The exercise-inducible bile acid receptor Tgr5 improves skeletal muscle function in mice

Abstract

TGR5 (also known as G protein–coupled bile acid receptor 1, GPBAR1) is a G protein–coupled bile acid receptor that is expressed in many diverse tissues. TGR5 is involved in various metabolic processes, including glucose metabolism and energy expenditure; however, TGR5's function in skeletal muscle is not fully understood. Using both gain- and loss-of-function mouse models, we demonstrate here that Tgr5 activation promotes muscle cell differentiation and muscle hypertrophy. Both young and old transgenic mice with muscle-specific Tgr5 expression exhibited increased muscle strength. Moreover, we found that Tgr5 expression is increased by the unfolded protein response (UPR), which is an adaptive response required for maintenance of endoplasmic reticulum (ER) homeostasis. Both ER stress response element (ERSE)- and unfolded protein response element (UPRE)-like sites are present in the 5′ upstream region of the Tgr5 gene promoter and are essential for Tgr5 expression by Atf6α (activating transcription factor 6α), a well known UPR-activated transcriptional regulator. We observed that in the skeletal muscle of mice, exercise-induced UPR increases Tgr5 expression, an effect that was abrogated in Atf6α KO mice, indicating that Atf6α is essential for this response. These findings indicate that the bile acid receptor Tgr5 contributes to improved muscle function and provide an additional explanation for the beneficial effects of exercise on skeletal muscle activity.

Introduction

Bile acids are essential for solubilizing lipids and fat-soluble vitamins to promote their absorption in the small intestine. In addition to these classical functions, bile acids are known to function as metabolic regulators (1). In the postprandial state, bile acids are reabsorbed in the ileum, and the bile acid concentration temporarily reaches a high level in blood; then bile acids are transported back to the liver and recycled. This cycle, called enterohepatic circulation, is achieved by bile acid transporters, which are exclusively expressed in the ileum and liver. TGR5 (also known as GPBAR1) is the only G protein–coupled receptor that has bile acids as its ligand. TGR5 is expressed in multiple tissues, such as brown adipose tissue (BAT),² small intestine and skeletal muscles (2, 3). Ligand-bound TGR5 induces intracellular cAMP production via the Gα_s subunit, resulting in the activation of the protein kinase A (PKA)–cAMP response element-binding protein (CREB) pathway (2–4), which regulates numerous biological processes (5). Previous reports have shown that TGR5 activation in enteroendocrine L cells and colon stimulates glucagon-like peptide-1 release and improves glucose homeostasis (6–9). Furthermore, TGR5 activation triggers energy expenditure via type 2 deiodinase expression in human and mouse BAT, as well as in primary myotubes of human skeletal muscle. However, this has not been observed in mouse skeletal muscle, because of extremely low expression of type 2 deiodinase (4, 10). Recent studies indicated that bile acid synthesis is increased by cold exposure in mice, and elevated bile acid circulating causes adaptive thermogenesis, possibly by BAT activation and beiging of subcutaneous white adipose tissue (WAT) (11, 12).Therefore, TGR5 activation shows promise for the treatment of diabetes, obesity, and associated metabolic disorders; however, the functions of TGR5 in skeletal muscle are not well understood.

The endoplasmic reticulum (ER) is an evolutionarily conserved subcellular organelle that regulates protein synthesis, folding, and assembly. ER stress, which is defined as the cell state in which ER accumulates unfolded or misfolded proteins, occurs when ER function is disrupted and causes metabolic disorders such as obesity and type 2 diabetes (13, 14). Cells that sense ER stress activate the unfolded protein response (UPR), which involves three major ER stress sensors: protein kinase R-like endoplasmic reticulum kinase, inositol-requiring enzyme 1, and ATF6α (activating transcription factor 6α) (15). To increase ER protein folding capacity and attenuate ER stress, activated UPR sensors produce a variety of transcription factors, such as ATF4, the spliced form of XBP-1 (XBP-1s), and the N terminus of cleaved ATF6α (ATF6α(N)). Previous studies have shown that exercise stimulates the UPR in human and mice skeletal muscles and, importantly, that UPR activation helps skeletal muscles adapted to exercise (16, 17). Furthermore, another recent report demonstrated that inhibition of ER stress leads to muscle wasting in Lewis lung carcinoma and Apc^Min/+ models of cancer cachexia, as well as naïve mice, indicating that ER stress and UPR pathways contribute to maintain skeletal muscle mass and strength (18). Taken together, ER stress is not always harmful but rather appears to assist in multiple physiological processes in skeletal muscle.

