MRG15 decline in aged/injured MuSCs hinders regeneration via differentiation defects

Abstract

Skeletal muscle aging is characterized by a functional decline in muscle stem cells (MuSCs), yet the key regulatory mechanisms driving this deterioration remain poorly understood. By integrating transcriptomic profiles from aged MuSCs with data from C2C12 cells exposed to spaceflight conditions (which mimic an aging-like phenotype), we identified MORF4-related gene on chromosome 15 (MRG15) as a putative epigenetic regulator involved in age-related myogenic decline. Using a MuSC-specific inducible knockout (iKO) mouse model, we found that loss of MRG15 severely compromises myogenic differentiation and muscle regeneration. Subsequent RNA sequencing of iKO MuSCs, combined with ChIP-seq analysis of histone modifications, revealed that MRG15 modulates the chromatin landscape of myogenic genes through interaction with MyoD, thereby facilitating transcriptional activation and differentiation. Our findings establish MRG15 as a critical epigenetic regulator that cooperates with MyoD to orchestrate chromatin remodeling, thereby promoting transcriptional activation of the myogenic program. Dysregulation of MRG15 may underlie impaired muscle regeneration during aging.

Supplementary Information

The online version contains supplementary material available at 10.1186/s13619-026-00279-9.

Background

Skeletal muscle regeneration is a highly orchestrated process requiring precise coordination of multiple cellular and molecular pathways. Serving as the primary cellular contributors of muscle repair and growth, skeletal muscle stem cells (MuSCs) normally maintain quiescence in homeostasis. Following injury, MuSCs undergo rapid activation, proliferative expansion as myoblasts, and subsequent cellular differentiation, ultimately fusing to form multinucleated myotubes that further mature into functional myofibers (Yin et al. 2013). Dysregulation at any critical process may inevitably compromise the overall regenerative process.

Aging impairs MuSC function and diminishes muscle regeneration capacity (Giza et al. 2022). This dysfunction disrupts skeletal muscle homeostasis, ultimately leading to atrophy (reduced muscle mass), and functional decline characterized by decreased strength and endurance. Collectively termed sarcopenia, these age-related changes closely resemble the effects of spaceflight on skeletal muscle (Cannavo et al. 2022; Juhl et al. 2021).

The dynamic transitions between MuSC functional states are tightly regulated by epigenetic modifications, particularly histone acetylation. In the quiescence state, low histone acetylation levels maintain transcriptional repression of activation genes, thereby preserving the stem cell reserves. Upon injury-induced activation, histone hyperacetylation opens chromatin to activate myogenic transcription (Liu et al. 2013; Massenet et al. 2021). It is also worth noting that, while global histone acetylation decreases during early myogenic differentiation, increases in histone acetylation at myogenic gene promoters drive myogenic differentiation (Asp et al. 2011; Yucel et al. 2019). Aging disrupts this epigenetic regulation, with aberrantly elevated histone hyper-acetylation in MuSCs, impairing stem cell maintenance and regenerative capacity (Blau et al. 2015; Dong et al. 2022; Liu et al. 2013; Zhou et al. 2019a). Although significant progress has been made, the mechanistic basis by which epigenetic regulators coordinate MuSC function and muscle regeneration remains elusive.

MORF4-related gene on chromosome 15 (MRG15) encodes a highly conserved epigenetic regulator that plays critical roles in diverse biological processes, including DNA repair, cellular differentiation, and immune responses (Bertram and Pereira-Smith 2001; Jiang et al. 2024). Functioning as a key modulator of chromatin architecture and transcriptional programs, MRG15 exerts its effects by influencing histone modifications and transcription factor activity. To be noted, MRG15 interacts with either histone acetyltransferases (HATs) or deacetylases (HDACs), thereby dynamically regulating chromatin structure through its association with these histone-modifying enzymes (Chen et al. 2009; Peña et al. 2011; Wei et al. 2020). While the importance MRG15 has been well-documented in various tissues, its specific contributions to MuSC function and muscle regeneration remain unexplored.

In this study, we have identified MRG15 as a regulator of myoblast differentiation. Its expression decreases accompanied by the impairment of myogenic function caused by aging and spaceflight. We elucidate the functional role of MRG15 in skeletal MuSCs and muscle regeneration by employing an MuSC-specific inducible knockout model combined with molecular profiling. Mechanistically, MRG15 orchestrates histone acetylation dynamics at the promoters of key myogenic genes through its direct interaction with MyoD, the master myogenic transcriptional factor. Collectively, our data suggests that decreased MRG15 expression may contribute to MuSC dysfunction in models of skeletal muscle impairment, including aging and spaceflight, highlighting its potential role in muscle homeostasis.

