Cardiac Atrophy, Dysfunction, and Metabolic Impairments

Muscle atrophy and weakness are prevalent features of cancer. Although extensive research has characterized skeletal muscle wasting in cancer cachexia, limited studies have investigated how cardiac structure and function are affected by therapy-naive cancer. Here, the authors used orthotopic, syngeneic models of epithelial ovarian cancer and pancreatic ductal adenocarcinoma, and a patient-derived pancreatic xenograft model, to define the impacts of malignancy on cardiac structure, function, and metabolism. Tumor-bearing mice develop cardiac atrophy and intrinsic systolic and diastolic dysfunction, with arterial hypotension and exercise intolerance. In hearts of ovarian tumor–bearing mice, fatty acid–supported mitochondrial respiration decreased, and carbohydrate-supported respiration increased—showcasing a substrate shift in cardiac metabolism that is characteristic of heart failure. Epithelial ovarian cancer decreased cytoskeletal and cardioprotective gene expression, which was paralleled by down-regulation of transcription factors that regulate cardiomyocyte size and function. Patient-derived pancreatic xenograft tumor–bearing mice show altered myosin heavy chain isoform expression—also a molecular phenotype of heart failure. Markers of autophagy and ubiquitin-proteasome system were upregulated by cancer, providing evidence of catabolic signaling that promotes cardiac wasting. Together, the authors cross-validate with two cancer types, evidence of the structural, functional, and metabolic cancer-induced cardiomyopathy, thus providing translational evidence that could impact future medical management strategies for improved cancer recovery in patients. Muscle atrophy and weakness are prevalent features of cancer. Although extensive research has characterized skeletal muscle wasting in cancer cachexia, limited studies have investigated how cardiac structure and function are affected by therapy-naive cancer. Here, the authors used orthotopic, syngeneic models of epithelial ovarian cancer and pancreatic ductal adenocarcinoma, and a patient-derived pancreatic xenograft model, to define the impacts of malignancy on cardiac structure, function, and metabolism. Tumor-bearing mice develop cardiac atrophy and intrinsic systolic and diastolic dysfunction, with arterial hypotension and exercise intolerance. In hearts of ovarian tumor–bearing mice, fatty acid–supported mitochondrial respiration decreased, and carbohydrate-supported respiration increased—showcasing a substrate shift in cardiac metabolism that is characteristic of heart failure. Epithelial ovarian cancer decreased cytoskeletal and cardioprotective gene expression, which was paralleled by down-regulation of transcription factors that regulate cardiomyocyte size and function. Patient-derived pancreatic xenograft tumor–bearing mice show altered myosin heavy chain isoform expression—also a molecular phenotype of heart failure. Markers of autophagy and ubiquitin-proteasome system were upregulated by cancer, providing evidence of catabolic signaling that promotes cardiac wasting. Together, the authors cross-validate with two cancer types, evidence of the structural, functional, and metabolic cancer-induced cardiomyopathy, thus providing translational evidence that could impact future medical management strategies for improved cancer recovery in patients. Cardiovascular disease (CVD) and cancer are the leading causes of death worldwide.1Khan M.A. Hashim M.J. Mustafa H. Baniyas M.Y. Al Suwaidi S.K.B.M. AlKatheeri R. Alblooshi F.M.K. Almatrooshi M.E.A.H. Alzaabi M.E.H. Al Darmaki R.S. Lootah S.N.A.H. Global epidemiology of ischemic heart disease: results from the Global Burden of Disease Study.Cureus. 