The Scientific World Journal

Volume 2013 (2013), Article ID 593267, 9 pages

http://dx.doi.org/10.1155/2013/593267

Review Article

Mitochondria as a Potential Regulator of Myogenesis

Akira Wagatsuma¹ and Kunihiro Sakuma²

¹Graduate School of Information Science and Technology, The University of Tokyo, 7-3-1 Hongo, Bunkyo-ku, Tokyo 113-8656, Japan

²Research Center for Physical Fitness, Sports and Health, Toyohashi University of Technology, 1-1 Hibarigaoka, Tenpaku-cho, Toyohashi 441-8580, Japan

Received 30 November 2012; Accepted 16 January 2013

Academic Editors: L. Guimarães-Ferreira, H. Nicastro, J. Wilson, and N. E. Zanchi

Copyright © 2013 Akira Wagatsuma and Kunihiro Sakuma. This is an open access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

Abstract.

Recent studies have shown that mitochondria play a role in the regulation of myogenesis. Indeed, the abundance, morphology, and functional properties of mitochondria become altered when the myoblasts differentiate into myotubes. For example, mitochondrial mass/volume, mtDNA copy number, and mitochondrial respiration are markedly increased after the onset of myogenic differentiation. Besides, mitochondrial enzyme activity is also increased, suggesting that the metabolic shift from glycolysis to oxidative phosphorylation as the major energy source occurs during myogenic differentiation. Several lines of evidence suggest that impairment of mitochondrial function and activity blocks myogenic differentiation. However, yet little is known about the molecular mechanisms underlying the regulation of myogenesis by mitochondria. Understanding how mitochondria are involved in myogenesis will provide a valuable insight into the underlying mechanisms that regulate the maintenance of cellular homeostasis. Here, we will summarize the current knowledge regarding the role of mitochondria as a potential regulator of myogenesis.

1. Introduction

Mitochondria generate most of the energy necessary for cellular function via oxidative phosphorylation (OXPHOS) as well as contribute to metabolism, Ca²⁺ signaling, and apoptosis. Besides, several lines of evidence suggest that mitochondrial function and activity are linked to cell differentiation, as have been shown in a wide variety of cell types including myoblasts [1–9]. When the myoblasts differentiate into myotubes, mitochondrial enzyme activity is drastically increased [10–12]. Likewise, muscle regeneration is also accompanied by an increased mitochondrial enzyme activity [13–15]. These findings suggest that the metabolic shift from glycolysis to OXPHOS as the major energy source occurs during myogenesis. The metabolic shift has been reported in embryonic stem cells (ESCs) [16, 17] and induced pluripotent stem cells (iPSCs) [18]. For example, iPSCs have low mitochondrial activity, relying predominantly on glycolysis for ATP generation and maintaining a state of dedifferentiation, while differentiation is accompanied by an increased mitochondrial activity [18]. Therefore, the metabolic shift may be a key event initiating cell differentiation. This shift requires an activation of mitochondrial biogenesis through coordinated expression of nuclear and mitochondrial genomes. Mitochondrial biogenesis is tightly controlled by transcriptional coactivators, transcription factors, and nuclear receptors [19–23]. Their expression is coordinately induced during myogenic differentiation [11, 12] and muscle regeneration [13, 15]. To further elucidate the relationship between mitochondria and cell differentiation, the effects of impairment of mitochondrial function and activity on myogenic cells have been investigated using antimycin [24], azide [24–27], chloramphenicol [4, 6–9], carbonyl cyanide m-chlorophenylhydrazone carbonyl (CCCP) [24], cyanide p-(trifluoromethoxy) phenylhydrazone (FCCP) [6], ethidium bromide (EtBr) [1–3, 24], myxothiazol [7], rhodamine 6G [28], rifampicin [1], rotenone [7], oligomycin [6, 7, 26], tetracycline [5], and valinomycin [24]. Overall, these antibiotics and chemicals can exert a negative influence on myogenesis. For example, respiration-deficient myoblasts devoid of mitochondrial DNA (rho° cells) by EtBr, an inhibitor of mtDNA replication and transcription, fail to differentiate into myotubes [1–3]. Rifampicin, which inhibits mitochondrial RNA synthesis, shows reversible inhibition of myotube formation [1]. Tetracycline, an inhibitor of mitochondrial protein synthesis, blocks myoblasts fusion [5]. Chloramphenicol, an inhibitor of mitochondrial protein synthesis, restricts myogenic differentiation [4, 6–9] and interferes with muscle regeneration [15]. Despite the data being accumulating, little is known about the molecular mechanisms underlying the regulation of myogenesis by mitochondria. In this paper, we will summarize the current knowledge regarding the role of mitochondria as a potential regulator of myogenesis.

2. Mitochondrial Biogenesis

Mitochondrial biogenesis (also referred to as mitochondriogenesis) is characterized as a vital process in the synthesis and degradation of the organelle [29, 30]. Therefore, this fundamental process comprehends (1) the synthesis import and incorporation of lipids and proteins to the existing mitochondrial reticulum; (2) the stoichiometric assembly of multisubunit protein complexes into a functional respiratory chain; (3) replication of the mitochondrial DNA (mtDNA); (4) selective degradation of mitochondria by autophagy (mitophagy) [22, 31, 32]. When it is not indicated, in this paper, mitochondrial biogenesis simply considers an increase in mitochondrial volume and changes in organelle composition per tissue or cell [31]. Mitochondrial biogenesis requires a coordination of expression of nuclear and mitochondrial genomes [20].

