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Cisplatin is currently the most effective antitumor agent available against bladder cancer. However, a majority of patients eventually relapse with cisplatin‐resistant disease. Chemoresistance thus remains a major obstacle in bladder cancer therapy. To clarify the molecular mechanisms underlying cisplatin resistance in bladder cancer, we established a cisplatin‐resistant subline from the human bladder cancer cell line HT1376 (HT1376‐CisR), and conducted large‐scale analyses of the expressed proteins using two‐dimensional (2D) gel electrophoresis coupled with mass spectrometry (MS). Comparative proteomic analysis of HT1376 and HT1376‐CisR cells revealed 36 differentially expressed proteins, wherein 21 proteins were upregulated and 15 were downregulated in HT1376‐CisR cells. Among the differentially regulated proteins, adseverin (SCIN), a calcium‐dependent actin‐binding protein, was overexpressed (4‐fold upregulation) in HT1376‐CisR, with the increase being more prominent in the mitochondrial fraction than in the cytosol fraction. SCIN mRNA knockdown significantly reduced cell proliferation with mitochondria‐mediated apoptosis in HT1376‐CisR cells. Immunoprecipitation analysis revealed voltage‐dependent anion channels (VDACs) to be bound to SCIN in the mitochondrial fraction. Our results suggest that the VDAC‐SCIN interaction may inhibit mitochondria‐mediated apoptosis in cisplatin‐resistant cells. Targeting the VDAC‐SCIN interaction may offer a new therapeutic strategy for cisplatin‐resistant bladder cancer.
Bladder cancer is the second most common malignancy of the genitourinary tract and the second leading cause of genitourinary cancer deaths in the United States of America (American Cancer Society, 2010). Although locally advanced or metastatic bladder cancer is generally a chemosensitive disease, the prognosis for these patients remains poor. Cisplatin‐based chemotherapy is the only modality that provides the potential for long‐term survival of bladder cancer patients. Recently, the combination of cisplatin and gemcitabine has been established as a standard treatment of metastatic bladder cancer. von der Maase et al. (2000) have reported the clinical response rate to be 49%, and our experience with Japanese patients showed a response rate of 63% (Tanji et al., 2010). However, a majority of these patients eventually relapse with cisplatin‐resistant disease. Cancer cells develop resistance to chemotherapy by inactivating apoptotic factors and enhancing survival pathways that antagonize apoptotic signals (Gonzalez et al., 2001). The acquisition of chemoresistance thus remains a major obstacle in cancer treatment, ultimately leading to death.
Cisplatin has been used to treat various cancers and, in particular, is accepted worldwide as a necessary anticancer agent in bladder cancer chemotherapy. The mechanisms underlying cisplatin resistance have been studied in depth. These mechanisms include reduced intracellular drug accumulation, increased detoxification of drug by thiol‐containing molecules, increased DNA damage repair, escape from reactive oxygen species‐mediated cytotoxicity, and the action of apoptosis mediators (Hour et al., 2010; Siddik, 2003; Tsunoda et al., 2005). However, the molecular mechanisms of cisplatin resistance acquisition in bladder cancer are not fully understood.
Proteins that are differentially expressed in cisplatin‐resistant bladder cancer are logical candidates as relevant biomarkers and therapeutic targets. The proteomic approach is a potent strategy that can provide insights into global protein changes involved in cisplatin multidrug resistance. Recently, quantitative proteomic technologies have been successfully used to identify differentially expressed proteins in several chemoresistant cancer cell lines (Dai et al., 2010; Sun et al., 2011; Zheng et al., 2010). Two‐dimensional gel electrophoresis (2‐DE) can quantify hundreds of proteins at once, and the development of matrix‐assisted laser desorption/ionization (MALDI) or electrospray ionization (ESI) mass spectrometry (MS) allows for highly sensitive identification of gel‐separated proteins even at low femtomolar levels. Recent advances in quantitative MS methods, such as multiple reaction monitoring (MRM), are promising tools for discovering protein markers of drug resistance from limited amounts of clinical samples. It has been demonstrated that by using these methods, proteins can be detected even at unprecedented low levels; moreover, these are high‐throughput assays (Picotti et al., 2010; Schmitz‐Spanke and Rettenmeier, 2011).
