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. 2015 Oct 9;157(1):4–15. doi: 10.1210/en.2015-1443

Thyroid Hormone Receptor α Plays an Essential Role in Male Skeletal Muscle Myoblast Proliferation, Differentiation, and Response to Injury

Anna Milanesi 1,✉, Jang-Won Lee 1, Nam-Ho Kim 1, Yan-Yun Liu 1, An Yang 1, Sargis Sedrakyan 1, Andrew Kahng 1, Vanessa Cervantes 1, Nikita Tripuraneni 1, Sheue-yann Cheng 1, Laura Perin 1, Gregory A Brent 1,✉
PMCID: PMC4701883  PMID: 26451739

Abstract

Thyroid hormone plays an essential role in myogenesis, the process required for skeletal muscle development and repair, although the mechanisms have not been established. Skeletal muscle develops from the fusion of precursor myoblasts into myofibers. We have used the C2C12 skeletal muscle myoblast cell line, primary myoblasts, and mouse models of resistance to thyroid hormone (RTH) α and β, to determine the role of thyroid hormone in the regulation of myoblast differentiation. T3, which activates thyroid hormone receptor (TR) α and β, increased myoblast differentiation whereas GC1, a selective TRβ agonist, was minimally effective. Genetic approaches confirmed that TRα plays an important role in normal myoblast proliferation and differentiation and acts through the Wnt/β-catenin signaling pathway. Myoblasts with TRα knockdown, or derived from RTH-TRα PV (a frame-shift mutation) mice, displayed reduced proliferation and myogenic differentiation. Moreover, skeletal muscle from the TRα1PV mutant mouse had impaired in vivo regeneration after injury. RTH-TRβ PV mutant mouse model skeletal muscle and derived primary myoblasts did not have altered proliferation, myogenic differentiation, or response to injury when compared with control. In conclusion, TRα plays an essential role in myoblast homeostasis and provides a potential therapeutic target to enhance skeletal muscle regeneration.

Thyroid hormone (TH) acts as a pleiotropic factor during development and regulates genes involved in growth and differentiation (1–3). The genomic actions of T3 are mediated by TH nuclear receptor (TR) α and β, which are ligand-inducible transcription factors (4, 5). TR α and β are expressed with distinct developmental patterns and tissue distribution. Pharmacologic and genetic approaches have demonstrated TR isoform–specific actions. Local ligand activation and inactivation in tissues by deiodinase enzymes is also critical for development (6). T3 has important actions in skeletal muscle and a number of T3-responsive genes coding for muscle structural proteins and ion transporters have been identified (7–9). These genes include myosin heavy chain and Serca1 (10).

Skeletal muscle is a striate tissue and it is composed of contractible multinuclear myofibers (10, 11). Myogenesis is required for normal skeletal muscle development and for maintenance and repair of adult myofibers. Vertebral skeletal muscle is derived from cells in the prechordal and somitic mesoderm. In myogenesis, myoblasts develop from mesenchymal precursor cells and through proliferation and differentiation progress to myogenic lineage. These cells then fuse to form multinucleated myofibers (11). Skeletal muscle myogenesis is disrupted in several pathological conditions, including diabetes, obesity (12, 13), muscular dystrophy (14) and mitochondrial myopathy (15). TH excess in humans is associated with proximal muscle weakness, likely due to both reduced muscle mass and an enhancement of type 2 fast-twitch muscle fibers (16, 17). A range of rodent models with TR isoform mutations and knockouts have been developed; however, relatively few studies of skeletal muscle in these models has been reported. Skeletal muscle isolated from TR α and β knockout mice showed a switch from type 2 fast-twitch muscle fibers to type 1 slow-twitch muscle fibers (18). Skeletal muscle from TRα-null mice had a 20–60% prolongation of contraction and relaxation times compared with muscle from TRβ-null and wild-type (WT) animals (19). TRα is important for metabolic regulation in liver and fat, and the various TRα mutation mouse models are associated with distinct metabolic phenotypes (20–22). Interestingly, increased metabolic rate in skeletal muscle has been described in humans with resistance to thyroid hormone (RTH) associated with dominant negative mutations in TRβ. This is primarily a result of elevated levels of circulating TH stimulating WT TRα in muscle and fat (23). In families with RTH due to TRα mutations, in which TRβ-mediated feedback to TSH is normal and TH levels are not elevated, there is evidence of reduced TH action in tissues (24, 25). The phenotypes in these individuals are variable, but manifestations of RTH-TRα mutations include delayed growth, constipation, and abnormal bone formation, as well as reduced metabolism.