2020; 12e9349Google Scholar,2Sung H. Ferlay J. Siegel R.L. Laversanne M. Soerjomataram I. Jemal A. Bray F. Global cancer statistics 2020: GLOBOCAN estimates of incidence and mortality worldwide for 36 cancers in 185 countries.CA Cancer J Clin. 2021; 71: 209-249Crossref PubMed Scopus (60760) Google Scholar Although generally thought of as distinct diseases, the intersectional pathophysiology of the cellular and molecular mechanisms between cancer and CVD have emerged. The field of cardio-oncology developed from this with the aim of improving the identification, monitoring, and treatment of cardiovascular complications in cancer patients during and after cancer therapy. Research has largely focused on how chemotherapeutics impair cardiovascular function.3Herrmann J. Adverse cardiac effects of cancer therapies: cardiotoxicity and arrhythmia.Nat Rev Cardiol. 2020; 17: 474-502Crossref PubMed Scopus (376) Google Scholar Indeed, the cumulative effects of cardiotoxic therapy and the presence of CVD risk factors leads to long-term morbidity and poor quality of life in cancer patients, even when cured of cancer.4Rothe D. Paterson I. Cox-Kennett N. Gyenes G. Pituskin E. Prevention of cardiovascular disease among cancer survivors: the role of pre-existing risk factors and cancer treatments.Curr Epidemiol Rep. 2017; 4: 239-247Crossref Google Scholar Yet, recent evidence suggests that cancer itself—in the absence of exposure to cardiotoxic therapeutics—also negatively impacts cardiovascular health, potentially compounding adverse effects of chemotherapy or impairing recovery.5Lena A. Wilkenshoff U. Hadzibegovic S. Porthun J. Rösnick L. Fröhlich A.K. Zeller T. Karakas M. Keller U. Ahn J. Bullinger L. Riess H. Rosen S.D. Lyon A.R. Lüscher T.F. Totzeck M. Rassaf T. Burkhoff D. Mehra M.R. Bax J.J. Butler J. Edelmann F. Haverkamp W. Anker S.D. Packer M. Coats A.J.S. von Haehling S. Landmesser U. Anker M.S. Clinical and prognostic relevance of cardiac wasting in patients with advanced cancer.J Am Coll Cardiol. 2023; 81: 1569-1586Crossref PubMed Scopus (0) Google Scholar Cancer cachexia, affecting approximately 50% to 80% of cancer patients, is a complex metabolic syndrome, characterized by the progressive loss of body mass, predominantly from skeletal muscle and fat, and multiple organs including the liver, kidney, spleen, gastrointestinal tract, and heart.6Fearon K. Strasser F. Anker S.D. Bosaeus I. Bruera E. Fainsinger R.L. Jatoi A. Loprinzi C. MacDonald N. Mantovani G. Davis M. Muscaritoli M. Ottery F. Radbruch L. Ravasco P. Walsh D. Wilcock A. Kaasa S. Baracos V.E. Definition and classification of cancer cachexia: an international consensus.Lancet Oncol. 2011; 12: 489-495Abstract Full Text Full Text PDF PubMed Scopus (3888) Google Scholar,7Argilés J.M. Busquets S. Stemmler B. López-Soriano F.J. Cancer cachexia: understanding the molecular basis.Nat Rev Cancer. 2014; 14: 754-762Crossref PubMed Scopus (951) Google Scholar Skeletal muscle weakness and atrophy are well-known consequences of cancer cachexia.8Delfinis L.J. Bellissimo C.A. Gandhi S. DiBenedetto S.N. Garibotti M.C. Thuhan A.K. Tsitkanou S. Rosa-Caldwell M.E. Rahman F.A. Cheng A.J. Wiggs M.P. Schlattner U. Quadrilatero J. Greene N.P. Perry C.G. Muscle weakness precedes atrophy during cancer cachexia and is linked to muscle-specific mitochondrial stress.JCI Insight. 2022; 7e155147Crossref PubMed Scopus (18) Google Scholar Cancer-induced cardiac atrophy and dysfunction have also been identified in cancer patients and preclinical cancer models,9Springer J. Tschirner A. Haghikia A. von Haehling S. Lal H. Grzesiak A. Kaschina E. Palus S. Pötsch M. von Websky K. Hocher B. Latouche C. Jaisser F. Morawietz L. Coats A.J. Beadle J. Argiles J.M. Thum T. Földes G. Doehner W. Hilfiker-Kleiner D. Force T. Anker S.D. Prevention of liver cancer cachexia-induced cardiac wasting and heart failure.Eur Heart J. 2014; 35: 932-941Crossref PubMed Scopus (159) Google Scholar, 10Tian M. Asp M.L. Nishijima Y. Belury M.A. Evidence for cardiac atrophic remodeling in cancer-induced cachexia in mice.Int J Oncol. 