In the present study, in order to further clarify the mechanisms underlying cisplatin resistance, we established a cisplatin‐resistant subline (HT1376‐CisR) derived from the human bladder cancer cell line HT1376. We applied a quantitative proteomic approach that combined 2‐DE and nano‐liquid chromatography (nanoLC) MS technology to identify proteins using differential expression profiles between HT1376 and HT1376‐CisR. In addition, functional analyses of the proteins expressed differentially in the cisplatin‐resistant cell line were conducted using biochemical and molecular biological techniques.
RPMI‐1640 and fetal bovine serum (FBS) for cell culture were supplied by Gibco (St. Louis, MO). Cisplatin was purchased from Sigma–Aldrich (Tokyo, Japan). Antibodies against adseverin (SCIN) and β‐actin were also obtained from Sigma–Aldrich. Antibodies against poly (ADP‐ribose) polymerase (PARP) and voltage‐dependent anion channel (VDAC) 1 were obtained from Cell Signaling Technology (Danvers, MA) and Lifespan Biosciences (Seattle, WA), respectively. Antibodies against VDAC2 and COXIV were purchased from Abcam (Tokyo, Japan).
The human bladder cancer cell line HT1376 used in this study was purchased from DS Pharma Biomedical (Osaka, Japan) in 2007. Cells were maintained in the RPMI‐1640 medium supplemented with 10% FBS in a humidified incubator at 37 °C in an atmosphere of 5% CO2 and 95% air. Cisplatin‐resistant cells (HT1376‐CisR) were obtained from the parental HT1376 cells using an intermittent stepwise selection protocol over 12 months, ending with exposure to 5 μM cisplatin.
Cells were seeded in 24‐well plates at 1.0 × 104 cells/well and cultured with graded concentrations of cisplatin in at least three replicate wells at 37 °C. At 72 h after cisplatin exposure, the cells were counted by Cell Counter (Invitrogen). The number of the untreated cells by cisplatin was considered to be 100%. The drug concentration that resulted in 50% growth inhibition (IC50) was determined from corresponding dose–response curve.
The cultured cells were washed twice with cold phosphate‐buffered saline (PBS) and lysed in a lysis solution containing 8 M urea, 4% CHAPS, 0.2% (w/v) Bio‐Lyte 3/10 (Bio‐Rad), and 1% (w/v) dithiothreitol. The amounts of proteins in the lysate were assayed using the DC/RC protein assay kit (Bio‐Rad). Extracted proteins were precipitated using the methanol–chloroform–water method (Wessel and Flügge, 1984) and resolubilized in 250 μL of lysis buffer. After centrifugation at 15,000× rpm for 10 min, the supernatants (180 μg total proteins) were loaded onto Immobiline Dry Strips (pH 3–10, 13 cm in length; GE Healthcare). Immobiline Dry Strips were rehydrated for 15 h at room temperature. Isoelectric focusing (IEF) was carried out using a CoolPhoreStar IEF apparatus (Anatech, Tokyo, Japan). Electrophoresis was performed as follows: 500 V for 1 h, 700 V for 15 min, 1000 V for 15 min, 1500 V for 15 min, 2000 V for 15 min, 2500 V for 15 min, 3000 V for 15 min, and 3500 V for 6 h. The strips were incubated for 30 min in SDS equilibration buffer [50 mM Tris–HCl (pH 8.8), 8.5 M urea, 30% (v/v) glycerol, 2% (w/v) sodium dodecyl sulfate (SDS), and 2% (w/v) dithiothreitol (DTT)] and transferred onto 12.5% SDS‐polyacrylamide gels. After IEF electrophoresis, the gels were stained with Bio‐Safe Coomassie (Bio‐Rad). The gel images were captured using a 16‐bit scanner GELSCAN (iMeasure, Nagano, Japan). Spot detection, matching, and quantitative analyses were performed with Progenesis software version 4.0 (Nonlinear Dynamics, UK).
Protein spots were excised from 2‐D gels and subjected to in‐gel tryptic digestion with sequence‐grade modified trypsin as described previously (Takemori and Yamamoto, 2009). After repeated extractions of tryptic digests from the gel with 50% (v/v) acetonitrile/5% (v/v) trifluoroacetic acid, the solution containing the extracted peptides was concentrated using a vacuum centrifuge. The tryptic peptide sample was reconstituted with 0.2% (v/v) trifluoroacetic acid and subjected to MS analysis.
