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Review

Skeletal Muscle Regeneration in Cardiotoxin-Induced Muscle Injury Models

by
Yanjie Wang
,
Jianqiang Lu
* and
Yujian Liu
*
School of Kinesiology, Shanghai University of Sport, Shanghai 200438, China
*
Authors to whom correspondence should be addressed.
Int. J. Mol. Sci. 2022, 23(21), 13380; https://doi.org/10.3390/ijms232113380
Submission received: 6 September 2022 / Revised: 27 October 2022 / Accepted: 28 October 2022 / Published: 2 November 2022
(This article belongs to the Section Molecular Biology)
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Abstract

:
Skeletal muscle injuries occur frequently in daily life and exercise. Understanding the mechanisms of regeneration is critical for accelerating the repair and regeneration of muscle. Therefore, this article reviews knowledge on the mechanisms of skeletal muscle regeneration after cardiotoxin-induced injury. The process of regeneration is similar in different mouse strains and is inhibited by aging, obesity, and diabetes. Exercise, microcurrent electrical neuromuscular stimulation, and mechanical loading improve regeneration. The mechanisms of regeneration are complex and strain-dependent, and changes in functional proteins involved in the processes of necrotic fiber debris clearance, M1 to M2 macrophage conversion, SC activation, myoblast proliferation, differentiation and fusion, and fibrosis and calcification influence the final outcome of the regenerative activity.

1. Introduction

Skeletal muscle, the main organ of systemic metabolism in the body, is composed of differentiated fibers and displays a strong ability to regenerate after injury. Skeletal muscle injuries occur frequently in daily life and exercise, and the capacity of regeneration is critical for the repair and functional maintenance of skeletal muscle. The regeneration of adult muscle is based on the activation of satellite cells (SCs), which are mononuclear progenitors of skeletal muscle and are located between the sarcolemma and basal lamina [1]. After injury, the regeneration of muscle occurs in three overlapping stages: in the first stage, inflammatory cells infiltrate into damaged sites, and necrotic fiber fragments are removed; in the second stage, SCs are activated and proliferate into myoblasts, thereafter differentiating and fusing to form new muscle cells and replace damaged fibers; the last stage involves the maturation of newly formed fibers and the remodeling of damaged muscle [2,3]. The processes of regeneration are highly coordinated, and the expression of genes involved in regeneration are spatially and temporally regulated [4,5]. Numerous studies have been conducted to investigate the molecular mechanisms underlying muscle regeneration. A comprehensive understanding of the events involved in muscle regeneration will facilitate the treatment of skeletal muscle diseases.
In order to achieve a better understanding of muscle regeneration following physiological injury, the innervation, tendons, vascularization, and SCs should not be injured in mouse models because they contribute to myogenesis following injury. Cardiotoxin (CTX), derived from Naja pallida, induces a transient and reproducible acute injury without affecting the vasculature or nerves, and then produces a consistent injury in the whole muscle followed by synchronized regeneration [6,7,8]. Its application also has the advantages of allowing molecular and biochemical analyses to be performed on the whole muscle in contrast to physiological injury models induced by exercise [9,10]. Additionally, CTX injury models have relatively low harmfulness for animals compared with other non-physiological models such as crushing models [11]. Due to these characteristics, the CTX-induced skeletal muscle injury model is a suitable model for exploring the mechanisms of skeletal muscle regeneration.
In this review, we summarize the effects and mechanisms of different mouse models for obesity, diabetes, exercise training, and nutrition on regeneration after CTX-induced muscle injuries. The results may provide therapeutic targets for the repair of damaged muscle in addition to new ideas for further studies.

2. The Characteristics and Positions of Injury in CTX-Induced Skeletal Muscle Injury Models

CTX, a natural amphiphilic peptide derived from Naja pallida, can affect membrane calcium binding sites, and lower the threshold of calcium-modulated calcium ion release from the sarcoplasmic reticulum, thereafter inducing the destruction of skeletal muscle [12,13]. Muscle injury occurs at days 1 to 2 after CTX injection, where inflammatory cells infiltrate and SCs are activated to proliferate; at days 3–5, the myoblasts are induced to differentiate; at days 5–7, the new fibers with a central nucleus begin to form; and at days 10–14, the major muscle structures are restored; at day 28, the damaged muscles have almost completely recovered [14,15,16]. Due to its characteristics of transience and reproducibility, the CTX-induced injury model has been widely used to explore the mechanisms of skeletal muscle regeneration.
In CTX-induced injury models, the damaged sites are hindlimb muscles, where injuries also often occur in humans. In the related literature, the tibialis anterior is the most widely studied site in CTX-induced injury models. This is because of its obvious location and the characteristics of having a mixture of fiber types. Additionally, as a highly heterogenous muscle, tibialis anterior has only one belly, which results in uniform injury. Gastrocnemius consists mainly of fast-twitch fibers and is a bicep muscle, which may result in nonuniform injury despite the obvious location. Furthermore, other hindlimb muscles were also used in the studies such as the extensor digitorum longus, soleus, and quadriceps. The characteristics of only one type of muscle fiber and the muscle group may lead to a preference for position.
Notably, there are still some limitations in CTX-induced skeletal muscle injury models. First, the skeletal muscles include antigravity (e.g., gastrocnemius, quadriceps) and non-antigravity muscles (e.g., tibialis anterior, biceps brachii) [17]. The mechanisms identified in CTX-induced non-antigravity muscle injury models may not apply directly to the CTX-induced antigravity muscles. Second, in CTX-induced injury models, it always does not affect the vasculature or nerves in muscles [8]. In contrast, the vasculature or nerve damage often occurs during the pathogenesis of human muscle injuries [18]. This discrepancy limits the exploration of the contribution of vasculature or nerves in muscle regeneration using CTX-induced muscle injury models. Third, CTX may induce a complete necrosis of the small muscles such as EDL when examined in cross-section 48 h after injection [19]. This may make it impossible to explore the mechanisms involved in the early stages of these muscles.

3. Skeletal Muscle Regeneration in Different Mouse Models after CTX-Induced Skeletal Muscle Injury

CTX has been used to induce skeletal muscle injury in many mouse models (Table 1) including that of diabetes, obesity, aging, exercise training, mechanical loading, and nutrition intervention, among others. Studies have shown that streptozocin and gene mutation-induced diabetes [20,21,22,23], high fat diet-induced obesity and ob/ob mice [22,24,25], cancer cachexia [26], aging [27,28], irradiation [29], elevated carbon dioxide (CO2) level [30], and hindlimb suspension [31,32] lead to impaired regeneration, whereas exercise training [33,34], microcurrent electrical neuromuscular stimulation [35], microelement zinc [36], and overloading [37,38] improve the regeneration of CTX-induced damaged muscle. The accumulation of mitochondrial DNA alterations activates muscle regeneration in myofibers during aging, but leads to reduced muscle mass [39].
Gender and sex hormone levels also influence the regeneration processes in CTX-induced muscle injuries. Males exhibit larger newly formed fibers than females at the same age after injury, whereas females show higher fat deposition than males during regeneration [40,44] and also remove necrotic tissue more rapidly [44]. Castration of males increases the cross-sectional areas (CSAs) of the newly formed fibers and fat accumulation, whereas ovariectomized mice exhibit inhibited regeneration and decreased adipocyte accumulation, and estrogen supplementation rescues regeneration in ovariectomized mice [41,44]. Lack of estrogen-related receptor α also impairs the recovery of mitochondrial energetic capacity and decreases the activity of adenosine 5′-moophpsphate (AMP)–activated protein kinase (AMPK), which then also leads to delayed regeneration [52].
Additionally, the studies also revealed that different mouse strains have similar regeneration processes with no significant morphological and functional differences. However, the mechanisms of skeletal muscle regeneration may be strain-dependent. For instance, toll-like receptor 4 (TLR4) plays distinct roles in the injured muscle of C57BL/6 and C3H/HeJ [49,53].
It was also reported that the regeneration of skeletal muscle is position-specific: after CTX injury in tibialis anterior and the masseter, head muscles recover slowly and eventually return to the base level, whereas limb muscles show quicker recovery and eventually excessive growth [50].

4. Mechanisms of Regeneration in CTX-Induced Injury Models

It has reported that the trajectories of skeletal muscle regeneration vary considerably despite achieving complete regeneration in different injury models [54], wherein the mechanisms of regeneration in damaged skeletal muscle depend on the injury models [54]. In the following section, we summarize the mechanisms of regeneration based on CTX-induced skeletal muscle injury.

4.1. Inflammatory Response in CTX-Induced Injury Models

Inflammatory response could play an important role in timely skeletal muscle regeneration after CTX-induced injury. In this section, we summarize the mechanisms of this process and its three stages: immune cell infiltration, M1 to M2 macrophage polarization, and the clearance of necrotic fiber debris.

4.1.1. The Mechanisms of Inflammatory Response in CTX-Induced Skeletal Muscle Injury

Upon injury, immune cells residing in the skeletal muscle are rapidly activated and then release tissue destruction factors to accelerate muscle injury [55]. Additionally, the immune cells also secrete cytokines such as tumor necrosis factor alpha (TNFα) and interleukin 6 (IL-6), recruiting neutrophils into damaged areas, which in turn stimulates the secretion of chemokines including monocyte chemoattractant protein 1 (MCP-1), macrophage inflammatory protein 1 alpha (MIP-1α), MIP-1β, and promotes the invasion of circulating monocytes [56]. Studies have shown that the mechanisms of inflammatory response involved in CTX-induced skeletal muscle regeneration are complex (Figure 1, Table 2).
It is reported that the lack of interleukin (IL-1) [77], CC chemokine receptor 2 (CCR2) [81], toll-like receptor 2 (TLR2) [79], and heat shock protein (Hsp70) [78] and the inactivation of IL-6/signal transducer and activator of transcription 3 (STAT3) signaling [82] and complement C3a-C3a receptor (C3aR)/CCL5 signaling [86] lead to reduced/delayed monocyte/macrophage infiltration, which then reduces the clearance of necrotic fiber debris and impairs myoblast proliferation, attenuating/delaying muscle regeneration. The endogenous conversion of n-6 to n-3 polyunsaturated fatty acids [76] and pretreatment with thymol [75] reduce macrophage infiltration and cell apoptosis, leading to increased SC migration and proliferation and improved muscle regeneration. Loss of Kruppel-like factor 2 (KLF2) [65], and the lack of plasminogen activator inhibitor (PAI-1) [2,23] and signal transducer and activator of transcription 1 (STAT1) in bone marrow-derived cells [66] and the inhibition of activin A [69] stimulate monocyte/macrophage recruitment, accelerate damaged muscle degradation, and promote myoblast proliferation, thereby improving muscle regeneration. Additionally, the accumulation of interleukin 17A (IL-17A)-producing T cells can also promote muscle regeneration in a microbiota-dependent way [97]. Lack of neuraminidase-1 (Neu1) [58] and estrogen signaling [59] and increased activation of calcium/calmodulin-dependent protein kinase IV (CaMKIV) [57] increase the inflammatory response but inhibit muscle regeneration. This may be because the lack of Neu1 leads to delayed myoblast differentiation and myofiber maturation [58]; however, the activation of CaMKIV and the lack of estrogen signaling increase the infiltration of pro-inflammatory macrophages, impair the transition of macrophages from M1 to M2, reduce the phagocytosis of macrophages, resulting in impaired muscle regeneration [57].
In addition, muscle cells are also involved in the immune response. Studies have shown that the inflammatory environment induced by interferon gamma (IFN-γ) stimulates the expression of major histocompatibility complex (MHC) and some co-stimulatory molecules from regenerated myofibers or cultured myoblasts and myotubes, which then contribute to the immune response [98]. Myofibers also mediate the inflammatory response through the activation of transforming growth factor beta (TGF-β)/IL-6 signaling, and direct Th17 and Treg cell responses [99]. Moreover, oxidative stress during the inflammatory response can also change the structure and function of proteins, which then regulates muscle regeneration in CTX-induced injury [100].

