Tenascin-C from the tissue microenvironment promotes muscle stem cell maintenance and function through Annexin A2

Abstract

Skeletal muscle regeneration occurs through the finely timed activation of resident muscle stem cells (MuSC). Following injury, MuSC exit quiescence, undergo myogenic commitment, and regenerate the muscle. This process is coordinated by tissue microenvironment cues, however the underlying mechanisms regulating MuSC function are still poorly understood. Here, we demonstrate that the extracellular matrix protein Tenascin-C (TnC) promotes MuSC self-renewal and function. Mice lacking TnC exhibit reduced number of MuSC, and defects in MuSC self-renewal, myogenic commitment, and repair. We show that fibro-adipogenic progenitors are the primary cellular source of TnC during regeneration, and that MuSC respond through the surface receptor Annexin A2. We further demonstrate that TnC declines during aging, leading to impaired MuSC function. Aged MuSC exposed to soluble TnC show a rescued ability to both migrate and self-renew in vitro. Overall, our results highlight the pivotal role of TnC during muscle repair in healthy and aging muscle.

Tenascin-C (TnC) produced by the fibro-adipogenic progenitors (FAPs) is required for MuSC maintenance and function. FAP-secreted TnC signals through Annexin-A2 on the MuSC surface to promote self-renewal and regeneration potential.

Introduction

Muscle stem cells (MuSC) are a muscle-resident stem cell population responsible for developmental and postnatal muscle growth as well as adult tissue repair¹. In healthy adult conditions, MuSC reside in a quiescent state in their sublaminar niche, between the muscle fiber membrane and the basal lamina. In response to damage, MuSC become activated and proliferate to either replenish the stem cell pool (self-renewal) or differentiate and repair the injured tissue (reviewed in ref. ²). Each stage of the MuSC differentiation process occurs in a timely and spatially defined manner to ensure proper muscle regeneration and a full restoration of its function.

The tissue microenvironment is a crucial regulator of muscle maintenance and repair³. The MuSC milieu contains a heterogeneous population of cells, extracellular matrix (ECM) proteins, and growth factors³. Among the tissue resident cells, fibro-adipogenic progenitors (FAPs) play a pivotal role in guiding MuSC-mediated tissue repair. It has been previously shown that FAPs assist the regeneration process by promoting MuSC differentiation^4,5. Muscle-specific ablation of FAPs results in a muscle that is unable to fully regenerate⁶. FAPs secret signaling molecules and deposit ECM components to promote muscle maintenance and repair and provide support to the MuSC niche^7–12. Although recent studies have provided evidence of a major effect of FAP signaling on MuSC in skeletal muscle, we still have a limited understanding of the local cues that mediate the interaction between these two cell types.

The ECM provides structural support for the physical localization of quiescent MuSC, and ECM proteins also mediate signaling through direct interaction with MuSC cell surface receptors¹³. In chronic conditions (e.g., aging, myopathies), changes in ECM composition can impair MuSC self-renewal and regeneration, impacting tissue maintenance¹⁴. We and other groups have previously shown that the ECM glycoprotein Tenascin-C (TnC) is expressed by MuSC during both embryonic development and adult tissue repair^15–18. In skeletal muscle, TnC is widely expressed during embryonic development, while in adulthood its expression is only transitional during regeneration or it is restricted to defined regions, such as tendons and ligaments^18,19. We have previously demonstrated that TnC regulates fetal MuSC development¹⁸. TnC can be produced by myofibers during necroptosis or by glial cells near NMJs upon muscle denervation^20–22, indicating that TnC is produced by multiple cellular sources in the injured muscle microenvironment^18,23. In addition, recent studies in other tissues have shown the requirement of TnC production for cell recruitment to the injury site for efficient regeneration²⁴. Genetic deletion of TnC induces a number of abnormalities, from behavioral (e.g., hyperlocomotion, coordination defects) to developmental (e.g., delayed olfactory detection) as well as defects in stem cell function, wound healing, and muscle atrophy^25–34, strengthening the hypothesis of its role in the skeletal muscle.

TnC accomplishes its different roles via signaling through multiple binding partners in a context-dependent manner. These include cell surface receptors (e.g., Annexin A2, Syndecan-4, EGFR, TLR4, integrins αVβ1/3/6, α2/7/8/9β1), ECM components (e.g., collagen, fibronectin), and soluble factors (e.g., FGF, TGFβ, VEGF, BMP)^35,36. Previous studies have demonstrated that TnC binds with high affinity to Ca²-dependent phospholipid-binding protein Annexin A2 across many cell types and both in mouse and human^37,38, and it has been demonstrated that the TnC:Annexin A2 interaction guides cell migration in diverse tissues^38,39. Interestingly, Annexin A2 is also involved in membrane repair of damaged muscle fibers, which makes Annexin A2 a potential target for ameliorative treatments in patients affected by inflammatory myopathies^40–42. While multiple TnC receptors have been identified in different contexts, the molecular mechanisms by which TnC signals to myogenic cells during muscle repair are still poorly understood.

There is extensive evidence of defective muscle regeneration occurring during aging, due to a disrupted stem cell niche and premature activation of MuSC⁴³. Changes in the ECM composition with age, such as increased stiffness and abnormal signaling from the microenvironment, cause the dysregulation of MuSC quiescence and activation, which leads to an overall diminished tissue regenerative capacity^14,44,45. These age-related ECM changes also correlate with the altered function of other resident cell populations, such as FAPs. During aging, FAPs exhibit reduced proliferation and secretion of signaling molecules, in favor of fibrogenic differentiation^46,47.

Here, we show that muscles lacking TnC exhibited impaired MuSC self-renewal, had reduced numbers of MuSC, and favored an increased number of committed myogenic progenitors. TnC-ablated MuSC also demonstrated impaired migration, which is rescued by soluble TnC treatment. We further provided evidence that FAPs are the predominant source of TnC during regeneration and that MuSC sense TnC through the cell surface receptor Annexin A2. Finally, we observed that young TnC-KO mice recapitulate the impaired regeneration phenotype of old wild-type mice following injury. Our results suggest TnC and its receptors on MuSC are potential candidates for pharmaceutical interventions to enhance muscle repair in aged tissue by restoring the signaling to a youthful state.

Results

Tenascin-C (TnC) regulates the quiescence and commitment of postnatal MuSC

We utilized Tenascin-C knockout (TnC-KO) mice⁴⁸ to evaluate the role of TnC in skeletal muscle. TnC-KO mice did not exhibit defects in skeletal muscle formation during embryogenesis and postnatal maturation. Tibialis anterior (TA) muscles from TnC-KO mice did not have significantly different weight (Fig. 1a) or myofiber cross-sectional area (CSA) (Fig. 1b) compared to wild-type (WT) controls (3–6 months of age). However, we observed a significant reduction in MuSC numbers in TA muscles of adult TnC-KO compared to age-matched WT controls (Fig. 1c). Histological analyses of TA muscles at E16.5, P14, and P30 showed that MuSC numbers are unaffected by TnC deletion at E16.5 or P14, while there was a ~50% reduction at P30, consistent with what we observed in adulthood (Supplementary Fig. 1a). While there was a decrease in TA myofiber CSA at P14, the difference was transient and not detectable either E16.5 or P30 (Supplementary Fig. 1b), further recapitulating what we detected in adulthood (Fig. 1c), suggesting a compensatory response. Furthermore, analysis of the diaphragm muscle at P14 and adult stages showed a reduced number of MuSC in TnC-KO mice (Supplementary Fig. 1c, d). The earlier appearance of MuSC reduction at P14 in the diaphragm compared to the TA could be due to the previously reported higher myonuclear turnover rate in the diaphragm⁴⁹. TnC-KO diaphragms also showed a transient reduction of CSA at P14, which was no longer detectable in adult mice, consistent with the TA phenotype, thus indicating that TnC effects not limited to the TA muscle (Supplementary Fig. 1c, d). Previous work has shown that as muscle reaches homeostasis during postnatal growth, MuSC progressively enter quiescence^18,50–53. Thus, we investigated whether the decline in MuSC numbers in TnC-KO mice was due to an impairment in their ability to regulate quiescence. While TnC-KO MuSC proliferation was not affected at any stage (E16.5, p14, and homeostatic adult muscle), as shown by the percentage of Pax7⁺Ki-67⁺/total Pax7⁺ cells (Supplementary Fig. 1e–g), we observed an increase in the number of MyoD⁺ cells in P14 TnC-KO TA muscles compared to WT controls (Fig. 1d), indicating a bias toward myogenic commitment. Indeed, the percentage of Pax7⁺MyoD⁺ was increased and the percentage Pax7⁺MyoD^- cells decreased in TA muscles at P14 and P30, which was no longer detectable in adulthood (Supplementary Fig. 2a–c). We further observed that P14 TnC-KO muscles exhibited an increase in the percentage of Pax7⁺ MuSC in the interstitial space compared to WT controls, suggesting that TnC plays a role in the proper localization of MuSC in the tissue (Fig. 1e). Interstitial Pax7⁺ cells were largely proliferating, while those in sublaminar position were non-dividing, as shown by the percentage of Pax7⁺Ki-67⁺/total Pax7⁺ cells (Supplementary Fig. 2d), consistent with our previous studies¹⁸. These findings suggest that MuSC in TnC-KO mice fail to properly enter the niche during postnatal muscle growth and undergo premature myogenic commitment at the expense of self-renewal. Recent studies have provided evidence that the MuSC quiescence is characterized by the presence of cellular projections, which are lost during activation^54,55. Thus, we investigated whether TnC-KO MuSC exhibit this feature of activation. Upon isolation of single myofibers from adult extensor digitorum longus (EDL) muscles, both projection length and number per single MuSC were significantly reduced in TnC-KO mice (Fig. 1f). To further investigate whether the premature commitment of MuSC in the absence of TnC leads to premature myogenic differentiation, we isolated MuSC from adult mice (Supplementary Data 1) and induced differentiation for 24 h in vitro. We selected 24 h as timepoint to evaluate the potential accelerated differentiation. Indeed, TnC-KO MuSC exhibited accelerated differentiation, as shown by the increased percentage of myogenin⁺ cells, differentiation index, and fusion index (Fig. 1g). Together, these findings indicate that TnC-KO MuSC exhibit a defect in quiescence in favor of premature myogenic commitment during postnatal muscle growth, leading to a decrease in Pax7⁺ MuSC numbers in TnC-KO adult skeletal muscles.