3. Transcriptional Regulation of Mitochondrial Biogenesis

Recent technological and scientific advances have allowed it to systematically identify the complement of over 1,000 different proteins that comprise the mammalian mitochondrial proteome [33]. The majority of the mitochondrial proteins are encoded by the nuclear genome and synthesized on cytoplasmic ribosomes [20, 34], whereas the minority are encoded and synthesized within the mitochondria. The mitochondrial genome contains 37 genes encoding 13 enzymes involved in OXPHOS, 22 types of transfer RNAs, and 2 types of ribosomal RNAs [35]. To maintain mitochondrial functionality, it is necessary for two genomes to be coordinately regulated [20]. It has been widely accepted that peroxisome proliferator-activated receptor gamma coactivator-1 alpha (PGC-1α) plays a central role in a regulatory network governing the transcriptional control of mitochondrial biogenesis [20–23]. PGC-1α works in concert with a wide variety of interacting partners, which are transcription factors and nuclear receptors [21, 22]. PGC-1α was discovered in a yeast two-hybrid screen for brown adipose-specific factors that interact with the nuclear receptor PPARγ and are dramatically induced by exposure to cold in brown fat and skeletal muscle [36]. Subsequently, two additional PGC-1 family members were identified, PGC-1-related coactivator (PRC) [37] and PGC-1β [38]. The three coactivators regulate expression of a broad set of mitochondrial genes and promote mitochondrial biogenesis [39]. Since these coactivators lack DNA-binding activity [40, 41], PGC-1 family coactivators exert their effects through interactions with transcription factors and nuclear receptors bound to specific DNA elements in the promoter region of genes. For example, nuclear respiratory factor-1 (NRF-1) and NRF-2 (GA-binding protein; GABP) were the first regulatory factors implicated in the global expression of multiple mitochondrial functions in vertebrates [23]. Both NRF-1 and NRF-2 are involved in the transcriptional control of nuclear and mitochondrial genes involved in OXPHOS, electron transport (complex I-V), mtDNA transcription/replication, heme biosynthesis, protein import/assembly, ion channels, shuttles, and translation [19]. For more complete details, excellent review articles are already available on this subject [19–23].

4. Mitochondrial Enzyme Activity and Function during Muscle Regeneration and Myogenesis In Vitro

Muscle regeneration, which partially recapitulates embryonic myogenesis [42], would stimulate mitochondrial biogenesis [13]. Muscle injury was induced by intramuscular injection of either bupivacaine (which induces Ca²⁺ release from the sarcoplasmic reticulum (SR) and simultaneously inhibits Ca²⁺ reuptake into the SR, resulting in persistently increased [Ca²⁺] levels and leads to myofiber death), notexin (which involves Ca²⁺ overload and activation of Ca²⁺-dependent proteases, resulting in tissue necrosis), or freezing (which causes uniform and complete necrosis of myofibers). Such acute muscle injury shows a rapid loss of the activities of citrate synthase [13–15], a mitochondrial matrix enzyme participating in the Krebs cycle, which is often used as marker for the mitochondria content of a tissue, after 2-3 days, a time when degenerative myofibers still persist and proliferating myoblasts reside [13, 15]. The activity of citrate synthase then is increased drastically between days 5 and 10, a time when myoblasts differentiate into myotubes [13, 15]. Similarly, the rate of state-3 respiration (respiratory rate during active phosphorylation of ADP) is recovered [13]. The rate of state-4 respiration (respiratory rate after exhaustion of ADP) is comparable between control and injured muscles [13]. Accordingly, the pattern of changes in the respiratory control ratio (RCR), a measure of the “tightness of coupling” between electron transport and oxidative phosphorylation, is calculated as the ratio of state-3/state-4 respiration [43], closely resembling the pattern obtained with citrate synthase activity [13]. Consistent with the in vivo findings, the activity of mitochondrial enzymes including citrate synthase, isocitrate dehydrogenase, 3-hydroxyacyl CoA dehydrogenase, cytochrome reductase, succinate dehydrogenase, and cytochrome oxidase is drastically increased during myogenic differentiation [10, 12]. The content of respiratory chain complexes is higher in myotubes than in myoblasts [11, 44]. Similarly, the rate of state-3 respiration exhibits the same trend observed in respiratory chain complexes [44]. Leary et al. examined the changes in metabolic rate during myogenic differentiation [25]. In proliferating myoblasts, approximately 30% of the ATP used by the cells is provided by OXPHOS, whereas terminally differentiated myotubes rely on mitochondrial respiration as their major source of metabolic energy (approximately 60%) [25]. Intriguingly, the total metabolic rate remains constant throughout the culture period, but there is a steady shift toward a greater reliance on mitochondrial pathways [25]. Taken together, these findings suggest that the metabolic shift from glycolysis to oxidative phosphorylation as the major energy source occurs during myogenesis.

5. Gene Expression Involved in Mitochondrial Biogenesis during Muscle Regeneration and Myogenesis In Vitro

An activation of mitochondrial biogenesis occurs through the coordinated expression of PGC-1 transcriptional coactivators, transcription factors, and nuclear receptors. Surprisingly, PGC-1α expression remains unchanged during muscle regeneration, whereas PRC and PGC-1β are upregulated 3 days after injury [15]. This finding may be in line with other studies using mouse myoblasts [37, 45, 46]. PGC-1α fails to detect in either myoblasts or myotubes, whereas PRC and PGC-1β are readily detectable [46]. In contrast, Duguez et al. have reported that PGC-1α is upregulated 10 days after injury [13]. Irrespective of whether PGC-1α is upregulated in injured muscle, these findings lead us to hypothesize that PRC and PGC-1β may contribute to the mitochondrial biogenesis, at least in part, at the early stage of muscle regeneration. It has been shown that PRC coactivates NRF-1 [37, 45], and PGC-1β also interacts with NRF-1 and estrogen-related receptor α (ERRα) [47], suggesting that both PRC and PGC-1β may functionally replace PGC-1α during muscle regeneration. In support of this hypothesis, several studies have revealed the potential roles of PRC and PGC-1β in mitochondrial biogenesis in myogenic cells. PRC-overexpressing myotubes show an elevated fatty acid oxidation and increased expression of mitochondrial genes [48]. Forced expression of PGC-1β in C2C12 cells results in increased mitochondrial biogenesis and oxygen consumption [49]. Skeletal muscle-specific PGC-1β transgenic mice exhibit increased mtDNA amount, mitochondrial content, mitochondrial enzyme activity, upregulation of mitochondrial genes, and enhanced exercise performance [50]. On the other hand, mice lacking PGC-1β show a reduced number of mitochondria, decreased respiration function, and decreased expression of mitochondrial genes [51]. However, the possibility cannot be excluded that PGC-1α may contribute to the mitochondrial biogenesis during muscle regeneration, as has been shown in gain-of-function and loss-of-function studies [52–56]. Accordingly, further studies are required to elucidate the role of PGC-1α in mitochondrial biogenesis during muscle regeneration.

Not only PGC-1 family coactiuators but also NRF-1, NRF-2, and mitochondrial transcription factor A (TFAM) are also upregulated during muscle regeneration [15]. This is in line with the findings that PGC-1 stimulates an induction of NRF-1 and NRF-2 gene expression and can also interact directly with and coactivate NRF-1 on the promoter for TFAM [57]. TFAM plays a key role in mammalian mtDNA transcription/replication [21]. Likewise, when myoblasts differentiate into myotubes, PGC-1α, NRF-1, and TFAM are upregulated, and mtDNA content and copy number are increased 2–4-fold in myotubes relative to myoblasts [11, 12]. Therefore, upregulation of these genes contributes to increase the template availability for transcription and translation of key mitochondrial proteins necessary for myogenesis.