In a model of skeletal muscle development, addition of T3 to the myogenic culture medium after induction of terminal differentiation induced a nearly 2-fold stimulation of myoblast differentiation (26). Interestingly, mice with knockout of the 5′-deiodinase 2 (D2) gene, the enzyme that converts the prohormone T4 to the active form T3, results in impaired in vitro differentiation of muscle-derived stem cells to myotubes and defective in vivo muscle regeneration after injury (27). More recently, satellite cell-specific ablation of the 5-deiodinase 3 gene, the enzyme that converts T3 to the inactive reverse T3, also impaired skeletal muscle regeneration (28). These findings support a role for T3 in muscle development, function, and adult muscle regeneration, with specific timing of T3 activation and inactivation required, as has been shown in sensory development (29). However, a recent study of mice with selective skeletal muscle myocyte D2 inactivation showed a minor effect on skeletal muscle T3 levels and T3-dependent gene expression (30).

In this study, we show that knockdown or mutation of TRα impairs myoblast proliferation and differentiation to the myogenic lineage. Understanding the mechanisms of T3 action in muscle regeneration may be important to identify novel therapeutic targets for disorders associated with impaired adult myogenesis as well as clinical management of TH excess and deficiency.

Materials and Methods

Primary myoblast preparation, cell culture and differentiation

Primary murine myoblasts were established using the previously described protocol with some modifications (31). Muscle was isolated from 6–8 week old male C57Bl6 (Jackson Laboratory) and 8 month male TRα1PV, TRβPV (32) and WT mice (33). The RTH mouse models are heterozygous for the RTH-associated PV mutation (a frame-shift mutation), introduced into TRα (TRα1PV) (33) or TRβ (TRβPV) (32). Animals were euthanized by cervical dislocation and hind-limb muscles were removed and minced with a razor blade. Isolated muscles were washed with two changes of PBS and placed into fresh PBS medium in a 60-mm dish. Clean muscle pieces were cut into 400-μm3 cubes with jeweler's forceps. A minimum of five pieces of muscle per well were placed into collagen coated six-well plates in 2 mL DMEM/F12 (Invitrogen) media containing 20% fetal bovine serum supplemented with 20 ng/mL each of EGF and basic fibroblast growth factor, and incubated at 37°C and 5% CO2. Once explant outgrowth was established, individual wells were fed by the addition of 500 μL increments of medium to adjust for medium acidification as a result of increased cell density. When the wells were full and cells near confluence, they were fed by replacement of 50% of the medium each time to ensure the maintenance of “conditioning” factors secreted by the cells.

C2C12 murine skeletal muscle myoblasts (from ATCC) and primary myoblasts were cultured in proliferating medium consisting of Dulbecco's Modified Eagle Medium and 15% fetal bovine serum and maintained at subconfluent density. To induce myogenic differentiation, the medium with serum was removed and a medium with Dulbecco's Modified Eagle Medium and 2% serum replacement supplement (Sigma) was added when the cells were approximately 70% confluent. The medium was changed three times per week.

T3, and a selective TRβ agonist (GC1), were used at 10nM concentration in differentiation experiments. GC1, compared with T3, has a comparable affinity for hTRβ1 and a 10-fold lower affinity for hTRα1 (34, 35). In these experiments, serum replacement with the addition of T3 or GC1 was used to eliminate the effects of any endogenous THin the serum. We did not use serum replacement or charcoal stripped serum in the proliferation experiments, because both block proliferation. In proliferation studies, cells were incubated with bromodeoxyuridine (BrdU; Sigma) for 3 hours before cell staining. In some experiments we costained for Ki67 and BrdU.

Gene Silencing

C2C12 cells and primary myoblasts were plated in six-well dish control plates and transfected with TRα shRNA plasmid (Santa Cruz Biotechnology) and control shRNA plasmid, using the recommendations of the manufacturer. We treated the cells with puromycin (2 μg/mL) to select a stable-transfected C2C12 cell line. No puromycin selection was used for primary myoblasts. Knockdown efficiency was evaluated by PCR for the targeted mRNA and Western blotting for the targeted protein after puromycin selection for C2C12, and 48 hours after knockdown for primary skeletal muscle myoblast.