Fig. 1. TnC regulates MuSC quiescence and myogenic commitment.

a Whole tibialis anterior (TA) muscles from wild-type (WT) and TnC knockout (TnC-KO) adult mice and quantification of their wet weight (mg) (WT n = 3; KO n = 4). Scale bar = 1 mm. b Representative immunofluorescence (IF) images of uninjured TA muscle cross-sections from WT and TnC-KO adult mice (Pax7, green; laminin, gray; DAPI, blue). Pax7⁺ are indicated with yellow arrows. Scale bar = 50 µm. Quantification of the myofiber cross-sectional area (CSA) in µm² (n = 3). c Quantification of the number of Pax7⁺ nuclei (yellow arrows in b) per mm² (WT n = 6; KO n = 5). d Representative IF images of cross-sections from P14 TA muscles of WT and TnC-KO mice (MyoD, red; laminin, gray; DAPI, blue). Scale bar = 50 µm. Quantification of the number of MyoD⁺ nuclei (yellow arrows) per mm² (n = 4). e Representative IF images of cross-sections from uninjured TA muscles of WT and TnC-KO P14 mice for quantification of Pax7⁺ cell localization (Pax7, red; laminin, gray; DAPI, blue). Scale bar = 10 µm. Quantification of Pax7⁺ cell localization under the basal lamina (white arrows) or in the interstitial space (yellow arrows) in WT versus TnC-KO muscles (WT n = 6; KO n = 4). f Representative IF images of WT and TnC-KO freshly isolated myofibers from adult mice (F-actin, green; Pax7, red; α-tubulin, magenta; DAPI, blue). Scale bar = 20 µm. Quantification of the distribution of quiescence projection length, and the number of projections per cell (n = 3). g Representative IF images of WT and TnC-KO differentiated cells from adult mice (Myogenin, red; myosin heavy chain, MHC, gray; DAPI, blue). Scale bar = 50 µm. Quantification of the percentage of myogenin⁺ cells, differentiation index, and fusion index (n = 3). Data are represented as mean ± SEM; *p < 0.05, ***p < 0.001, ****p < 0.0001, t-test (a, b, c, d, e, f); *p < 0.05, Two-way ANOVA (e, f, g). Data are represented as the median with quartiles; **p < 0.01, ****p < 0.0001, t-test (f, g).

TnC is required for adult skeletal muscle regeneration and MuSC self-renewal

TnC is transiently expressed in regenerating TA muscles of adult WT mice following tissue injury induced by barium chloride (BaCl₂) injection¹⁸. TnC begins to be detected at 3 days post injury (DPI) and peaks at 5 DPI, as shown by both whole muscle Western blot and histological analysis (Fig. 2a, b). At 5 DPI, we observed a decreased CSA of regenerating myofibers in TnC-KO muscles compared to WT controls, assessed by embryonic myosin heavy chain (eMyHC) (Fig. 2c), indicating an impairment in tissue repair. Consistent with uninjured tissues, the number of Pax7⁺ MuSC was significantly lower in injured TnC-KO mice than WT controls at the same regeneration stage (Fig. 2d). While MuSC in TnC-KO muscles were able to activate upon injury, the ratio of Pax7⁺ cells in 5 DPI versus uninjured was significantly decreased in TnC-KO compared to WT controls (Supplementary Data 2). We further observed an increased number of MyoD⁺ committed progenitors at 5 DPI (Fig. 2e), indicating increased myogenic commitment, as observed during postnatal growth (Fig. 1d). Indeed, the percentage of Pax7⁺MyoD⁺ cells was significantly increased in TnC-KO mice at 5 DPI compared to WT controls (Fig. 2f), while proliferation remained unaffected (Supplementary Fig. 3a), as observed during postnatal growth (Supplementary Fig. 2a–c). These findings indicate that the accelerated commitment at the expenses of MuSC expansion in TnC-KO mice results in lower number of progenitors able to contribute and fuse to form regenerating myofibers, resulting in smaller eMyHC⁺ myofibers. We next performed serial injury assays to investigate whether the absence of TnC impairs MuSC self-renewal. Injury was induced in TA muscles of both adult TnC-KO and WT mice; one group was analyzed at 30 DPI, while a second group was re-injured at 30 DPI in the same TA muscle and analyzed after an additional 30 days. MuSC numbers were further reduced after the second injury compared to the first injury in TnC-KO mice, demonstrating a defect in self-renewal (Fig. 2g). The CSA was not significantly different in TnC-KO mice between the first and second injury (Supplementary Fig. 3b), which might indicate that several rounds of injury are required to observe a reduction in CSA, as skeletal muscle is a low turnover tissue. The percentage of Pax7⁺MyoD⁺ cells was increased at 30 DPI after the first injury in TnC-KO mice compared to WT, while not detectable at 30 DPI after the second injury (Supplementary Fig. 3c), potentially due to changes in heterogeneity of MuSC compartment following repeated injuries⁵⁶. Overall, these findings indicate that the absence of TnC impairs adult MuSC self-renewal and leads to inefficient skeletal muscle repair.

Fig. 2. TnC is required for MuSC self-renewal and skeletal muscle regeneration.

a Representative Western blot of TnC expression kinetics in uninjured and regenerating whole muscle protein lysates from adult mice and quantification (n = 3). b Representative immunofluorescence (IF) images of cross-sections from uninjured and injured (5 DPI and 14 DPI) TA muscles of WT adult mice (TnC, red; DAPI, blue). Scale bar = 50 µm. c Representative IF images for embryonic myosin heavy chain (eMyHC) of injured TA muscles (5 DPI) in adult WT and TnC-KO mice (eMyHC, green; laminin, gray; DAPI, blue). Scale bar = 50 µm. Quantification of the cross-sectional area (CSA) (n = 4). d Representative IF images of Pax7⁺ cells (yellow arrows) in injured (5 DPI) TA muscles in adult WT and TnC-KO mice (Pax7, green; laminin, gray; DAPI, blue). Scale bar = 50 µm. Quantification of the number of Pax7⁺ nuclei per mm² (n = 4). e Representative IF images of MyoD⁺ cells in injured (5 DPI) TA muscles in adult WT and TnC-KO mice (MyoD, red; laminin, gray; DAPI, blue). Scale bar = 50 µm. Quantification of MyoD⁺ cells per mm² (n = 3). f Representative IF images of Pax7⁺ and MyoD⁺ cells in injured (5 DPI) TA muscles in adult WT and TnC-KO mice (Pax7, red; MyoD, green; laminin, gray; DAPI, blue). Scale bar = 50 µm. Quantification of percentage double positive Pax7⁺ MyoD⁺ (yellow arrows) and Pax7⁺ MyoD^- (white arrows) cells (n = 3). g Representative IF images of Pax7⁺ cells (yellow arrows) in injured (30 DPI post 1st injury) or double-injured (30 DPI post 2nd injury) TA muscles in adult WT and TnC-KO mice (Pax7, green; laminin, red; DAPI, blue) (Scale bar = 50 µm) and quantification per mm² (n = 3). Data are represented as mean ± SEM; *p < 0.05, one-way ANOVA (a); *p < 0.05, **p < 0.01, t-test (c, d, e); *p < 0.05, ***p < 0.001, Two-way ANOVA (f, g).

