BioMed Research International

BioMed Research International / 2014 / Article
Special Issue

Nutrients and Muscle Disease

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Review Article | Open Access

Volume 2014 |Article ID 121254 | 13 pages | https://doi.org/10.1155/2014/121254

Vitamin D Signaling in Myogenesis: Potential for Treatment of Sarcopenia

Academic Editor: Giuseppe D’Antona
Received25 Apr 2014
Accepted03 Jun 2014
Published30 Jun 2014

Abstract

Muscle mass and strength progressively decrease with age, which results in a condition known as sarcopenia. Sarcopenia would lead to physical disability, poor quality of life, and death. Therefore, much is expected of an effective intervention for sarcopenia. Epidemiologic, clinical, and laboratory evidence suggest an effect of vitamin D on muscle function. However, the precise molecular and cellular mechanisms remain to be elucidated. Recent studies suggest that vitamin D receptor (VDR) might be expressed in muscle fibers and vitamin D signaling via VDR plays a role in the regulation of myoblast proliferation and differentiation. Understanding how vitamin D signaling contributes to myogenesis will provide a valuable insight into an effective nutritional strategy to moderate sarcopenia. Here we will summarize the current knowledge about the effect of vitamin D on skeletal muscle and myogenic cells and discuss the potential for treatment of sarcopenia.

1. Introduction

Muscle wasting is observed in various disease states, in conditions of reduced neuromuscular activity, with ageing. Age-related muscle wasting is referred to as “sarcopenia” coined by Irwin H. Rosenberg from the Greek words sarx (meaning flesh) and penia (meaning loss) [1, 2]. There has been no consensus about definition of sarcopenia suitable for use in research and clinical practice [3]. Therefore, some studies [4, 5] suggest a working definition of sarcopenia: sarcopenia is a syndrome characterized by progressive and generalized loss of skeletal muscle mass and strength with a risk of adverse outcomes such as physical disability, poor quality of life, and death. Sarcopenia is characterized by the fact that it progresses very slowly throughout several decades. Muscle mass fairly consistently decreases at a rate of approximately 0.5–1%/year beginning at 40 years of age [6, 7] and the rate dramatically accelerates after the age of 65 years [8]. Muscle strength appears to decline more rapidly than muscle mass. Muscle strength declines at a rate of 3-4% per year in men and 2.5–3% per year in women aged 75 years [9]. Although the precise molecular and cellular mechanisms underlying age-related loss of muscle mass and strength have remained unknown [10, 11], multiple contributing factors have been proposed. The development and progress of sarcopenia have been thought to be mediated by the combination of these contributing factors.

Based on large-scale studies [1216], on average, it is estimated that the prevalence of sarcopenia reaches 5–13% in those aged 60–70 years and ranges from 11 to 50% in those aged over 80 years [17]. In USA in 2000, it was estimated that direct healthcare costs related to sarcopenia were $18.5 billion ($10.8 billion in men, $7.7 billion in women), which represented approximately 1.5% of total healthcare expenditures for that year [18]. Globally, the number of people aged over 60 years is 600 million in the year 2000 [19]. It is predicted that people aged over 65 years will double by 2020 and will triple by 2050 [20]. Therefore, sarcopenia is being recognized as not only a serious healthcare problem but also a social problem. Much is expected of an effective intervention for sarcopenia. Nutritional interventions would be a promising candidate in combating sarcopenia.

Epidemiologic, clinical, and laboratory evidence provide an effect of vitamin D on muscle function. Numerous studies have investigated the effect of vitamin D supplementation on muscle strength and physical performance in elderly people. However, the precise molecular and cellular mechanisms remain to be elucidated. Immunohistochemical studies have demonstrated that vitamin D receptor (VDR) might be localized in human muscle fibers [2123] with some contradictions [24, 25]. In addition, recent studies have reported that vitamin D signaling via VDR plays a role in the regulation of myoblast proliferation and differentiation [2632]. Understanding how vitamin D signaling contributes to myogenesis will provide a valuable insight into an effective nutritional strategy to moderate sarcopenia. Here we will summarize the current knowledge about effect of vitamin D on skeletal muscle and myogenic cells and discuss the potential for treatment of sarcopenia.

2. Vitamin D Signaling Pathways: Genomic and Nongenomic Pathways

Vitamin D signaling has been extensively investigated in a variety of cell types. During the past two decades, considerable progress has been made in understanding the action of 1α,25-dihydroxyvitamin D3 [1α,25(OH)2D3] on myogenic cells. The biological effect of 1α,25(OH)2D3 is exerted through genomic or nongenomic mechanisms (for reviews see [3335]). Better understanding of the molecular and cellular mechanisms of vitamin D action on skeletal muscle will enable us to develop an effective intervention for sarcopenia. We will focus on 1α,25(OH)2D3 signaling via VDR in genomic and nongenomic mechanism related to myogenic cells, although rapid alteration in intracellular calcium, which is nongenomically regulated by 1α,25(OH)2D3, has been well demonstrated both in vivo and in vitro; for details, excellent review article is already available on this subject [36].

An active form, 1α,25(OH)2D3, acts by binding to VDR [33]. The binding affinity of 25(OH)D3 for human vitamin D receptor (VDR) is approximately 500 times less than that of 1α,25(OH)2D3 but the circulating level of 25(OH)D3 is approximately 1000 times higher than that of 1α,25(OH)2D3 [37, 38]. In genomic mechanism, 1α,25(OH)2D3 binds to VDR and is transported to the nucleus [35]. VDR is heterodimerized with 9-cis-retinoic acid receptor (RXR) and VDR:RXR complex modulates gene expression via binding to specific target gene promoter regions, known as vitamin D response elements (VDREs), to activate or suppress their expression [35]. In general, VDREs possess either a direct repeat of two hexanucleotide half-elements with a spacer of three nucleotides (DR3) or an everted repeat of two half-elements with a spacer of six nucleotides (ER6) motif, with DR3s being the most common [39]. Wang et al. [40] investigated direct 1α,25(OH)2D3-target genes on a large scale by using a combined approach of microarray analysis and in silico genome-wide screens for DR3 and ER6-type VDREs. Microarray analyses, performed with RNA from human SCC25 cells treated with 1α,25(OH)2D3 and cycloheximide, an inhibitor of protein synthesis, revealed 913 regulated genes [40]. Of the 913 genes, 734 genes were induced and 179 genes were repressed by treatment of 1α,25(OH)2D3 [40]. In addition, a screening of the mouse genome identified more than 3000 conserved VDREs, and 158 human genes containing conserved elements were 1α,25(OH)2D3-regulated on microarrays [40]. These results support their broad physiological actions of 1α,25(OH)2D3 in a variety of cell types.

With respect to several genes related to myogenesis, we will describe them in more detail. For example, 1α,25(OH)2D3 induced expression of the gene encoding Foxo1 [40], which is a member of the FOXO subfamily of forkhead/winged helix family of transcription factors, governs muscle growth, metabolism, and myoblast differentiation. When transfected C2C12 cells with adenoviral vector encoded a constitutively active Foxo1 mutant, they effectively blocked myoblast differentiation [47]. This was partly rescued by inhibition of Notch signaling [47], which inhibits myoblast differentiation [48]. In addition, loss of Foxo1 function precluded Notch signaling-mediated inhibition of myoblast differentiation [47]. To elucidate the possible role of Notch signaling in Foxo1-mediated inhibition of myoblast differentiation, by combining coculture system, transfection assay, chromatin immunoprecipitation assay, and short interfering RNA (siRNA) technology, authors showed that Foxo1 physically and functionally interacted with Notch by promoting corepressor clearance from DNA binding protein, CSL [CBF1/RBPjk/Su(H)/Lag-1], leading to inhibition of myoblast differentiation through activation of Notch target genes [47]. Another gene, Id (inhibitor of differentiation) gene, is also known target of 1α,25(OH)2D3 [49]. Id mRNA was constitutively expressed in rat osteoblastic osteosarcoma ROS17/2.8 cells and its level was transcriptionally suppressed by 1α,25(OH)2D3 [50]. 1α,25(OH)2D3 exerted its negative effect on Id1 gene transcription via the 57 bp upstream response sequence (−1146/−1090) [49]. Id proteins (Id1, Id2, Id3, and Id4) dimerize and neutralize the transcriptional activity of basic helix-loop-helix (bHLH) proteins [51]. It has been shown that Id inhibits MyoD activity either by forming transcriptionally inactive complexes of MyoD-Id or by forming heterodimers with E-proteins and effectively blocking the formation of active MyoD/E-protein complexes [52]. At this time, there are only limited data available on Id expression and vitamin D during muscle development. For example, in VDR knockout mice with abnormal muscle development, there were no differences in expression levels of Id1 and Id2 [46]. Therefore, we cannot conclude whether vitamin D regulates myogenesis by modulating Id expression.

A nongenomic response to 1α,25(OH)2D3 is characterized by a rapid (the seconds to minutes range) activation of signaling cascades and an insensitivity to inhibitors of transcription and protein synthesis [34]. The rapid response to 1α,25(OH)2D3 has been hypothesized to elicit the classic VDR translocation to the plasma membrane. When treating chick myoblasts with 1α,25(OH)2D3, translocation of VDR from the nucleus to the plasma membrane rapidly occurred within 5 min after the addition of 1α,25(OH)2D3 [53]. This translocation was blocked by colchicine, suggesting the possible role of the intracellular microtubular transport system in the distribution of VDR [53]. The VDR translocation appears to depend on intact caveolae that are specialized plasmalemmal microdomains originally studied in numerous cell types for their involvement in the transcytosis of macromolecules [54]. Confocal microscopy revealed that 1α,25(OH)2D3-induced VDR translocation to the plasma membrane was abolished by methyl-beta-cyclodextrin, a reagent to disrupt the caveolae structure [55]. Both disruption of caveolae and siRNA-mediated silencing of caveolin-1 suppressed 1α,25(OH)2D3-dependent activation of protooncogene c-Src (cellular Src) with tyrosine-specific protein kinase activity [55]. Immunocytochemical analysis provided evidence that caveolin-1 colocalized with c-Src near the plasma membrane under basal conditions [55]. When treated with 1α,25(OH)2D3, the colocalization of caveolin-1 and c-Src was disrupted and they were redistributed into cytoplasm and nucleus [55]. On the basis of these results, it can be hypothesized that (1) interaction caveolin-1/c-Src inactivates the kinase under basal conditions and (2) when 1α,25(OH)2D3 stimulates VDR translocation to the plasma membrane, it dissociates the caveolin-1/c-Src complex allowing c-Src activation [55]. Non-genomic action of 1α,25(OH)2D3 might be required for a reciprocal interaction between c-Src and caveoline-1. Besides the classical VDR, it has been identified as a potential candidate as an alternate membrane-associated receptor for 1α,25(OH)2D3: 1,25D3-MARRS (membrane-associated, rapid response steroid binding) also known as ERp57, GRp58, ERp60, and Pdia3 [56]. Since 1,25D3-MARRS has been shown to function in various cell types [57], it also may potentially mediate vitamin D signaling in myogenic cells.

