Impaired stem cell migration and divisions in Duchenne muscular dystrophy revealed by live imaging

Domain: neuroscience
Published: 2026-01-28
Editorial outcome: minor_revision

Abstract
Dysregulation of stem cell properties is a hallmark of many pathologies, but the dynamic behaviour of stem cells in their microenvironment during disease progression remains poorly understood. Using the mdx mouse model of Duchenne Muscular Dystrophy, we developed innovative live imaging of muscle stem cells (MuSCs) in vivo, and ex vivo on isolated myofibres. We show that mdx MuSCs have impaired migration and precocious differentiation through unbalanced symmetric divisions, driven by p38 and PI3K signalling pathways, in contrast to the p38-only dependence of healthy MuSCs. Cross-grafting shows that MuSC fate decisions are governed by fibre-independent cues, whereas their migration behaviour is determined by the myofibre niche. This study provides the first dynamic analysis of dystrophic MuSC properties in vivo, reconciling conflicting reports on their function. Our findings establish DMD as a MuSC disease with niche dysfunctions, offering strategies to restore stem cell functions for improved muscle regeneration.

Introduction
Duchenne Muscular Dystrophy (DMD) is a severe X-linked genetic disease characterised by muscle wasting, physical incapacitation, and death around the age of 20-30. Affecting ~1/5000 male births, DMD arises from mutations in the DMD gene encoding Dystrophin, a structural protein essential for myofibre integrity. Dystrophin deficiency leads to chronic muscle degeneration, inflammation, fibrosis, and impaired regeneration1. While ex vivo models suggest that muscle stem cell (MuSC) dysfunction contributes to DMD pathology2, in vivo dynamics of MuSCs and early myogenic cell perturbations remain unexplored.

MuSCs reside in a niche between the myofibre and basement membrane, maintaining quiescence during homeostasis. Upon activation (e.g. muscle injury or isolation), MuSCs proliferate and differentiate to generate or repair muscle fibres, with a subset self-renewing. Myogenic progression involves temporal expression of transcription factors PAX7 (stem), MYF5 and MYOD (commitment), and MYOGENIN (MYOG; differentiation). Symmetric and asymmetric cell divisions (SCD/ACD) regulate this process3. Impaired ACD has been implicated in mdx mice (DMD mouse model), with MuSC hyperplasia and deficient progenitor generation4. Although SCDs are predominant in regenerating muscles5, their role in normal and mdx models remains unexplored. Additionally, conflicting studies reported increased6–8 or decreased4,9 differentiation of dystrophic myogenic cells, leaving the balance of proliferation and differentiation through ACDs and SCDs in mdx MuSCs unresolved.

MuSC differentiation integrates pathways such as p38 mitogen-activated protein kinases (MAPK)10 and PI3K signalling11. p38α/β inhibitors (SB203580) or Mapk14 deletion (p38α-encoding gene) reduced inflammation and prevented myofibre death in mdx mice12, however, the role of p38 in mdx MuSC differentiation remains to be explored. Furthermore, increased Akt activation was reported in mdx mouse muscles and in DMD patients13,14, yet elevated PI3K inhibitor PTEN was reported in golden retriever muscular dystrophy dogs and mdx mouse myoblasts15,16. Targeting PI3K/Akt signalling may mitigate DMD progression17, but its role in mdx MuSC differentiation is poorly studied.

Cell-based therapeutic strategies have been explored for muscle wasting diseases including DMD, however, clinical trials failed, in part due to limited cell migration (reviewed in refs. 18,19). MuSCs are immobile during homeostasis in vivo20, and they migrate along damaged fibres during regeneration following Erk signalling activation and interaction with stromal cells21–23. While in vitro studies suggest reduced chemotaxis and impaired migration6, in vivo migration and positioning of dystrophic MuSCs remain unstudied24.

MuSC properties arise from complex interactions between intrinsic and extrinsic cues20, as evidenced by transcriptome and epigenome reprogramming after transplantation to different anatomical or age-specific environments25. For dystrophy therapies, tissue dispersion of engrafted myogenic cells has been enhanced by boosting intrinsic migration capacity and modulating the environment18. Understanding these cues is essential for developing effective therapeutic strategies for healthy and dystrophic MuSCs.

We developed novel quantitative pipelines to track MuSC migration, proliferation and differentiation in vivo and ex vivo on their myofibre niche, in healthy and mdx contexts. mdx MuSCs showed impaired migration kinetics, increased symmetric divisions towards terminal differentiation, and reduced symmetric stem cell renewal. Transplant experiments revealed that fate decisions are primarily myofibre niche-independent while migration is largely influenced by myofibre-derived cues, indicating that these processes are uncoupled in mdx mice. Further, we identified differential roles for p38 MAPK and PI3K signalling in normal and dystrophic differentiation.

Results
Precocious differentiation and impaired migration of dystrophic MuSCs revealed by intravital imaging
Intravital imaging was used to investigate myogenic cell properties during muscle homeostasis and regeneration in vivo in wild-type (WT) and mdx mice. To do so, MuSCs and their descendants were genetically labelled with a membrane-GFP (mGFP) using an inducible Pax7CreERT2 26 and a R26mTmG 27 reporter in adult Dmd+/Y (WT) and Dmdmdx-βGeo/Y 28 (mdx) mice (Fig. 1a). The hindlimb Flexor digitorum brevis (FDB) muscle was imaged intravitally by two-photon microscopy due to its accessibility (Fig. 1b and Figure S1). In uninjured WT, MuSCs displayed long projections but remained stationary (Fig. S2a, b, Movie S1 and S2) as reported29,30. In uninjured mdx muscles, two distinct myogenic populations were observed: a quiescent non-mobile group resembling WT MuSCs, and a distinct mobile and dividing population in fibre-depleted regions, indicative of ongoing spontaneous regeneration (Fig. S2c, Movie S1 and S2). These mobile MuSCs represented ~24.1% of total GFP+ MuSCs (Fig. S2d). We also noted GFP+ fibres in mdx muscles (~15.9% of total), a sign of regeneration occurring between the tamoxifen treatment and analysis (7 days) (Fig. S2e). Together, these findings confirm that the FDB muscle recapitulates DMD-like chronic regeneration, exhibiting homeostatic quiescence in WT and ongoing turnover in mdx.

To circumvent the non-synchronous nature of myofibre damage and repair in mdx mice, we induced acute muscle injury in the FDB muscle by injecting cardiotoxin, a snake venom, to trigger MuSC activation. Myogenic cells were visualised at 3 days post-injury (dpi) when they undergo active divisions and migration22,23. WT myogenic cells were mononucleated, showed an elongated/mesenchymal-like morphology with few protrusions, actively proliferated and migrated, and had a uni- or bi-polar morphology (Fig. 1c, Movie S3 and S4), as reported29–31. In contrast, mdx mice showed the presence of round and immobile mononucleated cells and multinucleated mGFP+ myotubes (Fig. 1d, e, Movie S3 and S4), suggesting faster differentiation and fusion kinetics compared with the WT. Next, we quantified migration speed and directionality (reflected by straightness and turning angle) (Fig. 1f–i, Fig. S2f, Movies S5 and S6). WT myogenic cells migrated at 28.5 μm/h and exhibited a straight trajectory (mean straightness = 0.75) (Fig. 1g, h), indicating active migration along the longitudinal axis of damaged muscle fibres as reported previously22,23. In contrast, mdx MuSCs showed overall reduced speed (26.2 μm/h, p=6.1e-05), and reduced directionality (mean straightness = 0.70, p=1.13e-08 and turning angle, mean mdx=47.7° vs mean WT=38.1°, p=7.8e-16) (Fig. 1g–i), and the presence of round and immobile cells (Fig. 1d, Movie S4).

To analyse cell morphology parameters, MuSCs were segmented in 4D using the SAM plugin32 (Fig. S2g), that allowed analysis of morphological parameters such as mean area and circularity. Circularity ranged from 0 (elongated/irregular) to 1 (round). mdx MuSCs displayed smaller mean area and a bimodal circularity distribution (Figure S2h, i), with more cells being rounded. As validation, cells visually identified as small and round (Fig. 1d and Movie S4) mapped precisely within the high-circularity, low-area population (Fig. S2h, i). We next analysed morphology/mobility relationships (area vs. speed, circularity vs. speed, Fig. S2j, k). No global linear correlation emerged, likely due to trajectories alternating between movement and pause phases. Nevertheless, round cells were consistently the slowest, indicating a correlation between extreme morphology and low mobility. Of note, MuSCs were in sub-laminal position in WT and mdx mice, in uninjured and injured muscles (Fig. S2l–n).

After mitosis and completion of cytokinesis, WT cells separated rapidly from each other and migrated in opposing directions (>95%, 30–60 min after mitosis), whereas a fraction mdx cells migrated in the same direction shortly after mitosis (~20%, 30–60 min after mitosis) (Fig. 1j–l).

To investigate further the differentiation dynamics of MuSCs, we used a MyogntdTom33 knock-in reporter for differentiation together with Pax7CreERT2 and R26YFP34 alleles for labelling MuSCs and their progeny (Fig. 1m). Cytometry analyses at 3 dpi showed a higher differentiation index of mdx MuSCs compared with WT (9.6% vs 2.7% respectively, p=1.0e-03, Fig. 1n, o), but no difference in absolute numbers of mononucleated myogenic cells (Fig. 1p).

