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Myostatin Is a Negative Regulator of the Muscle Mass
Myostatin is an autocrine and paracrine hormone produced by muscle cells that inhibits muscle differentiation and growth. Knockout mice without myostatin and certain breeds of cattle (Belgian Blue and Piedmontese) that lack effective myostatin are “double muscled.” Because myostatin also targets adipocytes, these animals also lack adipogenesis (formation of fat) and so they are lean as well. A report was published about a young boy who had a myostatin mutation that is associated with gross muscle hypertrophy (New England Journal of Medicine 350:2682–2688, 2004). Its mode of action is illustrated in Figure 3.7.7.
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Myostatin is a secreted protein that negatively regulates skeletal muscle mass determining both muscle fiber number and size. The myostatin pathway is conserved and regulates muscle mass in a number of animal species ranging from fish to humans. Inhibition of myostatin using a variety of therapeutic approaches can increase muscle mass in a number of animal models of human disease, including muscular dystrophy.
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Myostatin, or growth and differentiation factor-8 (GDF-8), was cloned in 1997 by its homology to members of the transforming growth factor-β (TGF-β) superfamily [1]. Over the last seven years, myostatin has been implicated in the regulation of skeletal muscle mass and has emerged as a novel therapeutic target for treatment of a number of muscle degenerative diseases, including sarcopenia (see Glossary) and muscular dystrophy. It is now clear that inhibition of myostatin activity can lead to increased skeletal muscle mass in adult mammals. Thus, myostatin serves as a negative regulator of skeletal mass and acts in part by inhibiting the activation of muscle satellite cells. In this review we highlight some of the advances in this field over the past two years and focus on the validation of myostatin as a therapeutic target, the expression and regulation of myostatin activity, the identification of the first human example of myostatin deficiency, and the clinical application of myostatin inhibitors.
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8 Surrogates for gene replacement using a muscle building strategy
Blocking the myostatin pathway shows dramatic muscle building properties that are highly conserved across species, including mice, sheep, cattle, canines, and humans [24,25,34,39]. Myostatin is a member of the transforming growth factor (TGF)-β superfamily and numerous studies have demonstrated that it is a negative regulator of skeletal muscle growth. An antibody to myostatin (MYO-29) was not beneficial to patients in a clinical trial [42]. Follistatin is a powerful inhibitor of myostatin plus having the advantage of controlling muscle mass through pathways independent of the myostatin signaling cascade [21]. In studies of the myostatin knockout mice crossed to mice carrying a follistatin transgene there was a quadrupling of muscle mass compared with the doubling of muscle mass that is observed from lack of myostatin alone.
In preparation for taking this to clinical trial, we identified an alternatively spliced isoform of follistatin (FS344) that can be packaged in AAV1. Upon intramuscular injection, FS344 undergoes post-translational modification and generates a stable, FS315 isoform with minimal toxicity. In normal C57Bl/6 mice and in the mdx mouse, intramuscular injections of AAV1 carrying the FS344 gene increased muscle mass (histological measure) and quantitative strength, as assessed by hind- and forelimb grip and measures of force generation and protection against eccentric contractions [12]. The benefits persist for more than two years, and in older mice not treated until 210 days of age, muscle strength increased for 60 days post gene delivery and persisted long-term, throughout the 560 days of evaluation. Study of the muscle and multiple organs from these mice revealed no significant abnormalities. In addition, we have tested the safety and efficacy of AAV1.CMV.FS344 in non-human primates showing no organ system toxicity for as long as 15 months and an ability to increase quadriceps muscle size and strength following direct muscle injection [19]. This occurred without adverse effects. Necropsies showed no abnormalities in heart, lungs, liver, kidneys, or gonads. We also monitored clinical chemistries and hormone levels related to gonadal function including follicle stimulating hormone, luteinizing hormone, testosterone, and estradiol. No abnormalities were encountered.
