Evidence for Vitamin D Receptor Expression and Direct Effects of 1α,25(OH)₂D₃ in Human Skeletal Muscle Precursor Cells

Abstract

Presence of the vitamin D receptor and direct effects of vitamin D on the proliferation and differentiation of muscle precursor cells have been demonstrated in animal models. However, the effects and mechanisms of vitamin D actions in human skeletal muscle, and the presence of the vitamin D receptor in human adult skeletal muscle, remain to be established. Here, we investigated the role of vitamin D in human muscle cells at various stages of differentiation. We demonstrate that the components of the vitamin D-endocrine system are readily detected in human muscle precursor cells but are low to nondetectable in adult skeletal muscle and that human muscle cells lack the ability to convert the inactive vitamin D-metabolite 25-hydroxy-vitamin D₃ to the active 1α,25-dihydroxy-vitamin D₃ (1α,25(OH)₂D₃). In addition, we show that 1α,25(OH)₂D₃ inhibits myoblast proliferation and differentiation by altering the expression of cell cycle regulators and myogenic regulatory factors, with associated changes in forkhead box O3 and Notch signaling pathways. The present data add novel information regarding the direct effects of vitamin D in human skeletal muscle and provide functional and mechanistic insight to the regulation of myoblast cell fate decisions by 1α,25(OH)₂D₃.

Vitamin D deficiency is a widely spread and highly prevalent condition associated with skeletal muscle weakness and small muscle fiber size (1). In animal models, the skeletal muscle dysfunction observed in vitamin D deficiency is reversed by vitamin D repletion, whereas vitamin D supplementation in humans has been concluded to increase skeletal muscle strength (2, 3). In vitamin D-deficient and vitamin D receptor (VDR) knockout animal models, skeletal muscle abnormalities are independent of secondary metabolic changes (3–5). This have led researchers to suggest that the effects of vitamin D are mediated by direct actions of vitamin D in skeletal muscle, supported by the identification of the VDR in both rodent and human muscle cells (6–8). Challenging this notion, the finding that VDR protein was undetectable in skeletal muscle when using the stringently validated VDR D-6 antibody has led investigators to question the presence of the VDR in skeletal muscle (9). Consequently, direct effects of vitamin D in skeletal muscle remain controversial (9–11).

In contrast to adult skeletal muscle, VDR expression at both the mRNA and protein level has been repeatedly shown in rodent muscle precursor cells (12–15). Together with the demonstration of direct effects of vitamin D (12–15), these findings provide compelling evidence for the presence of functional VDR protein in these cells. Notably, studies investigating the expression level of the VDR in rodent skeletal muscle have established markedly higher levels of the VDR in neonatal and regenerating skeletal muscle than in adult skeletal muscle (12, 13). Collectively, the current body of data indicates differences in the expression level of the VDR throughout the various stages of muscle cell differentiation.

Rodent muscle precursor cells are responsive to both the inactive vitamin D-metabolite 25-hydroxy-vitamin D₃ (25(OH)D₃) and the active 1α,25-dihydroxy-vitamin D₃ (1α,25(OH)₂D₃) (12–16). This suggests that the 25-hydroxyvitamin D3-1-alpha-hydroxylase (CYP27B1), responsible for the conversion of the inactive 25(OH)D₃ into the active 1α,25(OH)₂D₃, is expressed and functional in these cells. Indeed, the presence of functionally active CYP27B1 has been confirmed in rodent muscle precursor cells at different stages of differentiation, and its expression verified in regenerating rodent muscle fibers (12–14). This is intriguing in that it expands the potential role of vitamin D in skeletal muscle to include localized vitamin D actions and regulation within this tissue. However, the presence and relative abundance of the CYP27B1, the 1α,25(OH)₂D₃-inactivating 1,25-dihydroxyvitamin D3 24-hydroxylase (CYP24A1), and the VDR at different stages of human muscle cell differentiation and in human adult skeletal muscle remain to be established.

Although the vast majority of studies performed in animal models have established an inhibitory effect of vitamin D on myoblast proliferation (7, 13–15, 17, 18), the effect on myoblast differentiation is ambiguous. This is demonstrated by the inconsistent findings of previous studies performed in animal muscle cells regarding the effects of vitamin D on myotube formation (14, 15, 18, 19) and expression of myogenic regulatory factors (4, 14, 17, 18, 20). Notably, recent studies have shown an increase in myotube size after myoblast differentiation in the presence of vitamin D, irrespective of differences regarding the effects of vitamin D on myoblast differentiation (14, 15). The associated decrease in myostatin expression led these authors to suggest that the effects of vitamin D on myotube size and differentiation may be mediated by a reduction in endogenous myostatin expression (14, 15). This hypothesis is intriguing in that it suggests a modulatory role of vitamin D in the local production of a well-established negative regulator of skeletal muscle mass. Additionally, these studies demonstrate that vitamin D may regulate skeletal muscle size independent of its effects on myoblast differentiation.

Here, we compared the expression of the components of the vitamin D-endocrine system in human muscle precursor cells at different stages of differentiation and in human adult skeletal muscle tissue. Human myoblast proliferation and differentiation into myotubes after exposure to 1α,25(OH)₂D₃ and 25(OH)D_3, and associated changes in the endogenous expression of myostatin, were studied. Finally, global gene expression analysis was conducted to identify plausible downstream mediators of 1α,25(OH)₂D₃-actions in human muscle precursor cells. We demonstrate that in human skeletal muscle, components of the vitamin D-endocrine system are predominantly expressed in muscle precursor cells, indicating a preferential role for vitamin D in these cells. In addition, we show that 1α,25(OH)₂D₃ inhibits both the proliferation and differentiation of human myoblasts without associated changes in endogenous myostatin expression, but by altering the expression of other key regulators of cell cycle progression and myogenic differentiation. This establishes a regulatory role for vitamin D in human myoblast cell fate decisions and has implications for our understanding of vitamin D actions in human skeletal muscle tissue.

Materials and Methods

Study participants and ethical approval

Nine recreationally active healthy individuals, 5 males and 4 females, age 20–27 years were recruited for the experiments. The planned experiments and procedures were explained before subjects gave their written informed consent to participate. The study was approved by the Regional Ethical Review Board in Stockholm (Dnr 2012/173-31/3).