In the present study, we investigate the role of TGR5 in the control of skeletal muscle function. Interestingly, Tgr5 KO mice exhibited lower muscle mass and strength and decreased the expression of several muscle hypertrophy and differentiation-related genes in skeletal muscle. As expected from these findings, TGR5-expressing C2C12 myoblasts and muscle-specific TGR5 transgenic (Tg) mice clearly show that TGR5 activation enhances muscle cell differentiation and muscle hypertrophy. Furthermore, we identify Tgr5 as a novel UPR target gene and reveal that exercise induces muscle Tgr5 expression in an Atf6α-dependent manner. These findings show that TGR5 plays an important role in regulating muscle function and provide a new explanation for the beneficial effects of exercise.

Results

Tgr5 KO mice exhibit lower muscle mass and strength

Because TGR5 is known to activate CREB via cAMP production (2, 3), we measured the mRNA expression of CREB target genes in the skeletal muscle of Tgr5 KO mice to evaluate the function of muscle Tgr5 in vivo (5, 19) (Fig. 1A). As expected, gene expressions of several CREB targets were decreased in the soleus of Tgr5 KO mice compared with WT littermates (Fig. 1A). Decreased genes included Sik1 (salt-inducible kinase 1), which plays an important role in muscle cell survival and differentiation (20, 21), and the hypertrophy-related genes Nr4a1 and Pgc-1α4 (22, 23). We also observed increased mRNA expression of the muscle atrophy-related E3-ubiquitin ligases Musa1 and Smart in Tgr5 KO mice (24, 25) (Fig. 1A). These findings indicate that TGR5 works as a muscle mass regulator. There were no differences in body weights between WT and Tgr5 KO mice under the standard chow diet, which is compatible with a previous report (26) (Fig. 1B). However, Tgr5 KO mice exhibited lower soleus and quadriceps mass compared with WT littermates (Fig. 1C). The same results applied to the ratio of muscle weights to body weight (Fig. 1D). Consistent with these findings, Tgr5 KO mice showed significantly lower grip strength than WT littermates (Fig. 1E). These data demonstrate that TGR5 is involved in regulating muscle mass and strength in vivo. Next, we measured food intake, energy expenditure, the respiratory exchange ratio (RER), and locomotor activity in Tgr5 KO and WT mice. Energy expenditure and RER were calculated from O₂ consumption (VO₂) and CO₂ production (VCO₂) by using a mass spectrometer. We found no differences in these measurements, showing that muscular atrophy observed in Tgr5 KO mice was not caused by changes in energy metabolism or physical activity (Fig. 1, F–I).

Figure 1.

The effects of Tgr5 knockout on skeletal muscle. A, the soleus was isolated from 8-week-old male Tgr5 KO mice and WT littermates, and the mRNA levels of Tgr5 and CREB target genes were measured (n = 9). B, body weights of 8-week-old male Tgr5 KO mice and WT littermates (n = 10–12). C and D, muscle weight (C) and percentage of body weight (D) (n = 10–12). E, grip strength was measured at 8 weeks of age (n = 30–37). F–I, food intake (F), energy expenditure (G), RER (H), and act count (I) of 8-week-old male Tgr5 KO mice and WT littermates were monitored over 96 h (n = 6–8). The data are means ± S.E. Statistical analyses were done using two-tailed unpaired Student's t test. *, p < 0.05; **, p < 0.01.

TGR5 enhances muscle cell differentiation in C2C12 myoblasts

SIK1 phosphorylates class II histone deacetylases, resulting in the activation of MEF2 (myocyte enhancer factor 2), which is considered to act as a linchpin for muscle development and differentiation (20, 27). Therefore, based on our current findings that Tgr5 KO mice had decreased Sik1 expression and muscle mass, we aimed to determine the effect of TGR5 activation on muscle differentiation. Because of low Tgr5 expression in C2C12 myoblasts, we use hTGR5 overexpressing C2C12 myoblasts and confirmed that TGR5 activation with taurolithocholic acid (TLCA), one of the most potent endogenous ligands for TGR5, induces CRE reporter activation (Fig. S1A and Fig. 2A). In addition, TLCA treatment in C2C12 myoblasts overexpressing TGR5 with adenovirus increased the mRNA levels of Nr4a2, a known CREB-responsive gene, and Sik1 (2.7- and 3.5-fold, respectively), whereas TLCA treatment in LacZ-expressing C2C12 myoblasts only slightly increased these mRNA levels (1.2-fold each) (Fig. 2B). Similar results were also observed when using cholic acid or deoxycholic acid instead of TLCA (Fig. S1, B and C). To evaluate the effect of TGR5 on muscle differentiation, LacZ or TGR5 overexpressing C2C12 myoblasts were differentiated with or without TLCA, and the mRNA levels of Sik1 and several differentiation marker genes, including MyoD, Myogenin, Muscle creatine kinase (MCK), Mef2a, and MyHC, were measured. TGR5/TLCA C2C12 myoblasts showed the highest mRNA expression levels of Sik1 and differentiation marker genes, particularly on day 2 (48 h after TLCA treatment) (Fig. 2C). The measurement of the differentiation index (i.e. the percentage of nuclei in MyHC-positive cells above total nuclei) also revealed that TLCA significantly promoted the differentiation of TGR5-expressing C2C12 myoblasts (1.5-fold). LacZ-expressing C2C12 myoblasts showed a similar tendency (1.3-fold), although it was not statistically significant (Fig. 2D). Endogenous Sik1 silencing using two types of siRNA attenuated the mRNA expression of several muscle differentiation marker genes in TGR5/TLCA C2C12 myotubes, indicating the importance of Sik1 for TGR5-mediated muscle differentiation (Fig. 2E, the details of statistical analysis were shown in Table S1). These results demonstrated that TGR5 activation leads to muscle cell differentiation caused in part b the up-regulation of Sik1 expression.