The c-Src tyrosine kinase induced by 1α,25(OH)2D3 is required for activation of mitogen-activated protein kinases (MAPKs), ERK1/2 (extracellular signal-regulated kinase 1/2) [58], and p38 [59]. 1α,25(OH)2D3 rapidly promoted phosphorylation of ERK1/2 through c-Src activation [58], Raf-1/Ras/MEK (MAPK/ERK kinase), and PKCα (protein kinase C alpha) [60]. In addition to ERK1/2 activation, 1α,25(OH)2D3 rapidly stimulated MKK3/MKK6 (mitogen-activated protein kinase kinases 3/6)/p38 MAPK through c-Src activation [59]. Although another MAPK family member, JNK1/2 (c-Jun NH2-terminal kinase 1/2), was also activated by 1α,25(OH)2D3 [59], an upstream mediator of 1α,25(OH)2D3-dependent JNK1/2 activation was characterized less than that of ERK/1/2 and p38. The molecular links between JNK and c-Src have been shown in Drosophila melanogaster. The JNK homolog Basket (Bsk) is required for epidermal closure [61]. Src42A, a Drosophila c-Src protooncogene homolog functions in epidermal closure during both embryogenesis and metamorphosis [61]. The severity of the epidermal closure defect in the Src42A mutant depended on the Bsk activity. These results suggest the possibility that JNK activation in mammals may also be required for Src tyrosine kinase activity. These MAPK signaling pathways have been shown to contribute to myogenesis [6265]. For example, inactivation of the Raf-1/MEK1/2/ERK1/2 pathway in MM14 cells through the overexpression of dominant negative mutants of Raf-1 blocked ERK1/2 activity and prevented myoblast proliferation [62]. Pharmacological blockade of p38α/β kinases by SB203580 inhibited myoblast differentiation [6365]. JNK was involved in regulating myostatin signaling [66], which is known as a member of tumor growth factor β family and functions as a negative regulator of muscle growth [67]. MAPK signaling pathways function at different stages of myogenesis.

Apart from MAPKs, PI3K (phosphatidyl inositol 3-kinase)/Akt signaling pathway, which is essential for initiation of myoblast differentiation [68], also seems to be activated by 1α,25(OH)2D3. After exposure to 1α,25(OH)2D3, Akt phosphorylation was enhanced through PI3K in C2C12 cells [68]. Intriguingly, suppression of c-Src activity by PP2, a specific inhibitor for all members of the Src family, and knockdown of c-Src expression by siRNA decreased Akt phosphorylation in 1α,25(OH)2D3-treated C2C12 cells [28]. In addition, when treating C2C12 cells with 1α,25(OH)2D3 in the presence of U0126 or SB203580 to inhibit ERK1/2 and p38 MAPK, respectively, SB203580 but not U0126 markedly blocked both basal and 1α,25(OH)2D3-induced Akt phosphorylation. These results suggest that 1α,25(OH)2D3-induced Akt phosphorylation may occur through c-Src and p38 MAPK [28]. Taken together, 1α,25(OH)2D3 can simultaneously activate multiple signaling pathways in myogenic cells but their relative contribution to myogenesis remains to be established.

3. Effects of Ageing on Serum Concentration of Vitamin D, Muscle Morphology, and Muscle Fiber Type

Vitamin D status varies with age [69]. Serum levels of 25(OH)D3 are qualitatively categorized as deficiency (<20 ng/L or <50 nM), insufficiency (21–29 ng/L or 50–75 nM), and normal (30 ng/L or >75 nM) [70]. van der Wielen et al. [69] measured wintertime serum 25(OH)D3 concentrations in 824 elderly people from 11 European countries [69]. They reported that 36% of men and 47% of women had 25(OH)D3 concentrations below 30 nM [69]. Vitamin D deficiency in elderly is thought to occur mainly due to restricted sunlight exposure, reduced dietary vitamin D intake, and decreased capacity of the skin to produce vitamin D [69]. MacLaughlin and Holick [71] examined the effects of ageing on the capacity of the skin to produce previtamin D3 in the skin by comparing young subjects (8 and 18 years old) with aged subjects (77 and 82 years old). They showed that ageing decreased the capacity less than half of young subjects [71], suggesting that elderly people are potentially at risk for vitamin D insufficiency/deficiency.

Vitamin D deficiency appears to be associated with changes in muscle morphology. For example, patients with osteomalacic myopathy associated with vitamin D deficiency show degenerative changes such as opaque fibers, ghost-like necrotic fibers, regenerating fibers, enlarged interfibrillar spaces, infiltration of fat, fibrosis, glycogen granules, and type II muscle fiber atrophy [72]. As is the case with vitamin D-deficient patients, it is well known that elderly people show aberrant muscle morphology. Scelsi et al. [73] performed histochemical and ultrastructure analysis using biopsies taken from the vastus lateralis of healthy sedentary men and women aged 65–89 years. They observed myofibrillar disorganization, streaming of Z-line, rod formation, intracellular lipid droplets, lysosomes, and type II muscle fiber atrophy [73]. The very elderly people had “flattened” or “crushed” shaped muscle fibers, whereas the young people had mature-appearing polygonal muscle fibers [74]. These aberrant changes were much more pronounced in the type II muscle fibers than in type I muscle fibers [74]. Although the precise mechanisms remain to be elucidated, it can be speculated that specific type II muscle fiber atrophy with ageing may be associated with a muscle fiber type-specific reduction in satellite cell content. Satellite cells are essential for normal muscle growth [75]. Verdijk et al. [76] examined whether satellite cells could specifically decrease in type II muscle fibers in the elderly people. Biopsies were taken from the vastus lateralis of elderly (average age: 76 years) and young (average age: 20 years) healthy males [76]. They found significant reduction in the proportion and mean cross-sectional area of the type II muscle fibers and the number of satellite cells per type II muscle fiber in elderly subjects compared to young subjects [76]. This study is the first to show type II muscle fiber atrophy in elderly people to be associated with a muscle fiber type-specific decline in satellite cell content. It remains unknown whether vitamin D supplementation specifically attenuates atrophy of type II muscle fibers with recruitment of satellite cell. Whether vitamin D has positive effects on myoblast proliferation and differentiation is currently under debate. Recent studies [27] suggest that vitamin D treatment enhances fast type (type IIa) MyHC expression in fully differentiated C2C12 myotubes. Type II muscle fibers contain type IIa MyHC [77]. Therefore, vitamin D could potentially contribute to the changes in phenotype of existing muscle fibers and/or the maintenance of type II muscle fibers.

4. Effects of Ageing on Expression of VDR

Bischoff-Ferrari et al. [22] investigated the effect of ageing on VDR expression in human skeletal muscle. Biopsies were taken from the gluteus medius of 20 female patients undergoing total hip arthroplasty (average age: 71.6 years) and from the transversospinalis muscle of 12 female patients with spinal operations (average age: 55.2 years). Immunohistochemical analysis revealed that the number of VDR-positive myonuclei decreased with ageing [22]. Importantly, VDR expression was not affected by 25(OH)D3 or 1α,25(OH)2D3 levels [22]. Buitrago et al. [78] showed that silencing of VDR expression in C2C12 myoblasts suppressed p38 MAPK phosphorylation and decreased ERK1/2 activation induced by 1α,25(OH)2D3. Tanaka et al. [31] demonstrated that knockdown of VDR expression resulted in downregulation of MyHC mRNA in differentiating C2C12 myoblasts when treated with 1α,25(OH)2D3. Therefore, it is possible that decreased expression of VDR observed in elderly people might reduce the functional response of the muscle fibers to 1α,25(OH)2D3.

5. Effects of Vitamin D Supplementation on Muscle Injury

The regenerative potential of skeletal muscle decreases with age [7981]. Satellite cells are absolutely required for muscle regeneration [82]. Satellite cells are defined anatomically by their position beneath the basal lamina and adhered to muscle fibers [83]. They, traditionally considered as a population of skeletal muscle-specific committed progenitors, play a crucial role in the postnatal maintenance, repair, and regeneration [75]. Under normal physiological conditions, they remain in a quiescent and undifferentiated state [75, 84]. However, when skeletal muscle is damaged by unaccustomed exercise or mechanical trauma, they are activated to proliferate, differentiate, and fuse with the already existing muscle fibers or fuse to form new muscle fibers [75, 84]. Few studies have examined the effects of vitamin D treatment on muscle injury. Stratos et al. [85] investigated whether systemically applied vitamin D could restore muscle function and morphology after trauma. Rats were injected subcutaneously with 7-dehydrocholesterol (332,000 IU/kg) immediately after crush injury and muscle samples were collected at days 1, 4, 14, and 42 after injury [85]. Vitamin D treatment increased cell proliferation and inhibited occurrence of apoptosis at day 4 compared to control rats [85]. In addition, a faster recovery of contraction forces was observed at day 42 in vitamin D-treatment group compared to control group [85]. Notably, the number of satellite cells was not influenced by vitamin D [85], suggesting the possibility that vitamin D supplementation has relatively little effect on satellite cell function in vivo. It is necessary to scrutinize thoroughly efficacy, duration, optimal dose, and side effects in relation to vitamin D treatment. Srikuea et al. [29] demonstrated that VDR was highly expressed in the nuclei of regenerating muscle fibers, indicating a potential role for vitamin D in muscle regeneration following injury. Relationship of vitamin D signaling and myogenesis will be discussed below in Section 10.

