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TOXICOLOGY

Protein Kinase C Mediates Repair of Mitochondrial and Transport Functions after Toxicant-Induced Injury in Renal Cells

Grayna Nowak

Department of Pharmaceutical Sciences, University of Arkansas for Medical Sciences, Little Rock, Arkansas

Received February 8, 2003; accepted March 27, 2003.

	Abstract

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Previously, we have shown that renal proximal tubular cells (RPTCs) recover physiological functions after injury induced by the oxidant tert-butylhydroperoxide (TBHP), but not by the nephrotoxic cysteine conjugate dichlorovinyl-L-cysteine (DCVC). This study examined the role of protein kinase C (PKC) in the repair of RPTC functions after sublethal injury produced by these toxicants. Total PKC activity decreased 65 and 86% after TBHP and DCVC exposures, respectively, and recovered in TBHP-injured but not in DCVC-injured RPTCs. Mitochondrial function, active Na⁺ transport, and Na⁺-dependent glucose uptake decreased after toxicant exposure and recovered in TBHP- but not in DCVC-injured RPTCs. PKC inhibition decreased the repair of RPTC functions after TBHP injury. PKC activation promoted recovery of mitochondrial function and active Na⁺ transport in TBHP- and DCVC-injured RPTCs but had no effect on recovery of Na⁺-dependent glucose uptake. We conclude that in RPTCs, 1) total PKC activity decreases after TBHP and DCVC injury and recovers after TBHP but not after DCVC exposure, 2) recovery of PKC activity precedes the return of physiological functions after oxidant injury, 3) PKC inhibition decreases recovery of physiological functions, and 4) PKC activation promotes recovery of mitochondrial function and active Na⁺ transport but not Na⁺-dependent glucose uptake. These results suggest that the repair of renal functions is mediated through PKC-dependent mechanisms and that cysteine conjugates may inhibit renal repair, in part, through inhibition of PKC signaling.

The kidney has the potential for complete recovery from ARF (Toback, 1992). Using an in vitro model of primary cultures of RPTCs, we have shown that RPTCs proliferate and recover physiological functions after sublethal injury induced by the oxidant TBHP (Nowak et al., 1998). However, the repair of RPTC functions does not occur in sublethally injured RPTCs after DCVC exposure (Nowak et al., 1999). DCVC also decreases synthesis of extracellular matrix proteins of the RPTC basement membrane, including collagen IV, and disrupts localization of collagen binding integrins (Nowak et al., 2000; Nony et al., 2001; Nony and Schnellmann, 2001). Interestingly, epidermal growth factor and pharmacological concentrations of ascorbic acid promote the repair of mitochondrial function and active Na⁺ transport in DCVC-injured RPTCs (Nowak et al., 1999, 2000; Nony et al., 2001). The promotion of RPTC repair by ascorbic acid is mediated, in part, by collagen IV and is associated with the relocalization of collagen-binding integrins to the basement membrane (Nony et al., 2001).

Protein kinase C (PKC), a family of serine/threonine protein kinases, controls 70% of the phosphorylating activity in renal proximal tubules (Kobryn and Mandel, 1994) and regulates numerous physiological functions of renal epithelial cells, including gluconeogenesis, Na⁺/K⁺-ATPase activity, and the transport of amino acids, glucose, sodium, potassium, chloride, phosphate, water, and organic anions and cations (Dempsey et al., 2000). Recent studies suggest that PKC is also involved in the regulation of cell survival and drug-induced cell injury (Dempsey et al., 2000). PKC is a target for some toxicants. Short-time exposure to oxidants activates PKC in some cell types (O'Brian et al., 1988; Palumbo et al., 1992; Abe et al., 1998). In contrast, longer exposures to oxidants inactivate PKC by modifying its catalytic domain (Gopalakrishna and Anderson, 1987).

PKC signaling regulates the regenerative processes in the liver after partial hepatectomy (Daller et al., 1994; Tessitore et al., 1995). PKC also has been implicated in the renal recovery after ARF (Alberti et al., 1993; La Porta and Commoli, 1993) and in wound healing after mechanically induced injury in renal tubular epithelial cells (Sponsel et al., 1995). Renal regeneration after toxicant exposure is associated with differential modulation of PKC isozymes. Activation of PKC, PKC, and PKC, but not PKC or PKC occurs during compensatory renal hypertrophy induced by unilateral nephrectomy (Dong et al., 1993). Regeneration after folic acid-and DCVC-induced injury in the kidney is associated with down-regulation of PKC and PKC (Dong et al., 1993; Zhang et al., 1993). Although PKC seems to be involved in renal recovery after toxic insult, it is not known what functions and processes are regulated by this kinase in RPTCs. Therefore, the aim of this study was to determine whether PKC plays a role in the recovery of mitochondrial function, active Na⁺ transport, and Na⁺-coupled glucose uptake in RPTCs after sublethal injury induced by two different toxicant, the oxidant TBHP and the model halocarbon DCVC.

