Senescent‐Like Myofibers Contribute to Anti‐Regenerative Cytokine Signaling in Duchenne Muscular Dystrophy

ABSTRACT

Duchenne muscular dystrophy (DMD) is a genetic muscular disease characterized by progressive muscle degeneration. p16 is expressed in skeletal muscles and induces cellular senescence in a rat model of DMD, whereas its ablation enhances muscle regeneration. However, the mechanism underlying this phenomenon remains unclear. This study aimed to elucidate the mechanism for p16‐induced DMD exacerbation. RNA‐seq analysis revealed p16‐dependent upregulation of cytokine gene expression in DMD rat skeletal muscles, which also altered the systemic blood cytokine profile. Furthermore, the effect of an altered humoral environment on muscle regeneration was assessed using the transplanted extensor digitorum longus muscle. Regeneration of grafted muscles from wild‐type rats was suppressed in DMD rats but was significantly improved by p16 ablation. Notably, p16 was expressed in the myofibers of DMD rats, and enzymatically isolated myofibers from DMD rats also showed p16‐dependent cytokine expression. Thus, cytokines secreted by senescent‐like myofibers mediate the anti‐regenerative niche in DMD rats, uncovering a novel mechanism for disease progression and potential therapeutic targets.

p16 expression in DMD model rats induces a senescent‐like state in myofibers accompanied by p16 and cytokine gene expression, reshaping systemic humoral environments. This cytokine‐rich milieu suppresses skeletal muscle regeneration, as demonstrated by impaired regeneration of transplanted wild‐type muscle. Genetic ablation of p16 attenuates cytokine gene expression, and markedly improves muscle regeneration, revealing a myofiber‐driven, cytokine‐mediated mechanism underlying DMD progression.

1. Introduction

The skeletal muscle is composed of multinuclear myofiber bundles. In mammals, myofibers are postmitotic, and mononuclear satellite cells contribute to myofiber regeneration upon muscle damage [1]. As myofibers degenerate, macrophages infiltrate the damaged site to phagocytose cellular debris, whereas quiescent satellite cells are activated to form myoblasts. Subsequently, proliferated myoblasts differentiate and fuse to form multinuclear myotubes. Finally, the myotubes mature to regenerate terminally differentiated myofibers.

Duchenne muscular dystrophy (DMD) is a devastating genetic disease caused by out‐of‐frame mutations in dystrophins. In patients with DMD, the deficiency of dystrophin, a component of the dystrophin‐glycoprotein complex which connects the cytoskeleton to the basement membrane, results in repeated degeneration and regeneration of myofibers, causing progressive muscle weakness. In DMD, muscle regeneration is progressively impaired, and muscle tissue is substituted by extensive fibrosis and fat accumulation [2]. We previously reported that in a rat model of DMD, p16 expression is upregulated with age, and cellular senescence is induced in skeletal muscles [3].

Cellular senescence is an irreversible cell cycle arrest that occurs in response to damaging stimuli. Senescent cells accumulate in older individuals and in progressive disease models [4]. p16, encoded by CDKN2A, is a cell cycle inhibitor that binds and inhibits cyclin‐dependent kinases (CDK) 4 and 6 [5]. CDKN2A also encodes p19^ARF, which induces cellular senescence via activation of another CDK inhibitor, p21. p16 plays a key role in inducing cellular senescence, and inhibiting cellular senescence by knockout of the p16 gene in DMD rats significantly alleviates the progressive decline in muscle regenerative capacity and inhibits fibrosis and fat accumulation in the skeletal muscles of DMD rats, thereby improving muscle strength [3].

However, the mechanism by which p16 expression impairs muscle regeneration and exacerbates DMD remains unclear. One possibility is via the depletion of satellite cells, which are responsible for muscle regeneration. Cell cycle arrest induced by p16 is known to limit the activation and self‐renewal of satellite cells in old mice [6]. Satellite cells in rat models of DMD also express p16, suggesting that impaired regeneration may be due to cell cycle arrest in myogenic cells [3, 7]. In contrast, senescent cells can secrete factors such as pro‐inflammatory and pro‐fibrotic cytokines, which are called senescence‐associated secretory phenotypes (SASP). The removal of senescent cells has improved muscle regeneration in wild‐type (WT) mice, mdx mice, and DMD rats [3, 8, 9], although conflicting evidence indicates that senescent cells may also contribute to regeneration [10, 11]. Therefore, the presence of senescent cells can negatively affect muscle regeneration, suggesting that a mechanism other than stem cell depletion may be involved in the p16‐induced exacerbation of DMD in rats.

In the present study, we aimed to elucidate the mechanism of p16‐induced DMD exacerbation.

2. Materials and Methods

2.1. Animals and Treatments

WT, DMD, p16‐null DMD (dKO) rats [3, 12], and Becker muscular dystrophy (BMD) model rats [13] were generated on a Wistar‐Imamichi background. GFP rats (LEW‐Tg [CAG‐EGFP] 1Ys; NBRP Rat No. 0297) were obtained from the National BioResource Project for Rats in Japan and backcrossed with Wistar‐Imamichi rats. The rats were maintained under controlled environmental conditions, at 23°C, with a light/dark (12/12 h) cycle, and ad libitum access to food and water. All animal experiments were performed in accordance with the Guide for the Care and Use of Laboratory Animals of the University of Tokyo and were approved (P18‐125) by the Institutional Animal Care and Use Committee of the University of Tokyo. The study was conducted in compliance with the Animal Research: Reporting of In Vivo Experiments guidelines.

For EDL (extensor digitorum longus) transplantation, EDL muscles of 9‐week‐old Wistar‐Imamichi rats purchased from the Institute for Animal Reproduction (Ibaraki, Japan) were excised from both tendons under isoflurane anesthesia and transplanted as previously described by Lawson‐Smith and McCeach [14]. The excised EDLs were grafted on the bilateral tibialis muscles of 8 to 10‐month‐old WT, DMD, dKO, and 5‐month‐old GFP rats under medetomidine‐midazolam‐butorphanol or isoflurane anesthesia. The tendons of the transplanted EDLs were fixed to the host patellar tendon or tibialis anterior muscle tendon. The transplanted EDLs were dissected from euthanized rats after 1, 3, 6, 10, 14, and 30 d of transplantation. Samples without histologically detectable regeneration after day 10 were excluded from the analysis (three out of nine WT, one out of eight DMD, and three out of six dKO rats were excluded on day 10, and two out of eight WT rats were excluded on days 14 and 30).