6. Vitamin D and Type 2 Diabetes Mellitus

Although the incidence of type 2 diabetes mellitus increases with age [86], the precise underlying mechanisms are still not fully understood. Skeletal muscle is the primary target for insulin action and glucose disposal. Therefore, elderly people with excessive loss of muscle mass are at risk for development of type 2 diabetes mellitus [87]. Meta-analysis reveals that vitamin D supplementation has beneficial effects among patients with glucose intolerance or insulin resistance at baseline [88]. However, an explanation for the beneficial role of vitamin D supplementation in the lowering of glycemia in diabetes mellitus remains to be determined. Skeletal muscle can increase glucose uptake through insulin-dependent and muscle contraction-dependent mechanisms [89]. Insulin and muscle contractions stimulate glucose transport in skeletal muscle via translocation of intracellular glucose transporter type 4 (GLUT4) to the cell surface. Manna and Jain [90] examined the mechanism by which vitamin D supplementation regulates glucose metabolism in 3T3L1 adipocytes. When 3T3L1 adipocytes were treated with high glucose in the presence of 1α,25(OH)2D3, it increased expression of GLUT4 and its translocation to cell surface, glucose uptake, and glucose utilization [90]. 1α,25(OH)2D3 also enhanced cystathionine-γ-lyase (CSE) activation and H2S formation [90], which is an important signaling molecule produced mainly by CSE in the cardiovascular system [91]. Furthermore, the effect of 1α,25(OH)2D3 on GLUT4 translocation and glucose utilization was prevented by chemical inhibition or silencing of CSE [90]. In muscle cells, it is currently not known whether CSE may be associated with 1α,25(OH)2D3-induced glucose metabolism. Therefore, further studies are required to elucidate the physiological role of CSE in regulation of glucose metabolism in skeletal muscle. Tamilselvan et al. [92] examined the effect of calcitriol (1,25, dihydroxycholecalciferol) on the expression of VDR, insulin receptor (IR), and GLUT4 in L6 cells when exposed to high glucose and high insulin which mimics type 2 diabetic model [92]. Calcitriol partially restored VDR, IR, and GLUT4 expression in type 2 diabetic model [92], raising the possibility that vitamin D could contribute to improving insulin signaling in type 2 diabetes mellitus. In a recent study, the effect of 1α,25(OH)2D3 on glucose uptake in rat skeletal muscle is investigated [93]. 1α,25(OH)2D3 stimulated glucose uptake with increased expression of GLUT4 protein and enhanced translocation of GLUT4 to the plasma membrane not through PI3K-signaling pathway [93], which is essential for insulin-stimulated GLUT4 translocation and glucose transport [94]. In addition, 1α,25(OH)2D3-stimulated glucose uptake was suppressed concomitantly with downregulation of GLUT4 protein by treatment with cycloheximide [93], suggesting that it may be mediated by genomic signaling of vitamin D. Taken together, vitamin D may improve glucose metabolism in skeletal muscle by modulating GLUT4 expression and translocation through insulin-dependent and/or insulin-independent mechanisms.

7. Vitamin D Receptor Expression in Skeletal Muscle and Myogenic Cells

VDR is known to be expressed in a wide variety of tissues including bone, bronchus, intestine, kidney, mammary gland, pancreas, parathyroid, pituitary gland, prostate gland, spleen, testis, and thymus [25]. However, there has been still some controversy as to whether VDR is expressed in skeletal muscle [24, 29, 32]. For example, some studies have failed to detect VDR in skeletal muscle [24, 25, 9597]; other studies have shown that VDR protein and/or mRNA are detectable in skeletal muscle [2123, 29, 46, 98] and myogenic cells [26, 27, 29, 31, 32, 42, 45, 46, 53, 55, 98104]. In brief, Wang et al. [105] call into question the specificity of various commercially available VDR antibodies. They systematically characterized these antibodies in terms of their specificity and immunosensitivity using negative control samples from VDR knockout mice [105]. They demonstrated that the mouse monoclonal VDR antibody against the C-terminus of human VDR, D-6 (Santa Cruz Biotechnology), possesses high specificity, high sensitivity, and versatility [105]. They showed that VDR protein was not detected in skeletal muscle by immunohistochemical analysis using this antibody and that VDR mRNA was detectable only at extremely low levels by quantitative RT-PCR assay [24]. By contrast, Kislinger et al. [106] used large scale gel-free tandem mass spectrometry to monitor global proteome alterations throughout the myogenic differentiation program in C2C12 cells. They observed upregulation of VDR protein during early stage of myoblast differentiation [106]. Srikuea et al. [29] provided strong evidence for the presence of VDR in myogenic cells, by combining immunoblot assay, immunocytochemical analysis, PCR-based cloning, and DNA sequencing to validate the expression of VDR in C2C12 cells. They showed that the full-length VDR mRNA transcript could be isolated from myoblasts and myotubes and VDR protein was primarily localized in the nucleus of myoblasts and in the cytoplasm of myotubes [29]. In addition, they examined the localization of VDR protein using a model of myogenesis in vivo. BaCl2 treatment was used to induce regeneration and immunohistochemical analysis was performed on sections from control and regenerating muscle. In control muscle, VDR was detected in muscle fibers but levels were very low, whereas in regenerating muscle, VDR expression was detected in the central nuclei of newly regenerating muscle fibers [29]. More recently, Girgis et al. [32] demonstrated that VDR protein was detectable in C2C12 myoblasts by immunoblot assay using VDR antibody (D-6). The discrepancy among studies may be explained, at least in part, by the difference in the expression of VDR during the stages of muscle development. For example, Endo et al. [46] reported that VDR mRNA was detected in skeletal muscle from 3-week-old wild-type mice but not 8-week-old wild-type mice. Wang and DeLuca [24] showed that VDR protein was undetectable in skeletal muscle from 6- to 7-week-old C57BL/6 mice. A similar result was also reported by Srikuea et al. [29] using 12-week-old C57BL/6 mice. Therefore, VDR expression may be dependent on the context of muscle development. It requires further clarification whether VDR is expressed in muscle fibers.

8. The Conversion of 25 (OH)D3 to 1,25(OH)2D3 Might Occur in Myogenic Cells

Vitamin D, in the form of vitamin D3, is synthesized from 7-dehydrocholesterol in the skin through the action of ultraviolet irradiation [33]. Alternatively, vitamin D, in the form of either vitamin D2 or vitamin D3, can also be taken in the diet [33]. An active form, 1α,25(OH)2D3, is synthesized from vitamin D3 through two hydroxylation steps [33]. Vitamin D3 is converted to 25-hydroxyvitamin D3 [25(OH)D3] in the liver by 25-hydroxylases (encoded by the gene CYP27A1) [33]. The generated 25(OH)D3 is further hydroxylated to 1α,25(OH)2D3 by 25-hydroxyvitamin D3 1α-hydroxylase (encoded by the gene CYP27B1) in the kidney [33]. However, CYP27B1 has been detected in various extrarenal tissues [107, 108], raising the possibility that 1α,25(OH)2D3 might be locally synthesized and activate VDR in myogenic cells [29, 32]. Inactive form of vitamin D3, 25(OH)D3, could inhibit cell proliferation in a similar manner to 1α,25(OH)2D3 [29, 32], indicating that the conversion of 25(OH)D3 to 1α,25(OH)2D3 by CYP27B1 occurs in myogenic cells. Girgis et al. [32] confirmed this possibility using luciferase reporter assay system that luciferase activity results from 1α,25(OH)2D3 binding to GAL-4-VDR and subsequent activation of the UASTK luciferase gene via its GAL4 promoter. They transfected C2C12 cells with GAL4-VDR (switch) and UASTK luciferase reporter with treatment of 25(OH)D3 and showed that luciferase activity increased in a dose-dependent manner, suggesting the conversion of 25(OH)D3 to 1α,25(OH)2D3 by CYP27B1 and the subsequent activation of luciferase expression via 1,25(OH)2D-bound GAL4-VDR [32]. Srikuea et al. [29] confirmed that C2C12 cells express the full-length CYP27B1 mRNA transcript and CYP27B1 protein could be detected in the cytoplasm of myoblasts, exhibiting partially overlapping with the mitochondria to which CYP27B1 has been reported to be typically localized [109]. Furthermore, they showed that siRNA-mediated knockdown of CYP27B1 could alleviate inhibitory effects of 25(OH)D3 on cell proliferation [29]. These observations provide direct evidence that CYP27B1 is biologically active in myogenic cells and mediates to convert 25(OH)D3 to 1α,25(OH)2D3. However, it should be noted that the agonistic action of 25(OH)D3 has been demonstrated in cells derived from CYP27B1 knockout mice [110]. Although further studies are needed to elucidate the basic mechanisms, locally synthesized 1α,25(OH)2D3 in myogenic cells might act through autocrine/paracrine mechanisms via VDR.

9. The Role of VDR in Muscle Development

Since the process of myogenesis has been extensively studied both in vivo and in vitro, substantial progress has been made in understanding the molecular and cellular mechanisms. The myogenic regulatory factors, a group of basic helix-loop-helix transcription factors, consisting of MyoD, Myf5, myogenin, and MRF4, play critical roles in myogenesis [111]. MyoD and Myf5 have redundant functions in myoblast specification [112, 113], whereas myogenin [114, 115] and either MyoD or MRF4 [116] are required for differentiation. These myogenic factors can form heterodimers in combination with less specific factors such as members of E12/E47 [117], which are generated by alternative splicing of the E2A gene [118], leading to activation of muscle-specific gene transcription [117].

VDR knockout mice model has provided insight into the possible physiological roles of vitamin D signaling via its receptor in muscle development [46]. VDR null mice recapitulate a human disease of vitamin D resistance, vitamin D-dependent rickets type II [119]. VDR null mice grow normally until weaning and thereafter develop various metabolic abnormalities including hypocalcemia, hypophosphatemia, secondary hyperparathyroidism, and bone deformity [46, 119]. Muscle fiber diameter of VDR null mice was approximately 20% smaller and fiber size was more variable than that of the wild-type mice at 3 weeks of age (before weaning). By 8 weeks of age, these morphological changes were more prominent in the VDR null mice compared to the wild-type mice, suggesting either a progressive nature of the abnormalities caused by the absence of VDR or additive effects of systemic metabolic changes already present at this age [46]. Although there are neither degenerative nor necrotic changes in VDR null mice, the aberrant myofibers were observed diffusely without any preference to type I or type II fibers [46]. Based on these results, they suggest that the absence of VDR induces these abnormalities probably in late stages of fiber maturation and/or in metabolism of mature muscle fibers. Tanaka et al. [31] showed that siRNA-mediated knockdown of VDR inhibited myotube formation concomitantly with downregulation of MyoD and myogenin using C2C12 and G8 cells. These results demonstrate that a substantial level of signaling via VDR is required for normal muscle development and myogenesis in vitro.

