2022 Volume 45 Issue 6 Pages 780-792
Muscle atrophy is commonly observed during cisplatin chemotherapy, leading to a reduced QOL in cancer patients. Reduced skeletal muscle mass caused by cisplatin treatment results from the activation of ubiquitin ligases–Atrogin-1 and MuRF1, but the precise mechanisms are poorly understood. In this study, we investigated the possible involvement of mitochondrial dysfunction, including reactive oxygen species (ROS) generation and ATP production, in cisplatin-induced muscle atrophy. Skeletal C2C12 myotubes were treated with cisplatin, and gene and protein expression were evaluated. Mitochondrial mass, membrane potential, and ROS levels were measured using fluorescent dyes. Mitochondrial respiratory function, ATP production rates, and glycolytic capacity were also analyzed using an extracellular flux analyzer. Metabolomic analyses were performed using gas chromatography-tandem mass spectrometry. Cisplatin treatment reduced myosin heavy chain expression by activating the ubiquitin-proteasome system. Increased ROS production was observed after cisplatin treatment, followed by significant changes in apoptosis-related gene expression and decrease in mitochondrial mass, membrane potential, respiration, and ATP production. Glycolytic capacity and tricarboxylic acid (TCA) cycle metabolite levels were reduced with cisplatin treatment. Mitochondria-targeted antioxidant mitoquinone mesylate prevented up-regulation of Atrogin-1 gene expression and restored myosin heavy chain levels, accompanied by a decrease in ROS generation, but not mitochondrial ATP production. We concluded that cisplatin-induced myotube atrophy was associated with mitochondrial dysfunction. Reducing ROS generation, rather than promoting ATP production, could be a useful therapeutic strategy for preventing cisplatin-induced muscle atrophy.
Muscle wasting is commonly observed in cancer cachexia and chemotherapy. Sarcopenia associated with chemotherapy results in poor QOL and is a poor prognostic factor for the overall survival of patients with cancer.1) Cisplatin is an anticancer drug that is clinically used for the treatment of a variety of cancers, such as gastrointestinal, lung, head, and neck, testicular, bladder, ovarian, and cervical cancers. The anticancer mechanism of cisplatin primary involves intra-strand crosslinking of DNA via binding to the guanine-N-7 position of DNA, resulting in the inhibition of DNA synthesis, replication, and translation; the inhibition of cell division and the induction of apoptosis in tumor cells is also reported.2–4) Although it is commonly accepted that the fundamental anticancer mechanism of cisplatin involves DNA damage, recent investigations have revealed several alternative mechanisms, including effects on cell metabolism, immunomodulation, and a wide variety of intracellular proteins and organelles; interference with the cellular communication between tumor cells and their microenvironment is also observed.5) Cisplatin chemotherapy can have serious adverse effects, such as nephrotoxicity, ototoxicity, and neurotoxicity.6,7) Recently, there has also been much attention focusing on the occurrence of skeletal muscle damage caused by chemotherapeutic agents.8–10) It has become evident that cisplatin chemotherapy frequently causes secondary sarcopenia, which negatively affects the QOL of patients.11,12)
Muscle atrophy involves a diverse set of signaling pathways that regulate protein degradation and synthesis. Cisplatin reportedly decreases skeletal muscle mass by activating the muscle-specific ubiquitin ligases, Atrogin-1 and MuRF1, which are key regulators of muscle atrophy referred to as “atrogenes.”8,10,13–16) The activation of these ubiquitin ligases is regulated by the Forkhead box-O (FOXO) family of transcriptional factors in many cases, and accumulating evidence indicates that FOXO1 and/or FOXO3a are involved in cisplatin-induced muscle atrophy.13,16–18) However, the molecular mechanism underlying the activation of the atrogenes by cisplatin remains to be elucidated.
Mitochondria play an important role in regulating many cellular functions, including ATP production, reactive oxygen species (ROS) generation, and apoptosis induction.19) Decline in these functions has been shown to be involved in age-related sarcopenia20–22) and disuse muscle atrophy.23,24) In a recent animal study, Sirago et al. reported that mitochondrial disorder occurs during cisplatin treatment.25) However, the group did not reveal any direct evidence of mitochondrial dysfunction contributing to the activation of MuRF1 and Atrogin-1. Furthermore, the causal relationship between mitochondrial dysfunction and muscle atrophy has not yet been investigated.
In the current study, we aimed to elucidate the effects of cisplatin on mitochondrial functions, i.e., ATP production, cellular energy metabolism, ROS generation, and regulation of apoptosis in C2C12 myotubes. Further, the efficacy of the mitochondria-targeted antioxidant mitoquinone mesylate (MitoQ)26–29) was examined to investigate the mechanism underlying mitochondrial disorder in cisplatin-induced myotube atrophy.
Mouse C2C12 myoblasts were purchased from KAC Co., Ltd. (Kyoto, Japan). The C2C12 myoblasts were cultured in Dulbecco’s modified Eagle’s medium (DMEM; Thermo Fisher Scientific, Waltham, MA, U.S.A.) supplemented with 10% fetal bovine serum (JRH Biosciences, Inc., Lenexa, KS, U.S.A.) and penicillin–streptomycin (Thermo Fisher Scientific) incubated at 37 °C with 5% CO2. To differentiate confluent C2C12 myoblasts into myotubes, the culture medium was replaced with differentiation medium consisting of DMEM +2% horse serum (Thermo Fisher Scientific) and incubated for 5 d. During differentiation induction, the differentiation medium of the myoblasts was replaced with fresh medium approximately once every two days. After differentiation, the myotubes were treated with cisplatin (479306; Sigma-Aldrich, St. Louis, MO, U.S.A.) or vehicle (0.02% dimethyl sulfoxide) and subsequently subjected to analysis. Unless otherwise stated, the C2C12 myotubes were incubated with 50 µmol/L cisplatin for 24 h. Mitoquinone mesylate (MitoQ; HY-100116 A) was purchased from MedChemExpress (Monmouth Junction, NJ, U.S.A.). MitoQ (0.16, 0.4 µmol/L) was added 30 min before the addition of cisplatin.