Altogether, our observations point to precocious differentiation and impaired migration kinetics of mdx MuSCs in vivo during muscle regeneration. To further explore the cellular and molecular mechanisms underlying these defects, we developed an ex vivo model designed to: i) recapitulate in vivo phenotypes, ii) enable dynamic tracking of migration and fate decisions, and iii) facilitate perturbation studies.

Impaired symmetric divisions, differentiation and migration kinetics of dystrophic MuSCs in their myofibre niche
Myogenic cell fate decisions have been extensively studied using ex vivo cultures of MuSCs on primary myofibres in suspension, a system that partially preserves the MuSC niche20, followed by static analysis of sister cell fates4,35–38. However, myogenic cells are highly motile on cultured myofibres39, thereby creating uncertainty in identifying daughter cells by static imaging. To overcome these limitations, we designed a confinement system with microwells that enables suspension culture, live imaging and retrospective immunostaining of single myofibres (Fig. 2a, b and Fig. S3a). FDB myofibres were selected due to their small size (length ∼700 μm), which facilitates their manipulation and full-length imaging. FDB fibres from adult Pax7CreERT2; R26mTmG mice in microwells were filmed for 72 h to monitor MuSC activation kinetics (Fig. 2c and Movie S7). MuSCs transiting from quiescence to early activation were mostly immobile and showed dynamic cellular protrusions, as previously reported29,30,40. They migrated more rapidly upon activation than during quiescence, proliferated, and underwent cell divisions where daughter cells were tracked continuously for up to five consecutive divisions (Movie S7). Therefore, our culture and imaging system allowed quantitative assessment of multiple parameters from quiescence to differentiation of each cell in the lineage on the myofibre niche.

To assess the properties of dystrophic myogenic cells, we isolated FDB fibres from 4-month-old Pax7CreERT2; R26YFP WT or mdx mice in microwells, tracked YFP+ MuSCs and their progeny, recorded cell migration, and immunostained for MYOG to reconstruct the lineage and modes of cell divisions retrospectively (Fig. 2a, d). Of note, mdx fibres exhibited centrally located nuclei (46.9%, vs 3.2% in WT) (Fig. S3b, c) in resting FDBs, confirming that they had undergone regeneration (Fig. S2c–e). The initial number of MuSCs per fibre was similar between WT and mdx mice (mean WT=1.8, mean mdx=2.3, p=0.055, Figure S3d). Further, sampling of the MuSC population by partial recombination of the R26YFP allele through Pax7CreERT2 and low-dose tamoxifen (see Methods) yielded similar results between WT and mdx (mean recombination WT=57.4%, mdx=60.6%, p=0.88, Fig. S3e), indicating no overt bias due to heterogeneities in behaviour among clonally labelled WT and mdx MuSCs. As previously reported39,41, WT MuSCs executed their first division on average around 47.3 h, then divided approximately every 8.5 h (Fig. 2e), further validating the fidelity of our culture system. Of note, daughter cells took about 2.3 h to fully disconnect post-mitosis (Fig. S3f), highlighting this time window for accurate identification of sister cells and their fate (a)symmetry4,35–38. Further, except for the first cell cycle where there was no difference in cell cycle kinetics, mdx MuSCs divided more slowly than WT cells (Fig. 2e).

Importantly, mdx MuSCs showed increased differentiation (mdx=76%, WT=38%, p=4.31e-06, Fig. 2f) and reduced proliferation (mdx=3.5, WT=7.9, p=8.88e-11, Fig. 2g) indexes compared to WT, thereby confirming our in vivo observations (Fig. 1). Further, we observed symmetric proliferative (SCDp; sister cells MYOG-, Fig. 2h top) and differentiative (SCDd, sister cells MYOG+, Fig. 2h middle) divisions, and asymmetric cell divisions (one sister MYOG+, Fig. 2h bottom), as reported previously5,41,42. WT MuSCs divided mostly symmetrically (ACD=12.9%; SCDd = 32.5%, SCDp = 54.5%, Fig. 2i), as in vivo5. Unexpectedly, mdx MuSCs showed no impairment of ACD4 (14.4%, p-value vs WT=0.77) but drastic alterations of symmetric divisions, with increased SCDd (57.7%, p-value vs WT=3.3e-05) and decreased SCDp (27.9%, p-value vs WT=1.2e-05) (Fig. 2i). Further, the mode of cell division was reported to be impacted by the extrinsic microenvironment and the orientation of the mitotic spindle apparatus (see43). To explore this possibility, we determined if the orientation of cell division relative to the myofibre was related to the modes of cell divisions, as early ACD decisions were reported to result from differential exposure of sister cells to basal lamina and myofibre-derived cues4,35–38. We observed planar (68.0%, Fig. 2j left and 2k) and perpendicular (division axis perpendicular to fibre at mitosis, 18.9%, Fig. 2j right and 2k) divisions of myogenic cells on myofibres, in agreement with previous reports36,39. However, analysis of planar and perpendicular cell divisions during live imaging showed no significant differences in fate outcomes for WT (Fig. 2l) and mdx myogenic cells (Fig. S3g) that were related to the orientation of cell division plane.

Next, we quantified migration parameters of WT and mdx MuSCs by imaging ex vivo. While WT myogenic cells migrated actively, a fraction of mdx cells showed low migration capacity. We then stratified the data into ‘mobile’ and ‘static’ populations, to exclude static cells that can generate confounding effects when measuring speed and straightness. A migration speed threshold of 0.41 μm/min (see Methods) was used to discriminate between mobile and static fractions (Figure S3h, Movie S8) for all ex vivo migration analyses (Figs. 2–4), and we observed that mdx myogenic cells exhibited a lower mobile fraction (mean = 0.19) than WT cells (mean = 0.52, p=3.33e-19, Fig. 2m). Further, the mobile fraction of mdx myogenic cells showed a lower migration speed (mean: mdx=36.4 μm/h, WT=46.4 μm/h, p=5.85e-12, Fig. 2n) and lower net displacement (Fig. S3i) than WT cells. The migration straightness was similar between the mobile fractions of WT and mdx cells (Fig. 2o), unlike our in vivo observations (Fig. 1h), albeit the in vivo analysis was performed on the total cell population. At the total population level (mobile + static), mdx myogenic cells showed a lower migration straightness ex vivo (mean: mdx=0.15, WT=0.21, p=1.35e-03 Fig. S3j) and higher turning angle (mean: mdx=97.6°, WT=81.4°, p=5.21e-13 Fig. S3k) than WT cells, as was observed in vivo (Fig. 1h, i).

Altogether, our ex vivo pipeline allowed high-throughput quantification and validation of precocious differentiation through SCD and impaired migration kinetics of mdx myogenic cells observed by intravital imaging during muscle regeneration.

Cross-grafting assay uncouples myofibre niche contribution to fate and migration phenotypes in dystrophic MuSCs
Although DYSTROPHIN is a well-known structural protein of muscle fibres, it was reported to be expressed in myoblasts as a regulator of ACD4. Therefore, perturbed myogenic cell fate decisions and migration in mdx could result from lack of DYSTROPHIN in myoblasts and/or myofibres. To discriminate between fibre-dependent vs -independent cues, we developed an ex vivo cross-transplantation assay where the behaviour of MuSCs of WT or mdx origin could be examined on myofibres of the opposite genotype and monitored continuously by live imaging.

WT or mdx MuSCs from Pax7CreERT2; R26YFP; MyogntdTom mice were engrafted to mdx or WT receiving FDB myofibres (Fig. 3a–c, see Methods, hereafter referred to as ‘WT_to_mdx graft’ and ‘mdx_to_WT graft’). The transplantation of WT MuSCs to WT fibres (WT Ctrl) and of mdx MuSCs to mdx fibres (mdx Ctrl) were used to monitor possible effects of the grafting procedure (Fig. 2 and Fig. S3).

We first measured cell cycle progression for all grafting conditions. Interestingly, mdx Ctrl cells executed their first division (mean 36.5 h) before WT Ctrl cells (mean 43.5 h, p=1.32e-05, Fig. 3d), unlike WT and mdx MuSCs in their endogenous myofibre niche (Fig. 2e). Upon fibre isolation and ex vivo culture, MuSCs exited from their niche and transited from below the basal lamina (sub-laminal) to above (supra-laminal) within 24 h (Fig. S4a–c). Upon ex vivo grafting, MuSCs adhered directly to the myofibre basal lamina (supra-laminal position, Fig. S4d, e). The difference in cell cycle entry between WT and mdx MuSCs observed upon grafting (Fig. 3d) might reflect mdx MuSCs poised for activation, masked by a delay in niche exit in the endogenous condition (Fig. 2e).

mdx Ctrl cells cycled slower than WT Ctrl cells in subsequent cell cycles, as observed with the endogenous condition. Notably, WT_to_mdx cells had a cell cycle progression like WT Ctrl cells but distinct from mdx Ctrl cells, whereas mdx_to_WT cells had a cell cycle progression like mdx Ctrl cells but different from WT Ctrl cells (Fig. 3d and related Fig. S4a–e), suggesting that proliferation kinetics of MuSCs is mostly independent from the myofibre environment.

