1 Division of Nephrology, Departments of Medicine and 2 Molecular Physiology and Biophysics, Veterans Affairs Medical Center and Vanderbilt University School of Medicine, Nashville, Tennessee 37232-2372
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ABSTRACT |
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Renal medullary interstitial cells (MICs) are a major site of cyclooxygenase (COX)-mediated PG synthesis. These studies examined the role of COX in MIC survival. Immunoblot and nuclease protection demonstrate that cultured MICs constitutively express COX2, with little constitutive COX1 expression. SC-58236, a COX2-selective inhibitor, but not SC-58560, a COX1 inhibitor, preferentially blocks PGE2 synthesis in MICs. Transduction with a COX2 antisense adenovirus reduced MIC COX2 protein expression and also decreased PGE2 production. Antisense downregulation of COX2 was associated with MIC death, whereas a control adenovirus was without effect. Similarly, the COX2-selective inhibitor SC-58236 (30 µM) and several nonselective COX-inhibiting nonsteroidal anti-inflammatory drugs (NSAIDs), including sulindac, ibuprofen, and indomethacin, all caused MIC death. In contrast, SC-58560, a COX1-selective inhibitor, was 100-fold less potent for inducing MIC death than its structural congener SC-58236. NSAID-induced MIC death was associated with DNA laddering and nuclear fragmentation, consistent with apoptosis. These results suggest that COX2 plays an important role in MIC survival. COX2 inhibition may contribute to NSAID-associated injury of the renal medulla.
papillary necrosis; adenovirus; antisense; analgesic nephropathy
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INTRODUCTION |
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CYCLOOXYGENASE (COX) is a key enzyme regulating the formation of PGs from arachidonate. COXs are also major therapeutic targets for nonsteroidal anti-inflammatory drugs (NSAIDs; see Refs. 17, 18, 40). Two isoforms of COX, designated COX1 and COX2, have been identified (13, 15, 26, 38, 47). These derive from different genes but share ~60% amino acid identity (20, 26). The expression pattern of COX1 and COX2 genes is quite different. COX1 is constitutively expressed in several tissues and is thought to participate in housekeeping functions, including the maintenance of gastric epithelial integrity. Conversely, COX2 is undetectable in most tissues, but its expression can be induced by a variety of cytokines and growth factors (19, 21, 35, 44- 46). Constitutive expression of COX2 has recently been reported in certain tissues, including kidney, lung, and brain (10, 17, 23, 25). In the kidney, COX2 is constitutively expressed in cortical thick ascending limb and medullary interstitial cells (MICs), whereas COX1 mRNA appears to be mainly expressed in the collecting duct (17, 18). The significance of MIC COX2 expression remains uncertain.
NSAIDs are widely used because of their high efficacyand over-the-counter availability. However, NSAID abuse may cause serious kidney damage, which may be associated with papillary necrosis and progressive renal structural and functional deterioration (1, 29). The mechanism is poorly understood. Experimental studies suggest that renal MICs are an early target of injury in analgesic nephropathy (27), and a similar syndrome has been reported with chronic NSAID use (34). COX2 has been demonstrated to play an important role in cell proliferation and survival (5, 36, 37). Because NSAIDs inhibit COX, we considered the possibility that they interfere with MIC survival by inhibiting COX2. The present studies examine the role of COX2 in MIC survival.
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METHODS |
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Materials
Ibuprofen, acetaminophen, and phenacetin were purchased from Sigma (St. Louis, MO). The COX1 inhibitor SC-58560 and COX2 inhibitor SC-58236 were generously provided by Drs. Peter Isakson and Karen Siebert, Monsanto/Searle Research and Development (St. Louis, MO). These compounds were dissolved in DMSO. When applied to cultured cells, the concentration of DMSO was <0.1% (vol/vol).Cell Culture
Rabbit MICs were cultured by the method of Muirhead et al. (28). Briefly, female New Zealand White rabbits were anesthetized (44 mg/kg ketamine and 10 mg/kg xylazine im). The left kidney was removed, and the medulla was dissected and minced with a razor blade under sterile conditions, in 5 ml of sterile RPMI 1640 plus 10% (vol/vol) FBS (Hyclone, Logan, Utah). This homogenate was injected subcutaneously in the abdominal wall using a 14-gauge needle. Twenty days postsurgery, subcutaneous nodules appeared. The rabbits were reanesthetized and killed by decapitation, and the nodules were removed under sterile conditions. The nodules were minced into 1-mm fragments and explanted in 75-cm2 circular tissue culture plates. Cells were cultured in RPMI 1640 tissue culture medium supplemented with 10% (vol/vol) FBS and streptomycin and penicillin. Cultures were incubated at 37°C in 95% O2-5% CO2. Tissue culture medium was changed every 48-72 h. These cells display characteristic abundant oil red O positive lipid droplets (17), which typifies type I MICs (28, 49). Cells were typically studied in their third to fourth passages. After MICs achieved confluence, and just before experimentation, the 10% serum medium was replaced with 0.5% serum for 24-48 h. Interstitial cells were then treated with PMA (100 nM) for 4 h or were treated with NSAIDs for 48 h.Rabbit collecting duct cells were immunodissected using a monoclonal antibody and were cultured as previously described (30). Female New Zealand White rabbits were killed as above. The kidney was perfused with Krebs-Ringer and harvested. The renal cortex was separated from the capsule and medulla via gross dissection and was passed through a tissue press. The dispersed tissue was digested with collagenase (0.1%), hyaluronidase (0.1%), DNase (100 U/ml), and soybean trypsin inhibitor (1,000 U/ml) in Krebs-Ringer at 37°C. This suspension was then poured over plates precoated with a monoclonal antibody specific for rabbit collecting duct (3G10) and was incubated at room temperature for 10 min. Nonadherent cells were removed by washing with PBS-BSA (1% BSA in PBS). The adherent cells were collected by scraping and cultured in DMEM plus 10% FBS.