Tryptic digests of protein extracts from HT1376 and HT1376 Cis‐R were prepared as follows: precipitated proteins (30 μg) in 30 μL of 6 M urea and 2.5 mM EDTA in 0.5 M Tris–HCl buffer (pH 8.8) were reduced by adding 10 μL of 40 mM DTT solution for 2 h at 37 °C and alkylated by adding 10 μL of 250 mM acrylamide solution for 30 min at room temperature. The protein solutions were diluted 4‐fold with 50 mM ammonium bicarbonate solution and digested with 0.5 μg of sequence‐grade trypsin (Promega, Madison, WI) at 37 °C for 12 h. Digestion was terminated by adding 5% trifluoroacetic acid (TFA), and the tryptic peptides were desalted and concentrated on a self‐prepared STAGE tip (Rappsilber et al., 2003).
Tandem MS (MS/MS) and MRM analysis were performed using the QTRAP 5500 hybrid triple quadrupole/linear ion trap system (AB‐SCIEX; Foster, CA) coupled with Prominence NanoLC (Shimadzu, Kyoto, Japan). Peptide separations were performed at a constant flow rate of 300 nL/min with a fused silica capillary column packed with C18 resin (75 μm × 15 cm). Mobile phases used for separation were 0.1% formic acid (A) and 80% acetonitrile with 0.1% formic acid (B). A gradient (5–60% mobile phase B) was applied for 25 min, followed by a 10‐min wash at 100% mobile phase B, followed by equilibration for 15 min with 5% mobile phase B. The MRM channel was selected using the MRMPilot Software (AB‐SCIEX). The Q1 and Q3 resolutions were set to unit resolution (0.7 Da). Relative quantization was performed using MultiQuant software (AB‐SCIEX), and protein identification was performed using ProteinPilot 4.0 software (AB‐SCIEX). Database searches were performed against the IPI human database version 20100624.
Protein samples were prepared by lysing cells in ice‐cold lysis buffer and a protease inhibitor cocktail mix (Sigma–Aldrich). Cell lysates were centrifuged at 12,000×g for 10 min at 4 °C. Equal amounts of protein were denatured in a sample buffer, subjected to SDS‐PAGE analysis, and then transferred to polyvinylidene fluoride (PVDF) membranes. After treating with skimmed milk, blots were incubated with specific antibodies at 4 °C overnight followed by appropriate horseradish peroxidase (HRP)‐conjugated secondary antibodies at room temperature for 1 h. Signals were visualized by using a chemiluminescence ECL reagent (GE Healthcare, Giles, UK).
For siRNA experiments, an SCIN siRNA cocktail was obtained from B‐Bridge International (Cupertino, CA) after being functionally annotated. The siRNA cocktail was as follows:
sense 1: 5′‐GGAUGAUGGUUCUGGCAAATT‐3′,
antisense 1: 5′‐UUUGCCAGAACCAUCAUCCTT‐3′,
sense 2: 5′‐GGAGAAAGGAGCAGAGUAUTT‐3′,
antisense 2: 5′‐AUACUCUGCUCCUUUCUCCTT‐3′,
sense 3: 5′‐GGGAAAAGCUUUUUGCUUATT‐3′,
antisense 3: 5′‐UAAGCAAAAAGCUUUUCCCTT‐3′.
For the control, we used non‐targeting siRNAs. The non‐targeting siRNAs included the following: 5′‐ATCCGCGCGATAGTACGTA‐3′, 5′‐TTACGCGTAGCGTAATACG‐3′, and 5′‐TATTCGCGCGTATAGCGGT‐3′.
siRNAs were transfected into HT‐1376‐CisR cells using Lipofectamine RNAiMAX (Invitrogen, Carlsbad, CA) and Optimem I (Invitrogen) at a concentration of 40 nmol/L according to the manufacturer's instructions. In general, each assay was carried out 48 h after the transfection.
Cells were seeded in 24‐well plates at 1 × 104 cells/well and cultured with 5 μM cisplatin; then, the cells were counted by Cell Counter (Invitrogen) at the indicated times.
After washing cells with PBS, mitochondrial and cytosolic extracts were isolated using Mitochondria/Cytosol Fractionation kits (BioVision, Mountain View, CA) according to the manufacturer's instructions.