To induce skeletal muscle injury, 0.75% bupivacaine hydrochloride (BPVC; Sigma‐Aldrich, MO, USA)/saline was injected into the tibialis anterior muscle under isoflurane anesthesia. The tibialis anterior muscles were dissected from the euthanized rats 14 days after BPVC injection. Uninjected contralateral tibialis anterior muscles were used as intact controls.

For histological analysis, EDL and tibialis anterior muscles were frozen in isopentane, chilled with liquid nitrogen, and the frozen sections (7 μm thick) were used for hematoxylin and eosin staining, immunostaining, and in situ hybridization (ISH).

2.2. Semi‐Quantitative Reverse Transcription‐PCR

The excised tissue was homogenized in TRIzol reagent (Thermo Fisher Scientific), and RNA was extracted. The RNA was reverse transcribed into cDNA using SuperScript II reverse transcriptase (Thermo Fisher Scientific). Quantitative reverse transcription‐PCR (qPCR) was performed on a Light Cycler 2.0 (Roche, Basel, Switzerland) or Step One Plus Real‐Time PCR system (Thermo Fisher Scientific) using Thunderbird SYBR qPCR Mix (TOYOBO, Osaka, Japan). The following primer sets were used:

Ccl2 5′‐CGT GCT GTC TCA GCC AGA T‐3′ (forward) and 5′‐GGA TCA TCT TGC CAG TGA ATG‐3′ (reverse) (NM_031530.1; 135–205).

Cxcl1 5′‐CCA AAC CGA AGT CAT AGC CAC‐3′ (forward) and 5′‐TCC ATT ACT TGG GGA CAC CC‐3′ (reverse) (NM_030845.2; 231–346).

Il1b 5′‐TGT GAT GAA AGA CGG CAC AC‐3′ (forward) and 5′‐CTT CTT CTT TGG GTA TTG TTT GG‐3′ (reverse) (NM_031512.2; 617–686).

Il6 5′‐CCT GGA GTT TGT GAA GAA CAA CT‐3′ (forward) and 5′‐GGA AGT TGG GGT AGG AAG GA‐3′ (reverse) (NM_012589.2; 433–574).

Ccn2 5′‐GGT GAC CTA GAG GAA AAC ATT AAG A‐3′ (forward) and 5′‐CCG GTA GGT CTT CAC ACT GG‐3′ (reverse) (NM_022266.2; 950–1058).

Mmp2 5′‐CAC CAC CGA GGA TTA TGA CC‐3′ (forward) and 5′‐CAC CCA CAG TGG ACA TAG CA‐3′ (reverse) (NM_031054.2; 1245–1315).

Tgfb1 5′‐CCT GGA AAG GGC TCA ACA C‐3′ (forward) and 5′‐CAG TTC TTC TCT GTG GAG CTG A‐3′ (reverse) (NM_021578.2; 937–1023).

p16 (exon 1α) 5′‐TTC ACC AAA CGC CCC GAA CA‐3′ (forward) and 5′‐CAG GAG AGC TGC CAC TTT GAC‐3′ (reverse) (NM_031550.2; 129–210).

p16 (3′ UTR) 5′‐AAG TTA GGC CTC AGC CCT CCT TTT T‐3′ (forward) and 5′‐AAA GCC ATA TGC TAG TCT CGC GTT‐3′ (reverse) (NM_031550.2; 524–593).

p19 5′‐GTG TTG AGG CCA GAG AGG AT‐3′ (forward) and 5′‐TTG CCC ATC ATC ATC ACC T‐3′ (reverse) (AY679727.1; 159–230).

p21 5′‐GAC ATC TCA GGG CCG AAA‐3′ (forward) and 5′‐GGC GCT TGG AGT GAT AGA AA‐3′ (reverse) (NM_080782.6; 549–610).

Hprt1 5′‐CCA TCA CAT TGT GGC CCT CT‐3′ (forward) and 5′‐TAT GTC CCC CGT TGA CTG GT‐3′ (reverse) (NM_012583.2; 243–408).

2.3. RNA‐Seq

Total RNA was extracted from the tibialis anterior muscles of 9‐month‐old WT, DMD, and dKO rats, and 3‐ and 18‐month‐old WT rats using the RNeasy Micro Kit (Qiagen, Venlo, Netherlands). RNA sequencing library preparation and sequencing were performed by BGI Genomics (Shenzhen, China). Briefly, cDNA libraries were prepared using the TruSeq RNA Library Prep Kit (Illumina, San Diego, CA, USA) and sequenced on an Illumina HiSeq 4000 (Illumina) with paired‐end reads of 100 bp. The sequenced reads were processed using Trimmomatic [15] (version 0.39‐1b), followed by quality control using Fast QC. The reads were then mapped to mRatBN7.2 (Ensembl release 108) using STAR [16] (version 2.7.10b). Gene expression levels were quantified using the RSEM [17] (version 1.3.3). The transcripts per million values generated by RSEM were scaled and used for principal component analysis (PCA) and heat map generation. Raw count data generated by RSEM were imported into the DESeq2 [18] R package to calculate p‐values and log2 fold change. Genes were ranked by log2 fold change, and gene set enrichment analyses (GSEA) of Kyoto Encyclopedia of Genes and Genomes (KEGG) were performed using the clusterProfiler [19] R package.

2.4. Cytokine Array

Blood was collected from the abdominal aorta of 9‐month‐old WT and DMD rats. Serum cytokine levels were evaluated using a Cytokine Array‐Rat Cytokine Antibody Array (Membrane, 34 Targets) (ab133992; Abcam, Cambridge, UK).

2.5. Immunohistochemistry

For immunostaining, the following primary antibodies were used: anti‐laminin (1:200, rabbit polyclonal, L9393; Sigma‐Aldrich), anti‐embryonic myosin heavy chain (eMHC; 1:100, mouse monoclonal, F1.652; Developmental Studies Hybridoma Bank, IA, USA), anti‐CD68 (1:100, mouse monoclonal, ED1; Bio‐Rad, CA, USA), anti‐Pax7 (1:100, mouse monoclonal, P3U1; Developmental Studies Hybridoma Bank), and anti‐bromodeoxyuridine (BrdU; 1:200, sheep polyclonal, ab1893; abcam).

Cryosections were fixed with 4% paraformaldehyde (PFA) in phosphate‐buffered saline (PBS), blocked with 5% normal donkey serum or 5% normal goat serum/PBS. The sections were incubated with primary antibodies overnight at 4°C, followed by incubation with AlexaFluor‐conjugated secondary antibodies (1:500, donkey polyclonal; Jackson ImmunoResearch Laboratory, PA, USA) or AlexaFluor‐conjugated anti‐rat IgG antibody (1:500, goat polyclonal; Thermo Fisher Scientific) for 1 h, and counterstaining the nuclei with Hoechst 33258. Photographs were taken using a UplanSApo ×4 objective lens, a UPlanApo ×20 objective lens, and a fluorescence microscope (BX53; Olympus, Tokyo, Japan) equipped with a digital camera (DP73, Olympus) or a Plan Fluorite 20× LD PH lens and a fluorescence microscope system (BZ‐X800; Keyence, Osaka, Japan).

