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Originally published In Press as doi:10.1074/jbc.M405666200 on July 22, 2004

J. Biol. Chem., Vol. 279, Issue 41, 43157-43167, October 8, 2004
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Osmotic Diuretics Induce Adenosine A1 Receptor Expression and Protect Renal Proximal Tubular Epithelial Cells against Cisplatin-mediated Apoptosis*

Sandeep C. Pingle{ddagger}§, Snigdha Mishra¶§, Adriana Marcuzzi{ddagger}, Satyanarayan G. Bhat{ddagger}, Yuko Sekino||, Leonard P. Rybak{ddagger}, and Vickram Ramkumar{ddagger}**

From the {ddagger}Department of Pharmacology, Southern Illinois University School of Medicine, Springfield, Illinois 62702, the Vollum Institute, Oregon Health Sciences University, Portland, Oregon 97201, and the ||Department of Neurobiology and Behavior, Gunma University School of Medicine, Maebashi 371-8511, Japan

Received for publication, May 20, 2004 , and in revised form, July 20, 2004.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Osmotic diuretics are used successfully to alleviate acute tubular necrosis (ATN) produced by chemotherapeutic agents and aminoglycoside antibiotics. The beneficial action of these agents likely involves rapid elimination of the nephrotoxic agents from the kidney by promoting diuresis. Adenosine A1 receptor (A1AR) subtype present on renal proximal tubular epithelial and cortical collecting duct cells mediates the antidiuretic and cytoprotective actions of adenosine. These receptors are induced by activation of nuclear factor (NF)-{kappa}B, a transcription factor reported to mediate hyperosmotic stress-induced cytoprotection in renal medullary cells. In this study, we tested the hypothesis that induction of the A1AR in renal proximal tubular cells by NF-{kappa}B contributes to the cytoprotection afforded by osmotic diuretics. Exposure of porcine renal proximal tubular epithelial (LLC-PK1) cells to mannitol or NaCl produced a significant increase in A1AR. This increase was preceded by adenosine release and NF-{kappa}B activation. Expression of an I{kappa}B-{alpha} mutant, which acts as a superrepressor of NF-{kappa}B, abrogated the increase in A1AR. Cells exposed to mannitol demonstrated increased reactive oxygen species (ROS) generation, which was attenuated by inhibiting xanthine oxidase with allopurinol. Allopurinol attenuated both the increase in A1AR expression and NF-{kappa}B activation produced by osmotic diuretics, indicating a role of adenosine metabolites in these processes. Treatment of LLC-PK1 cells with cisplatin (8 µM) resulted in apoptosis, which was attenuated by mannitol but exacerbated by selective A1AR blockade. Administration of mannitol to mice increases A1AR expression and activation of NF-{kappa}B in renal cortical sections. Taken together, these data provide novel mechanisms of nephroprotection by osmotic diuretics, involving both activation and induction of the A1AR, the latter mediated through activation of a xanthine oxidase pathway leading to ROS generation and promoting activation of NF-{kappa}B.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Osmotic diuretic agents, such as mannitol are used in the prophylaxis of acute renal failure induced by antineoplastic agents (1), even though they have been supplemented recently by other agents. The beneficial action of osmotic diuretics derives from a reduction in time the nephrotoxic drugs are in contact with the renal tubules (2, 3) and, in the case of mannitol, through an additional direct antioxidant action (4). Osmotic diuretics also induce a number of genes that aid in cell survival, including those for heat shock proteins, and genes involved in the synthesis of osmolytes, such as aldose reductase, betaine transporter, and the sodium-dependent myo-inositol transporter (5, 6). The induction of these genes is mediated by components of the mitogen-activated protein (MAP) kinase signaling pathways, such as c-Jun N-terminal kinase and p38 MAP kinase (7, 8), and in the case of aldolase reductase, through activation of an osmotic response element, crucial for its regulation by hypertonicity (9). A more recent study also indicates a role for NF-{kappa}B-dependent cyclooxygenase-2 (COX-2)1 expression in protecting interstitial fibroblasts from hypertonic stress (10). In general, the studies described above have all used high osmotic stressors to induce proteins of interest in renal medullary cells, since these cells are subjected to a relatively high osmotic environment in vivo during periods of water deprivation.

The renal proximal tubules represent a primary site of action of osmotic diuretic agents. It is highly permeable to water, and as such, reabsorption of water is essentially isotonic. Chemotherapeutic agents, such as cisplatin, are actively transported into proximal tubular cells (11) and concentrate in the P3/S3 pars recta segment (12). The concentration of the antineoplastic agent, cisplatin, in the proximal tubular epithelial cells exceeds plasma concentrations by a factor of 5 (13), rendering this segment most susceptible to injury. We have recently shown that cisplatin administration increased the expression of the adenosine A1 receptor (A1AR) in different regions of the kidney, including the proximal tubule (14), presumably via activation of NF-{kappa}B (15).

Adenosine receptor (AR) subtypes show differential localization in the kidney. Specific receptor expression is demonstrated in different nephron segments, such as the glomeruli, the thick ascending limb and collecting duct (16) and the proximal tubules (17). In the proximal tubule and the cortical collecting duct, adenosine stimulates the transport of sodium and phosphate via the apical surface (18), that of sodium and bicarbonate via the basolateral symporters (19) and stimulates chloride channel activity (20). As such, a primary physiological role of renal adenosine is believed to be antidiuresis, mediated through activation of proximal tubule A1AR coupled to reabsorptive transport processes (21, 22).

The present study was performed to determine whether exposure to osmotic diuretics could induce A1AR in proximal tubular epithelial cells and whether activation of A1ARs could contribute to the protective action of mannitol against drug-induced nephrotoxicity.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Cell Culture
LLC-PK1 cells were cultured in renal epithelial growth media (REGM) supplemented with 0.5% serum, specifically formulated for renal epithelial cells. Cultures were maintained as monolayers on 75-cm2 tissue culture flasks/well plates at 37 °C, in 5% CO2, 95% ambient air, with the medium being replaced every 2-3 days.

Animals
Male C57BL/6 mice (6-8-week-old) obtained from Harlan Laboratory (Indianapolis, IN) and were maintained on pulverized food and water. Animals were used according to the guidelines approved by the Laboratory Animal Care and Use Committee of the Southern Illinois University School of Medicine.

Drug Treatment and Sample Collection
Mice were anesthetized by isoflurane inhalation, followed by retro-orbital sinus injection of mannitol (0.8 g/ml/kg body weight) or an equal volume of saline (controls). Animals were sacrificed after 2 h (for NF-{kappa}B activity) or 20 h (for A1AR expression) by isoflurane inhalation followed by cervical dislocation. Kidneys were removed, and the cortices were dissected out, rapidly frozen in liquid nitrogen and stored at -80 °C.