	Materials and Methods

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Animals. Female New Zealand White rabbits (2.0–2.5 kg) purchased from Myrtle's Rabbitry (Thompson Station, TN) were used in this study.

Chemicals and Reagents. DCVC was synthesized according to the method of Moore and Green (1988). L-Ascorbic acid-2-phosphate magnesium salt was purchased from Wako Bioproducts (Richmond, VA). Protein kinase C assay kit, phorbol 12-myristate 13-acetate (PMA), and cell culture media were obtained from Invitrogen (Carlsbad, CA). Cell culture hormones, tert-butylhydroperoxide, and protease and phosphatase inhibitors were obtained from Sigma-Aldrich (St. Louis, MO). Calphostin C was purchased from Calbiochem (La Jolla, CA). Cell-permeable PKC peptide inhibitor, myristoylated protein kinase C fragment (20–28) (myr-Phe-Ala-Arg-Lys-Gly-Ala-Leu-Arg-Gln; pseudosubstrate sequence from PKC) was supplied by BIOMOL Research Laboratories (Plymouth Meeting, PA). [-³²P]ATP (specific activity 3000 Ci/mmol) and methyl -D-glucopyranoside, [glucose-¹⁴C(U)] (specific activity 282 mCi/mmol) were purchased from Amersham Biosciences, Inc. (Piscataway, NJ) and PerkinElmer Life Sciences (Boston, MA), respectively. PKC-, PKC-, PKC-, and PKC- antibodies were obtained from BD Transduction Laboratories (San Diego, CA). Antibodies against phosphorylated forms of PKC- and PKC- were supplied by Upstate Biotechnology (Lake Placid, NY). Anti-mouse IgG coupled to horseradish peroxidase was supplied by Kirkegaard and Perry Laboratories (Gaithersburg, MD) and SuperSignal chemiluminescent substrate by Pierce Chemical (Rockford, IL). The sources of the other reagents have been described previously (Nowak and Schnellmann, 1996).

Isolation of Proximal Tubules and Culture Conditions. Rabbit renal proximal tubules were isolated by iron oxide perfusion method and grown in 35-mm culture dishes in improved conditions as described previously (Nowak and Schnellmann, 1996). The purity of the renal proximal tubular S₁ and S₂ segments isolated by this method is approximately 96%. The culture medium was a 50:50 mixture of Dulbecco's modified Eagle's medium and Ham's F-12 nutrient mix without phenol red, pyruvate, and glucose, supplemented with 15 mM NaHCO₃, 15 mM HEPES, and 6 mM lactate (pH 7.4, 295 mOsM/kg). Human transferrin (5 µg/ml), selenium (5 ng/ml), hydrocortisone (50 nM), bovine insulin (10 nM), and L-ascorbic acid-2-phosphate (0.05 mM) were added to the medium immediately before daily media change (2 ml/dish).

Toxicant Treatment of RPTC Monolayer. RPTC cultures reached confluence within 6 days and were treated with toxicants on the 7th day of culture. RPTCs were treated with 0.2 mM TBHP for 45 to 50 min or with 0.2 mM DCVC for 1.5 h to obtain approximately 25% cell death and loss. After toxicant exposure, the remaining cellular monolayer was washed with fresh culture medium and cultured for the following 4 days. PKC inhibitors [100 nM calphostin C and 20 µM myristoylated protein kinase C (20–28) peptide] were added daily starting with the media change after TBHP exposure. PKC activator, 100 nM PMA, was added 30 min before toxicant and was present in the medium for 24 h after toxicant exposure. Samples of RPTCs were taken at various time points after toxicant exposure for measurements of cellular functions.

Isolation of Cytosolic and Membrane Fractions. RPTC samples were harvested at various time points during recovery period after toxicant exposure. Monolayers were washed four times with ice-cold phosphate-buffered saline (pH 7.4) to remove all nonviable cells, cells scraped from the dishes, suspended in 1 ml of phosphate-buffered saline, and pelleted by centrifugation. RPTC pellet was resuspended in ice-cold isolation buffer (20 mM Tris-HCl, pH 7.5, containing 10 mM MgCl₂, 80 mM -glycerophosphate, 2 mM EGTA, 2 mM EDTA, 2 mM dithiothreitol, 1 mM phenylmethylsulfonyl fluoride, 20 µg/ml aprotinin, 5 µg/ml pepstatin, and 5 µg/ml leupeptin). The cells were briefly sonicated (3 x 5 s) on ice and centrifuged at 1000g for 5 min to remove cell debris and nuclei. The supernatant was spun down at 100,000g for 30 min at 4°C, and the supernatant resulting from this centrifugation represented cytosolic fraction. The pellet was resuspended in the isolation buffer containing 1% Triton X-100, incubated on ice for 30 min, and centrifuged at 100,000g for 30 min at 4°C. The supernatant resulting from this spin represented the particulate fraction.

