New Therapeutic Concept of NAD Redox Balance for Cisplatin Nephrotoxicity

Oh, Gi-Su; Kim, Hyung-Jin; Shen, AiHua; Lee, Su-Bin; Yang, Sei-Hoon; Shim, Hyeok; Cho, Eun-Young; Kwon, Kang-Beom; Kwak, Tae Hwan; So, Hong-Seob

doi:https://doi.org/10.1155/2016/4048390

BioMed Research International

On this page

Abstract Introduction Conclusion References Copyright Related Articles

Special Issue

Novel Developments in Acute Kidney Injury

View this Special Issue

Review Article | Open Access

Volume 2016 | Article ID 4048390 | https://doi.org/10.1155/2016/4048390

New Therapeutic Concept of NAD Redox Balance for Cisplatin Nephrotoxicity

Gi-Su Oh,¹Hyung-Jin Kim,¹AiHua Shen,¹Su-Bin Lee,¹Sei-Hoon Yang,²Hyeok Shim,²Eun-Young Cho,²Kang-Beom Kwon,³Tae Hwan Kwak,⁴and Hong-Seob So¹

Academic Editor: Jeremiah R. Brown

Received22 Oct 2015

Accepted09 Dec 2015

Published05 Jan 2016

Abstract

Cisplatin is a widely used chemotherapeutic agent for the treatment of various tumors. In addition to its antitumor activity, cisplatin affects normal cells and may induce adverse effects such as ototoxicity, nephrotoxicity, and peripheral neuropathy. Various mechanisms such as DNA adduct formation, mitochondrial dysfunction, oxidative stress, and inflammatory responses are closely associated with cisplatin-induced nephrotoxicity; however, the precise mechanism remains unclear. The cofactor nicotinamide adenine dinucleotide (NAD⁺) has emerged as a key regulator of cellular energy metabolism and homeostasis. Recent studies have demonstrated associations between disturbance in intracellular NAD⁺ levels and clinical progression of various diseases through the production of reactive oxygen species and inflammation. Furthermore, we demonstrated that reduction of the intracellular NAD⁺/NADH ratio is critically involved in cisplatin-induced kidney damage through inflammation and oxidative stress and that increase of the cellular NAD⁺/NADH ratio suppresses cisplatin-induced kidney damage by modulation of potential damage mediators such as oxidative stress and inflammatory responses. In this review, we describe the role of NAD⁺ metabolism in cisplatin-induced nephrotoxicity and discuss a potential strategy for the prevention or treatment of cisplatin-induced adverse effects with a particular focus on NAD⁺-dependent cellular pathways.

1. Introduction

cis-Diamminedichloroplatinum II (CDDP, cisplatin) is a widely used chemotherapeutic drug for the treatment of various solid tumors in the head and neck, bladder, lung, ovaries, testicles, and uterus [1–6]. The various adverse effects of cisplatin during the course of chemotherapy include ototoxicity, nephrotoxicity, myelosuppression, and peripheral neuropathy. Cisplatin accumulates in renal tissues and cells, which are primary sites for drug filtration, concentration, and excretion. Even if blood concentrations are held at nontoxic levels during chemotherapy, concentrations may reach toxic levels in the kidneys [4]. In general, cisplatin concentrations in tubular epithelial cells of kidney tissues are five times higher than those in blood, and the elevated concentration of cisplatin therein causes nephrotoxicity. Clinical signs of kidney damage are a decrease in renal plasma flow and glomerular filtration rate, an increase of serum creatinine and blood urea nitrogen, and a reduction of serum magnesium and potassium levels [7]. Cisplatin-induced nephrotoxicity is dose-dependent and therefore limits the potential to increase dosage for optimal cancer therapy [8]. Even though the establishment of cisplatin-induced nephrotoxicity can be alleviated by diuretics and prehydration of patients, the prevalence of cisplatin nephrotoxicity is still high, occurring in approximately one-third of patients who have undergone cisplatin therapy [6]. Cisplatin nephrotoxicity can present in a number of ways, including acute kidney injury, hypomagnesemia, hypocalcemia, hyperuricemia, distal renal tubular acidosis, proximal tubular dysfunction, and chronic renal failure [7]. However, the most serious and one of the more common clinical features of cisplatin nephrotoxicity is acute kidney injury which occurs in 20–30% of patients. Recent studies have demonstrated that the cellular process of nephrotoxicity can be attributed to local accumulation of cisplatin inside the proximal tubule by membrane transportation, and intracellular conversion of the drug into toxic metabolites. Furthermore, various mechanisms such as DNA adduct formation, mitochondrial dysfunction, oxidative stress, inflammatory responses, and activation of apoptotic pathways are closely associated with cisplatin-induced nephrotoxicity [7].

The cofactor nicotinamide adenine dinucleotide (NAD⁺) has emerged as a key regulator of cellular energy metabolism and homeostasis. Recently, it has been reported that intracellular NAD⁺/NADH ratios are decreased in various pathological conditions such as diabetes [9], cisplatin-induced cochlear and kidney damage [10, 11], and in many tissues of aged animals and humans [12, 13]. Recent studies have demonstrated that a disturbance in intracellular NAD⁺ levels is linked to the progression of various diseases through the production of reactive oxygen species (ROS) and inflammation [10, 11, 14]. Furthermore, reduction of the intracellular NAD⁺/NADH ratio is critically involved in cisplatin-induced acute kidney damage; increasing the cellular NAD⁺/NADH ratio by pharmacological agents suppresses cisplatin-induced acute kidney damage by downregulation of potential damage mediators such as oxidative stress and inflammatory responses [11]. The decrease in the NAD⁺/NADH ratio has been attributed to hyperactivation of the NAD⁺-consuming poly(ADP-ribose) polymerase 1 (PARP-1) induced by oxidative damage due to altered redox mechanisms and consequent DNA damage [10, 11]. Since silent mating type information regulation 2 homolog 1 (sirtuin 1, SIRT1) deacetylase activity is influenced by the NAD⁺/NADH ratio [15], a significant reduction in the NAD⁺/NADH ratio causes a concomitant decrease in SIRT1 deacetylase activity, which is critically involved in diverse biological functions. In addition, the α-ketoglutarate dehydrogenase (α-KGDH) complex, an enzyme complex of the Krebs cycle in mitochondria, facilitates the generation of ROS after NAD⁺/NADH reduction [16]. Decreased NAD⁺/NADH also favors ROS generation in the respiratory chain complex I [17]. Therefore, maintenance of adequate NAD⁺ levels may be a critical factor for normal cellular function and could emerge as a useful strategy for treating many diseases.

Although there are review articles each focusing on cisplatin-mediated nephrotoxicity or beneficial role of NAD⁺, there is lack of effort illustrating a potential therapeutic or preventive strategy of modulatory NAD⁺ levels for treating cisplatin-associated nephrotoxicity. Therefore, we aimed to review a critical issue related to cisplatin-induced nephrotoxicity which can potentially be overcome by modulation of cellular NAD⁺ levels. We searched PubMed for published articles using separate search terms “cisplatin-mediated nephrotoxicity’’ and “NAD⁺-modulation and disease” and included only most recent and relevant publication including original research articles and reviews but excluded repetitive illustrations. In this review, we describe the mechanisms of cisplatin-mediated nephrotoxicity and the role of NAD⁺ metabolism therein and discuss a potential strategy for prevention of the adverse effects of cisplatin through targeting of NAD⁺-dependent cellular pathways.

2. Kidney-Specific Toxicity of Cisplatin: Cisplatin Transport and Biotransformation

Cisplatin is primarily cleared by the kidneys through both glomerular filtration and tubular secretion, whereas the biliary and intestinal excretions of this drug are negligible. During the excretion process, cisplatin is highly concentrated in the kidneys, which suggests an active accumulation of this drug by renal parenchymal cells, thereby explaining the particular damage caused by this drug to the kidneys compared to other organs. The toxic effects of cisplatin occur primarily in the renal proximal tubules, predominantly in the epithelial tubular cells of the S-3 segment [18]. Though the high concentrations of cisplatin in the kidneys favor its cellular uptake by passive diffusion, recent studies have demonstrated two different membrane transporters capable of facilitating the transport of cisplatin into cells. Copper transporter 1 (Ctr1) is highly expressed and localized in the basolateral membrane of the proximal tubule in the adult kidney [19]. Even though the role of Ctr1 in cisplatin nephrotoxicity in vivo has not been examined, cisplatin uptake and cytotoxicity were decreased by downregulation of Ctr1 expression in kidney cells in vitro, suggesting that Ctr1 is required for cisplatin uptake in these cells. In addition, the organic cation transporter 2 (OCT2) is specifically expressed in the basolateral membranes of the kidney renal proximal tubule cells, contributing to the etiology of the organ-specific toxicity of cisplatin. OCT2 expression is critical for the development of cisplatin-induced nephrotoxicity, and introduction of the OCT2 substrate cimetidine, a competing factor for transport, reduces nephrotoxicity [20], suggesting that OCT2 is critically involved in cisplatin uptake and its toxicity in these cells. In addition to specific expression of cisplatin transporter proteins in the kidney, many studies have demonstrated that cisplatin undergoes metabolic activation in the kidney to a more potent nephrotoxin. This process is initiated with the biotransformation of cisplatin-glutathione (GSH) conjugates by glutathione-S-transferase in the circulation [21]. When the cisplatin-GSH conjugate reaches and passes through the kidney, it is cleaved to a nephrotoxic metabolite primarily by the action of gamma-glutamyl transpeptidase, an enzyme principally located on the surface of the kidney proximal tubule cells. This metabolite is a highly reactive thiol/platinum compound that interacts with macromolecules, eventually leading to renal damage [22].

