INTRODUCTION

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AFTER ISCHEMIA- or toxicant-induced injury, the kidney has the potential for complete recovery from the acute loss of function. Renal proximal tubular cells (RPTC) are the predominant cell type injured by ischemia and toxicants, and the return of RPTC functions is critical for the restoration of normal renal function (10). The return of proximal tubular functions depends on the ability of sublethally injured RPTC to repair or to proliferate, migrate, differentiate and reline the damaged epithelium (1, 8, 29, 37). Unfortunately, the mechanisms that regulate RPTC repair and proliferation are poorly understood.

Ischemia/reperfusion and toxicants are thought to contribute to renal dysfunction through their ability to produce sublethal injury in RPTC, that is, RPTC physiological functions are diminished following the insult, but cell death does not occur (37). Under this condition, the RPTC must undergo cell repair, such that normal physiological functions return. For example, following in vivo toxicant or ischemia/reperfusion injury, loss and/or internalization of the brush-border membrane (BBM) microvilli and ATP depletion occur, resulting in the inhibition of proximal tubular reabsorption of sodium and glucose (2, 16, 25, 27, 36, 38, 39). Over time, these pathological changes are repaired, and normal physiological function is observed (9, 26, 36, 39).

The renal dysfunction associated with ischemia/reperfusion injury has been postulated to occur through oxidative stress (5). In addition, oxidative stress has been implicated in the renal toxicity of a number of drugs and chemicals, including halocarbons, platinum compounds, cyclosporine, and mercuric chloride (11, 13, 14, 19, 22, 28, 40). t-Butylhydroperoxide (TBHP) is used as a model compound to induce oxidant injury, and exposure of RPTC to TBHP results in the formation of reactive oxygen species, lipid peroxidation, decreases in mitochondrial and lysosomal membrane potential, mitochondrial dysfunction, and impaired basolateral membrane (BLM) transport functions (4, 6, 19, 28, 35).

Examination of the mechanisms regulating RPTC repair following injury requires an appropriate in vitro model that allows for morphological regeneration of the monolayer and recovery of physiological functions in sublethally injured RPTC. Recently, we have demonstrated that oxidative metabolism (oxygen consumption and utilization of oxidative substrates), active Na⁺ transport, and Na⁺-dependent glucose transport are upregulated in primary cultures of RPTC grown in improved conditions in the presence of L-ascorbic acid-2-phosphate (31). RPTC grown in these conditions undergo complete morphological regeneration following TBHP exposure, and this process is due to both proliferation and migration/spreading (32). However, the effects of oxidants on RPTC physiological functions following sublethal injury and the ability of RPTC to restore physiological functions have not been examined. Furthermore, the temporal relationships between morphological regeneration of the monolayer and the recovery of RPTC functions following injury are not known. Therefore, the aim of this study was 1) to determine the effect of an oxidant on mitochondrial function, substrate utilization, and BLM and BBM functions in sublethally injured RPTC; 2) to examine the recovery of mitochondrial and transport functions in RPTC following sublethal oxidant injury; and 3) to determine the temporal relationships among morphological regeneration of the monolayer, return of mitochondrial function, and the recovery of transport capabilities following sublethal oxidant injury.

	MATERIALS AND METHODS

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Materials. Female New Zealand White rabbits (1.5-2.0 kg) were purchased from Myrtle's Rabbitry (Thompson Station, TN). L-Ascorbic acid-2-phosphate magnesium salt and TBHP were obtained from Wako BioProducts (Richmond, VA) and Sigma Chemical (St. Louis, MO), respectively. Methyl--D-glucopyranoside (MGP), U-[¹⁴C]glucose (sp act, 282 mCi/mmol), and [³H]thymidine (sp act, 25 Ci/mmol) were purchased from DuPont-NEN (Boston, MA). Amino acids standards and o-phthaldehyde-3-mercaptopropionic acid (OPA) were obtained from Hewlett Packard (Palo Alto, CA). Sodium acetate, trifluoracetic acid, triethylamine, methanol, and acetonitrile were purchased from J. T. Baker (Philipsburg, NJ). Tetrahydrofuran was supplied by Bioanalytical Systems (West Lafayette, IN). The source of the other reagents has been described previously (30, 31).

