Oxidative stress resulting from unilateral ureteral obstruction (UUO) may be aggravated by increased production of ROS. Previous studies have demonstrated increased cyclooxygenase (COX)-2 expression in renal medullary interstitial cells (RMICs) in response to UUO. We investigated, both in vivo and in vitro, the role of ROS in the induction of COX-2 in rats subjected to UUO and in RMICs exposed to oxidative and mechanical stress. Rats subjected to 3-day UUO were treated with two mechanistically distinct antioxidants, the NADPH oxidase inhibitor diphenyleneiodonium (DPI) and the complex I inhibitor rotenone (ROT), to interfere with ROS production. We found that UUO-mediated induction of COX-2 in the inner medulla was attenuated by both antioxidants. In addition, DPI and ROT reduced tubular damage and oxidative stress after UUO. Moreover, mechanical stretch induced COX-2 and oxidative stress in RMICs. Likewise, RMICs exposed to H2O2 as an inducer of oxidative stress showed increased COX-2 expression and activity, both of which were reduced by DPI and ROT. Similarly, ROS production, which was increased after exposure of RMICs to H2O2, was also reduced by DPI and ROT. Furthermore, oxidative stress-induced phosphorylation of ERK1/2 and p38 was blocked by both antioxidants, and inhibition of ERK1/2 and p38 attenuated the induction of COX-2 in RMICs. Notably, COX-2 inhibitors further exacerbated the oxidative stress level in H2O2-exposed RMICs. We conclude that oxidative stress as a consequence of UUO stimulates COX-2 expression through the activation of multiple MAPKs and that the induction of COX-2 may exert a cytoprotective function in RMICs.
- ureteral obstruction
- oxidative stress
- reactive oxygen species
obstructive nephropathy is an important cause of renal insufficiency in both children and adults. Unilateral ureteral obstruction (UUO) is a well-established experimental rodent model that mimics the severe renal injury found in obstructive nephropathy (19).
The hydrostatic pressure increase as a result of obstruction causes massive tubular dilation, interstitial inflammatory infiltration, apoptotic tubular cell deletion, and progressive tubulointerstitial fibrosis (8). Importantly, oxidative stress plays an important role in the pathogenesis of UUO (16, 17). Several markers of oxidative stress, such as the oxidative stress response molecule heme oxygenase (HO)-1 and heat shock protein 27, are increased in UUO kidneys (16, 29).The general paradigm is that oxidative stress occurs when the production of ROS is greater than the ability of cells to detoxify the produced ROS; indeed, increased concentrations of ROS have been observed in the obstructed kidney (35). Although oxidative stress is involved in the UUO model, little is known about the source of the stress.
In vivo studies have demonstrated the induction of cyclooxygenase (COX)-2 in the inner medulla in response to both unilateral and bilateral ureteral obstruction (26–28), and immunohistochemical analysis showed marked labeling of COX-2 in renal medullary interstitial cells (RMICs) in the obstructed kidney (27). COXs, which are bifunctional enzymes that catalyze the conversion of arachidonic acid into PGs, can be divided into two isoforms, namely, COX-1 and COX-2 (5). COX-1 is constitutively expressed in most tissues and is thought to be responsible for the production of PGs involved in the regulation of normal “housekeeping” cellular processes, whereas COX-2 is undetectable in most tissues under normal physiological conditions (5). However, COX-2 can be rapidly and transiently induced by local osmotic, inflammatory, and mechanical stimuli, in addition to its homeostatic role (5).
Previous in vitro studies using collecting duct cells have demonstrated that COX-2 expression is increased by hypertonic stress in a ROS- and MAPK-dependent signaling pathway (34). In addition, it has also been shown that hyperglycemia leads to increases in mitochondrial ROS production as well as COX-2 expression and activity in human mesangial cells (18). However, it remains unclear whether ROS production plays a role in the induction of COX-2 in RMICs of rats subjected to UUO. In this study, we hypothesized that ROS production might increase the expression and activity of COX-2 in rat RMICs in response to UUO and play an important role in the pathophysiology of obstructive nephropathy.