6. Possible Role of Mitochondria in Regulating Muscle Regeneration

Recent studies have extended our knowledge of the potential role of mitochondrial biogenesis in muscle regeneration [15, 58]. It has been reported that muscle regeneration is impaired when mitochondrial protein synthesis is inhibited with chloramphenicol [15]. Chloramphenicol inhibits protein synthesis in mitochondria but not in mammalian cytoplasmic ribosomal systems [59] since mammalian mitochondrial ribosomes are susceptible to peptidyl-transferase inhibition by it [60]. Chloramphenicol reversibly binds to the 50S subunit of the 70S ribosome and blocks prokaryotic protein translation primarily by inhibiting peptidyl-transferase and blocking elongation [61]. Consequently, chloramphenicol inhibits the proper assembly of 4 out of 5 respiratory chain complexes within mitochondria and therefore potentially attenuates mitochondrial biogenesis in mammalian cells. Mice were intramuscularly injected with chloramphenicol at days 3, 5, and 7 after the initial freeze injury, and the muscle specimens were histochemically analyzed at day 10. Impairment of mitochondrial activity induced by chloramphenicol results in poor muscle regeneration with small myofibers and increased connective tissues [15]. Overall, this supports in vitro data that show that chloramphenicol blocks myogenic differentiation [4–9]. Therefore, in vivo data, when combined with the previous data in vitro, suggests a role for mitochondrial biogenesis for sustaining muscle regeneration. However, the molecular mechanisms remain unknown although chloramphenicol downregulates myogenin, which is required for terminal differentiation and myotube formation, in an avian QM7 myoblast [6, 8] and mouse C2C12 myoblast [9].

It has been reported that muscle regeneration is effectively accelerated using a method for complex mediated delivery to intracellular mitochondria [58]. The method is based on the mitochondriotropism of a multisubunit RNA import complex (RIC) [62]. Muscle injury was induced by piercing repeatedly with a 26-gauge hypodermic needle at an angle of ~45° to the longitudinal axis of the fiber, resulting in ~3000 myofibers being damaged at each insertion [58]. When a combination of polycistronic RNAs encoding the guanine-rich heavy-strand (H-strand) of the mitochondrial genome is administrated to injured muscle, it rejuvenates mitochondrial mRNA levels, organellar translation, respiratory capacity, and intramuscular ATP levels with reduced intracellular reactive oxygen species levels [58]. It increases proliferative potential of satellite cells and differentiation capacity of myoblasts concomitantly with upregulation of myogenic regulatory factors including Myf5, MyoD, myogenin, and MRF4, promoting muscle regeneration with the recovery of muscle contractility [58]. One of the most intriguing aspects of RIC-mediated transfection strategy, MyoD, and Numb-positive cells are detected and attached to old myofibers at the injury site [58]. This may provide new insight into the possible mechanism regulating muscle regeneration through enhancing mitochondrial activity. Numb protein has been generally considered to be a negative regulator of Notch signaling [63], which inhibits myogenic differentiation [63]. Numb segregates asymmetrically in dividing adult mouse muscle satellite cells [64, 65]. Attenuation of Notch signaling by Numb overexpression leads to the commitment of progenitor cells to the myoblast cell fate with increased expression of Myf5 and desmin [64]. Therefore, RIC-induced Numb protein may play a certain role in regulating muscle regeneration by modulating Notch signaling. However, recent evidence suggests that although forced expression of Numb in myogenic progenitors does not abrogate canonical Notch signaling, it can stimulate the self-renewal of myogenic progenitors [66]. Therefore, a role of Numb in regulating muscle regeneration remains to be elucidated. Furthermore, it is unknown how mitochondrial activity modulates Notch signaling at the present time.

7. Do Mitochondria Act as a Potential Regulator of Myogenesis?

Korohoda et al. [4] have reported that chloramphenicol inhibits the fusion of myoblasts isolated from chick embryo skeletal muscle. This is among the first study to show the effect of chloramphenicol on myogenesis. They show that tryptose phosphate broth and nucleosides can restore the cell capacity to proliferate but not to fuse and differentiate in the presence of chloramphenicol [4]. Subsequently, it has been demonstrated that mitochondrial activity is an important regulator of myogenic differentiation in quail myoblasts of the QM7 cell line and mouse myoblasts of the C2C12 cell line using chloramphenicol [6, 8, 9]. Chloramphenicol-treated myoblasts proliferate at a slower rate than control myoblasts without inducing any alteration of cell viability [6]. When chronically exposed to chloramphenicol throughout the culture period, it severely suppresses myogenic differentiation [6, 8, 9]. The possibility can be excluded that intracellular ATP depletion induced by chloramphenicol could be responsible for the inhibition of myoblast differentiation for the following the reasons: (1) glycolysis fully compensates for mitochondrial impairment just before and during terminal differentiation, as shown in a marked accumulation of L-lactate in the culture medium [6], and this has been already reported in C2C12 cells using tetracycline [5]; (2) differentiation of myoblast is repressed especially when exposing to chloramphenicol at the onset of terminal differentiation [6]. These findings indicate that mitochondrial activity regulates myogenic differentiation independently of their implication in ATP synthesis [6].

Chloramphenicol inhibits myogenic differentiation by downregulating myogenin but not MyoD and Myf5 [6, 8]. Intriguingly, this downregulation is commonly observed in FCCP, myxothiazol [7], rotenone [7], and oligomycin [6, 7], which affect mitochondria at different levels. These findings suggest that myogenin could be an important target of mitochondrial activity. Chloramphenicol has no effect on myogenin mRNA stability [6], suggesting that mitochondrial activity could regulate myogenin expression at the transcriptional level [6]. Unexpectedly, overexpression of neither myogenin nor MyoD fails to restore differentiation capacity in chloramphenicol-treated myoblasts [6]. This indicates that mitochondrial activity could regulate myogenic differentiation by decaying ability of myogenic regulatory factors via other negative regulators. Chloramphenicol has no effect on the expression of MEF2C (myocyte enhancer factor 2C) and Id (inhibitor of differentiation) [8]. Seyer et al. have identified c-Myc (cellular myelocytomatosis oncogene) gene, which could be a target gene regulated by mitochondrial activity [8]. c-Myc is a proto-oncogene encoding a transcription factor [67], which plays a role in regulating myogenesis [68–74]. Impairment of mitochondrial activity by chloramphenicol abrogates the downregulation of c-Myc normally occurring at the induction of differentiation in control cells [8]. Overexpression of c-Myc mimics the influence of mitochondrial activity inhibition on myogenic differentiation [8]. A triiodothyronine-dependent mitochondrial transcription factor (p43) overexpression, which stimulates mitochondrial activity, downregulates c-Myc expression [8]. These findings suggest the possibility that c-Myc could be a primary target of mitochondrial activity. Indeed, the endogenous c-Myc is downregulated within the first 24 h after switching to a differentiation medium [70]. Ectopic expression of c-Myc in quail myoblasts fails to form myotubes and downregulates MyoD, myogenin, and Myf5 expression [73]. Cotransfection of c-Myc with MyoD and myogenin in NIH 3T3 cells inhibits myogenic differentiation [71].