Radioligand Binding Assays
Cells were treated with mannitol for 24 h and subsequently harvested for radioligand binding assay. Cells were detached in ice-cold phosphate-buffered saline (PBS) containing 5 mM EDTA and resuspended in 50 mM Tris-HCl buffer (pH 7.4), containing 10 mM MgCl2 and 1mM EDTA, 10 µg/ml soybean trypsin inhibitor, 10 µg/ml benzamidine, and 2 µg/ml pepstatin (Buffer A). This was followed by homogenization with a Polytron homogenizer (Brinkmann Instruments, Westbury, NY) at setting 7 for 40 s at 4 °C. Membranes were obtained by centrifugation of the homogenates at 2,000 x g for 10 min, followed by centrifugation of the supernatant at 40,000 x g for 15 min. The final pellet was resuspended in Buffer A to yield a protein concentration of 0.5 mg/ml. The membrane suspensions were then treated with adenosine deaminase (5 units/ml) and incubated at 37 °C for 10 min to eliminate endogenously released adenosine.

Quantitation of A1AR was performed using the tritiated antagonist 8-cyclopentyl-1, 3-dipropylxanthine ([3H]DPCPX) and the iodinated agonist radioligand N6-(4-aminobenzyl)-9-[5-(methylcarbonyl)-{beta}-D-ribofuranosyl] adenine (125I-AB-MECA). For these assays, membrane preparations (~40 µg of protein) were incubated for 1 h at 37 °C with either radioligand in the absence or presence of theophylline (1 mM) (for [3H]DPCPX binding) or 10 µM DPCPX (for 125I-AB-MECA binding) in order to define nonspecific binding. The total incubation volume was 250 µl. Samples were then filtered through polyethyleneimine-treated Whatman GF/B glass-fiber filters using a cell harvester (Brandel, Gaithersburg, MD) and washed with 9 ml of ice cold Buffer A (without protease inhibitors), containing 0.01% CHAPS. Bound radioactivity was determined using either a scintillation or gamma counter.

RNA Preparation, Polymerase Chain Reactions, and Northern Blotting
Isolation of total RNA was performed using TRIzol reagent kit (Invitrogen) and selection of poly (A+) messenger RNA, using oligo (dT)-cellulose was performed as described previously (23). For PCR studies total RNA (1 µg each) was reverse-transcribed using a first strand cDNA synthesis kit (Amersham Biosciences) in a total volume of 15 µl. Five microliters of each of the reaction volumes were used for PCR amplification. Primers used include the canine A1AR consensus protein sequence ILGNULU (sense) and FALCWLP (antisense) and predictably generated a 770-bp fragment (24). PCR were performed in a total volume of 50 ml using 2.5 mM MgCl2, using 36 amplification cycles. The amplified products were resolved on 1.2% agarose gels, which were subsequently denatured, neutralized and transferred to nylon filters for Southern blot analysis. Filters were UV cross-linked and prehybridized for 4 h at 42 °C in a mixture containing 50% formamide, 6x SSC (20x SSC = 175 g NaCl, 88 g sodium citrate, pH 7.0), 5x Denhardt's (50x = 0.25 volumes of 4% bovine serum albumin, 0.25 volumes of 4% polyvinylpyrolidine, 0.25 volumes of 4% Ficoll, 0.25 volumes of dH2O, 0.5% sodium pyrophosphate), 0.1% SDS, and 0.1 mg/ml salmon sperm DNA, using 1 x 106 cpm/ml of 32P-labeled A1AR cDNA probe. Hybridizations were performed by shaking blots in a waterbath at 42 °C for 16-20 h. Following hybridization, blots were washed twice (15 min each) at room temperature in 2x SSC and 0.1% SDS and twice (20 min each) with 0.1x SSC and 0.1% SDS at 62 °C. The relative band intensities were determined by densitometric scanning on the GS-250 Molecular Imager (Bio-Rad) after exposing the blots to the imager screen for 1 h.

For Northern blotting experiments, poly(A+) RNA samples (5 µg) were electrophoresed on a 1% agarose/MOPS/formamide gel, transferred to nylon membranes, and cross-linked in Stratagene UV crosslinker. Prehybridization mixtures contained 5x SSC, 2x Denhardt's, 0.1% SDS, 0.2 mg/ml salmon sperm DNA, and 50% formamide. Hybridization mixtures (10 ml) were essentially the same, except for the Denhardt's concentration being 2.5x, and the added 32P-radiolabeled canine cDNA probes encoding the A1AR at concentrations of 0.5-1 x 106 cpm/ml. Hybridization and washing conditions were similar as described for Southern blotting. Image was visualized and quantitated using a densitometric scanner (as above). These blots were stripped and reprobed with labeled cDNA probe encoding the human glyceraldehyde phosphate dehydrogenase (GAPDH) for normalization.

H2DCFDA Fluorescence
Intracellular production of free radicals (reactive oxygen and nitrogen species) was detected in LLC-PK1 using 2', 7'-dichlorodihydrofluorescein diacetate (H2DCFDA, Calbiochem, San Diego, CA) (25-27). Cells were plated on sterile 12-mm glass coverslips at 400 cells/mm2 in individual wells of 24-well tissue culture plates. The cells were treated with 100 mM mannitol (Sigma), in the absence and presence of 250 µM allopurinol (Sigma) for 24 h. Coverslips were washed with PBS and the cells loaded with H2DCFDA by incubating in 5 µM H2DCFDA for 20 min at 37 °C and washed with PBS. The cultures were analyzed for green fluorescence 1 h later using an Olympus fluorview confocal laser-scanning microscope using argon laser and a x40 objective.

Luciferase Assay
LLC-PK1 cells were cultured to ~20-40% confluency and transfected with a mixture containing 100-250 ng of plasmid DNA, 500-650 ng of carrier DNA and 3 µl/g DNA of N-[1-(2,3-dioleoyloxy)propyl]-N,N,N-trimethylammonium methylsulfate (Lipofectin) in a volume of 50 µl of Opti-MEM (Invitrogen). The mixtures were incubated for 45-60 min at room temperature and then added to the culture plate. After ~6 h, regular renal epithelial growth media (supplemented with 0.5% serum medium) were added to the plate, and it was returned to the incubator for 24 h. For luciferase assays, cells were then lysed using 50 µl of reporter lysis buffer (Promega, Madison, WI) and centrifuged at 4 °C in a microcentrifuge at 12,000 x g. The extract was used immediately or stored at -70 °C. Twenty microliters of cell extract was mixed with 100 µl of luciferase assay reagent at room temperature and the chemiluminescent signal was determined in a luminometer using 1 min counts.

