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Glutamine protects against cisplatin-induced nephrotoxicity by decreasing cisplatin accumulation
Journal of Pharmacological Sciences, Volume 127, Issue 1, January 2015, Pages 117-126
Cisplatin is a chemotherapeutic drug but induces acute kidney injury (AKI). Cisplatin-induced AKI depends on several signaling pathways leading to apoptosis in tubular epithelial cells. Glutamine is a substrate for the synthesis of glutathione, the most abundant intracellular thiol and antioxidant, and plays an important role in protecting cells from apoptosis induced by different stimuli. In the present study, we investigated the protective effect of glutamine on cisplatin-induced AKI. Rats were divided into control, glutamine, cisplatin, and cisplatin plus glutamine groups. Glutamine ameliorated renal dysfunction, tissue injury, and cisplatin-induced apoptosis. Cisplatin increased cell death, caspase-3 cleavage, activation of MAPKs and p53, oxidative stress, and mRNA expression of TNF-αand TNFR1 in HK-2 cells. Glutamine treatment reduced cisplatin-induced these changes in HK-2 cells. Notably, glutamine reduced the cisplatin-induced expression of organic cation transporter 2 (OCT2) and cisplatin accumulation. Our results suggest that the protective effect of glutamine on cisplatin is specific for proximal tubular cells and the initial effects may be related to attenuation of cisplatin uptake. Thus, glutamine administration might represent a new strategy for the treatment of cisplatin-induced AKI.
Keywords: Cisplatin, Nephrotoxicity, Glutamine, Acute kidney injury, Cisplatin uptake.
Cisplatin is one of the most effective and potent anticancer drugs for the treatment of solid tumors such as those in lung, head and neck, ovarian, and testicular cancer (1) . Acute kidney injury (AKI), however, is the main side effect observed after cisplatin treatment. Indeed, this side effect has limited the clinical use of cisplatin in 25–30% of patients, even after the first dose. Although nephrotoxicity is a serious consequence of aggressive therapy with cisplatin, the mechanism by which cisplatin selectively damages proximal tubule cells remains unclear (2) . While multiple mechanisms have been documented to contribute to cisplatin nephrotoxicity, how these signaling pathways are integrated to induce renal pathology is largely unknown (3) . Several investigators have demonstrated that cisplatin induces oxidative stress in renal epithelial cells primarily by decreasing antioxidant enzyme activity and depleting intracellular concentrations of glutathione(4) and (5). Numerous studies have now documented the beneficial effects of various antioxidants in cisplatin-induced nephrotoxicity.
Glutamine is a nonessential amino acid, but it is heavily used as a major metabolic fuel as well as a precursor for nucleotide synthesis in fibroblasts, lymphocytes, and macrophages (6) . It has long been known that, under certain physiological circumstances, glutamine serves as a major fuel for the gut, kidney, and immune system. Many cultured mammalian cells use glutamine, within the context of glucose metabolism, as their major carbon source to meet energetic and biosynthetic needs(7), (8), (9), and (10). Glutamine is also a substrate for the synthesis of glutathione (GSH), the most abundant intracellular thiol and antioxidant (11) . Thus, the antioxidant properties of glutamine could contribute to beneficial effects in cells experimentally (12) and glutamine might play an important role in protecting cells from apoptosis induced by different stimuli.
Tubular damage is recognized as a major pathogenic factor in cisplatin-induced nephrotoxicity. The highest concentration of cisplatin in the kidney is reached in the proximal tubule, where cisplatin leads to renal toxicity, tubular injury, and cell death, time and dose dependently(13) and (14). Research has revealed multiple signaling pathways that are responsible for tubular cell injury and death during cisplatin nephrotoxicity(3) and (15). Thus, proximal tubule cell death is considered to be the major pathophysiological mechanism underlying cisplatin nephrotoxicity and AKI and is a limiting factor for the clinical use of cisplatin (3) . Renal tubular cell death afterin vitroexposure to cisplatin has been well documented(16) and (17). However, much less is known concerning the mechanism underlying cisplatin uptake by tubular cells. High sensitivity of tubular cells to cisplatin is partly attributed to the high uptake of cisplatin by these cells.