4.1.2. The Mechanism of Macrophage Polarization in CTX-Induced Skeletal Muscle Injury

Macrophages are the main inflammatory cells, and macrophage polarization is involved in the regulation of regeneration [79]. Infiltrated monocytes differentiate into pro-inflammatory M1 macrophages, secreting proinflammatory cytokines, cleaning up necrotic fiber debris, and maintaining an inflammatory environment. Upon the removal of necrotic fiber debris, M1 macrophages switch to M2 macrophages, secreting anti-inflammatory factors and stimulating regeneration [101]. Research shows that (Figure 1, Table 2) a preexisting inflammatory environment [91], irradiation [29], transglutaminase 2 (TG2) deficiency [102], and the excessive activation of calmodulin-dependent signaling [64] delay or impair the M1 to M2 macrophage conversion, which then delays or impairs muscle regeneration. On the other hand, estrogen signaling [59], extracellular vesicles (EVs) derived from mesenchymal stem cells (MSCs) [73], peroxisome proliferator-activated receptor γ coactivator 1α (PGC-1α) [56], and scavenger receptor class B1 (SRB1) [90] stimulate macrophage polarization, eliminate necrotic fibers, reduce fibrosis, and then induce muscle regeneration. In addition, the lack of progranulin prolongs the existence of M2 macrophages and increases the size of newly formed fibers [72]. Increased activation of peroxisome proliferator-activated receptor beta (PPARβ) promotes the recruitment of M2 macrophages and accelerates the regeneration processes [70].

4.1.3. The Mechanism of Necrotic Fiber Debris Clearance in CTX-Induced Skeletal Muscle Injury

The phagocytotic ability of macrophages plays an important role in the elimination of necrotic fiber debris (Figure 1). The lack of retinol saturase (RetSat) in macrophages results in less milk fat globule-epidermal growth factor-factor-8 (MFG-E8) produced and impaired efferocytosis [103]. However, skeletal muscle regeneration in RetSat-null mice is normal following CTX-injury. This is because other cell types participating in muscle regeneration such as myoblasts compensate for the impaired macrophage function, which leads to normal muscle regeneration in RetSat-null mice [93]. Supplementation with balenine promotes the infiltration of immune cells into damaged muscle, and increases the phagocytotic ability of macrophages, leading to improved regeneration [74].
Additionally, the genes involved in the clearance of necrotic muscle fiber debris are highly expressed in immune cells (Table 2). Disintergrin and metalloprotease (ADAM) 8 expression in neutrophils reduces the expression of P-selectin glycoprotein ligand-1 (PSGL-1) on the surface of neutrophils, which then increases the ability of neutrophils to infiltrate into damaged areas and contribute to the removal of fiber debris [88]. Tyro3/Axl/Mer (TAM) kinase signaling mediated by Axl/Mer (AM) receptor Mer expressed in CD45+ cells and SRB1 expressed in macrophages could also facilitate the elimination of necrotic fibers and stimulate macrophage transition from M1 to M2 [89,90].

4.2. SC Activation and Myoblast Proliferation, Differentiation, and Fusion in CTX-Induced Injury Models

Upon injury, the regeneration capacity of the skeletal muscle is due to the SCs, and the critical steps such as SC activation and myoblast proliferation, differentiation, and fusion determine the extent of regeneration. Additionally, myotube maturation and the self-renewal of SCs also influence regeneration.

4.2.1. Mechanisms of SCs Activation and Myoblast Proliferation in CTX-Induced Injury Models

In normal adult muscle, SCs are in a quiescent state (Figure 2) and express paired box 7 (Pax7); once injury occurs, SCs begin to express myogenic differentiation 1 (MyoD) and myogenic factor 5 (Myf5) and are activated and then enter the cell cycle [104,105,106]. During this process, the basement membrane plays an important role in triggering the activation of SCs. After injury, the components of the basement membrane that mediate the contacts of the basal lamina to SCs and myofibers are degraded, and key components of the basement membrane (collagen IV alpha 1, laminin gamma-1, nidogen-2, and heparane sulfate proteoglycan-2) are downregulated, which further leads to a release of growth factors from the dismantling basement membrane and increased the elasticity of the SC niche, thus providing a suitable environment for SC proliferation [107]. As shown in Table 3, Xin and insulin-like 6 (Insl6) are involved in the activation of SCs. Insl6 overexpression in muscle facilitates SC activation and proliferation through the reduction in cell apoptosis upon CTX injury [108]. Xin, which increased in SCs within 12 h following the CTX-induced injury, maintains the activation of SCs, and the downregulation of endogenous Xin leads to the increased proliferation and migration of myoblasts [109]. Other research has shown that Xin deficiency reduces the activation and proliferation of SCs, and muscle regeneration is then impaired through the reduction in primary myoblasts and increased apoptosis of SCs [110,111]. Additionally, studies have also reported that the over-activation of myostatin/TGF-β receptor/pSmad3 signaling in diabetic mice [43] inhibits the activation of SCs. Nevertheless, the inhibition of TGF-β signaling by simultaneous knockout of TGF-β type I receptor (Tgfbr1) and activin receptor type 2B (Acvr1b) accelerates the myogenic process and improves skeletal muscle regeneration [112], whereas knockout of TGF-β receptor II (TGF-βr2) increases the inflammatory response by affecting T-cell function and the withdrawal at the later stage of muscle regeneration. The lack of nuclear factor (erythroid-derived) like 2 (Nrf2) [113] and lipocalin-2 (LCN2) [114] also inhibits the activation of SCs. These may be associated with the pro-oxidation state, reactive oxygen species (ROS) accumulation, and reduced matrix metalloproteinase-9 (MMP-9) expression, which then lead to delayed or impaired regeneration [113,114].
The capacity for myoblast proliferation is influenced by numerous cytokines. Pretreatment with acetylated myostatin 1 (Ac-MIF1) and acetylated and amidated myostatin 2 (Ac-MIF2-NH2) stimulates muscle regeneration by increasing the capacity of myoblasts for proliferation and differentiation [242]. Chemokines including MCP-1, MIP-1α, or MIP-1β (CCL4) induce extracellular regulated protein kinase (ERK) 1/2 phosphorylation through a Gαi subunit-dependent manner, which promotes myoblast proliferation [170]. Gαi2 also promotes myoblast differentiation through the protein kinase C (PKC)/glycogen synthase kinase 3β(GSK3β)/miR-1 pathway or the histone deacetylase (HDAC) inhibition [160]. Nitric oxide (NO) stimulates the proliferation of SCs via a cyclic guanosine 3′,5′-monophosphate (GMP)-dependent pathway [145]. Double homeobox gene (Duxbl) [151] and factor for adipocyte differentiation 24 (fad24) [172] promote myoblast proliferation via increasing the cell cycle. The increased expression of anti-oxidant superoxide dismutase 1 (Sod1) and catalase (Cat) genes also facilitates the potential of proliferation and differentiation [153]. Additionally, the lack of H19 [169], hippo inhibition [117], and lack of heme oxygenase-1 (Hmox1) [67], fibroblast growth factor 6 (FGF6) [157], p38α, and p38γ [148] have also been found to promote myoblast proliferation, although the mechanisms remain unknown. Tweak/Fn14 also contributes to myoblast proliferation but inhibits their differentiation and delays their regeneration [168]. In contrast, increased inflammation, cell cycle inhibition, the destruction of membrane integrity, and iron accumulation all lead to attenuated myoblast proliferation. The results show that the lack of mitogen-activated protein kinase phosphatase-1 (MKP-1) [63] and heat shock transcription factor 1 (HSF1) [128] increased the inflammation and secretion of proinflammatory cytokines, the lack of α7 integrin destroyed the sarcolemmal integrity [165], and impaired fibroblast growth factor (FGF) responsiveness induced by deficiency of Cdon or fibroblast growth factor receptor 1 (FGFR1) led to the impairment of myoblast proliferation [164,174]. Lack of early growth response3 (Egr3) and the overexpression of calcium/calmodulin-dependent protein kinase kinase 2 (CaMKK2) might induce cell cycle arrest by the inactivation of nuclear factor kappa-B (NF-κB) and the activation of AMPK/p-cdc2-Tyr15 signaling, respectively [159,171]. Peroxisome proliferator-activated receptor δ (PPARδ) deficiency reduced forkhead box protein O1 (FOXO1) expression, which then impaired proliferation; FOXO1 overexpression also induced the expression of cell cycle inhibitors p57 and Gadd45α, which decreased the capacity for myoblast proliferation [127,167]. Iron accumulation and ROS production induced by transferrin receptor 1 (Tfr1) deletion also led to defective myoblast proliferation via the Tfr1–Slc39a14–iron axis [162]. Additionally, the deletion of Notch1 and/or Notch2 [141] and lack of nuclear T3 receptor TRα1 (p43) [158] also inhibited myoblast proliferation. However, the lack of Nur77 did not impair muscle regeneration even though it inhibited myoblast proliferation [150].
MicroRNAs are important regulators in SC activation and myoblast proliferation. The overexpression of miR-378 attenuates the activation and differentiation of SCs in an insulin-like growth factor 1 receptor (IGF1R)-dependent manner, which then delays the regeneration [118]. The expression of miR-29a, induced by fibroblast growth factor 2 (FGF2), reduces the expression of basement membrane members, which results in dismantling of the basement membrane, further providing a suitable environment for myoblast proliferation [107].
Epigenetic regulation is also involved in muscle regeneration. DNA methyltransferases 3a (Dnmt3a) ablation in SCs leads to hypomethylation of p57Kip2, which further induces a higher expression of p57Kip2, and then impairs the SC proliferation and attenuates the regeneration of damaged skeletal muscle [163]. Mixed lineage leukemia protein-1 (MLL1) facilitates the proliferation of myoblasts and Pax7+ SCs by epigenetically increasing the expression of Myf5 by mediating the trimethylation of lysine 4 on the histone H3 protein subunit (H3K4me3) enriched on the Myf5 promoter [123].

