Cisplatin-Mediated Upregulation of APE2 Binding to MYH9 Provokes Mitochondrial Fragmentation and Acute Kidney Injury

Cisplatin (75 mg/m²) is standard chemotherapy in the treatment of various solid tumors. However, at this dose, significant acute kidney injury (AKI) occurs, often necessitating dose reduction or withdrawal (1). Because intracellular chloride concentration is typically low, the chlorine atoms in cisplatin are quickly replaced by water molecules, and the resulting product is actually the species that reacts with glutathione in the cytoplasm and with DNA in the nucleus (2). In rapidly dividing tumor cells, cisplatin-DNA intrastrand and interstrands crosslinking arrests DNA replication, cell proliferation, resulting in further cytotoxicity (3). Within 24 hours of administration, 50%–80% of cisplatin is excreted via the urine, and the platinum concentration in the renal cortex is several-fold higher than in other organs (4). Cisplatin predominantly damages the proximal tubule cells (PTC), decreasing the glomerular filtration rate (5). Even though nonproliferating PTCs are generally less sensitive to other DNA-damaging agents, cisplatin selectively injures PTCs, as manifested by both necrosis and apoptosis (6). Second only to the heart, kidneys have the greatest mitochondrial content and oxygen consumption rate (OCR). Moreover, kidney PTCs contain more mitochondria than any other cellular organelle because this organ requires sufficient energy to reabsorb 80% of the filtrate that passes through the glomeruli, including toxins such as cisplatin (7). Thus, PTC vulnerability to cisplatin may reflect the kidney's ability to accumulate and retain platinum to a greater degree than other organs.

DNA damage repair mechanisms are abundant in the nucleus but sparse in mitochondria (8). Intrastrand and interstand adducts in nuclear DNA and mitochondrial DNA (mtDNA) are the most prevalent modification after cisplatin treatment. DNA adducts are disastrous to DNA replication and transcription because they prevent the DNA strand separation required for polymerase function. In the nucleus, several mechanisms exist that can repair the full spectrum of Pt-DNA adducts. Mitochondria, however, lack the full complement of these repair pathways, base excision repair (BER) being the only Pt-DNA damage repair pathway available (9), though they are able to make use of nuclear proteins that translocate into mitochondria, such as UNG, POLG, APE2, and LIG3 (10). Interestingly, humans possess two AP endonucleases, APE1 and APE2, which cleave the phosphodiester backbone 5′ to the AP site to initiate BER. APE1 retains the strong APE action and weak 3′-phosphodiesterase and 3′-5′ exonuclease activities of nuclear DNA damage and is the major AP endonuclease in human cells with a nuclear localization signal. Compared with APE1, APE2 has relatively weaker AP endonuclease activity, while retaining the robust proliferating-cell nuclear antigen (PCNA)-stimulated 3′-5′ exonuclease and 3′-phosphodiesterase activities associated with DNA damage. Murine mitochondria contain APE2 (11), and APE2-null mice exhibit growth retardation and dyslymphopoiesis accompanied by G₂–M arrest, suggesting that APE2 is important for proper cell-cycle progression (12). APE2 is also critical in normal B-cell development, including recovery from chemotherapy-related DNA damage (13), and it was recently suggested (14) that high expression of APE2 but not APE1 in germinal centers promotes error-prone repair and mutations during somatic hypermutation (14). Importantly, APE2 is a key player in PCNA-dependent repair of oxidative DNA damage (15). Although enzymic activities and other molecular events associated with DNA repair and oxidative stress have been characterized in many laboratory models, a detailed mechanistic understanding of conserved APE2 functions in AKI is lacking.

In the present work, we identified a novel function for APE2, namely that APE2 mediates cisplatin-induced AKI (C-AKI) even in the absence of cisplatin-induced Pt-DNA adducts in kidney PTCs. Our studies have revealed that cisplatin increases APE2 protein responses to DNA damage in PTCs. APE2 transgenic mice phenotypically recapitulated the pathophysiological features of C-AKI without cisplatin treatment. APE2 binds myosin heavy-chain 9 (MYH9) protein in mitochondria after cisplatin treatment. Human MYH9-related disorder is caused by MYH9 mutations that ultimately lead to nephritis, macrothrombocytopenia, and deafness. This constellation of symptoms even without cisplatin treatment is similar to the toxicity profile of cisplatin. MYH9 knockout mice (16) and mice with MYH9 mutations (mimic homologous to human R702C, D1424N, E1841K; ref. 17) developed AKI that mimicked the cisplatin toxicity phenotype of C-AKI mice. Moreover, C-AKI is known to be attenuated in APE2 knockout mice (12). Taken together, therefore, these findings suggest that cisplatin promotes AKI development by upregulating APE2, which leads to the subsequent MYH9 dysfunction in PTC mitochondria. It is important to note (i) that this theory differs profoundly from the generally held previous view that C-AKI is due to direct cisplatin-induced nuclear DNA damage, and (ii) that it offers a new and novel opportunity for prophylaxis of this serious disease.

