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Logo of nihpaAbout Author manuscriptsSubmit a manuscriptNIH Public Access; Author Manuscript; Accepted for publication in peer reviewed journal;
Cancer Ther. Author manuscript; available in PMC Jan 9, 2008.
Published in final edited form as:
Cancer Ther. 2003; 1: 47–61.
PMCID: PMC2180401

Cisplatin nephrotoxicity: molecular mechanisms


Cisplatin is one of the most widely used chemotherapeutic agents for the treatment of several human malignancies. The efficacy of cisplatin is dose dependent, but the significant risk of nephrotoxicity frequently hinders the use of higher doses to maximize its antineoplastic effects. Several advances in our understanding of the biochemical and molecular mechanisms underlying cisplatin nephrotoxicity have recently emerged, and are reviewed in this article. Evidence is presented for distinct mechanisms of cisplatin toxicity in actively dividing tumor cells versus the normally quiescent renal proximal tubular epithelial cells. The unexpected role of gamma-glutamyl transpeptidase in cisplatin nephrotoxicity is elucidated. Recent studies demonstrating the ability of proximal tubular cells to metabolize cisplatin to a nephrotoxin are reviewed. The evidence for apoptosis as a major mechanism underlying cisplatin-induced renal cell injury is presented, along with the data exploring the role of specific intracellular pathways that may mediate the programmed cell death. The information gleaned from this review may provide critical clues to novel therapeutic interventions aimed at minimizing cisplatin-induced nephrotoxicity while enhancing its antineoplastic efficacy.

Keywords: cisplatin, nephrotoxicity, molecular mechanisms, thiol compounds, apoptosis, death receptor, caspases, oxidative stress

I. Introduction

Cisplatin is a potent antitumor drug. Cisplatin-based combination chemotherapy regimens are currently used as front-line therapy in the treatment of testicular cancer, ovarian germ cell tumors, epithelial ovarian cancer, head and neck cancer, advanced cervical cancer, bladder cancer, mesothelioma, endometrial cancer, non-small cell lung cancer, malignant melanoma, carcinoids, penile cancer, adrenocorticol carcinoma and carcinoma of unknown primary (Langerak and Dreisbach 2001). Cisplatin-based chemotherapy is used with radiation therapy in the treatment of esophageal cancer, localized cervical cancer and head and neck cancer (Curran 2002). It is used as consolidation therapy for many types of solid tumors that have failed standard treatment regimens. The therapeutic effects of cisplatin are significantly improved by dose escalation. However, high-dose therapy with cisplatin is limited by its cumulative nephrotoxicity and neurotoxicity (O’Dwyer et al, 1999). Its dose-limiting toxicities have spurred the development of the non-nephrotoxic derivative carboplatin and other platinum-based drugs. However, cisplatin is still the drug of choice in many platinum-based therapy regimens, and remains one of the most commonly used chemotherapy drugs. The structure of cisplatin is shown in Figure 1.

Figure 1
Structure of cisplatin.

Cisplatin is toxic to the renal proximal tubules (Gonzales-Vitale et al, 1977). The severity of toxicity in early clinical trials called into question the use of cisplatin as a chemotherapy agent (DeConti et al, 1973). Hydration protocols were developed that reduced the nephrotoxicity and allowed dose escalation to therapeutic levels (Cornelison and Reed 1993). However, even with vigilant hydration, approximately one-third of patients treated with cisplatin have transient elevation of blood urea nitrogen levels or other evidence of kidney damage in the days following cisplatin treatment (Meyer and Madias 1994). Reports of accidental overdoses, all of which have led to renal failure, confirm the potency of cisplatin as a renal toxin in humans (Chu et al, 1993).

Nephrotoxicity is an unusual side effect of chemotherapy in general. Most chemotherapy drugs target pathways that are essential to dividing cells. The rapidly dividing cells in the bone marrow are sensitive to these agents. The dose-limiting toxicity of carboplatin is bone-marrow suppression with cumulative anemia (McKeage 2000). Carboplatin can be administered at doses 5-fold higher than cisplatin without evidence of nephrotoxicity or neurotoxicity. Both cisplatin and carboplatin bind DNA, killing dividing tumor cells (Fink and Howell 2000; Perez 1998). The toxicity of cisplatin towards the quiescent renal proximal tubular cells indicates that there are at least two distinct mechanisms by which cisplatin kills cells, as illustrated in Figure 2.

Figure 2
Proposed biochemical mechanisms of cisplatin nephrotoxocity. See text for details. GGT = gamma-glutamyl transpeptidase; AP-N = diaminopeptidase N.

II. Cisplatin uptake and DNA binding

Cisplatin is transported into cells by the copper transporter Ctr1 (Ishida et al, 2002; Lin et al, 2002). Once inside the cell the chloride ions dissociate from the platinum due to the low intracellular chloride concentrations. The positively charged platinum ion binds cellular nucleophiles in DNA, RNA and proteins (Cohen and Lippard 2001). Experimental evidence indicates that platinum-DNA adducts are the lesion that is toxic to dividing cells, as shown in Figure 2 (Eastman 1999).

