Department of Veterinary Physiology, College of Veterinary Medicine, Chonnam National University, Gwangju, Korea 500-757
Submitted 2 February 2004 ; accepted in final form 2 June 2004
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ABSTRACT |
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kidney; mitogen-activated protein kinase; phospholipase A2
A convenient way to evaluate the effect of oxalate on renal PTCs is by means of in vitro studies with differentiated cell cultures. The primary rabbit renal PTC culture system that was utilized in this study is well recognized to retain in vitro the differentiated phenotype typical of the renal proximal tubule, including a polarized morphology (38), apical membrane proteins (leucine aminopeptidase and -glutamyltranspeptidase), distinctive proximal tubule transport systems including the Na+-glucose cotransport system (21), as well as hormone responses (20). Therefore, PTCs in hormonally defined, serum-free culture conditions would be a powerful tool for studying the effect of oxalate on cell proliferation (31). Thus the present study was performed to investigate the effect of oxalate on renal PTC proliferation and to identify specific intracellular signaling pathways that are targeted by oxalate. We demonstrate here for the first time that oxalate inhibits 3H-labeled thymidine incorporation through increase of AA release via oxidative stress, p38 MAPK/c-Jun NH2-terminal kinase (JNK), and cPLA2 activation in primary cultured renal PTCs.
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MATERIALS AND METHODS |
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Isolation of rabbit renal proximal tubules and culture conditions. All procedures were performed in accordance with the "Guiding Principles for Research Involving Animals and Human Beings" of the Korean Association for Laboratory Animal Science. Male New Zealand White rabbits (1.52.0 kg) were used for this experiment. Primary rabbit kidney PTC cultures were prepared by a modification of the method of Chung et al. (12). The PTCs were grown in 2 ml of a 1:1 mixture of Dulbecco's modified Eagle's medium (DMEM) and F-12 with 15 mM HEPES buffer (pH 7.4) and 20 mM sodium bicarbonate (pH 7.4). Immediately before use of the medium, three growth supplements (5 µg/ml insulin, 5 µg/ml transferrin, and 5 x 108 M hydrocortisone) were added. Kidneys were perfused via the renal artery, first with phosphate-buffered saline (PBS), and subsequently with DMEM-F-12 containing 0.5% iron oxide (wt/vol) until the kidney turned gray-black in color. Renal cortical slices were prepared by cutting the renal cortex and then homogenized with four strokes of a sterile glass homogenizer. The homogenate was poured through first a 253-µm and then a 83-µm mesh filter. Tubules and glomeruli on top of the 83-µm filter were transferred into sterile DMEM-F-12 containing a magnetic stirring bar. Glomeruli (containing iron oxide) were removed with a magnetic stirring bar. The remaining proximal tubules were incubated briefly in DMEM-F-12 containing 60 µg/ml collagenase (class IV) and 0.025% soybean trypsin inhibitor. The dissociated tubules were then washed by centrifugation, resuspended in DMEM-F-12 containing the three growth supplements, and transferred into tissue culture dishes. PTCs were maintained at 37°C in a 5% CO2 humidified environment in DMEM-F-12 medium containing the three supplements. The medium was changed 1 day after plating and every 3 days thereafter.
Experimental protocol. PTCs were exposed to oxalate and other reagents by exchanging the medium for hormonally defined, serum-free DMEM-F-12 containing three growth supplements and the agents of interest, and the incubation continued for the indicated period at 37°C under an atmosphere of 5% CO2-95% air. Where indicated, sodium oxalate was added at a concentration of 0.1, 0.5, 1, 2, or 4 mM (total), which provided a free oxalate level of 38, 190, 380, 760, or 1,520 µM, respectively, and corresponded to a relative supersaturation level (RSS) for calcium oxalate of 24.9. Estimates of free oxalate and RSS were obtained with the EQUIL program (43). Incubation of cell-free tissue culture media with 1 mM oxalate did not result in the formation of calcium oxalate crystals. This concentration was taken as the metastable limit for the conditions described here. In the studies testing the effect of inhibitors, the PTCs were preincubated for 30 min before oxalate addition.
[3H]thymidine incorporation. When the cells were 7080% confluent, a final media change was done. Thymidine incorporation experiments were conducted as described by Brett et al. (8). Cells were incubated in medium in the absence or presence of oxalate for 24 h and were pulsed with 1 µCi of [methyl-3H]thymidine for 24 h at 37°C. The cells were then washed twice with PBS and were fixed in 10% trichloroacetic acid (TCA) at room temperature for 15 min and then washed twice in 5% TCA. The acid-insoluble material was dissolved in 2 N NaOH at room temperature and counted for radioactivity by liquid scintillation counter (LS 6500; Beckman Instruments, Fullerton, CA). All experiments were performed in triplicate. Values were converted from absolute counts to the percentage of control to allow for comparison between experiments.