The kidney is one of the most important organs for maintaining the homeostasis of organic cations (18) . Na+-independent polyspecific organic cation transporters (OCTs) have been postulated for epithelia in the intestine, liver, kidney, and the brain (19) . Regulation of OCTs is important for the secretion of multiple cationic endogenous substances, drugs, and other xenobiotics. A common feature of OCTs is the presence of several potential protein kinase phosphorylation sites in the intracellular loops, suggesting that their activity can be regulated. OCT2 is the main OCT in the human kidney (20) and is located in the basolateral membrane in the proximal tubules (21) . Both basolateral OCT1 and OCT2 and the copper transporter (Ctr1) are involved in cisplatin transport, accumulation, and toxicity(22) and (23). Unfortunately, few studies have reported that antioxidants are involved in cisplatin uptake in a cisplatin-induced nephrotoxicity model.
Multiple pathways are responsible for tubular cell apoptosis during cisplatin-induced nephrotoxicity and spontaneous inhibition of those pathways is usually necessary for global therapeutic effects. Although manyin vivoandin vitrostudies have been proposed to reduce cisplatin-induced nephrotoxicity(24), (25), (26), and (27), the protection is partial, highlighting the need for combined or novel strategies. Thus, alternative strategies are needed that are less toxic that produce more global therapeutic effects. The aim of the present study was to evaluate the role of glutamine as a protective agent against cisplatin-induced cytotoxicityin vivoandin vitro.
2. Materials and methods
2.1. Animal treatment
Animal studies were conducted according to the Gyeongsang National University Guide for Care and Use of Laboratory Animals. Male Sprague Dawley rats (200–230 g) were maintained in a 12/12-h light/dark cycle in a temperature- and humidity-controlled facility. Standard rat chow and water were provided ad libitum. Animals were divided into four groups. The control group received a single injection of saline, intraperitoneally (n = 3). The glutamine group received a single subcutaneous injection of glutamine (1 g/kg body weight) (n = 3). The cisplatin group received a single intraperitoneal injection of cisplatin (10 mg/kg body weight) and then an equal volume of phosphate-buffered saline (PBS) instead of glutamine (n = 10). The cisplatin plus glutamine group first received cisplatin, and then glutamine at day 1 after cisplatin injection (n = 10). All animals were sacrificed at day 3 after cisplatin injection. Blood and tissue samples were collected for analysis of renal function and tissue damage.
2.2. Assessment of renal function
Serum samples were examined for blood urea nitrogen (BUN) and serum creatinine (Bayer, Pittsburgh, PA) in an Autoanalyzer using standard diagnostic kits.
2.3. Renal histology and damage scoring
Sections (5 μm) were stained with hematoxylin and eosin. Tubular injury was defined as tubular epithelial necrosis, cast formation, intratubular debris, and loss of the brush border. Tubular injury was scored by grading the percentage of affected tubules under ×400 magnification: 0, 0%; 0.5, <10%; 1, 10–25%; 2, 26–50%; 3, 51–75%; and 4, 75–100%. To score injured tubules, whole tubular numbers per field were considered to be standard under ×400 magnification. The grading percentage was calculated in each field as follows; injury score (%) = (number of injured tubules/number of whole tubules) × 100. At least 10 areas in the cortex per slide were randomly selected.
2.4. Terminal deoxynucleotidyl transferase dUTP nick end labeling (TUNEL) assay
Detection of DNA fragmentation was performed using a commercial kit (Roche, Indianapolis, IN). Semi-quantitative analysis was performed by counting the number of TUNEL-positive cells per field, in renal tissue, at ×400 magnification. At least 10 areas in the cortex per slide were randomly selected.
2.5. Cell culture
HK-2 human kidney proximal tubular cells (American Type Culture Collection, Manassas, VA) were cultured in renal epithelial basal medium (GIBCO BRL, Grand Island, NY) with manufacture-provided supplements and were subcultured using a 0.05% trypsin–EDTA solution (GIBCO BRL). Cells were incubated with various concentrations of cisplatin or vehicle (PBS) for 15, 30, 60, or 120 min with PBS or glutamine, and then were used for various analyses, such as cell viability assays, measurement of cellular GSH, reverse transcription-polymerase chain reaction (RT-PCR), immunoblotting, and mitochondrial damage assays.
