Skip to main content
Download PDF

FTO-dependent m6A regulates muscle fiber remodeling in an NFATC1–YTHDF2 dependent manner

Clinical Epigenetics volume 15, Article number: 109 (2023) Cite this article

Abstract

Background

Adolescent idiopathic scoliosis (AIS) is characterized by low lean mass without vertebral deformity. The cause-and-effect relationship between scoliosis and paraspinal muscle imbalance has long puzzled researchers. Although FTO has been identified as a susceptibility gene for AIS, its potential role in the asymmetry of paraspinal muscles has not been fully elucidated.

Methods

We investigated the role of Fto in murine myoblast proliferation, migration, and myogenic differentiation. We examined its precise regulatory influence on murine muscle fiber remodeling in vitro and in vivo. We identified the downstream target gene of Fto by screening key regulators of murine muscle fiber remodeling and identified its m6A reader. Deep paraspinal muscle samples were obtained from the concave and convex sides of AIS patients with or without Schroth exercises, and congenital scoliosis served as a control group. We compared the content of type I fibers, expression patterns of fast- and slow-type genes, and levels of FTO expression.

Results

FTO contributed to maintain the formation of murine slow-twitch fibers both in vitro and in vivo. These effects were mediated by the demethylation activity of FTO, which specifically demethylated NFATC1 and prevented YTHDF2 from degrading it. We found a significant reduction in type I fibers, mRNA levels of MYH7 and MYH7B, and expression of FTO on the concave side of AIS. The percentage of type I fibers showed a positive correlation with the expression level of FTO. The asymmetric patterns observed in AIS were consistent with those seen in congenital scoliosis, and the asymmetry of FTO expression and fiber type in AIS was largely restored by Schroth exercises.

Conclusions

FTO supports the formation of murine slow-twitch fibers in an NFATC1–YTHDF2 dependent manner. The consistent paraspinal muscle features seen in AIS and congenital scoliosis, as well as the reversible pattern of muscle fibers and expression of FTO in AIS suggest that FTO may contribute to the muscle fiber remodeling secondary to scoliosis.

Introduction

Adolescent idiopathic scoliosis (AIS) is one of the most common spinal deformities, affecting millions of children with an incidence of about 3–4% worldwide [1]. A curve magnitude greater than 45 degrees is the surgical threshold for treating AIS [2]. Surgical treatment may induce trauma and psychological effects, such as neurovascular injury, internal fixation failure, and restriction of back movement [3]. Unlike congenital scoliosis (CS), which is characterized by hemivertebral deformity, AIS is distinguished by low lean mass without vertebral deformities. Additionally, low lean mass of the back muscles is associated with a higher risk of curve progression in AIS [4]. Although several genes have been associated with AIS, their specific roles in the development of the condition remain unclear. Recent research has given increasing attention to the role of epigenetics in the development of AIS [5,6,7]. While FTO has been identified as a susceptibility locus for AIS based on a genome-wide association analysis involving 79,211 people, its specific role in the etiology of AIS remains unclear [8].

AIS is believed to be caused by an imbalance of the paraspinal muscles, with the concave paraspinal muscles having a lower percentage of type I fibers than the convex [9]. The slow-twitch and fast-twitch fiber types of skeletal muscle are different from one another in terms of contractile protein composition, oxidative capacity, and preferred substrate for ATP synthesis [10]. Muscle fiber-type switching depends on the exact coordination of metabolic and contractile gene expression programs to control fiber-type specification and ensure contractile function [11]. FTO is well-known for its connection to obesity and its crucial role in controlling transcriptome-wide m6A modification [12]. Previous studies have demonstrated that FTO is necessary for myogenesis [13, 14]. However, the relevance of FTO-dependent m6A demethylation in fiber type-specific patterns and its role in skeletal muscle fiber remodeling have not been discussed.

To investigate the role of the FTO susceptibility gene in AIS and its potential function in skeletal muscle fiber remodeling via the FTO-dependent m6A epitranscriptome, we used paraspinal muscle samples from AIS patients, a mouse model of muscle injury, and murine myoblast culture methods. We found that FTO promotes the development of murine slow-twitch fibers both in vitro and in vivo, primarily through its demethylation of NFATC1 and prevention of YTHDF2-mediated degradation. Additionally, we demonstrated that the asymmetry in muscle fibers and FTO expression was not limited to AIS and could be reversed.

Materials and methods

Human samples

The current study adhered to the ethical guidelines outlined in the 1964 Declaration of Helsinki and its subsequent amendments after receiving approval from the Institutional Review Board. Participants in the study were scoliosis patients who met the following inclusion and exclusion criteria: (1) confirmed diagnosis of either AIS or CS and scheduled for one-stage posterior spinal surgery; (2) primary right thoracic curve; (3) female; (4) aged between 12 and 18 years; (5) Han ethnicity; (6) informed consent from guardians. Exclusion criteria included: (1) spinal deformities other than AIS or CS, such as neuromuscular scoliosis; (2) history of spinal surgery; (3) mental or psychological conditions preventing compliance with treatment.

The study enrolled twenty AIS and five CS patients who underwent posterior spinal surgery, as well as five additional AIS patients who had undergone Schroth exercises for more than three months. Deep paraspinal muscle biopsies were taken from the concave and convex sides of the curve apex during spinal surgery, and these samples were subsequently mounted for fiber typing or flash-frozen in liquid nitrogen for RNA and protein extraction. To minimize any confounding factors from individual variability that may impact our analysis results, we utilized a strict paired research design in our case selection. Although the sample size for the three groups (AIS, CS, and AIS with Schroth exercises) used for subsequent immunofluorescence and Western blot analyses was small, consisting of only five individuals each, all participants were female with a right thoracic curve and exhibited no significant differences in age or primary Cobb angle. Patient characteristics are provided in Additional file 1: Table S1.

Animal samples

We used 6-week-old C57/BL mice to study the role of Fto in muscle fiber remodeling in vivo after receiving ethics committee approval. In brief, 50 µl of cardiotoxin (CTX, 10 μM, Latoxan) was intramuscularly injected into the soleus to induce acute muscle damage. To prevent the impact of Fto on the proliferation of satellite cells, we utilized FB23-2 (a specific inhibitor of FTO m6A demethylase) five days after CTX injection. 50 µl of FB23-2 (5 μM, AbMole) was injected into the soleus every three days. After five rounds of FB23-2 injections, we collected soleus biopsies for further immunofluorescence and quantitative real-time PCR analysis.