4.2.2. Mechanisms of Self-Renewal of SC Pool in CTX-Induced Injury Models

During regeneration, the self-renewal of SCs is also essential for the repair of damaged muscle (Figure 2). The activated SCs downregulate MyoD expression, and then replenish the SC pool through both symmetric cell division and asymmetric cell division [104,143]. In this process, primary cilium, harbored on the surface of quiescent SCs, has been shown to be an intrinsic factor controlling the self-renewal of SCs. Upon SC activation, primary cilia disassemble, and SCs enter the cell cycle. Upon exit from the cell cycle, the primary cilia reassemble again at the surface of a minority of SCs that are committed to self-renewal [143]. Disruption of the cilia reassembly impairs the self-renewal of SCs [143]. Additionally, the lack of selenoprotein N (SelN) [140], angiotensin II/Ang II AT1 receptor (AT1R) [137], and thyroid hormone receptor alpha (TRa) deficiency [139] resulted in a reduced SC pool and impaired regeneration of damaged muscle. Mammalian target of rapamycin complex 2 (mTORC2) depletion does not affect the myogenic function of SCs but impairs the replenishment of the SC pool upon repeated CTX injury [136]. Lack of collagen VI reduces the self-renewal capacity of SCs and impairs muscle regeneration [146].
In contrast, using the CRISPR/Cas9 mutagenesis technique, Sincennes et al. abolished Pax7 acetylation in mice and demonstrated that the lack of Pax7 acetylation led to reduced numbers of asymmetric stem cell divisions, expansion of the SC pool, and increased numbers of oxidative II A myofibers [124]. PKCθ deficiency upregulates Pax7 expression and activates Notch signaling for the maintenance of the self-renewal capacity of SCs in CTX-injured mdx mice [121]. Retinoblastoma (Rb) ablation in SCs increases the cell cycle re-entry of quiescent SCs and promotes the expansion of SCs. However, sustained retinoblastoma 1 (Rb1) loss impairs muscle fiber formation [135]. In addition, Klotho rejuvenates aged SCs and maintains the function of SCs by inhibiting the Wnt signaling pathway [129].

4.2.3. Mechanisms of Myoblast Differentiation in CTX-Induced Injury Models

The mechanisms of myoblast differentiation in Figure 2 and Table 3 show that the upregulation of myogenic regulatory factors (MRFs) such as MyoD and myogenin is associated with the increased capacity of myoblast differentiation, which further contributes to skeletal muscle regeneration in CTX-induced injury models. Research suggests that A-kinase anchoring protein 6 (AKAP6) [195], andrographolide [218], mouse double minute 2 homolog (Mdm2)/CCAAT/enhancer-binding protein β (C/EBPβ) [214], apolipoprotein B mRNA editing enzyme catalytic polypeptide 2 (APOBEC2) knockout [8] induces the expression of MyoD, myogenin, MyoG, and desmin. Leucine-rich repeats and transmembrane domains 1 (LRTM1) inhibit the recruitment of p52Shc to FGFR1 and inhibit the activation of ERK, further reducing the inhibition of cyclin dependent kinase 4 (CDK4) on the transcriptional activity of MyoD [196]. The increased MyoD interacts with its targets transcription elongation factor A-like 7 (Tceal7) and R3h domain containing-like (R3hdml) and promotes myoblast differentiation [1]. Additionally, cyclin D-type binding-protein 1 (Ccndbp1) can bind to MyoD and regulate muscle differentiation [3,186]. The energy metabolism in cells also influences myoblast differentiation. Micropeptide in mitochondria (MPM) increases oxygen consumption and adenosine triphosphate (ATP) synthesis and promotes myoblast differentiation [197]. The expression of the type 1 canonical subfamily of transient receptor potential channels (Trpc1) promotes the influx of calcium in myoblasts during differentiation and activates the phosphatidylinositol-3-kinase (PI3K)/AKT/mTOR/p70S6K pathway [173]. The decrease in chondroitin sulfate (CS) also stimulates the activation of PI3K/AKT signaling [181], which leads to faster regeneration [243]. Additionally, trimetazidine modulates the metabolic shift from free fatty acid β-oxidation to glucose oxidation by stimulating AMPK/PGC1α, and inducing autophagy, both of which contribute to myoblast differentiation [179]. Furthermore, the lack of signal transducer and activator of transcription 6 (STAT6) increases myoblast differentiation in an IL-4-independent way [219]. Angiotensin type 2 receptor (AT2R) inhibits the activation of ERK1/2 signaling to promote myoblast differentiation and fusion [185]. Inositol requiring enzyme 1 (IRE1) suppresses the expression of myostatin through its RNase-dependent RIDD activity, which then promotes the differentiation of myoblasts [188].
In contrast, the increased oxidation state impairs myoblast differentiation. Nonalcoholic fatty liver disease (NAFLD) reduces the SC pool and impairs SC differentiation, leading to attenuated skeletal muscle regeneration. This may be associated with TNF-α upregulation and increased levels of oxidative stress marker nicotinamide adenine dinucleotide phosphate (NADPH) oxidase-2 (NOX 2) [47]. The inhibition of carbonyl reductase1 (CBR1) leads to increased ROS levels and diminishes myoblast differentiation [216]. Iron overload impairs myoblast differentiation through oxidative stress-induced inactivation of the mitogen-activated protein kinase (MAPK) signaling pathway [45]. Lack of Hsp70 [16] and TNF-α receptors p55 and p75 [187,209] also downregulate p38MAPK activation, which further impairs myoblast differentiation. Moreover, decreased expression of MRFs such as MyoD and myogenin also impairs differentiation. Overexpression of ladybird homeobox 1 (Lbx1) [213] and the tripartite motif domain of myospryn [203] inhibit the expression of MyoD and myogenin. High levels of cardiotrophin-1 (CT-1) repress the expression of the MRFs such as MyoD through the activation of mitogen-activated protein kinase kinase (MEK)-MAPK signaling [199]. Protein-activated kinase 1 (PAK1) inhibitor IPA-3 decreases the expression of myogenin and reduces p38 phosphorylation [193]. Teashirt-3 (Tshz3) cooperates with BRG1-associated factor 57 (BAF57) and inhibits the MYOD-dependent activation of Myog [201]. Additionally, HS 6-O-endosulfatases (Sulfs) mutation and lack of tensin lead to reduced withdrawal from the cell cycle and delayed myoblast differentiation [191,211]. The deletion of RNA binding motif protein 24 (Rbm24) also regulates the alternative splicing of myogenic associated genes such as myocyte enhancer factor 2d (Mef2d), Rho-associated protein kinase 2 (Rock2), further inhibiting myoblast differentiation [106].
MicroRNAs are involved in the regulation of myoblast differentiation. The overexpression of miR-351 protects differentiating myoblasts from apoptosis by regulating the target gene E2f3 and contributes to myoblast differentiation [152]. In a normoxic state, miR-210 induces myoblast differentiation in a hypoxia-inducible factor 1-α (Hif1a)-dependent manner [239]. Overexpression of Linc-smad7 increases the expression of smad7 and insulin-like growth factor 2 (IGF2), which then induces myoblast differentiation [178]. In addition, miR-431 directly interacts with the 3′ untranslated region of Smad4. Ectopic miR-431 injection greatly reduces Smad4 levels and improves muscle regeneration in CTX-induced skeletal muscle injury models, whereas the inhibition of miR-431 significantly represses myoblast differentiation [192]. The inhibition of miR-188 reduces the expression of myogenic regulator factor 4 (MRF4) and Mef2c and impairs myoblast differentiation, whereas the overexpression of miR-188 has the opposite effect [155]. Knockdown of transactivating response RNA-binding protein (Trbp) downregulates the expression of miR-1a and miR-133a and reduces myotube formation [244]. Intriguingly, MyoR, a muscle-restricted basic helix–loop–helix transcription factor that antagonizes the actions of MyoD, is found to be anticorrelated with miR-378 during CTX-induced muscle regeneration. MyoD binds to the miR-378 gene and causes both transactivation and chromatin remodeling, thus upregulating miR-378 during myogenic differentiation. The 3′ untranslated region of MyoR contains a direct binding site for miR-378. The presence of this binding site significantly reduces the ability of MyoR and prevents the MyoD-driven transdifferentiation of fibroblasts [180].
Epigenetic regulation is also involved in myoblast differentiation. Histone- and protein arginine methyl transferases 5 (PRMT5)-associated protein COPR5 is required for cell cycle exit and myoblast differentiation. The silencing of COPR5 reduces PRMT5 recruitment to the promoters of p21 and MYOG by hindering interaction with the Runt-related transcription factor 1 (RUNX1)-core binding factor-β (CBFβ), which then inhibits the expression of p21 and MYOG and further impairs myoblast differentiation [200]. IGF-1 induces the phosphorylation and activation of ATP citrate lyase (ACL) through the PI3K/AKT pathway. The activated ACL catalyzes the conversion of citrate into oxaloacetate and acetyl-CoA, and acetyl-CoA can be further utilized by histone acetylases to acetylate H3 (K9/14) and H3 (K27) at the MyoD locus to increase MyoD expression, thereby promoting myoblast differentiation [184].

4.2.4. Mechanisms of Myoblast Fusion in CTX-Induced Injury Models

Differentiated myoblasts fuse with damaged fibers or new myotubes by cell–cell recognition, adhesion, migration, and membrane fusion, subsequently forming multinucleated myotubes [106,230]. This is a dynamic and coordinated process involving many proteins (Figure 2, Table 3).
In terms of stimulating fusion, anoctamin 5 (ANO5) stimulates the repair of the sarcolemmal membrane and facilitates myoblast fusion [224]. Stabilin-2 activates the G-protein coupled receptor (GPCR) activity of BAI3 and then recruits Elmo to the membrane to stimulate myoblast fusion [226]. NADPH oxidase 4 (Nox4) induces the expression of myomarker fusion protein (Tmeme8c) via Nox4-mediated ROS production and then contributes to myoblast fusion [229]. The activation of phospholipase D1 (PLD1) on the plasma membrane facilitates mononucleated myoblast fusion with nascent myotubes [230]. Inhibition of the hierarchical non-clustered miRNA network including highly active (miR-29a), moderately active (let-7), and mildly active (miR-125b, miR-199a, miR-221) networks, stimulates the activation of focal adhesion kinase and AKT and MAPK signaling, and leads to the formation of myotubes [189]. Transient receptor potential cation channel vanilloid I (TRPV I) can be activated by IL-4 and calcium signaling, which then facilitates myoblast fusion instead of proliferation [232]. Syncytin contributes to myoblast fusion, and this effect is male-specific [222].
The mechanism of inhibition of myoblast fusion involves the upregulation of TGF-β via calpain-3 (CAPN3) deficiency, thus leading to defective myoblast fusion [234]. C1q-like 1-4 interacts with BAI3 to repress myoblast fusion [226]. Lack of IGF-1 receptor (IGF-1R) signaling leads to reduced fiber fusion via growth hormone receptor-independent signaling [220]. The expression of (Pro)renin receptor ((P)RR) activates the Wnt/β-catenin and Yes-associated protein (YAP) signaling pathways, and decreases myoblast fusion [228]. In addition, transglutaminase 2 (TG2) [102], constitutive expression of c-Myb lacking its 3′ untranslated region (3′ UTR) [223], inhibition of 3-hydroxy 3-methylglutaryl coenzyme A reductase (HMGR) [225], myasthenia gravis [46], and the lack of ste20-like kinase (SLK) [221] also impair the capacity of myoblast fusion and decrease the fusion index.

4.2.5. Mechanisms of Myotube Maturation in CTX-Induced Injury Models

Fused multinucleated myotubes undergo terminal differentiation and eventually become mature myofibers. Kruppel-like factor (Klf5) (Table 3) is shown to interact with MyoD and Mef2 to regulate terminal differentiation [3]. Doublecortin (Dcx) facilitates myofiber maturation [235]. Sema4C stimulates the phosphorylation of p38 and activates the p38/MAPK signaling pathway to promote terminal differentiation [176]. The expression of clathrin heavy chain like 1 (CHC22) in CTX-induced injury muscle, however, diminishes glucose transporter 4 (Glut4) response and further impairs fiber maturation [237].