Fibro-adipogenic progenitors (FAPs) are the major source of TnC during skeletal muscle repair

To assess which specific cell types contribute to TnC deposition in skeletal muscle, we interrogated a publicly available dataset of single-cell RNA-sequencing (scRNAseq) from mouse skeletal muscle at multiple timepoints after injury from Oprescu et al.⁵⁷. At 5 DPI, three muscle-resident cell types were identified expressing TnC: FAPs (Pdgfra⁺, Ly6a⁺, Tnmd^-), activated MuSC (Pax7⁺, Myod1⁺), and tenocytes (Tnmd⁺, Scx⁺) (Fig. 3a), according to Oprescu et al.⁵⁷ cell type classification. MuSC and FAPs exhibited a dynamic expression of the TnC transcript (Fig. 3b, c), while expression of TnC in tenocytes remained high and did not change between uninjured and repairing muscles (Supplementary Fig. 4a). In MuSC, TnC expression was detected at 0.5 DPI then returned to uninjured levels by day 21 (Fig. 3b). In FAPs, TnC expression began at 0.5 DPI to be maintained at higher levels than MuSC and declined by 10 DPI (Fig. 3c). We validated these data by performing qPCR at intermediate timepoints 3 and 7 DPI and indeed observed higher levels of TnC mRNA in FAPs compared to MuSC, but with a decline of expression at an earlier timepoint than MuSC (Fig. 3d, e). To investigate cell–cell communication in the Tenascin signaling network between cell types in skeletal muscle, we utilized CellChat on the Oprescu et al. scRNAseq dataset^57–59. In uninjured skeletal muscle, we identified tenocytes and FAPs as the highest probability senders of Tenascin signaling and MuSC as the dominant receivers of Tenascin signaling in uninjured tissue, along with milder contributions from tenocytes, FAPs, and immune cells (Fig. 3f, h). FAPs were identified as the primary senders of Tenascin signaling at 5 DPI. We also observed a net increase in Tenascin signaling sent from MuSC and FAPs (Fig. 3g, i). During repair, many more cell types are implicated as receivers of Tenascin signals; most notably, this includes immune cells (M1, M2, Monocytes, and Proliferating Immune Cells), MuSC, FAPs, and tenocytes (Fig. 3g, i). At 21 DPI, the sender–receiver dynamics returned to a state comparable with uninjured tissue (Supplementary Fig. 4b, c). Although Tenascin signaling is present in both uninjured and regenerating tissue, the relative contribution of specific ligands at each stage is unique. In uninjured tissue, a large part of the Tenascin signaling occurs through Tenascin-X (Tnxb), a member of the Tenascin family which is constitutively expressed in homeostatic conditions in multiple adult tissues, including skeletal muscle⁶⁰, which shifts toward TnC at 5 DPI before returning primarily to Tnxb at 21 DPI (Supplementary Data 3). Next, we performed transplantation assays to understand TnC cell autonomous versus non-cell autonomous functions in adult muscles. MuSC were isolated by FACS from hindlimb muscles of adult WT mice and infected overnight with a lentivirus expressing red fluorescent protein, RFP. After infection, 2000 RFP⁺ MuSC were transplanted into TA muscles of pre-irradiated adult TnC-KO or WT recipient mice, as previously described¹⁸. At day 21 post-transplantation, we observed significantly smaller numbers of donor-derived RFP⁺ myofibers in TnC-KO host mice compared to WT hosts, indicating the functional relevance of microenvironment-derived TnC during regeneration (Fig. 3j). We next investigated the role of FAP-produced TnC on MuSC behavior, due to the importance of FAPs during skeletal muscle regeneration^4–6. MuSC and FAPs were isolated from adult WT and TnC-KO mice by FACS and co-cultured for 72 h to assess the effect of cell-produced TnC on MuSC identity maintenance in vitro. Immunofluorescence analysis showed higher intensity of the TnC signal in WT FAPs compared to WT MuSC, both in co-cultures and monocultures (Fig. 3k and Supplementary Fig. 4d), indicating that FAPs express higher levels of TnC protein and that its expression is preserved after 72 h in culture. FAP secreted higher levels of TnC compared to MuSC, as shown by ELISA (Supplementary Fig. 4e). We observed a greater number of Pax7⁺ cells when MuSC were co-cultured with WT FAPs, regardless of the MuSC genotype, compared to WT MuSC monocultures and MuSC co-cultured with TnC-KO FAPs (Fig. 3l and Supplementary Fig. 4f). However, there were fewer Pax7⁺ cells when MuSC were co-cultured with TnC-KO FAPs, which was comparable to what observed in WT MuSC monoculture (Fig. 3l and Supplementary Fig. 4f). Proliferation of either MuSC or FAP was not significantly different among the different co-culture groups, as shown by the percentage of Ki-67⁺ cells, indicating that cell division is not impacted in these conditions (Supplementary Fig. 4g). These data indicate that MuSC-FAP interaction or FAP-derived TnC promote the maintenance of Pax7 expression in MuSC in vitro.

Fig. 3. TnC from the tissue microenvironment is required for MuSC function.

a scRNAseq analysis of TnC expression at 5 DPI TA muscles of adult WT mice. Data originally from Oprescu et al.⁵⁷ (n = 3). b, c TnC mRNA expression dynamics in MuSC and FAPs from uninjured and regenerating TA muscles at different DPIs (0.5–21). Data originally from Oprescu et al.⁵⁷ (n = 3). d, e TnC mRNA expression dynamics in MuSC and FAPs in uninjured and at 3 and 7 DPIs by qPCR (n = 3). f, g Chord diagrams representing the number of inter-cluster interactions within the TA muscle in uninjured and 5 DPI. Data originally from Oprescu et al.⁵⁷ (n = 3). h, i Sender–Receiver Probability Heatmap of the Tenascin signaling network in uninjured and 5 DPI TA muscles. Data originally from Oprescu et al.⁵⁷ (n = 3). j Representative immunofluorescence (IF) images representing the contribution of transplanted RFP-labeled WT MuSC from adult 3-month-old donor mice to regenerating TA muscles of WT or TnC-KO age-matched recipient mice (donor-derived fibers – RFP, red; DAPI blue) (Scale bar = 50 µm) and quantification of RFP⁺ myofibers per section (WT n = 4; KO n = 3). k Representative IF images of TnC expression in WT MuSC:WT FAP co-cultures from adult mice (TnC, red; Pax7, green; DAPI, blue). Yellow arrows indicate FAPs. Scale bar = 50 µm. Quantification of TnC fluorescent signal intensity in WT MuSC and WT FAPs normalized on TnC-KO FAPs (n = 3). l Representative IF images of MuSC and FAPs co-cultures from WT and TnC-KO adult mice (Pax7, red; PDGFRα, green; DAPI, blue) (Scale bar = 50 µm). Quantification of Pax7⁺ cells normalized on WT MuSC monoculture (n = 5). Data are represented as mean ± SEM; **p < 0.01, ***p < 0.001, one-way ANOVA (d, e), *p < 0.05, ****p < 0.0001, t-test (j, k), *p < 0.05, **p < 0.01, one-way ANOVA (l).

TnC signals through surface protein Annexin A2 to regulate MuSC function

Although previous studies have described several cell surface receptors mediating TnC signaling, including Annexin A2 (AnxA2), Syndecan-4, TLR4, EGFR, integrin αV/β1, α8/β1, and α9/β1 in multiple cell types³⁶, these interactions are less understood in skeletal muscle regeneration. scRNAseq analysis in MuSC from the Oprescu et al.⁵⁷ dataset for these known TnC receptors, both before injury and during regeneration, revealed that αV/8/9 integrin transcripts were not detected. While Sdc4 progressively decreased during MuSC activation, Tlr4 and Egfr mRNA levels were not notably changed across the repair process (Supplementary Data 4). Quantitative qPCR analysis on TnC receptors validated these findings and confirmed that Anxa2 is the one upregulated transcript upon injury (5 DPI) in MuSC (Supplementary Fig. 5a). We observed that Anxa2 expression transiently increases in MuSC at 3.5 and 5 DPI (Fig. 4a). In contrast, there was no evident change to the expression of Anxa2 in FAPs throughout the muscle regeneration process, except for a minor decrease at 21 DPI (Fig. 4a). These data also show that TnC expression begins earlier and at higher levels in FAPs than MuSC during regeneration (Supplementary Fig. 5b), consistent with our protein analysis (Fig. 3k and Supplementary Fig. 4d, e). In freshly isolated MuSC at 72 h after isolation, >80% of the cells express Annexin A2 (Fig. 4b). Next, we detected the physical interaction of TnC with Annexin A2 from 5 DPI adult whole TA muscles through co-immunoprecipitation assays (Fig. 4c). We further performed proximity ligation assay (PLA) in WT MuSC cultured with recombinant TnC starting at 24 h for additional 48 h and indeed observed a close proximity of the two proteins (Supplementary Fig. 5c). We then analyzed the percentage of Anxa2⁺ or Anxa2^- MuSC (Pax7⁺) or FAPs (Pdgfra⁺) expressing TnC and the relative expression of TnC in these populations throughout regeneration from the Oprescu et al.⁵⁷ dataset. The mRNA expression patterns for each cell type indicate that not only is the average expression of TnC increasing per cell, but also that a greater proportion of Anxa2⁺ cells express TnC during regeneration in both MuSC and FAP populations (Fig. 4d). We also investigated the Anxa2^- fraction of MuSC and FAPs and observed a low number of analyzed cells (raw data of cell numbers) per condition (Supplementary Fig. 5d), and that Anxa2^- MuSC express low levels of TnC during regeneration, while Anxa2^- FAPs follow a similar, but reduced trend in comparison to their Anxa2⁺ counterparts (Fig. 4d). A weaker trend was also observed in committed Myod1⁺Anxa2⁺ MuSC, with reduced raw cell count compared to the corresponding Pax7⁺ populations (Supplementary Fig. 5e). These data suggests that the subpopulation of Anxa2⁺ cells is responsible for driving the majority of TnC signaling in both MuSC and FAPs. To assess the requirement of Annexin A2 to mediate the effects of TnC on MuSC, we performed a knockdown (KD) by infecting freshly isolated mouse MuSC with lentivirus carrying an shRNA for Anxa2. After infection (24 h), MuSC were further cultured in growth conditions either in the presence or absence of recombinant TnC protein for 48 h. The efficiency of KD for Annexin A2 was >75% at 48 h post infection (Supplementary Fig. 5f). We observed that the treatment with TnC promoted an increase in the number of Pax7⁺ cells compared to non-treated GFP-control MuSC, while Annexin A2 KD abrogated the effect of TnC (Fig. 4e). Additionally, the TnC treatment showed a specific effect on Pax7⁺ MuSC, while leaving the numbers of MyoD⁺ myogenic progenitors unaffected in both control and KD in vitro (Supplementary Fig. 5g). Proliferation was not affected in either group in these conditions (Supplementary Fig. 5h). Therefore, these results indicate that Annexin A2 in MuSC is required for TnC signaling.