Measurement of PKC Activity. The activity of PKC was based on measurement of the phosphorylation of a synthetic peptide derived from the myelin basic protein (MBP) sequence (4–14) as described by Yasuda et al. (1990). The N-terminal glutamine of this peptide has been acetylated to maintain the peptide's stability [Ac-MBP(4–14)]. Specificity of the phosphorylation of Ac-MBP(4–14) by PKC was confirmed by using the PKC pseudosubstrate inhibitor peptide PKC(19–36), which acts as a potent inhibitor for this substrate. PKC activity was measured in the reaction mixture that contained 20 mM Tris (pH 7.5), 20 mM MgCl₂, 1.0 mM CaCl₂, 0.28 mg/ml phosphatidyl-L-serine, 10 µM PMA, 1 mM dithiotreitol, pepstatin, leupeptin, aprotinin (25 µg/ml each), 1 mM phenylmethylsulfonyl fluoride, 40 mM -glycerophosphate, 0.3% Triton X-100, 50 µM Ac-MBP(4–14), 20 µM [-³²P]ATP, and sample. Specificity of the phosphorylation of Ac-MBP(4–14) by PKC was measured in the reaction mixture containing all above-mentioned components and 20 µM PKC(19–36) peptide inhibitor. The reaction was carried out for 5 min at 30°C and terminated by spotting an aliquot of reaction mixture onto P-81 phosphocellulose discs. After washing the discs, -³²P incorporation into substrate was determined by liquid scintillation spectrometry. Specific PKC activity was calculated as the PKC(19–36) inhibitor-sensitive activity. Nonspecific activity (background) was determined in samples incubated in the absence of activators (phosphatidylserine, PMA, and CaCl₂).

Immunoblotting. Immunoblot analysis was used for the measurement of protein levels of PKC-, PKC-, PKC-, and PKC- in the cytosolic and particulate fractions of RPTCs and for assessment of levels of phosphorylated forms of PKC- and PKC- in total RPTC homogenates. Samples of cytosolic and particulate fractions and cell homogenates were lysed and boiled for 10 min in Laemmli sample buffer (60 mM Tris-HCl, pH 6.8, containing 2% SDS, 10% glycerol, 100 mM -mercaptoethanol, and 0.01% bromphenol blue) (Laemmli, 1970). Proteins were separated using SDS-polyacrylamide gel electrophoresis. After electroblotting of the proteins to a nitrocellulose membrane, blots were blocked for 1 h in Tris-buffered saline buffer containing 0.5% casein and 0.1% Tween 20 (blocking buffer), and incubated overnight at 4°C in the presence of primary antibodies diluted in the blocking buffer. After washing with Tris-buffered saline containing 0.05% Tween 20, the membranes were incubated for 1 h with anti-rabbit or anti-mouse IgG coupled to horseradish peroxidase and washed again. The SuperSignal chemiluminescent system was used for protein detection. The results were quantified using scanning densitometry.

Oxygen Consumption. RPTC monolayers were washed with 37°C culture medium and gently detached from the dishes with a rubber policeman, suspended in warm (37°C) culture medium and transferred to the oxygen consumption (QO₂) measurement chamber. QO₂ was measured polarographically in RPTCs suspended in the culture medium using Clark type electrode as described previously (Nowak and Schnellmann, 1996; Nowak et al., 1998, 1999, 2000). Basal QO₂ was used as a marker of entire mitochondrial function. Oligomycin-sensitive QO₂ was used as a maker of oxidative phosphorylation. Oligomycin-sensitive QO₂ was determined after addition of oligomycin (1 µg/ml) and was calculated as the difference between basal and oligomycin-insensitive QO2. Uncoupled QO₂ was used as a marker of integrity of the electron transport chain and was measured after the addition of carbonyl cyanide p-trifluoromethoxyphenyl-hydrazone (1.5 µM).

Active Na⁺ Transport. Active Na⁺ transport was measured using the ouabain-sensitive QO₂ as a marker as described previously (Nowak and Schnellmann, 1996; Nowak et al., 1998, 1999, 2000). Ouabain-sensitive QO₂ was measured as the difference between basal QO₂ and QO₂ in the presence of 1 mM ouabain.

Na⁺-Coupled Glucose Uptake. Na⁺-coupled glucose uptake was assessed using the nonmetabolizable glucose analog methyl -D-glucopyranoside as described previously (Nowak and Schnellmann, 1996). Methyl -D-glucopyranoside uptake was measured in glucose-free medium used for RPTC culture and corrected for Na⁺-independent (phlorizin-insensitive) and zero time uptakes.

