No evidence of the presence of dystrophin protein in rodent muscle stem cells

No evidence of the presence of dystrophin protein in rodent muscle stem cells | Research Square window.SnipcartSettings = { analytics: { enabled: false } }; (function() { var accessVector = localStorage.getItem('access_vector') || ''; window.dataLayer = window.dataLayer || []; if (accessVector) { window.dataLayer.push({ user: { profile: { profileInfo: { snid: accessVector } } } }); } })(); (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],j=d.createElement(s),dl=l!='dataLayer'?'&l='+l:'';j.async=true;j.src='https://www.googletagmanager.com/gtm.js?id='+i+dl;f.parentNode.insertBefore(j,f);})(window,document,'script','dataLayer','GTM-K279D39R'); Browse Preprints In Review Journals COVID-19 Preprints AJE Video Bytes Research Tools Research Promotion AJE Professional Editing AJE Rubriq About Preprint Platform In Review Editorial Policies Our Team Advisory Board Help Center Sign In Submit a Preprint Cite Share Download PDF Research Article No evidence of the presence of dystrophin protein in rodent muscle stem cells Meriem Matouk, Adrien Morin, Lucie Royer, Gaspard Macaux, Aris Gaci, and 12 more This is a preprint; it has not been peer reviewed by a journal. https://doi.org/ 10.21203/rs.3.rs-7997649/v1 This work is licensed under a CC BY 4.0 License Status: Under Review Version 1 posted 9 You are reading this latest preprint version Abstract The polarized distribution of dystrophin in muscle stem cells (MuSCs) has been found to regulate asymmetric cell division and maintain the balance between stem cell self-renewal and myogenic commitment. In Duchenne muscular dystrophy, which is associated with dystrophin deficiency, MuSC dysfunction is believed to contribute to fatal muscle wasting. This study investigated the dynamics of the polarized dystrophin distribution during MuSC activation. To this end, we used live fluorescence imaging to visualize EGFP-labeled dystrophin in muscle fibers and MuSCs in the Dmd EGFP reporter mouse model. We also investigated this phenomenon in rats to assess the existence of interspecies consistency. Dystrophin was nearly absent on the apical side of quiescent MuSCs, whereas it rapidly accumulated near a subset of activated mouse MuSCs and 100% of rat MuSCs following enzymatic myofiber isolation and culture. Surprisingly, dystrophin that accumulated near MuSCs accumulated in membranes, and it was always found in continuity with the sarcolemma. Live imaging revealed that MuSCs could move from the condensed dystrophin, which remained attached to the myofiber. Additionally, we detected no dystrophin protein in MuSCs using different techniques, including immunocytochemistry capillary electrophoresis and fluorescence-activated cell sorting, whereas it was present in primary myoblasts and myotubes. Our findings indicate that dystrophin attached to activated MuSCs represents condensed sarcolemmal membranes. Our data suggest that dystrophin protein in MuSCs is not required for normal muscle regeneration. Dystrophin MuSCs Pax7 sarcolemma Figures Figure 1 Figure 2 Figure 3 Figure 4 Significance Statement Duchenne muscular dystrophy results from the absence of dystrophin, leading to progressive muscle wasting. Although dystrophin is essential for myofiber stability, its role in muscle stem cells (MuSCs) has remained unclear. Using live imaging in dystrophin-EGFP mice and cross-species analysis, we show that the dystrophin signal in MuSCs originates from sarcolemmal membranes rather than intrinsic expression. These findings demonstrate that dystrophin is dispensable for MuSC regenerative function and resolve a long-standing controversy about its cellular role. Introduction Progressive skeletal muscle wasting in Duchenne muscular dystrophy (DMD) is traditionally attributed to the inherited absence of sarcolemmal dystrophin, which decreases the resistance of myofibers to mechanical stress, resulting in fiber necrosis, inflammation, chronic cycles of fiber degeneration and regeneration, and fibrotic and fatty transformation of muscle [ 1 – 2 ]. In addition to this classical view, inefficient muscle regeneration has been suspected to contribute to muscle wasting in DMD attributable to defective muscle stem cell (MuSC) function [ 3 – 4 ]. MuSCs are located between the sarcolemma and basal lamina of myofibers, and they are generally quiescent in mature muscle [ 5 ]. Myofiber injury, such as that in DMD, activates MuSCs, which begin to proliferate and eventually differentiate to replace damaged muscle fibers or self-renew to replenish the MuSC pool. The proper balance between self-renewal and myogenic progression is critical for ensuring the lifelong regenerative potential of skeletal muscle [ 6 ], [ 7 ]. Self-renewal in MuSCs requires asymmetric divisions, in which one daughter cell retains stemness and the other undergoes myogenic commitment, whereas symmetric divisions expand the cell pool by generating identical daughter cells [ 8 ], [ 9 ], [ 10 ]. Asymmetric division requires an apicobasal orientation of the dividing MuSCs relative to the myofiber, whereas symmetric division occurs in a planar orientation [ 11 ], [ 12 ]. Only the presence of the full-length dystrophin isoform Dp427 and the short ubiquitously expressed Dp71 have been documented in MuSCs [ 13 , 14 ]. A polarized distribution of full-length dystrophin in activated MuSCs is critical for proper MuSC self-renewal [ 14 ]. The spectrin-like repeat domain of dystrophin has been demonstrated to bind to MAP/microtubule affinity-regulating kinase 2 [ 14 , 15 ] and regulate polarity establishment for asymmetric versus symmetric self-renewal MuSC division [ 16 , 17 ]. The polarity of the dystrophin distribution in activated MuSCs has been primarily studied in isolated myofibers from murine models, but its polarity has also been detected in human fiber cultures derived from postmortem tissue [ 14 , 18 ]. Dystrophin is absent in Dmd mdx/y mice that harbor a nonsense mutation in Dmd exon 23 and lack the full-length dystrophin isoform Dp427 [ 14 ]. The loss of dystrophin impairs the establishment of MuSC polarity and results in the loss of apicobasal asymmetric cell division in favor of planar symmetric division, thereby altering the balance between MuSC self-renewal and myogenic commitment. This loss in polarity affects MuSCs proliferation, thereby impairing lineage progression of myoblasts and explaining the regeneration deficit of DMD muscle [ 14 ]. We previously generated a Dmd EGFP reporter mouse line harboring an EGFP-tag fused to the C-terminal end of dystrophin. The resulting dystrophin-EGFP fusion protein allows in vivo and ex-vivo live imaging of dystrophin isoforms carrying the C-terminal region [ 19 ]. Using live fluorescence imaging, we investigated the dynamics of dystrophin polarization in activated MuSCs. Our data illustrated that MuSCs themselves were devoid of dystrophin. In fact, dystrophin, which accumulated near activated MuSCs, belonged to the sarcolemma. We therefore systematically analyzed the relationship between dystrophin and MuSCs in murine and rat animal models. Results Live observation of dystrophin in the MuSC niche We examined skeletal muscle from Dmd EGFP/y ; Pax7 CreERT2/+ ; Rosa tdT/+ reporter mice to visualize native dystrophin-EGFP fusion protein and tdTomato + MuSCs following tamoxifen treatment. Dystrophin-EGFP and tdTomato signals were easily identified in vivo , ex vivo , and in vitro , as well as by immunohistochemistry. A limitation of the system was that tdTomato faintly photoconverted to the GFP spectrum, and this effect was increasingly noticeable following prolonged excitation (Figure S1 A) . This sometimes caused a faint GFP signal in the cytoplasm of MuSCs, which we considered non-specific and unrelated to dystrophin-EGFP. In vivo multiphoton microscopy of the TA muscle of adult Dmd EGFP/y ; Pax7 CreERT2/+ ; Rosa tdT/+ mice revealed that MuSCs had an elongated shape with long tdTomato + projections in the expected region near the dystrophin-EGFP + sarcolemma of myofibers ( Fig. 1 A ) . Curiously, dystrophin-EGFP was continuously present at homogeneous levels along the sarcolemma of one adjacent myofiber, whereas sarcolemmal dystrophin-EGFP protein was either reduced or absent from the other bordering myofiber ( Fig. 1 B ) . We next immediately fixed EDL muscle of adult Pax7 CreERT2/CreERT2 ; Rosa tdT/tdT mice in PFA following dissection and then mechanically isolated myofibers, which were immunostained to visualize dystrophin. Confocal microscopy revealed that tdTomato + MuSCs were attached to the muscle fiber, and they featured cell projections in accordance with the elongated shape observed in vivo. Sarcolemmal dystrophin was absent on the apical side of MuSCs (side of MuSCs facing the sarcolemma), in line with previously published observations [ 30 ] ( Fig. 1 C ) . Native fluorescence on the cryosections of PFA-fixed TA muscles from adult Dmd EGFP/y ; Pax7 CreERT2/+ ; Rosa tdT/+ mice confirmed the strong reduction or absence of sarcolemmal dystrophin-EGFP on one side of all MuSCs ( Fig. 1 D ) , in line with previous findings [ 31 ]. Similarly, in Pax7 CreERT2/+ ; Rosa tdT/+ mice, another member of the dystrophin associated protein complex (DAPC), β-sarcoglycan, was also absent from the same side of MuSCs (Figure S1 B) . Next, we enzymatically isolated myofibers (a procedure that lasts about 2–3 hours), thereby permitting live ex-vivo observation of tdTomato + MuSCs and dystrophin-EGFP during culture. Once isolated, the muscle fiber was detached from its tendinous attachments. We first sought to clarify whether isolation influenced the length of myofiber sarcomeres by comparing fibers isolated via enzymatic and mechanical isolation. In our previously published work, we recorded a sarcomere size of 3.3 µm in Dmd EGFP mice in vivo [ 32 ]. In this study, the sarcomere length was 2.36 µm in mechanically isolated fibers, versus 1.82 µm following enzymatic fiber isolation (Figure S1 C) . Taken together, the data indicate that myofibers contracted by approximately 23% during enzymatic isolation, and this likely decreased sarcolemmal tension. In addition, most MuSCs retracted their cell projections and exhibited a round morphology, and these retracted MuSCs usually remained attached to the fibers. The MuSCs displayed varying levels of dystrophin-EGFP accumulation. Based on their orientation, we distinguished three regions: an apical side facing the fiber’s sarcolemma, lateral sides on either edge, and a basal side oriented toward the basal lamina. Upon isolation (T0), we frequently observed increased dystrophin-EGFP accumulation on the apical side (sarcolemmal side) of these retracted MuSCs ( Fig. 2 A ) , which appeared in continuation of the sarcolemma and contrasted the findings in vivo , after cryosectioning, or following mechanical fiber isolation. Of note, this apically increased dystrophin was similar to the diameter of MuSCs of approximately 8 µm (Figure S2A) , being 10-fold smaller than the basal sarcolemmal dystrophin unit (BSDU), which is the sarcolemmal extension of a myonuclear domain [ 32 ]. This localized and rapid dystrophin accumulation suggests that this event is not driven by a myonuclear process. The apical condensation of dystrophin increased at T36 and T48 during myofiber culture and often extended laterally, whereas dystrophin-EGFP rarely accumulated on the basal side of MuSCs (Figure S2B) . We quantified the number of MuSCs in respect to dystrophin-EGFP accumulation and performed time-lapse imaging of cultured myofibers isolated from Dmd EGFP mice. We used PAX7 immunochemistry to visualize MuSCs. At T0, dystrophin-EGFP accumulated apicolaterally in 8.8% of MuSCs, and this value increased to 22.9% at T48. Dystrophin accumulation on the basal side was found in 0.6% of MuSCs at T12, 3.6% of cells at T24, 4.1% of cells at T36, and 4.4% of cells at T48, matching previous observations [ 14 ] (Figure S2B) . We next followed the movements of tdTomato + MuSCs on isolated fibers from Dmd EGFP/y ; Pax7 CreERT2/+ ; Rosa tdT/+ mice using time-lapse microscopy beginning at 44 h of culture ( Fig. 2 B ) . Images were acquired every 10 min for 6 h. At T44, dystrophin-EGFP often accumulated, as observed in the previous experiment, on the apical, lateral, or basal side of MuSCs. However, we unexpectedly found that dystrophin-EGFP accumulation did not follow the migration of MuSCs. As indicated by the example of Cell 1, MuSCs had moved from the apically accumulated dystrophin-EGFP at T46 to the other side of the fiber. Meanwhile, the cell left a visible footprint of dystrophin-EGFP at its original position at T44, which remained visible at T52 ( Fig. 2 B ) . Conversely, dystrophin-EGFP accumulated basally (Cell 2) or laterally of the MuSCs (Cell 3). Over time, we observed that the MuSCs squeezed through this dystrophin accumulation, and we subsequently lost view of the cells. Meanwhile, the dystrophin-EGFP accumulation visible at T44 remained at the same position at T52. As illustrated in the example 4, the MuSC partially moved from the basally accumulated dystrophin-EGFP. These data were confirmed by time-lapse videos, illustrating that MuSCs moved from the area of MuSC-associated dystrophin accumulation (Movie S1) . MuSC-associated dystrophin is of sarcolemmal origin These surprising observations prompted us to hypothesize that dystrophin accumulation associated with MuSCs might be of sarcolemmal origin. We previously reported that increased dystrophin-EGFP accumulation at myotendinous junctions resulted from sarcolemmal folding, which we could image in isolated fibers using the fluorescent membrane probe MemBright [ 33 ], a finding that we reproduced using isolated myofibers from Dmd EGFP/y ; Pax7 CreERT2/+ ; Rosa tdT/+ mice following 48 h of culture ( Fig. 3 A ) . Upon analyzing the MuSCs of these fibers, we found that MemBright fluorescence colocalized with the high accumulation of dystrophin-EGFP at the MuSC level, and this accumulation was continuous with the MemBright staining of the sarcolemma ( Fig. 3 A ) . These observations demonstrate that dystrophin-EGFP accumulation resulted indeed from condensed/stacked membranes; however, this experiment did not uncover the cellular origin of the dystrophin-EGFP + membranes. We next determined the frequency of dystrophin-EGFP positivity among tdTomato + cells. We first confirmed that the spectral emissions of EGFP and tdTomato in fibers freshly isolated from Dmd EGFP/y ; Pax7 CreERT2/+ ; Rosa tdT/+ reporter mice did not overlap, allowing high-quality spectral acquisition (Figure S3A) . Moreover, the EGFP spectrum was absent in tdTomato + MuSCs (Figure S3A) . This allowed us to differentiate dystrophin-EGFP and MuSC-emitted tdTomato fluorescence. Using FACS, we sorted tdTomato + MuSCs from the muscles of adult Dmd EGFP/y ; Pax7 CreERT2/+ ; Rosa tdT /+ mice and Dmd EGFP−mdx/y ; Pax7 CreERT2/+ ; Rosa tdT/+ reporter mice. FACS-sorted GFP + MuSCs from Tg:Pax7nGFP mice served as positive controls, and a GFP + cell population was easily identified ( Fig. 3 B ) . tdTomato + MuSCs from Pax7 CreERT2/CreERT2 ; Rosa tdT/tdT mice served as negative controls, and as expected, this population was GFP − , confirming the spectral specificity (Figure S3B) . No GFP + population was detected in FACS-sorted MuSCs from Dmd EGFP/y ; Pax7 CreERT2/+ ; Rosa tdT/+ muscles or Dmd EGFP−mdx/y ; Pax7 CreERT2/+ ; Rosa tdT /+ muscles, confirming the absence of dystrophin-EGFP protein in the MuSCs of these mice (Figure S3B) . To confirm the biological significance of this observation across species, we cultured isolated myofibers from adult Sprague–Dawley rats for 48 h and performed co-immunostaining against dystrophin and PAX7. MuSCs had often already undergone cell division, and they appeared in doublets or small clusters. Strikingly, dystrophin highly accumulated around all single MuSCs or in juxtaposition to all MuSCs doublets/clusters. Dystrophin accumulated erratically at the MuSC level, either apically, basally, laterally, or between MuSCs, with no preferential localization ( Figs. 3 C and S2 C ) . Sarcolemmal dystrophin was organized in costameres, which appeared tethered toward the dystrophin accumulation associated with MuSCs (Figure S3D) . Airyscan microscopy revealed dystrophin + membrane stacks in continuation with the sarcolemma ( Figs. 3 C and S3 E ) . Presence of Dmd transcripts but absence of dystrophin protein in MuSCs We then assessed the dynamics of Dmd expression and dystrophin protein synthesis during myogenic lineage progression. For this purpose, we generated primary myoblast cultures using MuSCs from 1-week-old wild-type CD-1 mice. Previous research demonstrated that such cultures form mature myotubes in which dystrophin is located at the sarcolemmal level [ 32 ]. ddPCR revealed the strong presence of Pax7 in freshly derived MuSCs (T0), Myod1 expression in MuSC-derived myoblasts after 3 days of culture, and strong Myh1 expression in myotubes following 13 days of culture, confirming the myogenic line progression of cells in these primary cultures (Figure S4A) . Dmd transcripts were already detectable in isolated MuSCs at T0 ( Fig. 4 A ) , and Dmd RNA was unexpectedly more abundant in MuSCs than in myoblasts and myotubes, corroborating previous findings [ 14 ]. We also performed