Trypan blue exclusion assay. Cells were grown to confluence in 35-mm dishes as described above. Monolayers were washed twice with PBS. The cells were detached from the culture dishes with a 0.05% trypsin-0.5 mM EDTA solution, and proteolytic action was then inhibited by soybean trypsin inhibitor (0.05 mg/ml). Trypan blue solution (0.4% wt/vol, 500 µl) was then added to the cell suspension and the cells were counted, keeping a separate count of blue cells with a hemocytometer under light microscopy. Cells failing to exclude the dye were considered nonviable; the data are expressed as percentage of viable cells.
LDH release. Cell injury was assessed by LDH activity. The level of LDH activity in the medium was measured with a LDH assay kit. For measurement of LDH activity, PTCs were treated with different concentrations of oxalate for 24 h. LDH activity was expressed as percentage of control.
H2O2 release.
The levels of H2O2 were determined by a modification of the method of Zhou et al. (48). The cells were washed twice with ice-cold PBS, and cells were harvested by microcentrifugation and resuspended in a Krebs-Ringer phosphate solution (KRPG; in mM: 145 NaCl, 5.7 sodium phosphate, 4.86 KCl, 0.54 CaCl2, 1.22 MgSO4, and 5.5 glucose, pH 7.35). One hundred microliters of the reaction mixture [50 µM Amplex Red reagent containing 0.1 U/ml horseradish peroxidase (HRP) in KRPG] was added to each microplate well and then prewarmed at 37°C for 10 min. After this, the reaction was started by adding resuspended cells in 20 µl of KRPG. Fluorescence readings became stable within 30 min of the start of the reaction. The fluorescence intensities of reaction mixtures were measured at 30 min with a fluorescence microplate reader (Multiskan; Thermo Labsystems, Franklin, MA) equipped for absorbance at 560 nm.
AA release. [3H]AA release experiments were performed by a modification of the method of Xing et al. (46). Confluent monolayers of PTC cultures were incubated for 24 h in DMEM-F-12 medium containing [3H]AA (0.5 µCi/ml) as well as the three growth supplements. The monolayers were then washed three times with PBS (pH 7.4) and incubated (at 37°C) for 1 h in DMEM-F-12 medium containing the specified agents at appropriate concentrations. At the end of the incubation period, the incubation medium was removed by aspiration and transferred to ice-cold tubes containing 100 µl of 55 mM EGTA and EDTA (final concentration 5 mM each). The uptake buffer was then centrifuged at 12,000 g to eliminate cell debris. To determine the level of radioactivity in the supernatant, the samples were placed in scintillation fluid and the radioactivity was counted with a liquid scintillation counter. The cells that remained attached to the plate were scraped into 1 ml of 0.1% SDS. Nine hundred microliters of the resulting cell lysate were used for scintillation counting. The remaining 100 µl of the cell lysate were used for protein determinations. For each condition, the quantity of [3H]AA that had been released (determined as described above) was first standardized with respect to protein. Subsequently, this standardized level of released [3H]AA was compared to the percentage of the total level of [3H]AA that had been incorporated into the cells at the beginning of the incubation period (or the total released radioactivity plus the total cell-associated radioactivity at the end of the stimulation period).
PGE2 assay. PGE2 levels in the culture medium were measured by radioimmunoassay using a general assay procedure adapted from Cetta and Goetz (9). In preliminary studies, PG was recovered from culture medium with extraction fluid (ethyl acetate-isopropanol-HCl, 0.05 N, 3:3:1) and the recovery rate was relatively constant (92 ± 3%, n = 9). When PGE2 was assayed in increasing aliquots of unextracted medium (25, 50, 100 µl), potency estimates were parallel to a linearly transformed dose-response curve. Thus the medium samples were assayed directly without extraction. Each sample was quantified with a liquid scintillation counter. Duplicate hormone standards (51,000 pg) were included in each assay. The between- and within-assay coefficients of variation for PGE2 were 7.3% and 6.5%, respectively.
