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INTRODUCTION
Skeletal muscle is composed of multinuclear cells called myofibers, which are formed by the fusion of myoblast during development [1,2]. When the muscle is injured, it responds by activating a complex response leading to repair and regeneration of the injured tissue. Skeletal muscle regeneration is primarily mediated by satellite cells, which give rise to myoblasts, the precursor cells of skeletal muscle fibers. Upon activation, these cells proliferate, differentiate into myoblasts, and ultimately fuse to form new myofibers [3]. Myogenic regulatory factors (MRFs), including myoblast determination protein 1 (MyoD), myogenic factor 5 (Myf5), myogenin, and myogenic regulatory factor 4 (MRF4), are essential transcription factors that govern the progression of satellite cells through quiescence, activation, commitment, and differentiation [4,5]. The temporal and spatial expression of these MRFs determines lineage specification and the activation of muscle-specific gene programs. In addition to transcriptional control, post-transcriptional regulation by alternative splicing, RNA-binding proteins, and non-coding RNAs fine tune gene expression during myogenesis [3,6]. The importance of epigenetic modifications such as DNA methylation and histone modifications also has been highlighted in modulating muscle-specific genes [7].
In recent years, microRNAs (miRNAs) and circular RNAs (circRNAs) have emerged as important regulators of gene expression and cellular processes including muscle development and differentiation [8,9]. miRNAs are small, non-coding RNAs that post-transcriptionally regulate gene expression by binding to complementary sequences in the 3' untranslated region (UTR) of target mRNAs leading to translational repression or mRNA degradation [10]. In the context of skeletal muscle, miR-133 has been reported to regulate the expression of genes involved in muscle development and function [11]. miR-143-3p also has been shown to modulate the expression of the slow muscle fiber gene in porcine skeletal muscle satellite cell [12] as well as playing a role in various biological processes, including cell proliferation, differentiation, and metabolism [13].
CircRNAs are a class of non-coding RNAs characterized by their covalently closed circular structure, which confers resistance to exonuclease degradation and increased stability compared to linear RNAs [14,15]. These molecules have been shown to act as miRNA sponges, sequestering miRNAs and thereby modulating their regulatory effects on target mRNAs [16,17]. One such circRNA, circMYBPC1, has recently been implicated in the regulation of mouse skeletal muscle differentiation [18].
The nicotinic acetylcholine receptor subunit alpha 1 (Chrna1) is one of the four types of subunits comprising muscle acetylcholine receptor (AChR), which is a component of the neuromuscular junction mediating signal transmission between motor neurons and skeletal muscle fibers [19]. Proper expression and function of AChR are essential for normal muscle development and contractile activity [20]. Mutations in the Chrna1 gene have been associated with congenital myasthenic syndromes, characterized by muscle weakness and fatigue due to impaired neuromuscular transmission [21]. In a mouse model of sarcopenia, the age-related loss of muscle mass and strength, Chrna1 induces sarcopenia through neuromuscular synaptic elimination [20]. Furthermore, AChRs have been found to be involved in the regulation of skeletal muscle fiber type composition [22]. Interestingly, AChR expression has also been linked to muscle regeneration and repair processes. AChRs have been found to be upregulated during muscle regeneration following injury, suggesting their involvement in the activation and differentiation of muscle satellite cells [23].
Recent studies have revealed complex regulatory networks involving circRNAs and miRNAs in muscle development [21,24]. The interaction between these regulatory molecules and key structural components like Chrna1 represents an emerging area of research with potential therapeutic implications [25,26]. Understanding these molecular mechanisms is particularly relevant given the role of AChR in various muscle disorders and age-related muscle conditions [27,28].
In this study, we aim to investigate the role of circAtxn10, miR-143-3p, and Chrna1 in skeletal muscle differentiation at the cellular and molecular levels. By elucidating the interactions between these molecules and their impact on muscle development, we hope to contribute to a better understanding of the complex regulatory networks governing skeletal muscle differentiation and identify potential targets for therapeutic interventions in muscle-related disorders.