snRNA-Seq of adult TA muscles, revealing that all Pax7 + myonuclei were also Dmd + (Figures S4B and C) . We next performed capillary electrophoresis of parallel myoblast and myotube cultures from 7-day-old CD-1 mice as used for the aforementioned PCR experiments. The full-length dystrophin isoform Dp427 was present at low levels in myoblasts and myotubes, contrasting the high dystrophin levels in TA muscles from adult wild-type CD-1 mice. However, dystrophin was not detected in MACS-sorted MuSCs from 2-week-old mice using antibodies against both the rod domain and C-terminal domain of dystrophin ( Figs. 4 B and S4 D ) . Similarly, in wild-type adult Sprague-Dawley rats, capillary electrophoresis revealed that full-length dystrophin was absent in MACS-sorted MuSCs, whereas it was highly abundant in skeletal muscle, which served as a positive control, and absent in muscle from the DMD rat model (harboring a frameshift deletion of Dmd exon 52), which served as negative control (Figure S4E, and F) . Finally, immunocytochemistry of parallel myoblast and myotube cultures from 7-day-old CD-1 mice revealed that dystrophin protein was first detected in a predominantly scattered perinuclear position in myoblasts and thereafter along the sarcolemma in myotubes ( Fig. 4 C ) . Discussion The polarized distribution of dystrophin protein in MuSCs is critical for establishing cell polarity, regulating asymmetric versus symmetric cell division, and enabling correct muscle regeneration [ 14 ]. In this study, we investigated the dynamics of the establishment of this polarized dystrophin distribution in MuSCs. We used reporter mouse lines to directly image the native dystrophin-EGFP fusion protein, which we previously employed to study the compartmentalization of dystrophin in skeletal muscle at subcellular levels [ 32 ]. We found that dystrophin protein was undetectable in quiescent adult MuSCs. In addition, sarcolemmal dystrophin facing MuSCs was virtually absent, confirming published data [ 31 ]. This absence of dystrophin in the niche of quiescent MuSCs contrasted the rapid accumulation of the protein in juxtaposition to activated MuSCs following the culture of isolated myofibers, in line with previous observations [ 14 , 18 ]. As previously demonstrated [ 14 ], we found that despite the absence of dystrophin protein, Dmd was highly transcribed according to bulk PCR, confirming previous data [ 14 ]. Furthermore, snRNA-Seq revealed that transcripts were present in all MuSCs, and we found no evidence of a subpopulation with disproportionately strong expression. Of note, enzymatic isolation of myofibers requires approximately 2 h, and it can induce the activation of quiescent MuSCs [ 34 ]. However, the timeframe of isolation is insufficient for significant transcription of the Dmd gene (2.4 million base pairs in length), as this process requires approximately 16 h [ 35 ]. Nevertheless, because high levels of Dmd transcripts are already present in quiescent MuSCs, it is possible that rapid translation could provide an explanation for the rapid appearance of dystrophin protein near MuSCs following isolation. However, dystrophin accumulated as membranous stacks in the niche of activated MuSCs and appeared in a continuum with the sarcolemma. This suggests a sarcolemmal origin of MuSC-associated dystrophin. Sarcolemmal dystrophin is organized in nuclear domains, also termed BSDUs, spanning approximately 80 µm in length [ 32 ]. It is therefore unlikely that the observed rapid and locally restricted accumulation of dystrophin in the MuSC niche is a myonuclear driven process. Could already existing dystrophin locally accumulate near activated MuSC? Sarcolemmal dystrophin is organized in costameres [ 36 ]. We demonstrated in this study that costamere-organized sarcolemmal dystrophin accumulated in these stacks. Do MuSCs actively gather sarcolemma? It was previously observed that MuSCs move along the length of an isolated fiber in culture [ 37 ]. Our time-lapse experiments illustrated that MuSCs moved from their niche but left dystrophin stacks that remained attached to the sarcolemma, providing further evidence that dystrophin represented condensed sarcolemma. Interestingly, myofibers contracted by approximately 23% during enzymatic isolation. In addition, following myofiber isolation, we and others observed that MuSCs retracted their cell membrane extensions and adopted a rounded morphology [ 30 ]. We suggest that the sarcolemma becomes floppy once fibers are no longer stretched between tendons, and retracting and mobile MuSCs shuffle these floppy sarcolemmata together. Our results imply that dystrophin accumulation near MuSCs occurs only when enzymatically isolating and culturing myofibers. In mice, we observed these events in approximately 25% of MuSCs either on the apical, apical–lateral, and (more rarely) basal sides of MuSCs. These results support previous findings [ 14 ]. Curiously, dystrophin accumulated in juxtaposition to all MuSCs in cultured myofibers isolated from rat muscles. The dystrophin accumulation appeared rather erratically at any position around or between MuSCs with no distinct morphological feature. Finally, we demonstrated that dystrophin protein first emerged in primary myoblasts in a perinuclear distribution before it became localized at the sarcolemmal level in myotubes. This perinuclear position is compatible with the expected endoplasmic and ribosomal translation sites. We used reporter mice in which EGFP was directly tagged to the C-terminus of dystrophin, permitting visualization of all dystrophin isoforms [ 25 ]. The absence of dystrophin-EGFP in MuSCs suggests the absence of different dystrophin isoforms despite prior evidence of Dp71 in myoblasts [ 13 ]. Although our findings do not contradict the occurrence of MuSC polarization or the critical role of balancing asymmetric and symmetric divisions in muscle regeneration, they challenge the previously proposed involvement of dystrophin in this process. Our data do not question the accumulating evidence that MuSCs function is compromised in DMD and DMD animal models [ 38 , 39 , 40 ]. However, our results do not support the direct involvement of dystrophin, as it was absent in MuSCs. Other mechanisms, such as an altered extracellular matrix, altered muscle metabolism, altered signaling pathways, altered vascularization, and precocious MuSCs senescence, must be considered [ 41 , 42 ]. Thus, several pathophysiological processes that are well described in DMD could hamper MuSC-dependent muscle regeneration. An alternative perspective is the possibility of post-transcriptional regulation of dystrophin, with miRNAs potentially modulating mRNA stability or translation. Supporting this idea, several studies highlighted the role of specific miRNAs in this process. For instance, miR-31, which is upregulated in Dmd mdx/y mice, has been identified as a potential repressor of dystrophin [ 43 ]. Additionally, intronic DMD RNA sequences might act as non-coding RNA (ncRNA) regulators [ 44 ], and DMD mutations could disrupt this function. However, the existence of this regulatory role for DMD ncRNA in MuSCs remains purely theoretical at this stage. Satellite cell-opathies are increasingly recognized clinical entities that are best illustrated in patients with PAX7 mutations [ 45 ]. We previously described the distinct clinical phenotype of a patient with PAX7 mutation [ 46 ], which is entirely different from that of DMD. Nevertheless, MuSCs could represent a useful target for dystrophin-restoring therapies such as gene editing. Correction of the mutated DMD gene could progressively replace dystrophin-incompetent myonuclei with dystrophin-competent myonuclei during muscle regeneration and thereby overcome the dystrophic phenotype. The function of high DMD RNA levels in MuSCs remains unclear. Are high DMD RNA levels required for rapid dystrophin synthesis and sarcolemmal stability upon MuSC fusion into the growing myofiber syncytium? If this is the case, then it would be important to restore the reading frame of mutated DMD RNA in differentiated muscle and MuSCs, and the effect of current exon skipping therapies on MuSCs should be tested. Materials and Methods Animal models All animal experiments were conducted according to the National and European legislation and institutional guidelines for the care and use of laboratory animals approved by the French government (Ministère de l’Enseignement Supérieur et de la Recherche, autorisation APAFiS #44987-2023092714211861 v4 and #34462-2021121716045314 v6). Mice were bred in the Plateforme 2Care animal facility, UFR des Sciences de la Santé, Université de Versailles Saint- Quentin-en-Yvelines and maintained under a standard 12-h light/12-h dark photoperiod with free access to food and water. Mice were weaned after 4–5 postnatal weeks, and 2–5 individuals were housed per cage. To generate the Dmd EGFP/y ; Pax7 CreERT2/+ ; Rosa tdT/+ mouse line, we crossed homozygous female Dmd EGFP/EGFP mice [ 19 ] with heterozygous male Pax7 CreERT2/+ ; R 26 LoxP − STOP−LoxP−TdTom ( i.e. , Rosa tdT/+ ) mice [ 20 , 21 ]. Similarly, we generated Dmd EGFP−mdx/y Pax7 CreERT2/+ ; Rosa tdT/+ reporter mice (carrying the mdx -23 mutation and C-terminal EGFP-tag in cis in the Dmd locus[ 22 ]). Tg:Pax7-nGFP mice [ 23 ] were used to express nuclear green fluorescent protein (nGFP) under the control of the Pax7 promoter. Activation of the tdTomato transgene in Pax7 -expressing MuSCs was induced by either 1 week of tamoxifen-enriched diet (TAM400-IRRADIATED, Envigo, Indianapolis, IN, USA) feeding or daily oral gavage of tamoxifen (Sigma-Aldrich, St. Louis, MO, USA) dissolved in corn oil at 25 mg/ml and administered at a dose of 8 µl/g body weight for 5 consecutive days. Wild-type CD-1® IGS mice and wild-type Sprague-Dawley rats were purchased from Charles River Laboratories (Wilmington, MA, USA). Muscles from Sprague-Dawley rats carrying Dmd exon 52 deletion ( R-DMDdel52 , also called DMD rats) were kindly gifted by Valentina Taglietti [ 24 ]. This study was performed in juvenile (1–2 weeks old) and adult animals (2–4 months old). Mice were euthanized by cervical dislocation, whereas rats were euthanized by intracardiac administration of Euthasol (0.1 mL/kg, Centravet, France), in accordance with approved institutional and ethical guidelines. All animal experimentation was conducted and reported in accordance with the ARRIVE 2.0 guidelines. Myofiber isolation, culture, and live imaging Isolated muscle fibers were prepared from extensor digitorum longus (EDL) muscle isolates from mice and rats as previously described [ 25 , 26 ]. The isolated live myofibers were either directly imaged in DMEM Fluorobright (Thermo Fisher Scientific, Waltham, MA, USA) supplemented with 10% l -glutamine (Thermo Fisher Scientific), 1% sodium pyruvate (Thermo Fisher Scientific) and 1% penicillin–streptomycin (Thermo Fisher Scientific) in eight-well glass-bottom dishes (Ibidi, Gräfelfing, Germany) or cultured up to 48 h as previously described [ 25 ]. For myofibers fixed prior to isolation, EDL muscle was fixed in 4% PFA for 15–20 min directly after dissection, followed by two washes with 1× PBS. The fibers were carefully separated under a binocular stereomicroscope using fine glass needles with a rounded tip. For live microscopy, the isolated myofibers were mounted following 1 h of incubation at 37°C with MemBright (1:1000 dilution, 550 Lipilight by MemBright, Idylle, Paris, France), SiR-actin (1:1000 dilution, Spirochrome, Stein am Rhein, Switzerland) or Hoechst (1:1000 dilution, Sigma-Aldrich) in isolation medium (DMEM supplemented with 1% penicillin–streptomycin, 4 mM l -glutamine, and 1% sodium pyruvate). Muscle preparation for histology Tibialis anterior (TA) and triceps brachii muscles were dissected following euthanasia (cervical dislocation for mice and anesthetic overload with Euthazol [Virbec, Carros, France] 400 mg/ml for rats). The muscles were mounted on 6% tragacanth gum (Sigma-Aldrich), snap frozen in liquid nitrogen-cooled isopentane, and stored at − 80°C. Cryosections were prepared using a NX70 Cryostar cryostat (Thermo Fisher Scientific) for native fluorescence imaging and subsequently used for RNA and protein extraction. Cell culture Cells were grown as described by Pimentel et al. [ 27 ]. In summary, hindlimb muscles from 7-day-old wild-type CD-1 mice were dissected and incubated in an enzymatic mixture as previously described [ 27 ]. The cell suspension was pre-plated at 37°C for 30 min to remove fibroblasts. The supernatant was then collected and plated on Matrigel-coated dishes (1:100 dilution, Corning, NY, USA) using proliferation medium (IMDM with GlutaMAX [Thermo Fisher Scientific], 0.1% gentamicin [Thermo Fisher Scientific], 20% fetal bovine serum [Thermo Fisher Scientific], and 1% chicken embryo extract [Thermo Fisher Scientific). After 3 days of culture, the medium was removed, and ice-cold Matrigel (1:2 dilution, Corning) was added before adding differentiation medium (IMDM with GlutaMAX, 2% horse serum, and agrin 100 ng/ml). The medium was refreshed every 2 days, and the culture was stopped after 13 days when well-differentiated myotubes had formed. Cells were harvested after 3 and 13 days for further digital droplet PCR (ddPCR), immunocytochemistry, and capillary electrophoresis experiments. Immunostaining on cells and myofibers For immunostaining, muscle fiber, myoblast, and myotube cultures were fixed in 4% PFA for 10 min. Immunostaining was performed as previously described [ 25 ] using primary antibodies targeting the C-terminal region of dystrophin (1:10 dilution, NCL-DYS2, mouse IgG1, Leica Biosystems, Nussloch, Germany; or 1:2,000 dilution, ab15277, Abcam), β-sarcoglycan (1:50 dilution, Leica), PAX7 (1:10 dilution, DHSB), MYOD (1:50 dilution, Agilent, Santa Clara, CA, USA), and MHC (1:200 dilution, DHSB) via overnight incubation at 4°C, followed by incubation for 1 h with fluorochrome-labeled secondary antibodies (1:400 dilution, Alexa Fluor goat anti-mouse IgG1-488/555/633, goat antimouse IgM-488/555, and goat antirabbit IgG (H + L)-488/555/633) at room temperature. Nuclei were stained using DAPI (1:1000 dilution, Sigma-Aldrich), and the fibers were subsequently mounted on glass slides with Fluoromount-G (Southern Biotech, Birmingham, AL, USA). Fluorescence imaging Fluorescent images of live or immune-stained single myofibers and cells were acquired using an epifluorescence microscope or confocal microscope as specified in the figure legends. For epifluorescence imaging, a Zeiss Axio Imager microscope (Carl Zeiss, Oberkochen, Germany) equipped with an Orca camera (Hamamatsu Photonics, Hamamatsu, Japan) and ZEN 3.2 Blue software (Carl Zeiss) was used. For live confocal microscopy, myofibers were maintained at 37°C with 5% CO 2 in the microscope’s incubation chamber, and imaging was performed using a Leica SP8 white light laser-scanning confocal microscope with LAS X software (Leica Microsystems). For the bleaching experiments, myofibers were cultured in eight-well glass-bottom dishes and imaged in DMEM Fluorobright (Thermo Fisher Scientific, Waltham, MA, USA) supplemented with 10% L-glutamine (Thermo Fisher Scientific), 1% sodium pyruvate (Thermo Fisher Scientific), and 1% penicillin–streptomycin (Thermo Fisher Scientific). Images were obtained before and after photobleaching in the 490- and 555-nm channels using a confocal microscope with standardized settings to ensure consistency. tdTomato fluorescence was bleached by increasing the laser power at 555 nm and targeting the region of interest (ROI), i.e., the satellite cell cytoplasm, for 15 iterations, ensuring an accurate and reproducible bleaching event. For time-lapse imaging, myofibers were cultured in eight-well glass-bottom dishes using the aforementioned culture medium. Images were automatically captured every 15 min using a confocal microscope and the “xyzt” mode of LAS X software. Intravital muscle fluorescence images were acquired using a combined confocal/multiphoton point-scanning microscope (SP8 MP, Leica Microsystems) as previously described [ 25 ]. For intravital imaging, the mice were anesthetized with 2.5% isoflurane (150–250 ml/min, Piramal Critical Care, Bethlehem, PA, USA) in oxygen for induction, after which a longitudinal skin incision was made on the lateral side of the lower hindlimb to expose the muscle. The mice were maintained at 37°C in the microscope’s incubation chamber with the exposed muscle securely placed on a coverslip attached to the sample holder to minimize motion artifacts. For confocal imaging of fixed myofibers, a confocal microscope or spinning-disk microscope (Leica Microsystems), both equipped with LAS X software, was used. Post-imaging analysis was conducted using ImageJ (US National Institutes of Health, Bethesda, MD, USA), LAS X, and Imaris software (Oxford Instruments, Oxford, UK). Quantification of satellite cells MuSCs located near accumulated dystrophin-EGFP protein were counted on isolated myofibers from Dmd EGFP/y ; Pax7 CreERT2/+ ; Rosa tdT/+ reporter mice directly after isolation (T0) or after up to 48 h of culture (T48). Measure of sarcomere length and MuSC size To measure sarcomere length, mechanically and enzymatically isolated myofibers were incubated with SiR-actin (1:1000 dilution) for 1 h and then mounted on slides. Images