2011; 39: 1321-1326PubMed Google Scholar, 11Tian M. Nishijima Y. Asp M.L. Stout M.B. Reiser P.J. Belury M.A. Cardiac alterations in cancer-induced cachexia in mice.Int J Oncol. 2010; 37: 347-353PubMed Google Scholar, 12Cosper P.F. Leinwand L.A. Cancer causes cardiac atrophy and autophagy in a sexually dimorphic manner.Cancer Res. 2011; 71: 1710-1720Crossref PubMed Scopus (169) Google Scholar yet few studies have investigated cancer-induced cardiomyopathy in a therapy-naive setting with insidious, clinically late-presenting cancers: ovarian or pancreatic, where cachexia is a frequent and prominent feature.13Ubachs J. Ziemons J. Minis-Rutten I.J.G. Kruitwagen R.F.P.M. Kleijnen J. Lambrechts S. Olde Damink S.W.M. Rensen S.S. Van Gorp T. Sarcopenia and ovarian cancer survival: a systematic review and meta-analysis.J Cachexia Sarcopenia Muscle. 2019; 10: 1165-1174Crossref PubMed Scopus (113) Google Scholar,14Mitsunaga S. Kasamatsu E. Machii K. Incidence and frequency of cancer cachexia during chemotherapy for advanced pancreatic ductal adenocarcinoma.Support Care Cancer. 2020; 28: 5271-5279Crossref PubMed Scopus (24) Google Scholar Improving survival and quality of life for these cancer patients could be possible with a fuller understanding of the pathology. Muscle wasting is attributed to protein degradation and low protein synthesis, due to elevated systemic inflammatory cytokines from activated immune cells of both tumor and host.15Vudatha V. Devarakonda T. Liu C. Freudenberger D.C. Riner A.N. Herremans K.M. Trevino J.G. Review of mechanisms and treatment of cancer-induced cardiac cachexia.Cells. 2022; 11: 1-16Crossref Scopus (8) Google Scholar These cytokines promote hyperactivation of ubiquitin-proteasome system (UPS)-mediated protein degradation, leading to the breakdown of myofibrillar proteins causing atrophy.7Argilés J.M. Busquets S. Stemmler B. López-Soriano F.J. Cancer cachexia: understanding the molecular basis.Nat Rev Cancer. 2014; 14: 754-762Crossref PubMed Scopus (951) Google Scholar Autophagic flux is up-regulated in conditions of nutrient or growth factor deprivation and contributes to skeletal muscle depletion in cancer cachexia.16Penna F. Ballarò R. Martinez-Cristobal P. Sala D. Sebastian D. Busquets S. Muscaritoli M. Argilés J.M. Costelli P. Zorzano A. Autophagy exacerbates muscle wasting in cancer cachexia and impairs mitochondrial function.J Mol Biol. 2019; 431: 2674-2686Crossref PubMed Scopus (4) Google Scholar,17Penna F. Costamagna D. Pin F. Camperi A. Fanzani A. Chiarpotto E.M. Cavallini G. Bonelli G. Baccino F.M. Costelli P. Autophagic degradation contributes to muscle wasting in cancer cachexia.Am J Pathol. 2013; 182: 1367-1378Abstract Full Text Full Text PDF PubMed Scopus (195) Google Scholar Subsequently, proteins and organelles are targeted for lysosomal degradation and the resulting molecular components are recycled as substrates to support metabolic demand. Although both autophagic and UPS activation are implicated in the development of skeletal muscle wasting in cancer, the proportional contributions in various cancer types to cancer-induced cardiomyopathy are uncertain. Cardiac transcription factors regulate the expression of genes encoding structural and regulatory proteins in the myocardium to maintain functional homeostasis between atrophy and hypertophy.18Akazawa H. Komuro I. Roles of cardiac transcription