METHODS
RNA-seq and circRNA selection
RNA-seq and circRNA selection were done as described previously [24]. Briefly, total RNA was prepared from human skeletal muscle satellite cells, human skeletal muscle myoblast and human skeletal muscle cells (#3500, #3510, #3520, Sciencell Research Laboratories Inc.) using TRIzol reagent (#15596026, ThermoFisher Scientific). Ribosomal RNAs were removed from total RNA with Ribo-Zero Gold rRNA Removal Kit (MRZG126, Illumina), and TruSeq Stranded Total RNA Kit (20020596, Illumina) was used to construct an RNA-seq library. The library was sequenced using NovaSeq6000 (20012850, Illumina) in the paired-end mode with 100 sequencing cycles, and FastQC was used for the initial quality check of the sequencing results. The reads with low quality were removed with Trimmomatic (http://www.usadellab.org/cms/?page=trimmomatic). The expression counts were calculated using the STAR aligner with DCC algorithm followed by normalization by the total counts of circRNAs in each sample. We selected circRNAs with a mean read count of at least 1 across all samples. The RNA-seq dataset including raw sequencing reads and processed data has been deposited in the GEO database under accession number GSE281859.
circRNA and miRNA target prediction
Prediction of miRNAs that bind to circRNA was performed on miRDB (http://mirdb.org/index.html), and putative targets of miR were predicted on TargetScan (https://www.targetscan.org/vert_80/). miRs and mRNAs related to myoblast and muscle cell were selected, and the expression of which was monitored after myogenic stimulation for final selection.
miR mimic, miR inhibitor, small interfering RNA (siRNA), and antibodies
SiRNA nucleotides for knockdown including custom designed circAtxn10 siRNA, predesigned Chrna1 siRNA, miR-143-3p mimic, miR-143-3p inhibitor, control miR, and control siRNA were purchased from Bioneer Corp. The sequences for circAtxn10 siRNA oligo were as shown Supplementary Table 1. Antibodies against Chrna1 (sc-65829, Santa Cruz Biotechnology), MyoD (sc-32758, Santa Cruz Biotechnology), Myogenin (ab1835, Abcam), muscle creatine kinase (MCK) (ab155901, Abcam), skeletal muscle alpha-actin (Acta1) (sc-58671, Santa Cruz Biotechnology), and glyceraldehyde 3-phosphate dehydrogenase (GAPDH) (G9545, Sigma) were used at 1:1,000 dilution.
Cell cultures
Mouse skeletal myoblasts C2C12 cells were cultured in Dulbecco’s Modified Eagle’s Medium (DMEM) (LM001-05, Welgene) with 15% fetal bovine serum (FBS) (S001-07, Welgene) and antibiotics (15240062, ThermoFisher Scientific). Cells were maintained in a humidified incubator with 5% CO2 at 37°C.
Induction of myogenesis in vitro
To induce myogenesis differentiation, C2C12 cells were induced by supplementing cell DMEM with 2% heat-inactivated horse serum (S200, Vector Laboratories). The medium was changed every 2 days for up to 6 days. During differentiation, the expression of myogensis markers such as MyoD and Myogenin increased notably by day 2. For experiments assessing induction of myogenesis, C2C12 cells differentiated for 2–4 days were used, while cells differentiated for 6 days were used to evaluate anti-myogenesis effects.
Transfection
3 × 10⁵ C2C12 cells were seeded in 60 mm dishes, and 2.5 μg of DNA was transfected using Lipofectamine (18324-012, Invitrogen) and Plus reagent (11514-015, Invitrogen). For RNA transfection, 2 × 10⁴ C2C12 cells were seeded in 60 mm dishes, and 10 nM of each RNA was transfected using Lipofectamine RNAiMAX (13778-100, Invitrogen).
Quantitative real-time polymerase chain reaction (qRT-PCR)
Total RNA was extracted and contaminating DNA was removed using NucleoSpin RNA/protein (740933.250, Macherey-Nagel) following manufacturer's protocol. cDNAs from mRNAs and miRNAs were reverse transcribed using either a SuperScript first-strand synthesis system for RT-PCR (11904018 Invitrogen) or a TaqMan advanced miRNA cDNA synthesis kit (A28007, Applied Biosystems), respectively. The cDNA was analyzed by qRT-PCR using QuantiTect SYBR Green PCR Kit (204141, Qiagen) and Rotor gene Q realtime PCR cycler (9001550, Qiagen). The RNA expression levels were normalized to GAPDH. The primer information used in PCR is provided in the Supplementary Tables 2 and 3.
