Chemogenetic Mitochondrial H2O2 Generation Triggers Dose-Dependent Skeletal Muscle Wasting Signatures

Chemogenetic Mitochondrial H2O2 Generation Triggers Dose-Dependent Skeletal Muscle Wasting Signatures | bioRxiv /* */ /* */ <!-- <!-- /*! * yepnope1.5.4 * (c) WTFPL, GPLv2 */ (function(a,b,c){function d(a){return"[object Function]"==o.call(a)}function e(a){return"string"==typeof a}function f(){}function g(a){return!a||"loaded"==a||"complete"==a||"uninitialized"==a}function h(){var a=p.shift();q=1,a?a.t?m(function(){("c"==a.t?B.injectCss:B.injectJs)(a.s,0,a.a,a.x,a.e,1)},0):(a(),h()):q=0}function i(a,c,d,e,f,i,j){function k(b){if(!o&&g(l.readyState)&&(u.r=o=1,!q&&h(),l.onload=l.onreadystatechange=null,b)){"img"!=a&&m(function(){t.removeChild(l)},50);for(var d in y[c])y[c].hasOwnProperty(d)&&y[c][d].onload()}}var j=j||B.errorTimeout,l=b.createElement(a),o=0,r=0,u={t:d,s:c,e:f,a:i,x:j};1===y[c]&&(r=1,y[c]=[]),"object"==a?l.data=c:(l.src=c,l.type=a),l.width=l.height="0",l.onerror=l.onload=l.onreadystatechange=function(){k.call(this,r)},p.splice(e,0,u),"img"!=a&&(r||2===y[c]?(t.insertBefore(l,s?null:n),m(k,j)):y[c].push(l))}function j(a,b,c,d,f){return q=0,b=b||"j",e(a)?i("c"==b?v:u,a,b,this.i++,c,d,f):(p.splice(this.i++,0,a),1==p.length&&h()),this}function k(){var a=B;return a.loader={load:j,i:0},a}var l=b.documentElement,m=a.setTimeout,n=b.getElementsByTagName("script")[0],o={}.toString,p=[],q=0,r="MozAppearance"in l.style,s=r&&!!b.createRange().compareNode,t=s?l:n.parentNode,l=a.opera&&"[object Opera]"==o.call(a.opera),l=!!b.attachEvent&&!l,u=r?"object":l?"script":"img",v=l?"script":u,w=Array.isArray||function(a){return"[object Array]"==o.call(a)},x=[],y={},z={timeout:function(a,b){return b.length&&(a.timeout=b[0]),a}},A,B;B=function(a){function b(a){var a=a.split("!"),b=x.length,c=a.pop(),d=a.length,c={url:c,origUrl:c,prefixes:a},e,f,g;for(f=0;f<d;f++)g=a[f].split("="),(e=z[g.shift()])&&(c=e(c,g));for(f=0;f<b;f++)c=x[f](c);return c}function g(a,e,f,g,h){var i=b(a),j=i.autoCallback;i.url.split(".").pop().split("?").shift(),i.bypass||(e&&(e=d(e)?e:e[a]||e[g]||e[a.split("/").pop().split("?")[0]]),i.instead?i.instead(a,e,f,g,h):(y[i.url]?i.noexec=!0:y[i.url]=1,f.load(i.url,i.forceCSS||!i.forceJS&&"css"==i.url.split(".").pop().split("?").shift()?"c":c,i.noexec,i.attrs,i.timeout),(d(e)||d(j))&&f.load(function(){k(),e&&e(i.origUrl,h,g),j&&j(i.origUrl,h,g),y[i.url]=2})))}function h(a,b){function c(a,c){if(a){if(e(a))c||(j=function(){var a=[].slice.call(arguments);k.apply(this,a),l()}),g(a,j,b,0,h);else if(Object(a)===a)for(n in m=function(){var b=0,c;for(c in a)a.hasOwnProperty(c)&&b++;return b}(),a)a.hasOwnProperty(n)&&(!c&&!--m&&(d(j)?j=function(){var a=[].slice.call(arguments);k.apply(this,a),l()}:j[n]=function(a){return function(){var b=[].slice.call(arguments);a&&a.apply(this,b),l()}}(k[n])),g(a[n],j,b,n,h))}else!c&&l()}var h=!!a.test,i=a.load||a.both,j=a.callback||f,k=j,l=a.complete||f,m,n;c(h?a.yep:a.nope,!!i),i&&c(i)}var i,j,l=this.yepnope.loader;if(e(a))g(a,0,l,0);else if(w(a))for(i=0;i (function(w,d,s,l,i){w[l]=w[l]||[];w[l].push({'gtm.start':new Date().getTime(),event:'gtm.js'});var f=d.getElementsByTagName(s)[0];var j=d.createElement(s);var dl=l!='dataLayer'?'&l='+l:'';j.src='//www.googletagmanager.com/gtm.js?id='+i+dl;j.type='text/javascript';j.async=true;f.parentNode.insertBefore(j,f);})(window,document,'script','dataLayer','GTM-M677548'); Skip to main content Home About Submit ALERTS / RSS Search for this keyword Advanced Search New Results Chemogenetic Mitochondrial H 2 O 2 Generation Triggers Dose-Dependent Skeletal Muscle Wasting Signatures View ORCID Profile Roberto Meneses-Valdés , View ORCID Profile Samantha Gallero , Jonva Hentze , Sascha Friis Nielsen , View ORCID Profile Joachim Nielsen , View ORCID Profile Enrique Jaimovich , View ORCID Profile Carlos Henríquez-Olguín , View ORCID Profile Thomas E Jensen doi: https://doi.org/10.1101/2025.08.19.671046 Roberto Meneses-Valdés 1 The August Krogh Section for Human and Molecular Physiology, Department of Nutrition , Exercise and Sports, University of Copenhagen. 2 Center for Exercise, Metabolism and Cancer, ICBM, Universidad de Chile , 8380453, Santiago, Chile Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Roberto Meneses-Valdés Samantha Gallero 1 The August Krogh Section for Human and Molecular Physiology, Department of Nutrition , Exercise and Sports, University of Copenhagen. Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Samantha Gallero Jonva Hentze 1 The August Krogh Section for Human and Molecular Physiology, Department of Nutrition , Exercise and Sports, University of Copenhagen. Find this author on Google Scholar Find this author on PubMed Search for this author on this site Sascha Friis Nielsen 1 The August Krogh Section for Human and Molecular Physiology, Department of Nutrition , Exercise and Sports, University of Copenhagen. Find this author on Google Scholar Find this author on PubMed Search for this author on this site Joachim Nielsen 3 Department of Sports Science and Clinical Biomechanics, University of Southern Denmark , Odense, Denmark Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Joachim Nielsen Enrique Jaimovich 2 Center for Exercise, Metabolism and Cancer, ICBM, Universidad de Chile , 8380453, Santiago, Chile Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Enrique Jaimovich Carlos Henríquez-Olguín 1 The August Krogh Section for Human and Molecular Physiology, Department of Nutrition , Exercise and Sports, University of Copenhagen. 4 Center for Exercise Physiology and Metabolism, Department of Kinesiology, Faculty of Medicine, Universidad Finis Terrae , Santiago, Chile Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Carlos Henríquez-Olguín For correspondence: col{at}nexs.ku.dk tejensen{at}nexs.ku.dk Thomas E Jensen 1 The August Krogh Section for Human and Molecular Physiology, Department of Nutrition , Exercise and Sports, University of Copenhagen. Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Thomas E Jensen For correspondence: col{at}nexs.ku.dk tejensen{at}nexs.ku.dk Abstract Full Text Info/History Metrics Supplementary material Preview PDF Abstract Mitochondrial hydrogen peroxide (mtH 2 O 2 ) has long been implicated in skeletal muscle atrophy, yet its direct role in vivo has remained unresolved due to methodological constraints. Here, we aimed to establish a chemogenetic platform for precise, compartment-specific induction of mtH 2 O 2 in adult skeletal muscle and to investigate how graded redox stress impacts on muscle proteostasis in vivo. Using mitochondria-targeted D-amino acid oxidase (mtDAAO), we show that prolonged and/or high mtH 2 O 2 exposure progressively activates proteolytic and denervation-associated pathways, culminating in myofiber damage and regeneration. Remarkably, even low exposure to mtH 2 O 2 is sufficient to acutely suppress protein synthesis and induce disuse-like atrophy, without structural damage or overt oxidative stress. This approach provides a powerful in vivo framework to dissect subcellular redox-controlled signaling in muscle and identifies mtH 2 O 2 as a modulator of muscle proteostasis, with therapeutic relevance for muscle-wasting conditions. 1. INTRODUCTION Loss of skeletal muscle mass, known as muscle wasting, is a hallmark of many diseases and conditions, including aging, cancer, and prolonged immobilization, affecting millions of people worldwide 1 , 2 . Beyond impairing mobility and strength, muscle wasting is associated with metabolic disorders such as insulin resistance, cognitive decline (including dementia), and increased all-cause mortality 3 – 6 . Despite its widespread prevalence 7 and serious clinical consequences, there is currently no approved pharmacological treatment to prevent or reverse muscle loss. Muscle wasting occurs when muscle fiber proteolysis chronically exceeds protein synthesis and/or when fibers undergo degeneration/regeneration 5 , with at least three overlapping but distinct responses. Firstly, disuse (e.g., limb immobilization, bed rest) reduces contractile activity and loading. In rodents, hindlimb immobilization or suspension typically reduces muscle fiber CSA by 20–40% within a week 8 , 9 . Disuse atrophy acutely impairs protein synthesis within the first day, preceding enhanced proteolysis 9 . Secondly, denervation induces disuse but also distinct molecular changes, including upregulation of NCAM1 10 , 11 and Akt 12 . In rodents, surgical or toxin-induced denervation causes similar or greater atrophy than disuse (∼30–50% CSA loss in 7 days) 12 – 16 , strongly upregulates proteolytic activity 14 , 17 , yet fibers remain structurally intact for at least three weeks 13 . Thirdly, myofiber death and degeneration occur in trauma or genetic myopathies. Rodent models include dystrophy/myopathy 18 – 23 or acute chemical/physical damage 