Department of Pharmaceutical Sciences, University of Arkansas for Medical Sciences, Little Rock, Arkansas 72205
Submitted 2 June 2003 ; accepted in final form 29 February 2004
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ABSTRACT |
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renal proximal tubular cells; mitochondria; F1F0-ATPase
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 F1F0-ATPase activity, suggesting an important role of PKC-
in the regulation of F1F0-ATPase activity (25).
F1F0-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, F1F0-ATPase operates in a reverse mode and hydrolyzes ATP. The enzyme has multiple subunits forming two major complexes, F0 and F1 (3, 4). F1 is a water-soluble catalytic complex made up of five subunits (
3
3
), with the catalytic site located on the
-subunit. F0 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 F1F0-ATPase activity. It has been shown that platelet-derived growth factor induces tyrosine phosphorylation of the
-subunit of F1F0-ATPase (42). Indeed, our search of the amino acid sequence of F1F0-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.
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MATERIALS AND METHODS |
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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 NaHCO3, 15 mM HEPES, and 6 mM lactate (pH 7.4, 295 mosmol/kgH2O). 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 (QO2) measurement chamber. QO2 was measured using a Clark-type electrode as described previously (25, 29, 30). Oligomycin-sensitive QO2 was used as a marker of oxidative phosphorylation and was calculated as a difference between basal QO2 and oligomycin-insensitive QO2, which was measured in the presence of oligomycin (0.6 µg/ml). Uncoupled QO2 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 KH2PO4, 10 mM HEPES, 1 mM MgSO4, 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 MgCl2, 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 F1F0-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 Na3VO4, 1 mM NaF, and the protease and phosphatase inhibitors cocktail; pH 7.4) and used for immunoblot analysis.
Measurement of F1F0-ATPase activity. F1F0-ATPase activity was determined in freshly isolated RPTC mitochondria by measuring the release of Pi from ATP according to Law et al. (21) with modifications described previously (23).
Isolation of F1F0-ATPase. Freshly isolated rabbit renal cortical mitochondria were used for the isolation of F1F0-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 (50250 mM; pH 7.5) containing 5.0 mM EDTA at a flow rate of 2 ml/min. The fractions eluted by 150200 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 F1F0-ATPase by PKC-.
In vitro phosphorylation of isolated F1F0-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 CaCl2, 20 mM MgCl2, 6 mM
-mercaptoethanol, 10 µM PMA, 280 µg/ml L-
-phosphatidyl-L-serine, 2.4 µg F1F0-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 [
-32P]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 32P 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 32P incorporation was quantified by liquid scintillation spectrometry.
To determine the identities of phosphorylated proteins, immunoblot analysis of the isolated F1F0-ATPase (2 µg) was performed using antibodies against various subunits of F1F0-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 F1F0-ATPase catalytic activity, the phosphorylation assay was performed as described above using nonradioactive ATP and the enzymatic activity of F1F0-ATPase was determined in the reaction mixture by measuring the release of Pi 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 F1F0-ATPase subunits in total RPTC homogenates, mitochondria, and F1F0-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 F1F0-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 F1F0-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.
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RESULTS |
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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. 1D). Go6976 abolished the activation of PKC-
in DCVC-treated RPTC at all time points studied (Fig. 1A). DCVC-induced injury had no effect on phosphorylation of PKC-
(Fig. 1C) and PKC-µ (data not shown).
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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. 4A). 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. 4B). Go6976 had no effects on
m in control or DCVC-injured RPTC (Fig. 4B). 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-
.
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Oxidative phosphorylation.
Oxidative phosphorylation and ATP synthesis are the fundamental functions of mitochondria. Oligomycin-sensitive QO2 was used as a marker of oxidative phosphorylation in RPTC. Oligomycin-sensitive QO2 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 QO2 at 4 h but promoted, in part, the return of oligomycin-sensitive QO2 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.
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Phosphorylation of F1F0-ATPase by PKC- in vitro.
PKC, a serine/threonine kinase, phosphorylates a wide range of intracellular substrates. Several F1F0-ATPase subunits, including the
-subunit (Fig. 8), have PKC consensus amino acid motifs. An in vitro assay was performed to determine whether F1F0-ATPase subunits are phosphorylated by major PKC isozymes present in RPTC. Phosphorylation of F1F0-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 F1F0-ATPase in vitro (Fig. 9A).