Ten-week-old wild-type C57BL/6J mice weighing 25–28 g were purchased from the Jackson Laboratory. Mice were treated with a single intraperitoneal dose of cisplatin (15 mg/kg body weight, 1 mg/mL in 0.9% NaCl), carboplatin (40 mg/kg body weight, 2 mg/mL in 0.9% NaCl) or oxaliplatin (40 mg/kg body weight, 2 mg/mL in 0.9% NaCl). An identical volume of saline (0.9% NaCl) was administered intraperitoneally to control mice. APE2 KO mice and C57BL/6J wild-type mice received a single intraperitoneal dose of freshly dissolved cisplatin in saline (15 mg/kg body weight, 1 mg/mL in 0.9% NaCl; Sigma).

APE2 conditional transgenic mice were generated by inserting a transcription–translation STOP cassette flanked by LoxP sites (LSL) at the start of the APE2 locus. An FRT-mediated recombination ColA1 3′ untranslated region locus was used first to generate APE2^LSL mice. The STOP cassette prevents expression of the APE2 gene. To remove the STOP cassette and activate APE2 expression in vivo, the APE2^LSL mice were mated with Rosa26-Cre-Estrogen Receptor-T2 (CreERT2^+/−) mice (18; #8463, The Jackson Laboratory), in which the ERT moiety retains Cre recombinase activity in the cytoplasm until tamoxifen administration releases this block and promotes recombination of genomic LoxP sites. Three-week-old APE2^LSL/+CreERT2^+/− mice and CreERT2^+/− mice were injected intraperitoneally with 10 mg/kg 4-OH-tamoxifen for 3 consecutive days. Creatinine and EPO were measured using mouse creatinine (Crystal Chem) and EPO (R&D Systems) ELISA kits. APE2 KO mice were provided by Drs. D. Tsuchimoto and Y. Nakabeppu (Department of Immunobiology and Neuroscience, Medical Institute of Bioregulation, Kyushu University, Fukuoka, Japan; refs. 12, 19). We used heterozygous male (APE2^−/Y) mice (12) bearing a mutated APE2 allele on the X chromosome, in which a genomic region from the 3′-region of its intron 5 to the 5′-region in exon 6 was replaced by a neo cassette by crossing heterozygous APE2 female (APE2^−/+) mice with wild-type C57BL/6 mice. We did not use female APE2 knockout (APE2^−/−) mice because this can only be accomplished by crossing male APE2 knockout (APE2^−/Y) with heterozygous APE2 female (APE2^−/+) mice. Using this breeding strategy, there is the potential risk and uncertainty of interference with the germline DNA in the male and female APE2 knockout progeny because APE2 has already been knocked out in the male parent (APE2^−/Y) and because the function of APE2 is to be responsible for DNA damage repair (13). All mice were of C57B6 genetic background. All experiments involving animals were pre-approved by the Cleveland Clinic IACUC (Institutional Animal Care and Use Committee).

Kidney specimens were obtained from healthy donors and from cisplatin-induced AKI patients in accordance with Cleveland Clinic Foundation Institutional Review Board approval requirements, and written informed consent was obtained in compliance with the Helsinki.

HEK293T and HK2 cells were purchased from the ATCC. Mouse anti–Prominin-1 MicroBeads (130-092-333, Miltenyi Biotec) were used to isolate PTCs from APE2^LSL/− CreERT2^+/− mice, Cre-ERT2^+/− mice, C57B6 mice, and APE2 KO mice. The primary PTCs were cultured in DMEM/Nutrient Mixture F-12 (DMEM/F12) containing 10% FBS, 100 U/mL penicillin, and 100 μg/mL streptomycin. HEK293T and HK2 cells were routinely genotyped using the Human Cell Line Genotyping System (Promega) and were used within their 6 times passage. Mouse PTCs were genotyped using the primer for transgenic mice and were used within their 3 times passage. All cell lines were used Mycoplasma tested negative.

