Slug deletion causes a defect in muscle regeneration

The transcription factor Slug is expressed in a variety of normal tissues in the adult mouse13, indicating its important roles in development. In agreement with this notion, Slug knockout mice showed numerous abnormalities such as smaller body size and weight (Supplementary Fig. 1a,b). Since skeletal muscle accounts for ~40% of adult body weight9, we examined if the reduced body weight in Slug-deficient mice is due to a reduction of muscle mass. Hindlimb muscles of adult Slug-mutant mice were reduced by 21–36% in weight (Supplementary Fig. 1c). To some extent, the lost muscle mass was caused by reduction in myofiber size in Slug-deficient mice (Supplementary Fig. 1d, e and Fig. 1e). However, when normalized to body weight, none of the relative weights of these muscles were affected by the absence of Slug (Supplementary Fig. 1f). Changes in muscle mass may be resulted from changes in protein or cell turnover14. The latter reflects the balance between myonuclear accretion and loss. Proliferation and fusion of SCs increases the number of myonuclei within the muscle fibers. Therefore, we determined the effect of Slug deficiency on MuSC maintenance. Unexpectedly, Slug-deficient mice had even slightly higher fraction of SCs in non-lineage cell subpopulation (Fig. 1a). The total yield of SCs calculated as per milligram of harvested hindlimb muscle was also increased in Slug knockout mice (Fig. 1b). Such Slug ablation-induced increases in MuSC frequency and number were further confirmed by staining of Pax7+ nuclei on freshly prepared TA muscle cryosections (Fig. 1c, d).

Fig. 1 Slug deficiency repairs regenerative capacity of SCs during serial muscle damage. a Representative flow cytometric analysis of the frequency of SCs (CD45−/CD11b−/CD31−/Sca1−/Integrin-α7+/CD34+) subpopulation in Slug+/+ and Slug−/− mice. b Yield of SCs per milligram (mg) of muscle from Slug+/+ and Slug−/− mice (n = 6 mice per group). *p < 0.05 by student’s t-test. c Representative IHC images of Pax7+ SCs within the intact tibialis anterior (TA) muscles of Slug+/+ and Slug−/− mice. DAPI was used as nuclear counterstaining. Scale bar, 100 μm. d Quantification of Pax7+ SC numbers in c. *p < 0.05 by student’s t-test. e H&E staining of the intact and injured TA muscles in Slug+/+ and Slug−/− mice (n = 5 mice per group). For single injury, TA muscles were harvested at day 10 after BaCl 2 injection. For double injury, mice were recovered for 1 month after the first BaCl 2 injection at TA muscles. A second BaCl 2 injection was administered thereafter. TA muscles were harvested 10 days after the second BaCl 2 injection. Scale bar, 100 μm. f Quantification of the mean myofiber cross-sectional area (CSA, μm2) of the intact and BaCl 2 -injured TA muscles of Slug+/+ and Slug−/− mice. g Ratio of myofiber CSA in the intact and BaCl 2 -injured TA muscles between Slug+/+ and Slug−/− mice. **p < 0.01 by student’s t-test (n.s., not significant). All these experiments were independently repeated three times with similar results. Data are presented as mean ± SEM. Also see Supplementary Fig. 1. Source data are provided as a Source Data file Full size image

SCs is essential for the maintenance and regeneration of skeletal muscle. Thus, we determined how Slug null-induced increase of SCs affects muscle regeneration upon injury. H&E staining showed that Slug-deficient muscles regenerated as well as that of wild-types after a single injury (Fig. 1e–g). Strikingly, upon second injury, Slug-deficient muscles exhibited severely impaired regeneration when compared with wild-type counterparts (Fig. 1e–g). Collectively, these data demonstrated that Slug is essential for efficient muscle repair during continuous muscle regeneration, suggesting a key role of Slug in regulation of SC function.

SC-specific Slug loss impairs skeletal muscle regeneration

Skeletal muscle regeneration is a highly coordinated process involving the activation of various cellular and molecular responses15. We first decided to examine the expression pattern of Slug in SCs in view of their critical role in muscle regeneration. By comparing with several other muscle resident cell types including fibro-adipogenic progenitors (FAPs), pan-lymphocytes (LCs), and epithelial cells (ECs), in which the role of Slug is well characterized, we found that Slug is most highly expressed in quiescent SCs (Fig. 2a), and its expression was slightly reduced in activated SCs and markedly decreased after SCs were differentiated into myotubes (Fig. 2b).