24 , 25 , which stimulate regenerative myogenesis 26 – 29 . The inflammatory response differs across types: low and transient in disuse 30 – 32 , progressively larger in denervation 33 , and greatest in degeneration 24 . Thus, disuse, denervation and degeneration share but also differ in molecular hallmarks. Mitochondria are a major source of reactive oxygen species (ROS) such as superoxide and its dismutation product H O 34 . Traditionally, mitochondrial ROS are linked to oxidative stress, where ROS production surpasses antioxidant defenses and damage proteins and lipids 35 , 36 . These processes have long been implicated in muscle wasting 35 – 37 . In disuse, several rodent and human studies observed increased mtH 2 O 2 , oxidative stress, and lipid peroxidation 31 , 37 – 41 , though some human studies found no changes 42 . Some rodent studies demonstrated protection against disuse atrophy via ROS-lowering interventions 38 , 39 , 43 , but others did not 40 . Denervation consistently elevates mtH 2 O 2 and oxidative stress markers in rodents 13 – 15 , 44 . Blocking cPLA2, 12/15 lipoxygenase or lipid peroxidation reduced denervation atrophy 14 , 16 , 44 , while overexpressing mitochondrial Catalase or Peroxiredoxin 3 did not prevent mtH 2 O 2 generation 16 . However, these interventions protected against muscle loss in SOD1KO sarcopenia mice 35 , 45 , though GPx4 overexpression reduced mtH 2 O 2 without preventing atrophy 46 . Similar modest benefits were reported in aging models 36 , 47 , suggesting that mtH 2 O 2 contributes to denervation but extra- mitochondrial factors are also involved. In degeneration, elevated oxidative stress and mtH 2 O 2 were reported in human myopathies 48 , 49 , mouse myopathy 22 , 50 and injury models 24 . In Duchenne muscular dystrophy, general antioxidants (Vitamin E, N-acetyl cysteine) were protective 51 , 52 , as were mitochondria-targeted strategies such as Catalase overexpression 53 , Olesoxime (blocking permeability transition pore) 54 , and mitochondrial transplantation 55 . Overall, many studies support that mitochondrial ROS and oxidative stress contribute to different types of muscle wasting. However, the specific mitochondrial ROS and their dose-dependent mechanistic roles remain unclear. To deepen our understanding of the spatial and temporal contribution of mtH2O2 to muscle wasting, we employed a chemogenetic model expressing D-amino acid oxidase in the mitochondrial matrix of skeletal muscle. This tool dose-dependently generates H 2 O 2 after administration of the DAAO substrate D-alanine in vitro and in vivo 56 – 59 allowing us, for the first time, to study the sufficiency of different degrees of mtH2O2 to trigger hallmarks of muscle wasting. Our findings indicate that mtH2O2 is sufficient to trigger molecularly distinct muscle wasting hallmarks at different exposure levels and acutely and selectively suppresses muscle protein synthesis at low concentrations without causing structural damage. 2. RESULTS 2.1 Mitochondrial H 2 O 2 manipulation via chemogenetic mtDAAO in skeletal muscle To determine whether mtH 2 O 2 per se is sufficient to cause skeletal muscle atrophy and to evaluate its dose dependency, we administered an adeno-associated virus (AAV) serotype 6 encoding a biosensor, HyPer3 and mtDAAO construct (HyPer3-mtDAAO) by intramuscular injection into the gastrocnemius and flexor digitorum brevis (FDB) muscles of adult C57BL/6N mice ( Fig. 1A ). At 25 days post-injection, muscles strongly expressed HyPer3-mtDAAO protein compared to the contralateral gelofusine-injected leg ( Fig. 1B ). Moreover, mtDAAO-transfected FDB single muscle fibers were isolated for confocal live-cell imaging ( Fig. 1C ). In these fibers, HyPer3 fluorescence visually ( Fig. 1D ) and quantitatively ( Fig. 1E, F ) overlapped with the signal of the hydrophobic mitochondria membrane potential (𝚫𝛙 m ) specific dye, PKmito orange, supporting mitochondrial localization of mtDAAO. Download figure Open in new tab Figure 1. Mitochondrial H 2 O 2 manipulation via chemogenetic mtDAAO in skeletal muscle. A) Schematic illustration of the experimental model and design. B) Representative western blot of GFP protein content in gastrocnemius muscles. C) Experimental outline of adult myofibers culture from Flexor Digitorum Brevis transduced with AAV6-mtDAAO. D) Representative confocal microscopy images of mitochondrial network (mitochondrial probe: PKmito) and GFP signal from genetically encoded biosensor HyPer3 fusion with DAAO enzyme (Scale bar 10μm). F) 2D histogram of colocalization analysis of PKmito and GFP and Manders’ coefficient of PKmito and GFP. G) Real-time images from isolated myofibers transfected with mtDAAO and stimulated with 10 mM of D-alanine. H) Isolated adult myofibers were stimulated with different concentrations of D-alanine and L-alanine, and the H 2 O 2 generated was measured using a genetically encoded biosensor HyPer3 fusion with DAAO enzyme. Representative quantification of ratiometric data from myofibers treated with 5mM and 10mM of D-alanine and 10mM of L-alanine for 15min. I) Representative image of transduction efficiency of AAV6 in gastrocnemius muscles (Scale bar 1000μm (top) and 100μm (bottom)). Because mtDAAO is fused to the H2O2-specific genetically encoded biosensor, HyPer3, we used its ratiometric fluorescence to monitor mtH2O2 generation in living single myofibers ( Fig. 1G ). Addition of D-alanine, but not L-alanine, led to a dose-dependent increase of HyPer3 signal, demonstrating both functionality and stereospecificity of the DAAO enzyme ( Fig. 1H ). We observed that HyPer3-mtDAAO expression in gastrocnemius muscle cryosections was regionally heterogeneous, with the highest levels observed close to the injection sites ( Fig. 1I ). Therefore, to enrich transduced tissue for downstream analyses, we dissected well-transduced areas of the gastrocnemius muscle during tissue collection, easily visible under a green fluorescent lamp (Fig. S1). Overall, these results demonstrate effective mitochondrial targeting and functional expression of mtDAAO-HyPer3 in mature skeletal muscle, enabling spatially and temporally controlled generation and real-time monitoring of mtH 2 O 2 within muscle fibers. 2.2 Graded H 2 O 2 exposure via mtDAAO reveals temporal progression of oxidative stress and muscle atrophy To determine whether sustained mtH 2 O 2 generation is sufficient to induce muscle wasting in vivo, we first performed a time-course experiment in which mice transduced unilaterally with mtDAAO in the gastrocnemius muscle were given 0.4M D-alanine in the drinking water for 1, 4, or 8 days ( Fig. 2A ). D-alanine did not alter body weight or fat mass, and led to a modest, time-dependent shift in whole-body lean mass, slightly reduced at day 1 and increased by day 8 ( Fig. 2B ). Food intake remained unchanged at concentrations up to 1M D-alanine (Fig. S2). Histological analysis of muscle cryosections revealed significant muscle fiber atrophy at days 4 (∼32%) and 8 (∼35%) in the mtDAAO-transduced leg compared to the contralateral leg ( Fig. 2C, D ). Download figure Open in new tab Figure 2. Graded H 2 O 2 exposure via mtDAAO reveals temporal progression of oxidative stress and muscle wasting. A) Experimental design of chronic 0.4 Molar D-alanine time course in mice unilaterally injected with mtDAAO. B) Body composition from day 0 to termination on day 8. C) Representative images of gastrocnemius muscles stained with WGA (Scale bar 100μm). D) Mean of the cross-sectional area (CSA) of gastrocnemius muscle after 1, 4, and 8 days of D-alanine administration (n=3 each group). E) Expression of 4-HNE and HO-1. F) Representative blots. H) Protein content of MuRF1, HDAC4, Ubiquitination, Akt, p62, NCAM1, αII-spectrin and Calpastatin (C-leg: contralateral leg). I) Representative blots. Data are shown as means and individual values. Statistical analyses were conducted using two-way ANOVA, and multiple comparisons were made with Tukeýs post hoc test. *p < 0.05, **p < 0.01, and ***p < 0.001, ****p < 0.0001. We next performed a detailed biochemical characterization of gastrocnemius muscle lysates from the 0.4M time-course using western blotting. The lipid peroxidation marker 4-hydroxynonenal (4- HNE) and the oxidative stress–responsive enzyme heme oxygenase-1 (HO-1) increased significantly by day 1 and continued to rise over time ( Fig. 2E, F ). Whereas a canonical hallmark of muscle atrophy, the MuRF1/Trim63 E3 ubiquitin ligase, showed a transient and early increase on day 1, the muscle atrophy-responsive protein Histone Deacetylase 4 (HDAC4) was strongly elevated from day 1 onwards ( Fig. 2G, H ). Additional markers of proteolysis, including ubiquitination, αII-spectrin (a substrate of redox-sensitive calpain proteases), and the autophagy- marker p62 were also elevated at day 1 and further increased over time ( Fig. 2G, H ). Calpastatin, the endogenous inhibitor of calpains 1 and 2, remained unchanged ( Fig. 2G, H ). Production of H 2 O 2 by mtDAAO stimulated markers of denervation, with NCAM showing a time-dependent increase at days 4 and 8 compared to the contralateral leg, while Akt, another denervation-responsive marker, was elevated only at day 8 ( Fig. 2G, H ). Thus, these data show that mtDAAO-induced mtH 2 O 2 production is sufficient to trigger early muscle oxidative stress, followed by time- dependent muscle atrophy, activation of multiple proteolytic systems, and induction of denervation- associated markers. 