To determine the effect of APE2 on PTC growth, PTCs from APE2^LSL/−CreERT2^+/− mice and Cre-ERT2^+/− mice were seeded into 12-well plates. Cell growth was assessed using an IncuCyte ZOOM live-cell image monitoring system for 4 days (Essen BioScience).

PMEV-3FLAG-hAPE2 plasmid was constructed by using hAPE2 cDNA cloned into PMEV-3FLAG-empty vector, and HEK293T cells were transfected with the plasmid. Flag tag antibody was then used to pull down APE2-binding proteins. Anti-IgG was used as control. Pulldown samples were run on SDS-PAGE gels. The bands from the gel were cut out, washed/de-stained in 50% ethanol containing 5% acetic acid, dehydrated in acetonitrile, reduced with dithiothreitol, and alkylated with iodoacetamide before digestion. All bands were completely digested in-gel by using trypsin 5 μL (10 ng/μL) in 50 mmol/L ammonium bicarbonate and incubating overnight at room temperature. Peptides were extracted from the polyacrylamide in two aliquots of 30 μL 50% acetonitrile containing 5% formic acid. The extracts were combined and evaporated to <10 μL in a Speedvac and then resuspended in 1% acetic acid to make up a final volume of approximately 30 μL for LC-MS analysis.

The LC-MS system was a LTQ-Obitrap Elite hybrid mass spectrometer system (Thermo Fisher Scientific) and a Dionex 15 cm ×75 μmol/L id Acclaim Pepmap C18, 2 μm, 100 Å reversed-phase capillary chromatography column. Extracts were injected in 5 μL volumes, and the peptides were eluted with an acetonitrile/0.1% formic acid gradient at a flow rate of 0.25 μL/min introduced into the mass spectrometer source. The microelectrospray ion-source was operated at 2.5 kV. The digest was analyzed using the data-dependent multi-task capability of the instrument, acquiring full-scan mass spectra in the Orbitrap at a resolution of 60,000 to determine peptide molecular weights and production spectra in the ion trap to enable determination of the amino acid sequence in sequential scans. Data were analyzed by using all the collected CID spectra and searching the NCBI human reference sequence database (March 2015, with 99,739 entries) with the search programs Mascot (version 2.3.0) and SEQUEST (version 2.2). The data were uploaded into Scaffold (version 4.0) for protein and peptide validation. To identify proteins, a threshold of at least five CID spectra (spectral counts) was set; the proteins identified in APE2 pull-down/control pull-down samples at a level greater than 2.5-fold were collected by filtration and marked as APE2-binding proteins for further analysis.

Immunofluorescence staining was performed as described previously (20). The slides of HK2 cells were incubated sequentially with rabbit anti-APE2 antibody (GeneTex), goat anti-rabbit Alexa Fluor 488–conjugated secondary antibody (Abcam), mouse anti-MYH9 antibody (GeneTex), goat anti-mouse Alexa Fluor 647–conjugated secondary antibody (Abcam), and rabbit anti-COXIV antibody Alexa Fluor 5469–conjugated antibody (Santa Cruz Biotechnology). For immunofluorescence staining of PTCs from transgenic mice, slides of mouse PTCs were incubated sequentially with rabbit anti-APE2 antibody (kindly provided by Dr. Y. Nakabeppu) and goat anti-rabbit Alexa Fluor 488–conjugated secondary antibody (Abcam); human anti-MYH9 antibody (GeneTex) and goat anti-human Alexa Fluor 546–conjugated secondary antibody (Abcam); rabbit anti-COXIV Alexa Fluor 647–conjugated antibody (Santa Cruz Biotechnology). Images were captured by confocal microscopy (Leica TCS SP8) at ×630 magnification.

The functional activity of PTC mitochondria was measured by using a Seahorse XF24 Analyzer and a Seahorse XF Cell Mito Stress Test Assay (Agilent Technologies). Mitochondrial functionality was examined by direct measurement of the OCR of cells according to the manufacturer's instructions. Briefly, 8 × 10⁴ PTCs were seeded into 24-well plates and allowed to attach overnight. The cells were subjected to OCR detection by sequential injections of oligomycin, carbonyl cyanide p-trifluoromethoxyphenylhydrazone (FCCP), and a combination of rotenone and antimycin A. Basal OCR = measured OCR level before the injection of oligomycin. Adenosine triphosphate (ATP) synthesis–linked OCR (ATP-linked) = basal OCR − OCR following oligomycin injection. Maximum OCR = OCR following the injection of FCCP. Reserve respiration (Reserve resp) = maximum respiration − basal OCR. Proton leak-linked OCR = uncoupled OCR following oligomycin − nonmitochondrial OCR following injection of rotenone and antimycin A. Nonmitochondrial OCR (Non-mito) = OCR following the injection of rotenone and antimycin A.

