 F 0 -ATPase probed with antibody against the -subunit. Immunoblot analysis was performed to determine the identities of phosphorylated proteins. Purified F 1 F 0 -ATPase (2 µg) was loaded onto a 15% Tris-glycine gel and subjected to electrophoresis simultaneously with proteins shown in C. E : F 1 F 0 -ATPase activity following in vitro phosphorylation by PKC-. Results are the average ± SE of 3 independent assays. Values with dissimilar superscripts ( a and b ) are significantly different ( P < 0.05) from each other.
To determine which subunits of F 0 F 1 -ATPase are phosphorylated by PKC-, the reaction mixture was subjected to SDS-PAGE and autoradiography following the phosphorylation assay in vitro. The F 1 F 0 -ATPase isolated for this study consisted of three major (molecular mass 58, 56, and 26 kDa) and two minor (molecular mass 46 and 40 kDa) proteins as revealed by SDS-PAGE and silver staining ( Fig. 9 B ). Autoradiography demonstrated that PKC- phosphorylated two of these proteins (molecular mass 56 and 26 kDa; Fig. 9 C ). To determine whether the 56-kDa protein represented the -subunit of F 1 F 0 -ATPase (molecular mass 56.6 kDa), the F 1 F 0 -ATPase preparation was subjected to SDS-PAGE simultaneously with the proteins following the phosphorylation assay and subjected to immunoblotting. As shown in Fig. 9 D, the antibody to the -subunit of F 1 F 0 -ATPase detected a 56-kDa protein corresponding to the radioactive 56-kDa band. The identity of the phosphorylated 26-kDa band is not known at present. Thus these data demonstrate that the -subunit of F 1 F 0 -ATPase is a PKC- but not PKC- or PKC- substrate.
An in vitro assay was carried out to determine whether phosphorylation of F 1 F 0 -ATPase by PKC- alters F 1 F 0 -ATPase activity. Figure 9 E shows that the activity of F 1 F 0 -ATPase decreased 53% in the presence of active PKC- and ATP. These results suggest that phosphorylation by PKC- has an inhibitory effect on F 1 F 0 -ATPase.
Phosphorylation of F 1 F 0 -ATPase by PKC- in DCVC-injured RPTC. Immunoprecipitation was used to determine 1 ) whether phosphorylation of the -subunit of F 1 F 0 -ATPase occurs in DCVC-injured RPTC and 2 ) whether the -subunit of F 1 F 0 -ATPase is phosphorylated by PKC- following DCVC exposure. The -subunit of F 1 F 0 -ATPase was immunoprecipitated from RPTC lysates and the serine and threonine phosphorylation of the -subunit was examined at various time points following DCVC injury. Phosphothreonine levels in the -subunit were not affected by DCVC and Go6976 treatments throughout the recovery period ( Fig. 10 B, data not shown). Phosphoserine PKC substrate antibody used for immunoblotting detects only phosphorylated serine residues that are within a PKC consensus motif. The phosphoserine levels on the -subunit of F 1 F 0 -ATPase were not altered at 1, 2, and 4 h but decreased at 8 h of the recovery period in DCVC-injured RPTC ( Fig. 10 A, data not shown). However, on day 4 of the recovery period, phosphoserine levels on the -subunit of F 1 F 0 -ATPase increased in DCVC-injured RPTC ( Fig. 10 A ). PKC- inhibitor Go6976 decreased the serine phosphorylation of the -subunit of F 1 F 0 -ATPase on day 4 of the recovery period ( Fig. 10 A ). Thus these results show that PKC- activation in RPTC during late recovery following DCVC exposure increases phosphorylation of serine(s) but not threonine(s) on the -subunit of F 1 F 0 -ATPase.
Fig. 10. Phosphorylation status of the -subunit of F 1 F 0 -ATPase in RPTC during recovery after DCVC-induced injury. A : serine phosphorylation of the -subunit of F 1 F 0 -ATPase immunoprecipitated from RPTC following DCVC-induced injury. Phosphoserine PKC substrate antibody detects phosphorylated serine residues only when they are within PKC consensus motif. B : threonine phosphorylation of the -subunit of F 1 F 0 -ATPase immunoprecipitated from RPTC following DCVC-induced injury. C : protein levels of the -subunit of F 1 F 0 -ATPase immunoprecipitated from RPTC following DCVC-induced injury. Data are representative of 3 independent immunoprecipitations (RPTC isolations).