Cell culture, transfection and tracking

C2C12 cells were cultured at 37 °C with 5% CO2 in growth media (DMEM containing 10% fetal bovine serum and 1% penicillin/streptomycin). To induce myogenic differentiation, C2C12 cells were grown to approximately 80% confluence and then switched to a differentiation medium (DMEM containing 2% horse serum and 1% penicillin/streptomycin). All cellular experiments were performed with biological replicates in 3–5 independent wells. Three independent siRNAs for Fto or Ythdf2 were transfected into cells using Lipofectamine™ RNAiMAX Reagent (Invitrogen) according to the manufacturer’s instructions. The siRNAs were designed and synthesized by RIBOBIO. The target sequences of siRNAs are provided in Additional file 2: Table S2.

Live cell tracking was performed according to the manufacturer’s guidelines of Harmony High-Content Imaging and Analysis Software (PerkinElmer). Briefly, after 24 h of cell seeding, cells were placed into the high-content imaging system with conditions set at 37℃ and 5% CO2. Under bright field conditions, the software could identify all cells within the visual range. Cell recording was conducted at 15-min intervals with a total tracking time of 6 h. By continuously capturing cellular motion for 6 h and using the software to segment cells, various indicators of cell migration, including the accumulated distance, displacement per track, and average speed per track, were calculated. Objects detected at only one time point were treated as detection errors, and the software automatically corrected them through splitting or merging procedures.

Immunofluorescence analysis

Paraspinal muscle biopsies from scoliosis patients and mice were frozen in precooled isopentane for immunofluorescence staining, and the frozen tissues were sectioned into 8 µm thick sections. After fixation with 4% paraformaldehyde and blocked with 5% BSA (Biofroxx), the sections were incubated with primary antibodies against MHC1 (1:20, DSHB) overnight at 4 °C. The sections were then incubated with secondary antibodies (Alexa Fluor 488 Anti-Mouse IgG or Alexa Fluor 594 Anti-Mouse IgG; 1:200, Yeasen) at room temperature for 2 h in the dark.

C2C12 cells and differentiated myotubes were fixed with 4% paraformaldehyde and then treated with 0.5% Triton® X-100 (Biofroxx). After blocking, cells were stained with primary antibodies against Ki67 (1:200, Proteintech), FTO (1:200, Proteintech), MHC1 (1:20, DSHB), and MyoG (1:100, Santa Cruz). Cells were then incubated with secondary antibodies (Alexa Fluor 488 Anti-Mouse IgG, Alexa Fluor 594 Anti-Rabbit IgG, or Alexa Fluor 594 Anti-Mouse IgG; 1:200, Yeasen) at room temperature for 2 h. Nuclei were stained with DAPI for 5 min.

The fluorescence microscope used for imaging was an ECLIPSE Ti2 (Nikon), and the positive signal threshold was determined by staining with negative control IgG. Five fields were randomly selected for each independent sample, and the sample sizes for each experimental group were specified in the figure legends. Image quantification was performed using ImageJ software (Version 1.52 V). The proportion of type I fibers was calculated as the percentage of type I fibers among the total muscle fibers in the muscle tissue cross-section. The fusion index of type I fibers was calculated by dividing the number of nuclei in the MHC1-positive region by the total number of nuclei in the field of view. The Ki67 and MyoG positivity rates were calculated as the percentage of positively stained cells in the field of view out of the total number of cells in the field of view, respectively. Individual data of immunofluorescence analysis were derived from three repeated measurements.

Quantitative real-time PCR and Western blot assay

After the muscle samples were pulverized using a mortar and pestle in liquid nitrogen or the cells were washed with PBS, RNAiso (Takara) was added, and total RNA was isolated with Qiagen RNAeasy Mini Kits (Qiagen) according to the manufacturer’s guidelines. cDNA was synthesized using the PrimeScript™ RT reagent kit (Takara). In brief, 1 μg total RNA was mixed with 1 μl of gDNA Eraser and RNase free dH2O was added to a final volume of 10 μl for genomic DNA removal. The mixture was incubated at room temperature for 5 min. Additionally, a 10 μl Master Mix was prepared, comprising 1 μl PrimeScript RT Enzyme Mix I, 1 μl RT Primer Mix, 4 μl 5 × PrimeScript Buffer 2, and 4 μl RNase free dH2O. The Master Mix was then mixed with the total RNA. Amplification was carried out in a SimpliAmp™ Thermal Cycler (Thermo Fisher Scientific) under the following amplification conditions: incubation at 37 °C for 15 min followed by incubation at 85 °C for 5 s. PowerUp™ SYBR™ green master mix (Thermo Fisher Scientific) was added to detect the expression of related genes. In brief, a 10 μl component was prepared for the PCR reactions, including 5 μl 2X PowerUp™ SYBR™ Green Master Mix, 0.5 μl forward primer, 0.5 μl reverse primer, and 4 μl DNA template with RNase free dH2O. Quantitative real-time PCR was performed using the standard reaction mode with an ABI QuantStudio 3 machine (Thermo Fisher Scientific), including UDG activation at 50 °C for 2 min, activation at 95 °C for 2 min, denature at 95 °C for 15 s, extend at 60 °C for 1 min. We used delta delta Ct method to quantitation, and the internal reference gene GAPDH was chosen as the normalization gene to standardize the expression levels of the target gene relative to the reference gene in the samples. The gene-specific primers are listed in Additional file 3: Table S3 and Additional file 4: Table S4.

The total protein was extracted using a RIPA lysis solution containing 1% PMSF. Protein quantification was achieved using the BCA Protein Assay Kit (Biosharp). The extracted protein was mixed with SDS PAGE protein sample buffer in a 1:1 volume ratio and heated in a water bath at 100 °C for 15 min to fully denature the protein. The sample volume for the loading of each lane was calculated based on the protein quantification result, with 20 μg of protein loaded per lane with a sample volume of 15 μl. Gel electrophoresis and membrane transfer were conducted in accordance with standard processes. After blocking with 5% BSA to prevent nonspecific antigen binding, the membranes were incubated overnight with primary antibodies including FTO (1:1000, Proteintech), NFATC1 (1:1000, Proteintech), and GAPDH (1:5000, Proteintech). The membranes were incubated at room temperature for 2 h following the secondary antibody incubation (HRP-conjugated goat anti-rabbit IgG secondary antibody; 1:10,000, Yeasen). After incubation, the membrane was washed 3 times with TBST buffer for 15 min each time. Signal detection was conducted using the Enhanced ECL chemiluminescence detection kit (Vazyme), and image acquisition was completed using the ChemiDocTM Imaging System. The gray value for the target protein was analyzed using ImageJ software (Version 1.52v), with individual data derived from three repeated measurements.