Fig. 2 SC-specific Loss of Slug Impairs Muscle Regeneration. a Slug expression in different muscle resident cells. Left, representative flow cytometric gating of SCs (CD31−CD45−Scal1−Vcam I+), pan-lymphocytes (LCs, CD45+), epithelial cells (ECs, CD31+), and fibro-adipogenic progenitors (FAPs, CD31−CD45−Scal1+) from freshly prepared skeletal muscle cells. Right, qPCR analysis of Slug expression. *p < 0.05, **p < 0.01, ***p < 0.001 by student’s t-test. b Quantification of Slug expression in undifferentiated and differentiated SCs. Left, representative images of SCs and myotubes. Scale bar, 100 µm. Right, qPCR analysis of Slug expression. *p < 0.05, ***p < 0.001 by student’s t-test. QSC, quiescent satellite cell; ASC, activated satellite cells upon culture in growth medium for 3 days; MT, myotube. c Gene targeting strategy for generation of SC-specific Slug knockout mice. d Diagram of Slug-specific primer design for genotyping PCR. M. DNA marker; Ng negative control for PCR. e Comparison of adult Slugfl/flPax7Cre/+ (SlugcKO) and Slugfl/+Pax7Cre/+ (Ctrl). f Immunofluorescence staining of Slug in SCs of SlugcKO and Ctrl mice (n = 3 mice). Scale bar, 100 µm. g Frequency of SCs in SlugcKO and Ctrl mice. Similar results were seen from three independent flow cytometric analyses. h Yield of SCs per mg of muscle from Ctrl and SlugcKO mice (n = 3 mice for each genotype). *p < 0.05 by student’s t-test. i H&E staining of intact and injured TA muscles in SlugcKO and Ctrl mice (n = 5 mice per group). For single injury, TA muscles were harvested at day 10 after BaCl 2 injection. For consecutive injury, mice with primary injury were recovered for 1 month followed by a second BaCl 2 injection at the same sites. TA muscles were harvested 10 days after each injury. Scale bar, 100 μm. The experiment was repeated independently for three times with similar results. j Quantification of the myofiber CSA (µm2) shown in i. *p < 0.05 by student’s t-test (n.s., not significant). Data are shown as mean ± SEM of three independent replicates. Also see Supplementary Fig. 2 and 3. Source data are provided as a Source Data file Full size image

To determine whether the impaired muscle regeneration in global Slug knockout mice is a SC-driven defect, we generated SC-specific Slug knockout mouse line using the Cre/loxP system (Fig. 2a–d and Supplementary Fig. 2). Unlike the global Slug knockout mice, Slugfl/flPax7Cre/+ (designated as SlugcKO) mice showed no apparent differences with the control animals (Slugfl/+/Pax7Cre/+) in body size and weight (Fig. 2e). Furthermore, SCs from Ctrl but not SlugcKO mice displayed positive-staining for Slug protein, indicating that Slug is efficiently deleted in SCs in SlugcKO mice (Fig. 2f).

Next, we investigated the effect of SC-specific Slug deletion on the maintenance and regenerative capacity of SCs. Consistently, both the overall frequency and total yield of SCs calculated as per milligram of muscles were increased in SlugcKO mice (Fig. 2g, h). Although individual myofiber diameters of the intact TA and cross-sectional area of TA on day 10 post single BaCl 2 injury were not different between Ctrl and mutant littermates, more severely compromised muscle regeneration with a large increase of both necrotic fibers and fibrotic tissue was observed in SlugcKO mice administered with double and triple muscle damages (Fig. 2i, j). The impaired secondary but not primary muscle regeneration was also detected in Slugfl/flPax7CreER mice, in which Slug was deleted specifically in adult SCs by administering tamoxifen prior to muscle injury (Supplementary Fig. 3). Taken together, these results demonstrated that the presence of Slug in SCs is essential for SC-driven skeletal muscle regeneration, and Slug regulates muscle regeneration via its SC-specific function.

Slug-null SCs fail to replenish SC pool after activation

Maintaining SC pool size is crucial for constant muscle turnover/injuries and ongoing repair16. We showed that muscle regeneration after first injury was normal, indicating that Slug-deleted SCs in intact muscles were functional in terms of activation and differentiation upon injury. Indeed, primary Slug−/− SCs showed robust potential for differentiation upon in vitro induction and in vivo transplantation (Supplementary Fig. 4). However, the consequent regeneration became severely compromised in the absence of Slug (Figs. 1e, 2i and Supplementary Fig. 3e). These results prompted the question whether Slug-deficient SCs were capable of self-renewing and replenishing the stem cell pool after activation. SlugcKO displayed a sharp decrease in SC number of regenerated TA muscle when compared with Ctrl littermates after single and double injuries (Fig. 3a), indicating that lack of Slug attenuates ability of SCs to properly maintain the stem cell pool in repaired muscle after injury. This was confirmed by an observation of about three-fold decrease in SC number per 100 myonuclei in regenerated TAs of SlugcKO mice when compared with control mice (Fig. 3b, c).