Results

MRG15 is associated with skeletal myogenesis and muscle regeneration during aging

To identify key epigenetic regulatory factors governing the impaired myogenic program in aging, we first analyzed existing transcriptomic profiles of activated MuSCs (ASC) isolated post-injury from young and old mice (Dong et al. 2022). GO pathway analysis of downregulated genes in old mice revealed significant enrichment in chromatin remodeling and transcriptional regulation pathways (Fig. 1A). Notably, transcriptomic datasets from our lab’s C2C12 cultures in orbit exhibited parallels to chromatin remodeling patterns observed in aged ASCs (Fig. 1B). Gene set enrichment analysis (GSEA) of down-regulated genes in C2C12 cultured under orbital conditions further demonstrated enrichment of downregulated genes in skeletal muscle development (Fig. 1C), consistent with the previous finding that the decline of myogenic differentiation under microgravity conditions (Calzia et al. 2020; Kim et al. 2024).

Fig. 1.

MRG15 is associated with skeletal myogenesis and muscle regeneration during aging. A Down-regulated genes GO enrichment analysis showing top ten pathway alterations in active MuSCs of young and old mice. B Down-regulated genes GO enrichment analysis showing top ten pathway alterations in C2C12 cells cultured in ground and orbit. C Gene set enrichment analysis (GSEA) revealed that the muscle tissue development gene set was significantly enriched among down-regulated genes in C2C12 cells cultured in orbit. D Venn diagram shows 14 overlapping genes among three sets: decreased chromatin remodeling genes in aged activated MuSCs (green, 126), those in orbit-cultured C2C12 cells (red, 144), and known transcription factors/co-factors (blue, 2581). E qPCR analysis of Mrg15 and myogenic gene expression in differentiated MuSCs isolated from young and old mice. F qPCR analysis of Mrg15 expression in tibialis anterior (TA) muscles during regeneration after BaCl₂-induced injury (at day 4) compared to uninjured TA muscles. G qPCR analysis of Mrg15 expression in active MuSC (ASCs) isolated from injured muscle, compared to freshly isolated MuSC (FISC) from non-injured muscle (n = 3). H The expression pattern of MRG15 and early-differentiation marker gene during myogenesis in vitro. Means ± SEM are shown. * p<0.05; *** p < 0.001, ns denotes not significant

To further identify the key factors implicated in impaired myogenic differentiation, we intersected the chromatin remodeling-associated gene sets (identified via GO enrichment analysis of ASCs from young and old mice and C2C12 cells cultured under orbital microgravity) with established databases of transcription factors and co-factors (Fig. 1D) (Shen et al. 2023). Prioritizing candidates by expression abundance, we identified chromatin regulatory factor MRG15—not previously characterized in myogenesis. Our integrated analysis suggests that reduced Mrg15 levels correlate with impaired myogenic differentiation in both aging and spaceflight contexts. This notion was further supported by an in vitro culture system using MuSCs isolated from aged mice. Results demonstrated that, Mrg15 expression was significantly lower in aged cells compared to young cells, accompanied by a marked downregulation of the key myogenic differentiation marker Mef2c (Fig. 1E). This aligns with a previous report that aged myogenic cells exhibit slower differentiation, characterized by reduced transcription of Mef2c (Kimmel et al. 2021). Furthermore, overexpression of Mrg15 expression in aged cells effectively restored Mef2c transcript levels (Fig. 1E). These findings suggest that the decline in MRG15 may contribute to the functional impairment of MuSCs during aging.

To determine whether MRG15 plays a functional role in MuSCs and muscle regeneration, we employed a well-established muscle injury model involving BaCl₂-induced acute muscle damage, which reliably triggers the muscle regenerative process (Morton et al. 2019). Intriguingly, Mrg15 transcription levels were significantly upregulated in injured muscle during the early regeneration phase, exhibiting a temporal expression pattern similar to the myogenic regulators Myod1 and Myog (Fig. 1F, Fig. S1A-B). These results suggested a potential involvement of MRG15 in the initiation of myogenesis during muscle repair.

Further supporting this notion, we found that Mrg15 mRNA levels were elevated progressively in ASC isolated from injured muscle compared to those freshly isolated MuSC (FISC) from non-injured muscle (Fig. 1G). This finding is consistent with existing transcriptomic data (Chen et al. 2021), demonstrating Mrg15 upregulation during the transition of MuSCs from quiescence to activation, as well as in the time-course of cultured MuSCs at various post-isolation time points (Fig. S1C).

To further delineate the temporal regulation of MRG15 expression during in vitro myogenesis, we induced differentiation in C2C12 myoblasts and observed a marked upregulation of MRG15 protein expression during differentiation. Concordantly, the early differentiation marker MyoD exhibited sustained induction during this process (Fig. 1H). Collectively, these results strongly implicate MRG15 in the regulation of active myogenic differentiation programs.