Immunoblotting
Cultured MICs or collecting ducts were washed with PBS and harvested in SDS-PAGE lysis buffer (0.125 M Tris · HCl, pH 6.8, 4% SDS, and 20% glycerol) followed by repetitive aspiration using a 25-gauge needle. The lysate was heated in boiling water for 2 min. Protein concentration was determined by bicinchoninic acid protein assay (Sigma). Thirty micrograms of each protein extract were loaded in each lane of a 10% SDS-PAGE minigel and run at 120 V. The proteins were transferred to a nitrocellulose membrane at 22 V overnight at 4°C. The membrane was washed three times with 50 mM Tris, pH 7.5, 150 mM NaCl, and 0.05% Tween 20 and then incubated in blocking buffer (150 mM NaCl, 50 mM Tris, 0.05% Tween 20, and 5% Carnation nonfat dry milk, pH 7.5) for 1 h at room temperature. The membrane was then incubated with an anti-human COX2 (1:300, C-20; Santa Cruz Biotechnology, Santa Cruz, CA) or an anti-COX1 (1:300, C-20, Santa Cruz) antibody in blocking buffer overnight at 4°C. After being washed (3 times), the membrane was incubated with a horseradish peroxidase-conjugated secondary antibody (1:20,000; Jackson Immuno-Research Laboratories) for 1 h at room temperature, followed by three 15-min washings. Antibody labeling was visualized by addition of chemiluminescence reagent (Renaissance; DuPont NEN, Boston, MA), and the membrane was exposed to Kodak XAR-5 film.For immunoprecipitation of COX, 3 ml of ice-cold RIPA buffer (PBS, 0.1% SDS, 0.5% sodium deoxycholate, and 1% Nonidet P-40) were added to the cell culture and incubated at 4°C for 10 min. Cells were disrupted by repeated aspiration through a 25-gauge needle. Cellular debris was pelleted by centrifugation at 3,000 g at 4°C for 15 min. Supernatant was transferred to a new centrifuge tube, and the lysate was precleared by mixing with 20 µl of agarose conjugate, which was then pelleted by centrifugation at 2,500 g at 4°C for 5 min. Supernatant (1 ml cell lysate) was transferred to another centrifuge tube at 4°C. Ten micrograms of anti-human COX2 antibody (Santa Cruz C-20) were added to the lysate and incubated for 1 h at 4°C. Twenty microliters of agarose conjugate (Protein G PLUS) were added and incubated at 4°C on a rotating device overnight. Immunoprecipitate was collected by centrifugation at 2,500 g for 5 min at 4°C. The pellet was washed four times with 1.0 ml RIPA buffer. After the final wash, the supernatant was aspirated and discarded. The pellet was resuspended in 40 µl of electrophoresis sample buffer. Samples were boiled for 1 min and analyzed by SDS-PAGE as described above. An anti-human COX2 antibody (PG27 1:1,000; Oxford Biomedical Research, Oxford, MI) and anti-human COX1 antibody (C-20, 1:300; Santa Cruz) were used for immunoblotting.
Nuclease Protection
A 387-bp riboprobe for nuclease protection was generated from a portion of the 3' untranslated region of the full-length COX2 cDNA obtained by subcloning a Rsa I digestion fragment into pBluescript SKConstruction of AdrCOX2-AS and Ad-GL
The full-length rabbit COX2 cDNA (17), a green fluorescent protein cDNA (Green Lantern or GL; GIBCO Life Technologies, Gaithersburg, MD), was used to construct two adenoviral vectors: AdrCOX2-AS and Ad-GL. GL was subcloned from pGL into pBluescript II (Stratagene, La Jolla, CA). The GL fragment was excised with Kpn I and Xba I and subcloned into the Kpn I/Xba I sites of shuttle plasmid pACCMV, yielding the sense GL construct. For construction of a COX2 antisense adenovirus (AdrCOX2-AS), the full-length rabbit COX2 cDNA fragment was excised from pcDNA3.1 using Hind III and Xba I and subcloned into the Xba I/Hin III sites of pACCMV (2), yielding a COX2 antisense transcript. The pACCMV shuttle plasmid contains the cytomegalovirus immediate early enhancer and promoter and the SV40 polyadenylation sequence. The resulting shuttle plasmids containing sense GL or antisense COX2 inserts were cotransfected into HEK 293 cells with the pJM17 vector by standard calcium phosphate coprecipitation methods (2). AdrCOX2-AS and Ad-GL generated by a homologous recombination event resulted in plaque formation in the HEK-293 cells. The resulting infectious adenovirus was plaque purified. For infection of MICs, 200 µl of virus [106 plaque-forming units (pfu)/cm2] were added to each culture dish. After 2 h of incubation, 2 ml of DMEM with 10% FBS were added, without removing the virus. Twelve hours later, the medium was replaced.Cell Viability and Apoptosis
Cell viability was assessed by the following methods.Method 1. Cell death was assessed using trypan blue exclusion. All floating and attached cells were pelleted (after trypsinization to detach adherent cells) and resuspended in culture medium loaded with 0.2% trypan blue. The percentage of dead cells was determined by counting total and blue- stained cells (dead) with a hemocytometer.