DISCUSSION
After nephrotoxicant exposure, the sublethally injured RPTC must undergo repair process to restore the physiological functions of the kidney. The repair process is composed of several stages including loss of the lethally injured cells, dedifferentiation of the sublethally injured cells, proliferation and morphological recovery of the damaged area, and differentiation to restore the physiological functions of renal proximal tubules ( 39 ). Despite the fact that numerous studies have been performed in the past to seek molecules that may accelerate regeneration following acute renal failure, the mechanisms regulating the recovery of renal functions are largely unknown. The aim of the present study was to determine whether PKC- is involved in these mechanisms.
Previously, we showed that DCVC produces significant RPTC death and loss. The remaining sublethally injured RPTC survive the insult but neither proliferate nor regain differentiated functions ( 29 ). Our present and previous studies show that DCVC exposure produces sustained mitochondrial dysfunction in sublethally injured RPTC and that mitochondrial function does not recover within 6 days following DCVC exposure ( 23, 26, 28, 29 ). In contrast, RPTC injury induced by an oxidant is followed by full recovery of mitochondrial and transport functions ( 27 ).
The decrease in cellular functions of the surviving RPTC was accompanied by sustained activation of PKC- on days 2 and 4 of the recovery period following DCVC treatment. The finding that the PKC- inhibitor Go6976 (10 nM) blocked DCVC-induced activation of PKC- but was not toxic in RPTC validated the use of this inhibitor in our study. It is known that, at higher concentrations, Go6976 can also inhibit other PKC isozymes such as PKC- and PKC-µ. However, these isozymes were not activated in RPTC at any time point during the injury or the recovery of DCVC-injured RPTC. Inibition of PKC- activation by Go6976 did not affect the DCVC-induced cell loss or decreases in mitochondrial functions, which showed that Go6976 is not toxic in RPTC. The initial (at 4 h) decreases in mitochondrial function in DCVC-injured RPTC were not accompanied by PKC- activation, which suggests that PKC- does not play a role in the initial decline of mitochondrial function. However, our results demonstrated that the lack of mitochondrial repair was associated with the activation of PKC- and that attenuation of PKC- activation during the 4-day recovery period promoted the return of overall mitochondrial function (improved basal Q O 2 ). Specifically, the inhibition of PKC- activation promoted recovery of the electron transfer rate and oxidative phosphorylation (improved uncoupled Q O 2 and oligomycin-sensitive Q O 2 ). These results suggest that PKC- is involved in the inhibition of recovery of these functions in sublethally injured RPTC. Similar to the previous studies ( 7, 38 ), we show that DCVC decreases m in RPTC. In contrast to other mitochondrial functions, m recovered on day 4 following DCVC injury. Furthermore, inhibition of PKC- activation did not affect the regain of m, suggesting that PKC- inhibits the repair of mitochondrial functions by mechanisms independent of m.
Because the inhibition of PKC- activation promoted the recovery of oligomycin-sensitive Q O 2 in DCVC-injured RPTC, we hypothesized that PKC- may target the F 1 F 0 -ATPase. Oligomycin-sensitive Q O 2 and F 1 F 0 -ATPase activity decreased following DCVC treatment and remained at the reduced levels throughout the recovery period. Inhibition of PKC- activation did not affect the initial reductions in F 1 F 0 -ATPase activity following DCVC treatment but promoted the recovery of the enzyme activity on day 4. Therefore, our data suggest that PKC- does not play a role in the initial inhibition of F 1 F 0 -ATPase but negatively regulates the enzyme activity during the repair period after DCVC exposure. This conclusion is consistent with the observation that PKC- activation in mitochondria does not occur during the early time points following DCVC exposure.