Furthermore, Myf5, myogenin, and E2A but not MyoD and MRF4 were aberrantly and persistently upregulated at the protein and/or mRNA levels in VDR null mice at 3 weeks of age [46]. Consistent with the deregulated expression of MRFs that control muscle phenotype, VDR null mice showed aberrantly increased expression of embryonic and neonatal MyHC isoforms but not type II (adult fast twitch) MyHC isoform [46]. These findings observed in VDR null mice may reflect compensatory response to a reduction in muscle fiber size. For example, it can be hypothesized that, in VDR null mice, satellite cells may be anomalously activated, proliferate, and differentiate to form new myonuclei that fuse with existing fibers to restore normal fiber size. Finally, they examined whether 1α,25(OH)2D3 could directly downregulate MRFs and neonatal MyHC gene expression in C2C12 myoblasts. 1α,25(OH)2D3 (10 nM) decreased the steady-state expression levels of these genes [46]. Overall, these results support a role of VDR in the regulation of muscle development, but the precise mechanisms remain to be elucidated and the interpretation is further complicated since negative vitamin D response elements [120122] in the promoter region of genes encoding Myf5 and myogenin have not been identified.

10. Effects of 1,25(OH)2D3 on Myoblast Proliferation and Differentiation

As referred to above, decline of intrinsic regenerative potential of skeletal muscle is a hallmark of ageing [7981] and may be due to age-related changes in satellite cell function. If vitamin D treatment does lead to improvements in muscle function in elderly people, more attention should be directed to the effect of vitamin D3 on myoblast proliferation and differentiation. Research on effect of 1α,25(OH)2D3 on myogenesis has been performed using an in vitro cell culture system. The effects of 1α,25(OH)2D3 on myoblast proliferation and differentiation are summarized in Table 1. Early studies [41, 43] have reported that 1α,25(OH)2D3 stimulates proliferation of myogenic cells. Giuliani and Boland [41] reported that 1α,25(OH)2D3 (0.13 nM) increased cell density of chick myoblasts. Drittanti et al. [43] showed that 1α,25(OH)2D3 (0.1 nM) had biphasic effects on DNA synthesis. 1α,25(OH)2D3 exhibited a mitogenic effect in proliferating chick myoblasts followed by an inhibitory effect during the subsequent stage of myoblast differentiation. Capiati et al. [44] showed that 1α,25(OH)2D3 (1 nM) increases the rate of [3H] thymidine incorporation into DNA in chick myoblasts. In addition, they investigated the role of PKC in mediating the effect of 1α,25(OH)2D3 using a PKC inhibitor. PKC activity increased after treatment of 1α,25(OH)2D3 [44]. The specific PKC inhibitor, calphostin, suppressed 1α,25(OH)2D3 stimulation of DNA synthesis in proliferating myoblasts [44]. Finally, they examined 1α,25(OH)2D3-dependent changes in the expression of PKC isoforms α, β, δ, ε, and ζ [44]. They identified PKCα as main isoform correlated with the early stimulation of myoblast proliferation by 1α,25(OH)2D3 [44]. By contrast, several studies suggest that, overall, 1α,25(OH)2D3 or 25(OH)D3 appears to have antiproliferative effect on myogenic cells [26, 27, 29, 32, 42]. 1α,25(OH)2D3 (1–100 nM) inhibited proliferation of C2C12 myoblasts in a dose-dependent manner [27, 32] without inducing necrotic and apoptotic cell death [32]. Okuno et al. [27] showed that 1α,25(OH)2D3 arrested the cells in the G0/G1 phase concomitantly with induction of cyclin-dependent kinase (CDK) inhibitors, that facilitates cell cycle withdrawal [123] and that inhibits a wide range of CDKs essential for cell cycle progression [124]. Girgis et al. [32] also reported the increased expression of genes involved in G0/G1 arrest including Rb (retinoblastoma protein) and ATM (ataxia telangiectasia mutated) and decreased expression of genes involved in G1/S transition, including c-myc (cellular myc) and cyclin-D1. In addition, they found reduced c-myc protein and hypophosphorylated Rb protein [32]. The active form, hypophosphorylated Rb, blocks entry into S-phase by inhibiting the E2F transcriptional program [125, 126]. In summary, the effects of 1α,25(OH)2D3 on myoblast proliferation remain inconclusive. The discrepancy may be due to the differences in the experimental settings. For example, different cell type (primary cells or immortalized cell lines): 1α,25(OH)2D3 concentration, serum concentration, duration of cell culture, and duration of treatment are employed. Further studies are needed to clarify the role of 1α,25(OH)2D3 on myoblast proliferation.


Muscle cell typeConcentration  
1 ,25(OH)2D3
ProliferationDifferentiationMethod of VDR detectionReference

Myoblast 
(chick)
0.13 nMNIGiuliani and Boland 1984 [41]
G83–300 nMNIEquilibrium binding assay, chromatographySimpson et al., 1985 [42]
Myoblast 
(chick)
0.1 nMNIDrittanti et al., 1989 [43]
Myoblast 
(chick)
1 nMNICapiati et al., 1999 [44]
C2C121 nMNDNIImmunoblotStio et al., 2002 [45]
C2C1210 nMNINDRT-PCREndo et al., 2003 [46]
C2C12100 nMRT-PCR, immunoblot, and immunocytochemistryGarcia et al., 2011 [26]
C2C121–100 nMRT-PCROkuno et al., 2012 [27]
C2C121 nM NIBuitrago et al., 2012 [28]
C2C1220 nM 
2  M 25(OH)D3
RT-PCR, PCR cloning, DNA sequencing, immunocytochemistry, and immunoblotSrikuea et al., 2012 [29]
C2C120.1 pM–10  MNINIRyan et al., 2013 [30]
C2C12, G81–100 nMNIRT-PCRTanaka et al., 2013 [31]
C2C121–100 nM 
1–100 nM 25(OH)D3
RT-PCR, immunoblotGirgis et al., 2014 [32]

Promote (↑); inhibit (↓); no difference between vehicle and treatment (ND); not investigated (NI).

Some studies [43, 44] reported that 1α,25(OH)2D3 (0.1 or 1 nM) had inhibitory effects on DNA synthesis in differentiating chick myoblasts, with an increased MyHC expression, increased myofibrillar and microsomal protein synthesis, and an elevation of creatine kinase activity. Garcia et al. [26] reported that prolonged treatment of C2C12 myoblasts with 1α,25(OH)2D3 (100 nM) enhanced myoblast differentiation by inhibiting cell proliferation and modulating the expression of promyogenic and antimyogenic growth factors using a culture system without reducing serum concentration to initiate cell differentiation. They showed that 1α,25(OH)2D3 downregulated insulin-like growth factor-I (IGF-I) and myostatin expression and upregulates IGF-II and follistatin expression [26]. Follistatin antagonizes myostatin-mediated inhibition of myogenesis [127]. Intriguingly, inhibition of myostatin is characterized by increased expression of IGF-1 and IGF-II [128133], which are known to be potent stimulus of myogenesis [134, 135]. Therefore, it can be hypothesized that 1α,25(OH)2D3 may contribute to myogenesis by inducing IGF-II expression through modulation of myostatin-follistatin system. It should be noted, however, that in these culture conditions, only small thin myotubes with few nuclei were observed on day 10 [26]. This may not recapitulate normal C2C12 myoblast differentiation as previously reported [136].

In general, C2C12 myoblasts normally proliferate and are mononucleated when kept subconfluently in high-mitogen medium (e.g., 10–20% fetal bovine serum). To initiate cell cycle exit and myogenic differentiation, by switching from high-mitogen medium to low-mitogen medium (e.g., 2% horse serum), they fuse and differentiate into postmitotic, elongated, and multinucleated myotubes. Using this C2C12 myoblast differentiation system, Buitrago et al. [28] showed that 1α,25(OH)2D3 (1 nM) enhanced the expression of MyHC and myogenin at 72 h after treatment. By contrast, Okuno et al. [27] investigated the effects of 1α,25(OH)2D3 (1–100 nM) on differentiating and differentiated stage of C2C12 myoblasts. In differentiating phase, 1α,25(OH)2D3 treatment downregulated the expression of neonatal myosin heavy chain and myogenin and inhibited myotube formation in a dose-dependent manner (1–100 nM) [27]. They showed that the expression of fast MyHC isoform increased when fully differentiated myotubes were treated with 1 and 10 nM 1α,25(OH)2D3 [27]. Girgis et al. [32] investigated the prolonged treatment of 1α,25(OH)2D3 (100 nM) on C2C12 myoblast differentiation. When myoblast was treated with 1α,25(OH)2D3 throughout myogenesis including proliferative, differentiating, and differentiated stages, myotube formation was delayed by day 10 concomitantly with downregulation of Myf5 and myogenin [32]. However, intriguingly, myotubes treated with 1α,25(OH)2D3 exhibit larger cell size than nontreated myotubes [32]. These results suggest that 1α,25(OH)2D3 may biphasically act in the process of early and late myoblast differentiation. Furthermore, they showed that the hypertrophic effect of 1α,25(OH)2D3 on myotubes is accompanied with downregulation of myostatin [32]. Several studies have provided evidence that myostatin acts as a negative regulator of the Akt/mammalian target of rapamycin (mTOR) signaling pathway [137140], which plays a key role in the regulation of protein synthesis [141]. For example, Trendelenburg et al. [139] show that myostatin reduces Akt/mTOR signaling complex 1 (TORC1)/p70 S6 kinase (p70S6K) signaling, inhibiting myoblast differentiation and reducing myotube size. In addition, 1α,25(OH)2D3 induced Akt phosphorylation in differentiating C2C12 cells [28]. Intriguingly, 1α,25(OH)2D3 sensitizes the Akt/mTOR signaling pathway to the stimulating effect of leucine and insulin, resulting in a further activation of protein synthesis in C2C12 myotubes [104]. Taken together, 1α,25(OH)2D3 may have an anabolic effect on myotubes by modulating Akt/mTOR signaling probably through genomic and nongenomic mechanisms.

11. Conclusions

The randomized-controlled studies and meta-analysis support a role of vitamin D in improving the age-related decline in muscle function. However, the effect remains inconclusive. Girgis et al. [32] emphasize that large studies employing standardized, reproducible assessments of muscle strength and double-blinded treatment regimens are required to identify the effect of vitamin D supplementation on muscle function and guide the recommended level of vitamin D intake. Although it remains intensely debated whether VDR is expressed in skeletal muscle, research on VDR null mice provides insight into the physiological roles of vitamin D in muscle development and suggests that a substantial level of signaling via VDR is required for normal muscle growth. VDR expression seems to be affected by ageing, suggesting that this might reduce the functional response of the muscle fibers to vitamin D. Vitamin D appears to function in primary myoblasts and established myoblast cell lines. Despite limited evidence available at the time, vitamin D might have an anabolic effect on myotubes by modulating multiple intracellular signaling pathways probably through genomic and nongenomic mechanisms. However, not all studies support this result. Further studies on the potential impact of vitamin D on muscle morphology and function are required to develop the effective intervention for sarcopenia.