While these findings are compelling, a role of c-Myc should be carefully considered. First, irreversible repression of c-Myc is not required for terminal myogenic differentiation, and its expression is insufficient to suppress the differentiated phenotype, since nuclear runoff transcription assay demonstrates that c-Myc and skeletal muscle-specific genes could be simultaneously transcribed in both biochemically differentiated cells (no fusion) and terminally differentiated cells [69]. The c-Myc- transformed C2C12 cells retain the ability to undergo commitment and biochemical differentiation, but they are strikingly unable to fuse into multinucleated myotubes with no change in the expression of MyoD, myogenin, and myosin heavy chain [72]. These findings lead us to rethink how c-Myc modulates myogenic differentiation. Secondly, c-Myc represses expression through transcriptional activator, Miz-1- (c-Myc interacting zinc-finger protein 1-) dependent interaction with core promoter [75]. In addition, c-Myc interacts with Miz-1 and recruits DNA methyltransferase 3A to promoter to silence p21 transcription [76]. The expression of is known to be a key event triggering the withdrawal of myoblasts from the cell cycle to G₀, a prerequisite to myogenic differentiation [77]. Indeed, chloramphenicol and overexpression of c-Myc decrease the proportion of myoblasts in the G₀-G₁ phase, whereas overexpression of p43 exerts opposite influence [8]. These findings suggest the possibility that mitochondrial activity could regulate myoblast cell cycle withdrawal by modulating expression of through c-Myc/Miz-1 complex. Thirdly, Myc is a member of the Myc/Max (Myc-associated factor X)/Mad (MAX dimerization protein) transcriptional network that comprises a group of widely expressed transcription factors [78]. c-Myc/Max heterodimers transactivate its downstream genes by binding to the E-box sequence 5′-CACGTG-3′ in the target promoter, whereas Mad/Max heterodimers act as transcriptional repressors at the same E-box-related DNA-binding sites [78]. Therefore, c-Myc/Max heterodimers function by competing with Mad/Max heterodimers, resulting in controlling the expression of their target genes. Intriguingly, a switching from c-Myc/Max to Mad/Max heterodimers occurs when leukemia cells differentiate into monocyte/macrophage [79, 80]. These findings lead us to hypothesize that mitochondrial activity may be involved in this switching during myogenic differentiation. It requires additional studies to validate this observation in myogenic cells. Finally, a new mode of Myc regulation has been recently reported in myogenic differentiation [81]. Myc protein is cleaved by a calpain to generate a cytoplasmic form, “Myc-Nick,” which retains Myc box regions but lacks nuclear localization sequence and the basic helix-loop-helix/leucine zipper domains essential for heterodimerization with Max and DNA binding activity [81]. During myogenic differentiation, while the full-length Myc decreases, Myc-nick is increased. Ectopic expression of Myc-nick in human primary myoblasts, human rhabdomyosarcoma (RD) cells, and mouse C2C12 myoblasts accelerates their differentiation and increases expression of skeletal muscle-specific markers [81]. Taken together, the mechanisms underlying the regulation of biological function of c-Myc are complicated. Therefore, further studies are needed to elucidate the role of c-Myc in the regulation of myogenesis by mitochondria.

To further understand the molecular mechanisms underlying the regulation of myogenic differentiation by mitochondria, Seyer et al. [9] conducted a comprehensive differential display analysis using total RNA from control and chloramphenicol-treated myoblasts to search for other gene modulating by mitochondrial activity [9]. They identified calcineurin (also referred to as protein phosphatase 2B) as another candidate molecule [9], in which serine/threonine protein phosphatase under the control of a eukaryotic Ca²⁺- and calmodulin plays a critical role in the coupling of Ca²⁺ signals to cellular responses [82]. It is a heterodimeric enzyme consisting of a 60 kDa catalytic A subunit (calcineurin A) and 19 kDa calcium-binding regulatory B subunit (calcineurin B) [82]. Calcineurin signaling has been implicated in regulating myogenesis [83–90]. Chloramphenicol attenuates the differentiation-induced upregulation of calcineurin A, whereas overexpression of p43 increases calcineurin A expression in proliferating myoblasts [9]. Based on these findings, they suggest that calcineurin could be a novel target regulated by mitochondrial activity. Intriguingly, expression of a constitutively active form of calcineurin upregulates the expression of myogenin [85]. Calcineurin regulates expression of the myogenin gene at the transcriptional level by activating MEF2 and MyoD transcription factors [87]. Taken together, mitochondrial activity may regulate myogenesis through calcineurin-mediated myogenin expression. On the other hand, it has been shown that calcineurin A and its direct downstream transcriptional effector, NFATc (nuclear factor of activated T-cells), are upregulated concomitantly with a modest increase in calcineurin B in mtDNA-depleted cells (only ~20% of the mtDNA content compared with normal untreated cells) [24]. Biswas et al. developed myogenic cell lines with partially depleted mtDNA when chronically exposed to EtBr for many passages to investigate the mechanism of mitochondrial-nuclear crosstalk [24]. The mtDNA-depleted cells have an elevated steady-state cytosolic Ca²⁺ level ([Ca²⁺]i), as shown in other mitochondrial inhibitors including antimycin, azide, CCCP, and valinomycin [24]. Therefore, increased cytosolic Ca²⁺ may stimulate the expression of calcineurin-related molecules in the myoblasts treated with these drugs. It is to be noted that increased expression of calcineurin is observed by mtDNA depletion or acute treatment (30 min) with high amounts of mitochondrial inhibitors. As already described, mtDNA-depleted myoblasts by EtBr fail to differentiate into myotubes [1–3], and NFAT is not an essential downstream target of calcineurin during myogenesis [85]. Therefore, the activation of calcineurin pathway induced by impairment of mitochondrial function and activity could not contribute to myogenesis.

