Oxalate inhibits renal proximal tubule cell proliferation via oxidative stress, p38 MAPK/JNK, and cPLA2 signaling pathways

Ho Jae Han, Min Jin Lim, and Yun Jung Lee

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


    ABSTRACT
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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 REFERENCES
 
Exposure of renal proximal tubule cells to oxalate may play an important role in cell proliferation, but the signaling pathways involved in this effect have not been elucidated. Thus the present study was performed to examine the effect of oxalate on 3H-labeled thymidine incorporation and its related signal pathway in primary cultured rabbit renal proximal tubule cells (PTCs). The effects of oxalate on [3H]thymidine incorporation, lactate dehydrogenase (LDH) release, Trypan blue exclusion, H2O2 release, activation of mitogen-activated protein kinases (MAPKs), and 3H-labeled arachidonic acid (AA) release were examined in primary cultured renal PTCs. Oxalate inhibited [3H]thymidine incorporation in a time- and dose-dependent manner. However, its analogs did not affect [3H]thymidine incorporation. Oxalate (1 mM) significantly increased H2O2 release, which was blocked by N-acetyl-L-cysteine (NAC) and catalase (antioxidants). Oxalate significantly increased p38 MAPK and stress-activated protein kinase (SAPK)/c-Jun NH2-terminal kinase (JNK) activity, not p44/42 MAPK. Oxalate stimulated [3H]AA release and translocation of cytosolic phospholipase A2 (cPLA2) from the cytosolic fraction to the membrane fraction. Indeed, oxalate significantly increased prostaglandin E2 (PGE2) production compared with control. Oxalate-induced inhibition of [3H]thymidine incorporation and increase of [3H]AA release were prevented by antioxidants (NAC), a p38 MAPK inhibitor (SB-203580), a SAPK/JNK inhibitor (SP-600125), or PLA2 inhibitors [mepacrine and arachidonyl trifluoromethyl ketone (AACOCF3)], but not by a p44/42 MAPK inhibitor (PD-98059). These findings suggest that oxalate inhibits renal PTC proliferation via oxidative stress, p38 MAPK/JNK, and cPLA2 signaling pathways.

kidney; mitogen-activated protein kinase; phospholipase A2


PREVIOUS STUDIES on LLC-PK1 cells demonstrated that oxalate, a simple dicarboxylic acid, acts as a mitogen for these renal epithelial cells. Exposure to oxalate initiates DNA synthesis and stimulates proliferation of quiescent cultures of LLC-PK1 cells (24). With toxic challenges such as exposure to high levels of oxalate, proximal tubule cells (PTCs) may be damaged and release their contents into the urine (16). However, the possible role of oxalate as a modulator of renal cells has not been fully appreciated. A recent study demonstrated that oxalate binds to specific receptors on the renal tubule cell surface (26). The receptors may be only minimally exposed under normal circumstances and increased in number under a variety of conditions that lead to cellular stress and injury (26, 40). Nonetheless, oxalate interaction with renal epithelial cells results in a program of events, including alterations in gene expression, initiation of DNA synthesis, cell growth, and death depending on the levels of oxalate (18). These considerations led us to investigate whether oxalate has an effect on the proliferation of renal tubular epithelial cells. Recent studies have shown that exposure to high levels of oxalate produces oxidative stress in renal epithelial cells (23). From the observation that oxalate can increase production of free radicals (36), it follows that arachidonic acid (AA) metabolism catalyzed by cytosolic phospholipase A2 (cPLA2) may be involved in the mechanism by which oxalate elicits renal cellular injury. Moreover, the stress- and mitogen-activated protein kinase (SAPK and MAPK) pathways play critical roles in responding to DNA synthesis by oxalate (5, 44). However, the specific signaling pathways activated in renal PTCs after oxalate exposure have not been elucidated.

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 {gamma}-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.


    MATERIALS AND METHODS
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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Materials. Adult New Zealand White male rabbits (1.5–2.0 kg) were purchased from Dae Han Experimental Animal (Chungju, Korea). Class IV collagenase and soybean trypsin inhibitor were purchased from Life Technologies (Grand Island, NY). Oxalate, succinic acid, citric acid, formic acid, Trypan blue, mepacrine, arachidonyl trifluoromethyl ketone (AACOCF3), SB-203580, N-acetyl-L-cysteine (NAC), catalase, SP-600125, prostaglandin E2 (PGE2), haloenol lactone suicide substrate (HELSS), and oleyloxyethyl phosphorylcholine (OPC) were obtained from Sigma (St. Louis, MO). PD-98059 was purchased from Calbiochem (La Jolla, CA). [3H]thymidine, [3H]AA, 14C-labeled {alpha}-methyl-D-glucopyranoside ({alpha}-MG), 32P, [3H]alanine, L-[3H]arginine, and PGE2 [5,6,8,11,12,14,1-3H(N)-prostaglandin E2] were purchased from NEN (Boston, MA). The lactate dehydrogenase (LDH) assay kit was obtained from Iatron Lab (Tokyo, Japan). Antibodies to p44/42, p38, SAPK/JNK, and cPLA2 were purchased from Santa Cruz Biotechnology (Santa Cruz, CA), and all other reagents were of the highest purity commercially available. Liquiscint was obtained from National Diagnostics (Atlanta, GA).

