Introduction
As a common clinical trauma, peripheral nerve injury can lead to severe skeletal muscle atrophy. Denervation-induced muscle atrophy progresses rapidly. Although it is possible to repair peripheral nerve defects, for long-distance nerve defects, due to the slow rate of nerve regeneration, irreversible atrophy often occurs before skeletal muscle is reinnervated, making it difficult to maintain skeletal muscle mass and function before reinnervation [
1‐
3]. Therefore, it is crucial to find ways to alleviate skeletal muscle atrophy. Though, remarkable advance has been made in the investigation of underlying mechanisms of muscle atrophy for the last decades. The trigger factors and specific molecules in the muscle atrophy have not been fully elucidated.
As is well known, the essence of skeletal muscle atrophy is an imbalance between protein synthesis and protein degradation. During denervation-induced muscle atrophy, protein degradation pathways such as ubiquitin–proteasome system (UPS), autophagy-lysosome pathway (ALP), caspase pathway and calpain pathway are significantly activated [
3‐
6]. Among them, Muscle RING Finger 1 (MuRF1) and Muscle Atrophy F-box (MAFbx, also known as Atrogin-1) are two key muscle specific E3 ubiquitin ligases that are significantly upregulated in various muscle atrophy models [
7‐
10]. Transcriptome analysis found that in addition to the changes in protein expression related to protein synthesis and degradation pathways, there were also changes in many mRNA molecules during the process of denervation-induced muscle atrophy, involving epigenetics, angiogenesis, energy metabolism, oxidative stress, inflammation, polyamine metabolism, etc. [
11‐
17]. Recent studies have found that many non-coding RNAs play an important role in denervation-induced muscle atrophy by regulating synthesis and degradation pathways [
18‐
26]. In addition, alternative splicing (AS) is also involved in the denervation-induced muscle atrophy process, and multiple genes involved in degradation pathway have been reported to undergo splicing changes during the process of denervation-induced muscle atrophy [
27]. From this, it can be seen that RNA metabolic disorders may be the trigger event for start transcriptional activity of proteolytic pathways in denervation-induced muscle atrophy.
It is worth noting that all RNA can be modified. Among the more than 150 known RNA modifications, N(6)-methylation of adenosine (m6A) is the most abundant RNA modification [
28]. The m6A modification is a dynamic and reversible post-transcriptional modification, which mediated by 'writers' (methylase, adding methyl groups, methyltransferase-like3 (METTL3), METTL14, Wilms tumor 1associated protein (WTAP), and Vir Like m6A methyltransferase associated (VIRMA)) and ‘erasers’ [demethylase, deleting methyl groups, FAT mass and obesity-associated protein (FTO) and ALKB homologue5 protein (ALKBH5)]. In addition to writer and eraser, transcripts containing m6A are also recognized and managed (determining fate) by m6A binding proteins (Reader), including YTH domain-containing family protein 1–3 (YTHDF1/2/3) and YTH domain-containing protein 1–2 (YTHDC1/2) [
29‐
31]. The m6A is involved in almost all metabolic processes of RNA from neogenesis to decay [
32]. Therefore, it is easy to understand that m6A methylation plays a crucial role in many biological processes, such as development, aging, cell fate determination, and cancer occurrence [
29]. In skeletal muscle, the deletion of muscle fiber specific conditional genes in the m6A writer METTL3 causes spontaneous muscle atrophy [
33]. In skeletal muscle atrophy caused by denervation, the overloaded m6A eraser ALKBH5 activates the atrophy pathway through the HDAC4-FoxO3 axis, ultimately leading to muscle atrophy [
34]. Although it is known that m6A changes driven by regulators can control skeletal muscle mass, it is not clear to what extent m6A changes are caused during the process of denervation-induced muscle atrophy and to what extent they are involved in the atrophy process.
In this study, we performed the m6A immunoprecipitation sequencing (MeRIP-seq) on skeletal muscles at multiple time points after denervation and described the dynamic changes in m6A modification. In addition, we manipulated the content of m6A in skeletal muscle using two drugs and found a negative correlation between m6A levels and skeletal muscle mass. Mechanistically, we found that m6A transcripts altered in the late stage of denervation-induced muscle atrophy are mainly associated with ubiquitin–proteasome pathway. These data provide a new perspective on the molecular mechanism of denervation-induced muscle atrophy, and also provide a basis for future anti-atrophy therapy targeting m6A.