Formalin-fixed paraffin-embedded tissue sections were de-paraffinized and then incubated with rabbit anti-APE2 polyclonal antibody (Bioss, bs-6587R), rabbit anti-cleaved caspase-3monoclonal antibody (Cell Signaling Technology, #9664S), rabbit anti-MYH9 antibody (GeneTex GTX113236), or rabbit anti–phosphor-histone H2A.X(Ser139) polyclonal antibody (Cell Signaling Technology, #2577) at 4°C overnight. After incubation with horseradish peroxidase (HRP)–conjugated goat anti-rabbit secondary antibody, signal was detected using a DAB Substrate kit (Abcam, ab64238) according the manufacturer's instructions. Images were obtained by means of a phase-contrast microscope (Leica DM2000 LED) and a digital camera (Leica DMC 2900).

Total protein samples were isolated with RIPA buffer, fractionated on 4%–12% Bis-Tris polyacrylamide gels (Thermo Fisher Scientific), and electroblotted onto PVDF membranes (Millipore). Membranes were blocked with 5% nonfat milk and incubated overnight with rabbit anti-APE2 polyclonal antibody (Bioss, #bs-6587R), rabbit anti-APE1 polyclonal antibody (Abnova, #H00000328-D02), mouse anti-LIG3 monoclonal antibody (Santa Cruz Biotechnology, #sc-56089), rabbit anti-MRE11 monoclonal antibody (Cell Signaling Technology, #4847), rabbit anti-CtIP polyclonal antibody (Bethyl Laboratories, #300-488A), or goat anti–β-actin polyclonal antibody (Santa Cruz Biotechnology, #sc-1615) overnight at 4°C. Membranes were then washed and incubated with HRP-linked anti-rabbit IgG secondary antibody (Cell Signaling Technology, #7074) or HRP-linked anti-mouse IgG secondary antibody (Cell Signaling Technology, #7076S). The specific protein–antibody complex was detected with HRP-conjugated rabbit anti-mouse IgG. Detection by chemiluminescence was carried out by means of a SuperSignal West Femto Maximum Sensitivity Substrate kit (Thermo Fisher Scientific) with β-actin as a loading control.

Mouse kidney biopsy specimens were submerged in 2.5% EM grade glutaraldehyde in 2.5% glutaraldehyde/4% paraformaldehyde in 0.2 mol/L sodium cacodylate buffer (pH 7.4) at 4°C immediately after sectioning and fixed at 4°C overnight. After washing three times for 5 minutes in sodium cacodylate buffer (0.2 mol/L, pH 7.3), kidney fragments were fixed in 1% aqueous osmium tetroxide for 60 minutes at 4°C, then washed twice for 5 minutes with sodium cacodylate buffer and rinsed once with maleate buffer (pH 5.1, 5 minutes). After changing to 1% uranyl acetate in maleate buffer, the samples were stained for 60 minutes; the uranyl acetate was removed. The samples were then washed three times for 5 minutes with maleate buffer and then dehydrated with ascending grades of ethanol, and finally embedded in Epon resin (Electron Microscopy Science). Ultrathin sections (85 nm) were cut by means of an EM UC7 ultramicrotome (Leica Microsystems), then successively stained with uranyl acetate and lead citrate, and examined with a transmission electron microscopy (TEM) instrument at 80 kV (Tecnai G2 SpiritBT, FEI).