Next, we measured migration parameters (mobile fraction, speed, straightness, displacement), for all or separated cell cycles (Fig. 3e–g and Fig. S4f–k) upon cross-grafting. The mobile fraction (Fig. 3e), migration speed (Fig. 3f) and net displacement (Fig. S4j) of the WT Ctrl and mdx Ctrl differed (p=4.69e-14, 9.68e-04 and 5.50e-07 respectively across all cell cycles), while their migration straightness (Fig. 3g and Fig. S4i) was similar when analysing the mobile fraction (p=0.69 across all cell cycles) but different at the total population level (Fig. S4k, p=2.48e-05 across all cell cycles), showing that cross-transplantations can recapitulate the differential migration phenotypes of WT and mdx MuSCs observed in their endogenous niches.

Strikingly, the mobile fraction and migration speed of the WT_to_mdx graft were different from the WT Ctrl (p=6.55e-14 and 4.05e-03 respectively across all cell cycles) but like the mdx Ctrl (p=0.62 and 0.92 respectively across all cell cycles) (Fig. 3e, f). Conversely, the mobile fraction and migration speed of the mdx_to_WT graft were different from the mdx Ctrl (p=3.24e-10 and 2.93e-03 respectively across all cell cycles) but like the WT Ctrl (p=0.33 and 1.0 respectively across all cell cycles) (Fig. 3e, f). At the total population level, the migration straightness of the WT_to_mdx graft was like the WT Ctrl (p=1.0) and the mdx_to_WT graft was like the mdx Ctrl (p=0.64) (Fig. S4k). Altogether, these observations indicate that the migration properties of MuSCs, and their alterations in mdx mice, are largely determined by myofibre-derived signals.

We then assessed division modes upon cross grafting. Consistent with our observations above (Fig. 2i), myogenic cells divided mostly through proliferative symmetric divisions in the WT Ctrl graft (ACD=8.3%; SCDd = 17.3%, SCDp = 74.4%, Fig. 3h), while they showed impaired SCDs in the mdx Ctrl control (ACD=14.6%; SCDd = 60.7%, SCDp = 24.7%, p-value vs WT Ctrl=1.09e-13), thereby validating the cross-transplantation assay. Notably, division modes in the WT_to_mdx graft (Fig. 3h) were like the WT Ctrl (p=0.27) but different from the mdx Ctrl (p=2.22e-09). Further, the division modes in the mdx_to_WT graft (Fig. 3h) were like the mdx Ctrl control (p=0.12) but different from the WT Ctrl (p=1.35e-08). These observations show that the division modes of MuSCs are not differentially affected by healthy vs dystrophic myofibre niches.

Finally, we measured differentiation and proliferation indexes (Fig. 3i, j) at the population level following cross-transplantations. As expected, the differentiation and proliferation indexes were different between the WT Ctrl and mdx Ctrl (p=3.6e-11 and 5.8e-06 respectively). The differentiation index of the WT_to_mdx graft (Fig. 3i) was like the WT Ctrl (p=0.62) but different from the mdx Ctrl control (p=5.0e-09), and the differentiation index of the mdx_to_WT graft (Fig. 3i) was like the mdx Ctrl (p=0.32) but different from the WT Ctrl (p=7.5e-08). These results indicate that the precocious differentiation of mdx myogenic cells was not rescued by a WT myofibre, and that the differentiation of WT myogenic cells was not affected by the dystrophic environment. This was also the case for proliferation indexes which were found to be independent of the myofibre niche (Fig. 3j). Altogether, our results show that proliferation and differentiation decisions through ACD/SCDs are largely independent of the respective myofibre niches and appear to be driven by MuSC-intrinsic cues.

In addition, we used the MyogntdTom live reporter mice to monitor cell fate decisions for each cell cycle over consecutive divisions (Fig. 3c and Movie S9). Most of the Myog-ntdTOM expression appeared ~9 h post-mitosis (Fig. S5a), with rare examples (9.1%) of Myog-ntdTOM-positive cells performing one more cell division (Fig. S5b), as reported33, and most SCDd cells originated from a Myog-ntdTOM-negative mother cell (90.9%; Fig. S5b).

mdx MuSCs lack Dystrophin expression4 but are also exposed to chronic environmental stress (inflammation, fibrosis, activation, etc). To assess whether prior regenerative history of the muscle alters resident MuSC properties, we grafted WT MuSCs from either uninjured (qMuSCs) or regenerated (>28 dpi; PA_qMuSCs. PA=Pre-Activated) FDB muscles onto WT uninjured fibres and quantified activation (first-mitosis timing), proliferation, and differentiation (Fig. S5c–g). Pre-activated MuSCs divided earlier (32.6 h vs 39.8 h for qMuSCs) and exhibited a higher differentiation index (PA_qMuSCs 34.3% vs qMuSCs 22.6%), while proliferation indices were similar (Fig. S5e–g). Some phenotypes of pre-activated MuSCs resembled those of mdx (faster activation, precocious differentiation). We then compared grafting of PA_qMuSCs or qMuSCs to grafting of WT or mdx MuSCs on WT fibres (Fig. 3). The fold-change in activation time between qMuSCs and qPA_MuSCs (1.22; 39.8/32.7) closely mirrored that between WT and mdx MuSCs in the same assay (1.17; 43.5/37.1; Fig. 3d). This indicates that prior activation can account significantly for the faster activation of mdx MuSCs. For differentiation, the fold-change between pre-activated and qMuSCs (1.5; 34.3/22.6) accounted for ~50% of the difference between mdx and WT MuSCs (2.6; 62.0/24.0; Fig. 3i), suggesting that environmental history explains roughly half of the precocious differentiation phenotype in mdx. However, pre-activation did not reproduce the reduced proliferation phenotype of mdx MuSCs.

Altogether, these results show that certain aspects of MuSC behaviour (e.g. activation timing, differentiation) can be modulated by environmental cues, whereas proliferation defects likely arise from other cell-intrinsic factors.

Excessive differentiation of dystrophic MuSCs is driven by the coordinated activity of p38 MAPK and PI3K pathways
To explore the molecular mechanisms behind the fibre-dependent migration defects in mdx MuSCs, we analysed RNA-seq data from FDB fibres of 2- and 5-month-old WT and mdx mice (Fig. 4a–c, Fig. S6a)44 and scRNA-seq data from gastrocnemius muscle of 2-month-old WT and mdx mice7 (Fig. S6b). Gene Ontology analysis revealed significant enrichment of terms related to migration and differentiation in mdx fibres and MuSCs (Fig. 4b, Fig. S6a, b). Differentially expressed genes included Parva, Lrp1, and Adipoq (negative regulators of migration45–47 and upregulated in mdx), and Nexn (positive regulator of migration48 and downregulated in mdx) (Fig. 4c). Additionally, Tspan and Emilin1 (upregulated in mdx, Fig. 4c) and integrins (upregulated in mdx, Fig. 4b) were implicated in migration inhibition49,50.

To investigate the mechanisms of the precocious differentiation of mdx MuSCs, cells from hindlimb muscles of Pax7CreERT2/+; R26YFP/+; MyogntdTom/+; WT and mdx mice were cultured with p38 MAPK and/or PI3K inhibitors (SB203580 [SB] and LY294002 [LY], respectively) and live-imaged continuously to track proliferation, differentiation, and fusion (Fig. 4d–n, Fig. S6c–f and Movie S10). As observed previously, vehicle-treated (DMSO) mdx cells showed earlier differentiation (Fig. 4e) and reduced proliferation compared to WT (Fig. 4g), leading to fewer cells (in total and differentiated, reaching a plateau after 60 h). Temporal analysis of differentiation index confirmed the precocious differentiation phenotype of mdx cells (Fig. 4i), later reaching a maximal index of 75%, as WT cells.

In mdx cells, combined SB and LY treatment nearly abolished differentiation (88.9% reduction, p=4.61e-07), while in WT cells, SB alone suppressed differentiation by 82.6% (p=1.19e-02), with LY having no significant effect (Fig. 4f, Fig. S6c). Similar observations were obtained when analysing the differentiation index (Fig. 4j). Interestingly, dynamic analysis of the differentiation index with inhibitors (Fig. S6f) revealed three phenotypic classes: i) mdx-DMSO showed the most precocious differentiation, ii) WT-DMSO, WT-LY, mdx-LY, mdx-SB showed that WT are insensitive to PI3K inhibition and single inhibitors rescue mdx toward WT levels, iii) WT-SB, mdx-SB+LY, WT-SB+LY showed that dual p38/PI3K inhibition is required for mdx to reach the single-p38 inhibition effect in WT. These results indicate that mdx differentiation depends on both p38 and PI3K pathways, whereas WT differentiation relies primarily on p38.

SB-treated WT cells showed increased proliferation (p=3.89e-02), while LY alone or combined with SB had no effect (Fig. 4h and Fig. S6d). In mdx cells, neither SB nor LY affected proliferation, indicating that the proliferation defect of mdx cells is not a direct consequence of their precocious differentiation but rather a distinct phenotype, as SB and/or LY treated mdx cells did not differentiate but still failed to proliferate. Possible mechanisms include cytokinesis defects as suggested by centrosomal anomalies4 and related Gene Ontology-terms enrichment (Fig. S6b).