Materials and methods
Animal feeding and management
The animal experiments involved in this project have been approved by the Animal Protection and Utilization Committee of Nantong University and the Jiangsu Provincial Animal Protection Ethics Committee (No. S20200312-003). All animals were kept in a specific pathogen free (SPF) level barrier system at the Experimental Animal Center of Nantong University. Animals were placed in an environment with optimal temperature (24 ± 2 ℃) and automatic control of light cycle, allowing for free access to food and water.
Construction of the model of denervation-induced muscle atrophy
Thirty healthy adult male Sprague Dawley (SD) rats weighing 210 g ± 5 g were provided by the Experimental Animal Center of Nantong University. Rats were randomly divided into 10 groups (n = 3) to establish a sciatic nerve dissociation model. Preoperative weighing was performed and rats were anesthetized with 1% pentobarbital sodium at a dose of 0.3 ml/100 g body weight. Cut off 1.5–2 cm long nerves from the sciatic nerves of rats at the same location, then suture the skin with sutures soaked in 75% ethanol, and finally treat the surgical wound with iodophor. Starting from the time of nerve disconnection, the rats were euthanized using spinal dislocation method at 0 h, 0.25 h, 0.5 h, 1 h, 3 h, 6 h, 12 h, 24 h, 36 h, and 72 h, respectively. The tibialis anterior (TA) muscle was taken and weighed. The muscle tissue was soaked in liquid nitrogen and stored at − 80 °C for RNA extraction. The healthy adult male ICR mice weighing 30 g ± 2 g were provided by the Experimental Animal Center of Nantong University. All ICR mice were randomly divided into 5 groups (0–3 days), 6 groups (0–14 days) and 4 groups (drug treatment). The sciatic nerve dissociation models were established. Anesthetize mice with 1% pentobarbital sodium at a dose of 0.2 ml/10 g body weight. After preparing the corresponding samples, measure and calculate the muscle mass/body mass ratio (mg/g), the muscle wet weight ratio (denervated/contralateral mass ratio), and the muscle fiber cross-sectional area. Other methods refer to rats.
Construction of MeIP library
MeRIP-Seq completed at OE Biotech (Shanghai, China). In short, the total RNA (400 μg, Qualified quality inspection) was purified twice using magnetic beads carrying oligo dT to capture mRNA with ploy (A) tail (polyadenylate). Fragmentation of Poly (A) RNA was performed in a buffer containing 10 mM ZnCl2. Fragmented RNA was divided into two parts, of which 10% was used as input control, and the rest RNA was subjected to m6A RNA immunoprecipitation (IP). RNA was first incubated with m6A polyclonal antibodies, followed by binding to pre-equilibrated m6A-Dynabeads. After multiple elutions, m6A-positive RNA was obtained using phenol–chloroform extraction and ethanol precipitation methods. Evaluate the quality of MeIP library using the BioAnalyzer 2100 system. Sequencing was performed on Illumina Hiseq to obtain a 150 bp double ended reading.
For raw data (raw reads) generated in high-throughput sequencing, firstly Trimmomatic software [
35] was used to remove the connectors, then low-quality reads were filtered out, and finally high-quality clean reads were obtained. SortMeRNA software is used to remove ribosomal RNA [
36]. Use the default parameters of HISAT2 software to compare clean reads to the reference genome of rats, while retaining the unique comparison reads for subsequent analysis. Using input samples as a control, MeTDiff software [
37] was used for peak detection and differential analysis, with parameters of ‘FRAGMENT_LENGTH = 200, PEAK_CUTOFF_PVALUE = 0.01, PEAK_CUTOFF_FDR = 0.05’. The ChIPseeker software was used to annotate the detected peaks. MEME and DREME software were used to detect motifs in peak sequences, while Tomtom software was used to compare the obtained motif sequences with known motif databases and annotate them accordingly using known motifs. The GO database (
www.geneontology.org) was used for GO analysis of differentially m6A genes. The rMATS software was used for variable splicing analysis.