RNase R digestion
Total RNA samples were incubated with RNase R (RNR07250, Lucigen) at 37°C for 20 min. RNase R was then inactivated by heating the sample at 95°C for 3 min following the manufacturer's protocol.
Cloning
DNA fragment corresponding to circAtxn10 was synthesized (CosmoGeneTech) and cloned into Zkscan1 MCS exon vector (Addgene, #69901) for overexpression in C2C12 cells. To check whether circAtxn10 acts as sponge for miR-143-3p, DNA fragment corresponding to circAtxn10 was cloned into pGL3UC vector, a modified pGL3 Luciferase Report Vector (E1751, Promega) with CMV promoter sequences and multiple cloning sites, for luciferase assay. Also, to check whether miR-143-3p binds to Chrna1 mRNA, DNA fragment corresponding to the 3′-UTR of Chrna1 mRNA containing the putative binding sites for miR-143-3p was PCR amplified from mouse genomic DNA and cloned into pGL3UC. The coding sequence of mouse Chrna1 was PCR amplified from mouse cDNA and cloned into pcDNA6/myc-His vector (V22120, ThermoFisher Scientific) for overexpression of Chrna1 in C2C12 cells. The primer sequences used for cloning in overexpression and luciferase reporter assays was listed in Supplementary Tables 4 and 5, respectively.
Western blot analysis
Cellular proteins were prepared with RIPA buffer (EBR001-500, Enzynomics) supplemented with 1 mM dithiothreitol (DTT), 1 mM phenylmethylsulfonyl fluoride, 1 mM Na3PO4, and protease inhibitor (11697498001, Hoffmann-La Roche). The proteins were separated by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) and then transferred to a polyvinylidene difluoride membrane (Millipore). The membrane was blocked with 5% skim milk (232100, BD Difco) in TRIS-buffered saline-Tween 20 (20605, ThermoScientific Fisher) (TBST) and then incubated with primary antibodies overnight at 4°C on a rocker. After three washes in TBST, the membranes were incubated with horseradish peroxidase-conjugated secondary antibodies (7076, 7074, Cell Signaling) for 1 h at room temperature. The peroxidase activity was visualized using Western Blotting Luminol Reagent (sc-2048 Santa Cruz Biotechnology) in a FUJIFILM Luminescent Image Analyzer LAS-3000 (Fujifilm Life Science). Quantification of the chemiluminescence signal was done after retrieving the density of the bands using Scion Image software (Scion Corporation).
Immunocytochemistry
C2C12 cells grown on microscope slide cover slips were fixed with 4% paraformaldehyde, permeabilized with 0.1% Triton X-100, and blocked with 5% bovine serum albumin (BSA) solution. Cells were then incubated with primary antibodies overnight at 4°C followed by Alexa Fluor-conjugated secondary antibodies (Cat # A-11001, 11005, Invitrogen) for 1 h at room temperature. Nuclei were counterstained with DAPI. Images were acquired using a Leica DM3000 microscope with a Nikon DS-Ri2 camera and Nikon NIS-Elements AR software (Nikon Instruments Korea Co., Ltd.).
Luciferase assay
The luciferase vector with either circAtxn10 or 3′-UTR of Chrna1 mRNA sequence was co-transfected with 10nM miR-143-3p mimic into C2C12 cells using LipofectaminLTX (Invitrogen). Luciferase activity was measured using the Luciferase Assay System (E1500, Promega) following the manufacturer's protocols. The pSV-β-Galactosidase Control Vector (E1081, Promega) was also co-transfected to normalize the luciferase signal with β-Galactosidase Enzyme Assay System (E2000, Promega).
Statistical analysis
Analyses were conducted using IBM SPSS Statistics 26 (IBM). Data normality was verified through the Shapiro–Wilk test, and either a two-tailed Student's t-test or analysis of variance (ANOVA) with subsequent post-hoc analysis was applied for the data confirmed as normally distributed dependent on the number of groups being either two or more. In case of evaluating more than two groups, we examined variance homogeneity between groups with Levene's test after ANOVA. Depending on homogeneity, either Dunnett's T3 or the post-hoc Tukey's honestly significant difference test/Bonferroni's test was used. Non-parametric methods: the Kruskal–Wallis test with a Bonferroni adjustment or the Mann–Whitney U-test, predicated on whether there were more than two groups in consideration, were used for non-normally distributed data. Significance was confirmed for a p-value of less than 0.05. Correlation analyses were performed using Pearson's correlation coefficient. All experiments were performed at least three times independently, and data are presented as mean ± standard error of the mean (SEM).