The present study was designed to elucidate the role of ROS in the expression and activity of COX-2 and to characterize the signal transduction pathway responsible for the regulatory mechanism in RMICs. To achieve this, we investigated, both in vivo and in vitro, the effects of two antioxidants that interfere with different ROS production sites (i.e., NADPH oxidase and mitochondria) on the regulation of COX-2 as well as oxidative stress in rats subjected to UUO for 3 days and in RMICs exposed to oxidative stress induced by H2O2 and mechanical stress produced by stretching the cells.
MATERIALS AND METHODS
Experiments were performed on male Munich-Wistar rats initially weighing 220 g. Animal protocols were approved by the board of the Institute of Clinical Medicine, Aarhus University, according to the licenses for use of experimental animals issued by the Danish Ministry of Food, Agriculture and Fisheries. Animals had ad libitum access to rodent diet (Altromin, Lage, Germany) and tab water. Rats were kept in cages with a 12:12-h light-dark cycle, a temperature of 21 ± 2°C, and a humidity of 55 ± 5%.
Animals were treated with sevoflurane (Abbott Scandinavia, Solna, Sweden) under anesthesia and placed on a heating pad to maintain the rectal temperature at 37–38°C. Left ureter was exposed by making a midline abdominal incision were occluded with a silk ligature. The abdominal incision was closed. Age- and time-matched sham-operated control animals were prepared and observed in parallel with each UUO group in the following protocols.
UUO was induced for 3 days. In group 1, rats were treated subcutaneously with diphenyleneiodonium (DPI; 1.5 mg·kg−1·day−1, Sigma-Aldrich, Brøndby, Denmark) dissolved in isotonic glucose starting 1 day before the operation and throughout the experiment. DPI was administrated in the afternoon, which would minimize the time before rats were active and started eating, thereby minimizing the chance for hypoglycemia. The dose was chosen on the basis of work performed by Cooper et al. (9). At the end of the experiment, rats were euthanized for either immunoblot analysis (n = 6) or immunohistochemistry (n = 4). In group 2, rats were treated subcutaneously with isotonic glucose 1 day before the experiment and throughout the experiment. Rats were euthanized for either immunoblot analysis (n = 6) or immunohistochemistry (n = 4). Sham-operated rats were prepared in parallel.
UUO was induced for 3 days. In group 1, rats were fed rotenone (ROT; Sigma-Aldrich) mixed into their food (600 mg/kg food) starting 1 day before the operation and throughout the experiment. Given the important role of mitochondria in energy production, ROT is suspected to induce some degree of toxicity. The dose was chosen on the basis of work performed by Zhang et al. (36) and the National Toxicology Program (25a), where rats tolerated ROT at doses of 600 mg/kg for up to 2 yr. At the end of the experiment, rats were euthanized for either immunoblot analysis (n = 6) or immunohistochemistry (n = 4). In group 2, rats were fed with normal rodent diet. Rats were euthanized for either immunoblot analysis (n = 6) or immunohistochemistry (n = 4). Sham-operated rats were prepared in parallel.
Blood Sampling and Kidney Removal
After rats were reanesthetized with sevoflurane, the midline incision was reopened, and the aortic bifurcation was dissected free. Blood samples (5–7 ml) were collected. Rapidly thereafter, the kidneys were removed, and the inner medulla was dissected. Blood samples were analyzed to determine the levels of plasma creatinine and urea by a Roche Cobas 6000 analyzer (Roche Diagnostic).