Live/Dead Cell AssaysLive and dead cell rates were determined using a Live/Dead Cell Staining Kit II (PK-CA707-30002; PromoCell, Heidelberg, Germany). C2C12 myotubes (1.5 × 104 cells) were treated with 6.25–50 µmol/L cisplatin for 24 h. The cells were then incubated for 30 min at room temperature in a 2 µmol/L Calcein-AM/4 µmol/L EthD-III staining solution, according to the manufacturer’s protocol provided with the kit. Calcein-AM stains viable cells green (Ex/Em = 485/525 nm) while EthD-III stains dead cells red (Ex/Em = 544/612 nm). Fluorescence was measured using a Flex station 3 multimode microplate reader (Molecular Devices, San Jose, CA, U.S.A.).
MTS AssaysCell viability was determined using MTS reagent provided with the CellTiter 96 AQueous One Solution Cell Proliferation Assay kit (G3580; Promega Corporation, Madison, WI, U.S.A.). C2C12 myotubes (5 × 103 cells) were treated with 6.25–50 µmol/L cisplatin for 24 h. MTS reagent was then added according to the manufacturer’s protocol. The absorbance of 490 nm was measured using an Infinite 200 PRO microplate reader (Tecan, Grödig, Austria).
Caspase 3/7 AssaysC2C12 myotubes (1.5 × 104 cells) were treated with 12.5–100 µmol/L cisplatin for 24 h. Caspase 3/7 activity was determined using an Amplite™ Fluorimetric Caspase 3/7 Assay Kit (13502; AAT Bioquest, Sunnyvale, CA, U.S.A.) according to the manufacturer’s protocol. Fluorescence was measured at Ex/Em = 350/450 nm using the Infinite 200 PRO microplate reader (Tecan).
Immunofluorescence StainingC2C12 myotubes (7 × 104 cells) were treated with 12.5–50 µmol/L cisplatin for 24 h and then fixed with 4% paraformaldehyde-phosphate buffer for 30 min at room temperature. After washing three times with phosphate-buffered saline, the cells were incubated with 10 µg/mL mouse anti-human myosin heavy chain (MyHC) monoclonal antibody (MAB4470; R&D Systems, Inc., Minneapolis, MN, U.S.A.) as primary antibody for 3 h at room temperature. The fluorescently labeled secondary antibody Alexa Fluor 488 anti-mouse immunoglobulin G (IgG) (A-11029; Thermo Fisher Scientific) at a concentration of 2 µg/mL was added and incubated for 1 h at room temperature. Images of MyHC staining were acquired using a BZ-9000 HS all-in-one fluorescence microscope (Keyence Corporation, Osaka, Japan). Twenty-four myotubes were randomly selected from each well of each treatment group and their short axis lengths were measured using ImageJ (https://imagej.nih.gov/ij/), and the mean value per well calculated.
Western Blot AnalysisProteins were isolated from C2C12 myotubes (7 × 104 cells) after 0.5, 2, 6, or 24 h of 50 µmol/L cisplatin treatment using Cell Lysis Buffer (10×) (9803; Cell Signaling Technology, Danvers, MA, U.S.A.), which contains protease inhibitors and serine phosphatase inhibitors. The expression of MyHC was assessed using the Jess Simple Western System (ProteinSimple, San Jose, CA, U.S.A.), which is an automated capillary-based size separation and nano-immunoassay system. The analysis was performed according to the manufacturer’s standard method unless otherwise noted. Briefly, an equal volume of cell lysate (approximately 900 µg/mL) was mixed with 0.1× sample buffer and Fluorescent 5× Master mix (ProteinSimple) at a ratio of 8 : 12 : 5 and then denatured at 95 °C for 5 min. The isolated proteins were separated using a 12–230-kDa Jess Separation Module (SM-W004; ProteinSimple), probed with 5 µg/mL anti-MyHC antibody (MAB4470; R&D Systems) for 30 min. Bound MyHC was detected using the Anti-Mouse Detection Module (DM-002; ProteinSimple). Primary and secondary antibodies were then removed using RePlex™ Module (RP-001; ProteinSimple) and the blots re-probed using 10 µg/mL of anti-glyceraldehyde-3-phosphate dehydrogenase (GAPDH) antibody (AF5718; Novus Biologicals, LLC, Centennial, CO, U.S.A.) for 30 min. Bound anti-GAPDH antibody was detected using Anti-Goat Detection Module (DM-006; ProteinSimple). The peak area was automatically calculated using Compass Simple Western software, version 5.0.1 (ProteinSimple) but baseline was altered in manual settings and the expression of MyHC was normalized to that of GAPDH.