Preparation of Nuclear Extracts
Nuclear extracts were prepared from the cells and renal cortices as described previously (15). Briefly, the samples were suspended in Buffer B (10 mM HEPES, pH 7.9, 10 mM KCl, 0.1 mM EDTA, 0.4% Nonidet P-40, 1 mM dithiothreitol, and 1 mM phenylmethanesulfonyl fluoride). The mixtures were centrifuged at 5000 x g for 30 s, and the cytosolic extract was separated. The nuclear pellet was washed with excess volume of Buffer B and then resuspended in Buffer C (20 mM HEPES, pH 7.9, 400 mM NaCl, 1 mM EDTA, 1 mM dithiothreitol, and 1 mM phenylmethanesulfonyl fluoride). After incubating for 5 min at 4 °C with rotation, the extracts were centrifuged (5000 x g, 1 min), and the supernatant was used for DNA binding activity analyses.

Electrophoretic Mobility Shift Assay
Nuclear extracts were incubated with double strand-specific {kappa}B oligonucleotide (5'-ATGTGAGGGGACTTTCCCAGGC-3') (28). Similar electrophoretic mobility shift assays were performed using a labeled oligonucleotide probe (5'-CGCTTGATGAGTCAGCCGGAA-3') for the AP-1 transcription factor binding sequence. Incubations were performed at room temperature for 30 min in a total volume of 15 µl of buffer containing 12% glycerol, 12 mM HEPES-NaOH (pH 7.9), 60 mM KCL, 1 mM EDTA, and 1 mM dithiothreitol, 1.0 µg of poly(dI-dC), and 10,000 cpm of the labeled probe. The DNA-protein complexes were resolved on nondenaturing 5% polyacrylamide gels, performed with 0.5x Tris borate/EDTA buffer (4.5 mM Tris, 4.5 mM boric acid, 1.0 mM EDTA, pH 8.0). Bands were imaged using the Phosphorimager and analyzed using Optiquant software.

Apoptosis Detection
AnnexinV-FITC Assay—Cells were washed with PBS and harvested using 1 ml of 0.5% of trypsin/EDTA solution at 37 °C. Cells were then centrifuged at 220 x g for 5 min and immediately resuspended in the medium. Cells (~5 x 10-5 cells/ml) were then incubated in the dark for 15 min at room temperature with 10 µl of media binding agent and 1.25 µl of a FITC conjugate of Alexa/5. Cells were next centrifuged at 1,000 x g for 5 min and resuspended in 0.5 ml of cold 1x binding buffer (HEPES, NaCl, CaCl2, MgCl2, bovine serum albumin). Propidium iodide (10 µl) was then added, and samples were placed on ice away from light and analyzed immediately. Quantification of Annexin V-FITC and propidium iodide signals was performed by a flow cell based bench top FACS Calibur Cytometer (BD Biosciences) with excitation wavelength of 488 nm and transmission wavelengths of 515-545 nm. Early apoptotic cells are reported in the lower right-hand quadrant of the dot plot, while necrotic or late apoptotic cells are reported in the upper right-hand quadrant of the plot. LLC-PK1 cells were also examined by fluorescence microscopy (using a Olympus fluorview confocal laser-scanning microscope), equipped with Argon (488 nm) and Krypton (568 nm) lasers for green and red fluorescence, respectively. Phase contrast images were observed using transmitted light. Apoptotic cells were detected as bright apple green, while necrotic cells appear with various intensities of yellow-red throughout the cytoplasm. Viable cells remain unstained.

TUNEL Assay—Cisplatin-induced apoptosis was confirmed using the TUNEL assays, as detailed by the manufacturer. For this assay, cells were grown on poly-L-lysine-coated coverslips, pretreated with either vehicle or mannitol, and then treated with either vehicle or cisplatin for an additional 24 h. The monolayers were then fixed with 4% formaldehyde in 1x Tris-buffered saline (TBS) for 10 min at room temperature. Cells were then permeabilized by addition of 100 µl of 20 µg/ml proteinase K solution (in Tris buffer at pH 8) at room temperature for 5 min, followed by incubation with 100 µl of 0.3% H2O2 at room temperature for 5 min to inactivate endogenous peroxidase. Monolayers were incubated with 100 µl of 1x terminal deoxythymidine (TdT) equilibration buffer at room temperature for 10-30 min, followed by incubation with TdT enzyme mix for 1.5-2 h at 37 °C in a humidified chamber. The reaction was terminated by addition of 100 µl of stop solution (0.5 M EDTA, pH 8) at room temperature for 5 min, followed by rinsing the cells with 1x TBS. Cells were then incubated for 10 min at room temperature with 100 µl of phosphate-buffered saline containing 4% bovine serum albumin, followed by addition of 100 µl of 1x dilution of peroxidase strepavidin for 30 min at room temperature in humidified chamber. Coverslips were then incubated with 100 µl of 3',3'-diaminobenzidine solution for 5 min and then rinsed with distilled water. Methyl green counterstain solution was then added for 3 min, the excess stain drained off, and the samples rinsed three times with distilled water. DNase (1 µg/µl) in 1x TBS/1 mM MgSO4 was used to generate a true control. For the negative control, treatment with the TdT reaction mix was replaced by distilled H2O.

Immunocytochemistry for A1AR
LLC-PK1 cells were cultured and treated as described in the figure legends. Following specific treatments, cultures were washed twice with warm phosphate-buffered saline and fixed with 4% paraformaldehyde for 10 min. After two more washes with phosphate-buffered saline, nonspecific binding was reduced by exposing cover slips for 5 min with a solution containing 5% normal donkey serum and 0.05% Triton X-100. The cells were treated with A1AR monoclonal antibody, diluted 1:20 in 5% normal donkey serum along with 0.05% Triton X-100 in PBS) and incubated overnight at 4 °C. After rinsing four times in phosphate-buffered saline, cells were treated for 1 h with donkey anti-mouse IgG labeled with rhodamine (Jackson Immunochemicals, West Grove, PA) diluted 1:100 in 5% normal donkey serum, 0.05% Triton X-100 in PBS. After four rinses with phosphate-buffered saline, the coverslips were mounted on glass microscope slides using Aquamount. The cells were observed for red color on an Olympus confocal microscope using Krytpon (568 nm) laser and a x40 objective.

Protein Determination
Protein concentrations were determined by the established Bradford protein assay (29) using bovine serum albumin to prepare standard curves.