For the quantification of myofiber size, eMHC‐positive myofibers or centrally nucleated myofibers were histologically considered regenerating myofibers, and their diameters were measured based on laminin staining using ImageJ [20] (version 1.54f).

2.6. Fluorescence‐Activated Cell Sorting Analysis

The number of macrophages was evaluated using a BD FACS Verse Flow Cytometer (651 153; BD Biosciences, Franklin Lakes, NJ, USA).

The dissected grafted muscles were minced and digested for 1 h at 37°C in collagenase type II (2 mg/mL, Worthington, NJ, USA). The cells were resuspended in 2% fetal bovine serum (FBS; Sigma‐Aldrich)/PBS and filtered through 70 μm cell strainers (Corning, NY, USA). CD163‐positive cells were stained with an FITC‐conjugated anti‐CD163 antibody (1:100, mouse monoclonal, ED2; Bio‐Rad) for 20 min on ice, followed by Alexa488‐conjugated secondary antibody (1:500, donkey polyclonal; Jackson ImmunoResearch Laboratory) for 20 min on ice. Dead cells were stained using the LIVE/DEAD Fixable Red Dead Cell Stain Kit (Thermo Fisher Scientific), followed by fixation using a BD Cytofix/Cytoperm Plus Fixation/Permeabilization Solution Kit with BD GolgiStop (Becton Dickinson and Company). CD68‐ and CD86‐positive cells were stained with Alexa647‐conjugated anti‐CD68 antibody (1:100, mouse monoclonal, ED1; Bio‐Rad) and PE‐conjugated anti‐CD86 antibody (1:50, mouse monoclonal, 24F; Becton Dickinson and Company) for 20 min on ice. The number of cells was calibrated using Count Bright Absolute Counting Beads (Thermo Fisher Scientific).

2.7. Myofiber Isolation

Under isoflurane anesthesia, EDLs were excised at both tendon ends and digested with collagenase type I (2 mg/mL, Worthington) in 10% FBS/Dulbecco's modified Eagle medium at 37°C with gentle agitation for 2.5–5 h until separation of myofibers was observed. Myofibers were manually isolated under a stereomicroscope using micropipettes and collected from dishes coated with 5% BSA. Photographs were captured using a UplanFL N × 4 objective lens and an inverted optical microscope (IX71, Olympus) equipped with a digital camera (DP70, Olympus).

2.8. Immunoblotting

For immunoblotting the following primary antibodies were used: anti‐p16 (1:4000, rabbit monoclonal, ab211542; abcam), anti‐ platelet‐derived growth factor receptor α (PDGFRα; 1:1000, rabbit polyclonal, SAB4502140; Sigma‐Aldrich), and anti‐fast‐type myosin (1:1000, mouse monoclonal, MY‐32; Sigma‐Aldrich).

Frozen muscle tissue and isolated myofibers were homogenized in radioimmunoprecipitation assay buffer. Protein concentrations were semi‐quantified and equalized using the bicinchoninic acid assay kit (Wako, Osaka, Japan). The extracts were denatured at 95°C for 5 min with 2× Laemmli sample buffer containing 2‐mercaptoethanol and were separated on a 15% SDS–polyacrylamide gel. Following this, the proteins were electroblotted onto a polyvinylidene fluoride membrane, followed by staining with Ponceau S staining solution (0.1% (w/v) Ponceau S in 5% acetic acid). The membrane was blocked with 5% skim milk/0.1% Tween 20/Tris‐buffered saline and incubated with the primary antibody at 4°C overnight. The next day, the membranes were incubated with horseradish peroxidase‐conjugated anti‐mouse IgG (1:10000) or anti‐rabbit IgG (1:8000) antibody (goat polyclonal, Jackson ImmunoResearch Laboratory). Signals were detected using an ECL Prime western blotting analysis system (GE Healthcare, IL, USA).

2.9. In Situ Hybridization

RNAscope 2.5HD Assay Kit (Advanced Cell Diagnostics) was used for the ISH of Cdkn2a mRNA (Probe‐Rn‐CDKN2A, Advanced Cell Diagnostics). Briefly, cryosections were fixed with 4% PFA/PBS, blocked for endogenous peroxidase activity, hybridized with the Cdkn2a probe [positive control probe (313 921; Advanced Cell Diagnostics), or negative control probe (310 043; Advanced Cell Diagnostics)], and subjected to signal amplification and detection. Immunohistochemistry was performed after detection with Diaminobenzidine.

2.10. Statistical Analyses

Statistical analyses were performed using the R software (version 4.3.1). Significant differences between groups were calculated using unpaired Student's or Welch's two‐tailed t‐test for experiments comparing two groups, Tukey–Kramer test for experiments comparing three or more groups, and Fisher's test for experiments comparing the rate of Cdkn2a expression in BrdU‐positive and ‐negative myonuclei and in eMHC‐positive and ‐negative myofibers. For GSEA, p‐values were calculated using the clusterProfiler package with Benjamini–Hochberg correction. All data are presented as mean ± standard error of the mean. p < 0.05 was considered statistically significant (*p < 0.05, **p < 0.01).

3. Results

3.1. DMD Skeletal Muscle Expresses Senescence‐Associated Cytokines

In addition to the skeletal muscle, dystrophin is expressed in the heart and central nervous system (CNS). In DMD rats, myocardial degeneration and fibrosis have been observed in the heart, and some neurons exhibit blunted excitability in the CNS [21, 22]. Examination of multiple organs in DMD rats for p16 expression revealed that p16 was highly expressed in the skeletal muscles (Figure 1A). To elucidate the mechanism by which p16 expression in skeletal muscle exacerbates muscle pathology, we performed RNA‐seq of skeletal muscle from 9‐month‐old WT, DMD, and p16‐null DMD (dKO) rats. The PCA plot revealed distinct separation in the transcriptomes of WT and DMD rats, whereas no clear difference was observed between DMD and dKO rats (Figure 1B). GSEA was then used to identify gene expression pathways differing between the skeletal muscles of DMD and dKO rats. The altered pathways primarily included those for cytokine suppression, such as interleukin (IL)‐17 and TNF signaling pathways, with the top hit of “Cytokine‐cytokine receptor interaction” pathway (Figure 1C,D), which was also significantly altered in DMD and WT rats (Figure 1E, Table S1). The expressions of many genes in this pathway were increased in DMD rats and decreased in dKO rats, compared to those in WT rats (Figure 1F). Thus, p16‐dependent expression of cytokines, which are considered to be SASP, occurs in DMD rat skeletal muscles. These SASP factors may be potentially involved in the development of muscle pathology in DMD rats and in ameliorating muscle pathology in dKO rats.