Thiols such as the sulfur of GSH will bind to the platinum molecule, replacing one of the chloride ions and preventing binding to other cellular nucleophiles (Berners-Price and Kuchel 1990). Increased intracellular GSH concentrations correlate with decreased platinum-DNA binding in freshly isolated peripheral blood mononuclear cells (Sadowitz et al, 2002). Studies of tumor cell lines have shown a correlation between increased levels of intracellular GSH and resistance to cisplatin (Chen et al, 1995; Hrubisko et al, 1993; Godwin et al, 1992). GSTs are a family of enzymes that catalyze the conjugation of GSH to a variety of substrates. GSTs can also bind the electrophilic substrates and thereby inactivate them (Oakley et al, 1999). Several isoforms of GST have been shown to bind cisplatin in vivo (Sadzuka et al, 1994). There are conflicting data in the literature as to which GST isozymes correlate with cisplatin resistance (Puchalski and Fahl 1990; Leyland-Jones et al, 1991; Nakagawa et al, 1990; Ban et al, 1996; Townsend et al, 1992; Nishimura et al, 1998; Cheng et al, 1997; Hamada et al, 1994; Kigawa et al, 1998; Germain et al, 1996; van der Zee et al, 1992; Murphy et al, 1992). Of the studies in which increased resistance to cisplatin was observed, none determined whether the inactivation of cisplatin was due to GST binding to the cisplatin or catalyzing its conjugation to GSH.

III. Evidence for two distinct mechanisms of cisplatin toxicity

A series of studies on the role of the enzyme gamma-glutamyl transpeptidase (GGT) in cisplatin toxicity revealed that in tumor cells GGT expression increased resistance to cisplatin, while in kidney GGT expression made the cells sensitive to cisplatin toxicity (Hanigan et al, 1999; Hanigan et al, 2001). GGT is a cell surface enzyme that cleaves the gamma-glutamyl bonds. GGT cleaves extracellular GSH into glutamic acid and cysteinyl-glycine (Figure 2). Cysteinyl-glycine is cleaved into cysteine and glycine by diaminopeptidase N (McIntyre and Curthoys 1982). Thus, by initiating the cleavage of extracellular GSH into its constituent amino acids, GGT provides the cell with an increased supply of cysteine (Hanigan and Ricketts 1993). In rapidly dividing cells, cysteine can become limiting for cell growth and for intracellular GSH synthesis. Transfection of human prostate tumor cells with GGT increased their growth rate in nude mice and increased their resistance to cisplatin (Hanigan et al, 1999). In contrast, the high level of GGT expression in renal proximal tubular cells renders them sensitive to cisplatin toxicity. Inhibition of GGT blocked the nephrotoxicity of cisplatin in both rats and mice (Hanigan et al, 1994; Townsend and Hanigan 2002). Cisplatin is not toxic to the kidneys in GGT knockout mice (Hanigan et al, 2001). The disparate roles of GGT in the antitumor activity and nephrotoxicity of cisplatin suggest that the mechanism by which cisplatin kills tumor cells is distinct from the mechanism by which it kills the proximal tubular cells in the kidney.

IV. Role of GGT in the nephrotoxicity of halogenated alkenes

Cisplatin is not unique in its requirement for GGT activity to exert its nephrotoxic effects. The nephrotoxic halogenated alkenes, which kill the proximal tubular cells in the kidney, also require GGT activity for their metabolism to a nephrotoxin (Dekant 2001). These compounds include hexachlorobutadiene, dichloroacetylene and trichloroethylene. The halogenated alkenes are conjugated to glutathione (GSH) in the liver and the kidney (Lash et al, 1998). The GSTs that catalyze the conjugation have not been identified, but the reaction rates of several of the nephrotoxic halogenated alkenes are higher with the microsomal GSTs than with the cytosolic GSTs (Wolf et al, 1984). Hexachlorobutadiene is conjugated to GSH predominantly in the liver and the GSH-conjugate is excreted into the bile (Nash et al, 1984). The metabolite is further processed through a hepatobiliary route (Anders and Dekant 1998). Bile duct cells express high levels of GGT and aminopeptidase N on their cell surface. GSH-conjugates in the bile are cleaved to cysteinyl-glycine (cys-gly)-conjugates by GGT and to cysteine-S-conjugates by aminodipeptidase N. The cysteine-S-conjugate is reabsorbed by the intestine and transported in the serum to the kidney. The kidney also contains GSTs that can metabolize the halogenated alkenes (Hassall et al, 1984). Renal GSH-conjugates would have to be excreted from the cell to undergo further processing by GGT and diaminopeptidase N expressed the surface of the proximal tubular cells. The cysteine-S-conjugates are transported into the renal proximal tubular cells by a carrier-mediated process (Wright et al, 1998), and are metabolized to a reactive thiol by a PLP-dependent cysteine-S-conjugate-beta-lyase. The enzyme that catalyzes this reaction has not been identified, although cysteine-S-conjugate beta-lyase activity has been found in the cytosolic and mitochondrial fractions of the kidney (Cooper et al, 2002). The thioacylating metabolites produced by cysteine-S-conjugate beta-lyase are highly reactive, and will bind to proteins in the proximity of the beta-lyase (Bruschi et al, 1993). Mitochondrial aspartate aminotransferase, which has cysteine-S-conjugate beta-lyase activity, is modified by the thioacylating products and is one of the likely candidates for the cysteine-S-conjugate beta-lyase that metabolizes the halogenated alkenes (Bruschi et al, 1998). The mitochondria of the proximal tubular cells are the primary target of haloalkene-induced toxicity as has been observed with cisplatin (Hayden et al, 1991; Brady et al, 1990; Lash et al, 1986; Hayden and Stevens 1990; Chen et al, 2001; Anders 1995). The cysteine-conjugates of the halogenated alkenes have been shown to induce either apoptosis or necrosis in LLC-PK1 cells depending on the chemical structure of the compound and the antioxidant status of the cell (Zhan et al, 1999; van de Water et al, 2001).