Analysis of myotube production over time revealed that mdx cells generated myotubes earlier but failed to sustain production at later stages (Fig. 4k–n and Fig. S6e). Blocking p38 or PI3K pathways did not rescue this defect (Fig. S6e).

Ex vivo fibre experiments (Fig. S6g–i) showed that the differentiation of mdx cells was reduced by p38 and PI3K inhibitors, and the dual p38/PI3K inhibition appears synergistic albeit not statistically significant. WT cells showed no detectable change upon PI3K inhibition, although p38 blockade tended to decrease differentiation. In both WT and mdx cells, inhibitor treatments had no significant effect on proliferation, whereas p38 inhibition stimulated WT proliferation in vitro (Fig. 4h), where all other treatments showed no effect. In addition, SB treatment did not affect MuSC motility (Fig. S6j-l).

In summary, mdx MuSCs exhibit p38- and PI3K-dependent precocious differentiation, contrasting with WT cells’ p38-only dependence, together with proliferation defects. These defects lead to premature myotube formation and insufficient production of fusion-competent cells at later stages.

Methods
Mousestrains and genotyping
Animals used in this study were handled according to national and European community guidelines, and protocols were approved by the ethics committee (CETEA, comité d'éthique en expérimentation animale) at Institut Pasteur (Licence 2015-0008 and DAP220077). Animals were housed at 22±2°C, with 30-70% humidity and under a 14:10h light:dark cycle. Pax7CreERT226, R26mTmG27, R26YFP34, MyogntdTom33 and Dmdmdx-βGeo28 mouse breeding was performed on a mixed B6D2F1/JRj background. 4-month-old male littermates were used, hemizygous for Dmd and heterozygous for other alleles. Study was investigated in males only as Duchenne Muscular Dystrophy is an X-linked disease affecting males only. Mice were genotyped at 3 weeks by PCR from an ear punch biopsy. PCR primers and conditions are listed in Table S1.

Tamoxifen treatment and muscle injury
Tamoxifen (T5648, Sigma) was reconstituted at 25 mg/ml in corn oil/ethanol (2.5%) and stored at -20°C. Tamoxifen was administered (200 μl/injection) by intragastric injection, once or 5X over 5 days, followed by 1 week of chase. Muscle injury was done as described65. Briefly, mice were anesthetized with ketamine (Imalgene 1000®, 100 mg/kg in NaCl 0.9%) and xylazine (Rompun 2%®, 12 mg/kg in NaCl 0.9%). The FDB muscle was injected with 20 μl of 10 μM cardiotoxin (L8102, Latoxan) in NaCl 0.9% using a 30G insulin syringe (BD, Micro-Fine 324826).

Intravital imaging
4-month-old male littermates Pax7CreERT2/+; R26mTmG/+; Dmd+/Y or Dmdmdx-βGeo/Y were treated once with tamoxifen, followed by 1 week of chase, to label MuSCs and their progeny (myoblasts and newly formed myofibres) with mGFP, and the surrounding tissue with mTOMATO. FDB muscles were injured (see below), one day apart for WT and mdx mice as intravital imaging sessions allowed to process 1 mouse/day.

On the day of imaging, the mouse was treated with buprenorphine (intraperitoneal injection, 0.1 mg/kg, Vetergesic®). After 30 min, the mouse was anaesthetised in an induction chamber with isofluorane (Iso-vet) 3% and O2 2 L/min in an incubation chamber, transferred onto a heat pad (Minerve) and anaesthesia was maintained using a rodent-adapted mask (Minerve) with 1-1.5% isoflurane and O2 0.6 L/min delivered through a gas humidifier. Ophthalmic gel (Ocry-gel®) was applied to the eyes to prevent corneal drying. The mouse foot was stabilised with tape, cleaned with 70% ethanol, and a 2×3 mm area of skin was carefully removed to expose the FDB. The FDB was immediately gently pressed against a glass-coverslip attached (silicone) to a 3D-printed custom coverslip holder, immobilised with tape and sealed (silicone) to the coverslip. After polymerisation of the silicone, the foot was turned upside-down to have the coverslip and the FDB facing up, and the coverslip holder was secured to the heating pad (silicone). The mouse was rehydrated with pre-warmed 0.9% NaCl (0.5 ml per hour) through a subcutaneous catheter (SV*S23NL30, Terumo) (Figure S1).

Imaging was done using a multiphoton microscope (TrimScope Matrix, Miltenyi Biotec). Excitation was performed at 950 nm with 3% laser power (Insight X3, Spectra Physics) and photons were collected with a 25x/NA 1.05 water immersion objective (Evident, Olympus) and detected on Gaasp detectors (Hammamatsu) at 510/20 nm (GFP) and 580/25 nm (tdTOMATO).

Imaging was performed by capturing tiles of 393×393 μm (1024×1024 pixels) on a 2.2×0.8 mm mosaic, with a z-stack of 80 μm (z-step 3 μm). Images were acquired every 10 min for 8-10 h. Mouse breathing was monitored every hour and anaesthesia adapted if necessary. At the end of the imaging session, the mouse was euthanised by cervical dislocation.

Assembling of movies was done with ImageJ with custom-made macros and the Stitching plugin66.

Immunohistochemistry
Adult FDB muscles were dissected and fixed for 2 h in PBS PFA 4% Triton X-100 0.1% at 4°C with gentle rolling. After an overnight PBS wash, samples were equilibrated in PBS sucrose 15% for 4 h at 4°C, followed by overnight incubation in PBS sucrose 30%. Tissues were then embedded in OCT compound (Sakura Finetek, Cat.# 4583) and frozen in isopentane cooled with liquid nitrogen. Cryosections (15 μm) were permeabilised with PBS Triton 0.5% for 5 min at RT and blocked for 1 h at RT in blocking solution (10% Goat serum, 1% BSA and 0.5% Triton X-100 in PBS). Primary antibodies were added in blocking buffer overnight at 4°C. Following washes in PBS-T (0.1% Tween 20 in PBS), secondary antibodies and Hoechst were diluted in blocking solution and incubated for 45 min at RT. Primary and secondary antibodies used are listed in Table S2. Images were acquired by Zeiss LSM800 confocal microscope.

MuSC analysis, isolation, culture and live imaging
FDB or hindlimb muscles were dissected and minced in a drop of cold collagenase type 2 (1000 U/ml, WOLS04177, Serlabo) in Muscle Dissociation Buffer (MuDB, Nutrient mixture Ham F10 (N6635, Sigma), 10% heat-inactivated Horse serum (HS, 11510516, Thermo Fisher), supplemented with NaHCO3 to pH 7.4 in distilled water). Samples were incubated in 10 ml of a collagenase 2/MuDB for 90 min at 37°C in a water bath under gentle agitation (70 rpm). Samples were mechanically dissociated (25 ml pipette), MuDB was added up to 50 ml and samples were spun (10 min, 500g at RT). The supernatant was removed up to 20 ml, cell pellet was resuspended and 1 ml of Dispase (1.8 U/ml in MuDB, 17105-041, Gibco) and 20 μl DNase I (10 mg/ml in DMEM, 11284932001, Roche) were added. Samples were incubated for 90 min at 37°C in a water bath under gentle agitation (70 rpm). Samples were passed through a syringe (Agani Needle 20G, 050109B Fisher Scientific) 10 times, filtered through 40 μm strainer (352235, Dutscher), spun (10 min, 500g at 4°C) and resuspended in 500 μl of DMEM/2% HS before cytometry.

Using Fluorescence Activated Cell Sorting (FACS), samples were analysed (CytoFLEX, Beckman Coulter, Fig. 1m-p) and MuSCs were isolated (Aria III, BD Biosciences, Fig. 4d-n) based on cell size, granularity and YFP fluorescence (70 μm nozzle) (Figure S8). Cells from Pax7CreERT2/+; R26YFP/+ mice were used to determine the positivity threshold of Myog-ntdTOM. Cells were collected in differentiation medium (88.5% DMEM, 10% HS, 1% Penicillin/Streptomycin, 0.5% Chick Embryo Extract (MD-004D-UK, Life Science Production)) at 4°C. Isolated MuSCs were plated at 4500 cells/cm2 in differentiation medium in 96-well plates (PhenoPlate (6055300, Revvity), well surface = 0.32 cm2) coated with fibronectin (F1141, Sigma; 40 μg/ml in NaHCO3 0.1 M pH 8.3 for 30 min at RT followed by 3 PBS washes). p38 MAPK inhibitor (SB203580 5 μM, 559389, Sigma), PI3K inhibitor (LY294002, 10 μM, 9901S, Cell Signaling) or vehicle (DMSO) were added at 21 h post plating. Cells were incubated at 37°C, 3% O2, 5% CO2. Cells were live-imaged with a Zeiss Observer Z1 equipped with a Plan-Apochromat 20x/0.8 M27 objective, Colibri 7 LEDs, a Hamamatsu Orca Flash 4 camera and piloted with Zen software (Carl Zeiss), at 37°C, 5% CO2, 3% O2 (Pecon incubation chamber). Each well was imaged every 3 h in 2 channels (YFP, DsRed).