3. Mechanisms of Cisplatin Nephrotoxicity

3.1. Oxidative Stress in Cisplatin Nephrotoxicity

Oxidative stresses, including superoxide anions, hydrogen peroxide, and hydroxyl radicals, are unavoidable by-products of cellular respiration. Oxidative stress is also closely involved in renal injury after cisplatin administration. In particular, production of ROS and antioxidant system dysfunction are associated with cisplatin-induced nephrotoxicity [23]. Due to their unstable and highly reactive nature, ROS may attack and modify multiple target molecules such as lipids, proteins, and DNA, producing cellular stress. ROS also activate important signaling pathways, including an apoptotic pathway, which leads to cell death in the event of cisplatin-induced nephrotoxicity [24]. Although the role of oxidative stress in renal damage is well established, its source is poorly understood. Potential sources of ROS include the mitochondrial electron transport chain system [25], xanthine oxidase [26], cytochrome P450 enzymes [27], and NADPH oxidase [28]. Cisplatin may produce ROS in microsomes via the cytochrome P450 system (CYP). In vitro and in vivo tests have demonstrated that CYP is an important source of catalytic iron for the generation of ROS during cisplatin treatment. Furthermore, the cisplatin-induced increase of ROS and kidney damage were attenuated in CYP2E1^−/− mice [29]. Mitochondria have also been reported to be a major source of ROS. Disturbance of the mitochondrial electron transport chain system, which was accompanied by loss of mitochondrial membrane potential, an indicator of mitochondrial dysfunction, is a well-recognized mechanism responsible for the generation of ROS [25]. Interestingly, mitochondria themselves are particularly vulnerable to oxidative stress. Oxidative damage to mitochondria causes the impairment of mitochondrial function and subsequent cell death via apoptosis and necrosis [30]. Thus, ROS-mediated oxidative damage to mitochondria favors the generation of additional ROS, resulting in a vicious cycle. Many studies have demonstrated a clear association between mitochondrial ROS generation and cisplatin nephrotoxicity [6]. Membrane NAD(P)H oxidases (NOXs) are also one of the major sources for ROS generation. Especially in phagocytic cells such as neutrophils, superoxide is generated by NOX enzyme complexes [31]. However, many studies have recently found that superoxide-generating NOX expression is not restricted to phagocytic cells but is present in a wide variety of nonphagocytic cells and tissues [32]. In particular, it has been reported that superoxide-generating NOXs are expressed in the inner ear and in kidney tissues and that their expression is increased by exposure to cisplatin, thereby causing oxidative stress that leads to cisplatin-mediated ototoxicity and nephrotoxicity [11, 33, 34].

3.2. Inflammation in Cisplatin Nephrotoxicity

In addition to direct cellular toxicity, inflammation is closely associated with the pathogenesis of cisplatin nephrotoxicity. Over the last decade, it has been found that a number of mediators of inflammation, including TNF-α, IL-1β, TGF-β, RANTES, MIP2, and MCP1, are increased in cisplatin-induced renal injury. Inflammation contributes to the development of renal tissue damage and renal failure under pathological conditions. However, evidence for a functional role in renal damage for many of these cytokines still remains to be identified, with the exception of TNF-α [35, 36]. The proinflammatory cytokine TNF-α plays a central role in many infectious and inflammatory diseases. Relevantly, the functional involvement of TNF-α in the pathogenesis of cisplatin-induced acute renal failure was determined in mice treated with cisplatin in the presence or absence of TNF-α production inhibitors, as well as in TNF-α knockout mice. Treatment with TNF-α production inhibitors reduced cisplatin-induced renal damage and also reduced histologic evidence of injury. TNF-deficient mice were also resistant to cisplatin nephrotoxicity. These results indicated an important role for TNF-α in the pathogenesis of cisplatin nephrotoxicity [37]. Furthermore, this study showed that pharmacological inhibitors and antibodies against TNF-α markedly suppressed the induction of other cytokines during cisplatin nephrotoxicity, suggesting that TNF-α might be a key upstream regulator of the inflammatory response triggered by cisplatin. These observations have been confirmed and extended by other studies [38–40]. TNF-α can be produced by a variety of both immune and nonimmune cells. However, Zhang et al. were able to determine the source of the TNF-α that was responsible for cisplatin-induced renal damage [39]. They created chimeric mice in which TNF-α could be produced by resident kidney cells or by circulating immune cells and evaluated kidney function, histology, and cytokine expression in these chimeric mice following cisplatin administration. In this study, they demonstrated that the local production of TNF-α by resident kidney cells, probably the renal epithelial cells themselves, was crucial to cisplatin-induced nephrotoxicity [39].

The next question, then, became: how does TNF-α stimulate the inflammatory response and contribute to cisplatin nephrotoxicity? The biological activities of TNF-α are primarily mediated by two functionally distinct receptors, TNFR1 and TNFR2, to induce a variety of cellular responses ranging from inflammation to cell death. TNFR1 and TNFR2 are also upregulated by cisplatin. While TNFR1 directly induces the extrinsic apoptotic pathway, TNFR2 is primarily associated with the inflammatory response, which amplifies the TNFR1 effects. Furthermore, as the TNFR2 protein does not contain the death domain necessary to trigger apoptosis, TNFR2, unlike TNFR1, would appear to indirectly induce apoptosis and necrosis in renal tubular epithelial cells [41, 42]. Conflictingly, Tsuruya et al. showed that TNFR1-deficient mice and renal tubular cells were more resistant to cisplatin-induced renal injury and apoptosis compared with wild type mice [43], whereas Ramesh and Reeves recently showed that cisplatin-induced tubular cell death and renal injury were clearly attenuated in TNFR2-deficient, but not in TNFR1-deficient, mice [41]. Although the cause of the inconsistency between these two studies has not been elucidated, together they suggest that TNF-α signaling plays a critical role for cisplatin nephrotoxicity. The production of TNF-α after cisplatin administration is highly dependent upon ROS, NF-κB activation, and activation of p38 MAPK. In fact, TNF-α both is an inducer of ROS and is induced by ROS generated by cisplatin [37]. ROS activates the transcription factor NF-κB, which in turn induces the production of proinflammatory cytokines such as TNF-α [42]. NF-κB activation is pivotal in the expression of proinflammatory cytokines and other mediators involved in acute inflammatory responses and other conditions associated with increased ROS generation [44]. In addition to direct oxidative damage to lipids, DNA, and proteins [23], ROS generated by cisplatin activates p38 MAPK through the induction of p38 MAPK phosphorylation, which mediates the synthesis of TNF-α. Ramesh and Reeves demonstrated that inhibition of p38 MAPK reduced TNF-α production and protected against cisplatin-induced renal damage in vivo [45]. Activation of p38 MAPK led to the degradation of IκB (an inhibitor of NF-κB), thereby promoting translocation of NF-κB to the nucleus and the consequent stimulation of proinflammatory cytokine production, including TNF-α [46].

Toll-like receptors (TLRs) are a family of pattern recognition receptors that detect pathogenic elements such as viral RNA, bacterial DNA, lipopolysaccharides, or proteins, called pathogen-associated molecular patterns (PAMPs). TLRs play a pivotal role in host defense against infection by sensing the invasion of organisms and initiating both innate and adaptive immune responses [47]. TLRs also detect and respond to certain endogenous molecules such as high-mobility group box protein 1 (HMGB1), heat shock proteins (HSPs), and extracellular matrix components, termed damage-associated molecular pattern molecules (DAMPs). DAMPs are generally released by damaged or stressed tissues to “alert” the immune system to tissue injury or impending danger [48]. Cisplatin increases the expression of TLRs, including TLR4 in murine peritoneal macrophages in vitro, and subsequent stimulation by individual TLR-related ligands induces the production of proinflammatory cytokines such as TNF-α, IFN-γ, IL-1β, and IL-12 [49]. In addition, Zhang et al. have demonstrated that TLR4 is essential to the initiation of intrarenal inflammatory cytokine production associated with cisplatin-induced nephrotoxicity [50]. Ramesh et al. also have demonstrated that the combination of cisplatin and lipopolysaccharides, which are specific ligands for TLR4, acts synergistically to produce inflammatory cytokines such as TNF-α, IL-6, MCP-1, KC, and GM-CSF, thereby inducing nephrotoxicity in an acute renal failure model [51].