RNA extraction and Real time fluorescence quantitative PCR
According to instructions, total RNA was extracted from muscle tissue using TRIzol reagent (Vazyme, Nanjing, China), and the first strand cDNA was synthesized using a reverse transcription kit (Absin, Shanghai, China). Real time fluorescence quantitative PCR (qPCR) detection was performed using ChamQ Universal SYBR qPCR Master Mix (Vazyme, Nanjing, China) and BIO-RAD system (BIO-RAD-96CFX). Using 18 s as the internal reference, the relative mRNA expression level was detected using the 2
−∆∆Ct method. The mRNA primers were designed and synthesized by Shanghai Shenggong Biology Co., Ltd. The primer information is listed in Additional file
1: Table S1.
Detecting m6A in muscle tissues
The EpiQuik m6A RNA Methylation Quantification Kit (GEPIGENTEK, Farmingdale, New York, USA) was used to analyze the total amount of m6A in the TA muscle of mice. Follow the manufacturer’s instructions, in short, mix the total RNA with a special binding solution to ensure that m6A modified RNA was captured by specific antibodies. After signal enhancement, the m6A RNA-antibody complex was determined by colorimetric quantification at 450 nm using an enzyme-linked immunosorbent assay (Bio-Tek, Vermont, USA).
Immunohistochemistry
Mouse TA muscles were fixed with 4% paraformaldehyde (Beyotime, Shanghai, China) for 12 h, followed by sucrose gradient dehydration. Subsequently, the tissue was cut into 10 µm thick frozen sections using freezing microtome (CM3050S, Leica, Mannheim, Germany) and placed overnight in a 37 ℃ incubator. Slices were rinsed with PBS three times for 5 min each time. After adding an appropriate amount of blocking solution (Beyotime, Shanghai, China), the slices were sealed at room temperature for 1 h. They were incubated overnight with the first antibody anti-laminin antibody (1:1000, Abcam, Cambridge, UK) at 4 ℃, and then incubated with the second antibody Goat anti-mouse IgG H&L (1:500, Abcam, Cambridge, UK) at room temperature in dark for 2 h. Slices were sealed with sealing solution containing DAPI. An upright fluorescence microscope (Zeiss, Oberkochen, Germany) was used to capture fluorescence signals and obtain images. ImageJ software was used to measure the cross-sectional area of muscle fibers in various fields of view.
Western blotting
Tissue samples were lysed with protein lysis buffer (Beyotime, Shanghai, China) added with phosphatase inhibitor mixture (Beyotime, Shanghai, China) and protease inhibitor mixture (Beyotime, Shanghai, China). The BCA detection kit (Beyotime, Shanghai, China) was used to determine protein concentration. Extracted samples (20 μg total proteins per lane) was separated by 10% sodium dodecyl sulfate polyacrylamide gel electrophoresis, then transferred to PVDF membrane (Millipore, Massachusetts, USA), sealed with Tris buffer saline containing 5% skimmed milk powder, and incubated with the primary antibody overnight at 4 ℃. The primary antibodies used in this study include anti-Myosin Heavy Chain/MHC antibody (1:1000, Abcam, Cambridge, UK), anti-Fbx32 antibody (1:1000, Abcam, Cambridge, UK), anti-beta Tubulin (1:1000, Abcam, Cambridge, UK), anti-MuRF1 antibody (1:1000, Abcam, Cambridge, UK), anti-ALKBH5 antibody (1:1000, Proteintech, UK), and anti-FTO antibody (1:1000, Proteintech, UK). The PVDF membrane was eluted multiple times in buffer saline containing 0.1% Tween-20, and incubated with goat anti-rabbit IgG H&L as secondary antibodies at room temperature for 2 h. Immunoblotting was visualized by High-sig ECL Western Blotting Substrate (Tanon, Shanghai, China). Use ImageJ software to analyze contrast and normalize it with reference bands. ImageJ software was used to analyze the grayscale values of immunoblotting bands and normalize the target band with the reference band.
Drug treatment
3-deazidenosine (Daa) is an inhibitor of S-adenosylhomocysteine hydrolase, which has been proved to reduce m6A methylation in vivo [
38,
39]. R-2-hydroxyglutarate (R-2HG) is a tumor metabolite, which can increase intracellular m6A levels by inhibiting the activity of FTO [
40,
41]. 5% DMSO was used to dissolve the drug, with a final concentration of 0.04 mg/μl for Daa and 0.0012 mg/μl for R-2HG. TA muscle intramuscular multipoint injection were performed immediately after the sciatic nerve transection in drug treatment group mice. Each mouse was injected with 15 μl per day for a total of 7 days.