RNA Extraction and Quantitative RT-PCRC2C12 myotube cells (7 × 104 cells) were treated with 6.25–50 µmol/L cisplatin for 2, 6, or 24 h and then lysed by adding a mixture of Buffer RLT and 2-mercaptoethanol provided in an RNeasy Mini Kit (#74106; QIAGEN, Valencia, CA, U.S.A.). The cell lysate was mixed by vortexing and homogenized using a QIAshredder spin column (79656; QIAGEN). RNA extraction was performed using the RNeasy Mini Kit. The RNA extracted from the C2C12 myotubes were diluted to approximately 110 ng/µL, incubated at 70 °C for 15 min, and then cooled to 4 °C. The RNA was then reverse transcribed into cDNA using a high-capacity cDNA Reverse Transcription Kit (4368813; Thermo Fisher Scientific). Real-time quantitative PCR assays were performed using TaqMan Fast Advanced Master Mix (4444557; Thermo Fisher Scientific) and gene-specific TaqMan primer/probes (Thermo Fisher Scientific) and a QuantStudio 7 Flex Real-Time PCR System (Thermo Fisher Scientific). The primer and probe information are provided in Supplementary Table 1. Analysis of 18s ribosomal RNA (rRNA) was used as an endogenous control and mRNA expression levels were calculated using the ΔΔCt method.
Mitochondrial Mass Measurements and Membrane Potential AssaysC2C12 myotubes (1.5 × 104 cells) were treated with 50 µmol/L cisplatin for 6 or 24 h, as indicated. Amounts of mitochondria were assessed using MitoGreen (PK-CA707-70054; PromoCell), which is a green fluorescent (Ex/Em = 485/525 nm), mitochondria-staining dye. Briefly, 50 nmol/L MitoGreen was added to the cisplatin-treated cells and incubated at 37 °C for 30 min. Mitochondrial membrane potential was measured using Image-iT TMRM Reagent (I34361; Thermo Fisher Scientific). TMRM is a cell-permeable fluorescent dye (Ex/Em = 544/590 nm) that accumulates in active mitochondria without any abnormal membrane potential. The cells were incubated with 100 nmol/L TMRM for 30 min at 37 °C. For both assays, fluorescence intensity was measured using a Flex station 3 multimode microplate reader (Molecular Devices).
Assessment of ROS LevelsC2C12 myotubes (1.5 × 104 cells) were treated with 50 µmol/L cisplatin for 0.5, 2, 6, or 24 h. Mitochondrial superoxide levels were assessed after cisplatin exposure by incubating the cells with 1 µmol/L MitoSOX (M36008; Thermo Fisher Scientific) for 10 min at 37 °C. After incubation with the fluorescent probe, the fluorescence intensity was measured at Ex/Em = 510/580 nm using an Infinite 200 PRO microplate reader (Tecan). Intracellular ROS production was measured using 5-(and-6)-chloromethyl-2′,7′-dichlorodihydrofluorescein diacetate, acetyl ester (CM-H2DCFDA) (C6827; Thermo Fisher Scientific). Briefly, 5 µmol/L CM-H2DCFDA was added to the cells after exposure to cisplatin and then incubated at 37 °C for 30 min. The fluorescence intensity was measured at Ex/Em = 485/525 nm using a Flex station 3 multimode microplate reader (Molecular Devices).
ATP AssaysIntracellular ATP production was assessed using a Luminometric ATP Assay Kit (IC2 100; TOYO B-Net Co., Tokyo, Japan). Briefly, C2C12 myotubes (1.5 × 104 cells) were treated with 50 µmol/L cisplatin for 2, 6, or 24 h and then subjected to the luminometric assay according to the manufacturer’s protocol. Luminescence intensity was measured using an Infinite 200 PRO microplate reader (Tecan).
Measurement of XFp Extracellular FluxThe oxygen consumption rate (OCR) and extracellular acidification rate (ECAR) of C2C12 myotubes were measured using an XFp Extracellular Flux Analyzer (Agilent Technologies, Santa Clara, CA, U.S.A.). Mitochondrial function was evaluated using a Seahorse XFp Mito Stress Test Kit (103010-100; Agilent Technologies). A Seahorse XFp Glycolysis Stress Test Kit (103017-100; Agilent Technologies) was used to evaluate glycolysis. The rate of ATP production from mitochondrial respiration and from glycolysis were measured using a Seahorse XFp Real-Time ATP Rate Assay Kit (103591-100; Agilent Technologies). The sensor cartridges were hydrated in water the day before the experiment and incubated overnight at 37 °C in a CO2-free incubator. Two hours prior to the experiment, the water was replaced with calibration solution and placed back in the CO2-free incubator at 37 °C. C2C12 myoblasts were seeded into XFp miniplates at a density of 1.2 × 104 cells and differentiated into myotubes as described above. After treatment with 50 µmol/L cisplatin for 24 h, the cells were subjected to the subsequent assays described below. The data obtained were corrected for protein content, which was determined using a Pierce BCA Protein Assay (23225; Thermo Fisher Scientific).
i) Mitochondria EvaluationEvaluation of the mitochondria was performed using the Seahorse XFp Mito Stress Kit. This is a reagent kit that can be used to evaluate mitochondrial function. On the day of the experiment, 1 mmol/L pyruvate, 2 mmol/L glutamine, and 10 mmol/L glucose were added to Seahorse XFp DMEM medium, pH 7.4 (103575-100; Agilent Technologies) to prepare the assay medium. The C2C12 myotube cell culture medium was replaced with the assay medium and the cells placed at 37 °C in a CO2-free incubator for 1 h. For measuring OCR, ATP synthase inhibitor oligomycin, deconjugate carbonyl cyanide-4 (trifluoromethoxy) phenylhydrazone (FCCP), and a mixture of mitochondrial complex I inhibitor rotenone/mitochondrial complex III inhibitor antimycin A were added according to the manufacturer’s instructions and protocol.