Mouse kidney biopsy specimens from control and transgenic mice were fixed 4% PFA with 0.05% glutaraldehyde overnight at 4°C and then subjected to dehydration in 30% to 100% ethanol for 10 minutes at room temperature. The tissues were treated with 1:1 100% ethanol:LR white resin overnight, 100% LR white resin at 6 hours, and then re-embedded with LR white resin at 50°C to complete the polymerization. Ultrathin sections were sliced with a diamond knife (90-nm thickness) and mounted on nickel grids coated with formvar. The mounted grids were each washed three times with PBS for 5 minutes and then blocked with PBS containing 1% BSA for 30 minutes at room temperature. Diluted rabbit anti-mouse APE2 antibody (kindly provided by Dr. Y. Nakabeppu, Department of Immunobiology and Neuroscience, Medical Institute of Bioregulation, Kyushu University; refs. 12, 19) at 1:50 dilution and human anti mouse MYH9 antibody (#33939, GeneTex) at 1:100 dilution in PBS with 1% BSA were added and incubation was performed overnight at 4°C. The grids were washed with PBS three times for 5 minutes each and then blocked with 0.1% BSA in PBS for 10 minutes at room temperature. Diluted 5-nm gold-conjugated anti-rabbit IgG and 15-nm gold-conjugated anti-human IgG with 0.1% BSA in PBS (1:10 dilution), then added to each grid a 30 μL drop of GAM for 60 minutes at room temperature and the grid was then washed with PBS three times (5 minutes per wash). The antibodies were fixed for 10 min with 1% glutaraldehyde in PBS. After being washed with water three times (5 minutes per wash), the grids were stained with UA and lead citrate. After air dried, the grids were observed with a Tecnai G2 SpiritBT electron microscope operated at 80 kV.

A two-tailed student t test was performed using SPSS (version 17.0) to compare independent pairs of groups. In our mouse model, the time to death was determined by the Kaplan–Meier method, with Cox proportional hazard regression analysis for group comparisons. A P value of ≤0.05 was statistically significant.

The majority of kidney cisplatin toxicity studies in rodents found marked histological changes, including proximal and distal tubular necrosis and glomerular toxicity, which are the clinical features of C-AKI disease in humans. We found as a prelude to the present work that mice treated with cisplatin (15 mg/kg) developed proximal and distal tubular necrosis and glomerular toxicity as described in the literature (21). Analysis of BER pathway protein expression in cisplatin-treated mouse kidney revealed that APE2 is the most highly upregulated protein (Fig. 1A), whereas APE1, LIG3, MRE11, and CTIP are slightly upregulated and PCNA is downregulated. APE2 is a key player in repair of mtDNA damage (15, 22). Because the ability of cisplatin to induce nuclear DNA damage per se is not sufficient to explain its toxic effects in post-mitotic normal tissues, we believe that upregulated APE2 may be responsible for cytotoxicity in PTCs. Carboplatin and oxaliplatin, are thought to be less nephrotoxic than cisplatin because they lack chloride atoms and display decreased organic cation transporter-2 (OCT-2) cellular uptake in PTCs than in other types of cells. When we treated mice with carboplatin and oxaliplatin a slight upregulation of APE2 in kidney cells was observed (Supplementary Fig. S1). In the clinic, cisplatin-induced AKI is much more common than carboplatin and oxaliplatin but very few patients are likely to be biopsied. We studied APE2 expression in two patients with C-AKI using IHC staining for APE2 and found the protein to be dramatically upregulated in their PTCs in comparison with their biopsied normal kidneys (Fig. 1B).

To determine whether C-AKI develops in mice as a consequence of cisplatin-mediated effects via APE2 overexpression, we generated APE2 conditional transgenic mice (Fig. 2A). To mimic the effect of cisplatin in all organs, we crossed APE2^LSL/− mice with CreERT2^+/− mice, in which Cre-induced expression of APE2 is controlled by the CAG promoter, a well-established strategy for generating transgenic mice with high expression of APE2 in nearly all organs after tamoxifen administration (Fig. 2B; ref. 18). We then evaluated the APE2^LSL/− CreERT2^+/− mice for the development of typical cisplatin treatment-related pathology. Two weeks after tamoxifen-induced transgene expression, body and kidney weight in 5-week-old APE2^LSL/− CreERT2^+/− mice were reduced to a remarkable degree (Fig. 2C and D). We observed C-AKI–like pathologic features (Fig. 2D) in APE2^LSL/− CreERT2^+/− mice. In molecular level, we observed similar changes as in C-AKI mice (Fig. 1A) except that CTIP expression was lower in APE2^LSL/− CreERT2^+/− mice as compared with CreERT2^+/− control mice (Fig. 2E).