NF-κB activation is a critical bridge to the expression of inflammatory cytokines and other mediators involved in inflammatory responses through TLR signaling. After dimerization of TLR4 through engagement with its ligand, adapter molecules such as TIRAP and TRAM are recruited on the cytoplasmic domain of TLR4, which further interacts with MyD88 and TRIF, respectively, and then transduces a signal to the nucleus. MyD88 is critical for signaling by all TLRs except TLR3. After stimulation, MyD88 associates with the cytoplasmic portion of the TLR and recruits IL-1R-associated kinase- (IRAK-) 4 and -1 through a homophilic interaction of the death domains. Subsequently, TRAF6, TAK1, and NF-κB are activated, and then NF-κB is translocated into the nucleus where it regulates the genes for proinflammatory cytokines among others [52]. In contrast, in TLR4 knockout mice, the activation of p38, which is critical for cisplatin-induced TNF-α production, was significantly blunted [45]. Finally, the released nuclear protein HMGB1 has been shown to activate TLR4 in various pathologic conditions [48, 53]; however, the role of HMGB1 and other DAMPs in TLR4 activation associated with cisplatin nephrotoxicity remains to be elucidated.

3.3. Role of NAD Redox Balance in Cisplatin Nephrotoxicity

NAD is a metabolic cofactor that is present in cells either in its oxidized (NAD⁺) or in its reduced (NADH) form. NAD⁺ or NADH functions as a cofactor for a multitude of enzymatic reactions and therefore critically regulates cellular energy metabolism and homeostasis. As NAD⁺ is critical for a variety of enzymatic reactions, including glycolysis, the NAD redox balance, represented as the NAD⁺/NADH ratio, is tightly regulated [54], and its disruption has been associated with multiple clinical disorders and pathologies. For example, pellagra is caused by NAD⁺ deficiency subsequent to poor dietary intake of NAD⁺ biosynthesis precursors and can be easily cured by providing dietary nicotinic acid [55]. Pathological conditions such as diabetes and oxidative stress are also well correlated with decreased cellular NAD⁺ levels [9, 56]. It has also been recently reported that the cellular NAD⁺ level in many tissues declines with age [12, 13], implying the importance of maintaining optimal intracellular NAD⁺ levels to prevent age-associated cellular dysfunction. Furthermore, cisplatin-induced cochlear and kidney damage are highly associated with the decreases of NAD⁺/NADH ratios that accompany inflammation and oxidative stress [10, 11]. Cisplatin treatment resulted in a decrease of NAD⁺/NADH ratio in renal tissue without significant changes of NADH level [11]. This suggests that the decrease of NAD⁺/NADH ratio by cisplatin is mainly caused by reduction of NAD⁺ level. Of note, β-lapachone coadministration with cisplatin also restored NAD⁺/NADH ratio to control level through elevation of NAD⁺ level, but not by decrease of NADH level. Together, these findings suggest that maintenance of the NAD redox balance is very important for general health.

3.4. Role of NAD⁺-Dependent Enzymes in Cisplatin Nephrotoxicity

NAD⁺ acts as a cofactor for numerous enzymes including SIRTs, PARPs, and cyclic ADP- (cADP-) ribose synthases [57]; therefore, NAD⁺ might exert its biological effect through these enzymes. The mammalian sirtuin family consists of seven enzymes, SIRT1–7 [58], that are ubiquitously expressed yet show specific cellular localizations and functions. SIRT1, SIRT6, and SIRT7 are generally localized in the nuclei of cells, whereas SIRT3, SIRT4, and SIRT5 are localized in the mitochondria [59]. SIRT1 and SIRT5 act exclusively as deacetylases [60, 61], whereas SIRT2, SIRT3, SIRT4, and SIRT6 might also have a mono-ADP-ribosyl transferase activity [60, 62–64]. SIRT1 is the most widely studied sirtuin and has a Km for NAD⁺ that lies within the range of the physiological changes in intracellular NAD⁺ content. This suggests that sirtuin activity could be modulated by the physiological changes in intracellular NAD⁺ levels [64]. Considering that the intracellular NAD⁺/NADH ratios are decreased in various pathological conditions, including cisplatin-induced nephrotoxicity and ototoxicity [9–11, 56], SIRT1 activity might be reduced in the damaged tissues as well. In particular, Hasegawa et al. demonstrated that SIRT1 protects against oxidative stress-induced apoptosis in the kidney by inducing catalase, which catalyzes the decomposition of the ROS hydrogen peroxide, via deacetylation of FOXO3 in cultured proximal tubular cells [65]. Furthermore, Hasegawa et al. also reported that renal proximal tubular cell-specific SIRT1 transgenic mice showed resistance to cisplatin-induced renal tubular cell injuries such as apoptosis by maintaining peroxisome number and function, concomitant with upregulation of catalase and elimination of renal ROS [66]. In addition, it has been demonstrated that SIRT1 activation by resveratrol reduced cisplatin-induced proximal tubular cell apoptosis through deacetylation of p53 [67]. In contrast, Kim et al. and Oh et al. demonstrated that the reduction of intracellular NAD⁺/NADH ratio in cisplatin-injected kidney and cochlear tissues was critically associated with the decline of SIRT1 activity, which thereby caused cisplatin-induced nephrotoxicity and ototoxicity through inflammation and oxidative stress [10, 11]. However, SIRT1 activation through the increase of the cellular NAD⁺/NADH ratio suppressed the adverse effects of cisplatin by downregulation of potential damage mediators such as oxidative stress factors and inflammatory responses.

SIRT1 regulates diverse biological functions through direct interaction with and subsequent deacetylation of its targets, including p53 and NF-κB, which are closely related to its function in cisplatin-induced nephrotoxicity [7]. As described previously, the transcription factor NF-κB is one of the key regulators of inflammation. NF-κB activation is attained by either IκB phosphorylation and subsequent degradation or an IκB-independent pathway through posttranslational modifications of the NF-κB Rel proteins, including acetylation of the NF-κB p65 subunit. NF-κB p65 can be acetylated at five specific lysine residues (Lys-122, Lys-123, Lys-218, Lys-221, and Lys-310). In particular, acetylation of the Lys-310 residue is required for the transcriptional activity of NF-κB, whereas the other acetylation sites are involved in DNA binding [68]. SIRT1 physically interacts with the nuclear translocated NF-κB p65 subunit and deacetylates it at Lys-310, thereby inhibiting the transcriptional activity of NF-κB [69]. An assortment of recent evidence indicates that SIRT1 regulates inflammatory response through NF-κB p65 deacetylation. In cisplatin-induced nephrotoxicity and ototoxicity, Kim et al. demonstrated that SIRT1 activation was critically associated with the deacetylation status of the NF-κB p65 subunit [10, 11]. In addition, it has been demonstrated that SIRT1 knockdown leads to inflammatory pathway activation with increased inflammatory gene expression, whereas SIRT1 activation produces anti-inflammatory effects [70].

The tumor suppressor p53 is another crucial transcription factor in the cellular stress response [71]. A number of posttranslational modifications can occur in p53 that have critical effects on its stability and function, including phosphorylation, acetylation, sumoylation, neddylation, and methylation [72]. Cytosolic p53 is bound to Mdm2, a RING finger E3 ubiquitin ligase that facilitates protein degradation under normal conditions. Cellular stress, including DNA damage, hypoxia, or oxidative stress, induces rapid mitochondrial translocation of p53 and its posttranslational modification such as acetylation by p300/CBP or PCAF acetyltransferase [73]. The p53 is acetylated at lysine residues, including Lys-370, Lys-372, Lys-382, and Lys-386 in the carboxy-terminal region. Because acetylated p53 cannot bind to Mdm2, increased p53 acetylation levels strongly correlate with protein stabilization and activation in response to cellular stress [74]. Both nuclear SIRT1 and mitochondrial SIRT3 regulate p53 function through direct interaction and subsequent deacetylation of p53 [75]. In the nucleus, acetylation of p53 stimulates its sequence-specific DNA binding and subsequent recruitment of other transcription cofactors to promoter regions and thereby enhances transcription of target genes [76–78] such as the p53-upregulated modulator of apoptosis (PUMA), NADPH activator A (NOXA), and p53-induced gene 3 (PIG3), all of which are involved in the production of ROS through mitochondrial dysfunction or apoptosis. Deacetylation of p53 by nuclear-localized SIRT1 inactivates its sequence-specific transcriptional activity and represses p53-mediated cell growth arrest and apoptosis in response to DNA damage and oxidative stress [74]. Mitochondrial-localized SIRT3 deacetylates and activates several enzymes that are critical in maintaining cellular ROS levels and for apoptosis. Though it is not well understood whether acetylated p53 in mitochondria might have other functions, mitochondrial p53 interacts with anti- and proapoptotic Bcl-2 family members to either inhibit or activate them, thereby promoting apoptosis through robust mitochondrial outer membrane permeabilization and subsequent cytochrome c release [79, 80]. Deacetylation of p53 by mitochondrial-localized SIRT3 also represses p53-mediated cell growth arrest and apoptosis in response to DNA damage and oxidative stress [75]. On the other hand, Kim et al. demonstrated that cisplatin treatment led to substantial elevation of acetylated p53 levels in the kidney and cochlear tissues compared to those of untreated normal control mice [10, 11].