V. Evidence that cisplatin is metabolized to a nephrotoxin

There is compelling evidence from both in vivo studies and cell culture that cisplatin is metabolized to a nephrotoxin through a GSH-conjugate intermediate as are the halogenated alkenes. The proposed pathway is shown in Figure 2. Platinum-GSH conjugates may be formed in either the liver or the kidney. Studies with 195mPt-labeled cisplatin show that following injection the highest levels of platinum are found in the liver and kidney (Lange et al, 1973; Benard et al, 1983). Platinum accumulation in the liver is transient. Biliary excretion of cisplatin accounts for approximately 1% of the administered dose in 6 hr (Siddik et al, 1987). The structures of the platinum-containing compounds that are excreted into the bile have not been identified. Metabolic intermediates of cisplatin have been identified in rat serum. Seven platinum-containing species were resolved by HPLC from serum within 15 min of cisplatin injection (Daley-Yates and McBrien 1984). The mixture was more nephrotoxic than equimolar cisplatin. Inhibiting the conjugation of cisplatin to GSH has been shown to reduce cisplatin nephrotoxicity. In mice, systemic inhibition of GSTs with ketoprofen reduced cisplatin nephrotoxicity (Sadzuka et al, 1994a; Sadzuka et al, 1994). In rats, inhibition of GSH synthesis also protected against the nephrotoxicity of cisplatin (Mayer et al, 1987). The nephrotoxic platinum-conjugates are very unstable relative to the metabolic intermediates of the halogenated alkenes. The GSH and cysteine-conjugates of the halogenated alkenes can be synthesized and purified whereas the nephrotoxic platinum-conjugates are unstable in solution (Kramer et al, 1987; Gaskin et al, 1995; Townsend et al, 2003). At least some of the platinum-GSH conjugates may be formed and metabolized within the kidney (Mistry et al, 1989). Cisplatin-GSH conjugates can be transported out of the cell by MRP2 (cMOAT), a member of the multidrug resistance-associated protein family of efflux pumps (Ishikawa et al, 1996; Borst et al, 2000). There is evidence that other pumps that have not yet been defined can also serve this function (Ueda et al, 1999). GSH-conjugates are metabolized extracellularly to cysteinyl-glycine-conjugates by GGT, which is a cell surface enzyme. Inhibition of GGT has been shown to block the nephrotoxicity of cisplatin in both rats and mice (Hanigan et al, 1994; Townsend and Hanigan 2002). Cisplatin is not nephrotoxic in GGT-knockout mice (Hanigan et al, 2001). Cysteinyl-glycine conjugates are cleaved by aminopeptidase N which is also on the cell surface (Hughey et al, 1978; McIntyre and Curthoys 1982). AOAA, an inhibitor of PLP-dependent enzymes, blocks the nephrotoxicity of cisplatin (Townsend and Hanigan 2002). AOAA also blocks the final enzymatic step in the bioactivation of the nephrotoxic halogenated alkenes. AOAA blocks the beta-elimination reaction that converts the cysteine-S-conjugates of the halogenated alkenes to reactive thiols (Lash et al, 1994; Elfarra et al, 1986). These data demonstrate that cisplatin can undergo enzymatic activation to a metabolite that is more toxic than the parent compound.

Confluent cultures of proximal tubular cells have been used to study cisplatin nephrotoxicity (Montine and Borch 1988; Kroning et al. 1999; Legallicier et al. 1996; Park et al. 2002; Townsend et al. 2003). The toxicity of cisplatin toward confluent monolayers of proximal tubular cells suggests that cells express the enzymes and transporters required in each step of the bioactivation of cisplatin to a reactive thiol. However, the efficiency with which they conjugate cisplatin to GSH may not be optimal. Preincubating cisplatin with equimolar cisplatin for up to 30 min potentiated the toxicity of cisplatin toward confluent monolayers of LLC-PK1 cells (Townsend et al. 2003). Examination of the incubation mixtures revealed a monoplatinum-mono-GSH conjugate and a diplatinum-monoGSH conjugate that formed spontaneously in the solution (Townsend et al. 2003). With prolonged incubation, the cisplatin-GSH solution became non-toxic in parallel with increased formation of the diplatinum-monoGSH conjugate. Solutions containing the monoplatinum-monoGSH were more toxic than equimolar cisplatin. The increased toxicity due to the presence of the platinum-GSH conjugate could be blocked by inhibiting GGT (Townsend et al. 2003). Preincubation of cisplatin with equimolar N-acetyl-cysteine (NAC) also potentiated the toxicity of the cisplatin, and this correlated with the formation of a monoplatinum-monoNAC conjugate (Townsend et al. 2003). NAC is deacetylated to cysteine (Commandeur et al. 1991). The transporter of the platinum-cystiene conjugate into the cell has not been identified. In vivo, AOAA inhibits the nephrotoxicity of cisplatin (Townsend and Hanigan 2002). In LLC-PK1 cells, AOAA blocks the toxicity of the platinum-NAC conjugate, as well as the platinum-conjugates that are upstream in the metabolic pathway (Townsend et al. 2003). Prolonged incubation of cisplatin with NAC inactivated solution and correlated with the formation of diplatinum-monoNAC conjugates. As detailed below, the molecular mechanism by which cisplatin kills renal cells is dependent on the concentration of cisplatin and the antioxidant status of the cell (Lee et al. 2001; Park et al. 2002).