were acquired at random locations along each fiber with a confocal microscope, with three independent ROIs and measurements taken per fiber using the draw scalebar tool of LAS X. To measure MuSC size, the diameter of tdTomato-positive MuSCs attached to enzymatically isolated myofibers (T0) was measured using the “draw scalebar” tool of LAS X. Muscle dissociation and cell preparation Adult and juvenile limb muscles were harvested from mice and rats, minced, and incubated with a mix of 0.5 mg/ml collagenase A (Sigma-Aldrich) and 3.5 mg/ml Dispase II (Roche, Basel, Switzerland) or PBS at 37°C and 60 rpm in a shaking water bath for 2 h. Digestion was stopped with IMDM supplemented with 20% fetal bovine serum and GlutaMAX. The cell suspension was filtered through a 70-µm cell strainer (Thermo Fisher Scientific) and then spun at 1300 rpm for 10 min at 4°C to remove large tissue fragments. Before fluorescence-activated cell sorting (FACS), cells were resuspended in HamF10 medium (Sigma-Aldrich) supplemented with 10% horse serum (Gibco). For magnetic-activated cell sorting (MACS), mouse and rat limb muscles were collected, minced, and incubated according to the manufacturer’s protocol using a skeletal muscle dissociation kit for mice and rats (Miltenyi Biotec, Bergisch Gladbach, Germany). Satellite cell isolation Prior to FACS, cell suspensions were prepared following the protocol described by Gayraud-Morel et al. [ 28 ]. The final cell pellet was resuspended in cold HamF10 supplemented with 10% horse serum and filtered through a 40-µm strainer (Thermo Fisher Scientific). MuSCs were sorted using the MoFlo Astrios cell sorter (Beckmann Coulter, Brea, CA, USA) and identified using either GFP or tdTomato staining. Post-isolation, mononuclear cells were collected in HamF10 containing 10% horse serum. For MACS, mouse and rat MuSCs were purified via magnetic separation according to the manufacturer’s protocol using a satellite cell isolation kit for mice and rats (Miltenyi Biotec). MuSCs were isolated by depleting Sca1 + , CD45 + , CD11b + , and CD31 + cells followed by positive selection with anti-alpha-7 integrin microbeads (Miltenyi Biotec). Briefly, muscles from 4–8 mice were harvested, cut carefully into small pieces, and then resuspended in skeletal muscle dissociation kit. The muscles were incubated for 1 h at 37°C, ground using the GentleMACS™ tissue dissociator (Miltenyi Biotec), and incubated for an additional 30 min. Following two washes, 100 µl of a biotinylated antibody cocktail were added per 1 × 10 8 cells, followed by the addition of antibiotin microbeads (200 µl per 1 × 10 8 cells, Miltenyi Biotec). The cells were then incubated for 15 min at 4°C. Finally, the cells were placed on an “LS Column” (Miltenyi Biotec) under the magnetic field of a MACS separator (QuadroMACS™, Miltenyi Biotec). The MuSCs were eluted, whereas the remaining cells were retained in the column. An additional purification step was performed using an “MS Column” (Miltenyi Biotec) positioned within the magnetic field of a MACS separator (OctoMACS™, Miltenyi Biotec), after which the cells were incubated with of anti-alpha 7 integrin microbeads (50 µl per 1 × 10 8 cells). MuSCs were collected following their extraction from the column. Subsequent phenotyping was conducted to ensure their purity. RNA analysis by digital droplet PCR Total RNA was extracted using TRIzol reagent (Thermo Fisher Scientific) in accordance with the manufacturer’s protocol. cDNA was synthesized in 20-µl reactions containing 1000 ng of total RNA and conducted using the LunaScript™ RT SuperMix Kit (New England Biolabs, Ipswich, MA, USA). ddPCR was conducted using 2 µl of cDNA in a 20-µl reaction volume, which included 0.5 µl of the relevant TaqMan probe, 10 µl of ddPCR Supermix for probes (Bio-Rad, Hercules, CA, USA), and 7.5 µl of DNase/RNase-free H 2 O. Each 20-µl reaction mixture was deposited into an eight-channel disposable droplet-generating cartridge. Additionally, 70 µl of droplet-generating oil (Bio-Rad) were loaded into the adjacent oil wells of the cartridge, and the microfluidic chip was placed into the QX200 droplet generator (Bio-Rad). The generated droplets were then transferred to a semi-skirted 96-well PCR plate (Bio-Rad). The plate was heat-sealed using a PX1 PCR plate sealer (Bio-Rad) and amplified on a T100 thermal cycler (Bio-Rad) using the following PCR protocol: initial denaturation at 95°C for 10 min, 40 cycles of 94°C for 30 s and 60°C for 1 min, and final deactivation at 98°C for 10 min. The plate containing the amplified droplets was then analyzed using the QX200 droplet reader and QuantaSoft software (Bio-Rad). Differentiation between target-negative and target-positive droplets was achieved by setting a manual fluorescence amplitude threshold for each TaqMan assay. Glyceraldehyde-3-phosphate dehydrogenase (GAPDH) served as the internal control for normalizing transcript concentrations, with the results reported as target/GAPDH ratios. The primers and probes (Integrated DNA Technologies, Coralville, IA, USA) are provided in Table 1 . Table 1 List of primers used for ddPCR Dmd (Mm.PT.58.42993407) Primer 1: CTT TAC ACA GAG AAA TGA TGC CA Primer 2: GTG TCT CAA CTG GCT TCT CAA Probe: /56-FAM/TT ATG ATA C/ZEN/G GGA CGA ACA GGG AGG A/3IABkFQ/ Pax7 (Mm.PT.58.12398641) Primer 1: GAA GAA GTC CCA GCA CAG C Primer 2: GCT ACC AGT ACA GCC AGT ATG Probe: /56-FAM/CC AAA AAC G/ZEN/T GAG CCT GTC CAC AC/3IABkFQ/ Myod1 (Mm.PT.58.8193525) Primer 1: GAC ACA GCC GCA CTC TT Primer 2: GCT CTG ATG GCA TGA TGG AT Probe: /56-FAM/AC GAC ACC G/ZEN/C CTA CTA CAG TGA GG/3IABkFQ/ Myh1 (Mm.PT.58.8856176) Primer 1: CTG GAT CTT GCG GAA TTT GG Primer 2: GGA CAA ACT GCA ATC AAA GGT C Probe: /56-FAM/AA GCT GAG G/ZEN/A AGC GGA GGA ACA AT/3IABkFQ/ Gapdh (Mm.PT.39a.1) Primer 1: GTG GAG TCA TAC TGG AAC ATG TAG Primer 2: AAT GGT GAA GGT CGG TGT G Probe: /56-FAM/TG CAA ATG G/ZEN/C AGC CCT GGT G/3IABkFQ/ Single-nuclei RNA sequencing Nuclei were isolated using the Chromium protocol kit (CG000505, Rev A, 10X Genomics, Pleasanton, CA, USA) from the TA muscles of Dmd EGFP/wt heterozygous mice using a previously reported protocol with modifications [ 29 ]. A filtration step was added to the protocol using 70-µm filters immediately after tissue lysis. Single-nuclei gel bead-in-emulsions (GEMs) were produced using a Chromium Controller (10X Genomics). Sequencing libraries were created using Chromium Single Cell 3′ Reagent Kit v3.1 (10x Genomics). Nuclei were hybridized with probe barcodes and then partitioned into nanoliter-scale GEMs using a microfluidic chip in the iX Chromium (10X Genomics) to target 20,000 nuclei. The sequencing data were processed into transcript count tables using Cell Ranger Single Cell Software Suite 1.3.1 (10x Genomics). Raw base call files from the Nextseq 500 system (Illumina, San Diego, CA, USA) were demultiplexed using the Cell Ranger mkfastq pipeline into library-specific FASTQ files. The FASTQ files for each library were then processed independently with the Cell Ranger pipeline. Once aligned, barcodes associated with these reads, namely cell identifiers and Unique Molecular Identifiers, were subjected to filtering and correction. Reads associated with retained barcodes were quantified and used to build a transcript count table. The resulting data for each sample were then aggregated using the Cell Ranger aggr pipeline, which performed a between-sample normalization step and concatenated the two transcript count tables. The subsequent visualizations, clustering, and differential expression tests were performed in R (v 3.4.3, The R Foundation for Statistical Computing, Vienna, Austria) using the standard Seurat pipeline (v5). Quality control on aligned and counted reads was performed to retain nuclei with > 200 and < 2500 nFeature RNA and < 5% mitochondrial genes. In total, 9946 nuclei were processed for this analysis. We detected the expression of approximately 1500 genes per nucleus in each of these muscles. Normalization was performed using the LogNormalize algorithm, and determination of variable features using the vst method (2000 features). We performed linear dimensional reduction and constructed a K-nearest neighbor graph using 15 principal components. Nuclei were then clustered using the Louvain algorithm with a resolution of 0.5, and satellite cells were identified according to PAX7 expression. Protein extraction and quantification Total protein was extracted from the cells and muscle sections of TA muscles using RIPA buffer (25 mM Tris-HCl, pH 7.6, 150 mM NaCl, 1% NP-40, 1% sodium deoxycholate) with 5% SDS and 1× protease inhibitors (Thermo Fisher Scientific). Cell lysates were incubated on ice for 30 min and centrifuged at 13,000 rpm for 10 min. The supernatant was collected and stored at − 80°C until capillary electrophoresis. Tissues were grounded in tubes with beads using a homogenizer grinder (Precellys, Bertin Technologies, Montigny-le-Bretonneux, France) at 10,000 rpm for two cycles of 20 s with a 10-s pause between cycles. The lysates were then centrifuged, heated for 3 min at 100°C, and centrifuged again at 13,000 rpm for 10 min at 4°C. The supernatant was collected and stored as previously mentioned. The total protein concentration was determined using the BCA protein assay kit (Thermo Fisher Scientific). Capillary electrophoresis Capillary electrophoresis was conducted on a Jess system (ProteinSimple, San Jose, CA, USA) using a 66–440 kDa separation module and anti-rabbit signal detection for dystrophin. Proteins extracted from lysed cells or muscle tissue collected from 14-day-old Swiss CD-1 mice, adult wild-type rats, and DMD rats were diluted to a concentration of 0.5 µg/µl in 10-fold diluted sample buffer (ProteinSimple). The samples were then mixed with Fluorescent Master Mix (ProteinSimple) and heated at 95°C for 5 min. Samples, blocking reagent, primary antibodies, HRP-conjugated secondary antibodies, and chemiluminescent substrate (ProteinSimple) were placed in a plate from the capillary Western blot kit (ProteinSimple). The default settings included stacking and separation at 475 V for 30 min, blocking for 5 min, primary and secondary antibody incubation for 30 min, and chemiluminescence detection for 15 min (exposure for 1, 2, 4, 8 16, 32, 64, 128, and 512 s). Calibration curves required chemiluminescence peaks exceeding 150,000 units and R2 greater than 0.95. Statistical analysis The data were analyzed using GraphPad Prism 7 software (GraphPad, Boston, MA, USA). The number of replicates is specific in each figure legend as necessary. Student’s t -test was used for comparisons between two groups, whereas one- and two- way ANOVA followed by multiple comparison tests was employed to compare three or more groups. Significance was denoted by P < 0.05. Declarations Ethics, Consent to Participate, and Consent to Publish declarations All animal experiments were conducted according to the National and European legislation and institutional guidelines for the care and use of laboratory animals approved by the French government (Ministère de l’Enseignement Supérieur et de la Recherche, autorisation APAFiS #44987-2023092714211861 v4 approved on February 5, 2024 “Evaluation of gene therapy approaches and antisense strategies for genetic diseases” and #34462-2021121716045314 v6 Approved on March 15, 2022 Study of dystrophin expression in the muscle stem cell niche). No human participants were involved in this study; therefore, consent to participate and consent to publish are not applicable. The authors declare that they have not use AI-generated work in this manuscript. All authors read the final version of the manuscript and gave their permission for publication. Data availability statement RNA sequencing data have been deposited in the Gene Expression Omnibus (GEO) under accession number GSE309529: https://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE309529. All data generated or analyzed during this study are included in the manuscript and supporting files; source data files have been provided for all figures. Acknowledgements We thank Valentina Taglietti and Frédéric Relaix from Institut Mondor de Recherche Biomédicale (IMRB, Creteil) for providing the Pax7 CreERT2 ;Rosa tdT mouse line and the DMD rats muscles. We thank Shahragim Tajbakhsh for his valuable scientific advice and constructive feedback. Funding This work was supported by the Association Française contre les Myopathies (AFM, #23802 and #29169 France), and the Agence Nationale de la Recherche (ANR-23-CE13-0032-03 “DYSCO”, France). Funders were not involved in the design, analysis and reporting of the study. Authors’ contributions HA, PM, and SF designed the study; MM, AM, LR, GM, AG, AJM, BE, LS, AS, TM, VM, SF and HA performed the research; MVP and MS, created the Dmd EGFP and Dmd EGFP-mdx reporter mouse lines; MM, AM, GM, AG, BE, LS, AS, HA, and LG analyzed data; MM, AM and HA wrote the manuscript. MM–Meriem Matouk AM–Adrien Morin LR–Lucie Royer GM–Gaspard Macaux AG–Aris Gaci AJM–Aude Jobart-Malfait BE–Brendan Evano LS–Liza Sarde AS–Amalia Stantzou MVP–Mina V. Petkova TM–Tudor Manoliu MS–Markus Schuelke LG–Luis Garcia PM–Pascal Maire VM–Vincent Mirouse SF–Sestina Falcone HA–Helge Amthor Competing interests The authors do not state any financial nor nonfinancial competing interest. References Davies KE, Nowak KJ. 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Supplementary Files MovieS1.mp4 SIAppendix.docx Cite Share Download PDF Status: Under Review Version 1 posted Editorial decision: Revision requested 11 Feb, 2026 Reviews received at journal 07 Feb, 2026 Reviews received at journal 01 Feb, 2026 Reviewers agreed at journal 23 Jan, 2026 Reviewers agreed at journal 21 Jan, 2026 Reviewers invited by journal 21 Jan, 2026 Editor assigned by journal 13 Jan, 2026 Submission checks completed at journal 21 Dec, 2025 First submitted to journal 19 Dec, 2025 You are reading this latest preprint version Research Square lets you share your work early, gain feedback from the community, and start making changes to your manuscript prior to peer review in a journal. As a division of Research Square Company, we’re committed to making research communication faster, fairer, and more useful. We do this by developing innovative software and high quality services for the global research community. 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Also discoverable on Platform About Our Team In Review Editorial Policies Advisory Board Help Center Resources Author Services Accessibility API Access RSS feed Manage Cookie Preferences © Research Square 2026 | ISSN 2693-5015 (online) Privacy Policy Terms of Service Do Not Sell My Personal Information {"props":{"pageProps":{"initialData":{"identity":"rs-7997649","acceptedTermsAndConditions":true,"allowDirectSubmit":false,"archivedVersions":[],"articleType":"Research Article","associatedPublications":[],"authors":[{"id":578339390,"identity":"675baeab-9f1a-47e9-a3d8-bf61957b71ba","order_by":0,"name":"Meriem Matouk","email":"","orcid":"","institution":"Université Paris-Saclay, Université de Versailles Saint-Quentin-en-Yvelines","correspondingAuthor":false,"prefix":"","firstName":"Meriem","middleName":"","lastName":"Matouk","suffix":""},{"id":578339391,"identity":"a0039a1c-62c2-4f6a-aae1-78baf62e63b7","order_by":1,"name":"Adrien Morin","email":"","orcid":"","institution":"Université Paris-Saclay, Université de Versailles Saint-Quentin-en-Yvelines","correspondingAuthor":false,"prefix":"","firstName":"Adrien","middleName":"","lastName":"Morin","suffix":""},{"id":578339400,"identity":"ca1fb014-568d-4856-8e11-0046159d6ecb","order_by":2,"name":"Lucie Royer","email":"","orcid":"","institution":"Université Paris-Saclay, Université de Versailles Saint-Quentin-en-Yvelines","correspondingAuthor":false,"prefix":"","firstName":"Lucie","middleName":"","lastName":"Royer","suffix":""},{"id":578339402,"identity":"034b3dcb-5679-4429-b194-cf0dda1a95c0","order_by":3,"name":"Gaspard Macaux","email":"","orcid":"","institution":"Université de Paris Cité, Institut Cochin, INSERM, CNRS","correspondingAuthor":false,"prefix":"","firstName":"Gaspard","middleName":"","lastName":"Macaux","suffix":""},{"id":578339411,"identity":"f01be8ec-fe3b-4a69-80b0-33819d0ea515","order_by":4,"name":"Aris Gaci","email":"","orcid":"","institution":"Université Paris-Saclay, Université de Versailles Saint-Quentin-en-Yvelines","correspondingAuthor":false,"prefix":"","firstName":"Aris","middleName":"","lastName":"Gaci","suffix":""},{"id":578339418,"identity":"51c54551-6b94-4aaf-8b7f-d02832c7853d","order_by":5,"name":"Aude Jobart-Malfait","email":"","orcid":"","institution":"UFR Simone Veil - Université de Versailles Saint-Quentin-en-Yvelines","correspondingAuthor":false,"prefix":"","firstName":"Aude","middleName":"","lastName":"Jobart-Malfait","suffix":""},{"id":578339421,"identity":"6cbc0b9e-e4d0-4b70-a580-e63bec17cdcf","order_by":6,"name":"Brendan Evano","email":"","orcid":"","institution":"Institut Pasteur","correspondingAuthor":false,"prefix":"","firstName":"Brendan","middleName":"","lastName":"Evano","suffix":""},{"id":578339429,"identity":"70bd1221-76a0-494d-896d-b00861a3a7ab","order_by":7,"name":"Liza Sarde","email":"","orcid":"","institution":"Institut Pasteur","correspondingAuthor":false,"prefix":"","firstName":"Liza","middleName":"","lastName":"Sarde","suffix":""},{"id":578339433,"identity":"4309328b-7e58-4e6e-9047-5fd2a48a2dff","order_by":8,"name":"Amalia Stantzou","email":"","orcid":"","institution":"Université Paris-Saclay, Université de Versailles Saint-Quentin-en-Yvelines","correspondingAuthor":false,"prefix":"","firstName":"Amalia","middleName":"","lastName":"Stantzou","suffix":""},{"id":578339436,"identity":"7f92a847-6b35-4560-987f-ea050f274365","order_by":9,"name":"Mina V. 