Statistical Analyses
Saturation curves and competition curves were analyzed by a computer-assisted curve fitting program (Graph PAD Prism Software, San Diego, CA). Statistical analyses were performed by the analysis of variance.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
Characterization of the Adenosine A1 Receptor in LLC-PK1 Cells—The presence of the A1 and A2AAR in LLC-PK1 cells have been determined by functional assays (30), but no studies to date have demonstrated the existence of AR subtypes in these cells by radioligand binding experiments or molecular biology techniques. Therefore, initial studies characterized the A1AR in membrane preparations obtained from LLC-PK1 cells by radioligand binding assays using the agonist radioligand 125I-N6-(4-aminobenzyl)-9-[5-(methylcarbonyl)-{beta}-D-ribofuranosyl]adenine (AB-MECA), along with 10 µM DPCPX to define nonspecific binding. While this radioligand interacts with both the A1 and A3AR (31), albeit with lower affinity for the latter receptor, the use of DPCPX, a selective antagonist to block the A1AR, provides a simple method for distinguishing this receptor subtype from the A3AR. The data were best fitted according to a one-site model by Graph Pad Prism Software (San Diego, CA), which indicates the receptor number (Bmax) of 42.4 ± 3.1 fmol/mg protein and equilibrium dissociation constant (Kd) of 1.1 ± 0.3 nM (Fig. 1A). In competition experiments, 125IABMECA binding was inhibited by DPCPX, with an inhibitory constant (Ki) of 1.6 ± 0.1 nM, characteristic of the interaction of this drug with the A1AR (Fig. 1B). The inability of DPCPX to completely inhibit 125I-ABMECA binding is indicative of this radioligand also interacting with other ARs (such as the A3AR), which are not targets of DPCPX. Radioligand binding was also inhibited completely by R-phenylisopropyladenosine (R-PIA), a relatively selective A1AR agonist, with a Ki of 1.2 ± 0.1 nM (data not shown). This relatively high affinity interaction of R-PIA is also characteristic of its preferential interaction with the A1AR, at lower concentrations, as opposed to the A3AR subtype. Similar to other G-protein-coupled receptors, addition of GTP{gamma}S to membrane preparations dramatically reduced the population of the A1AR in the high affinity state, and therefore the level of 125I-AB-MECA. The binding of 125I-AB-MECA was reduced by 37.5 ± 5.3% and 62.2 ± 7.0%, upon incubation of membranes with 0.1 and 10 µM of GTP{gamma}S, respectively (data not shown). The presence of A1AR in LLC-PK1 cells was also confirmed by Northern blotting studies (Fig. 1C, upper), using poly(A)+ preparations and a labeled bovine A1AR cDNA probe for detection of the transcript. Polymerase chain reactions were performed, using forward and reverse sequences derived from the canine A1AR cDNA. Primers used include amino acid sequences ILGNVLV (sense) and FALCWLP (antisense) common to the A1AR in several species (24). Sequences were identified as the A1AR by Southern blotting using a labeled canine A1AR cDNA probe (Fig. 1C). The figure shows the predicted 770-bp PCR fragments derived from the canine A1AR cDNA (lane 1), water blank (lane 2), LLC-PK1 cells (lane 3), rat kidney (lane 4), and testes (lane 5). Immunocytochemical assays, using a monoclonal antibody (32), were performed to further determine the presence of A1AR on LLC-PK1 cells. As shown in Fig. 1D, the presence of the A1AR was detected as a fluorescent halo, using confocal microscopy.



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  		FIG. 1.
Expression of A1AR in LLC PK1 cells. Membranes prepared from LLC-PK1 cells were used for 125I-AB MECA binding assays as described under "Experimental Procedures." A, saturation curve is fitted by Graph Pad Prism Software using a 1-site fit model. B, competition curves for DPCPX and R-PIA. Membranes were labeled with 1 nM 125I-AB MECA in the presence of increasing concentrations of DPCPX. Points were computer fitted according to Graph Pad Prism Software. C, detection of A1AR mRNA by PCR. Reverse-transcribed RNA from different tissues or canine A1AR cDNA were amplified using A1AR primers using 36 cycles. Lane 1 represents the canine A1AR cDNA, lane 2 represents the water blank, lane 3 represents LLC-PK1 cells, lane 4 represents rat kidney, and lane 5 represents rat testes. D, A1AR immunocytochemistry using a monoclonal antibody against the A1AR (32). Arrows indicate localization of staining to the cell membrane.

 
Mannitol Increases Expression of the Adenosine A1 Receptor in LLC-PK1 Cells—To determine whether the expression of the A1AR is modulated by hypertonicity, LLC-PK1 cells were exposed to 100 mM mannitol for 24 h, and A1AR levels were determined by radioligand binding assay. Saturation curves performed using 125I-AB-MECA indicate an increase in the number of A1AR in cells exposed to mannitol for 24 h (Fig. 2A). In cells exposed to mannitol, the Bmax was increased from 39.7 ± 1.7 to 65.8 ± 6.0 fmol/mg protein, with no significant change in Kd values. Similar increases in A1AR was obtained when the antagonist radioligand ([3H]DPCPX) was used to quantitate receptor levels (data not shown). LLC-PK1 cells exposed to increasing concentrations of mannitol for 24 h showed dose-dependent increases in receptor number by 90 ± 17, 110 ± 15, and 157 ± 16%, following administration of 50, 100, and 200 mM mannitol, respectively (Fig. 2B). Time course studies indicate significant increases in A1AR at 16 h after addition of mannitol, with further increases observed by 24 h (data not shown). In another series of experiments, addition of NaCl to separate cultures for 24 h resulted in a dose-dependent increase in A1AR expression, which was optimal at 150 mM (data not shown). Additional confirmation of an increase in A1AR was provided by immunocytochemistry (Fig. 2C). Results obtained show increases in A1AR expression in these cells following exposure of LLC-PK1 cells to mannitol (100 mM). Quantitation of the immunoreactivity by confocal microscopy indicated significant elevations in A1AR by ~80%, following exposure of cells to mannitol.



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  		FIG. 2.
Osmotic diuretics increase expression of A1AR. A, saturation curves were performed on membranes obtained from control LLC-PK1 cells and those treated mannitol (100 mM) for 24 h. Plots are generated by the Graph Pad Prism program using a 1-site fit model. The Bmax and Kd values were 39.7 ± 3.7 fmol/mg protein and 2.2 ± 0.2 nM for control cells and 65.8 ± 13.1 fmol/mg protein and 2.3 ± 0.8 nM for mannitol-treated cells. B, dose response curve for mannitol normalized to control cells. * indicates p < 0.05. C, immunocytochemistry for A1AR showing increased membrane-localized receptor expression following mannitol treatment. D, mannitol increases the steady state levels of A1AR mRNA. Cells were treated without or with mannitol for 24 h, poly(A+) RNA prepared and used in Northern blot analysis. For normalization, blots were stripped and probed with a 32P-labeled cDNA probe for the GAPDH. Band intensities were quantified by a PhosphorImager and show a 68 ± 23% increase after mannitol treatment (mean ± S.E. of six samples per treatment group).