ii) Glycolysis EvaluationThe Seahorse XFp Glycolysis Stress Test Kit was used to evaluate glycolysis. This is a reagent kit that can be used to evaluate the glycolytic capacity according to sugar uptake into cells. On the day of the experiment, 2 mmol/L glutamine was added to Seahorse XFp DMEM medium to prepare the assay medium. The C2C12 myotube culture medium was replaced with the assay medium and the cells placed at 37 °C in a CO2-free incubator for 1 h. For the measurement of ECAR, glucose, oligomycin, and the glycolysis inhibitor 2-deoxy-D-glucose were added according to the manufacturer’s instructions and protocol.
iii) Evaluation of ATP Production Rate from Mitochondrial Respiration and from GlycolysisThe ATP production rate of the mitochondria and glycolysis were measured using the Seahorse XFp Real-Time ATP Rate Assay Kit. This kit can be used to quantify the rate of ATP production from mitochondrial respiration and glycolysis in living cells in real time. On the day of the experiment, 1 mmol/L pyruvate, 2 mmol/L glutamine, and 10 mmol/L glucose were added to Seahorse XFp DMEM medium to prepare the assay medium. The C2C12 myotube culture medium was replaced with the assay medium and the cells placed at 37 °C in a CO2-free incubator for 1 h. To measure the rate of ATP production from mitochondrial respiration and glycolysis, oligomycin and rotenone/antimycin A mixture were added according to the manufacturer’s instructions and protocol and the cultures were then monitored for OCR and ECAR. The OCR based on mitochondrial respiration, ECAR based on glycolysis, the mitoATP production rate, glycoATP production rate, and total ATP production rate were calculated using Wave software (Agilent Technologies).
Metabolome AnalysisThe metabolome of C2C12 myotubes was evaluated using an SGI-M100 automated derivatization system (AiSTI SCIENCE, Wakayama, Japan) and gas chromatography–tandem mass spectrometry (GC-MS/MS). One milliliter of 80% acetonitrile containing 5 µg/mL 2-isopropylmalic acid, which was used as an internal standard, was added to C2C12 cells (7 × 104 cells). After 10 min of incubation on ice, the extract was collected and centrifuged. The supernatant was transferred to a vial and subjected to metabolome analysis. The analysis was performed based on a previously described method.30,31) The extracted samples were loaded onto an ion-exchange Solid Phase Extraction (SPE) cartridge and the target metabolites were retained in the SPE. Derivatization was performed using methoxyamine/pyridine and N-methyl-N-(trimethylsilyl) trifluoroacetamide, which were sequentially added directly to the SPE. The derivatized samples were then subjected to GC-MS/MS analysis, which was performed using a GCMS-TQ8040 system (Shimadzu, Kyoto, Japan) and a 30 m ×0.25 µm BPX5 fused silica capillary column with a film thickness of 0.25 µm (SGE, Melbourne, Australia). The analytical conditions, chromatogram acquisition, and waveform processing were previously summarized by Kitagawa et al.30) The peak intensity of each quantified ion was calculated and normalized to that of 2-isopropylmalic acid. Further analyses were performed using the normalized values. Clustering analysis of the metabolome data was conducted using MetaboAnalyst (https://www.metaboanalyst.ca)32) with Ward’s linkage being used as the clustering algorithm.
Data and Statistical AnalysisStatistical analysis was performed using GraphPad Prism software, version 7 (GraphPad Software Inc., San Diego, CA, U.S.A.). Data were expressed as mean ± standard deviation (S.D.). Statistical analysis between two groups was performed using the Student’s t-test. Differences in the means of multiple groups were assessed using Dunnett’s test or two-way ANOVA, followed by Tukey’s test. The level for statistical significance was set at p < 0.05.
We evaluated the effects of cisplatin on C2C12 myotube survival and atrophy. Treatment with 6.25–50 µmol/L cisplatin for 24 h resulted in a 15% decrease in calcein AM-positive live cells in the group treated with the highest dose of cisplatin (50 µmol/L) compared to that in the vehicle-treated group (Fig. 1A); however, there was no increase in the number of EthD-III-positive dead cells (Fig. 1B). Cell viability was evaluated using the MTS-based metabolic viability assays findings, which is proportional to mitochondrial dehydrogenase activity. There was a 16% reduction in metabolic viability after treatment with 50 µmol/L cisplatin (Fig. 1C). Meanwhile, caspase 3/7 activity was unaffected by a 24 h treatment with cisplatin at concentrations of 50 µmol/L or less (Fig. 1D). Treatment with 50 µmol/L cisplatin significantly decreased the short axis length of C2C12 myotubes to 72.6% compared to that of vehicle-treated C2C12 myotubes (p < 0.0001; Fig. 1E) and reduced the levels of MyHC protein (p < 0.05; Fig. 1F). In cell viability assays, slight decreases in calcein AM-positive cells and MTS signal after cisplatin treatment were observed; however, they may be attributed to cisplatin-induced reduction in cellular volumes and mitochondrial function in the C2C12 myotubes. Therefore, the findings suggest that muscular atrophy can be induced without adversely affecting cell viability.