4.3. Fibrosis in CTX-Induced Injury Models

Fibrosis is an important stage for regeneration (Figure 3). In this process, the temporary extracellular matrix (ECM) components serve as a scaffold for new fibers and stabilize muscle tissue [89]. In damaged skeletal muscle, fibro/adipogenic progenitors (FAPs) are considered as the main source of fibroblasts [245]. After injury, FAPs are activated and begin to proliferate. This increases FAPs in the necrotic area, which need to be removed in time. Failure to clear FAPs will result in their differentiation into fibroblasts and adipocytes [246]. Fibroblasts secrete extracellular matrix proteins and growth factors and then differentiate into myofibroblasts to increase α-smooth muscle actin (α-SMA) expression and ECM synthesis, finally resulting in fibrosis [247]. Studies on the regulation of FAPs show that (Table 4) IL-4 secreted by infiltrated eosinophils stimulates the activation of FAPs in an IL-4-dependent way. IL-4/IL-13 signaling in FAPs contributes to proliferation and adipogenic differentiation of FAPs is inhibited to facilitate regeneration [55]. IL-1α and IL-1β inhibit the adipogenic differentiation of FAPs, and epidermal growth factor (EGF) and betacellulin (BTC) stimulate the proliferation of FAPs [248]. Lack of TGF-β1 in macrophages inhibits FAP proliferation and reduces fibrosis [249]. Inactivation of retinoic acid (RA) signaling in FAPs leads to adipogenic differentiation, which then impairs regeneration [246].
Additionally, studies have also shown that increases in miR-199a-5p [247], growth differentiation factor 11 (GDF11) [254], and platelet-derived growth factor receptor beta (PDGFRβ) [245], together with the lack of GDF-associated serum protein-1 (Gasp1) and/or Gasp2 [256] and a prior burst of double homeobox 4 (DUX4) [251], induced the deposition of collagen and contributed to fibrosis. In contrast, laminin-111 reduced fibrosis and facilitated skeletal muscle regeneration [165]. Losartam therapy also reduced fibrosis by inhibiting the TGF-β signaling pathway [250].

4.4. Calcification in CTX-Induced Injury Models

Calcification occurs after muscle injury. Under normal conditions, calcification can be resorbed. While in a pathological state, continuous calcification can induce chronic inflammation and/or loss of muscle function [264]. Studies have revealed that (Table 4) after CTX injection, Tie2-expressing endothelial precursors are the main contributor to calcification in a mouse model of dysregulated bone morphogenetic protein (BMP) signaling [258]. Moreover, the inflammatory microenvironment induced by CTX injection is also necessary for calcification in injured muscle [258]. At the early stage after injury, calcific nodules are present in mitochondria, which are mediated by cell death and can be cleared by infiltrated macrophages [257]. Additionally, calcification in damaged muscle may also occur in connection with reduced plasmin, and this is independent of its canonical fibrinolytic function [255]. The hypoxia state induced by CTX injury can induce osteogenic differentiation and mineralization of muscle resident stromal cells and further stimulate the formation of myofiber calcification [259].

4.5. Angiopoiesis and Neurogenesis in CTX-Induced Injury Models

In CTX-induced injury models, the capillaries are destroyed, and endothelial cells are activated to repair the skeletal muscle endothelium. It is reported that (Table 5) macrophage cells derived from bone marrow can express endothelium-related markers such as Tie2 and CD31 to promote angiogenesis [265]. Angiotensin II derived from differentiated muscle myoblasts stimulates the migration of endothelial cells, which also further facilitates angiopoiesis [266]. In contrast, CCR2 deficiency leads to decreased vascular endothelial growth factor (VEGF) production and delayed angiogenesis in injured muscle, which then impairs regeneration [81].
In terms of neurogenesis, M2 macrophages infiltrate damaged muscle, produce hepatocyte growth factor (HGF), and then stimulate the expression of semaphorin 3A (Sema 3A) in myoblasts to regulate the regeneration of motor innervation in injured muscle [270,271]. Pre-activation of satellite cells delays the maturation of the neuromuscular junction by reducing the expression of semaphoring (Sema) 3A and S100B [269]. Lack of desmin leads to disrupted neuromuscular connections [238].

4.6. Other Regeneration-Related Genes in CTX-Induced Injury Models

In addition to the mechanism described above, there are a large number of genes involved in skeletal muscle regeneration in CTX-induced injury models such as Tsukushi, Dicer, mesoderm specific transcript (Mest), filamin C, LYVE-1, and so on (Table 6). However, in these studies, the special role of these genes has not been explored. Further experiments are needed to elucidate their function.

4.7. Non-SC Stem Cells Regulate Regeneration in CTX-Induced Injury Muscle

Non-SC stem cells are also involved in the regulation of regeneration (Figure 4). The results (Table 7) show that bone marrow-derived cells [308,309], pulp cells [310], bone marrow-derived human MSCs [311], hematopoietic stem cells [312], muscle precursor cells [313,314], capillary stem cells [315], adipose-derived mesenchymal stem cells [316], and human amniotic fluid stem cells [317,318] settle in the injured sites and differentiate into myogenic cells to stimulate the skeletal muscle regeneration in CTX-induced injury models. Additionally, mobilization of bone marrow stem cells also accelerates the muscle regeneration [319].

5. Conclusions

Skeletal muscle has a tremendous capacity for regeneration after injury. This is largely due to muscle SCs. In order to learn about the mechanisms of regeneration, skeletal muscle regeneration has been studied for decades in numerous injury models. However, differences in injury exist among the different models, which makes their comparison difficult. In the CTX-induced injury model, a transient and reproducible acute injury is induced without affecting the vasculature or nerves, and this allows for the possibility of performing molecular and biochemical analyses of the whole muscle. Additionally, CTX injury models have a relatively low level of harm for animals in contrast to crushing models, which are invasive and associated with the risk of infection. This explains why CTX-induced injury models have been widely used in exploring the mechanisms of muscle regeneration. To understand the regeneration mechanisms in CTX-induced injury models, we explored all the studies and summarized the characteristics and injury positions, different models of CTX injury, and functional factors involved in the process of regeneration. The results show that the process of regeneration is similar in different mouse strains but that differences exist between gender. Regeneration is impaired in obese, diabetic, and aging mice, whereas exercise, electrical stimulation, and overloading facilitate the regeneration of damaged muscle. Non-SCs transplanted in damaged muscle following CTX injury can also differentiate into myogenic cells and facilitate myogenesis. The emphasis throughout was on the process of regeneration, the changes in the functional proteins involved in the processes of clearance of necrotic fiber debris, M1 to M2 macrophage conversion, SC activation, myoblast proliferation, differentiation and fusion, and fibrosis and calcification, which influence the final outcome of the regenerative activity. However, the inflammatory process in muscle injury and repair is complex, with different effects on muscle regeneration observed in various studies. Additionally, angiopoiesis and neurogenesis also influence the outcome of regeneration, which are easily ignored. Thus, further experiments are needed to explore the mechanisms of inflammatory response during muscle regeneration.

Author Contributions

Writing—original draft preparation, Y.W., J.L. and Y.L.; writing—review and editing, Y.W., J.L. and Y.L. All authors have read and agreed to the published version of the manuscript.

Funding

This work was supported by Natural Science Foundation of China (NO.31971102, NO.32271180, NO.31971101) and Shanghai Frontiers Science Research Base of Exercise and Metabolic Health.

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Conflicts of Interest

The authors declare that they have no competing interests.