We have now taken this to clinical trial in two patient groups who could specifically benefit by increasing size and strength of the quadriceps muscle similar to our proof of principle studies in mdx mice and cynomolgus macaques (http://clinicaltrials.gov/). In Becker muscular dystrophy and in sporadic inclusion body myositis the quadriceps muscle is targeted by the disease process leading to frequent falls and ultimately loss of ambulation. The gene delivery protocol approved by the FDA calls for injection of 12 sites in the quadriceps muscle divided evenly between rectus femoris, vastus medialis, and vastus lateralis. Vector administration includes a low dose cohort receiving intramuscular injections on one side only (2 × 1011 vg/kg); intermediate dosing (6 × 1011 vg divided between extremities) and high level dosing (1.2 × 1012 vg/kg divided between extremities). Three patients in the low dose cohort have been injected at the time of this writing. No adverse effects have been encountered. Muscle biopsies will be done at 6 months in all patients to delineate fiber size. MRI of the quadriceps muscle will also be used to assess muscle volume. Additional testing includes the 6 min walk test, climbing 4 standard steps, timed to get up from a chair, and strength of knee extensors using quantitative myometry.
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4 Discussion
To our knowledge, this is the first meta-analysis investigating the effects of resistance training on myostatin and follistatin. We show that resistance training is effective for reducing myostatin and increasing follistatin, and these changes may contribute to beneficial effects on muscle strength and mass regardless of age.
Myostatin is one of the most well-known proteins which binds to the ActRIIA/B complex for controlling muscle atrophy [4]. Furthermore, it is well-established that myostatin mRNA expression is higher in sarcopenic adults compared with non-sarcopenic adults, and can be a major contributor of sarcopenia [64]. Despite the important role of myostatin as a negative regulator of muscle mass, evidence regarding the relationship between ages and circulating myostatin is sparse and unclear. Reports of circulating myostatin suggest it can either increase, decrease, or remain unchanged with aging [65–69]. For example, a cross-sectional study showed that serum myostatin increased with advancing age, and was inversely correlated with muscle mass [69]. Yet, higher concentrations of myostatin in younger as compared with older men were reported in another study, whereas older, including sarcopenic women had higher myostatin concentrations than younger women [65]. One other cross-sectional study indicated that low serum myostatin was associated with low skeletal muscle in men, but not in women [70]. Despite the clinical importance of myostatin signaling, further studies are needed to clarify the role of circulating myokines with aging. This protein is also involved in various conditions including diabetes, insulin resistance, inflammation, myocardial infarction, muscle wasting, and cancer cachexia [71,72]. The mechanism by which myostatin is reduced following resistance training is likely to be due to reduced production, synthesis and secretion from skeletal muscle, plus reduced stability and enhanced reuptake from circulation [13]. Resistance training increases muscle mass and strength, and prevents sarcopenia, linked to lower myostatin [73,74]. Myostatin is also an important regulator of insulin resistance [75,76] for which an improvement could occur with a reduction in myostatin.
Follistatin is an important plasma monomeric glycoprotein expressed in tissues such as the liver [77], which binds to activins to combat muscle wasting [78], and acts as a negative regulator of myostatin, increasing skeletal muscle hypertrophy [77]. It also appears to be a downstream target for testosterone function [79] and opposes the inhibition of MHC II expression induced by TGF-β [80]. Follistatin plays a pivotal role in controlling muscle mass and function and could be a therapeutic target on skeletal muscle wasting conditions such as age-related muscle degeneration [81,82]. The primary site of exercise-induced follistatin secretion is uncertain [31,83–86], but hepatic production of follistatin have been linked to changes in glucagon and insulin [84]. In addition to the effects of follistatin on myostatin and muscle mass regulation, its action on members of the TGF-β superfamily ligands to ameliorate insulin actions could be effective in reducing muscle wasting caused induced by insulin resistance [87]. We performed a Pearson's correlation to test the relationship between the changes in myostatin and follistatin. In agreement with independent effects for myostatin and follistatin, a negative correlation was observed; however, the correlation was not statistically significant (r = −0.23, p = 0.30). It should be noted that we were only able to use mean data as we did not have access to raw data.