Statistical analysis
All statistical analyses were performed using Prism 9 software (GraphPad, LaJolla, CA). All data are expressed as means ± standard deviation (SD) and analyzed using a one-way analysis of variance. Intergroup differences were detected using Tukey’s multiple comparisons test. A value of p < 0.05 was considered statistically significant.
Discussion
During denervation-induced muscle atrophy, the changes of transcriptome were dynamically regulated and highly coordinated by multiple mechanisms. Recently, m6A methylation has been confirmed to be involved in the development and management of skeletal muscle in multiple species [
33,
34,
54‐
57]. In this study, we described the dynamic landscape of the m6A transcriptome in denervation-induced muscle atrophy. The results showed that m6A levels showed a decreasing trend during denervation-induced muscle atrophy. Moreover, m6A methylation was negatively correlated with gene expression, and a large number of upregulated ubiquitin–proteasome pathway related genes carried m6A modifications. More importantly, we confirmed that increasing m6A methylation was a potential treatment strategy for denervation-induced muscle atrophy.
This study designed dense time points for sequencing, which fully described the dynamics of m6A during denervation-induced muscle atrophy as much as possible. Furthermore, we focused on the m6A characteristics of pre-atrophic skeletal muscles, identified m6A modified genes, and analyzed their functions. Overall, these high-quality data provided valuable resources for further exploration of the role of m6A methylation in skeletal muscle atrophy.
The main effect of m6A on mRNA is to affect its stability [
58,
59]. However, the relationship between m6A and gene expression has not been well explored during denervation-induced muscle atrophy. Our results showed that the relationship between m6A and gene expression was negatively correlated in both healthy and atrophic skeletal muscles, further confirming the role of m6A in degrading mRNA. The correlation in skeletal muscle is different from that in myocardium, indicating that m6A plays different roles in different tissues [
60]. Nearly half of the differentially expressed m6A methylated genes undergone expression changes in the late stages of denervation skeletal muscle atrophy. They were negatively correlated with m6A abundance and can be significantly enriched in pathways such as the ubiquitin proteasome, and thus m6A also acts as a brake to avoid hyper activation of the proteolytic pathways. However, differential m6A methylation genes did not overlap with differentially expressed genes at early stages of denervation suggesting a differential role for m6A at different stages of denervation. In the late stage, m6A mediated mRNA degradation is more involved, and m6A modified in 3′ UTR is more involved in mRNA decay [
61,
62]. Our results showed that 3′ UTR contained an increasing number of m6A peaks as muscle atrophy progresses, which was consistent with above.
Research has shown that the level of m6A controls skeletal muscle size. Increased expression of m6A writer Mettl3 using genetic approaches, and concomitantly m6A overload, caused hypertrophy of skeletal muscle and conversely caused spontaneous muscle atrophy [
33]. The upregulation of m6A eraser ALKBH5 expression and reduction of m6A levels are associated with skeletal muscle atrophy caused by denervation [
34]. Consistent with report, our study provided three direct evidences to confirm a positive correlation between the total level of m6A in skeletal muscle and skeletal muscle mass. Firstly, the level of m6A decreased after 3 days of denervation in skeletal muscle. Secondly, R-2HG increased m6A levels to rescue denervation-induced muscle atrophy. Thirdly, Daa reduced m6A levels in healthy skeletal muscles, leading to occurrence of skeletal muscle atrophy. Our results once again confirmed the positive correlation between m6A levels and skeletal muscle mass. Moreover, these also suggested that m6A decrease might be a common feature during all types of skeletal muscle atrophy.
Skeletal muscle contains multiple cell types, and it is not yet clear which cell type of m6A changes affect muscle atrophy. Muscle stem cells (MuSC) are crucial for skeletal muscle homeostasis and muscle regeneration after injury. Numerous studies have focused on the effects of m6A on the fate of muscle stem cells and muscle regeneration. METTL3 has high expression levels in proliferation and differentiation stages of MuSC, indicating that METTL3 induced high levels of m6A are involved in myogenesis [
63]. Gheller et al. demonstrated that overall m6A levels increased in the early stage of skeletal muscle regeneration in vivo, and knocking down m6A methyltransferase METTL3 levels downregulated overall m6A levels, leading to premature differentiation of C2C12 myoblasts [
42]. MuSC specific knockout of METTL3 significantly inhibited MuSC proliferation and blocked muscle regeneration after injury in mice, but the knockin of METTL3 promoted them [
64]. Contrary to METTL3, the silencing of FTO inhibited the differentiation of MuSC, leading to impaired skeletal muscle development [
65]. In addition, m6A reader Ythdc1 has been reported to be crucial for MuSC proliferation, but it is unclear whether this is mediated by m6A changes [
56,
66]. In the present study, however, it is not clear which cell type contributes to the m6A changes in the tibialis anterior muscle tissue; it could be MuSC, myofibers or others. This interesting issue deserves to be explored in the future.