Protein Assay. Protein concentration in all samples was determined using bicinchoninic acid assay with bovine serum albumin as the standard.

Statistical Analysis. Data are presented as means ± S.E. and were analyzed for significance using analysis of variance. Multiple means were compared using Student-Newman-Keuls test. Statements of significance were based on P < 0.05. Renal proximal tubules isolated from an individual rabbit represented a separate experiment (n = 1) consisting of data obtained from two plates.

	Results

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Effect of TBHP and DCVC Exposures on PKC Activity in the Cytosolic and Particulate Fractions of RPTCs. Total PKC activity decreased by 60 and 63% in the cytosolic and particulate fractions of sublethally injured RPTCs, respectively, during the 24 h after TBHP exposure (Fig. 1, A and B). PKC activity in the cytosol returned to control levels on day 2 of the recovery period (Fig. 1A). However, PKC activity in the particulate fraction of RPTCs did not recover until day 4 after TBHP-induced injury (Fig. 1B).

View larger version (23K): [in this window] [in a new window]   Fig. 1. Time-dependent changes in total PKC activity in cytosolic (A) and particulate (B) fractions of RPTCs after TBHP-induced injury (0.2 mM; 45 min). Samples of RPTCs were collected at different time points after TBHP exposure and PKC activity was measured in cytosolic and particulate fractions as described under Materials and Methods. Data are presented as means ± S.E., n = 4 separate RPTC cultures. *, P < 0.05, significantly different from controls.

DCVC exposure decreased PKC activity in both the cytosolic and particulate fractions of RPTCs, but the DCVC-induced decreases in PKC activity were slower than those induced by TBHP. At 24 h after the DCVC treatment, PKC activity was decreased by 86% in both fractions (Fig. 2, A and B). In contrast to TBHP-induced injury, PKC activity in RPTCs did not recover after DCVC injury (Figs. 2 and 6).

View larger version (23K): [in this window] [in a new window]   Fig. 2. Time-dependent changes in total PKC activity in cytosolic (A) and particulate (B) fractions of RPTCs after DCVC-induced injury (0.2 mM; 90 min). Samples of RPTCs were collected at different time points after DCVC exposure and PKC activity measured in cytosolic and particulate fractions as described under Materials and Methods. Data are presented as means ± S.E., n = 5 separate RPTC cultures. *, P < 0.05, significantly different from controls.

View larger version (37K): [in this window] [in a new window]   Fig. 6. Decrease in the recovery of active Na+ transport by inhibition of PKC in TBHP-injured RPTCs. Two dissimilar inhibitors of PKC were used in these studies: myristoylated protein kinase C(20–28) peptide, 20 µM (A) and calphostin C, 100 nM (B). Data are presented as means ± S.E., n = 3 separate RPTC cultures. Values with dissimilar superscripts on a given day are significantly different (P < 0.05) from each other.

PKC and the Recovery of Mitochondrial Function. Basal, uncoupled, and oligomycin-sensitive QO₂s were used as markers of mitochondrial function in RPTCs. Specifically, uncoupled QO₂ served as a marker of the electron transport rate, whereas oligomycin-sensitive QO₂ was used as a marker of the oxidative phosphorylation. TBHP treatment decreased basal, uncoupled, and oligomycin-sensitive QO₂s by 41, 52, and 48%, respectively, at 4 h after the exposure (Figs. 3 and 4). Respiratory functions of RPTCs returned to control levels on day 4 of the recovery period (Figs. 3 and 4). The return of mitochondrial function followed the recovery of PKC activity in the cytosol and was accompanied by the recovery of PKC activity in the particulate fraction (Figs. 1, 3, and 4).

View larger version (32K): [in this window] [in a new window]   Fig. 3. Effect of PKC inhibition (calphostin C; 100 nM) on the recovery of overall mitochondrial function in RPTCs after TBHP-induced injury (0.2 mM; 45 min). Data are presented as means ± S.E., n = 3 separate RPTC cultures. Values with dissimilar superscripts on a given day are significantly different (P < 0.05) from each other.

View larger version (36K): [in this window] [in a new window]   Fig. 4. Effect of PKC inhibition [myristoylated protein kinase C(20–28) peptide derived from pseudosubstrate sequence of PKC, 20 µM] on the recovery of overall mitochondrial function (A), oxidative phosphorylation (A), and electron transport rate (C) in RPTCs after TBHP-induced injury (0.2 mM; 45 min). Data are presented as means ± S.E., n = 3 separate RPTC cultures. Values with dissimilar superscripts on a given day are significantly different (P < 0.05) from each other.