Sarcopenia, prevalent among older adults, is a condition in which muscle mass and function decrease, and are linked to are inactivity and imbalance between myostatin and follistatin [88]. By changing this imbalance in favor of increasing follistatin, resistance training could be a therapeutic method for preventing and treating sarcopenia [89]. Our subgroup analysis showed that resistance training is effective for reducing myostatin and increasing follistatin regardless of age. This finding is clinically important and may further indicate the beneficial effects of resistance training by increasing muscle mass and strength as well as improving metabolic outcomes. However, there are several potential moderators such as participant characteristics including age, muscle mass, and health status, as well as exercise characteristics including intensity and mode that may influence myostatin and follistatin changes following resistance training [15]. In this regard, our results showed a larger effect size in subgroup analysis of the younger versus older subgroups. It is possible that the magnitude of resistance used in strength training interventions and lower muscle mass might explain the smaller effect size in older adults. Furthermore, it should be noted that the differences in timing of the myostatin sampling, plus laboratory measurement techniques and methodologies may have resulted in heterogeneity in results, making the potential of myostatin as a biomarker of muscle wasting less certain [15]. In this regard, among the studies included in the meta-analysis, Willoughby and et al. [36] reported significant increases in myostatin following resistance training. However, blood samples were taken immediately after the last exercise session which may have confounded the results.
The current systematic review and meta-analysis had several limitations that should be considered when interpreting the results. There was significant heterogeneity among studies which may be due to differences in participant ages, BMIs, health statuses of participants, and resistance training characteristics. Significant publication bias was assessed by Egger's tests. Finally, the number of subjects in the included was small.
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Muscle as a therapeutic target in treatment of bone disorders
Targeting muscle-fat-bone communication axis could also be a potential therapeutic strategy, as evidence is accumulating that myokines are able to influence bone metabolism.
As previously discussed, myostatin is a myokine down-regulated following exercise and is negatively correlated to bone health. Systemic administration of a myostatin antibody has been shown to protect diet-induced obese rats from the bone loss and microarchitectural deterioration associated with a high-fat diet, and reduce rheumatoid arthritis associated bone loss in a mouse model [38,196]. While in young adult mice a soluble myostatin decoy receptor increased bone mass [197], although this effect was not seen in aged mice [198]. More study is required in this area.
Very limited amount of studies have explored the effect of irisin treatment in bone, although systemic administration of irisin has been known to have a positive effect on bone [39,40], and it increased endothelial progenitor cell function in diabetic mice [199], suggesting it may be a future drugable target.
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As a member of the TGFβ family, myostatin is expected to exert its effects through a transmembrane serine/threonine kinase heterotetramer receptor family, activation of which enhances receptor transphosphorylation, leading to stimulation of serine/threonine kinase activity. Lee and McPherron [14] established that an activin receptor, ActRIIB, was able to mediate the influence of myostatin in vivo, as expression of a dominant-negative form of ActRIIB (dnActRIIB) in mice mimics myostatin gene knockout (Figure 1).
Figure 1. Putative elements of the myostatin pathway. Myostatin can be found in serum or locally in an inactive state when bound to FLRG, GASP-1, hSGT, T-cap, follistatin or the myostatin propeptide. The active myostatin dimer binds to the activine type II receptor (ActRIIB), which then recruits and activates by transphosphorylation the type I receptor (ALK4 or ALK5). Smad2 and Smad3 are subsequently activated: they form aggregates with Smad4 and are then translocated to the nucleus, activating target gene transcription. Two inhibitors of this signalisation have been identified: Smad7 and Smurf1. Smad7 represses myostatin signal by the binding of its MH2 domain to activated receptors, thus preventing recruitment and activation of R-Smads. Smurf1 is an E3 ubiquitin ligase that mediates ubiquitination and consequent degradation of the R-Smads. FLRG, follistatin-related gene; GASP1, growth and differentiation factor-associated serum protein-1; hSGT, human small glutamin-rich tetratricopeptide repeat-containing protein Reproduced with permission of [8].