factors in cardiac hypertrophy.Circ Res. 2003; 92: 1079-1088Crossref PubMed Scopus (331) Google Scholar Several transcription factors—GATA binding protein (GATA), Myocyte enhancing factor 2 (MEF2), Nuclear factor of activated T cells (NFAT) and Serum response factor (SRF)—regulate the cardiac gene program in embryonic development and in response to mitogenic or hemodynamic stimuli.19Pikkarainen S. Tokola H. Kerkelä R. Ruskoaho H. GATA transcription factors in the developing and adult heart.Cardiovasc Res. 2004; 63: 196-207Crossref PubMed Scopus (333) Google Scholar The up-regulation of these transcription factors in adults elicits hypertrophic growth, though each is regulated by varying stimuli. Thus, whether cardiac transcription factors are regulating cancer-induced cardiomyopathy is unknown. The etiology of cardiac disease in cancer patients is complex and our understanding of how this cardiac phenotype manifests in patients is confounded by a multitude of factors—aging, comorbidities, therapeutics, direct effects of tumors—influencing cardiac health. Thus, the use of preclinical cancer models to study cancer-induced cardiomyopathy is critical, particularly for investigating cellular and molecular mechanisms that define how tumors directly promote cardiac disease progression. Research in this area has primarily relied upon ectopic cancer models, where cancer cells or organoids are implanted to a different anatomical site rather than what would be the clinical origin. Such models show rapid formation of large primary tumors and were used generally to study the etiology of cancer cachexia, but do not reflect the early disruption of the oncogenic organ and don't mirror phenotypic characteristics of human cancer. Orthotopic cancer models involve the injection of malignant cells or organoids—ideally, and where possible, derived directly from patients—into the same anatomical location they originate; thus model more closely the local and systemic cancer environments. Here, the authors assessed cardiac structure and function in preclinical orthotopic cancer models as: i) cell-based murine epithelial ovarian cancer (EOC), ii) cell-based murine pancreatic ductal adenocarcinoma (PDAC), and iii) patient-derived pancreatic cancer xenograft (PDX). To investigate the underlying mechanisms of cancer-induced cardiomyopathy, the authors measured cardiac substrate metabolism and oxidative stress levels in the myocardium, quantified the cardiac gene program, and quantified the catabolic pathways that regulate protein degradation in the heart. The authors discovered that ovarian and pancreatic cancer cause hemodynamic dysfunction and cardiac atrophy, metabolic dysregulation, capillary rarefaction, and exercise intolerance. The authors distinguish the atrophy as autophagic dominant in ovarian cancer and, comparatively, both autophagic and proteasomal in pancreatic cancer. The authors provide here the first investigation of orthotopic patient-xenografted cancer-induced cardiomyopathy. C57BL/6 (Charles River Laboratories, Wilmington, MA) and NOD-SCID IL2 receptor γ chain knockout (NSG) mice (Jackson Laboratories, Bar Harbor, ME) were housed in a temperature-controlled, 12 hours light:dark cycle, and given food and water ad libitum. NSG mice were housed in virus-free conditions. All procedures were approved by the Animal Care Committee at the University of Guelph and in compliance with Canadian Council on Animal Care guidelines or guidelines from the Virginia Commonwealth University Institutional Animal Care and Use Committee. Spontaneously transformed murine ovarian surface epithelial cells (ID8; generously donated by Drs. K. Roby and P. Terranova, Kansas State University, Manhattan, KS) were cultured