NAD⁺ is consumed not only by sirtuins, but also by PARPs [81]. Cisplatin accumulation in target tissues produces ROS that deplete the cellular antioxidant defense factors necessary to reduce oxidative stress and DNA damage. Cisplatin also directly binds to DNA, resulting in the disruption of the synthesis of key proteins and leading to cell injury and cell death. Furthermore, accumulation of DNA damage can lead to cell cycle arrest or genomic instability. The removal of DNA damaged by oxidative stress is mediated by single-strand DNA break repair, which is facilitated by PARPs. PARP-1 is the most critical protein-modifying nuclear enzyme involved in DNA repair. PARP-1 is a major NAD⁺ consumer, wherein the ADP-ribose moiety is transferred to PARP-1 itself or to other acceptor proteins in order to build the poly(ADP-ribose) polymer (PAR) [82]. PARP-1 is strongly activated by DNA damage and oxidative stress. Under physiological conditions, mild activation of PARP-1 can regulate several cellular processes, including DNA repair, cell cycle progression, cell survival, chromatin remodeling, and genomic stability [83]. However, hyperactivation of PARP-1 upon severe oxidative damage causes rapid depletion of intracellular NAD⁺ and ATP levels and eventually leads to cell death and related pathological conditions [84, 85]. Kim et al. have demonstrated that hyperactivation of PARP-1 in cisplatin-treated cochlea led to a decline in intracellular NAD⁺ levels and SIRT1 activity, thereby causing cochlear damage [10]. It is well established that PARP-1 and SIRT1 activity are interdependent as they compete for a limited pool of cellular NAD⁺. However, the Km of PARP-1 for NAD⁺ is two to ten times lower than that of SIRT1, which falls within the physiological range of cellular NAD⁺ concentrations [86]. Thus, PAPR-1 activation might critically influence SIRT1 activity by reducing NAD⁺ bioavailability. This model was further supported by recent studies wherein genetic depletion of PARP-1 or pharmacological inhibition of PARP-1 activity increased intracellular NAD⁺ level and subsequent SIRT1 activity [10].

4. Therapeutic Considerations of Cisplatin Nephrotoxicity

The main protective actions currently employed in clinical practice to reduce nephrotoxicity during cisplatin chemotherapy are based on avoiding the extreme exposure of the kidneys to the drug. This is managed primarily by hydration/diuretics, monitoring of renal function by serum creatinine clearance, and decreasing cisplatin doses upon manifestation of renal dysfunction [87, 88]. However, even with aggressive hydration, renal toxicity occurs. Therefore, more effective preventative strategies without attenuation of tumoricidal activity need to be developed, taking into consideration the mechanisms underlying the adverse effects of cisplatin exposure. Although the exact mechanism responsible for cisplatin-associated cellular damage is still to be elucidated, numerous studies have indicated that ROS and increased inflammation are important factors. The roles of these two factors seem to be closely related, and thus their abnormal regulation impacts overlapping cellular processes. Accordingly, pharmacological interventions that reduce systemic inflammation and/or oxidative stress might prevent or alleviate the development and progression of cisplatin-induced nephrotoxicity. However, these effects need to be explored in vivo [7]. A better option might be to focus on maintaining a proper level of intracellular NAD⁺. The decrease of cellular NAD⁺ level in the kidney and cochlear tissues after cisplatin exposure [10, 11] implies the therapeutic potential of intracellular NAD⁺ level modulation for cisplatin-associated adverse effects. The role of NAD⁺ in the prevention and cure of diseases was first recognized in the 1930s by Conrad Elvehjem, who demonstrated the therapeutic effect of the vitamin nicotinic acid on pellagra in dogs [55]. Since then, the therapeutic potential of NAD⁺ has been further evidenced by several studies. Araki et al. showed that addition of exogenous NAD⁺ to neurons delayed axonal degeneration in response to mechanical or chemical damage [89]. Ying et al. also demonstrated that intranasal administration of NAD⁺ profoundly decreased brain injury in a rat model of transient focal ischemia [90]. Pillai et al. showed that exogenous NAD⁺ blocked cardiac hypertrophic response [91].

As NAD⁺ regulates SIRTs that are involved in various cellular processes, the beneficial effects observed following enhanced SIRT activity might be attributed to increased intracellular NAD⁺ levels. Since both PARPs and SIRTs are NAD⁺-consuming enzymes and thus compete for NAD⁺, selective blockage of NAD⁺-consuming PARPs might also potentially be a good strategy to increase NAD⁺ levels. Consistent with this notion, targeted PARP inactivation has been shown to increase NAD⁺ levels and increase SIRT1 activity [92], suggesting that the modulation of PARP activity could be a therapeutic strategy for the treatment of cisplatin-associated adverse effects. In addition, approaches aimed at increasing NAD⁺ levels by supplementing NAD⁺ precursors through the activation of de novo and salvage pathways (Figure 1) for NAD⁺ biosynthesis have demonstrated cytoprotective effects against cellular damage. In fact, this specific strategy has been shown to increase NAD⁺ levels both in vitro and in vivo. For example, the administration of nicotinamide, a NAD⁺ precursor, showed a protective effect against oxidative stress and glucose deprivation in vitro and also alleviated tissue damage in animal models of ischemia [93, 94], spinal cord injury [95], and multiple sclerosis [96]. Similarly, nicotinic acid, another NAD⁺ precursor, has also been used to treat hyperlipidemia [97], indicating the therapeutic potential of NAD⁺ precursors. Although NAD⁺ treatment has not been tested extensively for its cytoprotective effects, a recent report suggested that it might reduce brain damage by protecting against PARP-1-induced cell death [98, 99].

Figure 1

Possible pathways for mammalian NAD⁺ biosynthesis. The biosynthesis of NAD⁺ through de novo, salvage, NQO1 pathways, and specific inhibition for PARPs. ATP: adenosine triphosphate, FAD: flavin adenine dinucleotide, IDO: indoleamine 2,3-dioxygenase, Na: nicotinic acid, NaAD: nicotinic acid adenine dinucleotide, NAD: nicotinamide adenine dinucleotide, NADS: NAD synthetase, Nam: nicotinamide, NaMN: nicotinic acid mononucleotide, NaPRT: nicotinic acid phosphoribosyl transferase, NMN: nicotinamide mononucleotide, NMNAT: nicotinamide mononucleotide adenylyltransferase, NQO1: NAD(P)H:quinone oxidoreductase 1, NR: nicotinamide riboside, NRK1,2: nicotinamide riboside kinase 1, 2, NamPRT: nicotinamide phosphoribosyltransferase, NMNAT: nicotinamide mononucleotide adenyltransferase, QA: quinolinic acid, QPRT: quinolinate phosphoribosyltransferase, TDO: tryptophan 2,3-dioxygenase, AFA: arylformamidase, KMO: kynurenine 3-monooxygenase, KYN: kynureninase, HAD: 3-hydroxy-anthranilate 3,4-dioxygenase, and PARPs: poly(ADP-ribose) polymerases.

As shown in Figure 1, another strategy in addition to de novo and salvage NAD⁺ biosynthesis pathways for regulating cellular NAD⁺ levels might be to utilize the cytosolic flavoprotein NADH : quinone oxidoreductase 1 (NQO1) that normally participates in reduction of quinone compounds in exchange for NADH oxidation [100, 101]. NQO1 catalyzes the reduction of quinones to hydroquinones by utilizing NADH as an electron donor, which consequently increases intracellular NAD⁺ levels. Therefore, it is plausible that endogenous factors or chemical agents that potentially activate NQO1 enzymatic activity or act as strong substrates of NQO1 might be beneficial for protection against cisplatin-induced toxicity by increasing intracellular NAD⁺ levels. In addition, there is evidence that NQO1 also plays a role in other biological activities, including anti-inflammatory processes, the scavenging of superoxide anion radicals, and the stabilization of p53 and other tumor suppressor proteins [102–106]. As shown in Figure 2, several substrates of NQO1 enzyme, including mitomycin C, RH1, AZQ, Coenzyme Q10, and idebenone, have been identified [107, 108], of which β-lapachone (3,4-dihydro-2,2-dimethyl-2H-naphtho[1,2-b]pyran-5,6-dione) is recently well studied as a strong substrate of NQO1 [109, 110]. β-Lapachone was first isolated from the bark of the Lapacho tree and was reported to inhibit tumor growth [111]. Several reports have indicated that pharmacological substrates of NQO1 ameliorate phenotypic manifestations associated with pathological conditions in rodent models. In particular, metabolic diseases such as obesity and spontaneous hypertension were shown to be reversed upon NQO1 enzymatic action using β-lapachone [112, 113], and pathological conditions such as arterial restenosis due to tissue injury and cisplatin-associated nephrotoxicity were also ameliorated by NQO1 enzymatic action using this substrate [11, 114]. As summarized in Figure 3, the increase of the cellular NAD⁺/NADH ratio by β-lapachone prevents cisplatin-induced kidney damage by modulation of potential damage mediators such as oxidative stress and inflammatory responses. Furthermore, β-lapachone did not interfere with the tumoricidal effect of cisplatin in vivo [11].