RNA isolation and PCR/RT-PCR analysis

Total RNA was extracted using RNeasy Minikit (Quiagen), according to the manufacturer's instructions. cDNA was prepared using superscript reverse transcriptase (Invitrogen). cDNA samples derived from 50 ng of total RNA were analyzed by PCR and/or qRT-PCR using SYBR green dye with QuantiTect SYBR Green RT-PCR kit (QIAGEN). Primers used for PCR are listed in Supplemental Table 1. Basic Local Alignment Search Tool (BLAST) searches were conducted on all primer sequences to ensure gene specificity. PCR was performed with cycles at 95°C for 30 seconds, 60°C for 30 seconds, and 72°C for 2 minutes for a total of 30 cycles. Each amplification reaction for the qRT-PCR was performed in triplicate and included the addition of the SYBR Green Master Mix (Applied Biosystems) in iCycler (Bio-Rad) following the manufacturer's protocol. Relative quantitative analysis was performed following 2−ddCT. Housekeeping genes for RT-PCR control, TRα, and TRβ primers were purchased from QIAGEN (mouse β-actin, NM_007393; 18 S rRNA, NR_003278.3; mouse TRα, NM_178060; mouse TRβ, NM_001113417).

Western blotting

Nuclear and cytoplasmic extracts were made from C2C12 and primary myoblasts following the nuclear extract kit manufacturer's instructions (Active Motif). The protein concentrations within each lysate were determined by the Bradford protein assay (Bio-Rad). The fractionated extracts were mixed with Laemmli sample buffer and equal quantities of the samples were loaded and separated by SDS-PAGE, transferred to polyvinylidene difluoride membranes (Bio-Rad), and blocked with 5% nonfat milk. The membranes were incubated in primary antibody overnight at 4°C. The membranes were then washed in TBST (10mM Tris, 140mM NaCl, and 0.1% Tween-20 [pH, 7.6]), incubated with the appropriate secondary antibody, and washed again in TBST. Bands were visualized by enhanced chemiluminescence and exposed with Chemidoc (Bio-Rad). Western blot analysis was performed using the primary and the horseradish peroxidase-conjugated secondary antibodies (Cell Signaling) listed in Supplemental Table 2.

Immunohistochemistry

Cells were fixed with 2% paraformaldehyde and treated with 0.2% triton. Cells prepared for BrdU staining were pretreated with hydrogen chloride. Skeletal muscle was collected 14 days after skeletal muscle injury and fixed overnight with 4% paraformaldehyde at 4°C. The tissue was then dehydrated in graded ethanol, cleared in xylene, and finally embedded in paraffin. For immunohistochemical staining we used primary antibodies listed in Supplemental Table 2. After washing with PBS, detection of bound primary antibodies was carried out with appropriate secondary antibodies conjugated with Alexa Fluor 488 or 568 (Life Technologies). Nuclear DNA was counterstained with 4′,6-diamidino-2-phenylindole (DAPI; Vector Lab).

Animal and muscle injury

The generation of TRα1PV (33) and TRβPV (32) mice was previously described. We used 8–10-month-old male TRα1PV, TRβPV, and WT mice and injected cardiotoxin (CTX; from Sigma: 25 μL, 0.03 mg/mL) in the right tibialis anterior muscle (TAM) to induce muscle injury. The left TAM was used as an internal control and a volume of saline equal to the CTX was injected. TRβPV mice have elevated TH levels, compared with WT. To normalize TH levels in the mutant and WT mice, we induced hypothyroidism in the TRβPV mice with iodine-deficient diet (0.15% propylthiouracil, from Teklad) for 4 weeks and replaced to physiological levels of TH (T3 7 ng/g ip) daily (22) for the duration of the experiment. All mice were euthanized 14 days after the injury and the TAMs were collected, embedded in paraffin, and sectioned for histopathological studies. All animal procedures were performed in accordance with approved protocols by the Institutional Animal Care and Use Committee at VA Greater Los Angeles Healthcare System and Children's Hospital–Los Angeles.

Confocal microscopy and histopathological analysis and image quantification

Images were acquired with a fluorescent microscope (Model Upright, Zeiss). Scanning confocal images for immunofluorescence analysis were obtained by a laser scanning confocal microscope (Leica Microsystems SP5). Muscle fiber size and counting was performed using Aperio Scanscope AT Turbo and analyzed using Aperio ImageScope software (Leica Biosystems). The in vivo fusion index was determined as previously described (36). Briefly, at least five hematoxylin and eosin–stained cross-sectional sections were used for each mouse. Myofibers and central nuclei counting was performed as described above. The in vivo fusion index was calculated as the percentage of tibialis anterior myofibers containing multiple central located nuclei vs the total number of myofibers. Satellite cell abundance was assessed by paired box transcription factor 7 (Pax7) staining and identifying cells that were PAX7+ and DAPI+ were counted.