FIGURE 1.

Cytokine gene expression was p16‐dependently upregulated in Duchenne muscular dystrophy (DMD) rat muscle. (A) p16 mRNA quantification in 8‐month‐old wild‐type (WT) and DMD rats (n = 1, 3). Undetected values were treated as zero. (B) Principal component analysis of gene expression profile using RNA‐seq of 9‐month‐old WT, DMD, and p16‐null DMD (dKO) rats (n = 3, 3, and 6, respectively). (C) Significantly upregulated or downregulated Kyoto Encyclopedia of Genes and Genomes (KEGG) terms using gene set enrichment analysis between DMD and dKO. KEGG terms with adjusted p‐values less than 0.01 are shown. (D) Enrichment plot of “Cytokine‐cytokine receptor interaction” between DMD and dKO rats. (E) Enrichment plot of “Cytokine‐cytokine receptor interaction” between WT and DMD rats. (F) Heatmap of gene expression in “Cytokine‐cytokine receptor interaction”. Genes with significant differences (p < 0.05) in expression between DMD and dKO were selected. NES, normalized enrichment score; FDR, false discovery rate.

Subsequently, we quantified the serum cytokine levels by performing cytokine arrays for WT and DMD rats (Figure 2A) and determined that WT and DMD rats exhibited different expression profiles (Figure 2B). Thus, cytokine expression in the skeletal muscles of DMD rats may have systemic effects.

FIGURE 2.

Cytokine profiles in blood are altered in DMD rats. (A) Heatmap of cytokine protein levels in the sera of WT and DMD rats (n = 4 each). (B) Principal component analysis of cytokine protein levels. The ellipses indicate a 90% confidence interval using the t‐distribution.

3.2. Systemic Environment in DMD Rats Disrupts Muscle Regeneration in a p16‐Dependent Way

EDL muscle transplantation was performed to examine the effects of systemic changes, including serum cytokine profiles, on muscle regenerative potential. In this model, EDLs are grafted onto the hind legs of another rat, and degeneration and regeneration of myofibers are induced in the grafted muscles [14] (Figure 3A). To evaluate the effects of the systemic environment in DMD rats on the regeneration of WT skeletal muscles, WT EDLs were grafted onto WT, DMD, and dKO rats. When the transplanted muscles were sampled 10 d after transplantation during the regeneration process, their weight tended to be lower in DMD rats than that in WT or dKO rats (Figure 3B). Therefore, we quantified the diameters of eMHC‐positive and centrally nucleated regenerating myofibers in cross sections and found that the diameter of the regenerating myofibers was relatively smaller in DMD rats (Figure 3C), but was similar between WT and dKO rats (Figure 3D,E). Thus, the systemic environment in DMD rats may inhibit muscle regeneration in a p16‐dependent manner.

FIGURE 3.

p16‐dependent systemic environment in DMD rats impaired muscle regeneration in the grafted muscle. (A) Representative hematoxylin and eosin staining of grafted extensor digitorum longus (EDL) muscles transplanted to WT rats. Scale bar = 100 μm. (B) Weight of grafted EDLs transplanted to WT, DMD, and dKO rats (n = 6, 7, and 3, respectively). Data are compared using Tukey–Kramer's test. (C) Representative immunofluorescence images of grafted EDLs in WT, DMD, and dKO rats. Scale bar = 50 μm. (D) Distributions of regenerating myofibers' diameters of grafted EDLs in WT, DMD, and dKO rats (n = 6, 7, and 3, respectively). (E) Quantification of median for the diameters of regenerating myofibers for each grafted EDL in WT, DMD, and dKO rats (n = 6, 7, and 3, respectively). Data are compared using Tukey–Kramer's test.

Next, we examined whether host cells infiltrated the transplanted muscle by performing EDL grafting in GFP rats. GFP‐positive infiltrating cells were observed in the transplanted muscles. The GFP signal was not observed in the regenerating eMHC‐positive myofibers but was detected predominantly in CD68‐positive cells (Figure 4A,B). Therefore, all cells that infiltrated the transplanted muscle from the host were macrophages, and the regenerated myofibers expressing dystrophin were presumably derived from the transplanted muscle. Therefore, we hypothesized that differences in the characteristics of the macrophages infiltrating the transplanted muscle might affect the size of the regenerating myofibers. Hence, we quantified the number of infiltrating macrophages using flow cytometry at 1, 3, 10, 14, and 30 d following grafting. The number of CD68‐positive cells showed no significant difference between the DMD and WT rats at any post‐transplantation time point (Figure 4C). Furthermore, when comparing macrophage polarization, the proportion of CD86‐positive M1 macrophages was higher, whereas that of CD163‐positive M2 macrophages was lower at 14 days post‐transplantation in DMD rats than in WT rats (Figure 4D,E). However, this change could not explain the reduction in regenerating myofiber size observed in muscles grafted in DMD rats at 10 d post‐graft. The delayed transition of macrophages from the pro‐inflammatory M1 to the regenerative M2 phenotype, typically observed during muscle regeneration [1], by 14 days post‐graft is attributable to the earlier delay in muscle regeneration.

FIGURE 4.

The infiltrating macrophages were unaltered in the grafted muscle. (A, B) Representative immunofluorescence images of grafted EDL muscles transplanted to GFP rats. Arrowheads indicate GFP+ cells. Scale bar = 50 μm. (C–E) Number of CD68+, CD86+ (marker of M1 macrophage), and CD163+ (marker of M2 macrophage) cells in the grafted EDLs was determined using flow cytometry in WT and DMD rats at 1, 3, 6, 10, 14, and 30 d after graft (WT: N = 3 each; DMD: N = 3, 3, 3, 4, 4, and 4). Data are compared using an unpaired Student's t‐test.