RESULTS
Characterization of circAtxn10 in C2C12 cells
To investigate the role of circRNAs in skeletal muscle differentiation, we performed RNA-seq analysis on differentiating human myocytes at various stages, including satellite cells, myoblasts, and adult muscle tubes. Total RNA was isolated using Ribozero to remove ribosomal RNA, followed by sequencing and data analysis. Reads were aligned, and circRNA expression was quantified as illustrated in Fig. 1A.
Initial screening identified eight circRNAs with significantly increased expression during myogenic differentiation using a stringent threshold (Max deviation > 3) (Fig. 1B). To validate these candidate circRNAs, we designed divergent primers specific to the back-splicing junctions and successfully confirmed their expression profiles through PCR, including additional circRNAs identified using a relaxed threshold (Max deviation > 2) (Fig. 1C). Among the validated circRNAs, four—circAtxn10, circLdlrad3, circSfmbt2, and circPrdm5—were selected for further analysis by RT-PCR in differentiated C2C12 cells. Notably, circAtxn10 and circLdlrad3 exhibited particularly robust expression in differentiated myotubes (Fig. 1D).
To confirm the circular nature of these transcripts, we performed RNase R treatment, which selectively degrades linear RNAs while preserving circRNAs. As expected, linear control transcripts (e.g., GAPDH) were effectively degraded, while two candidate circRNAs, circAtxn10 and circLdlrad3, demonstrated strong resistance to RNase R digestion. These circRNAs showed remarkable stability, with minimal degradation following RNase R treatment (Fig. 1E). Additionally, circAtxn10 shares 82% homology between human and mouse species (Supplementary Fig. 1). This stability, coupled with their differential expression patterns during myogenesis, highlighted circAtxn10 as a promising candidate for further functional studies.
The role of circAtxn10 in skeletal muscle differentiation
To validate the existence of circAtxn10, we performed Sanger sequencing with divergent primers and confirmed the presence of intact back-splicing junction sequences, highlighted by red arrows (Fig. 2A). Using Zkscan1 mammalian expression vector containing the full circAtxn10 sequence, we achieved approximately 2.5-fold overexpression in C2C12 cells (Fig. 2B). This overexpression significantly enhanced myotube formation (Fig. 2C) and dramatically increased the expression of key myogenic markers such as MyoD, Acta1, MCK, and Myogenin compared to mock-transfected controls (Fig. 2D).
Next, to evaluate the functional role of circAtxn10, we designed two siRNAs targeting its splicing junction for knockdown of circAtxn10 (Supplementary Fig. 2A). Both siRNAs effectively suppressed circAtxn10 expression without altering linear Atxn10 mRNA levels (Fig. 2E, Supplementary Fig. 2B). Knockdown of circAtxn10 significant impaired myogenic differentiation, evidenced by a decrease in myotube formation (Fig. 2F, Supplementary Fig. 2C). Additionally, the expression levels of myogenic marker genes, including MyoD, Acta1, MCK, and Myogenin, were reduced upon circAtxn10 deficiency (Fig. 2G). Immunofluorescence staining for Myogenin further demonstrated that circAtxn10 knockdown markedly reduced both the number and size of multinucleated myotubes (Fig. 2H). These finding collectively indicatet that circAtxn10 positively regulates muscle differentiation by enhancing myogenic marker expression and promoting myotube formation.
CircAtxn10 acts as a miR-143-3p sponge
To determine the subcellular localization of circAtxn10, we isolated nuclear and cytosolic fractions from C2C12 cells. circAtxn10 expression was exclusively observed in the cytosolic fraction (Fig. 3A). Previous studies have reported that cytosolic circRNAs often act as miRNAs sponges, binding to specific miRNAs to modulate their regulatory activity [10]. We next sought to identify potential mRNA targets regulated by circAtxn10. Using bioinformatic analysis using miRDB identified miR-143-3p as a potential binding partner of circAtxn10, with a predicted interaction site at nucleotides 726-731 containing a complementary sequence (Fig. 3B). To validate this interaction, we performed luciferase reporter assays using constructs containing the circAtxn10 sequence. Co-transfection of the circAtxn10 luciferase reporter with miR-143-3p mimic resulted in a significant reduction in luciferase activity, demonstrating direct binding between circAtxn10 and miR-143-3p (Fig. 3C).