RMICs (a gift from Dr. C. Maric, University of Mississippi Medical Center) were obtained from fresh renal medullary tissues of Sprague-Dawley rats (80–90 g) using a modified version of the method previously described by Fontoura et al. (11). These cells show features that are characteristic of papillary interstitial cells, including elongated cellular outlines, vacuolated cytoplasm, and cytoplasmic lipid droplets (22). Cells were grown in RPMI-1640 supplemented with 10% FBS, 4 mM l-glutamine, penicillin (100 U/ml), and streptomycin (100 mg/ml). Cells were incubated at 37°C in a 5% CO2-95% air humidified atmosphere. Culture media were changed every 48 h. Cells were passaged at confluence. By passage 10, homogeneous cell populations were generally reached, and cells between passages 10 and 20 were used in the experiments. Cultures were 80–90% confluent at the start of experiments, and media were replaced with serum-free media at 24 h before the experiments. The ERK1/2 inhibitor PD-98059, the p38 MAPK inhibitor SB-202190 (Sigma-Aldrich), and the COX-2 inhibitor SC-236 (Cayman Chemical, Ann Arbor, MI) were added into the serum-free medium and incubated for 1 h before the stimulation experiments.
The effect of stretch on RMICs was studied in vitro using the Flexcell FX-5000T system (Dunn Labortechnik, Asbach, Germany), which applies stretch to adhesive cell types. RMICs were cultured on collagen-coated BioFlex plates (six-well plates, Dunn Labortechnik) and exposed to uniform static stretch for 2, 6, and 12 h. To determine the optimal condition, we applied different amounts of static stretch to RMICs and increase the attached cell surface area by 10%, 15%, and 20%. As a control, nonstretched cells were used. The complete system was placed in a CO2 incubator to maintain the temperature, humidity, and atmosphere during the stretch experiment. In the optimal condition, stretch of 0% (control) and 20% was applied to RMICs for 2 h.
Membrane Fractionation for Immunoblot Analysis
RMICs were collected and lysed using M-PER Mammalian Protein Extraction Reagent (Thermo Scientific, Vedbaek, Denmark). Cell suspensions were centrifuged at 14,000 g at room temperature for 10 min. Tissues were prepared for immunoblot analysis by homogenization for 30 s at 1,250 rpm in dissection buffer (0.3 M sucrose, 25 mM imidazole, and 1 mM EDTA, pH 7.2) containing Complete mini protease inhibitor cocktail tablets (Roche) followed by centrifugation at 4,000 g for 15 min at 4°C. Gel samples were prepared from supernatants mixed with Laemmli sample buffer containing 2% SDS. The Pierce BCA Protein Assay Kit (Roche) was used to determine the total protein concentration of homogenates.
Electrophoresis and Immunoblot Analysis
Gel samples were run on 12% polyacrylamide minigels (Bio-Rad Mini Protean II, Bio-Rad, Copenhagen, Denmark). For each gel intended for Western blot analysis, an identical gel was run before blotting and subjected to Coomassie staining to ensure identical protein loading. β-Actin was used as a loading control for normalization.
Protein samples run on 12% polyacrylamide minigels were transferred to nitrocellulose membranes (Hybond ECL RNP 3032D, GE Healthcare Europe, Brøndby, Denmark). Blots were blocked in 5% skim milk dissolved in PBS with Tween 20 (80 mM Na2HPO4, 20 mM NaH2PO4, 100 mM NaCl, and 0.1% Tween 20, adjusted to pH 7.4). After washes with PBS-Tween 20, blots were incubated with primary antibodies overnight at 4°C. Antigen-antibody complexes were visualized with horseradish peroxidase-conjugated secondary antibodies (P0448 or P0447, 1:3,000, DAKO, Glostrup, Denmark) using the ECL system (GE Healthcare Europe).
For semiquantitative immunoblot analysis and immunohistochemistry, the following previously characterized monoclonal and polyclonal antibodies were used: COX-1 (Cayman Chemical), COX-2 (Abcam, Cambridge, UK), HO-1 (ENZO Life Sciences, Farmingdale, NY), β-actin (Sigma-Aldrich, St. Louis, MO), and p38, phospho-p38, ERK1/2, phospho-ERK1/2, JNK, phospho-JNK, Bcl-2, and Bax (Cell Signal Technology, Danvers, MA).
Histology and Immunohistochemistry
Kidneys from UUO rats and sham-operated control rats were fixed by retrograde perfusion via the abdominal aorta with 4% paraformaldehyde in 0.01 M PBS buffer. Next, organs were fixed for an additional hour and washed three times (10 min) with 0.01 M PBS buffer. Fixed kidneys were then dehydrated, embedded in paraffin, and cut into 2-μm sections on a rotary microtome (Leica Microsystems, Herlev, Denmark).