Conflict of Interests

The authors declare that there is no conflict of interests regarding the publication of this paper.

Acknowledgment

This research was supported by MEXT (the Ministry of Education, Culture, Sports, Science and Technology) (Grant in Aid for Scientific Research (C), 25350882), Japan.

References

  1. I. H. Rosenberg, “Summary comments: epidemiological and methodological problems in determining nutritional status of older persons,” The American Journal of Clinical Nutrition, vol. 50, no. 5, pp. 1231–1233, 1989. View at: Google Scholar
  2. I. H. Rosenberg, “Sarcopenia: origins and clinical relevance,” Journal of Nutrition, vol. 127, no. 5, supplement, pp. 990S–991S, 1997. View at: Google Scholar
  3. A. J. Cruz-Jentoft, J. P. Baeyens, J. M. Bauer et al., “Sarcopenia: European consensus on definition and diagnosis: report of the European Working Group on Sarcopenia in Older People,” Age Ageing, vol. 39, no. 4, pp. 412–423, 2010. View at: Google Scholar
  4. B. H. Goodpaster, S. W. Park, T. B. Harris et al., “The loss of skeletal muscle strength, mass, and quality in older adults: The Health, Aging and Body Composition Study,” The Journals of Gerontology A; Biological Sciences and Medical Sciences, vol. 61, no. 10, pp. 1059–1064, 2006. View at: Google Scholar
  5. M. J. Delmonico, T. B. Harris, J.-S. Lee et al., “Alternative definitions of sarcopenia, lower extremity performance, and functional impairment with aging in older men and women,” Journal of the American Geriatrics Society, vol. 55, no. 5, pp. 769–774, 2007. View at: Publisher Site | Google Scholar
  6. J. F. Aloia, D. M. McGowan, A. N. Vaswani, P. Ross, and S. H. Cohn, “Relationship of menopause to skeletal and muscle mass,” The American Journal of Clinical Nutrition, vol. 53, no. 6, pp. 1378–1383, 1991. View at: Google Scholar
  7. V. A. Hughes, W. R. Frontera, R. Roubenoff, W. J. Evans, and M. A. Fiatarone Singh, “Longitudinal changes in body composition in older men and women: role of body weight change and physical activity,” The American Journal of Clinical Nutrition, vol. 76, no. 2, pp. 473–481, 2002. View at: Google Scholar
  8. K. S. Nair, “Aging muscle,” The American Journal of Clinical Nutrition, vol. 81, no. 5, pp. 953–963, 2005. View at: Google Scholar
  9. W. K. Mitchell, J. Williams, P. Atherton, M. Larvin, J. Lund, and M. Narici, “Sarcopenia, dynapenia, and the impact of advancing age on human skeletal muscle size and strength; a quantitative review,” Frontiers in Physiology, vol. 3, article 260, 2012. View at: Publisher Site | Google Scholar
  10. H. Augustin and L. Partridge, “Invertebrate models of age-related muscle degeneration,” Biochimica et Biophysica Acta—General Subjects, vol. 1790, no. 10, pp. 1084–1094, 2009. View at: Publisher Site | Google Scholar
  11. F. Demontis, R. Piccirillo, A. L. Goldberg, and N. Perrimon, “Mechanisms of skeletal muscle aging: insights from Drosophila and mammalian models,” Disease Models & Mechanisms, vol. 6, no. 6, pp. 1339–1352, 2013. View at: Google Scholar
  12. R. N. Baumgartner, K. M. Koehler, and D. Gallagher, “Epidemiology of sarcopenia among the elderly in New Mexico,” American Journal of Epidemiology, vol. 147, no. 8, pp. 755–763, 1998. View at: Google Scholar
  13. I. Janssen, S. B. Heymsfield, and R. Ross, “Low relative skeletal muscle mass (sarcopenia) in older persons is associated with functional impairment and physical disability,” Journal of the American Geriatrics Society, vol. 50, no. 5, pp. 889–896, 2002. View at: Publisher Site | Google Scholar
  14. F. Lauretani, C. R. Russo, S. Bandinelli et al., “Age-associated changes in skeletal muscles and their effect on mobility: an operational diagnosis of sarcopenia,” Journal of Applied Physiology, vol. 95, no. 5, pp. 1851–1860, 2003. View at: Google Scholar
  15. Y. Rolland, V. Lauwers-Cances, M. Cournot et al., “Sarcopenia, calf circumference, and physical function of elderly women: a cross-sectional study,” Journal of the American Geriatrics Society, vol. 51, no. 8, pp. 1120–1124, 2003. View at: Publisher Site | Google Scholar
  16. I. Janssen, “Influence of sarcopenia on the development of physical disability: the cardiovascular health study,” Journal of the American Geriatrics Society, vol. 54, no. 1, pp. 56–62, 2006. View at: Publisher Site | Google Scholar
  17. S. von Haehling, J. E. Morley, and S. D. Anker, “An overview of sarcopenia: facts and numbers on prevalence and clinical impact,” Journal of Cachexia, Sarcopenia and Muscle, vol. 1, no. 2, pp. 129–133, 2010. View at: Publisher Site | Google Scholar
  18. I. Janssen, D. S. Shepard, P. T. Katzmarzyk, and R. Roubenoff, “The healthcare costs of sarcopenia in the United States,” Journal of the American Geriatrics Society, vol. 52, no. 1, pp. 80–85, 2004. View at: Publisher Site | Google Scholar
  19. “What Works: Healing the Healthcare Staffing Shortage,” Pricewaterhouse-Coopers Health Research Institute, 2007. View at: Google Scholar
  20. World Population Ageing 1950–2050, Department of Economic and Social Affairs, United Nations, 2001, http://www.un.org/esa/population/publications/worldageing19502050/.
  21. H. A. Bischoff, M. Borchers, F. Gudat et al., “In situ detection of 1,25-dihydroxyvitamin D3 receptor in human skeletal muscle tissue,” The Histochemical Journal, vol. 33, no. 1, pp. 19–24, 2001. View at: Publisher Site | Google Scholar
  22. H. A. Bischoff-Ferrari, M. Borchers, F. Gudat, U. Dürmüller, and W. Dick, “Vitamin D receptor expression in human muscle tissue decreases with age,” Journal of Bone and Mineral Research, vol. 19, no. 2, pp. 265–269, 2004. View at: Publisher Site | Google Scholar
  23. L. Ceglia, M. da Silva Morais, L. K. Park et al., “Multi-step immunofluorescent analysis of vitamin D receptor loci and myosin heavy chain isoforms in human skeletal muscle,” Journal of Molecular Histology, vol. 41, no. 2-3, pp. 137–142, 2010. View at: Publisher Site | Google Scholar
  24. Y. Wang and H. F. DeLuca, “Is the vitamin D receptor found in muscle?” Endocrinology, vol. 152, no. 2, pp. 354–363, 2011. View at: Publisher Site | Google Scholar
  25. Y. Wang, J. Zhu, and H. F. DeLuca, “Where is the vitamin D receptor?” Archives of Biochemistry and Biophysics, vol. 523, no. 1, pp. 123–133, 2012. View at: Publisher Site | Google Scholar
  26. L. A. Garcia, K. K. King, M. G. Ferrini, K. C. Norris, and J. N. Artaza, “1,25(OH)2 vitamin D3 stimulates myogenic differentiation by inhibiting cell proliferation and modulating the expression of promyogenic growth factors and myostatin in C2C12 skeletal muscle cells,” Endocrinology, vol. 152, no. 8, pp. 2976–2986, 2011. View at: Publisher Site | Google Scholar
  27. H. Okuno, K. N. Kishimoto, M. Hatori, and E. Itoi, “1α,25-dihydroxyvitamin D3 enhances fast-myosin heavy chain expression in differentiated C2C12 myoblasts,” Cell Biology International, vol. 36, no. 5, pp. 441–447, 2012. View at: Publisher Site | Google Scholar
  28. C. G. Buitrago, N. S. Arango, and R. L. Boland, “1α,25(OH)2D3-dependent modulation of Akt in proliferating and differentiating C2C12 skeletal muscle cells,” Journal of Cellular Biochemistry, vol. 113, no. 4, pp. 1170–1181, 2012. View at: Publisher Site | Google Scholar
  29. R. Srikuea, X. Zhang, O.-K. Park-Sarge, and K. A. Esser, “VDR and CYP27B1 are expressed in C2C12 cells and regenerating skeletal muscle: potential role in suppression of myoblast proliferation,” American Journal of Physiology—Cell Physiology, vol. 303, no. 4, pp. C396–C405, 2012. View at: Publisher Site | Google Scholar
  30. K. J. Ryan, Z. C. Daniel, L. J. Craggs, T. Parr, and J. M. Brameld, “Dose-dependent effects of vitamin D on transdifferentiation of skeletal muscle cells to adipose cells,” Journal of Endocrinology, vol. 217, no. 1, pp. 45–58, 2013. View at: Publisher Site | Google Scholar
  31. M. Tanaka, K. N. Kishimoto, H. Okuno, H. Saito, and E. Itoi, “Vitamin D receptor gene silencing effects on differentiation of myogenic cell lines,” Muscle & Nerve, vol. 49, no. 5, pp. 700–708, 2014. View at: Google Scholar
  32. C. M. Girgis, R. J. Clifton-Bligh, N. Mokbel, K. Cheng, and J. E. Gunton, “Vitamin D signaling regulates proliferation, differentiation, and myotube size in C2C12 skeletal muscle cells,” Endocrinology, vol. 155, no. 2, pp. 347–357, 2014. View at: Google Scholar
  33. G. Jones, S. A. Strugnell, and H. F. DeLuca, “Current understanding of the molecular actions of vitamin D,” Physiological Reviews, vol. 78, no. 4, pp. 1193–1231, 1998. View at: Google Scholar
  34. R. Lösel and M. Wehling, “Nongenomic actions of steroid hormones,” Nature Reviews Molecular Cell Biology, vol. 4, no. 1, pp. 46–56, 2003. View at: Publisher Site | Google Scholar
  35. K. K. Deeb, D. L. Trump, and C. S. Johnson, “Vitamin D signalling pathways in cancer: potential for anticancer therapeutics,” Nature Reviews Cancer, vol. 7, no. 9, pp. 684–700, 2007. View at: Publisher Site | Google Scholar