Caspase activation was visualized by immunoblot analysis for cleavage of PARP using a rabbit anti‐PARP polyclonal antibody and an HRP‐conjugated goat anti‐rabbit IgG antibody (Cell signaling technology). Changes in the mitochondrial membrane potential were examined using a fluorescent microscope and the MitoCapture™ Mitochondrial Apoptosis Detection Kit (BioVision). In brief, after washing cells with PBS, cells were incubated in 1 mL of the diluted MitoCapture solution containing 1 μL of MitoCapture dye at 37 °C for 20 min in a CO2 incubator. Thereafter, cells were observed under a fluorescence microscope with a bandpass filter (OLYMPUS, Tokyo, Japan). In apoptotic cells, MitoCapture, which is a cationic dye, cannot aggregate in the mitochondria because of altered mitochondrial transmembrane potential, and remains in the cytoplasm in its monomer form.
The samples were incubated with anti‐VDAC1 or anti‐VDAC2 antibody overnight at 4 °C, then added to 50 μL of Immobilized Protein A resin slurry (Thermo). The samples were gently mixed and incubated for 2 h at room temperature. The complex‐bound resin was washed with water and centrifuged at 2500×g for 3 min. The precipitations were added to a 2X electrophoresis loading buffer and incubated for 5 min at 95 °C. They were then centrifuged at 2500×g, and the supernatants subjected to SDS‐PAGE. Proteins were blotted onto PVDF membranes and incubated in blocking solution (5% skim milk in PBS with 0.05% Tween 20) for 1 h at room temperature. For SCIN expression, membranes were then probed with anti‐SCIN polyclonal antibody for 1 h at room temperature. The specificity of the interaction between VDAC and SCIN was verified with immunoblot of immunoprecipitated samples. HRP‐conjugated secondary‐antibodies were used within 1 h at room temperature. Proteins were visualized by using an ECL Western blotting reagent (GE Healthcare).
Each value represents the mean ± standard deviation (SD) of at least three independent results. Statistical significance (P < 0.05) was determined by using Student's t‐test using the IBM SPSS 18.0 software program (Armonk, NY).
First, to confirm the acquisition of cisplatin resistance in HT1376‐CisR cells, the cytotoxicity of cisplatin was examined Figure 1 shows the relative number of surviving cells in various concentrations of cisplatin in HT1376 and HT1376‐CisR cells. The IC50 values for cisplatin treatment in HT1376 and HT1376‐CisR cells were 3.0 × 102 and 1.1 × 105 nM, respectively. The IC50 for HT1376‐CisR was thus 367‐fold higher than that of HT1376 cells, indicating that the cisplatin‐resistant cell line was successfully established.
IC50 values of cisplatin in parental and HT1376‐CisR cells. IC50 represents the concentration of drug that is required for 50% inhibition in vitro. HT1376 and HT1376‐CisR cells were treated with various concentrations of cisplatin ...
We identified >500 protein spots in 2‐DE, which separated extracted proteins from HT1376 or HT1376‐CisR cells, and established the reference proteome maps for both cell lines (Figure 2A). Progenesis software (Nonlinear) was used to compare the global protein profiles of HT1376 and HT1376‐CisR cells. Protein spots demonstrating consistent differences (≥2‐fold) between the two cell lines in triplicate experiments were chosen as differential protein spots, and we depicted the protein expression profiles using a scattergram. Protein spots that were significantly changed (≥2‐fold) between the two cell lines were indicated as red spots (Figure 2B). We queried 43 differential protein spots, in which 22 proteins were upregulated and 21 were downregulated in HT1376‐CisR cells as compared with HT1376 cells (Table 1). In Figure 2C, we have shown representative gel images of areas containing protein spots that were significantly upregulated and downregulated. Protein identification for each spot was conducted using nanoLC‐MS/MS, and the quantity of each target protein was verified by the MRM assay. Based on the tandem MS information obtained from tryptic digests of the proteins of interest, we selected Q1 (precursor)/Q3 (fragment) ion pairs for these peptides (Figure 3A), and measured the expression levels in the MRM mode (Figure 3B). In HT1376‐CisR, we noted significantly higher expression of galectin‐3 and CBR1, which are known to be associated with drug resistance (Fukumori et al., 2007; Plebuch et al., 2007). However, we also observed the increased expression of SCIN, which has not been previously described for HT1376‐CisR cells. Thus, of the proteins upregulated in the HT1376‐CisR cells, we focused on SCIN.