Method 2. This method involved
propidium iodide exclusion (3, 33). Cell viability was assessed using
spectrofluorometry to measure propidium iodide uptake. Propidium iodide
is impermeable to living cell membranes and was added to cell culture
dishes, yielding a final concentration of 50 µM. After 20 min,
fluorescence (F) at excitation = 530 nm and emission
= 620 nm
was measured using a Cytofluor II (PerSeptive BioSystems, Framingham,
MA). Digitonin (4 µM) was then added, and, after an additional 20 min, fluorescence was again determined [maximal fluorescence
(Fmax)]. Background
fluorescence (blank) was determined from cell-free wells containing 1.5 ml of 50 µM propidium iodide. Percent cell viability was estimated by
the formula 100
[(F
blank)/(Fmax
blank)] × 100.
Method 3. Apoptosis was assessed by DNA laddering and nuclear fragmentation. DNA laddering was assessed after agarose gel electrophoresis of genomic DNA isolated using an anion-exchange resin column (QIAGEN, Chatsworth, CA). Ten microliters of DNA were resolved on 1% agarose gels and visualized by ultraviolet transillumination. Nuclear fragmentation was assessed by staining nuclear DNA using Hoechst 33342. Floating and attached cells were pelleted and washed with PBS. Cells were fixed in 3% paraformaldehyde in PBS for 10 min at room temperature. Cells were washed with PBS and resuspended in 15 µl of PBS containing 16 µg Hoechst 33342 for 15 min at room temperature. A 10-µl aliquot was placed on a glass slide, and 1,000 cells/slide were scored for the incidence of nuclear fragmentation using a fluorescence microscope. Cells with three or more chromatin fragments were considered apoptotic (43).
PGE2 Measurement
PGE2 production by cultured MICs or cultured collecting ducts (CCDs) was determined by enzyme immunoassay (PGE2 enzyme immunoassay kit; Oxford Biomedical Research, Oxford, MI). The activity of COX1- and COX2-selective NSAIDs on COX activity was determined in CCDs and MICs, respectively. COX2 activity was enhanced by treating MIC cultures with phorbol myristate acetate (PMA, 100 nM) as previously described (8, 17, 32). After 4 h of incubation with PMA and COX inhibitors, MIC supernatant was collected for determination of PGE2 measurement. In studies of COX1 activity in collecting ducts, CCDs were first incubated with COX inhibitors for 4 h, and then medium was removed. PGE production was assessed after the addition of 10 µM arachidonic acid for 1 h at 37°C as previously described (8, 11, 32). IC50 values were calculated from a nonlinear regression plot using a computerized program (Prism; ISI software). ![]() |
RESULTS |
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COX Isoform Expression in Cultured Rabbit MICs
COX-selective antibodies immunoprecipitated COX2 but not COX1 from cultured rabbit MICs, whereas COX1 but not COX2 was the predominant isoform immunoprecipitated from CCDs (Fig. 1).1 This differential expression of COX1 and COX2 in cultured MICs versus collecting ducts was confirmed by RNase protection using riboprobes selective for COX1 and COX2 (Fig. 1 and Ref. 17). Treatment with PMA (100 nM, 4 h) markedly increased COX2 immunoreactivity but had no effect on COX1 expression in MICs. Furthermore, although faint COX2 immunoreactivity was detected in collecting ducts (Fig. 1A), phorbol esters did not significantly induce its expression in these cells.
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COX1- and COX2-selective inhibitors also differentially inhibited
PGE2 production in cultured MICs
and CCDs. The COX1 inhibitor SC-58560 is ~10,000-fold more potent in
inhibiting PGE2 synthesis in CCDs
(IC50: 1.6 × 1012 M) than in MICs
(IC50: 2 × 10
8 M; Fig.
2). Conversely, the COX2 inhibitor SC-58236
is ~106-fold more potent in MICs
(IC50: 5.7 × 10
11 M) than in CCDs
(IC50: 2.7 × 10
5 M). This supports the
above findings that COX2 is the major isoform in MICs, whereas COX1 is
the major isoenzyme in collecting ducts. Even at the highest NSAID
concentration tested, small amounts of
PGE2 production were still
detected in MICs. Acetaminophen (1 mM) failed to inhibit
PGE2 synthesis in either cell
type.