Quantification of m6A modification and RNA stability assays

For quantification of m6A in human and mouse skeletal muscle tissue as well as in C2C12 cells during differentiation, an EpiQuik m6A RNA Methylation Quantification Kit (Epigentek) was used following the manufacturer’s protocol. The percentage of m6A in total RNA was calculated with a colorimetric quantification method by reading the absorbance at 450 nm.

C2C12 cells were transfected with Fto siRNAs when they reached approximately 60% confluence. After being cultured in complete medium for an additional 24 h, the cell density reached around 80%. Then, the growth medium was replaced with a differentiation medium for subsequent experiments. Cells were then treated with actinomycin D (5 μg/mL, AbMole) to inhibit global mRNA transcription, and cells were harvested at 0, 3, and 6 h to assess RNA stability. The expression levels of target genes were detected with quantitative real-time PCR.

Statistical analysis

Data were analyzed and plotted using GraphPad Prism (Version 8.4.3). The data for individual samples was obtained by taking the average of three measurements. For samples in the same case series, we further process the quantified indicators of individual samples as the mean ± s.d. (e.g., In the case of a patient with AIS, we conducted three repeated measurements of their Cobb angle, resulting in an average Cobb angle of 53.4°. We then processed the Cobb angles of 5 AIS patients and summarized the data for this case series as 47.4° ± 3.9°). Non-parametric tests were performed due to the small sample size in each series. For the detection of para-vertebral muscle on the concave and convex sides of the same patient, paired t-tests were utilized, while independent samples were tested using non-paired t-tests. After passing the normality test, Pearson correlation analysis was used to explore the correlation between FTO mRNA expression levels and the proportion of type I fibers. One-way ANOVA was used to compare the data among the three groups. A P value of < 0.05 was considered statistically significant.

Results

Fto regulates the proliferation and migration of murine myoblasts

In order to investigate the potential role of Fto in murine myoblasts, we utilized siRNAs against Fto (si-Fto) in C2C12 cells to lose its function. The knockdown efficiency of si-Fto was verified with quantitative real-time PCR, and Western blot analysis (Fig. 1A–D). Murine myoblasts treated with si-Fto showed a considerably lower proportion of Ki67-positive cells (Fig. 1E–F). Similarly, we found that the total cell number of murine myoblasts was significantly decreased after si-Fto treatment (Fig. 1G). To assess the effect of Fto on murine myoblast migration, we used a live cell tracking technique to minimize the influence of Fto-regulated cell proliferation and low serum-induced myoblast differentiation. We cultured C2C12 cells in growth media and tracked live cells for 6 h using a high-content screening instrument (Fig. 1H). The accumulated distance, displacement per track, and average speed per track were significantly decreased in si-Fto treated myoblasts (Fig. 1I–K). In conclusion, these data indicate that Fto maintains the proliferation and migration of murine myoblasts.

Fig. 1
figure 1

Fto regulates the proliferation and migration of murine myoblasts. A Representative Western blot analysis of si-Fto interference in C2C12 cells. B Western blot quantification verified the knock-down efficiency of si-Fto interference. n = 3 in each case; data are represented as mean ± s.d. C Quantitative real-time PCR analysis verified the knock-down efficiency of si-Fto interference. n = 5 in each case; data are represented as mean ± s.d. D The m6A level was significantly increased after si-Fto interference. n = 5 in each case; data are represented as mean ± s.d. E Immunofluorescence staining of Ki67 after si-Fto interference in myoblasts. Scale bar, 50 μm. F Interference with si-Fto decreased the percentage of Ki67-positive cells. n = 5 in each case; data are represented as mean ± s.d. G Cell count analysis showed consistent trends with immunofluorescence analysis of Ki67. n = 5 in each case; data are represented as mean ± s.d. H Representative images of the live cell tracking technique. Scale bar, 200 μm. I The accumulated distance was significantly decreased in si-Fto-treated myoblasts. n = 5 in each case; data are represented as mean ± s.d. J Displacement per track was significantly decreased in si-Fto treated myoblasts. n = 5 in each case; data are represented as mean ± s.d. K Average speed per track was significantly decreased in si-Fto treated myoblasts. n = 5 in each case; data are represented as mean ± s.d. *P < 0.05, **P < 0.01, ***P < 0.001

Full size image

Fto maintains the expression of murine slow-twitch fiber related genes

During our preliminary experiments, we found that the first three days of myogenic differentiation exhibited the most pronounced fluctuations in gene expression, particularly for Fto and various myogenic regulatory factors. Meanwhile, on days 4 and 5, as the myogenic differentiation program completed, these regulatory factors tended to stabilize (data not shown). Therefore, we chose to perform various assays on the third day of differentiation to investigate the beneficial function of Fto during myogenic differentiation. We quantified the m6A level and the expression of Fto during the myogenic differentiation process. We found gradually increased Fto expression but decreased m6A level (Fig. 2A), suggesting that Fto was a key regulator of m6A level. In addition, the expression of m6A-related genes was unchanged after the knockdown of Fto (Fig. 2B). Immunofluorescence revealed that the fusion index of MHC1 and myotube width were significantly decreased in si-Fto treated murine myoblasts (Fig. 2C–E). However, neither the proportion of MyoG-positive nuclei (Fig. 2F, G) nor the expression of myogenic regulatory factors (Myog, Myod1, Myf5, and Myf6) showed any discernible changes (Fig. 2H). It was interesting to note that si-Fto treatment reduced the expression of Myh7 and Myh7b, which were primarily expressed in murine slow-twitch fibers (Fig. 2I). Additional experiments were conducted to test whether Fto affects the stability of RNA transcripts of Myh7 and Myh7b, which did not show any significant differences in si-Fto treated murine myoblasts (Fig. 2J–K). These findings imply that Fto maintains the expression of genes relevant to murine slow-twitch fibers in an indirect way.