(A, B) Cell viability rates (A: Calcein-AM-positive cells) and cell death rates (B: EthD-III-positive cells) of C2C12 myotubes were measured 24 h after cisplatin treatment using a fluorescent live/dead assay (n = 6). (C) Viability of C2C12 myotubes was assessed using MTS (n = 3). (D) Caspase 3/7 activity was measured 24 h after C2C12 myotubes were treated with 12.5–100 µmol/L cisplatin using a Fluorimetric Caspase 3/7 Assay Kit (n = 6). (E) Immunofluorescent staining of MyHC in cisplatin-treated myotubes. Bar = 100 µm. The graph shows the comparison of the short axis length between vehicle-treated and cisplatin-treated C2C12 myotubes (Twenty-four myotubes were randomly selected from each well of each treatment group and their short axis lengths were measured. n = 3). (F) Expression of MyHC protein in C2C12 myotubes assessed using an automated western immunoblotting system (n = 4). The values are expressed as the mean ± S.D. * p < 0.05, ** p < 0.01, *** p < 0.001, and **** p < 0.0001 vs. the vehicle group.
Expression levels of mRNAs for Atrogin-1 and MuRF1, two muscle-specific ubiquitin ligases that mediate muscle protein degradation and muscle atrophy, were significantly elevated after 24 h treatment with 25 and 50 µmol/L cisplatin (Figs. 2A, B). In addition, Foxo3a and Foxo4 mRNA levels, but not Foxo1 mRNA levels, were increased remarkably after 24 h cisplatin treatment (Fig. 2C). These results suggest that muscle atrophy occurs via transcription by FOXO signaling.
(A, B) mRNA expression levels of the ubiquitin ligases Atrogin-1 and MuRF1 in C2C12 myotubes treated with 6.25–50 µmol/L cisplatin for 2, 6, or 24 h (n = 3–4). (C) mRNA expression of the forkhead family of transcription factors Foxo1, Foxo3a, and Foxo4 in C2C12 myotubes after cisplatin treatment (n = 3). The values are expressed as the mean ± S.D. * p < 0.05, ** p < 0.01, *** p < 0.001, and **** p < 0.0001 vs. the vehicle group.
Both the mitochondrial mass and membrane potential were decreased in cells treated with cisplatin for 24 h, but these parameters were not altered after 6 h of cisplatin treatment (Figs. 3A, B). Meanwhile, treatment with 6.25–50 µmol/L cisplatin for 6 or 24 h increased the expression levels of the mitochondrial pro-apoptotic regulators B-cell lymphoma 2 (Bcl-2)-associated X protein (Bax), and decreased Bcl-2 expression levels in a concentration- and time-dependent manner (Fig. 3C). We also observed increased gene expression levels of peroxisome proliferator-activated receptor, gamma coactivator-1α (Pgc-1α), mitochondrial transcription factor A (Tfam), and mitochondrially localized superoxide dismutase 2 (Sod2), which possibly suggest a compensatory mechanism for the pro-apoptotic signaling induced by cisplatin treatment (Fig. 3D, Supplementary Figs. 1A–C).
C2C12 myotubes were treated with vehicle or 6.25–50 µmol/L cisplatin for 2, 6, or 24 h. (A, B) Mitochondrial mass (A) and mitochondrial membrane potential (B) were measured using MitoGreen and TMRM fluorescent dyes, (n = 6–8). (C) Evaluation of mRNA expression of mitochondrial apoptotic regulators Bax and Bcl-2 in cisplatin-treated C2C12 myotubes (n = 3–4). (D) Evaluation of mRNA expression of mitochondria-related genes Pgc-1α, Tfam, and Sod2 in cisplatin-treated C2C12 myotubes (n = 4). The values are presented as the mean ± S.D. * p < 0.05, ** p < 0.01, *** p < 0.001, and **** p < 0.0001 vs. the vehicle group.
We then investigated the association between mitochondrial ROS generation and muscle atrophy by measuring mitochondrial superoxide levels using MitoSOX and intracellular ROS levels using CM-H2DCFDA. Mitochondrial superoxide levels in C2C12 cells were temporarily increased 1.3-fold after 2 h of cisplatin treatment, but had returned to basal values at 6 h and 24 h (Fig. 4A). Increased intracellular ROS levels were detected after 2 h of cisplatin treatment and remained elevated until 24 h post cisplatin treatment (Fig. 4B). Meanwhile, intracellular ATP levels increased after 6 h of cisplatin treatment, but decreased significantly at 24 h post cisplatin treatment compared to that of vehicle-treated cells (p < 0.001; Fig. 4C).
(A) Mitochondrial superoxide production was measured using the MitoSOX fluorescent probe after C2C12 myotubes were treated with vehicle or 50 µmol/L cisplatin for 0.5, 2, 6, and 24 h (n = 5–6). (B) Intracellular ROS production was measured using oxidized CM-H2DCFDA after C2C12 myotubes were treated with vehicle or 50 µmol/L cisplatin for 0.5, 2, 6, and 24 h (n = 6). (C) Intracellular ATP level was assessed using a Luminometric ATP assay (n = 6). The values are expressed as the mean ± S.D. * p < 0.05, ** p < 0.01, and *** p < 0.001 vs. the vehicle group.
The OCR was measured as an indicator of mitochondrial oxidative phosphorylation (Fig. 5A(I)). Cisplatin treatment reduced the basal respiratory volume (p < 0.01; Fig. 5A(II)), the spare respiratory capacity of the electron-transport system (p < 0.05; Fig. 5A(III)) and ATP-producing capacity (p < 0.01; Fig. 5A(IV)), and limited proton leak (p < 0.05; Fig. 5A(V)). Meanwhile, the ECAR was subsequently measured as an indicator of glycolysis (Fig. 5B(I)). Cisplatin significantly reduced glycolysis (p < 0.05; Fig. 5B(II)) and the maximal glycolytic capacity (p < 0.05; Fig. 5B(III)). However, no change in the glycolytic reserve was observed (Fig. 5B(IV)). In the ATP rate assay, the mitochondrial ATP production rate, glycolytic ATP production rate, and total ATP production rate were reduced in the cisplatin-treated group compared to those in the vehicle-treated group by approximately 40, 50, and 40%, respectively (Figs. 5C(I)–(III)).