  36. C. M. Girgis, R. J. Clifton-Bligh, M. W. Hamrick, M. F. Holick, and J. E. Gunton, “The roles of vitamin D in skeletal muscle: form, function, and metabolism,” Endocrine Reviews, vol. 34, no. 1, pp. 33–83, 2013. View at: Publisher Site | Google Scholar
  37. R. Bouillon, W. H. Okamura, and A. W. Norman, “Structure-function relationships in the vitamin D endocrine system,” Endocrine Reviews, vol. 16, no. 2, pp. 200–257, 1995. View at: Google Scholar
  38. Y.-R. Lou, S. Qiao, R. Talonpoika, H. Syvälä, and P. Tuohimaa, “The role of vitamin D3 metabolism in prostate cancer,” The Journal of Steroid Biochemistry and Molecular Biology, vol. 92, no. 4, pp. 317–325, 2004. View at: Publisher Site | Google Scholar
  39. M. R. Haussler, P. W. Jurutka, M. Mizwicki, and A. W. Norman, “Vitamin D receptor (VDR)-mediated actions of 1α,25(OH)2 vitamin D3: genomic and non-genomic mechanisms,” Best Practice & Research: Clinical Endocrinology and Metabolism, vol. 25, no. 4, pp. 543–559, 2011. View at: Publisher Site | Google Scholar
  40. T.-T. Wang, L. E. Tavera-Mendoza, D. Laperriere et al., “Large-scale in silico and microarray-based identification of direct 1,25-dihydroxyvitamin D3 target genes,” Molecular Endocrinology, vol. 19, no. 11, pp. 2685–2695, 2005. View at: Publisher Site | Google Scholar
  41. D. L. Giuliani and R. L. Boland, “Effects of vitamin D3 metabolites on calcium fluxes in intact chicken skeletal muscle and myoblasts cultured in vitro,” Calcified Tissue International, vol. 36, no. 2, pp. 200–205, 1984. View at: Google Scholar
  42. R. U. Simpson, G. A. Thomas, and A. J. Arnold, “Identification of 1,25-dihydroxyvitamin D3 receptors and activities in muscle,” The Journal of Biological Chemistry, vol. 260, no. 15, pp. 8882–8891, 1985. View at: Google Scholar
  43. L. Drittanti, A. R. de Boland, and R. Boland, “Modulation of DNA synthesis in cultured muscle cells by 1,25-dihydroxyvitamin D-3,” Biochimica et Biophysica Acta—Molecular Cell Research, vol. 1014, no. 2, pp. 112–119, 1989. View at: Google Scholar
  44. D. A. Capiati, M. T. Téllez-Iñón, and R. L. Boland, “Participation of protein kinase C α in 1,25-dihydroxy-vitamin D3 regulation of chick myoblast proliferation and differentiation,” Molecular and Cellular Endocrinology, vol. 153, no. 1-2, pp. 39–45, 1999. View at: Publisher Site | Google Scholar
  45. M. Stio, A. Celli, and C. Treves, “Synergistic effect of vitamin D derivatives and retinoids on C2C12 skeletal muscle cells,” IUBMB Life, vol. 53, no. 3, pp. 175–181, 2002. View at: Publisher Site | Google Scholar
  46. I. Endo, D. Inoue, T. Mitsui et al., “Deletion of vitamin D receptor gene in mice results in abnormal skeletal muscle development with deregulated expression of myoregulatory transcription factors,” Endocrinology, vol. 144, no. 12, pp. 5138–5144, 2003. View at: Publisher Site | Google Scholar
  47. T. Kitamura, Y. I. Kitamura, Y. Funahashi et al., “A Foxo/Notch pathway controls myogenic differentiation and fiber type specification,” The Journal of Clinical Investigation, vol. 117, no. 9, pp. 2477–2485, 2007. View at: Publisher Site | Google Scholar
  48. M. F. Buas and T. Kadesch, “Regulation of skeletal myogenesis by Notch,” Experimental Cell Research, vol. 316, no. 18, pp. 3028–3033, 2010. View at: Publisher Site | Google Scholar
  49. Y. Ezura, O. Tournay, A. Nifuji, and M. Noda, “Identification of a novel suppressive vitamin D response sequence in the 5′-Flanking region of the murine Id1 gene,” The Journal of Biological Chemistry, vol. 272, no. 47, pp. 29865–29872, 1997. View at: Publisher Site | Google Scholar
  50. N. Kawaguchi, H. F. Deluca, and M. Noda, “Id gene expression and its suppression by 1,25-dihydroxyvitamin D3 in rat osteoblastic osteosarcoma cells,” Proceedings of the National Academy of Sciences of the United States of America, vol. 89, no. 10, pp. 4569–4572, 1992. View at: Google Scholar
  51. A. Lasorella, R. Benezra, and A. Iavarone, “The ID proteins: master regulators of cancer stem cells and tumour aggressiveness,” Nature Reviews. Cancer, vol. 14, no. 2, pp. 77–91, 2014. View at: Google Scholar
  52. Y. Jen, H. Weintraub, and R. Benezra, “Overexpression of Id protein inhibits the muscle differentiation program: in vivo association of Id with E2A proteins,” Genes & Development, vol. 6, no. 8, pp. 1466–1479, 1992. View at: Google Scholar
  53. D. Capiati, S. Benassati, and R. L. Boland, “1,25(OH)2-vitamin D3 induces translocation of the vitamin D receptor (VDR) to the plasma membrane in skeletal muscle cells,” Journal of Cellular Biochemistry, vol. 86, no. 1, pp. 128–135, 2002. View at: Publisher Site | Google Scholar
  54. P. W. Shaul and R. G. Anderson, “Role of plasmalemmal caveolae in signal transduction,” American Journal of Physiology—Lung Cellular and Molecular Physiology, vol. 275, no. 5, pp. L843–L851, 1998. View at: Google Scholar
  55. C. Buitrago and R. Boland, “Caveolae and caveolin-1 are implicated in 1α,25(OH)2-vitamin D3-dependent modulation of Src, MAPK cascades and VDR localization in skeletal muscle cells,” The Journal of Steroid Biochemistry and Molecular Biology, vol. 121, no. 1-2, pp. 169–175, 2010. View at: Publisher Site | Google Scholar
  56. I. Nemere, M. C. Farach-Carson, B. Rohe et al., “Ribozyme knockdown functionally links a 1,25(OH)2D3 membrane binding protein (1,25D3-MARRS) and phosphate uptake in intestinal cells,” Proceedings of the National Academy of Sciences of the United States of America, vol. 101, no. 19, pp. 7392–7397, 2004. View at: Publisher Site | Google Scholar
  57. R. C. Khanal and I. Nemere, “The ERp57/GRp58/1,25D3-MARRS receptor: multiple functional roles in diverse cell systems,” Current Medicinal Chemistry, vol. 14, no. 10, pp. 1087–1093, 2007. View at: Publisher Site | Google Scholar
  58. C. Buitrago, R. Boland, and A. R. de Boland, “The tyrosine kinase c-Src is required for 1,25(OH)2-vitamin D3 signalling to the nucleus in muscle cells,” Biochimica et Biophysica Acta—Molecular Cell Research, vol. 1541, no. 3, pp. 179–187, 2001. View at: Publisher Site | Google Scholar
  59. C. G. Buitrago, A. C. Ronda, A. R. de Boland, and R. Boland, “MAP kinases p38 and JNK are activated by the steroid hormone 1α,25(OH)2-vitamin D3 in the C2C12 muscle cell line,” Journal of Cellular Biochemistry, vol. 97, no. 4, pp. 698–708, 2006. View at: Publisher Site | Google Scholar
  60. C. G. Buitrago, V. González Pardo, A. R. de Boland, and R. Boland, “Activation of Raf-1 through Ras and protein kinase Cα mediates 1α,25(OH)2-vitamin D3 regulation of the mitogen-activated protein kinase pathway in muscle cells,” The Journal of Biological Chemistry, vol. 278, no. 4, pp. 2199–2205, 2003. View at: Publisher Site | Google Scholar
  61. M. Tateno, Y. Nishida, and T. Adachi-Yamada, “Regulation of JNK by Src during Drosophila development,” Science, vol. 287, no. 5451, pp. 324–327, 2000. View at: Publisher Site | Google Scholar
  62. N. C. Jones, Y. V. Fedorov, R. S. Rosenthal, and B. B. Olwin, “ERK1/2 is required for myoblast proliferation but is dispensable for muscle gene expression and cell fusion,” Journal of Cellular Physiology, vol. 186, no. 1, pp. 104–115, 2001. View at: Google Scholar
  63. A. Cuenda and P. Cohen, “Stress-activated protein kinase-2/p38 and a rapamycin-sensitive pathway are required for C2C12 myogenesis,” The Journal of Biological Chemistry, vol. 274, no. 7, pp. 4341–4346, 1999. View at: Publisher Site | Google Scholar
  64. A. Zetser, E. Gredinger, and E. Bengal, “p38 mitogen-activated protein kinase pathway promotes skeletal muscle differentiation: participation of the MEF2C transcription factor,” The Journal of Biological Chemistry, vol. 274, no. 8, pp. 5193–5200, 1999. View at: Publisher Site | Google Scholar
  65. Z. Wu, P. J. Woodring, K. S. Bhakta et al., “p38 and extracellular signal-regulated kinases regulate the myogenic program at multiple steps,” Molecular and Cellular Biology, vol. 20, no. 11, pp. 3951–3964, 2000. View at: Publisher Site | Google Scholar
  66. Z. Huang, D. Chen, K. Zhang, B. Yu, X. Chen, and J. Meng, “Regulation of myostatin signaling by c-Jun N-terminal kinase in C2C12 cells,” Cellular Signalling, vol. 19, no. 11, pp. 2286–2295, 2007. View at: Publisher Site | Google Scholar
  67. A. C. McPherron, A. M. Lawler, and S.-J. Lee, “Regulation of skeletal muscle mass in mice by a new TGF-β superfamily member,” Nature, vol. 387, no. 6628, pp. 83–90, 1997. View at: Google Scholar
  68. E. M. Wilson and P. Rotwein, “Selective control of skeletal muscle differentiation by Akt1,” The Journal of Biological Chemistry, vol. 282, no. 8, pp. 5106–5110, 2007. View at: Publisher Site | Google Scholar
  69. R. P. van der Wielen, M. R. Lowik, H. van dan Berg et al., “Serum vitamin D concentrations among elderly people in Europe,” The Lancet, vol. 346, no. 8969, pp. 207–210, 1995. View at: Google Scholar
  70. M. F. Holick, “Vitamin D deficiency,” The New England Journal of Medicine, vol. 357, no. 3, pp. 266–281, 2007. View at: Publisher Site | Google Scholar