To determine whether PKC plays a role in the recovery of mitochondrial function after TBHP-induced injury, RPTCs were treated with a specific PKC inhibitor, calphostin C (100 nM), and the return of basal QO₂ examined during the recovery period. Calphostin C had no effect on basal QO₂ in control RPTCs or on the decrease in QO₂ in TBHP-treated RPTCs, but it inhibited the recovery of this function in sublethally injured RPTC (Fig. 3). Likewise, the treatment of RPTCs with another specific PKC inhibitor [PKC(20–28)], a peptide derived from pseudosubstrate sequence of PKC, prevented the recovery of basal QO₂ (Fig. 4A). Furthermore, PKC(20–28) inhibited the return of oligomycin-sensitive and uncoupled QO₂s in TBHP-injured RPTCs (Fig. 4, B and C).

Experiments were also performed to determine whether activation of PKC accelerates the repair of mitochondrial function after TBHP injury. PMA (100 nM) was used to activate PKC in RPTCs before toxicant-induced injury. Protein levels of PKC and PKC in the particulate fraction of RPTCs increased 1.3- and 2.5-fold, respectively, within 2 to 5 min of PMA treatment (Fig. 5A). Protein levels of PKC and PKC in the cytosolic fraction of PMA-treated RPTCs decreased concomitantly (Fig. 5A). The levels of phosphorylated forms of PKC and PKC were increased 2.5 and 4.4-fold, respectively, in RPTC treated for 5 min with 100 nM PMA, whereas the total levels of PKC and PKC remained unaffected (Fig. 5B). These data show that PMA induces phosphorylation and translocation of PKC and PKC from the cytosol to the particulate fraction, which suggests that PMA activates these PKC isozymes. No phosphorylation and translocation of PKC and PKC occurred in the presence of PMA in RPTCs (data not shown). PMA treatment before TBHP exposure did not protect against the decrease in mitochondrial function after the exposure (Fig. 5B). However, the recovery of mitochondrial function in PMA-treated RPTCs occurred on day 2 after TBHP injury and was accelerated in comparison with the recovery of this function in RPTCs pretreated with the vehicle (day 4) (Figs. 5B and 4A).

View larger version (42K): [in this window] [in a new window]   Fig. 5. Effect of PKC activation (PMA; 100 nM) on protein levels of PKC and PKC in cytosolic and particulate fractions of RPTCs (A) and phosphorylated and total protein levels of PKC and PKC in total cell homogenates after 5-min exposure of RPTCs to 100 nM PMA (B). C, controls; E, RPTCs treated with diluent (0.1% ethanol); P, RPTCs treated with 100 nM PMA for 5 min. C, effect of PKC activation (PMA; 100 nM) on the recovery of overall mitochondrial function in RPTCs after TBHP-induced injury. D, effect of PKC activation (PMA; 100 nM) on the recovery of overall mitochondrial function in RPTCs after DCVC-induced injury. Data are presented as means ± S.E., n = 3 separate RPTC cultures. Values with dissimilar superscripts on a given day are significantly different (P < 0.05) from each other.

DCVC exposure decreased basal QO₂ in sublethally injured RPTCs by 33 and 47% at 4 and 24 h, respectively. Oligomycin-sensitive QO₂ decreased by 77% (21.7 ± 1.1 versus 5.1 ± 3.2 nmol O₂/min/mg protein in control and DCVC-injured RPTCs, respectively) and uncoupled QO₂ decreased 63% (82.3 ± 5.6 versus 30.1 ± 3.7 nmol O₂/min/mg protein in control and DCVC-injured RPTCs, respectively) at 4 h after the exposure. No repair of basal QO₂ occurred after DCVC exposure (Fig. 5C). However, treatment with PMA before DCVC exposure resulted in the return of basal QO₂ within 2 days in injured RPTCs (Fig. 5C).

These results demonstrate that inhibition of PKC prevents the repair of mitochondrial function after toxicant injury in RPTCs and that activation of PKC before toxicant exposure promotes recovery of respiratory functions. Thus, these data suggest that PKC plays a role in the repair of mitochondrial function after toxic insult in RPTCs.

PKC and the Recovery of Active Na⁺ Transport. Active Na⁺ transport was used as a marker of the basolateral membrane function and was assessed by measurement of ouabain-sensitive QO₂. Active Na⁺ transport is an ATP-consuming process because it is driven by Na⁺/K⁺-ATPase. The consumption of oxygen associated with production of ATP required for maintaining the Na⁺/K⁺-ATPase activity and active Na⁺ transport accounts for approximately 50% of basal oxygen consumption in RPTCs. Ouabain, a specific Na⁺/K⁺-ATPase inhibitor, decreases oxygen consumption by the amount of oxygen associated with the synthesis of ATP that is consumed by Na⁺/K⁺-ATPase and active Na⁺ transport. Therefore, ouabain-sensitive portion of QO₂ is an indirect indicator of active Na⁺ transport.