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Figure 1. The inflammatory response in CTX-induced skeletal muscle injury. (I) Infiltration of the immune cells. Upon injury, the immune cells are recruited into the damaged area, then induce inflammatory response in the damaged muscle. In this process, the lack of Klf2, STAT1, Hmox1, MKP-1 and upregulation of adiponectin (APN), activin A, and calmodulin (CaM) signaling increase, while the lack of IL-1a/β, Hsp70, CCR2, and IL-6/STAT3 and so on, inhibits the inflammatory infiltration. (II) Elimination of necrotic muscle fibers. The monocyte infiltrated into the damaged site, and become pro-inflammatory macrophages (M1 macrophages). M1 macrophages could secrete proinflammatory cytokines, maintain the inflammatory environment, then clean the necrotic debris. In this process, RetSat/MFG-E8 is required in the regulation of efferocytosis. ADAM8/PSGL-1 signaling, the TAM kinase signaling pathway, and the expression of SRB1 facilitate the elimination of necrotic fibers. Additionally, supplementation of balenine also increases the phagocytosis ability of macrophages. (III) Polarization of the macrophages. As the clearance of the muscle debris, the pro-inflammatory macrophages switch to anti-inflammatory macrophages (M2 macrophages), then secrete anti-inflammatory factors and stimulate regeneration. TG2 deficiency and the excessive calmodulin-dependent signaling delay/impair, while PGC-1α, SRB1, and so on, stimulate the polarization of the macrophages.
Figure 1. The inflammatory response in CTX-induced skeletal muscle injury. (I) Infiltration of the immune cells. Upon injury, the immune cells are recruited into the damaged area, then induce inflammatory response in the damaged muscle. In this process, the lack of Klf2, STAT1, Hmox1, MKP-1 and upregulation of adiponectin (APN), activin A, and calmodulin (CaM) signaling increase, while the lack of IL-1a/β, Hsp70, CCR2, and IL-6/STAT3 and so on, inhibits the inflammatory infiltration. (II) Elimination of necrotic muscle fibers. The monocyte infiltrated into the damaged site, and become pro-inflammatory macrophages (M1 macrophages). M1 macrophages could secrete proinflammatory cytokines, maintain the inflammatory environment, then clean the necrotic debris. In this process, RetSat/MFG-E8 is required in the regulation of efferocytosis. ADAM8/PSGL-1 signaling, the TAM kinase signaling pathway, and the expression of SRB1 facilitate the elimination of necrotic fibers. Additionally, supplementation of balenine also increases the phagocytosis ability of macrophages. (III) Polarization of the macrophages. As the clearance of the muscle debris, the pro-inflammatory macrophages switch to anti-inflammatory macrophages (M2 macrophages), then secrete anti-inflammatory factors and stimulate regeneration. TG2 deficiency and the excessive calmodulin-dependent signaling delay/impair, while PGC-1α, SRB1, and so on, stimulate the polarization of the macrophages.
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Figure 2. SC activation, myoblast proliferation, differentiation and fusion, and the myotube maturation in CTX-induced injury models. (I) SC activation. In the damaged muscle, the quiescent SCs are activated to repair the skeletal muscle. Conversion of n-6 to n-3 polyunsaturated fatty acids (PUFAs), Salvador, testosterone, upregulation of Insl6, and so on, promote, while the lack of Nrf2, LCN2, and miRNA-501 inhibit the activation of SCs. (II) Myoblast proliferation. Activated SCs begin to proliferate and become myoblast. FGF2, RING finger protein13 (RNF13) derived from macrophages, Sod1/Cat, lack of p21, and so on facilitate, while FOXO1/p57/Gadd45α and CaMKK2/AMPK, lack of Gαi2 and H19/Igf2, etc., inhibit myoblast proliferation. (III) Myoblast differentiation and fusion. The expended myoblasts experience differentiation and fuse with other muscle fibers to form multinuclear myotubes. Lack of SLK and APOBEC2, upregulation of miR-206, miR-378, MMP-13, and Ccndbp1 enhance; Parkin, tension, Rbm24, and Hsp70 deficiency inhibit the capacity of myoblast differentiation. Lack of ANO5, HMGR, BAI3, and miR-188, and so on, inhibit, upregulation of Nox4/ROS, PLD1, Mustn1, and TRPV1 stimulate myoblast fusion. (IV) Maturation of myofibers. Lack of Dcx, PTEN, and desmin influence the myofiber maturation. (V) Self-renewal of SCs. The activated SCs proliferate to replenish the SC pool of the skeletal muscle. In this process, Angiotensin II, lack of mTORC2, AIF, SelN, and Notch1/Notch2 impair, while Lgr5, laminin-111, lack of PKC θ, Rb1, and Pax7 acetylation stimulate SC self-renewal or SC pool replenishment.
Figure 2. SC activation, myoblast proliferation, differentiation and fusion, and the myotube maturation in CTX-induced injury models. (I) SC activation. In the damaged muscle, the quiescent SCs are activated to repair the skeletal muscle. Conversion of n-6 to n-3 polyunsaturated fatty acids (PUFAs), Salvador, testosterone, upregulation of Insl6, and so on, promote, while the lack of Nrf2, LCN2, and miRNA-501 inhibit the activation of SCs. (II) Myoblast proliferation. Activated SCs begin to proliferate and become myoblast. FGF2, RING finger protein13 (RNF13) derived from macrophages, Sod1/Cat, lack of p21, and so on facilitate, while FOXO1/p57/Gadd45α and CaMKK2/AMPK, lack of Gαi2 and H19/Igf2, etc., inhibit myoblast proliferation. (III) Myoblast differentiation and fusion. The expended myoblasts experience differentiation and fuse with other muscle fibers to form multinuclear myotubes. Lack of SLK and APOBEC2, upregulation of miR-206, miR-378, MMP-13, and Ccndbp1 enhance; Parkin, tension, Rbm24, and Hsp70 deficiency inhibit the capacity of myoblast differentiation. Lack of ANO5, HMGR, BAI3, and miR-188, and so on, inhibit, upregulation of Nox4/ROS, PLD1, Mustn1, and TRPV1 stimulate myoblast fusion. (IV) Maturation of myofibers. Lack of Dcx, PTEN, and desmin influence the myofiber maturation. (V) Self-renewal of SCs. The activated SCs proliferate to replenish the SC pool of the skeletal muscle. In this process, Angiotensin II, lack of mTORC2, AIF, SelN, and Notch1/Notch2 impair, while Lgr5, laminin-111, lack of PKC θ, Rb1, and Pax7 acetylation stimulate SC self-renewal or SC pool replenishment.
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Figure 3. Fibrosis in CTX-induced injury models. After injury, FAPs are activated and begin to proliferate and then differentiate into fibroblasts and adipocytes. Fibroblasts secrete extracellular matrix proteins and growth factors, and differentiate into myofibroblasts to increase α-SMA expression and ECM synthesis, which may provide scaffolds for the skeletal muscle regeneration while abnormal deposition of ECM finally results in fibrosis. In CTX-induced injury muscle, increases in miR-199a-5p, growth differentiation factor 11 (GDF11), platelet-derived growth factor receptor beta (PDGFRβ), lack of GDF-associated serum protein-1 (Gasp1) and/or Gasp2, and a prior burst of double homeobox 4 (DUX40) induce the deposition of collagen and contribute to fibrosis. In contrast, laminin-111 and Losartam therapy reduce fibrosis. Additionally, IL-4/IL-13 signaling, EGF and BTC, and the RA signaling pathway also regulate the fate of FAPs in skeletal muscle regeneration.
Figure 3. Fibrosis in CTX-induced injury models. After injury, FAPs are activated and begin to proliferate and then differentiate into fibroblasts and adipocytes. Fibroblasts secrete extracellular matrix proteins and growth factors, and differentiate into myofibroblasts to increase α-SMA expression and ECM synthesis, which may provide scaffolds for the skeletal muscle regeneration while abnormal deposition of ECM finally results in fibrosis. In CTX-induced injury muscle, increases in miR-199a-5p, growth differentiation factor 11 (GDF11), platelet-derived growth factor receptor beta (PDGFRβ), lack of GDF-associated serum protein-1 (Gasp1) and/or Gasp2, and a prior burst of double homeobox 4 (DUX40) induce the deposition of collagen and contribute to fibrosis. In contrast, laminin-111 and Losartam therapy reduce fibrosis. Additionally, IL-4/IL-13 signaling, EGF and BTC, and the RA signaling pathway also regulate the fate of FAPs in skeletal muscle regeneration.
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Figure 4. Non-SC stem cells regulate regeneration in CTX-induced injury muscle. Transplantation of hematopoietic stem cells, adipose-derived mesenchymal stem cells, bone marrow-derived mesenchymal stem cells, and amniotic fluid stem cells stimulate regeneration in the damaged skeletal muscle.
Figure 4. Non-SC stem cells regulate regeneration in CTX-induced injury muscle. Transplantation of hematopoietic stem cells, adipose-derived mesenchymal stem cells, bone marrow-derived mesenchymal stem cells, and amniotic fluid stem cells stimulate regeneration in the damaged skeletal muscle.
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Table
Table 1. Different mice models on CTX-induced regeneration in skeletal muscle.
Table 1. Different mice models on CTX-induced regeneration in skeletal muscle.
Author, YearInjury PortionsMouse ModelsTargetsRegeneration
(Impair/Improve)		Ref.
Mouisel E, 2010Right tibialis anterior5 weeks,5, 12 and 18–24 months mdx miceDifferent agesImpair[28]
Fearing CM, 2016Right hind limb anterior and post compartmentsMale and female C57BL/6J mice; young:4-6 months, middle:12–19 months, old 25–30 months and very old:32–33 monthsAges and sexImpair; ——[40]
Takahashi Y, 2021Left tibialis anterior22 weeks C57BL/6-WT, C57BL/6-Akita, KK/Ta-WT, and KK/Ta-Akita male miceDiabetesImpair[20]
Vignaud A, 2007Right tibialis anterior3–4 months STZ-treated Swiss and Akita male miceDiabetesImpair[21]
Chaiyasing R, 2021Right tibialis anterior12 weeks ovariectomized and normal C5BL/6JJcl female miceEstrogenImpair[41]
Rebalka IA, 2017Tibialis anterior10–12W STZ-treated and normal male C57BL/6J miceFluvastatinImpair[42]
Nguyen MH, 2011Extensor digitorum longus14–16 weeks leptin-deficient, leptin receptor-mutant mice and a group of C57BL/6 mice fed a high-fat dietob/ob and db/dbImpair[22]