With aging, all APE2^LSL/− CreERT2^+/− mice developed irreversible C-AKI–like disease, weight loss (Fig. 3A), and pale/stiff kidneys (Fig. 3B). Dissection revealed a small, pale (Fig. 3B), and necrotic (Fig. 3C) kidney but no necrotic changes in other organs (Supplementary Fig. S2). Also noted were late-stage C-AKI pathologic features, and ongoing apoptosis in PTCs isolated from APE2^LSL/− CreERT2^+/− mice kidney (Fig. 3D). All the APE2^LSL/− CreERT2^+/− mice died of kidney failure at 15 weeks of age, but none of the CreERT2^+/− control mice (Fig. 3E) showed these changes or died. More importantly, APE2^LSL/− CreERT2^+/− mice developed clinical features of C-AKI, including high urine albumin (Fig. 3F) and blood creatinine (Fig. 3G) and decreased EPO (Fig. 3H). Furthermore, at one week after tamoxifen injection, increased urine albumin was seen in APE2^LSL/− CreERT2^+/− mice but not in CreERT2^+/− mice, indicating the kidney injury to be primary rather than secondary, and to be due to some other organ failure (e.g., of the heart or to tamoxifen toxicity). Furthermore, the pathologic features of APE2^LSL/− CreERT2^+/− mice is known to differ from the renal glomerular disease induced by hypertension or diabetes, mimicking instead C-AKI kidney damage. In the present work, we observed apoptotic PTCs in APE2^LSL/− CreERT2^+/− mice and DNA damage (Fig. 4A). Electron microscopy revealed marked mitochondrial fragmentation in APE2^LSL/− CreERT2^+/− mouse PTCs but not in control CreERT2^+/− mouse PTCs (Fig. 4B).

To rule out the possibility of PTC death due to APE-induced microenvironmental dysregulation, we isolated and cultured PTCs from APE2^LSL/− CreERT2^+/− mice and CreERT2^+/− control mice. After we added 4-OH tamoxifen, we found that PTCs from APE2^LSL/− CreERT2^+/− mice died (Fig. 5A) much faster (Fig. 5B) than those from CreERT2^+/− control mice, indicating that APE2 overexpression in PTCs induces cell death directly rather than via secondary changes in the kidney microenvironment. We also found the OCR to be dramatically reduced in the APE2 transgenic mice as compared with control mice (Fig. 5C), pointing to mitochondrial dysfunction induced by overexpression of APE2.

To explore the mechanism underlying APE2 induction of AKI disease, we performed an APE2 IP pulldown experiment, followed by liquid chromatography-mass spectrometry (LC-MS) proteomic analysis using Flag-APE2 overexpressed in HEK293T cells. The most abundant APE2 pulldown proteins were found to be MYH9/10/14 (Supplementary Table S1). Patchy expression of MYH9/10/14 has previously been reported to be ubiquitous in all renal tubule segments and in glomerulus cells (23). We chose to focus on MYH9 protein because only MYH9 knockout and MYH9 R702C-, D1424N-, E1841K-mutated mice develop a phenotype similar to that of APE2 transgenic mice (17). To further explore the subcellular localization pattern of APE2 and MYH9 in PTCs during cisplatin treatment, HK2 and primary mouse PTCs were treated with cisplatin and subjected to immunofluorescence analysis. Immunofluorescence was used to identify the subcellular localization of APE2 and MYH9 in primary mouse PTCs, with COX IV as a mitochondria marker. As shown in Fig. 6A and B, in control HK2 cells and mouse PTCs, APE2 was localized in both the nucleus and the cytoplasm, and was also partially colocalized with MYH9 in the cytoplasm. When the cells were treated by cisplatin, APE2 was similarly enriched in the cytoplasm, especially within mitochondria.

In PTCs from CreERT2^+/− control mice, APE2 was localized in both the nucleus and the cytoplasm, whereas MYH9 localized only in the cytoplasm (Fig. 6C). APE2 was also partially colocalized with MYH9 in the cytoplasm. When APE2 was ectopically overexpressed in APE2 transgenic mice, it was granularly enriched in the mitochondria without cisplatin treatment, mimicking cisplatin-induced APE2 overexpression and enrichment in mitochondria. Colocalization of APE2 and MYH9 in mouse kidney mitochondria was further confirmed by immunoelectron microscopy. As shown in Fig. 6D, APE2 and MYH9 were colocalized in the mitochondria. Furthermore, we also observed MYH9 enrichment in the cytosol of PTCs from C-AKI patients (Fig. 6E).