in Dulbecco's Modified Eagle Medium (Wisent Inc., Saint-Jean-Baptiste, QC, Canada) with 10% fetal bovine serum and 1% antibiotic/antimycotic (Wisent Inc.). KrasLSL-G12D; Trp53LSL-R172H; Pdx1-Cre (KPC) cells (generously donated by Dr. Steven Gallinger, University of Toronto, Toronto, ON, Canada)20Lutz V. Hellmund V.M. Picard F.S.R. Raifer H. Ruckenbrod T. Klein M. Bopp T. Savai R. Duewell P. Keber C.U. Weigert A. Chung H.R. Buchholz M. Menke A. Gress T.M. Huber M. Bauer C. IL18 receptor signaling regulates tumor-reactive CD8+ T-cell exhaustion via activation of the IL2/STAT5/mTOR pathway in a pancreatic cancer model.Cancer Immunol Res. 2023; 11: 421-434Crossref PubMed Scopus (0) Google Scholar, 21Horvat N.K. Karpovsky I. Phillips M. Wyatt M.M. Hall M.A. Herting C.J. Hammons J. Mahdi Z. Moffitt R.A. Paulos C.M. Lesinski G.B. Clinically relevant orthotopic pancreatic cancer models for adoptive T cell transfer therapy.J Immunother Cancer. 2024; 12e008086Crossref PubMed Scopus (2) Google Scholar, 22Michaelis K.A. Zhu X. Burfeind K.G. Krasnow S.M. Levasseur P.R. Morgan T.K. Marks D.L. Establishment and characterization of a novel murine model of pancreatic cancer cachexia.J Cachexia Sarcopenia Muscle. 2017; 8: 824-838Crossref PubMed Scopus (88) Google Scholar were cultured in RPMI (Wisent Inc.) with 10% fetal bovine serum, 1% antibiotic/antimycotic, and 1% sodium pyruvate (Wisent Inc.). Cell cultures were maintained on 100-mm cell culture plates (Corning Inc., Corning, NY) by passaging cells once they reached 70% to 80% confluency. In female C57BL/6 mice at 9 to 10 weeks of age, ovarian tumors were induced as an orthotopic, syngeneic mouse model of EOC as described previously.23Greenaway J. Petrik J. Ovarian cancer methods and protocols methods in molecular biology 1049.Methods Mol Biol. 2013; 1049: 409-423Crossref PubMed Scopus (0) Google Scholar, 24Matuszewska K. Ten Kortenaar S. Pereira M. Santry L.A. Petrik D. Lo K.M. Bridle B.W. Wootton S.K. Lawler J. Petrik J. Addition of an Fc-IgG induces receptor clustering and increases the in vitro efficacy and in vivo anti-tumor properties of the thrombospondin-1 type I repeats (3TSR) in a mouse model of advanced stage ovarian cancer.Gynecol Oncol. 2022; 164: 154-169Abstract Full Text Full Text PDF PubMed Scopus (0) Google Scholar, 25Russell S. Duquette M. Liu J. Drapkin R. Lawler J. Petrik J. Combined therapy with thrombospondin-1 type I repeats (3TSR) and chemotherapy induces regression and significantly improves survival in a preclinical model of advanced stage epithelial ovarian cancer.FASEB J. 2015; 29: 576-588Crossref PubMed Scopus (54) Google Scholar, 26Matuszewska K. Santry L.A. van Vloten J.P. AuYeung A.W.K. Major P.P. Lawler J. Wootton S.K. Bridle B.W. Petrik J. Combining vascular normalization with an oncolytic virus enhances immunotherapy in a preclinical model of advanced-stage ovarian cancer.Clin Cancer Res. 2019; 25: 1624-1638Crossref PubMed Scopus (56) Google Scholar Briefly, a dorsal incision was made on isoflurane-anesthetized mice, and 1.0 × 106 ID8 cells in 5 μL of sterile saline were injected under the bursa of the left ovary. Sham mice followed the same procedure with injection of only sterile saline. In this model, large primary tumors form approximately 60 days after tumor cell injection, followed the development of abdominal ascites and peritoneal metastasis via transcoelomic spread, where lesions are characterized by TP53175H gain-of-function mutations, at which point the model has disease characteristics that correspond to a clinical profile of stage III EOC.25Russell S. Duquette M. Liu J. Drapkin R. Lawler J. Petrik J. Combined therapy with thrombospondin-1 type I repeats (3TSR) and chemotherapy induces regression and significantly improves survival in a preclinical model of advanced stage epithelial ovarian cancer.FASEB J. 2015; 29: 576-588Crossref PubMed Scopus (54) Google Scholar,27Greenaway J.B. Virtanen C. Osz K. Revay T. Hardy D. Shepherd T. DiMattia G. Petrik J. Ovarian tumour growth is characterized by mevalonate pathway gene signature in an orthotopic, syngeneic model of epithelial ovarian cancer.Oncotarget. 