2.3 Muscle damage and regeneration induced by chemogenetic mtH 2 O 2 generation at increased substrate concentrations To investigate whether sustained mtH 2 O 2 generation elicits muscle damage and whether these changes are reversible through established physiological muscle regeneration mechanisms, we examined muscle histology in animals exposed to either 0.4 M or 1.0 M D-alanine for 8 days. Histological analyses revealed extensive muscle damage at both concentrations, with no statistically significant difference between doses, suggesting that 0.4 M elicits similar levels of muscle damage as 1.0 M under these conditions (Fig. S3A–C). Among molecular markers, NCAM1 protein content was significantly increased at 0.4 M and showed a non-significant trend at 1.0 M, likely reflecting increased variability in the mtDAAO-transduced limb. Cleaved caspase-3 levels were robustly elevated at both doses, supporting activation of mtH 2 O 2 -induced apoptotic signaling. Calpastatin protein content was significantly increased only at 1.0 M. Finally, αII spectrin protein expression was significantly elevated at 0.4 M and trended upward at 1.0 M (Fig. S3C). Given the pronounced degeneration at higher doses of D-alanine, we next tested whether this damage was reversible and triggered a physiological regenerative response. For this, mice were unilaterally transduced with mtDAAO in the gastrocnemius muscle and administered 0.4 M D-alanine in drinking water for 8 days, followed by a recovery phase on regular drinking water for either 7 or 14 days ( Fig. 3A ). Histological analysis revealed that ∼50% of the muscle-area was damaged compared with the contralateral control leg after 8 days of D-alanine, followed by progressive recovery at days 7 and 14 of the washout phase ( Fig. 3B, C ). Protein marker analysis suggested that regeneration was still ongoing at day 14 ( Fig. 3D, E ). Download figure Open in new tab Figure 3. Muscle damage and regeneration induced by chemogenetic mitochondrial H 2 O 2 generation at increased substrate concentrations. A) Experimental design: mice were unilaterally injected with mtDAAO received first 0.4 Molar D-alanine via drinking water for 8 days, then regular drinking water (RDW) for 7 days and 14 days. B) Representative images of Hematoxylin and Eosin (H&E) staining showing the damaged area (yellow dotted line) at day 0 and 8 of chronic D-alanine in drinking water and day 7 and 14 after return to regular drinking water (Scale bar 100μm). C) Quantification of damaged area per total area (n=4 each group). D) Protein content of MyoD, Myogenin, Akt, Hsp27, Hsp27, STAT3 (C-leg: contralateral leg). E) Representative blots. Data are shown as means and individual values. Statistical analyses were conducted using two-way ANOVA, and multiple comparisons were made with Tukey post hoc test. *p < 0.05, **p < 0.01, and ***p < 0.001, ****p < 0.0001. Chemogenetically induced muscle damage reduced the early differentiation marker MyoD, similar to what has been shown before in human muscle-derived myoblasts 60 . MyoD levels remained low at day 7 and returned to baseline by day 14. In contrast, a late differentiation marker, Myogenin 60 , was unchanged across all time points. The satellite-cell marker Pax7 61 , 62 increased markedly after 8 days of mtH 2 O 2 exposure and remained elevated through day 14 of recovery. The redox responsive chaperones Hsp27 and Hsp90, both implicated in muscle regeneration 63 – 65 , were significantly elevated after 8 days of D-alanine treatment. Upon switching to D-alanine free water, both Hsp27 and Hsp90 peaked at day 7 and returned to baseline by day 14 of recovery. STAT3, a transcription factor responsive to inflammation signaling and muscle repair in myocytes and satellite cells 33 , 62 , 66 , 67 , increased markedly throughout the exposure and early recovery phases. Finally, the denervation markers NCAM1 68 – 70 and Akt 12 were significantly elevated after 8 days of mtH2O2 exposure, remained high at day 7 of recovery and returned to basal levels by day 14. Thus, chemogenetic mtH2O2 production at high substrate concentrations induces pronounced, yet reversible myofiber degeneration. This mtDAAO model provides a powerful experimental tool to dissect how mtH2O2 us involved in muscle degeneration and regeneration in adult skeletal muscle. 2.4 Low chemogenetic mtH 2 O 2 generation triggers muscle atrophy without structural damage An emerging concept is that ROS, in particular the diffusible and selectively reactive H2O2, can act as redox second messengers via specific and reversible protein cysteine oxidation 71 . This so-called redox signaling is often framed within the concept of redox hormesis where low H2O2 triggers beneficial adaptation at low exposure 72 – 74 and causes damage at high exposure 75 (e.g. in response to exercise) but specific redox signaling might also contribute negatively to muscle wasting. Therefore, we also characterized the effect of lowering the D-alanine dose to 0.2M, taking the muscle at day 4 ( Fig. 4A ). Here, a 36% reduction in CSA was observed in the transfected vs. non- transfected contralateral control muscle, with no visible structural damage ( Fig. 4B, C ) or ultrastructural changes in muscle fiber and mitochondrial morphology ( Fig. 4D ). Western blot analysis revealed a small and variable ∼ 30% increase in 4-HNE (p=0.04) by mtDAAO activation, whereas HO-1 was not different ( Fig. 4E, F ), suggesting the presence of oxidative stress, albeit to a lesser extent than with higher D-alanine doses, and an absent antioxidant defense response. Effects on atrogene and proteolysis markers were similarly modest, with significant increases in ubiquitination (∼20%) and calpastatin (∼15%), and notably no increases in the denervation- associated NCAM or Akt ( Fig. 4G, H ). Activation of mtDAAO did not increase markers of stress kinase activation, including the phosphorylation of the integrated stress response pathway marker eIF2α Ser51, the energy-stress kinase AMPK Thr172, the mTORC1 substrate p70S6K Thr389 or the stress-kinase p38 MAPK Thr180/Tyr182 (Fig. S5A-B). The magnitude of atrophy resembled that observed after 5 days of unilateral hindlimb immobilization (U-Imm) using a 3D-printed splint model, which reduced CSA by 27% (Fig. S4A–C), and caused modest but significant induction of oxidative stress and canonical atrophy markers (Fig. S4D-G). Thus, chronic mtH 2 O 2 generation at lower concentrations triggers a degree of muscle fiber atrophy reminiscent of immobilization without accompanying structural damage, antioxidant induction, general stress kinase activation, or denervation-hallmarks. Download figure Open in new tab Figure 4. Chronic chemogenetic mtH 2 O 2 generation at a non-damaging substrate concentration causes muscle wasting resembling disuse. A) Experimental design: Mice unilaterally injected with mtDAAO drank 0.2 Molar D-alanine water for 4 days. B) Representative images of gastrocnemius muscles stained with WGA (Scale bar 100μm). C) Mean cross-sectional area (CSA) of mtDAAO-transfected vs. contralateral (C-leg) gastrocnemius muscle after 4 days (n=15). D) Representative electron microscopy images of gastrocnemius muscles (Scale bar 2μm). E) Expression of 4-HNE and HO-1. F) Representative blots. G) Protein content of MuRF1, HDAC4, Ubiquitination, Akt, p62, NCAM1, αII-spectrin and Calpastatin. H) Representative blots. I) Experimental design: Mice underwent either unilateral immobilization or mice expressing mtDAAO unilaterally received 0.2 Molar of D-alanine in drinking water for 24 hours, after which protein synthesis was measured for 15 min using Puromycin. J) Quantified puromycin incorporation in the different conditions. K) Representative blots. Data are shown as means and individual values. Statistical analyses were conducted using a Wilcoxon matched-pairs test. *p < 0.05, **p < 0.01, and ***p < 0.001, ****p < 0.0001. 