Total RNA was extracted from HK-2 cells using the TRIzol method according to the manufacturer's protocol (GIBCO BRL). Equal amounts (5 μg) of DNA-free total RNA from each sample were converted to cDNA using 200 U of SuperScript II RT (GIBCO BRL) in a 20 μL reaction volume. Reverse transcription was performed using the following conditions: 22 °C for 10 min, followed by 42 °C for 45 min, and finally 95 °C for 5 min. Reaction products (2.0 μL) were subjected to PCR amplification using 1.25 U ofTaqDNA polymerase (Promega, Madison, WI) in a 50 μL reaction volume. Primer sequences for RT-PCR are followings; TAAAGATAATGACCACATCGC (sense) and TATGATGTTTAACCAGTGCAGG (antisense) for OCT-1, AGTTGCCTATACAGTTGGGCTC (sense) and CAGGGCAGAGTAGAAATCC (antisense) for OCT-2, GACCAAGGATTTGAGAAAGTTG (sense) and AGGGAATCTGTGGCTCTACG (antisense) for OCT-3, GACAAGCCTGTAGCCCATGTTGTA (sense) and CAGCCTTGGCCCTTGAAGA (antisense) for TNF-α, GACTGGTTCCTTCTCTTGGT (sense) and GGTGTTCTGTTTCTCCTTAC (antisense) for TNFR1, CCACCCATGGCAAATTCCATGGCA (sense) and TCTAGACGGCAGGTCAGGTCCACC (antisense) for GAPDH. PCR was performed using the BioRad thermal cycler according to the manufacturer's instructions. Equal volumes of the amplification products were analyzed by 1.5%-agarose gel electrophoresis with 0.5 mg/mL of ethidium bromide staining.
2.7. Cell viability assay
Cell viability was determined using the WST-1 assay. Briefly, HK-2 cells (4 × 103cells in 200 μL) were treated with various concentrations of cisplatin or vehicle for each time point with various concentrations (1, 2, 5, or 10 mM) of glutamine, which was administered simultaneously with cisplatin treatment. WST-1 reagents, 2-(4-iodophenyl)-3-(4-nitrophenyl)-5-(2,4-disulfophenyl)-2H-tetrazolium monosodium salt and 1-methoxy-5-methylphenazinium methylsulfate (Dojin Laboratories), were added to cells and incubated for 3 h at 37 °C. Using an enzyme-linked immunoabsorbent assay autoreader, cell viability was determined by measuring the difference between absorbances at 450 nm.
2.8. Measurement of renal GSH levels
GSH content in HK-2 cells was measured using Glutathione Assay kits (Sigma) according to the manufacturer's instructions. Briefly, HK-2 cells were homogenized in 0.5 mL of glutathione reaction buffer containing 0.1 mL of 5% sulfosalicylic acid. To generate NADPH, 20 μL of NADPH Generation Mix and 140 μL of glutathione reaction buffer was mixed and incubated at room temperature for 10 min. Next, 20 μL of either the GSH standard solution or sample solution was added, followed by incubation at room temperature for 5–10 min and a further addition of 20 μL of substrate solution. A microplate reader was used to measure the absorbance at 405 nm (Molecular Devices Corp., Sunnyvale, CA).
2.9. Immunoblot analysis
Kidneys were removed and homogenized in lysis buffer (ProPrep, iNtTON Biotechnology, Inc., Sungnam, Korea). Protein concentration was determined using the bicinchoninic acid reagent from Pierce. Aliquots of protein extracts (60 and 20 μg, kidney tissue and HK cells, respectively) were loaded in each lane for electrophoresis under reducing conditions and were subsequently electroblotted onto nitrocellulose membranes. Blots were incubated in a blocking buffer containing 1% BSA and 2% fat-free milk, and then exposed to the primary antibodies (rabbit anti-caspase-3 (Cell Signaling), OCT-2 (Santa Crutz), p-JNK (Cell Signaling), JNK (Cell Signaling), pERK (Cell Signaling), ERK (Cell Signaling), p-p38 (Cell Signaling), p38 (Cell Signaling), p-p53 (Cell Signaling) and p53 (Cell Signaling)) overnight at 4 °C. Finally, blots were incubated with horseradish peroxidase-conjugated secondary antibody, and antigens on the blots were detected using the enhanced chemiluminescence kit from Pierce.
2.10. Determination of platinum contents
100 mg of tissue sample was dissolved in 250 μL of benzethonium hydroxide and incubated at 55 °C overnight. An aliquot of 50 μL of solubilized tissue sample was diluted with 0.15 N HCl solution to a final volume of 5 mL. The platinum levels in tissue samples were determined using a ICP-AES method. Tissue concentrations are expressed as μg platinum/g tissue.