 
Additional experiments were performed to test whether the increase in A1AR could be explained by an increase in the steady state level of A1AR-specific mRNA, using Northern blotting assays. Blots probed with the canine A1AR cDNA probe showed a statistically significant increase in the steady-state levels of A1AR-specific mRNA by 68 ± 23% (Fig. 2D). One possible explanation for this observation is that mannitol increases the promoter activity of the A1AR gene. To test this possibility, LLC-PK1 cells were transiently transfected with plasmid pBLPnif/PmtA, which contains the firefly luciferase reporter gene driven by the A1AR promoter (33). Subsequent exposure of these cells to mannitol (100 mM) for 24 h resulted in a 3.4 ± 0.3-fold increase in luciferase activity compared with untreated control cells (data not shown), suggesting that the increase in A1AR mRNA could be caused by the increase in transcription of the A1AR gene.

Induction of A1AR Expression by Mannitol Involves Activation of NF-{kappa}B—Because previous studies have shown that hypertonicity increases NF-{kappa}B activity in renal medullary cells (10), we tested whether the increase in A1AR expression by these agents was due to activation of NF-{kappa}B in LLC-PK1 cells. Cells treated with mannitol (100 mM) demonstrated a significant increase in NF-{kappa}B activation, which was reduced substantially following co-incubation of mannitol with, either sodium salicylate (100 µM) or dexamethasone (100 nM), drugs known to inhibit this transcription factor (34) (Fig. 3A). Furthermore, infection of LLC-PK1 cells with an adenoviral vector expressing a mutant form of I{kappa}B-{alpha}, which acts as a superrepressor of NF-{kappa}B (mI{kappa}B-{alpha}), abrogated the induction of A1AR by mannitol (Fig. 3A). Infection of LLC-PK1 cells was visualized by fluorescence microscopy, as the I{kappa}B-{alpha} gene used was tagged with a green fluorescent protein. Further, the expression of mI{kappa}B-{alpha} in these cells was confirmed by electrophoretic mobility shift assay to determine nuclear translocation of NF-{kappa}B and Western blotting for I{kappa}B-{alpha} (data not shown). Taken together, these data provide strong support for the involvement of NF-{kappa}B in the induction of A1AR expression by mannitol. Additional studies showed that inhibition of the MAPK-ERK kinase (MEK) pathway by PD98059 failed to block the induction of the A1AR by mannitol (data not shown), indicating that this latter pathway does not contribute significantly to this process.



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  		FIG. 3.
Up-regulation of the A1AR by mannitol involves activation of NF- {kappa}B. A, LLC-PK1 cells were pretreated with vehicle, sodium salicylate (100 mM), or dexamethasone (100 nM) 2 h prior to the administration of mannitol. Treatment with viral vector encoding the super-repressor of NF-{kappa}B (mI{kappa}B-{alpha}) was carried out 15 h prior to the addition of mannitol. The levels of A1AR were determined by radioligand binding assay using 125I-ABMECA (2 nM). * indicates statistical significance from cells not exposed to mannitol (p < 0.05). ** indicates the degree of suppression of the mannitol response was statistically significant (p < 0.05). B, mannitol and NaCl increased the activity of NF-{kappa}B in LLC-PK1 cells. Cells were exposed to vehicle, mannitol (100 mM), or NaCl (100 mM) for 1 h, and nuclear preparations were made and used in electrophoretic mobility shift assays. The two bands indicating NF-{kappa}B were shown to increase by mannitol over that observed for the control treatment. C, super-shift assay to determine subunit composition of NF-{kappa}B in mannitol-treated cells. Shifted bands were detected by using antibodies against p65 and c-Rel (200 ng of antibody/reaction mix).

 
Further confirmation of mannitol-induced activation of NF-{kappa}B in LLC-PK1 cells was provided using electrophoretic mobility shift assays to determine nuclear translocation of NF-{kappa}B in LLC-PK1 cells. As shown in Fig. 3B, exposure of LLC-PK1 cells to mannitol (100 mM) or NaCl (100 mM) for 30 min-induced activation of NF-{kappa}B by 2-3-fold, as determined by increased retention of the 32P-labeled {kappa}B oligonucleotide probe. There was a greater increase in NF-{kappa}B activity in cell treated with 100 mM NaCl than with 100 mM mannitol, as would be expected from the higher osmolarity produced by 100 mM NaCl. To determine the subunit composition of the NF-{kappa}B complex, aliquots of the nuclear preparation were each incubated with preimmune serum or with polyclonal antibodies against distinct subunits, p50, p52, RelA, cRel, or RelB. The binding of the antibody to a specific subunit results in a higher molecular weight protein-DNA complex, which appears "supershifted" on a non-denaturing polyacrylamide gel. Our experiments indicate supershifted bands only in the preparations treated with anti-p65 and anti-cRel, suggesting that the NF-{kappa}B complex detected in the nucleus contained predominantly the p65 and c-Rel proteins (Fig. 3C).

Role of Reactive Oxygen Species in Mannitol-induced Activation of NF-{kappa}B—Previous studies from our laboratory indicate that the generation of ROS in response to cisplatin administration is the prime mediator of NF-{kappa}B activation by this chemotherapeutic agent, leading to induction of A1AR expression (15). To detect ROS, we utilized the reagent H2DCFDA, which fluoresces on binding with superoxides and peroxynitrite (25-27). In cells treated with mannitol, an increase in ROS generation was observed. The addition of allopurinol alone did not alter the basal ROS production. However, ROS production in response to mannitol was blocked by pretreatment with 250 µM allopurinol (Fig. 4A). Since allopurinol inhibits xanthine oxidase activity, a major source of free radical production in cells, this finding suggests that mannitol-induced ROS generation is mediated primarily by the xanthine oxidase pathway. Moreover, free radical production by this pathway could serve as the trigger for activation of NF-{kappa}B. In support of this notion, we showed that mannitol-induced activation of NF-{kappa}B was attenuated following exposure of cells to allopurinol (Fig. 4B). Since adenosine serves as a precursor for substrates of the xanthine oxidase pathway, we determined the levels of adenosine released following administration of mannitol. Cells treated with 100 mM mannitol showed ~2-fold increase in extracellular adenosine levels, suggesting that this could serve as a source of free radical production by the xanthine oxidase pathway.