Fig. 2
figure 2

Fto maintains expression of murine slow-twitch fiber related genes in vitro. A During myogenic differentiation, the mRNA expression of Fto was gradually increased, and the m6A level gradually decreased. n = 5 in each case; data are represented as mean ± s.d. B Quantitative real-time PCR analysis showed that the expression of m6A-related genes was undisturbed after Fto knock-down. n = 5 in each case; data are represented as mean ± s.d. C Representative immunohistochemical images of MHC1 (red) during myogenic differentiation. Scale bar, 100 μm. D Immunofluorescence revealed that the MHC1 fusion index was significantly decreased in si-Fto treated myoblasts. n = 5 in each case; data are represented as mean ± s.d. E Immunofluorescence revealed that myotube width was significantly decreased in si-Fto treated myoblasts. n = 5 in each case; data are represented as mean ± s.d. F Representative immunohistochemical images of MyoG (green) during myogenic differentiation. Scale bar, 100 μm. G Immunofluorescence revealed no significant difference for the MyoG-positive nucleus index in si-Fto treated myoblasts. n = 5 in each case; data are represented as mean ± s.d. H Quantitative real-time PCR analysis showed that the expression of myogenic regulatory factors was undisturbed after Fto knock-down. n = 5 in each case; data are represented as mean ± s.d. I Quantitative real-time PCR analysis revealed that si-Fto interference restrained the expression of slow-twitch fiber related genes, including Myh7 and Myh7b. n = 5 in each case; data are represented as mean ± s.d. J Myh7 mRNA stability showed no difference in si-Fto treated myoblasts. n = 5 in each case; data are represented as mean ± s.d. K Myh7b mRNA stability showed no difference in si-Fto treated myoblasts. n = 5 in each case; data are represented as mean ± s.d. *P < 0.05, **P < 0.01, ***P < 0.001,#P < 0.05, ##P < 0.01

Full size image

Fto contributes to maintain the formation of murine slow-twitch fiber in vivo

To address the potential function of Fto in murine muscle fiber remodeling in vivo, we induced acute muscle injury in mice. FB23-2, a selective inhibitor of the FTO m6A demethylase, was administered into the soleus every 3 days (Fig. 3A). Immunofluorescence revealed that the percentage of type I muscle fibers was significantly decreased in the FB23-2-treated soleus (Fig. 3B, C). In addition, we detected significantly decreased levels of Myh7 and Myh7b but decreased levels of Myh1 and Tnnt3 expression in this soleus (Fig. 3D). Collectively, these results reveal that Fto contributes to maintain murine slow-twitch fiber formation in vivo.

Fig. 3
figure 3

Fto maintains the formation of murine slow-twitch fiber in vivo. A Flow chart of the experiment on C57/BL mice. B Representative immunohistochemical images of MHC1 (green) in the soleus. Scale bar, 100 μm. C Immunofluorescence revealed that the percentage of type I muscle fiber was significantly decreased after treatment with FB23-2. n = 5 in each case; data are represented as mean ± s.d. D Quantitative real-time PCR analysis revealed that FB23-2 treatment decreased the mRNA expression of Myh7 and Myh7b in the soleus. n = 5 in each case; data are represented as mean ± s.d. *P < 0.05

Full size image

Fto regulates murine muscle fiber remodeling in an Nfatc1–Ythdf2-dependent manner

To identify the m6A demethylating targets of Fto, we examined the mRNA levels of several key regulators known to be involved in muscle fiber remodeling, including Pgc-1 (Ppargc1a, Nrf1, and Tfam), Mef2s (Mef2a, Mef2b, Mef2c, and Med2d), and Nfats (Nfatc1, Nfatc2, Nfatc3, Nfatc4, and Nfat5) [15, 16], and murine myoblasts treated with si-Fto substantially reduced the expression of Nfatc1 (Fig. 4A). Recent research studies have indicated that nuclear accumulation of Nfatc1 may trigger the activation of muscle remodeling genes. Moreover, Nfatc1 governs the composition of fiber type and is necessary for the transformation of fast-to-slow fiber type in response to exercise in vivo [17, 18]. We confirmed the decreased level of NFATC1 with Western blot analysis (Fig. 4B, C), and the mRNA stability of Nfatc1 in si-Fto treated myoblasts was significantly reduced (Fig. 4D). Additionally, we measured the level of NFATC1 in the concave paravertebral muscles of 20 AIS patients, and we discovered a positive correlation between FTO and NFATC1 (Fig. 4E). These findings demonstrate that Fto regulates the expression of Nfatc1 in a demethylation-dependent way. YTHDF2 has been shown to destabilize mRNAs as the RNA m6A reader [19]. To investigated whether Fto-mediated Nfatc1 mRNA degradation relied on m6A reader protein YTHDF2, we performed the rescue experiment by silencing endogenous Ythdf2 during myogenic differentiation after FB23-2 was added. We found the expression of Nfatc1, Myh7, and Myh7b was restored after the treatment with si-Ythdf2 (Fig. 4F). In addition, immunofluorescence showed that murine myoblasts treated with si-Ythdf2 had a considerably increased MHC1 fusion index (Fig. 4G, H). Collectively, these data suggest that Fto regulates murine muscle fiber remodeling in an Nfatc1–Ythdf2-dependent manner.

Fig. 4
figure 4

FTO regulates murine slow-twitch fiber related genes in a NFATC1–YTHDF2- dependent manner. A Quantitative real-time PCR analysis revealed that the mRNA expression of Nfatc1 was significantly decreased in si-Fto treated myoblasts during myogenic differentiation. n = 5 in each case; data are represented as mean ± s.d. B Representative Western blot of NFATC1 in si-Fto treated myoblasts during myogenic differentiation. C Western blot quantification showed that NFATC1 was significantly decreased in si-Fto treated myoblasts. n = 3 in each case; data are represented as mean ± s.d. D The mRNA stability of Nfatc1 was significantly decreased si-Fto treated myoblasts. n = 5 in each case; data are represented as mean ± s.d. E The mRNA expression of FTO was positively correlated with the expression level of NFATC1 in the paraspinal muscle of adolescent idiopathic scoliosis. n = 20 in each case; data are represented as mean. F Quantitative real-time PCR analysis revealed that mRNA expression of Ythdf2 was significantly decreased but Nfatc1, Myh7, and Myh7b were increased in si-Ythdf2 silenced myoblasts with FB23-2 treatment. n = 5 in each case; data are represented as mean ± s.d. G Representative immunohistochemical images of MHC1 (red) in si-Ythdf2 silenced myoblasts with FB23-2 treated during myogenic differentiation. Scale bar, 100 μm. H Immunofluorescence revealed that the MHC1 fusion index was significantly increased in si-Ythdf2 silenced myoblasts with FB23-2 treated. n = 5 in each case; data are represented as mean ± s.d. *P < 0.05 and **P < 0.01

Full size image

Asymmetric expression of FTO in AIS paravertebral muscles

We collected AIS paraspinal muscle biopsies from the concave and convex sides of the curve apex during surgery (Fig. 5A). Immunofluorescence revealed that the percentage of type I muscle fibers was significantly decreased on the concave side (Fig. 5B, C). In addition, quantitative real-time PCR analysis showed that the mRNA levels of MYH7 and MYHB, which were predominantly expressed in slow-twitch fibers, were decreased on the concave side, while the mRNA levels of MYH1, MYH4, TNNI2, TNNC2, and TNNT3, which were predominantly expressed in fast-twitch fibers, were increased on the concave side (Fig. 5D). Collectively, our findings show that AIS paravertebral muscles have an asymmetric fiber type.