Fig. 3 SCs deficient in Slug exhibit intrinsic defects in self-renewal in vivo. a Ratio of SCs between SlugcKO and Ctrl mice under indicated conditions. ***p < 0.001 by student’s t-test. b IHC for Pax7+ SCs in TAs of SlugcKO and Ctrl mice (n = 3 mice per group) at day 30 post single injury. Laminin indicates the boundaries of myofibers. Scale bar, 100 μm. Three independent experiments were performed with similar results. c Quantification of Pax7+ SC numbers in b **p < 0.01 by student’s t-test. d Scheme of SC transplantation. 3000 SCs of indicated genotype were transplanted into each side of pre-injured TA muscles of mdx recipients (n = 6), respectively. Donor-derived SCs (GFP+) in total recipient MuSCs (CD45−CD31−Sca1−Vcam I+) were analyzed 4 weeks after transplantation. e Representative flow cytometric analysis of the frequency of donor-derived SCs (GFP+) within total recipient MuSCs. Similar results were seen in three independent transplantation experiments. f Percent of donor-derived SCs (GFP+) in total recipient MuSCs. *p < 0.05 by student’s t-test. g Diagram of competitive MuSC repopulation assay. SCs from Slug+/+GFP- and Slug−/−GFP-transgenic mice were equally mixed for genomic DNA extraction and transplantation, respectively. The remaining cells were then transplanted into pre-injured TA muscles of mdx recipients. Four weeks after transplantation, GFP+ cells were sorted from total recipient MuSCs for genotyping PCR analysis. h Determination of relative ratio of repopulated MuSCs. Left, representative flow cytometric plotting of GFP+ cells from total recipient MuSCs. Right, PCR determining the ratio of SCs before and after transplantation. M DNA ladder, MBT mixed SCs before transplantation, MAT mixed SCs after transplantation, Ng negative control. i Scheme of AraC treatment indicating self-renewal. j Pax7 and Ki67 immunostaining in cultured myofibers as treated in i. Scale bar, 50 µm. Similar results were seen from three independent experiments. k Quantification of the quiescent daughter SCs stained in j (n = 3 mice, >20 myofibers per condition). ***p < 0.001 by student’s t-test. Data are shown as mean ± SEM of three independent experiments. Unprocessed gel blots are provided in the Supplementary Fig. 1. Also see Supplementary Fig. 4-6. Source data are provided as a Source Data file Full size image

Injection of tractable adult SCs into pre-injured adult muscle followed with fluorescence-activated cell sorting (FACS) analysis provides a quantitative assay for SC self-renewal17. By this assay, we analyzed the stem cell repopulation of donor-derived SCs (GFP+) as a fraction of the total SC population from the primary recipients (Fig. 3d). Because a small portion of SCs remain cycling 1 month after transplantation18, we analyzed SCs with distinct immunophenotype (CD31−CD45−Sca1−Vcam I+), which consist of both quiescent and activated MuSCs19. Via this analysis, we found that the frequency of Slug−/− SCs was about three-fold lower than that of Slug+/+ SCs (Fig. 3e, f). Since the compromised self-renewing capability of SCs from Slug−/−GFP mice may be due to embryonic absence of Slug, we induced an acute Slug knockout in SCs from Slugfl/flPax7-zsgreen mice by infecting with retrovirus expressing Cre recombinase (Supplementary Fig. 5a,b). Cre-expressing SCs yielded about five-fold less GFP+ fraction in the total SC population from the recipients compared to that of Ctrl virus-infected SCs (Supplementary Fig. 5c,d). Consistently, by zsgreen staining we identified considerably less Cre retrovirus-infected SCs in the SC niche, beneath the basal lamina and atop myofibers (p < 0.01 by student’s t-test) (Supplementary Fig. 5e,f). These data indicated that deletion of Slug decreases SC self-renewing and regenerative ability.

Competitive repopulation assay has been widely used as the gold standard for testing self-renewal capacity of hematopoietic stem cells20. Here, we adopted a similar competitive repopulation assay for SCs (Fig. 3g). Technically, this assay overcomes the potential variations due to different SC niches in individual recipients and uneven injection of myotoxins and SCs at different sites. By this assay, we showed that although donor SCs from Slug+/+GFP and Slug−/−GFP mice were equally mixed before transplantation, only the DNA band corresponding to wild-type alleles of Slug was predominantly amplified from the gDNA of the donor-derived SC mixtures in recipients (Fig. 3h and Supplementary Fig. 6), indicating a markedly attenuated self-renewing capability in Slug−/− SCs.