VI. Protection from cisplatin nephrotoxicity by thiol compounds

Thiol compounds have been used clinically to reduce the nephrotoxicity of cisplatin. High doses of GSH injected intravenously within 30 min of cisplatin administration is protective (Tedeschi et al. 1991; Smyth et al. 1997). The amount of GSH that is necessary to achieve this protective effect in humans is 30 to 40-fold higher than the dose of cisplatin. These data may appear to contradict the hypothesis that the formation of a cisplatin-GSH conjugate activates cisplatin to a nephrotoxin, but high concentrations of GSH can protect against cisplatin nephrotoxicity by serving as a competitive inhibitor of GGT activity (Hanigan et al. 1994). GSH is the major physiologic substrate for GGT (Curthoys and Hughey 1979; Hanigan and Pitot 1985). GGT is localized to the cell surface and would be inhibited by high levels of GSH in the extracellular fluid (Hanigan and Frierson Jr. 1996). By inhibiting GGT activity, GSH would reduce the metabolism of the cisplatin-GSH to a cisplatin-cysteinyl-glycine conjugate. A large number of sulfur-containing compounds such as procainamide, diethyldithiocarbamate, N-methyl-D-glucaminedithio carbamate, methimazole, sulfathiazole and the prodrug Amifostine have been shown to reduce the nephrotoxicity of cisplatin without inhibiting its antitumor effect (Jones et al. 1992; Borch et al. 1980; Jones et al. 1992; Yee et al. 1994; Korst et al. 1998; Osman et al. 2000). Procainamide is an antiarrhythmic drug that binds cisplatin forming a procainamide-cisplatin complex (Esposito et al. 1996). In the presence of procainamide more platinum is bound to DNA, which would explain the maintenance of the antitumor activity (Viale et al. 2000). The binding of procainamide to the cisplatin may prevent the formation of a cisplatin-GSH complex and thereby protect against the metabolism of cisplatin to a nephrotoxin. Other thiol compounds may also be binding cisplatin in a complex that does not prevent the binding of the platinum to DNA but does prevent the formation of a GSH-cisplatin conjugate. In contrast, sodium thiosulfate and biotin inhibit both the nephrotoxicity and antitumor activity of cisplatin (Howell and Taetle 1980; Uozumi et al. 1984; Jones et al. 1992).

VII. Evidence for cisplatin-induced apoptosis in renal tubular cells

It has long been recognized that in acute renal failure induced by nephrotoxins or other causes, renal tubular cells suffer a spectrum of cytotoxic injuries, ranging from mild sublethal changes to a catastrophic necrotic death characterized by swelling and rupture of cells and activation of an inflammatory response (Thadani et al. 1996). The ensuing clinical syndrome has conventionally been designated by the term acute tubular necrosis. However, it is now known that at least two distinct mechanisms may be responsible for renal tubular cell death following injury, depending primarily on the extent and severity of the insult. While extensive injury can lead to necrotic cell death, increasing evidence has indicated that the less severe renal injuries most commonly encountered in modern clinical practice are associated predominantly with patchy apoptosis of tubular epithelial cells (Ueda et al. 2000; Levine and Lieberthal 2001). Apoptosis or programmed cell death is characterized by distinct morphologic changes consisting of cell shrinkage, nuclear condensation, and internucleosomal DNA fragmentation (Kerr et al. 1972). It is well known that induction of programmed cell death is a common mechanism by which cytotoxic drugs such as cisplatin kill tumor cells (Friesen et al. 1999). In recent years, kidney tubular cell apoptosis has been detected in an increasing array of renal disorders, and is emerging as a final common pathway in response to a variety of cellular stresses applied at an intensity below the threshold for necrosis (Ueda et al, 2000; Levine and Lieberthal 2001). This observation especially holds true for cisplatin nephrotoxicity, in which necrotic cell death is encountered with higher doses whereas lower concentrations induce apoptosis (Lieberthal et al. 1996; Lau 1999).

