Glutaminolysis is Essential for Myofibroblast Persistence and In Vivo Targeting Reverses Fibrosis and Cardiac Dysfunction in Heart Failure

HomeCirculationVol. 145, No. 21Glutaminolysis is Essential for Myofibroblast Persistence and In Vivo Targeting Reverses Fibrosis and Cardiac Dysfunction in Heart Failure Free AccessLetterPDF/EPUBAboutView PDFView EPUBSections ToolsAdd to favoritesDownload citationsTrack citationsPermissions ShareShare onFacebookTwitterLinked InMendeleyReddit Jump toFree AccessLetterPDF/EPUBGlutaminolysis is Essential for Myofibroblast Persistence and In Vivo Targeting Reverses Fibrosis and Cardiac Dysfunction in Heart Failure Andrew A. Gibb, PhD, Emma K. Murray, PhD, Anh T. Huynh, BS, Ryan B. Gaspar, Tori L. Ploesch, Ken Bedi, BS, Alyssa A. Lombardi, MD, PhD, Pawel K. Lorkiewicz, PhD, Rajika Roy, PhD, Zolt Arany, MD, PhD, Daniel P. Kelly, MD, Kenneth B. Margulies, MD, Bradford G. Hill, PhD and John W. Elrod, PhD Andrew A. GibbAndrew A. Gibb https://orcid.org/0000-0003-2770-430X Center for Translational Medicine, Department of Cardiovascular Sciences, Lewis Katz School of Medicine at Temple University, Philadelphia, PA (A.A.G., E.K.M., A.T.H., R.B.G., T.L.P., A.A.L., R.R., J.W.E.). , Emma K. MurrayEmma K. Murray https://orcid.org/0000-0001-5871-0564 Center for Translational Medicine, Department of Cardiovascular Sciences, Lewis Katz School of Medicine at Temple University, Philadelphia, PA (A.A.G., E.K.M., A.T.H., R.B.G., T.L.P., A.A.L., R.R., J.W.E.). , Anh T. HuynhAnh T. Huynh Center for Translational Medicine, Department of Cardiovascular Sciences, Lewis Katz School of Medicine at Temple University, Philadelphia, PA (A.A.G., E.K.M., A.T.H., R.B.G., T.L.P., A.A.L., R.R., J.W.E.). , Ryan B. GasparRyan B. Gaspar https://orcid.org/0000-0002-7922-3893 , Tori L. PloeschTori L. Ploesch https://orcid.org/0000-0001-5089-3255 , Ken BediKen Bedi Cardiovascular Institute and Cardiovascular Medicine Division, Department of Medicine, Perelman School of Medicine at the University of Pennsylvania, Philadelphia (K.B., Z.A., D.P.K., K.B.M.). , Alyssa A. LombardiAlyssa A. Lombardi Center for Translational Medicine, Department of Cardiovascular Sciences, Lewis Katz School of Medicine at Temple University, Philadelphia, PA (A.A.G., E.K.M., A.T.H., R.B.G., T.L.P., A.A.L., R.R., J.W.E.). , Pawel K. LorkiewiczPawel K. Lorkiewicz https://orcid.org/0000-0002-1755-9826 Department of Chemistry (P.K.L.), University of Louisville, KY. , Rajika RoyRajika Roy Center for Translational Medicine, Department of Cardiovascular Sciences, Lewis Katz School of Medicine at Temple University, Philadelphia, PA (A.A.G., E.K.M., A.T.H., R.B.G., T.L.P., A.A.L., R.R., J.W.E.). , Zolt AranyZolt Arany https://orcid.org/0000-0003-1368-2453 Cardiovascular Institute and Cardiovascular Medicine Division, Department of Medicine, Perelman School of Medicine at the University of Pennsylvania, Philadelphia (K.B., Z.A., D.P.K., K.B.M.). , Daniel P. KellyDaniel P. Kelly https://orcid.org/0000-0002-3811-9491 Cardiovascular Institute and Cardiovascular Medicine Division, Department of Medicine, Perelman School of Medicine at the University of Pennsylvania, Philadelphia (K.B., Z.A., D.P.K., K.B.M.). , Kenneth B. MarguliesKenneth B. Margulies https://orcid.org/0000-0002-8093-4465 Cardiovascular Institute and Cardiovascular Medicine Division, Department of Medicine, Perelman School of Medicine at the University of Pennsylvania, Philadelphia (K.B., Z.A., D.P.K., K.B.M.). , Bradford G. HillBradford G. Hill Division of Environmental Medicine (B.G.H.), University of Louisville, KY. and John W. ElrodJohn W. Elrod Correspondence to: John W. Elrod, PhD, Center for Translational Medicine, Lewis Katz School of Medicine at Temple University, 3500 N Broad Street, MERB 949, Philadelphia, PA 19140. Email E-mail Address: [email protected] https://orcid.org/0000-0003-3925-2224 Center for Translational Medicine, Department of Cardiovascular Sciences, Lewis Katz School of Medicine at Temple University, Philadelphia, PA (A.A.G., E.K.M., A.T.H., R.B.G., T.L.P., A.A.L., R.R., J.W.E.). Originally published23 May 2022https://doi.org/10.1161/CIRCULATIONAHA.121.057879Circulation. 