Fig. 4. TnC signals MuSC through Annexin A2.

a Violin plots depicting the different expression dynamics Anxa2, Pax7, and Pdgfra at different DPIs (0, 3.5, 5, 21) in MuSC and FAPs. Data originally from Oprescu et al.⁵⁷ (n = 3). b Validation by immunofluorescence (IF) of Annexin A2 expression in WT MuSC in vitro (Annexin A2, red; Pax7, green; DAPI, blue) (Scale bar = 50 µm). Quantification of the percentage of Pax7⁺ cells Annexin A2⁺ or Annexin A2^- (n = 3). c Western blot analysis for Annexin A2 on TnC-immunoprecipitated 5 DPI TA muscle protein lysates. d Dot plot representing the changes in the relative expression to the threshold of TnC and the percentages of TnC expressing cell populations within Pax7⁺Anxa2⁺ and Pax7⁺Anxa2^- MuSC and Pdgfra⁺Anxa2⁺ and Pdgfra⁺Anxa2^- FAPs in uninjured TA muscles and across different timepoints post injury (3.5, 5, 21 DPI). Data originally from Oprescu et al.⁵⁷ (n = 3). e Representative IF images of GFP control and Annexin A2 (AnxA2) knockdown (KD) being treated or not with recombinant TnC (48 h treatment) (Pax7, red; GFP, green; DAPI, blue) (scale bar = 50 µm) and quantification of Pax7⁺ cells normalized on non-treated GFP-control MuSC (n = 3). Data are represented as mean ± SEM; ****p < 0.0001, one-way ANOVA (e).

TnC promotes MuSC migration

The ability of MuSC to migrate upon injury is an essential feature to reach the damaged area and initiate regeneration. TnC has been previously shown to promote migration in multiple cell types including fibroblasts, endothelial and smooth muscle cells, astrocytes, and in metastatic cancer^61–63. Thus, we asked whether TnC regulates MuSC migration by performing time-lapse microscopy on freshly isolated MuSC from TnC-KO and WT mice cultured in growth conditions for 24 h. Both the total traveled distance and the average velocity of the cells were significantly reduced in TnC-KO MuSC compared to WT controls (Fig. 5a–c). We implemented transwell assays to further assess MuSC migration capacity and rule out the observed results were due to a specific migration assay; we observed that the lack of TnC reduced MuSC migration after 48 h in culture (Fig. 5d). We confirmed that the increased number of cells migrated through the transwell was indeed due to migration and not to differences in MuSC proliferation (Fig. 5e). Next, we administered soluble recombinant TnC to WT and TnC-KO MuSC, which further promoted migration compared to controls (Fig. 5f). Consistent with in vivo data, MuSC proliferation was not altered by the presence of recombinant TnC (Fig. 5f). Together, these findings indicate that the deletion of TnC limits MuSC migration in vitro.

Fig. 5. TnC promotes MuSC migration in vitro.

a Representative images of time-lapse microscopy to track cultured MuSC from adult WT and TnC-KO mice. Scale bar = 50 µm. b Quantification of total distance covered (n = 3, N_cell ≥ 75). c Quantification of average velocity of cells (n = 3, N_cell ≥ 75). d Quantification of migrated TnC-KO cells through transwell matrix normalized to WT (n = 6). e Quantification of the percentage of proliferating cells by EdU staining (timepoint = 48 h) (n = 3). f Representative immunofluorescence (IF) images of migrated WT and TnC-KO MuSC from adult mice with or without recombinant TnC treatment (EdU, gray; DAPI, blue) (scale bar = 50 µm). Quantification of the number of migrated WT and TnC-KO cells through transwell matrix after vehicle or TnC treatment (per mm²) (timepoint = 48 h) and quantification of the percentage of proliferating cells through EdU staining (timepoint = 48 h) (n = 3). Data are represented as mean ± SEM; *p < 0.05, t-test (b, c, d), *p < 0.05, **p < 0.01, one-way ANOVA (f).

TnC-KO mice exhibit a premature aging phenotype in skeletal muscle

In light of the role of TnC during muscle regeneration and the widely studied impairment of regeneration in aged skeletal muscle, we asked whether TnC expression changes during skeletal muscle aging. We observed decreased levels of TnC both via Western blot (Fig. 6a) and immunofluorescence in aged regenerating muscles compared to young regenerating controls (Fig. 6b). We further showed that MuSC numbers were reduced in aged muscles compared to young controls (Supplementary Fig. 6a, b), consistent with previous literature^64–66. TnC deletion significantly reduced the wet weight of uninjured TA muscles from aged TnC-KO mice compared to both young and aged WT TA controls (Supplementary Fig. 6c). We further observed reduced myofiber CSA in uninjured aged TnC-KO mice compared to WT young, TnC-KO young, and WT aged mice (Fig. 6c). A comparable reduction in CSA was also observed in the diaphragm (Supplementary Fig. 6d), indicating that the observed phenotype is not limited to hindlimb muscles. Fibrosis accumulation was <2% muscle section area, and a slight reduction was observed in aged TnC-KO TA muscle (Supplementary Fig. 6e), consistent with the previously reported role of TnC in tissue fibrosis⁶⁷, and did not observe accumulation of adipose tissue across genotypes and ages (Supplementary Fig. 6f). We also detected a significantly decrease in the number of Pax7⁺ cells in aged TnC-KO mice compared to the age-matched WT counterparts (Fig. 6d). Following tissue injury, aged TnC-KO muscles undergoing regeneration showed a severe decline in the percentage of eMyHC⁺ area and regenerating myofiber CSA compared to controls (Fig. 6e and Supplementary Fig. 6g). We then used MuSC migration as a readout to further investigate this regeneration phenotype in aged muscle; we observed that aged MuSC exhibited a defect in cell migration compared to young controls (Fig. 6f). When aged MuSC were exposed to TnC in the transwell assay, the treatment rescued their migratory defect (Fig. 6g). Cell proliferation was not affected in aged MuSC exposed or not to recombinant TnC in transwell assays, as shown by the percentage of EdU⁺ cells (Supplementary Fig. 6h). Finally, while we observed a decrease in the expression of Annexin A2 protein in aged MuSC compared to their young counterparts (Supplementary Fig. 6i), exposure to soluble TnC induced an increase in the number of Pax7⁺ cells in aged MuSC, indicating a rescue of the phenotype, while the number of MyoD⁺ cells was not affected (Fig. 6h and Supplementary Fig. 6j). Together, these data demonstrate that the lack of TnC in aged mice further impairs the regeneration phenotype, and that the impaired self-renewal of Pax7⁺ population and reduced migration observed in aged muscle are rescued by treating aged MuSC with exogenous TnC.

Fig. 6. TnC declines with aging and is required for muscle maintenance and regeneration of aged skeletal muscles.

a Western blot for TnC on young and old WT mice and quantification of TnC protein relative expression (n = 4). b Representative immunofluorescence (IF) images of 5 DPI TA muscles of young and old mice (TnC, red; laminin, gray; DAPI, blue) (n = 3). Scale bar = 50 µm. c Representative IF images of cross-sections of uninjured TA muscles from young and old WT and TnC-KO mice (laminin, green; DAPI, blue) (scale bar = 50 µm) and quantification of the cross-sectional area (CSA) in µm² (n = 3). d Representative IF images of cross-sections of uninjured TA muscles from old WT and TnC-KO mice (Pax7, green; laminin, red; DAPI, blue) (scale bar = 50 µm) for Pax7⁺ cell quantification. Pax7⁺ cells are indicated with yellow arrows (n = 3). e Representative IF images of cross-sections of injured (5 DPI) TA muscles from young and old WT and TnC-KO mice (laminin, green; eMyHC, red; DAPI, blue) (scale bar = 50 µm) and quantification of the percentage of regenerating area (eMyHC⁺) (n = 3). f Quantification of migrated aged WT MuSC through transwell matrix (48 h culture) normalized to WT young mice (WT young n = 6; WT old n = 4). g Representative IF images of migrated aged WT MuSC with or without recombinant TnC treatment (timepoint= 48 h) (DAPI, gray) (scale bar = 50 µm) and quantification of migrated aged WT MuSC through transwell matrix after TnC treatment normalized to control non-treated cells (n = 4). h Representative IF images of Pax7⁺ young and old WT MuSC cells (yellow arrows) treated with or without recombinant TnC (timepoint = 48 h) (Pax7, red; DAPI, blue) (scale bar = 50 µm) and quantification of Pax7⁺ cells after TnC treatment normalized on control non-treated WT young cells (n = 3). Data are represented as mean ± SEM; *p < 0.05, **p < 0.01, t-test (a, d, f, g), *p < 0.05, **p < 0.005, ***p < 0.001, ****p < 0.0001, one-way ANOVA (c, e, h).