Isolation of proximal tubules and culture conditions. Rabbit renal proximal tubules were isolated and grown in improved conditions, as described previously (30, 31). The culture medium was a 50:50 mixture of Dulbecco's modified Eagle's medium (DMEM) and Ham's F-12 nutrient mix without phenol red, pyruvate, and glucose, supplemented with 15 mM NaHCO₃, 15 mM N-2-hydroxyethylpiperazine-N'-2-ethanesulfonic acid, 0.44 mM L-alanine, 0.44 mM glycine, 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 (0.05 mM) were added to the medium immediately before daily media change.

Toxicant treatment of RPTC monolayer. RPTC cultures reached confluence within 4-5 days and were treated with TBHP on the 6th day. RPTC were treated with 0.2 mM TBHP or the vehicle (dimethyl sulfoxide, 0.3% final concentration) for 50-60 min to obtain ~25% cell death and loss. After toxicant or vehicle exposure, the remaining cellular monolayer was washed with fresh culture medium. Samples of control and TBHP-treated RPTC were taken at various time points after exposure for measurements of cellular functions.

[³H] thymidine incorporation. [³H]thymidine incorporation by RPTC was measured starting at 4 h and at 1, 4, and 6 days following TBHP treatment. Control and TBHP-treated RPTC were cultured for 4 h in the presence of 2 µCi of [³H]thymidine. After the incubation, the culture media were aspirated, monolayers were washed twice with ice-cold phosphate-buffered saline (PBS) and suspended in PBS, and proteins were precipitated with perchloric acid (3% final concentration). The precipitates were washed with 3% perchloric acid and solubilized in 0.1 M tris(hydroxymethyl)aminomethane hydrochloride buffer (pH 7.5) containing 0.15 M NaCl and 0.05% (vol/vol) Triton X-100. The amount of [³H]thymidine incorporated was determined by liquid scintillation spectrometry and normalized to cellular protein.

Oxygen consumption. RPTC monolayers were washed with 37°C culture medium and gently detached from the dishes with a rubber policeman, suspended in 37°C culture medium, and transferred to the oxygen consumption (QO₂) measurement chamber. QO₂ was measured polarographically using a Clark-type electrode, as described previously (30, 31). For measurements of ouabain-sensitive and uncoupled QO₂, 0.1 mM ouabain and 1 µM carbonyl cyanide p-trifluormethoxyphenylhydrazone (final concentrations) were used, respectively.

ATP content. After medium aspiration, RPTC monolayers were washed twice with ice-cold PBS, suspended in 0.5 ml PBS, precipitated with 0.5 ml of 6% perchloric acid, and centrifuged at 10,000 g for 1 min. Supernatants were neutralized to pH 7.0 and stored at 70°C until analyzed. Intracellular ATP contents were measured using reverse-phase high-performance liquid chromatography (HPLC), as described previously (12).

Net lactate and amino acid consumption. Net lactate and amino acid consumption were measured 24 h after media change. Lactate content in the culture medium was determined spectrophotometrically using Sigma kit no. 826-UV, as described previously (20). Amino acid contents in the culture medium were determined by HPLC using Hewlett Packard 1090L series II liquid chromatograph diode-array system (338 nm), equipped with a TosoHaus TSK-GEL super ODS column and a Hewlett Packard 1046A fluorescence detector (340 nm excitation/450 nm emission). Mobile phase A consisted of 20 mM sodium acetate buffer containing 0.018% triethylamine and 0.3% tetrahydrofuran (pH 7.2). Mobile phase B consisted of 100 mM sodium acetate buffer, pH 7.2, 40% acetonitrile and 40% methanol. The initial column conditions consisted of 95% mobile phase A and 5% mobile phase B. After sample injection, a linear gradient was started that reached 30% mobile phase A/70% mobile phase B at 10 min, maintained at this rate for 0.1 min, and immediately changed to 100% mobile phase B for 1.9 min. OPA derivatives were prepared by an automated pre-column sample preparation system. The sensitivity of the method was 10-20 pmol/µl.

Na⁺-coupled glucose uptake. Na⁺-coupled glucose uptake was assessed using the nonmetabolizable glucose analogue, MGP, as described previously (31). MGP uptake was corrected for Na⁺-independent (phloridzin-insensitive) and zero time uptakes.