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\u003cem\u003eDmd\u003c/em\u003e\u003csup\u003e\u003cem\u003eEGFP/y\u003c/em\u003e\u003c/sup\u003e\u003cem\u003e;Pax7\u003c/em\u003e\u003csup\u003e\u003cem\u003eCreERT2/+\u003c/em\u003e\u003c/sup\u003e\u003cem\u003e;Rosa\u003c/em\u003e\u003csup\u003e\u003cem\u003etdT\u003c/em\u003e\u003c/sup\u003e\u003csup\u003e/+\u003c/sup\u003e mice. After tamoxifen treatment, tdTomato\u003csup\u003e+\u003c/sup\u003e MuSCs (red) appeared in the same area as dystrophin-EGFP (green).\u003cstrong\u003e (A)\u003c/strong\u003e Two MuSCs (yellow arrows) displaying an elongated shape that are positioned in contact to the dystrophin-EGFP\u003csup\u003e+\u003c/sup\u003e sarcolemmata of neighboring myofibers (white arrowheads). The three images present the tdTomato signal, dystrophin-EGFP signal, and merged signal, respectively. \u003cstrong\u003e(B) \u003c/strong\u003eA representative quiescent MuSC (left image, yellow arrow) with its cellular extensions, sarcolemmal dystrophin-EGFP (middle image), and merged view (right image). Notably, dystrophin-EGFP was evenly distributed along the sarcolemma of one adjacent myofiber, whereas its expression was reduced or absent in the other ajacent myofiber (white arrowheads). Scale bars: 25 and 10 µm. \u003cstrong\u003e(C)\u003c/strong\u003e \u003cem\u003eEx-vivo\u003c/em\u003e confocal microscopy of a representative quiescent MuSC and its cellular extensions (yellow arrows) on mechanically isolated EDL myofibers from \u003cem\u003ePax7\u003c/em\u003e\u003csup\u003e\u003cem\u003eCreERT2/CreERT2\u003c/em\u003e\u003c/sup\u003e\u003cem\u003e;Rosa\u003c/em\u003e\u003csup\u003e\u003cem\u003etdT/tdT\u003c/em\u003e\u003c/sup\u003e mice. Sarcolemmal dystrophin on the apical side of the MuSC was markedly diminished (white arrow).\u003cstrong\u003e \u003c/strong\u003eMyofibers were stained with anti-dystrophin antibody (green, white arrow), SiR-actin (magenta), and DAPI (cyan) to visualize nuclei. Scale bar: 10 µm.\u003cstrong\u003e (D)\u003c/strong\u003e Confocal microscopy of TA muscle cryosections from \u003cem\u003eDmd\u003c/em\u003e\u003csup\u003e\u003cem\u003eEGFP/y\u003c/em\u003e\u003c/sup\u003e\u003cem\u003e;Pax7\u003c/em\u003e\u003csup\u003e\u003cem\u003eCreERT2/+\u003c/em\u003e\u003c/sup\u003e\u003cem\u003e;Rosa\u003c/em\u003e\u003csup\u003e\u003cem\u003etdT/+\u003c/em\u003e\u003c/sup\u003e mice highlights sarcolemmal dystrophin-EGFP and tdTomato\u003csup\u003e+\u003c/sup\u003e MuSCs. In examples 1 and 2, dystrophin-EGFP is present along the sarcolemma of one myofiber but is notably absent from the adjacent fiber precisely at the satellite cell’s location (framed area, white arrowheads). An enlarged views of the framed regions are presented in the rightmost images. In total, 113 MuSCs from five different individuals were used. Scale bar: 10 µm.\u003c/p\u003e","description":"","filename":"image1.jpeg","url":"https://assets-eu.researchsquare.com/files/rs-7997649/v1/7c0bef20c8f5060ed131ed40.jpeg"},{"id":101170627,"identity":"8d19d977-b95f-4adc-8cb7-2ff7e31d2485","added_by":"auto","created_at":"2026-01-27 00:04:27","extension":"jpeg","order_by":2,"title":"Figure 2","display":"","copyAsset":false,"role":"figure","size":352411,"visible":true,"origin":"","legend":"\u003cp\u003e\u003cem\u003e\u003cstrong\u003eEx-vivo\u003c/strong\u003e\u003c/em\u003e\u003cstrong\u003e imaging of MuSCs.\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003e\u003cem\u003e\u003cstrong\u003e(A\u003c/strong\u003e\u003c/em\u003e\u003cem\u003e) Ex-vivo\u003c/em\u003e confocal microscopy of live myofibers from \u003cem\u003eDmd\u003c/em\u003e\u003csup\u003e\u003cem\u003eEGFP/y\u003c/em\u003e\u003c/sup\u003e\u003cem\u003e;Pax7\u003c/em\u003e\u003csup\u003e\u003cem\u003eCreERT2/+\u003c/em\u003e\u003c/sup\u003e\u003cem\u003e;Rosa\u003c/em\u003e\u003csup\u003e\u003cem\u003etdT/+\u003c/em\u003e\u003c/sup\u003e mice immediately following enzymatic dissociation (T0). Red fluorescence represents tdTomato\u003csup\u003e+\u003c/sup\u003e MuSCs, whereas green fluorescence denotes dystrophin-EGFP. Dystrophin-EGFP accumulated on the apical side (white arrowheads) of MuSCs (yellow arrow), as observed in the frontal (left) and lateral (right) views (15 MuSCs from four mice). \u003cstrong\u003e(B)\u003c/strong\u003e Examples of time-lapse confocal microscopy to track individual MuSCs attached to isolated myofibers. \u003cstrong\u003eCell 1:\u003c/strong\u003e Dystrophin-EGFP accumulated on the apical side (white arrowheads) of the MuSC (yellow arrow) at T44. The MuSC then migrated to the opposite side of the myofiber, and it was traceable at a different Z-position at T46 (yellow arrow; 23.2 µm Z difference versus the previous image [29 Z-steps of 0.8 μm each]), leaving behind a footprint of dystrophin-EGFP that persisted at T52 (white arrowheads). \u003cstrong\u003eCell 2:\u003c/strong\u003e Dystrophin-EGFP accumulated on the basal side (white arrowheads) of the MuSC (yellow arrow) at T46. The MuSC squeezed through the basally located dystrophin-EGFP (white arrowheads) at T48 and T50, but it was no longer visible at T52. However, the detected dystrophin-EGFP remained in place (white arrowheads). \u003cstrong\u003eCell 3:\u003c/strong\u003e Dystrophin-EGFP accumulated on the apical and lateral sides (white arrowheads) of the MuSC (yellow arrow) at T44, but it was subsequently no longer visible, leaving behind a footprint of dystrophin-EGFP that persisted up to T52 (white arrowheads). \u003cstrong\u003eCell 4:\u003c/strong\u003e Dystrophin-EGFP accumulated on the basal side (white arrowheads) of the MuSC (yellow arrow) at T46. The MuSCs moved, but it was out of focus at T52. Conversely, the detected dystrophin-EGFP remained in place (white arrowheads) Scale bars: 10 µm.\u003c/p\u003e","description":"","filename":"image2.jpeg","url":"https://assets-eu.researchsquare.com/files/rs-7997649/v1/73fece5e6a897f3ecfe5d87e.jpeg"},{"id":101206149,"identity":"2e77c2b0-be0f-4d68-9943-948954625b10","added_by":"auto","created_at":"2026-01-27 09:55:26","extension":"jpeg","order_by":3,"title":"Figure 3","display":"","copyAsset":false,"role":"figure","size":499314,"visible":true,"origin":"","legend":"\u003cp\u003e\u003cstrong\u003eSarcolemmal origin of MuSC-associated dystrophin.\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e(A)\u003c/strong\u003e\u0026nbsp;\u003cem\u003eEx-vivo\u003c/em\u003e\u0026nbsp;confocal imaging of isolated myofibers from adult \u003cem\u003eDmd\u003c/em\u003e\u003csup\u003e\u003cem\u003eEGFP/y\u003c/em\u003e\u003c/sup\u003e\u003cem\u003e;Pax7\u003c/em\u003e\u003csup\u003e\u003cem\u003eCreERT2/+\u003c/em\u003e\u003c/sup\u003e\u003cem\u003e;Rosa\u003c/em\u003e\u003csup\u003e\u003cem\u003etdT/+\u003c/em\u003e\u003c/sup\u003e mice stained with the membrane marker MemBright (magenta). Dystrophin-EGFP, green; tdTomato, red. Dystrophin-EGFP and MemBright accumulated and colocalized on the apical side (white arrowheads) of MuSCs (yellow arrows) and at the myotendinous junction (yellow arrowheads; 16 MuSCs from seven myofibers). \u003cstrong\u003e(B)\u003c/strong\u003e\u0026nbsp;Frequency analysis of GFP\u003csup\u003e+\u003c/sup\u003e and tdTomato\u003csup\u003e+\u003c/sup\u003e cell populations following FACS of MuSCs from \u003cem\u003eTg:Pax7-nGFP\u003c/em\u003e mice (positive GFP control, upper diagram) and \u003cem\u003eDmd\u003c/em\u003e\u003csup\u003e\u003cem\u003eEGFP/y\u003c/em\u003e\u003c/sup\u003e\u003cem\u003e;Pax7\u003c/em\u003e\u003csup\u003e\u003cem\u003eCreERT2/+\u003c/em\u003e\u003c/sup\u003e\u003cem\u003e;Rosa\u003c/em\u003e\u003csup\u003e\u003cem\u003etdT/+\u003c/em\u003e\u003c/sup\u003e mice (lower diagram). MuSCs only positive for GFP clustered in the “GFP” quadrant, whereas MuSCs only positive for tdTomato clustered in the “tdTomato” quadrant. Double-positive MuSCs, expected in the R6 quadrant, were absent. \u003cstrong\u003e(C)\u003c/strong\u003e\u0026nbsp;Airyscan microscopy of isolated myofibers from adult rats after 48 h of culture. Co-immunocytochemistry depicts dystrophin (green), PAX7- (orange), and DAPI-stained nuclei (blue). A MuSC doublet was present in an apical-to-basal orientation in which only the basally located MuSC was PAX7\u003csup\u003e+\u003c/sup\u003e (yellow arrow). Dystrophin accumulated (white arrows) around the apically located MuSC. The same cells are presented in the four images. The top right image depicts an overexposure of the dystrophin signal. The lower images depict a 3D reconstruction, with the bottom left image combining dystrophin and PAX7 staining and the bottom right image combining dystrophin and DAPI staining. Dystrophin accumulated in continuation with the sarcolemma (white arrows). N = 16 MuSCs. Scale bars: 20 µm.\u003c/p\u003e","description":"","filename":"image3.jpeg","url":"https://assets-eu.researchsquare.com/files/rs-7997649/v1/0c49fa3d8bd37a51d75fe350.jpeg"},{"id":101206478,"identity":"108310a4-8412-41e2-aa75-57555ce0a55d","added_by":"auto","created_at":"2026-01-27 09:56:22","extension":"jpeg","order_by":4,"title":"Figure 4","display":"","copyAsset":false,"role":"figure","size":261629,"visible":true,"origin":"","legend":"\u003cp\u003e\u003cstrong\u003eEmergence of \u003c/strong\u003e\u003cem\u003e\u003cstrong\u003eDmd\u003c/strong\u003e\u003c/em\u003e\u003cstrong\u003e transcripts and dystrophin protein during myogenic lineage progression.\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eMuSCs were freshly derived from 7-day-old wild-type mice or cultured for 3 and 13 days to obtain myoblasts (MB) and myotubes (MT), respectively. \u003cstrong\u003e(A)\u003c/strong\u003eDiagram of ddPCR analysis depicts \u003cem\u003eDmd\u003c/em\u003e transcript levels in MuSCs, MB, MT (3–7 mice; seven culture replicates), and TA muscle (n = 3). Statistical significance was assessed using \u003cstrong\u003eone-way ANOVA\u003c/strong\u003e followed by \u003cstrong\u003eTukey’s multiple comparisons test. (B)\u003c/strong\u003e Diagram of capillary electrophoresis depicts the quantity of dystrophin protein in MuSCs, MB, MT (6–7 mice; 5–7 culture replicates) and positive-control muscle (TA; n = 2) using an antibody targeting the C-terminal region of dystrophin.\u003c/p\u003e","description":"","filename":"image4.jpeg","url":"https://assets-eu.researchsquare.com/files/rs-7997649/v1/2fd9763e5915f9050d3695c5.jpeg"},{"id":101297444,"identity":"d77e285c-ddba-4810-8f24-762f4de2856b","added_by":"auto","created_at":"2026-01-28 09:27:13","extension":"pdf","order_by":0,"title":"","display":"","copyAsset":false,"role":"manuscript-pdf","size":2943403,"visible":true,"origin":"","legend":"","description":"","filename":"manuscript.pdf","url":"https://assets-eu.researchsquare.com/files/rs-7997649/v1/6db2591e-fa67-4f36-b98d-d64dbb1554c1.pdf"},{"id":101205835,"identity":"caf0e702-32a3-4481-a962-d1ba3aaaed08","added_by":"auto","created_at":"2026-01-27 09:50:21","extension":"mp4","order_by":1,"title":"","display":"","copyAsset":false,"role":"supplement","size":7927888,"visible":true,"origin":"","legend":"","description":"","filename":"MovieS1.mp4","url":"https://assets-eu.researchsquare.com/files/rs-7997649/v1/45cb603baf7854891dc4fe32.mp4"},{"id":101206346,"identity":"6a0f8228-9beb-4c81-81fe-535a51da40f5","added_by":"auto","created_at":"2026-01-27 09:56:01","extension":"docx","order_by":2,"title":"","display":"","copyAsset":false,"role":"supplement","size":9540857,"visible":true,"origin":"","legend":"","description":"","filename":"SIAppendix.docx","url":"https://assets-eu.researchsquare.com/files/rs-7997649/v1/f084946fb3416030384c25d2.docx"}],"financialInterests":"No competing interests reported.","formattedTitle":"No evidence of the presence of dystrophin protein in rodent muscle stem cells","fulltext":[{"header":"Significance Statement","content":"\u003cp\u003eDuchenne muscular dystrophy results from the absence of dystrophin, leading to progressive muscle wasting. Although dystrophin is essential for myofiber stability, its role in muscle stem cells (MuSCs) has remained unclear. Using live imaging in dystrophin-EGFP mice and cross-species analysis, we show that the dystrophin signal in MuSCs originates from sarcolemmal membranes rather than intrinsic expression. These findings demonstrate that dystrophin is dispensable for MuSC regenerative function and resolve a long-standing controversy about its cellular role.\u003c/p\u003e"},{"header":"Introduction","content":"\u003cp\u003eProgressive skeletal muscle wasting in Duchenne muscular dystrophy (DMD) is traditionally attributed to the inherited absence of sarcolemmal dystrophin, which decreases the resistance of myofibers to mechanical stress, resulting in fiber necrosis, inflammation, chronic cycles of fiber degeneration and regeneration, and fibrotic and fatty transformation of muscle [\u003cspan citationid=\"CR1\" class=\"CitationRef\"\u003e1\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR2\" class=\"CitationRef\"\u003e2\u003c/span\u003e]. In addition to this classical view, inefficient muscle regeneration has been suspected to contribute to muscle wasting in DMD attributable to defective muscle stem cell (MuSC) function [\u003cspan citationid=\"CR3\" class=\"CitationRef\"\u003e3\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR4\" class=\"CitationRef\"\u003e4\u003c/span\u003e].\u003c/p\u003e \u003cp\u003eMuSCs are located between the sarcolemma and basal lamina of myofibers, and they are generally quiescent in mature muscle [\u003cspan citationid=\"CR5\" class=\"CitationRef\"\u003e5\u003c/span\u003e]. Myofiber injury, such as that in DMD, activates MuSCs, which begin to proliferate and eventually differentiate to replace damaged muscle fibers or self-renew to replenish the MuSC pool. The proper balance between self-renewal and myogenic progression is critical for ensuring the lifelong regenerative potential of skeletal muscle [\u003cspan citationid=\"CR6\" class=\"CitationRef\"\u003e6\u003c/span\u003e], [\u003cspan citationid=\"CR7\" class=\"CitationRef\"\u003e7\u003c/span\u003e]. Self-renewal in MuSCs requires asymmetric divisions, in which one daughter cell retains stemness and the other undergoes myogenic commitment, whereas symmetric divisions expand the cell pool by generating identical daughter cells [\u003cspan citationid=\"CR8\" class=\"CitationRef\"\u003e8\u003c/span\u003e], [\u003cspan citationid=\"CR9\" class=\"CitationRef\"\u003e9\u003c/span\u003e], [\u003cspan citationid=\"CR10\" class=\"CitationRef\"\u003e10\u003c/span\u003e]. Asymmetric division requires an apicobasal orientation of the dividing MuSCs relative to the myofiber, whereas symmetric division occurs in a planar orientation [\u003cspan citationid=\"CR11\" class=\"CitationRef\"\u003e11\u003c/span\u003e], [\u003cspan citationid=\"CR12\" class=\"CitationRef\"\u003e12\u003c/span\u003e].\u003c/p\u003e \u003cp\u003eOnly the presence of the full-length dystrophin isoform Dp427 and the short ubiquitously expressed Dp71 have been documented in MuSCs [\u003cspan citationid=\"CR13\" class=\"CitationRef\"\u003e13\u003c/span\u003e, \u003cspan citationid=\"CR14\" class=\"CitationRef\"\u003e14\u003c/span\u003e]. A polarized distribution of full-length dystrophin in activated MuSCs is critical for proper MuSC self-renewal [\u003cspan citationid=\"CR14\" class=\"CitationRef\"\u003e14\u003c/span\u003e]. The spectrin-like repeat domain of dystrophin has been demonstrated to bind to MAP/microtubule affinity-regulating kinase 2 [\u003cspan citationid=\"CR14\" class=\"CitationRef\"\u003e14\u003c/span\u003e, \u003cspan citationid=\"CR15\" class=\"CitationRef\"\u003e15\u003c/span\u003e] and regulate polarity establishment for asymmetric versus symmetric self-renewal MuSC division [\u003cspan citationid=\"CR16\" class=\"CitationRef\"\u003e16\u003c/span\u003e, \u003cspan citationid=\"CR17\" class=\"CitationRef\"\u003e17\u003c/span\u003e].\u003c/p\u003e \u003cp\u003eThe polarity of the dystrophin distribution in activated MuSCs has been primarily studied in isolated myofibers from murine models, but its polarity has also been detected in human fiber cultures derived from postmortem tissue [\u003cspan citationid=\"CR14\" class=\"CitationRef\"\u003e14\u003c/span\u003e, \u003cspan citationid=\"CR18\" class=\"CitationRef\"\u003e18\u003c/span\u003e]. Dystrophin is absent in \u003cem\u003eDmd\u003c/em\u003e\u003csup\u003e\u003cem\u003emdx/y\u003c/em\u003e\u003c/sup\u003e mice that harbor a nonsense mutation in \u003cem\u003eDmd\u003c/em\u003e exon 23 and lack the full-length dystrophin isoform Dp427 [\u003cspan citationid=\"CR14\" class=\"CitationRef\"\u003e14\u003c/span\u003e]. The loss of dystrophin impairs the establishment of MuSC polarity and results in the loss of apicobasal asymmetric cell division in favor of planar symmetric division, thereby altering the balance between MuSC self-renewal and myogenic commitment. This loss in polarity affects MuSCs proliferation, thereby impairing lineage progression of myoblasts and explaining the regeneration deficit of DMD muscle [\u003cspan citationid=\"CR14\" class=\"CitationRef\"\u003e14\u003c/span\u003e].