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1 Introduction
Skeletal muscle mass and strength are important for healthy aging, and given long lifespans and sedentary lifestyles, targets for prevention. Myokines are cytokines synthesized by myocytes and released during muscular contraction which play important roles in regulating muscle function [1,2]. Myostatin is a member of the transforming growth factor-β family, that has a negative role on skeletal muscle mass by suppressing cellular growth and differentiation [3]. It binds to activin receptor types IIA and IIB (ActRIIA and ActRIIB) to reduce the activity of the protein kinase B (Akt) pathway and increase the ubiquitin proteasome system that mediates anabolic and catabolic responses [4]. Follistatin is a myostatin antagonist, that inhibits several members of the transforming growth factor (TGF-β) superfamily and promotes skeletal muscle mass development [5,6], by inhibiting myostatin dependent pathways [7,8] and decapentaplegic homolog 4 (Smad3), as well as mammalian target of rapamycin mTOR signaling independently of myostatin [9]. Elevated follistatin is also associated with increased risks of type 2 diabetes mellitus (T2DM) [10]. Several studies have reported the presence of myostatin and follistatin proteins in blood [11–14], and some have suggested that circulating myostatin and follistatin might be related to skeletal muscle mass regulation and whole body metabolism, suggesting that blood concentrations of these proteins may be important markers of muscle wasting and metabolism [10,11,15–18].
Resistance training is an effective exercise that improves muscular fitness and other health outcomes across a wide range of participants [19–21]. It enhances muscle mass and strength, and can prevent and treat sarcopenia [22,23], improving some markers of cardiometabolic health including inflammation, body fat, lipid profiles, blood pressure, and glycemia [24–28]. The benefits of resistance training on muscle strength are partly mediated by proteins/hormones produced from skeletal muscle or related tissues during exercise [29,30]. Myostatin is a myokine that acts on muscle cells to inhibit muscle hypertrophy and myogenesis [30]. Follistatin is an inhibitor of myostatin, thought to be a contraction-induced hepatokine [31]. Given the important roles of myostatin and follistatin in regulating muscle mass, resistance training induced changes, might play a role in exercise induced improvements in muscle mass [13]. However, the reported effects of resistance training on myostatin and follistatin have been mixed [32–36], that currently no meta-analysis has examined, which was the aim of our study.
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3.7 Additional contributors to muscle atrophy
A number of other findings related to the molecular features of muscle atrophy require mention, although how they fully integrate with the aspects described above remains to be established.
Myostatin (growth and differentiation factor 8, GDF8) is a member of the transforming growth factor beta (TGF-β) family of growth factors/cytokines, and is a secreted ligand that acts as a negative regulator of muscle growth. Genetic deletion of myostatin causes hypermuscularity primarily through myofiber hyperplasia (increased myofiber number), and to a lesser extent hypertrophy (increased myofiber size) (McPherron et al., 1997). Myostatin acts as a dimer on type II TGF-β receptor kinases, leading to intracellular phosphorylation of the transcription factors SMAD2 and SMAD3 which together translocate to the nucleus to regulate gene expression, as well as through activation of MAPK signaling (MacDonald and Cohn, 2012). Its overexpression in muscle causes severe muscle atrophy, and its action on SMAD3 may act synergistically with FOXO transcription factors (Gomis et al., 2006; Seoane et al., 2004) to induce expression of Trim63 and Fbxo32, although it has also been suggested that SMAD2 and SMAD3 may induce atrophy via mechanisms independent of TRIM63. Similar to its regulation of FOXO transcription factors, AKT is capable of phosphorylating SMAD3, leading to its cytoplasmic sequestration (Hoogaars and Jaspers, 2018). Myostatin is also thought to be capable of antagonizing AKT signaling (McFarlane et al., 2006; Morissette et al., 2009; Trendelenburg et al., 2009), which may contribute to FOXO-dependent induction of atrogenes (McFarlane et al., 2006; Morissette et al., 2009). Despite these apparent intersections with intracellular signaling pathways known to mediate atrophy, however, myostatin inhibition via interval intraperitoneal delivery of soluble ActRIIB (activin receptor IIB), a myostatin receptor fused to an Fc domain that inhibits myostatin signaling, did not protect against atrophy of TA muscle at 3 weeks post-denervation in a sciatic nerve resection mouse model (MacDonald et al., 2014).