2.11. Statistical analysis
Data are expressed as means ± SD. Statistical analyses were conducted using the GraphPad Prism software (GraphPad Software, San Diego, CA). Data were evaluated using one-way ANOVA with Tukey's multiple comparison test (for comparison all groups). All statistical testes usedp < 0.05 to indicate significance.
3.1. Glutamine protects against renal functional impairment induced by cisplatin
Glutamine (Cis+GLN) markedly protected cisplatin-treated rats from renal function impairment, with significantly lower BUN and serum creatinine levels at day 3 compared with rats that received saline (Cis) (CisvsCis+GLN, 153 mg/dLvs118 mg/dL, respectively; **p < 0.001 for BUN; CisvsCis+GLN, 2.2 mg/dLvs1.3 mg/dL, **p < 0.001 for serum creatinine; Fig. 1 ). These results indicate that glutamine protected kidneys against cisplatin-induced nephrotoxicity and ameliorated partially renal dysfunction.
3.2. Reduced cisplatin-induced renal tubular injury following glutamine administration
The control (Con) and glutamine-only (GLN) groups demonstrated no renal structural changes ( Fig. 2 A and B). The kidneys of cisplatin-treated rats that received saline showed AKI-associated tubular lesions at day 3, consisting of tubular necrosis, hyaline casts, loss of brush border, debris of the intratubular and flattening of the tubular epithelium (Cis, Fig. 2 A and B). Kidneys from rats that received glutamine had fewer necrotic tubules and tubular casts compared with kidneys from cisplatin group (Cis+GLN, Fig. 2 A and B). These results show that glutamine administration protected kidney from renal injury after cisplatin injection.
3.3. Reduced cisplatin-induced renal tubular apoptosis following glutamine administration
A key target in cisplatin-induced AKI is the tubular epithelium, which undergoes detachment and apoptosis. We used TUNEL staining to investigate whether glutamine could exert anti-apoptotic activity. The number of TUNEL-positive cells in the tubular epithelium was significantly increased in cisplatin-treated rats at day 3, and the extent of cell death was significantly reduced in rats that had received glutamine ( Fig. 3 A). To strengthen these morphological observations and further investigate the ability of glutamine to prevent tubular apoptosis, we measured the expression of cleaved caspase-3, as known an index of apoptosis, in the kidney. Glutamine significantly reduced the expression of cleaved caspase-3 ( Fig. 3 B).
3.4. Reduced mRNA and protein expression of hOCT2 by glutamine in HK cells
Cisplatin-induced nephrotoxicity is known to be mediated via OCT2 in the human kidney(22) and (23). We performed western blot for OCT2 protein expression and measured the Platinum (Pt) concentration for cisplatin accumulation in the kidney at day 3 after cisplatin injection. Expression of OCT2 was increased in cisplatin-treated kidneys while the expression was decreased by glutamine administration in cisplatin-treated rats ( Fig. 4 A). Pt concentration was well correlated with OCT2 expression in the kidneys given to cisplatin. High level of Pt concentration in the kidneys given to cisplatin was significantly reduced in glutamine-treated cisplatin group ( Fig. 4 B). These data suggest that glutamine treatment might be involved in reduced OCT2 expression, and is thought to provide protection against the cisplatin uptake in the kidneys.
3.5. Effect of glutamine on cisplatin-induced cell viability in HK-2 cells
Proximal tubular cell death is considered to be the main pathophysiological event underlying cisplatin nephrotoxicity (3) . To investigate effects of glutaminein vitro, we cultured HK-2 cells. Loss of cell viability was noted after 24 h incubations with various concentrations of cisplatin (10, 20, 30, or 50 μM) ( Fig. 5 A) and glutamine (1, 2, 5, or 10 mM) ( Fig. 5 B). Based on these data, 30 μM cisplatin with 50% cell death was selected for further study. For glutamine, 2 mM was chosen as the concentration in the conventional culture medium; this concentration had no significant effect on the growth rate and cell morphology of HK-2 cells. To confirm the effect of 30 μM cisplatin and/or 2 mM glutamine concentrations, cell viability was assessed in cells treated with cisplatin plus glutamine for 24 h. Cell viability was increased significantly in the cisplatin plus glutamine (Cis+GLN) compared with the cisplatin alone (Cis) group ( Fig. 5 C). To study whether apoptotic-like nuclear morphology was present, HK-2 cells were treated with 30 μM cisplatin and 30 μM cisplatin plus 2 mM glutamine for 24 h, and then stained with 4′,6-diamidino-2-phenylindole (DAPI). Nuclear fragmentation (apoptotic nuclei) was clearly detected in cisplatin-treated cells and glutamine treatment decreased this fragmentation ( Fig. 5 D). Taken together, these results suggest that glutamine attenuated cisplatin-induced apoptosis of proximal tubular cells.