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 (1.5–2.0 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 10–8 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 70–80% 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 (5–1,000 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).

{alpha}-MG uptake. {alpha}-MG uptake experiments were conducted according to the method of Sakhani et al. (34). To study {alpha}-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 {alpha}-MG and [14C]{alpha}-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]{alpha}-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.


    RESULTS
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
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Effect of oxalate on [3H]thymidine incorporation. To determine the time- and dose-response effect of oxalate on [3H]thymidine incorporation, PTCs that were 70–80% confluent were incubated in the presence of various concentrations of oxalate (0–4 mM total, 3.8–1,520 µM free) for different times (0–72 h). When PTCs were treated with 1 mM oxalate, [3H]thymidine incorporation was inhibited from 2 h and this effect was significantly increased over 48 h in a time-dependent manner (Fig. 1A). [3H]thymidine incorporation was also inhibited in the presence of >1 mM oxalate for 24 h (Fig. 1B). In Trypan blue exclusion and LDH (nonspecific cell injury marker enzyme) release studies to examine the extent of cell injury with 1 mM oxalate treatment for 24 h, there were no significant changes in cell death for control or oxalate treatment (Table 1). In addition, oxalate did not affect fructose, {alpha}-MG, L-arginine, L-alanine, and Pi uptake, functions of apical transporters (Table 2). Moreover, the effect on [3H]thymidine incorporation appeared to be unique to oxalate because structurally similar compounds (succinic acid, formic acid, and citric acid; all added at 1 mM) had no effect in PTCs (Fig. 2). Thus all further experiments were performed with 1 mM oxalate for 24 h.



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Fig. 1. Time- and dose-dependent effects of oxalate on 3H-labeled thymidine incorporation. A: proximal tubule cells (PTCs) were incubated in the presence of oxalate (1 mM total, 380 µM free) for different time periods (0–72 h) and subsequently pulsed with 1 µCi of [3H]thymidine for 24 h before counting. B: PTCs were incubated for 24 h in the absence or presence of oxalate (0.1–4 mM total, 38–1,520 µM free) and pulsed with 1 µCi of [3H]thymidine for 24 h. Values are means ± SE of 4 independent experiments with triplicate dishes. *P < 0.05 vs. control.

 

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Table 1. Effect of oxalate on LDH and cell viability

 

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Table 2. Effect of oxalate on fructose, {alpha}-MG, L-arginine, L-alanine, and Pi uptake

 


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Fig. 2. Effects of succinic acid, formic acid, and citric acid on [3H]thymidine incorporation. PTCs were incubated in the presence of oxalate, succinic acid, formic acid, and citric acid (1 mM) for 24 h and pulsed with 1 µCi of [3H]thymidine for 24 h. Values are means ± SE of 4 independent experiments with triplicate dishes. *P < 0.05 vs. control.

 
Involvement of oxidative stress in oxalate-induced inhibition of [3H]thymidine incorporation. To examine the relationship of oxidative stress to the oxalate-induced inhibition of [3H]thymidine incorporation, we examined the effect of oxalate on H2O2 release. A statistically significant increase in H2O2 release was found from 30 min after the start of the 1 mM oxalate incubation (Fig. 3A). The greatest effect of oxalate-induced stimulation of H2O2 release was observed at 240 min, and prolongation of the incubation period for >480 min did not further increase the stimulation. One, two, or four millimolar oxalate for 120 min significantly increased H2O2 release, whereas 0.1 mM oxalate did not (Fig. 3B). These oxalate-induced increases of H2O2 release and inhibition of [3H]thymidine incorporation were significantly blocked by pretreatment with the antioxidants NAC (1 µM) and catalase (600 U/ml) (Fig. 4). H2O2 (1 µM) plus oxalate further inhibited [3H]thymidine incorporation compared with oxalate alone (Fig. 4B).



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Fig. 3. Time- and dose-dependent effects of oxalate on H2O2 release. PTCs were treated with oxalate (1 mM total, 380 µM free) for different times (0–480 min) (A) and incubated for 120 min in the absence or presence of oxalate (0.1–4 mM total, 380–1,520 µM free) (B). Values are means ± SE of 4 independent experiments with triplicate dishes. *P < 0.05 vs. control.