To further assess the intrinsic self-renewing propensity of Slug null SCs, we performed myofiber-associated SCs culture in the presence of arabinosylcytosine (AraC) that eliminates cycling cells by inhibiting DNA synthesis. It was previously demonstrated that a small population of AraC-resistant, myofiber-associated Pax7+ cells arose following the first SC division in culture by self-renewing and behaved as quiescent SCs6,21. We treated myofibers with AraC from day 3–5 after isolation (Fig. 3i) and detected a lower number of surviving Pax7+Ki67− SC daughters from SlugcKO mice compared to those from Ctrl mice (Fig. 3j, k). Together, these results provide convincing evidence that Slug-deficient SCs have an intrinsic defect in self-renewal following muscle regeneration.

Slug directly represses p16 Ink4a transcription in SCs

To explore intrinsic factors responsible for losing self-renewal capacity of activated SCs in Slug-deficient mice, we initially retrieved and interrogated the global gene expression data (GEO accession: GSM38236) generated from Slug-silenced primary myoblasts22. Gene ontology enrichment analysis of biological processes (GOBP) identified that Slug silencing derepressed sets of genes related to transcription regulation, cell proliferation, and skeletal muscle differentiation processes (Fig. 4a, left panel). Notably, the genes downregulated upon Slug silencing identified enrichment of genes among various categories of cellular defense responses (Fig. 4a, right panel). Interestingly, p16Ink4a was listed in the most upregulated genes ranked in negative regulation of cell proliferation. It was previously reported that self-renewal capacity of aged stem cells (hematopoietic stem cells23, intestinal stem cells24, and skeletal MuSCs5) was attenuated. Therefore, we postulated p16Ink4a as a potential mediator for SC self-renewing defect caused by Slug deficiency.

Fig. 4 p16Ink4a is a direct target gene of Slug in SCs. a GOBP analysis of genes modulated in Slug-silenced myoblasts (GEO accession: GSE38236). The term of negative regulation of cell proliferation being highly relevant to observed self-renewal defect of Slug-deficient SCs in our study was highlighted in red. b Hierarchical clustering and heatmap representation of 168 differentially expressed genes separating Slug+/+ and Slug−/− SCs. Color pattern represents row Z-score. c Bubble chart showing results of GOBP analysis. Bubble size indicated number of genes associated with each term. GOBP ranked by fold enrichment score associated with upmodulated (green bubbles) and downregulated (red bubbles) signatures were selected for significance by using a false discovery rate cutoff of 5%. d qPCR analysis of p16Ink4a and p19Arf in primary SCs from Slug+/+ and Slug−/− mice (n = 3). n.s. not significant; ***p < 0.001 by student’s t-test. e Schematic diagram of the INK4a/ARF. The consensus Slug-binding site (E-box) is located in its promoter. f ChIP analysis of Slug occupancy at the p16Ink4a promotor. Primers targeting the 3’ untranslated region (without E-box elements) was used as a negative control for the ChIP assay. Relative binding affinity of Slug on the putative E-box element was quantified relative to the non-specific binding. ***p < 0.001 by student’s t-test. g Quantification of p16Ink4a in quiescent, ex-vivo culture- and in vivo injury-activated SCs from Slug+/+ and Slug−/− mice. For activation ex-vivo, SCs were cultured for 7 days; while for activation in vivo, SCs were harvested at day 10 after injury. *p < 0.05, ***p < 0.001 by student’s t-test. h IHC for p16Ink4a and Pax7 protein in SCs within the TA muscles of SlugcKO and Ctrl mice at day 30 post injury (n = 3 mice per genotype). Scale bar, 100 μm. i Percentage of Pax7 and p16Ink4a double positive SCs in h. ***p < 0.001 by student’s t-test. The experiments g–i were independently repeated three times with similar results. Data are shown as mean ± SEM of three independent experiments. Also see Supplementary Fig. 7-11. Source data are provided as a Source Data file Full size image

We then performed microarrays analysis to compare the genome-wide gene expression profiles of wild-type and Slug-deficient SCs. Unsupervised hierarchical clustering analysis separated the samples into their respective genotypes (Fig. 4b). We identified 168 differentially expressed genes, of which 108 genes were upregulated (fold change > 2, p value < 0.05) and 60 genes were downregulated (fold change < −2, p value < 0.05) in Slug−/− SC compared to control cells (Fig. 4b). Slug deletion enriched GO categories related to mitochondria metabolism and cell cycle regulators (Fig. 4c). To identify pathways enriched in Slug−/− SCs, gene set enrichment analysis (GSEA)25 was performed. Notably, signaling pathways involved in cell metabolism including glycolysis, PI3K-AKT-MTOR, oxidative phosphorylation, and reactive oxygen species were induced in quiescent Slug−/− SCs (Supplementary Fig. 7a,b), suggesting a switched metabolic reprogramming with relatively higher energy-consuming status in Slug null SCs. Meanwhile, E2F targets and G2M checkpoints signaling signatures were also enriched upon Slug deletion (Supplementary Fig. 7c,d). These gene sets enrichment data indicated that loss of Slug in SCs might disturb cell cycle progression and the balance between self-renewal and differentiation after activation.