2.5 Low chemogenetic mtH 2 O 2 generation acutely suppresses protein synthesis Decreased protein synthesis is believed to be the principal determinant of disuse muscle atrophy in humans 76 . In mice, 6-24h of hindlimb immobilization rapidly inhibits protein synthesis prior to the upregulation of proteolysis 9 . Consistent with this, we observed a 14% reduction in puromycin labeling in U-Imm vs. contralateral control muscles after 24 h ( Fig. 4J, K ). Because our 0.2 M D- alanine intervention over 4 days produced a 36% CSA reduction with only mildly increased ubiquitination, we next asked whether low-level mtH 2 O 2 production was sufficient to suppress protein synthesis. Indeed, 24h exposure to 0.2M D-alanine significantly decreased protein synthesis ( Fig. 4J, K ). These data indicate that non-damaging low-level mtH2O2 generation is sufficient to acutely suppress protein synthesis, an effect that likely contributes to the muscle atrophy observed after 4 days. 3. DISCUSSION This study leveraged a chemogenetic model to achieve precise, compartment-specific control of mtH 2 O 2 production, for the first time, in adult skeletal muscle of living mice. By titrating DAAO activity within the mitochondrial matrix, we were able to assess for the first time how graded elevations in mtH 2 O 2 influence muscle fiber size and proteostasis in vivo. Strikingly, we observed that even the lowest dose of mtH 2 O 2 was sufficient to rapidly impair protein synthesis and reduce muscle fiber size without inducing obvious structural damage or overt oxidative stress. Increasing levels of mtH 2 O 2 triggered molecular signatures consistent with denervation, proteolysis, and eventually fiber degeneration, followed by regenerative myogenesis. These findings support the concept that mtH 2 O 2 can act as a dose-dependent regulator of distinct muscle remodeling programs. The etiology of human disuse atrophy is consistently linked to decreased protein synthesis rather than increased proteolysis with interventions up to 14 days 42 , 77 , 78 . This pattern resembles what is observed within the first day after immobilization in mice 9 . Similarly, the rate of disuse atrophy is much faster in rodents than in humans (CSA decreases of ∼4% day −1 in rodents vs. ∼0.4% day −1 in humans 42 , 76 – 78 ) . Hence, our demonstration that low mtH2O2 suppresses protein synthesis after 24h might be highly relevant mechanism to human disuse prevention and treatment. Earlier studies in humans measured only relatively insensitive oxidative stress markers 31 , 42 . Furthermore, daily ingestion of an anti-inflammatory/antioxidative cocktail significantly lowered protein carbonylation but did not protect against muscle atrophy after 60 days 79 . However, our present results linking mitochondrial redox-changes to acute inhibition of protein synthesis with only mild oxidative stress in mice, highlight the need for further studies to determine whether more subtle mitochondrial redox signaling inhibits protein synthesis in early human disuse and thereby contributes to muscle atrophy. How might mitochondrial matrix H2O2 inhibit cytosolic protein synthesis? Overall, H2O2 produced in the mitochondrial matrix could either act directly on cytosolic protein synthesis or indirectly via secondary mediators. Supporting the first notion, studies in permeabilized muscle fibers have proposed that H2O2 can be released from the mitochondria into the cytosol 80 . Furthermore, cytosolic H2O2 and other ROS are known to directly inhibit multiple regulatory steps of protein translation in different eukaryotic cell types 81 . In permeabilized muscle fibers from different rodent skeletal muscle atrophy models, mtH2O2 release was observed using ex vivo mitochondrial respirometry with detection by Amplex Red 16 , 38 , 50 , 82 . Whether such diffusion occurs in intact cells remains unclear, as mitochondrial matrix antioxidant systems are likely compromised to some extent in permeabilized muscle fibers 80 , and Amplex Red may not be specific to H 2 O 2 but also detect lipid peroxides 16 , 83 . In intact live cells, the degree of mtH 2 O 2 release into the cytosol differs between different cell types and appears tightly linked to the cytosolic Peroxiredoxin-thioredoxin reductase antioxidant defense capacity 84 . It is unclear at what dosage mtH 2 O 2 escapes skeletal muscle mitochondria in vivo, warranting further investigation into whether mtH 2 O 2 might act as a direct signal in proximity to mitochondria to suppress protein synthesis. In terms of indirect mechanisms, mitochondrial matrix H2O2 might stimulate one or more mitochondrial stress-response pathways to inhibit protein synthesis either via lowering of cytosolic protein translation or by retrograde signaling to the nucleus 84 . However, with 0.2M DAAO stimulation protocol, we did not detect differences in any markers of cellular stress pathways. Chowdhury et al. (2020) reported in three different cell types including C2C12 myoblasts that mitochondria-targeted paraquat (MitoPQ) and metformin (MitoMet) induce mitochondrial superoxide production, leading to dose-dependent activation of the Ca²⁺/calcineurin pathway, while having no significant impact on the HIF1α or AMPK retrograde pathways 85 . This suggests that different mitochondrial stress-response pathways have different sensitivities to ROS. Calpains are Ca 2+ and redox-sensitive proteases activated by disuse in rodents 86 . Interestingly, the suppression of protein synthesis after 12h of mechanical ventilation in rat diaphragm muscle was rescued by overexpression of the calpain 1 and 2 inhibitor calpastatin via protection against cleavage of aminoacyl-tRNA synthetase 87 . Thus, low-level DAAO activation might suppress protein synthesis via non-canonical mitochondrial stress signaling. Sustained moderate mtH2O2 exposure increased skeletal muscle NCAM and Akt expression in our study, suggestive of muscle fiber denervation. Previous studies in different animal models suggest that denervation provokes a rapid and sustained increase in both presynaptic and post-synaptic mitochondrial ROS production 35 , 88 – 90 . This might occur because of synaptic inactivity, lowering ATP demand and thereby increasing mitochondrial proton motive force and electron transport chain slippage 88 . Our data show that mtH2O2 release may both be a cause and an effect of denervation. Interestingly, prolonged human bedrest causes NMJ instability in humans and this response appears to be preceded by altered NMJ-specific mitochondrial phenotype after 10 days 91 . In this context, inactivity-associated NMJ-specific mtH2O2 production might be a trigger of denervation. With high and prolonged mtDAAO stimulation, we observed marked fiber degeneration accompanied by regenerative myogenesis, with robust induction of Pax7, MyoD, STAT3, and redox-responsive chaperones (Hsp27 and Hsp90). Given the many mechanistic links between mitochondria and cell death 92 , it is fully expected that high exposure to a mitochondrial-targeted stressor provokes this response. For reference, other in vivo studies performed chronic DAAO stimulation in the cytosol of vascular endothelial cells, brain neurons or cardiomyocytes using 1M D-alanine administration in drinking water, similar to our highest to link H2O2-production to different pathological changes 58 , 59 , 93 . In contrast, another study expressing DAAO in the nucleus of mouse cardiomyocytes used a much lower 55mM concentration to identify an H2O2-regulated mitochondrial redox signaling mechanism regulating energy metabolism 94 . Depending on the purpose of the study (studying a non-specific oxidative stress response vs. specific H2O2 redox signaling), the level of H2O2 exposure should be considered. Nonetheless, the current degeneration- regeneration model may be useful to understand the isolated role of mitochondrial redox changes in triggering or exacerbating different types of cell death (apoptosis, pyroptosis, ferroptosis and necroptosis) and regeneration, particularly in diseases where the involvement of mtH2O2 has been proposed, e.g. different myopathies. Limitations of the study While the mtDAAO system enables unique spatial and temporal control of H 2 O 2 production and supports a contralateral control design, several limitations should be acknowledged. Firstly, transduction was regionally heterogeneous within the transduced gastrocnemius muscle, although we circumvented this shortcoming by restricting our analysis to well-transduced areas (Fig. S1). Secondly, while AAV6 does not transduce satellite cells 95 , it may affect other non-myofiber populations such as endothelial or immune cells which may contribute to the observed phenotype. Thirdly, DAAO is expressed in select mammalian tissues, and although endogenous D-amino acids are scarce 96 and D-alanine had minimal effects in non-transduced mice, off-target activation of endogenous DAAO cannot be fully excluded. Finally, DAAO activity also generates ammonia and pyruvate alongside H 2 O 2 . Although these products are unlikely to perturb bulk metabolism or pH 97 – 99 in skeletal muscle fibers, their compartment-specific effects