The past decade has witnessed an explosion of information on the molecular and cellular biology of programmed cell death, and specific intracellular proteases belonging to the caspase family have emerged as crucial effectors of apoptosis (Thornberry and Lazebnik 1998). Members of this family (now totaling at least 14) are expressed as pro-enzymes and require activation by upstream signal transduction pathways to commit a cell into the execution phase of apoptosis. It is convenient to classify the major intracellular apoptotic pathways according to the type of pro-caspase that is activated. Activation of the initiator pro-caspase 8 results predominantly from signaling via integral membrane death receptor proteins such as Fas and TNFR1 (Ashkenazi and Dixit 1996). On the other hand, activation of the initiator pro-caspase 9 is dependent primarily on mitochondrial signaling pathways regulated by members of the Bcl-2 family of proteins (Adams and Cory 1996; Brenner and Kroemer 2000). Activation of pro-apoptotic Bcl-2 family members such as Bax can trigger a sequence of events that leads to alterations in mitochondrial permeability, release of mitochondrial cytochrome c into the cytosol, and activation of pro-caspase 9 (Korsmeyer et al. 2000; Goldstein et al. 2000). The anti-apoptotic Bcl-2 family members such as Bcl-2 itself play a pivotal protective role by preserving mitochondrial function and preventing release of cytochrome c (Adams and Cory 1996). Several levels of cross-talk exist between the caspase 8- and 9-dependent pathways. First, initial activation of caspase 8 via death receptor pathways can induce the mitochondrial translocation of BID, a pro-apoptotic member of the Bcl-2 family, with resultant cytochrome c release and activation of caspase 9 (Luo et al. 1998). Second, the p53 gene is a potent transcription factor that regulates apoptosis most notably by activating pro-apoptotic Bcl-2 family members as well as the Fas-FADD axis (Burns and El-Deiry 1999). Third, both pathways culminate in the activation of caspase 3, with subsequent entry into the “execution” phase of apoptosis (Thornberry and Lazebnik, 1996). In this review, the evidence for cisplatin-induced apoptosis in renal tubular cells will first be presented. In the subsequent sections, the role of each of these pathways in cisplatin-induced apoptosis of renal tubular cells will be reviewed.

Studies from a number of laboratories over the past few years have demonstrated that cisplatin can induce apoptosis in renal tubular cells both in animal models and in cell culture systems. Using an established mouse model of cisplatin nephrotoxicity (intraperitoneal injection of cisplatin 20 mg/kg body weight), the appearance of apoptotic epithelial cells by Tunel assay was first shown predominantly in the distal tubular and collecting duct (Megyesi et al. 1998). Subsequent studies have confirmed and extended these findings. Apoptosis in this model has now been documented by a variety of additional methods (including hematoxylin-eosin staining, DNA laddering, and electron microscopy) to occur in both distal and proximal tubular cells, predominantly in the outer medullary region (Shiraihi et al. 2000; Tsuruya et al. 2003). Apoptosis was evident within 3 days of cisplatin injection, temporally correlating with the onset of renal dysfunction. Several analogous studies have also been completed in a rat model (cisplatin 5 mg/kg intraperitoneally), with comparable results (Zhou et al. 1999; Miyaji et al. 2001; Huang et al. 2001; Nishikawa et al. 2001; Chang et al. 2002). Importantly, several maneuvers aimed at attenuating the extent of tubular cell apoptosis have also resulted in amelioration of renal dysfunction (Megyesi et al, 1998; Zhou et al, 1999; Shiraishi et al, 2000, Miyaji et al, 2001; Nishikawa et al. 2001; Chang et al. 2002; Tsuruya et al. 2003). These findings have provided support for the notion that inhibition of apoptosis may represent a novel and powerful therapeutic strategy for the prevention and treatment of cisplatin nephrotoxicity, and have stimulated recent efforts at identifying the programmed cell death pathways induced by cisplatin. These investigations have been facilitated by the establishment and elucidation of cell culture models. Cisplatin was first shown to cause apoptosis in cultured mouse proximal tubular cells (Lieberthal et al. 1996; Takeda et al. 1997; Takeda et al. 1998; Fukuoka et al. 1998). Several subsequent studies have now documented the ability of cisplatin to induce apoptosis in pig proximal tubular (Kruidering et al. 1998; Lau 1999; Okuda et al. 1999; Zhan et al. 1999; Kaushal et al. 2001; Park et al. 2002), human proximal tubular (Razzaque et al. 1999; van de Water et al. 2000; Nowak 2002; Cummings and Schnellmann 2002), and even collecting duct cells (Liu et al. 1998; Lee et al. 2001). A recurrent theme gleaned from these works is that cisplatin induces apoptosis in a dose- and duration-dependent manner, and that while this agent activates programmed cell death at lower (10–100 μM) doses, it can also result in necrotic cell death at higher (200–800 μM) concentrations.