TWEAK (TNF-like weak inducer of apoptosis, TNFS12) is a transmembrane cytokine and member of the TNF superfamily that is cleaved intracellularly to a soluble form capable of extracellular release and activation of the Fn14 (Fibroblast growth factor-inducible 14, TNFRSF12A) receptor (Winkles, 2008). Fn14 levels are upregulated in response to numerous causes of tissue injury including muscle denervation, and TWEAK-Fn14 engagement may activate several intracellular signaling cascades, most prominently NF-κB signaling (Winkles, 2008). ERK, JNK, MAPK and PI3K/AKT signaling pathways have also been described as downstream targets of TWEAK-Fn14 engagement (Pascoe et al., 2020; Winkles, 2008). Differentiated C2C12 myotubes treated with soluble TWEAK showed reduced mean diameter compared to untreated cultures, which corresponded with increased expression of Trim63 and Fbxo32 as well as ubiquitination and loss of myosin heavy chain fast type (MyHCf) (Dogra et al., 2007). TWEAK was shown to decrease AKT phosphorylation and activation, with a corresponding decrease in phosphorylation of downstream targets including FOXO1a, and the TWEAK-induced degradation of MyHCf was shown to be dependent on NF-KB activation (Dogra et al., 2007). Finally, chronic in vivo administration of TWEAK caused atrophy of mouse TA and soleus muscle, with evidence of increased NF-κB activity, and mice with muscle-specific transgenic overexpression of TWEAK showed significantly reduced hindlimb muscle sizes and perinatal or neonatal lethality (Dogra et al., 2007). The role of TWEAK in neurogenic atrophy was investigated using conventional TWEAK-KO mice, in which TA and soleus muscle showed partial protection from atrophy at 10 days post-sciatic nerve resection (Mittal et al., 2010). The same muscles from transgenic mice with skeletal muscle-specific overexpression of TWEAK (TWEAK-Tg, derived from founders different from the line described by Dogra et al. (2007) and capable of surviving to adulthood) showed significantly increased atrophy. These changes were shown to result from increased NF-κB-dependent induction of Trim63, but not Fbxo32, in TWEAK-Tg mice, with no apparent effect on AKT signaling. Furthermore, TA and soleus muscle from wild type mice treated with a neutralizing antibody against TWEAK showed partial protection from atrophy at 14 days post-denervation (Mittal et al., 2010). TWEAK immunodepletion or genetic ablation of Tweak in the SOD1G93A mouse model of ALS provided partial protection against TA muscle atrophy (no comparison to wild type mice was made), but did not prolong survival (Bowerman et al., 2015).