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COX2 Antisense Adenovirus Downregulates COX2 Expression and Kills MICs
The role of COX2 in MIC survival was examined using a rabbit COX2 antisense adenoviral vector (AdrCOX2-AS). Adenoviral transduction of cultured MICs was highly efficient, with >85-95% of the cells expressing green fluorescent protein (data not shown). This was substantially more efficient than liposome-mediated transfection, which yielded an efficiency of <5%. Fluorescent intensity of the reporter green fluorescent protein, Green Lantern, plateaued 4-7 days after infection and remained stable for >20 days. At the viral dose used (106 pfu/cm2), Ad-GL-treated MIC did not exhibit any morphological changes or changes in COX immunoreactivity.Immunodetectable COX2 was markedly reduced 48 h after transduction of
MICs with AdrCOX2-AS compared with COX2 expression in Ad-GL-infected
cells (Fig.
3A).
Furthermore, PMA (100 nM, 4 h) induction of COX2 immunoreactivity was
blocked by AdrCOX2-AS. Similarly, induction of
PGE2 synthesis by PMA was blocked
by AdrCOX2-AS but not with Ad-GL (Fig.
3B).
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The effect of adenoviral transduction on MIC viability was determined
by propidium iodide uptake. Treatment of confluent monolayers of MICs
with AdrCOX2-AS for 72 h significantly increased MIC death compared
with treatment with the control adenovirus, Ad-GL (Fig. 4). This effect was dose dependent, and
increasing viral concentration was associated with significantly more
MIC death. More than 54% of the MICs died when treated with AdrCOX2-AS
(2.5 × 106
pfu/cm2) versus only 16% cell death after treatment with a
comparable dose of Ad-GL.
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Effects of NSAIDs on MICs
The effect of NSAIDs on MIC viability was assessed. Treatment of MICs for 48 h with ibuprofen, sulindac sulfide, indomethacin, aspirin, SC-58236 (COX2 inhibitor), or SC-58560 (COX1 inhibitor) induced cell death in a concentration-dependent manner. The minimal lethal NSAID concentration, as assessed by propidium iodide or trypan blue exclusion assay, varied considerably for these different COX inhibitors: for SC-58236 it was 30 µM, sulindac 40 µM, ibuprofen 100 µM, and indomethacin 500 µM. The COX1 inhibitor SC-58560 and relative COX1-selective inhibitor aspirin failed to cause MIC death until concentrations equaled or exceeded 3,000 µM. The analgesics acetaminophen and phenacetin also failed to cause MIC cell death at concentrations up to 1,000 µM (Figs. 5 and 6).
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Treatment of MICs with NSAIDs induced apoptosis. Agarose gel electrophoresis of genomic DNA extracted from ibuprofen-treated (100 µM, 48 h) MICs exhibited distinctive "laddering" (Fig. 6A). Morphological studies of cells stained with Hoechst 33342 demonstrated fragmentation of nuclear DNA, also consistent with apoptosis (Fig. 6, B and C). Nuclear fragmentation was also observed in AdrCOX2-AS-, SC-58236-, and indomethacin-treated MICs as assessed by Hoechst 33342 or acridine orange staining (data not shown).
Effect of NSAIDs on COX Expression in MICs
Treatment of cultured MICs with SC-58236 (20-100 µM) or ibuprofen (50-500 µM) for 48 h increased the amount of COX2 detected by immunoblot. In contrast, neither COX1 nor COX2 expression was affected by this dose of the COX1-selective inhibitor SC-58560 (100 µM, Fig. 7).
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DISCUSSION |
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The renal medulla displays one of the highest rates of endogenous COX-mediated PG synthesis. The medulla is comprised of three cellular compartments: vascular cells, epithelial cells, and interstitial cells. The predominant interstitial cells are lipid droplet-rich type I MICs (28, 49). MICs are relatively unique in that they constitutively express COX2 in vivo (17). In contrast, COX1 appears to be predominately expressed in collecting ducts (18). We now show that MICs also retain selective COX2 expression in culture, whereas primary cultures of collecting ducts predominantly express COX1. These results contrast with studies of transformed collecting duct cell lines (e.g., M1 cells) in which COX2 is detected (48). These differences may be due to the heterologous expression of the SV40 large-T antigen in these cell lines (41), which may drive COX2 expression. The significance of this constitutive and selective expression of COX2 in MICs remains uncertain. These studies utilized cultured MIC to examine the potential role of COX2 in MIC survival.
The present results provide two independent lines of evidence pointing to a role for COX2 in MIC survival: one provided by antisense studies and one using COX1- and COX2-selective inhibitors. Inhibition of COX2 expression using an antisense adenovirus dose-dependently killed MICs. COX2 downregulation was confirmed by reduction of both basal and phorbol ester-stimulated COX2 immunoreactivity and PGE2 synthesis, effects not seen using a control (green fluorescent protein) adenovirus. Cell death does not seem to be the result of adenovirus infection per se, since a similar dose of adenovirus encoding green fluorescent protein did not kill these cultured interstitial cells. This suggests that the lethal effects of COX2 antisense are via COX2 downregulation.