Membrane preparation for cPLA2 blotting. Medium of confluent PTCs was exchanged 1 day before the experiment. The medium was then removed, and the cells were washed twice with ice-cold PBS, scraped, harvested by microcentrifugation, and resuspended in buffer A [in mM: 137 NaCl, 8.1 Na2HPO4, 2.7 KCl, 1.5 KH2PO4, 2.5 EDTA, 1 dithiothreitol, and 0.1 PMSF with 10 µg/ml leupeptin (pH 7.5)]. The resuspended cells were then mechanically lysed on ice by trituration with a 21.1-gauge needle. The lysates were first centrifuged at 1,000 g for 10 min at 4°C. The supernatants were centrifuged at 100,000 g for 1 h at 4°C to prepare cytosolic and total particulate fractions. The particulate fractions, which contained the membrane fraction, were washed twice and resuspended in buffer A containing 1% (vol/vol) Triton X-100. The protein in each fraction was quantified with a Bradford procedure (7).
Western blot analysis. Cell homogenates (20 µg of protein) were separated using 10% SDS-polyacrylamide gel electrophoresis and transferred to nitrocellulose paper. Blots were then washed with H2O, blocked with 5% skimmed milk powder in TBST [10 mM Tris·HCl (pH 7.6), 150 mM NaCl, and 0.05% Tween 20] for 1 h and incubated with the appropriate primary antibody at dilutions recommended by the supplier. The membrane was then washed, primary antibodies were detected with goat anti-rabbit-IgG (1:5,000) conjugated to HRP, and the bands were visualized with enhanced chemiluminescence (Amersham Pharmacia Biotech).
-MG uptake.
-MG uptake experiments were conducted according to the method of Sakhani et al. (34). To study
-MG uptake, the culture medium was removed by aspiration and monolayers were gently washed twice with the uptake buffer (in mM: 136 NaCl, 5.4 KCl, 0.41 MgSO4, 1.3 CaCl2, 0.44 Na2HPO4, 0.44 KH2PO4, 5 HEPES, and 2 glutamine, with 0.5 µg/ml BSA, pH 7.4). After the washing procedure, the monolayers were incubated at 37°C for 30 min in an uptake buffer that contained 0.5 mM
-MG and [14C]
-MG (0.5 µCi/ml). At the end of the incubation period, the monolayers were again washed three times with ice-cold uptake buffer and the cells were solubilized in 1 ml 0.1% SDS. To determine the [14C]
-MG incorporated intracellularly, 900 µl of each sample was removed and counted in a liquid scintillation counter. The remainder of each sample was used for protein determination by a Bradford method (7). The radioactivity counts in each sample were then normalized with respect to protein and were corrected for zero time uptake per milligram of protein. All uptake measurements were made in triplicate. Pi, L-arginine, fructose, and L-alanine uptake experiments were conducted as described by Rabito (33), Acevedo et al. (1), Corpe et al. (13), and Nishida et al. (28), respectively.
Statistical analysis. Results are expressed as means ± SE. The difference between two mean values was analyzed by the nonparametric Wilcoxon sign test or ANOVA. Differences were considered statistically significant when P < 0.05.
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RESULTS |
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DISCUSSION |
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The present study shows that the inhibition of [3H]thymidine incorporation induced by oxalate exposure was associated with increased H2O2 production. It suggests that superoxide or another reactive oxygen species (ROS) may mediate oxalate actions. It has been proposed that renal epithelial cells are stimulated by high levels of oxalate and, as a result, produce certain molecules favoring crystal attachment on cell surface membranes (2). Thus we suggest that oxalate inhibited PTC proliferation via a process dependent on reactive oxygen intermediates. Results of the present studies also demonstrate that the pretreatment of PTCs with the antioxidants NAC and catalase protects oxalate-induced inhibition of [3H]thymidine incorporation. These data confirm previous findings by other laboratories that oxalate increases superoxide production in LLC-PK1 and MDCK cells and that catalase and superoxide dismutase as well as peroxidase prevent oxalate-induced cell injury (2, 36). In addition, the addition of catalase caused a significant decrease in H2O2, demonstrating that the cellular effect on oxalate exposure was a result of the production of ROS (3, 19). Bhandari et al. (4) also demonstrated that oxalate increases superoxide production in HK-2 cells, as measured by nitro blue tetrazolium assay.