VIII. Activation of death receptor pathways

Renal tubular epithelial cells upregulate Fas-dependent pathways and undergo apoptosis following ischemic injury both in vitro (Feldenberg et al, 1999) and in vivo (Nogae et al, 1998), and activation of the Fas pathway is a common mechanism by which cytotoxic drugs such as cisplatin induce apoptosis in tumor cells (Friesen et al, 1999). A possible role for this pathway in cisplatin nephrotoxicity was first suggested by experiments done in cultured human proximal tubular epithelial cells, in which cisplatin (20–80 μM) resulted in apoptosis which was temporally correlated with an increased expression of Fas (Razzaque et al, 1999). A subsequent detailed analysis in mouse and rat kidney as well as in cultured murine proximal tubular cells has recently provided substantial support for this mechanism (Tsuruya et al, 2003). In this study, wild-type mice subjected to cisplatin displayed renal dysfunction, tubular cell apoptosis, and an increase in mRNAs encoding Fas and Fas ligand in the kidneys, whereas Fas-mutant B6-lpr/lpr mice exhibited an attenuated response. Proximal tubular cells cultured from wild-type mice responded to cisplatin by upregulation of Fas mRNA and protein, increase in caspase 8 activity, and apoptosis, all of which were blunted in cells from kidneys of Fas-mutant mice (Tsuruya et al, 2003). Furthermore, the cells derived from wild-type mice exhibited increased TNF-α secretion following cisplatin exposure, and kidneys of TNFR1-deficient mice displayed an attenuated functional and apoptotic response to cisplatin (Ramesh and Reeves 2002; Tsuruya et al, 2003). Taken together, these results suggest that cisplatin induces renal epithelial cell apoptosis at least in part via activation of death receptor pathways, as illustrated in Figure 3. However, it is worth emphasizing that following cisplatin treatment, both Fas-mutant and TNFR1-deficient animals continued to exhibit a significant number of Tunel-positive apoptotic tubular cells (albeit about 50% less than in wild-type mice), and displayed only a partial protection of renal function (a rise in BUN of approximately 50% compared to wild-type). These observations indicate that other pathways leading to apoptosis must additionally be invoked in cisplatin nephrotoxicity. Also, the precise mechanism by which cisplatin (or its toxic metabolite) activates either the Fas- or the TNFR1-dependent pathways is unknown. Based on results obtained from other cell types, a possible role for p53 (Miyashita and Reed 1995; Muller et al, 1998; Burns and El-Deiry 1999; Chandel et al, 2000) and oxidative stress (Bauer et al, 1998) as inducers of Fas-mediated apoptosis in this situation has been postulated (Figure 3). These notions are detailed in subsequent sections.

Figure 3
Proposed apoptotic pathways in cisplatin nephrotoxicity. See text for details. ROM = reactive oxygen metabolites; Casp = caspase. Death receptor pathways are shown in green, and mitochondrial pathways are in purple. All arrows indicate stimulatory influences, ...

IX. Activation of mitochondrial pathways

Substantial evidence is now available to implicate the mitochondrial apoptotic pathways in cisplatin-induced tubular cell death. Initial studies completed in cultured proximal tubular cells demonstrated that forced overexpression of Bcl-2 rendered the cells partially resistant to cisplatin-induced apoptosis (Takeda et al, 1997; Zhan et al, 1999). This observation has been confirmed in rats in which pretreatment with uranyl acetate caused a significant upregulation of Bcl-2 in the kidney, and ameliorated cisplatin-induced tubular cell apoptosis as well as the ensuing renal dysfunction (Zhou et al, 1999).

According to the “dueling dimers” prediction, Bcl-2 inhibits apoptosis primarily by opposing pro-apoptotic molecules in the mitochondrial pathway (Oltvai and Korsmeyer 1994). Indeed, cisplatin nephrotoxicity has been associated with increased renal expression of Bax in vivo (Huang et al, 2001), and with translocation of Bax from the cytosol to a membrane fraction in cultured cells (Lee et al, 2001). A detailed analysis of mitochondrial pathways activated by cisplatin has recently been completed in cultured proximal tubular cells (Park et al, 2002). Cisplatin-induced apoptosis was associated with increased caspase 9 activity, and the DNA laddering was inhibited by pretreatment with specific caspase 9 inhibitors, thereby implicating the mitochondrial mechanisms. Cisplatin also triggered a duration-dependent translocation of Bax from the cytosol to the mitochondria, induction of mitochondrial permeability transition, and release of cytochrome c into the cytosol (Park et al, 2002). Collectively, these studies suggest a major role for mitochondrial pathways in cisplatin-induced apoptosis, at least in cultured renal epithelial cells, as shown in Figure 3. The relative contribution of these pathways in the analogous in vivo situation remains under active investigation. Also, the precise mechanisms by which cisplatin activates the mitochondrial apoptotic pathways is unclear.

It has been known for a long time that cisplatin-induced nephrotoxicity is accompanied by alterations in mitochondrial function and structure (Gordon and Gattone 1986).

This issue has recently been revisited, especially in reference to injury induced by oxidative stress (Nishikawa et al, 2001; Chang et al, 2002), and is reviewed in the section on role of oxidative damage. At any rate, cisplatin-induced oxidative stress in renal mitochondria has been postulated to result in the release of cytochrome c into the cytosol, with resultant activation of caspase 9 (Nishikawa et al, 2001). However, this does not explain the well-documented observation of Bax translocation from the cytosol to the mitochondria following cisplatin exposure (Lee et al, 2001; Park et al, 2002).

X. Activation of caspases

Since caspases represent the final mediators of apoptosis in most situations, several authors have sought evidence for caspase activation following cisplatin exposure. A role for the “executioner” caspase 3 was first suggested in cultured proximal tubular cells, which responded to cisplatin exposure by increasing caspase 3 activity in a dose- and duration-dependent manner (Fukuoka et al, 1998; Lau 1999). Cell-permeant inhibitors of caspase 3, such as zVAD.fmk and DEVD.CHO, were effective in protecting against cisplatin-induced DNA damage and cell death (Zhan et al, 1999; Kaushal et al, 2001). Subsequent studies searching for activation of the more proximate caspases have revealed a dramatic activation of caspase 9, and to a lesser extent of caspase 8 (Kaushal et al, 2002; Park et al, 2002). These findings are in agreement with the collective previous data implicating both the mitochondrial and the death receptor pathways in cisplatin-induced nephrotoxocity. However, more detailed analyses have shown that while specific inhibition of caspase 9 (with LEHD-CHO) largely prevented cisplatin-induced DNA fragmentation in cultured cells, a specific inhibitor of caspase 8 (IETD.fmk) was much less efficacious (Park et al, 2002). Furthermore, recent results have indicated that cisplatin-induced apoptosis in cultured renal proximal tubular cells proceeds via both caspase-dependent and caspase-independent pathways, and that inhibition of the executioner caspase 3 blocks only about 50% of cisplatin-induced apoptosis (Cummings and Schnellmann 2002; Nowak 2002). The potential efficacy of caspase inhibition in a complex in vivo system such as the kidney in response to cisplatin is currently unknown.