Skeletal muscle biopsy

Resting skeletal muscle biopsies were collected from the vastus lateralis muscle via the percutaneous needle biopsy technique as described previously (21). After sterilization (chlorhexidine alcohol cutaneous solution, 5 mg/mL; Fresenius Kabi) and local anesthesia (Carbocain, 10 mg/mL; Astra Zeneca) a small insertion in the skin was made with a scalpel. The muscle biopsy was obtained using the Bergström needle to penetrate the muscular fascia and connective tissue. Biopsy material for myoblast isolation was immediately put in sterile PBS (Gibco Invitrogen, Life Technologies) containing 1% antibiotic-antimycotic (ABAM) (Gibco Invitrogen, Life Technologies), whereas muscle tissue for protein and RNA analysis was snap frozen in isopentane and stored at −80°C.

Myoblast isolation and cell culture

All reagents used for myoblast isolation as well as human myoblast and monocytic cell culture were from Gibco Invitrogen, Life Technologies. Myoblasts were extracted from fresh muscle tissue as previously described (22), with some modifications. Immediately after the biopsy procedure, approximately 100 mg of muscle tissue were placed in PBS solution containing 1% ABAM and incubated at 4°C overnight. The next day, the muscle biopsy was incubated in 5 mL TrypLE Express Enzyme (1×) at 37°C, 5% CO₂ with gentle agitation for 20 minutes. Undigested tissue was allowed to settle for 5 minutes at room temperature, and the supernatant containing the myogenic cells was collected in 5 mL DMEM-F12 GlutaMAX, containing 20% fetal bovine serum (FBS) and 1% ABAM. Digestion of the slurry was repeated twice. Isolated myoblasts were cultured in DMEM-F12 GlutaMAX containing 20% FBS and 1% ABAM (high-serum media) at 37°C, 5% CO₂. Culture dishes were coated with collagen I (collagen I, bovine 5 mg/mL) diluted to a final concentration of 50 μg/mL in 0.02M acetic acid according to the manufacturer's instructions. Myoblasts were taken through serial passages to increase cell numbers before experimentation. All myoblasts were used for experimentation at passage 4–5. For experimentation, myoblasts were harvested and transferred to collagen I-coated 6-well plates at a density of 8 × 10⁴ cells per well, or to collagen I-coated glass cover slides at the same density (for morphological analysis) and allowed to settle in high-serum media for at least 24 hours before experimentation. Myotube differentiation was promoted by substituting the proliferation media with DMEM-F12 GlutaMAX containing 2% FBS and 1% ABAM (low-serum media). THP1 cells (Sigma-Aldrich) were cultured in RPMI 1640 medium GlutaMAX supplemented with 1% ABAM, 10% FBS, 1mM sodium pyruvate, and 0.05mM 2-mercaptoethanol.

Immunomagnetic cell separation

Enrichment of the cell population for myogenic cells was accomplished by a combination of preplating and magnetic-activated cell sorting (MACS) separation as has previously been reported to produce a high yield of myogenic cells (23). MACS separation of myogenic and nonmyogenic cells was carried out as previously described for human muscle-derived cells (24), with some modifications. Muscle-derived cells plated in T75 flasks were incubated with a mouse antihuman antibody for CD56 (MY31; BD Biosciences) dissolved in DMEM-F12 GlutaMAX for 30 minutes at 37°C, 5% CO₂. Cells were subsequently pelleted and resuspended in PBS containing 0.1% FBS and antimouse IgG microbeads (Miltenyi Biotech) according to the manufacturer's instructions. Cell suspension was incubated in the dark at 4°C for 15 minutes before being rinsed with PBS containing 0.1% FBS and repelleted. Cells were magnetically separated using a midiMACS magnet and LS columns (Miltenyi Biotech). The cells that were bound to the anti-CD56 microbead complex were maintained in the column and constituted the positive (myogenic) fraction of cells. This fraction was subsequently plated and cultured as described above.

Validation of myogenic origin

At the time of plating cells for experimentation, a fraction of myoblasts was collected for confirmation of myogenic origin. Cells were spun down onto a cover glass for subsequent immunofluorescent staining for the myogenic marker desmin (ab15200; Abcam). The fraction of desmin-positive cells in the cell population was analyzed by dividing with the total number of nuclei counterstained with 4',6-diamidino-2-phenylindole dihydrochloride (Molecular Probes) within each field. In the current study, 97% (±0.8) of subconfluent myoblasts were positive for desmin.

Cell proliferation analysis

Cell proliferation was analyzed by incorporation of bromodeoxyuridine (BrdU) by a commercially available kit (Roche Diagnostics GmbH). Myoblasts were plated in 96-well plates at a density of 3 × 10³ cells per well in high-serum media. After 24 hours, media were changed to low-serum media containing 1nM 1α,25(OH)₂D₃ (Sigma-Aldrich), 100nM 1α,25(OH)₂D₃, 100nM 25(OH)D₃ (Sigma-Aldrich), or vehicle (dimethyl sulfoxide) (Sigma-Aldrich). Stimulation media were changed after 24 hours of treatment, and stimulation media containing BrdU were added for another 24 hours. BrdU ELISA was performed according to the manufacturer's instructions. The rationale for analyzing proliferation in low-serum media were based upon previous studies addressing proliferation in C2C12 cells, where high-serum media have been demonstrated to produce unacceptably high background levels of proliferation (25).

Apoptosis analysis

Myoblasts stimulated with 1nM 1α,25(OH)₂D₃, 100nM 1α,25(OH)₂D₃, or vehicle for 48 hours were analyzed for apoptosis and necrosis using the FITC Annexin V Apoptosis Detection kit (BD Biosciences) according to the manufacturer's instructions. In brief, after dissociation using TrypLE Express Enzyme (1×), the myoblasts were rinsed twice in PBS. After suspending the myoblasts in Binding buffer (1×), Annexin V and propidium iodide (PI) were added, and the cells were incubated for 20 minutes in the dark at room temperature. Sample acquisition was performed within an hour on a FACSAria cell sorter with FACSDiva 6.1 software (BD Biosciences). Unstained cells and cells stained with either Annexin V or PI were used to set up compensation and quadrants.

Cell stimulation protocol

For analysis of the effects of vitamin D metabolites on myoblast gene and protein expression and differentiation, high-serum media were changed to low-serum media containing 1nM or 100nM 1α,25(OH)₂D₃, 100nM 25(OH)D₃, or vehicle alone. After 24 hours of stimulation, RNA and protein were collected for the 24-hour analysis, whereas remaining cells were allowed to differentiate for another 7 days in low-serum media. In order to minimize the number of cells needed and thus keep the number of passages required at a minimum, the highest concentration of 1α,25(OH)₂D₃ was only continued in cells plated for morphological analysis at the 8-day time point.