Figure 2.

TGR5 enhances muscle cell differentiation in C2C12 myoblasts. A, CRE reporter assay in empty vector- or TGR5 expression vector-transfected C2C12 myoblasts. TLCA (50 μm) was treated for 5 h (n = 3). B, C2C12 myoblasts infected with adenovirus-expressing TGR5 or LacZ were treated with TLCA (50 μm) for 3 h. Nr4a2 and Sik1 mRNA levels were determined by RT–PCR (n = 3). C and D, C2C12 myoblasts were infected with adenovirus containing LacZ or TGR5 before 36 h of differentiation. These cells were differentiated with or without TLCA (20 μm). The media were refreshed every 24 h. C, samples were collected at the indicated time points, and mRNA expressions were determined by RT–PCR (n = 3). D, cells were immunostained for MyHC after 2 days of differentiation (left panels) and then quantified for the differentiation index as a percentage of nuclei in myosin heavy chain-positive cells (right panel). E, C2C12 myoblasts were transfected with Sik1 siRNA or control (ctrl) siRNA. After 12 h, these cells were infected with adenovirus-expressing TGR5 or LacZ. 36 h later, the cells were differentiated with or without TLCA (20 μm) for 48 h. mRNA levels were determined by RT–PCR (n = 3). The data are means ± S.E. Statistical analyses were done using two-tailed unpaired Student's t test or one-way ANOVA (Tukey's post hoc test). *, p < 0.05; **, p < 0.01; n.s., not significant.

TGR5 induces muscle hypertrophy in mice

To evaluate the function of TGR5 in differentiated mouse skeletal muscle, we generated muscle-specific FLAG-hTGR5 Tg mice under the control of the MCK promoter and enhancer. Among four transgenic lines obtained (lines A–D), line A and B showed higher FLAG-hTGR5 expression levels than lines C and D (Fig. S2A). Thus, unless otherwise noted, line A was used in the following experiments. FLAG-hTGR5 expression was observed in skeletal muscles but not in the heart, liver, kidney, epididymal WAT, or BAT (Fig. S2B). FLAG-hTGR5 was strongly expressed in white muscle tissue such as that of the gastrocnemius and quadriceps, whereas it was weakly expressed, as expected, in the soleus, a typical red muscle, using the MCK promoter (Fig. S2C).

We observed no differences in the daily food intake between Tg mice and WT littermates fed a normal diet at 8 weeks old (Fig. 3A). However, Tg mice were slightly but significantly heavier (Fig. 3B). Consistent with our finding that Tgr5 KO mice have muscle atrophy, Tg mice exhibited 9.5 and 14.6% increases in gastrocnemius and quadriceps mass, respectively, where sufficient levels of FLAG-hTGR5 protein were found (Fig. 3, C and D, and Fig. S2C). The same results applied to the ratio of gastrocnemius and quadriceps weights to body weight (7.5 and 12.5%, respectively) (Fig. S2D). In contrast, tissue weights did not increase in the soleus, liver, or WAT (Fig. 3C). Similar changes in body and tissue weights were also seen in female Tg mice (Fig. S2, E and F). In addition, the quantitative analysis of gastrocnemius fiber size revealed an increased average cross-sectional area in Tg mice (Fig. 3, E and F); particularly, TGR5 appeared to drive an increase in the frequency of very large fibers (>2400 μm²), which was uncommon in WT littermates. As expected, this muscle mass increment was accompanied by an increase in grip strength in Tg mice at 2, 6, 9, 12, 15, 18, and 21 months old (Fig. 3G). However, because there was no difference in the degree of muscle weakness caused by aging and muscle atrophy caused by denervation, TGR5 seems to increase muscle mass by inducing muscle hypertrophy rather than suppressing muscle atrophy (Fig. 3G and Fig. S2G). Intramuscular TLCA injection to the quadriceps of Tg mice immediately promoted the mRNA expression of Sik1 and the hypertrophy-related genes Nr4a1 and Pgc-1α4, which were decreased in the skeletal muscle of Tgr5 KO mice (Figs. 1A and 3H). These responses were not strong in WT littermates; however, we observed a mild increment of these genes 3 and 8 h after TLCA injection. Previous reports have shown that NR4A1- and PGC-1α4-induced muscle hypertrophy is mediated by the growth-promoting gene IGF1, and growth-limiting genes, such as Myostatin, Atrogin1, and MuRF1 (22, 23). Consistent with these reports, TLCA-injected Tg mice showed significantly higher Igf1 and lower Atrogin1 mRNA expressions compared with the vehicle group 8 h after injection, although Myostatin and MuRF-1 were not affected. Moreover, Musa1, which is required for protein breakdown in atrophying muscles, was negatively regulated by TGR5 activation in contrast to Tgr5 KO mice (24) (Figs. 1A and 3H). Phosphorylation of Akt was also accelerated as Igf1 expression increased (Fig. 3I). These results suggest molecular evidence that activation of TGR5 in matured skeletal muscle induces muscle hypertrophy and increases muscle strength.