in mitochondria cannot be ruled out. Thus, while this system offers precise temporal and spatial control of mtH 2 O 2 in vivo, these potential confounders should be kept in mind. CONCLUSION This study presents a novel in vivo chemogenetic approach that allows precise control of mitochondrial matrix H 2 O 2 levels in adult skeletal muscle, offering the first direct evidence that graded mitochondrial elevations of H 2 O 2 can modulate muscle size and proteostasis. We show that even subtoxic increases in mtH 2 O 2 are sufficient to rapidly suppress protein synthesis and induce a disuse-like atrophic phenotype without causing structural damage. In contrast, higher exposures activate proteolytic and denervation-associated pathways and ultimately muscle fiber degeneration and regeneration. By enabling tightly controlled redox manipulation in vivo, this platform provides a powerful tool to explore redox-regulated mechanisms of muscle plasticity and guide the search for therapeutic strategies in muscle-wasting disorders. MATERIALS AND METHODS Animals Mice were single-housed, maintained on a 12:12-h light-dark cycle, at 21 °C, and received a standard rodent chow diet (Altromin no. 1324; Chr. Pedersen, Denmark) and water ad libitum. All experiments were approved by the Bioethics Committee of the Danish Animal Experimental Inspectorate (license: 2022-15-0201-01319) and complied with the “European Convention for the Protection of Vertebrate Animals Used for Experiments and Other Scientific Purposes.” Unilateral immobilization model C57BL/6N male mice, aged 10-12 weeks old, were randomized into control and immobilized groups and single-housed. Hindlimbs were unilaterally immobilized at both the knee and ancle joint in a plantarflexed position using a 3D-printed splint for 1 and 5 days. Control mice were sham- handled but not treated. At the end of each experiment, animals were euthanized with pentobarbital anesthesia and cervical dislocation, and the gastrocnemius was dissected, rapidly frozen in liquid N 2 , and stored at −80°C until processing. The muscles were dissected and embedded in an optimum cutting temperature medium from Tissue-Tek, frozen in melting isopentane and kept at −70 °C until processing. 3D printed splints The splints were printed using a FLASHFORGE FDM 3D Printer Adventure 3 with 1.75mm transparent PLA filament. AVV6-mtDAAO-HyPer3 transduction via direct intramuscular injection Mice were anesthetized with 2.5% isoflurane. The fur of the hindlimb to be injected was removed using clippers. The hindlimb was held firmly distal to the injected area, and the needle was inserted into the center of the muscle group from the distal end. Mice were unilaterally injected with the AVV6-mtDAAO-HyPer3 construct diluted in Gelofusine, while the contralateral limb received an equal volume of Gelofusine alone as vehicle control. Gastrocnemius and FDB muscles were injected with 35-40 μl and 10-15 μl volumes per muscle, respectively, delivering a total dose of 4 x 10^ 10 viral genomes using an ultra-fine insulin syringe with a half-unit scale 31G x 6 mm, 3/10 mL, as previously described 100 . DAAO-transduced muscle fibers were visualized using a NIGHTSEA Royal Blue light source (excitation: 440–460 nm) combined with a green filter (ISO 100, aperture ƒ/1.8, and exposure time 1/100 s). In vivo and in vitro mtDAAO activation For chemogenetic in vivo and in vitro activation of mtDAAO, the substrates D-alanine (Sigma- Aldrich, CAS: 338-69-2) was dissolved either in drinking water at final concentrations of 0.2 M, 0.4 M, or 1.0 M, or in sterile Dulbecco’s Phosphate-Buffered Saline (Gibco™ Cat #15326239), respectively 57 – 59 , 101 – 103 . Mice had ad libitum access to the D-alanine–supplemented water for defined experimental durations ranging from 1 to 8 days. Water bottles were replaced every 2–3 days to ensure substrate stability and consistent intake. L-alanine (Sigma-Aldrich, CAS: 56-41-7) was prepared and administered the same. Body composition Total body mass, fat mass, and lean tissue assessments were performed through nuclear magnetic resonance using an EchoMRI 4-in-1 500 (EchoMRI, USA). FDB fiber isolation FDB muscle fibers were obtained through enzyme digestion of whole muscle using collagenase type II (3 mg/ml) (ThermoFisher, Cat #17101015) for 120 minutes at 37 °C, followed by mechanical dissociation with fire-polished Pasteur pipettes. Isolated fibers were seeded in ECM Gel-coated (Sigma-Aldrich, Cat #E6909) cell culture dishes in Alpha MEM supplemented with 10% fetal bovine serum and 1% antibiotic-antimycotic solution. After 16 hours of seeding, the fibers were utilized for experimentation. Live imaging FDB fibers were imaged in a Hanks′ Balanced Salt Solution (Sigma-Aldrich, Cat #D8537) supplemented with 15 mM HEPES while kept at 95% O2 and 5% CO2 at 37 °C using a Pecon Lab- Tek S1 heat stage. Fibers were loaded with PKmito Orange fluorescence (Spirochrome, Switzerland) for 30 minutes before imaging following the manufacturer’s instructions Imaging and image analysis For live-imaging experiments, confocal images were collected using a ×63 1.4 NA oil immersion objective lens on an LSM 980 confocal microscope (Zeiss). PKmito ORANGE fluorescence was detected using the excitation-emission λ591–608 nm. For the HyPer3 biosensor images, raw data of the λ420-nm and 500-nm laser lines were exported to ImageJ as 16-bit TIFFs for further analysis. Data are presented as fluorescence ratio (λ500/420 nm) normalized to the resting WT group. For fiber size stainings, images were collected using a dry ×20 0.5 NA Plan-Neofluar objective on an AXIO Imager.M2 (Zeiss). The excitation laser line used was 488 nm. Images were exported to ImageJ. The images were analyzed in Fiji software and Cellpose as described by Waisman et al. 104 . Four areas in one mid-muscle-belly cryosection were analyzed. On average > 50 fibers/area were quantified. All image collections and quantifications were performed blinded. Cryosections and stainings Gastrocnemius muscles were embedded in Tissue-Tek, frozen in liquid nitrogen-cooled isopentane and stored at −70 °C. Frozen muscles were transferred to a cryostat chamber and allowed to equilibrate to −20 °C. Transverse sections 10μm were cut and mounted on positively charged glass slides (ThermoFisher Scientific). Samples were fixed in 4% paraformaldehyde for 10 min, followed by three washes in phosphate-buffered saline (PBS). Green fluorescence-conjugated wheat germ agglutinin (WGA-Alexa fluor 488, ThermoFisher #W11261) (1:100 dilution), and Laminin (1:300) along with goat-anti-rabbit IgG2b, conjugated with Alexa 647 fluorophore were used to stain the surface membranes. Electron microscopy Gastrocnemius muscle specimens were prepared for transmission electron microscopy as described by Jensen et al. (2022) 105 . Briefly, a small part of the muscle (<mm 3 ) was fixed in 2.5% glutaraldehyde in 0.1 M sodium cacodylate buffer (pH 7.3) for 24 h. It was then washed four times (15 min between each wash) in 0.1 M sodium cacodylate buffer (pH 7.3) and stored at 5 °C until further preparation. The specimens were then post-fixed with 1% osmium tetroxide (OsO4) and 1.5% potassium ferrocyanide (K4Fe(CN)6) in 0.1 M sodium cacodylate buffer (pH 7.3), dehydrated, and embedded in resin. Ultra-thin (60-70 nm) sections were cut of longitudinally oriented fibers and contrasted with uranyl acetate and lead citrate. Images were acquired using a Philips CM100 transmission electron microscope (Philips) and an Olympus Veleta camera (Olympus Soft Imaging Solutions). Western Blots Frozen gastrocnemius muscle tissue was homogenized in lysis buffer (0.05 M Tris Base pH 7.4, 0.15 M NaCl, 1 M EDTA and EGTA, 0.05 M; sodium fluoride, 5 mM sodium pyrophosphate, 2 mM sodium orthovanadate, 1 mM benzamidine, 0.5% protease inhibitor cocktail (P8340, Sigma Aldrich), and 1% NP-40) supplemented with 100 mM NEM to prevent cysteine oxidation. The homogenates were rotated end over end for 30 minutes at 4 °C and then subsequently centrifuged at 18,320g for 20 minutes at 4 °C. Supernatants were transferred to new tubes and the total protein concentration measured using the bicinchoninic acid assay method. The samples were diluted in dH2O and a non-reducing 6x Laemmli sample buffer (Tris Base pH 6.8 340 mM, SDS 11%, Glycerol 20%, Bromophenol blue 0.05%) to a protein concentration of 1-2 mg/ml. The samples were then loaded onto an SDS-PAGE stacking gel (4%) cast upon an SDS-PAGE running gel (5- 12%) and kept in running buffer (Tris-base pH=7.4 25 mM, Glycine 190 mM, SDS 10%). For stacking gel alignment, the gel was run at 90V and 0,06A for approximately 15 minutes. After alignment, the gel was run at 120-130V and 0,06A for approximately 1 hour, until a loaded Precision Plus Standard All Blue Marker (Bio-Rad) indicated a good separation of proteins. After