Real-time quantitative PCR

Total RNA was prepared by the TRIzol method (Invitrogen, Life Technologies) and quantified spectrophotometrically by absorbance at 260 nm. One microgram of total RNA was reverse transcribed by Superscript reverse transcriptase (Life Technologies) using random hexamer primers (Roche Diagnostics) in a total volume of 20 μL. Detection of mRNA was performed on an ABI-PRISM7700 Sequence Detector (PerkinElmer, Applied Biosystems). Primer and probe for VDR, CYP24A1, CYP27B1, myogenin, myogenic differentiation 1 (MyoD1), myostatin, ubiquitin B (UBB), ubiquitin C (UBC), ribosomal protein 18 (RPS18), glyceraldehyde-3-phosphate dehydrogenase (GAPDH), eukaryotic 18S rRNA (18S), hairy and enhancer of split 1 (HES1), and forkhead box O3 (FOXO3) were ordered as assay on demand (VDR, Hs00172113_m1; CYP24A1, Hs00167999_m1; CYP27B1, Hs00168017_m1; myogenin, Hs01072232_m1; MyoD1, Hs02330075_g1; myostatin, Hs00976237_m1; UBB, Hs00430290_m1; UBC, Hs00824723_m1; RPS18, Hs01375212_g1; GAPDH, 4352934E; 18S, 4310893E, HES1, Hs00172878_m1; and FOXO3, Hs00818121_m1; PerkinElmer, Applied Biosystems). To identify the optimal reference gene for normalization the stability of the following reference genes were analyzed: UBB, UBC, RPS18, GAPDH, and 18S. GAPDH was identified as the most stable reference gene for comparison of gene expression between vitamin D-stimulated myoblasts and myotubes, whereas the geometrical mean of RPS18 and UBB was identified as the most stable reference for comparison of gene expression between myoblasts, myotubes, and adult skeletal muscle tissue according to the NormFinder algorithm (26). Target gene expression was subsequently reported as a ratio of the respective reference genes by the 2^−ΔCT formula.

Immunofluorescence and microscopy

Cells cultured on glass slides were washed twice with PBS and fixed in 4% formaldehyde for 20 minutes. Cells were subsequently permeabilized and blocked in 0.2% Triton X-100 and 2% FBS in PBS solution for 30 minutes and stained for desmin (ab15200; Abcam) diluted 1:200 in PBS solution. Secondary antibody Alexa Fluor 568 (Life Technologies) diluted 1:800 in PBS solution was added to enable fluorescent detection. ProLong Gold antifade reagent with 4',6-diamidino-2-phenylindole dihydrochloride (Life Technologies) was used to counterstain nuclei and mount onto microscope slides. The sections were visualized using a Leica DFC450 fluorescent microscope with manufacturer's software. Alexa Fluor 568 was excited at 561 nm and emitted at 565 nm. A total of 5 fields per subject and condition were captured with a cell imaging system at ×10 magnification and analyzed using ImageJ (National Institutes of Health; available at http://rsb.info.nih.gov/ij). Morphology was assessed in 4 subjects by determining the ratio of the total number of myotube nuclei and the total nuclei per field of view (ie, fusion index), and the number of myotubes per view. Myotubes were defined as desmin-positive cells containing more than or equal to 3 nuclei encapsulated within cellular structures, so to avoid counting of single cells undergoing mitosis.

Western blotting and densitometry analysis

Cells were lysed and tissue homogenized in Pierce RIPA buffer (Thermo Scientific) with the addition of 0.5M EDTA solution (Thermo Scientific) and Halt Protease & Phosphatase Inhibitor Cocktail (Thermo Scientific) according to the manufacturer's instructions. In a subset of experiments, the effect of different homogenization protocols on the detection of VDR protein in human adult skeletal muscle tissue was investigate by comparing the detection of VDR protein in skeletal muscle tissue prepared in regular lysis buffer with that prepared in hyperosmolar lysis buffer (HLB) as previously described (12). Protein concentration was assessed by Bradford protein assay (Bio-Rad). Protein was separated at the stated concentrations by SDS-PAGE. 10% Mini-PROTEAN TGX gels were used (Bio-Rad), and the membrane was blocked in Fluorescent Blocker (Merck Millipore) for 1 hour at room temperature. Primary antibodies were applied overnight at 4°C at a concentration of 1:500 for the VDR (VDR D-6, sc-131333; Santa Cruz Biotechnology, Inc) and 1:2000 for GAPDH (ab9485; Abcam). Washed membranes were incubated for 1 hour at room temperature with secondary antibodies at a concentration of 1:15000. A final series of washes were then performed before scanning the membranes (Odyssey SA Infrared Imaging System; LI-COR Bioscience). The blots were subsequently quantified using ImageJ. For comparisons between myoblasts and myotubes, GAPDH was used as loading control. Comparison of the basal protein quantity of the VDR between myoblasts, myotubes, adult muscle and monocytic cells was performed by normalizing the protein quantity of the VDR to the total protein abundance detected by coomassie staining as previously described (27). HeLa whole-cell lysate (Santa Cruz Biotechnology, Inc) was used as a positive control for the VDR. A list of primary antibodies, the dilutions used and manufacturers' details are included in Table 1.

Table 1

Antibody Table

Peptide/Protein Target	Antigen Sequence (if Known)	Name of Antibody	Manufacturer, Catalog Number, and/or Name of Individual Providing the Antibody	Species Raised in; Monoclonal or Polyclonal	Dilution Used
VDR	Amino acids 344–424 of VDR of human origin	VDR antibody (D-6)	Santa Cruz Biotechnology, Inc, sc-13133	Mouse; monoclonal	1:500
GAPDH	Full-length native protein from human erythrocytes	Anti-GAPDH antibody, loading control	Abcam, ab9485	Rabbit; polyclonal	1:2000
Desmin	Synthetic peptide. Human, C terminal	Antidesmin antibody	Abcam, ab15200	Rabbit; polyclonal	1:200