2016; 7: 47343-47365Crossref PubMed Scopus (30) Google Scholar,28Greenaway J.B. Virtanen C. Osz K. Revay T. Hardy D. Shepherd T. DiMattia G. Petrik J. Epithelial-stromal interaction increases cell proliferation, survival and tumorigenicity in a mouse model of human epithelial ovarian cancer.Gynecol Oncol. 2008; 108: 385-394Abstract Full Text Full Text PDF PubMed Scopus (79) Google Scholar To evaluate cardiac health in advanced-stage ovarian cancer, tumors were allowed to develop for approximately 75 to 90 days after cell implantation, then in vivo experiments were performed and tissues were collected. At 9 to 10 weeks of age, pancreatic tumors were induced as an orthotopic, syngeneic, immunocompetent mouse model of PDAC. Briefly, male and female C57BL/6 mice were anesthetized with isoflurane, a dorsal incision was made and 1.0 × 105 KPC cells were injected into the tail of the pancreas. Sham mice followed the same procedure with injection of only sterile saline. To evaluate cardiac physiology in advanced-stage pancreatic cancer, tumors were allowed to develop for approximately 28 to 30 days after cell implantation, then cardiac hemodynamic and histological analyses were performed. Tumor tissue was obtained from patients with pancreatic ductal adenocarcinoma at the time of surgery (University of Florida, IRB 201600873). Written consent was obtained from the subject, and research was conducted according to the International Ethical Guidelines for Biomedical Research Involving Human Subjects. The PDX (G160) utilized in this study was obtained from a 63-year-old male patient with T1N1M0 PDAC. The tumor was 1.5 cm in size, poorly differentiated with lymphovascular and perineural invasion, and has a KRASG12D mutation. A viable 2 × 2-mm portion of tissue was isolated from a surgically resected primary pancreatic cancer specimen with minimal ischemia time. Tissue was implanted subcutaneously into male (NSG) mice (Jackson Laboratory). Xenografts were allowed to grow to a maximum diameter of 1.5 cm before passage. Herein, the authors defined passage as explantation of a pancreatic cancer xenograft and orthotopic implantation into the pancreas of a new host. Xenografts were allowed to grow for approximately 20 weeks, then mice were euthanized and hearts were collected for histological and molecular analyses. For voluntary wheel running assessments, mice were individually housed in cages with access to an exercise wheel, and a cycle computer (VDO, Hanover, Germany) was used to record running distance. Mice were given a 24-hour acclimation period, then data recorded for two consecutive 24-hour periods were averaged. Mice were anesthetized with an isoflurane/oxygen mix (2%:100%) and body temperature maintained at 37.2°C to 37.5°C. M-Mode echocardiography was performed on the left ventricle (LV) using the Vevo2100 system (VisualSonics Inc., Toronto, ON, Canada) with a MS550D transducer. Mice were anesthetized with an isoflurane/oxygen mix (2.5%:100%) and body temperature maintained at 37.2°C to 37.5°C. The right carotid artery was isolated, and a 1.2-Fr pressure catheter (Transonic Scisense Inc., London, ON, Canada) was inserted and advanced into the LV. Hemodynamic signals were sampled at a rate of 2 kHz and analysis performed with Spike2 software (Cambridge Electronic Design, Ltd., Cambridge, UK). Mice