Decrease in the expression of muscle-specific miRNAs, miR-133a and miR-1, in myoblasts with replicative senescence

Kaori Shintani-Ishida; Riko Tsurumi; Hiroshi Ikegaya

doi:10.1371/journal.pone.0280527

Abstract

Muscles that are injured or atrophied by aging undergo myogenic regeneration. Although myoblasts play a pivotal role in myogenic regeneration, their function is impaired with aging. MicroRNAs (miRNAs) are also involved in myogenic regeneration. MiRNA (miR)-1 and miR-133a are muscle-specific miRNAs that control the proliferation and differentiation of myoblasts. In this study, we determined whether miR-1 and miR-133a expression in myoblasts is altered with cellular senescence and involved in senescence-impaired myogenic differentiation. C2C12 murine skeletal myoblasts were converted to a replicative senescent state by culturing to a high passage number. Although miR-1 and miR-133a expression was largely induced during myogenic differentiation, expression was suppressed in cells at high passage numbers (passage 10 and/or passage 20). Although the senescent myoblasts exhibited a deterioration of myogenic differentiation, transfection of miR-1 or miR-133a into myoblasts ameliorated cell fusion. Treatment with the glutaminase 1 inhibitor, BPTES, removed senescent cells from C2C12 myoblasts with a high passage number, whereas myotube formation and miR-133a expression was increased. In addition, primary cultured myoblasts prepared from aged C57BL/6J male mice (20 months old) exhibited a decrease in miR-1 and miR-133a levels compared with younger mice (3 months old). The results suggest that replicative senescence suppresses muscle-specific miRNA expression in myoblasts, which contributes to the senescence-related dysfunction of myogenic regeneration.

Citation: Shintani-Ishida K, Tsurumi R, Ikegaya H (2023) Decrease in the expression of muscle-specific miRNAs, miR-133a and miR-1, in myoblasts with replicative senescence. PLoS ONE 18(1): e0280527. https://doi.org/10.1371/journal.pone.0280527

Editor: Abdul Qadir Syed, Northwest University, UNITED STATES

Received: September 9, 2022; Accepted: December 29, 2022; Published: January 17, 2023

Copyright: © 2023 Shintani-Ishida et al. This is an open access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.

Data Availability: All relevant data are within the paper and its Supporting Information files.

Funding: This work was supported by JSPS KAKENHI Grant Number JP20K10558 (to KS). The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

Competing interests: The authors have declared that no competing interests exist.

Introduction

The regeneration of skeletal muscle involves repairing muscles that have been injured or atrophied by aging or disease. Muscle regeneration following myotrauma is triggered by the activation of resident muscle satellite (stem) cells (MuSCs). Activated MuSCs proliferate as myoblasts and induce the expression of muscle-specific genes, including myoblast determination protein 1 (MyoD1) [1], myogenin [2], and myosin heavy chain (MHC) [3], to promote myocyte differentiation. The myocytes migrate and fuse with other myocytes to form multinucleated myotubes that mature into myofibers or with damaged myofibers. The regenerative capacity of skeletal muscle declines with advancing age, and the impaired function of senescent MuSCs is partly responsible [4–6].

MicroRNAs (miRNAs) are short (~22 n) non-coding RNAs that post-transcriptionally regulate gene expression by binding to the 3′ untranslated region of target mRNAs [7, 8]. Over 1000 miRNAs are present in the human genome, and many exhibit tissue- or developmental stage-specific expression patterns [9, 10]. Muscle-specific or muscle-enriched miRNAs include miRNA (miR)-1, miR-133a, miR-133b, miR-206, miR-208a, miR-208b, miR-486, and miR-499 [11]. MiR-1, miR-133a/b, and miR-206 control the proliferation and differentiation of myoblasts [12]. MiR-1 accelerates myoblast differentiation by reducing the expression of histone deacetylase 4 (HDAC4), which inhibits MyoD1 [13]. MiR-206, which also targets HDAC4 [14], is skeletal muscle-specific, while miR-1 is both skeletal and cardiac muscle-specific. MiR-133a/b promotes myoblast proliferation by downregulating the serum response factor (SRF) [15]. MiR-1/miR-133a or miR-206/miR-133b is co-transcribed in the bicistronic transcript [15]. The MiR-206/133b cluster is likely to be dispensable for skeletal muscle regeneration because of the overlapping functions of the related miR-1/133a clusters [16]. MiR-208b and miR-499 are intronic miRNAs encoded in the myosin genes Myh7 and Myh7B, respectively, and they control muscle gene expression and performance [17]. Although miR-486 is expressed in other tissues as well (muscle-enriched miRNA), it is upregulated during myoblast differentiation and downregulates Pax7 as well as miR-1/miR-206 [18]. MiR-208a is not expressed in skeletal muscles (cardiac muscle-specific expression).

There are several reports that miR-1 and miR-133a, among these muscle-specific/enriched miRNAs, vary during aging. In the right atrial appendage samples from patients undergoing coronary artery bypass grafting, miR-1 and miR-133 expression levels were lower in aged patients (≥65 years old) than in adult patients (<65 years old) [19]. Quantitative analysis of miRNAs from the tibialis anterior (TA) muscle of adult mice (6 months old) and aged mice (24 months old) revealed that several miRNAs, including miR-133a, decreased with aging [20]. However, miR-1 and miR-133a alterations in MuSCs during the aging process have not been reported.

In this study, we determined whether miR-1 and miR-133a expression in MuSCs is altered with aging and examined the mechanism through which altered miRNA expression contributes to aging-impaired regeneration; C2C12 murine skeletal myoblasts with a high passage number were used as an aging model [21].

Materials and methods

Cell culture and passage

Mouse C2C12 cells were provided by the RIKEN BioResource Research Center through the National BioResource Project of the MEXT/AMED (Ibaraki, Japan). The cells were cultured in the growth medium (GM) consisting of Dulbecco’s modified Eagle medium (DMEM) (Invitrogen, Life technologies Japan, Tokyo, Japan) supplemented with 10% fetal bovine serum substitute EquaFETAL (Atlas Biologicals, Fort Collins, CO, USA), at 37°C and 5% CO₂ until the cells reached 60%–70% confluency. For each passage, the cells were trypsinized, diluted ten times, and seeded into new dishes for 72 h. Cells from the second passage were stored in a serum-free cell preservation medium Bambanker® (GC Lymphotec, Tokyo, Japan) at −80°C. For the experiments, the stock cells were thawed (passage 0) and passaging was repeated until a high passage number (passage 10 or passage 20) was achieved. Myogenic differentiation was induced by changing the GM to a differentiation medium (DM) consisting of 2% horse serum (Invitrogen^TM) with DMEM. The DM was changed every other day. After 7 days, myotube formation was evaluated.