To confirm the functional relevance of this interaction in myogenic differentiation, we examined the regulatory relationship between circAtxn10 and myogenic markers. While circAtxn10 overexpression enhanced myogenic marker expression, this effect was significantly attenuated by co-transfection with miR-143-3p mimic (Fig. 3D). This rescue experiment provides strong evidence that circAtxn10's pro-myogenic effects are mediated through its ability to sequester miR-143-3p.
The role of miR-143-3p in skeletal muscle differentiation
To investigate the role of miR-143-3p in muscle differentiation, we examined its effects on C2C12 myoblasts. Overexpression of miR-143-3p via treatment with a miR-143-3p mimic led to a significant reduction in myotube formation, as evidenced by diminished muscle fiber development (Fig. 4A). In control cells undergoing differentiation, myogenin expression typically increased approximately 20-fold by day 4. However, in cells transfected with the miR-143-3p mimic, this induction was significantly inhibited, resulting in a less than 5-fold increase in myogenin levels at the same time point (Fig. 4B). This suppression was accompanied by marked decreases in the expression of myogenic markers such as Acta1 and MCK, as confirmed by Western blot analysis (Fig. 4C).
Immunofluorescence staining further validated these findings, showing reduced expression of Myogenin and Acta in miR-143-3p mimic-treated cells (Fig. 4D). Additionally, miR-143-3p overexpression impaired myoblast fusion, reducing the formation of multinucleated myotubes (Fig. 4E). Conversely, inhibition of miR-143-3p through a specific inhibitor enhanced myotube formation (Fig. 4F). These results suggest that miR-143-3p negatively regulates muscle differentiation by downregulating critical myogenic markers and inhibiting myoblast fusion.
Chrna1 is a target of miR-143-3p during skeletal muscle differentiation
By binding to complementary sequences in the 3’ UTR of target mRNA, miRNAs can suppress gene expression either by promoting mRNA degradation or inhibiting translation [13]. In our study, bioinformatic analysis revealed Chrna1 as target of miR-143-3p, with three potential miR-143-3p binding sites in its 3'-UTR (Fig. 5A). Luciferase reporter assays confirmed that miR-143-3p directly binds to the Chrna1 3'-UTR, leading to a significant reduction in luciferase activity (Fig. 5B). During C2C12 differentiation, Chrna1 expression increases progressively, reaching approximately 6-to 8- fold elevation by day 2 and day 4. However, transfection of miR-143-3p mimic substantially impaired this induction throughout the differentiation time course (Fig. 5C). This temporal pattern of regulation suggests that the circAtxn10/miR-143-3p/Chrna1 axis is particularly important during the early-to-mid stages of myogenic differentiation.
Role of Chrna1 in skeletal muscle differentiation
Chrna1 is traditionally known for its role in neuromuscular junction formation and function [29]. However, emerging evidence suggests that Chrna1 may also contribute directly to myogenesis, independent of its role in neuromuscular signaling [30]. To investigate the functional significance of Chrna1 in skeletal muscle differentiation, we conducted both gain- and loss-of-function experiments.
In gain-of-function experiments, we transfected C2C12 cells with pcDNA6-Chrna1-myc-His, resulting in a substantial increase in Chrna1 expression compared to mock-transfected controls (Fig. 6A). Elevated Chrna1 levels significantly enhanced the expression of myogenic markers, including Acta1, MCK, and Myogenin, suggesting that Chrna1 acts as a positive regulator of myogenic differentiation (Fig. 6B).
To assess the necessity of Chrna1 for myogenesis, we performed siRNA-mediated knockdown of Chrna1. Transfection with Chrna1 siRNA effectively reduced Chrna1 expression compared to the scramble control (Fig. 6C). Morphological analysis following 6 days of differentiation revealed distinct differences in myotube formation. Control cells successfully fused to form mature myotubes with strong Acta1 staining, while Chrna1 knockdown significantly impaired myotube formation, with reduced Acta1 staining intensity (Fig. 6D, E). This decrease in Chrna1 expression impaired myogenic differentiation, as demonstrated by reduced levels of key myogenic markers, including Acta1, MCK, and Myogenin (Fig. 6F). These findings were further supported by Western blot analysis, which confirmed reductions in the protein expression of Acta1, MCK, and Myogenin upon Chrna1 knockdown (Fig. 6G).