separation, the gel was cut to molecular weight regions of interest. The proteins were then transferred from the separation gel to PVDF membrane (Millipore), which had been cut to accommodate the gel, ethanol-activated and soaked in transfer buffer (Tris-base pH=9 50 mM, Glycine 40 mM, SDS 0.015%, 20% Ethanol). The protein transfer was performed using a Trans- Blot Turbo Machine (Transfer System, Bio-Rad) running at 20V for 30 minutes. To prevent unspecific antibody binding, the membranes were blocked for 1 hour at room temperature in 2-3% skimmed milk powder diluted in Tris-buffered saline (Tris-base pH=7.4, NaCl 150 mM) with 0,05% Tween-20 (TBST). The membranes were incubated overnight at 4 °C with the primary antibody (see Table 1). Membranes were washed 3 x 15 minutes in TBS-T and incubated for 1 hour minutes in horseradish-peroxidase conjugated secondary antibody diluted 1:5000 in 3% skimmed milk and TBS-T. Membranes were washed 3 times, 15 minutes in TBST. Horseradish peroxidase ECL substrate was added to the membrane for 1 minute, and visualization of the protein performed using a ChemiDocTM MP Image System (Bio-Rad). Hematoxylin & Eosin staining Gastrocnemius muscles were embedded in Tissue-Tek, frozen in liquid nitrogen-cooled isopentane and stored at −70 °C. Frozen muscles were transferred to a cryostat chamber and allowed to equilibrate to −20 °C. Ten μm thick cryosections were cut and mounted on positively charged glass slides (ThermoFisher Scientific). Briefly, samples were deparaffinized and hydrated with distilled water. Samples were then submerged in Hematoxylin solution for 5 minutes, followed by two rinses in distilled water. The samples were then incubated in bluing reagent for 10-15 seconds and rinsed in distilled water twice. Afterwards, samples were dipped in absolute alcohol before submersion in Eosin solution for 3 minutes. Finally, the samples were rinsed in absolute alcohol, dehydrated through three changes of absolute alcohol, and mounted in synthetic resin. Statistical analysis The experimental data were presented as means overlayed on the individual data points of several experiments indicated as (n) or representative results from at least three independent determinations. Statistical comparisons between groups were carried out using the Wilcoxon matched-pairs test or Two-way ANOVA as applicable. Tukeýs post hoc test was performed for multiple group-wise comparisons when ANOVA revealed significant main effects or interactions. GraphPad Prism 10 software was used for statistical analysis. (GraphPad Software, San Diego, CA). A value p<0.05 was considered a statistically significant difference. S1. Whole-muscle transfection visualized by GFP fluorescence under a fluorescent lamp. A) PBS injected and AAV6-HyPer3-DAAO transduced muscles exposed to a green fluorescent lamp. S2. Body composition and food intake of mice exposed to high doses of D-alanine. A) Experimental design of 1.0 M L-alanine and D-alanine in mice unilaterally injected with mtDAAO. A) Body composition and C) Food intake of mice exposed to 1.0M L-alanine and D-alanine drinking water. S3. High doses of D-alanine induced massive damage in muscle in vivo. A) Experimental design: Mice unilaterally expressing mtDAAO drank 1.0 and 0.4M D-alanine water for 8 days. B) Representative images of Hematoxylin and Eosin staining showing the damaged area (yellow dotted line) at day 0 and day 8 of receiving 1.0M or 0.4M D-alanine (Scale bar 100μm). C) Quantification of damaged area per total. Statistical analyses were conducted using a Mann- Whitney test. *p < 0.05. D) Protein expression of NCAM1, Pro-Caspase3, Cleaved-Caspase3, Calpastatin, αII-spectrin (C-leg: contralateral leg). E) Representative blots. Data are shown as means and individual values. Statistical analyses were conducted using a Wilcoxon matched- pairs test. *p < 0.05, **p < 0.01, and ***p < 0.001, ****p < 0.0001. S4. Characterization of the unilateral immobilization disuse atrophy model. A) Experimental design: Mice were unilaterally immobilized using a 3D printed plastic splint for 5 days. B) Representative images of gastrocnemius muscles stained with WGA (Scale bar 100μm). C) Mean cross-sectional area (CSA) of immobilized vs. contralateral gastrocnemius muscle after 5 days (n=5). D) Protein content of 4-HNE and HO-1. E) Representative blot. F) Expression of MuRF1, HDAC4, Ubiquitination, Akt, p62, NCAM1, αII spectrin and Calpastatin (C-leg: contralateral leg). Data are shown as means and individual values. Statistical analyses were conducted using a Wilcoxon matched-pairs test. *p < 0.05, **p < 0.01, and ***p < 0.001, ****p < 0.0001. S5. Low dosage of chronic mtH2O2 generation does not stimulate stress kinases. A) Experimental design: Mice unilaterally expressing mtDAAO received 0.2 Molar D-alanine for 4 days. B) Phosphorylation of eIF2α Ser51, AMPK Thr172, p70S6K Thr389, p38 MAPK Thr180/Tyr182. C) Representative blots. Data are shown as means and individual values. Statistical analyses were conducted using a Wilcoxon matched-pairs test. *p < 0.05, **p < 0.01, and ***p < 0.001, ****p < 0.0001. Acknowledgements The electron microscopy was performed at the Core Facility for Integrated Microscopy, Faculty of Health and Medical Sciences, University of Copenhagen. References 1. ↵ Evans , W. J . Skeletal muscle loss: cachexia, sarcopenia, and inactivity . Am J Clin Nutr 91 , 1123S – 1127S ( 2010 ). OpenUrl Abstract / FREE Full Text 2. ↵ Kim , T. N. & Choi , K. M . Sarcopenia: definition, epidemiology, and pathophysiology . J Bone Metab 20 , 1 – 10 ( 2013 ). OpenUrl CrossRef PubMed 3. ↵ 3. Cavalcante , B. R. , Falck , R. S. & Liu-Ambrose , T . Editorial: ‘May the Force (and Size) Be with You’: Muscle Mass and Function Are Important Risk Factors for Cognitive Decline and Dementia . J Nutr Health Aging 27 , 926 – 928 ( 2023 ). OpenUrl 4. Cui , M. , Zhang , S. , Liu , Y. , Gang , X. & Wang , G . Grip Strength and the Risk of Cognitive Decline and Dementia: A Systematic Review and Meta-Analysis of Longitudinal Cohort Studies . Front Aging Neurosci 13 , 625551 ( 2021 ). 5. ↵ Sartori , R. , Romanello , V. & Sandri , M . Mechanisms of muscle atrophy and hypertrophy: implications in health and disease . Nat Commun 12 , 330 ( 2021 ). 6. ↵ Wilburn , D. , Ismaeel , A. , Machek , S. , Fletcher , E. & Koutakis , P . Shared and distinct mechanisms of skeletal muscle atrophy: A narrative review . Ageing Res Rev 71 , 101463 ( 2021 ). 7. ↵ Iannuzzi-Sucich , M. , Prestwood , K. M. & Kenny , A. M . Prevalence of sarcopenia and predictors of skeletal muscle mass in healthy, older men and women . J Gerontol A Biol Sci Med Sci 57 , M772 – 777 ( 2002 ). OpenUrl CrossRef PubMed Web of Science 8. ↵ You , J.-S. , Anderson , G. B. , Dooley , M. S. & Hornberger , T. A . The role of mTOR signaling in the regulation of protein synthesis and muscle mass during immobilization in mice . Dis Model Mech 8 , 1059 – 1069 ( 2015 ). OpenUrl Abstract / FREE Full Text 9. ↵ Lin , K.-H. et al. A deep analysis of the proteomic and phosphoproteomic alterations that occur in skeletal muscle after the onset of immobilization . J Physiol 599 , 2887 – 2906 ( 2021 ). OpenUrl CrossRef 10. ↵ Sirago , G. et al. Upregulation of Sarcolemmal Hemichannels and Inflammatory Transcripts with Neuromuscular Junction Instability during Lower Limb Unloading in Humans . Biology (Basel ) 12 , 431 ( 2023 ). 11. ↵ Murgia , M. et al. Signatures of muscle disuse in spaceflight and bed rest revealed by single muscle fiber proteomics . PNAS Nexus 1 , pgac086 ( 2022 ). 12. ↵ Li , Z. et al. Chemical denervation using botulinum toxin increases Akt expression and reduces submaximal insulin-stimulated glucose transport in mouse muscle . Cell. Signal . 53 , 224 – 233 ( 2019 ). OpenUrl 13. ↵ Scalabrin , M. et al. Redox responses in skeletal muscle following denervation . Redox Biol 26 , 101294 ( 2019 ). 14. ↵ Brown , J. L. et al. Lipid hydroperoxides and oxylipins are mediators of denervation induced muscle atrophy . Redox Biol 57 , 102518 ( 2022 ). 15. ↵ Pollock , N. , Staunton , C. A. , Vasilaki , A. , McArdle , A. & Jackson , M. J . Denervated muscle fibers induce mitochondrial peroxide generation in neighboring innervated fibers: Role in muscle aging . Free Radic. Biol. Med . 112 , 84 – 92 ( 2017 ). OpenUrl 16. ↵ Pharaoh , G. et al. Targeting cPLA2 derived lipid hydroperoxides as a potential intervention for sarcopenia . Sci Rep 10 , 13968 ( 2020 ). 17. ↵ You , J.-S. , Kim , K. , Steinert , N. D. , Chen , J. & Hornberger , T. A . mTORC1 mediates fiber type-specific regulation of protein synthesis and muscle size during denervation . Cell Death Discov 7 , 74 ( 2021 ). 18. ↵ Massopust , R. T. et al. Lifetime analysis of mdx skeletal muscle reveals a progressive pathology that leads to myofiber loss . Sci Rep 10 , 17248 ( 2020 ). 