Ouabain-sensitive QO₂ decreased (51%) at 4 h after TBHP exposure and returned on day 4 of the recovery period (Fig. 6A). Inhibition of PKC [using calphostin C or PKC(20–28)] did not affect decreases in ouabain-sensitive QO₂ at 4 h after TBHP injury but prevented the return of this function on day 4 of the recovery period (Fig. 6, A and B). In contrast, the return of ouabain-sensitive QO₂ in RPTCs treated with PMA before TBHP exposure occurred on day 2 and was accelerated in comparison with RPTCs exposed to TBHP only (Fig. 7A).

View larger version (41K): [in this window] [in a new window]   Fig. 7. Promotion of the recovery of active Na+ transport by PKC activation (PMA; 100 nM) in TBHP-injured (A) and DCVC-injured (B) RPTCs. Data are presented as means ± S.E., n = 3 separate RPTC cultures. Values with dissimilar superscripts on a given day are significantly different (P < 0.05) from each other.

DCVC exposure in RPTCs resulted in 40% decrease in ouabain-sensitive QO₂ at 4 h of the recovery period (Fig. 7B). Ouabain-sensitive QO₂ did not recover over time (Fig. 7B). However, treatment with PMA before DCVC exposure resulted in the return of ouabain-sensitive QO₂ in RPTCs within 2 days after the exposure (Fig. 7B).

These data show that PKC inhibition prevents the recovery of active Na⁺ transport after oxidant injury in RPTCs and that PKC activation promotes the repair of active Na⁺ transport after toxicant exposure.

Interestingly, inhibition of PKC by PKC(20–28) peptide decreased ouabain-sensitive QO₂ at 4 h, whereas calphostin C had no effect on this function in control RPTCs (Fig. 6). This could be explained by dissimilar sensitivity of different PKC isozymes to these two inhibitors. PKC(20–28) is more specific and general PKC inhibitor, whereas calphostin C is thought to inhibit primarily novel (Ca²⁺-independent and PMA-sensitive) isozymes of PKC.

PKC and the Recovery of Na⁺-Dependent Glucose Uptake. Na⁺-dependent glucose uptake was used as marker of the brush-border membrane function in RPTCs. TBHP and DCVC decreased Na⁺-dependent glucose uptake by 66 and 63%, respectively, at 4 h after the exposure (Fig. 8, A and B). Na⁺-dependent glucose uptake recovered after TBHP, but not after DCVC-induced injury (Fig. 8, A and B). Inhibition of PKC with calphostin C before TBHP exposure inhibited the return of Na⁺-dependent glucose uptake in regenerating RPTCs (Fig. 8A). Furthermore, 4-day treatment of RPTCs with calphostin C inhibited Na⁺-dependent glucose uptake by 89% (Fig. 8A).

View larger version (29K): [in this window] [in a new window]   Fig. 8. Decrease in the recovery of Na+-dependent glucose uptake by inhibition of PKC (calphostin c; 100 nM) in TBHP-injured RPTCs (A). Lack of an effect of PKC activation (PMA, 100 nM) on the recovery of Na+-dependent glucose uptake in TBHP- and DCVC-injured RPTCs (B). Data are presented as means ± S.E., n = 3 separate RPTC cultures. Values with dissimilar superscripts on a given day are significantly different (P < 0.05) from each other.

Activation of PKC using PMA did not accelerate the repair of Na⁺-dependent glucose uptake in TBHP-injured RPTCs (Fig. 8B). Likewise, treatment of RPTCs with PMA before DCVC injury did not promote the recovery of Na⁺-dependent glucose uptake (Fig. 8B).

	Discussion

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Toxicant-induced renal dysfunction is often caused by chemically produced sublethal injury to RPTCs, which decreases the physiological functions of renal proximal tubules without producing apparent cell death. It has been also proposed that some toxicants cause renal dysfunction not only by inducing injury but also by inhibiting RPTC regeneration (Counts et al., 1995). Our previous study showed that RPTCs in primary culture recover their functions after sublethal injury induced by an oxidant (TBHP) and that this repair is mediated by autocrine mechanisms (Nowak et al., 1998). In contrast, physiological functions do not recover in RPTCs after injury produced by the halocarbon nephrotoxicant DCVC (Nowak et al., 1999). This suggested that DCVC disrupts the autocrine mechanisms of RPTC repair. The repair of mitochondrial function and active Na⁺ transport but not Na⁺-dependent glucose transport in DCVC-injured RPTCs are promoted by epidermal growth factor, pharmacological concentrations of ascorbic acid, and collagen IV (Nowak et al., 1999, 2000; Nony et al., 2001). Therefore, our previous data suggested that different toxicants affect the mechanisms of RPTC repair in a differential manner. The present study examined the role of one of the signaling molecules, PKC, in the recovery of RPTC functions after toxicant-induced injury.