3.3. DMD Myofibers Express SASP Genes

Skeletal muscle myofibers function as endocrine organs since they are a key source of cytokines secreted following exercise or upon lipopolysaccharide injection [23, 24]. Therefore, cytokines in the skeletal muscles of DMD rats may be derived from myofibers. However, in the skeletal muscles of DMD rats, the prevalence of inflammatory and mesenchymal progenitor cells is increased [3], strongly suggesting that these stromal cells could also be a source of cytokines. To investigate the contribution of DMD rat myofibers to cytokine expression in skeletal muscles, we enzymatically isolated myofibers from 9‐month‐old DMD rats after collagenase digestion and characterized them (Figure 5A). After isolation, the relative amount of fast‐type myosin protein in the myofibers increased compared to that in bulk muscle, and PDGFRα, a marker of stromal mesenchymal progenitor cells and fibroblasts, was not detected in the myofibers, confirming that the isolated myofibers are devoid of attached stromal cells (Figure 5B).

FIGURE 5.

Myofibers in DMD rats expressed SASP and senescence marker genes. (A) Representative images of isolated myofibers from the EDL muscles of 9‐month‐old WT and DMD rats. Scale bar = 200 μm. (B) Immunoblotting analysis of platelet‐derived growth factor receptor α (PDGFRα) and fast‐type myosin in bulk EDLs and isolated myofibers from EDLs. (C‐E) mRNA levels of SASP genes, cellular senescence marker genes in isolated myofibers from 9‐month‐old WT, DMD, and dKO rats (n = 7, 7, 6). Undetected values were treated as zero. Data are compared using Tukey‐Kramer's test. (F) Immunoblotting analysis of p16 in the isolated myofibers of 3‐, 6‐, and 9‐month‐old WT and DMD rats (n = 1 and 3, respectively, for each age).

We then quantified the inflammatory SASP factors with p16‐induced upregulation in gene expressions in DMD rat skeletal muscle (Figure 1F; Ccl2, Cxcl1, Il1b, Il6) as well as other previously reported p16‐dependent SASP factors (Ccn2, Mmp2, Tgfb1) in DMD rat muscle [3]. All of the quantified cytokines showed significantly higher expression in myofibers isolated from DMD rats than in those isolated from WT rats (Figure 5C,D), suggesting that myofibers contribute to the SASP in the skeletal muscles of DMD rats. While the inflammatory SASP factors (Ccl2, Cxcl1, Il1b, Il6) showed higher gene expression in the myofibers from DMD rats compared to those from WT, the expressions are comparable between those of WT and those of dKO (Figure 5C). In contrast, among the pro‐fibrotic SASP factors (Ccn2, Mmp2, Tgfb1), the expressions of Ccn2 and Tgfb1 were significantly upregulated in the myofibers from both DMD and dKO rats compared with WT rats (Figure 5D). These results suggest that p16 is responsible for the upregulation of most SASP‐associated genes in myofibers, except for Ccn2 and Tgfb1.

Recent reports have shown that postmitotic cells in the brain, retina, heart, and skeletal muscle also exhibit these cellular senescence markers, and these “senescence‐like” phenomena are called postmitotic cellular senescence [25, 26]. Therefore, SASP factor expression in myofibers might be due to postmitotic cell senescence. In myofibers isolated from DMD rats, p16 expression was confirmed at both the gene and protein levels (Figure 5E,F), suggesting its senescence‐like state. Gene expressions of other senescence markers (p19 and p21) were also upregulated in myofibers from DMD rats compared to WT rats. However, p19 and p21 showed higher gene expression in myofibers of dKO rats compared to WT rats (Figure 5E). Taken together, the myofibers of DMD rats underwent postmitotic cell senescence and secreted SASP factors.

3.4. DMD Myofibers Express p16

We further investigated the characteristics of “postmitotic senescent” myofibers, which contribute to the SASP in DMD rat skeletal muscles. ISH of Cdkn2a mRNA was performed to examine p16 localization in the skeletal muscles of DMD rats. While no signal was observed in WT skeletal muscle, p16 was expressed in the myofibers, satellite cells, and other stromal cells in DMD rat skeletal muscle (Figure 6A), which is consistent with a previous report that satellite and mesenchymal progenitor cells express p16 [3]. In DMD rat sections, from 3‐month‐old, when the pathology is relatively mild, and myofiber necrosis and regeneration are in process, to 9‐month‐old, when adipogenesis and significant muscle weakness are observed [3], the proportion of p16‐positive satellite cells and myofibers increased as the muscle pathology progressively worsened (Figure 6B,C). Necrotic and regenerating myofibers, which are rare in WT rats, are frequently present in DMD rats [12]. Therefore, we hypothesized that necrotic or regenerating myofibers express p16. Following ISH, we immunohistochemically stained necrotic myofibers for rat IgG. However, IgG‐positive myofibers did not coincide with p16‐positive myofibers (Figure 6D). Furthermore, immunostaining for eMHC was performed to identify immature regenerating myofibers. Although p16 was observed in both eMHC‐positive and ‐negative myofibers, its expression was higher in eMHC‐positive myofibers than in eMHC‐negative myofibers at all ages (Figure 6E,F).

FIGURE 6.

DMD myofibers express p16. (A) Representative images after Cdkn2a mRNA in situ hybridization using RNAscope and immunofluorescence of Pax7. White arrowheads indicate Pax7+ cells. Black and red arrows indicate Diaminobenzidine (DAB) signals inside and outside myofibers, respectively. Scale bar = 50 μm. (B) Ratios of Cdkn2a + Pax7+ cells per Pax7+ cells in the EDL muscles of 3‐, 6‐, and 9‐month‐old DMD rats (n = 3 each). Data are compared using Tukey–Kramer's test. (C) Ratios of Cdkn2a + myofibers in EDLs of 3‐, 6‐, and 9‐month‐old DMD rats (n = 3 each). Data are compared using Tukey–Kramer's test. (D) Tibialis anterior muscle sections were subjected to Cdkn2a mRNA in situ hybridization using RNAscope and immunofluorescence of rat IgG. White arrowheads indicate IgG+ cells. Black arrows indicate DAB signals. Scale bar = 100 μm. (E) Representative images after Cdkn2a mRNA in situ hybridization using RNAscope and immunofluorescence of embryonic myosin heavy chain (eMHC). White arrowheads indicate eMHC+ myofibers. Black arrows indicate DAB signals. Scale bar = 50 μm. (F) Ratios of Cdkn2a + myofibers per eMHC+ or eMHC‐ myofibers in the EDLs of 3‐, 6‐, and 9‐month‐old DMD rats (n = 3 each). Data are compared using unpaired Student's t‐tests. (A, D, E) The lower images were generated by averaging bright‐field images with DAB signals of Cdkn2a and fluorescent images.s.