19. Hughes , M. C. et al. Mitohormesis during advanced stages of Duchenne muscular dystrophy reveals a redox- sensitive creatine pathway that can be enhanced by the mitochondrial-targeting peptide SBT-20 . Redox Biol 76 , 103319 ( 2024 ). 20. Rando , T. A. , Disatnik , M. H. , Yu , Y. & Franco , A . Muscle cells from mdx mice have an increased susceptibility to oxidative stress . Neuromuscul Disord 8 , 14 – 21 ( 1998 ). OpenUrl CrossRef PubMed Web of Science 21. Sugihara , H. et al. Cellular senescence-mediated exacerbation of Duchenne muscular dystrophy . Sci Rep 10 , 16385 ( 2020 ). 22. ↵ Xiao , Y. et al. ROS-related mitochondrial dysfunction in skeletal muscle of an ALS mouse model during the disease progression . Pharmacol Res 138 , 25 – 36 ( 2018 ). OpenUrl CrossRef PubMed 23. ↵ Klickovic , U. et al. Muscle MRI quantifies disease progression in amyotrophic lateral sclerosis . J Neurol Neurosurg Psychiatry jnnp-2024-335571 ( 2025 ) doi: 10.1136/jnnp-2024-335571 . OpenUrl Abstract / FREE Full Text 24. ↵ Ramadasan-Nair , R. et al. Mitochondrial alterations and oxidative stress in an acute transient mouse model of muscle degeneration: implications for muscular dystrophy and related muscle pathologies . J Biol Chem 289 , 485 – 509 ( 2014 ). OpenUrl Abstract / FREE Full Text 25. ↵ Hardy , D. et al. Comparative Study of Injury Models for Studying Muscle Regeneration in Mice . PLoS One 11 , e0147198 ( 2016 ). OpenUrl CrossRef PubMed 26. ↵ Sicherer , S. T. , Venkatarama , R. S. & Grasman , J. M . Recent Trends in Injury Models to Study Skeletal Muscle Regeneration and Repair . Bioengineering (Basel ) 7 , 76 ( 2020 ). 27. Ikeda , T. et al. Degenerative and regenerative features of myofibers differ among skeletal muscles in a murine model of muscular dystrophy . J Muscle Res Cell Motil 37 , 153 – 164 ( 2016 ). OpenUrl 28. Anderson , J. E. , Bressler , B. H. & Ovalle , W. K . Functional regeneration in the hindlimb skeletal muscle of the mdx mouse . J Muscle Res Cell Motil 9 , 499 – 515 ( 1988 ). OpenUrl CrossRef PubMed Web of Science 29. ↵ Boldrin , L. , Zammit , P. S. & Morgan , J. E . Satellite cells from dystrophic muscle retain regenerative capacity . Stem Cell Res 14 , 20 – 29 ( 2015 ). OpenUrl CrossRef PubMed 30. ↵ Hirata , Y. et al. A Piezo1/KLF15/IL-6 axis mediates immobilization-induced muscle atrophy . J Clin Invest 132 , 1 – 13 ( 2022 ). OpenUrl CrossRef 31. ↵ Eggelbusch , M. et al. The impact of bed rest on human skeletal muscle metabolism . Cell Rep Med 5 , 101372 ( 2024 ). 32. ↵ Caron , A. Z. , Drouin , G. , Desrosiers , J. , Trensz , F. & Grenier , G . A novel hindlimb immobilization procedure for studying skeletal muscle atrophy and recovery in mouse . J Appl Physiol (1985) 106 , 2049 – 2059 ( 2009 ). OpenUrl CrossRef PubMed Web of Science 33. ↵ Huang , Z. et al. Inhibition of IL-6/JAK/STAT3 pathway rescues denervation-induced skeletal muscle atrophy . Ann Transl Med 8 , 1681 ( 2020 ). OpenUrl 34. ↵ Henriquez-Olguin , C. , Meneses-Valdes , R. & Jensen , T. E . Compartmentalized muscle redox signals controlling exercise metabolism - Current state, future challenges . Redox Biol 101473 ( 2020 ) doi: 10.1016/j.redox.2020.101473 . OpenUrl CrossRef 35. ↵ Xu , H. , Ranjit , R. , Richardson , A. & Van Remmen , H . Muscle mitochondrial catalase expression prevents neuromuscular junction disruption, atrophy, and weakness in a mouse model of accelerated sarcopenia . J Cachexia Sarcopenia Muscle 12 , 1582 – 1596 ( 2021 ). OpenUrl 36. ↵ Tang , H. et al. Overexpression of thioredoxin-2 attenuates age-related muscle loss by suppressing mitochondrial oxidative stress and apoptosis . JCSM Rapid Communications 5 , 130 – 145 ( 2022 ). OpenUrl 37. ↵ Kondo , H. , Miura , M. & Itokawa , Y . Oxidative stress in skeletal muscle atrophied by immobilization . Acta Physiol. Scand . 142 , 527 – 528 ( 1991 ). OpenUrl CrossRef PubMed Web of Science 38. ↵ Min , K. et al. Mitochondrial-targeted antioxidants protect skeletal muscle against immobilization-induced muscle atrophy . J Appl Physiol (1985) 111 , 1459 – 1466 ( 2011 ). OpenUrl CrossRef PubMed 39. ↵ Talbert , E. E. et al. Immobilization-induced activation of key proteolytic systems in skeletal muscles is prevented by a mitochondria-targeted antioxidant . J. Appl. Physiol . 115 , 529 – 538 ( 2013 ). OpenUrl CrossRef PubMed 40. ↵ Eshima , H. et al. Neutralizing mitochondrial ROS does not rescue muscle atrophy induced by hindlimb unloading in female mice . J. Appl. Physiol . ( 2020 ) doi: 10.1152/japplphysiol.00456.2019 . OpenUrl CrossRef PubMed 41. ↵ Gram , M. et al. Skeletal muscle mitochondrial H2 O2 emission increases with immobilization and decreases after aerobic training in young and older men . J. Physiol. (Lond .) 593 , 4011 – 4027 ( 2015 ). OpenUrl CrossRef PubMed 42. ↵ Glover , E. I. , Yasuda , N. , Tarnopolsky , M. A. , Abadi , A. & Phillips , S. M . Little change in markers of protein breakdown and oxidative stress in humans in immobilization-induced skeletal muscle atrophy . Appl Physiol Nutr Metab 35 , 125 – 133 ( 2010 ). OpenUrl CrossRef PubMed Web of Science 43. ↵ Powers , S. K. et al. Mitochondrial-targeted antioxidants protect against mechanical ventilation-induced diaphragm weakness . Crit Care Med 39 , 1749 – 1759 ( 2011 ). OpenUrl CrossRef PubMed Web of Science 44. ↵ Bhattacharya , A. et al. Denervation induces cytosolic phospholipase A2-mediated fatty acid hydroperoxide generation by muscle mitochondria . J Biol Chem 284 , 46 – 55 ( 2009 ). OpenUrl Abstract / FREE Full Text 45. ↵ Ahn , B. et al. Scavenging mitochondrial hydrogen peroxide by peroxiredoxin 3 overexpression attenuates contractile dysfunction and muscle atrophy in a murine model of accelerated sarcopenia . Aging Cell 21 , e13569 ( 2022 ). OpenUrl 46. ↵ Xu , H. , Czyżowska , A. , Van Remmen , H. & Brown , J. L . Modulation of sarcopenia phenotypes by glutathione peroxidase 4 overexpression in mice . J Physiol 601 , 5277 – 5293 ( 2023 ). OpenUrl CrossRef PubMed 47. ↵ Czyżowska , A. et al. Elevated phospholipid hydroperoxide glutathione peroxidase (GPX4) expression modulates oxylipin formation and inhibits age-related skeletal muscle atrophy and weakness . Redox Biol 64 , 102761 ( 2023 ). 48. ↵ Petrillo , S. et al. Oxidative stress in Duchenne muscular dystrophy: focus on the NRF2 redox pathway . Hum Mol Genet 26 , 2781 – 2790 ( 2017 ). OpenUrl 49. ↵ Haycock , J. W. , MacNeil , S. , Jones , P. , Harris , J. B. & Mantle , D . Oxidative damage to muscle protein in Duchenne muscular dystrophy . Neuroreport 8 , 357 – 361 ( 1996 ). OpenUrl CrossRef PubMed Web of Science 50. ↵ Hughes , M. C. et al. Early myopathy in Duchenne muscular dystrophy is associated with elevated mitochondrial H2 O2 emission during impaired oxidative phosphorylation . J Cachexia Sarcopenia Muscle 10 , 643 – 661 ( 2019 ). OpenUrl CrossRef PubMed 51. ↵ Mâncio , R. D. et al. Vitamin E treatment decreases muscle injury in mdx mice . Nutrition 43–44 , 39 – 46 ( 2017 ). OpenUrl 52. ↵ Whitehead , N. P. , Pham , C. , Gervasio , O. L. & Allen , D. G . N-Acetylcysteine ameliorates skeletal muscle pathophysiology in mdx mice . J Physiol 586 , 2003 – 2014 ( 2008 ). OpenUrl CrossRef PubMed Web of Science 53. ↵ Selsby , J. T . Increased catalase expression improves muscle function in mdx mice . Exp Physiol 96 , 194 – 202 ( 2011 ). OpenUrl CrossRef PubMed Web of Science 54. ↵ Bellissimo , C. A. et al. Mitochondrial creatine sensitivity is lost in the D2.mdx model of Duchenne muscular dystrophy and rescued by the mitochondrial-enhancing compound Olesoxime . Am J Physiol Cell Physiol 324 , C1141 – C1157 ( 2023 ). OpenUrl CrossRef PubMed 55. ↵ Dubinin , M. V. et al. Mitochondrial Transplantation Therapy Ameliorates Muscular Dystrophy in mdx Mouse Model . Biomolecules 14 , 316 ( 2024 ). 56. ↵ Stein , K. T. , Moon , S. J. , Nguyen , A. N. & Sikes , H. D . Kinetic modeling of H2O2 dynamics in the mitochondria of HeLa cells . PLoS Comput Biol 16 , e1008202 ( 2020 ). OpenUrl 57. ↵ Erdogan , Y. C. et al. Complexities of the chemogenetic toolkit: Differential mDAAO activation by d-amino substrates and subcellular targeting . Free Radic Biol Med 177 , 132 – 142 ( 2021 ). OpenUrl CrossRef PubMed 58. ↵ Steinhorn , B. et al. Chemogenetic generation of hydrogen peroxide in the heart induces severe cardiac dysfunction . Nat Commun 9 , 4044 ( 2018 ). OpenUrl CrossRef PubMed 59. ↵ Das , A. A. et al. Differential aortic aneurysm formation provoked by chemogenetic oxidative stress . J Clin Invest 135 , e188743 ( 2025 ). OpenUrl 60. ↵ Mackey , A. L. , Magnan , M. , Chazaud , B. & Kjaer , M . Human skeletal muscle fibroblasts stimulate in vitro myogenesis and in vivo muscle regeneration . J Physiol 595 , 5115 – 5127 ( 2017 ). OpenUrl CrossRef PubMed 61. ↵ von Maltzahn , J. , Jones , A. E. , Parks , R. J. & Rudnicki , M. A . Pax7 is critical for the normal function of satellite cells in adult skeletal muscle . Proc Natl Acad Sci U S A 110 , 16474 – 16479 ( 2013 ). OpenUrl Abstract / FREE Full Text 62. ↵ Tierney , M. T. et al. STAT3 signaling controls satellite cell expansion and skeletal muscle repair . Nat Med 20 , 1182 – 1186 ( 2014 ). OpenUrl CrossRef PubMed 63. ↵ Dodd , S. L. , Hain , B. , Senf , S. M. & Judge , A. R . Hsp27 inhibits IKKbeta-induced NF-kappaB activity and skeletal muscle atrophy . FASEB J 23 , 3415 – 3423 ( 2009 ). OpenUrl CrossRef PubMed Web of Science 64. He , M. Y. , et al. Hsp90β interacts with MDM2 to suppress p53-dependent senescence during skeletal muscle regeneration . Aging Cell 18 , e13003 ( 2019 ). 