Transfection

A 5 μM miRNA mimic, synthesized for miR-1 or miR-133a (AccuTarget^TM Human miRNA mimic, Bioneer Daejeon, Korea), was transfected into cells using Lipofectamine RNAiMAX Transfection Reagent (Invitrogen) following the manufacturer’s instructions. AccuTarget^TM miRNA mimic Negative Control #1 (Bioneer) was used as control miRNA.

Senolysis

Cells were seeded into 60-mm culture dishes and 96-well black plates. Cells at 60%–70% confluence were treated with GM containing 10 μM BPTES [Bis-2-(5-phenylacetamido-1,3,4-thiadiazol-2-yl)ethyl sulfide] (AdipGen, Füellinsdorf、Switzerland) for 48 h. The 96-well plates were used to determine the number of viable cells; the dishes were used for miRNA quantitation. Cells on the other dishes were re-seeded into new 60-mm culture dishes at 5 × 10⁵ and grown to 60%–70% confluence and differentiation was induced by DM exchange. The cells were collected for Western blot analysis and immunostaining with antibodies against MHC after incubation with DM for 7 days.

Immunostaining

The cells were fixed by incubation in cold methanol for 2 min and washed three times with phosphate-buffered saline (PBS). They were subsequently incubated with PBS containing 1% bovine serum albumin (BSA) (Sigma-Aldrich, St. Louis, MO, USA) at room temperature for 30 min followed by incubation with an anti-MHC monoclonal antibody (Clone MF20, R&D Systems, Minneapolis, MN, USA) at 4°C overnight. The antibody was diluted 1000-fold with PBS containing 1% BSA. The cells were washed and then incubated with Alexa Fluor 568–conjugated secondary antibody (ab175701, abcam, Cambridge, UK) at room temperature for 1 hour. After washing, they were mounted with a medium containing 4′,6-Diamidino-2-phenylindole dihydrochloride (DAPI) (Vector Laboratories, Burlingame, CA, USA). Using a fluorescence microscope (IX73, Olympus, Tokyo, Japan), four images were randomly selected from each well. The nuclei were counted using imaging software (cellSens, Olympus). The percentage of nuclei in the myotubes with MHC-positive staining was calculated as the fusion index.

Quantitative reverse transcription (RT)-PCR

The relative quantities of miR-1 and miR-133 were determined using quantitative RT-PCR. Total RNA was extracted from cells using ISOGEN extraction reagent (Nippon Gene, Tokyo, Japan). Quantitative RT-PCR for miR-1 and miR-133a was performed using a TaqMan™ MicroRNA Assay kit (assay ID: 002222 for hsa-miR-1, 002246 for hsa-miR-133a) according to the manufacturer’s protocol. U6 snRNA was used as an endogenous control (assay ID: 001973). The threshold cycle (CT) values for the genes were determined with a StepOnePlus™ Real-Time PCR System (Thermo Fisher Scientific, Tokyo, Japan). The amount of the target gene relative to the endogenous control gene was normalized with each control sample by the ΔΔCT method.

Western blot analysis

Cells were sonicated in an ice-cold solution of 320 mM sucrose, 10 mM Tris–HCl (pH 7.4), 1 mM EDTA, 50 mM NaF, 1 mM Na₃VO₄, and a protease inhibitor cocktail (P8340, Sigma-Aldrich). The lysates were solubilized in Laemmli sample buffer and subjected to 10% and 15% SDS-PAGE for MHC, MyoD, Myogenin, and β-tubulin and for p16^INK4a and p21, respectively. After transfer, the membranes were incubated with antibodies specific to MHC (Clone MF20, R&D Systems), MyoD (18943-1-AP, Proteintech, Tokyo, Japan), β-tubulin (ab179513, abcam), p16^INK4a (10883-1-ap, Proteintech), or p21 (28248-1-AP, Proteintech). The primary antibodies were diluted 1000-fold with 1% BSA in Tris-buffered saline containing Tween-20 (BSA/TBS-T). The secondary antibody, horseradish peroxidase-conjugated goat anti-mouse or anti-rabbit IgG (Promega, Madison, WI), was diluted 5000-fold with 1% BSA/TBS-T. The immune complexes were detected with chemiluminescent reagents (Western Lightning-ECL; PerkinElmer, Waltham, MA). The band intensities were measured using the ChemiDoc Touch imaging system (BioRad Laboratories, Hercules, CA, USA).

Plate assay

SPiDER-βGal cellular senescence plate assay kit (Dojindo Laboratories, Kumamoto, Japan) was used to determine senescence-associated β-galactosidase activity according to the manual instructions.

CellTiter-Glo® 2.0 Cell Viability Assay (Promega) was used to determine the number of viable cells after senolysis.

Isolation of primary myoblasts

Primary myoblasts were isolated from the skeletal muscles of C57BL/6J male mice [22]. All methods were carried out in accordance with the Guidelines for Proper Conduct of Animal Experiments by the Science Council of Japan. The experimental protocol used in this study was reviewed and approved by the experimental animal committee of the Kyoto Prefectural University of Medicine (Permit No. M2020-1). The study was carried out in compliance with the ARRIVE guidelines. Skeletal muscle tissues of the hindlimbs were digested with collagenase II (Sigma-Aldrich Japan, Tokyo, Japan). Dissociated cells were suspended in Ham’s F-10 medium (InvitrogenTM) containing 20% EquaFETAL (Atlas Biologicals) and 2 ng/mL of human recombinant basic fibroblast growth factor (bFGF) (Nacalai Tesque, Kyoto, Japan). They were seeded onto noncoated dishes and incubated at 37°C in 5% CO₂ overnight to remove the fibroblasts. The next day, the supernatant was collected and centrifuged. The pellets were resuspended in Ham’s F-10 medium, which contains 20% EqualFETAL and 2 ng/mL of bFGF, and cultured on collagen-coated dishes until 70%–80% confluent.

Statistics

The quantitative data are presented as the means ± SE. Data were analyzed using the Tukey–Kramer test, Dunnett′s test, or a t-test. p < 0.05 was considered statistically significant.

Results

Replicative senescence suppresses the myogenic differentiation of C2C12 cells

To determine the effect of replicative senescence on the proliferation and differentiation of C2C12 cells, we repeated cell passage 10 or 20 times and induced myogenic differentiation by culturing the cells with DM for 7 days. The senescence markers p16^INK4a (Fig 1A) and p21 (Fig 1B) were induced by the 10th passage. Senescence-associated β-galactosidase was also activated by the 10th passage (Fig 1C). The infusion index decreased by 63% by passage 10 and by 88% by passage 20 compared with the control (passage 1) (Fig 2A and 2B). The cell numbers decreased by passage 10 but returned to those of the control level (passage 1) by passage 20. The expression of the MyoD1 and myogenin proteins, markers for the early phase of myogenic differentiation, and MHC, a marker for myotubes, was induced by DM incubation in passage 1 (Fig 3) However, the induction of MyoD1 and myogenin was decreased by passage 20 (Fig 3). The induction of MHC was suppressed in a stepwise manner by passages 10 and 20 (Fig 3). These results indicated that replicative senescence suppressed myogenic differentiation and that the effect was likely to expand with developing differentiation.