Microarray

For 6 of the recruited subjects, 3 male and 3 female, global gene expression was analyzed using microarray on myoblasts exposed to 100nM 1α,25(OH)₂D₃ or vehicle alone for 24 hours in low-serum media. Total RNA was extracted using TRIzol reagent as described above. In vitro transcription was performed using the Bioarray high yield RNA transcript labeling kit (P/N 900182; Affymetrix). Unincorporated nucleotides from the in vitro transcription reaction were removed using the RNeasy column (QIAGEN). Hybridization, washing, staining, and scanning of the arrays were performed according to the manufacturer's instructions (Affymetrix). As a means to control the quality of the individual arrays, all arrays were examined using hierarchical clustering and Normalized Unscaled Standard Error (a variance based metric to identify outliers before statistical analysis), in addition to the standard quality assessments, including scaling factors and chip-housekeeper 5′ to 3′ ratios. All chips passed these quality control steps. The probe set level intensities of the arrays were normalized using the robust multiarray analysis method implemented within the R statistical software environment using the “affy” package (Bioconductor project). Raw data and robust multiarray analysis normalized expression values are publicly available through Gene Expression Omnibus accession number GSE69698. Before differential expression analysis probe sets without available annotation and probe sets displaying low-variance across samples were excluded from further analysis. Annotation was carried out in the R environment using “hgu133plus2cdf,” “annotation,” and “genefilter” packages (Bioconductor). Differential expression was estimated through pairwise-statistical analysis of microarrays on the R-platform. A total of 2315 probe sets with a false discovery rate (FDR) of less than or equal to 5% were categorized as differentially expressed (1364 up and 951 down). Differential expression was validated by quantitative real-time PCR for 3 up-regulated (CYP24A1, FOXO3A, and HES1) and 2 down-regulated (myogenin and MyoD1) genes demonstrating agreement with microarray data (see Figures 2 and 5 below and Supplemental Figure 1). Significantly up- and down-regulated probe sets were analyzed for gene ontology (GO) enrichment using the web-based Database for Annotation, Visualization and Integrated Discovery (DAVID) version 6.7 (28, 29). Probe sets with a FDR of less than or equal to 5% and a fold change of more than or equal to 1.5-fold were analyzed for enrichment of “biological functions.” To correct for possible selection bias of genes, only probe sets used in the statistical analysis of microarray analysis were used as background. Enriched ontologies with a FDR of less than or equal to 1% were categorized as significant. For analysis, we used the web-based bioinformatics tool Ingenuity Pathway Analysis (IPA) (QIAGEN). IPA is a knowledge database generated from peer-reviewed scientific publications that enables discovery of biological networks in gene expression data and determines the biological functions most significant to those networks. HUGO gene name identifiers were uploaded into IPA and queried against the verified IPA knowledge database.

Figure 2

VDR and CYP24A1 expression is induced in primary human myoblasts after exposure to 1α,25(OH)₂D₃ but not 25(OH)D₃. VDR mRNA (A) and protein quantity, including a representative Western blotting for the VDR using the VDR D-6 antibody (top bands) and GAPDH (lower bands) (B), in primary human myoblasts. CYP24A1 mRNA quantity in primary human myoblasts after a 24-hour exposure to 1nM or 100nM 1α,25(OH)₂D₃, 100nM 25(OH)D₃, or vehicle alone (C). Data are presented as the mean ± SEM (n = 6); *, P < .05; **, P < .01; ***, P < .001. 1,25D (1α,25(OH)₂D₃), 25D (25(OH)D₃).

Figure 5

1α,25(OH)₂D₃ exposure modulates the expression of key regulators of myogenic proliferation and differentiation. FOXO3A (A) and HES1 mRNA quantity (B) in human myoblasts exposed to 1nM or 100nM 1α,25(OH)₂D₃ or vehicle alone for 24 hours. Data are presented as the mean ± SEM (n = 6); **, P < .01; ***, P < .001. Visualization of the inhibited biological function differentiation of myoblasts, one of the top inhibited biological functions identified using IPA network analysis (C). 1,25D (1α,25(OH)₂D₃), 25D (25(OH)D₃).

Statistical analysis

For analysis of microarray data the statistical software R was employed (see details above). The statistical software SigmaPlot version 13.0 (Systat Software, Inc) was used for all other statistical analysis. All variables were log-transformed before statistical analysis. Statistically significant differences when comparing 2 groups were evaluated by the use of an unpaired or paired Student's t test for independent and dependent variables, respectively. Statistically significant differences when comparing more than 2 groups were evaluated by the use of a one-way ANOVA or a one-way repeated measures ANOVA. Tukey's post hoc test was used to locate differences in mean values. Differences were considered significant at P < .05. Data are presented as the mean ± SEM unless stated otherwise.

Results

Components of the vitamin D-endocrine system are readily detected in human muscle precursor cells while low to nondetectable in adult skeletal muscle

In order to compare the expression level of the components of the vitamin D-endocrine system at different levels of muscle cell differentiation, the mRNA and protein quantities of the VDR and the mRNA quantity of CYP27B1 and CYP24A1 were analyzed in human myoblasts, myotubes and adult skeletal muscle. The mRNA quantity of the VDR was higher in myoblasts (P = .012) and myotubes (P < .001) than in adult skeletal muscle and increased (P < .001) after differentiation of myoblasts into myotubes (Figure 1A. VDR protein was readily detected in both myoblast and myotubes and increased (P = .005) after differentiation of myoblasts into myotubes (Figure 1B. A comparison of the VDR protein quantity in myotubes with that in the human monocytic cell line THP1 revealed similar levels of VDR protein in these cell types (Figure 1C.

Figure 1

Components of the vitamin D-endocrine system are readily detected in human muscle precursor cells while low to nondetectable in human adult skeletal muscle. Basal VDR mRNA (A) and protein quantity, including a representative Western blotting for the VDR using the VDR D-6 antibody (top band at normal exposure and bottom band deliberately overexposed) (B), in human primary myoblasts, myotubes, and adult skeletal muscle (25 μg, n = 3). C, VDR protein quantity was analyzed in human myotubes (25 μg, n = 2), in adult skeletal muscle prepared in regular lysis buffer (RLB) and HLB (50 μg, n = 2), in HeLa whole-cell lysates (Pos ctrl) and in human monocytic cells (THP1 cell line) (25 μg, n = 1). Basal CYP27B1 (D) and CYP24A1 (E) mRNA quantity in human primary myoblasts, myotubes, and adult skeletal muscle (n = 3). Data are presented as the mean ± SEM; *, P < .05; **, P < .01; ***, P < .001. (ND), nondetectable; (overexp.), overexposed.