The transmission of intracellular signals is mediated by a network of interacting proteins that regulate a variety of cellular processes and functions. PKC, a family of serine/threonine protein kinases, is a critical element of this network (Dempsey et al., 2000). Toxicants have been implicated in the alterations in PKC activity. Our results show that TBHP injury in RPTCs is associated with decreases in total PKC activity. These data remain in contrast to early and transient activation of PKC in neuronal and endothelial cells in response to short-term exposure to an oxidant (O'Brian et al., 1988; Palumbo et al., 1992; Abe et al., 1998). In contrast, longer exposure to oxidative stress modifies the catalytic domain of PKC and inactivates PKC (Gopalakrishna and Anderson, 1987), which is consistent with our results in RPTCs. DCVC-induced decreases in total PKC activity in our model are in agreement with the down-regulation of renal PKC after an exposure to DCVC in vivo and with the inhibition of PKC activity in the kidney by exposure to folic acid in vivo (Dong et al., 1993).

PKC has also been implicated in regeneration of different tissues and organs. Liver regeneration after partial hepatectomy and carbon tetrachloride injury is associated with the activation of PKC, PKC, PKC, and PKC (Daller et al., 1994; Tessitore et al., 1995). In the kidney, the activations of PKC, PKC, and PKC, but not PKC or PKC occur during compensatory renal hypertrophy induced by unilateral nephrectomy (Dong et al., 1993). PKC activation also plays a role in the acceleration of wound healing after mechanical injury in renal tubular epithelial cells (Sponsel et al., 1995). In our RPTC model, decreased PKC activity after TBHP- and DCVC-induced injury was accompanied by major decreases in the mitochondrial function, active Na⁺ transport, and Na⁺-dependent glucose uptake. The repair of RPTC functions after TBHP-induced injury was preceded by the recovery of total PKC activity. On the other hand, the lack of repair of RPTC functions in DCVC-injured RPTCs was associated with the lack of return of the PKC activity. Furthermore, inhibition of PKC prevents the return of mitochondrial function, active Na⁺ transport and Na⁺-dependent glucose uptake in TBHP-injured RPTCs, which suggests that PKC plays a role in the repair of RPTC functions after the oxidant injury.

Two dissimilar PKC inhibitors have been used in this study, the chemical inhibitor calphostin C and a short peptide derived from PKC and PKC pseudosubstrate sequence. PKC(20–28) inhibitor decreased, whereas calphostin C had no effect on mitochondrial function and active Na⁺ transport in control RPTCs, which could be explained by higher specificity of PKC(20–28) toward PKC and PKC. The fact that calphostin C, which is rather an inhibitor of novel PKC isozymes (, , , µ, and ), did not have any effect on these functions in control RPTCs suggests that the maintenance of these functions is dependent on classical PKC isozymes.

The lack of functional repair in RPTCs treated with PKC inhibitors was not due to toxic effects produced by these compounds. These inhibitors did not potentiate decreases in mitochondrial function and active Na⁺ transport in TBHP-injured RPTCs. Furthermore, both inhibitors produced a similar degree of inhibition of the repair of mitochondrial function and active Na⁺ transport. Interestingly, a 4-day treatment of control RPTCs with calphostin C decreased Na⁺-dependent glucose uptake in RPTCs by 89%, which suggests that functional PKC is necessary to maintain Na⁺-dependent glucose transport in control RPTCs.

Because our data suggested that functional PKC is a critical for the recovery of RPTC functions after a toxic insult, we hypothesized that activation of PKC before toxicant exposure may prevent the decreases in RPTC functions or promote the repair of some or all RPTC functions. Activation of PKC is associated with redistribution from the cytosol to cellular membranes and organelles. This event is necessary for phosphorylation of various intracellular proteins targeted by PKC. Previous studies suggested that renal regeneration after toxicant exposure is associated with differential modulation of PKC isozymes, which may be associated with different functions of these isozymes in the process of RPTC regeneration (Dong et al., 1993). At the present, however, the role of individual PKC isozymes in the repair of these functions is unknown. In our RPTC model, among four major PKC isozymes present in RPTCs (PKC, PKC, PKC, and PKC), only PKC and PKC were phosphorylated and redistributed from the cytosol to the particulate fraction in response to PMA treatment. In contrast, PMA had no effect on the activation of ERK1/2 (data not shown). This suggested that only PKC and PKC were activated by PMA. PMA pretreatment had no effect on the decrease in mitochondrial function induced by TBHP or DCVC, which demonstrates that PKC activation does not protect against mitochondrial dysfunction caused by these nephrotoxicants. However, the recovery of mitochondrial function was accelerated in TBHP-injured RPTCs pretreated with PMA. Furthermore, PKC activation abolished inhibition of the repair of mitochondrial function in DCVC-injured RPTCs. These results show that PKC plays an important role in the repair of mitochondrial function and suggest that PKC and/or PKC are involved in these processes.