Statistical analysis

Values are presented as mean ± SD. For all quantitative analyses, a minimum of three replicates were performed in independent experiments or in individual mice. Significant difference between means was determined using a two-tailed Student t test or one-way ANOVA followed by the Student-Newman-Keuls test.

Results

T3 stimulates myoblast differentiation

We used C2C12 cells, a well-characterized murine skeletal muscle myoblastic cell line, derived from satellite cells, that retains properties of the progenitor lineage (37) and primary myoblasts isolated from murine skeletal muscle. Both C2C12 cells and primary myoblasts were successfully differentiated to myotubes after serum withdrawal (2% serum replacement). Under these myogenic conditions, myotube formation was observed at day 3 and spontaneously contracting myotubes appeared around day 7 (Figure 1A; Supplemental Video 1). The influence of T3 on myogenic differentiation was tested by adding T3, which stimulates TRα and TRβ, or GC1, a selective agonist of TRβ. We observed spontaneous myofiber contraction as early as day 5 in C2C12 cells treated with T3, and a significant increase above baseline at day 6, 7, and 8 (Figure 1B). In contrast, with GC1 treatment, a significant increase in myofibers above baseline was seen only on day 8. The myofiber number was significantly higher in T3 treated, compared with GC1, at each time point (Figure 1B). The finding of a robust response to T3, and a delayed and modest response to GC1, suggest that stimulation of TRα is essential for normal differentiation. TRα mRNA is highly expressed in proliferating C2C12 compared with the low expression of TRβ mRNA (Figure 1C), suggesting that TRα is the dominant thyroid receptor isoform in myoblasts. We found a similar high expression of TRα mRNA and low TRβ mRNA expression in primary myoblast cultures (Figure 1D).

Figure 1.

Figure 1.

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In vitro differentiation of mouse C2C12 myoblasts, and skeletal muscle myoblasts from primary culture, into skeletal muscle myotubes after serum withdrawal. A, Light microscopic analysis of myotubes from C2C12 and primary myoblasts at day 7. B, Number of spontaneously contracting myotubes derived from C2C12, per microscopic visual high-powered field, after treatment with T3 (10nM), GC1 (10nM), or no hormonal treatment (control), shown from d 5–8 (mean of three replicates). C, Expression of TR isoform mRNA from proliferating C2C12 cells prior to differentiation. D, Expression of thyroid TR isoform mRNA in proliferating primary myoblasts prior to differentiation. Scale bar = 50 μM; *, P < .01; **P < .001.

Knockdown of TRα impairs myoblast proliferation and differentiation

To investigate the role of TRα in myoblasts, we selectively reduced TRα mRNA expression using small hairpin RNA for TRα (shTRα) in both C2C12 cells and primary myoblasts (Figure 2). shTRα C2C12 cells had almost complete knockdown of TRα mRNA after selection with puromycin (Figure 2A). These cells displayed a marked reduction in proliferation capacity compared with control C2C12 cells, as shown by a marked reduction in cell growth after 72 hours in the growth medium (Figure 2B). Impaired proliferation in shTRα -C2C12 cells was confirmed by a significant reduction in bromodeoxyuridine (BrdU) incorporation following a 3-hour incubation (Figure 2, C and D). A similar result of reduced proliferation after TRα mRNA knockdown was obtained in primary myoblast culture (Figure 2, E–H). The efficiency of TRα knockdown in primary culture is shown in Figure 2E. Primary myoblasts showed significant reduction in cell growth (Figure 2 F) and in BrdU incorporation after a 3-hour BrdU pulse (Figure 2, G and H) compared with control cells.

Figure 2.

Figure 2.

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Role of TRα in myoblast proliferation in C2C12 cell line and primary myoblasts. A, TRα mRNA in C2C12 cells after transfection with shTRα compared with control. B, Number of cells cultured in growth medium and collected per plate at specific time points. Graph represents the average cell number ± SD of five independent experiments. C, Immunostaining of control and shTRα-transfected C2C12 cells after 3 h in culture, BrdU (thymidine analog stains green), and DAPI (stains nuclei blue). D, Percentage of C2C12 BrdU-positive nuclei relative to the total, indicating proliferating cells. E, TRα RNA in primary myoblasts after transfection with shTRα, compared with control. F, Number of primary myoblasts cultured in growth medium and collected per plate at specific time points. Graph represents the average cell number ± SD of five independent experiments. G, Immunostaining of control and shTRα-transfected primary myoblasts after 3 h in culture; BrdU (green), nuclei (blue). H, Percentage of BrdU-positive nuclei in primary myoblasts. Scale bar = 50μ; *, P < .01; **, P < .001.