MRG15 is a pro-myogenic factor during myoblast differentiation

The observed upregulation of MRG15 during myoblast differentiation led us to hypothesize that MRG15 might function as a pro-myogenic factor. To investigate this possibility, we generated MRG15 deficient myoblasts using CRISPR-Cas9 gene editing with a single guide RNA targeting Mrg15 (sgMrg15, Fig. 2A) (Wei et al. 2020). While MRG15 depletion did not significantly affect proliferation rates, as measured by EdU incorporation (Fig. 2B) or cell number (Fig. S2A) under growth conditions, it caused pronounced differentiation defects, as evidenced by impaired myotube formation (Fig. 2C, S2B) and diminished or delayed the expression of essential myogenic markers, including MyoG and MyHC (Fig. 2D). Together, these results suggest that MRG15 plays a critical role in promoting myogenic differentiation without affecting myoblast proliferation. Both the p38MAPK and AKT-mTOR pathways are critical for myoblast differentiation (Cong et al. 2020; Lluís et al. 2006; Serra et al. 2007). We therefore asked if MRG15 regulates these signaling pathways during differentiation. Interestingly, MRG15 knockdown led to a concurrent increase in both total and phosphorylated proteins of p38MAPK, AKT, and mTOR downstream effector S6K (Fig. S2C), yet the normalized phosphorylation levels remained comparable to controls (Fig. S2D). These results imply a homeostatic compensatory response, which maintains pathway activity despite MRG15 reduction. Nevertheless, this compensatory activation failed to rescue the differentiation impairment, suggesting that MRG15 may act as a critical downstream effector within these pathways or play a distinct role in driving differentiation.

Fig. 2.

MRG15 is a pro-myogenic factor during myoblast differentiation. A Western blot analysis of MRG15 protein levels in C2C12 cells modified by CRISPR-Cas9 with deletion of MRG15 (sgMrg15) or a non-targeting control (sgCtrl). B EdU labeling for 3 h in above cells was performed. The percentage of EdU⁺ cells was quantified (n = 4). C Immunofluorescence (IF) staining of MF20 (green) was performed on sgMrg15 or sgCtrl-C2C12 after induction of differentiation and the percentage of MF20⁺ nuclei/total nuclei was quantified (n = 4). D The expression pattern of MRG15 and differentiation marker genes in sgMrg15 or sgCtrl-C2C12 during myogenesis in vitro. Scale bar, 50 μm. Means ± SEM are shown. *** p < 0.001. ns denotes not significant

Loss of MRG15 impairs injury-induced muscle regeneration in vivo

To extend our findings from myoblasts to an in vivo animal model, we generated an MRG15-inducible knockout (iKO) mice by crossing Mrg15^flox/flox control (Ctrl) mice with Pax7-Cre^ERT mice, enabling specific deletion of MRG15 in MuSCs (Fig. 3A). Quantitative mRNA analysis confirmed nearly complete ablation of Mrg15 in MuSCs isolated from iKO mice compared to Ctrl mice (Fig. 3B). This was achieved by intraperitoneal (i.p.) injection of adult mice with tamoxifen (TMX) for five consecutive days to induce MRG15 deletion in MuSCs, which were subsequently purified using fluorescence-activated cell sorting (FACS) (Fig. 3C).

Fig. 3.

Loss of MRG15 impairs injury-induced muscle regeneration in vivo. A Breeding scheme for generating the Control (Ctrl) and inducible MRG15 knockout (iKO) mice. B qPCR analysis of Mrg15 expression in FISC from Ctrl and iKO mice in 3 days after Tamoxifen injection (n = 3). C Representative fluorescence-activated cell sorting plots and the percentage of MuSC isolated from the uninjured muscles of Ctrl and iKO mice were presented. D Numbers of PAX7⁺ cells per 100 myofibers was quantified (n = 6) in uninjured muscles. E H&E staining was performed in uninjured (0dpi) and 5 days post-injury (5dpi) TA muscles. Scale bar, 50 μm. F IF staining for eMyHC (red) were performed on TA muscles 5 days post-injury. Numbers of fibers per field was quantified (n = 6). G Protein expression of p38MAPK and AKT-mTOR pathways in muscle from Ctrl and iKO mice at 5 days post-injury. Scale bar, 50 μm. Means ± SEM are shown. *** p < 0.001. ns denotes not significant

Notably, MRG15 iKO mice exhibited normal body weight and muscle mass, and gross morphology with no apparent physical abnormalities (Fig. S3A-D). Furthermore, three days post-TMX injection, immunohistochemical assessment of muscle sections showed comparable MuSC numbers in both groups (Fig. 3D, S3E), consistent with this observation, FACS analysis revealed no significant difference in MuSC quantity between Ctrl and MRG15 iKO littermates (Fig. 3C), indicating that acute MRG15 loss does not impair skeletal muscle homeostasis or MuSC maintenance.