To test whether APE2 KO can prevent cisplatin-induced AKI, we used APE2-null male mice (12) and wild-type male mice to perform the study. The APE2-null male mice showed moderate growth retardation (12). Their body weight was only approximately 80% of the weight of their wild-type littermates at birth, and this decrease persisted into adulthood, indicating that all developing APE2-null embryos, infants, and adults are somehow retarded in their growth. We treated 6-week-old wild-type (WT) and APE2 KO mice with cisplatin and then toxicity was evaluated at one-week post-treatment. Renal damage was most severe in WT mice, which also had decreased kidney weight. In contrast, APE2 KO mice (Fig. 7A) had only minor kidney damage (Fig. 7B) and less mitochondrial fragmentation (Fig. 7C), suggesting that APE2 KO in cisplatin-treated mice can prevent C-AKI development. To examine mitochondrial functional activity in PTCs from WT and APE2 KO mice, PTCs were isolated and subjected to Cell Mito Stress assay by using Seahorse. As shown in Fig. 7D, the OCR was similar in nontreated WT and APE2 KO PTCs. After the mice were treated by cisplatin, however, OCR in PTCs was significantly increased in APE2 KO mice (Fig. 7E).

Because the molecular mechanism of action of C-AKI has not been determined until now, we still have no genetic engineered mouse model that recapitulates the clinical features of C-AKI. Currently, only cisplatin-induced AKI (C-AKI) mouse model is widely used. Many potential kidney-protective therapies have been tested in this laboratory model and have been found to provide some therapeutic benefit, but unfortunately all of them failed in early-stage clinical trials (24). If we had available a functionally engineered transgenic mouse model based on the mechanism underlying C-AKI and accurately reflecting human pathogenesis, we could determine not only the etiology and pathology of C-AKI, but also develop preventive strategies and decrease the need for cisplatin dose adjustment or cessation. Cisplatin treatment caused the proximal and distal tubular necrosis in the mouse kidney. Notably, the kidney proteins related to the BER pathway, such as APE2, APE1, and LIG3, homologous recombination (HR) pathway, such as MRE11 and CTIP, were increased whereas PCNA was decreased after treatment with cisplatin. In APE2-overexpressing transgenic mice, similar pathological features were found in mouse kidney. However, expression of additional kidney proteins (MRE11 and CTIP) was observed. The discrepancy of MRE11 and CTIP expression found between C-AKI and APE2-AKI mice models may be ascribed to the fact that MRE11 and CTIP are related to HR pathway but not BER pathway, which is only responsively upregulated in C-AKI mice but not in APE2-AKI mice. The most attractive feature of our APE2 AKI mouse model is that it mechanistically recapitulates cisplatin-induced kidney damage but excludes other cisplatin-induced DNA damage repair response such as the HR pathway.

From our mechanistic studies, it may be concluded that when APE2 overexpression is induced by cisplatin, the APE2 that is enriched in mitochondria binds to MYH9 and subsequently interferes with mitochondrial fission (Fig. 7F). MYH9 mutations are associated with several human syndromes, currently grouped as MYH9-related disorders, which are characterized by kidney disease, hearing loss, thrombocytopenia, and cataracts (25, 26). Three hexameric isoforms—IIA, IIB, and IIC—are encoded by MYH9, MYH10, and MYH14, respectively, all of which have a similar structure. The MYH9 isoform has important roles in cell adhesion, migration, proliferation, and differentiation (27). Some patients with MYH9 mutations develop progressive proteinuria, glomerulosclerosis, and kidney failure (28). MYH9 SNPs have been proposed as a major genetic risk locus for non-diabetic end-stage kidney disease (26). Functional studies of the three isoforms in mice have elucidated both unique and redundant functions for MYH9 and MYH10 (29). The importance of MYH14 is unknown, as global MYH14 knockout in mice has no obvious phenotype (30). The variable occurrence of the kidney‐specific phenotype in patients with MYH9‐related disorders is possibly due to the type of MYH9 mutation, along with variable compensation by MYH10 and MYH14 (23). Other potential explanations include defects in endocytosis and receptor‐mediated transport in tubule epithelial cells and defects in glomerular filtration, but the exact mechanism underlying MYH9-related disease is not clear. Pathological mutations in MYH9 cause kidney dysfunction, hearing loss, and hematopoietic disorders in humans, clinical features that are also seen with cisplatin treatment, suggesting that the underlying pathophysiological mechanisms may be similar.