The SAPK and MAPK pathways play critical roles in responding to cellular stress and promoting cell growth and survival. Therefore, we investigated the effect of oxalate on MAPK signaling pathways. Our results demonstrate that oxalate stimulates p38 MAPK and SAPK/JNK. In contrast, p42/44 MAPK was not affected by oxalate. Pretreatment of cells with SB-203580 and SP-600125, but not PD-98059, blocked oxalate-induced inhibition of [3H]thymidine incorporation. These findings suggest that the inhibition of [3H]thymidine incorporation by oxalate was mediated by activation of p38 MAPK and SAPK/JNK. The activation of p38 MAPK by oxalate in LLC-PK1 cells is rapid and robust (27). Activation of the p38 MAPK pathway results in a plethora of changes in transcription, protein synthesis, cell surface receptor expression, and cytoskeletal structure, ultimately affecting cell survival or leading to programmed cell death (29, 38). Thus activation of the p38 MAPK cascade is suggestive of a functional role of this kinase cascade in mediating cellular actions of oxalate. With regard to p38 MAPK and serine-threonine protein kinases, as well as the JNK family, the major stress-activated signaling pathway is activated by a number of cellular stresses (37, 44). We found that oxalate caused activation of JNK in PTCs. JNK is activated by osmotic stress and during ischemia-reperfusion of the kidney (47). Recent reports suggest the involvement of JNK in apoptotic signals (30). It has also been shown that extracellular stress-related kinase-1/2 activation inhibits apoptosis, whereas JNK mediates apoptosis induced by cytokine (45). Oxalate did not cause activation of p42/44 MAPK in PTCs. Oxalate did not cause activation of p42/44 MAPK in LLC-PK1 cells (10). p42/44 MAPK are activated by mitogens, and a common view is that they are essentially shared elements in mitogenic signaling. However, DNA synthesis can occur independently of p42/44 MAPK activation (17). Our results also demonstrate that oxalate-induced inhibition of [3H]thymidine incorporation does not involve the p42/44 MAPK.
In the present study, oxalate increased AA release from PTCs by a process involving cPLA2. In addition, on the basis of evidence obtained by using a selective inhibitor of this isoform, it would appear that the activity of this enzyme is responsible, at least in part, for the cellular effects of oxalate. Not all cells succumb to oxalate toxicity; however, in those cells that do not succumb, ROS and lipid-signaling molecules induce changes in gene expression that allow them to survive and adapt to the toxic insult (22). Indeed, PLA2 has been implicated in the processes leading to cellular injury in many cell types (15, 41), including renal epithelial cells (35). ROS can promote phosphorylation of cPLA2 (11). Because oxalate produces ROS, it is possible that oxalate-induced AA release in PTCs is secondary to the generation of ROS. Many or all oxalate-induced responses are blocked by antioxidants and can be mimicked by PLA2 agonist. Neither OPC, a selective inhibitor for sPLA2, nor HELSS, a selective inhibitor for calcium-independent PLA2, blocked oxalate-induced [3H]AA release. These findings strongly support the role of cPLA2 as the enzyme primarily responsible for oxalate-induced AA release from PTCs, although we cannot entirely rule out a contribution from other forms of PLA2, given the limited selectivity and efficacy of the pharmacological tools available. These results suggest links between oxalate-induced increase in oxidant stress and subsequent molecular responses that may eventuate in renal cell injury.
The MAPKs p44/42 and p38 can both contribute to the activation of AA release. One type of MAPK, p44/42, was proposed early on to cause the phosphorylation and activation of cPLA2 (39). Later studies in smooth muscle cells and certain cell lines using the p38 MAPK inhibitor SB-203580 have instead suggested more prominent roles for p38 MAPK in the activation of cPLA2 induced by endothelin-1 (14). We have investigated the hypothesis that p44/42 and p38 MAPK, together or independent of one another, play roles in the regulation of AA release in PTCs responding to oxalate. The present results suggest that oxalate causes p38 MAPK cPLA2 activation and AA release in PTCs. Consistent with our results, collagen-induced phosphorylation of cPLA2 was shown in the presence of SB-203580 in human platelets and was not altered in the presence of PD-98059 (6). In addition, in human neutrophils treated with tumor necrosis factor-, the activation of cPLA2 appeared to be regulated by p38 rather than p44/42 MAPK (42). The present results also demonstrate that oxalate can induce cPLA2-mediated [3H]AA release and inhibit proliferation of PTCs. Such changes may play a role in the development and/or progression of renal dysfunctions through oxalate exposure of PTCs, although further studies are required to assess the steps involved in oxalate's action. In conclusion, oxalate inhibits renal PTC proliferation via oxidative stress, p38 MAPK/JNK, and cPLA2 signaling pathways.
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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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