Paraffin-embedded sections were stained with hematoxylin and eosin to assess the grade of tubular damage. Under high magnification (×200), 10 nonoverlapping fields from each section of the renal cortex were photographed. The tubular luminal area of each section was measured using image-analysis software (ImageJ, National Institutes of Health). A grid containing sampling points was superimposed on each photograph. Points falling on glomerular structures and large vessels were excluded from the total count. The tubular dilatation score was determined by the number of points overlying dilated tubular spaces and then converted to a percentage. All analyses were performed blind.
For immunoperoxidase labeling, sections were deparaffinized, rehydrated, and processed for immunolabeling using previously characterized antibodies as described elsewhere (27).
Paraffin-embedded sections were permeabilized in 0.1% Triton X-100 and 0.1% sodium citrate in PBS and stained for apoptosis using an in situ cell detection kit (POD, Roche) followed by a counterstain in hematoxylin. Under magnification (×40), five to seven nonoverlapping sections were imaged from each inner medulla. The amount of TUNEL-positive cells was counted in Adobe Photoshop CS5 and normalized to the number of images per inner medulla and expressed as number of TUNEL-positive cells per section.
Measurements of ROS
Intracellular ROS generation in RMICs was quantified using 2′,7′-dichlorodihydrofluorescein diacetate (Sigma-Aldrich). Briefly, 1 day before the experiment, cells were seeded in 96-well plates and incubated for 6 h with H2O2 in the presence or absence of inhibitors of ROS production. Next, cells were washed twice with HBSS without phenol red and then incubated with 10 μM 2′,7′-dichlorodihydrofluorescein diacetate in HBSS for 30 min at 37°C. Finally, 2,7-dichlorofluorescein fluorescence was measured with excitation at 485 nm and emission at 520 nm.
Levels of PGE2 in culture media were measured using the PGE2 EIA kit (monoclonal, Cayman Chemical), according to the manufacturer's protocol. Briefly, RMICs were subjected to H2O2 treatment for 6 h with or without inhibitors of ROS production followed by PGE2 measurements.
Data are expressed as means ± SE. Statistical comparisons were analyzed by an unpaired Student's t-test when two groups were compared or by one-way ANOVA followed by a post hoc unpaired Student's t-test with the Bonferroni correction when several groups were compared. P values of <0.05 were taken as significant.
Effects of DPI and ROT on Tissue Damage and Oxidative Stress in Rats Subjected to 3-Day UUO
To examine the effects of two potential antioxidants that affect ROS production on tubular damage, oxidative stress, and apoptosis after UUO injury, we administered the NADPH oxidase inhibitor DPI and the mitochondrial respiratory chain complex I inhibitor ROT to rats subjected to 3-day UUO. To analyze apoptosis, both TUNEL staining and protein expression levels of proapoptotic Bax and antiapoptotic Bcl-2 were measured, whereas the oxidative stress marker HO-1 was used as an indicator of the oxidative stress level. As shown in Fig. 1A, obstructed kidneys had massive tubular dilatation and injury compared with kidneys from sham-operated control rats. Apoptosis assessed by both TUNEL staining (Fig. 1B) and the ratio of apoptosis-related proteins Bax to Bcl-2 (Bax/Bcl-2 ratio; Fig. 1C) as well as HO-1 expression (Fig. 1D) were significantly increased in the inner medulla of rats subjected to 3-day UUO compared with sham-operated rats. Notably, both tubular damage and oxidative stress were significantly attenuated in both DPI- and ROT-treated UUO rats. DPI also attenuated the increase in apoptosis levels in 3-day UUO rats, as measured by the Bax/Bcl-2 ratio. TUNEL staining did not show significant differences between 3-day UUO and DPI- or ROT-treated groups (Fig. 1B), indicating that DPI only affected the Bax/Bcl-2 ratio in response to 3-day UUO.