PKC- has a variety of intracellular targets and regulates multiple processes including gene transcription and translation ( 11 ). It is likely that PKC- inhibits transcription and translation of F 1 F 0 -ATPase subunits in DCVC-injured RPTC. Therefore, we examined whether DCVC injury alters the protein levels of the catalytic ( ) subunit of F 1 F 0 -ATPase in RPTC. The protein levels of the -subunit of F 1 F 0 -ATPase in both cell homogenates and mitochondria in DCVC-treated RPTC remained unchanged during the whole recovery period. This indicated that the decreases in F 1 F 0 -ATPase activity following DCVC injury were not due to decreased protein levels of this enzyme. Thus, our data suggested that F 1 F 0 -ATPase activity may be regulated by posttranslational modification of one or more subunits of F 1 F 0 -ATPase leading to alterations in catalytic activity. Protein phosphorylation, a posttranslational modification producing changes in conformation and enzymatic activity, may represent a mechanism regulating F 1 F 0 -ATPase activity. Indeed, tyrosine phosphorylation of the -subunit of F 1 F 0 -ATPase was described previously in cortical neurons following treatment with platelet-derived growth factor, but the functional consequences of this phosphorylation are unknown ( 42 ).
We performed analysis of amino acid sequences on several F 1 F 0 -ATPase subunits. The results suggested the existence of several PKC consensus motifs on yeast subunit, human subunit, coupling factor 6, and -subunit from Chaetosphaeridium globosum. Therefore, we tested whether PKC targets F 1 F 0 -ATPase directly by phosphorylating one or more subunits. Our results show that recombinant PKC- catalyzed incorporation of 32 P to the -subunit of F 1 F 0 -ATPase under in vitro conditions. In contrast, PKC- and PKC- did not phosphorylate F 1 F 0 -ATPase under the same in vitro conditions. Thus our data suggest that PKC- -mediated phosphorylation of the catalytic -subunit plays a role in the regulation of F 1 F 0 -ATPase activity. To determine whether the -subunit of F 1 F 0 -ATPase is phosphorylated by PKC- following DCVC exposure in RPTC, we carried out an immunoprecipitation of this subunit from RPTC lysates and examined its phosphorylation status. Phosphoserine levels on the -subunit of F 1 F 0 -ATPase were increased in DCVC-injured RPTC on day 4 but not during the early recovery period. PKC- inhibition abolished phosphorylation of the -subunit on day 4, which is consistent with the activation and involvement of PKC- in the inhibition of F 1 F 0 -ATPase activity in the late recovery period. In contrast, the phosphothreonine levels remained unchanged in DCVC-injured RPTC. Thus our results show that PKC- activation in RPTC during late recovery following DCVC exposure results in phosphorylation of serine(s) but not threonine(s) on the -subunit of F 1 F 0 -ATPase. The data also suggest that PKC- -mediated phosphorylation negatively regulates F 1 F 0 -ATPase activity. Indeed, catalytic activity of F 1 portion of F 1 F 0 -ATPase is decreased following phosphorylation by PKC- in vitro. However, the effect of phosphorylation on the activity of F 1 F 0 -ATPase in the cell is not known at present. Our data suggest that F 1 F 0 -ATPase phosphorylation decreases enzymatic activity and contributes to the lack of recovery of oxidative phosphorylation in DCVC-injured RPTC and that PKC- is the isozyme responsible for the modification of F 1 F 0 -ATPase activity in our model. Because our results are based on the use of a pharmacological PKC- inhibitor, another approach such as overexpression of dominant positive and negative PKC- is necessary to confirm the role of PKC- in phosphorylation and regulation of F 1 F 0 -ATPase activity in DCVC-injured RPTC.
The recovery of oxidative phosphorylation and overall mitochondrial function depends on the repair of electron transport. Previous reports demonstrated that DCVC alters the concentration of several citric acid cycle intermediates, reduces the activity of isocitrate dehydrogenase, and inhibits succinate:cytochrome c oxidoreductase and succinate-linked state 3 respiration, suggesting that DCVC targets electron transport in RPTC ( 1, 15 ). Furthermore, our previous data showed that electron transport rate does not recover in DCVC-injured RPTC ( 29 ). These results show that attenuation of PKC- activation partially promotes the recovery of the electron transport rate in DCVC-injured RPTC. Thus our results suggest that, in addition to phosphorylating F 1 F 0 -ATPase, PKC- inhibits the repair of mitochondrial function also by decreasing the electron transport rate. Although inhibition of PKC- activation promoted the recovery of mitochondrial respiration and ATP production, it did not restore the intracellular ATP content in RPTC. The reason for this apparent dilemma is unknown at present. Because the intracellular ATP content reflects a net difference between synthesis and consumption of ATP, it is conceivable that the inhibition of PKC- activation also promoted the repair of energy-consuming processes such as active ion transport and thus increased ATP consumption.