Proteomic analysis of differentially expressed proteins using two‐dimensional gel electrophoresis. A) Representative 2‐D gel images of HT1376 and HT1376 Cis‐R cell extracts. Whole‐cell extracts were separated according ...
Large‐scale MRM analysis of differentially expressed proteins between HT1376 and HT1376 Cis‐R using hybrid triple quadrupole/linear ion trap mass spectrometer. A) Representative MS/MS spectrum of SCIN tryptic peptide: SLGGQAVQIR (doubly ...
Differentially expressed proteins between HT1376 and HT1376‐CisR cells in 2‐D gel based proteomics analysis
The SCIN tryptic peptide SLGGQAVQIR was detected in HT1376 and HT1376‐CisR whole‐cell lysates using the MRM assay (Figure 3B). Further, SCIN expression in HT1376‐CisR was revealed to be 4 times as high as that in HT1376 (Table 2). We also assessed the expression levels of SCIN protein by western blotting; these were significantly higher in HT1376‐CisR cells than in the parental cells (Figure 4A). Moreover, although differences in the expression levels of SCIN were detectable between HT1376‐CisR and HT1376 cells in the cytosol and mitochondrial fractions both, the differences in the mitochondrial fractions were more prominent than those in the cytosol fractions (Figure 4B).
Analysis of the functions and the location of SCIN protein. A) Western blot analysis was performed to measure SCIN expression in HT1376 and HT1376‐CisR cells. β‐Actin was included as a loading control. B) Mitochondrial and cytosolic ...
MRM assay for quantitative verification of target proteins in HT1376 and HT11376‐CisR cells
We transiently transfected lipofectamine RNAiMAX with SCIN or non‐targeting siRNA. HT1376‐CisR cells (1 × 105) were transfected with 50 nM of each siRNA. After 48 h, SCIN protein expression was noted to be markedly decreased in comparison to cells treated with control siRNA as well as untreated cells (Figure 4C). Then, to clarify the involvement of SCIN in cell proliferation, we assessed siRNA‐mediated growth inhibition. At 48 h after transfection, 5 μM cisplatin was added to the medium, and the cells were cultured for 7 d. As shown in Figure 4D, SCIN siRNA transfection significantly inhibited cell proliferation. Subsequently, to clarify the mechanism underlying SCIN siRNA‐mediated inhibition of HT1376‐CisR cell growth, we assessed apoptosis in HT1376‐CisR cells following SCIN‐knockdown. Mitochondrial membrane integrity was analyzed using the MitoCapture™ apoptosis detection kit. HT1376‐CisR cells transfected with either negative control or SCIN siRNAs were incubated for 48 h. Then, cisplatin (5 μM) was added to the transfected cells; all cells were analyzed at 48 h after cisplatin exposure. The number of cells labeled with yellow, indicating surviving cells, were significantly decreased in the SCIN‐knockdown HT1376‐CisR cell culture suggesting that mitochondrial transmembrane potential was considerably disrupted in these cells (Figure 4E and F). The cleavage of the nuclear enzyme PARP was examined in HT1376‐CisR cells and SCIN‐knockdown HT1376‐CisR cells following cisplatin exposure by western blotting, with HT1376 cells being used as a positive control. PARP cleavage was only faintly observed in HT1376‐CisR cells following cisplatin exposure; however, it was prominently noted in the SCIN‐knockdown HT1376‐CisR cells (Figure 4G).
To investigate the function of SCIN in mitochondria, we assessed the behavior of VDAC1 and VDAC2 proteins located on the mitochondrial outer membrane. The results from western blotting analyses demonstrated that VDAC1 and VDAC2 proteins existed mainly in the mitochondrial fraction; further, the levels of both proteins in HT1376‐CisR cells were slightly—but not significantly—higher than those in HT1376 cells (Figure 5A). To examine the interactions between SCIN and VDAC expression, we performed immunoprecipitation using mitochondrial fractions from HT1376 and HT1376‐CisR cells. Mitochondrial extracts previously immunoprecipitated with VDAC1 or VDAC2 were reacted with anti‐SCIN antibodies. Our results suggested that VDAC1 and VDAC2 were bound to SCIN in the mitochondrial fraction particularly in HT1376‐CisR cells (Figure 5B).