A second line of evidence suggesting that COX2 plays an important role in MIC survival is provided by studies using COX2- and COX1-selective inhibitors. The COX2 inhibitor SC-58236 and the COX1 inhibitor SC-58560 are thought to act like other NSAIDs occupying the substrate channel within COX, thereby blocking substrate access to the active site (16, 22). These inhibitors are structurally related but display >1,000-fold difference in potency for inhibiting COX2 (31, 39). Similar selectivity of these COX2 and COX1 inhibitors was seen for inhibition of PGE2 synthesis by cultured rabbit MICs, where the COX2 inhibitor SC-58236 was 100-fold more potent than SC-58560. This finding is consistent with the predominant expression of COX2 in MICs. Treatment of MICs with a COX2-selective inhibitor, as well as several structurally dissimilar nonselective COX-inhibiting NSAIDs, induced apoptosis of MICs. Lethal effects of NSAIDs have been previously reported in cancer cells, where COX2 may play a role promoting resistance to apoptosis (43). This is in contrast to the mouse intestinal crypt stem cells, where COX1 but not COX2 appears to play a role in survival after radiation injury (9). The present studies now suggest a role of COX2 for survival of these nontransformed MICs.
Interestingly, the relative potency of these nonselective NSAIDs in causing MIC death correlates well with their relative potency in blocking rabbit MIC PGE2 production (49), with a rank order of relative potency of ibuprofen > indomethacin > aspirin. Importantly, the present studies also show that a structurally related COX1 inhibitor was markedly less potent in causing MIC death than the COX2 inhibitor. The functional activity of the COX1 inhibitor was confirmed by studies showing it is a highly potent inhibitor of PGE2 production in CCDs where COX1 is the predominant isoform. This correlation between the potency of these structurally dissimilar compounds for inhibiting MIC PGE2 production and causing MIC death suggests a link between their lethal effect and COX inhibition. This latter finding complements the antisense studies and points to a specific role for COX2 function in maintaining MIC viability.
The mechanism by which COX2 contributes to MIC viability is unclear. The present results do not suggest a role for expression of the COX2 protein per se, since COX2 inhibitors increased protein expression, whereas antisense adenovirus decreased protein expression, yet both resulted in cell death. These results instead suggest that COX2 function is an important factor in maintaining MIC viability.
COX inhibition could result in cell death either by reducing PG
synthesis or by elevating levels of free arachidonic acid and shunting
it to another pathway (6, 42). As previously observed in other cells,
the concentrations of NSAIDs required to induce apoptosis were higher
than those required to maximally inhibit the appearance of
PGE2 in the culture media.
Nevertheless, even at the highest NSAID concentration tested, a small
amount of PGE2 production was
still detected. It may be that intracellular levels of PGs remain high
enough to maintain MIC viability until the NSAID concentrations are
well above the IC50 values. In
this regard, the existence of nuclear PG receptors, including
peroxisome proliferator-activated receptor- for
PGD2,
PGJ2, and
PGI2 (14, 24) and
PGE2 EP1 receptor (4), supports a
role for intracellular PGs. MICs produce several PGs in addition to
PGE2 (49), and the complete
profile of these compounds, which might contribute to MIC viability,
remains to be determined. Alternatively, COX2 inhibition might reduce
MIC survival by increasing free cellular arachidonate, which, in turn,
stimulates the conversion of sphingomyelin to ceramide, a known
mediator of apoptosis (6).
Prolonged NSAID use is associated with necrosis of the renal papillae (34), and analgesic nephropathy is associated with early drop out of type I interstitial cells, suggesting that MICs may be particularly sensitive to this form of toxic injury (27). The present studies show that NSAIDs can directly induce apoptosis in renal MICs. In contrast, phenacetin and acetaminophen, which are typically associated with analgesic nephropathy, did not directly cause cell death in cultured MICs. In this regard, their combination with COX-inhibiting concentrations of aspirin may be crucial for their toxic effect (12).
In conclusion, COX2 is the predominant isoform constitutively expressed in cultured renal MICs. Both antisense downregulation of COX2 and COX2 (but not COX1)-selective inhibitors induced cell death in these cells. These results suggest that COX2 is important for survival of MICs. By blocking COX2 activity, prolonged NSAID use could directly injure interstitial cells of the renal medulla.
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ACKNOWLEDGEMENTS |
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We thank Drs. Karen Siebert and Peter Isakson from Monsanto/Searle who generously provided the SC-58236 and SC-58560 used in these studies.
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FOOTNOTES |
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M. D. Breyer is the recipient of a Veterans Affairs Career Development award. This study was supported by National Institute of Diabetes and Digestive and Kidney Diseases Grants 1-P50-DK-39261 and DK-37097, the George M. O'Brien Kidney Center, and by funding from Searle/Monsanto Pharmaceuticals. M. Kömhoff is a recipient of a research grant (KO 1855/1-1) from the Deutsche Forschungsgemeinschaft.
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. §1734 solely to indicate this fact.