  71. J. MacLaughlin and M. F. Holick, “Aging decreases the capacity of human skin to produce vitamin D3,” The Journal of Clinical Investigation, vol. 76, no. 4, pp. 1536–1538, 1985. View at: Google Scholar
  72. S. Yoshikawa, T. Nakamura, H. Tanabe, and T. Imamura, “Osteomalacic myopathy,” Endocrinologia Japonica, vol. 26, supplement, pp. 65–72, 1979. View at: Google Scholar
  73. R. Scelsi, C. Marchetti, and P. Poggi, “Histochemical and ultrastructural aspects of M. vastus lateralis in sedentary old people (age 65–89 years),” Acta Neuropathologica, vol. 51, no. 2, pp. 99–105, 1980. View at: Google Scholar
  74. J. L. Andersen, “Muscle fibre type adaptation in the elderly human muscle,” Scandinavian Journal of Medicine & Science in Sports, vol. 13, no. 1, pp. 40–47, 2003. View at: Publisher Site | Google Scholar
  75. S. B. P. Chargé and M. A. Rudnicki, “Cellular and molecular regulation of muscle regeneration,” Physiological Reviews, vol. 84, no. 1, pp. 209–238, 2004. View at: Publisher Site | Google Scholar
  76. L. B. Verdijk, R. Koopman, G. Schaart, K. Meijer, H. H. C. M. Savelberg, and L. J. C. Van Loon, “Satellite cell content is specifically reduced in type II skeletal muscle fibers in the elderly,” American Journal of Physiology—Endocrinology and Metabolism, vol. 292, no. 1, pp. E151–E157, 2007. View at: Publisher Site | Google Scholar
  77. W. Scott, J. Stevens, and S. A. Binder-Macleod, “Human skeletal muscle fiber type classifications,” Physical Therapy, vol. 81, no. 11, pp. 1810–1816, 2001. View at: Google Scholar
  78. C. Buitrago, V. G. Pardo, and R. Boland, “Role of VDR in 1α,25-dihydroxyvitamin D3-dependent non-genomic activation of MAPKs, Src and Akt in skeletal muscle cells,” The Journal of Steroid Biochemistry and Molecular Biology, vol. 136, no. 1, pp. 125–130, 2013. View at: Publisher Site | Google Scholar
  79. A. S. Brack and T. A. Rando, “Intrinsic changes and extrinsic influences of myogenic stem cell function during aging,” Stem Cell Reviews, vol. 3, no. 3, pp. 226–237, 2007. View at: Publisher Site | Google Scholar
  80. S. Carosio, M. G. Berardinelli, M. Aucello, and A. Musarò, “Impact of ageing on muscle cell regeneration,” Ageing Research Reviews, vol. 10, no. 1, pp. 35–42, 2011. View at: Publisher Site | Google Scholar
  81. Y. C. Jang, M. Sinha, M. Cerletti, C. Dall'osso, and A. J. Wagers, “Skeletal muscle stem cells: effects of aging and metabolism on muscle regenerative function,” Cold Spring Harbor Symposia on Quantitative Biology, vol. 76, pp. 101–111, 2011. View at: Publisher Site | Google Scholar
  82. F. Relaix and P. S. Zammit, “Satellite cells are essential for skeletal muscle regeneration: the cell on the edge returns centre stage,” Development, vol. 139, no. 16, pp. 2845–2856, 2012. View at: Publisher Site | Google Scholar
  83. A. Mauro and W. R. Adams, “The structure of the sarcolemma of the frog skeletal muscle fiber,” The Journal of Biophysical and Biochemical Cytology, vol. 10, no. 4, supplement, pp. 177–185, 1961. View at: Google Scholar
  84. T. J. Hawke and D. J. Garry, “Myogenic satellite cells: physiology to molecular biology,” Journal of Applied Physiology, vol. 91, no. 2, pp. 534–551, 2001. View at: Google Scholar
  85. I. Stratos, Z. Li, P. Herlyn et al., “Vitamin D increases cellular turnover and functionally restores the skeletal muscle after crush injury in rats,” American Journal of Pathology, vol. 182, no. 3, pp. 895–904, 2013. View at: Publisher Site | Google Scholar
  86. P. Zimmet, K. G. Alberti, and J. Shaw, “Global and societal implications of the diabetes epidemic,” Nature, vol. 414, no. 6865, pp. 782–787, 2001. View at: Publisher Site | Google Scholar
  87. S. W. Park, B. H. Goodpaster, J. S. Lee et al., “Excessive loss of skeletal muscle mass in older adults with type 2 diabetes,” Diabetes Care, vol. 32, no. 11, pp. 1993–1997, 2009. View at: Publisher Site | Google Scholar
  88. J. Mitri, M. D. Muraru, and A. G. Pittas, “Vitamin D and type 2 diabetes: a systematic review,” European Journal of Clinical Nutrition, vol. 65, no. 9, pp. 1005–1015, 2011. View at: Publisher Site | Google Scholar
  89. N. Jessen and L. J. Goodyear, “Contraction signaling to glucose transport in skeletal muscle,” Journal of Applied Physiology, vol. 99, no. 1, pp. 330–337, 2005. View at: Publisher Site | Google Scholar
  90. P. Manna and S. K. Jain, “Vitamin D up-regulates glucose transporter 4 (GLUT4) translocation and glucose utilization mediated by cystathionine-γ-lyase (CSE) activation and H2S formation in 3T3L1 adipocytes,” The Journal of Biological Chemistry, vol. 287, no. 50, pp. 42324–42332, 2012. View at: Publisher Site | Google Scholar
  91. O. Kabil and R. Banerjee, “Redox biochemistry of hydrogen sulfide,” The Journal of Biological Chemistry, vol. 285, no. 29, pp. 21903–21907, 2010. View at: Publisher Site | Google Scholar
  92. B. Tamilselvan, K. G. Seshadri, and G. Venkatraman, “Role of vitamin D on the expression of glucose transporters in L6 myotubes,” Indian Journal of Endocrinology and Metabolism, vol. 17, supplement 1, pp. S326–S328, 2013. View at: Google Scholar
  93. A. J. Castro, M. J. Frederico, L. H. Cazarolli et al., “Betulinic acid and 1, 25(OH)2 vitamin D3 share intracellular signal transduction in glucose homeostasis in soleus muscle,” The International Journal of Biochemistry & Cell Biology, vol. 48, pp. 18–27, 2014. View at: Google Scholar
  94. S. Lund, G. D. Holman, O. Schmitz, and O. Pedersen, “Contraction stimulates translocation of glucose transporter GLUT4 in skeletal muscle through a mechanism distinct from that of insulin,” Proceedings of the National Academy of Sciences of the United States of America, vol. 92, no. 13, pp. 5817–5821, 1995. View at: Publisher Site | Google Scholar
  95. W. E. Stumpf, M. Sar, F. A. Reid, Y. Tanaka, and H. F. DeLuca, “Target cells for 1,25-dihydroxyvitamin D3 in intestinal tract, stomach, kidney, skin, pituitary, and parathyroid,” Science, vol. 206, no. 4423, pp. 1188–1190, 1979. View at: Google Scholar
  96. K. Colston, M. Hirst, and D. Feldman, “Organ distribution of the cytoplasmic 1,25-dihydroxycholecalciferol receptor in various mouse tissues,” Endocrinology, vol. 107, no. 6, pp. 1916–1922, 1980. View at: Google Scholar
  97. M. E. Sandgren, M. Brönnegärd, and H. F. DeLuca, “Tissue distribution of the 1, 25-dihydroxyvitamin D3 receptor in the male rat,” Biochemical and Biophysical Research Communications, vol. 181, no. 2, pp. 611–616, 1991. View at: Google Scholar
  98. S. B. Zanello, E. D. Collins, M. J. Marinissen, A. W. Norman, and R. L. Boland, “Vitamin d receptor expression in chicken muscle tissue and cultured myoblasts,” Hormone and Metabolic Research, vol. 29, no. 5, pp. 231–236, 1997. View at: Google Scholar
  99. E. M. Costa, H. M. Blau, and D. Feldman, “1,25-dihydroxyvitamin D3 receptors and hormonal responses in cloned human skeletal muscle cells,” Endocrinology, vol. 119, no. 5, pp. 2214–2220, 1986. View at: Google Scholar
  100. R. Boland, A. Norman, E. Ritz, and W. Hasselbach, “Presence of a 1,25-dihydroxy-vitamin D3 receptor in chick skeletal muscle myoblasts,” Biochemical and Biophysical Research Communications, vol. 128, no. 1, pp. 305–311, 1985. View at: Google Scholar
  101. C. Buitrago, G. Vazquez, A. R. De Boland, and R. L. Boland, “Activation of Src kinase in skeletal muscle cells by 1, 1, 25-(OH2)-vitamin D3 correlates with tyrosine phosphorylation of the vitamin D receptor (VDR) and VDR-Src interaction,” Journal of Cellular Biochemistry, vol. 79, no. 2, pp. 274–281, 2000. View at: Google Scholar
  102. C. Buitrago, G. Vazquez, A. R. De Boland, and R. Boland, “The vitamin D receptor mediates rapid changes in muscle protein tyrosine phosphorylation induced by 1,25(OH)2D3,” Biochemical and Biophysical Research Communications, vol. 289, no. 5, pp. 1150–1156, 2001. View at: Publisher Site | Google Scholar
  103. R. Boland, A. R. De Boland, C. Buitrago et al., “Non-genomic stimulation of tyrosine phosphorylation cascades by 1,25(OH)2D3 by VDR-dependent and -independent mechanisms in muscle cells,” Steroids, vol. 67, no. 6, pp. 477–482, 2002. View at: Publisher Site | Google Scholar
  104. J. Salles, A. Chanet, C. Giraudet et al., “1,25(OH)2-vitamin D3 enhances the stimulating effect of leucine and insulin on protein synthesis rate through Akt/PKB and mTOR mediated pathways in murine C2C12 skeletal myotubes,” Molecular Nutrition & Food Research, vol. 57, no. 12, pp. 2137–2146, 2013. View at: Google Scholar
  105. Y. Wang, B. R. Becklund, and H. F. DeLuca, “Identification of a highly specific and versatile vitamin D receptor antibody,” Archives of Biochemistry and Biophysics, vol. 494, no. 2, pp. 166–177, 2010. View at: Publisher Site | Google Scholar
  106. T. Kislinger, A. O. Gramolini, Y. Pan, K. Rahman, D. H. MacLennan, and A. Emili, “Proteome dynamics during C2C12 myoblast differentiation,” Molecular and Cellular Proteomics, vol. 4, no. 7, pp. 887–901, 2005. View at: Publisher Site | Google Scholar