(a)

(b)

(c)

Figure 3

Role of NAD⁺ and NAD⁺-dependent enzymes in cisplatin-induced nephrotoxic mechanisms. Ctr1: copper transporter 1, OCT2: organic cation transporter 2, NOXs: NADPH oxidoreductases, Mt: mitochondria, ROS: reactive oxygen species, PFL: positive feedback loop, ER: endoplasmic reticulum, TLRs: toll-like receptors, HMGB1: high-mobility group box protein 1, TNF-α: tumor necrosis factor-α, NF-κB: nuclear factor-κB, NQO1: NAD(P)H:quinone oxidoreductase 1, and PARPs: poly(ADP-ribose) polymerases.

5. Conclusion

In conclusion, a NAD redox balance is critically important for sustaining a healthy condition, and maintenance of adequate NAD redox balance may show therapeutic benefits in various diseases through the regulation of NAD⁺-dependent enzymes and their downstream targets including SIRTs, PARPs, NF-κB, and p53. In this review we strongly suggest for the first time that direct modulation of a cellular NAD redox balance by pharmacological agents could be a promising therapeutic approach for the treatment of various diseases, including cisplatin nephrotoxicity.

Conflict of Interests

All authors declare no competing interests.

Acknowledgment

This work was supported by National Research Foundation of Korea (NRF) grants funded by the Korean government (MSIP): no. 2011-0028866 and no. 2011-0030130.