The nuclear factor-κB (NF-κB) functions as a negative regulator of myogenesis [91]. NF-κB is a heterodimeric or homodimeric complex formed from five distinct subunits: RelA (p65), RelB, c-Rel, NF-κB1 (p50/p105), and NF-κB2 (p52/p100) [91]. Only RelA, c-Rel, and RelB possess C-terminal transcriptional transactivation domains, whereas NF-kB1 and NF-kB2 lack intrinsic transactivating properties and instead function as homodimeric transcriptional repressors or modulators of transactivating dimer partners [91]. When stimulated by a wide variety of different stimuli, IκB is phosphorylated by IκB kinase (IKK) complex and subsequently degraded by the proteasome, allowing NF-κB to translocate into the nucleus where they regulate target gene expression [91]. Respiration-deficient myoblasts devoid of mitochondrial DNA by EtBr show a decreased expression of RelA, increased expression of IκB and p50, and unchanged expression of RelB and p52 [24]. Intriguingly, other mitochondrial inhibitors also have same effects on their expression [24]. These findings suggest that mitochondrial activity can modulate NF-κB transcriptional activity although it is required for measuring its DNA binding activity, for example, by an electrophoretic mobility shift assay.

8. Conclusion

This paper provides the current knowledge about the role for mitochondria as a potential regulator of myogenesis. Several studies have highlighted that mitochondria play a role in regulating myogenic differentiation possibly through a number of mechanisms. In particular, myogenin, c-Myc, and calcineurin have been identified as candidate molecules of mitochondrial target [6, 8, 9]. Together with previous data [8, 9, 87], a hypothetical model involving c-Myc and calcineurin in the regulation of myogenic differentiation by mitochondrial activity is presented in Figure 1. In this model, when myoblasts are induced to differentiate in the presence of mitochondrial inhibitors, downregulation of c-Myc could be inhibited, which depresses the activity of MyoD and myogenin, resulting in blocking myogenic differentiation. Decreased calcineurin signaling by inhibiting mitochondrial activity could contribute to myogenin expression through modulating MyoD and MEF2 activity. Understanding how mitochondria are involved in myogenesis will provide a valuable insight into the underlying mechanisms that regulate the maintenance of cellular homeostasis. Recently, it has been reported that the transgenic mice with skeletal muscle-specific expression of PGC-1α preserve mitochondrial function as well as neuromuscular junctions and muscle integrity during ageing [92], and mitochondrial gene therapy may be effective in the treatment of muscle injury [58]. These efforts may facilitate to understand the molecular mechanisms of mitochondrial disorders.

Figure 1: Hypothetic model of mitochondrial activity in myogenic differentiation.

Acknowledgment

This research was supported by the Ministry of Education, Culture, Sports, Science and Technology (MEXT) (Grant-in-Aid for Scientific Research (C), 22500658), Japan.

References

C. F. Brunk and D. Yaffe, “The reversible inhibition of myoblast fusion by ethidium bromide (EB),” Experimental Cell Research, vol. 99, no. 2, pp. 310–318, 1976.

N. H. Herzberg, R. Zwart, R. A. Wolterman et al., “Differentiation and proliferation of respiration-deficient human myoblasts,” Biochimica et Biophysica Acta, vol. 1181, no. 1, pp. 63–67, 1993.

N. H. Herzberg, E. Middelkoop, M. Adorf et al., “Mitochondria in cultured human muscle cells depleted of mitochondrial DNA,” European The Journal of Cell Biology, vol. 61, no. 2, pp. 400–408, 1993.

W. Korohoda, Z. Pietrzkowski, and K. Reiss, “Chloramphenicol, an inhibitor of mitochondrial protein synthesis, inhibits myoblast fusion and myotube differentiation,” Folia Histochemica et Cytobiologica, vol. 31, no. 1, pp. 9–13, 1993.

N. Hamai, M. Nakamura, and A. Asano, “Inhibition of mitochondrial protein synthesis impaired C2C12 myoblast differentiation,” Cell Structure and Function, vol. 22, no. 4, pp. 421–431, 1997.

P. Rochard, A. Rodier, F. Casas et al., “Mitochondrial activity is involved in the regulation of myoblast differentiation through myogenin expression and activity of myogenic factors,” The Journal of Biological Chemistry, vol. 275, no. 4, pp. 2733–2744, 2000.

P. Pawlikowska, B. Gajkowska, J. F. Hocquette, and A. Orzechowski, “Not only insulin stimulates mitochondriogenesis in muscle cells, but mitochondria are also essential for insulin-mediated myogenesis,” Cell Proliferation, vol. 39, no. 2, pp. 127–145, 2006.

P. Seyer, S. Grandemange, M. Busson et al., “Mitochondrial activity regulates myoblast differentiation by control of c-Myc expression,” Journal of Cellular Physiology, vol. 207, no. 1, pp. 75–86, 2006.

P. Seyer, S. Grandemange, P. Rochard et al., “P43-dependent mitochondrial activity regulates myoblast differentiation and slow myosin isoform expression by control of Calcineurin expression,” Experimental Cell Research, vol. 317, no. 14, pp. 2059–2071, 2011.

C. D. Moyes, O. A. Mathieu-Costello, N. Tsuchiya, C. Filburn, and R. G. Hansford, “Mitochondrial biogenesis during cellular differentiation,” American Journal of Physiology, vol. 272, no. 4, pp. C1345–C1351, 1997.

A. H. V. Remels, R. C. J. Langen, P. Schrauwen, G. Schaart, A. M. W. J. Schols, and H. R. Gosker, “Regulation of mitochondrial biogenesis during myogenesis,” Molecular and Cellular Endocrinology, vol. 315, no. 1-2, pp. 113–120, 2010.

E. Barbieri, M. Battistelli, L. Casadei et al., “Morphofunctional and biochemical approaches for studying mitochondrial changes during myoblasts differentiation,” Journal of Aging Research, vol. 2011, Article ID 845379, 16 pages, 2011.

S. Duguez, L. Féasson, C. Denis, and D. Freyssenet, “Mitochondrial biogenesis during skeletal muscle regeneration,” American Journal of Physiology, vol. 282, no. 4, pp. E802–E809, 2002.

E. Fink, D. Fortin, B. Serrurier, R. Ventura-Clapier, and A. X. Bigard, “Recovery of contractile and metabolic phenotypes in regenerating slow muscle after notexin-induced or crush injury,” Journal of Muscle Research and Cell Motility, vol. 24, no. 7, pp. 421–429, 2003.

A. Wagatsuma, N. Kotake, and S. Yamada, “Muscle regeneration occurs to coincide with mitochondrial biogenesis,” Molecular and Cellular Biochemistry, vol. 349, no. 1-2, pp. 139–147, 2011.