Fig. 5
figure 5

Reduced FTO in the concave paraspinal muscle of adolescent idiopathic scoliosis. A Magnetic resonance imaging showed convex and concave sides of adolescent idiopathic scoliosis. B Representative immunohistochemical images of MHC1 (red) in paraspinal muscle. Scale bar, 100 μm. C Immunofluorescence showed that the percentage of type I fibers was significantly reduced in concave paraspinal muscle. n = 5 in each case; data are represented as mean ± s.d. D Quantitative real-time PCR analysis showed that mRNA expression of MYH7 and MYH7B was decreased, but MYH1, MYH4, TNNI2, TNNC2 and TNNT3 mRNA levels were increased in concave paraspinal muscle. n = 5 in each case; data are represented as mean ± s.d. E The m6A level was significantly increased in concave paraspinal muscle. n = 5 in each case; data are represented as mean ± s.d. F Quantitative real-time PCR analysis revealed that mRNA expression of m6A-related genes, and only FTO was significantly decreased in concave paraspinal muscle. n = 5 in each case; data are represented as mean ± s.d. G Representative Western blot of FTO in paraspinal muscle. H Western blot quantification showed that FTO was significantly decreased in concave paraspinal muscle. n = 5 in each case; data are represented as mean ± s.d. I The level of mRNA expression of FTO was positively correlated with the proportion of type I fiber. n = 20 in each case; data are represented as mean. J The mRNA expression level of FTO was negatively correlated with the Cobb angle of scoliosis. n = 20 in each case; data are represented as mean. *P < 0.05, **P < 0.01

Full size image

We attempted to further explain the fiber type discrepancy by evaluating the m6A methylation level in AIS paravertebral muscles, and we discovered that the concave side had a significantly higher level of m6A than the convex side (Fig. 5E). We further assessed the mRNA levels of many known regulators linked to m6A methylation writers (METTL3, METTL4, METTL14, and WTAP), erasers (FTO and ALKBH5), and readers (YTHDF1, YTHDF2, YTHDF3, YTHDC1, and YTHDC2). The expression of FTO was considerably downregulated in the concave side of AIS paravertebral muscles, according to quantitative real-time PCR and Western blot analysis (Fig. 5F–H). Additionally, we measured the mRNA levels of FTO in the concave paravertebral muscles of 20 AIS patients. We found a positive correlation between the mRNA level of FTO and the fraction of type I fibers, and a negative correlation between the mRNA level of FTO and the Cobb angle of scoliosis (Fig. 5I, J). These findings suggest that FTO might be involved in muscle fiber remodeling.

Asymmetric expression of FTO in CS paravertebral muscles

In CS, a clear spinal deformity could be found (Fig. 6A), and the asymmetric vertebral body was the primary factor contributing to the development of scoliosis [20]. We, therefore, examined the paravertebral muscles in CS to ascertain whether the asymmetry of the muscle fibers was specific to AIS. Similar to AIS, we discovered that the concave side had a much lower percentage of type I muscle fibers (Fig. 6B, C). Additionally, quantitative real-time PCR analysis revealed that MYH7 and MYHB expression levels were lower on the concave side, whereas MYH1 and TNNC2 expression levels were higher (Fig. 6D). We quantified the m6A methylation level in CS paravertebral muscles, and we found a considerably higher level of m6A on the concave side compared to the convex side (Fig. 6E). The expression of FTO was markedly downregulated in the concave side paravertebral muscles, according to quantitative real-time PCR and Western blot analysis (Fig. 6F–H). These data reveal that the asymmetric features of muscle fibers and FTO expression were consistent in AIS and CS.

Fig. 6
figure 6

Reduced FTO in the concave paraspinal muscle of congenital scoliosis. A Magnetic resonance imaging showed convex and concave curves of congenital scoliosis, and the yellow arrow showed hemivertebra deformity. B Representative immunohistochemical images of MHC1 (green) in paraspinal muscle. Scale bar, 100 μm. C Immunofluorescence showed that the percentage of type I muscle fiber was significantly reduced in concave paraspinal muscle. n = 5 in each case; data are represented as mean ± s.d. D Quantitative real-time PCR analysis showed that mRNA expression of MYH7 and MYH7B was decreased, but MYH1 and TNNC2 mRNA levels were increased in concave paraspinal muscle. n = 5 in each case; data are represented as mean ± s.d. E The m6A level was significantly increased in concave paraspinal muscle. n = 5 in each case; data are represented as mean ± s.d. F Representative Western blot of FTO in paraspinal muscle. G Western blot quantification showed that FTO was significantly decreased in concave paraspinal muscle. n = 5 in each case; data are represented as mean ± s.d. H Quantitative real-time PCR analysis showed that FTO mRNA expression was decreased in concave paraspinal muscle. n = 5 in each case; data are represented as mean ± s.d. *P < 0.05, **P < 0.01

Full size image

Physical exercise restores the asymmetric expression of FTO

Decreased electromyographic amplitudes have been found in the concave side paraspinal muscles of AIS [21, 22], and it is widely known that endurance training facilitates the change from fast to slow fiber types [23]. Since the Schroth method was first developed by Katharina Schroth in the 1920s, it has been widely used in scoliosis patients [24]. To investigate the beneficial effects of Schroth exercises on muscle fiber remodeling, we recruited five AIS patients who performed Schroth exercises for longer than 3 months (Fig. 7A). Immunofluorescence revealed that the percentage of type I muscle fibers was comparable between the convex and concave sides (Fig. 7B, C). In addition, quantitative real-time PCR analysis showed that the expression of MYH7, MYH7B, MYH1, and TNNI2, which were asymmetric in untrained AIS, was also restored (Fig. 7D). Interestingly, there was no difference in the m6A level or FTO expression (Fig. 7E–G). These data indicate that Schroth exercises partially restore the asymmetric expression of FTO and the percentage of type I muscle fibers. The reversible pattern of muscle fibers in AIS and the consistent paraspinal muscle features in AIS and CS suggest that the asymmetry of muscle fibers and FTO expression might not be the fundamental cause of AIS. Perhaps something intrinsic other than the susceptibility gene FTO is what causes AIS.