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Fig 1. Induction of replicative senescence by high passage number in C2C12 cells.

Panels A and B show representative western blots of p16^INK4a and p21 as senescence markers and tubulin as an internal control, respectively. Cells were subjected to passage once (passage 1), 10 times (passage 10), or 20 times (passage 20). The band density of p16^INK4a or p21 was normalized with that of tubulin. The normalized density was compared with the average of the normalized density in passage 1. Data are presented as means ± SE (n = 4 dishes). *P <0.05 (Dunnett′s test). Panel C shows the relative β-galactosidase activity. Data are presented as means ± SE (n = 6 wells). *P < 0.05, **P < 0.01 (Tukey–Kramer test).

https://doi.org/10.1371/journal.pone.0280527.g001

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Fig 2. Effect of replicative senescence on myogenic differentiation of C2C12 cells.

Panel A shows representative images of C2C12 cells after culture with differentiation medium for 7 days. Myogenic differentiation was induced in cells subjected to passage once (passage 1), 10 times (passage 10), or 20 times (passage 20). Red fluorescence of myosin heavy chain (MHC), blue fluorescence of nuclei, and phage contrast images were merged. Panel B shows the infusion index and the number of cells after myogenic differentiation for 7 days. Data are presented as means ± SE (n = 8 wells). *P < 0.05, **P < 0.01; n.s., not significant (Tukey–Kramer test).

https://doi.org/10.1371/journal.pone.0280527.g002

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Fig 3. Effect of replicative senescence on myogenic marker expression during myogenic differentiation of C2C12 cells.

Panel A shows representative Western blots of MyoD1, Myogenin, myosin heavy chain (MHC), and tubulin (as an internal control) in C2C12 cells after myogenic differentiation. Cells were collected 7 days after differentiation for MyoD and MHC or 5 days after differentiation for myogenin. Panel B shows the time course of MyoD1, myogenin, and MHC protein expression after changing the medium for myogenic differentiation. The band density of MyoD, myogenin, or MHC was normalized with that of tubulin. The normalized density was compared with the average of the normalized density in passage 1 just before myogenic differentiation (day 0) in the MyoD and myogenin expressions. Because MHC expression was undetectable on day 0, the normalized density was compared with the average of the normalized density in passage 1, 7 days after myogenic differentiation. Data are presented as means ± SE (n = 4 dishes). *P < 0.05, **P < 0.01 vs. passage 1 at each time point; †P < 0.05, ††P < 0.01 vs. passage 10 at each time point (Tukey–Kramer test).

https://doi.org/10.1371/journal.pone.0280527.g003

Replicative senescence suppresses miR-1 and miR-133a expression

Expression of either miR-1 or miR-133a gradually increased during culture with DM (Fig 4A). Expression of another muscle-specific miRNA cluster, miR-206/miR-133b, also increased during culture with DM (S1 Fig). The levels of miR-1 and miR-133a increased by 186-fold and 22-fold, respectively, 7 days after cell culture in DM (Fig 4A), while those of miR-206 and miR-133b increased by 15-fold and 11-fold, respectively (S1 Fig). The effects of replicative senescence on the expression levels of miR-1/miR-133a were examined. The induction of miR-1 expression was suppressed in cells at passage 20, but not in cells at passage 10. The induction of miR-133a was suppressed in cells at passage 10 and further in passage 20. These suppressive effects in replicative senescent cells had already appeared before myogenic differentiation was induced (Fig 4B).

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Fig 4. Effect of replicative senescence on miR-1 and miR-133a expression.

Panel A shows the time course of miR-1 and miR-133a levels after changing the medium for myogenic differentiation. Myogenic differentiation was induced in cells subjected to passage once (passage 1), 10 times (passage 10), or 20 times (passage 20). Data are presented as means ± SE (n = 6 dishes). *P < 0.05, **P < 0.01 vs. passage 1 at each time point (Tukey–Kramer test). Panel B shows the relative levels of miR-1 and miR-133a in cells just before changing the medium for myogenic differentiation (day 0 in Fig 4A). Data are presented as means ± SE (n = 6 dishes). **P < 0.01 vs. passage 1 (Dunnett’s test).

https://doi.org/10.1371/journal.pone.0280527.g004

Overexpression of miRNAs ameliorates the differentiation capacity of senescent C2C12 cells

To determine whether the upregulation of miR-1 or/and miR-133a ameliorates the deterioration of myotube formation in senescent cells, miR-1 mimic, miR-133a mimic, or both were transfected into C2C12 cells with high passage numbers (passage 10) and the fusion index was determined. Because cells at the 20th passage could not survive lipofectamine treatment for miRNA transfection, the experiments on the 10th passage cells which were tolerant to transfection were carried out. MiR-1 and/or miR-133a were overexpressed in the transfected cells (S2 Fig). Transfection with miR-133a or miR-1 increased the infusion index compared with the negative control miRNA (Fig 5A). Co-transfection with miR-133a and miR-1 further improved the infusion index. Although there was no difference in MyoD1 and myogenin expression between mock cells and muscle-specific miRNA-transfected cells (Fig 5B–5D), the cells transfected with miR-1 or cells co-transfected with miR-133a and miR-1 exhibited increased MHC levels, whereas cells transfected with miR-133a did not express different MHC levels (Fig 5B and 5E). These results showed that the overexpression of both miR-1 and miR-133a was required to ameliorate the deterioration of myotube formation in senescent cells.

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Fig 5. Effect of miRNA mimic transfection on myogenic differentiation in replicative senescent C2C12 cells.

Panel A shows the infusion index after 7 days of differentiation in cells transfected with miR-133a, miR-1, or both. Mock cells were transfected with a negative control miRNA. Data are presented as means ± SE (n = 10 dishes). **P < 0.01 (Tukey–Kramer test). Panels B–E show representative blots of MyoD1, myogenin, MHC, and tubulin (B), and quantitative data (C–E) in cells after differentiation for 7 days. The band density of MyoD1, myogenin, or myosin heavy chain (MHC) was normalized with that of tubulin. The normalized density was compared with the average of the normalized density in the mock group. Data are presented as means ± SE (n = 5 dishes). *P < 0.05 (Tukey–Kramer test).

https://doi.org/10.1371/journal.pone.0280527.g005

Senolysis by glutaminase inhibitor increases miR-133a expression

To determine whether the expression of miR-1 and miR-133a decreased as cellular senescence accelerated in C2C12 cells, senescent cells were removed by glutaminolysis inhibition and miRNA expression levels were measured in the surviving cells. Treatment with the glutaminase 1 inhibitor, BPTES [bis-2-(5-phenyl-acetamido-1,3,4-thiadiazol-2-yl) ethyl sulfide] [23], for 2 days removed 24% of the cells at passage 10 and 42% of the cells at passage 20, whereas it had no effect on the cells at passage 1 (Fig 6A). Cells that survived senolysis decreased p21 expression to 60% and increased differentiation capacity by 3-fold (Fig 6C) and MHC expression by 30-fold (Fig 6D) compared with untreated cells at passage 20. In surviving cells, the expression of miR-1 was slightly higher than that in untreated cells, whereas miR-133a levels increased two-fold (Fig 6E). These results indicated that cellular senescence decreased miR-1 and miR-133a expressions as well as differentiation capacity.