The present study shows that the inhibition of PKC- only partially promoted the recovery of mitochondrial function, indicating that other mechanisms are also involved in the lack of the repair of mitochondrial functions in DCVC-injured RPTC. The other mechanisms may include the growth factor receptor-mediated mechanisms that result in activation of tyrosine kinase and phosphorylation of tyrosine residues in target proteins ( 40 ). Indeed, tyrosine phosphorylation of the -subunit of F 0 F 1 -ATPase was described in cortical neurons treated with platelet-derived growth factor ( 42 ). Furthermore, our previous studies demonstrating that epidermal growth factor promotes recovery of mitochondrial function and active Na + transport following DCVC injury support the role of growth factor receptor- and tyrosine kinase-mediated mechanisms in the repair of mitochondrial function ( 29 ). Thus it is very likely that the recovery of mitochondrial function is under control of more than one signaling pathway.
In conclusion, the lack of repair of mitochondrial function following DCVC exposure is associated with activation of PKC-, including mitochondrial PKC-. PKC- activation during the repair period in DCVC-injured RPTC mediates the inhibition of the recovery of the electron transfer rate, oxidative phosphorylation, ATP production, and F 1 F 0 -ATPase activity. Furthermore, our data show that PKC- phosphorylates the -subunit of F 1 F 0 -ATPase both under in vitro conditions and in recovering RPTC and that in vitro phosphorylation of F 1 F 0 -ATPase decreases its catalytic activity. These results suggest that phosphorylation of the catalytic -subunit of F 1 F 0 -ATPase by PKC- represents a mechanism regulating F 1 F 0 -ATPase activity and the repair of mitochondrial function following toxicant-induced injury in RPTC.
GRANTS
This work was supported by National Institutes of Health National Institute of Diabetes and Digestive and Kidney Diseases Grant R01-DK-59558.
【参考文献】
Anders MW. Mitochondrial bioactivation of cysteine S-conjugate and 4-thiaalkanoates: implications for mitochondrial dysfunction and mitochondrial diseases. Biochim Biophys Acta 1271: 51-57, 1995.
Borkan SC, Emami A, and Schwartz JH. Heat stress protein-associated cytoprotection of inner medullary collecting duct cells from rat kidney. Am J Physiol Renal Fluid Electrolyte Physiol 265: F333-F341, 1993.
Boyer PD. A research journey with ATP synthase. J Biol Chem 277: 39045-39061, 2002.
Boyer PD. The ATP synthase-a splendid molecular machine. Annu Rev Biochem 66: 717-749, 1997.
Catterall WA and Pedersen PL. Adenosine triphosphatase from rat liver mitochondria. I. Purification, homogeneity, and physical properties. J Biol Chem 246: 4987-4994, 1971.
Chen Q, Jones TW, and Stevens JL. Early cellular events couple covalent binding of reactive metabolites to cell killing by nephrotoxic cysteine conjugates. J Cell Physiol 161: 293-302, 1994.
Chen Y, Cai J, Anders MW, Stevens JL, and Jones DP. Role of mitochondrial dysfunction in S -(1,2-dichlorovinyl)- L -cysteine-induced apoptosis. Toxicol Appl Pharmacol 170: 172-180, 2001.
Dempsey EC, Newton AC, Mochly-Rosen D, Fields AP, Reyland ME, Insel PA, and Messing RO. Protein kinase C isozymes and the regulation of diverse cell responses. Am J Physiol Lung Cell Mol Physiol 279: L429-L438, 2000.
Dong L, Stevens JL, Fabbro D, and Jaken S. Regulation of protein kinase C isozymes in kidney regeneration. Cancer Res 53: 542-549, 1993.