Fig. 7
figure 7

Physical exercise restores paraspinal muscle asymmetry. A Concave muscle activation was stretched after Schroth exercises. B Representative immunohistochemical images of type I muscle fiber (green) in paraspinal muscle from AIS patients who performed Schroth exercises longer than 3 months. C Immunofluorescence showed that the percentage of type I muscle fiber was comparable between convex and concave sides. n = 5 in each case; data are represented as mean ± s.d. Scale bar, 100 μm. D Quantitative real-time PCR analysis showed that MYH7, MYHB, MYH1, and TNNI2, which had an asymmetric expression in untrained AIS, were restored. n = 5 in each case; data are represented as mean ± s.d. E Representative Western blot for FTO expression between convex and concave sides. F Western blot quantification showed no significant difference in FTO expression between convex and concave sides. n = 5 in each case; data are represented as mean ± s.d. G The m6A level was comparable between convex and concave paraspinal muscle. n = 5 in each case; data are represented as mean ± s.d. *P < 0.05

Full size image

Discussion

The asymmetry of paraspinal muscle in AIS has been demonstrated in several dimensions, including histology, electrophysiological activity, and magnetic resonance imaging. The asymmetric composition of muscle fibers is thought to be the driving force behind AIS. However, the present study suggests that FTO may not be the intrinsic factor driving the etiology of AIS. Instead, the related muscle fiber remodeling is more likely a secondary change after scoliosis. After comparing the paraspinal muscles of patients with AIS and CS, we observed a downregulation in the expression of FTO in the concave paraspinal muscles of both types of scoliosis. In AIS patients, we found a positive correlation between the mRNA level of FTO and the proportion of type I fibers, and Schroth exercises could restore the symmetrical muscle fiber ratio and FTO expression. Mechanistically, we provided experimental evidence from in vivo mouse models and in vitro mouse cell lines, demonstrating that FTO selectively interacts with NFATC1 to prevent its degradation and promote the expression of murine slow-type fiber genes in an NFATC1–YTHDF2-dependent manner.

Researchers have been baffled by the cause-and-effect link between AIS and the asymmetry of its paraspinal muscles for a long time [25]. In a previous study, a reduced proportion of type I muscle fibers was found on the concave side of AIS [22]. We also identified the downregulation of type I muscle fibers and mRNA expression of MYH1, MYH4, TNNI2, TNNC2, and TNNT3 on the concave side of AIS. In addition, as a reference standard, we examined the paravertebral muscles in CS with a definite hemivertebral deformity. Surprisingly, the characteristics of paraspinal muscle asymmetry between these two types of scoliosis were highly consistent. Moreover, we investigated the paraspinal muscles of AIS patients who performed Schroth exercises, and the asymmetric muscle fibers were restored. Exercise enhances muscle performance and endurance by increasing the proportion of slow oxidative fibers, but muscle disuse causes type I fibers atrophy with a slow-to-fast fiber type shift [23, 26]. This reverse pattern suggests that the asymmetry of muscle fibers may not be the primary cause of AIS and that a possible explanation for the decreased proportion of slow-twitch fibers in concave paraspinal muscles is a secondary adaptation due to its chronic low-load demand.

The distinctive feature of AIS Is a low lean mass with no discernible vertebral deformities [27, 28]. The locus rs12149832 near FTO has been identified as significantly associated with AIS [8]. The risk allele of rs12149832 is associated with decreased expression of FTO [8, 27], and previous studies have shown that FTO is required for myogenesis [13, 14]. In this study, we investigated whether FTO was involved in the asymmetry of paraspinal muscle, which could indicate a potential role in the etiology of AIS. Consistent with the asymmetric pattern of muscle fiber proportions, the expression of FTO was significantly decreased on the concave side in AIS and CS, and Schroth exercises partially restored the asymmetric expression of FTO in AIS. These results suggest that perhaps something intrinsic other than the susceptibility gene FTO is what causes AIS.

Muscle fiber remodeling is a reversible process, and it depends on the exact coordination of metabolic and contractile gene expression programs to control fiber type specification [15]. Interestingly, we found that the mRNA level of FTO was positively correlated with the proportion of type I fibers in AIS. FTO has a reversible function in relevant RNA modifications [29], and its coordinated changes with muscle fiber asymmetry suggest that FTO may be involved in fiber type remodeling. In our study, the knockdown of Fto did not affect the mRNA expression of myogenic regulatory factors, such as Myog, Myod1, Myf5, and Myf6. It was noteworthy that slow-twitch fiber development was inhibited by Fto knockdown both in vitro and in vivo, and that Myh7 and Myh7b expression—which was predominantly expressed in slow-twitch fibers—was regulated. These findings imply that FTO promotes the expression of genes relevant to murine slow-twitch fibers.

Muscle fiber remodeling has been studied in extensive detail in the past, and mitochondrial dysfunction could impair the function of skeletal muscle [30]. In addition, endurance training increases the intracellular calcium concentration, which activates calcineurin/NFATs and MEF2s. These two sets of transcription factors are primarily responsible for fiber remodeling and muscle formation [31]. In the present study, FTO promoted the expression of Myh7 and Myh7b, but mRNA stability analysis indicated that they were not directly regulated by FTO demethylation. We further screened the core regulators of muscle fiber remodeling, including mitochondrial biogenesis, NFATs, and MEF2s [32], and NFATC1 was identified as the m6A demethylating target of FTO during myogenic differentiation. NFATC1, which controls fiber type composition, is required for rapid to slow fiber type switching in response to exercise [18]. Moreover, YTHDF2 was identified as the m6A reader to degrade NFATC1 mRNA. Mechanistically, we provided evidence from in vivo mouse models and in vitro mouse cell lines, showing that FTO participates in the murine muscle fiber remodeling process in an NFATC1–YTHDF2-dependent way.

It is essential to recognize its limitations. Firstly, due to the constraints in sample availability, we were unable to investigate the composition of paraspinal muscles in early AIS cases with a curvature range of 10°–20°. Secondly, we implemented a rigorous selection criterion and a paired methodology that inevitably led to a smaller sample size. Lastly, our mechanism study utilized a mouse tibialis anterior injury model and a mouse cell line rather than using myotubes sourced from patients with AIS. These limitations should be taken into consideration when interpreting our results.

Conclusion

FTO supported the formation of murine slow-twitch fibers in an NFATC1–YTHDF2-dependent manner. The consistent paraspinal muscle features in AIS and CS and reversible pattern of muscle fibers in AIS suggest that the asymmetry of muscle fibers might not be the fundamental cause of AIS. Perhaps something intrinsic other than the susceptibility gene FTO is what causes AIS. Instead, FTO may contribute to the muscle fiber remodeling secondary to scoliosis.

Availability of data and materials

The datasets used and/or analysed during the current study are available from the corresponding author on reasonable request.