were anesthetized with an isoflurane/oxygen mix (2.5%/100%). Hearts were excised and rinsed in ice-cold phosphate buffered saline. The aorta was mounted on a 20-ga cannula, and hearts were retrograde-perfused at a constant pressure of 70 to 75 mm Hg with warmed (37°C) and oxygenated (95% O2:5% CO2) Krebs-Henseleit buffer (pH 7.4), containing (in mmol/L) 118 NaCl, 4.7 KCl, 1.2 MgSO4, 1.2 KH2PO4, 0.5 C3H3NaO3, 0.05 EDTA, 11 glucose, and 2 CaCl2. After stable perfusion rates were achieved, the left atrial appendage was removed, and a deflated balloon connected to a pressure transducer was inserted into the LV to record pressure. The balloon was inflated in a stepwise manner (approximately 2 mm Hg change in end-diastolic pressure) until a set end-diastolic pressure of 5 to 8 mm Hg was reached. The maximal rates of LV contraction (dP/dt Max) and relaxation (dP/dt Min) and maximal LV pressure were obtained from raw pressure tracings using Spike2. This technique was adapted from previous methods described elsewhere.29Hughes M.C. Ramos S.V. Turnbull P.C. Edgett B.A. Huber J.S. Polidovitch N. Schlattner U. Backx P.H. Simpson J.A. Perry C.G.R. Impairments in left ventricular mitochondrial bioenergetics precede overt cardiac dysfunction and remodelling in Duchenne muscular dystrophy.J Physiol. 2020; 598: 1377-1392Crossref PubMed Scopus (26) Google Scholar Briefly, the heart was removed and placed in ice-cold BIOPS, containing (in mmol/L): 50 MES hydrate, 7.23 K2EGTA, 20 imidazole, 0.5 dithiothreitol, 20 taurine, 5.77 ATP, 15 PCr, and 6.56 MgCl2·6H2O (pH 7.2). The LV was isolated and gently separated along the longitudinal axis to form fiber bundles (PmFBs), which were blotted and weighed (0.8 to 2.1 mg wet weight) in 1.5 mL of tared, prechilled BIOPS to ensure PmFBs remained relaxed and hydrated. These bundles were treated with 40 μg/mL saponin in BIOPS on a rotor for 30 minutes at 4°C to selectively permeabilize the cell membrane. PmFBs that were used for mH2O2 were also treated with 35 μmol/L 2,4-dinitrochlorobenzene (CDNB) during the permeabilization step to deplete glutathione and allow for detectable rates of mH2O2.30Fisher-Wellman K.H. Gilliam L.A.A. Lin C.T. Cathey B.L. Lark D.S. Darrell Neufer P. Mitochondrial glutathione depletion reveals a novel role for the pyruvate dehydrogenase complex as a key H2O2-emitting source under conditions of nutrient overload.Free Radic Biol Med. 2013; 65: 1201-1208Crossref PubMed Scopus (93) Google Scholar All PmFBs were then washed in Buffer Z (containing, in mmol/L: 105 K-MES, 50 KCl, 10 KH2PO4, 5 MgCl2·6 H2O, 1 EGTA, and 5 mg/mL bovine serum albumin; pH 7.2) on a rotator for 15 minutes at 4°C to remove the cytoplasm. High-resolution oxygen consumption measurements were conducted in 2 mL of respiration media (Buffer Z) using the Oxygraph-2k (Oroboros Instruments, Innsbruck, Austria) while stirring at 750 rpm at 37°C. Buffer Z contained 20 mmol/L creatine to saturate mitochondrial creatine kinase (mtCK) and promote phosphate shuttling through mtCK.31Perry C.G. Kane D.A. Lin C.T. Kozy R. Cathey B.L. Lark D.S. Kane C.L. Brophy P.M. Gavin T.P. Anderson E.J. Neufer P.D. Inhibiting myosin-ATPase reveals a dynamic range of mitochondrial respiratory control in skeletal muscle.Biochem J. 2011; 437: 215-222Crossref PubMed Scopus (135) Google Scholar For carbohydrate-supported ADP-stimulated respiratory kinetics, 5 mmol/L pyruvate and 2 mmol/L malate were added as complex I substrates (via generation of NADH to saturate electron entry into complex I) followed by titrations of submaximal ADP (25 μmol/L, 100 μmol/L, 500 μmol/L, and 1000 μmol/L) and maximal ADP (5000 μmol/L). Glutamate (10 mmol/L) was also added to the assay medium to further saturate complex I with