Next, we examined expression of p16Ink4a and p19Arf, the two gene products of the INK4A/ARF locus, by qPCR in SCs of adult wild-type and Slug−/− mice, respectively. As shown in Fig. 4d, p16Ink4a was about two-fold higher in Slug-deficient SCs when compared with the wild-type counterparts, indicating a derepression of p16Ink4a in the absence of Slug in resting SCs in vivo. In contrast, there was no apparent difference in p19Arf expression between the two types of SCs. Furthermore, we examined the proximal promoter region of p16Ink4a and identified potential Slug-binding sites (E-box)26,27 (Fig. 4e). To facilitate assessing the occupancy of endogenous Slug at the promoter region of p16Ink4a by chromatin immunoprecipitation (ChIP) assay, we performed Slug affinity tagging at its C-terminus in mouse myoblasts by CRISPR/Cas9-mediated gene tagging (Supplementary Fig. 8). ChIP-qPCR analysis displayed enriched binding in anti-Flag antibody immunoprecipitated gDNA fragments but not in anti-IgG control (Fig. 4f), indicating that Slug occupies the promoter region of p16Ink4a in vivo. Such direct regulation of p16Ink4a promoter by Slug was further confirmed by p16Ink4a-driven luciferase reporter assay in SC-derived myobalsts (Supplementary Fig. 9a,b). Importantly, this E-box element is also present in human p16INK4A promoter (Supplementary Fig. 9c). Knockout or overexpression of SLUG in primary human myoblasts significantly derepressed or suppressed p16INK4A transcript (p < 0.001 by student’s t-test) (Supplementary Fig. 9d-g), respectively, indicating a highly conserved role of Slug in regulating p16Ink4a.

Of note, loss of Slug caused a more robust dysregulation of p16Ink4a in ex vivo cultured myoblasts22 compared to resting SCs in vivo. Normally, p16Ink4a expression is elevated during aging and replicative senescence28. Myoblasts are a type of proliferating myogenic progenitor cells derived from quiescent SCs under stimulating conditions. Therefore, we asked whether derepression of p16Ink4a in Slug-deficient SCs would be exacerbated under stress conditions, including ex vivo culture and muscle damage. Strikingly, p16Ink4a expression increased by over 8-fold in ex vivo cultured Slug−/− SCs and five-fold in in vivo injury-activated Slug−/− SCs, respectively, when compared with their corresponding wild-type controls (Fig. 4g) Notably, derepression of p16Ink4a in cultured myoblasts was accompanied with graduate decline of Slug expression (Supplementary Fig. 10).

In spite of the increased p16Ink4a transcription, primary muscle regeneration was normal in Slug-deficient mice. This is a departure from what was reported in geriatric mice model, i.e., that resting SCs fail to activate and expand to regenerate the muscle on injury due to derepression of p16Ink4a5. We suspected that p16Ink4a mRNA but not protein was increased in resting Slug-deficient SCs, since p16Ink4a mRNA could be induced to decay by an RNA-binding protein in early passage of fibroblasts but accumulated at protein level during late-passage of culture28. Indeed, p16Ink4a protein was undetectable in undamaged muscle cryosections from both Ctrl and SlugcKO mice (Supplementary Fig. 11). Instead, there was an apparent co-expression of p16Ink4a with Pax7 in muscle tissue harvested from SlugcKO mice 30-day post injury (Fig. 4h, i). Taken as whole, these results suggested that Slug deficiency leads to an increase in p16Ink4a transcription in SCs, and replicative stress signaling triggered by SC activation and proliferation concurrently increases the stability of p16Ink4a protein.

Slug loss promotes senescence in SCs during proliferation

In terms of the elevated p16Ink4a protein in activated SCs in regenerating skeletal muscles of Slug-deficient mice, we hypothesized that Slug-ablated SCs acquired features of cellular senescence at late stages of regeneration during the transition of SCs from activation to quiescence or differentiation. To test this notion, we first used an in vitro reserve cell culture system29. As shown in Fig. 5a, reserve cells from wild-type mice robustly proliferated while cells from Slug−/− mice were only sporadically distributed by day 7 of subculture (Fig. 5a). Compared to their wild-type control cells, significantly higher proportion of progeny from Slug−/− reserve cells was positive for p16Ink4a staining (p < 0.001 by student’s t-test) (Supplementary Fig. 12). SA-β-Gal staining showed that over 50% of Slug-deficient reserve cells-expanded cells was SA-β-Gal+, a marker of senescence, whereas few SA-β-Gal+ cells were detected in the control group (Fig. 5a, b).