\u003c/p\u003e \u003cp\u003eWe previously generated a \u003cem\u003eDmd\u003c/em\u003e\u003csup\u003e\u003cem\u003eEGFP\u003c/em\u003e\u003c/sup\u003e reporter mouse line harboring an EGFP-tag fused to the C-terminal end of dystrophin. The resulting dystrophin-EGFP fusion protein allows \u003cem\u003ein vivo\u003c/em\u003e and \u003cem\u003eex-vivo\u003c/em\u003e live imaging of dystrophin isoforms carrying the C-terminal region [\u003cspan citationid=\"CR19\" class=\"CitationRef\"\u003e19\u003c/span\u003e]. Using live fluorescence imaging, we investigated the dynamics of dystrophin polarization in activated MuSCs. Our data illustrated that MuSCs themselves were devoid of dystrophin. In fact, dystrophin, which accumulated near activated MuSCs, belonged to the sarcolemma. We therefore systematically analyzed the relationship between dystrophin and MuSCs in murine and rat animal models.\u003c/p\u003e"},{"header":"Results","content":"\u003cdiv id=\"Sec3\" class=\"Section2\"\u003e \u003ch2\u003eLive observation of dystrophin in the MuSC niche\u003c/h2\u003e \u003cp\u003eWe examined skeletal muscle from \u003cem\u003eDmd\u003c/em\u003e\u003csup\u003e\u003cem\u003eEGFP/y\u003c/em\u003e\u003c/sup\u003e;\u003cem\u003ePax7\u003c/em\u003e\u003csup\u003e\u003cem\u003eCreERT2/+\u003c/em\u003e\u003c/sup\u003e;\u003cem\u003eRosa\u003c/em\u003e\u003csup\u003e\u003cem\u003etdT/+\u003c/em\u003e\u003c/sup\u003e reporter mice to visualize native dystrophin-EGFP fusion protein and tdTomato\u003csup\u003e+\u003c/sup\u003e MuSCs following tamoxifen treatment. Dystrophin-EGFP and tdTomato signals were easily identified \u003cem\u003ein vivo\u003c/em\u003e, \u003cem\u003eex vivo\u003c/em\u003e, and \u003cem\u003ein vitro\u003c/em\u003e, as well as by immunohistochemistry. A limitation of the system was that tdTomato faintly photoconverted to the GFP spectrum, and this effect was increasingly noticeable following prolonged excitation \u003cb\u003e(Figure \u003cspan refid=\"MOESM1\" class=\"InternalRef\"\u003eS1\u003c/span\u003eA)\u003c/b\u003e. This sometimes caused a faint GFP signal in the cytoplasm of MuSCs, which we considered non-specific and unrelated to dystrophin-EGFP.\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003cp\u003e \u003cem\u003eIn vivo\u003c/em\u003e multiphoton microscopy of the TA muscle of adult \u003cem\u003eDmd\u003c/em\u003e\u003csup\u003e\u003cem\u003eEGFP/y\u003c/em\u003e\u003c/sup\u003e;\u003cem\u003ePax7\u003c/em\u003e\u003csup\u003e\u003cem\u003eCreERT2/+\u003c/em\u003e\u003c/sup\u003e;\u003cem\u003eRosa\u003c/em\u003e\u003csup\u003e\u003cem\u003etdT/+\u003c/em\u003e\u003c/sup\u003e mice revealed that MuSCs had an elongated shape with long tdTomato\u003csup\u003e+\u003c/sup\u003eprojections in the expected region near the dystrophin-EGFP\u003csup\u003e+\u003c/sup\u003e sarcolemma of myofibers \u003cb\u003e(\u003c/b\u003eFig.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e1\u003c/span\u003eA\u003cb\u003e)\u003c/b\u003e. Curiously, dystrophin-EGFP was continuously present at homogeneous levels along the sarcolemma of one adjacent myofiber, whereas sarcolemmal dystrophin-EGFP protein was either reduced or absent from the other bordering myofiber \u003cb\u003e(\u003c/b\u003eFig.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e1\u003c/span\u003eB\u003cb\u003e)\u003c/b\u003e. We next immediately fixed EDL muscle of adult \u003cem\u003ePax7\u003c/em\u003e\u003csup\u003e\u003cem\u003eCreERT2/CreERT2\u003c/em\u003e\u003c/sup\u003e;\u003cem\u003eRosa\u003c/em\u003e\u003csup\u003e\u003cem\u003etdT/tdT\u003c/em\u003e\u003c/sup\u003e mice in PFA following dissection and then mechanically isolated myofibers, which were immunostained to visualize dystrophin. Confocal microscopy revealed that tdTomato\u003csup\u003e+\u003c/sup\u003e MuSCs were attached to the muscle fiber, and they featured cell projections in accordance with the elongated shape observed \u003cem\u003ein vivo.\u003c/em\u003e Sarcolemmal dystrophin was absent on the apical side of MuSCs (side of MuSCs facing the sarcolemma), in line with previously published observations [\u003cspan citationid=\"CR30\" class=\"CitationRef\"\u003e30\u003c/span\u003e] \u003cb\u003e(\u003c/b\u003eFig.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e1\u003c/span\u003eC\u003cb\u003e)\u003c/b\u003e. Native fluorescence on the cryosections of PFA-fixed TA muscles from adult \u003cem\u003eDmd\u003c/em\u003e\u003csup\u003e\u003cem\u003eEGFP/y\u003c/em\u003e\u003c/sup\u003e;\u003cem\u003ePax7\u003c/em\u003e\u003csup\u003e\u003cem\u003eCreERT2/+\u003c/em\u003e\u003c/sup\u003e;\u003cem\u003eRosa\u003c/em\u003e\u003csup\u003e\u003cem\u003etdT/+\u003c/em\u003e\u003c/sup\u003e mice confirmed the strong reduction or absence of sarcolemmal dystrophin-EGFP on one side of all MuSCs \u003cb\u003e(\u003c/b\u003eFig.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e1\u003c/span\u003eD\u003cb\u003e)\u003c/b\u003e, in line with previous findings [\u003cspan citationid=\"CR31\" class=\"CitationRef\"\u003e31\u003c/span\u003e]. Similarly, in \u003cem\u003ePax7\u003c/em\u003e\u003csup\u003e\u003cem\u003eCreERT2/+\u003c/em\u003e\u003c/sup\u003e;\u003cem\u003eRosa\u003c/em\u003e\u003csup\u003e\u003cem\u003etdT/+\u003c/em\u003e\u003c/sup\u003e mice, another member of the dystrophin associated protein complex (DAPC), β-sarcoglycan, was also absent from the same side of MuSCs \u003cb\u003e(Figure \u003cspan refid=\"MOESM1\" class=\"InternalRef\"\u003eS1\u003c/span\u003eB)\u003c/b\u003e.\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003cp\u003eNext, we enzymatically isolated myofibers (a procedure that lasts about 2\u0026ndash;3 hours), thereby permitting live \u003cem\u003eex-vivo\u003c/em\u003e observation of tdTomato\u003csup\u003e+\u003c/sup\u003e MuSCs and dystrophin-EGFP during culture. Once isolated, the muscle fiber was detached from its tendinous attachments. We first sought to clarify whether isolation influenced the length of myofiber sarcomeres by comparing fibers isolated \u003cem\u003evia\u003c/em\u003e enzymatic and mechanical isolation. In our previously published work, we recorded a sarcomere size of 3.3 \u0026micro;m in \u003cem\u003eDmd\u003c/em\u003e\u003csup\u003e\u003cem\u003eEGFP\u003c/em\u003e\u003c/sup\u003e mice \u003cem\u003ein vivo\u003c/em\u003e [\u003cspan citationid=\"CR32\" class=\"CitationRef\"\u003e32\u003c/span\u003e]. In this study, the sarcomere length was 2.36 \u0026micro;m in mechanically isolated fibers, versus 1.82 \u0026micro;m following enzymatic fiber isolation \u003cb\u003e(Figure \u003cspan refid=\"MOESM1\" class=\"InternalRef\"\u003eS1\u003c/span\u003eC)\u003c/b\u003e. Taken together, the data indicate that myofibers contracted by approximately 23% during enzymatic isolation, and this likely decreased sarcolemmal tension. In addition, most MuSCs retracted their cell projections and exhibited a round morphology, and these retracted MuSCs usually remained attached to the fibers.\u003c/p\u003e \u003cp\u003eThe MuSCs displayed varying levels of dystrophin-EGFP accumulation. Based on their orientation, we distinguished three regions: an apical side facing the fiber\u0026rsquo;s sarcolemma, lateral sides on either edge, and a basal side oriented toward the basal lamina. Upon isolation (T0), we frequently observed increased dystrophin-EGFP accumulation on the apical side (sarcolemmal side) of these retracted MuSCs \u003cb\u003e(\u003c/b\u003eFig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e2\u003c/span\u003eA\u003cb\u003e)\u003c/b\u003e, which appeared in continuation of the sarcolemma and contrasted the findings \u003cem\u003ein vivo\u003c/em\u003e, after cryosectioning, or following mechanical fiber isolation. Of note, this apically increased dystrophin was similar to the diameter of MuSCs of approximately 8 \u0026micro;m \u003cb\u003e(Figure S2A)\u003c/b\u003e, being 10-fold smaller than the basal sarcolemmal dystrophin unit (BSDU), which is the sarcolemmal extension of a myonuclear domain [\u003cspan citationid=\"CR32\" class=\"CitationRef\"\u003e32\u003c/span\u003e]. This localized and rapid dystrophin accumulation suggests that this event is not driven by a myonuclear process. The apical condensation of dystrophin increased at T36 and T48 during myofiber culture and often extended laterally, whereas dystrophin-EGFP rarely accumulated on the basal side of MuSCs \u003cb\u003e(Figure S2B)\u003c/b\u003e. We quantified the number of MuSCs in respect to dystrophin-EGFP accumulation and performed time-lapse imaging of cultured myofibers isolated from \u003cem\u003eDmd\u003c/em\u003e\u003csup\u003e\u003cem\u003eEGFP\u003c/em\u003e\u003c/sup\u003e mice. We used PAX7 immunochemistry to visualize MuSCs. At T0, dystrophin-EGFP accumulated apicolaterally in 8.8% of MuSCs, and this value increased to 22.9% at T48. Dystrophin accumulation on the basal side was found in 0.6% of MuSCs at T12, 3.6% of cells at T24, 4.1% of cells at T36, and 4.4% of cells at T48, matching previous observations [\u003cspan citationid=\"CR14\" class=\"CitationRef\"\u003e14\u003c/span\u003e] \u003cb\u003e(Figure S2B)\u003c/b\u003e.\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003cp\u003eWe next followed the movements of tdTomato\u003csup\u003e+\u003c/sup\u003e MuSCs on isolated fibers from \u003cem\u003eDmd\u003c/em\u003e\u003csup\u003e\u003cem\u003eEGFP/y\u003c/em\u003e\u003c/sup\u003e;\u003cem\u003ePax7\u003c/em\u003e\u003csup\u003e\u003cem\u003eCreERT2/+\u003c/em\u003e\u003c/sup\u003e;\u003cem\u003eRosa\u003c/em\u003e\u003csup\u003e\u003cem\u003etdT/+\u003c/em\u003e\u003c/sup\u003e mice using time-lapse microscopy beginning at 44 h of culture \u003cb\u003e(\u003c/b\u003eFig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e2\u003c/span\u003eB\u003cb\u003e)\u003c/b\u003e. Images were acquired every 10 min for 6 h. At T44, dystrophin-EGFP often accumulated, as observed in the previous experiment, on the apical, lateral, or basal side of MuSCs. However, we unexpectedly found that dystrophin-EGFP accumulation did not follow the migration of MuSCs. As indicated by the example of Cell 1, MuSCs had moved from the apically accumulated dystrophin-EGFP at T46 to the other side of the fiber. Meanwhile, the cell left a visible footprint of dystrophin-EGFP at its original position at T44, which remained visible at T52 \u003cb\u003e(\u003c/b\u003eFig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e2\u003c/span\u003eB\u003cb\u003e)\u003c/b\u003e. Conversely, dystrophin-EGFP accumulated basally (Cell 2) or laterally of the MuSCs (Cell 3). Over time, we observed that the MuSCs squeezed through this dystrophin accumulation, and we subsequently lost view of the cells. Meanwhile, the dystrophin-EGFP accumulation visible at T44 remained at the same position at T52. As illustrated in the example 4, the MuSC partially moved from the basally accumulated dystrophin-EGFP. These data were confirmed by time-lapse videos, illustrating that MuSCs moved from the area of MuSC-associated dystrophin accumulation \u003cb\u003e(Movie S1)\u003c/b\u003e.\u003c/p\u003e \u003c/div\u003e\n\u003ch3\u003eMuSC-associated dystrophin is of sarcolemmal origin\u003c/h3\u003e\n\u003cp\u003eThese surprising observations prompted us to hypothesize that dystrophin accumulation associated with MuSCs might be of sarcolemmal origin. We previously reported that increased dystrophin-EGFP accumulation at myotendinous junctions resulted from sarcolemmal folding, which we could image in isolated fibers using the fluorescent membrane probe MemBright [\u003cspan citationid=\"CR33\" class=\"CitationRef\"\u003e33\u003c/span\u003e], a finding that we reproduced using isolated myofibers from \u003cem\u003eDmd\u003c/em\u003e\u003csup\u003e\u003cem\u003eEGFP/y\u003c/em\u003e\u003c/sup\u003e;\u003cem\u003ePax7\u003c/em\u003e\u003csup\u003e\u003cem\u003eCreERT2/+\u003c/em\u003e\u003c/sup\u003e;\u003cem\u003eRosa\u003c/em\u003e\u003csup\u003e\u003cem\u003etdT/+\u003c/em\u003e\u003c/sup\u003e mice following 48 h of culture \u003cb\u003e(\u003c/b\u003eFig.\u0026nbsp;\u003cspan refid=\"Fig5\" class=\"InternalRef\"\u003e3\u003c/span\u003eA\u003cb\u003e)\u003c/b\u003e. Upon analyzing the MuSCs of these fibers, we found that MemBright fluorescence colocalized with the high accumulation of dystrophin-EGFP at the MuSC level, and this accumulation was continuous with the MemBright staining of the sarcolemma \u003cb\u003e(\u003c/b\u003eFig.\u0026nbsp;\u003cspan refid=\"Fig5\" class=\"InternalRef\"\u003e3\u003c/span\u003eA\u003cb\u003e)\u003c/b\u003e. These observations demonstrate that dystrophin-EGFP accumulation resulted indeed from condensed/stacked membranes; however, this experiment did not uncover the cellular origin of the dystrophin-EGFP\u003csup\u003e+\u003c/sup\u003e membranes.\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003cp\u003eWe next determined the frequency of dystrophin-EGFP positivity among tdTomato \u003csup\u003e+\u003c/sup\u003e cells. We first confirmed that the spectral emissions of EGFP and tdTomato in fibers freshly isolated from \u003cem\u003eDmd\u003c/em\u003e\u003csup\u003e\u003cem\u003eEGFP/y\u003c/em\u003e\u003c/sup\u003e;\u003cem\u003ePax7\u003c/em\u003e\u003csup\u003e\u003cem\u003eCreERT2/+\u003c/em\u003e\u003c/sup\u003e;\u003cem\u003eRosa\u003c/em\u003e\u003csup\u003e\u003cem\u003etdT/+\u003c/em\u003e\u003c/sup\u003e reporter mice did not overlap, allowing high-quality spectral acquisition \u003cb\u003e(Figure S3A)\u003c/b\u003e. Moreover, the EGFP spectrum was absent in tdTomato\u003csup\u003e+\u003c/sup\u003e MuSCs \u003cb\u003e(Figure S3A)\u003c/b\u003e. This allowed us to differentiate dystrophin-EGFP and MuSC-emitted tdTomato fluorescence. Using FACS, we sorted tdTomato\u003csup\u003e+\u003c/sup\u003e MuSCs from the muscles of adult \u003cem\u003eDmd\u003c/em\u003e\u003csup\u003e\u003cem\u003eEGFP/y\u003c/em\u003e\u003c/sup\u003e;\u003cem\u003ePax7\u003c/em\u003e\u003csup\u003e\u003cem\u003eCreERT2/+\u003c/em\u003e\u003c/sup\u003e;\u003cem\u003eRosa\u003c/em\u003e\u003csup\u003e\u003cem\u003etdT\u003c/em\u003e/+\u003c/sup\u003e mice and \u003cem\u003eDmd\u003c/em\u003e\u003csup\u003e\u003cem\u003eEGFP\u0026minus;mdx/y\u003c/em\u003e\u003c/sup\u003e;\u003cem\u003ePax7\u003c/em\u003e\u003csup\u003e\u003cem\u003eCreERT2/+\u003c/em\u003e\u003c/sup\u003e;\u003cem\u003eRosa\u003c/em\u003e\u003csup\u003e\u003cem\u003etdT/+\u003c/em\u003e\u003c/sup\u003e reporter mice. FACS-sorted GFP\u003csup\u003e+\u003c/sup\u003e MuSCs from \u003cem\u003eTg:Pax7nGFP\u003c/em\u003e mice served as positive controls, and a GFP\u003csup\u003e+\u003c/sup\u003e cell population was easily identified \u003cb\u003e(\u003c/b\u003eFig.\u0026nbsp;\u003cspan refid=\"Fig5\" class=\"InternalRef\"\u003e3\u003c/span\u003eB\u003cb\u003e)\u003c/b\u003e. tdTomato\u003csup\u003e+\u003c/sup\u003e MuSCs from \u003cem\u003ePax7\u003c/em\u003e\u003csup\u003e\u003cem\u003eCreERT2/CreERT2\u003c/em\u003e\u003c/sup\u003e;\u003cem\u003eRosa\u003c/em\u003e\u003csup\u003e\u003cem\u003etdT/tdT\u003c/em\u003e\u003c/sup\u003e mice served as negative controls, and as expected, this population was GFP\u003csup\u003e\u0026minus;\u003c/sup\u003e, confirming the spectral specificity \u003cb\u003e(Figure S3B)\u003c/b\u003e. No GFP\u003csup\u003e+\u003c/sup\u003e population was detected in FACS-sorted MuSCs from \u003cem\u003eDmd\u003c/em\u003e\u003csup\u003e\u003cem\u003eEGFP/y\u003c/em\u003e\u003c/sup\u003e;\u003cem\u003ePax7\u003c/em\u003e\u003csup\u003e\u003cem\u003eCreERT2/+\u003c/em\u003e\u003c/sup\u003e;\u003cem\u003eRosa\u003c/em\u003e\u003csup\u003e\u003cem\u003etdT/+\u003c/em\u003e\u003c/sup\u003e muscles or \u003cem\u003eDmd\u003c/em\u003e\u003csup\u003e\u003cem\u003eEGFP\u0026minus;mdx/y\u003c/em\u003e\u003c/sup\u003e;\u003cem\u003ePax7\u003c/em\u003e\u003csup\u003e\u003cem\u003eCreERT2/+\u003c/em\u003e\u003c/sup\u003e;\u003cem\u003eRosa\u003c/em\u003e\u003csup\u003e\u003cem\u003etdT\u003c/em\u003e/+\u003c/sup\u003e muscles, confirming the absence of dystrophin-EGFP protein in the MuSCs of these mice \u003cb\u003e(Figure S3B)\u003c/b\u003e.\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003cp\u003eTo confirm the biological significance of this observation across species, we cultured isolated myofibers from adult Sprague\u0026ndash;Dawley rats for 48 h and performed co-immunostaining against dystrophin and PAX7. MuSCs had often already undergone cell division, and they appeared in doublets or small clusters. Strikingly, dystrophin highly accumulated around all single MuSCs or in juxtaposition to all MuSCs doublets/clusters. Dystrophin accumulated erratically at the MuSC level, either apically, basally, laterally, or between MuSCs, with no preferential localization \u003cb\u003e(\u003c/b\u003eFigs.\u0026nbsp;\u003cspan refid=\"Fig5\" class=\"InternalRef\"\u003e3\u003c/span\u003eC and \u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003eS2\u003c/span\u003eC\u003cb\u003e)\u003c/b\u003e. Sarcolemmal dystrophin was organized in costameres, which appeared tethered toward the dystrophin accumulation associated with MuSCs \u003cb\u003e(Figure S3D)\u003c/b\u003e. Airyscan microscopy revealed dystrophin\u003csup\u003e+\u003c/sup\u003e membrane stacks in continuation with the sarcolemma \u003cb\u003e(\u003c/b\u003eFigs.