Microwell plate assembly
The microwell plate is composed of two Plexiglas plates (bottom plate: 128 mm×85 mm x 10 mm; top plate: 123 mm×81 mm x 10 mm) in which 4 rectangles (55.5 mm×34 mm x 10 mm) have been laser-cut (Speedy 300, Trotec), cleaned and attached together (Figure S3a) with silicone (Mold Star 20T, Smooth-on). 4 glass coverslips (Claritx Coverglass #1, Eloïse-SARL) were cut to 58 mm×38 mm, cleaned and sealed (Figure S3a) with silicone to the outer part of the bottom Plexiglas plate to form 4 glass-bottom chambers. The lid of a 6-well dish (TPP) was used to close the microwell plate on the top Plexiglas plate.

Silicone sheets with microwells were cleaned and placed on glass coverslips (Fig. S3a) and fixed to the glass with additional silicone on the edges. Microwells were pierced manually (biopsy ear punch in 1 mm-thick silicone; microwell volume ∼3 μl. 50 microwells/glass-bottom chamber. Figure 2b and Fig. S3a) or formed using a 3D-printed mold (3 mm-thick silicone; microwell volume ∼6 μl. 200 microwells/glass-bottom chamber Fig. S3a). Spinning-disk confocal live-imaging (Fig. 2c and Movie S7) was carried out using a 2 well glass bottom (80287, Ibidi) in which two silicone sheets with microwells were sealed with silicone.

All the material (Plexiglas plates, coverslips, silicone microwells) was cleaned and sterilized with soap, water and ethanol 70% before use. After assembly of the microwell plate, microwells were washed with PBS and equilibrated with culture medium (5 ml per chamber, final volume 10 ml) to prevent adhesion of muscle fibres. Air bubbles in microwells were removed with a P200 pipette and single FDB fibres were loaded in microwells. As the height of the microwells (1 or 3 mm) is 20–60 times larger than the thickness of an FDB fibre (50 μm), this microwell design allows confinement, culture and live-imaging of single FDB fibres in non-adherent conditions. All subsequent manipulations after fibre loading (medium exchange, live-imaging, immunostaining) were done as usual when manipulating a 6-well plate.

Single myofibre isolation, culture and live imaging
A solution of 0.2% collagenase Type 1 (C0130, Sigma) was prepared in DMEM GlutaMAX (31966, Thermo Fisher), filtered (0.22 μm) and kept at 37°C. Mice were sacrificed by cervical dislocation. The foot was cleaned with 70% ethanol and the skin covering the Flexor Digitorum Brevis (FDB) was cut from the base of the heel to the toes. The FDB muscle was dissected away from the surrounding tissue, being handled carefully by the proximal tendon to avoid muscle damage and contraction. The FDB muscle was incubated in 5 ml of collagenase solution in Sterilin 7 mL Polystyrene Bijou Containers (11399133, Fisher) for 2.5 h at 37°C.

Individual muscle fibres were released by mechanical dissociation (glass Pasteur pipette) of FDB in Sterilin petri deep dishes (10655821, Fisher) filled with pre-warmed DMEM. The majority of the FDB fibres were released after five rounds of 10 vigorous but careful back-and-forth pipetting, separated by 5-min pauses at 37°C. After each round, the FDB was transferred to a new Sterilin dish with pre-warmed DMEM for further dissociation. Large debris were removed with a plastic Pasteur pipette. When most fibres were dissociated, two-thirds of the DMEM volume were removed from each Sterilin dish. Muscle fibres were then pooled in a new Sterilin dish filled with pre-warmed DMEM, washed with DMEM and distributed to 5 Sterilin dishes containing culture medium and caps of 15 ml Falcon tubes in their centre (fixed with silicone) to prevent fibres from concentrating and aggregating in the centre of the dishes. Sterilin dishes and Pasteur pipettes were coated with horse serum (HS, 11510516, Fisher Scientific) before use to prevent adhesion of muscle fibres. Muscle fibres were loaded in the microwell plate in bulk and/or manually under the binocular loupe using a P20 pipette.

Muscle fibres were cultured in differentiation medium at 37°C, 5% CO2, 3% O2. For p38 inhibition experiments, p38 MAPK inhibitor (SB203580 5 μM, 559389, Sigma) or vehicle (DMSO) were added directly to the culture medium ~39 h post-isolation of muscle fibres.

Spinning-disk confocal live-imaging of FDB fibres (Fig. 2c and Movie S7) was performed immediately after isolation using a Spinning Disk Confocal Yokogawa CSU-W1 on a Nikon Ti2E equipped with a 20x dry objective (numerical aperture 0.75, working distance 1 mm), a motorized XY stage (with a Z piezostage, 200 μm range), a Hamamatsu Orca Flash 4 camera (pixel size 6.5 μm, 2048 × 2044 pixels, quantum efficiency 82%), and piloted with Nikon software (NIS Element) at 37°C, 5% CO2 and 3% O2 with humidity control and objective heater (Okolab). Each fibre was imaged every 10 min in 488 nm (300 ms, 1.5% laser power) and 561 nm (200 ms, 0.8% laser power), with a Z-stack of 71 μm (1 μm slices).

Widefield live-imaging of FDB fibres (Fig. 2d–o, Fig. S3, Fig. 3 and Fig. S4f–m, Fig. S5) in microwell plates was performed between ~17 h and 64 h post-isolation with a Zeiss Observer Z1 (see above). Each fibre was imaged every 10-14 min in 2 or 3 channels (Brightfield, YFP, DsRed), with a Z-stack of 50 μm (5 μm slices). Stage parameters were set at 50% speed, 5% acceleration.

Immunocytochemistry
Muscle fibres were fixed with 4% paraformaldehyde (PFA, 15710, Euromedex) in PBS (D1408, Sigma) for 10 min at room temperature (RT), washed twice 5 min in PBS, permeabilized in cold permeabilization buffer (Hepes 20 mM pH 7.4, MgCl2 3 mM, NaCl 50 mM, Sucrose 300 mM, 0.5% Triton X-100) for 15 min at 4°C and washed three times in PBS. Fibres were blocked with 10% Goat Serum (GS, 11540526, Thermo Fisher) in PBS (0.22 μm-filtered) for 1 h at RT and washed once in 2% GS in PBS (0.22 μm-filtered). Excess of liquid between microwells was removed, leaving each microwell with 3 μl of 2% Goat Serum solution. Primary antibodies (Table S2) were diluted in 2% GS to a 4x concentration compared with their final concentration. 1 μl of diluted antibodies was added to each microwell (3 μl) and incubated overnight at 4°C. After three washes with 2% GS for 5 min at RT, the excess of liquid was removed. Secondary antibodies (Table S2) were diluted in 2% GS to a 4x concentration compared with their final concentration. 1 μl of diluted antibodies was added to each microwell (3 μl), for 1 h at RT. Fibres were washed with PBS (5 min at RT), incubated with 1 μg/ml Hoechst 33342 (H1399, Thermo Fisher) in PBS (5 min at RT), washed twice with PBS (5 min at RT) and stored at 4°C.

Immunostainings were analysed with Zeiss LSM800 confocal microscope. Each fibre was imaged in 3 to 5 channels (Brightfield, 488, 555, 633, Hoechst), with a centred Z-stack of 42 μm (1.9 μm slices) with a Plan-Apochromat 20X/0.8 M27 objective. Our confinement system allowed filming and staining the same fibres (Fig. 2a-b, Figure S3a and Movie S7), despite their culture in floating conditions.

Cross-transplantation assays
Two 4-month-old male littermates Pax7CreERT2/+; R26YFP/+; MyogntdTom/+; Dmd+/Y or Pax7CreERT2/+; R26YFP/+; MyogntdTom/+; Dmdmdx-βGeo/Y were treated 5X with tamoxifen, followed by 1 week of chase, and served as donors of WT or mdx YFP-labelled MuSCs. The MyogntdTom allele harbours a nuclear tdTOMATO reporter for Myogenin expression, in the 3’ UTR of the Myogenin locus. Two 4-month-old male littermates Dmd+/Y or Dmdmdx-βGeo/Y served to generate WT or mdx recipient FDB fibres (Fig. 3b and Figure S7).

For each recipient mouse, fibres were isolated (see above) from 1 FDB. Half of the isolated recipient fibres were split into two Sterilin dishes (with caps of 15 ml Falcon tubes fixed in the centre and culture medium) and then transferred into two 15 ml Falcon tubes with the base cut (to form a cylinder) and attached to the lid of a Sterilin dish (silicone). This setup allowed to load a large volume (~12 ml) in the Falcon tube and to concentrate the recipient fibres in a small surface (cross section area of 15 ml tube, ~1.8 cm2) after sedimentation to maximise contact between recipient fibres and grafted MuSCs (see below). The recipient fibres were then incubated at 37°C, 5% CO2 and 3% O2 while the following steps were carried out.