3.6. Reduced mRNA of hOCT2 in renal tubular cells
Cisplatin is taken up into the tubular cells by OCTs(22) and (23). To investigate the involvement of glutamine in the regulation of OCTs expression, we performed RT-PCR.hOCT1mRNA was slightly induced by cisplatin treatment but the level was not changed by glutamine and hOCT3-positive signals were quite strong but the level was not changed by glutamine, similar tohOCT1expression (data not shown). Compared withhOCT1andhOCT3mRNA expression, glutamine decreased level of thehOCT2mRNA expression induced by cisplatin ( Fig. 6 A). Moreover, to verify the onset of OCT2 protein expression by cisplatin treatment, we tried to find the level of OCT2 protein in various times. The levels of OCT2 protein was started to induce at 60 min, peaked at 120 min, and then downed at 24 h after cisplatin treatment. Unfortunately, our results were also not correlated with mRNA ( Fig. 6 A) and protein ( Fig. 6 B) levels for hOCT-2 at 24 h after cisplatin treatment. Both at 60 min and 24 h, the levels seemed to decrease by glutamine but not significant. However, the level was significantly reduced by glutamine at 120 min. Thus, this result suggests that this earlier inhibition of cisplatin uptake by glutamine may be resulted in other cellular signaling events such as MAPKs and p53 activation.
3.7. Effect of glutamine on GSH levels reduced by cisplatin in HK-2 cells
Because caspase-3 cleavage was actively induced by cisplatin, we decided to study whether glutamine modified other steps upstream in the extrinsic apoptosis pathway. As an oxidative stress index, the cellular GSH levels were measured in HK-2 cells treated with 30 μM cisplatin for 24 h. In Fig. 7 , GSH levels were decreased by cisplatin in HK-2 cells and in the presence of glutamine, the levels were significantly increased.
3.8. Effect of glutamine on cisplatin-induced MAPK activation in HK-2 cells
MAPKs activation was augmented in HK-2 cells incubated with cisplatin. Activation of JNK, ERK, and p38 was significantly higher in cisplatin-stimulated cells than in control cells. However, in cells incubated with glutamine and cisplatin, only a slight increase was detected despite cisplatin stimulation ( Fig. 8 ).
3.9. Effect of glutamine on cisplatin-induced p53 activation and upregulation of TNF-α and TNFR1 mRNA expression in HK-2 cells
The signaling pathways that lead to tubular cell apoptosis during cisplatin treatment are complex and remain to be clarified(15) and (28). In rat proximal tubular cells, p53 is activated during cisplatin treatment, and p53 inhibition by pharmacological and molecular approaches ameliorates cisplatin-induced apoptosis(16), (29), (30), and (31). To determine whether p53 was involved in the cytoprotective effect of glutamine, we examined p53 phosphorylation by immunoblot analysis. Activation of p53 by cisplatin was suppressed when cells were treated with glutamine ( Fig. 9 A). The result suggests that p53 inactivation by glutamine may result in less apoptotic death during cisplatin treatment. The death receptor-mediated apoptotic pathway is involved in the pathogenesis of cisplatin-induced AKI(32) and (33). The mRNA level of TNF-αincreased in the cells with 30 μM cisplatin and this increase was inhibited by treatment with 2 mM glutamine. The mRNA level of tumor necrosis factor receptor 1 (TNFR1) was also increased with 30 μM cisplatin and decreased by glutamine treatment ( Fig. 9 B).
We found that cisplatin induced a rapid loss of renal function, as indicated by increases in BUN and serum creatinine levels, which were partially, but significantly, reduced in animals post-treated with glutamine. Consistently, glutamine ameliorated tubular damage and apoptotic cell death in renal tissues. Additionally, inin vitrostudies, we showed that glutamine attenuates cisplatin-induced renal tubular cell injury, most likely in different ways. Glutamine alone had no significant effect on growth, cell confluence, or the monolayer morphology of HK-2 cells. According to our results, glutamine seemed to protect tubular cells against cisplatin toxicity in a sequential manner, initially through regulation of cisplatin accumulation, followed by oxidative stress, and finally through the death receptor-mediated signaling pathway.