Figure 3.

Skeletal muscle-specific TGR5 Tg mice have muscle hypertrophy. A–C, food intake (A), body weight (B), and tissue weight (mg) (C) of 8-week-old male littermates of the indicated genotype (n = 8–10). D, dorsal view of WT and Tg mice. E, hematoxylin and eosin staining of the gastrocnemius (Gastro) muscle. F, cross-sectional area frequency distribution of gastrocnemius and average (n = 4). G, grip strength was measured at 2, 6, 9, 12, 15, 18, and 21 months of age (n = 21–26). H, saline (10% BSA) or TLCA (0.4 mg/100 μl, saline with 10% BSA) was administrated to the quadriceps (Quad) of 18 h fasted Tg and WT littermates by single intramuscular injections (100 μl/20 g body weight). After 3 or 8 h, quadriceps were isolated, and mRNA levels were measured (n = 6–8). I, Akt phospho- and total protein and FLAG protein were measured by Western blotting in WT and Tg quadriceps 8 h after TLCA intramuscular injection. The data are means ± S.E. Statistical analyses were done using two-tailed unpaired Student's t test. *, p < 0.05; **, p < 0.01.

To evaluate the possibility that TGR5 also promotes muscle hypertrophy in humans, TGR5 overexpressing human skeletal muscle myotubes were treated with TLCA, and mRNA expression levels were measured. TGR5 activation increased SIK1, NR4A1, NR4A3, PGC-1α4, and IGF1 mRNA expression, as well as these in TLCA-treated Tg mice, and decreased Myostatin, MuRF1, and MUSA1 mRNA expression significantly in human skeletal muscle myotubes (Fig. 4). We next performed an analysis of gene expression in human skeletal muscle using RNA-sequence data from the Genotype-Tissue Expression project (28). Interestingly, TGR5 mRNA expression showed moderate positive correlations with that of IGF1 (Pearson's r = 0.354), although very weak or no correlations were observed with that of muscle atrophy-related E3 ligases and Myostatin (|r| < 0.2) (Fig. S3). These data suggest that TGR5 has the potential to regulate muscle mass in humans similar to mice.

Figure 4.

The effect of TGR5 activation on HSMM gene expression. Human skeletal muscle myotubes infected with adenovirus-expressing TGR5 or LacZ were treated with TLCA (50 μm) for 3 or 8 h. mRNA levels were determined by RT–PCR (n = 3). The data are means ± S.E. Statistical analysis were done using one-way ANOVA (Tukey's post hoc test). *, p < 0.05; **, p < 0.01.

Tgr5 expression was up-regulated by the UPR in muscle cells

As observed above, alterations in TGR5 expression level had effects on muscle mass and muscle cell differentiation. However, little is known regarding the molecular mechanisms that regulate endogenous TGR5 expression until now. Interestingly, we found that thapsigargin (an ER-specific calcium ATPase inhibitor) and tunicamycin (an N-glycosylation inhibitor), both well known UPR inducers, increased Tgr5 mRNA levels in C2C12 myotubes (2.1- and 2.5-fold, respectively) similarly to several UPR marker genes, including Atf4, Atf6α, Bip, total Xbp-1 (Xbp-1t), and Xbp-1s (Fig. 5A). The increase in Tgr5 and Bip mRNA expression by UPR inducers was accompanied by their increased protein levels (Fig. 5B). A similar response was observed in mouse skeletal muscle when thapsigargin was intramuscularly injected (1.6-fold for Tgr5) (Fig. 5C). Consistent with these results, a reporter gene assay using the mouse Tgr5 (mTgr5) promoter region also showed that the promoter activity was increased by tunicamycin treatment (2.5-fold) (Fig. 5D). These results strongly suggest that Tgr5 is a novel UPR target gene.