Enzyme assays. Na⁺-K⁺-adenosinetriphosphatase (Na⁺-K⁺-ATPase) activity was determined in cellular lysates by measuring the difference between total ATPase activity and ouabain-insensitive ATPase activity, as described previously (31). -Glutamyltranspeptidase (GGT) activity (BBM marker) was determined in cell lysates, according to Meister et al. (23). Protein was measured by the method of Lowry et al. (21). DNA content was determined by the method of Labarca and Paigen (20), as described previously (30).

Statistical analysis. Data are presented as means ± SE and were analyzed for significance using two-way analysis of variance or Student's t-test for paired data, where appropriate. 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 three plates.

	RESULTS

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Monolayer regeneration. Treatment of confluent RPTC monolayers with 0.2 mM TBHP for 50-60 min resulted in a 24% RPTC loss at 4 h following the exposure (19.3 ± 3.0 vs. 25.4 ± 2.7 µg DNA/plate) and did not result in additional loss at 24 h (19.3 ± 3.0 vs. 18.7 ± 2.3 µg/plate at 4 and 24 h following TBHP exposure, respectively). DNA content in TBHP-treated monolayers returned to control values on day 4 following the treatment (Fig. 1A) and was accompanied by total recovery of monolayer confluence (data not shown). [³H]thymidine incorporation into DNA of sublethally injured RPTC was decreased by 83% at 4 h, increased on day 1, and, on day 4 after TBHP exposure, was 1.5-fold higher than controls (Fig. 1B). These changes in [³H]thymidine incorporation may reflect both DNA repair following oxidant damage and new DNA synthesis associated with proliferation. Protein-to-DNA ratios were not affected by TBHP treatment (73 ± 14 vs. 66 ± 11 in TBHP-treated and control RPTC, respectively, 4 h following exposure) or during regeneration (57 ± 9 vs. 64 ± 9 in TBHP-treated and control RPTC, respectively, 4 days following exposure), which demonstrates that regeneration of RPTC monolayers was not due to cellular hypertrophy. These data show that sublethally injured RPTC regenerate by proliferation and migration/spreading following oxidant exposure.

View larger version (16K): [in this window] [in a new window]   Fig. 1.   Recovery of monolayer DNA content (A) and [3H]thymidine incorporation (B) during renal proximal tubular cells (RPTC) regeneration following t-butylhydroperoxide (TBHP, ) (0.2 mM) exposure. Data are presented as means ± SE, n = 4-10 separate experiments; , control. * P < 0.05, significantly different from controls.

Mitochondrial function. Basal and uncoupled QO₂ and ATP content were used as markers of RPTC mitochondrial function. TBHP treatment resulted in 64 and 55% decreases in basal QO₂ in sublethally injured RPTC at 4 and 24 h, respectively, following the exposure (Fig. 2A). Basal QO₂ in TBHP-treated RPTC returned to control levels on day 6 following the exposure (Fig. 2A). Uncoupled QO₂ was used as a marker of electron transport chain integrity. Uncoupled QO₂ in sublethally injured RPTC was reduced by 71 and 61% at 4 and 24 h, respectively, following TBHP treatment and returned to control levels on day 4 after the exposure (Fig. 2B). Intracellular ATP content in sublethally injured RPTC decreased 63 and 54% at 4 and 24 h, respectively, following TBHP treatment (Fig. 3) and returned to control values on day 6 following exposure. These data show that TBHP treatment decreases mitochondrial function in sublethally injured RPTC and that mitochondrial function recovers over time following oxidant treatment. These results also demonstrate that the repair of the electron transport chain precedes the recovery of basal QO₂ and ATP content.

View larger version (16K): [in this window] [in a new window]   Fig. 2.   Recovery of basal oxygen consumption (QO2) (A) and uncoupled QO2 (B) during RPTC regeneration following TBHP (0.2 mM) exposure. Data are presented as means ± SE, n = 6-10. * P < 0.05, significantly different from controls. Symbols are as in Fig. 1.

View larger version (15K): [in this window] [in a new window]   Fig. 3.   Recovery of intracellular ATP content during RPTC regeneration following TBHP (0.2 mM) exposure. Data are presented as means ± SE, n = 4. * P < 0.05, significantly different from controls.