NADH. Last, cytochrome c was added to test for outer mitochondrial membrane integrity, and succinate (20 mmol/L) was added to saturate electron entry into complex II with FADH2. For fatty acid–supported ADP-stimulated respiratory kinetics, 5 mmol/L l-carnitine, 0.02 mmol/L palmitoyl-CoA, and 0.5 mmol/L malate were added as complex I, II, and electron transport chain (ETC) substrates (via generation of NADH and FADH2 from ß-oxidation) followed by titrations of submaximal ADP (100 μmol/L and 300 μmol/L) and maximal ADP (500 μmol/L). Cytochrome c was added to test for outer mitochondrial membrane integrity, and succinate (20 mmol/L) was added to saturate electron entry into complex II. All experiments were completed in the presence of 5 μmol/L BLEB in the assay medium to prevent ADP-induced contraction of PmFBs.31Perry C.G. Kane D.A. Lin C.T. Kozy R. Cathey B.L. Lark D.S. Kane C.L. Brophy P.M. Gavin T.P. Anderson E.J. Neufer P.D. Inhibiting myosin-ATPase reveals a dynamic range of mitochondrial respiratory control in skeletal muscle.Biochem J. 2011; 437: 215-222Crossref PubMed Scopus (135) Google Scholar mH2O2 was determined spectrofluorometrically (QuantaMaster 40; HORIBA Scientific, Kyoto, Japan) in a quartz cuvette with continuous stirring at 37°C, in 1 mL of Buffer Z supplemented with 10 μmol/L Amplex Ultra Red, 0.5 U/mL horseradish peroxidase, 1 mmol/L EGTA, 40 U/mL Cu/Zn-SOD1, 5 μmol/L BLEB, and 20 mmol/L Cr. Buffer Z contained (in mmol/L) 105 K-MES, 30 KCl, 10 KH2PO4, 5 MgCl2·6H2O, and 1 EGTA and 5 mg/mL bovine serum albumin (pH 7.2). State II mH2O2 (maximal emission in the absence of ADP) was induced using the complex I–supporting substrates (NADH) pyruvate (10 mmol/L) and malate (2 mmol/L) as described previously.32Hughes M.C. Ramos S.V. Turnbull P.C. Rebalka I.A. Cao A. Monaco C.M.F. Varah N.E. Edgett B.A. Huber J.S. Tadi P. Delfinis L.J. Schlattner U. Simpson J.A. Hawke T.J. Perry C.G.R. Early myopathy in Duchenne muscular dystrophy is associated with elevated mitochondrial H2O2 emission during impaired oxidative phosphorylation.J Cachexia Sarcopenia Muscle. 2019; 10: 643-661Crossref PubMed Scopus (84) Google Scholar Following the induction of state II mH2O2, a titration of ADP was employed to progressively attenuate mH2O2 as occurs when membrane potential declines during oxidative phosphorylation. The rate of mH2O2 emission was calculated from the slope (F/minute) using a standard curve established with the same reaction conditions and normalized to fiber bundle wet weight. LV tissue was homogenized in ice-cold buffer containing (in mmol/L) 20 Tris, 150 NaCl, 1 EDTA, 1 EGTA, 2.5 Na4O7P2, 1% Triton X-100 with PhosSTOP inhibitor tablet (MilliporeSigma, Burlington, MA) and protease inhibitor cocktail (MilliporeSigma). Seven to 10 μg of protein, measured by a BCA protein assay kit (ThermoFisher Scientific Inc., Waltham, MA), was subjected to 10% to 12% SDS-PAGE followed by transfer to polyvinylidene difluoride or nitrocellulose membrane. For ETC proteins, polyvinylidene difluoride membranes were blocked with Odyssey blocking buffer (Li-COR, Lincoln, NE) and immunoblotted overnight (4°C) with rodent OXPHOS cocktail monoclonal antibody (Abcam, Cambridge, UK; ab110413, 1:1000) to detect ETC proteins. Membranes were washed and incubated with an infrared fluorescent secondary antibody (LI-COR; 1:20,000). Immunoreactive proteins were detected by infrared imaging. For LC3 proteins, nitrocellulose membranes were blocked with 5% skim milk in Tris-buffered saline plus 0.1% Tween 20 and immunoblotted overnight (4°C) with rabbit monoclonal antibody for LC3-I/LC3-II proteins (1:1000, Cell Signaling Technology #12741; Cell Signaling Technology, Danvers, MA). Membranes were washed with Tris-buffered s