For each donor mouse, fibres were isolated (see above) from both FDBs. Fibres were washed twice in PBS, PBS was removed as much as possible and TrypLE Express (12604013, Thermo Fisher) was added (2 ml) for 15 min at 37°C. MuSCs were then dissociated mechanically using a P1000 pipette, filtered through a 30 μm filter (130-041-407, Miltenyi) into a 15 mL falcon, centrifuged at 500g for 15 min at 21°C and resuspended in 500 μl differentiation medium.

The medium in the 15 ml Falcon tubes containing the recipient fibres was removed to leave 1.5 ml, to which 250 μl of MuSCs from donor mice were added. WT and mdx MuSCs were incubated with both WT and mdx fibres, corresponding to 4 grafting conditions. MuSCs and recipient fibres were incubated for 5 h at 37°C, 5% CO2 and 3% O2. For each grafting condition, medium was added (10 ml) in the 15 ml Falcon tube, the cell suspension was transferred into two Sterilin petri dishes to facilitate handling and the fibres were loaded in microwells of 1 chamber of a custom-made microplate (see above). Fibres were loaded in bulk from one Sterilin dish and manually from the other to reach full occupancy of microwells. During loading of a chamber, the other chambers were covered with Parafilm to ensure sterile conditions.

The microplate was then incubated at 37°C, 5% CO2 and 3% O2 before live-imaging. Under these conditions, each chamber (i.e., grafting condition) of the microwell plate contained ~200 recipient fibres isolated in microwells, ~25% of them grafted with 1-2 YFP-labelled MuSCs from donor mice. For each experiment, ~20 fibres per grafting condition were imaged live.

Ex vivo cell tracking and modes of cell division
Widefield live-imaging data of MuSCs on FDB fibre ex vivo was converted to 8-bit Tiff with ImageJ (https://github.com/gletort/ImageJFiles/blob/master/convertFiles/convertCZIto8bTiff.ijm). MuSCs were tracked (cell body) manually using Phase and YFP channels for all timepoints with Trackmate67, and data was exported to .xml format for further analyses. All tracking procedures, developed softwares, detailed descriptions of the statistical analyses, together with the associated datasets and representative images and movies, have now been made available on GitLab: https://gitlab.pasteur.fr/hub/sarde_et_al_2025. Tracking was cross validated independently by two authors (LS, BE) through random sampling of raw data and duplicate manual tracking, which yielded identical outcomes.

The fate of sister MuSCs, identified by cell tracking, was determined by immunostaining at the end of the experiment (Fig. 2f, h, i, l and Fig. S3g) or continuously using live-imaging and the Myog-ntdTOM reporter (Fig. 3c, h, i and Fig. S5a, b).

Divisions were classified as symmetric proliferative (both sister cells MYOG or Myog-ntdTOM -negative), symmetric differentiative (both sister cells MYOG or Myog-ntdTOM-positive) or asymmetric (one sister cell MYOG or Myog-ntdTOM-positive, the other MYOG or Myog-ntdTOM -negative). Of note, comparison of WT Ctrl graft (Fig. 3h) with WT MuSCs on their own myofibre (Fig. 2i) showed a lower frequency of ACD (graft=8.3%, endogenous=12.9%) and SCDd (graft=17.3%, endogenous=32.5%) and higher frequency of SCDp (graft=74.4%, endogenous=54.2%), likely due to the lower sensitivity (short exposure times, slow maturation (t0.5 ~1 h) of tdTOMATO protein) of the Myog-ntdTOM live reporter compared to MYOG immunostaining).

The orientation of cell division was classified according to the position of sister cells at mitosis with respect to the FDB fibre: parallel if both sisters were in contact with the fibre, perpendicular if one of the sister cells had no contact with the fibre at mitosis. The orientation of some divisions could not be determined due to their dynamic nature and lack of spatiotemporal resolution with our imaging modality.

Analysis of cell migration
The migration of MuSCs (cell body) was analysed in vivo (Fig. 1f–l) and ex vivo on muscle fibres (Fig. 2m–o, Fig. S3h–k, Fig. 3e–g and Fig. S4f–m). Cells within dense areas in vivo (e.g., right part of Movie S3 where fusion has already begun) were excluded from analysis to avoid ambiguous segmentation. Areas analysed contained trackable cells with comparable densities in WT and mdx samples, as illustrated by tracking overlays (Fig. S2f, Movies S5 and S6). We first corrected the movements of regenerating muscles or FDB fibres to analyse MuSC migration in a static referential.

Displacement (translation, rotation, deformation) of FBD fibres ex vivo (due to culture in floating conditions) was corrected using a newly developed Fiji plugin: cellsOnFiber (distributed under the BSD-3 license and available in open source: https://gitlab.pasteur.fr/gletort/cellsonfiber/). Inputs are 3D movies of FDB fibres with TrackMate tracks of their associated MuSCs at every timepoints. This plugin detects the fibre from 2D projection of transmitted light images at each time frame, aligns it on the segmented fibre from the previous frame and corrects the TrackMate tracks accordingly. Outputs are 3D movies of FDB fibres and MuSCs TrackMate tracks corrected for fibre displacement.

Training dataset. We first imaged FDB fibres from R26mTmG mice in both the fluorescent channel (DsRed) and transmitted light and we obtained a ground-truth fibre segmentation by applying a simple threshold on the fluorescent channel. The dataset was composed of 97 filmed scenes imaged on 149 timepoints. For the training, we subsampled the dataset every 20 timepoints for each scene. Overall, around 750 images of size 352×352 pixels of fibre in transmitted light with its ground-truth based on the thresholded fluorescent channel were used for training with basic data augmentation (flippings, translation and histogram). The dataset was randomly split so that 20% of the fibres were used for validation and the rest for training. Test was performed visually on dataset without ground-truth.

Network architecture and training. We used a U-Net architecture68 to detect the fibre contour from transmitted light 2D images. After testing different size of the network, we built a U-Net with 5-layer blocks in the left part (condensing part) of the network and 16 initial features in the first layer. The implementation was done with the python Keras library. The loss was calculated with the Jaccard distance, and the final score used was the intersection over union. Training was performed on a local computer with 1 GPU, for 30 epochs with a batch size of 30. The trained network is accessible here: https://gitlab.pasteur.fr/gletort/cellsonfiber/-/blob/main/networks/FiberFromTransMatch.zip.

Registration and track correction. The plugin calculates the registration to apply to the movie on the segmented fibre images to obtain a stabilised fibre. The fibre was segmented in each Z-slice of the time frame and projected in 2D. The transformation to register the fibre is calculated in our plugin with TurboReg plugin69 on the projected movie of the segmented fibre. The resulting transformation is then applied to each Z-slice of the movie of each input channel and used to correct the TrackMate trajectories to the new registered movie. The obtained tracks correspond to the cell motion relative to the immobilised fibre in 3D.

Limitations. In 7% of the movies, FDB fibre movement or deformation impaired analysis, and the plugin could not correct for this motion. These tracks were not used in any motility analysis but could still be used in lineage analysis (e.g. cycle duration, modes of divisions).

To correct for movements of the tissue during intravital microscopy, a first step of image registration was performed using ~50 fixed points (mostly blood vessels) manually tracked with TrackMate Manual tracking67. We calculated the image registration on the channel containing the tissue information and applied it to channel containing the MuSCs. 3D registration was calculated with itk-elastix python module70. For each time frame, the registration was iteratively calculated to align to the previous frame. There was registration was optimised in this step to decrease the distance between the same point at two consecutive times. The rigid registration was applied at 4 levels of resolution with 1000 iterations and a size of the final spacing grid of 50 pixels. This first registration was used to compensate for large local translation/rotation. Then a finer step was applied with b-spline based transformation to account for local deformation of the tissue, again with itk-elastix library. This step did not use the reference point and calculated the registration based on the intensity Mutual Information. The spline registration was performed at 2 resolution levels, with 1000 iterations and a size of the final spacing grid of 200 pixels. Finally, the calculated transformations were applied on all the channels of the original movie. All these steps were implemented in a Napari plugin napari-3dtimereg (distributed under the BSD-3 license and available in open source here: https://gitlab.pasteur.fr/gletort/napari-3dtimereg).

Next, we aimed at analysing the speed and directionality of migrating cells. Cell trajectories and lineage information were extracted from TrackMate files with a customed Jython script. The trajectories were then analysed with R scripts to measure different motility parameters for each cell.

Some MuSCs did not actively migrating on ex vivo FDB fibres, notably for mdx MuSCs, and some cells exhibited irregular trajectories, with phases of very small movements around the same point and more effective phases with more directional and faster motion. To analyse migration parameters on actively migrating cells, we fractionated the population into mobile and static subsets (Fig. S4l, m). The threshold value between mobile and static was determined by analysing the turning angle (see below) and the instantaneous speed (see below) between each consecutive time frames for WT and mdx MuSCs in the endogenous (dataset of Fig. 2 and S3) (Fig. S4l) and grafting condition (dataset of Fig. 3 and S4, control conditions WT Ctrl and mdx Ctrl) (Fig. S4m). WT cells (pooled from endogenous and WT Ctrl grafting datasets) showed two subpopulations (mobile with high speed/low turning angle and static with low speed/high turning angle), with a median log speed of 0.41 μm/min allowing to discriminate these subpopulations. mdx cells (pooled from endogenous and mdx Ctrl grafting datasets) showed mostly one population (low speed/high turning angle), with a median log speed of 0.18 μm/min. We then used a threshold value of 0.41 μm/min to discriminate between mobile and static subsets for all ex vivo datasets (Fig. S4l, m). Intravital imaging during muscle regeneration also identified some cells with low migration capacity, notably for mdx mice. Nevertheless, we could not identify a specific parameter to discriminate clearly mobile and static subsets. The analysis of cell migration properties in vivo was then performed on the total population of cells.