\u0026nbsp;\u003cspan refid=\"Fig5\" class=\"InternalRef\"\u003e3\u003c/span\u003eC and \u003cspan refid=\"Fig6\" class=\"InternalRef\"\u003eS3\u003c/span\u003eE\u003cb\u003e)\u003c/b\u003e.\u003c/p\u003e \u003cp\u003e \u003cb\u003ePresence of\u003c/b\u003e \u003cb\u003eDmd\u003c/b\u003e \u003cb\u003etranscripts but absence of dystrophin protein in MuSCs\u003c/b\u003e\u003c/p\u003e \u003cp\u003eWe then assessed the dynamics of \u003cem\u003eDmd\u003c/em\u003e expression and dystrophin protein synthesis during myogenic lineage progression. For this purpose, we generated primary myoblast cultures using MuSCs from 1-week-old wild-type CD-1 mice. Previous research demonstrated that such cultures form mature myotubes in which dystrophin is located at the sarcolemmal level [\u003cspan citationid=\"CR32\" class=\"CitationRef\"\u003e32\u003c/span\u003e]. ddPCR revealed the strong presence of \u003cem\u003ePax7\u003c/em\u003e in freshly derived MuSCs (T0), \u003cem\u003eMyod1\u003c/em\u003e expression in MuSC-derived myoblasts after 3 days of culture, and strong \u003cem\u003eMyh1\u003c/em\u003e expression in myotubes following 13 days of culture, confirming the myogenic line progression of cells in these primary cultures \u003cb\u003e(Figure S4A)\u003c/b\u003e. \u003cem\u003eDmd\u003c/em\u003e transcripts were already detectable in isolated MuSCs at T0 \u003cb\u003e(\u003c/b\u003eFig.\u0026nbsp;\u003cspan refid=\"Fig7\" class=\"InternalRef\"\u003e4\u003c/span\u003eA\u003cb\u003e)\u003c/b\u003e, and \u003cem\u003eDmd\u003c/em\u003e RNA was unexpectedly more abundant in MuSCs than in myoblasts and myotubes, corroborating previous findings [\u003cspan citationid=\"CR14\" class=\"CitationRef\"\u003e14\u003c/span\u003e]. We also performed snRNA-Seq of adult TA muscles, revealing that all \u003cem\u003ePax7\u003c/em\u003e\u003csup\u003e\u003cem\u003e+\u003c/em\u003e\u003c/sup\u003e myonuclei were also \u003cem\u003eDmd\u003c/em\u003e\u003csup\u003e\u003cb\u003e+\u003c/b\u003e\u003c/sup\u003e \u003cb\u003e(Figures S4B and C)\u003c/b\u003e.\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003cp\u003eWe next performed capillary electrophoresis of parallel myoblast and myotube cultures from 7-day-old CD-1 mice as used for the aforementioned PCR experiments. The full-length dystrophin isoform Dp427 was present at low levels in myoblasts and myotubes, contrasting the high dystrophin levels in TA muscles from adult wild-type CD-1 mice. However, dystrophin was not detected in MACS-sorted MuSCs from 2-week-old mice using antibodies against both the rod domain and C-terminal domain of dystrophin \u003cb\u003e(\u003c/b\u003eFigs.\u0026nbsp;\u003cspan refid=\"Fig7\" class=\"InternalRef\"\u003e4\u003c/span\u003eB and \u003cspan refid=\"Fig8\" class=\"InternalRef\"\u003eS4\u003c/span\u003eD\u003cb\u003e)\u003c/b\u003e. Similarly, in wild-type adult Sprague-Dawley rats, capillary electrophoresis revealed that full-length dystrophin was absent in MACS-sorted MuSCs, whereas it was highly abundant in skeletal muscle, which served as a positive control, and absent in muscle from the DMD rat model (harboring a frameshift deletion of \u003cem\u003eDmd\u003c/em\u003e exon 52), which served as negative control \u003cb\u003e(Figure S4E, and F)\u003c/b\u003e. Finally, immunocytochemistry of parallel myoblast and myotube cultures from 7-day-old CD-1 mice revealed that dystrophin protein was first detected in a predominantly scattered perinuclear position in myoblasts and thereafter along the sarcolemma in myotubes \u003cb\u003e(\u003c/b\u003eFig.\u0026nbsp;\u003cspan refid=\"Fig7\" class=\"InternalRef\"\u003e4\u003c/span\u003eC\u003cb\u003e)\u003c/b\u003e.\u003c/p\u003e"},{"header":"Discussion","content":"\u003cp\u003eThe polarized distribution of dystrophin protein in MuSCs is critical for establishing cell polarity, regulating asymmetric versus symmetric cell division, and enabling correct muscle regeneration [\u003cspan citationid=\"CR14\" class=\"CitationRef\"\u003e14\u003c/span\u003e]. In this study, we investigated the dynamics of the establishment of this polarized dystrophin distribution in MuSCs. We used reporter mouse lines to directly image the native dystrophin-EGFP fusion protein, which we previously employed to study the compartmentalization of dystrophin in skeletal muscle at subcellular levels [\u003cspan citationid=\"CR32\" class=\"CitationRef\"\u003e32\u003c/span\u003e]. We found that dystrophin protein was undetectable in quiescent adult MuSCs. In addition, sarcolemmal dystrophin facing MuSCs was virtually absent, confirming published data [\u003cspan citationid=\"CR31\" class=\"CitationRef\"\u003e31\u003c/span\u003e]. This absence of dystrophin in the niche of quiescent MuSCs contrasted the rapid accumulation of the protein in juxtaposition to activated MuSCs following the culture of isolated myofibers, in line with previous observations [\u003cspan citationid=\"CR14\" class=\"CitationRef\"\u003e14\u003c/span\u003e, \u003cspan citationid=\"CR18\" class=\"CitationRef\"\u003e18\u003c/span\u003e].\u003c/p\u003e \u003cp\u003eAs previously demonstrated [\u003cspan citationid=\"CR14\" class=\"CitationRef\"\u003e14\u003c/span\u003e], we found that despite the absence of dystrophin protein, \u003cem\u003eDmd\u003c/em\u003e was highly transcribed according to bulk PCR, confirming previous data [\u003cspan citationid=\"CR14\" class=\"CitationRef\"\u003e14\u003c/span\u003e]. Furthermore, snRNA-Seq revealed that transcripts were present in all MuSCs, and we found no evidence of a subpopulation with disproportionately strong expression. Of note, enzymatic isolation of myofibers requires approximately 2 h, and it can induce the activation of quiescent MuSCs [\u003cspan citationid=\"CR34\" class=\"CitationRef\"\u003e34\u003c/span\u003e]. However, the timeframe of isolation is insufficient for significant transcription of the \u003cem\u003eDmd\u003c/em\u003e gene (2.4\u0026nbsp;million base pairs in length), as this process requires approximately 16 h [\u003cspan citationid=\"CR35\" class=\"CitationRef\"\u003e35\u003c/span\u003e]. Nevertheless, because high levels of \u003cem\u003eDmd\u003c/em\u003e transcripts are already present in quiescent MuSCs, it is possible that rapid translation could provide an explanation for the rapid appearance of dystrophin protein near MuSCs following isolation. However, dystrophin accumulated as membranous stacks in the niche of activated MuSCs and appeared in a continuum with the sarcolemma. This suggests a sarcolemmal origin of MuSC-associated dystrophin. Sarcolemmal dystrophin is organized in nuclear domains, also termed BSDUs, spanning approximately 80 \u0026micro;m in length [\u003cspan citationid=\"CR32\" class=\"CitationRef\"\u003e32\u003c/span\u003e]. It is therefore unlikely that the observed rapid and locally restricted accumulation of dystrophin in the MuSC niche is a myonuclear driven process.\u003c/p\u003e \u003cp\u003eCould already existing dystrophin locally accumulate near activated MuSC? Sarcolemmal dystrophin is organized in costameres [\u003cspan citationid=\"CR36\" class=\"CitationRef\"\u003e36\u003c/span\u003e]. We demonstrated in this study that costamere-organized sarcolemmal dystrophin accumulated in these stacks. Do MuSCs actively gather sarcolemma? It was previously observed that MuSCs move along the length of an isolated fiber in culture [\u003cspan citationid=\"CR37\" class=\"CitationRef\"\u003e37\u003c/span\u003e]. Our time-lapse experiments illustrated that MuSCs moved from their niche but left dystrophin stacks that remained attached to the sarcolemma, providing further evidence that dystrophin represented condensed sarcolemma. Interestingly, myofibers contracted by approximately 23% during enzymatic isolation. In addition, following myofiber isolation, we and others observed that MuSCs retracted their cell membrane extensions and adopted a rounded morphology [\u003cspan citationid=\"CR30\" class=\"CitationRef\"\u003e30\u003c/span\u003e]. We suggest that the sarcolemma becomes floppy once fibers are no longer stretched between tendons, and retracting and mobile MuSCs shuffle these floppy sarcolemmata together.\u003c/p\u003e \u003cp\u003eOur results imply that dystrophin accumulation near MuSCs occurs only when enzymatically isolating and culturing myofibers. In mice, we observed these events in approximately 25% of MuSCs either on the apical, apical\u0026ndash;lateral, and (more rarely) basal sides of MuSCs. These results support previous findings [\u003cspan citationid=\"CR14\" class=\"CitationRef\"\u003e14\u003c/span\u003e]. Curiously, dystrophin accumulated in juxtaposition to all MuSCs in cultured myofibers isolated from rat muscles. The dystrophin accumulation appeared rather erratically at any position around or between MuSCs with no distinct morphological feature.\u003c/p\u003e \u003cp\u003eFinally, we demonstrated that dystrophin protein first emerged in primary myoblasts in a perinuclear distribution before it became localized at the sarcolemmal level in myotubes. This perinuclear position is compatible with the expected endoplasmic and ribosomal translation sites.\u003c/p\u003e \u003cp\u003eWe used reporter mice in which EGFP was directly tagged to the C-terminus of dystrophin, permitting visualization of all dystrophin isoforms [\u003cspan citationid=\"CR25\" class=\"CitationRef\"\u003e25\u003c/span\u003e]. The absence of dystrophin-EGFP in MuSCs suggests the absence of different dystrophin isoforms despite prior evidence of Dp71 in myoblasts [\u003cspan citationid=\"CR13\" class=\"CitationRef\"\u003e13\u003c/span\u003e].\u003c/p\u003e \u003cp\u003eAlthough our findings do not contradict the occurrence of MuSC polarization or the critical role of balancing asymmetric and symmetric divisions in muscle regeneration, they challenge the previously proposed involvement of dystrophin in this process. Our data do not question the accumulating evidence that MuSCs function is compromised in DMD and DMD animal models [\u003cspan citationid=\"CR38\" class=\"CitationRef\"\u003e38\u003c/span\u003e, \u003cspan citationid=\"CR39\" class=\"CitationRef\"\u003e39\u003c/span\u003e, \u003cspan citationid=\"CR40\" class=\"CitationRef\"\u003e40\u003c/span\u003e]. However, our results do not support the direct involvement of dystrophin, as it was absent in MuSCs. Other mechanisms, such as an altered extracellular matrix, altered muscle metabolism, altered signaling pathways, altered vascularization, and precocious MuSCs senescence, must be considered [\u003cspan citationid=\"CR41\" class=\"CitationRef\"\u003e41\u003c/span\u003e, \u003cspan citationid=\"CR42\" class=\"CitationRef\"\u003e42\u003c/span\u003e]. Thus, several pathophysiological processes that are well described in DMD could hamper MuSC-dependent muscle regeneration.\u003c/p\u003e \u003cp\u003eAn alternative perspective is the possibility of post-transcriptional regulation of dystrophin, with miRNAs potentially modulating mRNA stability or translation. Supporting this idea, several studies highlighted the role of specific miRNAs in this process. For instance, miR-31, which is upregulated in \u003cem\u003eDmd\u003c/em\u003e\u003csup\u003e\u003cem\u003emdx/y\u003c/em\u003e\u003c/sup\u003e mice, has been identified as a potential repressor of dystrophin [\u003cspan citationid=\"CR43\" class=\"CitationRef\"\u003e43\u003c/span\u003e]. Additionally, intronic \u003cem\u003eDMD\u003c/em\u003e RNA sequences might act as non-coding RNA (ncRNA) regulators [\u003cspan citationid=\"CR44\" class=\"CitationRef\"\u003e44\u003c/span\u003e], and \u003cem\u003eDMD\u003c/em\u003e mutations could disrupt this function. However, the existence of this regulatory role for \u003cem\u003eDMD\u003c/em\u003e ncRNA in MuSCs remains purely theoretical at this stage.\u003c/p\u003e \u003cp\u003eSatellite cell-opathies are increasingly recognized clinical entities that are best illustrated in patients with \u003cem\u003ePAX7\u003c/em\u003e mutations [\u003cspan citationid=\"CR45\" class=\"CitationRef\"\u003e45\u003c/span\u003e]. We previously described the distinct clinical phenotype of a patient with \u003cem\u003ePAX7\u003c/em\u003e mutation [\u003cspan citationid=\"CR46\" class=\"CitationRef\"\u003e46\u003c/span\u003e], which is entirely different from that of DMD. Nevertheless, MuSCs could represent a useful target for dystrophin-restoring therapies such as gene editing. Correction of the mutated \u003cem\u003eDMD\u003c/em\u003e gene could progressively replace dystrophin-incompetent myonuclei with dystrophin-competent myonuclei during muscle regeneration and thereby overcome the dystrophic phenotype. The function of high \u003cem\u003eDMD\u003c/em\u003e RNA levels in MuSCs remains unclear. Are high \u003cem\u003eDMD\u003c/em\u003e RNA levels required for rapid dystrophin synthesis and sarcolemmal stability upon MuSC fusion into the growing myofiber syncytium? If this is the case, then it would be important to restore the reading frame of mutated \u003cem\u003eDMD\u003c/em\u003e RNA in differentiated muscle and MuSCs, and the effect of current exon skipping therapies on MuSCs should be tested.\u003c/p\u003e"},{"header":"Materials and Methods","content":"\u003cdiv id=\"Sec7\" class=\"Section2\"\u003e \u003ch2\u003eAnimal models\u003c/h2\u003e \u003cp\u003e All animal experiments were conducted according to the National and European legislation and institutional guidelines for the care and use of laboratory animals approved by the French government (Minist\u0026egrave;re de l\u0026rsquo;Enseignement Sup\u0026eacute;rieur et de la Recherche, autorisation APAFiS #44987-2023092714211861 v4 and #34462-2021121716045314 v6). Mice were bred in the Plateforme 2Care animal facility, UFR des Sciences de la Sant\u0026eacute;, Universit\u0026eacute; de Versailles Saint- Quentin-en-Yvelines and maintained under a standard 12-h light/12-h dark photoperiod with free access to food and water. Mice were weaned after 4\u0026ndash;5 postnatal weeks, and 2\u0026ndash;5 individuals were housed per cage. To generate the \u003cem\u003eDmd\u003c/em\u003e\u003csup\u003e\u003cem\u003eEGFP/y\u003c/em\u003e\u003c/sup\u003e;\u003cem\u003ePax7\u003c/em\u003e\u003csup\u003e\u003cem\u003eCreERT2/+\u003c/em\u003e\u003c/sup\u003e;\u003cem\u003eRosa\u003c/em\u003e\u003csup\u003e\u003cem\u003etdT/+\u003c/em\u003e\u003c/sup\u003e mouse line, we crossed homozygous female \u003cem\u003eDmd\u003c/em\u003e\u003csup\u003e\u003cem\u003eEGFP/EGFP\u003c/em\u003e\u003c/sup\u003e mice [\u003cspan citationid=\"CR19\" class=\"CitationRef\"\u003e19\u003c/span\u003e] with heterozygous male \u003cem\u003ePax7\u003c/em\u003e\u003csup\u003e\u003cem\u003eCreERT2/+\u003c/em\u003e\u003c/sup\u003e;\u003cem\u003eR\u003c/em\u003e26\u003csup\u003e\u003cem\u003eLoxP\u0026thinsp;\u0026minus;\u0026thinsp;STOP\u0026minus;LoxP\u0026minus;TdTom\u003c/em\u003e\u003c/sup\u003e (\u003cem\u003ei.e.\u003c/em\u003e, \u003cem\u003eRosa\u003c/em\u003e\u003csup\u003e\u003cem\u003etdT/+\u003c/em\u003e\u003c/sup\u003e) mice [\u003cspan citationid=\"CR20\" class=\"CitationRef\"\u003e20\u003c/span\u003e, \u003cspan citationid=\"CR21\" class=\"CitationRef\"\u003e21\u003c/span\u003e]. Similarly, we generated \u003cem\u003eDmd\u003c/em\u003e\u003csup\u003e\u003cem\u003eEGFP\u0026minus;mdx/y\u003c/em\u003e\u003c/sup\u003e\u003cem\u003ePax7\u003c/em\u003e\u003csup\u003e\u003cem\u003eCreERT2/+\u003c/em\u003e\u003c/sup\u003e;\u003cem\u003eRosa\u003c/em\u003e\u003csup\u003e\u003cem\u003etdT/+\u003c/em\u003e\u003c/sup\u003e reporter mice (carrying the \u003cem\u003emdx\u003c/em\u003e-23 mutation and C-terminal EGFP-tag \u003cem\u003ein cis\u003c/em\u003e in the \u003cem\u003eDmd\u003c/em\u003e locus[\u003cspan citationid=\"CR22\" class=\"CitationRef\"\u003e22\u003c/span\u003e]). \u003cem\u003eTg:Pax7-nGFP\u003c/em\u003e mice [\u003cspan citationid=\"CR23\" class=\"CitationRef\"\u003e23\u003c/span\u003e] were used to express nuclear green fluorescent protein (nGFP) under the control of the \u003cem\u003ePax7\u003c/em\u003e promoter. Activation of the \u003cem\u003etdTomato\u003c/em\u003e transgene in \u003cem\u003ePax7\u003c/em\u003e-expressing MuSCs was induced by either 1 week of tamoxifen-enriched diet (TAM400-IRRADIATED, Envigo, Indianapolis, IN, USA) feeding or daily oral gavage of tamoxifen (Sigma-Aldrich, St. Louis, MO, USA) dissolved in corn oil at 25 mg/ml and administered at a dose of 8 \u0026micro;l/g body weight for 5 consecutive days. Wild-type CD-1\u0026reg; IGS mice and wild-type Sprague-Dawley rats were purchased from Charles River Laboratories (Wilmington, MA, USA). Muscles from Sprague-Dawley rats carrying \u003cem\u003eDmd\u003c/em\u003e exon 52 deletion (\u003cem\u003eR-DMDdel52\u003c/em\u003e, also called DMD rats) were kindly gifted by Valentina Taglietti [\u003cspan citationid=\"CR24\" class=\"CitationRef\"\u003e24\u003c/span\u003e]. This study was performed in juvenile (1\u0026ndash;2 weeks old) and adult animals (2\u0026ndash;4 months old). Mice were euthanized by cervical dislocation, whereas rats were euthanized by intracardiac administration of Euthasol (0.1 mL/kg, Centravet, France), in accordance with approved institutional and ethical guidelines. All animal experimentation was conducted and reported in accordance with the ARRIVE 2.0 guidelines.