Elfarra AA, Jakobson I, and Anders MW. Mechanisms of S -(1,2-dichlorovinyl) glutathione-induced nephrotoxicity. Biochem Pharmacol 35: 283-288, 1986.
Franklin RB, Zou J, Ma J, and Costello LC. Protein kinase C, and AP-1 mediate prolactin regulation of mitochondrial aspartate aminotransferase expression in the rat lateral prostate. Mol Cell Endocrinol 170: 153-161, 2000.
Ilinskaja O and Vamvakas S. Alterations of the renal function in the isolated perfused rat kidney system after in vivo and in vitro application of S -(1,2-dichlorovinyl)- L -cysteine and S -(2,2-dichlorovinyl)- L -cysteine. Arch Toxicol 70: 224-229, 1996.
Laemmli UK. Cleavage of structural proteins during the assembly of the head bacteriophage T4. Nature 227: 680-685, 1970.
Lash LH and Anders MW. Cytotoxicity of S -(1,2-dichlorovinyl)glutathione and S -(1,2-dichlorovinyl)- L -cysteine in isolated rat kidney cells. J Biol Chem 261: 13076-13081, 1986.
Lash LH and Anders MW. Mechanism of S -(1,2-dichlorovinyl)- L -cysteine- and S -(1,2-dichlorovinyl)- L -homocysteine-induced renal mitochondrial toxicity. Mol Pharmacol 32: 549-556, 1987.
Lash LH, Elfarra AA, and Anders MW. Renal cysteine conjugate -lyase. Bioactivation of nephrotoxic cysteine S -conjugates in mitochondrial outer membrane. J Biol Chem 261: 5930-5935, 1986.
Lash LH, Hueni SE, and Putt DA. Apoptosis, necrosis and cell proliferation induced by S -(1,2-dichlorovinyl)- L -cysteine in primary cultures of human proximal tubular cells. Toxicol Appl Pharmacol 177: 1-16, 2001.
Lash LH, Qian W, Putt DA, Hueni SE, Elfarra AA, Krause RJ, and Parker JC. Renal and hepatic toxicity of trichloroethylene and its glutathione-derived metabolites in rats and mice: sex-, species-, and tissue-dependent differences. J Pharmacol Exp Ther 297: 155-164, 2001.
Lash LH and Sall JM. Mitochondrial isolation from liver and kidney: strategy, techniques, and criteria for purity. In: Methods in Toxicology. Orlando, FL: Academic, 1993, vol. 2, pp. 8-28.
Lash LH, Xu Y, Elfarra AA, Duescher RJ, and Parker JC. Glutathione-dependent metabolism of trichloroethylene in isolated liver and kidney cells of rats and its role in mitochondrial and cellular toxicity. Drug Metab Dispos 23: 846-853, 1995.
Law RHP, Manon S, Devenish RJ, and Nagley P. ATP synthase from Saccharomyces cerevisiae. Methods Enzymol 260: 133-190, 1995.
Mischak H, Pierce JH, Goodnight J, Kazanietz MG, Blumberg PM, and Mushinski JF. Phorbol ester-induced myeloid differentiation is mediated by protein kinase C- and - and not by protein kinase C- II, -, -, and -. J Biol Chem 268: 20110-20115, 1993.
Nony PA, Nowak G, and Schnellmann RG. Collagen IV promotes repair of renal cell physiological functions after toxicant injury. Am J Physiol Renal Physiol 281: F443-F453, 2001.
Nony PA and Schnellmann RG. Interactions between collagen IV and collagen-binding integrins in renal cell repair after sublethal injury. Mol Pharmacol 60: 1226-1234, 2001.
Nowak G. Protein kinase C- and ERK1/2 mediate mitochondrial dysfunction, decreases in active Na + transport, and cisplatin-induced apotosis in renal cells. J Biol Chem 277: 43377-43388, 2002.
Nowak G. Protein kinase C mediates repair of mitochondrial and transport functions following toxicant-induced injury in renal cells. J Pharmacol Exp Ther 306: 1-9, 2003.
Nowak G, Aleo MD, Morgan JA, and Schnellmann RG. Recovery of cellular functions following oxidant injury. Am J Physiol Renal Physiol 274: F509-F515, 1998.