S. Chung, P. P. Dzeja, R. S. Faustino, C. Perez-Terzic, A. Behfar, and A. Terzic, “Mitochondrial oxidative metabolism is required for the cardiac differentiation of stem cells,” Nature Clinical Practice Cardiovascular Medicine, vol. 4, supplement 1, pp. S60–S67, 2007.

S. Chung, P. P. Dzeja, R. S. Faustino, and A. Terzic, “Developmental restructuring of the creatine kinase system integrates mitochondrial energetics with stem cell cardiogenesis,” Annals of the New York Academy of Sciences, vol. 1147, pp. 254–263, 2008.

C. D. L. Folmes, T. J. Nelson, A. Martinez-Fernandez et al., “Somatic oxidative bioenergetics transitions into pluripotency-dependent glycolysis to facilitate nuclear reprogramming,” Cell Metabolism, vol. 14, no. 2, pp. 264–271, 2011.

D. P. Kelly and R. C. Scarpulla, “Transcriptional regulatory circuits controlling mitochondrial biogenesis and function,” Genes and Development, vol. 18, no. 4, pp. 357–368, 2004.

M. T. Ryan and N. J. Hoogenraad, “Mitochondrial-nuclear communications,” Annual Review of Biochemistry, vol. 76, pp. 701–722, 2007.

R. C. Scarpulla, “Transcriptional paradigms in mammalian mitochondrial biogenesis and function,” Physiological Reviews, vol. 88, no. 2, pp. 611–638, 2008.

M. B. Hock and A. Kralli, “Transcriptional control of mitochondrial biogenesis and function,” Annual Review of Physiology, vol. 71, pp. 177–203, 2009.

R. C. Scarpulla, “Metabolic control of mitochondrial biogenesis through the PGC-1 family regulatory network,” Biochimica et Biophysica Acta, vol. 1813, no. 7, pp. 1269–1278, 2011.

G. Biswas, O. A. Adebanjo, B. D. Freedman et al., “Retrograde Ca²⁺ signaling in C2C12 skeletal myocytes in response to mitochondrial genetic and metabolic stress: a novel mode of inter-organelle crosstalk,” The EMBO Journal, vol. 18, no. 3, pp. 522–533, 1999.

S. C. Leary, B. J. Battersby, R. G. Hansford, and C. D. Moyes, “Interactions between bioenergetics and mitochondrial biogenesis,” Biochimica et Biophysica Acta, vol. 1365, no. 3, pp. 522–530, 1998.

S. C. Leary, B. C. Hill, C. N. Lyons et al., “Chronic treatment with azide in situ leads to an irreversible loss of cytochrome c oxidase activity via holoenzyme dissociation,” The Journal of Biological Chemistry, vol. 277, no. 13, pp. 11321–11328, 2002.

Z. Yun, Q. Lin, and A. J. Giaccia, “Adaptive myogenesis under hypoxia,” Molecular and Cellular Biology, vol. 25, no. 8, pp. 3040–3055, 2005.

C. Sobreira, M. P. King, M. M. Davidson, H. Park, Y. Koga, and A. F. Miranda, “Long-term analysis of differentiation in human myoblasts repopulated with mitochondria harboring mtDNA mutations,” Biochemical and Biophysical Research Communications, vol. 266, no. 1, pp. 179–186, 1999.

D. A. Hood, “Invited review: contractile activity-induced mitochondrial biogenesis in skeletal muscle,” Journal of Applied Physiology, vol. 90, no. 3, pp. 1137–1157, 2001.

A. Terman, T. Kurz, M. Navratil, E. A. Arriaga, and U. T. Brunk, “Mitochondrial turnover and aging of long-lived postmitotic cells: the mitochondrial-lysosomal axis theory of aging,” Antioxidants and Redox Signaling, vol. 12, no. 4, pp. 503–535, 2010.

D. A. Hood, I. Irrcher, V. Ljubicic, and A. M. Joseph, “Coordination of metabolic plasticity in skeletal muscle,” Journal of Experimental Biology, vol. 209, no. 12, pp. 2265–2275, 2006.

I. Kim, S. Rodriguez-Enriquez, and J. J. Lemasters, “Selective degradation of mitochondria by mitophagy,” Archives of Biochemistry and Biophysics, vol. 462, no. 2, pp. 245–253, 2007.

S. E. Calvo and V. K. Mootha, “The mitochondrial proteome and human disease,” Annual Review of Genomics and Human Genetics, vol. 11, pp. 25–44, 2010.

S. Goffart and R. J. Wiesner, “Regulation and co-ordination of nuclear gene expression during mitochondrial biogenesis,” Experimental Physiology, vol. 88, no. 1, pp. 33–40, 2003.

S. Anderson, A. T. Bankier, B. G. Barrell et al., “Sequence and organization of the human mitochondrial genome,” Nature, vol. 290, no. 5806, pp. 457–465, 1981.

P. Puigserver, Z. Wu, C. W. Park, R. Graves, M. Wright, and B. M. Spiegelman, “A cold-inducible coactivator of nuclear receptors linked to adaptive thermogenesis,” Cell, vol. 92, no. 6, pp. 829–839, 1998.

U. Andersson and R. C. Scarpulla, “PGC-1-related coactivator, a novel, serum-inducible coactivator of nuclear respiratory factor 1-dependent transcription in mammalian cells,” Molecular and Cellular Biology, vol. 21, no. 11, pp. 3738–3749, 2001.

J. Lin, P. Puigserver, J. Donovan, P. Tarr, and B. M. Spiegelman, “Peroxisome proliferator-activated receptor γ coactivator 1β (PGC-1β), a novel PGC-1-related transcription coactivator associated with host cell factor,” The Journal of Biological Chemistry, vol. 277, no. 3, pp. 1645–1648, 2002.

C. Handschin and B. M. Spiegelman, “Peroxisome proliferator-activated receptor γ coactivator 1 coactivators, energy homeostasis, and metabolism,” Endocrine Reviews, vol. 27, no. 7, pp. 728–735, 2006.

P. Puigserver, G. Adelmant, Z. Wu et al., “Activation of PPARγ coactivator-1 through transcription factor docking,” Science, vol. 286, no. 5443, pp. 1368–1371, 1999.

B. N. Finck and D. P. Kelly, “Peroxisome proliferator-activated receptor γ coactivator-1 (PGC-1) regulatory cascade in cardiac physiology and disease,” Circulation, vol. 115, no. 19, pp. 2540–2548, 2007.