Metabolic substrate consumption. Lactate is the primary oxidative substrate in control RPTC (41). Net lactate consumption in sublethally injured RPTC decreased 71 and 46% at 4 and 24 h, respectively, following TBHP treatment and returned to control levels on day 4 after the exposure (Fig. 4A). In contrast, net consumption of glutamine (another oxidative substrate for cultured RPTC) increased 2.7-fold at 4 h following TBHP exposure (Fig. 4B), remained increased during the first 4 days of the recovery period, and returned to control values on day 6 (Fig. 4B). Under control conditions, there was net consumption of arginine, serine, and asparagine (0.42 ± 0.07, 0.15 ± 0.04, and 0.03 ± 0.01 µmol · day¹ · mg protein¹, respectively) and net production of glutamic acid, alanine, and glycine (2.87 ± 0.54, 1.53 ± 0.13, and 0.59 ± 0.14 µmol · day¹ · mg protein¹, respectively) in RPTC. No changes in the net consumption or production of these amino acids were observed following TBHP-induced injury (data not shown). These results show that lactate consumption is decreased in sublethally injured RPTC, which is consistent with the decrease in mitochodrial function. The decrease in mitochondrial function and lactate consumption is associated with an increase in glutamine consumption.

View larger version (15K): [in this window] [in a new window]   Fig. 4.   Recovery of net lactate (A) and net glutamine (B) consumptions during RPTC regeneration following TBHP (0.2 mM) exposure. Data are presented as means ± SE, n = 4-9. * P < 0.05, significantly different from controls.

Active Na⁺ transport. Ouabain-sensitive QO₂ and Na⁺-K⁺-ATPase activity were used to assess active Na⁺ transport in RPTC. Ouabain-sensitive QO₂ in sublethally injured RPTC was inhibited by 77 and 66% at 4 and 24 h, respectively, following TBHP treatment (Fig. 5A). Ouabain-sensitive QO₂ gradually increased during the recovery period and returned to control values on day 6 after the exposure (Fig. 5A). TBHP treatment resulted in an 88% inhibition of Na⁺-K⁺-ATPase activity in sublethally injured RPTC following a 4 h exposure (Fig. 5B). Na⁺-K⁺-ATPase activity increased during the recovery period and reached control values on day 6 after the treatment (Fig. 5B). These data demonstrate that TBHP treatment inhibits Na⁺-K⁺-ATPase activity and active Na⁺ transport in RPTC and that these functions recover during the repair of sublethally injured RPTC. The repair of active Na⁺ transport follows morphological regeneration of the RPTC monolayer, recovery of mitochondrial electron transport chain function, and lactate consumption and is associated with the recovery of ATP concentrations and basal QO₂.

View larger version (16K): [in this window] [in a new window]   Fig. 5.   Recovery of ouabain-sensitive QO2 (A) and Na+-K+-ATPase activity (B) during RPTC regeneration following TBHP (0.2 mM) exposure. Data are presented as means ± SE, n = 4-10. * P < 0.05, significantly different from controls.

Brush-border membrane function. Na⁺-dependent glucose uptake was used as a marker of BBM function. TBHP exposure resulted in 83 and 80% decreases in Na⁺-dependent glucose uptake in sublethally injured RPTC at 4 and 24 h, respectively, following the treatment. Na⁺-dependent glucose uptake gradually recovered during the repair period and returned to control levels on day 6 following TBHP exposure (Fig. 6). TBHP treatment had no effect on GGT activity in sublethally injured RPTC (614 ± 79 vs. 515 ± 38 mU/mg protein in TBHP-treated and control RPTC at 4 h following the exposure). These data show that Na⁺-dependent glucose uptake is inhibited following RPTC exposure and recovers following TBHP-induced injury with the recovery of active Na⁺ transport. In contrast, GGT activity remained unchanged following TBHP exposure and during the repair process. Thus GGT activity is not a target of TBHP.

View larger version (16K): [in this window] [in a new window]   Fig. 6.   Recovery of Na+-dependent glucose uptake during RPTC regeneration following TBHP (0.2 mM) exposure. Data are presented as means ± SE, n = 6-8. * P < 0.05, significantly different from controls.