In conclusion, lack of TnC causes a skeletal muscle phenotype characterized by fewer MuSC, impaired migration, and a decreased ability to repair tissues. We also showed that exposing TnC-KO MuSC to soluble recombinant TnC protein rescued their migratory capabilities. We demonstrated that both TnC-KO and WT MuSC are maintained in their Pax7⁺ state in the presence of WT FAPs in vitro, which are the primary source of TnC. Mechanistically, MuSC sense TnC signaling through Annexin A2. Upon aging, the TnC-KO mouse exhibits an exacerbated phenotype of MuSC at both steady state and during regeneration, and exposure of aged MuSC to soluble TnC rescues both their migratory ability as well as their maintenance in a Pax7-positive state.

Discussion

During mouse postnatal muscle growth, MuSC enter quiescence around one month of age^18,50–53. Our findings show that MuSC numbers are not affected during prenatal development in TnC-KO mice, but that TnC deletion reduced MuSC numbers during early postnatal muscle growth. We also provide evidence that during postnatal growth, TnC-KO mice have increased localization of MuSC outside of the sublaminar niche and in the interstitial space, and are actively dividing, consistent with our previous report¹⁸. In addition, in TnC-KO mice, we observed a higher number of Pax7⁺MyoD⁺ committed progenitors compared to WT controls, indicating a role of TnC in the postnatal development of skeletal muscle. MuSC lacking TnC did not exhibit a defect in cell proliferation in any of the conditions tested. A previous study reported that TnC promotes MuSC proliferation²⁰. The discrepancy among the findings could be due to different experimental settings, including the use of C2C12 necroptosis-inducible media, which might exhibit different properties than MuSC, and the use of a truncated version of Tenascin-C, lacking the C-terminus portion of the protein, which has been previously shown to interact with multiple cell surface receptors³⁵. Our findings suggest that during postnatal growth, TnC-KO MuSC are impaired in entering the sublaminar niche, fail to enter quiescence, and undergo premature myogenic commitment. Recent studies using intravital imaging have demonstrated that a morphological feature of quiescent MuSC is the presence of long cellular projections^68–70. Upon tissue injury, rearrangement of cytoskeletal components occurs, and the retraction of cellular projections is observed upon MuSC activation. This is mediated by mechano-sensing pathways triggered by a Rac-to-Rho switch and Piezo-1 signaling^54,55. Our data demonstrate that TnC-KO MuSC exhibit both reduced length and number of cellular projections, which further supports spontaneous activation. This phenotype is also associated with a defect in MuSC self-renewal in TnC-KO mice, as we demonstrated in serial injury assays. Our previous work has shown that fetal MuSC, which expresses TnC, undergoes both asymmetric division and symmetric expansion in culture, compared to adult MuSC, which mainly sustains symmetric depletion events¹⁸. We propose that the absence of TnC further biases MuSC toward symmetric depletion events, resulting in defective self-renewal, maintenance, and re-entry into quiescence following injury. We further show that MuSC isolated from TnC-KO mice exhibit impaired migration. We demonstrated that the administration of soluble TnC to both WT and TnC-KO MuSC enhanced their migration. This is consistent with the literature describing the role of TnC in regulating adhesion and promoting migration in multiple tissues and cell types, through inhibition of focal adhesion formation or engaging the Hippo pathway to promote regeneration^39,71–74. Whether the impaired migration in the absence of TnC is a cause or consequence of an activated MuSC phenotype is currently unclear, and future studies will shed light on this functional interdependence.

We demonstrate that TnC signaling in MuSC occurs through binding to Annexin A2. TnC has been previously reported to bind several cell surface receptors³⁶. Our analysis of the skeletal muscle scRNAseq dataset from Oprescu et al.⁵⁷ showed that the transcripts for Syndecan-4, EGFR, TLR4, and integrins are either not detected or do not significantly increase in MuSC during regeneration. Although we cannot rule out the role of these surface receptors in skeletal muscle repair, we focused on Annexin A2, due to its early-stage mRNA upregulation consistent with TnC expression patterns.

The role of Annexin A2 has been previously studied in the context of myofiber membrane repair^40,75. Annexins are a widely expressed family of Ca²⁺- and phospholipid-binding proteins that contribute to many cell functions at the plasma membrane, including membrane organization and trafficking, transmembrane channel activity, cell-ECM communication, and often act as a bridge for membrane-cytoskeleton interactions⁷⁶. In skeletal muscle, Annexin A2 trafficking to the injured site⁷⁷ and its interaction with proteins associated with sarcolemma repair, such as dysferlin^78–80, are required processes for efficient regeneration. Indeed, myofibers lacking Annexin A2 demonstrated defective repair upon muscle injury⁷⁹. Annexin A2 is upregulated in intestinal epithelial cells during migration and promotes cell spreading and positively regulates Rho activity⁸¹. Upon interaction with TnC, Annexin A2 promotes the loss of focal adhesions and cell migration in multiple cell types^39,82.

Our bioinformatics analysis shows transient upregulation of Anxa2 during repair. In culture, >80% of MuSC express Annexin A2 protein, which we showed through proximity ligation assay to be in close proximity with TnC in MuSC. Furthermore, we observed that TnC-KO MuSC exhibited defects in activation and migration, which is in line with previous evidence that Annexin A2 mediates actin rearrangements through Rho activation⁸³. We propose that the transient increase of MuSC-specific Annexin A2 expression during tissue repair promotes TnC-mediated MuSC activation and migration to the damaged area during the early stages of regeneration. This may occur through a feedback loop involving the reciprocal regulation of TnC:Annexin A2 interaction and the Rho/ROCK axis. Our expression analysis has highlighted that not all MuSC exhibit the same level of expression of Annexin A2, and it will be relevant in future studies to address whether these differences underlie distinct cell subpopulations or transitional cellular states.

We have previously shown that TnC is expressed by MuSC at fetal stages as well as in adult MuSC during tissue repair¹⁸. Downregulation of TnC in fetal MuSC, though not in adult MuSC, impairs MuSC contribution to skeletal muscle repair upon transplantation. This different behavior indicates that multiple cell types in the adult tissue microenvironment produce TnC for efficient regeneration^18,23. Our transplantation assays demonstrate that when WT adult MuSC are transplanted into TnC-KO mice, their contribution to tissue repair is impaired, indicating that TnC from the tissue microenvironment is required for MuSC function. Our analyses of scRNAseq from a publicly available dataset from Oprescu et al.⁵⁷ and qPCR data demonstrate that the major sources of TnC in skeletal muscle during tissue repair are FAPs, tenocytes, and MuSC. FAPs dynamically increase expression of TnC in the initial phases of tissue repair. A previous study has reported that myofibers during acute injury express TnC²⁰. However, analysis of publicly available scRNAseq datasets during muscle regeneration from Oprescu et al.⁵⁷ and Walter et al.⁸⁴ (Fig. 3a and Supplementary Data 3b) did not show any TnC expression in myonuclei during skeletal muscle regeneration (consistent with our previous observation that TnC is expressed in undifferentiated muscle cells and declines upon myogenic differentiation in culture¹⁸), while its expression in FAPs and MuSC was clearly detectable. While we cannot rule out the possibility of low TnC production in myofibers under specific conditions, our analyses indicate that FAP express higher levels of TnC compared to MuSC, both at the mRNA and protein level, and they secrete it in the extracellular space. Our co-culture experiments demonstrate that TnC released from FAPs is required for the maintenance of the MuSC pool. While it has been previously shown that co-cultivating FAPs and MuSC isolated from uninjured mouse muscle for 7-10 days increases the terminal differentiation of myogenic progenitors^4,5,85,86, our results indicate that upregulation of TnC in FAPs occurs specifically at early timepoints, suggesting that FAPs might promote different processes at different stages of tissue repair. We speculate that there may be an increase in autocrine MuSC TnC signaling during muscle repair, but that the signaling from the FAPs is the primary driver of MuSC self-renewal in this context. Our bioinformatic analysis of scRNAseq data further shows that 50–75% of FAPs upregulate TnC in response to injury, demonstrating the heterogeneity of this cell population, either through distinct cell subpopulations or transitional cellular states. TnC gene transcription has been shown to be induced by mechanical stress through α5β1 integrin, ILK, and Rho/ROCK signaling, inducing nuclear translocation of MLK1 (MRTF-A)^87,88. We suggest, to the best of our knowledge, a novel role for FAPs in regeneration through the secretion of TnC, where initially FAP-derived TnC is required to regulate the number of MuSC following injury, which is then followed by the known role of FAPs to induce differentiation of activated MuSC.