These results underscore the crucial role of Chrna1 in promoting skeletal muscle differentiation and suggest that it may be a target of miR-143-3p, adding a new dimension to its functional significance beyond the neuromuscular junction.
DISCUSSION
In this study, we identified a regulatory cascade involving circAtxn10, miR-143-3p, and Chrna1 that modulates skeletal muscle cell differentiation. The expressions of circAtxn10 and Chrna1 increase during myogenesis, and their overexpression promoted myogenic differentiation. Our findings demonstrate that CircAtxn10, derived from the back-splicing of exons 4-9 of the Atxn10 pre-mRNA, is upregulated in the cytosol of skeletal muscle cells. It functions as a miRNA sponge by sequestering miR-143-3p, which otherwise targets Chrna1. The circAtxn10/miR-143-3p/Chrna1 axis plays a crucial role in promoting myocyte differentiation (Fig. 7). Both circAtxn10 knockdown and miR-143-3p mimic downregulated myogenic differentiation. These results suggest that the upregulation of circAtxn10 during C2C12 differentiation promotes myogenesis by binding miR-143-3p, thereby preventing the downregulation of Chrna1.
The temporal dynamics of this regulatory circuit provide insights into the molecular mechanisms that link gene expression changes to morphological outcomes in myogenesis. Early circAtxn10 upregulation (2-fold by day 2) corresponds with the activation of myogenic factors that drive cytoskeletal remodeling. The subsequent upregulation of Chrna1 enhances acetylcholine receptor (AChR) clustering, which is followed by myogenic cell fusion [31]. Also nicotinic acetylcholine receptor antagonists inhibited spontaneous fusion of myoblasts [32].
The progression of this regulatory axis is particularly crucial during the day 2–4 differentiation window, a period marked by significant morphological changes. Chrna1-dependent calcium signaling activates calcium-dependent proteases essential for membrane fusion [22] and initiates the calcineurin-NFAT pathway, thereby enhancing the expression of fusion-related genes [23]. This creates a positive feedback loop wherein AChR signaling increases the expression of fusion-related proteins and additional myogenic factors, thereby reinforcing the differentiation program [24].
These findings have important implications for muscle disorders and regenerative medicine. Conditions like Duchenne muscular dystrophy and sarcopenia involve progressive muscle weakness and degeneration, stemming from mutations in genes critical for muscle development and maintenance [31,32], or from dysregulated myogenic signaling pathways [21]. The nicotinic acetylcholine receptor (AChR) and its alpha-1 subunit (Chrna1) are essential for neuromuscular junction function, muscle development, and regeneration following injury [16,18,33]. Our discovery of the circAtxn10/miR-143-3p/Chrna1 regulatory axis not only enhances our understanding of myogenic regulation but also highlights potential therapeutic targets. This pathway may be particularly relevant for treating conditions such as congenital myasthenic syndromes, which often involve CHRNA1 mutations [14]. Therapeutic modulation of this axis, whether through circAtxn10, miR-143-3p, or Chrna1, could offer new strategies for enhancing muscle regeneration in both degenerative conditions and age-related muscle wasting. The fact that Chrna1 downregulation, either by siRNA or miR-143-3p, impairs myotube formation and reduces myogenic marker expression further underscores the therapeutic potential of this pathway.
In conclusion, our study uncovers a novel regulatory axis involving circAtxn10, miR-143-3p, and Chrna1, which plays a crucial role in skeletal muscle differentiation. These findings contribute to the growing body of knowledge on the intricate molecular mechanisms governing myogenesis and identify potential targets for therapeutic interventions in muscle-related disorders. Future studies should aim to elucidate the downstream signaling pathways and transcriptional networks influenced by this regulatory axis and to explore its relevance in in vivo models of muscle development and disease.
SUPPLEMENTARY MATERIALS
Supplementary data including five tables and two figures can be found with this article online at https://doi.org/10.4196/kjpp.25.046
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