While body weight was unchanged among sham-operated control rats, untreated UUO rats, and DPI-treated UUO rats, ROT led to a slight decrease in the body weight of UUO rats. The obstructed kidneys from UUO rats appeared swollen and had an increased weight compared with those from sham-operated control rats. As shown in Table 1, the administration of DPI or ROT did not change the weight of obstructed kidneys. In addition, compared with sham-operated control rats, levels of plasma creatinine and urea were significantly increased in UUO rats but were not changed by either antioxidants. Together, these results show that inhibition of ROS production reduced tubular damage and oxidative stress after 3-day UUO injury, indicating an antioxidant role of DPI and ROT.
DPI and ROT Prevent the Induction of COX-2 in Rats Subjected to 3-Day UUO
To examine whether ROS production plays a role in the induction of COX-2 in response to ureteral obstruction, the expression and localization of COX-2 in the inner medulla of DPI- and ROT-treated UUO rats were investigated. The protein abundance of COX-2 was significantly increased in the inner medulla of rats subjected to 3-day UUO compared with sham-operated rats, and this induction was attenuated by DPI or ROT treatment (Fig. 2, A and B). COX-1 expression was decreased in UUO rats compared with sham-operated control rats; however, DPI and ROT did not change COX-1 levels in UUO rats (Fig. 2, A and C). Immunohistochemical analysis also showed strong COX-2 labeling in RMICs in the inner medulla of obstructed kidneys compared with control kidneys (Fig. 2, D and E). In contrast, weaker labeling was observed in DPI- or ROT-treated UUO rats (Fig. 2, F and G) compared with untreated UUO rats (Fig. 2E). In the cortex, there was no change in COX-2 protein levels among all four groups (data not shown). These results suggest that ROS play a role in the induction of COX-2 in RMICs of rats subjected to 3-day UUO.
DPI and ROT Reduce the Induction of COX-2 in RMICs Exposed to Oxidative Stress
Since ROS might play a role in the regulation of COX-2 expression in RMICs of rats subjected to 3-day UUO in vivo, we further investigated the signaling pathway responsible for the induction of COX-2 in RMICs exposed to 75 μM H2O2 to mimic oxidative stress in vitro. COX-2 mRNA and protein abundance in RMICs were found to be increased in a time-dependent manner upon H2O2 exposure (Fig. 3, A and B). In contrast, preincubation of RMICs with either 2.5–10 μM ROT or 10–25 μM DPI for 6 h led to a dose-dependent inhibition of COX-2 induction (Fig. 3C). Especially, treatment with DPI (10 μM) and ROT (2.5 μM) for 6 h effectively reduced H2O2-induced COX-2 protein abundance and PGE2 concentrations in RMICs (Fig. 3, D and E). Taken together, these in vitro observations are consistent with the in vivo results and support an important role for ROS in the increased COX-2 expression and activity in RMICs.
Roles of DPI and ROT in MAPK Pathway Activation in RMICs Exposed to Oxidative Stress
The mechanism linking ROS and COX-2 was further examined. Our previous study (6) demonstrated that the MAPK pathway plays a role in the regulation of COX-2 expression in RMICs subjected to mechanical stress; therefore, we investigated whether ROS may act via MAPK. To test this, we determined the effects of the antioxidants DPI and ROT on the activation of ERK1/2, p38, and JNK in RMICs exposed to H2O2. The activation of ERK1/2, p38, and JNK was determined by immunoblot analysis using phosphorylation-specific antibodies, as previously described. The results showed that exposure of RMICs to H2O2 increased the phosphorylation of both ERK1/2 and p38 compared with untreated cells (Fig. 4, A–D). The activation of both ERK1/2 and p38 by oxidative stress was attenuated by DPI and ROT, even though the regulation of p38 did not completely reach a significant level (P = 0.06 for DPI and P = 0.07 for ROT; Fig. 4, A–D). Notably, the abundance of total ERK1/2 and p38 as well as JNK and the active phosphorylated form of JNK was unchanged (Fig. 4, A–D).