Expression of microRNAs and other small noncoding RNAs that may confer posttranscriptional regulation has been examined in denervated mouse models and muscle tissue from patients with ALS (Di Pietro et al., 2017; Kovanda et al., 2018; Williams et al., 2009). miR-206 is a skeletal muscle-specific microRNA that is increased in TA muscle during the symptomatic stage of disease in the SOD1G93A mouse model of ALS, and in EDL, TA, and gastrocnemius/plantaris muscle of wild type mice subjected to sciatic nerve resection (Williams et al., 2009). It is also induced in muscle from patients with early-stage ALS (Di Pietro et al., 2017). Genetic ablation of miR-206 was associated with accelerated muscle atrophy and decreased survival in the SOD1G93A mouse model of ALS (Williams et al., 2009). Furthermore, miR-206 knockout mice subjected to sciatic nerve transection or crush showed delayed reinnervation of neuromuscular junctions (NMJs), with mutant NMJs displaying only partial reoccupation by regenerated nerves, disaggregation of presynaptic vesicles in regenerating nerve terminals, and abnormal motor axon sprouting beyond the NMJ interface (Williams et al., 2009). These observations are consistent with a role for miR-206 as a regulator of other muscle-derived factor(s)/signal(s) that mediate reinnervation. HDAC4, previously described to have a role in inducing myogenin expression (Cohen et al., 2007) and subsequently shown to contribute to muscle atrophy (Moresi et al., 2010), was then computationally predicted and demonstrated to be translationally repressed by miR-206 (Williams et al., 2009). Intriguingly, mice with muscle-specific deletion of HDAC4 showed enhanced NMJ reinnervation after sciatic nerve crush or transection, an effect that was proposed to occur through disinhibition of expression of the secreted FGF binding protein FGFBP1, which would typically be suppressed by HDAC4 (Da Cruz et al., 2012). FGFBP1 interacts with and appears to potentiate the effects of the muscle-derived factors FGF7, FGF10, and FGF22 (Williams et al., 2009), known to promote presynaptic differentiation (Fox et al., 2007). Direct injection of miR-206 into gastrocnemius of a rat model of sciatic nerve transection showed partial protection against atrophy, as reflected in increased myofiber cross-sectional area compared to saline-injected controls, although NMJ formation was not evaluated (Huang et al., 2016). Soares et al. also found that miRNA-206 (along with miRNA-21) are two of the most significantly induced microRNAs in gastrocnemius muscle from a mouse model of sciatic nerve transection (Soares et al., 2014). However, they found that transfection of miRNA-206 in denervated mouse TA muscle exacerbated atrophy at 10 days post-transfection (Soares et al., 2014). These discrepancies again likely raise awareness that caution in choice of experimental models, and having multiple models where possible, is important for drawing inferences.
Other differentially expressed microRNAs have also been described in denervated muscle. Skeletal muscle miR-351 expression is reduced in a mouse model of sciatic nerve transection, and repeated direct injection of a miR-351 agomir partially reduced TA muscle atrophy in this model (He et al., 2016). miR-351 was proposed to downregulate TRAF6 (see section 3.1) by targeting its 3’-UTR, thereby also negatively regulating its downstream effectors, Fbxo32 and Trim63 (He et al., 2016). An independent miRNA array analysis of denervated gastrocnemius muscle from a rat sciatic nerve resection model identified 15 miRNAs that were differentially expressed at 5 days post-injury, of which miR-29b was also confirmed to be elevated in atrophying muscle from mouse models of dexamethasone treatment, fasting, cancer cachexia, and aging (Li et al., 2017). Interestingly, IGF1 and PI3K were predicted and confirmed to be direct targets of miR-29b, as determined by a luciferase reporter assay employing the 3’-UTRs of these genes as well as transfection of a miR-29b mimic into C2C12 myotubes. C2C12 myotubes transfected with a miR-29b mimic showed reduced site-specific phosphorylation of AKT, FBXO3A, mTOR, and P70S6K, as well as increased expression of Trim63 and Fbxo32. Furthermore, direct injection of a miR-29b agomir into mouse gastrocnemius caused myofiber atrophy and induction of some atrogenes including Trim63 and Fbxo32. Intramuscular injection of a miR-29b “sponge” into mouse gastrocnemius followed by sciatic nerve resection yielded attenuated atrophy at 5-days post-transection (Li et al., 2017).