Figure 5.

The UPR increases Tgr5 expression in skeletal muscles. A and B, C2C12 myotubes were treated with thapsigargin (250 nm) or tunicamycin (2.5 μg/ml) for 9 h. A, mRNA levels were determined by RT–PCR (n = 3). B, immunoblot analysis of protein lysates using the indicated antibodies (n = 3, mixture) and the Tgr5 protein level normalized to α-tubulin (n = 3). C, saline (0.2% DMSO, 150 μl) or thapsigargin (20 μm, saline with 0.2% DMSO, 150 μl) was administrated to the quadriceps by single intramuscular injection. After 12 h, the quadriceps were isolated, and mRNA levels were measured (n = 5–6). D, C2C12 myoblasts were transfected with mouse the Tgr5 promoter-luciferase construct (1600 bp). 24 h later, the cells were treated with tunicamycin (2.5 μg/ml) for 12 h, and luciferase activity was measured and normalized against β-gal activity (n = 3). The data are means ± S.E. Statistical analyses were done using two-tailed unpaired Student's t test or one-way ANOVA (Tukey's post hoc test). *, p < 0.05; **, p < 0.01.

Atf6α activates Tgr5 transcription and is necessary for exercise-induced Tgr5 expression

UPR, which involves the ATF6α, inositol-requiring enzyme 1, and protein kinase R-like endoplasmic reticulum kinase pathways, is known to activate various transcription factors such as ATF2, ATF3, ATF4, ATF6α, CHOP, and XBP-1. To identify the direct factors that regulate TGR5 transcription, we performed reporter assays on the mTgr5 promoter and found that ATF6α(N) and XBP-1s were potent factors that promote mTgr5 promoter activity (Fig. 6A). A previously published report showed that both ATF6α and XBP-1 bind to the ER stress-response element (ERSE) in an NF-Y–dependent manner, whereas XBP-1 also binds to an unfolded protein response element (UPRE) without NF-Y (29). Consistent with this report, there is a combined ERSE-like and UPRE-like element in the 5′ upstream region of the mTgr5 promoter (Fig. S4). Interestingly, the assay on the mTgr5 promoter with mutations in these elements indicated that the 5′-GCAGT-3′ sequence, a potential NF-Y–binding site, and the 5′-CCACG-3′ sequence, a potential ATF-6α- and XBP-1–binding site, were necessary for ATF6α(N)-induced mTgr5 promoter activation (Fig. S4 and Fig. 6B). In contrast, XBP-1s–responsive promoter activation was strongly but not completely eliminated by the mutation in the 5′-GCAGT-3′ sequence, whereas the unaltered 5′-CCACG-3′ sequence was essential for this response (Fig. S5). These findings suggest that ATF6α(N) binds to the ERSE-like element, whereas XBP-1s recognizes both the ERSE-like and UPRE-like elements in the mTgr5 5′ upstream region. To test whether ATF6α(N) increases endogenous Tgr5 expression, C2C12 myotubes were transduced with adenovirus-expressing ATF6α(N). We observed a significant increase in Tgr5 mRNA (16.3-fold) in ATF6α(N)-expressing C2C12 myotubes, which was higher than that of Bip (6.9-fold), a major ATF6α target gene (Fig. 6C). Similarly, these protein levels were also increased by ATF6α(N) overexpression (Fig. 6D). A CRE-Luc assay revealed that ATF6α(N)-expressing C2C12 myoblasts reacted to TLCA more strongly than LacZ-expressing C2C12 myoblasts (Fig. 6E).

Figure 6.

Exercise increases Tgr5 mRNA expression in an Atf6α-dependent manner. A, C2C12 myoblasts were transfected with the described expression plasmids and the mouse Tgr5 promoter-luciferase construct. 24 h later, the cells were harvested, and luciferase activity was measured and normalized against β-gal activity (n = 3). B, C2C12 myoblasts were transfected with the indicated reporter constructs in the presence or absence of the ATF6α(N) expression plasmid. 24 h after transfection, luciferase assays were performed and normalized against β-gal. Promoter activities in the absence of ATF6α(N) were set at 1 (n = 3). C and D, C2C12 myotubes were infected with adenovirus-expressing ATF6α(N) or LacZ as a control. C, mRNA levels were determined by RT–PCR (n = 3). D, immunoblot analysis of protein lysates using the indicated antibodies (n = 3, mixture) and the Tgr5 protein level normalized to α-tubulin (n = 3). E, CRE reporter assay in LacZ or ATF6α(N) adenovirus-infected C2C12 myoblasts. TLCA (100 μm) was treated for 5 h. Luciferase activities in DMSO were set at 1 (n = 3). F, C2C12 myoblasts were transfected with expression plasmids for ATF6α(N) and/or PGC-1α, together with the mouse Tgr5 promoter-luciferase construct (2000 bp). The cells were differentiated for 2 days and harvested, and luciferase activity was measured and normalized against β-gal (n = 3). G, total RNA was isolated from the quadriceps of 8–10-week-old male Atf6α KO mice and WT littermates 5 h after treadmill running or sedentary control. mRNA levels were measured by RT–PCR (n = 6 per group). The data are means ± S.E. Statistical analyses were done using two-tailed unpaired Student's t test or one-way ANOVA (Tukey's post hoc test). *, p < 0.05; **, p < 0.01.