	DISCUSSION

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Although it is clear that the kidney can recover following ischemia/reperfusion and toxicant-induced injury, the mechanisms regulating these processes have not been resolved. Numerous investigators have shown RPTC injury, death, and detachment after ischemia or toxicant exposure, followed by proliferation and the return of normal RPTC morphology and function (1, 4, 5, 8, 11, 19, 35, 37). Under these circumstances, it is thought that RPTC adjacent to the denuded areas dedifferentiate, proliferate, migrate, differentiate, and reline the damaged nephron (1, 37). Often, however, renal failure is associated with cellular dysfunction without extensive necrosis of the epithelial cells (1, 37). Therefore, following an ischemic or toxicant/oxidant exposure, a number of RPTC may be sublethally injured and exhibit diminished physiological functions in the absence of cell death. Under this condition, the RPTC must undergo cell repair with the return of normal physiological functions.

A previous study from this laboratory (32) showed that RPTC monolayers regenerate following TBHP exposure through migration/spreading and proliferation. In the present study, we have specifically examined sublethal cell injury and repair of physiological functions, using an improved in vitro RPTC model that more closely expresses normal oxidative metabolism and transport functions (31) and the model oxidant TBHP. RPTC were treated with the TBHP such that 24% cell death and loss occurred 4 h after oxidant exposure, and the remaining RPTC were sublethally injured. Our data show that sublethally injured RPTC regenerate through proliferation and migration/spreading. The increase in [³H]thymidine incorporation into DNA between 4 h and 4 days following the treatment may reflect DNA repair and/or increased proliferation of sublethally injured RPTC following oxidant damage. The return of monolayer confluence and DNA content on day 4 following exposure suggests that both DNA repair and synthesis occurred after oxidant injury. Inducement of greater than 50% cell death and loss is followed by an increase in monolayer confluence but not by complete repair of physiological functions (data not shown).

The data illustrate that RPTC mitochondrial function, Na⁺-K⁺-ATPase activity, active Na⁺ transport, and Na⁺-dependent glucose transport are major targets of TBHP with extensive inhibition (64, 88, 77, and 83%, respectively) of these functions 4 h after exposure. The inhibition of RPTC oxidative metabolism (decreased basal and uncoupled QO₂ and intracellular ATP content) is in agreement with a previous study showing that TBHP decreases mitochondrial function in freshly isolated rabbit renal proximal tubules (35). This study demonstrates that the decrease in the respiration rate produced by TBHP results from disruption of mitochondrial electron transport chain integrity.

TBHP exposure had differential effects on metabolic substrate consumption in RPTC. The inhibition of net lactate consumption, a major oxidative substrate for renal proximal tubules (41), was accompanied by an increase in net consumption of glutamine. It is not known whether these changes were due to a decrease in lactate uptake, a shift of RPTC metabolism from lactate to glutamine oxidation, and/or increased demands for glutamine as a precursor for protein or purine and pyrimidine synthesis. The decrease in net lactate consumption may reflect impaired BBM uptake of lactate through the Na⁺-lactate transporter due to loss of the Na⁺ gradient, inhibition of the lactate transporter, or impaired utilization of lactate as a metabolic substrate. Because lactate utilization returned to control values (4 days) prior to the return of active Na⁺ transport and Na⁺-dependent glucose transport (6 days), the decrease in lactate consumption may be due, in part, to the inhibition of the lactate transporter itself.

The increased utilization of glutamine after TBHP exposure probably reflects a complex mixture of events. The generation of free radicals from TBHP by the mitochondrial electron transport chain may selectively impair certain stages of oxidative phosphorylation (18), leading to a switch in metabolic substrate utilization from lactate to glutamine during cell injury and repair. The protective effect of glutamine, a precursor of the tricarboxylic acid cycle intermediate -keto-glutarate, on preserving intracellular ATP content during oxidant (H₂O₂)-induced damage to endothelial cells when glucose-dependent pathways of ATP synthesis have been inhibited (15) would support this hypothesis. The need for glutamine may be selective, since oxidative injury dramatically reduces cellular glutamine synthetase activity (33). Enhanced utilization of glutamine may also be associated with the regenerative stage of the cell repair process. Glutamine is an important amino acid for proliferation and metabolism of growing cells, since it is an important intermediate in purine and pyrimidine synthesis (34). This may explain high glutamine consumption between days 1 and 4, when monolayer DNA content increased and returned to control levels.

