Skeletal muscle is important not only for locomotion but also for regulating metabolic function. Lahiri et al. studied the interactions between the gut microbiota and skeletal muscle in mice. They identified genes and signaling pathways involved in the regulation of skeletal muscle mass and function that responded to cues from the gut microbiota. Additional biochemical and functional analysis also revealed the influence of the gut microbiota on the function of neuromuscular junctions. These findings open the door to a better understanding of the role of the gut microbiota in the mechanisms underlying loss of muscle mass.

The functional interactions between the gut microbiota and the host are important for host physiology, homeostasis, and sustained health. We compared the skeletal muscle of germ-free mice that lacked a gut microbiota to the skeletal muscle of pathogen-free mice that had a gut microbiota. Compared to pathogen-free mouse skeletal muscle, germ-free mouse skeletal muscle showed atrophy, decreased expression of insulin-like growth factor 1, and reduced transcription of genes associated with skeletal muscle growth and mitochondrial function. Nuclear magnetic resonance spectrometry analysis of skeletal muscle, liver, and serum from germ-free mice revealed multiple changes in the amounts of amino acids, including glycine and alanine, compared to pathogen-free mice. Germ-free mice also showed reduced serum choline, the precursor of acetylcholine, the key neurotransmitter that signals between muscle and nerve at neuromuscular junctions. Reduced expression of genes encoding Rapsyn and Lrp4, two proteins important for neuromuscular junction assembly and function, was also observed in skeletal muscle from germ-free mice compared to pathogen-free mice. Transplanting the gut microbiota from pathogen-free mice into germ-free mice resulted in an increase in skeletal muscle mass, a reduction in muscle atrophy markers, improved oxidative metabolic capacity of the muscle, and elevated expression of the neuromuscular junction assembly genes Rapsyn and Lrp4. Treating germ-free mice with short-chain fatty acids (microbial metabolites) partly reversed skeletal muscle impairments. Our results suggest a role for the gut microbiota in regulating skeletal muscle mass and function in mice.

It has been established that the gut microbiota influences host health in part due to its coevolvement with the host to meet mutually beneficial biochemical and biological needs ( 4 ). There are many studies investigating how the gut microbiota influences the liver and intestinal metabolism, immunity, and behavior ( 5 – 8 ). However, few studies have reported how the gut microbiota regulates skeletal muscle, one of the dominant metabolic organs in the body. Here, we present evidence from germ-free (GF) and pathogen-free (PF) mice that the gut microbiota influences skeletal muscle mass and function.

Skeletal muscle function is regulated by the central nervous system through neurotransmission at neuromuscular junctions (NMJs). NMJs are highly specialized chemical synapses formed between motor neurons and skeletal muscle fibers ( 1 ). Skeletal muscle displays marked plasticity, being able to respond to a variety of environmental cues, such as exercise and nutrition. Skeletal muscle is also the major site of insulin-stimulated glucose uptake and fatty acid oxidation emphasizing its key role in metabolism. Reduced skeletal muscle mass and function are associated with metabolic disorders ( 2 ) and sarcopenia ( 3 ), underscoring the role of skeletal muscle in maintenance of health.