P. Zhao and E. P. Hoffman, “Embryonic myogenesis pathways in muscle regeneration,” Developmental Dynamics, vol. 229, no. 2, pp. 380–392, 2004.

A. J. Kanai, L. L. Pearce, P. R. Clemens et al., “Identification of a neuronal nitric oxide synthase in isolated cardiac mitochondria using electrochemical detection,” Proceedings of the National Academy of Sciences of the United States of America, vol. 98, no. 24, pp. 14126–14131, 2001.

D. Malinska, A. P. Kudin, M. Bejtka, and W. S. Kunz, “Changes in mitochondrial reactive oxygen species synthesis during differentiation of skeletal muscle cells,” Mitochondrion, vol. 12, no. 1, pp. 144–148, 2012.

N. Gleyzer, K. Vercauteren, and R. C. Scarpulla, “Control of mitochondrial transcription specificity factors (TFB1M and TFB2M) by nuclear respiratory factors (NRF-1 and NRF-2) and PGC-1 family coactivators,” Molecular and Cellular Biology, vol. 25, no. 4, pp. 1354–1366, 2005.

C. S. Kraft, C. M. R. LeMoine, C. N. Lyons, D. Michaud, C. R. Mueller, and C. D. Moyes, “Control of mitochondrial biogenesis during myogenesis,” American Journal of Physiology, vol. 290, no. 4, pp. C1119–C1127, 2006.

D. Shao, Y. Liu, X. Liu et al., “PGC-1β-Regulated mitochondrial biogenesis and function in myotubes is mediated by NRF-1 and ERRα,” Mitochondrion, vol. 10, no. 5, pp. 516–527, 2010.

A. Philp, M. Y. Belew, A. Evans et al., “The PGC-1α-related coactivator promotes mitochondrial and myogenic adaptations in C2C12 myotubes,” American Journal of Physiology, vol. 301, no. 4, pp. R864–R872, 2011.

J. St-Pierre, J. Lin, S. Krauss et al., “Bioenergetic analysis of peroxisome proliferator-activated receptor γ coactivators 1α and 1β (PGC-1α and PGC-1β) in muscle cells,” The Journal of Biological Chemistry, vol. 278, no. 29, pp. 26597–26603, 2003.

Z. Arany, N. Lebrasseur, C. Morris et al., “The transcriptional coactivator PGC-1β drives the formation of oxidative type IIX fibers in skeletal muscle,” Cell Metabolism, vol. 5, no. 1, pp. 35–46, 2007.

C. J. Lelliott, G. Medina-Gomez, N. Petrovic et al., “Ablation of PGC-1β results in defective mitochondrial activity, thermogenesis, hepatic function, and cardiac performance,” PLoS Biology, vol. 4, no. 11, article e369, 2006.

J. Lin, H. Wu, P. T. Tarr et al., “Transcriptional co-activator PGC-1α drives the formation of slow-twitch muscle fibres,” Nature, vol. 418, no. 6899, pp. 797–801, 2002.

T. C. Leone, J. J. Lehman, B. N. Finck et al., “PGC-1alpha deficiency causes multi-system energy metabolic derangements: muscle dysfunction, abnormal weight control and hepatic steatosis,” PLoS Biology, vol. 3, no. 4, article e101, 2005.

C. Handschin, S. C. Cheol, S. Chin et al., “Abnormal glucose homeostasis in skeletal muscle-specific PGC-1α knockout mice reveals skeletal muscle-pancreatic β cell crosstalk,” Journal of Clinical Investigation, vol. 117, no. 11, pp. 3463–3474, 2007.

C. Handschin, S. Chin, P. Li et al., “Skeletal muscle fiber-type switching, exercise intolerance, and myopathy in PGC-1α muscle-specific knock-out animals,” The Journal of Biological Chemistry, vol. 282, no. 41, pp. 30014–30021, 2007.

J. A. Calvo, T. G. Daniels, X. Wang et al., “Muscle-specific expression of PPARγ coactivator-1α improves exercise performance and increases peak oxygen uptake,” Journal of Applied Physiology, vol. 104, no. 5, pp. 1304–1312, 2008.

Z. Wu, P. Puigserver, U. Andersson et al., “Mechanisms controlling mitochondrial biogenesis and respiration through the thermogenic coactivator PGC-1,” Cell, vol. 98, no. 1, pp. 115–124, 1999.

S. Jash and S. Adhya, “Induction of muscle regeneration by RNA-mediated mitochondrial restoration,” FASEB Journal, vol. 26, no. 10, pp. 4187–4197, 2012.

M. E. Pullman and G. Schatz, “Mitochondrial oxidations and energy coupling,” Annual Review of Biochemistry, vol. 36, pp. 539–611, 1967.

Z. M. Chrzanowska-Lightowlers, A. Pajak, and R. N. Lightowlers, “Termination of protein synthesis in mammalian mitochondria,” The Journal of Biological Chemistry, vol. 286, no. 40, pp. 34479–34485, 2011.

E. Cundliffe and K. McQuillen, “Bacterial protein synthesis: the effects of antibiotics,” Journal of Molecular Biology, vol. 30, no. 1, pp. 137–146, 1967.

B. Mahata, S. Mukherjee, S. Mishra, A. Bandyopadhyay, and S. Adhya, “Functional delivery of a cytosolic tRNA info mutant mitochondria of human cells,” Science, vol. 314, no. 5798, pp. 471–474, 2006.

M. F. Buas and T. Kadesch, “Regulation of skeletal myogenesis by Notch,” Experimental Cell Research, vol. 316, no. 18, pp. 3028–3033, 2010.

I. M. Conboy and T. A. Rando, “The regulation of Notch signaling controls satellite cell activation and cell fate determination in postnatal myogenesis,” Developmental Cell, vol. 3, no. 3, pp. 397–409, 2002.

V. Shinin, B. Gayraud-Morel, D. Gomès, and S. Tajbakhsh, “Asymmetric division and cosegregation of template DNA strands in adult muscle satellite cells,” Nature Cell Biology, vol. 8, no. 7, pp. 677–682, 2006.

A. Jory, I. Le Roux, B. Gayraud-Morel et al., “Numb promotes an increase in skeletal muscle progenitor cells in the embryonic somite,” Stem Cells, vol. 27, no. 11, pp. 2769–2780, 2009.

C. V. Dang, K. A. O'Donnell, K. I. Zeller, T. Nguyen, R. C. Osthus, and F. Li, “The c-Myc target gene network,” Seminars in Cancer Biology, vol. 16, no. 4, pp. 253–264, 2006.