AKI is attributed to the accumulation of cisplatin in renal proximal tubular cells and cisplatin is known to permeate proximal tubular cells through apical and basolateral OCTs(13) and (34). Both basolateral OCT1 and OCT2 as well as Ctr1 are involved in cisplatin transport, accumulation, and toxicity(13), (22), and (23). In both cases, apoptosis and accumulation of cisplatin were decreased, but not eliminated, in the presence of competitive organic cations or transporter knockdown(13), (22), and (23). In particular, hOCT2 is a renal OCT isoform and hOCT1 is a hepatic isoform in humans(20) and (35), indicating that the organ-specific toxicity of cisplatin with toxic effects is in the kidney, not in the liver. Because the nephrotoxicity of cisplatin is caused by its uptake via OCT2, it is important whether glutamine treatment involves changes in OCT2 expression. Glutamine treatment affected OCT2 expression and cisplatin accumulation in the kidneys induced by cisplatin ( Fig. 4 ). This result was consistent with OCT2 mRNA expression in vitro ( Fig. 6 ) and a previous report, in which cisplatin interacted with hOCT2, but not with hOCT1 in hOCT1 and 2-transfected cells (13) . We suggest the following possibilities for a protective role of glutamine in cisplatin-induced nephrotoxicity. 1) OCTs have been described to have a big binding pocket with overlapping interaction domains for different substrates(36) and (37). Glutamine may act either to help the binding of a substrate or to upregulate the cellular level of a substrate that can bind in a binding pocket of hOCT2. This can result in less cisplatin uptake and reduced toxicity. Thus, our result suggests that glutamine may play a role in the regulation of cisplatin uptake into proximal tubular cells, although the effect is indirect to hOCT2, and finally reduce cisplatin-induced nephrotoxicity. 2) Regulation of OCTs may alter the secretion and absorption of multiple endogenous substances, drugs, and other xenobiotics, leading to changes in pharmacokinetics of these organic cations (38) . OCTs have several potential protein kinase phosphorylation sites in the intracellular loops, suggesting that their activity may be subject to regulation. Thus, glutamine may also be a factor regulating the activity of OCTs, focused on protein kinase phosphorylation. 3) Several signaling pathways are involved in the regulation of hOCT2-mediated OCT (38) . Whereas protein kinase A (PKA) stimulation and activation of the G receptor-coupled signaling pathway resulted in inhibition of hOCT2-mediated transport, endogenously active Ca2+/calmodulin (CaM) or the CaM-dependent kinases II (CaMKII) and myosin light-chain kinase (MLCK) led to activation of hOCT2-mediated transport. Thus, we cannot exclude the possibility that glutamine could inhibit hOCT2-mediated transport of cisplatin by stimulating PKA, activating the G receptor-coupled signaling pathway, or involvement in the inactivation of endogenous active CaM or the CaMKII and MLCK. However, these possibilities are all hypotheses and none suggest how glutamine could be involved in regulation of OCTs expression in cisplatin-induced nephrotoxicity. We believe that these hypotheses will provide motivation for further studies. As shown in Fig. 4 B, our result indicates that entry of cisplatin in the cell could be partially impaired after treatment with glutamine. Although glutamine is not a substrate of OCT, its administration for renal OCT2 targeting could allow the use of high-dose cisplatin in anticancer therapy without causing nephrotoxicity or maintaining higher cisplatin concentrations in blood.
Both preserving the cellular GSH level and inhibiting the reduction of the GSH level are important for the metabolic activation of cisplatin in the kidney to a more potent toxin. In the present study, the GSH level was decreased by cisplatin treatment. This was probably resulted in using glutathione for cisplatin to be converted to a cisplatin–glutathione conjugate through activation of GGT in tubular cells. However, our results suggest that glutamine can maintain glutathione at basal levels through decreasing cisplatin uptake.