Real-time measurements of OCR and ECAR in C2C12 myotubes treated with vehicle or 50 µmol/L cisplatin for 24 h. Measurements were made using an Extracellular Flux Analyzer and were normalized to protein contents. (A) Evaluation of mitochondrial function. OCR was measured under basal conditions and after sequential addition of oligomycin, FCCP, and rotenone/antimycin A at the indicated time points. The data show time courses for OCR (I), changes in basal respiration (II), spare respiratory capacity (III), ATP production (IV), and proton leak (V) in the cisplatin-treated group of C2C12 myotubes compared with those of the vehicle-treated group. (B) Evaluation of the glycolysis. ECAR was measured under basal conditions and after sequential injection of glucose, oligomycin, and the glycolysis inhibitor 2-deoxy-D-glucose at the indicated time points. The data show time courses for ECAR (I), changes in glycolysis (II), glycolytic capacity (III), and glycolytic reserve (IV) in the cisplatin-treated group of C2C12 myotubes compared with those of the vehicle-treated group. (C) Mitochondrial and glycolytic ATP production rate. OCR and ECAR were measured under basal conditions and after sequential addition of oligomycin and rotenone/antimycin A at the indicated time points. Mitochondrial ATP (mitoATP) and glycolytic ATP (glycoATP) levels were calculated from the OCR and ECAR data, respectively, using a validated algorithm. The data show the mitochondrial ATP production rate (I), glycolytic ATP production rate (II), and total ATP production rate (III) in the cisplatin-treated group of C2C12 myotubes compared with those of the vehicle-treated group. The values are expressed as the mean ± S.D. (n = 3). * p < 0.05 and ** p < 0.01 vs. the vehicle group.
The altering effects of cisplatin on intracellular metabolites in C2C12 myotubes were analyzed using GC-MS/MS, where targeted metabolomics of approximately 200 water-soluble metabolites, including sugars, amino acids, organic acids, and free fatty acids, was performed. Sixty metabolites were identified as different in C2C12 myotubes after being treated with 50 µmol/L cisplatin compared with that after being treated with vehicle. These 60 metabolites were used for hierarchical clustering analysis and heat map construction (Fig. 6A). Cisplatin treatment reduced intermediate metabolite levels of the tricarboxylic acid (TCA) pathway, such as succinate, fumaric acid, and malic acid, and also reduced levels of amino acids such as alanine, glutamic acid, and aspartic acid (Figs. 6A, B). Glucose and mannose levels were elevated in the cisplatin-treated group compared to that in the vehicle-treated group, while the level of lactic acid, the end product of glycolysis, was decreased (Figs. 6A, B). These observations suggest that cisplatin may suppress not only the TCA cycle but also the glycolytic pathway in C2C12 myotubes.
(A) Hierarchical clustering analysis and heat map of metabolites in C2C12 myotubes treated with vehicle or 50 µmol/L cisplatin for 24 h (Red: upregulation, blue: downregulation). Sixty metabolites were detected by metabolome analysis in the C2C12 myotubes. (B) Schematic of the amino acid metabolic pathway in C2C12 myotubes. The mean ratio of amino acid levels in cisplatin-treated C2C12 myotubes compared with that in vehicle-treated C2C12 myotubes is shown according to the color scale (Red: upregulation, blue: downregulation). Undetected amino acids and key metabolites are indicated with small black circles. The graphs show the relative changes for each amino acid metabolite. The values are expressed as the mean ± S.D. (n = 8). * p < 0.05, ** p < 0.01, *** p < 0.001, and **** p < 0.0001 vs. the vehicle group.
Pre-treatment of C2C12 myotubes with MitoQ ameliorated cisplatin-induced decrease in mitochondrial content (Fig. 7A) and increase in intracellular ROS levels (Fig. 7B) after 24 h cisplatin treatment. However, intracellular ATP levels (Fig. 7C) were not restored by pre-treatment with MitoQ. Changes in Bax and Bcl-2 levels caused by cisplatin treatment were mitigated by MitoQ pre-treatment (Figs. 7D, E). Foxo4 mRNA levels were increased with cisplatin and decreased with MitoQ pre-treatment (Fig. 7F). Moreover, MitoQ inhibited the cisplatin-induced increase in Atrogin-1 expression (Fig. 7G) and increased the amount of MyHC protein (Fig. 7H). These results indicate that excessive ROS production might be mainly involved in cisplatin-induced myotube atrophy.
C2C12 myotubes were treated with MitoQ (0.16, 0.4 µmol/L) for 30 min followed by treatment with 50 µmol/L cisplatin for 24 h. (A) Mitochondrial mass was measured after cisplatin treatment using MitoGreen regents (n = 5). (B) Intracellular ROS production was determined by measuring the fluorescence intensity of oxidized CM-H2DCFDA (n = 5). (C) Intracellular ATP levels were assessed using a luminometric ATP assay (n = 4). (D, E) mRNA expression levels of the apoptotic regulators Bax and Bcl-2 (n = 4). (F, G) mRNA expression of Fluorometric Foxo4 and Atrogin-1, regulators related to muscle protein degradation (n = 4). (H) MyHC protein expression in C2C12 myotubes (n = 8). The values are expressed as the mean ± S.D. #### p < 0.0001 vs. the vehicle group. * p < 0.05, ** p < 0.01, *** p < 0.001, and **** p < 0.0001 vs. the MitoQ 0 group.