References

B. Rosenberg, L. Van Camp, and T. Krigas, “Inhibition of cell division in Escherichia coli by electrolysis products from a platinum electrode,” Nature, vol. 205, no. 4972, pp. 698–699, 1965.
View at: Publisher Site | Google Scholar
D. Wang and S. J. Lippard, “Cellular processing of platinum anticancer drugs,” Nature Reviews Drug Discovery, vol. 4, no. 4, pp. 307–320, 2005.
View at: Publisher Site | Google Scholar
S. M. Cohen and S. J. Lippard, “Cisplatin: from DNA damage to cancer chemotherapy,” Progress in Nucleic Acid Research and Molecular Biology, vol. 67, pp. 93–130, 2001.
View at: Publisher Site | Google Scholar
I. Arany and R. L. Safirstein, “Cisplatin nephrotoxicity,” Seminars in Nephrology, vol. 23, no. 5, pp. 460–464, 2003.
View at: Publisher Site | Google Scholar
Z. H. Siddik, “Cisplatin: mode of cytotoxic action and molecular basis of resistance,” Oncogene, vol. 22, no. 47, pp. 7265–7279, 2003.
View at: Publisher Site | Google Scholar
N. Pabla and Z. Dong, “Cisplatin nephrotoxicity: mechanisms and renoprotective strategies,” Kidney International, vol. 73, no. 9, pp. 994–1007, 2008.
View at: Publisher Site | Google Scholar
R. P. Miller, R. K. Tadagavadi, G. Ramesh, and W. B. Reeves, “Mechanisms of cisplatin nephrotoxicity,” Toxins, vol. 2, no. 11, pp. 2490–2518, 2010.
View at: Publisher Site | Google Scholar
M. H. Hanigan and P. Devarajan, “Cisplatin nephrotoxicity: molecular mechanisms,” Cancer Therapy, vol. 1, pp. 47–61, 2003.
View at: Google Scholar
Y. Ido, “Pyridine nucleotide redox abnormalities in diabetes,” Antioxidants & Redox Signaling, vol. 9, no. 7, pp. 931–942, 2007.
View at: Publisher Site | Google Scholar
H.-J. Kim, G.-S. Oh, A. Shen et al., “Augmentation of NAD⁺ by NQO1 attenuates cisplatin-mediated hearing impairment,” Cell Death and Disease, vol. 5, no. 6, Article ID e1292, 2014.
View at: Publisher Site | Google Scholar
G.-S. Oh, H.-J. Kim, J.-H. Choi et al., “Pharmacological activation of NQO1 increases NAD⁺ levels and attenuates cisplatin-mediated acute kidney injury in mice,” Kidney International, vol. 85, no. 3, pp. 547–560, 2014.
View at: Publisher Site | Google Scholar
N. Braidy, G. J. Guillemin, H. Mansour, T. Chan-Ling, A. Poljak, and R. Grant, “Age related changes in NAD+ metabolism oxidative stress and Sirt1 activity in wistar rats,” PLoS ONE, vol. 6, no. 4, Article ID e19194, 2011.
View at: Publisher Site | Google Scholar
H. Massudi, R. Grant, N. Braidy, J. Guest, B. Farnsworth, and G. J. Guillemin, “Age-associated changes in oxidative stress and NAD+ metabolism in human tissue,” PLoS ONE, vol. 7, no. 7, Article ID e42357, 2012.
View at: Publisher Site | Google Scholar
H.-J. Kim, G.-S. Oh, S.-K. Choe, T. H. Kwak, R. Park, and H.-S. So, “NAD⁺ metabolism in age-related hearing loss,” Aging and Disease, vol. 5, no. 2, pp. 150–159, 2014.
View at: Publisher Site | Google Scholar
D. Chen, J. Bruno, E. Easlon et al., “Tissue-specific regulation of SIRT1 by calorie restriction,” Genes and Development, vol. 22, no. 13, pp. 1753–1757, 2008.
View at: Publisher Site | Google Scholar
V. Adam-Vizi and L. Tretter, “The role of mitochondrial dehydrogenases in the generation of oxidative stress,” Neurochemistry International, vol. 62, no. 5, pp. 757–763, 2013.
View at: Publisher Site | Google Scholar
M. P. Murphy, “How mitochondria produce reactive oxygen species,” The Biochemical Journal, vol. 417, no. 1, pp. 1–13, 2009.
View at: Publisher Site | Google Scholar
M. Werner, M. J. Costa, L. G. Mitchell, and R. Nayar, “Nephrotoxicity of xenobiotics,” Clinica Chimica Acta, vol. 237, no. 1-2, pp. 107–154, 1995.
View at: Publisher Site | Google Scholar
N. Pabla, R. F. Murphy, K. Liu, and Z. Dong, “The copper transporter Ctr1 contributes to cisplatin uptake by renal tubular cells during cisplatin nephrotoxicity,” American Journal of Physiology—Renal Physiology, vol. 296, no. 3, pp. F505–F511, 2009.
View at: Publisher Site | Google Scholar
G. Ciarimboli, “Membrane transporters as mediators of cisplatin side-effects,” Anticancer Research, vol. 34, no. 1, pp. 547–550, 2014.
View at: Google Scholar
D. M. Townsend, K. D. Tew, L. He, J. B. King, and M. H. Hanigan, “Role of glutathione S-transferase Pi in cisplatin-induced nephrotoxicity,” Biomedicine and Pharmacotherapy, vol. 63, no. 2, pp. 79–85, 2009.
View at: Publisher Site | Google Scholar
R. D. Wainford, R. J. Weaver, K. N. Stewart, P. Brown, and G. M. Hawksworth, “Cisplatin nephrotoxicity is mediated by gamma glutamyltranspeptidase, not via a C-S lyase governed biotransformation pathway,” Toxicology, vol. 249, no. 2-3, pp. 184–193, 2008.
View at: Publisher Site | Google Scholar
N. A. G. Santos, C. S. Catão, N. M. Martins, C. Curti, M. L. P. Bianchi, and A. C. Santos, “Cisplatin-induced nephrotoxicity is associated with oxidative stress, redox state unbalance, impairment of energetic metabolism and apoptosis in rat kidney mitochondria,” Archives of Toxicology, vol. 81, no. 7, pp. 495–504, 2007.
View at: Publisher Site | Google Scholar
R. Çetin, E. Devrim, B. Kilicoglu, A. Avci, Ö. Çandir, and I. Durak, “Cisplatin impairs antioxidant system and causes oxidation in rat kidney tissues: possible protective roles of natural antioxidant foods,” Journal of Applied Toxicology, vol. 26, no. 1, pp. 42–46, 2006.
View at: Publisher Site | Google Scholar
H. H. Szeto, “Mitochondria-targeted peptide antioxidants: novel neuroprotective agents,” The AAPS Journal, vol. 8, no. 3, pp. E521–E531, 2006.
View at: Publisher Site | Google Scholar
C. E. Berry and J. M. Hare, “Xanthine oxidoreductase and cardiovascular disease: molecular mechanisms and pathophysiological implications,” Journal of Physiology, vol. 555, no. 3, pp. 589–606, 2004.
View at: Publisher Site | Google Scholar
R. A. Gottlieb, “Cytochrome P450: major player in reperfusion injury,” Archives of Biochemistry and Biophysics, vol. 420, no. 2, pp. 262–267, 2003.
View at: Publisher Site | Google Scholar
A. A. Miller, G. J. Dusting, C. L. Roulston, and C. G. Sobey, “NADPH-oxidase activity is elevated in penumbral and non-ischemic cerebral arteries following stroke,” Brain Research, vol. 1111, no. 1, pp. 111–116, 2006.
View at: Publisher Site | Google Scholar
H. Liu and R. Baliga, “Cytochrome P450 2E1 null mice provide novel protection against cisplatin-induced nephrotoxicity and apoptosis,” Kidney International, vol. 63, no. 5, pp. 1687–1696, 2003.
View at: Publisher Site | Google Scholar
J. Bai and A. I. Cederbaum, “Mitochondrial catalase and oxidative injury,” Biological Signals and Receptors, vol. 10, no. 3-4, pp. 189–199, 2001.
View at: Publisher Site | Google Scholar
M. Geiszt, “NADPH oxidases: new kids on the block,” Cardiovascular Research, vol. 71, no. 2, pp. 289–299, 2006.
View at: Publisher Site | Google Scholar
M. T. Quinn, M. C. B. Ammons, and F. R. DeLeo, “The expanding role of NADPH oxidases in health and disease: no longer just agents of death and destruction,” Clinical Science, vol. 111, no. 1, pp. 1–20, 2006.
View at: Publisher Site | Google Scholar
B. Bánfi, B. Malgrange, J. Knisz, K. Steger, M. Dubois-Dauphin, and K.-H. Krause, “NOX3, a superoxide-generating NADPH oxidase of the inner ear,” The Journal of Biological Chemistry, vol. 279, no. 44, pp. 46065–46072, 2004.
View at: Publisher Site | Google Scholar
H.-J. Kim, J.-H. Lee, S.-J. Kim et al., “Roles of NADPH oxidases in cisplatin-induced reactive oxygen species generation and ototoxicity,” Journal of Neuroscience, vol. 30, no. 11, pp. 3933–3946, 2010.
View at: Publisher Site | Google Scholar
G. Ramesh and W. B. Reeves, “Inflammatory cytokines in acute renal failure,” Kidney International Supplement, vol. 66, no. 91, pp. S56–S61, 2004.
View at: Google Scholar
C. Y. Lu, J. Hartono, M. Senitko, and J. Chen, “The inflammatory response to ischemic acute kidney injury: a result of the ‘right stuff’ in the ‘wrong place’?” Current Opinion in Nephrology and Hypertension, vol. 16, no. 2, pp. 83–89, 2007.
View at: Publisher Site | Google Scholar
G. Ramesh and W. Brian Reeves, “TNF-α mediates chemokine and cytokine expression and renal injury in cisplatin nephrotoxicity,” The Journal of Clinical Investigation, vol. 110, no. 6, pp. 835–842, 2002.
View at: Publisher Site | Google Scholar
G. Ramesh, B. Zhang, S. Uematsu, S. Akira, and W. B. Reeves, “Endotoxin and cisplatin synergistically induce renal dysfunction and cytokine production in mice,” American Journal of Physiology—Renal Physiology, vol. 293, no. 1, pp. F325–F332, 2007.
View at: Publisher Site | Google Scholar
B. Zhang, G. Ramesh, C. C. Norbury, and W. B. Reeves, “Cisplatin-induced nephrotoxicity is mediated by tumor necrosis factor-α produced by renal parenchymal cells,” Kidney International, vol. 72, no. 1, pp. 37–44, 2007.
View at: Publisher Site | Google Scholar
Z. Dong and S. S. Atherton, “Tumor necrosis factor-α in cisplatin nephrotoxicity: a homebred foe?” Kidney International, vol. 72, no. 1, pp. 5–7, 2007.
View at: Publisher Site | Google Scholar
G. Ramesh and W. B. Reeves, “TNFR2-mediated apoptosis and necrosis in cisplatin-induced acute renal failure,” American Journal of Physiology—Renal Physiology, vol. 285, no. 4, pp. F610–F618, 2003.
View at: Publisher Site | Google Scholar