T. Sejersen, J. Sumegi, and N. R. Ringertz, “Density-dependent arrest of DNA replication is accompanied by decreased levels of c-myc mRNA in myogenic but not in differentiation-defective myoblasts,” Journal of Cellular Physiology, vol. 125, no. 3, pp. 465–470, 1985.

T. Endo and B. Nadal-Ginard, “Transcriptional and posttranscriptional control of c-myc during myogenesis: its mRNA remains inducible in differentiated cells and does not suppress the differentiated phenotype,” Molecular and Cellular Biology, vol. 6, no. 5, pp. 1412–1421, 1986.

N. Denis, S. Blanc, M. P. Leibovitch et al., “c-myc oncogene expression inhibits the initiation of myogenic differentiation,” Experimental Cell Research, vol. 172, no. 1, pp. 212–217, 1987.

J. H. Miner and B. J. Wold, “c-myc Inhibition of MyoD and myogenin-initiated myogenic differentiation,” Molecular and Cellular Biology, vol. 11, no. 5, pp. 2842–2851, 1991.

M. Crescenzi, D. H. Crouch, and F. Tatò, “Transformation by myc prevents fusion but not biochemical differentiation of C2C12 myoblasts: mechanisms of phenotypic correction in mixed culture with normal cells,” The Journal of Cell Biology, vol. 125, no. 5, pp. 1137–1145, 1994.

S. A. La Rocca, D. H. Crouch, and D. A. F. Gillespie, “c-Myc inhibits myogenic differentiation and myoD expression by a mechanism which can be dissociated from cell transformation,” Oncogene, vol. 9, no. 12, pp. 3499–3508, 1994.

N. M. Yeilding, W. N. Procopio, M. T. Rehman, and W. M. F. Lee, “c-myc mRNA is down-regulated during myogenic differentiation by accelerated decay that depends on translation of regulatory coding elements,” The Journal of Biological Chemistry, vol. 273, no. 25, pp. 15749–15757, 1998.

S. Wu, C. Cetinkaya, M. J. Munoz-Alonso et al., “Myc represses differentiation-induced p21CIP1 expression via Miz-1-dependent interaction with the p21 core promoter,” Oncogene, vol. 22, no. 3, pp. 351–360, 2003.

C. Brenner, R. Deplus, C. Didelot et al., “Myc represses transcription through recruitment of DNA methyltransferase corepressor,” The EMBO Journal, vol. 24, no. 2, pp. 336–346, 2005.

V. Andrés and K. Walsh, “Myogenin expression, cell cycle withdrawal, and phenotypic differentiation are temporally separable events that precede cell fusion upon myogenesis,” The Journal of Cell Biology, vol. 132, no. 4, pp. 657–666, 1996.

C. Grandori, S. M. Cowley, L. P. James, and R. N. Eisenman, “The Myc/Max/Mad network and the transcriptional control of cell behavior,” Annual Review of Cell and Developmental Biology, vol. 16, pp. 653–699, 2000.

D. E. Ayer and R. N. Eisenman, “A switch from Myc:Max to Mad:Max heterocomplexes accompanies monocyte/macrophage differentiation,” Genes and Development, vol. 7, no. 11, pp. 2110–2119, 1993.

D. Xu, N. Popov, M. Hou et al., “Switch from Myc/Max to Mad1/Max binding and decrease in histone acetylation at the telomerase reverse transcriptase promoter during differentiation of HL60 cells,” Proceedings of the National Academy of Sciences of the United States of America, vol. 98, no. 7, pp. 3826–3831, 2001.

M. Conacci-Sorrell, C. Ngouenet, and R. N. Eisenman, “Myc-nick: a cytoplasmic cleavage product of Myc that promotes α-tubulin acetylation and cell differentiation,” Cell, vol. 142, no. 3, pp. 480–493, 2010.

F. Rusnak and P. Mertz, “Calcineurin: form and function,” Physiological Reviews, vol. 80, no. 4, pp. 1483–1521, 2000.

U. Delling, J. Tureckova, H. W. Lim, L. J. de Windt, P. Rotwein, and J. D. Molkentin, “A calcineurin-NFATc3-dependent pathway regulates skeletal muscle differentiation and slow myosin heavy-chain expression,” Molecular and Cellular Biology, vol. 20, no. 17, pp. 6600–6611, 2000.

S. E. Dunn, E. R. Chin, and R. N. Michel, “Matching of calcineurin activity to upstream effectors is critical for skeletal muscle fiber growth,” The Journal of Cell Biology, vol. 151, no. 3, pp. 663–672, 2000.

B. B. Friday, V. Horsley, and G. K. Pavlath, “Calcineurin activity is required for the initiation of skeletal muscle differentiation,” The Journal of Cell Biology, vol. 149, no. 3, pp. 657–666, 2000.

B. B. Friday and G. K. Pavlath, “A calcineurin- and NFAT-dependent pathway regulates Myf5 gene expression in skeletal muscle reserve cells,” Journal of Cell Science, vol. 114, no. 2, pp. 303–310, 2001.

B. B. Friday, P. O. Mitchell, K. M. Kegley, and G. K. Pavlath, “Calcineurin initiates skeletal muscle differentiation by activating MEF2 and MyoD,” Differentiation, vol. 71, no. 3, pp. 217–227, 2003.

M. Oh, I. I. Rybkin, V. Copeland et al., “Calcineurin is necessary for the maintenance but not embryonic development of slow muscle fibers,” Molecular and Cellular Biology, vol. 25, no. 15, pp. 6629–6638, 2005.

B. M. Scicchitano, L. Spath, A. Musarò et al., “Vasopressin-dependent myogenic cell differentiation is mediated by both Ca²⁺/calmodulin-dependent kinase and calcineurin pathways,” Molecular Biology of the Cell, vol. 16, no. 8, pp. 3632–3641, 2005.

A. S. Armand, M. Bourajjaj, S. Martínez-Martínez et al., “Cooperative synergy between NFAT and MyoD regulates myogenin expression and myogenesis,” The Journal of Biological Chemistry, vol. 283, no. 43, pp. 29004–29010, 2008.

N. Bakkar and D. C. Guttridge, “NF-κB signaling: a tale of two pathways in skeletal myogenesis,” Physiological Reviews, vol. 90, no. 2, pp. 495–511, 2010.

T. Wenz, S. G. Rossi, R. L. Rotundo, B. M. Spiegelman, and C. T. Moraes, “Increased muscle PGC-1α expression protects from sarcopenia and metabolic disease during aging,” Proceedings of the National Academy of Sciences of the United States of America, vol. 106, no. 48, pp. 20405–20410, 2009.