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An in vitro assay was carried out to determine whether phosphorylation of F1F0-ATPase by PKC- alters F1F0-ATPase activity. Figure 9E shows that the activity of F1F0-ATPase decreased 53% in the presence of active PKC-
and ATP. These results suggest that phosphorylation by PKC-
has an inhibitory effect on F1F0-ATPase.
Phosphorylation of F1F0-ATPase by PKC- in DCVC-injured RPTC.
Immunoprecipitation was used to determine 1) whether phosphorylation of the
-subunit of F1F0-ATPase occurs in DCVC-injured RPTC and 2) whether the
-subunit of F1F0-ATPase is phosphorylated by PKC-
following DCVC exposure. The
-subunit of F1F0-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. 10B, 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 F1F0-ATPase were not altered at 1, 2, and 4 h but decreased at 8 h of the recovery period in DCVC-injured RPTC (Fig. 10A, data not shown). However, on day 4 of the recovery period, phosphoserine levels on the
-subunit of F1F0-ATPase increased in DCVC-injured RPTC (Fig. 10A). PKC-
inhibitor Go6976 decreased the serine phosphorylation of the
-subunit of F1F0-ATPase on day 4 of the recovery period (Fig. 10A). 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 F1F0-ATPase.
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DISCUSSION |
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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 QO2). Specifically, the inhibition of PKC-
activation promoted recovery of the electron transfer rate and oxidative phosphorylation (improved uncoupled QO2 and oligomycin-sensitive QO2). 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 QO2 in DCVC-injured RPTC, we hypothesized that PKC-
may target the F1F0-ATPase. Oligomycin-sensitive QO2 and F1F0-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 F1F0-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 F1F0-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 F1F0-ATPase subunits in DCVC-injured RPTC. Therefore, we examined whether DCVC injury alters the protein levels of the catalytic (
) subunit of F1F0-ATPase in RPTC. The protein levels of the
-subunit of F1F0-ATPase in both cell homogenates and mitochondria in DCVC-treated RPTC remained unchanged during the whole recovery period. This indicated that the decreases in F1F0-ATPase activity following DCVC injury were not due to decreased protein levels of this enzyme. Thus, our data suggested that F1F0-ATPase activity may be regulated by posttranslational modification of one or more subunits of F1F0-ATPase leading to alterations in catalytic activity. Protein phosphorylation, a posttranslational modification producing changes in conformation and enzymatic activity, may represent a mechanism regulating F1F0-ATPase activity. Indeed, tyrosine phosphorylation of the
-subunit of F1F0-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 F1F0-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 F1F0-ATPase directly by phosphorylating one or more subunits. Our results show that recombinant PKC-
catalyzed incorporation of 32P to the
-subunit of F1F0-ATPase under in vitro conditions. In contrast, PKC-
and PKC-
did not phosphorylate F1F0-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 F1F0-ATPase activity. To determine whether the
-subunit of F1F0-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 F1F0-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 F1F0-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 F1F0-ATPase. The data also suggest that PKC-
-mediated phosphorylation negatively regulates F1F0-ATPase activity. Indeed, catalytic activity of F1 portion of F1F0-ATPase is decreased following phosphorylation by PKC-
in vitro. However, the effect of phosphorylation on the activity of F1F0-ATPase in the cell is not known at present. Our data suggest that F1F0-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 F1F0-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 F1F0-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 F1F0-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 F0F1-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 F1F0-ATPase activity. Furthermore, our data show that PKC-
phosphorylates the
-subunit of F1F0-ATPase both under in vitro conditions and in recovering RPTC and that in vitro phosphorylation of F1F0-ATPase decreases its catalytic activity. These results suggest that phosphorylation of the catalytic
-subunit of F1F0-ATPase by PKC-
represents a mechanism regulating F1F0-ATPase activity and the repair of mitochondrial function following toxicant-induced injury in RPTC.
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GRANTS |
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FOOTNOTES |
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The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
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REFERENCES |
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