Nowak G, Carter CA, and Schnellmann RG. Ascorbic acid promotes recovery of cellular functions following toxicant-induced injury. Toxicol Appl Pharmacol 167: 37-45, 2000.
Nowak G, Keasler KB, McKeller DE, and Schnellmann RG. Differential effects of EGF on repair of cellular functions after dichlorovinyl- L -cysteine-induced injury. Am J Physiol Renal Physiol 276: F228-F236, 1999.
Nowak G and Schnellmann RG. L -Ascorbic acid regulates growth and metabolism of renal cells: improvements in cell culture. Am J Physiol Cell Physiol 271: C2072-C2080, 1996.
Moore RB and Green T. The synthesis of nephrotoxin conjugates of glutathione and cysteine. Toxicol Environ Chem 17: 153-162, 1988.
Pedersen PL and Amzel LM. ATP synthases. Structure, reaction center, mechanism, and regulation of one of nature's most unique machines. J Biol Chem 268: 9937-9940, 1993.
Perletti GP, Smeraldi C, Porro D, and Piccinni F. Involvement of the -isoenzyme of protein kinase C in the growth inhibition induced by phorbol esters in MH1C1 hepatoma cells. Biochem Biophys Res Commun 205: 1589-1594, 1994.
Saraste M. Oxidative phosphorylation at the fin de siecle. Science 283: 1488-1493, 1999.
Stevens JL, Ayoubi N, and Robbins JD. The role of mitochondrial matrix enzymes in the metabolism and toxicity of cysteine conjugates. J Biol Chem 263: 3395-3401, 1988.
Vamvakas S, Bittner D, Dekant W, and Anders MW. Events that precede and that follow S -(1,2-dichlorovinyl)- L -cysteine-induced release of mitochondrial Ca 2+ and their association with cytotoxicity to renal cells. Biochem Pharmacol 44: 1131-1138, 1992.
Van de Water B, Zoetewey PJ, de Bont HJGM, Mulder GJ, and Nagelkerke JF. The relationship between intracellular Ca 2+ and the mitochondrial membrane potential in isolated proximal tubular cells from rat kidney exposed to the nephrotoxin 1,2-dichlorovinyl-cysteine. Biochem Pharmacol 45: 2259-2267, 1993.
Van de Water B, Zoeteweij PJ, de Bont HJGM, Mulder GJ, and Nagelkerke FJ. Role of mitochondrial Ca 2+ in the oxidative stress-induced dissipation of the mitochondrial membrane potential. Studies in isolated proximal tubular cells using the nephrotoxin 1,2-dichlorovinyl- L -cysteine. J Biol Chem 269: 14546-14552, 1994.
Wallin A, Zhang G, Jones TW, Jaken S, and Stevens JL. Mechanism of the nephrogenic repair response. Studies on proliferation and vimentin expression after 35 S -1,2-dichlorovinyl- L -cysteine nephrotoxicity in vivo and in cultured proximal tubule epithelial cells. Lab Invest 66: 474-484, 1992.
Weinberg JM. The cell biology of ischemic renal injury. Kidney Int 39: 476-500, 1991.
Yasuda I, Kishimoto A, Tanaka S, Tominaga M, Sakurai A, and Nishizuka Y. A synthetic peptide substrate for selective assay of protein kinase C. Biochem Biophys Res Commun 166: 1220-1227, 1990.
Zhang FX, Pan W, and Hutchins JB. Phosphorylation of F 1 F 0 ATPase delta-subunit is regulated by platelet-derived growth factor in mouse cortical neurons in vitro. J Neurochem 65: 2812-2815, 1995.
Zhang G, Ichimura T, Maier JA, Maciag T, and Stevens JL. A role for fibroblast growth factor type-1 in nephrogenic repair. Autocrine expression in rat kidney proximal tubule epithelial cells in vitro and in the regenerating epithelium following nephrotoxic damage by S -(1,1,2,2-tetrafluoroethyl)- L -cysteine in vivo. J Biol Chem 268: 11542-11547, 1993.
作者单位：Department of Pharmaceutical Sciences, University of Arkansas for Medical Sciences, Little Rock, Arkansas 72205