Exercise has been reported to activate the UPR in mouse and human skeletal muscles (16, 17). Particularly, exercise-induced PGC-1α promotes UPR gene expression, in part by coactivating ATF6α (17). Indeed, we found that ATF6α and PGC-1α coordinately activated the mTgr5 promoter (Fig. 6F). Therefore, we subjected WT and Atf6α KO mice to treadmill running and analyzed gene expression in their quadriceps. The mRNA levels of Tgr5 and UPR marker genes increased significantly in WT mice after exercise (1.4-fold for Tgr5) (Fig. 6G). In contrast, the induction of Tgr5 and some UPR marker genes, such as Bip and Chop were completely repressed in Atf6α KO mice. These observations indicate that Tgr5 is a novel UPR target gene, and Atf6α is necessary for exercise-induced increases in muscle Tgr5 expression.

Discussion

Ligand-bound GPCRs activate specific signaling pathways depending on the type of coupled G-protein α-subunit. Because TGR5 is coupled with Gα_s, its activation enhances the cAMP–PKA–CREB pathway via adenylyl cyclase in various types of cells (2, 4, 30). Previously, several Gα_s-coupled GPCRs, such as β-adrenergic receptor (β-AR), Frizzled7, and CRFR2, were shown to induce muscle hypertrophy and attenuate muscle atrophy (31). For example, the prolonged administration of β-AR agonists increases muscle mass and attenuates sarcopenia (32, 33). β-AR–mediated muscle hypertrophy requires CREB and its coactivators, which regulate muscle mass regulatory genes, such as PGC-1α4, SIK1, and NR4A1 (34). Indeed, silencing Pgc-1α4 has been shown to blunt β-AR agonist-induced muscle hypertrophy in mouse primary myotubes (23). Consistent with these findings, the genetic deletion of Gα_s in skeletal muscle decreases total Pgc-1α expression and induces muscle atrophy (35). Another study showed that muscle-specific expression of dominant-negative CREB causes a severe dystrophic phenotype with decreased expression of Sik1 that can be rescued by exogenous Sik1 overexpression (20). In addition, a recent study has demonstrated the importance of Nr4a1 for muscle mass regulation by using both gain- and loss-of-function mouse models (22). These reports suggest that PGC-1α4, SIk1, and NR4A1 contribute to muscle hypertrophy induced by Gα_s-coupled GPCRs. As expected from these studies, Tgr5 KO mice exhibited lower muscle mass with decreased CREB target genes, including Sik1, Nr4a1, and Pgc-1α4 (Fig. 1, A, C, and D). Because the skeletal muscle reduction rate in Gα_s KO mice that loses reactivity to all Gα_s-coupled GPCRs is 20–40% (35), it is a surprising result that Tgr5 KO mice showed ∼10% reduction in skeletal muscle mass, suggesting that TGR5 plays an important role as a muscle mass regulatory GPCR (Fig. 1C). Conversely, muscle-specific TGR5 overexpression induces muscle hypertrophy and increases grip strength in young and old mice (Fig. 3, C–G). TLCA-induced TGR5 activation in skeletal muscle promptly raised Sik1, Nr4a1, and Pgc-1α4 expression, leading to the up-regulation of Igf1 expression and the down-regulation of Atrogin1 and Musa1 expression (Fig. 3H). Contrary to previous reports (20, 21), si Sik1#1 and #2 did not inhibit muscle cell differentiation in C2C12 myoblasts but rather increase differentiation marker genes in si Sik1#2 (Fig. 2E). It may have been caused by low knockdown efficiency and/or compensatory mechanisms in the muscle cell to maintain its function. Although our present study could not provide clear answers to this issue, we have confirmed that TGR5 activation promoted Sik1 expression and that TGR5-induced muscle cell differentiation is canceled by Sik1 knockdown (Fig. 2, C–E). These data indicate that Sik1 plays an important role in muscle differentiation by activation of TGR5. Therefore, the decrease in muscle mass observed in Tgr5 KO mice is considered to be the result of muscle cell development and growth disorder. Unlike other GPCRs, TGR5 does not interact with β-arrestin; thus, it does not desensitize, resulting in a long-lasting activation of cAMP signaling (36). This fact may explain the reason why Tgr5 KO mice exhibited significantly lower muscle mass and strength compared with WT mice in this study, despite endogenous Tgr5 showing just a weak immediate response to single TLCA injection in mouse skeletal muscle (Figs. 1, C and D, and 3H). Previously, we and others have identified several compounds as TGR5 ligands and confirmed that these compounds contribute to metabolic improvement (6, 37–41). Additionally, obacunone and ursolic acid, both of which are known to activate TGR5, have been shown to increase muscle mass in mice (42, 43). Although whether TGR5 is needed for this response has yet to be definitively determined, these results support our notion that TGR5 activation leads to muscle hypertrophy. By using RNA-Seq data, we found a positive correlation between TGR5 and IGF1 expression levels in human skeletal muscle, which is in agreement with the fact that IGF1 mRNA expression is increased by TGR5 activation in human skeletal muscle myotubes (Fig. 4 and Fig. S3). These results suggest that TGR5 has the potential to increase muscle mass in humans as well. Additional studies are needed to confirm whether TGR5 regulates human skeletal muscle functions.