Abbreviations

AIS:

Adolescent idiopathic scoliosis

CS:

Congenital scoliosis

FTO:

Fat mass and obesity-associated gene

CTX:

Cardiotoxin

References

  1. Wang W, Chen T, Liu Y, Wang S, Yang N, Luo M. Predictive value of single-nucleotide polymorphisms in curve progression of adolescent idiopathic scoliosis. Eur Spine J. 2022;31(9):2311–25.

    Article  PubMed  Google Scholar 

  2. Dunn J, Henrikson NB, Morrison CC, Blasi PR, Nguyen M, Lin JS. Screening for adolescent idiopathic scoliosis: evidence report and systematic review for the US Preventive Services Task Force. JAMA. 2018;319(2):173–87.

    Article  PubMed  Google Scholar 

  3. Bartley CE, Yaszay B, Bastrom TP, Shah SA, Lonner BS, Asghar J, Miyanji F, Samdani A, Newton PO. Perioperative and delayed major complications following surgical treatment of adolescent idiopathic scoliosis. J Bone Joint Surg Am. 2017;99(14):1206–12.

    Article  PubMed  Google Scholar 

  4. Cheng JC, Castelein RM, Chu WC, Danielsson AJ, Dobbs MB, Grivas TB, Gurnett CA, Luk KD, Moreau A, Newton PO, et al. Adolescent idiopathic scoliosis. Nat Rev Dis Primers. 2015;1:15030.

    Article  PubMed  Google Scholar 

  5. Faldini C, Manzetti M, Neri S, Barile F, Viroli G, Geraci G, Ursini F, Ruffilli A. Epigenetic and genetic factors related to curve progression in adolescent idiopathic scoliosis: a systematic scoping review of the current literature. Int J Mol Sci. 2022;23(11):5914.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  6. Pérez-Machado G, Berenguer-Pascual E, Bovea-Marco M, Rubio-Belmar PA, García-López E, Garzón MJ, Mena-Mollá S, Pallardó FV, Bas T, Viña JR, et al. From genetics to epigenetics to unravel the etiology of adolescent idiopathic scoliosis. Bone. 2020;140:115563.

    Article  PubMed  Google Scholar 

  7. Montemurro N, Ricciardi L, Scerrati A, Ippolito G, Lofrese G, Trungu S, Stoccoro A. The potential role of dysregulated miRNAs in adolescent idiopathic scoliosis and 22q11.2 deletion syndrome. J. Pers. Med. 2022;12(11).

  8. Kou I, Otomo N, Takeda K, Momozawa Y, Lu H-F, Kubo M, Kamatani Y, Ogura Y, Takahashi Y, Nakajima M, et al. Genome-wide association study identifies 14 previously unreported susceptibility loci for adolescent idiopathic scoliosis in Japanese. Nat Commun. 2019;10(1):3685.

    Article  PubMed  PubMed Central  Google Scholar 

  9. Liu L, Ding C, Fu T, Feng Z, Lee JE, Xiao L, Xu Z, Yin Y, Guo Q, Sun Z, et al. Histone methyltransferase MLL4 controls myofiber identity and muscle performance through MEF2 interaction. J Clin Invest. 2020;130(9):4710–25.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  10. Duan Y, Li F, Tan B, Yao K, Yin Y. Metabolic control of myofibers: promising therapeutic target for obesity and type 2 diabetes. Obes Rev. 2017;18(6):647–59.

    Article  PubMed  Google Scholar 

  11. Rahman FA, Quadrilatero J. Mitochondrial apoptotic signaling involvement in remodeling during myogenesis and skeletal muscle atrophy. Semin Cell Dev Biol. 2023;143:66–74.

    Article  PubMed  Google Scholar 

  12. Rivera M, Locke AE, Corre T, Czamara D, Wolf C, Ching-Lopez A, Milaneschi Y, Kloiber S, Cohen-Woods S, Rucker J, et al. Interaction between the FTO gene, body mass index and depression: meta-analysis of 13701 individuals. Br J Psychiatry. 2017;211(2):70–6.

    Article  PubMed  PubMed Central  Google Scholar 

  13. Wang X, Huang N, Yang M, Wei D, Tai H, Han X, Gong H, Zhou J, Qin J, Wei X, et al. FTO is required for myogenesis by positively regulating mTOR-PGC-1α pathway-mediated mitochondria biogenesis. Cell Death Dis. 2017;8(3):e2702.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  14. Deng K, Fan Y, Liang Y, Cai Y, Zhang G, Deng M, Wang Z, Lu J, Shi J, Wang F, et al. FTO-mediated demethylation of GADD45B promotes myogenesis through the activation of p38 MAPK pathway. Mol Ther Nucleic Acids. 2021;26:34–48.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  15. Qaisar R, Bhaskaran S, Van Remmen H. Muscle fiber type diversification during exercise and regeneration. Free Radic Biol Med. 2016;98:56–67.

    Article  CAS  PubMed  Google Scholar 

  16. Potthoff MJ, Olson EN, Bassel-Duby R. Skeletal muscle remodeling. Curr Opin Rheumatol. 2007;19(6):542–9.

    Article  PubMed  Google Scholar 

  17. Martins KJ, St-Louis M, Murdoch GK, MacLean IM, McDonald P, Dixon WT, Putman CT, Michel RN. Nitric oxide synthase inhibition prevents activity-induced calcineurin-NFATc1 signalling and fast-to-slow skeletal muscle fibre type conversions. J Physiol. 2012;590(6):1427–42.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  18. Ehlers ML, Celona B, Black BL. NFATc1 controls skeletal muscle fiber type and is a negative regulator of MyoD activity. Cell Rep. 2014;8(6):1639–48.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  19. Li J, Xie H, Ying Y, Chen H, Yan H, He L, Xu M, Xu X, Liang Z, Liu B, et al. YTHDF2 mediates the mRNA degradation of the tumor suppressors to induce AKT phosphorylation in N6-methyladenosine-dependent way in prostate cancer. Mol Cancer. 2020;19(1):152.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  20. Sebaaly A, Daher M, Salameh B, Ghoul A, George S, Roukoz S. Congenital scoliosis: a narrative review and proposal of a treatment algorithm. EFORT Open Rev. 2022;7(5):318–27.

    Article  PubMed  PubMed Central  Google Scholar 

  21. Schmid AB, Dyer L, Böni T, Held U, Brunner F. Paraspinal muscle activity during symmetrical and asymmetrical weight training in idiopathic scoliosis. J Sport Rehabil. 2010;19(3):315–27.