Since regenerating muscle transiently upregulates expression of p16 and other CDK inhibitors [10, 27], we considered the possibility that p16 may also be expressed in skeletal muscles during regeneration, but not specifically in DMD pathology. To test this hypothesis, we induced muscle injury in the tibialis anterior muscle using BPVC and quantified p16 gene expression. However, the relative p16 gene expression levels at 1, 2, 3, 4, 5, 6, 7, and 14 days after injury, as well as in intact skeletal muscle, were less than 1.5% of that in the tibialis anterior muscle of DMD rats, with no significant changes observed in response to injury (n = 2) (data not shown). We also examined whether p16 is transiently expressed in DMD rat myofibers as part of the differentiation process by labeling the regenerating myonuclei with BrdU over 2 weeks (SF. 1A). When comparing the proportion of Cdkn2a mRNA detected nearest to BrdU‐positive and BrdU‐negative myonuclei within eMHC‐positive fibers, p16 was more frequently expressed near BrdU‐negative nuclei than near BrdU‐positive nuclei which had recently differentiated from myoblasts (SF. 1B, C). Thus, p16 expression is not part of the regeneration process; instead, it is expressed at a higher rate in immature myofibers, where muscle regeneration has not yet progressed.

3.5. p16 Is Expressed in Myofibers of the BMD Model Rats and Aging Rats

We investigated whether p16 is also expressed in myofibers during other muscular disorders. We examined a rat model of BMD, a progressive muscle disease with a pathology similar to that of DMD. Our previously generated BMD rat model exhibits skeletal muscle necrosis and exercise intolerance due to reduced function and expression of dystrophin, adipogenesis, and pseudohypertrophy during their muscle pathogenesis with age [13]. Our findings revealed that p16 gene expression in the skeletal muscles of BMD rats was comparable at 6 months old between WT and BMD rats, whereas it was higher in BMD rats than in WT rats at 10 months old (Figure 7A). This increase in p16 expression coincides with a reported decline in the muscle regeneration capacity between 6 and 11 months of age in the tibialis anterior muscle of BMD rats [13]. ISH of p16 mRNA revealed that p16 was exclusively expressed in myofibers of BMD skeletal muscles (Figure 7B). Additionally, immunostaining for eMHC to detect immature myofibers, followed by ISH, showed that eMHC‐positive myofibers expressed p16 at a higher rate than eMHC‐negative fibers, which was consistent with the results in DMD rats (Figure 7C,D). We then considered the possibility that SASP factors might also be expressed in BMD rat skeletal muscle and quantified their gene expression levels. In BMD rats of the same age, the expression of Cxcl1, Mmp2, and Tgfb1 increased in the skeletal muscle (Figure 7E). Therefore, in rats with BMD, p16 is expressed in myofibers, along with a decline in muscle regenerative capacity and increased expression of SASP factors. Thus, myofiber‐p16 and SASP may impair muscle regeneration in DMD and BMD rats.

FIGURE 7.

p16 is expressed in the myofibers of BMD model rats. (A) mRNA levels of p16 in tibialis anterior muscles of 6‐ and 10‐month‐old WT and BMD rats (WT: N = 3, 3; BMD: N = 3, 6). Relative values compared to the tibialis anterior muscle of 10‐month‐old DMD rats. The undetected values were treated as zero. Data were compared using an unpaired Welch's t‐test. (B, C) Representative images of Cdkn2a mRNA in situ hybridization using RNAscope and immunofluorescence of Pax7 (B) and eMHC (V) in 10‐month‐old WT and BMD rats. White arrowheads indicate Pax7+ and eMHC+ myofibers. The lower images were generated by averaging the bright‐field images with the DAB signals of Cdkn2a and the fluorescent images. Black arrows indicate DAB signals inside and outside the myofibers. Scale bar = 50 μm. (D) Ratios of Cdkn2a + myofibers per eMHC+ and eMHC‐ myofibers in 10‐month‐old BMD rats. Numbers in parentheses indicate the number of myofibers. Data were compared using Fisher's exact test. (E) mRNA levels of SASP genes in tibialis anterior muscles of 10‐month‐old WT and BMD rats (n = 3, 6). Data were compared using an unpaired Welch's t‐test.

In rodent skeletal muscles, p16 expression increases with age, along with a decline in muscle function [28, 29]. Therefore, we injected BPVC into the tibialis anterior muscle of 3‐ to 4‐month‐old (young) and 24‐month‐old (aged) rats to induce muscle regeneration and compared the regenerative capacity by examining the diameter of the regenerated muscle fibers 14 days after injury (Figure 8A). Aged skeletal muscle demonstrated reduced diameter of the regenerated myofibers (Figure 8B,C), suggesting a decline in regenerative capacity in aged rats, which is consistent with previous reports [30]. Simultaneously, p16 gene expression was higher in the skeletal muscle of aged rats compared to that in younger rats 14 days post‐injury (Figure 8D). ISH revealed that p16 was expressed in the myofibers of injured and uninjured aged muscles (Figure 8E). Furthermore, eMHC‐positive myofibers were present in the control group of aged rats and p16 was highly expressed in these immature myofibers (Figure 8E). Next, we performed RNA‐seq on the tibialis anterior muscle of 3‐month‐old and 18‐month‐old rats; Cdkn2a (p16 and p19) gene expression was detected only in aged skeletal muscles (Figure 8F). Furthermore, the expression of genes involved in the “Cytokine‐cytokine receptor interaction” pathway was increased in the skeletal muscle of 18‐month‐old rats (Figure 8G, Table S2). These results suggest that p16 is also expressed in the immature myofibers of aged rats and may contribute to increased cytokine expression and decreased regenerative capacity in the skeletal muscle of aged rats.

FIGURE 8.

p16 is expressed in myofibers of aged rats. (A) Representative immunofluorescence images of the tibialis anterior muscles of 3‐ to 4‐month‐old (young) and 24‐month‐old (aged) rats, 14 days after bupivacaine hydrochloride (BPVC) injection. Scale bar = 100 μm. (B) Distribution of regenerating myofiber diameters in the BPVC‐injected muscles of young and aged rats (n = 4, 3). (C) Quantification of the median regenerating myofiber diameter of each BPVC‐injected muscle in young and aged rats (n = 4, 3). Data were compared using an unpaired Student's t‐test. (D) p16 mRNA levels in BPVC‐injected or untreated (intact) tibialis anterior muscles of young and aged rats (n = 4, 3). (E) Tibialis anterior muscle sections were subjected to Cdkn2a mRNA in situ hybridization using an RNAscope and eMHC immunofluorescence. The lower images were generated by averaging the bright‐field images with the DAB signals of Cdkn2a and the fluorescent images. White arrowheads indicate eMHC+ cells. Black arrows indicate the DAB signals. Scale bar = 50 μm. (F) The transcripts per million (TPM) values of Cdkn2a in RNA‐seq of the tibialis anterior muscles of 3‐ (young) and 18‐month‐old (aged) rats (n = 3, 3). (G) Enrichment plot of “Cytokine‐cytokine receptor interaction” between young and aged rats (n = 3, 3).