  107. D. Zehnder, R. Bland, M. C. Williams et al., “Extrarenal expression of 25-hydroxyvitamin D3-1α-hydroxylase,” Journal of Clinical Endocrinology & Metabolism, vol. 86, no. 2, pp. 888–894, 2001. View at: Publisher Site | Google Scholar
  108. J. S. Adams and M. Hewison, “Extrarenal expression of the 25-hydroxyvitamin D-1-hydroxylase,” Archives of Biochemistry and Biophysics, vol. 523, no. 1, pp. 95–102, 2012. View at: Publisher Site | Google Scholar
  109. Y. Nakamura, T.-A. Eto, T. Taniguchi et al., “Purification and characterization of 25-hydroxyvitamin D3 1α- hydroxylase from rat kidney mitochondria,” FEBS Letters, vol. 419, no. 1, pp. 45–48, 1997. View at: Publisher Site | Google Scholar
  110. Y. R. Lou, F. Molnár, M. Peräkylä et al., “25-Hydroxyvitamin D3 is an agonistic vitamin D receptor ligand,” The Journal of Steroid Biochemistry and Molecular Biology, vol. 118, no. 3, pp. 162–170, 2010. View at: Google Scholar
  111. K. Bismuth and F. Relaix, “Genetic regulation of skeletal muscle development,” Experimental Cell Research, vol. 316, no. 18, pp. 3081–3086, 2010. View at: Publisher Site | Google Scholar
  112. T. Braun, M. A. Rudnicki, H.-H. Arnold, and R. Jaenisch, “Targeted inactivation of the muscle regulatory gene Myf-5 results in abnormal rib development and perinatal death,” Cell, vol. 71, no. 3, pp. 369–382, 1992. View at: Publisher Site | Google Scholar
  113. M. A. Rudnicki, T. Braun, S. Hinuma, and R. Jaenisch, “Inactivation of MyoD in mice leads to up-regulation of the myogenic HLH gene Myf-5 and results in apparently normal muscle development,” Cell, vol. 71, no. 3, pp. 383–390, 1992. View at: Publisher Site | Google Scholar
  114. P. Hasty, A. Bradley, J. H. Morris et al., “Muscle deficiency and neonatal death in mice with a targeted mutation in the myogenin gene,” Nature, vol. 364, no. 6437, pp. 501–506, 1993. View at: Publisher Site | Google Scholar
  115. Y. Nabeshima, K. Hanaoka, M. Hayasaka et al., “Myogenin gene disruption results in perinatal lethality because of severe muscle defect,” Nature, vol. 364, no. 6437, pp. 532–535, 1993. View at: Publisher Site | Google Scholar
  116. A. Rawls, M. R. Valdez, W. Zhang, J. Richardson, W. H. Klein, and E. N. Olson, “Overlapping functions of the myogenic bHLH genes MRF4 and MyoD revealed in double mutant mice,” Development, vol. 125, no. 13, pp. 2349–2358, 1998. View at: Google Scholar
  117. A. B. Lassar, R. L. Davis, W. E. Wright et al., “Functional activity of myogenic HLH proteins requires hetero-oligomerization with E12/E47-like proteins in vivo,” Cell, vol. 66, no. 2, pp. 305–315, 1991. View at: Publisher Site | Google Scholar
  118. X.-H. Sun and D. Baltimore, “An inhibitory domain of E12 transcription factor prevents DNA binding in E12 homodimers but not in E12 heterodimers,” Cell, vol. 64, no. 2, pp. 459–470, 1991. View at: Google Scholar
  119. T. Yoshizawa, Y. Handa, Y. Uematsu et al., “Mice lacking the vitamin D receptor exhibit impaired bone formation, uterine hypoplasia and growth retardation after weaning,” Nature Genetics, vol. 16, no. 4, pp. 391–396, 1997. View at: Publisher Site | Google Scholar
  120. K. Sakoda, M. Fujiwara, S. Arai et al., “Isolation of a genomic DNA fragment having negative vitamin D response element,” Biochemical and Biophysical Research Communications, vol. 219, no. 1, pp. 31–35, 1996. View at: Publisher Site | Google Scholar
  121. T. Nishishita, T. Okazaki, T. Ishikawa et al., “A negative vitamin D response DNA element in the human parathyroid hormone-related peptide gene binds to vitamin D receptor along with Ku antigen to mediate negative gene regulation by vitamin D,” The Journal of Biological Chemistry, vol. 273, no. 18, pp. 10901–10907, 1998. View at: Publisher Site | Google Scholar
  122. S. Seoane, M. Alonso, C. Segura, and R. Pérez-Fernández, “Localization of a negative vitamin D response sequence in the human growth hormone gene,” Biochemical and Biophysical Research Communications, vol. 292, no. 1, pp. 250–255, 2002. View at: Publisher Site | Google Scholar
  123. G. P. Dotto, “p21(WAF1/Cip1): more than a break to the cell cycle?” Biochimica et Biophysica Acta—Reviews on Cancer, vol. 1471, no. 1, pp. M43–M56, 2000. View at: Publisher Site | Google Scholar
  124. C. J. Sherr and J. M. Roberts, “CDK inhibitors: positive and negative regulators of G1-phase progression,” Genes & Development, vol. 13, no. 12, pp. 1501–1512, 1999. View at: Google Scholar
  125. J. R. Nevins, “E2F: a link between the Rb tumor suppressor protein and viral oncoproteins,” Science, vol. 258, no. 5081, pp. 424–429, 1992. View at: Google Scholar
  126. C. J. Sherr, “Cancer cell cycles,” Science, vol. 274, no. 5293, pp. 1672–1677, 1996. View at: Publisher Site | Google Scholar
  127. H. Amthor, G. Nicholas, I. McKinnell et al., “Follistatin complexes Myostatin and antagonises Myostatin-mediated inhibition of myogenesis,” Developmental Biology, vol. 270, no. 1, pp. 19–30, 2004. View at: Publisher Site | Google Scholar
  128. H. Kocamis, S. A. Gahr, L. Batelli, A. F. Hubbs, and J. Killefer, “IGF-I, IGF-II, and IGF-receptor-1 transcript and IGF-II protein expression in myostatin knockout mice tissues,” Muscle & Nerve, vol. 26, no. 1, pp. 55–63, 2002. View at: Publisher Site | Google Scholar
  129. H. Gilson, O. Schakman, L. Combaret et al., “Myostatin gene deletion prevents glucocorticoid-induced muscle atrophy,” Endocrinology, vol. 148, no. 1, pp. 452–460, 2007. View at: Publisher Site | Google Scholar
  130. A. Marshall, M. S. Salerno, M. Thomas et al., “Mighty is a novel promyogenic factor in skeletal myogenesis,” Experimental Cell Research, vol. 314, no. 5, pp. 1013–1029, 2008. View at: Publisher Site | Google Scholar
  131. H. Gilson, O. Schakman, S. Kalista, P. Lause, K. Tsuchida, and J.-P. Thissen, “Follistatin induces muscle hypertrophy through satellite cell proliferation and inhibition of both myostatin and activin,” American Journal of Physiology—Endocrinology and Metabolism, vol. 297, no. 1, pp. E157–E164, 2009. View at: Publisher Site | Google Scholar
  132. M. Miyake, S. Hayashi, Y. Taketa et al., “Myostatin down-regulates the IGF-2 expression via ALK-Smad signaling during myogenesis in cattle,” Animal Science Journal, vol. 81, no. 2, pp. 223–229, 2010. View at: Publisher Site | Google Scholar
  133. N. G. Williams, J. P. Interlichia, M. F. Jackson, D. Hwang, P. Cohen, and B. D. Rodgers, “Endocrine actions of myostatin: systemic regulation of the igf and igf binding protein axis,” Endocrinology, vol. 152, no. 1, pp. 172–180, 2011. View at: Publisher Site | Google Scholar
  134. C. Duan, H. Ren, and S. Gao, “Insulin-like growth factors (IGFs), IGF receptors, and IGF-binding proteins: roles in skeletal muscle growth and differentiation,” General and Comparative Endocrinology, vol. 167, no. 3, pp. 344–351, 2010. View at: Publisher Site | Google Scholar
  135. N. Zanou and P. Gailly, “Skeletal muscle hypertrophy and regeneration: interplay between the myogenic regulatory factors (MRFs) and insulin-like growth factors (IGFs) pathways,” Cellular and Molecular Life Sciences, vol. 70, no. 21, pp. 4117–4130, 2013. View at: Publisher Site | Google Scholar
  136. X. Shen, J. M. Collier, M. Hlaing et al., “Genome-wide examination of myoblast cell cycle withdrawal during differentiation,” Developmental Dynamics, vol. 226, no. 1, pp. 128–138, 2003. View at: Publisher Site | Google Scholar
  137. A. Amirouche, A.-C. Durieux, S. Banzet et al., “Down-regulation of Akt/mammalian target of rapamycin signaling pathway in response to myostatin overexpression in skeletal muscle,” Endocrinology, vol. 150, no. 1, pp. 286–294, 2009. View at: Publisher Site | Google Scholar
  138. R. Sartori, G. Milan, M. Patron et al., “Smad2 and 3 transcription factors control muscle mass in adulthood,” American Journal of Physiology—Cell Physiology, vol. 296, no. 6, pp. C1248–C1257, 2009. View at: Publisher Site | Google Scholar
  139. A. U. Trendelenburg, A. Meyer, D. Rohner, J. Boyle, S. Hatakeyama, and D. J. Glass, “Myostatin reduces Akt/TORC1/p70S6K signaling, inhibiting myoblast differentiation and myotube size,” American Journal of Physiology—Cell Physiology, vol. 296, no. 6, pp. C1258–C1270, 2009. View at: Publisher Site | Google Scholar
  140. C. Lipina, H. Kendall, A. C. McPherron, P. M. Taylor, and H. S. Hundal, “Mechanisms involved in the enhancement of mammalian target of rapamycin signalling and hypertrophy in skeletal muscle of myostatin-deficient mice,” FEBS Letters, vol. 584, no. 11, pp. 2403–2408, 2010. View at: Publisher Site | Google Scholar
  141. D. D. Sarbassov, S. M. Ali, and D. M. Sabatini, “Growing roles for the mTOR pathway,” Current Opinion in Cell Biology, vol. 17, no. 6, pp. 596–603, 2005. View at: Publisher Site | Google Scholar

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