Fig. 5 Slug-deficient SCs acquire senescence properties during self-renewal. a SA-β-Gal staining. SCs of indicated genotype were differentiated for 21 days. 5000 zsGreen+ reserve cells were sub-cultured for another 7 days and subject to SA-β-Gal staining. Red arrows indicate SA-β-Gal+ cells. Scale bar, 100 µm. b Quantification of the percentage of SA-β-Gal+ cells stained in a. ***p < 0.001 by student’s t-test. c Cumulative population doubling level (CPDL) obtained in cultures of primary SC-derived myoblasts from Slug+/+ and Slug−/− mice. d Photographs (left) and SA-β-Gal staining (right) of SC-derived myoblasts on day 7 of culture at passage 3. Scale bar, 200 µm (left); 100 µm (right). e Percentage of the SA-β-Gal+ cells in d. **p < 0.01 by student’s t-test. f Representative images of H&E (upper) and SA-β-Gal staining (lower) on transverse TA muscle cryosections. TA muscles from SlugcKO and Ctrl mice (n = 3 per genotype) were harvested on day 10 post injury and subject to H&E and SA-β-Gal staining. Red arrows indicated SA-β-Gal+ cells. The window (lower panel, right picture) represents high magnification of dotted boxed area. Scale bar, 50 µm. g Quantification of the SA-β-Gal+ cell numbers stained in f. **p < 0.01 by student’s t-test. h Senescent SCs in injured mice of SlugcKO and Ctrl mice. SA-β-Gal staining was combined with IHC staining against Pax7 and Ki67 on TA muscle cryosections being treated as described in f. Scale bar, 50 µm. i IHC staining for Pax7 and Ki67 in TA muscle sections harvested at day 2.5 post the second BaCl 2 injury (n = 3 mice per group). Arrows indicated SCs in the section. Scale bar, 100 μm. j Percentage of cycling (Pax7+Ki67+) SCs stained in i. ***p < 0.001 by student’s t-test. Data are presented as mean ± SEM of three independent experiments. Also see Supplementary Fig. 12. Source data are provided as a Source Data file Full size image

Population doubling level (PDL) is an intrinsic measurement of the age of the particular culture of a cell line. In culture, an untransformed cell line has a finite life span expressed in the number of cumulative population doublings that can be achieved. To assess the accelerated senescence in Slug-deficient SCs under proliferative pressure in vitro, we determined the relative growth rates of control and Slug-mutant SCs by calculating the cumulative PDL. As shown in Fig. 5c, the growth rate of Slug-deficient SCs was clearly retarded at early passage 3 when wild-type SCs remained for exponential growth. SA-β-Gal staining demonstrated that about 40% of Slug-ablated myoblasts show strong positive SA-β-Gal staining while less than 5% of control cells were SA-β-Gal+ on passage 3 (Fig. 5d, e), indicating the existence of replicative senescence and growth retardation in early-passaged Slug-deficient myoblasts.

Next, we investigated whether cellular senescence in SCs would occur when Slug is specifically deleted in Pax7+ SCs during regeneration in vivo. Young adult mice were used in this study to exclude the process of regeneration from aging in geriatric mice5. We performed SA-β-Gal staining on cryosections of TA muscles from Ctrl and SlugcKO mice on day 10 post first injury when necrotic fibers were replaced by central-nucleated regenerated myofibers (Fig. 5f). Notably, despite the sporadic SA-β-Gal+ cells in control TA muscle sections, there was a four-fold increase in the number of SA-β-Gal+ cells in SlugcKO TA (Fig. 5f, g). Most of the SA-β-Gal+ cells were located beneath basal lamina and outside myofiber plasma membrane, a classical SC anatomical location (Fig. 5f). IHC results further demonstrated that these SA-β-Gal+ cells were also Pax7-positive. As expected, SA-β-Gal+Pax7+ cells were not in cycling (Ki67−), suggesting the status of cellular senescence. In contrast, SCs with negative SA-β-Gal staining (SA-β-Gal−/Pax7+) were on the occasion of proliferating (Ki67+). In agreement with this notion, the majority of SCs in SlugcKO mice failed to re-activate as indicated by markedly lowered percentage of Ki67+ SCs by 2.5-day post second BaCl 2 injury compared to Ctrl mice (Fig. 5i, j). Together, using multiple in vitro and in vivo assays we proved that lack of Slug facilitated entry of SCs into cellular senescence under proliferative pressure.

p16Ink4a loss restores impaired self-renewal of Slug −/− SCs

We demonstrated elevated p16Ink4a protein and acquired senescence feature in activated Slug-deficient SCs. At this point, a key question was whether p16Ink4a is causally involved in Slug loss-induced defects in SC self-renewing capacity and muscle regeneration. To answer this question, we assessed muscle regeneration in Slug+/+p16+/+, Slug−/−p16+/+, and Slug−/−p16−/− mice following serial damages. By H&E staining, we showed that although muscle regeneration was severely compromised in TAs of Slug−/−p16+/+ mice following double or triple injuries, muscle repair was greatly improved in Slug−/−p16−/− mice after injuries (Fig. 6a, b).