\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec8\" class=\"Section2\"\u003e \u003ch2\u003eMyofiber isolation, culture, and live imaging\u003c/h2\u003e \u003cp\u003eIsolated muscle fibers were prepared from extensor digitorum longus (EDL) muscle isolates from mice and rats as previously described [\u003cspan citationid=\"CR25\" class=\"CitationRef\"\u003e25\u003c/span\u003e, \u003cspan citationid=\"CR26\" class=\"CitationRef\"\u003e26\u003c/span\u003e]. The isolated live myofibers were either directly imaged in DMEM Fluorobright (Thermo Fisher Scientific, Waltham, MA, USA) supplemented with 10% \u003cspan type=\"SmallCaps\" class=\"SmallCaps\" name=\"Emphasis\"\u003el\u003c/span\u003e-glutamine (Thermo Fisher Scientific), 1% sodium pyruvate (Thermo Fisher Scientific) and 1% penicillin\u0026ndash;streptomycin (Thermo Fisher Scientific) in eight-well glass-bottom dishes (Ibidi, Gr\u0026auml;felfing, Germany) or cultured up to 48 h as previously described [\u003cspan citationid=\"CR25\" class=\"CitationRef\"\u003e25\u003c/span\u003e]. For myofibers fixed prior to isolation, EDL muscle was fixed in 4% PFA for 15\u0026ndash;20 min directly after dissection, followed by two washes with 1\u0026times; PBS. The fibers were carefully separated under a binocular stereomicroscope using fine glass needles with a rounded tip. For live microscopy, the isolated myofibers were mounted following 1 h of incubation at 37\u0026deg;C with MemBright (1:1000 dilution, 550 Lipilight by MemBright, Idylle, Paris, France), SiR-actin (1:1000 dilution, Spirochrome, Stein am Rhein, Switzerland) or Hoechst (1:1000 dilution, Sigma-Aldrich) in isolation medium (DMEM supplemented with 1% penicillin\u0026ndash;streptomycin, 4 mM \u003cspan type=\"SmallCaps\" class=\"SmallCaps\" name=\"Emphasis\"\u003el\u003c/span\u003e-glutamine, and 1% sodium pyruvate).\u003c/p\u003e \u003c/div\u003e\n\u003ch3\u003eMuscle preparation for histology\u003c/h3\u003e\n\u003cp\u003eTibialis anterior (TA) and triceps brachii muscles were dissected following euthanasia (cervical dislocation for mice and anesthetic overload with Euthazol [Virbec, Carros, France] 400 mg/ml for rats). The muscles were mounted on 6% tragacanth gum (Sigma-Aldrich), snap frozen in liquid nitrogen-cooled isopentane, and stored at \u0026minus;\u0026thinsp;80\u0026deg;C. Cryosections were prepared using a NX70 Cryostar cryostat (Thermo Fisher Scientific) for native fluorescence imaging and subsequently used for RNA and protein extraction.\u003c/p\u003e\n\u003ch3\u003eCell culture\u003c/h3\u003e\n\u003cp\u003eCells were grown as described by Pimentel et al. [\u003cspan citationid=\"CR27\" class=\"CitationRef\"\u003e27\u003c/span\u003e]. In summary, hindlimb muscles from 7-day-old wild-type CD-1 mice were dissected and incubated in an enzymatic mixture as previously described [\u003cspan citationid=\"CR27\" class=\"CitationRef\"\u003e27\u003c/span\u003e]. The cell suspension was pre-plated at 37\u0026deg;C for 30 min to remove fibroblasts. The supernatant was then collected and plated on Matrigel-coated dishes (1:100 dilution, Corning, NY, USA) using proliferation medium (IMDM with GlutaMAX [Thermo Fisher Scientific], 0.1% gentamicin [Thermo Fisher Scientific], 20% fetal bovine serum [Thermo Fisher Scientific], and 1% chicken embryo extract [Thermo Fisher Scientific). After 3 days of culture, the medium was removed, and ice-cold Matrigel (1:2 dilution, Corning) was added before adding differentiation medium (IMDM with GlutaMAX, 2% horse serum, and agrin 100 ng/ml). The medium was refreshed every 2 days, and the culture was stopped after 13 days when well-differentiated myotubes had formed. Cells were harvested after 3 and 13 days for further digital droplet PCR (ddPCR), immunocytochemistry, and capillary electrophoresis experiments.\u003c/p\u003e \u003cdiv id=\"Sec11\" class=\"Section2\"\u003e \u003ch2\u003eImmunostaining on cells and myofibers\u003c/h2\u003e \u003cp\u003eFor immunostaining, muscle fiber, myoblast, and myotube cultures were fixed in 4% PFA for 10 min. Immunostaining was performed as previously described [\u003cspan citationid=\"CR25\" class=\"CitationRef\"\u003e25\u003c/span\u003e] using primary antibodies targeting the C-terminal region of dystrophin (1:10 dilution, NCL-DYS2, mouse IgG1, Leica Biosystems, Nussloch, Germany; or 1:2,000 dilution, ab15277, Abcam), β-sarcoglycan (1:50 dilution, Leica), PAX7 (1:10 dilution, DHSB), MYOD (1:50 dilution, Agilent, Santa Clara, CA, USA), and MHC (1:200 dilution, DHSB) via overnight incubation at 4\u0026deg;C, followed by incubation for 1 h with fluorochrome-labeled secondary antibodies (1:400 dilution, Alexa Fluor goat anti-mouse IgG1-488/555/633, goat antimouse IgM-488/555, and goat antirabbit IgG (H\u0026thinsp;+\u0026thinsp;L)-488/555/633) at room temperature. Nuclei were stained using DAPI (1:1000 dilution, Sigma-Aldrich), and the fibers were subsequently mounted on glass slides with Fluoromount-G (Southern Biotech, Birmingham, AL, USA).\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec12\" class=\"Section2\"\u003e \u003ch2\u003eFluorescence imaging\u003c/h2\u003e \u003cp\u003eFluorescent images of live or immune-stained single myofibers and cells were acquired using an epifluorescence microscope or confocal microscope as specified in the figure legends. For epifluorescence imaging, a Zeiss Axio Imager microscope (Carl Zeiss, Oberkochen, Germany) equipped with an Orca camera (Hamamatsu Photonics, Hamamatsu, Japan) and ZEN 3.2 Blue software (Carl Zeiss) was used. For live confocal microscopy, myofibers were maintained at 37\u0026deg;C with 5% CO\u003csub\u003e2\u003c/sub\u003e in the microscope\u0026rsquo;s incubation chamber, and imaging was performed using a Leica SP8 white light laser-scanning confocal microscope with LAS X software (Leica Microsystems). For the bleaching experiments, myofibers were cultured in eight-well glass-bottom dishes and imaged in DMEM Fluorobright (Thermo Fisher Scientific, Waltham, MA, USA) supplemented with 10% L-glutamine (Thermo Fisher Scientific), 1% sodium pyruvate (Thermo Fisher Scientific), and 1% penicillin\u0026ndash;streptomycin (Thermo Fisher Scientific). Images were obtained before and after photobleaching in the 490- and 555-nm channels using a confocal microscope with standardized settings to ensure consistency. tdTomato fluorescence was bleached by increasing the laser power at 555 nm and targeting the region of interest (ROI), i.e., the satellite cell cytoplasm, for 15 iterations, ensuring an accurate and reproducible bleaching event. For time-lapse imaging, myofibers were cultured in eight-well glass-bottom dishes using the aforementioned culture medium. Images were automatically captured every 15 min using a confocal microscope and the \u0026ldquo;xyzt\u0026rdquo; mode of LAS X software. Intravital muscle fluorescence images were acquired using a combined confocal/multiphoton point-scanning microscope (SP8 MP, Leica Microsystems) as previously described [\u003cspan citationid=\"CR25\" class=\"CitationRef\"\u003e25\u003c/span\u003e]. For intravital imaging, the mice were anesthetized with 2.5% isoflurane (150\u0026ndash;250 ml/min, Piramal Critical Care, Bethlehem, PA, USA) in oxygen for induction, after which a longitudinal skin incision was made on the lateral side of the lower hindlimb to expose the muscle. The mice were maintained at 37\u0026deg;C in the microscope\u0026rsquo;s incubation chamber with the exposed muscle securely placed on a coverslip attached to the sample holder to minimize motion artifacts. For confocal imaging of fixed myofibers, a confocal microscope or spinning-disk microscope (Leica Microsystems), both equipped with LAS X software, was used. Post-imaging analysis was conducted using ImageJ (US National Institutes of Health, Bethesda, MD, USA), LAS X, and Imaris software (Oxford Instruments, Oxford, UK).\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec13\" class=\"Section2\"\u003e \u003ch2\u003eQuantification of satellite cells\u003c/h2\u003e \u003cp\u003eMuSCs located near accumulated dystrophin-EGFP protein were counted on isolated myofibers from \u003cem\u003eDmd\u003c/em\u003e\u003csup\u003e\u003cem\u003eEGFP/y\u003c/em\u003e\u003c/sup\u003e;\u003cem\u003ePax7\u003c/em\u003e\u003csup\u003e\u003cem\u003eCreERT2/+\u003c/em\u003e\u003c/sup\u003e;\u003cem\u003eRosa\u003c/em\u003e\u003csup\u003e\u003cem\u003etdT/+\u003c/em\u003e\u003c/sup\u003e reporter mice directly after isolation (T0) or after up to 48 h of culture (T48).\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec14\" class=\"Section2\"\u003e \u003ch2\u003eMeasure of sarcomere length and MuSC size\u003c/h2\u003e \u003cp\u003eTo measure sarcomere length, mechanically and enzymatically isolated myofibers were incubated with SiR-actin (1:1000 dilution) for 1 h and then mounted on slides. Images were acquired at random locations along each fiber with a confocal microscope, with three independent ROIs and measurements taken per fiber using the draw scalebar tool of LAS X. To measure MuSC size, the diameter of tdTomato-positive MuSCs attached to enzymatically isolated myofibers (T0) was measured using the \u0026ldquo;draw scalebar\u0026rdquo; tool of LAS X.\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec15\" class=\"Section2\"\u003e \u003ch2\u003eMuscle dissociation and cell preparation\u003c/h2\u003e \u003cp\u003eAdult and juvenile limb muscles were harvested from mice and rats, minced, and incubated with a mix of 0.5 mg/ml collagenase A (Sigma-Aldrich) and 3.5 mg/ml Dispase II (Roche, Basel, Switzerland) or PBS at 37\u0026deg;C and 60 rpm in a shaking water bath for 2 h. Digestion was stopped with IMDM supplemented with 20% fetal bovine serum and GlutaMAX. The cell suspension was filtered through a 70-\u0026micro;m cell strainer (Thermo Fisher Scientific) and then spun at 1300 rpm for 10 min at 4\u0026deg;C to remove large tissue fragments. Before fluorescence-activated cell sorting (FACS), cells were resuspended in HamF10 medium (Sigma-Aldrich) supplemented with 10% horse serum (Gibco). For magnetic-activated cell sorting (MACS), mouse and rat limb muscles were collected, minced, and incubated according to the manufacturer\u0026rsquo;s protocol using a skeletal muscle dissociation kit for mice and rats (Miltenyi Biotec, Bergisch Gladbach, Germany).\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec16\" class=\"Section2\"\u003e \u003ch2\u003eSatellite cell isolation\u003c/h2\u003e \u003cp\u003ePrior to FACS, cell suspensions were prepared following the protocol described by Gayraud-Morel et al. [\u003cspan citationid=\"CR28\" class=\"CitationRef\"\u003e28\u003c/span\u003e]. The final cell pellet was resuspended in cold HamF10 supplemented with 10% horse serum and filtered through a 40-\u0026micro;m strainer (Thermo Fisher Scientific). MuSCs were sorted using the MoFlo Astrios cell sorter (Beckmann Coulter, Brea, CA, USA) and identified using either GFP or tdTomato staining. Post-isolation, mononuclear cells were collected in HamF10 containing 10% horse serum.\u003c/p\u003e \u003cp\u003eFor MACS, mouse and rat MuSCs were purified \u003cem\u003evia\u003c/em\u003e magnetic separation according to the manufacturer\u0026rsquo;s protocol using a satellite cell isolation kit for mice and rats (Miltenyi Biotec). MuSCs were isolated by depleting Sca1\u003csup\u003e+\u003c/sup\u003e, CD45\u003csup\u003e+\u003c/sup\u003e, CD11b\u003csup\u003e+\u003c/sup\u003e, and CD31\u003csup\u003e+\u003c/sup\u003e cells followed by positive selection with anti-alpha-7 integrin microbeads (Miltenyi Biotec). Briefly, muscles from 4\u0026ndash;8 mice were harvested, cut carefully into small pieces, and then resuspended in skeletal muscle dissociation kit. The muscles were incubated for 1 h at 37\u0026deg;C, ground using the GentleMACS\u0026trade; tissue dissociator (Miltenyi Biotec), and incubated for an additional 30 min. Following two washes, 100 \u0026micro;l of a biotinylated antibody cocktail were added per 1 \u0026times; 10\u003csup\u003e8\u003c/sup\u003e cells, followed by the addition of antibiotin microbeads (200 \u0026micro;l per 1 \u0026times; 10\u003csup\u003e8\u003c/sup\u003e cells, Miltenyi Biotec). The cells were then incubated for 15 min at 4\u0026deg;C. Finally, the cells were placed on an \u0026ldquo;LS Column\u0026rdquo; (Miltenyi Biotec) under the magnetic field of a MACS separator (QuadroMACS\u0026trade;, Miltenyi Biotec). The MuSCs were eluted, whereas the remaining cells were retained in the column. An additional purification step was performed using an \u0026ldquo;MS Column\u0026rdquo; (Miltenyi Biotec) positioned within the magnetic field of a MACS separator (OctoMACS\u0026trade;, Miltenyi Biotec), after which the cells were incubated with of anti-alpha 7 integrin microbeads (50 \u0026micro;l per 1 \u0026times; 10\u003csup\u003e8\u003c/sup\u003e cells). MuSCs were collected following their extraction from the column. Subsequent phenotyping was conducted to ensure their purity.\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec17\" class=\"Section2\"\u003e \u003ch2\u003eRNA analysis by digital droplet PCR\u003c/h2\u003e \u003cp\u003eTotal RNA was extracted using TRIzol reagent (Thermo Fisher Scientific) in accordance with the manufacturer\u0026rsquo;s protocol. cDNA was synthesized in 20-\u0026micro;l reactions containing 1000 ng of total RNA and conducted using the LunaScript\u0026trade; RT SuperMix Kit (New England Biolabs, Ipswich, MA, USA). ddPCR was conducted using 2 \u0026micro;l of cDNA in a 20-\u0026micro;l reaction volume, which included 0.5 \u0026micro;l of the relevant TaqMan probe, 10 \u0026micro;l of ddPCR Supermix for probes (Bio-Rad, Hercules, CA, USA), and 7.5 \u0026micro;l of DNase/RNase-free H\u003csub\u003e2\u003c/sub\u003eO. Each 20-\u0026micro;l reaction mixture was deposited into an eight-channel disposable droplet-generating cartridge. Additionally, 70 \u0026micro;l of droplet-generating oil (Bio-Rad) were loaded into the adjacent oil wells of the cartridge, and the microfluidic chip was placed into the QX200 droplet generator (Bio-Rad). The generated droplets were then transferred to a semi-skirted 96-well PCR plate (Bio-Rad). The plate was heat-sealed using a PX1 PCR plate sealer (Bio-Rad) and amplified on a T100 thermal cycler (Bio-Rad) using the following PCR protocol: initial denaturation at 95\u0026deg;C for 10 min, 40 cycles of 94\u0026deg;C for 30 s and 60\u0026deg;C for 1 min, and final deactivation at 98\u0026deg;C for 10 min. The plate containing the amplified droplets was then analyzed using the QX200 droplet reader and QuantaSoft software (Bio-Rad). Differentiation between target-negative and target-positive droplets was achieved by setting a manual fluorescence amplitude threshold for each TaqMan assay. Glyceraldehyde-3-phosphate dehydrogenase (GAPDH) served as the internal control for normalizing transcript concentrations, with the results reported as target/GAPDH ratios. The primers and probes (Integrated DNA Technologies, Coralville, IA, USA) are provided in Table\u0026nbsp;\u003cspan refid=\"Tab1\" class=\"InternalRef\"\u003e1\u003c/span\u003e.\u003c/p\u003e \u003cp\u003e \u003cdiv class=\"gridtable\"\u003e\u003ctable float=\"Yes\" id=\"Tab1\" border=\"1\"\u003e \u003ccaption language=\"En\"\u003e \u003cdiv class=\"CaptionNumber\"\u003eTable 1\u003c/div\u003e \u003cdiv class=\"CaptionContent\"\u003e \u003cp\u003eList of primers used for ddPCR\u003c/p\u003e \u003c/div\u003e \u003c/caption\u003e \u003ccolgroup cols=\"2\"\u003e \u003cdiv align=\"left\" class=\"colspec\" colname=\"c1\" colnum=\"1\"\u003e\u003c/div\u003e \u003cdiv align=\"left\" class=\"colspec\" colname=\"c2\" colnum=\"2\"\u003e\u003c/div\u003e \u003cthead\u003e \u003ctr\u003e \u003cth align=\"left\" colname=\"c1\"\u003e \u003cp\u003e\u003cem\u003eDmd\u003c/em\u003e\u003c/p\u003e \u003cp\u003e\u003cem\u003e(Mm.PT.58.42993407)\u003c/em\u003e\u003c/p\u003e \u003c/th\u003e \u003cth align=\"left\" colname=\"c2\"\u003e \u003cp\u003ePrimer 1: CTT TAC ACA GAG AAA TGA TGC CA\u003c/p\u003e \u003cp\u003ePrimer 2: GTG TCT CAA CTG GCT TCT CAA\u003c/p\u003e \u003cp\u003eProbe: /56-FAM/TT ATG ATA C/ZEN/G GGA CGA ACA GGG AGG A/3IABkFQ/\u003c/p\u003e \u003c/th\u003e \u003c/tr\u003e \u003c/thead\u003e \u003ctbody\u003e \u003ctr\u003e \u003ctd align=\"left\" colname=\"c1\"\u003e \u003cp\u003e\u003cb\u003ePax7\u003c/b\u003e\u003c/p\u003e \u003cp\u003e\u003cem\u003e(Mm.PT.58.12398641)\u003c/em\u003e\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c2\"\u003e \u003cp\u003ePrimer 1: GAA GAA GTC CCA GCA CAG C\u003c/p\u003e \u003cp\u003ePrimer 2: GCT ACC AGT ACA GCC AGT ATG\u003c/p\u003e \u003cp\u003eProbe: /56-FAM/CC AAA AAC G/ZEN/T GAG CCT GTC CAC AC/3IABkFQ/\u003c/p\u003e \u003c/td\u003e \u003c/tr\u003e \u003ctr\u003e \u003ctd align=\"left\" colname=\"c1\"\u003e \u003cp\u003e\u003cb\u003eMyod1\u003c/b\u003e\u003c/p\u003e \u003cp\u003e\u003cem\u003e(Mm.PT.58.8193525)\u003c/em\u003e\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c2\"\u003e \u003cp\u003ePrimer 1: GAC ACA GCC GCA CTC TT\u003c/p\u003e \u003cp\u003ePrimer 2: GCT CTG ATG GCA TGA TGG AT\u003c/p\u003e \u003cp\u003eProbe: /56-FAM/AC GAC ACC G/ZEN/C CTA CTA CAG TGA GG/3IABkFQ/\u003c/p\u003e \u003c/td\u003e \u003c/tr\u003e \u003ctr\u003e \u003ctd align=\"left\" colname=\"c1\"\u003e \u003cp\u003e\u003cb\u003eMyh1\u003c/b\u003e \u003cem\u003e(Mm.PT.58.8856176)\u003c/em\u003e\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c2\"\u003e \u003cp\u003ePrimer 1: CTG GAT CTT GCG GAA TTT GG\u003c/p\u003e \u003cp\u003ePrimer 2: GGA CAA ACT GCA ATC AAA GGT C\u003c/p\u003e \u003cp\u003eProbe: /56-FAM/AA GCT GAG G/ZEN/A AGC GGA GGA ACA AT/3IABkFQ/\u003c/p\u003e \u003c/td\u003e \u003c/tr\u003e \u003ctr\u003e \u003ctd align=\"left\" colname=\"c1\"\u003e \u003cp\u003e\u003cb\u003eGapdh\u003c/b\u003e\u003c/p\u003e \u003cp\u003e\u003cem\u003e(Mm.PT.39a.1)\u003c/em\u003e\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c2\"\u003e \u003cp\u003ePrimer 1: GTG GAG TCA TAC TGG AAC ATG TAG\u003c/p\u003e \u003cp\u003ePrimer 2: AAT GGT GAA GGT CGG TGT G\u003c/p\u003e \u003cp\u003eProbe: /56-FAM/TG CAA ATG G/ZEN/C AGC CCT GGT G/3IABkFQ/\u003c/p\u003e \u003c/td\u003e \u003c/tr\u003e \u003c/tbody\u003e \u003c/colgroup\u003e \u003c/table\u003e\u003c/div\u003e \u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec18\" class=\"Section2\"\u003e \u003ch2\u003eSingle-nuclei RNA sequencing\u003c/h2\u003e \u003cp\u003eNuclei were isolated using the Chromium protocol kit (CG000505, Rev A, 10X Genomics, Pleasanton, CA, USA) from the TA muscles of \u003cem\u003eDmd\u003c/em\u003e\u003csup\u003e\u003cem\u003eEGFP/wt\u003c/em\u003e\u003c/sup\u003e heterozygous mice using a previously reported protocol with modifications [\u003cspan citationid=\"CR29\" class=\"CitationRef\"\u003e29\u003c/span\u003e]. A filtration step was added to the protocol using 70-\u0026micro;m filters immediately after tissue lysis.