In contrast to myoblasts and myotubes, VDR protein was undetectable in adult skeletal muscle (Figure 1B. Neither overexposing the membrane (Figure 1B or loading 130–150 μg of total protein (Supplemental Figure 2A) resulted in detection of the VDR in adult skeletal muscle. In rodent skeletal muscle, ameliorated detection of VDR protein was recently reported by using a HLB, believed to be necessary for VDR to separate from tight binding to genomic DNA (12). We therefore performed additional Western blot analysis on human adult skeletal muscle prepared in HLB but were unable to detect VDR protein also by this approach (Figure 1C. In cells expressing the VDR, most VDR protein is confined to the nucleus (11). Hence, we further explored the presence of VDR protein in adult skeletal muscle by Western blotting performed on nuclear extracts from whole skeletal muscle biopsies. However, results from these analyses did not lend support to the presence of VDR protein in adult skeletal muscle tissue, as VDR protein was still nondetectable (Supplemental Figure 2B).

CYP27B1 mRNA quantity was higher in both myoblasts (P = .008) and myotubes (P = .028) than in adult skeletal muscle tissue, but was similar (P = .545) between myoblasts and myotubes (Figure 1D. CYP24A1 mRNA quantity was higher in both myoblasts (P = .014) and myotubes (P = .003) than in adult skeletal muscle tissue, but was not significantly different (P = .342) between myoblasts and myotubes (Figure 1E.

1α,25(OH)₂D₃, but not 25(OH)D₃, exert direct effects in human myoblasts

To study the ability of human muscle precursor cells to respond to direct 25(OH)D₃ exposure and to confirm responsiveness to 1α,25(OH)₂D₃, myoblasts were exposed to 1nM or 100nM 1α,25(OH)₂D₃, 100nM 25(OH)D₃, or vehicle alone for 24 hours. VDR mRNA quantity was increased (P = .037) after exposure to 100nM 1α,25(OH)₂D₃ and VDR protein quantity demonstrated a dose-dependent increase after exposure to 1nM and 100nM 1α,25(OH)₂D₃ (P = .004 and P < .001, respectively) compared with vehicle alone (Figure 2, A and B). There was no change in VDR mRNA or protein quantity (P = .808 and P = .575, respectively) after exposure to 100nM 25(OH)D₃ (Figure 2, A and B). Exposure to 1α,25(OH)₂D₃ induced a dose-dependent increase (P < .001) in CYP24A1 mRNA quantity compared with vehicle alone (Figure 2C. There was no change (P = .980) in CYP24A1 mRNA quantity after exposure to 100nM 25(OH)D₃ (Figure 2C.

1α,25(OH)₂D₃ inhibits the proliferation of human myoblasts without affecting cellular apoptosis or necrosis

The effect of 1α,25(OH)₂D₃ and 25(OH)D₃ on myoblast proliferation was analyzed by BrdU incorporation, an indicator of cell proliferation, after a 48-hour exposure to 1nM or 100nM 1α,25(OH)₂D₃, 100nM 25(OH)D₃, or vehicle alone. BrdU incorporation was reduced after exposure to both 1nM (P = .014) and 100nM 1α,25(OH)₂D₃ (P < .001) compared with vehicle alone (Figure 3A. BrdU incorporation was similar (P = .595) in myoblasts exposed to 100nM 25(OH)D₃ and vehicle alone (Figure 3A.

Figure 3

1α,25(OH)₂D₃ inhibits primary human myoblast proliferation without affecting necrosis or apoptosis. BrdU incorporation presented as OD values in primary human myoblasts after a 48-hour exposure to vehicle alone, 1nM, or 100nM 1α,25(OH)₂D₃, or 100nM 25(OH)D₃ (A). Data are presented as the mean ± SEM (n = 6); *, P < .05; **, P < .01. Percentage of total cells in early apoptosis, late apoptosis, and necrosis based on flow cytometry analysis for Annexin V and PI on human myoblasts exposed to vehicle alone, 1nM, or 100nM 1α,25(OH)₂D₃ for 48 hours (B). Data are presented as the mean ± SEM (n = 3). Representative dot plots for Annexin V and PI staining of human myoblasts exposed to vehicle alone, 1nM, or 100nM 1α,25(OH)₂D₃ for 48 hours (C). The P2 gate denotes early apoptosis, P3 denotes late apoptosis, and P4 denotes necrosis. 1,25D (1α,25(OH)₂D₃), 25D (25(OH)D₃).

To establish that the effect of 1α,25(OH)₂D₃ on myoblast BrdU incorporation was not caused by more cells undergoing apoptosis or necrosis, flow cytometry analysis was performed for Annexin V and PI on myoblasts exposed to 1nM and 100nM 1α,25(OH)₂D₃ or vehicle alone. This analysis revealed the percentage of cells in early and late apoptosis and necrosis to be similar between the groups (P = .517, P = .882 and P = .165, respectively) (Figure 3B.

1α,25(OH)₂D₃ inhibits human myotube formation but does not alter endogenous myostatin expression

The effect of 1α,25(OH)₂D₃ and 25(OH)D₃ on myoblast differentiation was explored by differentiating myoblasts for 8 days in low-serum media containing 1nM or 100nM 1α,25(OH)₂D₃, 100nM 25(OH)D₃, or vehicle alone. Exposure to 1nM and 100nM 1α,25(OH)₂D₃ during the differentiation process resulted in less myotubes per field of view (P = .012 and P < .001, respectively) and a reduced fusion index (P = .019 and P = .001, respectively) as compared with exposure to vehicle alone (Figure 4, A–C). Fusion index and myotubes per field of view were similar (P = 1.000 and P = .792, respectively) after exposure to 100nM 25(OH)D₃ and vehicle alone (Figure 4, A–C).

Figure 4

1α,25(OH)₂D₃ inhibits human myotube formation without associated changes in endogenous myostatin expression. Typical example of desmin staining used to analyze myotube formation after 8 days of differentiation in low-serum media containing vehicle alone, 1nM, or 100nM 1α,25(OH)₂D₃, or 100nM 25(OH)D₃ (A). White arrows indicate myotubes (desmin-positive cells with ≥ 3 nuclei). Scale bar, 200 μm. Myotubes per field (B) and fusion index (C) after 8 days of differentiation in the presence of vehicle alone, 1nM, or 100nM 1α,25(OH)₂D₃, or 100nM 25(OH)D₃. Data are presented as the mean ± SEM (n = 4); *, P < .05; **, P < .01. Myostatin mRNA quantity after 8 days of differentiation in the presence of vehicle alone, 1nM 1α,25(OH)₂D₃, or 100nM 25(OH)D₃ (D). Data are presented as the mean ± SEM (n = 6). 1,25D (1α,25(OH)₂D₃), 25D (25(OH)D₃).