1 Several commercial and proprietary anti-COX antibodies were tested by our laboratory for specificity toward rabbit COX1 and COX2. No antibody tested was absolutely specific. In the present study, we used two anti-human COX2 antibodies for immunoblots (C-20 from Santa Cruz Biotechnology and PG27 from Oxford Biomedical Research). Preliminary studies show that anti-COX2 antibody from Santa Cruz Biotechnology (C-20) binds to rabbit COX2, but it cross-reacts with rabbit COX1. The antibody from Oxford (PG27) binds only to COX2 and not to COX1, but it displays several nonspecific bands on immunoblot. To circumvent these problems in specificity, we sequentially used Santa Cruz anti-COX2 antibody (C-20) to immunoprecipitate both COX1 and COX2 first and then probed COX2 with Oxford PG27 on an immunoblot (Fig. 1). COX2 immunoreactivity was only detected in MICs. Only faint COX2 could be detected in CCD. PMA treatment induced COX2 expression in MICs but not in CCDs. We then stripped the membrane and reprobed with COX1-specific antibody (Santa Cruz C-20); abundant COX1 immunoreactivity was present in CCDs and only faintly detected in MICs. PMA had no effect on COX1 expression (Fig. 1). In later studies, we took advantage of the fact that COX2 is the predominant isoform expressed in MICs, using only the anti-COX2 antibody from Santa Cruz (C-20) for immunoblotting to examine the changes in COX2. We confirmed that these changes were COX2 and not COX1 by reprobing the same membrane with COX1 antibody as shown in Figs. 3 and 7.
Address for reprint requests and other correspondence: M. D. Breyer, F427 ACRE Bldg., Veterans Affairs Medical Center, 1310 24th Ave. S., Nashville, TN 37212 (E-mail: Matthew.Breyer{at}mcmail.Vanderbilt.edu).
Received 4 February 1999; accepted in final form 12 May 1999.
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REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1.
Barrett, B. J.
Acetaminophen and adverse chrnic renal outcomes: an appraisal of the epidemiologic evidence.
Am. J. Kidney Dis.
28:
S14-S19,
1996[Medline].
2.
Becker, T.,
R. Noel,
W. Coats,
A. Gomez-Foix,
T. Alam,
R. Gerard,
and
C. Newgard.
Use of Recombinant Adenovirus for metabolic engineering of mammalian cells.
In: Methods in Cell Biology. New York: Academic, 1994, p. 161-189.
3.
Beletsky, I. P.,
and
S. R. Umansky.
A new assay for cell death.
J. Immunol. Methods
134:
201-205,
1990[Medline].
4.
Bhattacharya, M.,
K. G. Peri,
G. Almazan,
A. Ribeiro-da-Silva,
H. Shichi,
Y. Durocher,
M. Abramovitz,
X. Hou,
D. R. Varma,
and
S. Chemtob.
Nuclear localization of prostaglandin E2 receptors.
Proc. Natl. Acad. Sci. USA
95:
15792-15797,
1998
5.
Castano, E.,
M. Dalmau,
M. Marti,
F. Berrocal,
R. Bartrons,
and
J. Gil.
Inhibition of DNA synthesis by aspirin in Swiss 3T3 fibroblasts.
J. Pharmacol. Exp. Ther.
280:
366-372,
1997
6.
Chan, T. A.,
P. J. Morin,
B. Vogelstein,
and
K. W. Kinzler.
Mechanisms underlying nonsteroidal antiinflammatory drug-mediated apoptosis.
Proc. Natl. Acad. Sci. USA
95:
681-686,
1998
7.
Chomczynski, P.,
and
N. Sacchi.
Single-step method of RNA isolation by acid guanidinium thiocyanate-phenol chloroform extraction.
Anal. Biochem.
162:
126-159,
1987.
8.
Chulada, P.,
C. Loftin,
V. Winn,
D. Young,
H. Riano,
T. Eling,
and
R. Langenback.
Relative activities of retrovirally expressed murine prostaglandin synthase-1 and -2 depend on source of arachidonic acid.
Arch. Biochem. Biophys.
1996:
301-313,
1996.
9.
Cohn, S.,
S. Schloemann,
T. Tessner,
K. Seibert,
and
W. Stenson.
Crypt stem cell survival in the mouse intestinal epithelium is regulated by prostaglandins synthesized through cyclooxygenase-1.
J. Clin. Invest.
99:
1367-1379,
1997
10.
Crofford, L. J.
COX-1 and COX-2 tissue expression: implications and predictions.
J. Rheumatol.
4 Suppl. 49:
15-9,
1997.
11.
Currie, M.,
B. Cole,
K. DeSchryver-Kecskemeti,
S. Holmberg,
and
P. Needleman.
Cell culture of renal epithelium derived from rabbit microdissected cortical collecting tubules.
Am. J. Physiol.
244 (Renal Fluid Electrolyte Physiol. 13):
F724-F728,
1983[Medline].
12.
DeBroe, M.,
and
M. Elseviers.
Analgesic Nephropathy.
N. Engl. J. Med.
338:
446-452,
1998
13.
DeWitt, D.,
and
W. Smith.
Primary structure of prostaglandin G/H synthase from sheep vesicular gland determined from the complementary DNA sequence.
Proc. Natl. Acad. Sci. USA
85:
1412-1416,
1988[Abstract].
14.
Forman, B.,
P. Tontonoz,
J. Chen,
R. Brun,
B. Spiegelman,
and
R. Evans.
15-deoxy-12,14-Prostaglandin J2 is a ligand for the adipocyte determination factor PPAR-gamma.
Cell
83:
803-812,
1995[Medline].
15.
Funk, C.,
L. Funk,
M. Kennedy,
A. Pong,
and
G. Fitzgerald.
Human platelet/erythroleukemia cell prostaglandin G/H synthase: cDNA cloning, expression, and gene chromosomal assignment.
FASEB J.
5:
2304-2312,
1991
16.