Turning angle (°) measures the average angle between two consecutive cell displacements (Fig. 1f). This measure varies from 0° (straight in the forward direction) to 180° (backward direction). The average speed (μm/h) is the ratio between the total distance travelled and the trajectory duration. The net distance (μm) measures the effective distance (3D) between the first and the last point of the trajectory. The total distance (μm) measures the total distance (3D) travelled by the cell as the sum of the distances between each time frames. The straightness is the ratio of the net distance over the total distance (Fig. 1f). If the trajectory was totally straight, then these two parameters are equal and the straightness is equal to 1. Otherwise, the cell had travelled more in total (Total Distance) than the observed resulting distance (Net Distance), so the straightness is closer to 0.

For in vitro experiments, motility parameters were analysed for separated and pooled cell cycles.

In vitro analysis of proliferation, differentiation and myotube production
ZEN.czi files were analysed with ImageJ using the threshold tools to segment cells at each timepoint. Proliferation was assessed by measuring the number of YFP-positive objects per timepoint, differentiation by measuring the number of YFP and Myog-ntdTOM-positive objects, and myotube production by measuring the number of YFP-positive cells with an aspect ratio (Max diameter/Min diameter) superior to 2.

Software and analysis
All tracking procedures, developed softwares, detailed descriptions of the statistical analyses, together with the associated datasets and representative images and movies are available on GitLab: https://gitlab.pasteur.fr/hub/sarde_et_al_2025.

Statistical analyses
For cell migration, sister cell separation in vivo and in vitro analysis, linear mixed models were fitted using the nlme_3.1-168 under R version 4.5.0 and including the microwell identifier (nested within the experiment number) as random effects to test for the genotype [resp. inhibitor] factor. The mixed effects models included specific cell cycle variances when appropriate. The cell cycle variable was included in interaction with the genotype [resp. inhibitor] when testing for this effect for each cell cycle separately, otherwise the cell cycle was included as an additive main fixed effect. Pairwise comparisons were extracted from the linear mixed models using the emmeans R package. Statistical tests from linear mixed models were two-sided. When a model included two covariates as well as their interaction, all the pairwise comparisons were two-sided and adjusted for multiple testing using the Tukey’s method.

For FACS and ex vivo analysis, mean comparisons were performed using unpaired two-tailed Wilcoxon tests unless stated otherwise. Comparisons between more than two groups were performed using Kruskall-Wallis tests followed by a post-hoc pairwise comparison.

RNA-sequencing analyses
For analysis of bulk FDB fibres RNA-seq44, raw fastq files were downloaded from the Gene Expression Omnibus repository GSE162455 and aligned to the mm10 reference genome using STAR (v2.7.9a)71. Mapped reads were then quantitated using the RSEM pipeline (v1.3.3)72. Quality control plots and differential expression analyses were performed with the R package DESeq2 (v1.36.0)73.

For analysis of single-cell RNAseq7, Seurat objects were obtained from the authors and differentially expressed genes between conditions in MuSCs were identified using Wilcoxon tests implemented in the FindMarkers function from the R package Seurat (v4.4.0)74. Gene Set Enrichment Analyses were performed with R using the fgsea function with default parameters from the fgsea package (v1.24.0). For each Gene Ontology term included in the analysis, genes included in the term and its children’s terms were assigned to the specific term. Terms with less than 10 genes were discarded. Genes were ranked using -log10(adjusted p-value) * log2(fold change).