Given the pro-differentiation role of MRG15 in myoblasts in vitro (Fig. 2), we hypothesized that ablation of MRG15 in MuSCs might similarly impair muscle regeneration in vivo. To test this, we induced acute muscle injury via intramuscular injection of BaCl₂ into the tibialis anterior (TA) muscle of mice (Fig. S3F). By the fifth day post-injury (dpi), immune infiltration persisted but had not fully resolved. During this phase, we observed numerous regenerating muscle fibers characterized by central nucleation and expression of embryonic myosin heavy chain (eMyHC)—a transient marker of early regenerating myofibers (Schiaffino et al. 2015). Strikingly, histological analysis revealed fewer regenerating myofibers in MRG15 iKO mice (Fig. 3E-F). Consistently, the muscle tissue from iKO models exhibited a significant downregulation of MyHC protein expression, as well as a compensatory activation of the p38MAPK and ATK-mTOR signaling pathways (Fig. 3G), which mirrors the cellular data (Fig. S2C). Altogether suggested that MRG15 loss compromises limb muscle repair.

MRG15 regulates myogenic gene expression in MuSCs

To further elucidate whether impaired regeneration is MuSC-autonomous, we isolated MuSCs from iKO and Ctrl mice and cultured them for 6 days (Fig. 4A). When grown in culture, MuSCs enter the cell cycle and start dividing within 2 days of culture (Li et al. 2020). Within 6 days, they start differentiating and express myogenic markers like MyHC and troponin (Fig. S4A). Consistent with the phenotype observed in sgMrg15 myoblasts, MRG15-deficient MuSCs exhibited a significant reduction in myogenic genes, underscoring their diminished differentiation potential (Fig. S4A). These data suggest that MRG15 deletion in MuSCs can disrupt myogenic differentiation, ultimately leading to impaired muscle regeneration.

Fig. 4.

MRG15 regulates myogenic gene expression in MuSCs. A Schematic of MuSC RNA-seq after collection and culture in Ctrl or iKO mice. B Volcano plot showing changes of different expression genes (DEGs) in Ctrl or iKO mice. C Down-regulated genes GO enrichment analysis showing top eight pathway alterations in Ctrl or iKO MuSC. D Up-regulated genes GO enrichment analysis showing top eight pathway alterations in Ctrl or iKO MuSC. E Top eight KEGG pathway of down-regulated genes in Ctrl or iKO MuSC. F Top eight KEGG pathway of up-regulated genes in Ctrl or iKO MuSC

To elucidate the molecular mechanisms by which MRG15 regulates differentiation of MuSC, we performed RNA sequencing (RNA-seq) to analyze the transcriptomic alterations resulted from MRG15 depletion. Comparative analysis between MRG15 iKO and Ctrl MuSCs we identified 2,689 differentially expressed genes (DEGs), with 1,214 genes being significantly upregulated and 1,475 genes downregulated (Fig. 4B).

Gene Ontology (GO) enrichment analysis of downregulated genes revealed significant associations with key biological processes, including “muscle contraction”, “cell differentiation”, and “sarcomere organization” (Fig. 4C), which aligns with the observed impairment in myogenic differentiation upon MRG15 ablation (Fig. S4A). Intriguingly, upregulated genes were prominently enriched in processes such as “cell adhesion”, “inflammatory response”, and “signal transduction” (Fig. 4D).

Complementary KEGG pathway analysis demonstrated that downregulated genes were significantly enriched in pathways, including “cytoskeleton in muscle cells”, whereas upregulated genes markedly enriched in pathways, including “cell adhesion molecules” (Fig. 4E-F). Notably, among the downregulated genes, several muscle-specific genes exhibited the most pronounced reduction in expression levels (Fig. S4B). Collectively, these results support the hypothesis MRG15 plays a critical role in activating myogenic gene expression and facilitating myoblast differentiation.

MRG15 interacts with MyoD to regulate histone acetylation and transcription of myogenic genes

To investigate whether MRG15 cooperates with transcription factors (TFs) in regulating MuSC differentiation, we performed a bioinformatic analysis by using Metascape (Zhou et al. 2019b). Conspicuously, RUNX1, MEF2C, P300 and MyoD were among the top four most significant transcriptional regulators associated with the downregulated gene set (Fig. 5A). Previous studies have reported that MyoD modulate differentiation through epigenetic mechanisms, and P300 is required for MyoD-dependent cell cycle arrest and muscle-specific gene transcription (Puri et al. 1997; Wang et al. 2022). Given that MRG15 associates with acetyltransferases TIP60 which mediated acetylation of H4 (specifically K5/K12/K16) and H2A and H4K16ac is critical for myogenic activation (Devoucoux et al. 2022; Peña et al. 2011; Ryall et al. 2015), we integrated our MRG15 iKO RNA-Seq data with published H4K16ac ChIP-Seq data in MuSC (Ryall et al. 2015), and identified 216 downregulated genes marked by H4K16ac (Fig. 5B), and further analysis revealed that these gene were enriched in skeletal muscle development-related GO terms (Fig. 5C). Subsequent de novo motif analysis of H4K16ac-bound regions revealed strong enrichment for the MyoD binding motif (Fig. 5D), further suggesting a functional link between MRG15-mediated histone acetylation and MyoD-dependent transcription. Strikingly, among these downregulated genes, Mef2c and Myog—known downstream targets of MyoD (Byun et al. 2017; Cao et al. 2006; de Esteves Lima and Relaix 2021; Wang et al. 2001).