Our study revealed that treating C2C12 myotubes with 50 µmol/L cisplatin for 24 h induced cellular atrophy without causing apoptotic cell death, concomitant with a decrease in cellular ATP levels, inhibition of mitochondrial oxidative phosphorylation and glycolysis, and sustained increase in intracellular ROS levels. The mitochondria-targeted antioxidant MitoQ suppressed intracellular ROS production without restoring ATP levels, inhibited Atrogin-1 activation, and attenuated the reduction in myosin heavy chain levels. These results suggest that cisplatin-induced myotube atrophy was mediated primarily by intracellular ROS production rather than by metabolic disturbances, including decreased ATP levels, despite the correlation of these two pathologies to mitochondrial disturbances.
Previous clinical8,12,33,34) and laboratory studies15,16,35–37) have shown that cisplatin causes skeletal muscle atrophy, in which muscle proteolysis is enhanced owing to increased Atrogin-1 and/or MuRF1 levels. Atrogin-1 and MuRF1, originally discovered using the tail-suspension model, are skeletal muscle-specific E3 ubiquitin ligases that play key roles in muscle atrophy and ubiquitination, and assist in the degradation of myoblast determination protein 1 (MyoD) and muscle fiber constituent proteins, respectively.38,39) In the current study, the Atrogin-1 and MuRF1 activation after cisplatin treatment were observed in C2C12 myotubes similar to previous reports.13,36,40)
Mitochondria are fundamental for cellular energy production and are involved in muscle activity, development, and maintenance, as well as act as a crucial element in regulating skeletal muscle function. Mitochondrial impairment has been reported to be involved in age-related sarcopenia20–22) and disuse muscle atrophy.23,24) A recent study on cisplatin-induced cachexia in rats demonstrated that mitochondrial biosynthesis (PGC-1α, NRF-1, TFAM, mtDNA, and ND1), mitochondrial mass (porin and citrate synthase activity), and the fusion index are decreased in the skeletal muscle of cisplatin-treated rats25); however, the changes of fundamental mitochondrial functions such as ATP production, ROS generation, and apoptosis induction were not examined. Moreover, the causal relationship between mitochondrial dysfunction and muscle atrophy has not been proven. Thus, in the current study, we aimed to evaluate the changes in mitochondrial functions in C2C12 myotubes after cisplatin treatment and evaluate its association with myotube atrophy.
In this study, C2C12 myotubes treated with 50 µmol/L cisplatin for 24 h demonstrated cellular atrophy and decreased myosin levels without the induction of cell death. Subsequently, we also observed a decrease in the number of mitochondria and the mitochondrial membrane potential, along with a significant decrease in mitochondrial oxidative phosphorylation activity and ATP production, suggesting that mitochondrial energy production was disrupted.
Interestingly, this study proved that cisplatin treatment of C2C12 myotubes significantly reduced the glycolytic pathways and declined both the glycolytic capacity and glycolytic reserve. Metabolome analysis results also demonstrated a disruption of the glycolytic pathway by cisplatin. There is compelling evidence suggesting that glycolytic genes are suppressed by p53.41–44) Furthermore, recent studies have demonstrated that cisplatin activates p53 and then inhibits the glycolytic pathway in proximal tubular cells.45,46) These findings suggest that p53 might be involved in the cisplatin-induced reduction of glycolytic capacity in myotubes. Further studies are needed to confirm this possibility.
The regulation of intracellular ROS production is one of the critical functions in mitochondria.19) As it is widely known that oxidative stress commonly results from cisplatin cytotoxicity, and accumulation of ROS is observed in the mitochondria after cisplatin treatment,5,47,48) we focused on the association of mitochondrial impairment and ROS production in cisplatin-induced myotube atrophy. Here, a transient increase in mitochondrial ROS (superoxide anion) levels was observed immediately after 2 h treatment with cisplatin, followed by a sustained increase in intracellular ROS levels (mainly H2O2) for 24 h. These findings suggest that intracellular ROS played a crucial role in Atrogin-1 activation in cisplatin-treated C2C12 myotubes.
However, the present study did not reveal the target molecule involved in sustained ROS production following cisplatin treatment. The mitochondrial ROS production was transient, even though the increase in intracellular ROS was sustained 24 h after cisplatin treatment, suggesting the involvement of some ROS-producing enzymes, such as reduced nicotinamide adenine dinucleotide phosphate (NADPH) oxidase (NOX), other than mitochondria, in cisplatin-induced sustained ROS production. The Nox isozyme 2 (NOX2) reportedly contributes to doxorubicin-induced cardiac or gastrocnemius muscle atrophy in mice,49) and up-regulation of NOX4 in renal tissue is associated with cisplatin-induced nephrotoxicity.50,51) Currently, there is no evidence on the consequences of NOX in cisplatin-induced muscle atrophy. However, considering NOX1, 2, and 4 are expressed in C2C12 cells,52) NOXs could be target enzymes of cisplatin, and the NOX-derived ROS may be involved in mitochondrial impairment.