XI. Role of regulatory pathways

In order to reconcile the seemingly disparate results indicating activation of both death receptor and mitochondrial pathways in cisplatin-induced nephrotoxicity, investigators have examined the regulatory mechanisms that can account for “cross-talk” between the two. Studies have now documented the rapid activation and nuclear translocation of p53 in response to cisplatin both in kidneys (Miyaji et al, 2001) and in cultured renal proximal tubular cells (Cummings and Schnellmann 2002). Inhibition of p53 prior to cisplatin exposure blunted the apoptotic response by 50%, attesting to the importance of this regulatory pathway at least in vitro. It is well known that both cisplatin-induced DNA damage and cisplatin-induced oxidant stress are potent activators of p53 (Muller et al, 1998; Chandel et al, 2000), and that p53 can in turn activate both Bax as well as the Fas-FADD axis (Miyashita and Reed 1995; Burns and El-Deiry 1999). It is therefore likely that this regulatory mechanism may play a crucial role in cisplatin-induced apoptosis.

It is well known that one of the responses of the normally quiescent renal tubular epithelial cell to damage induced by cisplatin includes entry into the cell cycle with subsequent cell proliferation, which presumably represents a reparative event (Megyesi et al, 1995; Sano et al, 2000). However, cisplatin also results in DNA damage (Zamble and Lippard 1995), and uncontrolled proliferation of these cells would be expected to result in apoptotic and/or necrotic cell death. Fortunately, renal epithelial cells have evolved mechanisms to prevent further progression of the cell cycle, allowing time and opportunity for their DNA to be repaired and the cell to then complete the regeneration and replacement process (Megyesi et al, 1998; Megyesi et al, 2002). Candidate proteins that contribute to the cell cycle arrest required for DNA repair include p21 and 14-3-3σ. Several studies have now documented that cisplatin-induced nephrotoxicity is associated with upregulation of p21 mRNA (Megyesi et al, 1996; Huang et al, 2001) and protein (Megyesi et al, 1998; Miyaji et al, 2001; Megyesi et al, 2002). While it is well known that the p53 gene is a potent regulator of p21 (El-Deiry et al, 1993), induction of p21 in cisplatin nephrotoxicity appears to be p53-dependent as well as -independent (Megyesi et al, 1996). Mice lacking p21 develop normally, but respond to cisplatin with a more severe nephrotoxic injury, including a more rapid onset of renal failure, uncoordinated progression into S-phase of the cell cycle, and increased apoptosis (Megyesi et al, 1998). Recent work has suggested a role for another cell cycle inhibitor, 14-3-3σ. Following cisplatin exposure, there is a marked induction of 14-3-3σ mRNA and protein in the kidney tubular cells both in vivo and in vitro (Megyesi et al, 2002). Both p21 and 14-3-3σ are known to be induced following DNA-damaging injury, at least in part via a p53-dependent mechanism (Hermeking et al, 1997). Both are over-expressed in terminally differentiating epithelia, both are required for proper coordination of the cell cycle, and the absence of either of these factors can accelerate apoptosis (Megyesi et al, 2002).

It is recognized that apoptosis represents a default pathway in most cells, and can be activated by a relative deficiency of a variety of “survival factors” (Raff 1992). One example of a survival factor for kidney tubular cells following cisplatin injury is hepatocyte growth factor (HGF). Kidney mRNA levels for HGF are rapidly induced by ischemic or nephrotoxic injury (Liu et al, 1998), and administration of exogenous HGF ameliorates the renal dysfunction induced by cisplatin in vivo by enhancing tubular repair processes (Kawaida et al, 1994). It has recently been shown by a variety of assays that forced overexpression of HGF in cultured renal tubular cells partially inhibited the apoptotic response to cisplatin incubation (Liu et al, 1998). Whether HGF exerts its beneficial effects in vivo also by protecting tubular cells from cisplatin-induced apoptotic death is not known.