Two recent reports indicate that MYH9 and actin contribute to mammalian mitochondrial DNA integrity and fission (31, 32). Impaired mtDNA homeostasis may account for one or more of the clinical features associated with MYH9 mutation (31). Accumulated data also point to the involvement of MYH9 and/or actin in membrane morphogenesis events, such as mitochondrial fission (33). Mitochondrial fission is involved in a quality-control mechanism whereby damaged mitochondrial components are segregated from healthy ones, with subsequent mitochondrial fission and degradation of the damaged daughter mitochondrion (33). Mitochondrial fission requires action by the enzyme dynamin GTPase (DRP1; ref. 34). However, mechanisms controlling DRP1 assembly in mammalian mitochondria are unclear. Recent results from Korobova and colleagues (33) show that MYH9 inhibition decreases DRP1 association with mitochondria and plays an important role in mitochondrial fission. These authors proposed a mechanistic model in which actin polymerization leads to myosin II recruitment and constriction at the fission site, enhancing subsequent DRP1 accumulation and fission. Futher study the dynamic functional relationship between APE2, MYH9, actin, and DRP1 will further unravel the molecular mechanism and pathogenesis of C-AKI. The influence of gender on cisplatin tolerability in rodents and patients is not entirely certain. One study has found that male C57BL/6 mice possess approximately 10-fold higher levels of GSTP expression in the liver than females and are more susceptible to cisplatin nephrotoxicity (35), whereas another study has found female mice (C57BL/6J and 129Sv) to be more susceptible than males (36). In the present work, we found no differences in phenotype development between genders in our APE2 transgenic cohort. A larger age-matched cohort study and additional mechanism studies using patients' biopsy samples would be needed to address the issue of whether there exists a gender difference in the level of cisplatin-induced AKI.

In summary, we have developed a genetically engineered APE2 mouse model that affords a clearer understanding of cisplatin related nephrotoxicity. This knowledge may prove useful in guiding future drug development efforts aimed at reducing, or even entirely avoiding, acute cisplatin–induced kidney injury.

F. Anwer reports speaker bureau from InCyte. N.C. Munshi reports personal fees from BMS, Amgen, Jassen, Karyopharm, Takeda, Abbvie, Legend, Novartis, Adaptive, and BeiGene, as well as personal fees and other fees from Oncopep outside the submitted work. J. Zhao reports a patent for Targeting APE2 to treat acute kidney injury pending. No disclosures were reported by the other authors.

Y. Hu: Data curation, formal analysis, writing-review and editing. C. Yang: Data curation, formal analysis, investigation. T. Amorim: Data curation. M. Maqbool: Formal analysis. J. Lin: Investigation. C. Li: Data curation. C. Fang: Data curation. L. Xue: Data curation. A. Kwart: Data curation. H. Fang: Data curation. M. Yin: Data curation, visualization. A.J. Janocha: Resources, data curation, methodology. D. Tsuchimoto: Resources. Y. Nakabeppu: Resources. X. Jiang: Resources. A. Mejia-Garcia: Data curation, writing-review and editing. F. Anwer: Resources, writing-review and editing. J. Khouri: Writing-review and editing. X. Qi: Resources, methodology. Q. Zheng: Writing-review and editing. J. Yu: Conceptualization, resources. S. Yan: Data curation, validation. T. LaFramboise: Data curation, validation. K.C. Anderson: Resources. L.C. Herlitz: Resources, data curation, formal analysis, validation, writing-review and editing. N.C. Munshi: Conceptualization, resources, data curation, formal analysis, writing-review and editing. J. Lin: Conceptualization, resources, data curation, formal analysis, supervision, writing-original draft, writing-review and editing. J. Zhao: Conceptualization, resources, data curation, formal analysis, supervision, funding acquisition, validation, investigation, writing-original draft, project administration, writing-review and editing.

The authors thank the Lerner Research Institute proteomic, genomic and imaging cores for their assistance and support. The authors thank Dr. Mark R. Kelley from Indiana University School of Medicine, Drs. Thomas M. McIntyre, Thomas Egelhoff, Donna M. Driscoll, and Serpil C. Erzurum from Cleveland Clinic Lerner Research Institute for valuable discussions. This work was supported by grants (to J. Zhao) from the National Cancer Institute (R00 CA172292), start-up funds and two core utilization pilot grants from the Clinical and Translational Science Collaborative of Cleveland, a V Foundation Scholar Award, an American Society of Hematology (ASH) Bridge Grant, an American Cancer Society Institutional Research Grant Pilot Award, a National Institutes of Health training grant (T32 CA094186, “Training in Computational Genomic Epidemiology of Cancer” to J. Lin), a grant (R01 DC015111/NIDCD to Q.Y. Zheng) from National Institutes of Health, and a grant (R01HL60917 to Serpil C. Erzurum) from National Institutes of Health, a grant (5UL1TR002548) from the NIH National Center for Advancing Translational Sciences (NCATS), a component of the National Institutes of Health and NIH roadmap for Medical Research. The Orbitrap Elite instrument was purchased with the help of an NIH shared-instrument grant (1S10RR031537-01).

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