\u003c/p\u003e \u003cp\u003eSingle-nuclei gel bead-in-emulsions (GEMs) were produced using a Chromium Controller (10X Genomics). Sequencing libraries were created using Chromium Single Cell 3\u0026prime; Reagent Kit v3.1 (10x Genomics). Nuclei were hybridized with probe barcodes and then partitioned into nanoliter-scale GEMs using a microfluidic chip in the iX Chromium (10X Genomics) to target 20,000 nuclei. The sequencing data were processed into transcript count tables using Cell Ranger Single Cell Software Suite 1.3.1 (10x Genomics). Raw base call files from the Nextseq 500 system (Illumina, San Diego, CA, USA) were demultiplexed using the Cell Ranger mkfastq pipeline into library-specific FASTQ files. The FASTQ files for each library were then processed independently with the Cell Ranger pipeline. Once aligned, barcodes associated with these reads, namely cell identifiers and Unique Molecular Identifiers, were subjected to filtering and correction. Reads associated with retained barcodes were quantified and used to build a transcript count table. The resulting data for each sample were then aggregated using the Cell Ranger aggr pipeline, which performed a between-sample normalization step and concatenated the two transcript count tables.\u003c/p\u003e \u003cp\u003eThe subsequent visualizations, clustering, and differential expression tests were performed in R (v 3.4.3, The R Foundation for Statistical Computing, Vienna, Austria) using the standard Seurat pipeline (v5). Quality control on aligned and counted reads was performed to retain nuclei with \u0026gt;\u0026thinsp;200 and \u0026lt;\u0026thinsp;2500 nFeature RNA and \u0026lt;\u0026thinsp;5% mitochondrial genes. In total, 9946 nuclei were processed for this analysis. We detected the expression of approximately 1500 genes per nucleus in each of these muscles. Normalization was performed using the LogNormalize algorithm, and determination of variable features using the vst method (2000 features). We performed linear dimensional reduction and constructed a K-nearest neighbor graph using 15 principal components. Nuclei were then clustered using the Louvain algorithm with a resolution of 0.5, and satellite cells were identified according to PAX7 expression.\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec19\" class=\"Section2\"\u003e \u003ch2\u003eProtein extraction and quantification\u003c/h2\u003e \u003cp\u003eTotal protein was extracted from the cells and muscle sections of TA muscles using RIPA buffer (25 mM Tris-HCl, pH 7.6, 150 mM NaCl, 1% NP-40, 1% sodium deoxycholate) with 5% SDS and 1\u0026times; protease inhibitors (Thermo Fisher Scientific). Cell lysates were incubated on ice for 30 min and centrifuged at 13,000 rpm for 10 min. The supernatant was collected and stored at \u0026minus;\u0026thinsp;80\u0026deg;C until capillary electrophoresis. Tissues were grounded in tubes with beads using a homogenizer grinder (Precellys, Bertin Technologies, Montigny-le-Bretonneux, France) at 10,000 rpm for two cycles of 20 s with a 10-s pause between cycles. The lysates were then centrifuged, heated for 3 min at 100\u0026deg;C, and centrifuged again at 13,000 rpm for 10 min at 4\u0026deg;C. The supernatant was collected and stored as previously mentioned. The total protein concentration was determined using the BCA protein assay kit (Thermo Fisher Scientific).\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec20\" class=\"Section2\"\u003e \u003ch2\u003eCapillary electrophoresis\u003c/h2\u003e \u003cp\u003eCapillary electrophoresis was conducted on a Jess system (ProteinSimple, San Jose, CA, USA) using a 66\u0026ndash;440 kDa separation module and anti-rabbit signal detection for dystrophin. Proteins extracted from lysed cells or muscle tissue collected from 14-day-old Swiss CD-1 mice, adult wild-type rats, and DMD rats were diluted to a concentration of 0.5 \u0026micro;g/\u0026micro;l in 10-fold diluted sample buffer (ProteinSimple). The samples were then mixed with Fluorescent Master Mix (ProteinSimple) and heated at 95\u0026deg;C for 5 min. Samples, blocking reagent, primary antibodies, HRP-conjugated secondary antibodies, and chemiluminescent substrate (ProteinSimple) were placed in a plate from the capillary Western blot kit (ProteinSimple). The default settings included stacking and separation at 475 V for 30 min, blocking for 5 min, primary and secondary antibody incubation for 30 min, and chemiluminescence detection for 15 min (exposure for 1, 2, 4, 8 16, 32, 64, 128, and 512 s). Calibration curves required chemiluminescence peaks exceeding 150,000 units and R2 greater than 0.95.\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec21\" class=\"Section2\"\u003e \u003ch2\u003eStatistical analysis\u003c/h2\u003e \u003cp\u003eThe data were analyzed using GraphPad Prism 7 software (GraphPad, Boston, MA, USA). The number of replicates is specific in each figure legend as necessary. Student\u0026rsquo;s \u003cem\u003et\u003c/em\u003e-test was used for comparisons between two groups, whereas one- and two- way ANOVA followed by multiple comparison tests was employed to compare three or more groups. Significance was denoted by P\u0026thinsp;\u0026lt;\u0026thinsp;0.05.\u003c/p\u003e \u003c/div\u003e"},{"header":"Declarations","content":"\u003cp\u003e\u003cstrong\u003eEthics, Consent to Participate, and Consent to Publish declarations\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eAll animal experiments were conducted according to the National and European legislation and institutional guidelines for the care and use of laboratory animals approved by the French government (Ministère de l’Enseignement Supérieur et de la Recherche, autorisation APAFiS #44987-2023092714211861 v4 approved on February 5, 2024 “Evaluation of gene therapy approaches and antisense strategies for genetic diseases” and #34462-2021121716045314 v6\u0026nbsp;Approved on March 15, 2022 Study of dystrophin expression in the muscle stem cell niche). No human participants were involved in this study; therefore, consent to participate and consent to publish are not applicable. The authors declare that they have not use AI-generated work in this manuscript. All authors read the final version of the manuscript and gave their permission for publication.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eData availability statement\u0026nbsp;\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eRNA sequencing data have been deposited in the Gene Expression Omnibus (GEO) under accession number GSE309529: https://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE309529.\u003c/p\u003e\n\u003cp\u003eAll data generated or analyzed during this study are included in the manuscript and supporting files; source data files have been provided for all figures.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eAcknowledgements\u0026nbsp;\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eWe thank Valentina Taglietti and Frédéric Relaix from Institut Mondor de Recherche Biomédicale (IMRB, Creteil) for providing the \u003cem\u003ePax7\u003csup\u003eCreERT2\u003c/sup\u003e;Rosa\u003csup\u003etdT\u0026nbsp;\u003c/sup\u003e\u003c/em\u003emouse line and the DMD rats muscles. We thank \u0026nbsp;Shahragim Tajbakhsh for his valuable scientific advice and constructive feedback.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eFunding\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eThis work was supported by the Association Française contre les Myopathies (AFM, #23802 and #29169 France), and the Agence Nationale de la Recherche (ANR-23-CE13-0032-03 “DYSCO”, France). Funders were not involved in the design, analysis and reporting of the study.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eAuthors’ contributions\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eHA, PM, and SF designed the study;\u0026nbsp;\u003c/p\u003e\n\u003cp\u003eMM, AM, LR, GM, AG, AJM, BE, LS, AS, TM, VM, SF and HA performed the research;\u003c/p\u003e\n\u003cp\u003eMVP and MS, created the \u003cem\u003eDmd\u003csup\u003eEGFP\u003c/sup\u003e\u003c/em\u003e and \u003cem\u003eDmd\u003csup\u003eEGFP-mdx\u003c/sup\u003e\u003c/em\u003ereporter mouse lines;\u0026nbsp;\u003c/p\u003e\n\u003cp\u003eMM, AM, GM, AG, BE, LS, AS, HA, and LG analyzed data;\u003c/p\u003e\n\u003cp\u003eMM, AM and HA wrote the manuscript.\u003c/p\u003e\n\u003cp\u003eMM–Meriem Matouk\u003c/p\u003e\n\u003cp\u003eAM–Adrien Morin\u003c/p\u003e\n\u003cp\u003eLR–Lucie Royer\u003c/p\u003e\n\u003cp\u003eGM–Gaspard Macaux\u003c/p\u003e\n\u003cp\u003eAG–Aris Gaci\u0026nbsp;\u003c/p\u003e\n\u003cp\u003eAJM–Aude Jobart-Malfait\u003c/p\u003e\n\u003cp\u003eBE–Brendan Evano\u003c/p\u003e\n\u003cp\u003eLS–Liza Sarde\u0026nbsp;\u003c/p\u003e\n\u003cp\u003eAS–Amalia Stantzou\u003c/p\u003e\n\u003cp\u003eMVP–Mina V. Petkova\u003c/p\u003e\n\u003cp\u003eTM–Tudor Manoliu\u003c/p\u003e\n\u003cp\u003eMS–Markus Schuelke\u003c/p\u003e\n\u003cp\u003eLG–Luis Garcia\u003c/p\u003e\n\u003cp\u003ePM–Pascal Maire\u003c/p\u003e\n\u003cp\u003eVM–Vincent Mirouse\u0026nbsp;\u003c/p\u003e\n\u003cp\u003eSF–Sestina Falcone\u003c/p\u003e\n\u003cp\u003eHA–Helge Amthor\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eCompeting interests\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eThe authors do not state any financial nor nonfinancial competing interest.\u0026nbsp;\u003c/p\u003e"},{"header":"References","content":"\u003col\u003e\n \u003cli\u003e\u003cspan\u003eDavies KE, Nowak KJ. Molecular mechanisms of muscular dystrophies: old and new players. Nat Rev Mol Cell Biol. 2006;7(10):762\u0026ndash;73. \u003cspan class=\"ExternalRef\"\u003e\u003cspan class=\"RefSource\"\u003e10.1038/nrm2024\u003c/span\u003e\u003c/span\u003e.\u003c/span\u003e\u003c/li\u003e\n \u003cli\u003e\u003cspan\u003eRahimov F, Kunkel LM. 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Nat Commun. 2019;10(1):5776. \u003cspan class=\"ExternalRef\"\u003e\u003cspan class=\"RefSource\"\u003e10.1038/s41467-019-13650-z\u003c/span\u003e\u003c/span\u003e.\u003c/span\u003e\u003c/li\u003e\n\u003c/ol\u003e"}],"fulltextSource":"","fullText":"","funders":[],"hasAdminPriorityOnWorkflow":false,"hasManuscriptDocX":true,"hasOptedInToPreprint":true,"hasPassedJournalQc":"","hasAnyPriority":false,"hideJournal":false,"highlight":"","institution":"","isAcceptedByJournal":false,"isAuthorSuppliedPdf":false,"isDeskRejected":"","isHiddenFromSearch":false,"isInQc":false,"isInWorkflow":false,"isPdf":false,"isPdfUpToDate":true,"isWithdrawnOrRetracted":false,"journal":{"display":true,"email":"[email protected]","identity":"stem-cell-research-and-therapy","isNatureJournal":false,"hasQc":true,"allowDirectSubmit":false,"externalIdentity":"scrt","sideBox":"Learn more about [Stem Cell Research \u0026 Therapy](http://stemcellres.biomedcentral.com)","snPcode":"","submissionUrl":"https://www.editorialmanager.com/scrt/default.aspx","title":"Stem Cell Research \u0026 Therapy","twitterHandle":"@BioMedCentral","acdcEnabled":true,"dfaEnabled":true,"editorialSystem":"em","reportingPortfolio":"BMC/SO AJ","inReviewEnabled":true,"inReviewRevisionsEnabled":true},"keywords":"Dystrophin, MuSCs, Pax7, sarcolemma","lastPublishedDoi":"10.21203/rs.3.rs-7997649/v1","lastPublishedDoiUrl":"https://doi.org/10.21203/rs.3.rs-7997649/v1","license":{"name":"CC BY 4.0","url":"https://creativecommons.org/licenses/by/4.0/"},"manuscriptAbstract":"\u003cp\u003eThe polarized distribution of dystrophin in muscle stem cells (MuSCs) has been found to regulate asymmetric cell division and maintain the balance between stem cell self-renewal and myogenic commitment. In Duchenne muscular dystrophy, which is associated with dystrophin deficiency, MuSC dysfunction is believed to contribute to fatal muscle wasting. This study investigated the dynamics of the polarized dystrophin distribution during MuSC activation. To this end, we used live fluorescence imaging to visualize EGFP-labeled dystrophin in muscle fibers and MuSCs in the \u003cem\u003eDmd\u003c/em\u003e\u003csup\u003e\u003cem\u003eEGFP\u003c/em\u003e\u003c/sup\u003e reporter mouse model. We also investigated this phenomenon in rats to assess the existence of interspecies consistency. Dystrophin was nearly absent on the apical side of quiescent MuSCs, whereas it rapidly accumulated near a subset of activated mouse MuSCs and 100% of rat MuSCs following enzymatic myofiber isolation and culture. Surprisingly, dystrophin that accumulated near MuSCs accumulated in membranes, and it was always found in continuity with the sarcolemma. Live imaging revealed that MuSCs could move from the condensed dystrophin, which remained attached to the myofiber. Additionally, we detected no dystrophin protein in MuSCs using different techniques, including immunocytochemistry capillary electrophoresis and fluorescence-activated cell sorting, whereas it was present in primary myoblasts and myotubes. Our findings indicate that dystrophin attached to activated MuSCs represents condensed sarcolemmal membranes. Our data suggest that dystrophin protein in MuSCs is not required for normal muscle regeneration.\u003c/p\u003e","manuscriptTitle":"No evidence of the presence of dystrophin protein in rodent muscle stem cells","msid":"","msnumber":"","nonDraftVersions":[{"code":1,"date":"2026-01-27 00:04:22","doi":"10.21203/rs.3.rs-7997649/v1","editorialEvents":[{"type":"communityComments","content":0},{"type":"decision","content":"Revision requested","date":"2026-02-11T22:28:23+00:00","index":"","fulltext":""},{"type":"editorInvitedReview","content":"","date":"2026-02-07T14:30:28+00:00","index":"hide","fulltext":""},{"type":"editorInvitedReview","content":"","date":"2026-02-01T22:04:57+00:00","index":"hide","fulltext":""},{"type":"reviewerAgreed","content":"36948636274092886790476695298062167995","date":"2026-01-23T18:03:35+00:00","index":"hide","fulltext":""},{"type":"reviewerAgreed","content":"196228190092916923436664176383012043717","date":"2026-01-21T19:39:18+00:00","index":"hide","fulltext":""},{"type":"reviewersInvited","content":"","date":"2026-01-21T19:28:24+00:00","index":"","fulltext":""},{"type":"editorAssigned","content":"","date":"2026-01-13T07:51:05+00:00","index":"","fulltext":""},{"type":"checksComplete","content":"","date":"2025-12-22T04:32:26+00:00","index":"","fulltext":""},{"type":"submitted","content":"Stem Cell Research \u0026 Therapy","date":"2025-12-19T18:12:33+00:00","index":"","fulltext":""}],"status":"published","journal":{"display":true,"email":"[email protected]","identity":"stem-cell-research-and-therapy","isNatureJournal":false,"hasQc":true,"allowDirectSubmit":false,"externalIdentity":"scrt","sideBox":"Learn more about [Stem Cell Research \u0026 Therapy](http://stemcellres.biomedcentral.com)","snPcode":"","submissionUrl":"https://www.editorialmanager.com/scrt/default.aspx","title":"Stem Cell Research \u0026 Therapy","twitterHandle":"@BioMedCentral","acdcEnabled":true,"dfaEnabled":true,"editorialSystem":"em","reportingPortfolio":"BMC/SO AJ","inReviewEnabled":true,"inReviewRevisionsEnabled":true}}],"origin":"","ownerIdentity":"4e3d9371-770b-49c7-80c9-61d0770e6b54","owner":[],"postedDate":"January 27th, 2026","published":true,"recentEditorialEvents":[],"rejectedJournal":[],"revision":"","amendment":"","status":"under-review","subjectAreas":[],"tags":[],"updatedAt":"2026-05-05T20:08:18+00:00","versionOfRecord":[],"versionCreatedAt":"2026-01-27 00:04:22","video":"","vorDoi":"","vorDoiUrl":"","workflowStages":[]},"version":"v1","identity":"rs-7997649","journalConfig":"researchsquare"},"__N_SSP":true},"page":"/article/[identity]/[[...version]]","query":{"redirect":"/article/rs-7997649","identity":"rs-7997649","version":["v1"]},"buildId":"XKTyCvWXoU3ODBz1xrDgd","isFallback":false,"isExperimentalCompile":false,"dynamicIds":[84888],"gssp":true,"scriptLoader":[]}