Gierse, J.,
J. McDonald,
S. Hauser,
S. Rangwala,
C. Koboldt,
and
K. Seiber.
A single amino acid difference between cyclooxygenase-1 (COX-1) and -2 (COX-2) reverses the selectivity of COX-2 specific inhibitors.
J. Biol. Chem.
271:
15810-15814,
1996
17.
Guan, Y.,
M. Chang,
W. Cho,
Y. Zhang,
R. Redha,
L. Davis,
S. Chang,
R. N. Dubois,
C. Hao,
and
M. D. Breyer.
Cloning, expression, and regulation of rabbit cyclooxygenase-2 in renal medullary interstitial cells.
Am. J. Physiol.
273 (Renal Physiol. 42):
F18-F26,
1997
18.
Harris, R.,
J. McKanna,
Y. Akai,
H. Jacobson,
R. Dubois,
and
M. Breyer.
Cyclooxygenase-2 is associated with the macula densa of rat kidney and increases with salt restriction.
J. Clin. Invest.
94:
2504-2510,
1994[Medline].
19.
Herschman, H. R.
Prostaglandin synthase 2.
Biochim. Biophys. Acta
1299:
125-140,
1996[Medline].
20.
Hla, T.,
and
T. Maciag.
Cyclooxyenase gene expression is down-regulated by heparin-binding (acidic fibroblast) growth factor-1 in human endothelial cells.
J. Biol. Chem.
266:
24059-24063,
1991
21.
Hla, T.,
and
K. Neilson.
Human cyclooxygenase-2 cDNA.
Proc. Natl. Acad. Sci. USA
89:
7384-7388,
1992[Abstract].
22.
Kalgutkar, A.,
B. Crews,
S. Rowlinson,
C. Garner,
K. Seibert,
and
L. Marnett.
Aspirin-like molecules that covalently inactivate cyclooxygenase-2.
Science
280:
1268-1271,
1998
23.
Kaufmann, W.,
P. Worley,
J. Pegg,
M. Bremer,
and
P. Isakson.
Cox-2, a synaptically induced enzyme, is expressed by excitatory neurons at postsynaptic sites in rat cerebral cortex.
Proc. Natl. Acad. Sci. USA
93:
2317-2321,
1996
24.
Kliewer, S.,
S. Sundseth,
S. Jones,
P. Brown,
G. Weisely,
C. Koble,
P. Dvchand,
W. Wahli,
T. Willson,
J. Lenhard,
and
J. Lehmann.
Fatty acids and eicosanoids regulate gene expression through direct interactions with peroxisome proliferator-activated receptors and
.
Proc. Natl. Acad. Sci. USA
1997:
4318-4323,
1997.
25.
Komhoff, M.,
H. Grone,
T. Klein,
H. Seyberth,
and
R. Nusing.
Localization of cyclooxygenase-1 and -2 in adult and fetal human kidney: implication for renal function.
Am. J. Physiol.
272 (Renal Physiol. 41):
F460-F468,
1997
26.
Kujubu, D.,
B. Fletcher,
B. Varnum,
R. Lim,
and
H. Herschman.
TIS10, a phorbol ester tumor promoter-inducible mRNA from Swiss 3T3 cells, encodes a novel prostaglandin synthgase/cyclooxygenase homologue.
J. Biol. Chem.
266:
12866-12872,
1991
27.
Molland, E. A.
Experimental renal papillary necrosis.
Kidney Int.
13:
5-14,
1978[Medline].
28.
Muirhead, E.,
B. Brooks,
J. Pitcock,
and
P. Stephenson.
Renomedullary antihypertensive function in accelerated (malignant) hypertension: observations on renomedullary interstitial cells.
J. Clin. Invest.
51:
181-190,
1972[Medline].
29.
Murray, M. D.,
and
D. C. Brater.
Renal toxicity of the nonsteriodal anti-inflammatory drugs.
Annu. Rev. Pharmacol. Toxicol.
32:
435-65,
1993.
30.
Noland, T.,
C. Carter,
H. R. Jacobson,
and
M. Breyer.
PGE2 regulates cAMP generation in cultured rabbit cortical collecting duct cells: evidence for dual inhibitory pathways.
Am. J. Physiol.
263 (Cell Physiol. 32):
C1208-C1216,
1992
31.
Penning, T. D.,
J. J. Talley,
S. R. Bertenshaw,
J. S. Carter,
P. W. Collins,
S. Docter,
M. J. Graneto,
L. F. Lee,
J. W. Malecha,
J. M. Miyashiro,
R. S. Rogers,
D. J. Rogier,
S. S. Yu,
G. D. Anderson,
E. G. Burton,
J. N. Cogburn,
S. A. Gregory,
C. M. Koboldt,
W. E. Perkins,
K. Seibert,
A. W. Veenhuizen,
Y. Y. Zhang,
and
P. C. Isakson.
Synthesis and biological evaluation of the 1,5-diarylpyrazole class of cyclooxygenase-2 inhibitors: identification of 4-[5-(4-methylphenyl)-3- (trifluoromethyl)-1H-pyrazol-1-yl]benze nesulfonamide (SC-58635, celecoxib).
J. Med. Chem.
40:
1347-1365,
1997[Medline].
32.