Regarding TGR5 activity, the TGR5 expression level is an important factor for its downstream activation. For example, Tgr5 mRNA expression is strongly correlated with that of several estimated Tgr5 target genes, such as CoxVIa1 and ATP synthase subunits, in mouse liver (6). This is supported by our finding that TGR5 mRNA expression is positively correlated with that of IGF1 in human skeletal muscle (Fig. S3). However, the expression mechanism is not well understood. Notably, our study clearly showed that Tgr5 expression is regulated by the UPR in skeletal muscle cells (Fig. 5, A–D). mTgr5 promoter activity was up-regulated by both XBP-1 and ATF6α, which recognize combined ERSE-like and UPRE-like elements in the 5′ upstream region of the mTgr5 promoter (Fig. 6, A and B, and Fig. S5). Endogenous Tgr5 up-regulation by ATF6α(N) overexpression resulted in a strong response to TLCA in C2C12 cells (Fig. 6E). In accordance with previous reports (16, 17), treadmill exercise activated the UPR in mouse skeletal muscle, and we found that exercise increased Tgr5 expression in an Atf6α-dependent manner (Fig. 6G). These facts suggest that Atf6α-dependent Tgr5 up-regulation in skeletal muscle contributes to exercise-induced muscle hypertrophy. However, contrary to our expectation, basal Tgr5 expression level was increased in Atf6α KO mice. These data indicate that the contribution of Atf6α to resting muscle Tgr5 expression is small and suggest the presence of unknown regulation mechanism of Tgr5 expression. Because TGR5 is known to exert several adverse effects, such as gallstone formation and bile acid-induced itch and analgesia (19, 44), other than those beneficial on systemic metabolism and muscle function, tissue-specific regulation of TGR5 expression may be a valid approach to obtain benefit from TGR5 with fewer side effects.

In the postprandial state, farnesoid X receptor is activated by bile acids in ileal enterocytes to promote fibroblast growth factor 15/19 secretion, which regulates bile acid metabolism and metabolic homeostasis (45). Interestingly, a recent study shows that fibroblast growth factor 19 treatment induces skeletal muscle hypertrophy and ameliorates glucocorticoid, obesity, and aging-induced skeletal muscle atrophy in mice (46). Bile acids may be more closely related to skeletal muscle function than we expected.

In summary, we showed that TGR5 activation in skeletal muscles promotes muscle hypertrophy and differentiation. Moreover, we found that Tgr5 expression was regulated by the UPR in muscle cells, and exercise increased Tgr5 mRNA expression in an Atf6α-dependent manner in mouse skeletal muscle, suggesting a synergistic effect between feeding and exercise on Tgr5 activity by up-regulating both blood bile acid concentration and Tgr5 expression (Fig. 7). These results establish a new linkage between bile acid function and skeletal muscle metabolic adjustment and indicate that muscle TGR5 may be a feasible target for maintaining muscle function in the elderly.

Figure 7.

A proposed model for roles of the Tgr5 in skeletal muscle. Tgr5 expression is increased by Atf6α in muscle cells, and it is mimicked by exercise in vivo. Tgr5 activation promotes muscle hypertrophy and muscle cell differentiation.

Experimental procedures

Animals and diets

For a generation of skeletal muscle-specific TGR5 transgenic mice, 3× FLAG hTGR5 was cloned into MCK promoter plasmids, gifted from Ronald Kahn (Addgene plasmid no. 12528) (47); purified transgene was then injected into C57BL/6 oocytes. Atf6α KO mice were generated as described previously (48). Mice were housed with a 12:12-h light-dark cycle and given free access to water and food. All animal experiments were performed according to the Guideline for the Care and Use of Laboratory Animals of the University of Tokyo, under the approval of the Animal Usage Committee of University of Tokyo (approval numbers P13-812, P15-079, and P17-120).