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  		FIG. 4.
Mannitol increases ROS production in LLC-PK1 cells. A, cells were treated with either vehicle or allopurinol (250 µM) for 2 h, followed by the addition of vehicle or mannitol for 30 min. ROS production was measured by confocal microscopy using the indicator H2DCFDA (5 µM), with wavelength setting of 488 nm. ROS production was detected as an increase in green fluorescence. These experiments were repeated at least three times. B, mannitol-induced increase in NF-{kappa}B activity in LLC-PK1 cells was attenuated by pretreatment with allopurinol (250 µM), catalase (200 units/ml), or PDTC (50 µM), as detected by decreased band intensity in electrophoretic mobility shift assays. C, pretreatment of LLC-PK1 cells with allopurinol or catalase also attenuated mannitol-induced increase in A1AR expression, as observed by a decrease in the binding of the radioligand 125I-AB-MECA. D, treatment with hydrogen peroxide (200 µM) increased the expression of the A1AR. Pretreatment with catalase (200 units/ml) attenuated the increase in A1AR in response hydrogen peroxide.

 
Treatment of cells for 24 h with either catalase (200 units/ml) or allopurinol (250 µM) abolished the induction in A1AR expression induced by mannitol (Fig. 4C). Additional experiments were performed to determine whether increasing oxidative stress by the addition of H2O2 (200 µM) to the LLCPK1 cultures for 24 h mimics the response of mannitol. Results shown in Fig. 4D indicate that H2O2 induced A1AR and that this induction was reversed by catalase (200 units/ml) (Fig. 4D).

Mannitol Activates NF-{kappa}B and Increases Expression of A1ARs in Renal Cortices of Mice—To determine whether mannitol can induce expression of the A1AR in vivo, mice were administered mannitol by retro-orbital sinus injections (0.8 g/ml/kg mannitol). The diuretic effect of mannitol was evidenced by increased urination of the mice on the bedding in their cages and by increased water consumption. Kidneys were isolated after 20 h and the renal cortices were used to perform radioligand binding assay for the A1AR using [3H]DPCPX. We observed an ~45% increase in A1AR expression in kidneys of mannitol-treated mice (5.9 ± 0.61 fmol/mg protein) when compared with control mice (4.1 ± 0.12 fmol/mg protein) injected with normal saline (Fig. 5A). Separate kidneys were used to assess nuclear translocation of NF-{kappa}B by mannitol treatment. As shown in Fig. 5B, there was a 3-5-fold increase in NF-{kappa}B in cortices obtained from the mannitol-treated when compared with saline-treated animals (Fig. 5B).



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  		FIG. 5.
Mannitol increases A1AR expression and promotes NF-{kappa}B activation in the mouse renal cortex. A, renal cortices from mice treated with mannitol for 20 h were used for radioligand binding studies, using ~40-µg membrane preparation and 1 nM [3H]-DPCPX. B, mannitol increases NF-{kappa}B activity. Nuclear extracts were prepared from mouse renal cortices and used in electrophoretic mobility shift assays. Each lane was loaded with 10 µg of nuclear extracts. A 3-5-fold increase in activity was observed in cortical samples obtained from mannitol-treated mice as compared with the saline-treated controls.

 
Cytoprotection Provided by Mannitol Involves Up-regulation of the A1AR—Whereas the protective effects of mannitol on the kidneys have been widely reported (1-4), the mechanism underlying protection remains to be established. Previous studies in our laboratory have demonstrated a protective role of the A1AR in reducing oxidative stress induced by cisplatin in a hamster vas deferens smooth muscle clone (15). We therefore examined whether activation of the A1AR would render the LLC-PK1 cells more tolerant to cisplatin toxicity. The conventional dose of 20 mg/m2/day cisplatin intravenously (35) results in plasma levels of ~8 µM in a 70 kg man. Since cisplatin is concentrated in proximal tubular cells, it is expected that the levels achieved in these cells would be much higher than the plasma concentration of the drug (13). In order to correlate with a clinically effective plasma concentration of cisplatin, LLC-PK1 cells were pretreated with either vehicle or mannitol (100 mM) for 12 h, followed by administration of either vehicle or cisplatin (8 µM) for an additional 20 h. Flow cytometric analysis (Fig. 6, A and B) indicate that exposure to mannitol alone did not significantly affect apoptosis (upper panel, right). Exposure to cisplatin for 20 h resulted in a significant induction in apoptosis (Fig. 6A, upper panel, middle), with 40.7 ± 2.1% of cells staining positive for Alexa5-FITC. However, pretreatment with mannitol decreased the number of apoptotic cells (lower panel, left) to 1.2 ± 0.2% of the total cells. To test whether this protective effect of mannitol involved the A1AR, cells were pretreated with the DPCPX, a selective antagonist of this receptor subtype, prior to the administration of mannitol and cisplatin. These cells demonstrated increased apoptosis (49.2 ± 1.4% apoptotic cells, lower right panel) compared with that produced by cisplatin alone, indicating that the protective effect of mannitol is linked to activation of the A1AR. The addition of DPCPX alone did not change the level of apoptosis from that observed for control cells (lower panel, middle), suggesting a lack of tonic stimulation of these receptors under resting conditions.




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  		FIG. 6.
Mannitol protects against cisplatin-induced apoptosis. A, flow cytometry: cells were pretreated with 100 mM mannitol (panels c, d, and f) or with the selective A1AR antagonist (DPCPX, 1 µM) (panels e and f), along with mannitol (panels c, d, and f) for 12 h prior to the administration of cisplatin (8 µM) (panels b and d) in the culture medium for an additional 20 h. Apoptosis was determined by measuring the percentage of annexin-positive cells by flow cytometry and plotted in B. B, percentage of apoptotic cells for each treatment was determined in A and plotted as the mean ± S.E. of six independent experiments each. * indicates statistically significant change in apoptosis induced by cisplatin. ** indicates statistically significant potentiation of apoptosis by pretreatment of cells with DPCPX prior to addition of mannitol. *** indicates statistically significant change from cisplatin added alone. C, TUNEL assay: cells were pretreated with either vehicle, mannitol (100 mM), or NaCl (100 mM) for 12 h, followed by the addition of cisplatin (8 µM) for 20 h and then used for TUNEL assays. Apoptotic cells are detected as cells possessing dark brown to gray or black diaminobenzidine-stained nuclei. Cisplatin produced a significant increase in apoptotic cells, which was substantially reduced by pretreatment with mannitol and NaCl. Data are presented as the mean ± S.E. * indicates statistically significant difference from control (p < 0.05). ** indicates statistically significant suppression of cisplatin-induced apoptosis (p < 0.05).