UPS activation is a direct characteristic of skeletal muscle atrophy initiation. Previous studies had mostly focused on the upstream signaling mediated UPS transcriptional activation, such as ROS, IL6, etc., neglecting the activation mechanism of UPS itself [
14,
67,
68]. Our previous research found that the variable splicing changes of UPS related genes might lead to UPS activation during denervation-induced muscle atrophy [
11,
27]. In this study, we proposed a new mechanism for autonomous activation of UPS, which involved the reduction of m6A modifications in UPS mRNAs during denervation-induced muscle atrophy (especially in the late stage of atrophy), leading to the activation of UPS. It is worth noting that the common genes with differential m6A modification and differential expression are also involved in transcriptional activation related pathways. Transcription factors play crucial roles in the process of neurogenic muscle atrophy, with the Forkhead box O (FoxO) family being extensively studied [
69,
70]. FoxO3 can drive the expression of most atrogenes, including UPS and ALP [
71‐
73]. Our results suggested that the expressions of many transcriptional activation related genes were regulated by m6A, which might indirectly activate UPS. Although there is still a lack of experimental evidence, this may still be one of the important regulatory mechanisms of m6A on denervation-induced muscle atrophy.
The regeneration and decay of m6A are the results of expression changes of writers and erasers. During denervation-induced muscle atrophy, both writers and erasers showed upward expression trends, indicating that the m6A changes were quite complex over time and did not change in a single direction. The expression of m6A eraser FTO showed increased change, and the expression of ALKBH5 showed a gradual upward trend, which were consistent with the study by LIU et al
. [
34]. Based on the overall decrease in m6A, it seemed that the upregulation of ALKBH5 expression was driving the m6A decrease during denervation-induced muscle atrophy. LIU et al. also confirmed that ALKBH5 deletion alleviated denervation-induced muscle atrophy. Both FTO and ALKBH5 have been reported to affect the expression of multiple atrophy genes or the activation of atrophy pathways. ALKBH5 drives skeletal muscle mass reduction after denervation through direct demethylation of HDAC4 and activation of the FoxO3 pathway [
34]. FTO indirectly increases PGC-1α expression through demethylation of intermediate molecules, leading to fiber conversion in diet-induced skeletal muscle fiber remodeling [
74]. In addition, FTO and ALKBH5 cooperatedly activate the downstream FOXO signaling pathway, which in turn affects cell proliferation [
75]. Interestingly, the target of R-2HG is FTO, which does not affect the activity of ALKBH5, and still alleviates denervation-induced muscle atrophy. Although ALKBH5 and FTO both recognize RNA containing m6A, they exhibit key differences in substrate preference, intracellular localization, and the products [
76‐
78]. Therefore, we proposed that the protective effect of m6A on denervation-induced muscle atrophy was not enzyme dependent, but depended on its overall m6A level.
The m6A exhibits therapeutic potential in many diseases [
32]. R-2HG is an FTO targeted inhibitor that has been shown to significantly increase m6A levels and inhibit the development of glioma and leukemia [
40,
41]. This study demonstrated the potential application of R-2HG (or m6A targeted therapy) during denervation-induced muscle atrophy, in addition to anti-tumor therapy. In addition to R-2HG, other FTO inhibitors (or m6A activators), such as Meclofenamic acid, N-(5-Chloro-2,4-dihydroxyphenyl)-1-phenylcyclobutanecarboxamide, as well as some small molecules, also have potential in anti-muscular atrophy effect [
17,
79,
80], and are worth verifying in the future.
In summary, our research established a unique data resource for the study of denervation-induced muscle atrophy. Through data analysis combined with experimental verification, we not only analyzed the dynamic changes of m6A but also confirmed its role, which deepened our understanding of the molecular mechanism in denervation-induced muscle atrophy. Moreover, our data also emphasized that R-2HG was a potential treatment drug during denervation-induced muscle atrophy.
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