Treadmill exercise

8–10-week-old male Atf6α KO mice and WT littermates were acclimated to running on a treadmill (Muromachi Kikai) at a 10% incline for 60 min. Running speed was set at 5 m/min for the first 5 min and increased to 10 m/min for the next 5 min. Subsequently, speed was increased to 15 m/min for 20 min and finally 30 min was set at 18 m/min. Treadmill running was continued until the mice were exhausted (they remained on an electric stimulus grid for 6 s) or completed a 60-min running program.

Cell culture

C2C12 myoblasts were cultured in growth medium (DMEM supplemented with 10% fetal bovine serum). For differentiation to myotubes, the medium was change into differentiation medium (DMEM supplemented with 2% horse serum) as previously described (49). Human skeletal muscle myoblasts (HSMMs) were obtained from Lonza and cultured in accordance with the supplier's instructions. For differentiation to myotubes, the medium was changed into differentiation medium (DMEM/Ham's F-12 supplemented with 2% horse serum). C2C12 myoblasts and HSMM were infected with 2.5 × 10⁶ plaque-forming units/ml adenovirus culture overnight, and the medium was refreshed the next morning.

Immunostaining

C2C12 cells were washed by PBS and fixed with 4% paraformaldehyde in PBS for 15 min at room temperature. The cells were then permeabilized with 0.5% Triton X-100 in PBS for 5 min and blocked with 3% BSA in PBS. Subsequently, the cells were incubated with anti-MyHC antibody (MF20; R&D Systems) for 60 min and then incubated with secondary antibody (fluorescein (FITC)-AffiniPure donkey anti-mouse IgG; Jackson ImmunoResearch) for 30 min.

Immunoblotting

C2C12 cells and mouse tissues were lysed with radioimmune precipitation assay buffer (50 mm Tris·HCl, pH 7.4, 1 mm EDTA, 150 mm NaCl, 1% Nonidet P-40, and 0.25% sodium deoxycholate) supplemented with a protease inhibitor mixture (Nacalai Tesque) and a phosphatase inhibitor mixture (Sigma–Aldrich). The proteins were subjected to SDS–PAGE and analyzed by immunoblotting. Anti-α-tubulin (H-300) and anti-TGR5 (H-90) antibodies were obtained from Santa Cruz Biotechnology. Anti-BiP, anti-AKT, and anti-phospho-AKT antibodies were purchased from Cell Signaling Technology. Anti-FLAG (M2), and anti-β-actin (AC-15) antibodies were acquired from Sigma.

Quantitative RT–PCR

Total RNA was extracted using ISOGEN (Nippon Gene) according to the manufacturer's instructions. Total RNA was reverse-transcribed using a high capacity cDNA reverse transcription kit (Applied Biosystems) and subjected to quantitative RT–PCR analysis (TaqMan probe and SYBR Green) by using a StepOnePlus real time PCR system. The values were normalized to 18S. The primers used for the PCR analysis (SYBR green) are described in Table S2.

siRNA experiments

siRNAs for mouse Sik1 (AM16708-156369 and AM16708-156371) and control (AM4611) (ThermoFisher Scientific) were transfected using Lipofectamine RNAiMAX (Invitrogen) into C2C12 myoblasts (25 nm each) according to the manufacturer's instructions.

Physiological measurements

O₂ consumption and CO₂ production in 8-week-old male Tgr5 KO mice and WT littermates were measured using a ARCO-2000 mass spectrometer (ARCO System) with one mouse per chamber. The environment was maintained at 21 ± 3 °C with 50 ± 10% relative humidity. The grip strength test was performed using a MK-380M grip strength meter (Muromachi Kikai). The grip strength was measured seven times for each mouse. The same measurements were repeated 4 days later, and the average of the highest values was used.

Luciferase assay

C2C12 myoblasts were plated on 12-well plates at a density of 1 × 10⁵ cells/well. the cells were cultured with growth medium overnight and transfected with plasmids by a calcium phosphate method. After 24 h of incubation with growth medium or an additional 2 days of differentiation, the luciferase and β-gal activities were determined.

Statistical analysis

All results are expressed as means ± S.E. of at least three independent biological replicates. Two-tailed unpaired Student's t tests or one-way ANOVAs (Tukey's post hoc test) were used to determine p values. Statistical significance was defined as p < 0.05.

Author contributions

T. S., K. M., and R. S. conceptualization; T. S., M. S., and J. I. formal analysis; T. S., M. S., and R. S. funding acquisition; T. S., A. K., M. M., and S. M. investigation; T. S. writing-original draft; T. S. and R. S. project administration; K. M. resources; R. S. supervision; R. S. writing-review and editing.