XII. Role of oxidative stress and mitochondrial dysfunction

The mechanisms by which cisplatin activates the myriad of apoptotic pathways outlined above remain unclear. However, a role for cisplatin-induced oxidative stress may provide an attractive hypothesis (Baliga et al, 1997). Several studies have now documented the importance of reactive oxygen metabolites (ROM) in cisplatin-induced renal cell apoptosis (Ueda et al, 2000). It is well known that mitochondria continuously produce ROM such as superoxide (Richter et al, 1995). Mitochondria also continuously scavenge ROM via the action of antioxidant enzymes such as superoxide dismutase, glutathione peroxidase, catalase, and glutathione S-transferase (Richter et al, 1995). Cisplatin is known to accumulate in mitochondria of renal epithelial cells (Singh 1989; Gemba and Fukuishi 1991). Several investigators have demonstrated that cisplatin induces ROS in renal epithelial cells primarily by decreasing the activity of antioxidant enzymes and by depleting intracellular concentrations of GSH (Sadzuka et al, 1992; Kruidering et al, 1997; Husain et al, 1998; Huang et al, 2001). A large number of studies have now accumulated documenting the beneficial effects of a variety of antioxidants in cisplatin-induced nephrotoxicity. Agents such as superoxide dismutase, dimethylthiourea, and GSH have been shown to reduce the degree of renal failure and tubular cell damage when administered simultaneously with cisplatin in rats (McGinness et al, 1978; Sadzuka et al, 1992; Matsushima et al, 1998). Antioxidants such as GSH, superoxide dismutase, catalase, deferoxamine, probucol, and heme oxygenase-1 specifically provide partial protection against cisplatin-induced apoptosis in cultured renal epithelial cells (Lieberthal et al, 1996; Okuda et al, 2000; Shiraishi et al, 2000). Furthermore, significant attenuation of cisplatin-induced apoptosis and renal failure in animal models have resulted from maneuvers such as treatment with the hydroxyl radical scavenger DMTU (Zhou et al, 1999), targeted proximal tubular delivery of superoxide dismutase (Nishikawa et al, 2001), and pre-treatment with L-carnitine (Chang et al, 2002). Reactive oxygen molecules can trigger several of the apoptotic mechanisms activated by cisplatin (Figure 3). For example, ROM can induce Fas (Bauer et al, 1998), activate p53 (Chandel et al, 2000), alter mitochondrial permeability (Kruidering et al, 1997; Nowak 2002), release cytochrome c into the cytosol (Reed 1997), and even directly activate caspases (Higuchi et al, 1998).

However, one recent study has suggested that at least in cultured proximal tubular cells, the primary cause of cell death following cisplatin exposure is not ROM formation per se (Kruidering et al, 1996). Rather, cisplatin-induced mitochondrial dysfunction with consequent induction of cell death pathways appeared to be the underlying mechanism. Several studies have demonstrated that cisplatin causes mitochondrial dysfunction in kidney epithelial cells (Gordon and Gattone 1986; Brady et al, 1990; Brady et al, 1993; Kruiderink et al, 1996; Nowak 2002). The major mitochondrial targets of cisplatin appear to be the enzymatic complexes that comprise the electron transport chain, leading to a reduction in cellular ATP levels (Kruiderink et al, 1996; Nowak 2002). If the dose of cisplatin is high, ATP depletion is severe, and a rapid metabolic collapse and necrotic cell death would follow (Lieberthal et al, 1996). Lesser grades of ATP depletion associated with lower (pharmacologic) doses of cisplatin can induce apoptosis via release of mitochondrial cytochrome c, which has been documented in cultured renal epithelial cells exposed to cisplatin (Kruiderink et al, 1996; Park et al, 2002; Nowak 2002). It is currently not known exactly how cisplatin inhibits the enzymatic complexes of the respiratory chain. The mechanisms by which cisplatin causes release of cytochrome c also remain controversial, and include induction of mitochondrial permeability transition (Park et al, 2002) and increases in mitochondrial transmembrane potential (Nowak 2002). By analogy with studies of ATP depletion by alternative methods (exposure to inhibitors of oxidative phosphorylation such as antimycin A), it may be inferred that cisplatin-induced ATP depletion can also trigger death receptor-mediated apoptosis (Feldenberg et al, 1999) and the mitochondrial apoptotic pathways (Saikumar et al, 1998). Thus, partial ATP depletion may constitute a common biochemical pathway that leads to apoptosis following a variety of cellular stresses applied at an intensity below the threshold for necrosis.

If ATP depletion plays a central role in cisplatin-induced apoptosis, and the primary cause of cell death is not ROM formation, then how does one explain the encouraging results obtained from ROM inhibition as detailed above? One possibility is that these two pathogenetic processes co-exist, and are not mutually exclusive. Thus, while ATP-depletion may induce the primary cell injury and programmed cell death, this in turn may accelerate ROM formation by the damaged cells, which may contribute to an amplification loop leading to ROM-mediated cell death of the same cell or even the neighboring cells. ROM inhibition can limit this amplification loop, and may also alleviate nephrotoxicity by reducing the accompanying inflammatory response (Kruidering et al, 1997).

XIII: Summary and future perspectives

This review has focused on recent evidence for the paradoxical role of GGTP in cisplatin nephrotoxicity, the unexpected ability of proximal tubular cells to metabolize cisplatin to a nephrotoxin, and the evidence for apoptotic pathways and reactive oxygen metabolites as major mechanisms underlying cisplatin-induced renal cell injury. The information gleaned from this review may provide critical clues to novel therapeutic interventions aimed at minimizing cisplatin-induced nephrotoxicity while enhancing its antineoplastic efficacy. Such strategies may include inhibition of pathways leading to activation of cisplatin to a nephrotoxin, use of antioxidants to counter the ravages of reactive oxygen molecules, and targeted inhibition of apoptotic mechanisms activated by cisplatin specifically in kidney cells.


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Dr. Prasad Devarajan

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Dr. Marie H. Hanigan


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