To examine whether ROS-activated ERK1/2 and p38 pathways play a role in the induction of COX-2 in RMICs exposed to oxidative stress, RMICs were treated with H2O2 as well as inhibitors of ERK1/2 and p38 activation followed by an analysis of COX-2 and HO-1 expression. The immunoblot results showed that oxidative stress-induced COX-2 and HO-1 were attenuated by inhibition of the ERK1/2 and p38 pathway (Fig. 5, A and B). Taken together, these findings suggest a relationship among ROS, MAPK, and COX-2 in RMICs exposed to oxidative stress.
Effects of DPI and ROT on ROS Production and Apoptosis in RMICs Exposed to Oxidative Stress
Exposure of cells to H2O2 can induce both oxidative stress and apoptosis, depending on the cell type and H2O2 concentrations (30). H2O2-induced oxidative stress was examined by measuring ROS production and the oxidative stress marker HO-1. The effects of the antioxidants DPI and ROT were also investigated. As shown in Fig. 6A, treatment of RMICs with H2O2 (75 μM) for 6 h increased ROS production, which was significantly attenuated by the administration of both DPI and ROT. Similarly, the induction of HO-1 in RMICs upon H2O2 treatment was also attenuated by ROT and DPI (Fig. 6B). We found no significant increase in the Bax/Bcl-2 ratio between H2O2-exposed cells and control cells, nor did we observe any effect of either DPI or ROT (Fig. 6C). These findings suggest that the antioxidants DPI and ROT enhance the resistance of RMICs to oxidative stress.
Effects of COX-2 Inhibition on HO-1 Expression in RMICs Exposed to Oxidative Stress
It has previously been demonstrated that the ability of RMICs to tolerate H2O2 is dependent on COX-2 activity, since COX-2 inhibition with SC-58236 reduces cell viability and increases apoptosis (15). To test whether COX-2 activity play a role in the regulation of oxidative stress in RMICs, we treated RMICs with the selective COX-2 inhibitor SC-236 (5 μg) and measured the expression of HO-1. The results demonstrated that COX-2 inhibition increases H2O2-induced HO-1 expression (Fig. 7), indicating reduced cell resistance to oxidative stress.
Effects of DPI and ROT in the Regulation of Oxidative Stress and the MAPK Pathway in RMICs Exposed to Mechanical Stress
Tubule hydrostatic pressure increases in response to ureteral obstruction. To evaluate whether mechanical stress plays a role in the regulation of oxidative stress and the MAPK pathway in RMICs, we exposed cells to stretch and treated them with the antioxidants ROT and DPI. The results demonstrated that stretch for 2 h increases COX-2 and HO-1 expression in RMICs, indicating that mechanical stress plays a role in the regulation of COX-2 and oxidative stress. The administration of DPI attenuated the stretch-induced HO-1 expression (Fig. 8, A and B).
The MAPK pathway was studied by analyzing the activation of ERK1/2, p38, and JNK in RMICs exposed to stretch. Our data demonstrated increased ERK1/2 and p38 activity after 2 h of stretch, whereas the activation of JNK was unchanged (Fig. 8, A and B). The administration of ROT abolished the stretch-induced activation of both ERK1/2 and p38. In contrast, DPI treatment further increased the activation of ERK1/2 in RMICs exposed to stretch (Fig. 8, A and B). These findings suggest that mitochondrial respiration chain complex I and NADPH oxidase play a role for the upstream regulation of the MAPK signaling pathway in RMICs exposed to stretch.