Fig. 5.

MRG15 interacts with MyoD to regulate histone acetylation and transcription of myogenic genes. A Metascape databases were used to predict the transcription factors interacting with MRG15. B Venn diagrams showing the overlapping (216 genes) between the H4K16 acetylation (ChIP-Seq) target (9930) and the down-regulated genes (1460) among RNA-seq. C GO analysis of the above 216 genes revealed an extreme enrichment of skeletal muscle tissue development terms. D Enrichment of canonical MyoD motif in the above binding regions. E and F Co-IP assays showing the interactions between MRG15 and MyoD (E), or TIP60 (F) in C2C12. G ChIP-PCR analysis of H4K16ac occupancy at the promoter regions of myogenic genes in sgMrg15 versus sgCtrl-C2C12 cells. Data are presented as percentage of input (% Input). H qPCR analysis of myogenic genes expression in sgMrg15 or sgCtrl-C2C12 (n = 3). Means ± SEM are shown. * p < 0.05, ** p < 0.01, *** p < 0.001

Given the demonstrated role of MRG15 regulated differentiation both in vitro and in vivo, we hypothesized that MRG15 may functionally interact with MyoD to orchestrate epigenetic reprogramming of downstream target genes, thereby regulating myoblast differentiation. Subsequently, we first confirmed the MyoD/MRG15 association by western blotting following Co-immunoprecipitation (Co-IP) in differentiating C2C12 cells. The Co-IP assay showed that MRG15 was co-immunoprecipitated with MyoD, indicating a direct binding between MRG15 and MyoD (Fig. 5E). Since MRG15 serves as a key recruiter of the TIP60 HAT complex (Peña et al. 2011), which mediated acetylation of H4K16, and given that P300 (Puri et al. 1997) and TIP60 (Kim et al. 2011), well-characterized member of the HAT family, both directly interacts with MyoD to regulate myogenic differentiation. Therefore, to define the components within the HAT complex, we next sought to determine whether MRG15 associates with TIP60 or P300, two key acetyltransferase components, in C2C12 cells. The result demonstrates that MRG15 specifically interacts with TIP60 (Fig. 5F) but not with P300 (Fig. S5A).

To further directly link MRG15 loss to impaired histone acetylation and transcriptional silencing, we performed ChIP-qPCR to assess H4K16ac enrichment at potential target genes, and examined their mRNA levels in sgMrg15 cells. Our results show that loss of MRG15 significantly reduces H4K16ac deposition at these loci (Fig. 5G, S5B), which corresponds with a decrease in their transcription levels (Fig. 5H). Collectively, these findings further support a model wherein MRG15 recruits the histone acetyltransferase TIP60, to MyoD-targeted genomic regions to facilitates histone acetylation and transcriptional activation of differentiation-related genes (Fig. S5C).

Discussion

Our study provides comprehensive insights into the functional role of MRG15 in MuSCs and muscle regeneration, while uncovering its critical involvement in the epigenetic and transcriptional regulation of myogenesis. As depicted in Fig. S5 myogenic differentiation represents a pivotal stage in muscle regeneration, principally mediated by MuSCs. During the initiation of MuSC differentiation, MRG15 expression is upregulated and facilitates the recruitment of histone-modifying complexes such as TIP60 to MyoD-binding loci through its physical interaction with MyoD. This recruitment subsequently induces histone acylation such as H4K16ac, thereby activating myogenic gene expression and promoting MuSC differentiation.

The low levels of MRG15 in ASC during aging and C2C12 cultured in orbit suggest that the dysregulation of its expression may also contribute to the impairment of skeletal muscle regeneration caused by space flight and aging. Consequently, developing strategies to modulate MRG15 expression levels, such as through gene therapy or pharmacological agents, could emerge as a promising therapeutic approach to counteract the impaired muscle regeneration observed both in the elderly following injury and astronauts during long-duration spaceflight.

A direct interaction between MyoD, TIP60 and MRG15 was observed by Co-IP assay in our system. Since MRG15 serves as a key recruiter of the TIP60 HAT complex, which mediated acetylation of H4K16, and given that TIP60, a well-characterized member of the HAT family, directly interacts with MyoD to regulate myogenic differentiation(Kim et al. 2011; Peña et al. 2011; Puri et al. 1997; Yuan et al. 1996), it is plausible that MyoD also acts through MRG15 to recruit these HAT complexes to its target genomic loci. This MRG15-mediated recruitment would enhance local histone acetylation within enhancers or promoters of MyoD-target genes, thereby facilitating the transcription of myogenic genes and promoting cellular differentiation.