Interaction between SCIN and voltage‐dependent anion channel (VDAC) protein in HT1376‐CisR cells. A) The expressions of VDAC1, VDAC2, and COXIV were detected in the mitochondrial fraction by western blot analysis using the corresponding ...
Our present study has demonstrated for the first time the participation of the SCIN‐VDAC interaction in the mechanisms underlying cisplatin resistance in bladder cancer cells. Although cisplatin is widely used for the treatment of advanced and metastatic bladder cancer, a majority of the patients relapse with cisplatin‐resistant disease during chemotherapy. Bladder cancer generally responds well to combination chemotherapy with cisplatin and gemcitabine. Unfortunately, the initial response rate of up to 50% is not sustained, eventually resulting in a 5‐year patient survival rate of only 15%; this is primarily due to chemoresistance of tumor cells (Roberts et al., 2006; von der Maase et al., 2000). The acquisition of chemoresistance thus remains a major obstacle in the treatment of bladder and other cancers. The mechanisms of cisplatin resistance have been previously investigated, and potential mechanisms include reduced intracellular drug accumulation, increased detoxification of drug by thiol‐containing molecules, increased DNA damage repair, escape from reactive oxygen species‐mediated cytotoxicity, and involvement of apoptosis mediators (Hour et al., 2010; Siddik, 2003; Tsunoda et al., 2005). In general, chemoresistance is multifactorial, i.e., several mechanisms are simultaneously encountered within the same tumor cell (Eastman and Schulte, 1988; Rabik and Dolan, 2007; Richon et al., 1987; Teicher et al., 1987).
To clarify the possible mechanisms underlying cisplatin resistance in bladder cancer cells, we established a cisplatin‐resistant cell line, HT1376‐CisR, by exposing human bladder cancer HT1376 cells to a clinically relevant concentration of cisplatin (5 μM); the resulting cells displayed a 21‐fold increase in cisplatin resistance as compared to the parental cells. Then, we applied a quantitative proteomics approach to identify the molecules that underwent alterations following acquisition of cisplatin resistance. This approach has certain benefits over the cDNA microarray method that captures mRNA levels, since mRNA has the potential for high turnover with no protein expression; further, proteins may undergo a wide variety of post‐translational modifications that affect protein stability, localization, and function (Hodgkinson et al., 2010).
Here, we identified a total 36 proteins that displayed significant differences in the abundance levels between HT1376‐CisR and HT1376 cells. In this analysis, galectin‐3 was expressed at significantly higher levels in HT1376‐CisR cells. Recent studies have revealed that galectin‐3 demonstrates anti‐apoptotic effects that contribute to cell survival in several types of cancer cells (Fukumori et al., 2007). Intracellular galectin‐3 is also known to inhibit cell apoptosis induced by chemotherapeutic agents such as cisplatin and etoposide (Fukumori et al., 2006). Therefore, our result did not contradict the acquisition of chemoresistance. Out of these 36 candidate proteins, we assessed SCIN in detail for the following reasons. (a) The expression levels of SCIN were significantly increased in the HT1376‐CisR cells compared with the parental cells. (b) SCIN, which is a member of the calcium‐regulated villin family of actin‐severing and actin‐capping proteins, comprises 6 homologous domains that share 60% identity with gelsolin (Chumnarnsilpa et al., 2009). Gelsolin is known to enhance or inhibit apoptosis depending on the nature of pathological conditions (Kusano et al., 2000).
As cisplatin is a highly potent inducer of apoptosis, cisplatin resistance implies that tumor cells fail to undergo apoptosis. SCIN silencing by siRNA transfection induced apoptosis in HT1376‐CisR cells cultured with cisplatin. Thus, this is the first report to demonstrate that SCIN expression resulted in cisplatin resistance in bladder tumor cells.
Furthermore, SCIN expression was abundant in the mitochondrial fraction of HT1376‐CisR cells. Because, in general, SCIN is located mainly in the cytoplasm, we considered that the increase in SCIN expression in the cytoplasm and its translocation to the mitochondria may be involved in cisplatin resistance.