We further evaluated the mechanism underlying the severe impairment in proliferation after TRα knockdown. Knockdown of TRα in proliferating C2C12 cells dramatically reduced the expression of β-catenin, Cyclin D1, a downstream marker of Wnt/β-catenin signaling; and Wnt10b mRNA, important in regulating myogenic vs adipogenic lineage in the myoblast (38) (Figure 3A). β-catenin (Figure 3, B and C), and Cyclin D1 (Figure 3C) protein expression was significantly lower in shTRα C2C12 cells, compared with control. Moreover, the active form of β-catenin was similarly reduced in both nuclei and the cytoplasm of shTRα CC12 cells, compared with the control (Figure 3D).

Figure 3.

Figure 3.

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β-catenin expression in C2C12 myoblast cells. A, Fold change in β-catenin, cyclin D1, and Wnt10b mRNA in C2C12 myoblast cells after knockdown of TRα mRNA, shTRα, compared with control shRNA (control expression set at 1, broken black line). B, Staining of C2C12 myoblast–proliferating cells for β-catenin (green) and nuclei (blue) after transfection of control RNA and shTRα. C, Fraction (%) of C2C12 myoblast cell nuclei positive for β-catenin and cyclin D1 after transfection of control RNA (black bar) and shTRα (gray bar). D, Western blot of nuclear and cytoplasmic fraction of nonphosphorylated (active) β-catenin in C2C12 myoblast cells after transfection of control RNA and shTRα. Controls for the Western blot are β-actin for the nuclear and Histone H3 for the cytoplasmic fraction. scale bar = 50μ; *, P < .01; **, P < .001.

We further investigated the differentiation potential of C2C12 and primary myoblasts after TRα knockdown. TRα knockdown in primary myoblasts, as well as C2C12 cells, was associated with reduced expression of transcription factors important for the myogenic program, including PAX7, myoblast determination protein (MyoD), and myogenic factor 5 (MyF5) (Figure 4, A). In proliferating condition we observed more than 50% reduction in expression of Pax7, MyoD, and Myf5 protein after TRα knockdown (Figure 4, A and B). We did not detect troponin, which is a marker of terminal differentiation, in proliferating myoblasts. With induction of differentiation (d 2 after switching myoblast culture to differentiation medium) we observed a significant increase of both Myf5 and MyoD protein in the control myoblasts. MyoD and Myf5 are important myoblast regulatory factors that promote differentiation (39). After TRα knockdown in myoblasts, the increase in Myf5 and MyoD protein expression was significantly blunted, compared with control cells (Figure 4, A and B). Troponin protein expression was increased in both control and TRα knockdown cells 2 days after initiating differentiation with increased expression after TRα knockdown (Figure 4).

Figure 4.

Figure 4.

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Expression of Pax7, Myf5, MyoD, and troponin during proliferation and differentiation in primary myoblasts transfected with shTRα and control RNA. A, Western blot of skeletal muscle myoblasts treated with shTRα (shTRα) and control cells (−) during proliferation and 2 d after inducing differentiation. B, The relative level of Pax7 (B), Myf5 (C), MyoD (D), and troponin protein were estimated by normalizing OD units to β-actin density and expressing it as a fraction (%) of control protein. *, P < .05; **, P < .001.

The shTRα transfected C2C12 cells were able to form myotubes in vitro (Figure 5A); however, at day 7 in differentiation medium they showed a significantly lower number of myotubes compared with control, and failed to develop into functional spontaneous contracting myofibers (Figure 5A). shTRα-derived myotubes had a significantly lower fusion index, measured as the average nuclei per myotube (Figure 5B) and lower expression of skeletal muscle-related gene mRNA, such as Serca1, compared with control cells (Figure 5C). Moreover, we observed a progressive loss of shTRα-derived myotubes at 2 weeks, whereas multiple fully contracting myotubes were present in control cells.

Figure 5.

Figure 5.

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Role of TRα expression in C2C12 myoblast differentiation. A, Myogenic differentiation of C2C12 myoblasts after transfection of control RNA and shTRα RNA under light microscopy at d 7. B, Average nuclei per myotube (fusion index) at d 7 of differentiation. C, Sarcoendoplasmic reticulum calcium ATPase 1 (SERCA1) mRNA expression at d 7 during differentiation in control and shTRα-transfected C2C12 myoblasts. Magnification, 20×; **, P < .001.