2022;145:1625–1628After injury, cardiac fibroblasts (CFs) differentiate into highly specialized myofibroblasts, essential for maintaining myocardial structural integrity by contracting surrounding tissue and remodeling the extracellular matrix network.1 It was recently reported that withdrawal of stress stimuli is sufficient for some myofibroblasts to revert to a quiescent phenotype,2 suggesting the potential for myofibroblast reversion by targeting the molecular pathways maintaining the fibrotic phenotype. Metabolic reprogramming has become appreciated as a primary trigger of cell fate transition, directing metabolic intermediates into pathways necessary for bioenergetics, anabolism, and the synthesis of cofactors regulating epigenetic modifiers and downstream gene expression. We recently reported that glutaminolysis (glutamine→glutamate) was linked to histone demethylation mediating myofibroblast formation,3 implicating metabolic remodeling as a critical feature of the differentiation program. However, the greatest therapeutic potential of targeting the myofibroblast phenotype lies in reversing fibrosis by dedifferentiating activated myofibroblasts after injury under chronic stress stimuli.To determine whether glutaminolysis inhibition is sufficient to revert myofibroblasts to a quiescent phenotype and reverse tissue fibrosis, we targeted Gls1 (glutaminase-1), the replace with committed step in of glutaminolysis, following fibroblast activation in a murine model of pressure-overload heart failure (HF). Gls1fl/fl mice4 were crossed with a Periostin inducible-Cre mouse line, restricting Cre recombination to activated fibroblasts (PostniCre.)2 Because periostin is only expressed after fibroblast activation and the in vivo half-life of GLS1 is ~5.1 days,5 this strategy allows for deletion of Gls1 exclusively after myofibroblast differentiation. Given the difficulty in accurately identifying fibroblasts by surface antigens, Gls1fl/fl×PostniCre mice were subsequently crossed to a Cre-dependent reporter mouse line (tdTomato) to permit isolation and assessment of any activated fibroblasts previously or currently expressing periostin (Figure, A). Mice were fed tamoxifen chow (tamox; 40 mg/kg/d) 1 day after transverse aortic constriction (TAC) and for the remainder of the study. Maintenance on tamox allowed for tdTomato labeling and Gls1 deletion in all CFs activated during pressure overload. Cardiac function was monitored before and every 2 weeks after TAC (Figure, B). All animal work was performed in accordance with Institutional Animal Care and Use Committee approval by Temple University and followed American Association for Accreditation of Laboratory Animal Care guidelines. The data that support the findings of this study are available from the corresponding author upon reasonable request.Download figureDownload PowerPointFigure. Inhibition of glutaminolysis in activated myofibroblasts reverses cardiac functional decline and fibrosis after murine pressure overload and reverts myofibroblasts isolated from nonischemic cardiomyopathy patients to a less fibrotic phenotype. A, Gls1fl/fl mice were crossed with mice expressing a tamoxifen (tamox)–inducible, fibroblast-specific Cre recombinase (PostniCre), allowing for Gls1 (glutaminase-1)–specific deletion in activated myofibroblasts, after differentiation. In addition, these mice were crossed to a Cre-dependent reporter line to allow for isolation and assessment of the activated and Gls1-deleted fibroblast population. B, Schematic of experimental timeline. Mice were fed tamox chow 1 day after transverse aortic constriction (TAC) and maintained on tamox chow for the duration of the study. Echocardiography was assessed at baseline and every 2 weeks after surgery. C, Western blot of tdTomato+ cells collected through fluorescence-activated cell sorting to confirm GLS1 deletion (KGA, full length; GAC, splice variant isoform). D, Aortic pressures 1 week after TAC. E, Total TdTomato+ cells from hearts 16 weeks after TAC. F, Ratio of heart weight to tibia length (HW/TL) 16 weeks after TAC. G, Short-axis M-mode echo measurements of percent fractional shortening (FS%). Long-axis B-mode echo measurements of (H) end diastolic volume (EDV), (I) end systolic volume (ESV), and (J) percent ejection fraction (EF%) 16 weeks after TAC. K, Long-axis B-mode measurements of radial myocardial strain rates using speckle tracking echocardiography. L, Inverse strain rates indicative of the rate of myocardial relaxation. M, Representative images of hearts stained with picrosirius red