4. Discussion

This study establishes that myofibers in DMD rats undergo postmitotic cell senescence and acquire the SASP in a p16‐dependent manner, which may contribute to a systemic humoral environment that inhibits muscle regeneration.

We previously reported that cellular senescence is induced in the skeletal muscle of DMD rats, with the expression of p16 in mesenchymal progenitor cells and satellite cells [3, 31]. Traditionally, cellular senescence has been considered a phenomenon seen in mitotic cells. However, senescence‐like phenotypes, such as the expression of CDK inhibitors, SASP, lamin B1 reduction, and DNA damage, are also seen in postmitotic cells, a process referred to as postmitotic cell senescence [25, 26]. In aged mouse cardiomyocytes, DNA damage induces the expression of p16 and p21 [32]; primary cultured rat neurons also express p16 and SASP factors with lamin B1 reduction [33]; myofiber p16 is highly expressed in a model mouse with elevated DNA damage [34]. In mouse and human skeletal muscles, myofibers upregulate p16 or p21 expression with aging, and senescence‐like changes such as DNA damage, lamin B1 reduction, and SASP occur in myofibers [28, 35, 36]. In this study, we found that p16 was primarily expressed in the myofibers of DMD rat skeletal muscles, and the isolated myofibers expressed senescence markers such as p16, p19, p21, and SASP factors. This suggests that senescence‐like changes occur in the myofibers of DMD rats.

In the transplanted muscles of DMD rats, the diameter of the regenerating myofibers was reduced in a p16‐dependent manner. This suggests that regeneration processes, such as the removal of necrotic myofibers through phagocytosis, the growth and differentiation of satellite cells, and the fusion and maturation of myotubes, may be inhibited or delayed. These changes may be attributed to the altered characteristics of infiltrating macrophages. In the present study, no difference in the M1/M2 ratio of macrophages was observed in the transplanted muscles of DMD and WT rats until day 10 post‐transplantation, and no significant difference in macrophage numbers was observed. However, owing to the large variation in macrophage numbers, the possibility of increased or decreased infiltration affecting muscle regeneration cannot be ruled out.

Potential systemic changes, other than infiltrated macrophages, should affect the transplanted muscle; however, these changes are likely attributable to the skeletal muscle cytokine levels. Since p16 expression is localized in the skeletal muscle, p16‐dependent impaired regeneration originates from the host skeletal muscle. For example, C‐C motif chemokine ligand (CCL)‐2, whose expression is upregulated in DMD rat skeletal muscles, enhances monocyte chemotaxis and induces IL‐6 expression [37]. Thus, SASP factors from DMD rat skeletal muscles may have created an environment unfavorable for muscle regeneration by altering extramuscular cell function. Additionally, SASP factors derived from skeletal muscle may directly act on the transplanted muscle, inhibiting its regeneration. Co‐culturing senescent cells, which acquire SASP, is known to suppress myoblast division or myotube formation [9, 29]. Furthermore, transplanting senescent cells into skeletal muscle inhibits regeneration [9], thus validating that the SASP negatively impacts muscle regeneration. Among the SASP factors quantified in DMD rat myofibers, transforming growth factor (TGF)‐β [38, 39], cellular communication network factor 2 (CCN2) [40], IL‐1β [41], and CCL‐2 [42] are known to inhibit muscle differentiation, while C‐X‐C motif chemokine ligand (CXCL)‐1 inhibits muscle regeneration in vivo [42]. Therefore, the SASP of skeletal muscles directly or indirectly suppresses muscle regeneration in grafted EDL.

In addition, p16 is expressed in the skeletal muscle of patients with DMD, suggesting cellular senescence of skeletal muscle [3]. In patients with DMD, the expressions of TGF‐β1 [43], CCN2 [44], and chemokines [45] in the skeletal muscle are upregulated, along with elevated blood levels of TGF‐β1 [46], IL‐1β [47], IL‐6 [47, 48], CCL‐2 [49], and CXCL‐10 [49]. Research investigating the localization of these cytokines via immunohistochemistry has identified that they are primarily expressed in myofibers [45, 50, 51]. Specifically, CCL2 mRNA [50], CXCL‐1 [50], and CCN2 [44] are highly expressed in immature myofibers from patients with DMD, indicating that SASP due to postmitotic cell senescence in myofibers may frequently occur in immature myofibers from patients with DMD, as well as in DMD rats.

In this study, aged rat myofibers expressed p16, exhibited SASP factors in skeletal muscle, and showed decreased muscle regenerative capacity. In muscle transplantation experiments using 2‐ or 3‐month‐old and 24‐month‐old rats, muscle regeneration was significantly suppressed when the host, but not the donor, was aged [52]. Carlson and Faulkner suggested that the decrease in muscle regenerative capacity in aged rats is due to changes in the systemic environment rather than to alterations within skeletal muscle, such as those in satellite cells. However, changes in nerve reinnervation in aged individuals also contribute significantly to a decline in muscle regenerative capacity [53]. In aged humans, the number of p16‐positive myofibers also increases [28]. This suggests the possibility of a similar mechanism, involving reduced p16‐induced muscle regeneration in myofibers, in human sarcopenia. However, the level of p16 gene expression in aged rats was only a few percent of that in DMD rats, suggesting that the functional role of p16 may be limited in 24‐month‐old rats compared to that in DMD rats.

In DMD rats, we previously suggested that increased oxidative stress associated with chronic inflammation presumably causes cellular senescence [3]. Senescent‐like myofibers in DMD rats may result either from the postmitotic cell senescence of differentiated myofibers or the differentiation and fusion of senescent myoblasts. Myotubes, which are terminally differentiated cells, have lower DNA repair capacity than that of myoblasts, since they lack cell division and are more sensitive to oxidative stress [54]. Postmitotic senescence in cells after terminal differentiation is known and is induced by culturing postmitotic primary neurons in vitro [33]; the upregulated expression of p21 is induced by doxorubicin treatment of myotubes [36]. A recent study further shows that plasma membrane damage, which is frequent in dystrophic myofibers, can induce senescence‐like responses [55], suggesting that membrane injury may act together with oxidative stress to promote senescent‐like myofibers in DMD rats. Alternatively, the senescent‐like myofibers may originate from senescent myoblasts expressing p16 in rat models of DMD and patients with DMD [2, 3, 7]. Myotubes with reduced lipid metabolism are formed from senescent myoblasts, although myotube formation is impaired in senescent myoblasts [56], suggesting that postmitotic myofibers inherit an altered phenotype of senescent myoblasts. As p16 expression itself is known to promote myogenic differentiation [57], the possibility that mitotic myoblasts undergo cellular senescence, differentiate, and fuse to form postmitotic senescent myofibers cannot be disregarded. In aging rats, a similar explanation can be applied as both inflammageing and cellular senescence of satellite cells are reported in aged mice. Although similar senescent features are observed in aged muscles [6, 36], the distinction between disease‐associated and age‐associated contexts is not fully clear and is noted here.