Fig. 6 Removal of p16Ink4a partially rescues regeneration and self-renewal of Slug-deficient SCs. a H&E staining of the double and triple BaCl 2 -injured TA muscles from Slug+/+p16+/+, Slug−/−p16+/+, and Slug−/−p16−/− mice (n = 3–6 mice per group). Left panel, the diagram for consecutive injury. Right panel, representative images of H&E staining of muscle tissue sections from injured mice. Scale bar, 100 μm. b Ratio of myofiber CSA in the intact and BaCl 2 -injured TA muscles between Slug+/+p16+/+, Slug−/−p16+/+, and Slug−/−p16−/− mice. *p < 0.05, ***p < 0.001 by one-way ANOVA; n.s. not significant. c Scheme of SC transplantation. SCs isolated from Slug−/−p16+/+ and Slug−/−p16−/− mice were infected with GFP-expressing retroviruses, and injected into either side of the BaCl 2 -pre-injured TA muscle of mdx recipient, respectively. Total mononucleated muscle cells were isolated separately from either TA muscle 4 weeks after transplantation, and subjected to flow cytometric analysis for the fraction of donor-derived SCs (GFP+) in MuSC (CD45−CD31−Sca1−Vcam I+) subpopulation of recipient mice. d Representative flow cytometry plots showing the frequency of donor-derived SCs (GFP+) within the total recipient MuSC (CD45−CD31−Sca1−Vcam I+) subpopulation in mononucleated TA muscle cells. e Percent of donor-derived cells (GFP+) in total MuSC (CD45−CD31−Sca1−Vcam I+) subpopulation of mononucleated TA muscle cells from mdx recipients (n = 4–6). ***p < 0.001 by student’s t-test. f Representative SA-β-Gal staining of cultured primary SC-derived myoblasts. 104 SCs of indicated genotype were plated in Matrigel-coated wells of 24-well plate, and passaged weekly. Bright-field imaging and SA-β-Gal staining were performed in cells on day 7 of culture at passage 3. Scale bar, 100 µm. g Percentage of the SA-β-Gal+ cells in serially passaged myoblasts on day 7 of culture at passage 3. ***p < 0.01; by student’s t-test. n.s. not significant. Data are shown as mean ± SEM of three independent replicates. Source data are provided as a Source Data file Full size image

Next, we assessed how removal of p16Ink4a would affect the self-renewing ability of Slug-ablated SCs by the quantitative repopulating assay (Fig. 6c). As expected, the frequency of Slug−/−p16+/+ donor-derived SCs was low (~1.96%) (Fig. 6d, e). However, the frequency of Slug−/−p16−/− donor-derived SCs was increased by two-fold in the recipients (Fig. 6d, e). Mechanistically, enhanced repopulation of Slug−/−p16−/− SC in recipient muscles after deletion of p16Ink4a might be partially ascribed to reduced senescence under proliferative stress, as Slug−/−p16−/− SC-derived myoblasts displayed little or no SA-β-Gal staining at ex vivo passage 3 when majority of Slug−/−- cells were SA-β-Gal positive (Fig. 6f, g). In summary, p16Ink4a is a crucial mediator for the defects in SC self-renewal and muscle regeneration caused by Slug deficiency.

Slug overexpression restores self-renewal of aged SCs

Aging alters stem cell function in terms of the capacities of self-renewal, proper activation and/or proliferation as well as lineage commitment. Aged SCs were characterized by self-renewal defect6,7,16, inclined commitment of differentiation in low mitogen ex vivo culture condition3,6, and susceptibility to senescence upon mitogen exposure7,30. These aging-associated phenotypes resemble to some extent behaviors of Slug-deficient SCs in current study. Such similarities drew our attention to the role of Slug in SC aging. By interrogating the transcriptomes of young and aged SCs from different groups of mice4,31, we observed an age-associated downregulation of Slug in aged SCs (Fig. 7a). This is further confirmed in SCs from young and aged mice by PCR analysis (Fig. 7b–d).