VDAC1 and VDAC2 are located on the mitochondrial outer membrane and control the entry and exit of mitochondrial metabolites. VDACs have been suggested to function in mitochondria‐mediated apoptosis and as critical players in the release of apoptogenic proteins, such as cytochrome c, triggering caspase activation and apoptosis. Castagna et al. have shown that exposure of cisplatin to a cervical squamous cell carcinoma cell line induced the upregulation of VDAC1 (Castagna et al., 2004). In addition, Tajeddine et al. found that VDAC1 silencing by siRNA efficiently prevented cisplatin‐induced apoptosis and Bax activation in non‐small cell lung cancer cells (Tajeddine et al., 2008). These results indicate that VDAC is an essential protein for cisplatin‐induced apoptosis. However, until recently, how apoptotic initiators cross the mitochondrial outer membrane has remained unclear. Zalk et al. (2005) demonstrated that VDAC exists in a dynamic equilibrium between monomers and tetramers and that the oligomeric status of VDAC offers a route for such transfers. More recently, Keinan et al. (2010) suggested that the overexpression of VDAC1 resulted not only in VDAC oligomerization but also in apoptosis.
Based on several lines of experimental evidence, it has been shown that VDAC is the target of pro‐ and anti‐apoptotic members of the Bcl2 family or hexokinases (HKs) (Arbel and Shoshan‐Barmatz, 2010; Arzoine et al., 2009; Azoulay‐Zohar et al., 2004; Shi et al., 2003; Shoshan‐Barmatz et al., 2010; Tsujimoto and Shimizu, 2000). HKs bind to VDAC on the mitochondrial surface, with VDAC providing both metabolic benefits and apoptosis‐suppressive capacity, subsequently increasing cellular resistance to chemotherapy (Mathupala et al., 2009). Furthermore, the binding of apoptosis‐regulating proteins to the N‐terminal region of VDAC might hamper its motility, thereby inhibiting cytochrome c release (Abu‐Hamad et al., 2009; De Pinto et al., 2007). In addition, Kusano et al. (2000) have shown that the C‐terminal fragment of human gelsolin is capable of preventing mitochondria‐induced apoptotic changes by directly binding to VDAC. In the present study, we demonstrated that (a) SCIN was overexpressed in cisplatin‐resistant cells; (b) SCIN coexisted with VDACs in the mitochondrial fraction; and (c) SCIN‐knockdown resulted in apoptosis with disrupted mitochondrial transmembranes. We expect that in cisplatin‐sensitive conditions, VDAC exists in the monomeric state on the mitochondrial outer membrane with apoptotic signals such as cisplatin exposure inducing VDAC oligomerization on the membrane followed by the release apoptotic proteins via the VDAC oligomers (as shown in Figure 6). However, in cisplatin‐resistant conditions, the overexpressed SCIN possibly binds to VDAC oligomers on the surface, thereby preventing mitochondria‐induced apoptotic changes.
Schematic diagram of potential pathways and roles of SCIN in cisplatin‐resistance. A) Before apoptosis induction, VDAC1 lies in the monomeric state on the mitochondrial outer membrane. VDAC1 forms the oligomer with the apoptosis signal; subsequently, ...
To our knowledge, this is the first report to demonstrate the interaction between VDAC and SCIN, particularly in HT1376‐CisR cells that overexpressed SCIN; this binding possibly contributed to cisplatin resistance via the inhibition of mitochondria‐mediated apoptosis. Based on our results, we consider that the suppression of the SCIN‐VDAC interaction may offer a new therapeutic target for the treatment of cisplatin‐resistant bladder cancer.
Grant Support: This research was supported in part by Grants‐in‐Aid for scientific research from the Ministry of Education, Culture, Sports, Science and Technology of Japan (Program for Enhancing Systematic Education in Graduate School).
The authors thank Ms Ayako Takemori, Dr. Tomohisa Sakaue, Ms Kazumi Kanno, Ms Izumi Tanimoto, and Ms Maria Mori for their excellent technical assistance.
Miura Noriyoshi, Takemori Nobuaki, Kikugawa Tadahiko, Tanji Nozomu, Higashiyama Shigeki, Yokoyama Masayoshi, (2012), Adseverin: A novel cisplatin‐resistant marker in the human bladder cancer cell line HT1376 identified by quantitative proteomic analysis, Molecular Oncology, 6, doi: 10.1016/j.molonc.2011.12.002.