The skeletal muscle environment is comprised of many cell types in addition to myocytes, including muscle satellite cells, fibroblasts, PW1-expressing interstitial cells, vessel-associated cells, immune cells, and fibro-adipogenic progenitors (FAPs) (Farup et al., 2015; Mitchell et al., 2010; Pannerec et al., 2012; Uezumi et al., 2010; Uezumi et al., 2014). An understanding of the contribution of non-muscle cells to neurogenic atrophy is still in early stages. A recent study suggested that denervation induces proliferation of fibro-adipogenic progenitors (FAPs) and STAT3 activation in these cells, leading to production of IL6 (interleukin-6), which appeared to act in a paracrine manner to promote myofiber atrophy (Madaro et al., 2018). FAP proliferation was observed in TA muscle from mice subjected to sciatic nerve transection, as well as in muscle from the SMAΔ7 mouse model of spinal muscular atrophy, muscle from the SODG93A mouse model of ALS, and in human muscle biopsies from patients with ALS. This proliferation of FAPs appeared to be dependent on loss of muscle innervation, as similar changes were not observed in immobilization or disuse models of muscle atrophy or in aged muscle. Pharmacologic STAT3 inhibition or IL6 blockade using neutralizing antibodies partially reduced TA atrophy in denervated mice, and intriguingly transplantation of FAPs from denervated mice was sufficient to promote atrophy in TA muscle of wild type mice (Madaro et al., 2018). Further confirmation of these findings may provide a rationale to consider targeting IL6-STAT3 signaling as a means to reduce atrophy and fibrosis in denervating disease.
Skeletal muscle satellite cells have the capacity to proliferate and differentiate into myofibers in a variety of settings of muscle injury (Dumont et al., 2015), but their role in atrophy and hypertrophy remains controversial (Bamman et al., 2018; Pallafacchina et al., 2013). Madaro et al. did not observe a significant change in satellite cell content in atrophying mouse TA muscle at 3, 7, or 15 days post-denervation, although embryonic myosin heavy chain (eMyHC)-expressing myofibers were observed at the 7 day timepoint (Madaro et al., 2018). In contrast, a modest but significant increase in satellite cell content was described in TA muscle from a mouse model of sciatic nerve transection at 6 weeks post-denervation (Liu et al., 2015). Both of these observations appear consistent with earlier ultrastructural analyses of rodent soleus and/or EDL muscles examined 3 days to 25 months after sciatic nerve resection, wherein satellite cell content significantly increased in the first few weeks after denervation followed by a progressive decline (Borisov et al., 2005; Dedkov et al., 2001; Snow, 1983; Viguie et al., 1997). The role and fate of satellite cells residing in atrophying muscle that account for these population changes has also been controversial. In vivo time-lapse imaging of mouse EDL and soleus myofibers denervated by sciatic nerve resection showed no loss or accretion of myonuclei over the 4-week interval examined, but many satellite cell nuclei were reported to be TUNEL-positive suggesting apoptosis (Bruusgaard and Gundersen, 2008). Borisov et al. observed that approximately 20-25% of the myofiber population of rat TA and EDL muscles consisted of very small fibers at 10-18 months post-sciatic nerve transection, which were presumed to represent nascent satellite cell-derived myofibers and a reflection of abortive myogenesis, resulting in satellite cell “exhaustion” or depletion (Borisov et al., 2005).