Fig. 7 Slug Overexpression Restores Repopulating Capacity of Aged SCs. a Volcano plots demonstrating differentially expressed genes between freshly isolated young and aged SCs from the microarray data (GEO accession: GSE50821 and GSE72179) of indicated publications. Green dots indicate substantially increased while red dots indicated substantially decreased genes in aged SCs compared with young SCs. b Flow cytometric plotting of MuSCs (CD45−CD11b−CD31−Sca1−Integrin-α7+CD34+) in young and aged mice. c Qualitative RT-PCR showing up-regulation of p16Ink4a in aged SCs. HPRT was used as internal control for PCR. d qPCR analysis of relative mRNA levels of Slug in SCs from the young and aged mice (n = 3 mice per group). **p < 0.01 by student’s t-test. e Scheme of SCs repopulation experiment. SCs were passaged weekly. After 2 weeks of culture, SC-derived myoblasts were infected with GFP-control (pMIGR1) or Slug-expressing (pMIG-Slug) retroviruses, and then transplanted into either side of pre-injured TA muscle of mdx recipient. Total mononucleated muscle cells were isolated separately from either TA muscle 4 weeks after transplantation, and subjected to flow cytometric analysis for the frequency of donor-derived SCs (GFP+). f Quantification of relative mRNA levels of p16Ink4a in quiescent and cultured primary mouse SCs with or without overexpressing Slug. *p < 0.05, ***p < 0.001 by one-way ANOVA. g Representative flow cytometric analysis of the fraction of donor-derived SCs (pMIGR1 or pMIG-Slug) within the total recipient MuSC (CD45−CD31−Sca1−Vcam I+) subpopulation in mononucleated TA muscle cells. h Percent of donor-derived cells (GFP+) in total recipient MuSC (CD45−CD31−Sca1−Vcam I+) subpopulation of TA mononucleated muscle cells from mdx recipients (n = 3–4). ***p < 0.001 by student’s t-test. i Representative dystrophin immunostaining (Scale bar, 50 µm) on TA muscles transplanted with pMIGR1 or pMIGR1-Slug retrovirus-infected myoblasts. j Quantification of dystrophin-expressing myofibers in TA muscles of recipient mdx mice (n = 6 mice per group) shown in i. ***p < 0.001, Student’s t-test. Data are shown as mean ± SEM of three independent replicates. Unprocessed gel blots are provided in the Supplementary Fig. 14 and source data file. Also see Supplementary Fig. 13. Source data are provided as a Source Data file Full size image

We demonstrated that there was an increased bioenergetic requirement in Slug null SCs (Supplementary Fig. 7a). Interestingly, GSEA revealed a similarly altered metabolic reprogramming in old SCs (GSE81096), which was evidenced by activation of genes ranked in glycolysis and mitochondrial metabolism in MTORC1, oxidative phosphorylation, PI3K-Akt-MTOR, and reactive oxygen species signaling pathways (Supplementary Fig. 13a). Of note, 71 genes were overlapped in these metabolic pathways of both Slug-deleted and aged SCs (Supplementary Fig. 13b). In addition, E2F targets and G2M checkpoints signatures being activated in Slug−/− SCs were also enriched in old SCs in comparison to the young SCs (Supplementary Fig. 13c). Of those cell cycle regulators being upregulated in Slug−/− SCs, 29 of which were activated in aged SCs as well (Supplementary Fig. 13d). Together, these data indicated that Slug insufficiency might be an important factor in causing multiple regenerative defects in aged SCs.

It has been reported that replicative senescence can play a role in the regenerative defects during normal skeletal muscle aging32 and replicative senescence may be caused by p16Ink4a stress pathway33. Next, we applied ex vivo cultured SCs as a model to address whether forced expression of Slug could restore the intrinsic regenerative and self-renewing capacities of the aged SCs (Fig. 7e). Similar to what found in the geriatric SCs (Fig. 7c), a sharp increase in p16Ink4a expression was found in cultured SCs (Fig. 7f). Forced expression of Slug significantly suppressed stress-induced p16Ink4a (p < 0.001 by student’s t-test) (Fig. 7f). Furthermore, we demonstrated that Slug-overexpressing myoblasts gave rise to a substantially larger fraction of the total MuSC population in mdx recipients than myoblasts transduced with the control retrovirus (Fig. 7g, h). In addition to the self-renewal capacity, Slug overexpression also largely restored regenerative capability of SCs that underwent passaging and subculturing for weeks, as evidenced by an increase in the number of dystrophin-positive myofibers in mdx recipient (Fig. 7i, j).

Taken as a whole, these findings demonstrated that lack of sufficient Slug expression is an important factor for derepression of p16Ink4a in aged SCs, and that restoring Slug expression improves p16Ink4a-caused regenerative and self-renewing defects in SCs.