    Article  PubMed  Google Scholar 

  22. Park Y, Ko JY, Jang JY, Lee S, Beom J, Ryu JS. Asymmetrical activation and asymmetrical weakness as two different mechanisms of adolescent idiopathic scoliosis. Sci Rep. 2021;11(1):17582.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  23. Moreillon M, Conde Alonso S, Broskey NT, Greggio C, Besson C, Rousson V, Amati F. Hybrid fiber alterations in exercising seniors suggest contribution to fast-to-slow muscle fiber shift. J Cachexia Sarcopenia Muscle. 2019;10(3):687–95.

    Article  PubMed  PubMed Central  Google Scholar 

  24. Park JH, Jeon HS, Park HW. Effects of the Schroth exercise on idiopathic scoliosis: a meta-analysis. Eur J Phys Rehabil Med. 2018;54(3):440–9.

    Article  PubMed  Google Scholar 

  25. Xu L, Dai Z, Xia C, Wu Z, Feng Z, Sun X, Liu Z, Qiu Y, Cheng JC, Zhu Z. Asymmetric expression of Wnt/B-catenin pathway in AIS: primary or secondary to the curve? Spine (Phila Pa 1976) 2020;45(12):E677–83.

  26. Deshmukh AS, Steenberg DE, Hostrup M, Birk JB, Larsen JK, Santos A, Kjøbsted R, Hingst JR, Schéele CC, Murgia M, et al. Deep muscle-proteomic analysis of freeze-dried human muscle biopsies reveals fiber type-specific adaptations to exercise training. Nat Commun. 2021;12(1):304.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  27. Miyagi M, Saito W, Imura T, Nakazawa T, Shirasawa E, Kawakubo A, Uchida K, Akazawa T, Inage K, Ohtori S, et al. Body composition in japanese girls with adolescent idiopathic scoliosis. Spine Surg Relat Res. 2021;5(2):68–74.

    Article  PubMed  Google Scholar 

  28. Altaf F, Gibson A, Dannawi Z, Noordeen H. Adolescent idiopathic scoliosis. BMJ. 2013;346:f2508.

    Article  PubMed  Google Scholar 

  29. Jiang X, Liu B, Nie Z, Duan L, Xiong Q, Jin Z, Yang C, Chen Y. The role of m6A modification in the biological functions and diseases. Signal Transduct Target Ther. 2021;6(1):74.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  30. Rahman FA, Quadrilatero J. Mitochondrial network remodeling: an important feature of myogenesis and skeletal muscle regeneration. Cell Mol Life Sci. 2021;78(10):4653–75.

    Article  CAS  PubMed  Google Scholar 

  31. Wang T, Xu YQ, Yuan YX, Xu PW, Zhang C, Li F, Wang LN, Yin C, Zhang L, Cai XC, et al. Succinate induces skeletal muscle fiber remodeling via SUCNR1 signaling. EMBO Rep. 2021;22(6):e53027.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  32. Liu N, Nelson BR, Bezprozvannaya S, Shelton JM, Richardson JA, Bassel-Duby R, Olson EN. Requirement of MEF2A, C, and D for skeletal muscle regeneration. Proc Natl Acad Sci U S A. 2014;111(11):4109–14.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

Download references

Acknowledgements

Not applicable.

Funding

This study was sponsored by the National Natural Science Foundation of China (82202767) and the Nonprofit Central Research Institute Fund of the Chinese Academy of Medical Sciences (2019PT320025).

Author information

Authors and Affiliations

  1. Department of Orthopedics, The First Affiliated Hospital of Zhengzhou University, Zhengzhou, 450052, People’s Republic of China

    Wengang Wang

  2. Department of Gynaecology and Obstetrics, The Third Affiliated Hospital of Zhengzhou University, Zhengzhou, 450052, Henan Province, People’s Republic of China

    Xueming Du

  3. Department of Orthopedics, Zhongnan Hospital of Wuhan University, Wuhan, 430071, People’s Republic of China

    Ming Luo

  4. Department of Emergency, The First Affiliated Hospital of Zhengzhou University, Zhengzhou, 450052, People’s Republic of China

    Ningning Yang

Authors
  1. Wengang Wang
    View author publications

    You can also search for this author in PubMed Google Scholar

  2. Xueming Du
    View author publications

    You can also search for this author in PubMed Google Scholar

  3. Ming Luo
    View author publications

    You can also search for this author in PubMed Google Scholar

  4. Ningning Yang
    View author publications

    You can also search for this author in PubMed Google Scholar

Contributions

ML and NNY contributed to the study conception and design. WGW, XMD, and NNY developed the experiment methods and completed all the experiments. XMD conducted the statistical analysis. ML and WGW wrote the original draft, and NNY revised it. All the authors read and approved the final manuscript.

Corresponding authors

Correspondence to Ming Luo or Ningning Yang.

Ethics declarations

Ethics approval and consent to participate

This study was approved by the institutional research ethics board. Animal experiments were performed according to protocols approved by the Experimental Animal Welfare Ethics Committee of Zhongnan Hospital of Wuhan University.

Consent for publication

All authors give their consent for publication.

Competing interests

The authors declare that they have no competing interests.

Additional information

Publisher's Note

Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Supplementary Information

Additional file 1

. Table S1: The patient characteristics of adolescent idiopathic scoliosis and congenital scoliosis. AIS, adolescent idiopathic scoliosis; CS, congenital scoliosis.

Additional file 2

. Table S2: The target sequences of siRNAs for Fto and Ythdf2.

Additional file 3

. Table S3: The real time quantitative PCR primers for human.

Additional file 4

. Table S4: The real time quantitative PCR primers for mouse.

Rights and permissions

Open Access This article is licensed under a Creative Commons Attribution 4.0 International License, which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons licence, and indicate if changes were made. The images or other third party material in this article are included in the article's Creative Commons licence, unless indicated otherwise in a credit line to the material. If material is not included in the article's Creative Commons licence and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of this licence, visit http://creativecommons.org/licenses/by/4.0/. The Creative Commons Public Domain Dedication waiver (http://creativecommons.org/publicdomain/zero/1.0/) applies to the data made available in this article, unless otherwise stated in a credit line to the data.

Reprints and permissions

About this article

Check for updates. Verify currency and authenticity via CrossMark

Cite this article

Wang, W., Du, X., Luo, M. et al. FTO-dependent m6A regulates muscle fiber remodeling in an NFATC1–YTHDF2 dependent manner. Clin Epigenet 15, 109 (2023). https://doi.org/10.1186/s13148-023-01526-5

Download citation

  • Received:

  • Accepted:

  • Published:

  • DOI: https://doi.org/10.1186/s13148-023-01526-5

Keywords

Clinical Epigenetics

ISSN: 1868-7083

Contact us