RESULTS

Altered skeletal muscle mass in GF mice GF mice lacking a gut microbiota displayed reduced skeletal muscle weight compared to PF mice that had a gut microbiota (P < 0.01; Fig. 1A). Transplanting GF mice with the gut microbiota of pathogen-free mice [henceforth referred to as conventionalized GF (C-GF) mice] restored muscle mass in the transplanted C-GF animals (P < 0.05; Fig. 1A). Histological examination of the tibialis anterior, a fast oxidative muscle, revealed a trend toward fewer but larger muscle fibers (>2000- to 3000-μm2 cross-sectional fiber area) in GF mice compared to PF mice (fig. S1, A and B). In addition, reduced expression of the myosin heavy chain genes MyHCIIa, MyHCIIb, and MyHCIIx was observed in GF mouse tibialis anterior muscle compared to PF mouse tibialis anterior muscle (P < 0.05; Fig. 1B). A similar trend was also observed in the soleus, a slow oxidative muscle, and the extensor digitorum longus, a fast glycolytic muscle, in GF mice compared to PF mice (P < 0.01; fig. S1, C and D). Furthermore, we observed increased expression of Atrogin-1 and Murf-1 encoding E3 ubiquitin ligases, which are known to be involved in muscle atrophy, in the tibialis anterior muscle of GF mice compared to PF mice (P < 0.01; Fig. 1, C, E, and F). Reduced expression of Murf-1 and Atrogin-1 was observed in the anterior tibialis muscle of the transplanted C-GF animals in comparison to GF mice (P < 0.01; Fig. 1C). The expression of Atrogin-1 and Murf-1 is known to be regulated by FoxO transcription factors (9). Whereas elevated expression of FoxO3 was observed in the tibialis anterior muscle of GF mice in comparison to PF mice (P < 0.05; Fig. 1C), FoxO3 expression was normalized in the transplanted C-GF mice. The energy sensor adenosine 5′-monophosphate–activated protein kinase (AMPK) controls muscle fiber size by activating the FoxO-mediated protein degradation pathway (10). Further analysis of the tibialis anterior muscle from GF mice revealed an increase in phosphorylation of the AMPK catalytic domain (on Thr172) compared to PF mouse muscle (Fig. 1, E and F). It has been reported that Atrogin-1 expression regulates MyoD expression in skeletal muscle under atrophy conditions (11). We observed reduced expression of key muscle genes that regulate skeletal muscle differentiation, MyoD and Myogenin, in GF mouse skeletal muscle compared to PF mouse skeletal muscle (P < 0.01; Fig. 1D). Similar trends in increased expression of atrophy markers such as Atrogin-1 and Murf-1 with a concomitant down-regulation of MyoD expression were also observed in soleus (slow oxidative) muscle and extensor digitorum longus (fast glycolytic) muscle of GF mice compared to PF mice (P < 0.001; fig. S1, E to H). Fig. 1 Skeletal muscle mass and function in GF mice. (A) Weights of soleus, gastrocnemius, tibialis anterior (TA), quadriceps, and extensor digitorum longus (EDL) muscles from PF mice, GF mice, and C-GF mice. The number of mice used per experimental group is the following: Soleus muscle (PF, n = 13; GF, n = 14; C-GF, n = 10), gastrocnemius muscle (PF, n = 15; GF, n = 14; C-GF, n = 13), TA muscle (PF, n = 14; GF, n = 13; C-GF, n = 12), quadriceps muscle (PF, n = 15; GF, n = 14; C-GF, n = 13), and EDL muscle (PF, n = 13; GF, n = 12; C-GF, n = 13). (B) Shown are changes in expression of genes encoding myosin heavy chain (MyHC) isoforms in TA muscles of PF (n = 7), GF (n = 7), and C-GF (n = 10) mice. (C) Shown are changes in expression of Atrogin-1, Murf-1, and FoxO3 genes in TA muscles from PF (n = 7), GF (n = 7), and C-GF (n = 9) mice. (D) Shown are changes in expression of genes encoding the skeletal muscle–specific transcription factors MyoD and Myogenin in TA muscle samples from PF (n = 7), GF (n = 7), and C-GF (n = 9) mice. (E) Shown is immunoblot analysis of protein lysates from TA muscles harvested from PF, GF, and C-GF mice, indicating expression of Atrogin-1 (n = 4 mice per group), Murf-1 (n = 4 mice per group), and phosphorylated AMPK (p-AMPK; n = 5 mice per group). (F) The results in the histogram are expressed as the ratio of relative intensity of Atrogin-1 and Murf-1 protein expression normalized to tubulin as a loading control and the intensity of p-AMPK expression relative to total AMPK expression. Data are expressed as means ± SEM. Data were analyzed using ANOVA followed by Tukey’s post hoc test and were considered statistically significant at *P < 0.05, **P < 0.01, and ****P < 0.0001 between indicated groups. Glucocorticoids are known to induce skeletal muscle atrophy under various pathological conditions (12). The transcription factor Kruppel-like factor 15 (KLF15) is one of the target genes activated by the glucocorticoid receptor and is involved in the regulation of several metabolic processes in skeletal muscle including the up-regulation of branched-chain aminotransferase 2 (BCAT2), which in turn induces the degradation of branched-chain amino acids (BCAAs) (13). A surge in serum corticosterone concentrations was observed in GF mice compared to PF mice (P < 0.05; Fig. 2A), along with up-regulation of Klf15 expression in tibialis anterior muscle of GF mice compared to PF mice (P < 0.01; Fig. 2B). The observed muscle atrophy in GF mice was also associated with activation of enzymes involved in the BCAA catabolic pathway. BCAAs are transaminated by BCAT2 to generate branched-chain α-keto acids, which in turn are subjected to oxidative decarboxylation by branched-chain α-keto acid dehydrogenase (BCKDH) to produce coenzyme A esters. BCKDH activity is negatively regulated by the BCKDH kinase (BCKDK). Increased gene expression of Bcat2 and Bckdh was observed in tibialis anterior muscle of GF mice compared to PF mice, whereas the expression of the inhibitor Bckdk was reduced, suggesting increased BCAA catabolism in skeletal muscle of GF mice (P < 0.05; Fig. 2C). Fig. 2 Branched chain amino acid (BCAA) metabolism in skeletal muscle of GF mice. (A) Shown are measurements of serum corticosterone concentrations in PF (n = 17), GF (n = 16), and C-GF (n = 10) mice. (B) Shown is expression of the Klf15 gene in TA muscles from PF (n = 7), GF (n = 7), and C-GF (n = 9) mice. (C) Shown are changes in expression of genes involved in BCAA catabolism (Bcat2, Bckdk, and Bckdh) in TA muscles of PF (n = 6), GF (n = 7), and C-GF (n = 9) mice. (D) Shown are changes in the expression of the genes Igf1 and Igf-binding proteins (Igfbps) in TA muscle of PF mice (n = 7), GF mice (n = 7), and C-GF (n = 9) mice. Data are expressed as means ± SEM. Data are analyzed using ANOVA followed by Tukey’s post hoc test and were considered statistically significant at *P < 0.05, **P < 0.01, and ***P < 0.001 between indicated groups. Skeletal muscle mass is maintained by the balance between protein synthesis and protein degradation. To monitor a potential decline in the protein synthesis pathway, we assessed the Igf1-Akt-mTOR growth-promoting pathway. Whereas the expression of the insulin-like growth factor 1 gene (Igf1) was reduced in tibialis anterior muscle of GF mice compared to PF mice, Igf1 expression was normalized when GF mice were transplanted with the gut microbiota of pathogen-free mice (C-GF) (P < 0.01; Fig. 2D). The amount of Igf1 protein remained unaltered in the serum of GF mice (fig. S1I), suggesting additional Igf1 regulatory loops possibly involving Igf-binding proteins (IGFBPs) that are known to regulate Igf1 function. Of the six IGFBPs, we observed increased expression of the Igfbp3 gene in tibialis anterior muscle of GF mice compared to PF mice and C-GF mice (P < 0.01; Fig. 2D), in line with a previous observation (14). Increased expression of Igfbp3 is known to exert an inhibitory effect on skeletal muscle growth, suggesting one explanation for the reduced muscle mass of GF mice (15). However, the activation profile of Akt, the mammalian target of rapamycin (mTOR), and its downstream effector, the S6 ribosomal protein, were unaffected in GF mouse muscle (fig. S1, J and K).