to visualize and quantify tissue fibrosis after 16 weeks of TAC. Scale bar, 250 μm. N, Quantification of images in (M) as the percent of red stained area to total tissue area. O, Hydroxyproline content in left ventricular (LV) tissue samples. P, Schematic of cell acquisition, isolation, and experimental timeline to assess GLS1 inhibition on myofibroblast reversion under sustained stress stimuli (transforming growth factor–β [TGFβ]) in cardiac fibroblasts obtained from patients with nonischemic cardiomyopathy. Q and T, Representative images of cardiac fibroblasts chronically treated ± TGFβ for 48 hours. After the first 24-hour period, once myofibroblast formation had occurred, GLS1 inhibitor CB-839 (10 μM) was added and stained for α–smooth muscle actin (αSMA; red) and nuclear stain (DAPI; blue). Scale bar, 40 μm. R and U, Quantification of images in Q and T as the percent of cells expressing αSMA to the total number of cells (minimum of 80 cells quantified per replicate). S and V, Relative fold change in expression of myofibroblast genes (ACTA2, FN1, POSTN, PDGFRA, COL1A1, and COL3A1). RPS13 was used as the loading control. Data shown as mean ± SEM, n=2 to 15 per group, as indicated. For statistical considerations, unpaired t tests (D, E, and H–L), 1-way analyses of variance with the Tukey post hoc test (R, S, U, and V), 2-way analyses of variance with the Tukey post hoc test (F, N, and O), or a 2-way mixed-effects model analysis of variance with Sidak post hoc test for multiple comparisons (G) was used (*P<0.05, **P<0.01, ***P<0.001, ****P<0.0001). qPCR indicates quantitative polymerase chain reaction.Western blot analysis of tdTomato+ CFs isolated by fluorescence-activated cell sorting confirmed Gls1 deletion after TAC (Figure, C). Aortic pressures obtained 1 week after TAC indicated similar pressure gradients between control (PostniCre) and experimental (Gls1fl/fl×PostniCre) mice (Figure, D). Determination of the number of CFs that had been activated after TAC, ~3% of all gated cells, revealed no significant difference between groups, confirming we were temporally deleting Gls1 after fibroblast activation and differentiation (Figure, E). We did not observe any difference in cardiac hypertrophy (Figure, F), but we did observe preserved left ventricular fractional shortening from 10 to 16 weeks after TAC in Gls1fl/fl×PostniCre mice (Figure, G). To further examine cardiac contractility and relaxation, B-mode speckle-tracking echocardiography was performed. Ejection fraction rebounded in Gls1fl/fl×PostniCre mice 16 weeks after TAC (Figure, H–J). We also found left ventricular radial strain rates in Gls1fl/fl×PostniCre mice to be significantly greater than in controls, indicating increased systolic function (Figure, K). We also saw enhanced inverse radial strain rates, a measure of the rate of left ventricular relaxation or diastolic function (Figure, L), an important finding because diastolic dysfunction highly correlates with tissue fibrosis. Assessment of collagen deposition (% fibrosis) revealed a significant increase in left ventricular fibrosis in control mice 16 weeks after TAC, which was prevented by targeting Gls1 in activated myofibroblasts (Figure, M–O). It is important to note, collagen deposition was similar between groups at an earlier time point (6 weeks after TAC; Figure, O), confirming our hypothesis of fibrotic regression after loss of glutaminolysis in activated myofibroblasts. These in vivo results indicate the therapeutic potential of targeting glutaminolysis in activated myofibroblasts to reverse cardiac fibrosis and mitigate functional decline in HF.To further determine the translational potential of targeting glutaminolysis for myofibroblast reversion, we isolated CFs from heart tissue of patients with nonischemic cardiomyopathy and maintained these cells under disease conditions (transforming growth factor–β stimulation for 48 hours), treating with a GLS1 inhibitor (CB-839; 10 μM) at the 24-hour time point (Figure, P). Research use of human heart tissues in this study was approved by the University of Pennsylvania institutional review board with written informed consent obtained from each of the heart transplant recipients providing explanted heart tissue for this research. Analysis of α–smooth muscle actin+ cells (% αSMA+, myofibroblast number) and activation state of the fibrotic gene