 
The protective action of mannitol could also be demonstrated using TUNEL assays to quantitate the number of apoptotic cells. In this assay, apoptotic cells are detected as dark brown to gray or black diaminobenzidine-stained nuclei (Fig. 6C). Quantitation of TUNEL assay images showed 3 ± 1 TUNEL-positive cells (8 ± 3%) in the control fields (total cells per field = 37 ± 7) and 2 ± 1 TUNEL-positive cells (6 ± 3%) in the mannitol treatment group (total cells per field = 34 ± 7). Following cisplatin treatment for 20 h, the number of TUNEL positive cells increased to 25 ± 3 (64 ± 8%) (total cells per field = 39 ± 2), while pretreatment with mannitol blocked the induction of apoptosis to 5 ± 1 (16 ± 3%) (total cells per field = 31 ± 5) (Fig. 6D). A similar finding was observed after cells were pretreated with NaCl (100 mM). Pretreatment with NaCl resulted in no significant change in the number of apoptotic cells from control. A total of 2.6 ± 1.0 (10 ± 4%) TUNEL-positive cells were detected per field (total cells per field = 26 ± 1). Following pretreatment with 100 mM NaCl for 12 h, exposure to cisplatin (8 µM) for 20 h did not result in any appreciable change in the number of apoptotic cells, which averaged 5.3 ± 0.5 TUNEL-positive cells per field (16 ± 1%) (total cells per field = 34 ± 1) (data not shown).


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
Osmotic diuresis has commonly been used to alleviate nephrotoxicity due to chemotherapeutic agents (1) and other nephrotoxins. This mode of treatment to prevent nephrotoxicity was first introduced in the clinic by Ozols and Young (36). One factor which contributes to the beneficial action of agents like mannitol is an increase in single-nephron filtration due to an increase in glomerular plasma flow, which reduces the time the drug and renal tubule are in contact (2, 3). Mannitol also inhibits cisplatin-induced lipid peroxidation in rat renal slices, presumably via a direct antioxidant action (4). The administration of mannitol to the ischemic kidney increases renal blood flow, by decreasing the intrarenal vascular resistance through release of vasodilator substances such as prostaglandins or by washing out interstitial sodium, and reducing the sensitivity of the renal vasculature to ischemia-induced stimulation of the renin-angiotensin system (37). Indirect evidence from our laboratory indicates that blockade of the A1AR exacerbated cisplatin nephrotoxicity (14), implying a cytoprotective role of this receptor subtype under normal physiological conditions. However, the possibility that these receptors are also involved in mediating the beneficial actions of mannitol against cisplatin nephrotoxicity has not been studied.

Adenosine, acting via the A1AR, plays an important cytoprotective role against ischemic and chemical stressors, which increase ROS generation. Interestingly, ROS enhance the expression of the A1AR by NF-{kappa}B activation, which can then interact with consensus DNA sequences on the A1AR promoter (15). This suggests the presence of a feedback loop involving adenosine, ROS, NF-{kappa}B, and the A1AR, whereby the presence of ROS activates an NF-{kappa}B-dependent A1AR expression and the activation and induction of the A1AR reduces the toxicity of ROS. Adenosine controls the activity of a number of transport processes and ion channels in the kidney by interacting with renal tubular A1AR on the basolateral and apical membranes (18-22). Thus, up-regulation of the A1AR could serve as an additional mechanism of adaptation to osmotic stress. Recently published data have also shown that mannitol induces the expression of COX-2 in renal medullary interstitial cells through activation of NF-{kappa}B, and that this promotes cell survival by stimulating renal medullary blood flow (10). The induction of COX-2 was evident at higher concentrations (>200 mOsmol) of mannitol, unlike the induction of the A1AR, which was statistically significant at 50 mOsmol.

The data derived from this study clearly indicate upregulation of the A1AR in proximal tubular cell cultures by osmotic diuretics, which confers protection against cisplatin-induced apoptosis. The A1AR was induced in cells exposed to 50 and 100 mM mannitol, a concentration range that is achieved in the plasma after intravenous infusion (38) and which would be achieved in the lumen of the proximal tubules. Increased A1AR expression was also observed in mice injected with mannitol, indicating that this change might have beneficial application in vivo. The induction of A1AR was much slower than that of p53 (39) and heat shock proteins (40) by osmotic stress in the renal medullary epithelial cells. Thus, the A1AR might contribute to the slower adaptative response induced by osmotic stress.

Activation of NF-{kappa}B plays an integral role in the induction of the A1AR in LLC-PK1 cells by osmotic diuretics. This conclusion is supported by the observation that induction of A1AR was abolished following inhibition of NF-{kappa}B with sodium salicylate, dexamethasone, or by using a viral vector expressing a super-repressor of NF-{kappa}B. Furthermore, when LLC-PK1 cells were transiently transfected with a plasmid containing the A1AR promoter coupled to the luciferase reporter gene (15, 33), an increase in luciferase activity was observed upon exposure of cells to mannitol. In addition, exposure to mannitol resulted in an increase in the steady state levels of A1AR-specific mRNA.

The mechanism underlying the activation of NF-{kappa}B is presently unclear. In cells treated with mannitol, an increase in ROS generation was observed, which was blocked by co-administration of allopurinol. Since allopurinol inhibits xanthine oxidase activity, this finding directly implicates the generation of superoxides and subsequent H2O2 production in the activation of NF-{kappa}B. In support of this contention, we show that mannitol-induced activation of NF-{kappa}B was attenuated following exposure of cells to allopurinol. In preliminary studies, we observed a significant rise in the levels of extracellular adenosine (~2-fold increase) following exposure of LLC-PK1 cells to mannitol, suggesting that adenosine released could serve as a source for substrates by the xanthine oxidase pathway. A role of ROS in mediating the hypertonicity-mediated induction in A1AR expression was further supported by the observation that exposure of these cells to H2O2 mimicked the response to mannitol. Furthermore, the addition of catalase (to scavenge H2O2) reversed the induction in A1AR. In addition, our data show that allopurinol also inhibited the increase in A1AR induced by mannitol. Thus, our data suggest that exposure of kidney cells to osmotic diuretics leads to increased adenosine production, which is metabolized by adenosine deaminase to generate oxygen free radicals via the xanthine oxidase pathway. The generation of free radicals can, in turn, mediate NF-{kappa}B activation and increase A1AR expression. An important implication of this observation is that the nucleoside adenosine could likely regulate expression of its own receptors through generation of ROS and activation of NF-{kappa}B. However, the effect of hypertonic solutions to induce NF-{kappa}B activity seems to be cell-type specific, because a previous study from our laboratory demonstrates that, in smooth muscle cells, mannitol inhibits NF-{kappa}B activity induced by cytokines and LPS (41). A recent study support our contention of an NF-{kappa}B mediated upregulation of the A1AR in DDT1MF-2 cells by chronic hypoxia (42).