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "In this manuscript, Sarde et al extensively used live-cell imaging to examine the fate and behavior of muscle satellite cells (MuSCs) in mdx mice, a mouse model for Duchenne Muscular Dystrophy. The authors started by using intravital imaging to monitor the division and migration of GFP-labeled MuSCs 3 days after cardiotoxin-induced muscle injury. They found that MuSCs from mdx mice exhibited reduced proliferation and impaired migration as well as precocious differentiation in vivo. Subsequently, the authors imaged freshly isolated myofibers in custom-built microwells to study MuSC proliferation and differentiation over time. Using such an ex vivo system, they confirmed their findings in vivo by showing that MuSCs from mdx mice indeed had reduced proliferation and precocious differentiation. They further demonstrated that it is the symmetric cell division (SCD) instead of asymmetric division (ACD) that is impaired in MuSCs from mdx mice. They further designed an unique ex vivo cross-grating experiment by mixing freshly isolated MuSCs and myofibers from wildtype and mdx mice followed by live-cell imaging. They showed that the precocious differentiation and defective proliferation are intrinsic properties of MuSCs, while the migration of MuSCs was mainly influenced by myofibers. By applying kinase inhibitors for p38 MAPK and PI3K to myofibers in culture, they showed that the precocious differentiation of MuSCs from mdx mice is dependent on both p38 MAPK and PI3K, while the differentiation of MuSCs from wildtype mice is only dependent on p38 MAPK signaling.\nOverall, the experiments were well designed and controlled. The live-cell imaging of MuSCs genetically labeled with different fluorescent reporters allows the authors to track the cells over time and to pinpoint stage-specific defects in MuSCs from mdx mice. Moreover, the cross-grafting experiment allows the authors to determine which properties of MuSCs are intrinsically or extrinsically controlled. One caveat of the manuscript is that most studies were carried out in an ex vivo system. Thus, it remains to be confirmed whether some of the conclusions hold true in vivo.\nMy major comments are all related to Figure 4:\n1. For Figure 4d-i, the authors should use the same myofiber system as used in previous figures to monitor the behaviors of MuSCs with and without the inhibitors.\n2. Figure 4e, it is better to use the percentage of YFP+ Myog-ntdTOM+ cells/YFP+ cells for the Y axis. Based on the differentiation curve here, does it mean mdx MuSCs failed to undergo further differentiation after 60 h (as the number of Myog-ntdTOM+ cells plateaued after this time point)? What percentage of mdx MuSCs undergo differentiation at the end of the imaging?\n3. Figure 4g, based on the growth curve of mdx MuSCs, it seems mdx MuSCs only divided once (on average) within the imaging period (as YFP+ cell number changes from 500 to ~1,000). This contradicts with the data shown in Figures 1p, 2e, and 3c. How to explain this discrepancy?\n4. Figure 4f: the conclusion based on 4f that the differentiation of WT MuSCs/myoblasts does not depend on PI3K is not solid, as the analysis was carried out at a single time point (54 h) when very few WT cells undergo differentiation. At later time points, the differentiation of WT myoblasts was clearly inhibited by LY, the inhibitor of PI3K (Figure S4c, S4e).\n5. Figure S4c-e: SB appears to have dual effects on differentiation of both WT and mdx myoblasts: it inhibits differentiation at the early phase (between 50 and 70 h for WT myoblasts), but stimulates differentiation at the late phase (between 75-105 h for WT myoblasts). What is the underlying mechanism?\n6. It would be very novel if the authors can uncover the mechanisms underlying reduced proliferation or defective differentiation at the late phase for mdx MuSCs.", "reviewer_2": "The manuscript by Evano et al. reports on development of new technology to visualize and study the behavior of muscle stem cells (MuSCs) during repair of muscle injury in wild type and mdx mice (a model for Duchenne Muscular Dystrophy). Specifically, they have generated new intravital and ex vivo single myofiber live-imaging protocols. Such protocols already exist and are cited (Refs 21-23 and 38) but they are challenging and not widely used. The authors extend the intravital techniques to the accessible FDB muscle and the single myofiber technique to a microwell system. These techniques were also applied to mdx mice for the first time, revealing interesting properties in a dynamic manner. mdx MuSCs display aberrant behavior including diminished migration and precocious differentiation. A cleverly designed, ex vivo, single myofiber system of cross-grafting was also developed allowing analysis of cell-autonomous vs. myofiber niche-derived regulation of these phenotypes, with defects in differentiation being mainly cell-autonomous and migration problems mainly determined by the myofiber niche.\nThis paper has many strengths, including innovative new techniques and that the work appears to have been performed and reported very carefully. My comments derive from comparisons made in vivo vs. ex vivo and how MUSCs may behave differently in these conditions.\n1. In Figure 1, the authors study MuSCs 3 days post-CTX injury with intravital imaging, demonstrating perturbed migration and differentiation by mdx MuSCs. In Figures 2 and 3, they move to single myofiber analyses. Similar results were obtained but additional observations were made with the latter system. It has previously been shown that in wild type TA muscles 3 days post-CTX injury in vivo, MuSCs are present inside a remaining basal lamina (i.e., so-called “ghost fibers”) where they divide only along the axis of the myofiber (“planar” divisions, Refs 23 and 29). However, with single myofibers, by 24 hr of isolation, MuSCs have reversed their normal position to sit outside the basal lamina, and that is where cell divisions occur. Therefore, there is a major difference as to where the measured MuSC events occur. The authors’ results are largely consistent between the figures, so I don’t think the findings are invalidated, but the authors should comment on this being the case and what it may mean for interpreting the data.\n2. Along these lines, what is the location of MuSCs prior to injury and 3 days post-CTX injury in mdx mice? Are MuSCs present in ghost fibers in mdx mice in a similar way to wild type mice? Could many mdx MuSCs be outside the myofiber basal lamina and might that influence the results? Again, this is not invalidating, but the authors should consider whether they are truly making apples-to-apples comparisons.\n3. A similar issue occurs with the cross-grafting experiments. Do the grafted MuSCs remain outside the basal lamina?", "reviewer_3": "The authors describe the dysfunction of satellite cells (muscle stem cells) isolated from dystrophic muscle using innovative in vivo and ex vivo imaging systems. Their in vivo imaging is remarkable and their ex vivo culture experiments are technically impressive and informative. The authors demonstrate that SCs isolated from dystrophic muscle have greater differentiation propensity than WT isolated SCs and reduced proliferation and mobility. They demonstrate with an impressive SC-myofiber engraftment culture system that the altered differentiation/proliferation phenotype is intrinsic to the dystrophic SCs, but that the altered mobility is a consequence of the cellular environment (dystrophic myofiber). They provide some evidence that SCs from dystrophic muscle have excessive differentiation due to coordinated activity of p38 MAPK and PI3K pathways, which is distinct from WT derived SCs. This study is interesting and showcases innovative techniques, however there are some key conceptual limitations that should be addressed by discussion or ideally by additional experimentation.\nMajor comments:\n1. The authors conclude that SCs from dystrophic muscle have “intrinsic” changes and that “proliferation and differentiation decisions through ACD/SCDs are mostly driven by MuSC intrinsic cues, and largely independent of the respective myofiber niches.” This conclusion is made after an acute transfer experiment where SCs isolated from 4 month old healthy WT or sick DMD mice are introduced to either WT or DMD isolated myofibers and cultured for 20H-63H. While it is convincing that DMD derived SCs behave differently than WT derived SCs, and this difference persists after transplantation to myofibers, the authors do not address that DMD derived SCs have spent 4 months in a DMD environment/niche prior to transfer. During this time the SCs are exposed to inflammation, DAMPs, altered extracellular matrix, and have potentially gone through rounds of activation/division prior to isolation. Essentially the authors are likely comparing chronically activated DMD SCs to potentially naïve healthy SCs. While the data support that at the time of isolation there is an intrinsic difference between WT and DMD derived SCs, it is entirely possible this difference is caused by environmental signals/previous activation states of DMD derived SCs prior to isolation. I think it’s an over interpretation of the present data to conclude that this difference is intrinsic to SCs or due to Mdx-mutations in the DMD derived SCs rather than the disease state/environment the SCs are isolated from. This inability to separate intrinsic SC differences from the consequences of a chronic diseased environment is a major limitation of the study, and one that the authors need to explicitly address in their discussion, interpretation of results, and/or with experiments that tease out this difference. Resolving this issue would substantially increase the impact of this study.\n2. The definitive experiment would be to perform tissue-specific knockout of DmD or another dystrophy causing gene (such as Sgcd) with a Pax7-CRE or SKA-CRE such that the authors could examine dystrophic SCs raised in healthy muscle or wildtype SCs raised in dystrophic muscle. This would allow the authors to remove dystrophic environment as a confounding variable from the study and isolate intrinsic SC defects. This is more work than is fair to request for this revision and may not be realistic, since there is a frankly surprising shortage of cre/lox targeted dystrophic models. There may be a Sgcd-flox animal commercially available (https://en.gempharmatech.com/product/details100035_3700133.html), and I encourage the authors to consider doing this experiment in the future since they clearly have the expertise to perform these experiments (and I am curious to see the outcome).\n3. Is it possible for the authors to consider wildtype SCs isolated following injury/activation using their cell transplant model? Do wildtype SCs have altered function during cardiotoxin injury and activation? Do they behave differently after injury resolution? Re-activation? Perhaps this could be a way to test whether SCs are altered by environmental cues or previous activation states in a way that persists following transplantation.\n4. The authors show SC migration and activation in vivo in mouse feet following cardiotoxin treatment in wildtype and DMD mice (as in Figure 1). Do the authors have any information about SC activation/mobility in “uninjured” DMD mice? Some insight into SC function during baseline DMD injury would be valuable from this innovative imaging platform.\n5. The SC activation studies in Figure 4 are interesting but highly divorced from the physiological models used throughout the rest of the paper. Is it possible to test the inhibitors on the primary SC/Myofiber culture model and track differentiation index and symmetrical/asymmetrical divisions as in figures 1-3? It would be informative to know if this trend persists when in the physiological context.\nMinor Comments:\n1. L134: Migrated more rapidly than what? DMD SCs? Than previously reported?\n2. L209: I suggest rewriting this section for clarity; I found it a bit hard to read on first pass.", "reviewer_4": "* Overall assessment\nThis manuscript presents a live-imaging approach to analyze muscle stem cell behavior in the mdx model of Duchenne muscular dystrophy. The authors demonstrate impaired migration and skewed division patterns. The study is methodologically advanced and offers new mechanistic insights into MuSC dysfunction. However, improvements in the transparency of tracking methodology, statistical reporting, and data availability are needed.\n* Comments and Suggestions for tracking analysis\n1. Transparency of single-cell tracking methodology\n• The tracking procedure (e.g., via TrackMate) should be described in more detail.\n• Was tracking accuracy validated (e.g., by repeated tracking or inter-observer comparison)?\n• Was the nucleus used as the reference point? How were elongated or morphologically complex cells handled?\n• Was tracking performed in 2D or 3D? If 3D, was it done manually through z-stacks?\n• How were cells selected in densely populated areas (e.g., right part of Movie S1)?\n2. Visualization of tracking data\n• Movies S1 and S2 should include overlays of the tracking data to illustrate dynamics.\n• Fig. 1e: The meaning of different symbols is unclear – a legend is needed.\n3. Quantitative morphology analysis\n• The study would greatly benefit from quantitative assessment of cell morphology (cf.\nhttps://www.nature.com/articles/s41592-024-02580-4).\n• Can morphology and motility be correlated? Rounder cells appear less motile.\n4. Statistical reporting\n• The statistical tests used in Fig. 1g–i and Fig. 2e/g/m–o should be explicitly stated.\n• Fig. 2e: A Kruskal–Wallis test would be appropriate.\n• Fig. 2g: The pronounced differences are surprising given the similar cell cycle lengths—an explanation would be helpful.\n• Fig. 3d: Consider using stacked bars with a unified x-axis, as in the following panels.\n5. Data availability\n• A data availability statement is missing. All raw movies and tracking data should be made publicly available to enable reproducibility and algorithmic benchmarking.\n* General readability\nThe number of abbreviations should be reduced to improve clarity and accessibility, especially for interdisciplinary readers."}, "author_response": "We thank the Reviewers for their careful reading of our manuscript and for their positive comments, detailed suggestions, and appreciation of the novelty of our work. We have revised the manuscript according to the points raised by the Reviewers and our detailed point-by-point response is listed below.\nSummary of major revisions:\n• Text extensively revised throughout (highlighted in blue in manuscript).\n• New inhibitor experiments on isolated myofibres for both WT and mdx mice (Fig. S6).\n• New Laminin-staining analyses clarifying MuSC position relative to the basal lamina (Fig. S2 and S4).\n• New intravital imaging experiments of uninjured WT and mdx muscles (Fig. S2, Movies S1 and S2).\n• PA_qMuSCs vs qMuSCs (PA = Pre-activated) grafting assay added to test the impact of the environment (Fig. S5).\n• Tracking overlays and quantitative morphology analyses introduced (Fig. S2).\n• All new figure panels have been highlighted with light grey background for clarity.\n• All statistical tests explicitly stated and consolidated in Methods.\n• New GitLab repository for code, plugins, and representative datasets (link in manuscript)."}, {"round": 2, "reviewer_comments": {"reviewer_1": "In the revised manuscript, the authors have satisfactorily addressed all the issues and concerns I raised. I have no more issues with the revised manuscript.", "reviewer_2": "The authors have addressed my comments in a satisfactory way. I have no further comments and congratulate them on a beautiful study.", "reviewer_3": "The authors have satisfactorily addressed my main concerns and the manuscript is fit for publication.", "reviewer_4": "* The authors have adequately addressed all my points.\n* In the new movies S5 and S6, the cell tracks are now shown alongside the cells. For the visible cells, these tracks are meaningful. However, the movies also contain numerous tracks without any corresponding visible cell. The authors should clarify where these tracks originate, for example from deeper Z-layers, or use a maximum-intensity projection to make all cells visible."}, "author_response": "We thank the Reviewer for their valuable feedback on our manuscript.\nThe movies S5 and S6 were generated using the TrackMate plugin in ImageJ. However, the only available option for displaying cell tracks on a movie is to select a single Z-plane. Cells from different Z-planes are therefore not visible, and using a maximum-intensity projection to overlay cell tracks is not an available option in TrackMate."}]

Decision letter