 


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Fig. 4. Effects of antioxidants on oxalate-induced H2O2 release (A) and inhibition of [3H]thymidine incorporation (B). PTCs were treated with N-acetyl-L-cysteine (NAC, 1 µM) or catalase (600 U/ml) before treatment with oxalate (1 mM total, 380 µM free) or were incubated with oxalate alone or together with H2O2 (1 µM) for 24 h. Values are means ± SE of 3 independent experiments with triplicate dishes. Open bars, control; filled bars, oxalate. *P < 0.05 vs. control; **P < 0.05 vs. oxalate alone.

 
Involvement of MAPKs in oxalate-induced inhibition of [3H]thymidine incorporation. To examine the effects of MAPKs on oxalate-induced inhibition of [3H]thymidine incorporation, PTCs were treated with 1 mM oxalate for 24 h. Oxalate at 1 mM increased phosphorylation of p38 MAPK from the 15-min treatment of oxalate. Oxalate also increased SAPK/JNK activity at 10 min. However, p42/44 MAPK was not affected by oxalate (Fig. 5A). SB-203580 (a p38 MAPK inhibitor; 1 µM) and SP-600125 (a SAPK/JNK inhibitor; 1 µM) prevented oxalate-induced inhibition of [3H]thymidine incorporation, whereas PD-98059 (a p44/42 MAPK inhibitor; 1 µM) did not block it (Fig. 5B).



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Fig. 5. Time course of effect of oxalate on mitogen-activated protein kinase (MAPK) activity (A) and effects of SP-600125, SB-203580, or PD-98059 on oxalate-induced inhibition of [3H]thymidine incorporation (B). A: representative autoradiographs of time course of MAPK phosphorylation by oxalate. PTCs were treated at different time points (0–120 min) with oxalate. Phosphorylated p38, stress-activated protein kinase (SAPK)/c-Jun NH2-terminal kinase (JNK), and p44/42 MAPKs were then detected as described in MATERIALS AND METHODS. B: PTCs were treated with SP-600125 (SAPK/JNK inhibitor), SB-203580 (p38 MAPK inhibitor), or PD-98059 (p44/42 MAPK inhibitor) (1 µM) for 30 min before treatment with oxalate or were incubated with oxalate alone for 24 h. Values are means ± SE of 3 or 5 independent experiments with triplicate dishes. Open bars, control; filled bars, oxalate. *P < 0.05 vs. control; **P < 0.05 vs. oxalate alone.

 
Relation of PLA2 to oxalate-induced inhibition of [3H]thymidine incorporation. To examine the relation of PLA2 to oxalate-induced inhibition of [3H]thymidine incorporation, we examined the effect of oxalate on [3H]AA release. Figure 6A shows that 1-h incubation with 1 mM oxalate significantly increased [3H]AA release and PGE2 production. Oxalate resulted in translocation of cPLA2 from the cytosolic fraction to the membrane fraction (Fig. 6B). Oxalate-induced increase of [3H]AA release and inhibition of [3H]thymidine incorporation were blocked by AACOCF3 and mepacrine (cPLA2 inhibitors), whereas HELSS (a calcium-independent PLA2 inhibitor) or OPC [a secretory PLA2 (sPLA2) inhibitor] (1 µM) had no effect (Fig. 7).



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Fig. 6. Dose-dependent effect of oxalate on 3H-labeled arachidonic acid (AA) release (A) and effect of oxalate on cytosolic phospholipase A2 (cPLA2) translocation (B). A: after the incorporation of [3H]AA (0.5 µCi/ml) into the PTCs for 24 h, PTCs were washed 3 times with Dulbecco's modified Eagle's medium (DMEM)-F-12, pH 7.4, and incubated in the same medium at 37°C for 1 h. Oxalate (0.1–4 mM total, 38–1,520 µM free) was then used to treat the PTCs for 1 h. Inset: result of prostaglandin (PG)E2 assay after treatment with oxalate (1 mM total, 380 µM free) for 24 h. B: PTCs were treated with oxalate for 4 h. The amount of cPLA2 protein was determined by Western blot analysis of cytosolic fraction or membrane particulate fraction as described in MATERIALS AND METHODS. Arrow indicates the 110-kDa band corresponding to cPLA2. The example shown is a representative of 3 experiments. Values are means ± SE of 3 or 5 independent experiments with triplicate dishes. Open bars, control; filled bars, oxalate. *P < 0.05 vs. control.

 


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Fig. 7. Effects of PLA2 inhibitors on oxalate-induced [3H]AA release (A) and inhibition of [3H]thymidine incorporation (B). PTCs were treated with arachidonyl trifluoromethyl ketone (AACOCF3) and mepacrine (PLA2 inhibitors), haloenol lactone suicide substrate (HELSS; calcium-independent PLA2 inhibitor), and oleyloxyethyl phosphorylcholine [OPC; secretory (s)PLA2 inhibitor] (1 µM) for 30 min before treatment with oxalate or were incubated with oxalate alone for 1 h (A) and 24 h (B). Values are means ± SE of 3 or 4 independent experiments with triplicate dishes. Open bars, control; filled bars, oxalate. *P < 0.05 vs. control; **P < 0.05 vs. oxalate alone.