Our results show that TnC expression during tissue repair is reduced during aging. We further show that mice lacking TnC exhibit a premature aging phenotype, with a reduced number of MuSC and impaired MuSC self-renewal and overall tissue regeneration. Analysis of TnC-KO muscles upon injury shows that the defect is further exacerbated during aging, with a severe reduction in the number of MuSC and impaired tissue repair. Previous studies have reported an increased abundance of FAPs in aged muscles, while other studies show a decrease in the number of FAPs in favor of differentiation toward a fibrotic state^46,47,89. Thus, the reduced levels of TnC in aged WT muscles could be the result of either changes in TnC expression or changes in FAP abundance. Finally, we demonstrate that exposing aged MuSC to recombinant TnC rescues both the number of Pax7⁺ MuSC as well as their migratory defect.

Overall, our findings indicate that FAP-derived TnC is a major regulator of MuSC self-renewal and migration through Annexin A2, and that its reduced levels during aging contribute to the impaired MuSC function in aged muscles. Identification of this molecular mechanism is relevant for therapeutic purposes as it can be utilized as a tool to enhance MuSC self-renewal and function and improve tissue repair in aged tissues.

Methods

Contact for reagent and resource sharing

Further information and requests for resources and reagents should be directed to and will be fulfilled by the Lead Contact, Alessandra Sacco, Ph.D. (asacco@sbpdiscovery.org).

Experimental model and subject detail

Experimental Animals

Tenascin-C knockout (TnC-KO) mice were a kind gift from Drs. Faessler and Orend. All protocols were approved by the Sanford Burnham Prebys Medical Discovery Institute Animal Care and Use Committee (protocol numbers: AUF#13-007; AUF #24-088; AUF #24-092). Mice were housed according to institutional guidelines, in a controlled environment at a temperature of 22 °C ± 1 °C, under a 12-h dark-light period, and provided with standard chow diet and water ad libitum. Male and female TnC-KO and wild-type mice at ages E16.5, P14, P30, adult age (3–6 months old), and geriatric (18–29 months old) were used. All mice were maintained on an FVB background.

Method details

Cells Isolation

Muscle stem cells (MuSC) were isolated as described in Gromova et al. with minor revisions⁹⁰. Whole hindlimb (tibialis anterior, extensor digitorum longus, gastrocnemius, soleus, plantaris, vastus lateralis, quadriceps, and adductors) were minced and sequentially incubated in 800 units/mL collagenase type II solution (catalog number: 17101-015, Life technologies, Gibco®) and subsequent incubation with 80 units/mL collagenase II and 2 units/mL dispase II (catalog number: 04942078001, Roche) solution. Muscle tissues were then passed through a 10 mL syringe with a 20G needle and a 40 μm nylon filter. Primary and secondary antibodies incubation was performed in a 300 µl volume. Biotin-labeled lineage negative cells (CD45⁺, CD11b⁺, CD31⁺, Sca1⁺ cells) were either depleted using streptavidin beads (catalog number: 130-048-101, Miltenyi Biotec) through a magnetic field or excluded during sorting by using streptavidin-APC-Cy7. MuSC were isolated with BD Biosciences FACSAria II cell sorter as CD45^-, CD11b^-, CD31^-, Sca1^-, CD34⁺, and integrin α-7⁺ population. For the co-culture experiments, Fibro/adipogenic progenitors FAPs (Sca1⁺ and CD34⁺ population) were also isolated. The gating strategy can be found in the Supplementary Data 1.

MuSC culture

All cells were cultured in incubators at 37 °C and 5% CO₂. After isolation MuSC were plated on laminin (catalog number: 11243217001, Roche, 1:25 dilution in PBS) in growth medium (catalog number: 10313-21, Gibco), 15% FBS (catalog number: FB-11, Omega Scientific), 1% Pen/Strep (catalog number: 15140163, Life technologies, Gibco®), 2.5 μg/mL FGFb (catalog number: 100-18B, Peprotech). For the knockdown experiments, cells were seeded at 3000 cells/well and cultured for 72 h before fixation, as described in the paragraph below. In each condition, cells were further processed for immunostaining analysis.

Lentiviral infection and recombinant TnC treatment

Lentiviral particles were purchased from the MISSION shRNA library (Sigma). Tissue culture plates (96-well) were coated with laminin (20 mg/mL) for 1 h at 37 °C, followed by RetroNectin® (catalog number: T100A, Takara) (20 mg/mL in sterile PBS) for 2 h at 37 °C. Upon plating (cell density 3000 cells/well), MuSC were treated with cationic polymer (hexadimethrine bromide, catalog number: 50-187-2421, Fisher) (8 µg/mL) at 37 °C for 5 min immediately followed by infection with 100 MOI of either custom control lentiviral GFP or lentiviral GFP sh-Anxa2 (trcn0000110696) in growth medium for 4 h at 37 °C and 5% CO₂. At 4 h, the media containing the lentiviral particles were removed and replaced with growth medium. At 24 h from the infection, half of the control and sh-Anxa2 samples were incubated with recombinant TnC (catalog number: 3358TC050, R&D) (5 µg/mL) or vehicle (PBS) for 48 h in growth media at 37 °C and 5% CO₂. At the end of the experiment, samples were processed for: 1. mRNA extraction and qPCR to assess the knockdown (KD) efficiency; 2. Immunofluorescence. Pax7⁺ nuclei in GFP⁺ cells were counted and then normalized by the number of Pax7⁺ nuclei in control, non-treated cells.

Co-culture assay of MuSC and FAPs

After FACS isolation, cells were seeded on laminin-coated 96-well plates at 2000 cells/well per cell type (4000 cells/well total) for co-culture. Cells were cultured for 72 h and then fixed with 4% PFA for 10 min at room temperature. The cell samples were then either stored at 4 °C or prepared for immunofluorescence. The percentage of Pax7⁺ cells in each co-culture combination was obtained by dividing the number of Pax7⁺ nuclei by the number of total nuclei minus the number of Pdgfra⁺ cells. Control monocultures were used as a control for baseline expression of Pax7. All co-culture data was normalized on non-treated, monocultured WT MuSC. A minimum of 8 fields of view acquired with a 20× objective were analyzed.

Skeletal muscle injury

Muscle injury was performed in the tibialis anterior (TA)⁹¹. Mice were anesthetized with isofluorane and injected with 50 µl barium chloride (catalog number: 202738, Sigma) resuspended in PBS (to 1.2% w/v) in the proximal, medial, and distal locations of the TA to distribute the solution to the entire muscle. MuSC and FAPs were isolated at 3 and 7 days after performing the injury for qPCR, or TA muscles were harvested at different timepoints and included in OCT (catalog number: 4583, VWR) for sectioning and immunohistochemistry.

In vivo EdU treatment

Mice were treated with EdU to assess cell proliferation. With a 29G needle insulin syringe, mice were intraperitoneally administered with 200 μl of EdU (5 mg/mL). Muscle groups were then harvested 24 h after injection.

Myofiber isolation for quiescent projection length quantification

Myofibers from EDL muscles of TnC-KO and WT adult mice were isolated following the protocol described by Kann et al.⁵⁴, with minor adaptations. Mice were euthanized following the guidelines of IACUC-approved protocols. EDL muscles were harvested (by first severing the proximal tendon) and subjected to enzymatic dissociation (collagenase type II, catalog number: LS004177, Worthington) (700 units/mL) in low glucose (1 mg/mL) DMEM at 37 °C for 55 min. Dissociated single myofibers were manually collected and purified under a dissection microscope, then placed in the incubator for 10 min. Myofibers were then fixed in 2% PFA at room temperature for 10 min, subsequently washed twice in PBS, and stored in PBS at 4 °C. All steps were performed in the presence of Y-27632 Rock inhibitor (catalog number: 72304, Stem Cell Technologies) (final concentration 10 µM) as described by Kann et al.⁵⁴.

Immunofluorescence

Muscle tissues (TA muscles) were isolated from healthy and injured mice at different timepoints. Tissues were embedded in OCT and frozen in 2-methyl butane. Tissues were sectioned into 10-μm-thick slices and further processed by immunostaining. Fixation was performed with 4% PFA (catalog number: sc-281692, Santa Cruz Biotechnology). Tissue sections were washed in PBS twice, then permeabilized with 0.5% Triton X-100 (catalog number: 1003477329, EMD Millipore) in PBS and blocked in 10% goat serum (catalog number: 16210-072, Life Technologies, Gibco®) and 0.1% Triton X-100 in PBS at room temperature for 1 h. For Pax7 staining, samples were incubated with AffiniPure Fab fragment goat anti-mouse IgG (1:40, catalog number: 115-007-003, Jackson ImmunoResearch) solution in 0.2 µm filtered PBS at room temperature for 30 min, then washed with PBS, and incubated in antigen retrieval (1:100, antigen unmasking solution, citric acid based, catalog number: H-3300, Vector Laboratories) solution at 92 °C for 10 min. Incubation with the primary antibodies was conducted at room temperature for 1 h or at 4 °C overnight in blocking buffer. All washes after incubation with antibodies were done using PBS with 0.5% Triton X-100.

For Pax7/MyoD co-staining (respectively using catalog number: Pax7-c, Developmental Studies Hybridoma Bank (DSHB), 1:10; and M-318, catalog number: sc-760, SCBT, 1:50) in tissue sections, fixation was performed with 2% PFA for 20 min at room temperature, permeabilization with −20 °C methanol for 6 min at room temperature, blocking and antibody dilutions in 5% BSA in PBS 1 ×, while washes were performed in 0.1% BSA in PBS 1 ×.