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Fig 6. Effect of senolysis on the differentiation capacity of replicative senescent C2C12 cells.

Panel A shows the relative cell number after treatment with BPTES or 1% DMSO for 48 h following repeating passage once (passage 1), 10 times (passage 10), or 20 times (passage 20). Data are presented as means ± SE (n = 8 wells). **P < 0.01 vs. 1% DMSO at each passage number (Tukey–Kramer test). Panel B shows representative blots of p21 and tubulin, and quantitative data normalized with the tubulin in cells surviving BPTES treatment following repeated passage 20 times. Data are presented as means ± SE (n = 4 dishes). *P < 0.05 (t-test). Panel C shows representative myosin heavy chain (MHC) immunostaining images and the number of MHC-positive cells. Cells that survived BPTES treatment following repeated passage 20 times were subjected to differentiation for 7 days and were stained with DAPI (blue fluorescence) and an antibody against MHC (red fluorescence). Data are presented as means ± SE (n = 7 dishes). **P < 0.01 (t-test). Panel D shows representative MHC and tubulin blots and quantitative data normalized with tubulin in cells surviving BPTES treatment following repeated passage 20 times. Data are presented as means ± SE (n = 4 dishes). **P < 0.01 (t-test). Panel E shows miR-1 and miR-133a levels in cells surviving BPTES treatment following repeated passage 20 times. Data are presented as means ± SE (n = 4 dishes). **P < 0.01 (t-test).

https://doi.org/10.1371/journal.pone.0280527.g006

Expression of miR-1 and miR-133a in myoblasts decreases in aged mice

Myoblasts were isolated from young mice (3 months) and aged mice (20 months). The isolated myoblast fraction did not contain MHC-positive muscle cells (Fig 7A). The miR-1 and miR-133a levels in aged mice were only one tenth of those in young mice (Fig 7B) in the isolated myoblast fraction.

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Fig 7. Comparison of miRNA levels in isolated myoblasts between young and aged mice.

Panel A shows a representative immunocytochemical image of the isolated myoblast fraction. Green fluorescence of Pax 7, red fluorescence of myosin heavy chain (MHC), and blue fluorescence of nuclei images were merged. Panel B shows miR-1 and miR-133a levels in myoblasts isolated from young and aged mice. Data are presented as means ± SE (n = 4). **P <0.01 (t-test).

https://doi.org/10.1371/journal.pone.0280527.g007

Discussion

In this study, we showed for the first time that muscle-specific miRNA (miR-1 and miR-133a) expression in myoblasts is downregulated with replicative senescence using C2C12 myoblasts with a high passage number. These miRNAs in myoblasts from aged mice also decreased.

There have been several studies demonstrating the myogenic differentiation capacity of C2C12 cells declining at high passage number. Repeated passage up to 30 times reduced the length and area of myotubes formed from C2C12 cells [24]. Late passage up to 40 times impaired myocyte fusion and suppressed the expression of myogenic regulatory factors including MyoD1 [25]. In the present study, the infusion index began to decrease after the 10th passage (Fig 2), when replicative senescence was induced (Fig 1). Induction of MyoD1 and myogenin, early markers of myogenic differentiation, ceased by the 20th passage [25], whereas that of MHC was gradually suppressed after the 10th passage (Fig 3). These results suggest that myogenic differentiation becomes more senescent-sensitive with progression.

MiR-1 and miR-133a are organized as a bicistronic cluster and are thus transcribed together. Although miR-133a and miR-1 expressions were largely induced during myogenic differentiation, they decreased after the 10th and 20th passages, respectively. MiR-1 promotes myogenesis by inhibiting the expression of HDAC4, which represses the essential muscle-related transcriptional factors, MEF2C and MyoD1 [13]. At the same time, MyoD1 up-regulates miR-1 [26, 27]. Therefore, it is likely that both the induction of miR-1 and MyoD1 failed at the same passage number [25]. In contrast, the induction of miR-133a was suppressed after the 10th passage, when replicative senescence was induced and myogenic differentiation capacity deteriorated. In addition, the ameliorating effect of senolysis on miRNA expression was larger in miR-133a compared with miR-1 (Fig 6). Thus, it appeared that miR-133a was more sensitive to cellular senescence compared to miR-1. Expression of another muscle-specific miRNA cluster, miR-206/133b, also induced during myogenic differentiation, but the induction was less compared to that of the miR-1/133a cluster. In the miR-206/miR-133b knock-out mice, the deletion of the miR-206/133 cluster does not affect skeletal muscle regeneration in the Duchenne muscular dystrophy model mice [16]. The study suggested that the miR-206/133b cluster is likely to be dispensable for skeletal muscle regeneration because of the overlapping functions of the related miR-1/133a clusters [16]. The miR-1/133a cluster may be more dominant in myogenic differentiation.

Overexpression of either miR-133a or miR-1 ameliorated cell fusion in senescent C2C12 cells (Fig 5). Co-transfection with miR-133a and miR-1 further increased the fusion index. Misexpression of either miR-133a or miR-1 resulted in somite development defects in Xenopus laevis in vivo [15]. These findings suggest that the correct temporal expression and amount of both miR-133a and miR-1 are required for myogenic regeneration. In contrast, overexpression of miRNA-133a increased the infusion index without the induction of MHC expression (Fig 5). Tetraplex-binding porphyrin-treated C2C12 cells blocked MyoD1 transcription and ceased MHC induction but formed myotubes during myogenic differentiation [28]. MyoD1 induction was not repressed in cells at the 10th passage, and the transfection of miR-133a or miR-1 did not affect MyoD1 expression.

Sarcopenia is a condition characterized by the progressive and generalized loss of muscle mass, strength, and function [29, 30]. The prevalence of sarcopenia is related to skeletal muscle loss with aging [31]. Although the pathogenesis of sarcopenia remains unclear, sarcopenia is likely to be closely related to MuSC dysfunction [32]. MuSCs from 24-month-old mice exhibited a loss in migration speed, which is a key component of skeletal muscle regeneration [33]. The pool size and proliferation potential of MuSCs declined with age in mice [34]. In contrast, in satellite cell-depleted Pax7^CreER-DTA mice, lifelong satellite cell reduction did not affect age-related myofiber atrophy and weakness, whereas it exacerbated the age-related excessive accumulation of extracellular matrix (ECM) in hind limb muscles [35]. Satellite cell reduction increased muscle fibroblast content, resulting in ECM accumulation [36]. Mice with deleted Trbp, an HIV TAR RNA-binding protein, exhibited reduced expression of miR-1 and miR-133a, impaired muscle regeneration and enhanced fibrosis in response to muscle injury [37]. Injection of miR-133a-overexpressed cardiac progenitor cells markedly improved cardiac function in a rat myocardial infarction model by reducing fibrosis and hypertrophy [38]. The role of muscle-specific miRNAs in myoblasts, including not only maintaining myogenic regeneration, but also protecting muscle fibrosis warrant further study.