In this study, we investigated, both in vivo and in vitro, whether ROS play a role in the regulation of COX-2 in RMICs in response to UUO. Our main findings demonstrated ROS as an upstream regulatory mediator of COX-2 expression and activity in RMICs. To interfere with ROS production, we used two mechanistically distinct antioxidants, namely, the NADPH oxidase inhibitor DPI and the complex I inhibitor ROT. Our results show that UUO-induced tubular dilatation and oxidative stress were attenuated by the administration of DPI and ROT in obstructed kidneys. The induction of COX-2 in the inner medulla was also effectively inhibited by both antioxidants in rats subjected to 3-day UUO and in RMICs exposed to oxidative stress. Using an in vitro model that induces mechanical stress to mimic the in vivo situation in response to UUO, we demonstrated that stretch stimulates COX-2 and HO-1 expression as well as ERK1/2 and p38 activation in RMICs. In addition, oxidative stress-induced phosphorylation of ERK1/2 and p38 was partly attenuated by antioxidant treatment, whereas inhibition of ERK1/2 and p38 blocked the induction of COX-2 in RMICs. Notably, COX-2 inhibitors also exacerbated the oxidative stress level in H2O2-exposed RMICs. These results suggest that oxidative stress stimulates COX-2 expression through activation of the MAPK pathway and that the induction of COX-2 may exert a cytoprotective function in RMICs.
The mitochondrial respiration chain and NADPH oxidase have been considered as key sources of ROS production. Here, our data show that COX-2 expression and activity were induced by ROS derived from both NADPH oxidase and complex 1 in the mitochondrial respiration chain in RMICs in the inner medulla of rats subjected to 3-day UUO as well as in cultured RMICs exposed to oxidative stress. These findings are consistent with previous in vitro studies demonstrating that mitochondria and NADPH oxidase increased ROS production and COX-2 expression in a variety of cells, such as human mesangial cells incubated with glucose (18) and collecting duct cells exposed to hypertonic treatment (34). In the present in vivo study, the link between oxidative stress and COX-2 induction in an experimental model of UUO was examined. We assume that increased UUO-associated oxidative stress may be an integral component of the mechanism involved in the induction of COX-2 in RMICs, even though we cannot exclude other manifestations of the obstruction state as contributing factors to the induction of COX-2. Nevertheless, a previous in vivo study (25) has demonstrated a similar effects of DPI in the attenuation of COX-2 induction during ischemia-reperfusion damage in rat stomachs. In addition, Li et al. (21) showed that chronic administration of the antioxidant tempol prevented increased renal expression of COX-2 in streptozotocin-induced diabetic rats, indicating that oxidative stress resulted in the induction of COX-2 in diabetes.
We and others (6, 15, 24, 27) have observed increased COX-2 expression in the renal inner medulla in response to ureteral obstruction. There is increasing evidence supporting a cytoprotective role of COX-2 in the renal medulla and increased COX-2 expression as a prerequisite for RMIC survival from hypertonic stress (13, 14, 33). Based on these studies, one might speculate that oxidative stress-induced COX-2 expression in response to UUO contributes to protection against oxidant injury. However, COX-2 has also been linked to renal damage in UUO models, in which the COX-2 inhibitor etodolac reduced renal tubular damage and apoptosis (23). Therefore, prevention of COX-2 induction with antioxidants can similarly be expected to ameliorate renal damage related to UUO. These findings reveal a counterbalanced intrarenal handling of COX-2, that is, the detrimental versus beneficial effects of COX-2 may depend on the pathophysiological condition, again highlighting the importance of the enzyme in renal physiology.
To explore the possible underlying mechanisms that link ROS and COX-2 in RMICs, the MAPK cascade was examined. MAPK pathways mediate the stimulatory effects of different extracellular stimuli on COX-2 expression in a stimulus- and cell type-specific manner (6, 34), and studies have also demonstrated that oxidative and mechanical stress can stimulate MAPK pathways in various cell types (1, 3, 10, 31). Here, we studied different classes of MAPKs, including ERK1/2, p38, and JNK, and our data showed that the antioxidants DPI and ROT partly attenuated oxidative stress-induced phosphorylation of both ERK1/2 and p38 in RMICs. These results indicate that ROS play a role in the activation of MAPK cascades, particularly the ERK1/2 and p38 pathway, in RMICs. In addition, we found that inhibition of ERK1/2 and p38 activation suppressed COX-2 expression in RMICs in response to oxidative stress, suggesting that ERK1/2 and p38 may function as downstream effectors of ROS to transduce signals for the induction of COX-2. This observation is consistent with a previous study (34) that identified a distinct role of the ROS/MAPK/COX-2 pathway in the osmotic response in collecting duct cells.