65. ↵ Niu , M. et al. Inhibition of heat shock protein (HSP) 90 reverses signal transducer and activator of transcription (STAT) 3-mediated muscle wasting in cancer cachexia mice . Br J Pharmacol 178 , 4485 – 4500 ( 2021 ). OpenUrl 66. ↵ Wang , Y. , van Boxel-Dezaire , A. H. H. , Cheon , H. , Yang , J. & Stark , G. R . STAT3 activation in response to IL-6 is prolonged by the binding of IL-6 receptor to EGF receptor . Proc Natl Acad Sci U S A 110 , 16975 – 16980 ( 2013 ). OpenUrl Abstract / FREE Full Text 67. ↵ Madaro , L. et al. Denervation-activated STAT3-IL-6 signalling in fibro-adipogenic progenitors promotes myofibres atrophy and fibrosis . Nat Cell Biol 20 , 917 – 927 ( 2018 ). OpenUrl CrossRef PubMed 68. ↵ Jamali , A. A. , Afshar , P. , Abrams , R. A. & Lieber , R. L . Differential expression of neural cell adhesion molecule (NCAM) after tenotomy in rabbit skeletal muscle . J Orthop Res 20 , 364 – 369 ( 2002 ). OpenUrl CrossRef PubMed 69. Soendenbroe , C. et al. Key Components of Human Myofibre Denervation and Neuromuscular Junction Stability are Modulated by Age and Exercise . Cells 9 , 893 ( 2020 ). 70. ↵ Ruple , B. A. et al. The effects of resistance training on denervated myofibers, senescent cells, and associated protein markers in middle-aged adults . FASEB J 38 , e23621 ( 2024 ). OpenUrl 71. ↵ Sobotta , M. C. et al. Peroxiredoxin-2 and STAT3 form a redox relay for H 2 O 2 signaling . Nat Chem Biol 11 , 64 – 70 ( 2015 ). OpenUrl CrossRef PubMed 72. ↵ Henríquez-Olguin , C. et al. Cytosolic ROS production by NADPH oxidase 2 regulates muscle glucose uptake during exercise . Nat Commun 10 , 4623 ( 2019 ). OpenUrl CrossRef 73. Henríquez-Olguín , C. et al. Adaptations to high-intensity interval training in skeletal muscle require NADPH oxidase 2 . Redox Biol 24 , ( 2019 ). 74. ↵ Ristow , M. et al. Antioxidants prevent health-promoting effects of physical exercise in humans . Proc. Natl. Acad. Sci. U.S.A . 106 , 8665 – 8670 ( 2009 ). OpenUrl Abstract / FREE Full Text 75. ↵ Henríquez-Olguín , C. et al. The Emerging Roles of Nicotinamide Adenine Dinucleotide Phosphate Oxidase 2 in Skeletal Muscle Redox Signaling and Metabolism . Antioxid. Redox Signal . 31 , 1371 – 1410 ( 2019 ). OpenUrl 76. ↵ Lim , C. et al. Turning over new ideas in human skeletal muscle proteostasis: What do we know and where to from here? Exp Physiol ( 2025 ) doi: 10.1113/EP092353 . OpenUrl CrossRef 77. ↵ Phillips , S. M. , Glover , E. I. & Rennie , M. J . Alterations of protein turnover underlying disuse atrophy in human skeletal muscle . J Appl Physiol (1985) 107 , 645 – 654 ( 2009 ). OpenUrl CrossRef PubMed 78. ↵ Ferrando , A. A. , Lane , H. W. , Stuart , C. A. , Davis-Street , J. & Wolfe , R. R . Prolonged bed rest decreases skeletal muscle and whole body protein synthesis . Am J Physiol 270 , E627 – 633 ( 1996 ). OpenUrl CrossRef Web of Science 79. ↵ Arc-Chagnaud , C. et al. Evaluation of an Antioxidant and Anti-inflammatory Cocktail Against Human Hypoactivity-Induced Skeletal Muscle Deconditioning . Front Physiol 11 , 71 ( 2020 ). 80. ↵ Saks , V. A. et al. Permeabilized cell and skinned fiber techniques in studies of mitochondrial function in vivo . Mol Cell Biochem 184 , 81 – 100 ( 1998 ). OpenUrl CrossRef PubMed Web of Science 81. ↵ Moore , M. , Gossmann , N. & Dietz , K.-J . Redox Regulation of Cytosolic Translation in Plants . Trends in Plant Science 21 , 388 – 397 ( 2016 ). OpenUrl CrossRef PubMed 82. ↵ Sakellariou , G. K. et al. Mitochondrial ROS regulate oxidative damage and mitophagy but not age-related muscle fiber atrophy . Sci Rep 6 , 33944 ( 2016 ). 83. ↵ Murphy , M. P. et al. Guidelines for measuring reactive oxygen species and oxidative damage in cells and in vivo . Nat Metab 4 , 651 – 662 ( 2022 ). OpenUrl 84. ↵ Hoehne , M. N. et al. Spatial and temporal control of mitochondrial H2 O2 release in intact human cells . EMBO J 41 , e109169 ( 2022 ). OpenUrl CrossRef PubMed 85. ↵ Chowdhury , A. R. et al. Mitochondria-targeted paraquat and metformin mediate ROS production to induce multiple pathways of retrograde signaling: A dose-dependent phenomenon . Redox Biol 36 , 101606 ( 2020 ). 86. ↵ Smuder , A. J. , Kavazis , A. N. , Hudson , M. B. , Nelson , W. B. & Powers , S. K . Oxidation enhances myofibrillar protein degradation via calpain and caspase-3 . Free Radical Biology and Medicine 49 , 1152 – 1160 ( 2010 ). OpenUrl CrossRef PubMed 87. ↵ Hyatt , H. W. , Ozdemir , M. , Bomkamp , M. P. & Powers , S. K . Activation of Calpain Contributes to Mechanical Ventilation-Induced Depression of Protein Synthesis in Diaphragm Muscle . Cells 11 , 1028 ( 2022 ). OpenUrl 88. ↵ Sidlauskaite , E. et al. Mitochondrial ROS cause motor deficits induced by synaptic inactivity: Implications for synapse pruning . Redox Biol 16 , 344 – 351 ( 2018 ). OpenUrl 89. Stavrovskaya , I. et al. Mitochondrial ROS modulate presynaptic plasticity in the drosophila neuromuscular junction . Redox Biol 79 , 103474 ( 2024 ). 90. ↵ Su , Y. et al. Transgenic expression of SOD1 specifically in neurons of Sod1 deficient mice prevents defects in muscle mitochondrial function and calcium handling . Free Radic Biol Med 165 , 299 – 311 ( 2021 ). OpenUrl CrossRef PubMed 91. ↵ Motanova , E. et al. Mitochondrial alterations as potential early events in neuromuscular junction remodelling with muscle disuse . Experimental Physiology n/a , ( 2025 ). 92. ↵ Bock , F. J. & Tait , S. W. G . Mitochondria as multifaceted regulators of cell death . Nat Rev Mol Cell Biol 21 , 85 – 100 ( 2020 ). OpenUrl CrossRef PubMed 93. ↵ Kalinichenko , A. L. et al. Chemogenetic emulation of intraneuronal oxidative stress affects synaptic plasticity . Redox Biol 60 , 102604 ( 2023 ). 94. ↵ Nanadikar , M. S. et al. IDH3γ functions as a redox switch regulating mitochondrial energy metabolism and contractility in the heart . Nat Commun 14 , 2123 ( 2023 ). OpenUrl 95. ↵ Arnett , A. L. et al. Adeno-associated viral (AAV) vectors do not efficiently target muscle satellite cells . Mol Ther Methods Clin Dev 1 , 14038 - ( 2014 ). OpenUrl PubMed 96. ↵ Koga , R. , Miyoshi , Y. , Sakaue , H. , Hamase , K. & Konno , R . Mouse d-Amino-Acid Oxidase: Distribution and Physiological Substrates . Front Mol Biosci 4 , 82 ( 2017 ). 97. ↵ Constantin-Teodosiu , D. , Peirce , N. S. , Fox , J. & Greenhaff , P. L . Muscle pyruvate availability can limit the flux, but not activation, of the pyruvate dehydrogenase complex during submaximal exercise in humans . J Physiol 561 , 647 – 655 ( 2004 ). OpenUrl CrossRef PubMed 98. Stellingwerff , T. , Leblanc , P. J. , Hollidge , M. G. , Heigenhauser , G. J. F. & Spriet , L. L . Hyperoxia decreases muscle glycogenolysis, lactate production, and lactate efflux during steady-state exercise . Am J Physiol Endocrinol Metab 290 , E1180 – 1190 ( 2006 ). OpenUrl CrossRef PubMed Web of Science 99. ↵ Matlashov , M. E. , Belousov , V. V. & Enikolopov , G . How much H(2)O(2) is produced by recombinant D- amino acid oxidase in mammalian cells? Antioxid Redox Signal 20 , 1039 – 1044 ( 2014 ). OpenUrl CrossRef PubMed Web of Science 100. ↵ Gruntman , A. M. et al. Gene transfer in skeletal and cardiac muscle using recombinant adeno-associated virus . Curr Protoc Microbiol Chapter 14 , Unit 14D.3 ( 2013 ). 101. ↵ Saeedi Saravi , S. S. , et al. Differential endothelial signaling responses elicited by chemogenetic H2O2 synthesis . Redox Biol 36 , 101605 ( 2020 ). 102. Spyropoulos , F. et al. Metabolomic and transcriptomic signatures of chemogenetic heart failure . Am J Physiol Heart Circ Physiol 322 , H451 – H465 ( 2022 ). OpenUrl 103. ↵ den Toom , W. T. F. , et al. Oxygen-consumption based quantification of chemogenetic H2O2 production in live human cells . Free Radic Biol Med 206 , 134 – 142 ( 2023 ). OpenUrl CrossRef PubMed 104. ↵ Waisman , A. , Norris , A. M. , Elías Costa , M. & Kopinke , D . Automatic and unbiased segmentation and quantification of myofibers in skeletal muscle . Sci Rep 11 , 11793 ( 2021 ). 105. ↵ Jensen , R. , Ørtenblad , N. , di Benedetto , C. , Qvortrup , K. & Nielsen , J . Quantification of Subcellular Glycogen Distribution in Skeletal Muscle Fibers using Transmission Electron Microscopy . 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