Supporting information

S1 Fig.

Time course of miR-206 (panel A) and miR-133b (panel B) levels after changing the medium for myogenic differentiation. Myogenic differentiation was induced in cells subjected to passage once (passage 1). Data are presented as means ± SE (n = 4 dishes).

https://doi.org/10.1371/journal.pone.0280527.s001

(TIF)

S2 Fig.

The miR-1 (panel A) and miR-133a (panel B) expression levels in replicative senescent C2C12 cells transfected with miRNA mimic. Cells subjected to passage 10 times were transfected with miR-1, miR-133a, or both and collected before the induction of differentiation for miRNA quantitation. Mock cells were transfected with a negative control miRNA. Data are presented as means ± SE (n = 4 dishes). **P < 0.01 vs. mock (Dunnett′s test).

https://doi.org/10.1371/journal.pone.0280527.s002

(TIF)

S1 Raw images. Raw images of western blots for Figs 1, 3, 5 and 6.

https://doi.org/10.1371/journal.pone.0280527.s003

(PDF)

References

1. Sabourin LA, Girgis-Gabardo A, Seale P, Asakura A, Rudnicki MA. Reduced differentiation potential of primary MyoD-/- myogenic cells derived from adult skeletal muscle. J Cell Biol. 1999;144:631–643. pmid:10037786
- View Article
- PubMed/NCBI
- Google Scholar
2. Wright WE, Sassoon DA, Lin VK. Myogenin, a factor regulating myogenesis, has a domain homologous to MyoD. Cell. 1989;56:607–617. pmid:2537150
- View Article
- PubMed/NCBI
- Google Scholar
3. John HA, Patrinou-Georgoulas M, Jones KW. Detection of myosin heavy chain mRNA during myogenesis in tissue culture by in vitro and in situ hybridization. Cell. 1977;12:501–508. pmid:912755
- View Article
- PubMed/NCBI
- Google Scholar
4. Yamakawa H, Kusumoto D, Hashimoto H, Yuasa S. Stem Cell Aging in Skeletal Muscle Regeneration and Disease. Int J Mol Sci. 2020;21:1830–1848. pmid:32155842
- View Article
- PubMed/NCBI
- Google Scholar
5. Sousa-Victor P, García-Prat L, Muñoz-Cánoves P. Control of satellite cell function in muscle regeneration and its disruption in ageing. Nat Rev Mol Cell Biol. 2022;23:204–226. pmid:34663964
- View Article
- PubMed/NCBI
- Google Scholar
6. Blau HM, Cosgrove BD, Ho AT. The central role of muscle stem cells in regenerative failure with aging. Nat Med. 2015;21:854–862. pmid:26248268
- View Article
- PubMed/NCBI
- Google Scholar
7. Bartel DP. MicroRNAs: genomics, biogenesis, mechanism, and function. Cell. 2004;116:281–297. pmid:14744438
- View Article
- PubMed/NCBI
- Google Scholar
8. Lewis BP, Shih IH, Jones-Rhoades MW, Bartel DP, Burge CB. Prediction of mammalian microRNA targets. Cell. 2003;115:787–798. pmid:14697198
- View Article
- PubMed/NCBI
- Google Scholar
9. Grishok A, Pasquinelli AE, Conte D, Li N, Parrish S, Ha I, et al. Genes and mechanisms related to RNA interference regulate expression of the small temporal RNAs that control C. elegans developmental timing. Cell. 2001;106:23–34. pmid:11461699
- View Article
- PubMed/NCBI
- Google Scholar
10. Zhao Y, Srivastava D. A developmental view of microRNA function. Trends Biochem Sci. 2007;32:189–197. pmid:17350266
- View Article
- PubMed/NCBI
- Google Scholar
11. Horak M, Novak J, Bienertova-Vasku J. Muscle-specific microRNAs in skeletal muscle development. Dev Biol. 2016;410:1–13. pmid:26708096
- View Article
- PubMed/NCBI
- Google Scholar
12. Nakasa T, Ishikawa M, Shi M, Shibuya H, Adachi N, Ochi M. Acceleration of muscle regeneration by local injection of muscle-specific microRNAs in rat skeletal muscle injury model. J Cell Mol Med. 2010;14:2495–2505. pmid:19754672
- View Article
- PubMed/NCBI
- Google Scholar
13. Lu J, McKinsey TA, Zhang CL, Olson EN. Regulation of skeletal myogenesis by association of the MEF2 transcription factor with class II histone deacetylases. Mol Cell. 2000;6:233–244. pmid:10983972
- View Article
- PubMed/NCBI
- Google Scholar
14. Winbanks CE, Wang B, Beyer C, Koh P, White L, Kantharidis P, et al. TGF-beta regulates miR-206 and miR-29 to control myogenic differentiation through regulation of HDAC4. J Biol Chem. 2011;286:13805–13814. pmid:21324893
- View Article
- PubMed/NCBI
- Google Scholar
15. Chen JF, Mandel EM, Thomson JM, Wu Q, Callis TE, Hammond SM, et al. The role of microRNA-1 and microRNA-133 in skeletal muscle proliferation and differentiation. Nat Genet. 2006;38:228–233. pmid:16380711
- View Article
- PubMed/NCBI
- Google Scholar
16. Boettger T, Wüst S, Nolte H, Braun T. The miR-206/133b cluster is dispensable for development, survival and regeneration of skeletal muscle. Skelet Muscle. 2014;4:23. pmid:25530839
- View Article
- PubMed/NCBI
- Google Scholar
17. van Rooij E, Quiat D, Johnson BA, Sutherland LB, Qi X, Richardson JA, et al. A family of microRNAs encoded by myosin genes governs myosin expression and muscle performance. Dev Cell. 2009:17:662–673. pmid:19922871
- View Article
- PubMed/NCBI
- Google Scholar
18. Dey BK, Gagan J, Dutta A. MiR-206 and -486 induce myoblast differentiation by downregulating Pax7. Mol Cell Biol. 2011;31:203–214. pmid:21041476
- View Article
- PubMed/NCBI
- Google Scholar
19. Li YD, Hong YF, Yusufuaji Y, Tang BP, Zhou XH, Xu GJ, et al. Altered expression of hyperpolarization-activated cyclic nucleotide-gated channels and microRNA-1 and -133 in patients with age-associated atrial fibrillation. Mol Med Rep. 2015;12:3243–3248 pmid:26005035
- View Article
- PubMed/NCBI
- Google Scholar
20. Soriano-Arroquia A, House L, Tregilgas L, Canty-Laird E, Goljanek-Whysall K. The functional consequences of age-related changes in microRNA expression in skeletal muscle. Biogerontology. 2016;17:641–654. pmid:26922183