P. D. Sánchez-González, F. J. López-Hernández, J. M. López-Novoa, and A. I. Morales, “An integrative view of the pathophysiological events leading to cisplatin nephrotoxicity,” Critical Reviews in Toxicology, vol. 41, no. 10, pp. 803–821, 2011.
View at: Publisher Site | Google Scholar
K. Tsuruya, T. Ninomiya, M. Tokumoto et al., “Direct involvement of the receptor-mediated apoptotic pathways in cisplatin-induced renal tubular cell death,” Kidney International, vol. 63, no. 1, pp. 72–82, 2003.
View at: Publisher Site | Google Scholar
E. Abraham, “Nuclear factor-κB and its role in sepsis-associated organ failure,” Journal of Infectious Diseases, vol. 187, supplement 2, pp. S364–S369, 2003.
View at: Publisher Site | Google Scholar
G. Ramesh and W. B. Reeves, “p38 MAP kinase inhibition ameliorates cisplatin nephrotoxicity in mice,” American Journal of Physiology—Renal Physiology, vol. 289, no. 1, pp. F166–F174, 2005.
View at: Publisher Site | Google Scholar
T. Yamakawa, S. Eguchi, T. Matsumoto et al., “Intracellular signaling in rat cultured vascular smooth muscle cells: roles of nuclear factor-κB and p38 mitogen-activated protein kinase on tumor necrosis factor-α production,” Endocrinology, vol. 140, no. 8, pp. 3562–3572, 1999.
View at: Google Scholar
R. Medzhitov and C. A. Janeway Jr., “Decoding the patterns of self and nonself by the innate immune system,” Science, vol. 296, no. 5566, pp. 298–300, 2002.
View at: Publisher Site | Google Scholar
H. Kono and K. L. Rock, “How dying cells alert the immune system to danger,” Nature Reviews Immunology, vol. 8, no. 4, pp. 279–289, 2008.
View at: Publisher Site | Google Scholar
D. U. Ferreiro and E. A. Komives, “Molecular mechanisms of system control of NF-κB signaling by IκBα,” Biochemistry, vol. 49, no. 8, pp. 1560–1567, 2010.
View at: Publisher Site | Google Scholar
B. Zhang, G. Ramesh, S. Uematsu, S. Akira, and W. B. Reeves, “TLR4 signaling mediates inflammation and tissue injury in nephrotoxicity,” Journal of the American Society of Nephrology, vol. 19, no. 5, pp. 923–932, 2008.
View at: Publisher Site | Google Scholar
G. Ramesh, S. R. Kimball, L. S. Jefferson, and W. B. Reeves, “Endotoxin and cisplatin synergistically stimulate TNF-α production by renal epithelial cells,” American Journal of Physiology—Renal Physiology, vol. 292, no. 2, pp. F812–F819, 2007.
View at: Publisher Site | Google Scholar
A. F. McGettrick and L. A. J. O'Neill, “The expanding family of MyD88-like adaptors in Toll-like receptor signal transduction,” Molecular Immunology, vol. 41, no. 6-7, pp. 577–582, 2004.
View at: Publisher Site | Google Scholar
A. Tsung, J. R. Klune, X. Zhang et al., “HMGB1 release induced by liver ischemia involves Toll-like receptor 4–dependent reactive oxygen species production and calcium-mediated signaling,” Journal of Experimental Medicine, vol. 204, no. 12, pp. 2913–2923, 2007.
View at: Publisher Site | Google Scholar
R. H. Houtkooper, C. Cantó, R. J. Wanders, and J. Auwerx, “The secret life of NAD⁺: an old metabolite controlling new metabolic signaling pathways,” Endocrine Reviews, vol. 31, no. 2, pp. 194–223, 2010.
View at: Publisher Site | Google Scholar
C. A. Elvehjem, R. J. Madden, F. M. Strong, and D. W. Wolley, “The isolation and identification of the anti-black tongue factor. 1937,” The Journal of Biological Chemistry, vol. 277, no. 34, article e22, 2002.
View at: Google Scholar
W. Ying, “NAD+/NADH and NADP+/NADPH in cellular functions and cell death: regulation and biological consequences,” Antioxidants & Redox Signaling, vol. 10, no. 2, pp. 179–206, 2008.
View at: Publisher Site | Google Scholar
P. Belenky, K. L. Bogan, and C. Brenner, “NAD⁺ metabolism in health and disease,” Trends in Biochemical Sciences, vol. 32, no. 1, pp. 12–19, 2007.
View at: Publisher Site | Google Scholar
N. Dali-Youcef, M. Lagouge, S. Froelich, C. Koehl, K. Schoonjans, and J. Auwerx, “Sirtuins: the ‘magnificent seven’, function, metabolism and longevity,” Annals of Medicine, vol. 39, no. 5, pp. 335–345, 2007.
View at: Publisher Site | Google Scholar
E. Michishita, J. Y. Park, J. M. Burneskis, J. C. Barrett, and I. Horikawa, “Evolutionarily conserved and nonconserved cellular localizations and functions of human SIRT proteins,” Molecular Biology of the Cell, vol. 16, no. 10, pp. 4623–4635, 2005.
View at: Publisher Site | Google Scholar
B. J. North, B. L. Marshall, M. T. Borra, J. M. Denu, and E. Verdin, “The human Sir2 ortholog, SIRT2, is an NAD⁺-dependent tubulin deacetylase,” Molecular Cell, vol. 11, no. 2, pp. 437–444, 2003.
View at: Publisher Site | Google Scholar
H. Vaziri, S. K. Dessain, E. Ng Eaton et al., “hSIR2(SIRT1) functions as an NAD-dependent p53 deacetylase,” Cell, vol. 107, pp. 149–159, 2001.
View at: Google Scholar
M. C. Haigis, R. Mostoslavsky, K. M. Haigis et al., “SIRT4 inhibits glutamate dehydrogenase and opposes the effects of calorie restriction in pancreatic β cells,” Cell, vol. 126, no. 5, pp. 941–954, 2006.
View at: Publisher Site | Google Scholar
G. Liszt, E. Ford, M. Kurtev, and L. Guarente, “Mouse Sir2 homolog SIRT6 is a nuclear ADP-ribosyltransferase,” The Journal of Biological Chemistry, vol. 280, no. 22, pp. 21313–21320, 2005.
View at: Publisher Site | Google Scholar
S. Imai, F. B. Johnson, R. A. Marciniak, M. McVey, P. U. Park, and L. Guarante, “Sir2: an NAD-dependent histone deacetylase that connects chromatin silencing, metabolism, and aging,” Cold Spring Harbor Symposia on Quantitative Biology, vol. 65, pp. 297–302, 2000.
View at: Publisher Site | Google Scholar
K. Hasegawa, S. Wakino, K. Yoshioka et al., “Sirt1 protects against oxidative stress-induced renal tubular cell apoptosis by the bidirectional regulation of catalase expression,” Biochemical and Biophysical Research Communications, vol. 372, no. 1, pp. 51–56, 2008.
View at: Publisher Site | Google Scholar
K. Hasegawa, S. Wakino, K. Yoshioka et al., “Kidney-specific overexpression of Sirt1 protects against acute kidney injury by retaining peroxisome function,” The Journal of Biological Chemistry, vol. 285, no. 17, pp. 13045–13056, 2010.
View at: Publisher Site | Google Scholar
D. H. Kim, Y. J. Jung, J. E. Lee et al., “Sirt1 activation by resveratrol ameliorates cisplatin-induced renal injury through deacetylation of p53,” American Journal of Physiology—Renal Physiology, vol. 301, no. 2, pp. F427–F435, 2011.
View at: Publisher Site | Google Scholar
M. Neumann and M. Naumann, “Beyond IκBs: alternative regulation of NF-κB activity,” The FASEB Journal, vol. 21, no. 11, pp. 2642–2654, 2007.
View at: Publisher Site | Google Scholar
F. Yeung, J. E. Hoberg, C. S. Ramsey et al., “Modulation of NF-κB-dependent transcription and cell survival by the SIRT1 deacetylase,” The EMBO Journal, vol. 23, no. 12, pp. 2369–2380, 2004.
View at: Publisher Site | Google Scholar
T. Yoshizaki, J. C. Milne, T. Imamura et al., “SIRT1 exerts anti-inflammatory effects and improves insulin sensitivity in adipocytes,” Molecular and Cellular Biology, vol. 29, no. 5, pp. 1363–1374, 2009.
View at: Publisher Site | Google Scholar
A. J. Levine and M. Oren, “The first 30 years of p53: growing ever more complex,” Nature Reviews Cancer, vol. 9, no. 10, pp. 749–758, 2009.
View at: Publisher Site | Google Scholar
J.-P. Kruse and W. Gu, “Modes of p53 regulation,” Cell, vol. 137, no. 4, pp. 609–622, 2009.
View at: Publisher Site | Google Scholar
Y. Tang, W. Zhao, Y. Chen, Y. Zhao, and W. Gu, “Acetylation is indispensable for p53 activation,” Cell, vol. 133, no. 4, pp. 612–626, 2008.
View at: Publisher Site | Google Scholar
C. L. Brooks and W. Gu, “The impact of acetylation and deacetylation on the p53 pathway,” Protein and Cell, vol. 2, no. 6, pp. 456–462, 2011.
View at: Publisher Site | Google Scholar
S. Li, M. Banck, S. Mujtaba, M.-M. Zhou, M. M. Sugrue, and M. J. Walsh, “p53-Induced growth arrest is regulated by the mitochondrial SirT3 deacetylase,” PLoS ONE, vol. 5, no. 5, Article ID e10486, 2010.
View at: Publisher Site | Google Scholar
W. Gu and R. G. Roeder, “Activation of p53 sequence-specific DNA binding by acetylation of the p53 C-terminal domain,” Cell, vol. 90, no. 4, pp. 595–606, 1997.
View at: Publisher Site | Google Scholar
Y. Zhao, S. Lu, L. Wu et al., “Acetylation of p53 at lysine 373/382 by the histone deacetylase inhibitor depsipeptide induces expression of ${p 21}^{Waf 1 /Cip 1}$ ,” Molecular and Cellular Biology, vol. 26, no. 7, pp. 2782–2790, 2006.
View at: Publisher Site | Google Scholar
A. Kotsinas, V. Aggarwal, E.-J. Tan, B. Levy, and V. G. Gorgoulis, “PIG3: a novel link between oxidative stress and DNA damage response in cancer,” Cancer Letters, vol. 327, no. 1-2, pp. 97–102, 2012.
View at: Publisher Site | Google Scholar
J. D. Amaral, J. M. Xavier, C. J. Steer, and C. M. P. Rodrigues, “Targeting the p53 pathway of apoptosis,” Current Pharmaceutical Design, vol. 16, no. 22, pp. 2493–2503, 2010.
View at: Publisher Site | Google Scholar
A. V. Vaseva and U. M. Moll, “The mitochondrial p53 pathway,” Biochimica et Biophysica Acta (BBA)—Bioenergetics, vol. 1787, no. 5, pp. 414–420, 2009.
View at: Publisher Site | Google Scholar
L. Mouchiroud, R. H. Houtkooper, and J. Auwerx, “NAD⁺ metabolism: a therapeutic target for age-related metabolic disease,” Critical Reviews in Biochemistry and Molecular Biology, vol. 48, no. 4, pp. 397–408, 2013.
View at: Publisher Site | Google Scholar