Intriguingly, a recent study showed that muscle satellite cells become activated in denervated muscle and contribute to reinnervation of NMJs (Liu et al., 2015). Compared to controls, diphtheria toxin A (DTA)-activated depletion of satellite cells in a mouse model of sciatic nerve transection was associated with reduced TA myofiber size, increased fibrosis, increased fiber-type transitions, and reduced force generation capacity at 6 weeks post-injury. Lineage tracing of genetically GFP-labeled satellite cells suggested increased accretion within the vicinity of NMJs of TA and EDL muscles at 4 weeks post-sciatic nerve transection. Finally, compared to controls, DTA depletion of satellite cells was associated with a significantly higher proportion of partially or fully denervated NMJs in TA muscles at 6 weeks post-sciatic nerve transection. Altogether, these observations suggest that satellite cells contribute to muscle maintenance after denervation as well as to NMJ regeneration (Liu et al., 2015). Follow-up studies exploring the role of age-related NMJ remodeling and muscle atrophy (sarcopenia) found a significant decline in EDL muscle satellite cell content in the vicinity of NMJs of aged (18 and 24 month-old) mice. DTA depletion of satellite cells was associated with significantly reduced subsynaptic myonuclear number, exacerbated deterioration of NMJs (though not robust denervation), and myofiber atrophy in 12-month-old mice compared to controls, suggesting that satellite cell accretion is required for NMJ maintenance (Liu et al., 2017). Confirmation of these findings and establishing the molecular basis for this apparent requirement will be of interest.
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3.6 Peptides and other agents regulating irisin levels
Other metabolic pathways and peptides have also been reported to alter irisin circulating levels in both healthy and unhealthy subjects.
Myostatin is a myokine that inhibits myogenesis and is overexpressed in patients with obesity and DM [92]. A study on high-fat diet-induced obese mice showed that administration of anti-myostatin peptide resulted in improved glucose tolerance, insulin resistance and enhanced muscle growth [93]. Furthermore, inhibition of myostatin action stimulated the conversion of WAT into BAT through activating PPAR-γ and increasing irisin secretion. As a result, targeting myostatin arises as a potential therapeutic target to counteract insulin resistance and obesity via irisin pathway. Another study replicated this effect both in-vitro and in-vivo [94]. Loss of myostatin in both myoblasts cell lines and mice resulted in increased expression of FNDC5 and irisin and facilitated the irisin browning effect on WAT. Betatrophin is another peptide which was found to increase irisin levels in healthy individuals while it had no effect in patients with T2DM [98]. Irisin levels were found to be correlated with another peptide, urotensin II, in both in diabetic patients and patients with preeclampsia [101,102]. Oxytocin and irisin were also found to be correlated in young athletes which suggested a possible role for oxytocin in energy regulation and metabolism [108]. Interestingly, irisin administration was also found to increase its own expression in subcutaneous adipose tissues [83].
A correlation was also found between irisin and brain-derived neurotrophic factor (BDNF) in athletes [106]. In fact, this study concluded that both irisin and BDNF levels were elevated in athletic individuals compared to sedentary subjects which was responsible for the improved cognitive performance in athletes. Moreover, many other peptides and proteins were capable of regulating irisin levels. These peptides include follistatin, tumor growth factor β, plasma fatty acids and serum cytokines [116,118,105,104] (Table 4).
Table 4. Endogenous peptides and hormones and their effects on irisin levels.
Hormone/peptide Effect Reference Lower thyroxine Decrease [97] TSH No effect [61] Betatrophin (in patients with normal glucose tolerance) Increase [98] Betatrophin (in patients with T2DM) No effect [98] Myostatin inhibition Increase [99, 94] Adiponectin Decrease [100] Urotensin II No effect [101] Urotensin II Decrease [102] Sclerostin Increase [103] Serum cytokines Increase [104] Plasma Fatty acids Increase [105] Brain-derived neurotrophic factor Increase [106] Immunomodulatory and neuroprotective peptides Increase [107] Oxytocin Increase [108] Follistatin Increase [116] Irisin Increase [83] Myostatin Decrease [117] Tumor growth factor β (TGF-β) Decrease [118]
Rho-kinase-1 (ROCK1) is a kinase that is involved in various cellular events such as secretion, gene expression, proliferation and motility [95]. Activation of ROCK1 caused a reduction in UCP1 expression in both WAT and BAT and also reduced the circulating irisin concentration [96]. Upon inhibiting ROCK1 pathway, irisin levels were restored and caused improvements in glucose homeostasis and insulin resistance.
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