An altered expression of endogenous myostatin has been suggested to partly explain the effects of vitamin D on rodent myoblast differentiation and myotube size (14, 15). To investigate this possibility, the effect of 1α,25(OH)₂D₃ and 25(OH)D₃ on the mRNA quantity of myostatin after 24 hours and 8 days of differentiation was studied. Myostatin mRNA quantity was unchanged (P = .597) after a 24-hour exposure to 1nM 1α,25(OH)₂D₃, 100nM 1α,25(OH)₂D₃, or 100nM 25(OH)D₃ as compared with vehicle alone (data not shown). Equally, despite a marked reduction of myotube formation after exposure to 1nM 1α,25(OH)₂D₃, myostatin mRNA quantity was unaltered after 8 days of exposure to 1nM 1α,25(OH)₂D₃ and 100nM 25(OH)D₃ as compared with vehicle alone (Figure 4D.

1α,25(OH)₂D₃ exposure alters the expression of key regulators of cell cycle progression and myogenic differentiation

In order to identify plausible signaling pathways that may mediate the effects of 1α,25(OH)₂D₃ on myoblast proliferation and differentiation, the gene expression profile of myoblasts exposed to 100nM 1α,25(OH)₂D₃ for 24 hours was explored using microarray analysis. With statistical significance defined as probe sets with a FDR of less than or equal to 5% and using a cut-off at more than or equal to 1.5-fold change, 189 genes were identified as up-regulated and 99 genes as down-regulated by 1α,25(OH)₂D₃ (Supplemental Table 1). Consistent with a profound inhibition of myoblast proliferation, differential expression analysis identified the expression of cyclin D2 to be down-regulated by approximately 3-fold in myoblasts after exposure to 1α,25(OH)₂D₃. When expanding the differential expression analysis to include all probe sets with a FDR of less than or equal to 5%, the gene expression of cyclin A2, B1, D3, and E1 were all identified as down-regulated (Supplemental Table 1). In addition, the Cdk-inhibitors p27 and p21 were both identified as up-regulated (Supplemental Table 1).

Differential expression analysis also identified the myogenic transcription factors MyoD, myogenin, and myocyte enhancer factor 2C (MEF2C), to be down-regulated after exposure to 1α,25(OH)₂D₃ (Figure 5A and Supplemental Table 1). In line with this, the gene expression of muscle structural proteins, including type I troponin I, type I troponin T, and type I troponin C were identified as down-regulated after exposure to 1α,25(OH)₂D₃ as compared with vehicle alone (Table 2). In addition, when expanding the differential expression analysis to include all probe sets with a FDR of less than or equal to 5%, the gene expression of type II troponin C, type I myosin heavy chain, type IIa myosin heavy chain, perinatal myosin heavy chain, neonatal myosin heavy chain, desmin, myozenin-2, type II regulatory myosin light chain, type II alkali myosin light chain, myopalladin, embryonic myosin light chain, and titin were also identified as down-regulated (Table 2).

Table 2

Muscle Structural Proteins Identified as Differentially Expressed (FDR ≤ 5%) in Human Myoblasts After 24 Hours of Exposure to 1α,25(OH)₂D₃

Muscle Structural Protein	Gene ID	Score	Numerator	Denominator	Fold Change2
Type I troponin I	TNNI1	−2.57	−0.73	0.28	−1.66
Type I troponin T	TNNT1	−2.51	−0.74	0.29	−1.67
Type I troponin C	TNNC1	−3.02	−0.94	0.31	−1.92
Type II troponin C	TNNC2	−0.99	−0.27	0.28	−1.21
Type I myosin heavy chain	MYH7	−1.12	−0.29	0.26	−1.22
Type IIa myosin heavy chain	MYH2	−1.11	−0.34	0.31	−1.27
Perinatal myosin heavy chain	MYH8	−1.10	−0.40	0.36	−1.32
Neonatal myosin heavy chain	MYH3	−1.44	−0.42	0.29	−1.34
Desmin	DES	−1.93	−0.49	0.26	−1.41
Myozenin-2	MYOZ2	−1.93	−0.57	0.30	−1.48
Myosin light chain, phosphorylatable, fast skeletal muscle	MYLPF	−1.50	−0.40	0.27	−1.32
Embryonic myosin light chain	MYL4	−1.68	−0.52	0.31	−1.43
Myosin light chain, alkali, fast skeletal muscle	MYL1	−2.32	−0.58	0.25	−1.49
Myopalladin	MYPN	−1.79	−0.57	0.32	−1.49
Titin	TTN	−1.52	−0.58	0.38	−1.50

Furthermore, pathways demonstrated to be of importance for myoblast proliferation and differentiation were affected by 1α,25(OH)₂D₃ exposure. These included an up-regulation of FOXO3 and a modulation of components of the Notch signaling pathway, including an up-regulation of the Notch target gene HES1 as well as down-regulation of HES6, HEYL, and the Notch receptor ligand delta-like 1 (Supplemental Table 1). The effect on FOXO3A and HES1 gene expression was validated by quantitative real-time PCR after 24 hours of 1α,25(OH)₂D₃ exposure, demonstrating that both 1nM and 100nM 1α,25(OH)₂D₃ exposure increased (P < .001) FOXO3A mRNA and dose dependently increased (P < .001 and P = .014, respectively) HES1 mRNA quantity (Figure 5B.

1α,25(OH)₂D₃ exposure affects GOs and biological functions associated with cell fate commitment and differentiation in human myoblasts

The identified genes regulated by 1α,25(OH)₂D₃ were further analyzed for GO enrichment of biological functions using DAVID. Ontologies enriched with a FDR of less than or equal to 1% were categorized as significant. GO enrichment of up-regulated genes related to exposure to 1α,25(OH)₂D₃ identified GO categories involved in “cell fate commitment,” “macromolecule biosynthesis,” “response to oxygen levels,” “hematopoietic or lymphoid organ development,” and “response to vitamin” (Table 2). GO enrichment of down-regulated genes identified GO categories involved in “muscle organ development” (Table 3).