Our study revealed that relatively low levels, i.e., at 6.25–50 µmol/L of cisplatin exposure to C2C12 myotubes, failed to induce apoptotic cell death, despite a considerable increase in Bax/Bcl-2 ratio. The Bcl-2 family consists of anti- and pro-apoptotic proteins, and the increased Bax/Bcl-2 ratio corresponds with the mitochondrial outer membrane permeabilization, which is a prerequisite for inducing apoptosis.53) However, emerging evidence has suggested that Bcl-2 family proteins have other noncanonical functions, e.g., regulation of mitochondrial ATP production53) and mitochondrial ROS generation.54) Thus, increased Bax/Bcl-2 ratio might be related to decreased mitochondrial respiratory capacity and excessive ROS production in cisplatin-treated myotubes. Interestingly, the expression levels of Pgc-1a, Tfam, and Sod2 were increased after cisplatin treatment, suggesting that these regulatory mechanisms could help combat ROS overproduction and rescue from apoptotic cell death.
Previous reports have shown that FOXO3a17,18,36,55,56) or FOXO113,16) in muscle is activated with cisplatin treatment, indicating that the FOXO family plays a central role in activating MuRF1 and Atrogin-1. Although the FOXOs activities were not examined in this study, we found that the gene expression of Foxo3a and Foxo4, but not of Foxo1, increased with cisplatin treatment. FOXO3a activation upregulates the promotor activities of FOXO3a57) and FOXO458) but it does not affect FOXO1 in myotubes.59) Therefore, it is plausible to consider that FOXO3a was mainly activated in cisplatin-treated C2C12 myotubes in this study.
In the current study, cisplatin treatment resulted in elevated mitochondrial ROS (superoxide anion) and intracellular ROS (H2O2) levels in C2C12 myotubes. Intracellular ROS is known to promote nuclear translocation and transcriptional activity of FOXOs through the post-translational modifications by activating several kinds of protein kinases such as c-Jun N-terminal kinases (JNK) or mammalian sterile 20-like kinase 1(MST1) and by inhibiting Akt.60,61) FOXOs are also activated by depletion of intracellular ATP contents through the phosphorylation by the AMP-activated protein kinase (AMPK).62) Therefore, both elevated ROS levels and decreased intracellular ATP levels associated with mitochondrial impairment may contribute to the cisplatin-induced FOXO activation and the elevation of Atrogin-1/MuRF1 expression.
It has been reported that the induction of Atrogin-1 and MuRF1 are not necessarily observed with similar trends in muscle atrophy models; ROS (H2O2) stimulates up-regulation of Atrogin-1,63) while MuRF1 is mainly elevated under glucose starvation.64) In the present study, in cisplatin-treated myotubes, ROS (especially H2O2) production was important for the increase in atrogenes, which may be the main reason for the mild increase in MuRF1.
MitoQ is a mitochondria-targeted derivative of the antioxidant ubiquinone that can be localized in mitochondria and protect various cells from hydrogen peroxide-induced apoptosis.26–29) It has also been reported that MitoQ decreased intracellular ROS and mitochondrial ROS through in vitro studies.27,28) Moreover, MitoQ is used in animal studies and human trials for evaluating several diseases caused by mitochondrial oxidative damage.29) However, potential protective effects of MitoQ in cisplatin-treated C2C12 myotubes have not yet been determined.
In cisplatin-treated C2C12 myotubes, although Bcl-2 levels were still low, MitoQ attenuated the elevation of intracellular ROS levels and improved Bax/Bcl-2 ratio. It also reduced the gene expression levels of Foxo4 as well as Atrogin-1 and restored MyHC protein content.
Surprisingly, MitoQ failed to restore mitochondrial respiration and ATP production, but rather aggravated it. Previous reports have demonstrated that MitoQ caused biphasic effects on ROS production,27) and higher doses of MitoQ showed the pro-oxidant property.28) Moreover, it exhibited a non-specific effect on mitochondrial respiration,65–68) concluding that these unanticipated results might be due to the characteristic property of MitoQ.
As discussed previously, activation of FOXOs in cisplatin-treated C2C12 myotubes could be attributed to not only ROS overproduction but also energy disturbances. Our results regarding inhibition of expression of FOXO family by MitoQ and decreased ROS production without restoring ATP production strongly support the fact that ROS production has a major impact on cisplatin-induced myotube atrophy, and the mechanism of mitochondrial ATP reduction might have minimal effect.
Based on the overall results obtained in this study, we propose a mechanistic model of cisplatin-induced myotube atrophy (Fig. 8). Accordingly, cisplatin induces mitochondrial ROS production followed by an increase in Bax/Bcl-2 ratio, which further impairs mitochondrial integrity, disrupts its compensatory mechanism, and ultimately results in functional disturbances. Furthermore, the continuous ROS generation activates the FOXO signaling pathway and leads to the transcriptional activation of Atrogin-1/MuRF1.
Our study revealed that cisplatin generated excessive intracellular ROS, as well as reduced ATP production by mitochondrial oxidative phosphorylation and glycolysis, and induced myotube atrophy without apoptotic cell death. Further, the mitochondria-targeted antioxidant MitoQ decreased intracellular ROS generation and restored Atrogin-1 activation and decrease in myosin heavy chain protein content in cisplatin-treated C2C12 myotubes. Our results suggest that mitochondrial protection and/or ROS scavenging, but not ATP restoration, may be promising strategy for preventing muscle atrophy associated with cisplatin-based chemotherapy.
We are grateful to Prof. Masabumi Minami for his constructive comments and suggestions.
HT has been received Grant support from Tsumura & Co. CM, HS, MN, SM, KO, and NF are employees of Tsumura & Co., and this study was funded by this company.
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