Genotoxic stress induced by cisplatin is accompanied by oxidative stress, a widely studied phenomenon (39) and is related to several cellular signaling activators, including MAPKs. Glutamine certainly reduced the activation of MAPKs by cisplatin, although we cannot say which one is an advanced event between oxidative stress and MAPKs activation in HK-2 cells. Additionally, cisplatin is also related to p53 activation (40) . Pharmacological and genetic inhibition of p53 attenuates cisplatin-induced apoptosis in cultured tubular cells(29) and (31). Furthermore, less renal injury is observed when p53 is inhibited by siRNA in cisplatin-treated animals (41) . Previous studies have shown that p53 phosphorylation is induced within 30 min of cisplatin incubation, followed by p53 stabilization and accumulation in 2–4 h in renal proximal tubular cells(26) and (30). In our study, glutamine might be involved in reducing tubular cell apoptosis by cisplatin, as determined by decreased p53 phosphorylation.
The death receptor-mediated pathway is involved in the pathogenesis of cisplatin-induced AKI and it has been demonstrated that ablation of Fas and TNFR1 protects tubular cells against cisplatin toxicity (33) . Oxidative stress, including reactive oxygen species (ROS), was especially reported to induce renal tubular cell death through the activation of death receptor-mediated apoptotic pathways in cisplatin-induced renal tubular cell death (32) . Whereas death receptor-mediated pathways can be activated by ROS, several studies have reported that such cascades can also generate ROS. Specifically, ROS can act as second messengers during receptor-mediated apoptosis (42) . Thus, oxidative stress and death receptor-mediated apoptotic cascades are considered to interact synergistically. Together with reduced GSH levels, our data suggest that damage of mitochondrial function may be related to receptor-mediated cell death and glutamine may ameliorate cisplatin nephrotoxicity at these levels.
To evaluate the potential loss of the tumoricidal effect of cisplatin caused by the presence of glutamine, we applied cisplatin and glutamine to a tumor-derived cell line. In particular, cisplatin caused cell death in A549 cells, a typical cell type of clinical cisplatin use, as reflected in cell viability.
However, 2 mM of glutamine alone was not significant to cell viability as well as not to affect protection from cisplatin toxicity. The result indicates that the protective effect of glutamine on cell viability is specific to renal cells, compared with tumor cells.
There is only one study about effects of glutamine in cisplatin nephrotoxicity until now. Mora et al. (43) reported that the pretreatment with a single dose of glutamine inhibited the increase in renal glutathione 24 h after cisplatin injection and decreased the lipid peroxidation observed 7 days after cisplatin. However, oral administration of glutamine was not effective to protect the kidney from nephrotoxicity induced by cisplatin. Thus, it is noteworthy that cisplatin uptake, renal dysfunction, tissue damage, and tubular cell death by glutamine was decreased in the present study.
In conclusion, we showed that glutamine attenuated cisplatin-induced nephrotoxicity in different ways. Its contribution may likely start with decreasing cisplatin accumulation. The present study provides a new role for glutamine in decreasing cisplatin-induced nephrotoxicity and glutamine could be a useful treatment for patients with cisplatin-induced acute kidney injury and cancer.
Conflict of interest
This work was supported by Special Clinical Fund from Gyeongsang National University Hospital awarded to Hyun-Jung Kim , Se-Ho Chang [GNUHCRF-2009-001], and by a grant of the Korean Health Technology R&D Project, Ministry of Health and Welfare, Republic of Korea (A100603) awarded to Dong Jun Park.
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a Division of Nephrology, Department of Internal Medicine School of Medicine, Gyeongsang National University, Jinju, Gyeongnam, Republic of Korea
b Institute of Health Sciences, Gyeongsang National University, Jinju, Gyeongnam, Republic of Korea
c Biomedical Research Institute, Gyeongsang National University Hospital, Jinju, Gyeongnam, Republic of Korea
d Division of Hematology-Oncology, Department of Internal Medicine School of Medicine, Gyeongsang National University, Jinju, Gyeongnam, Republic of Korea
e College of Pharmacy and Research Institute of Pharmaceutical Sciences, Gyeongsang National University, Jinju, Gyeongnam, Republic of Korea
∗ Corresponding author. Division of Nephrology, Department of Internal Medicine, Gyeongsang National University Hospital, Gyeongsang National University, Chilam-dong 90, Jinju, Gyeongnam 660-751, Republic of Korea. Tel.: +82 55 750 9267; fax: +82 55 750 9255.
1 These authors contributed equally to this work.
Peer review under responsibility of Japanese Pharmacological Society.
© 2014 Japanese Pharmacological Society, Published by Elsevier B.V.