D’Souza DM, 2015Left gastrocnemius-plantaris, tibialis anterior, quadriceps16 weeks male C57BL/6J mice fed a high-fat dietDiet-induced obesityImpair[24]
Jinno N, 2014Right gastrocnemius8 weeks male C57BL/6 mice: a marginally zinc-deficient diet-fed group, a zinc-adequate diet-fed group and a zinc-high diet-fed groupZincImpair[36]
Matsuba Y, 2009Soleus musclegravitational unloadingGravitational unloadingImpair[32]
Jeong J, 2013Tibialis anterior and gastrocnemius2–4 months C57BL/6 male mice treated with STZ, 20–24 months C57BL/6 male mice and C57BL/6J-Ins2Akita miceDiabetesImpair[43]
Inaba S, 2018Tibialis anteriorcancer cachexiaCancer cachexiaImpair[26]
McHale MJ, 2012Right hind limb anterior and posterior compartment4–6 months ovariectomized, castrated and normal male and female C57BL/6J miceSex hormones——[44]
Patsalos A, 2017Tibialis anterior2–6 months BoyJ, C57BL/6J male mice with or without irradiation and Pax7 Cre-Rosa26 DTA miceIrradiationImpair[29]
Ikeda Y, 2019Gastrocnemius8 weeks male C57BL/6J mice with or without iron overloadIronImpair[45]
Attia M, 2017Right tibialis anterior10 weeks myasthenia gravis and normal female C57BL/6J miceMyasthenia gravisImpair[46]
Saliu TP, 2022Left tibialis anterior and gastrocnemius9 weeks normal and NAFLD male CD-1 miceNAFLDImpair[47]
Rahman FA, 2020Left tibialis anteriorYoung (3 months) and old (18 months) male C57BL/6J miceAgingImpair[48]
Kohno S, 2012SoleusC57BL/6J mice with or without tail suspensionUnloadingImpair[38]
Kimoloi S, 2022Tibialis anteriorBoth male and female K320Eskm and K320Emsc transgenic miceMitochondrial DNA alterationsImpair[39]
Paiva-Oliveira EL, 2017Right gastrocnemiusIsogenic 6–8 weeks C3H/HeJ (TLR4 defective), C3H/HeN(TLR4 WT), C57BL/6 (WT) and TLR-4 knockout male miceDifferent strains——[49]
Yoshioka K, 2021Tibialis anterior and masseterC57BL/6J and mdx micePosition specificity——[50]
Joanisse S, 2016Tibialis anterior24 months old male C57BL/6J mice with or without exercise and 8 weeks young male miceExercise and ageImprove; impair[33]
Nagata K, 2013Tibialis anterior8 weeks C57BL/6J mice with ultrasound exposureUltrasoundImprove[51]
Morioka S, 2008Soleus10 weeks male C57BL/6J mice with or without functional overloadingFunctional overloadingImprove[37]
Fujiya H, 2015Left tibialis anterior7 weeks male C57BL/6J mice with or without microcurrent electrical neuromuscular stimulation (MENS)MENSImprove[35]
Table
Table 2. The mechanism of inflammatory infiltration, conversion of macrophages, and elimination of the necrotic debris.
Table 2. The mechanism of inflammatory infiltration, conversion of macrophages, and elimination of the necrotic debris.
Author, YearInjury PortionsTarget Molecule/
Drug		Target ProcessExpressionEffects
(Positive/Negative)		Regeneration
(Impair/Improve)		Ref.
Shi D, 2018Tibialis anteriorCaMKIVInfiltration of macrophagesUp/DownPositive/NegativeImpair/improve[57]
Neves Jde C, 2015Right tibialis anteriorNeuraminidase-1Inflammatory response; myofiber maturationDownPositive; NegativeImpair[58]
Liao ZH, 2019Tibialis anteriorEstrogen signalingInflammation infiltration; conversion of macrophages from M1 to M2DownPositive; NegativeImpair[59]
Kohno S, 2011Tibialis anteriorCbl-bCytotoxic T-cell infiltrationDownPositiveImpair[60]
Wang H, 2014Right tibialis anterior and hindlimb posterior compartmentMonocyte/
macrophage		Monocyte and macrophages recruitment; conversion of macrophages from M1 to M2——Positive; NegativeImpair[61]
Park CY, 2010Gastrocnemius and soleusskNACInflammation infiltration and myonecrosisDownPositiveImpair[62]
Shi H, 2010Right tibialis anteriorMKP-1Inflammation; myoblast proliferation; differentiationDownPositive; Negative; PositiveImpair[63]
Hu J, 2019Tibialis anteriorCaM signalingInflammatory responseUpPositiveImprove[64]
Manoharan P, 2019GastrocnemiusKlf2Inflammatory responseDownPositiveImprove[65]
Gao Y, 2012Unilateral tibialis anteriorSTAT1Inflammatory responseDownPositiveImprove[66]
Kozakowska M, 2018GastrocnemiusHmox1Inflammation and SC proliferationDownPositiveImprove[67]
Koh, 2005Extensor digitorum longusPAI-1Macrophage and SC migrationDownPositiveImprove[2]
Zhang, 2020Tibialis anterior and gastrocnemiusIFN-γ/CXCL10/
CXCR3		Macrophages and myoblast proliferation——PositiveImprove[68]
Yaden BC, 2014Right GastrocnemiusActivin AMacrophage infiltrationUpPositiveImprove[69]
Mothe-Satney I, 2017Left Tibialis AnteriorPPARβMacrophage recruitmentUpPositiveImprove[70]
Tanaka Y, 2019Tibialis AnteriorAPNElimination of the necrotic fibersUpPositiveImprove[71]
Dinulovic I, 2016Tibialis AnteriorPGC-1αConversion of macrophages from M1 to M2Up/DownPositive; NegativeImprove/Impair[56]
Sugihara H, 2018Tibialis AnteriorPGRNProlonged Persistence of M2 MacrophagesDownPositiveImprove[72]
Lo Sicco, 2017Tibialis AnteriorExtracellular vesicles released by human adipose derived-MSCsConversion of macrophages from M1 to M2UpPositiveImprove[73]
Yang M, 2022tibialis anteriorBaleninePhagocytosis ability of macrophages——Positiveimprove[74]
Cardoso ES, 2016GastrocnemiusThymolInflammatory response——NegativeImprove[75]
Wang ZG, 2021GastrocnemiusConversion of n-6 to n-3 PUFAsInflammatory response; SC activationUpNegative; PositiveImprove[76]
Chaweewannakorn C, 2018Unilaterally tibialis anteriorIL-1a/βInflammatory responseDownNegativeImpair[77]
Senf SM, 2013Tibialis anteriorHsp70Inflammatory responseDownNegativeImpair[78]
Mojumdar K, 2016Tibialis anteriorTLR2Macrophage accumulation; elimination of the necrotic fibersDownNegative; NegativeImpair[79]
Varga T, 2013Tibialis anteriorNUR77Macrophage developmentDown——Impair[80]
Ochoa O, 2007Right anterior and posterior compartmentCCR2Macrophage recruitment and angiogenesis and VEGF productionDownNegativeImpair[81]
Zhang C, 2013Tibialis anterior and gastrocnemiusIL-6/STAT3Infiltration of macrophages and myoblast proliferationDownNegativeImpair[82]
Zhang J, 2014Tibialis anteriorCD8Macrophage recruitmentDownNegativeImpair[83]
Martinez CO, 2010Right tibialis anterior and hindlimb posterior compartmentCCR-2/MCP-1Macrophage recruitmentDownNegativeImpair[84]
Krause MP, 2013Left tibialis anterior and gastrocnemius-plantaris-soleusDiabetesMacrophages infiltration————Impair[23]
Cheng M, 2008Extensor digitorum longus and tibialis anteriorIFN-γMacrophages infiltration; myoblast proliferationDownNegativeImpair[85]
Zhang, 2017Tibialis anterior and gastrocnemiusC3aMonocyte/macrophage infiltrationDownNegativeImpair[86]
Sun D, 2009Right hind limb anterior and posterior compartmentCCR2Recruitment of macrophages and neutrophilsDownNegativeImpair[87]
Nishimura D, 2015Tibialis anteriorADAM8Elimination of the necrotic fibersDownNegativeImpair[88]
AI-Zaeed N, 2021Tibialis anteriorTAM kinase receptor MerElimination of the necrotic fibers and conversion of macrophage from M1 to m2sDownNegativeImpair[89]
Zhang J, 2019Tibialis anteriorSRB1Elimination of the necrotic fibers and conversion of macrophage from M1 to M2DownNegativeImpair[90]
Jin R M, 2018One or more hindlimb musclesPreexisting inflammatory environmentConversion of macrophage from M1 to M2——NegativeImpair[91]
Bronisz-Budzyńska I, 2020GastrocnemiusNrf2Inflammatory responseDownPositiveNo effects[92]
Tarban N, 2022Tibialis anteriorRetinol saturasePhagocytosis ability of macrophagesDownNegativeNo effects[93]
Dalle S, 2020Tibialis anteriorIbuprofenInflammatory response——NegativeNo effects[94]
Shen W, 2008GastrocnemiusMacrophage, TFG-β1 and COX-2Inflammatory response——————[95]
Rousseau AS, 2021Left tibialis anteriorPPARβ/δT cell dynamicDown————[96]
Table
Table 3. The mechanism of SC activation, myoblast proliferation, differentiation, and fusion.
Table 3. The mechanism of SC activation, myoblast proliferation, differentiation, and fusion.
Author, YearInjury PortionsTarget Molecule/
Drug		Target ProcessExpressionEffects
(Positive/Negative)		Regeneration
(Impair/Improve)		Ref.
Schaaf G J, 2018Quadriceps femoris and gastrocnemiusAcid alpha glucosidaseSC activationDownNegativeImpair[115]
Serra C, 2013Left tibialis anteriorTestosteroneSC activationUpPositiveImprove[116]
Liu Q, 2021Left tibialis anteriorSalvadorSC activation and angiogenesisDownPositiveImprove[117]
Zeng L, 2010Tibialis anterior or gastrocnemiusInsl6SC activation and proliferationUp/DownPositive/NegativeImprove/impair[108]
Shelar SB, 2016Tibialis anteriorNrf2SC activation and proliferationDownNegativeImpair[113]
Rebalka IA, 2018Left tibialis anterior or left gastrocnemiusLcn2SC activation and fibrosisDownNegativeImpair[114]
Zeng P, 2016Right tibialis anteriorMir-378/IGF1RSC activation and differentiationUpNegativeImpair[118]
Nissar AA, 2011Tibialis anteriorXinSC activationDownNegativeImpair[111]
Lagalice L, 2018Tibialis anteriorAcid alpha glucosidaseSC activationDownNegativeImpair[119]
Mizbani A, 2016Tibialis anteriorMirna-501SC activationDownNegativeImpair[120]
Fiore PF, 2020Tibialis anteriorPkcθSC self-renewalDownPositiveImprove[121]
Fortier M, 2013Tibialis anteriorS1pr3SC proliferationDownPositiveImprove[122]
Cai S, 2020Tibialis anteriorMll1/myf5SC proliferationUpPositiveImprove[123]
Sincennes MC, 2021Tibialis anteriorPax7 acetylationSC poolDownPositiveImprove[124]
Naito T, 2009Tibialis anteriorG-csfSC numberUpPositiveImprove[125]
Ohno Y, 2016Left soleusMstnSC numberDownPositiveImprove[31]
Price FD, 2014Tibialis anteriorJak/statSC numberDownPositiveImprove[126]
Hillege MMG, 2022Tibialis anteriorTGF-β signalingSC numberDownPositiveImprove[112]
Angione AR, 2011Tibialis anterior and gastrocnemiusPparδSC number and proliferationDownNegativeImpair[127]
Nishizawa S, 2013Left soleusHsf1SC number and proinflammatory responseDownNegativeImpair[128]
Ahrens HE, 2018Right tibialis anteriorKlothoSC number and functionDownNegativeImpair[129]
Sakamoto K, 2019Forearm muscleR3hdmlSC numberdownNegativeImpair[130]
Bye-A-Jee H, 2018Tibialis anteriorZFP36L1 and ZFP36L2SC numberDownNegativeImpair[131]
Tonami K, 2013Left tibialis anteriorCapn6Myoblast differentiationDownPositiveImprove[132]
Accornero F, 2014Tibialis anteriorTGF-βSC number and activity; decreased degenerationDownPositive; PositiveImprove[133]
Van Ry PM, 2014Tibialis anteriorLaminin-111SC pool; fibrosisUpPositive; NegativeImprove[134]
Hosoyama T, 2011Tibialis anteriorRb1SC pool; differentiationDownPositive; NegativeImpair[135]