- View Article
- PubMed/NCBI
- Google Scholar
21. Sharples AP, Al-Shanti N, Lewis MP, Stewart CE. Reduction of myoblast differentiation following multiple population doublings in mouse C2 C12 cells: a model to investigate ageing? J Cell Biochem. 2011;112:3773–3785. pmid:21826704
- View Article
- PubMed/NCBI
- Google Scholar
22. Rando TA, Blau HM. Primary mouse myoblast purification, characterization, and transplantation for cell-mediated gene therapy. J Cell Biol. 1994;125:1275–1287. pmid:8207057
- View Article
- PubMed/NCBI
- Google Scholar
23. Johmura Y, Yamanaka T, Omori S, Wang TW, Sugiura Y, Matsumoto M, et al. Senolysis by glutaminolysis inhibition ameliorates various age-associated disorders. Science. 2021;371:265–270. pmid:33446552
- View Article
- PubMed/NCBI
- Google Scholar
24. Chang YC, Liu HW, Chan YC, Hu SH, Liu MY, Chang SJ. The green tea polyphenol epigallocatechin-3-gallate attenuates age-associated muscle loss via regulation of miR-486-5p and myostatin. Arch Biochem Biophys. 2020;692:108511. pmid:32710883
- View Article
- PubMed/NCBI
- Google Scholar
25. Shahini A, Choudhury D, Asmani M, Zhao R, Lei P, Andreadis ST. NANOG restores the impaired myogenic differentiation potential of skeletal myoblasts after multiple population doublings. Stem Cell Res. 2018;26:55–66. pmid:29245050
- View Article
- PubMed/NCBI
- Google Scholar
26. Hirai H, Verma M, Watanabe S, Tastad C, Asakura Y, Asakura A. MyoD regulates apoptosis of myoblasts through microRNA-mediated down-regulation of Pax3. J Cell Biol. 2010;191:347–365. pmid:20956382
- View Article
- PubMed/NCBI
- Google Scholar
27. Sun Y, Ge Y, Drnevich J, Zhao Y, Band M, Chen J. Mammalian target of rapamycin regulates miRNA-1 and follistatin in skeletal myogenesis. J Cell Biol. 2010;189:1157–1169. pmid:20566686
- View Article
- PubMed/NCBI
- Google Scholar
28. Testa S, D’Addabbo P, Fornetti E, Belli R, Fuoco C, Bernardini S, et al. Myoblast myogenic differentiation but not fusion process is inhibited via MyoD tetraplex interaction. Oxid Med Cell Longev. 2018;2018:7640272. pmid:29854094
- View Article
- PubMed/NCBI
- Google Scholar
29. Rosenberg IH. Sarcopenia: origins and clinical relevance. J Nutr. 1997;127:990S–991S. pmid:9164280
- View Article
- PubMed/NCBI
- Google Scholar
30. Santilli V, Bernetti A, Mangone M, Paoloni M. Clinical definition of sarcopenia. Clin Cases Miner Bone Metab. 2014;11:177–180. pmid:25568649
- View Article
- PubMed/NCBI
- Google Scholar
31. Janssen I, Heymsfield SB, Ross R. Low relative skeletal muscle mass (sarcopenia) in older persons is associated with functional impairment and physical disability. J Am Geriatr Soc. 2002;50:889–896. pmid:12028177
- View Article
- PubMed/NCBI
- Google Scholar
32. Cai Z, Liu D, Yang Y, Xie W, He M, Yu D, et al. The role and therapeutic potential of stem cells in skeletal muscle in sarcopenia. Stem Cell Res Ther. 2022;13:28. pmid:35073997
- View Article
- PubMed/NCBI
- Google Scholar
33. Collins-Hooper H, Woolley TE, Dyson L, Patel A, Potter P, Baker RE, et al. Age-related changes in speed and mechanism of adult skeletal muscle stem cell migration. Stem Cells. 2012;30(6):1182–1195. pmid:22593017
- View Article
- PubMed/NCBI
- Google Scholar
34. Shefer G, Van DP, Richardson JB, Yablonka-Reuveni Z. Satellite-cell pool size does matter: defining the myogenic potency of aging skeletal muscle. Dev Biol. 2006;294:50–66. pmid:16554047
- View Article
- PubMed/NCBI
- Google Scholar
35. Fry CS, Lee JD, Mula J, Kirby TJ, Jackson JR, Liu F, et al. Inducible depletion of satellite cells in adult, sedentary mice impairs muscle regenerative capacity without affecting sarcopenia. Nat Med. 2015;21:76–80. pmid:25501907
- View Article
- PubMed/NCBI
- Google Scholar
36. Fry CS, Lee JD, Jackson JR, Kirby TJ, Stasko SA, Liu H, et al. Regulation of the muscle fiber microenvironment by activated satellite cells during hypertrophy. FASEB J. 2014;28:1654–1665. pmid:24376025
- View Article
- PubMed/NCBI
- Google Scholar
37. Ding J, Nie M, Liu J, Hu X, Ma L, Deng ZL, et al. Trbp is required for differentiation of myoblasts and normal regeneration of skeletal muscle. PLoS One. 2016;11:e0155349. pmid:27159388
- View Article
- PubMed/NCBI
- Google Scholar
38. Izarra A, Moscoso I, Levent E, Cañón S, Cerrada I, Díez -Juan A, et al. miR-133a enhances the protective capacity of cardiac progenitors cells after myocardial infarction. Stem Cell Reports. 2014;3:1029–1042. pmid:25465869
- View Article
- PubMed/NCBI
- Google Scholar

[ref1] 1. Sabourin LA, Girgis-Gabardo A, Seale P, Asakura A, Rudnicki MA. Reduced differentiation potential of primary MyoD-/- myogenic cells derived from adult skeletal muscle. J Cell Biol. 1999;144:631–643. pmid:10037786
View Article
PubMed/NCBI
Google Scholar

[2] View Article

[3] PubMed/NCBI

[4] Google Scholar

[ref2] 2. Wright WE, Sassoon DA, Lin VK. Myogenin, a factor regulating myogenesis, has a domain homologous to MyoD. Cell. 1989;56:607–617. pmid:2537150
View Article
PubMed/NCBI
Google Scholar

[6] View Article

[7] PubMed/NCBI

[8] Google Scholar

[ref3] 3. John HA, Patrinou-Georgoulas M, Jones KW. Detection of myosin heavy chain mRNA during myogenesis in tissue culture by in vitro and in situ hybridization. Cell. 1977;12:501–508. pmid:912755
View Article
PubMed/NCBI
Google Scholar

[10] View Article

[11] PubMed/NCBI

[12] Google Scholar

[ref4] 4. Yamakawa H, Kusumoto D, Hashimoto H, Yuasa S. Stem Cell Aging in Skeletal Muscle Regeneration and Disease. Int J Mol Sci. 2020;21:1830–1848. pmid:32155842
View Article
PubMed/NCBI
Google Scholar