Myofibers were blocked in 10% goat serum with 0.3% Triton X-100 buffer at room temperature for 1 h, followed by incubation with primary antibody blocking solution overnight at 4 °C. Isolated cells were fixed in 4% PFA, washed in PBS twice, permeabilized at room temperature for 8 min with 0.5% Triton X-100, and incubated in 4% BSA (catalog number: SH30574.02, HyClone) with 0.5% Triton X-100 blocking buffer at room temperature for 1 h or at 4 °C overnight. The primary antibodies used are the following: mouse anti-Pax7 (catalog number: Pax7-c, Developmental Studies Hybridoma Bank (DSHB), 1:50 dilution for cultured cells, 1:10 for tissue sections and myofibers), mouse anti-myogenin (catalog number: 556358, BD Biosciences; 1:100 dilution), mouse anti-MHC (catalog number: Mf20-c, DSHB,1:50 dilution), mouse anti-myosin (embryonic) (catalog number: F1.652, DSHB, 1:100 dilution), rabbit anti-tenascin-C (catalog number: AB19013, MilliporeSigma), mouse anti-Annexin A2 (catalog number: 3D5 sc-47696, SCBT, 1:50), rabbit anti-Annexin A2 (catalog number: 8235S, Cell Signaling Technology), rabbit anti-α-tubulin (catalog number: ab18251, Abcam), rabbit anti-Pdgfra (catalog number: 3174T, Cell Signaling), rabbit anti-MyoD (catalog number: sc-760, Santa Cruz, 1:100 dilution), mouse anti-MyoD (catalog number: sc-377460, Santa Cruz, 1:100), rabbit anti-laminin (catalog number: L9393, Sigma, 1:100 dilution), rat anti-laminin (catalog number: 05-206, Millipore, 1:100 dilution), chicken anti-GFP (catalog number: QAB10251, inquire Bio, 1:200 dilution). Alexa-conjugated secondary antibodies (Invitrogen, 1:500 dilution) were diluted in appropriate blocking buffer depending on the type of sample and incubated at room temperature for 45–60 min. Nuclear DNA was stained with DAPI (Catalog number: MBD0011, Sigma). F-actin was labeled by using Alexa Fluor^TM 488 Phalloidin (catalog number: A12379, Life Technologies). Tissue sections and myofibers were prepared for imaging in Fluoromount-G® (catalog number: 0100-01, SouthernBiotech) mounting solution.

Images were acquired with an Inverted IX81 Olympus Compound Fluorescence Microscope, XYZ Automated stage - ASI 2000 (Applied Scientific Instrumentation Inc.), with a Color/monochrome cooled CCD camera - Spot RT3 and MetaMorph 7.11 Software (UIC, Molecular Devices) at 10× or 20× magnification or using confocal scanning through Leica TCS SP8 and LAS X software at 20× or 63× magnification. The Leica DMi8 epifluorescent microscope was used for cell culture and tissue section imaging; a minimum of 8 fields of view acquired with a 20× objective were analyzed. Nikon A1R HD confocal (running Nikon Elements software Version 5.42.04) with oil immersion 63× objective was used to acquire images of Pax7⁺ cells on myofibers. All images were composed, edited, and modifications applied to the whole image using Adobe Photoshop 2025.

Migration assays

MuSC were isolated from WT young and old mice as well as TnC-KO young mice. 3000 cells were plated in a Costar™ HTS Transwell™-96 (porosity 8 µm) (catalog number: 07-200-657, Fisher Scientific) and maintained in growth conditions for 48 h. Membranes were pre-coated with either recombinant TnC or PBS, and the bottom of the well with laminin (as described in the “MuSC culture” section) prior to MuSC culture. Cells were then fixed with 4% PFA in PBS 1× at 48 h after plating. Images were obtained and analyzed in an automated manner by using the Cytation 5 Cell Imaging Multimode Reader (Agilent). For time-lapse microscopy, images were acquired every 20 min to monitor the cell total covered distance (µm), average distance (µm per 20 min), and velocity (µm/20 min) over a 26-h period before fixation.

Proximity ligation assay (PLA)

PLA was performed using MilliporeSigma Duolink® PLA as advised by the manufacturer. MuSC present physiologically low expression of TnC; thus, to maximize the presence of TnC on the cells, we treated MuSC (24 h after FACS isolation) with recombinant TnC (catalog number: 3358TC050, R&D) (5 µg/mL) or with the vehicle (PBS) for 48 h in growth media at 37 °C and 5% CO₂. Cells were then fixed in 4% PFA as described in the “Immunofluorescence” section. Permeabilization was performed in 0.2% Triton X-100 in PBS 1× for 8 min at room temperature. Cells were then washed 3 times with PBS 1× before blocking with Duolink® blocking solution 45 min at 37 °C and then 15 min at room temperature. Anti-Annexin A2 mouse antibody (catalog number: 3D5 sc-47696, SCBT, 1:50) and anti-Tenascin-C rabbit antibody (catalog number: AB19013, MilliporeSigma, 1:50) were diluted in Duolink® antibody diluent as the experimental condition. Controls for the PLA were: (1) Cells treated with vehicle incubated with both primary antibodies, (2) TnC-treated cells incubated with anti-Annexin A2 antibody only, (3) TnC-treated cells incubated with anti-Tenascin-C antibody only, (4) TnC-treated cells incubated with no antibody. All samples were incubated overnight at 4 °C. The following day, cells were washed twice with Buffer A (10 mM Tris, 150 mM NaCl, 0.05% Tween-20, pH 7.4) 5 min at room temperature. Duolink® PLUS and MINUS PLA probes (respectively, catalog numbers: DUO92002 and DUO92004, MilliporeSigma) were diluted in the Duolink® antibody diluent as recommended by the manufacturer and incubated 60 min at 37 °C. After two 5 min washes with Buffer A at room temperature, the ligase was diluted in Ligation Buffer (both contained in the Duolink® In Situ Detection Reagents Green catalog number: DUO92014, MilliporeSigma) as indicated by the manufacturer. Ligase was immediately applied to cells and incubated for 30 min at 37 °C. For amplification, polymerase was diluted in appropriate buffer (both contained in the Duolink® In Situ Detection Reagents Green catalog number: DUO92014, MilliporeSigma), then applied to the cells, after two 5 min washes with Buffer A at room temperature, and then incubated for 100 min at 37 °C in a humidified chamber and protected from light. The samples were washed twice with Buffer B 10 mM Tris, 200 mM NaCl, pH 7.4 (no detergent) for 10 min each at room temperature, followed by one wash with 0.01× Buffer B for 1 min at room temperature. Finally, DAPI 1:1000 was added for 5 min at room temperature, followed by 2 washes with PBS 1×. Samples were immediately imaged with Nikon A1R HD confocal (running Nikon Elements software Version 5.42.04) with a water immersion 60× objective.

Histology: Sirius red and oil red O staining of muscle sections

For Sirius red staining, tissue sections were re-hydrated for 2 min in distilled water and then incubated with Sirius red solution in picric acid for 1 h. A 10 min wash was done with distilled water and subsequently with an acetic acid-based solution in distilled water for 2 min. Three washes with 100% Ethanol were performed, each for 3 min. The ethanol was removed before adding xylene for 3 min. After removing the xylene sections were mounted with mounting solution. All steps were performed at room temperature.

For Oil Red O staining, fresh tissue sections were fixed in 10% formalin for 2–5 min, and then immediately rinsed with tap water for 2–3 min, followed by two changes of distilled water. Sections were then placed in absolute propylene glycol for 2 min to ensure no water was carried into the Oil Red O solution, which was subsequently performed at room temperature for 1 h. Sections were then incubated in 85% propylene glycol solution for 1 min to remove excess dye and then rinsed twice in distilled water. Hematoxylin counterstaining of the nuclei was then performed in Modified Mayer’s Hematoxylin for 30 s. Final washed with two changes of tap water and two changes of distilled water were done before mounting using Glycerin Jelly Mounting Medium.

Sirius red and Oil Red O staining visualization was obtained with the Aperio AT2 Leica slide scanning system. All images were edited and modifications applied to the whole image through Fiji or Photoshop CS4 and Photoshop 2024 (Adobe).

Quantification of muscle tissue cross-sectional area (CSA)

CSA quantification was performed in an automated manner using a Macro through ImageJ64⁹² or the Muscle Morphometry ImageJ plugin developed by Anthony Sinadinos (https://drive.google.com/drive/folders/0B_bBI7SbDQhCR1MxNEVXSlhiekE?resourcekey=0-8wdIKyTc0OKlB7WN67JqIw) by using the laminin fluorescent signal channel. CSA and the number of regenerating myofibers at 5 DPI in the experiments involving old mice were calculated by using the embryonic myosin heavy-chain (eMyHC) fluorescent signal. The area of each myofiber and their number in each field of view were obtained by converting the images into binary, then followed by the command “Analyze particles” limited to the set threshold value.