TRα1PV mice have reduced type 2 fast-muscle fibers and PAX7-expressing cells

To further investigate the role of TRα in myoblasts we used a mouse model with an RTH-associated mutation, PV (a frame-shift mutation), introduced into the TRα gene (TRα1PV) (33). We compared the TRα1PV mice to WT mice and another mouse model with the PV RTH-associated mutation introduced into the TRβ gene, TRβPV. Type 2 fast-muscle fibers were significantly decreased in the TAM of the TRα1PV mice compared with control and TRβPV mice (Figure 6, A and B). Moreover, the number of cells expressing PAX7 in the TAM of the TRα1PV mice was significantly reduced compared with both WT and TRβPV mice (Figure 7, A and B), suggesting a reduced number of quiescent satellite cells in the TRα1PV mice.

Figure 6.

Figure 6.

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type 2 fast-fiber expression in TAM of TRα1PV, TRβPV, and WT mice. A, Immunostaining for MHCII (myosin heavy-chain 2, red) in TAM of WT, TRβPV, and TRα1PV mice. B, Percentage (%) of fiber positive for MHCII in TAM of WT, TRβPV, and TRα1PV mice. Scale bar = 50 μm; **, P < .001.

Figure 7.

Figure 7.

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PAX7 expression in the TAM of TRα1PV, TRβPV, and WT mice. A, Immunostaining for PAX7, (green) in TAM of WT, TRβPV, and TRα1PV mice. Arrows suggest Pax7+ cells. B, Number of PAX7-positive cells per high-power field in TAM of WT, TRβPV, and TRα1PV mice. Nuclei: DAPI. Scale bar = 50 μm; **, P < .001.

TRα1PV mice have reduced skeletal muscle–derived myoblast proliferation and have impaired skeletal muscle regeneration after injury

TRα1PV-derived primary myoblasts had a very slow growth rate, compared with WT or TRβPV-derived myoblasts (Figure 8, A and B). There was a significant reduction in TRα1PV-derived myoblast proliferation, as shown by reduced BrdU incorporation, 3 hours in culture medium (Figure 8, A and C), as well as reduced Ki67 expression (Figure 8, A and D). Approximately 50% of the TRα1PV myoblasts were double negative (Figure 8A; arrows) for BrdU and Ki67, compared with approximately 1% of the WT myoblasts and 10% of the TRβPV myoblasts, indicating that a significant number of TRα1PV myoblasts did not enter the cell cycle. Moreover, we observed a reduction in expression of the myogenic transcription factors MyoD and Myf5 in TRα1PV myoblasts (Figure 9, A, C, and D), as previously shown with knockdown of TRα, but did not observe a significant reduction in PAX7 (Figure 9, A and B).

Figure 8.

Figure 8.

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Primary myoblasts from mice heterozygous for RTH-associated mutation, PV, introduced into TRα (TRα1PV) and TRβ (TRβ PV). A, Immunostaining for BrdU (green) after 3 h in culture and Ki67 (red), and nuclei (blue) in proliferating myoblasts from WT, TRα1PV, and TRβPV mice. Arrows indicate cells negative for BrdU and Ki67. B, Fold increase in total cell number after 5 d in culture. C, Percentage of BrdU-positive nuclei in myoblasts from WT, TRβPV, and TRα1PV mice. D, Percentage of Ki67-positive nuclei in myoblasts from WT, TRβPV, and TRα1PV mice. Scale bar = 50μ; *, P < .05; **, P < .001.

Figure 9.

Figure 9.

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Expression of protein markers of muscle differentiation in skeletal muscle from WT and RTH-associated TRβPV and TRα1PV mutant mice. A, Western blot demonstrating expression of Pax7, Myf5, MyoD, and β-actin proteins in skeletal muscle myoblasts from WT, TRβPV, and TRα1PV mice. The relative level of Pax7 (B), Myf5 (C), and MyoD (D) protein was determined by normalizing OD units to β-actin density and expressing it as a fraction (%) of control protein. **, P < .001.