The relevance of hypertonicity-mediated increase in A1AR is apparent when one examines the effect of mannitol on cisplatin-mediated cytotoxicity. Incubation of LLC-PK1 cells with mannitol resulted in significant protection of these cells against cisplatin-induced apoptosis. One explanation for this protective action of mannitol is that this agent serves as a scavenger of hydroxyl radicals (43-44) and thereby reduces oxidant-induced peroxidative damage (45). However, we observed that mannitol increased the generation of ROS in LLCPK1 cells. Similarly, our data indicate that exposure of cells to cisplatin was associated with increased evidence of lipid peroxidation (as indicated by increased levels of malondialdehyde), whereas mannitol treatment had no effect on malondialdehyde levels (data not shown). So, it is possible that exposure to hypertonicity induces a low level of oxidative stress in these cells, not associated with increased malondialdehyde levels or significant cytotoxicity. Another event that may contribute to the protective action of mannitol is induction of COX-2 expression in LLC-PK1 cells, as observed for the medullary epithelial cells (10), leading to increased cell survival. However, activation of the A1AR, probably by the increased adenosine release (as described above), appears critical for mediating cytoprotection against cisplatin, since blockade of this receptor by DPCPX abolished the protective response elicited by mannitol. The lack of toxic effect of mannitol in these cells is likely due to the low concentrations of mannitol used (50-100 mM) in our studies, observed in the plasma after mannitol infusion (46).

The exact role of A1AR in the kidneys has been controversial, and there is no consensus as to whether these receptors protect the kidneys or whether they mediate cytotoxicity. However, a recent study by Lee et al. (47) demonstrates convincingly that mice lacking the A1AR exhibit increased apoptosis and necrosis, secondary to renal ischemia and reperfusion. Another recent study (48) demonstrates a cytoprotective role of the A1AR in the kidneys. Adenosine has also been shown to protect human proximal tubular cells from severe ATP depletion injury (49). Furthermore, data from our laboratory (14) provide indirect evidence for a protective effect of A1AR activation, because AR antagonists potentiated the toxicity of cisplatin.

The beneficial effect of agonists which have been described above could be due to a direct action on renal tubular epithelial cells, such as those in the proximal tubules. However, nephrotoxicity might result from A1AR-dependent constriction of the renal afferent arterioles, thereby reducing blood flow to the kidney. It is likely that the net beneficial effect of A1AR activation results when the direct tubular beneficial effects outweigh the direct vasoconstrictor action. We propose that mannitol produces selective induction of A1AR in the renal proximal tubules and other nephron segments, but not the afferent arteriole. Such a scenario is possible since we have shown that mannitol inhibits NF-{kappa}B in cultured vascular smooth muscle cells (41), which would confer a reduction in A1AR expression in the afferent artioles.

Previous studies have demonstrated induction of apoptosis in LLC-PK1 cells by cisplatin (50). The mechanism(s) underlying this event is only recently being elucidated. Cisplatin-induced apoptosis is likely initiated by a number of proteins, which can "sense" DNA damage, such as nuclear excision repair proteins, mismatch repair proteins, DNA-dependent protein kinase, and high-mobility group proteins (51). DNA damage may then be communicated to other proteins involved in cell cycle arrest such as p53 (52), proapoptotic proteins such as Bax and Bak (53) and antiapoptotic protein Bcl-2 (54). Cisplatin-induced apoptosis also involves mitochondrial release of cytochrome c and sequential activation of caspase-8, caspase-3 and caspase-6 (55). Cells resistant to cisplatin toxicity demonstrate high levels of Bcl-2 and a reduction in the accumulation of p53. Bcl-2 suppresses the induction of Bax and thereby prolongs cell survival (56). The exposure of renal inner medullary cells to hyperosmotic stress induces the expression of heat shock proteins (57), p53 and MDM-2 (58), suggesting that these proteins might contribute to the protection afforded by the mannitol-induced hypertonicity. The induction of other anti-apoptotic proteins by hypertonic stress needs to be addressed in the future.

A number of early studies have indicated a beneficial action of mannitol therapy against cisplatin nephrotoxicity (for review see Ref 1). In addition, the use of mannitol is indicated in ischemic injury to the kidney (59). However, recent trends have moved more toward hydration as a treatment alternative to cisplatin-mediated nephrotoxicity (60). While the use of mannitol in this study might not be in line with current therapy for cisplatin-mediated nephrotoxicity, we believe that the present data provide good evidence that activation of NF-{kappa}B and the A1AR by different agents could provide a useful prophylaxis against nephrotoxic drugs. One point to consider is that the induction of the A1AR required longer term exposure of cells to hypertonicity, generally between 16-24 h. We also observed a significant increase in A1AR in vivo 20 h following administration of a single dose of mannitol. This suggests that a treatment strategy worth considering is the prophylactic use of mannitol ~24 h before chemotherapy to induce A1AR expression, in addition to its current use (or hydration) just prior to chemotherapy to enhance drug clearance.

In summary, the present study demonstrates that the A1AR gene could serve as an important target for modulation by osmotic diuretics. Our data suggest that osmotic diuretics, through activation of NF-{kappa}B, could induce expression of the A1AR. In addition, increased adenosine released following exposure to osmotic diuretics could further stimulate A1AR, thereby conferring additional protection to proximal tubular cells. We believe that the combined effect of A1AR activation and induction provides a novel mechanism by which osmotic diuretics protect renal proximal tubular cells against cisplatin-mediated nephrotoxicity.


   FOOTNOTES
 
* This study was funded by National Institutes of Health Grants HL56316-01 (to V. R.) and DC-02396 (to L. P. R.) and by funds from the Central Research Committee, Southern Illinois University School of Medicine (to V. R.). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

§ These authors contributed equally to the completion of this research. Back

** To whom correspondence should be addressed: SIU School of Medicine, Box 19230, Springfield, IL 62974-1222. Tel.: 217-785-2171; Fax: 217-545-0145; E-mail: vramkumar{at}siumed.edu.

1 The abbreviations used are: COX-2, cyclooxygenase-2; A1AR, adenosine A1 receptor; AB-MECA, N6-(4-aminobenzyl)-9-[5-(methylcarbonyl)-{beta}-D-ribofuranosyl]adenine; DPCPX, 8-cyclopentyl-1,3-dipropylxanthine; H2DCFDA, 2',7'-dichlorodihydrofluorescein diacetate; NF-{kappa}B, nuclear factor kappa B; R-PIA, R-phenylisopropyladenosine; CHAPS, 3-[(3-cholamidopropyl)dimethylammonio]-1-propanesulfonate; GTP{gamma}S, guanosine 5'-O-3-thiotriphosphate; PBS, phosphate-buffered saline; MOPS, 4-morpholinepropanesulfonic acid. Back


   ACKNOWLEDGMENTS
 
We thank Dr. Sanjay Maggirwar (Department of Microbiology and Immunology, University of Rochester Medical Center, Rochester, NY) for providing us with an adenovirus vector expressing the mutant form of I{kappa}B-{alpha}, which acts as a superrepressor of NF-{kappa}B.



   REFERENCES
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 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
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