Likewise, our data show that the repair of active Na⁺ transport after toxicant injury is controlled by PKC. In contrast to the lack of repair of active Na⁺ transport in DCVC-injured RPTCs, the recovery of this function occurred in DCVC-injured RPTCs pretreated with PMA. Moreover, PKC activation accelerated the repair of active Na⁺ transport after TBHP injury, which was not due to the protection against TBHP- and DCVC-induced decreases in this function because the degree of injury was equivalent in RPTCs treated with toxicants and RPTCs treated with both PMA and toxicants.

Active Na⁺-transport in RPTCs is mediated through Na⁺/K⁺-ATPase, which is regulated through phosphorylation by PKC and PKA (Bertorello and Aperia, 1989; Feschenko and Sweadner, 1994; Chibalin et al., 1997). Phosphorylation of Na⁺/K⁺-ATPase leads to internalization of the pump into endosomes and decreased expression on the basolateral membrane (Chibalin et al., 1997). Our data suggest that this is also true in our model as the short-term exposure to PMA (which results in PKC activation) decreased ouabain-sensitive QO₂ in control RPTCs. In contrast, long-term exposure to PMA (24 h), which does not activate PKC (Fig. 5A), had no effect on ouabain-sensitive QO₂. PKC activation seems to be necessary for promoting the recovery of active Na⁺ transport after toxicant exposure. Previously, it has been proposed that the endosomes may constitute reservoirs during Na⁺/K⁺ pump synthesis and degradation (Chibalin et al., 1997). It is likely that Na⁺/K⁺-ATPase can be either protected or repaired with higher efficiency in the endosomes and is, therefore, available for reassembly on the basolateral membrane sooner during the recovery process. It is also likely that the accelerated return of active Na⁺ transport by PKC activation is due to the promotion of mitochondrial function and ATP availability. Na⁺/K⁺-ATPase activity requires constant supply of ATP. The lack of recovery of active Na⁺ transport in DCVC-treated RPTCs is associated with decreased ATP levels (Nowak et al., 1999). Promotion of the recovery of mitochondrial function in DCVC-injured RPTCs treated with PMA may support the recovery of active Na⁺ transport by supplying ATP for Na⁺/K⁺-ATPase function.

In contrast, activation of PKC did not promote the repair of Na⁺-dependent glucose uptake in DCVC-injured RPTCs. These results suggest that the disruption of Na⁺ homeostasis after DCVC injury is not the exclusive mechanism responsible for the inhibition of repair of Na⁺-dependent glucose uptake. Furthermore, our present data suggest that the recovery of this function also involves PKC-independent mechanisms.

In conclusion, our results show that toxicant-induced injury in RPTC causes decreases in mitochondrial function, active Na⁺ transport and Na⁺-dependent glucose uptake and that these events are associated with decreases in PKC activity. PKC inhibition blocks whereas PKC activation promotes the repair of mitochondrial function and active Na⁺ transport. In contrast, activation of PKC does not stimulate the recovery of Na⁺-dependent glucose uptake. Thus, our data show that PKC mediates the repair of mitochondrial function and active Na⁺ transport following toxicant injury and suggest that PKC and/or PKC are involved in this regulation.

	Acknowledgements

 
We thank Diana Bakajsova (University of Arkansas for Medical Sciences) for help with the measurements of oxygen consumption and Dr. Thomas W. Petry (Upjohn Pharmacia, Kalamazoo, MI) for the generous gift of S-(1,2-dichlorovinyl)-L-cysteine.

	Footnotes

 
This work was supported by National Institutes of Health National Institute of Diabetes and Digestive and Kidney Diseases Grant R01 DK59558 and a Pilot Study Grant from Intramural Grant Programs at the University of Arkansas for Medical Sciences.

DOI: 10.1124/jpet.103.050336.

ABBREVIATIONS: RPTC, renal proximal tubular cell; TBHP, tert-butylhydroperoxide; ARF, acute renal failure; DCVC, S-(1,2-dichlorovinyl)-L-cysteine; PKC, protein kinase C; PMA, phorbol-12-myristate-13-acetate; MBP, myelin basic protein; QO₂, oxygen consumption.

Address correspondence to: Dr. Grayna Nowak, Department of Pharmaceutical Sciences, University of Arkansas for Medical Sciences, 4301 West Markham St., MS 522-3, Little Rock, AR 72205. E-mail: gnowak{at}uams.edu

	References

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