In the present study, the majority of the myofibers expressing p16 were eMHC‐positive and BrdU‐negative, which are immature but not recently differentiated, suggesting inhibition of the maturation process in these myofibers; however, p16 expression at the myoblast stage, which can cause cell cycle arrest, may also explain the BrdU negativity in p16‐positive myonuclei without inhibiting maturation. In the present study, the proportion of p16 expression in eMHC‐positive myofibers of DMD rats remained relatively constant regardless of age, whereas that in eMHC‐negative fibers increased with age. Thus, p16 might be initially expressed in immature myofibers during regeneration. As these myofibers slowly mature with persistent p16 expression, the p16‐positive rate increases in eMHC‐negative myofibers. As an example of immature myofibers, denervated myofibers continuously express eMHC [58] and eventually express p21 and factors such as TGF‐β and osteopontin, which are also known as SASP factors [58, 59]. Thus, immature myofibers are prone to postmitotic cellular senescence. Verily, myofiber populations with high expression of p21 and neonatal MHC, another immature form of MHC, are specifically present in aged individuals [36]. Conversely, p16 expression itself may delay myofiber maturation. The p16‐mediated inhibition of CDK4 in myofibers inhibits peroxisome proliferator‐activated receptor‐gamma coactivator (PGC)‐1α, leading to decreased mitochondrial function [60]. The deficiency of PGC‐1α is known to inhibit the formation of neuromuscular junctions, which is essential for myofiber maturation [61], and the upregulation of Mfn2, a PGC‐1α target gene, is necessary for myofiber maturation [62]. Further studies are required to investigate the effects of myofiber‐p16 on muscle regeneration.

In DMD rats, p16 is expressed in postmitotic myofibers. In mitotic cells, p16 is a tumor suppressor that induces cellular senescence by inhibiting cell cycle progression. The knockout of p16 in DMD rats results in more than 50% incidence of rhabdomyosarcoma [63]. However, p16 also functions independently of cell cycle arrest by inhibiting CDK4 and its downstream target, Rb [64, 65, 66, 67, 68, 69, 70, 71, 72, 73], suggesting a role for p16 in postmitotic myofibers. Notably, p16 is necessary for persistent SASP, independent of cell cycle arrest in senescent cells [74, 75]. Therefore, myofiber‐p16 may play a role in sustaining the SASP in DMD rats.

In the present study involving DMD rats, p16 expression in postmitotic myofibers exacerbated muscular pathology by secreting SASP factors, resulting in phenotypes similar to those observed in BMD and sarcopenia models. Additionally, myofibers may undergo cellular senescence‐like change during obesity [76], potentially leading to a similar decline in muscle regeneration. Since myofibers are large cells present throughout the body, the SASP in myofibers can not only impact muscle regeneration but also other muscle pathogeneses and even overall systemic health. Furthermore, cellular senescence is associated with phenotypes such as mitochondrial dysfunction and metabolic alterations, which may also be induced by postmitotic cell senescence in myofibers, thereby exacerbating DMD pathology.

This study does exclude the possibility that p16 expression in the myoblasts and satellite cells of DMD rats suppresses muscle regeneration by inhibiting muscle activation and proliferation. In addition, the extent to which myofibers and individual SASP factors contribute to the exacerbation of DMD pathogenesis remains unclear. Thus, future research may focus on demonstrating how much the suppression of SASP in myofibers improves the DMD pathology and identifying the SASP factors and further mechanisms that inhibit muscle regeneration.

Author Contributions

M. Ikeda and Y. Tanaka designed the study and conducted most of the experiments. M. Ikeda analyzed the data and wrote the original manuscript. K. Yamanouchi supervised the study and reviewed the manuscript. H. Sugihara prepared RNA‐seq samples. H. Sugihara, T. Matsuwaki, and K. Yamanouchi provided helpful suggestions.

Funding

This work was supported by MEXT | Japan Society for the Promotion of Science (JSPS), JP23H00359, JP20H03161, JP24KJ0705, JP22J20889, JP22KJ1027. Japan Science Society (JSS), 2022–4087.

Conflicts of Interest

The authors declare no conflicts of interest.

Supporting information

Figure S1: p16 is expressed by BrdU‐myonuclei in eMHC+ myofibers of DMD rats.

(A) Schematic of experimental design. After 2 weeks of feeding drinking water containing bromodeoxyuridine (BrdU, 0.8 mg/mL; Wako, Osaka, Japan), tibialis anterior muscles were collected from 12‐month‐old DMD rats (n = 2). (B) Representative images after Cdkn2a mRNA in situ hybridization using RNAscope and immunofluorescence of BrdU and embryonic myosin heavy chain (eMHC). The lower images were generated by averaging bright‐field images with Diaminobenzidine (DAB) signals of Cdkn2a and fluorescent images. White arrowheads indicate BrdU+ and BrdU‐ cells in an eMHC+ myofiber. The black arrow indicates the DAB signal. Scale bar = 50 μm. (C) Ratios of Cdkn2a + myofibers per BrdU+ and BrdU‐ myonuclei inside eMHC+ myofibers in 12‐month‐old DMD rats. Numbers in parentheses indicate the number of myonuclei counted. Data were compared using Fisher's exact test.

Table S1: Significantly upregulated or downregulated Kyoto Encyclopedia of Genes and Genomes (KEGG) terms were identified using gene set enrichment analysis (GSEA) between DMD and WT rats. KEGG terms with adjusted p‐values less than 0.01 are shown.

Supplementary Table 2. Significantly upregulated or downregulated KEGG terms identified using GSEA between 3‐ and 18‐month‐old rats. KEGG terms with adjusted p‐values less than 0.1 are shown.

Data Availability Statement

All data are available in the main text or Supporting Information, or from the corresponding author upon reasonable request. The RNA‐seq data generated in this study were deposited in the Gene Expression Omnibus database under the accession code GSE282700.