Myofibers from TRα1PV mice were smaller, compared with WT and age-matched TRβPV mice (Figure 10A). Two weeks after inducing skeletal muscle injury with CTX, regenerating muscle fibers from TRα1PV mice were significantly smaller, compared with regenerating muscle fibers from WT and TRβPV mice (Figure 10A). Histological analysis, 14 days after CTX, showed an immature skeletal muscle phenotype in all mice (TRα1PV, TRβPV, and WT), consisting of smaller muscle fibers in the regenerating muscle and centrally located nuclei (Figure 10A). This is consistent with the immature skeletal muscle phenotype describe in CTX-injured muscle, up to 4 months from the injury (40). However, there was a striking reduction in the number of nuclei per muscle fiber in regenerating TRα1PV, compared with WT and TRβPV mice. In vivo fusion index, defined as the percentage of regenerating fiber with more than one centralized nucleus in cross-sectional area, was dramatically lower in the TRα1PV mice compared with WT and TRβPV mice (Figure 10B). Moreover, the reduction in muscle fiber cross-sectional area (CSA) observed 14 days after CTX in TAM was significantly more pronounced in TRα1PV mice than WT and TRβPV mice (Figure 10C). In the WT mice and TRβPV mice we observed larger regenerating myofibers (3500–5000 μm2) 14 days after injury, similar to the contralateral uninjured muscle fibers (Figure 10D). In the TRα1PV mice, smaller myofibers (< 1000 μm2) with the absence of larger fibers (1200–2000 μm2), was seen compared with the contralateral uninjured muscle (Figure 10D). These data suggest an essential role for TRα in the optimal fusion and regeneration of myofibers after muscle injury.

Figure 10.

Figure 10.

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Skeletal muscle regeneration 14 d after CTX-induced muscle injury in WT, TRβPV, and TRα1PV mice. A, Hematoxylin and eosin stain of paraffin section from the TAM uninjured control and CTX-injured TAM in WT, TRβPV, and TRα1PV mice, 14 days after CTX injection. Insert panels show higher magnification of representative images. B, In vivo fusion index for regenerating myofibers in WT, TRβPV, and TRα1PV mice. C, Percent reduction in cross-sectional area (CSA) after CTX-induced skeletal muscle injury in WT, TRβPV, and TRα1PV mice. D, Frequency distribution of myofiber CSA (in μm2) in uninjured control TAM and post–skeletal muscle injury (post-CTX) in WT, TRβPV, and TRα1PV mice (dark blue area, uninjured TAM from TRα1PV; light blue area, injured TAM from TRα1PV; dark green area, uninjured TAM from WT; light green area, injured TAM from WT; pink area, uninjured TAM from TRβPV mice; purple area, injured TAM from TRβPV mice). Scale bar = 100μ; *, P < .05; **, P < .001.

Discussion

We investigated the role of TRα in the regulation of myoblast proliferation and differentiation as well as skeletal muscle regeneration in response to injury. We identified specific actions of TRα in proliferation and differentiation of skeletal muscle myoblasts. T3 has previously been shown to stimulate muscle growth by increasing the number and diameter of the muscle fibers (41), potentiating myoblast differentiation (26), regulating the transition between neonatal and adult myosin isoforms (42), and the contractile properties of adult myofiber (43). Similarly, our data showed that T3 treatment of myoblast cells increased myofiber differentiation. Treatment with the TRβ selective agonist, GC1, had a reduced effect on myofiber differentiation, compared with T3 treatment, consistent with the predominance of TRα isoform in myoblasts.

In limited previous studies, disruption of the TR α and β genes in mice were reported to modestly impair muscle development. The role of TR in adult myoblast differentiation and proliferation, in the context of muscle regeneration after injury, however, has not previously been reported. Recent studies have established a connection between TH signaling and the intrinsic pathways important in progenitor/stem cell physiology and homeostasis (44). TRα1 mediates TH signaling in the regulation of mouse intestinal progenitor/stem cell proliferation (45). Regeneration of the intestinal epithelium requires TRα1 expression (46). The mechanism involves a complex interplay of TRα1 with components of the Wnt/β-catenin pathway (47). The canonical Wnt pathway is activated when Wnt ligands bind to Frizzled receptor, allowing stabilization and nuclear translocation of β-catenin (48). In intestinal stem cells, TRα1 modulates the Wnt pathway through several mechanisms. TRα1 is a direct transcriptional regulator of β-catenin (49) and Frizzled-related sFRP2 (50), but also binds to Wnt targets. TRα1 may also act by stabilizing the β-catenin/Tcf4 complex when bound to DNA (47), increasing expression of Wnt-target genes and enhancing cell proliferation (51). A link between TR and the Wnt/β catenin pathway has also been described in chondrocytes differentiation and bone maturation (52–54). Similar to these reports, we found that TRα plays an important role in skeletal muscle myoblast by modulating the Wnt/β-catenin pathway. In murine myoblasts lacking TRα, we observed reduction of expression of β-catenin at the mRNA and protein levels, and reduction of the nuclear localization of β-catenin. This interplay between TRα and the Wnt canonic pathway may have an important role in regulating skeletal muscle myoblasts proliferation and differentiation.