Growing evidence has demonstrated that MAPK cascades, especially ERK1/2 and p38, play roles in cytoprotection and oxidative stress in different cell types (3, 10, 20). Our present study shows that inhibition of ERK1/2 and p38 attenuated the increased expression of the oxidative stress marker HO-1, supporting that the MAPK pathway may protect against oxidant injury in RMICs. A previous study (32) has also demonstrated that COX-2 is critical for the capability of medullary epithelial cells to survive under hypertonic stress. Consistently, our present results show that the COX-2 inhibitor SC-236 increased oxidative stress-induced HO-1 expression in RMICs, indicating that COX-2 mediates protection against oxidant injury. Furthermore, it has been previously demonstrated that the ability of RMICs to tolerate H2O2 is dependent on COX-2 activity, since COX-2 inhibition with SC-58236 reduces cell viability and increases apoptosis (15). Taking together, our observations in RMICs suggest that mitochondria and NAPDH oxidase may contribute to H2O2-induced ROS production, which transduces the signal to MAPKs and leads to COX-2 activation and protection against oxidant injury. The NADPH oxidase subunits involved in the oxidative response in RMICs were not directly addressed in the present study. However, both p22phox and p47phox have been shown to be involved in the development of oxidative stress in a number of animal models of diseases with renal involvement, and these subunits are also expressed in the renal medulla (12) and might play a role in the oxidative response in RMICs as well.
The tubule pressure increases in response to UUO, leading to renal tubular distention and cell deformation. To mimic this in vitro, we exposed RMICs to mechanical stress and demonstrated that stretch induces COX-2 and HO-1 expression as well as ERK1/2 and p38 activation in RMICs, indicating that stretch play a role in the regulation of COX-2, oxidative stress, and the MAPK signaling pathway. As a part of the investigation into the signaling pathways linking mechanical stretch to ERK1/2 and p38 activation in RMICs, we explore the possibility that signaling mediated by mitochondrial respiration chain complex I- or NADPH oxidase-derived ROS production may be involved. Previous studies (4, 7) have demonstrated that ROS are associated with mechanical stress-induced phosphorylation of ERK1/2 and p38 in endothelial cells and vascular smooth muscle cells. We found that stretch induced the activation of ERK1/2 and p38 signaling involved complex I stimulation. The NADPH oxidase inhibitor DPI increased the phosphorylation of ERK1/2 in RMICs exposed to stretch, whereas p38 phosphorylation was not changed. These data support the concept of different regulation pathways for individual MAPKs.
In summary, this study shows, both in vivo and in vitro, that ROS/oxidative stress might play a role in the induction of COX-2 in RMICs of rats subjected to ureteral obstruction. The activation of MAPK pathways might be involved in signal transduction, leading to increased COX-2 expression in RMICs. Overall, we conclude that oxidative stress as a consequence of ureteral obstruction might stimulate COX-2 expression through activation of the MAPK cascade pathway and that the induction of COX-2 may exert a cytoprotective function in RMICs.
This work was supported by the Danish Research Council for Health and Disease, the Lundbeck Foundation, the Helen and Ejnar Bjørnow Foundation, the NOVO Nordisk Foundation, and the A. P. Møller Foundation.
No conflicts of interest, financial or otherwise, are declared by the author(s).
Author contributions: M.O., M.C., L.N., and I.C. performed experiments; M.O., M.C., L.N., I.C., and R.N. analyzed data; M.O., M.C., L.N., I.C., and R.N. interpreted results of experiments; M.O., M.C., L.N., I.C., and R.N. prepared figures; M.O., M.C., L.N., I.C., J.F., and R.N. drafted manuscript; M.O., M.C., L.N., I.C., J.F., and R.N. edited and revised manuscript; M.O., M.C., L.N., I.C., J.F., and R.N. approved final version of manuscript; L.N., J.F., and R.N. conception and design of research.
The authors thank Gitte Skou, Gitte Kall, and Line V. Nielsen for expert technical assistance.
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