[14] View Article

[15] PubMed/NCBI

[16] Google Scholar

[ref5] 5. Sousa-Victor P, García-Prat L, Muñoz-Cánoves P. Control of satellite cell function in muscle regeneration and its disruption in ageing. Nat Rev Mol Cell Biol. 2022;23:204–226. pmid:34663964
View Article
PubMed/NCBI
Google Scholar

[18] View Article

[19] PubMed/NCBI

[20] Google Scholar

[ref6] 6. Blau HM, Cosgrove BD, Ho AT. The central role of muscle stem cells in regenerative failure with aging. Nat Med. 2015;21:854–862. pmid:26248268
View Article
PubMed/NCBI
Google Scholar

[22] View Article

[23] PubMed/NCBI

[24] Google Scholar

[ref7] 7. Bartel DP. MicroRNAs: genomics, biogenesis, mechanism, and function. Cell. 2004;116:281–297. pmid:14744438
View Article
PubMed/NCBI
Google Scholar

[26] View Article

[27] PubMed/NCBI

[28] Google Scholar

[ref8] 8. Lewis BP, Shih IH, Jones-Rhoades MW, Bartel DP, Burge CB. Prediction of mammalian microRNA targets. Cell. 2003;115:787–798. pmid:14697198
View Article
PubMed/NCBI
Google Scholar

[30] View Article

[31] PubMed/NCBI

[32] Google Scholar

[ref9] 9. Grishok A, Pasquinelli AE, Conte D, Li N, Parrish S, Ha I, et al. Genes and mechanisms related to RNA interference regulate expression of the small temporal RNAs that control C. elegans developmental timing. Cell. 2001;106:23–34. pmid:11461699
View Article
PubMed/NCBI
Google Scholar

[34] View Article

[35] PubMed/NCBI

[36] Google Scholar

[ref10] 10. Zhao Y, Srivastava D. A developmental view of microRNA function. Trends Biochem Sci. 2007;32:189–197. pmid:17350266
View Article
PubMed/NCBI
Google Scholar

[38] View Article

[39] PubMed/NCBI

[40] Google Scholar

[ref11] 11. Horak M, Novak J, Bienertova-Vasku J. Muscle-specific microRNAs in skeletal muscle development. Dev Biol. 2016;410:1–13. pmid:26708096
View Article
PubMed/NCBI
Google Scholar

[42] View Article

[43] PubMed/NCBI

[44] Google Scholar

[ref12] 12. Nakasa T, Ishikawa M, Shi M, Shibuya H, Adachi N, Ochi M. Acceleration of muscle regeneration by local injection of muscle-specific microRNAs in rat skeletal muscle injury model. J Cell Mol Med. 2010;14:2495–2505. pmid:19754672
View Article
PubMed/NCBI
Google Scholar

[46] View Article

[47] PubMed/NCBI

[48] Google Scholar

[ref13] 13. Lu J, McKinsey TA, Zhang CL, Olson EN. Regulation of skeletal myogenesis by association of the MEF2 transcription factor with class II histone deacetylases. Mol Cell. 2000;6:233–244. pmid:10983972
View Article
PubMed/NCBI
Google Scholar

[50] View Article

[51] PubMed/NCBI

[52] Google Scholar

[ref14] 14. Winbanks CE, Wang B, Beyer C, Koh P, White L, Kantharidis P, et al. TGF-beta regulates miR-206 and miR-29 to control myogenic differentiation through regulation of HDAC4. J Biol Chem. 2011;286:13805–13814. pmid:21324893
View Article
PubMed/NCBI
Google Scholar

[54] View Article

[55] PubMed/NCBI

[56] Google Scholar

[ref15] 15. Chen JF, Mandel EM, Thomson JM, Wu Q, Callis TE, Hammond SM, et al. The role of microRNA-1 and microRNA-133 in skeletal muscle proliferation and differentiation. Nat Genet. 2006;38:228–233. pmid:16380711
View Article
PubMed/NCBI
Google Scholar

[58] View Article

[59] PubMed/NCBI

[60] Google Scholar

[ref16] 16. Boettger T, Wüst S, Nolte H, Braun T. The miR-206/133b cluster is dispensable for development, survival and regeneration of skeletal muscle. Skelet Muscle. 2014;4:23. pmid:25530839
View Article
PubMed/NCBI
Google Scholar

[62] View Article

[63] PubMed/NCBI

[64] Google Scholar

[ref17] 17. van Rooij E, Quiat D, Johnson BA, Sutherland LB, Qi X, Richardson JA, et al. A family of microRNAs encoded by myosin genes governs myosin expression and muscle performance. Dev Cell. 2009:17:662–673. pmid:19922871
View Article
PubMed/NCBI
Google Scholar

[66] View Article

[67] PubMed/NCBI

[68] Google Scholar

[ref18] 18. Dey BK, Gagan J, Dutta A. MiR-206 and -486 induce myoblast differentiation by downregulating Pax7. Mol Cell Biol. 2011;31:203–214. pmid:21041476
View Article
PubMed/NCBI
Google Scholar

[70] View Article

[71] PubMed/NCBI

[72] Google Scholar

[ref19] 19. Li YD, Hong YF, Yusufuaji Y, Tang BP, Zhou XH, Xu GJ, et al. Altered expression of hyperpolarization-activated cyclic nucleotide-gated channels and microRNA-1 and -133 in patients with age-associated atrial fibrillation. Mol Med Rep. 2015;12:3243–3248 pmid:26005035
View Article
PubMed/NCBI
Google Scholar

[74] View Article

[75] PubMed/NCBI

[76] Google Scholar

[ref20] 20. Soriano-Arroquia A, House L, Tregilgas L, Canty-Laird E, Goljanek-Whysall K. The functional consequences of age-related changes in microRNA expression in skeletal muscle. Biogerontology. 2016;17:641–654. pmid:26922183
View Article
PubMed/NCBI
Google Scholar

[78] View Article

[79] PubMed/NCBI

[80] Google Scholar

[ref21] 21. Sharples AP, Al-Shanti N, Lewis MP, Stewart CE. Reduction of myoblast differentiation following multiple population doublings in mouse C2 C12 cells: a model to investigate ageing? J Cell Biochem. 2011;112:3773–3785. pmid:21826704
View Article
PubMed/NCBI
Google Scholar

[82] View Article

[83] PubMed/NCBI

[84] Google Scholar

[ref22] 22. Rando TA, Blau HM. Primary mouse myoblast purification, characterization, and transplantation for cell-mediated gene therapy. J Cell Biol. 1994;125:1275–1287. pmid:8207057
View Article
PubMed/NCBI
Google Scholar