 
Relationship among oxidative stress, MAPK, and cPLA2 in oxalate-induced inhibition of [3H]thymidine incorporation. We next examined the relationship among oxidative stress, MAPK, and cPLA2 in oxalate-induced inhibition of [3H]thymidine incorporation. SB-203580 (a p38 MAPK inhibitor), SP-600125 (a SAPK/JNK inhibitor), and AACOCF3 (a cPLA2 inhibitor) (1 µM) did not block the oxalate-induced increase of H2O2 release, whereas NAC (1 µM) blocked the increase of H2O2 induced by oxalate (Fig. 8). NAC (an antioxidant), but not AACOCF3, blocked oxalate-induced phosphorylation of p38 MAPK (Fig. 9). In the [3H]AA release experiment, NAC, catalase, SB-203580, and SP-600125, but not PD-98059, prevented the oxalate-induced increase of [3H]AA release (Fig. 10). These results were consistent with those of [3H]thymidine incorporation.



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Fig. 8. Effects of NAC, SB-203580, SP-600125, and AACOCF3 on oxalate-induced H2O2 release. PTCs were treated with NAC, SB-203580, SP-600125, or AACOCF3 (1 µM) for 30 min before treatment with oxalate or were incubated with oxalate alone for 24 h. Values are means ± SE of 4 independent experiments with triplicate dishes. Open bars, control; closed bars, oxalate. *P < 0.05 vs. control; **P < 0.05 vs. oxalate alone.

 


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Fig. 9. Effects of NAC and AACOCF3 on oxalate-induced phosphorylated p38 MAPK. PTCs were treated with NAC or AACOCF3 (1 µM) for 30 min before treatment with oxalate. Phosphorylated p38 MAPK was then detected as described in MATERIALS AND METHODS. The example shown is a representative of 3 experiments. Data are expressed as %basal value in each fraction and are means ± SE of 3 independent experiments. Open bar, control; filled bars, oxalate. *P < 0.05 vs. control; **P < 0.05 vs. oxalate alone.

 


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Fig. 10. Effects of antioxidants and MAPK inhibitors on oxalate-induced [3H]AA release. PTCs were treated with NAC, PD-98059, SB-203580, or SP-600125 (1 µM) or catalase (600 U/ml) for 30 min before treatment with oxalate or were incubated with oxalate alone for 1 h. Values are means ± SE of 4 independent experiments with triplicate dishes. Open bars, control; filled bars, oxalate. *P < 0.05 vs. control; **P < 0.05 vs. oxalate alone.

 

    DISCUSSION
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
In the present study, we demonstrated that oxalate inhibits proliferation of renal PTCs via oxidative stress, p38 MAPK/JNK, and cPLA2 signal pathways. Previous studies demonstrated that exposure to various concentrations of oxalate produces biphasic effects on PTC proliferation, with lower concentrations (80–320 µM) increasing and higher concentrations (>400 µM) decreasing the number of LLC-PK1 cells. Koul et al. (25) also demonstrated time- and concentration-dependent stimulation of DNA synthesis, with maximal stimulation after 48 h of exposure to 400 µM free oxalate. However, in our study concentration-dependent inhibition of renal PTC proliferation was observed monophasically. This discrepancy may be partially caused by differences in experimental model system [i.e., animal species (rabbit vs. rat), cell type (proximal tubules vs. smooth muscle or nerve cell), culture condition (serum vs. hormonally defined), or other experimental conditions]. It has been estimated that the oxalate concentrations in the glomerular filtrate range from 1 to 10 µM, rising 10- to 30-fold during tubular transit (32). In the present studies, sodium oxalate was added at a concentration of 1 mM (380 µM free), which is somewhat higher than expected to occur in the proximal tubule under normal physiological conditions. This concentration of oxalate inhibited [3H]thymidine incorporation, whereas cell viability and apical transporter activities were not affected. These findings support the concept that oxalate exposure of PTCs may stimulate multiple signal pathways, although these pathways remain to be explored.

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-{alpha}, 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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This work was supported by the Stem Cell Research Center of 21st Century Frontier Research Program funded by the Ministry of Science and Technology, Republic of Korea (contract grant no. SC14032).


    FOOTNOTES
 

Address for reprint requests and other correspondence: H. J. Han, Dept. of Veterinary Physiology, College of Veterinary Medicine, Chonnam National Univ., Gwangju, Korea 500-757 (E-mail: hjhan{at}chonnam.ac.kr)

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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