It is worth noting that previous studies have reported context-dependent roles of MRG15 in cell proliferation—either promoting proliferation through activation of cell cycle-related genes (e.g., Cdc2) or inhibiting proliferation via p21 upregulation in various cell types (Jiang et al. 2024)—our findings revealed distinct characteristics in myoblasts. Specifically, MRG15 depletion did not significantly affect myoblast proliferation, as evidenced by unaltered EdU incorporation rates (Fig. 2B) and unchanged cell numbers (Fig. S2A). Consistent with these observations, our RNA-seq data showed no significant changes in Cdc2 or Cdkn1a levels in MRG15-deficient MuSCs (data not shown). Taken together, these results suggest that MRG15 is dispensable for myoblast proliferation, and that its role in promoting myogenic differentiation—likely mediated through MyoD—may represent a tissue-specific function.

Methods

Mouse studies

All procedures that involved animal handling were approved by the Ethics Committee of the Shanghai Institute of Nutrition and Health (SINH), Chinese Academy of Sciences (CAS). All in vivo experiments described in this study followed institutional guidelines for animal care and use. Pax7-Cre^ERT (Wang et al. 2021) mouse was kindly provided by Prof. Ping Hu (Guangzhou Laboratory-Guangzhou Medical University). Mrg15^flox/flox strain (Wei et al. 2020) was kindly provided by Prof. Qiurong Ding (SINH, CAS). The MRG15-inducible conditional KO (iKO) strain (Ctrl: Pax7-Cre^ERT/+; Mrg15^+/+, iKO: Pax7-Cre^ERT/+; Mrg15^flox/flox mice) was generated by crossing Pax7-Cre^ERT/+ mice with Mrg15^flox/flox mice. Inducible conditional deletion of MRG15 was administered by tamoxifen (TMX; T5648, Sigma) intraperitoneally (i.p.) at 2 mg per 20 g body weight. All mice were housed at a temperature of 23 ± 3℃ and humidity of 35 ± 5% under a 12 h light/dark cycle in a specific pathogen-free animal facility. To induce muscle injury, 50 µl of 1.2% BaCl₂ was injected into the tibialis anterior (TA) muscle of adult mice. Muscles were harvested at designated time points for further analysis.

Cell culture

Mouse C2C12 myoblasts were cultured in growth medium, GM, DMEM medium (11995073, Gibco) with 10% fetal bovine serum, FBS (10270106, Gibco), 1% penicillin/streptomycin (P/S, 15140122, Gibco), or differentiation medium, DM, DMEM medium with 2% horse serum (16050114, Gibco), 1% P/S in incubator at 37 °C. Cells were treated with EdU (C10086, Thermo Fisher Scientific, 10 µM) for 3 h. The EdU staining was performed with a Click-iT™ imaging kit (C10337, Thermo Fisher Scientific).

MuSC isolation and culture

Hindlimb muscles from mice were digested with collagenase II (LS004177, Worthington, 1000 U/mL) for 90 min at 37 °C, the digested muscles were then washed in washing medium (Ham’s F-10 medium (N6635, Sigma) containing 10% horse serum, heat-inactivated (HIHS, 26050088, Gibco), 1% P/S) before cells were liberated by treating with Collagenase II (100 units/mL) and Dispase II (D4693-1G, Sigma, 1.1 U/mL) for 30 min. The suspensions were passed through a 20 G needle to release myofiber-associated cells. Mononuclear cells were filtered with a 40-μm cell strainer and incubated with the following primary antibodies: Vcam1-biotin (105704, BioLegend), CD31-FITC (102506, BioLegend), CD45-FITC (103108, BioLegend), and Sca1-Alexa647 (108118, BioLegend). The Vcam1 signal was amplified with Streptavidin-PE (554061, BD Biosciences). All antibodies were used at a dilution of 1:75. The BD FACSAria Fusion Cell Sorter (BD Biosciences) was used for cell sorting following the manufacturer’s instructions. Flowjo (version X.0.7, BD Biosciences) software is used to analysis of flow cytometry data. Coverslips and cultural wells were coated with poly-D-lysine solution (p0899, Sigma) at 37 °C for overnight and then coated with extracellular matrix (ECM) (E-1270, Sigma) at 4 °C for at least 6 h. FACS-isolated MuSCs were seeded in coated wells and cultured in Ham’s F10 medium with 10% HIHS, 5 ng/mL β-FGF (PHG0026, Thermo Fisher Scientific) and 1% P/S.