Reddy, S.,
and
H. Herschman.
Ligand-induced prostaglandin synthesis requires expression of the TIS10/PGS-2 prostaglandin synthase gene in murine fiborlbasts and macrophages.
J. Biol. Chem.
269:
15473-15480,
1994
33.
Sarafian, T. A.,
L. Vartavarian,
D. J. Kane,
D. E. Bredesen,
and
M. A. Verity.
bel-2 expression decreases methyl mercury-induced free-radical generation and cell killing in a neural cell line.
Toxicol. Lett.
74:
149-155,
1994[Medline].
34.
Segasothy, M.,
S. A. Samad,
A. Zulfigar,
and
W. M. Bennett.
Chronic renal disease and papillary necrosis associated with the long-term use of nonsteriodal anti-inflammatory drugs as the sole or predominant analgesic.
Am. J. Kidney Dis.
24:
17-24,
1994[Medline].
35.
Sheng, G. G.,
J. Shao,
H. Sheng,
E. B. Hooton,
P. C. Isakson,
J. D. Morrow,
R. J. Coffey, Jr.,
R. N. DuBois,
and
R. D. Beauchamp.
A selective cyclooxygenase 2 inhibitor suppresses the growth of H-ras- transformed rat intestinal epithelial cells.
Gastroenterology
113:
1883-1891,
1997[Medline].
36.
Sheng, H.,
J. Shao,
J. D. Morrow,
R. D. Beauchamp,
and
R. N. DuBois.
Modulation of apoptosis and Bcl-2 expression by prostaglandin E2 in human colon cancer cells.
Cancer Res.
58:
362-366,
1998[Abstract].
37.
Shiff, S. J.,
L. Qiao,
L.-L. Tsai,
and
B. Rigas.
Sulindac sulfide, an aspirin-like compound, inhibits proliferation, cause cell cycle quiescence, and induces apoptosis in HT-29 colon adenocarcinoma cells.
J. Clin. Invest.
96:
491-503,
1995[Medline].
38.
Simmons, D.,
W. Xie,
J. Chipman,
and
G. Evett.
Multiple cyclooxygenase: cloning of a mitogen-inducible form.
In: Prostaglandins, Leukotrienes, Lipoxins, and PAF, edited by J. Mailey. New York: Plenum, 1991, p. 67-78.
39.
Smith, C. J.,
Y. Zhang,
C. M. Koboldt,
J. Muhammad,
B. S. Zweifel,
A. Shaffer,
J. J. Talley,
J. L. Masferrer,
K. Seibert,
and
P. C. Isakson.
Pharmacological analysis of cyclooxygenase-1 in inflammation.
Proc. Natl. Acad. Sci. USA
95:
13313-13318,
1998
40.
Smith, W.,
and
G. Wilkin.
Immunochemistry of prostaglandin endoperoxide-forming cyclooxygenases: the detection of the cyclooxygenases in rat, rabbit and guinea pig kidneys by immnofluoresence.
Prostaglandins
13:
873-892,
1977[Medline].
41.
Stoos, B. A.,
A. Naray-Fejes-Toth,
O. A. Carretero,
S. Ito,
and
G. Fejes-Toth.
Characterization of a mouse cortical collecting duct cell line.
Kidney Int.
39:
1168-1175,
1991[Medline].
42.
Tang, D. G.,
Y. Q. Chen,
and
K. V. Honn.
Arachidonate lipoxygenases as essential regulators of cell survival and apoptosis.
Proc. Natl. Acad. Sci. USA
93:
5241-5246,
1996
43.
Tsujii, M.,
and
R. DuBois.
Alterations in cellular adhesion and apoptosis in epithelial cells overexpressing prostaglandin endoperoxide synthase 2.
Cell
83:
493-501,
1995[Medline].
44.
Tsujii, M.,
S. Kawano,
and
R. DuBois.
Cyclooxygenase-2 expression in human colon cancer cells increases metastatic potential.
Proc. Natl. Acad. Sci. USA
94:
3336-3340,
1997
45.
Vane, J. R.,
and
R. M. Botting.
Mechanism of action of antiinflammatory drugs.
Int. J. Tissue React.
20:
3-15,
1998[Medline].
46.
Williams, C.,
and
R. DuBois.
Prostaglandin endoperoxide synthase: why two isoforms?
Am. J. Physiol.
270 (Gastrointest. Liver Physiol. 33):
G393-G400,
1996
47.
Xie, W.,
J. Chipman,
D. Robertson,
R. Erikson,
and
D. Simmons.
Expression of a mitogen-responsive gene encoding prostaglandin synthase is regulated by mRNA splicing.
Proc. Natl. Acad. Sci. USA
88:
2692-2696,
1991[Abstract].
48.
Yang, T.,
I. Singh,
H. Pham,
D. Sun,
A. Smart,
J. Schnerman,
and
J. Briggs.
Regulation of cyclooxygenase expression in the kidney by dietary salt intake.
Am. J. Physiol.
274 (Renal Physiol. 43):
F481-F489,
1998
49.
Zusman, R. L.,
and
H. R. Keiser.
Prostaglandin biosynthesis by rabbit renomedullary interstitial cells in tissue culture. Stimulation by angiotensin II, bradykinin, and arginine vasopressin.
J. Clin. Invest.
60:
215-223,
1977[Medline].