Important myogenic transcription factors, such as Pax7, MyoD, and Myf5, were down-regulated during both proliferation and early differentiation in myoblasts after TRα knockdown. PAX7 is essential for satellite cell function and regulation of self renewal and expansion of adult skeletal muscle (55), and MyoD and Myf5 are important factor in promoting differentiation. It is well known that MyoD is transcriptionally stimulated by T3 (56). Moreover, it is interesting that in proliferating avian myoblasts, TRα1 physically interacts with MyoD and this interaction induces TR signaling, but represses MyoD. The MyoD/TRα complex is replaced by the TRα/RXR complex in the beginning of terminal differentiation, leaving MyoD able to stimulate gene expression (57). This suggests a time-sensitive interplay between TRα and MyoD during myogenic proliferation and differentiation. Gene knockout has shown that the absence of MyoD delays the early proliferation response of satellite cells and affects their differentiation potential (58, 59), implicating MyoD in both proliferation and differentiation phase of satellite cells. Moreover, the Myf5-null myoblast, has a precocious differentiation and expression of myogenin and troponin T (39). Although mice lacking MyoD or Myf5 have apparent normal skeletal muscle, mice lacking both genes have complete absence of skeletal muscle (60).

Skeletal muscle of mice with a mutation of the TRα gene, associated with RTH, showed a significantly smaller myofiber cross-sectional area, compared with the WT animal, and impairment in skeletal muscle regeneration after injury. The TRα1PV muscle phenotype at baseline is partially explained by the fact that TRα1PV mice are smaller than WT mice (33). In addition, TRα may be involved in prenatal muscle development and/or postnatal muscle growth. In support of this, it has been reported that TR-null mice (TRα−/−β−/−) have smaller muscle fiber size and less type 2 fast fiber compared with WT, and the TRα-null mouse has a more pronounced phenotype than TRβ-null mice (18) (4). Here we showed a similar phenotype in TRα1PV mice, where we observed a dramatic reduction of muscle fiber size and type 2 fiber number compared with TRβPV and WT mice (18) (4). Moreover, we showed that skeletal muscle myoblasts from TRα1PV mice exhibit impaired proliferation and significantly lower expression of important myogenic transcription factors, such as MyoD and Myf5. Considering that progenitor myoblasts are involved in skeletal muscle maintenance during adult life, we may conclude that the smaller muscle fibers in the TRαPV mice are likely secondary to the “underperforming” myoblasts. In addition, we show an impairment in skeletal muscle regeneration in TRα1PV mice, compared with WT mice, with a less efficient recovery after injury. This is likely the result of reduced proliferation, fusion, and differentiation capability of the skeletal muscle myoblasts. Moreover, the reduced expression of PAX7 in the skeletal muscle of the TRα1PV mice suggests a loss in quiescent satellite cells, which represents a crucial stem cell reserve for postnatal skeletal muscle regeneration and maintenance (61).

In conclusion, our data demonstrate that TRα plays an essential role in proliferation and differentiation of skeletal muscle myoblasts and in in vivo skeletal muscle regeneration after injury. These findings should lead to a better understanding of disorders associated with impaired adult myogenesis and identify novel therapeutic targets for these conditions.

Acknowledgments

The GC1 was generously provided by Dr Thomas Scanlan, Oregon Health & Science University.

This work was supported by National Institutes of Health (NIH) Grant No. 1K08DK097295 (to A.M.), and NIH RO1DK98576 (to G.A.B.). The research described was supported by NIH/National Center for Advancing Translational Science University of California, Los Angeles, Clinical and Translational Science Institute Grant No. UL1TR000124.

Disclosure Summary: The authors have nothing to disclose.

Footnotes

Abbreviations:
BrdU
bromodeoxyuridine
CSA
cross-sectional area
CTX
cardiotoxin
D2
5′-deiodinase 2
DAPI
4′,6-diamidino-2-phenylindole
MyF5
myogenic factor 5
MyoD
myoblast determination protein
Pax7
paired box transcription factor 7
RTH
resistance to thyroid hormone
shTRα
small-hairpin RNA for TRα
TAM
tibialis anterior muscle
TH
thyroid hormone
TR
thyroid hormone receptor
WT
wild type.

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