Generation of MRG15 knockout cell line

To delete MRG15 in C2C12, one target-specific guide RNA was provided by Prof. Ding Qiurong (SINH, CAS) and described previously (Wei et al. 2020). Transfect 293 T cells (40–60% confluent) with 3 plasmids: 11 μg of pLenti-sgMrg15 or pLenti-CRISPR-V2 plasmid, 8.5 μg of Pspax2 plasmid and 4.07 μg of pMD2G plasmid using Lipofectamine2000 transfection reagent (11668,019, Invitrogen) for each 10 cm dish. Collect the supernatant 2–3 days after transfection. Filter the virus-containing medium through 0.45 μm filter. Transduce C2C12 cells (6 cm dish, ~ 40–50% confluent) with 2 mL medium containing 32 µg Polybrene and 2 mL thawed virus. 48 h after transduction, cells were placed in puromycin (10 µg/mL) medium for stable selection.

Immunofluorescence

Cells were fixed in 4% PFA for 15 min and permeabilized with 0.5% NP-40 for 10 min. Then cells were blocked in 3% BSA for 1 h followed by incubating with primary antibodies overnight at 4 °C and secondary antibodies for one hour at room temperature (RT). Antibodies and dilutions were used as follows: mouse anti-MF20 (1:100, MF20, DSHB); goat anti-mouse IgG Alexa Fluor 488 (1:200, A11001, Invitrogen). All images were captured by a fluorescence microscope (Cycloud Bio).

Histology

For Haematoxylin and Eosin (H&E) staining, the OCT-embedded muscle was sectioned at 6 μm and then stained according to standard protocols. For immunofluorescence staining, slides were fixed with 4% PFA for 15 min at RT and permeabilized in ice-cold menthol for 6 min at −20 °C. Heat-mediated antigen retrieval with a 0.01 M citric acid (pH 6.0) was performed for 5 min. After blocking with 4% IgG-free BSA (001–000–162, Jackson) in PBS, the sections were further blocked with unconjugated AffiniPure Fab Fragment (1:100 in PBS; 115–007-003, Jackson) for 30 min. Primary antibodies and dilutions were used as follows: mouse anti-PAX7 (1:50, PAX7-s, DSHB), mouse anti-eMyHC (1:300, NCL-MHC-d, Leica) and rabbit anti-laminin (1:800, L9393, Sigma-Aldrich). The biotin-conjugated anti-mouse IgG (1:500 in 4% BBBSA, 115–065–205, Jackson), Cy3-Streptavidin (1:1250 in 4% BBBSA, 016–160-084, Jackson) and goat anti-rabbit IgG Alexa Fluor 488 or 594 (1:200, Invitrogen) were used as secondary antibodies. All images were captured with a microscope (Cycloud Bio).

RNA isolation and real-time PCR

Tissues or cells were immediately homogenized in TRIzol reagent (Invitrogen) to isolate the RNA, followed by the reverse transcription using RT Reagent Kit (Takara). Real-time PCRs were performed using a commercial master mix (Takara) and 7900 ABI Real-Time System (Applied Biosystems). Primers used in this study were provided (Supplementary Table 1). An average cycle threshold (Ct) value was calculated from the duplicate reactions and normalized to the expression of 18S, and the ΔΔCt value was then calculated.

Western blotting analysis

Protein from cells or tissues was extracted by using RIPA buffer (P0013B, Beyotime) supplemented with Protease Inhibitor Cocktail (5892791001, Roche) and Phosphatase Inhibitor Cocktail (4906837001, Roche) immediately before use and the protein concentration was determined using a Pierce™ Coomassie Plus (Bradford) assay kit (23238, Thermo Fisher Scientific). The following antibodies and dilutions were used for Western blot analysis: rabbit anti-MRG15 (1:1000, 55257–1-AP, Proteintech), mouse anti-MyoG (1:1000, F5D, DSHB), mouse anti-MyoD1 (1:1000, M3512, Dako), mouse anti-MYH (sc-376157, Santa Cruz), mouse anti-TIP60 (1:1000, 10827–1-AP, Proteintech), mouse anti-P300 (1:1000, 83078–5-RR, Proteintech), rabbit anti-AKT (1:1000, 9272S, CST), rabbit anti-Phospho-Akt (Ser473) (1:1000, 9271S, CST), rabbit anti-Phospho-P38 MAPK (Thr180/Tyr182) (1:1000, 8690 T, CST), rabbit anti-S6K (1:1000, 9202S, CST), rabbit anti-Phospho-S6K (Thr389) (1:1000, 9234S, CST), rabbit anti-HSP90(4874S, CST).

Immunoprecipitation assays

C2C12 myoblasts were cultured in differentiation medium for 3 days. Cells were lysed with lysis buffer (50 mM Tris–HCl, pH 8.0, 150 mM NaCl, 0.1% SDS, 0.5% sodium deoxycholate, and 1% NP-40). The whole-cell lysates obtained by centrifugation (with equal concentration of protein in different samples) were incubated with 1 μg of MyoD1 (M3512, Dako) or MRG15 (55257–1-AP, Proteintech), or TIP60 (10827–1-AP, Proteintech), antibody for 1.5 h at 4 °C with rotation followed by binding to Protein A/G PLUS-Agarose (sc-2003, Santa Cruz) for overnight at 4 °C. The immunocomplexes were then washed with washing buffer (10 mM Tris–HCl, pH 7.5, 150 mM NaCl, 1.0 mM EDTA, 1.0 mM EGTA, and 1% Triton X-100) four times and applied to SDS-PAGE.