R. Krishnakumar and W. L. Kraus, “The PARP side of the nucleus: molecular actions, physiological outcomes, and clinical targets,” Molecular Cell, vol. 39, no. 1, pp. 8–24, 2010.
View at: Publisher Site | Google Scholar
C. Thomas and A. V. Tulin, “Poly-ADP-ribose polymerase: machinery for nuclear processes,” Molecular Aspects of Medicine, vol. 34, no. 6, pp. 1124–1137, 2013.
View at: Publisher Site | Google Scholar
P. M. Goodwin, P. J. Lewis, M. I. Davies, C. J. Skidmore, and S. Shall, “The effect of gamma radiation and neocarzinostatin of NAD and ATP levels in mouse leukaemia cells,” Biochimica et Biophysica Acta (BBA)—General Subjects, vol. 543, no. 4, pp. 576–582, 1978.
View at: Publisher Site | Google Scholar
C. J. Skidmore, M. I. Davies, P. M. Goodwin et al., “The involvement of poly(ADP-ribose) polymerase in the degradation of NAD caused by gamma-radiation and N-methyl-N-nitrosourea,” European Journal of Biochemistry, vol. 101, no. 1, pp. 135–142, 1979.
View at: Google Scholar
C. Cantó and J. Auwerx, “NAD⁺ as a signaling molecule modulating metabolism,” Cold Spring Harbor Symposia on Quantitative Biology, vol. 76, pp. 291–298, 2011.
View at: Publisher Site | Google Scholar
V. Launay-Vacher, J.-B. Rey, C. Isnard-Bagnis, G. Deray, and M. Daouphars, “Prevention of cisplatin nephrotoxicity: state of the art and recommendations from the European Society of Clinical Pharmacy Special Interest Group on Cancer Care,” Cancer Chemotherapy and Pharmacology, vol. 61, no. 6, pp. 903–909, 2008.
View at: Publisher Site | Google Scholar
G. Losonczy, C. Máthé, V. Müller, K. Szondy, and J. Moldvay, “Incidence, risk factors and prevention of cisplatin-induced nephrotoxicity in patients with lung cancer,” Magyar Onkologia, vol. 54, no. 4, pp. 289–296, 2010.
View at: Publisher Site | Google Scholar
T. Araki, Y. Sasaki, and J. Milbrandt, “Increased nuclear NAD biosynthesis and SIRT1 activation prevent axonal degeneration,” Science, vol. 305, no. 5686, pp. 1010–1013, 2004.
View at: Publisher Site | Google Scholar
W. Ying, G. Wei, D. Wang et al., “Intranasal administration with NAD⁺ profoundly decreases brain injury in a rat model of transient focal ischemia,” Frontiers in Bioscience, vol. 12, no. 7, pp. 2728–2734, 2007.
View at: Google Scholar
V. B. Pillai, N. R. Sundaresan, G. Kim et al., “Exogenous NAD blocks cardiac hypertrophic response via activation of the SIRT3-LKB1-AMP-activated kinase pathway,” The Journal of Biological Chemistry, vol. 285, no. 5, pp. 3133–3144, 2010.
View at: Publisher Site | Google Scholar
P. Bai, C. Cantó, H. Oudart et al., “PARP-1 inhibition increases mitochondrial metabolism through SIRT1 activation,” Cell Metabolism, vol. 13, no. 4, pp. 461–468, 2011.
View at: Publisher Site | Google Scholar
I. A. Ayoub, E. Jian Lee, C. S. Ogilvy, M. F. Beal, and K. I. Maynard, “Nicotinamide reduces infarction up to two hours after the onset of permanent focal cerebral ischemia in Wistar rats,” Neuroscience Letters, vol. 259, no. 1, pp. 21–24, 1999.
View at: Publisher Site | Google Scholar
I. A. Ayoub and K. I. Maynard, “Therapeutic window for nicotinamide following transient focal cerebral ischemia,” NeuroReport, vol. 13, no. 2, pp. 213–216, 2002.
View at: Publisher Site | Google Scholar
K. L. Brewer and J. S. Hardin, “Neuroprotective effects of nicotinamide after experimental spinal cord injury,” Academic Emergency Medicine, vol. 11, no. 2, pp. 125–130, 2004.
View at: Publisher Site | Google Scholar
S. Kaneko, J. Wang, M. Kaneko et al., “Protecting axonal degeneration by increasing nicotinamide adenine dinucleotide levels in experimental autoimmune encephalomyelitis models,” Journal of Neuroscience, vol. 26, no. 38, pp. 9794–9804, 2006.
View at: Publisher Site | Google Scholar
A. A. Sauve, “NAD⁺ and vitamin B₃: from metabolism to therapies,” The Journal of Pharmacology and Experimental Therapeutics, vol. 324, no. 3, pp. 883–893, 2008.
View at: Publisher Site | Google Scholar
W. Ying, P. Garnier, and R. A. Swanson, “NAD⁺ repletion prevents PARP-1-induced glycolytic blockade and cell death in cultured mouse astrocytes,” Biochemical and Biophysical Research Communications, vol. 308, no. 4, pp. 809–813, 2003.
View at: Publisher Site | Google Scholar
W. Ying, C. C. Alano, P. Garnier, and R. A. Swanson, “NAD+ as a metabolic link between DNA damage and cell death,” Journal of Neuroscience Research, vol. 79, no. 1-2, pp. 216–223, 2005.
View at: Publisher Site | Google Scholar
A. Gaikwad, D. J. Long II, J. L. Stringer, and A. K. Jaiswal, “In vivo role of NAD(P)H:Quinone Oxidoreductase 1 (NQO1) in the regulation of intracellular redox state and accumulation of abdominal adipose tissue,” The Journal of Biological Chemistry, vol. 276, pp. 22559–22564, 2001.
View at: Publisher Site | Google Scholar
D. Ross, J. K. Kepa, S. L. Winski, H. D. Beall, A. Anwar, and D. Siegel, “NAD(P)H:quinone oxidoreductase 1 (NQO1): chemoprotection, bioactivation, gene regulation and genetic polymorphisms,” Chemico-Biological Interactions, vol. 129, no. 1-2, pp. 77–97, 2000.
View at: Publisher Site | Google Scholar
M. Gao, A. Singh, K. Macri et al., “Antioxidant components of naturally-occurring oils exhibit marked anti-inflammatory activity in epithelial cells of the human upper respiratory system,” Respiratory Research, vol. 12, article 92, 2011.
View at: Publisher Site | Google Scholar
D. K. Gessner, R. Ringseis, M. Siebers et al., “Inhibition of the pro-inflammatory NF-κB pathway by a grape seed and grape marc meal extract in intestinal epithelial cells,” Journal of Animal Physiology and Animal Nutrition, vol. 96, no. 6, pp. 1074–1083, 2012.
View at: Publisher Site | Google Scholar
R. Pazdro and J. R. Burgess, “The antioxidant 3H-1,2-dithiole-3-thione potentiates advanced glycation end-product-induced oxidative stress in SH-SY5Y cells,” Experimental Diabetes Research, vol. 2012, Article ID 137607, 8 pages, 2012.
View at: Publisher Site | Google Scholar
C. I. Jones III, H. Zhu, S. F. Martin, Z. Han, Y. Li, and B. R. Alevriadou, “Regulation of antioxidants and phase 2 enzymes by shear-induced reactive oxygen species in endothelial cells,” Annals of Biomedical Engineering, vol. 35, no. 5, pp. 683–693, 2007.
View at: Publisher Site | Google Scholar
O. Moscovitz, P. Tsvetkov, N. Hazan et al., “A mutually inhibitory feedback loop between the 20S proteasome and its regulator, NQO1,” Molecular Cell, vol. 47, no. 1, pp. 76–86, 2012.
View at: Publisher Site | Google Scholar
R. H. Haefeli, M. Erb, A. C. Gemperli et al., “NQo1-dependent redox cycling of idebenone: effects on cellular redox potential and energy levels,” PLoS ONE, vol. 6, no. 3, Article ID e17963, 2011.
View at: Publisher Site | Google Scholar
X. Huang, Y. Dong, E. A. Bey et al., “An NQO1 substrate with potent antitumor activity that selectively kills by PARP1-induced programmed necrosis,” Cancer Research, vol. 72, no. 12, pp. 3038–3047, 2012.
View at: Publisher Site | Google Scholar
A. B. Pardee, Y. Z. Li, and C. J. Li, “Cancer therapy with β-lapachone,” Current Cancer Drug Targets, vol. 2, no. 3, pp. 227–242, 2002.
View at: Publisher Site | Google Scholar
L. S. Li, E. A. Bey, Y. Dong et al., “Modulating endogenous NQO1 levels identifies key regulatory mechanisms of action of β-lapachone for pancreatic cancer therapy,” Clinical Cancer Research, vol. 17, no. 2, pp. 275–285, 2011.
View at: Publisher Site | Google Scholar
S. M. Planchon, S. Wuerzberger, B. Frydman et al., “β-Lapachone-mediated apoptosis in human promyelocytic leukemia (HL-60) and human prostate cancer cells: a p53-independent response,” Cancer Research, vol. 55, no. 17, pp. 3706–3711, 1995.
View at: Google Scholar
J. H. Hwang, D. W. Kim, E. J. Jo et al., “Pharmacological stimulation of NADH oxidation ameliorates obesity and related phenotypes in mice,” Diabetes, vol. 58, no. 4, pp. 965–974, 2009.
View at: Publisher Site | Google Scholar
Y.-H. Kim, J. H. Hwang, J.-R. Noh et al., “Activation of NAD(P)H:quinone oxidoreductase ameliorates spontaneous hypertension in an animal model via modulation of eNOS activity,” Cardiovascular Research, vol. 91, no. 3, pp. 519–527, 2011.
View at: Publisher Site | Google Scholar
S.-Y. Kim, N. H. Jeoung, C. J. Oh et al., “Activation of NAD(P)H:quinone oxidoreductase 1 prevents arterial restenosis by suppressing vascular smooth muscle cell proliferation,” Circulation Research, vol. 104, no. 7, pp. 842–850, 2009.
View at: Publisher Site | Google Scholar

Copyright

Copyright © 2016 Gi-Su Oh et al. This is an open access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

PDF Download Citation

Download other formats

Order printed copies

Views

2478

Downloads

1366

Citations

BioMed Research International

Novel Developments in Acute Kidney Injury

New Therapeutic Concept of NAD Redox Balance for Cisplatin Nephrotoxicity

Abstract

1. Introduction

2. Kidney-Specific Toxicity of Cisplatin: Cisplatin Transport and Biotransformation

3. Mechanisms of Cisplatin Nephrotoxicity

3.1. Oxidative Stress in Cisplatin Nephrotoxicity

3.2. Inflammation in Cisplatin Nephrotoxicity

3.3. Role of NAD Redox Balance in Cisplatin Nephrotoxicity

3.4. Role of NAD+-Dependent Enzymes in Cisplatin Nephrotoxicity

4. Therapeutic Considerations of Cisplatin Nephrotoxicity

5. Conclusion

Conflict of Interests

Acknowledgment

References

Copyright

3.4. Role of NAD⁺-Dependent Enzymes in Cisplatin Nephrotoxicity