Quantification of protein immunofluorescence

After fixation and immunostaining, cells were imaged using the fixed settings (e.g., exposure time, gain, laser power, averaging, and dwelling) across different cell types and culture conditions. The maximum of the fluorescent intensity signal was quantified on the raw images with the FIJI command “Measure”, using ROIs on single cells. A minimum of 50 cells per cell type/condition was acquired for a minimum of three biological replicates for the main figures, and a minimum of 2 biological replicates for supplementary data.

RNA isolation and quantitative PCR

Total RNA was isolated with the RNeasy Micro Kit (catalog number: 74004, Qiagen) following the manufacturer's instructions. RNA quantification was performed with Qubit RNA HS Assay Kit (catalog number: Q32852, Invitrogen). The samples for qPCR analysis were further converted into cDNA with SuperScript® VILO cDNA Synthesis Kit and Master Mix (catalog number: 11754050, Invitrogen) or High-Capacity cDNA Reverse Transcription Kit (catalog number: 4368814, Applied Biosystems) following manufacturer instructions. Real-time PCR was performed on LightCycler® 96 System (Roche) with Power SYBR® Green PCR Master Mix (catalog number: 4367659, Applied Biosystems), 5 µM primers concentration, and 0.5 ng of cDNA. Relative gene expression was calculated by dividing the Ct value of each gene by the Ct value of the control (Large Ribosomal Protein, Rplp0, or Ribosomal Protein L7, Rpl7). The used primers are listed in Supplementary Table 1.

Co-immunoprecipitation (Co-IP)

Mice were anesthetized with isofluorane, and barium chloride (concentration described in the “Skeletal muscle injury” section) was injected into the TA muscles. At 5 DPI, TA muscles were harvested, washed in 1× PBS for 5 min, snap frozen in liquid nitrogen, and immediately processed for protein lysate. Frozen TA muscles were lysed in lysis buffer (10×; 50 mM Tris, 100 mM NaCl, 1 mM EDTA, 0.5% NP40 + freshly added 1× phosphatases and proteases inhibitors, catalog numbers respectively: 04906837001 and 11836153001, Roche). Samples were kept at 4 °C for 15 min for the lysis step, followed by centrifugation at 4 °C for 15 min. The supernatant fraction was quantified by a BSA standard curve before performing co-IP. Four mg of protein lysate were pre-cleared with protein G beads (catalog number: 10004D, Invitrogen) at 4 °C for 1 h, followed by incubation with anti-TnC antibody 1.5 µg/IP (catalog number: 10337, Immuno-Biological Laboratories) conjugated magnetic protein G beads at 4 °C overnight on a rotator. Prior to performing the Western blot assay, beads were collected using a magnetic rack and washed 3 times with 1× lysis buffer and 1 time with ammonium bicarbonate 50 mM at 4 °C for 5 min on a rotator. Beads were resuspended in a 40 µl final volume of Western blot loading buffer, and a Western blot was performed. Uncropped gels can be found in Supplementary Data 5.

Western blot

Total protein lysates were prepared from cultures using lysis buffer (50 mM Tris HCl, 100 mM NaCl­, 1 mM EDTA, 1% Triton X-100, pH = 7.5) containing protease and phosphatase inhibitors. Cell membranes were removed by centrifugation, and protein concentration was determined by using a BSA standard curve. Protein lysates or co-IP processed samples were loaded onto a NuPAGE 4–12% Bis-Tris gel, and electrophoresis was performed in MOPS SDS running buffer. Proteins were transferred to a PVDF membrane and blocked with 5% BSA in PBST (PBS with 0.1% Triton X-100). Incubation with primary antibodies was performed at 4 °C overnight. The antibodies used were: mouse anti-tenascin (catalog number: 10337, Immuno-Biological Laboratories), rabbit anti-Annexin A2 (catalog number: 8235S, Cell Signaling Technology), mouse anti-vinculin (catalog number: sc-25336, Santa Cruz Biotechnology), and HRP-conjugated secondary antibodies (Santa Cruz), at the final concentration recommended by the manufacturers. Membranes were visualized with enhanced chemiluminescence (Pierce, Thermo Scientific Cat #32106) and developed on film. Uncropped gels can be found in Supplementary Data 5.

ELISA assay

Cells were cultured for 72 h in growth conditions on laminin-coated plates before medium collection. Upon collection, the media was spun down for 15 s at the highest speed (~16,000 rpm) with a tabletop centrifuge. Supernatant was transferred into sterile tubes and stored at −80 °C until analysis. Mouse Tenascin-C ELISA Kit (catalog number: ab303745, Abcam) was used for this assay. Antibody cocktail (capture antibody, detector antibody, 5BR antibody - anti-Tenascin-C) was prepared according to the manufacturer's directions. Undiluted sample, blank, and 6 serial dilutions of the sample were prepared and mixed with the antibody cocktail and then incubated in the provided plate for 1 h at room temperature on a shaker at 400 rpm. After incubation, three 5-min washes with wash buffer were performed before 10 min at room temperature incubation with TMB solution (in the dark). Immediately after, we added the stop solution and read at the plate reader (FilterMax F5, Molecular Devices, operating on SoftMax Pro 7.0.2) at 450 nm. To calculate the concentration of TnC expression per cell (in picograms, pg) after 72 h, we accounted for the proliferation rates and division timing of each cell type.

Analysis of single-cell RNA sequencing

scRNAseq data was downloaded from Oprescu et al.⁵⁷ using the accession number GSE138826. Bioinformatics analysis was performed using Seurat (version 5.0.2 and R version 4.3.3), dplyr, reshape2, tidyverse, and scCustomize. Visualizations were performed using the Seurat function (VlnPlot), the CellChat package (see “Cell–cell communication analysis” below), and the R packages ggplot2 and dittoSeq.

Cell–cell communication analysis

The cell–cell communication and visualization were performed using CellChat (version 2.1.0). CellChatDB is a manually curated database of literature-supported ligand-receptor interactions in multiple species (we sub-selected the CellChatDB.mouse database), including multi-subunit structures of ligand-receptor complexes and co-factors. We manually updated the CellChatDB.mouse interaction database to include several known interactions that were not included by default (TnC_Anxa2, TnC_Itga7_Itgb1, TnC_Egfr, TnC_Itgav_Itgb1, TnC_Itga2_Itgb1, and TnC_Tlr4) after conducting a thorough literature search. For the cell–interaction analysis, the expression levels were calculated relative to the total read mapping to the same set of coding genes in all transcriptomes. The expression values were averaged within each single-cell cluster/cell sample. Visualizations were performed using the following CellChat functions: netVisual_aggregate, netVisual_heatmap, netVisual_bubble, and netAnalysis_individual.

Statistics and reproducibility

All statistical analyses were performed using GraphPad Prism version 7 (GraphPad Software). Error bars in the figures represent the standard error of the mean (SEM). The number of biological replicates is indicated by n, while the number of cells or fields of view is indicated as N_cell in the figure legends. Normality of the data was assessed using the Shapiro-Wilk test. Based on the distribution, parametric tests (Student’s t-test, one-way ANOVA, or two-way ANOVA) were applied to normally distributed data, while non-parametric alternatives were used for non-normally distributed data. Statistical significance was defined as p < 0.05. Violin plots generated from the bioinformatics analysis are reported in Supplementary Data 6 as an overlay with box plots. Violin plots of experimental data and stack plots are reported in Supplementary Data 7 as, respectively, violin plots with individual datapoints and grouped graphs with individual datapoints. Individual values are reported in Supplementary Data 8.

A minimum of 3 biological replicates was chosen, which is based on preliminary data from the lab, to give statistical significance in the assays performed. Randomization was not performed using a formal randomization method. Mice were randomly assigned to treatment and control groups to avoid selection biases. Treatments and assessments were performed in a consistent order. Animals were housed under identical conditions, with cage positions rotated weekly to minimize environmental confounding. Blinding was not implemented during the analysis of the results as there was no specific preconception about the experimental outcome. Randomization was used to strengthen the objectivity of this study.

Reporting summary

Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.

Reporting summary

Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.

Author contributions

Conceptualization, A.C., M.L., and A.S.; Methodology, M.L., A.C., C.K., C.S., G.G., A.R., C.N., A.K., D.M., S.D., L.C. and X.W.; Investigation, M.L., A.C., C.K., C.S., G.G., A.R., A.K., D.M., S.D., L.C., X.W. and A.S.; Writing & Editing, A.C., M.L., C.K., A.Co., P.L.P. and A.S.; Funding Acquisition, A.S.; Resources, A.S.; Supervision, A.S.

Peer review

Peer review information

Communications Biology thanks Ande Marini and the other anonymous reviewer(s) for their contribution to the peer review of this work. Primary Handling Editors: Ngan Huang, George Inglis, and Mengtan Xing.

Data availability

The data analyzed during this study are included in this published article. The source data behind the graphs presented in the study, data not shown in the main figures or the Supplementary Figs., such as uncropped blots supporting the findings of this study, the FACS gating strategy, and complementary information, can be found in the Supplementary Data files. Publicly available RNAseq datasets were from Oprescu et al.⁵⁷ (accession number GSE138826) and Walter et al.⁸⁴ (accession numbers GSE143437, GSE159500, GSE162172, GSE232106). Further information for resources and reagents is available from the corresponding author upon reasonable request.

Competing interests

The authors declare no competing interests.

Supplementary information

The online version contains supplementary material available at 10.1038/s42003-025-09189-z.