Chromatin Immunoprecipitation (ChIP) assays

ChIP assays of sgMrg15 or sgCtrl-C2C12 cells were performed using a ChIP Assay Kit (Beyotime) according to the manufacturer’s protocol. The sgMrg15 or sgCtrl-C2C12 cells were cultured in differentiation medium for 3 days. Immunoprecipitation was performed using an anti-Histone H4K16 acetylation antibody (39,167, Active Motif) with rabbit IgG (2729, Cell Signaling Technology) as a negative control. PCR products were resolved by electrophoresis in a 2% Agarose-gel. Primer sequences for ChIP assay are provided in Supplementary Table 2.

RNA-seq and data analysis

MuSCs were cultured for 6 days before being lysed using the TRIzol reagent (Invitrogen). RNA-seq libraries were prepared using Illumina TruSeq Stranded mRNA Library Prep (Illumina). Samples were pooled for deep sequencing on Nova seq 6000 platforms. For RNA-seq data analysis, the raw paired-end reads were trimmed and quality controlled by SeqPrep and Sickle with default parameters. Then clean reads were separately aligned to the reference genome (GRCm39) with orientation mode using HISAT2 software. The mapped reads of each sample were assembled by StringTie in a reference-based approach. Differential expression analysis was performed using DESeq2. The data were analyzed through the free online platform of Majorbio Cloud Platform (www.majorbio.com). Heatmaps for indicated genes were obtained using OmicStudio tools at https://www.omicstudio.cn and free online platform of Majorbio Cloud Platform (www.majorbio.com). ClusterProfiler was used for the Gene Ontology (GO) analysis with Entrez gene IDs converted from DAVID tool as inputs. Functional enrichment analysis was performed with DAVID.

Motif enrichment analysis

Applying HOMER (Heinz et al. 2010) to predict potential TF binding motifs enriched within the 5 kb window of TSSs of genes (log₂ (Fold change) > 1 or <  − 1 with adjusted P value < 0.05).

Statistical analyses

Data were analyzed using GraphPad Prism (version 8; GraphPad Software, San Diego, CA). Results were represented as mean ± SEM. Statistical significance was assessed by Student’s t-test. Differences were considered statistically significant at P < 0.05.

Abbreviations

MuSCs

Muscle stem cells

MRG15

MORF4-related gene on chromosome 15

iKO

Inducible knockout

HATs

Histone acetyltransferases

HDACs

Histone deacetylases

ASC

Activated MuSCs

GSEA

Gene set enrichment analysis

FISC

Freshly isolated MuSC

Ctrl

Control

i.p.

Intraperitoneal

TMX

Tamoxifen

FACS

Fluorescence-activated cell sorting

TA

Tibialis anterior

dpi

Day post-injury

eMyHC

Embryonic myosin heavy chain

RNA-seq

RNA sequencing

DEGs

Differentially expressed genes

GO

Gene Ontology

TFs

Transcription factors

Co-IP

Co-immunoprecipitation

Authors’ contributions

Z.L., and Y.L. designed the experiments; Z.L. carried out most of the cell culture experiments; Z.L., M.M., and R.M. analyzed the data; S.S., W.K., and Y.W. provided the technical assistance, analyzed, and interpreted the data; Q.D., Y.L., and H.Y. contributed to the discussion and supervised the project; Z.L., Y.L., and H.Y. wrote the manuscript.

Funding

This work was supported by National Natural Science Foundation of China (NSFC, 32200686, 32371246 to Y.L.，32271170 to H.Y.); the National Key Research and Development Program of China (2023YFA1801100 to Q.D.&H.Y., 2025YFF0511500 to Y.L.); the China Manned Space Flight Technology Project Chinese Space Station (YYWT-0901-EXP-05) (H.Y.); the Key Research Program of Chinese Academy of Sciences, ZDBS-ZRKJZ-TLC004 (Y.L.); and the Shanghai Leading Talent Program of Eastern Talent (H.Y.). .

Data availability

Data supporting the findings of this study are available within the article and its Supplementary information files or from the corresponding author upon reasonable request. The RNA-seq and ChIP-seq datasets re-analyzed during the current study are available under accession code GSE189073, GSE175501 and GSE64379. The RNA-seq data of MRG15 iKO vs. control MuSCs are accessible in NODE (https://www.biosino.org/node) with the accession number OEP00006709.

Declarations

Ethics approval and consent to participate

All procedures that involved animal handling were approved by the Ethics Committee  of the Shanghai Institute of Nutrition and Health (SINH), Chinese Academy of Sciences (CAS).

Consent for publication

Not applicable.

Competing interests

The authors declare no conflict of interest related to this work.