Table 3

Enriched GO Categories in Human Myoblasts After 24 Hours of Exposure to 1α,25(OH)₂D₃ as Generated by DAVID

GO Term	List Hits	List Total	Population Hits	Population Total	Fold Enrichment	FDR (%)
Up-regulated processes
Cell fate commitment	11	162	70	5818	5.64	0.03
Positive regulation of macromolecule metabolic process	27	162	444	5818	2.18	0.31
Response to oxygen levels	10	162	77	5818	4.66	0.41
Hematopoietic or lymphoid organ development	13	162	140	5818	3.33	0.77
Response to vitamin	7	162	38	5818	6.62	0.91
Down-regulated processes
Muscle organ development	13	86	211	13 528	6.28	0.001

To reveal the possible biological interaction of specifically regulated genes, we performed network analysis using IPA. The analysis revealed 30 and 9 biological functions as activated and inhibited, respectively (Supplemental Table 2). Among the most highly activated biological functions were pathways annotated as “cell viability,” “migration of muscle cells,” “differentiation of mononuclear leukocytes,” “differentiation of lymphocytes,” “differentiation of blood cells,” and “differentiation of leukocytes.” In contrast, among the most highly inhibited biological functions were pathways annotated as “differentiation of myoblasts” and “differentiation of muscle cells” (Figure 5A and Supplemental Table 2).

Discussion

In the present study, a direct comparison of the expression level of components of the vitamin D-endocrine system in human muscle cells at different developmental stages, and direct effects of 1α,25(OH)₂D₃ and 25(OH)D₃ in human muscle cells, were studied. The major novel findings include: 1) a robust expression of components of the vitamin D-endocrine system, including VDR mRNA and protein, in human muscle precursor cells while low to nondetectable levels in human adult skeletal muscle; 2) 1α,25(OH)₂D₃, but not 25(OH)D₃, exert direct effects in human muscle precursor cells; and 3) 1α,25(OH)₂D₃ inhibits the proliferation and differentiation of human myoblasts without associated changes in endogenous myostatin expression, but by altering the expression of key regulators of cell cycle progression and myogenic differentiation.

We tested the hypothesis that in human skeletal muscle, VDR expression is predominantly located to muscle precursor cells. Using the stringently validated VDR D-6 antibody (9, 30), we demonstrate that VDR protein is readily detected in human myoblasts and myotubes while nondetectable in human adult skeletal muscle. In human myotubes, the VDR protein abundance is similar to that in human monocytic cells, which are well-established direct 1α,25(OH)₂D₃-targeted cells (31, 32). A predominant expression of the VDR in muscle precursor cells was verified at the mRNA level, as was the mRNA expression of the vitamin D activating and inactivating hydroxylases CYP27B1 and CYP24A1. These findings corroborate previous results from animal models, showing a predominant expression of the VDR in immature rather than adult muscle cells (12, 13). Although it cannot be excluded that VDR protein is expressed in adult skeletal muscle at levels below the detection limit of the methods employed in this study, the current results prove this expression to be markedly lower than in muscle precursor cells. These findings, together with previous findings in rodent skeletal muscle, indicate that the direct effects of vitamin D in adult skeletal muscle are likely to predominantly concern processes involving muscle precursor cells.

Previous studies in animal models have established that both 1α,25(OH)₂D₃ and 25(OH)D₃ exposure affects the proliferation and differentiation of muscle precursor cells (1, 7, 13–16, 18, 19, 33). The present findings expand this role of 1α,25(OH)₂D₃ to include effects on human muscle precursor cells. In contrast to 1α,25(OH)₂D₃, 25(OH)D₃ exposure did not affect human myoblast proliferation or differentiation, nor did it affect the expression of the VDR or the CYP24A1. This stands in contrast to previous studies performed in rodents and indicates that, whereas the vitamin D activating hydroxylase CYP27B1 is biologically active in rodent muscle precursor cells (13, 14), it lacks biological activity in human muscle cells.

We found 1α,25(OH)₂D₃ to have a marked dose-dependent inhibitory effect on human myoblast proliferation. This finding is in accordance with most studies performed in animal models (7, 13–15, 18). As such, it adds to the bulk of data establishing an inhibitory role of 1α,25(OH)₂D₃ on myoblast proliferation. In a previous study, a 48-hour exposure to 1α,25(OH)₂D₃ in C2C12 cells resulted in the reduction of c-myc and cyclin D1 and an increase of ataxia telangiectasia mutated and retinoblastoma 1 mRNA levels, whereas a reduction in phosphorylated retinoblastoma 1 protein and total c-myc protein levels was observed (14). In the present study, none of these genes were identified as differentially expressed in myoblasts exposed to 100nM 1α,25(OH)₂D₃ for 24 hours, even when expanding the differential expression analysis to include all probe sets with a FDR of less than or equal to 5%. However, consistent with an effect on the expression of genes that control the G₁/S restriction point, cyclin A2, D2, D3, and E1 were all found to be down-regulated in the current study. In fact, cyclin D2 was the second most highly down-regulated gene (approximately a 3-fold down-regulation). In addition, the Cdk-inhibitors p27 and p21 were both identified as up-regulated. Collectively, these findings establish a role for 1α,25(OH)₂D₃ in inhibiting human myoblast proliferation by modulating the gene expression of key regulators of cell cycle progression. Notably, the antiproliferative effect of 1α,25(OH)₂D₃ and modulation of the gene expression of cell cycle regulators demonstrated here are similar to those previously reported in human nonmuscle cells (34, 35). Thus, 1α,25(OH)₂D₃ appears to regulate proliferation and cell cycle progression by similar mechanisms in a multitude of human cell types.

The effects of 1α,25(OH)₂D₃ and 25(OH)D₃ on myoblast differentiation have been carefully investigated in previous studies performed in animal skeletal muscle models (14–18, 20). In the present study, we found a robust inhibitory effect of 1α,25(OH)₂D₃ on human myotube formation that was associated with a reduced expression of the myogenic regulatory factors MyoD, myogenin, and MEF2C. In accordance with the inhibitory effect of 1α,25(OH)₂D₃ on human myoblast differentiation, GO enrichment of down-regulated genes identified GO categories involved in muscle organ development and network analysis using IPA identified differentiation of myoblasts and differentiation of muscle cells among the most highly ranked inhibited biological functions. Even though vitamin D is often acknowledged to promote muscle cell differentiation (36), current literature demonstrates that this is a rather simplified view of vitamin D actions in muscle precursor cells. In fact, animal studies have demonstrated that 1α,25(OH)₂D₃ may both inhibit (14, 18, 19) and stimulate (15) myotube formation, as well as both down-regulate (4, 14, 18) and up-regulate the expression of myogenic regulatory factors (17, 20).