Rion N, 2019Unilaterally tibialis anteriormTORC2SC pool replenishmentDownNegativeImpair[136]
Yoshida T, 2013Unilateral gastrocnemiusAngiotensin IISC pool and proliferationUpNegativeImpair[137]
Armand AS, 2011Soleus or EDLAIFSC poolDownNegativeImpair[138]
Milanesi A, 2017Right tibialis anterior or quadriceps femorisThyroid hormone receptor alphaSC poolDownNegativeImpair[139]
Castets P, 2011Unilateral tibialis anterior and soleus musclesSelNSC poolDownNegativeImpair[140]
Fujimaki S, 2018Right tibialis anteriorNotch1/Notch2SC pool and proliferation; myoblast differentiation; fibrosisDownNegative; Positive; PositiveImpair[141]
Johnston AP, 2011Tibialis anteriorAng IISC number; myoblast differentiationDownPositive; NegativeImpair[142]
Jaafar M N, 2016Right tibialis anteriorPrimary ciliumSC self-renewal——————[143]
Leung C, 2020Tibialis anterior/Extensor digitorum longusLgr5SC replenish and myofiber formation——PositiveImprove[144]
Buono R, 2012Tibialis anterior and quadricepsNO signalingSC self-renewal and proliferationDownNegativeImpair[145]
Urciuolo A, 2014Tibialis anteriorCollagen VISC self-renewalDownNegativeImpair[146]
Alexeev V, 2014Left gastrocnemius muscleAdipose-derived stem cellsMigration of SCs——PositiveImprove[147]
Brien P, 2013Tibialis anterior/extensor digitorum longus muscle groupP38αMyoblast proliferation; differentiationDownPositive; NegativeImpair[148]
Hawke TJ, 2003Tibialis anteriorp21Myoblast proliferation; differentiationDownPositive; NegativeImpair[149]
Cortez-Toledo O, 2017Tibialis anteriorNur77Myoblast proliferationDownNegativeNo effects[150]
Alves, 2019Rectus femoral muscleKinin-B2 receptorMyoblast proliferation; differentiationDownPositive; NegativeImpair[4]
Wu, 2014Right tibialis anteriorDuxblMyoblast proliferation and differentiationUpPositive; NegativeImpair[151]
Chen Y, 2012Tibialis anteriormiR-351Myoblast proliferation and differentiationUp/DownPositive/NegativeImprove/impair[152]
Tseng C, 2019GastrocnemiusSod1/CatMyoblast proliferation and differentiationUpPositiveImprove[153]
Jia Y, 2012GastrocnemiusEPOProliferation and survival of the SCs——PositiveImprove[154]
Shibasaki H, 2019Tibialis anteriormiR-188Myoblast fusionUp/DownPositive/NegativeImprove/impair[155]
Hawke TJ, 2007Tibialis anteriorXinMyoblast proliferation and migrationDownPositiveImprove[109]
Meng, 2014Tibialis anteriorRNF13Myoblast proliferation and differentiationUpPositiveImprove[104]
Lee EJ, 2021Left gastrocnemiusGlycyrrhiza uralensis-extracted compoundsMyoblast proliferation and differentiation——PositiveImprove[156]
Galimov A, 2016Tibialis anteriorFGF2Myoblast proliferationUpPositiveImprove[107]
Armand, 2003SoleusFGF6Myoblast proliferationDown/UpNegative/PositiveImpair/improve[157]
Shi, 2010Tibialis anterior and gastrocnemiusTceal7Myoblast proliferation; differentiationUpNegative; Positive——[1]
Pessemesse L, 2019Right tibialis anteriorp43Myoblast proliferationDown/UpNegative/PositiveImpair/improve[158]
Ye, 2016GastrocnemiusCaMKK2Myoblast proliferation and differentiationUp/DownNegative/PositiveImpair/
Improve		[159]
Minetti GC, 2014Tibialis anteriorGαi2Myoblast proliferation, differentiation and fusionDownNegativeImpair[160]
Zhang CC, 2021Tibialis anteriorCyp4a14Myoblast proliferation and differentiation and inflammatory responseDownNegativeImpair[161]
Ding, 2021Tibialis anteriorTfr1Myoblast proliferation and differentiationDownNegativeImpair[162]
Naito M, 2016Tibialis anteriorDnmt3aMyoblast proliferationDownNegativeImpair[163]
Bae, 2020Tibialis anteriorCdonMyoblast proliferation and senescenceDownNegativeImpair[164]
Rooney JE, 2009Left tibialis anteriorα7 integrinMyoblast proliferation and differentiationDownNegativeImpair[165]
Katsushi, 2020Tibialis anteriorBach 1Myoblast proliferation and differentiationDownNegativeImpair[166]
Yamashita, 2016GastrocnemiusFOXO1Myoblast proliferationUpNegativeImpair[167]
Al-Sajee D, 2015Left tibialis anterior, gastrocnemius/
Plantaris/soleus, quadriceps muscles		XinMyoblast proliferationDownNegativeImpair[110]
Girgenrath, 2006Tibialis anteriorFn14Myoblast proliferationDownNegativeImpair[168]
Martinet C, 2016Tibialis anteriorH19Myoblast proliferationDownNegativeImpair[169]
Yahiaoui, 2008Tibialis anteriorMCP-1Myoblast proliferationUpNegativeImpair[170]
Kursaka, 2017One leg of tibialis anteriorEgr3Myoblast proliferationDownNegativeImpair[171]
Ochiai N, 2016Tibialis anteriorfad24Myoblast proliferationDownNegativeImpair[172]
Zanou, 2012Tibialis anterior and extensor digitorium longusTrpc1Myoblast migration and differentiationDownNegativeImpair[173]
Yablonka-Reuveni Z, 2015Unilateral tibialis anteriorFGFR1Myoblast proliferationDownNegativeNo effects[174]
Ohtsubo, 2017GastrocnemiusAPOBEC2Myoblast differentiation and fusionDownPositiveImprove[8]
He, 2019Extensor digitorum longusNicotineMyoblast differentiation——PositiveImprove[175]
Wu, 2007Tibialis anteriorSema4CMyoblast differentiationUp/DownPositive/NegativeImprove/impair[176]
Liu, 2012Tibialis anteriormiR-206Myoblast differentiationUpPositiveImprove[177]
Song, 2018Tibialis anteriorLinc-smad7Myoblast differentiationUpPositiveImprove[178]
Gatta L, 2017Right tibialis anteriorTrimetazidineMyoblast differentiation——PositiveImprove[179]
Gagan, 2011Tibialis anteriormiR-378Myoblast differentiationUppositiveImprove[180]
Mikami T, 2012Tibialis anteriorChondroitin sulfateMyoblast differentiationDownPositiveImprove[181]
Lee KP, 2015Hindlimb musclemiR-431Myoblast differentiationUppositiveImprove[27]
Storbeck CJ, 2013Tibialis anteriorSLKMyoblast differentiationDownPositiveImprove[182]
Lei, 2013One leg of tibialis anteriorMMP-13Myoblast migrationDown/UpNegative/PositiveImpair/improve[183]
Das, 2017Tibialis anteriorACLMyoblast differentiationDown/UpNegative/PositiveImpair/improve[184]
Yoshida T, 2014GastrocnemiusAT2RMyoblast differentiationDown/UpNegative/PositiveImpair/improve[185]
Huang Y, 2016Right tibialis anteriorCcndbp1Myoblast differentiationDown/UpNegative/PositiveImpair/
Improve		[186]
Chen SE, 2007SoleusTNF-αMyoblast differentiationDown/UpNegative/PositiveImpair/
Improve		[187]
He, 2021Tibialis anteriorIRE1aMyoblast differentiation and hypertrophyDown/UpNegative/ PositiveImpair/improve[188]
Luca E, 2020Tibialis anteriormiRNA networkMyoblast differentiationDownNegativeImprove[189]
Esteca MV, 2020Left tibialis anteriorParkinMyoblast differentiationDownNegativeImpair[190]
Ishii A, 2001Tibialis anteriorTensinMyoblast differentiation and fusionDownNegativeImpair[191]
Zhang M, 2020Tibialis anterior of one limbRbm24Myoblast differentiationDownNegativeImpair[106]
Lee, 2015Tibialis anteriormiR-431Myoblast differentiationDownNegativeImpair[192]
Fan, 2018Tibialis anteriorHsp70Myoblast differentiationDownNegativeImpair[16]
Cerquone, 2018Tibialis anteriorPAK1Myoblast differentiationDownNegativeImpair[193]
Lee, 2020Tibialis anteriorPHF20Myoblast differentiationUpNegativeImpair[194]
Hayashi, 2016Tibialis anteriorKlf5Myoblast differentiationDownNegativeImpair[3]
Lee, 2015Tibialis anteriorAKAP6Myoblast differentiationDownNegativeImpair[195]
Li, 2020Tibialis anteriorLRTM1Myoblast differentiationDownNegativeImpair[196]
Lin, 2019Left gastrocnemiusMPMMyoblast differentiationDownNegativeImpair[197]
Harada, 2018Tibialis anteriorH3mm7Myoblast differentiationDownNegativeImpair[198]
Tetsuaki, 2009Tibialis anteriorCT-1Myoblast differentiationUpNegativeImpair[199]
Paul, 2012Tibialis anteriorCOPR5Myoblast differentiationDownNegativeImpair[200]
Faralli, 2011Tibialis anterior and gastrocnemius
of one hind limb		Tshz3Myoblast differentiationUpNegativeImpair[201]
Liu N, 2014Tibialis anteriorMEF2A, C and DMyoblast differentiationDownNegativeImpair[202]
Kielbasa OM, 2011Unilateral tibialis anteriorMyosprynMyoblast differentiationUpNegativeImpair[203]
Verpoorten S, 2020Tibialis anteriorCD36Myoblast differentiationDownNegativeImpair[204]
Andrée B, 2002Right gastrocnemius and soleusPopMyoblast differentiationDownNegativeImpair[205]
Paolini A, 2018Tibialis anteriorAutophagyMyoblast differentiationDownNegativeImpair[206]
Clow C, 2010Tibialis anteriorBDNFMyoblast differentiationDownNegativeImpair[207]
Marshall JL, 2012Left quadricepsSSPNMyoblast differentiationDownNegativeImpair[208]
Chen SE, 2005SoleusTNF-αMyoblast differentiationDownNegativeImpair[209]
Ravel, 2014Tibialis anteriorStaufen1Myoblast differentiationUpNegativeImpair[210]
Langsdorf A, 2007Tibialis anteriorSulfsMyoblast differentiationDownNegativeImpair[211]
Liu H, 2011Tibialis anterior and soleusβ3-IntegrinMyoblast differentiationDownNegativeImpair[212]
Watanabe S, 2007Right tibialis anteriorLbx1Myoblast differentiationDownNegativeImpair[213]
Fu D, 2015Tibialis anteriorMdm2Myoblast differentiationDownNegativeImpair[214]
Schroer, 2019Tibialis anteriorRGS12A switch from myoblast proliferation to differentiationDownNegativeImpair[215]
Lim S, 2013Left tibialis anteriorCBR1Myoblast differentiationDownNegative——[216]
Mammen AL, 2009Right tibialis anteriorMi-2Myoblast differentiationUpNegative——[217]
Wu Z, 2020Tibialis anteriorAndrographolideMyoblast differentiation and fusion————Improve[218]
Kurosaka, 2021Left tibialis anteriorSTAT6Myoblast differentiation and fusionDown——Improve[219]
Budai Z, 2021Tibialis anteriorTG2Myoblast fusion and conversion of macrophages from M1 to M2DownNegativeImpair[102]
Vijayakumar A, 2013Unilateral tibialis anteriorIGF-1RMyoblast fusionDownNegativeImpair[220]
Pryce BR, 2017Unilateral tibialis anteriorSLKMyoblast fusionDownNegativeImpair[221]
Redelsperger, 2016One tibialis anteriorSyncytinMyoblast fusionDownNegativeImpair[222]
Kaspar, 2013Tibialis anterior3′ untranslated region of c-MybMyoblast fusionDownNegativeImpair[223]
Griffin, 2016Left tibialis anterior and gastrocnemiusANO5Myoblast fusionDownNegativeImpair[224]
Trapani L, 2012Right tibialis anteriorHMGRMyoblast fusionDownNegativeImpair[225]
Hamoud, 2018Tibialis anteriorBAI3Myoblast fusionDownNegativeImpair[226]
Tamilarasan K P, 2012GastrocnemiusLipid accumulationMyoblast fusionUpNegativeImpair[227]
Yoshida N, 2019Tibialis anterior(P)RRMyoblast fusionUpNegativeImpair[228]
Shibasaki H, 2019Tibialis anteriormiR-188Myoblast fusionUp/DownPositive/NegativeImprove/impair[155]
Youm TH, 2019tibialis anteriorNox4/ROSMyoblast fusion——PositiveImprove[229]