[86] View Article

[87] PubMed/NCBI

[88] Google Scholar

[ref23] 23. Johmura Y, Yamanaka T, Omori S, Wang TW, Sugiura Y, Matsumoto M, et al. Senolysis by glutaminolysis inhibition ameliorates various age-associated disorders. Science. 2021;371:265–270. pmid:33446552
View Article
PubMed/NCBI
Google Scholar

[90] View Article

[91] PubMed/NCBI

[92] Google Scholar

[ref24] 24. Chang YC, Liu HW, Chan YC, Hu SH, Liu MY, Chang SJ. The green tea polyphenol epigallocatechin-3-gallate attenuates age-associated muscle loss via regulation of miR-486-5p and myostatin. Arch Biochem Biophys. 2020;692:108511. pmid:32710883
View Article
PubMed/NCBI
Google Scholar

[94] View Article

[95] PubMed/NCBI

[96] Google Scholar

[ref25] 25. Shahini A, Choudhury D, Asmani M, Zhao R, Lei P, Andreadis ST. NANOG restores the impaired myogenic differentiation potential of skeletal myoblasts after multiple population doublings. Stem Cell Res. 2018;26:55–66. pmid:29245050
View Article
PubMed/NCBI
Google Scholar

[98] View Article

[99] PubMed/NCBI

[100] Google Scholar

[ref26] 26. Hirai H, Verma M, Watanabe S, Tastad C, Asakura Y, Asakura A. MyoD regulates apoptosis of myoblasts through microRNA-mediated down-regulation of Pax3. J Cell Biol. 2010;191:347–365. pmid:20956382
View Article
PubMed/NCBI
Google Scholar

[102] View Article

[103] PubMed/NCBI

[104] Google Scholar

[ref27] 27. Sun Y, Ge Y, Drnevich J, Zhao Y, Band M, Chen J. Mammalian target of rapamycin regulates miRNA-1 and follistatin in skeletal myogenesis. J Cell Biol. 2010;189:1157–1169. pmid:20566686
View Article
PubMed/NCBI
Google Scholar

[106] View Article

[107] PubMed/NCBI

[108] Google Scholar

[ref28] 28. Testa S, D’Addabbo P, Fornetti E, Belli R, Fuoco C, Bernardini S, et al. Myoblast myogenic differentiation but not fusion process is inhibited via MyoD tetraplex interaction. Oxid Med Cell Longev. 2018;2018:7640272. pmid:29854094
View Article
PubMed/NCBI
Google Scholar

[110] View Article

[111] PubMed/NCBI

[112] Google Scholar

[ref29] 29. Rosenberg IH. Sarcopenia: origins and clinical relevance. J Nutr. 1997;127:990S–991S. pmid:9164280
View Article
PubMed/NCBI
Google Scholar

[114] View Article

[115] PubMed/NCBI

[116] Google Scholar

[ref30] 30. Santilli V, Bernetti A, Mangone M, Paoloni M. Clinical definition of sarcopenia. Clin Cases Miner Bone Metab. 2014;11:177–180. pmid:25568649
View Article
PubMed/NCBI
Google Scholar

[118] View Article

[119] PubMed/NCBI

[120] Google Scholar

[ref31] 31. Janssen I, Heymsfield SB, Ross R. Low relative skeletal muscle mass (sarcopenia) in older persons is associated with functional impairment and physical disability. J Am Geriatr Soc. 2002;50:889–896. pmid:12028177
View Article
PubMed/NCBI
Google Scholar

[122] View Article

[123] PubMed/NCBI

[124] Google Scholar

[ref32] 32. Cai Z, Liu D, Yang Y, Xie W, He M, Yu D, et al. The role and therapeutic potential of stem cells in skeletal muscle in sarcopenia. Stem Cell Res Ther. 2022;13:28. pmid:35073997
View Article
PubMed/NCBI
Google Scholar

[126] View Article

[127] PubMed/NCBI

[128] Google Scholar

[ref33] 33. Collins-Hooper H, Woolley TE, Dyson L, Patel A, Potter P, Baker RE, et al. Age-related changes in speed and mechanism of adult skeletal muscle stem cell migration. Stem Cells. 2012;30(6):1182–1195. pmid:22593017
View Article
PubMed/NCBI
Google Scholar

[130] View Article

[131] PubMed/NCBI

[132] Google Scholar

[ref34] 34. Shefer G, Van DP, Richardson JB, Yablonka-Reuveni Z. Satellite-cell pool size does matter: defining the myogenic potency of aging skeletal muscle. Dev Biol. 2006;294:50–66. pmid:16554047
View Article
PubMed/NCBI
Google Scholar

[134] View Article

[135] PubMed/NCBI

[136] Google Scholar

[ref35] 35. Fry CS, Lee JD, Mula J, Kirby TJ, Jackson JR, Liu F, et al. Inducible depletion of satellite cells in adult, sedentary mice impairs muscle regenerative capacity without affecting sarcopenia. Nat Med. 2015;21:76–80. pmid:25501907
View Article
PubMed/NCBI
Google Scholar

[138] View Article

[139] PubMed/NCBI

[140] Google Scholar

[ref36] 36. Fry CS, Lee JD, Jackson JR, Kirby TJ, Stasko SA, Liu H, et al. Regulation of the muscle fiber microenvironment by activated satellite cells during hypertrophy. FASEB J. 2014;28:1654–1665. pmid:24376025
View Article
PubMed/NCBI
Google Scholar

[142] View Article

[143] PubMed/NCBI

[144] Google Scholar

[ref37] 37. Ding J, Nie M, Liu J, Hu X, Ma L, Deng ZL, et al. Trbp is required for differentiation of myoblasts and normal regeneration of skeletal muscle. PLoS One. 2016;11:e0155349. pmid:27159388
View Article
PubMed/NCBI
Google Scholar

[146] View Article

[147] PubMed/NCBI

[148] Google Scholar

[ref38] 38. Izarra A, Moscoso I, Levent E, Cañón S, Cerrada I, Díez -Juan A, et al. miR-133a enhances the protective capacity of cardiac progenitors cells after myocardial infarction. Stem Cell Reports. 2014;3:1029–1042. pmid:25465869
View Article
PubMed/NCBI
Google Scholar

[150] View Article

[151] PubMed/NCBI

[152] Google Scholar

Figures

Abstract

Introduction

Materials and methods

Cell culture and passage

Transfection

Senolysis

Immunostaining

Quantitative reverse transcription (RT)-PCR

Western blot analysis

Plate assay

Isolation of primary myoblasts

Statistics

Results

Replicative senescence suppresses the myogenic differentiation of C2C12 cells

Replicative senescence suppresses miR-1 and miR-133a expression

Overexpression of miRNAs ameliorates the differentiation capacity of senescent C2C12 cells

Senolysis by glutaminase inhibitor increases miR-133a expression

Expression of miR-1 and miR-133a in myoblasts decreases in aged mice

Discussion

Supporting information

S1 Fig.

S2 Fig.

S1 Raw images. Raw images of western blots for Figs 1, 3, 5 and 6.

References