The mechanisms whereby 1α,25(OH)₂D₃ may influence myoblast differentiation have recently received considerable attention and are suggested to include effects on the gene expression of the myogenic regulatory factors Myf5, MyoD, and myogenin as well as the TGF-β family member myostatin (14–18, 20). In the current study, myostatin mRNA levels were not changed after exposure to 1α,25(OH)₂D₃ for 24 hours or 8 days. Although an effect on myostatin gene expression at other time points are not possible to exclude, this finding questions the hypothesis that regulation of endogenous myostatin expression is responsible for the effects of 1α,25(OH)₂D₃ on myoblast differentiation. Considering that previous studies reporting an effect of 1α,25(OH)₂D₃ on endogenous myostatin expression were conducted in animal skeletal muscle, it is plausible that these conflicting results may derive from species differences. However, in accordance with previous animal studies, we found both MyoD and myogenin to be down-regulated in human muscle precursor cells after 1α,25(OH)₂D₃ exposure. In addition, we demonstrated MEF2C to be down-regulated after exposure to 1α,25(OH)₂D₃. MEF2C has previously been identified as a key transcription factor in the formation of mature sarcomeres by driving the expression of myosin and myomesin (37, 38). Indeed, in the present study, exposure of myoblasts to 1α,25(OH)₂D₃ resulted in a down-regulation of the expression of muscle structural proteins, including myosin heavy and light chains.

In order to maintain the satellite stem cell pool in adult skeletal muscle, satellite cells are normally kept in a quiescent state and, once activated, part of the myoblast progeny are returned to quiescence by the process of asymmetric cell division (39). These processes involve a reversible exit from the cell cycle and an inhibition of myoblast differentiation, and are critically controlled by the FOXO3 and Notch signaling pathways (39–41). In the present study, 1α,25(OH)₂D₃ exposure inhibited both myoblast proliferation and differentiation, which is consistent with the events promoting a return of myoblasts to quiescence. This was associated with an induction of FOXO3 and a modulation of the Notch signaling pathway, including an increased expression of the Notch target gene HES1. Notably, a previous in vivo study in rodents has suggested that vitamin D deficiency may contribute to age-related skeletal muscle atrophy by down-regulating the Notch signaling pathway as well as reducing the satellite cell proliferative and skeletal muscle regenerative potential (42). Based on the current findings, we suggest that 1α,25(OH)₂D₃ may play an important role in satellite cell self-renewal and quiescence, possibly through the modulation of FOXO3 and Notch signaling pathways (Figure 6).

Figure 6

Proposed model of 1α,25(OH)₂D₃ actions in human adult skeletal muscle. Activation and subsequent expansion of satellite cells by asymmetric division results in Pax7⁺/Myf5⁺ myogenic progenitor cells (MPCs) committed to myogenic differentiation and activated Pax7⁺/Myf5⁻ satellite cells that return to reversible quiescence to replenish the satellite stem cell pool. Direct effects of 1α,25(OH)₂D₃ in human adult skeletal muscle may include the promotion of self-renewal of activated satellite cells, possibly mediated by a modulation of Notch and FOXO3 signaling pathways. Indirect effects of 1α,25(OH)₂D₃ on adult muscle fibers are also probable and may be mediated by altered systemic levels of factors that influence adult skeletal muscle form and function. 1,25D (1α,25(OH)₂D₃) Mstn (myostatin).

In conclusion, this study demonstrates that the expression of components of the vitamin D-endocrine system and direct effects of 1α,25(OH)₂D₃ in human skeletal muscle are predominantly located to muscle precursor cells. In human myoblasts, inhibition of myoblast proliferation by 1α,25(OH)₂D₃ is associated with an altered expression of cell cycle regulators consistent with a block of cell cycle progression at the G₁/S restriction point. Myotube formation is markedly reduced by 1α,25(OH)₂D₃ without concomitant changes in endogenous myostatin expression, but associated with a reduced expression of myogenic regulatory factors. These effects are consistent with a promotion of myoblast self-renewal and maintenance of the satellite stem cell pool, possibly mediated by a modulation of the FOXO3 and Notch signaling pathways. Future studies investigating the role of vitamin D in skeletomuscular health and function should therefore be directed towards the role of muscle precursor cells in these processes.

Acknowledgments

Author contributions: K.O. designed and performed the experiments, collected human vastus lateralis biopsies, analyzed and interpreted the data, prepared the figures, and drafted the manuscript; A.Sa. and S.A. performed the human myoblast extractions and cell culture, myoblast and myotube RNA and protein extractions and analysis together with K.O., and helped with the drafting of the manuscript; M.L. contributed to the work involving myoblast and myotube RNA extractions and analysis and helped with the drafting of the manuscript; A.St. performed flow cytometry analysis, analyzed and interpreted the data, and helped with the drafting of the manuscript; E.R. contributed to the design of the experiments, performed microarray data analysis, contributed to the interpretation of the data, and helped with the drafting of the manuscript; T.G. was involved in the design and coordination of the study and critically revised the manuscript. The final manuscript was read and approved by all authors. All authors agree to be accountable for the accuracy or integrity of all parts of the work.

This work was supported by funding from the Swedish Research Council, the Swedish National Centre for Research in Sports, the Swedish Heart Lung Foundation, and the Marianne and Marcus Wallenberg Foundation. K.O. is supported by a Clinical Scientist Training Program fellowship from Karolinska Institutet.

Disclosure Summary: The authors have nothing to disclose.

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Abbreviations

 
• ABAM

  antibiotic-antimycotic

 
• BrdU

  bromodeoxyuridine

 
• CYP24A1

  cytochrome p450 24A1

 
• CYP27B1

  cytochrome p450 27B1

 
• DAVID

  Database for Annotation, Visualization and Integrated Discovery

 
• FBS

  fetal bovine serum

 
• FDR

  false discovery rate

 
• FOXO3

  forkhead box O3

 
• GAPDH

  glyceraldehyde-3-phosphate dehydrogenase

 
• GO

  gene ontology

 
• HES1

  hairy and enhancer of split 1

 
• HLB

  hyperosmolar lysis buffer

 
• IPA

  Ingenuity Pathway Analysis

 
• MACS

  magnetic-activated cell sorting

 
• MEF2C

  myocyte enhancer factor 2C

 
• MyoD1

  myogenic differentiation 1

 
• 25(OH)D₃

  25-hydroxy-vitamin D₃

 
• 1α,25(OH)₂D₃

  1α,25-dihydroxy-vitamin D₃

 
• PI

  propidium iodide

 
• RPS18

  ribosomal protein 18

 
• 18S

  eukaryotic 18S rRNA

 
• UBB

  ubiquitin B

 
• UBC

  ubiquitin C

 
• VDR

  vitamin D receptor.