Teng, 2015One tibialis anteriorPLD1Myoblast fusionUpPositiveImprove[230]
Krause MP, 2013Left tibialis anteriorMustn1Myoblast fusionUpPositiveImprove[231]
Kurosaka, 2016Tibialis anteriorTRPV IMyoblast fusionUpPositiveImprove[232]
Singhal N, 2015Gastrocnemius, quadriceps, tibialis anteriorGalgt1Myoblast fusion; SC apoptosisDownNegative; PositiveImpair[233]
Yalvac ME, 2017Left tibialis anterior and left gastrocnemiusCalpain-3Myoblast fusion; fibrosisDownNegative; PositiveImpair[234]
Ogawa, 2015Tibialis anteriorDcxMyofiber maturationDownNegativeImpair[235]
Ohno Y, 2019Right tibialis anteriorLactateMyotube formationUpPositiveImprove[236]
Hoshino S, 2013Right tibialis anteriorCHC22Myofiber maturationUpNegativeImpair[237]
Hu Z, 2010Unilateral tibialis anteriorPTENMyofiber maturation; fibrosisDownPositiveImprove[25]
Agbulut O, 2001Gastrocnemius and soleus or tibialis anteriorDesminMyofiber maturation; neuromuscular junctionsDownNegativeImpair[238]
Cicchillitti, 2012Tibialis anteriormiR-210Myoblast differentiationDown——No effects[239]
Piccioni A, 2014Tibialis anteriorShhActivated SCsUp——Improve[240]
Ceco E, 2021Left tibialis anteriorElevated CO2 exposureMyoblast differentiation and fusion————Impair[30]
Liu, 2017Tibialis anteriorTwist2Maintain SC state——————[241]
Table
Table 4. Fibrosis, calcification, and apoptosis.
Table 4. Fibrosis, calcification, and apoptosis.
Author, YearInjury PortionsTarget Molecule/
Drug		Target ProcessExpressionEffects (Positive/Negative)Regeneration
(Impair/Improve)		Ref.
Heredia JE, 2013Unilateral tibialis anteriorIL-4FAP proliferationDownNegativeImpair[55]
Vumbaca S, 2021Tibialis anterior, quadriceps, and gastrocnemiusIL1a/IL1β and extracellular vesiclesFAP proliferation and differentiationUpPositive——[248]
Zhao L, 2020Tibialis anteriorRA signalingFAP proliferationUppositiveImprove[246]
Zanotti S, 2018Tibialis anteriorExosome miR-199a-5p/CAV1FibrosisUpPositiveImpair[247]
Horii N, 2018Tibialis anteriorC1q/Wnt and resistance trainingFibrosisUppositiveImpair[34]
Murray, 2017Tibialis anteriorαV interginFibrosisDownNegativeImprove[245]
Burks, 2011Tibialis anteriorLosartamFibrosis——NegativeImprove[250]
Stepien DM, 2020Left tibialis anteriorTGF-β1FibrosisDownNegativeImprove[249]
Bosnakovski D, 2022Tibialis anteriora prior DUX4 burstFibrosis——Positive——[251]
Ding, 2016Tibialis anteriorTAR RNA-binding protein (Trbp)Fibrosis; myofiber formationDownPositive; NegativeImpair[244]
Ogasawara S, 2018Left gastrocnemiusCatKFibrosis; inflammation and cell apoptosisDownNegativeImprove[252]
Lee SJ, 2010GastrocnemiusFollistatinFibrosis; myofiber maturationDownNegativeImpair[253]
Rinaldi F, 2016Tibialis anteriorGDF11Collagen depositionUpPositiveNo effects[254]
Mignemi NA, 2017Posterior compartments of the lower extremitiesPlasminCalcificationDownNegativeImpair[255]
Lee YS, 2013Right gastrocnemiusGasp1 and/or Gasp2Calcified fibers and fibrosisDownPositiveImpair[256]
Zhao Y, 2009Tibialis anterior——Dystrophic calcification——————[257]
Lounev V Y, 2009QuadricepsTie2-expressing endothelial precursorsHeterotopic ossification——————[258]
Drouin G, 2019——Hypoxic stateHeterotopic ossification——————[259]
Arsic N, 2004Tibialis anteriorVEGFApoptosisUpNegativeimprove[260]
Sinha-Hikim I, 2007GastrocnemiusJNK and iNOS signalingCell apoptosisDownNegativeimprove[261]
Min K, 2017Gastrocnemius/
Soleus		MKP-5myofiber apoptosisDownNegativeimprove[262]
Tjondrokoesoemo A, 2016Tibialis anteriorserpina3nStabilization of myofiber plasm membraneUpPositiveImprove[263]
Table
Table 5. Angiogenesis and neurogenesis in CTX-induced injury models.
Table 5. Angiogenesis and neurogenesis in CTX-induced injury models.
Author, YearInjury PortionsTarget Molecule/
Drug		Target ProcessExpressionEffects
(Positive/Negative)		Regeneration
(Impair/Improve)		Ref.
Bellamy LM, 2010Unilateral tibialis anteriorAngiotensin IIAngiogenesisDownNegativeImpair[266]
Ieronimakis N, 2012Tibialis anterior, quadriceps, gastrocnemiusBone marrow-derived cellsAngiogenesis——PositiveImprove[265]
Mellows B, 2017Tibialis anteriorExtracellular vesicles-derived from amniotic fluid stem cellAngiogenesis——PositiveImprove[267]
Hosaka Y, 2002Right tibialis anteriorα1-SyntrophinHypertrophy and neuromuscular junctionsDownPositive; NegativeImprove[268]
Daneshvar N, 2020Left tibialis anteriorPremature satellite cell activationMaturation of neuromuscular junctionsUpNegativeImprove[269]
Kurosaka M, 2021Tibialis anteriorM2 macrophageMotor innervation regeneration————Improve[219]
Sawano S, 2014Tibialis anteriorM2 macrophageMotor innervation regeneration————Improve[270]
Randazzo D, 2019Unilaterally tibialis anteriorTubb6Microtubule organizationUpNegative——[15]
Table
Table 6. Single gene in the CTX-induced skeletal muscle injury model.
Table 6. Single gene in the CTX-induced skeletal muscle injury model.
Author, YearInjury PortionsTarget MoleculeExpressionRegeneration
(Impair/Improve)		Ref.
Kim DS, 2015Left tibialis anteriorTLR2DownImprove[272]
Oikawa S, 2019Tibialis anteriorDicerDownImpair[273]
Hiramuki Y, 2015Tibialis anteriorMestDownImpair[274]
Norton CR, 2013Tibialis anteriorSnai1/Snai3DownNo effects[275]
Call JA, 2017Left tibialis anterior and left flexor digitorum longusUlk1Down——[276]
Chaturvedi N, 2020Left gastrocnemiusS100A1Down——[277]
Parks CA, 2019Left tibialis anteriorTrim33DownNo effects[278]
Goetsch SC, 2005GastrocnemiusFilamin CUp——[279]
Wardrop KE, 2011Left tibialis anteriorLYVE-1Down——[280]
Merkulova T, 2000Extensor digitorum longus and tibialis anteriorβ enolaseUp——[281]
Yuasa K, 2008Tibialis anteriormiR-206Up——[282]
Casciola-Rosen L, 2012Right anterior tibiliasAldolase AUp——[283]
Mammen AL, 2011Right tibialis anteriorUFD2aUp——[284]
Nakamura K, 2010Right gastrocnemiusGNEUp——[285]
Sato Y, 2013Left gastrocnemiusSema3AUp——[286]
Garry, 2000Hind limbsMNFDownImpair[287]
Kemp MW, 2009Tibialis anteriorSyncoilinUp——[288]
Miura P, 2005Right tibialis anteriorUtrophin AUp——[289]
Wang Q, 2022Tibialis anteriorTsukushiDownImpair[290]
McCullagh KJ, 2008Unilateral tibialis anteriorSyncoilinDownNo effects[291]
Demonbreun AR, 2010Quadriceps or gastrocnemius/soleusFerlinUp——[292]
Maeda Y, 2017Right tibialis anteriorCXCL12UpImprove[293]
Darabi R, 2008Tibialis anteriorPax3UpImprove[294]
Di Rocco A, 2015Tibialis anteriorRARγDownImpair[295]
Bryer SC, 2007Extensor digitorum longus and tibialis anterioruPAR——No effects[296]
Bryan BA, 2005Tibialis anteriorGEFTUpImprove[297]
Mathes AL, 2011Tibialis anteriorTLR-3DownImpair[298]
Wu G, 2010Forelimb leg muscleChkbDownNo effects[299]
Yan Z, 2003Tibialis anteriorE2f1DownImpair[5]
Fujita R, 2014Left tibialis anteriorIL-6RDownImprove[300]
Wu G, 2009GastrocnemiusChkbDownImpair[301]
Wada E, 2019Tibialis anteriorEmerin and lamin A/CDownImpair[302]
Mofarrahi M, 2015Unilateral tibialis anteriorAng-1UpImprove[303]
Gattazzo F, 2014Tibialis anteriorCyclosporin AUpImprove[304]
Kim MH, 2011Unilaterally tibialis anteriorAktUpImprove[243]
Laziz I, 2007SoleusSpryDown——[305]
Armand AS, 2003Unilateral soleusFollistatin and myostatinUp/DownImprove[306]
Li C, 2013Right gastrocnemiusProsaposinUp——[307]
Table
Table 7. Non-SCs in the CTX-induced skeletal muscle injury models.
Table 7. Non-SCs in the CTX-induced skeletal muscle injury models.
Author, YearInjury PortionsTarget Molecule/DrugRegenerationRef
Kano K, 2020GastrocnemiusCapillary stem cellsImprove[315]
Liu Y, 2007Unilateral tibialis anteriorFlk-1+ AD-MSCsImprove[316]
Kim JA, 2013Left tibialis anteriorhAFS cells transfected with MyoDImprove[318]
Xuan W, 2021Tibialis anteriorPluripotent stem cells-induced skeletal muscle progenitor cells with givinostatImprove[320]
Naldaiz-Gastesi N, 2019Tibialis anteriorHuman cremaster muscle-derived precursor cellsImprove[321]
Mori J, 2008Tibialis anteriorCD45+: Sca-1+ hematopoietic stem cellsImprove[312]
Abedi M, 2007Tibialis anteriorHematopoietic stem cellsImprove[322]
Hwang Y, 2014Tibialis anteriorHuman embryonic stem cellsImprove[323]
Piccoli M, 2012Tibialis anterior mice transplanted with bone marrow or amniotic fluid stem cellsAmniotic fluid stem cellsImprove[324]
Yang R, 2010Right tibialis anteriorClones of ectopic stem cellsImprove[325]
Rousseau J, 2010EDLMuscle precursor cellsImprove[313]
Gang EJ, 2009Tibialis anteriorMesenchymal stem cellsImprove[314]
Jung JE, 2017Gastrocnemius and masseterPulp-derived cellImprove[310]
de la Garza-Rodea AS, 2011Tibialis anteriorBM-hMSCsimprove[311]
Bossolasco, 2004Tibialis anteriorHuman adult BMImprove[326]
Ma, 2012Tibialis anteriorHuman AF-amniotic fluid stem cellsImprove[317]
Fukada S, 2002Tibialis anteriorBone marrow and fetal liver cellsImprove[327]
Luth ES, 2008Tibialis anterior, quadriceps, and gastrocnemiusBone marrow side population cellsImprove[328]
Zheng JK, 2006Tibialis anteriorHuman embryonic stem cellsImprove[329]
Cížková D, 2011Right tibialis anteriorBMCsImprove[308]
Meeson AP, 2004HindlimbsSkeletal muscle side populationImprove[330]
Drapeau C, 2010Right tibialis anteriorMobilization of bone marrow stem cellsImprove[319]
Kowalski K, 2016GastrocnemiusSdf-1 and granulocyte-colony stimulating factorImprove[331]
Mitchell R, 2019Right tibialis anteriorADSC secretomeImprove[332]
Tobin S, 2021Tibialis anterior; quadriceps; gastrocnemiusYoung/aging macrophagesImprove/impair[333]
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Wang, Y.; Lu, J.; Liu, Y. Skeletal Muscle Regeneration in Cardiotoxin-Induced Muscle Injury Models. Int. J. Mol. Sci. 2022, 23, 13380. https://doi.org/10.3390/ijms232113380

AMA Style

Wang Y, Lu J, Liu Y. Skeletal Muscle Regeneration in Cardiotoxin-Induced Muscle Injury Models. International Journal of Molecular Sciences. 2022; 23(21):13380. https://doi.org/10.3390/ijms232113380

Chicago/Turabian Style

Wang, Yanjie, Jianqiang Lu, and Yujian Liu. 2022. "Skeletal Muscle Regeneration in Cardiotoxin-Induced Muscle Injury Models" International Journal of Molecular Sciences 23, no. 21: 13380. https://doi.org/10.3390/ijms232113380

APA Style

Wang, Y., Lu, J., & Liu, Y. (2022). Skeletal Muscle Regeneration in Cardiotoxin-Induced Muscle Injury Models. International Journal of Molecular Sciences, 23(21), 13380. https://doi.org/10.3390/ijms232113380

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