The inhibitory effect of TBHP on active Na⁺ transport in RPTC was primarily due to the inhibition of Na⁺-K⁺-ATPase activity and secondarily to the decrease in intracellular ATP content. The inhibition of the Na⁺-K⁺-ATPase by TBHP is consistent with previous studies that have demonstrated oxidant-induced inactivation of the Na⁺-K⁺-ATPase in renal cells (3, 17).

Associated with the reduction in active Na⁺ transport following TBHP exposure, was a decrease in BBM Na⁺-dependent glucose uptake. In contrast to the inhibition of glucose transport by TBHP, BBM GGT activity was not altered by TBHP exposure or during regeneration. Oxidant-induced reduction in Na⁺-dependent glucose uptake may be due to the decrease in the Na⁺ gradient or alterations in the Na⁺-dependent glucose carrier. A decrease in the Na⁺ gradient is not always associated with the inhibition of Na⁺-dependent transport processes in renal cells. For example, Molitoris and Kinne (27) showed that ischemia-induced membrane dysfunction results in decreased Na⁺-dependent glucose uptake but not Na⁺-dependent alanine uptake. The decrease in Na⁺-dependent glucose uptake in renal cortical cells following ischemia/reperfusion injury was suggested to be due to a reduced number of functional carrier units (27). It remains to be determined whether the decrease in Na⁺-dependent glucose uptake produced by TBHP is due to a decrease in the Na⁺ gradient, a direct effect on the glucose carrier, or both. However, since the return of Na⁺-dependent glucose uptake paralleled the recovery of Na⁺-K⁺-ATPase activity and active Na⁺ transport, the recovery Na⁺-dependent glucose uptake may depend, in part, on the return of active Na⁺ transport.

The present study demonstrates that RPTC grown under conditions that promote in vivo physiological functions have the potential for complete regeneration of the monolayer and restoration of oxidative metabolism and transport functions following TBHP exposure. However, the recovery occurred in a differential manner. Proliferation and migration/spreading and the recovery of the monolayer confluence were completed 4 days following TBHP exposure. In parallel, mitochondrial electron transport chain integrity and net lactate consumption were restored. Subsequently, intracellular ATP content, active Na⁺ transport, Na⁺-dependent glucose uptake, and net glutamine consumption returned to control levels. These data suggest that proliferation, monolayer regeneration, and the repair of mitochondrial function precede the return of transport functions.

Our data are in agreement with the observations of Spiegel and co-workers (36). After ischemia/reperfusion injury in vivo, morphological regeneration of renal proximal tubules occurred by day 3, whereas the return of Na⁺ and glucose reabsorption occurred on day 8. It is generally accepted that specialized functions of RPTC are dependent on maintenance of RPTC polarity and that ischemia- and toxicant-induced injury result in the loss of cell polarity (24-26). Reestablishment of epithelial polarity and basolateral localization of the Na⁺-K⁺-ATPase following an injury are necessary for the recovery of Na⁺ and glucose reabsorption, and RPTC with normal cellular morphology are unable to function in this manner until repolarization of the cell membrane occurs (16, 24). Loss of RPTC polarity following TBHP treatment may result from both oxidant damage to the cell and/or a decrease in cell number caused by cell death and loss. It is unlikely that a decrease in cell number resulted in the loss of RPTC polarity, since mechanically induced injury resulting in the removal of ~24% of the RPTC monolayer (7) did not alter QO₂, active Na⁺ transport, and Na⁺-dependent glucose transport (data not shown).

In conclusion, this study shows that 1) mitochondrial function, active Na⁺ transport, and Na⁺-dependent glucose uptake are major targets of TBHP in RPTC; 2) sublethally injured RPTC have the potential to repair physiological functions following oxidant exposure; 3) monolayer regeneration following oxidant-induced sublethal injury is due to proliferation and migration/spreading, is associated temporally with the return of mitochondrial functions, and precedes the repair of active Na⁺ transport and Na⁺-dependent glucose uptake; and 4) sublethal injury and subsequent regeneration of RPTC are associated with alterations in metabolic substrate requirements. Our results also suggest that oxidant-induced sublethal injury to RPTC in vivo may contribute to renal dysfunction.