program by quantitative polymerase chain reaction revealed pharmacologic inhibition of glutaminolysis in CFs isolated from patients with HF was sufficient to decrease myofibroblast number and revert the gene program to a less fibrotic phenotype (Figure, Q–V). These results provide direct evidence for therapeutically targeting glutaminolysis in patients with HF to positively alter the fibroblast population and curb active fibrosis.Our results identify glutaminolysis as a critical mediator of myofibroblast persistence in a murine model of HF and in CFs isolated directly from failing human hearts. This could be attributable to the essential role of glutamine as a carbon source for bioenergetics, anabolism, collagen biosynthesis, and epigenetic regulation of gene expression, all of which contribute to myofibroblast differentiation and the fibrotic phenotype. Small molecule inhibition of GLS1 is in phase 2 cancer clinical trials, which if found to be safe, could be repurposed rapidly for the treatment of fibrotic disease.Article InformationAcknowledgmentsThe authors thank Trevor Tierney and Alycia Hildebrand for management of the laboratory and mutant mouse colony. Author contributions are as follows: methodology: Drs Gibb and Elrod; data collection: Drs Gibb and Murray, A.T. Huynh, R.B. Gaspar, T.L. Ploesch, K. Bedi, and Drs Lombardi, Lorkiewicz, and Elrod; formal analysis: Drs Gibb, Lorkiewicz, Hill, and Elrod; resources: Drs Gibb, Elrod, Arany, Kelly, Margulies, and Hill; and manuscript writing: Drs Gibb and Elrod. All authors gave approval for the final version of the article.Sources of FundingThis work was supported in part by grants from the National Institutes of Health to Drs Elrod (grants R01HL123966, R01HL136954, R01HL142271, and P01HL134608), Gibb (grant F32HL145914), Murray (grant F31HL143913), Arany (grant R01HL152446), Kelly (grant R01HL058493), Margulies (grants R01HL149891 and R01HL105993), and Hill (grant R01HL130174) and from the American Heart Association to Dr Elrod (grant 20EIA35320226).Nonstandard Abbreviations and AcronymsCFcardiac fibroblastGLS1glutaminase-1HFheart failureTACtransverse aortic constrictionDisclosures Dr Kelly is a scientific consultant for Pfizer Inc and Amgen Inc. Dr Margulies is a consultant for Bristol Myers Squibb and has received research funding from Amgen, Sanofi-Aventis USA, and Merck Sharp & Dohme. Dr Elrod is a scientific consultant for Mitobridge Inc and Janssen and receives research funding from Mitobridge Inc.FootnotesCirculation is available at www.ahajournals.org/journal/circFor Sources of Funding and Disclosures, see page 1628.Correspondence to: John W. Elrod, PhD, Center for Translational Medicine, Lewis Katz School of Medicine at Temple University, 3500 N Broad Street, MERB 949, Philadelphia, PA 19140. Email [email protected]eduReferences1. Gibb AA, Lazaropoulos MP, Elrod JW. Myofibroblasts and fibrosis: mitochondrial and metabolic control of cellular differentiation.Circ Res. 2020; 127:427–447. doi: 10.1161/CIRCRESAHA.120.316958LinkGoogle Scholar2. Kanisicak O, Khalil H, Ivey MJ, et al. Genetic lineage tracing defines myofibroblast origin and function in the injured heart.Nat Commun. 2016; 7:12260. doi: 10.1038/ncomms12260CrossrefMedlineGoogle Scholar3. Lombardi AA, Gibb AA, Arif E, et al. Mitochondrial calcium exchange links metabolism with the epigenome to control cellular differentiation.Nat Commun. 2019; 10:4509. doi: 10.1038/s41467-019-12103-xCrossrefMedlineGoogle Scholar4. Mingote S, Masson J, Gellman C, et al. Genetic pharmacotherapy as an early CNS drug development strategy: testing glutaminase inhibition for schizophrenia treatment in adult mice.Front Syst Neurosci. 2015; 9:165. doi: 10.3389/fnsys.2015.00165MedlineGoogle Scholar5. Tong J, Harrison G, Curthoys NP. The effect of metabolic acidosis on the synthesis and turnover of rat renal phosphate-dependent glutaminase.Biochem J. 1986; 233:139–144. doi: 10.1042/bj2330139CrossrefMedlineGoogle Scholar Previous Back to top Next FiguresReferencesRelatedDetails May 24, 2022Vol 145, Issue 21 Advertisement Article InformationMetrics © 2022 American Heart Association, Inc.https://doi.org/10.1161/CIRCULATIONAHA.121.057879PMID: 35605036 Originally publishedMay 23, 2022 Keywordsmetabolismheart failureglutaminemyofibroblastsfibrosisPDF download Advertisement SubjectsFibrosisHeart FailureMetabolismRemodeling