Altered metabolism in skeletal muscle of GF mice Given that skeletal muscle of GF mice showed signs of atrophy, we next investigated the oxidative metabolic capacity of skeletal muscle. Histological staining revealed reduced activity of the mitochondrial enzyme succinate dehydrogenase (SDH; Fig. 3A) and reduced expression of the Sdh gene (P < 0.05; Fig. 3) in GF mouse muscle compared to muscle of PF mice. Mitochondrial DNA content was also reduced in GF mouse muscle but was restored when GF mice were transplanted with the gut microbiota of pathogen-free mice (C-GF) (P < 0.05; Fig. 3C). In addition, we observed reduced gene expression of mitochondrial biogenesis markers, such as peroxisome proliferator–activated receptor γ coactivator 1α (Pgc1α) and mitochondrial transcription factor A (Tfam) in skeletal muscle of GF mice compared to PF mice (P < 0.05; Fig. 3D). Moreover, we noticed reduced expression of genes encoding different mitochondrial oxidative phosphorylation complexes in skeletal muscle of GF mice compared to PF mice, including genes encoding cytochrome oxidase subunits of complex IV (CoxVa, CoxVIIb, and CytC) known to be involved in the electron transport chain (P < 0.05; Fig. 3E). Similar observations supporting dysfunctional mitochondrial biogenesis and oxidative capacity were also observed in two other skeletal muscle subtypes, the soleus (oxidative) and extensor digitorum longus (glycolytic) muscle of GF mice compared to PF mice (P < 0.01; fig. S2, A to D). The protein expression profile of the different oxidative phosphorylation complexes of the electron transport chain (fig. S6A) was also altered in muscle from GF mice compared to PF mice. Despite a possible reduction in oxidative metabolic capacity, GF mice performed as well as PF mice when challenged to exercise until exhaustion (fig. S2E). Fig. 3 Oxidative capacity of the skeletal muscle of GF mice. (A) Representative images of TA muscle sections from PF, GF, and C-GF mice stained for the enzyme SDH. (B) Shown is expression of the Sdh gene in TA muscle from PF (n = 7), GF (n = 7), and C-GF (n = 9) mice. (C) Quantitative analysis of the ratio of mitochondrial DNA (mtDNA) to nuclear DNA in gastrocnemius muscles from PF (n = 4), GF (n = 5), and C-GF (n = 5) mice. (D and E) Shown are changes in expression of the Pgc1α and Tfam genes (D), and the CoxVa, CoxVIIb, and CytC genes (E) in TA muscles of PF (n = 6), GF (n = 7), and C-GF (n = 9) mice. (F) Shown are changes in expression of genes involved in glucose metabolism (Pfk, Pk, Ldh, and Pdh) in TA muscles of PF (n = 6), GF (n = 7), and C-GF (n = 9) mice. (G) Shown are changes in expression of genes involved in the fatty acid oxidation pathway (Lcad, Mcad, and Cpt1b) in TA muscles of PF (n = 7), GF (n = 7), and C-GF (n = 9) mice. (H) Shown is the amount of glycogen in quadriceps muscles of PF (n = 4), GF (n = 3), and C-GF (n = 4) mice. Data are expressed as means ± SEM. Data were analyzed using ANOVA followed by Tukey’s post hoc test and were considered statistically significant at *P < 0.05, **P < 0.01, and ***P < 0.001 between indicated groups. These results prompted us to undertake further metabolic analyses in GF mice. Analysis of serum metabolic markers revealed reduced quantities of glucose and insulin in GF mice compared to PF mice (P < 0.05; fig. S2F). Further investigation of metabolic pathways for fatty acid and glucose metabolism in GF mouse muscle revealed reduced expression of glycolytic genes encoding the enzymes phosphofructokinase (Pfk), pyruvate kinase (Pk), lactate dehydrogenase (Ldh), and pyruvate dehydrogenase (Pdh) in tibialis anterior muscle from GF mice compared to PF mice (P < 0.05; Fig. 3F). Expression of the genes encoding these enzymes was restored when GF mice were transplanted with the gut microbiota of pathogen-free mice (C-GF) (P < 0.01; Fig. 3F). A similar trend of reduced expression of glycolytic genes was also observed in the soleus and extensor digitorum longus muscles of GF mice compared to PF mice (P < 0.01; fig. S2, G and H). No differences in expression of genes encoding medium-chain acyl–coenzyme A dehydrogenase (Mcad) and muscle-specific carnitine palmitoyltransferase 1b (mCpt1b) involved in fatty acid oxidation (Fig. 3G) or phosphorylation of acetyl–coenzyme A carboxylase (fig. S2, I and J) were observed in tibialis anterior muscle of GF mice compared to PF mice. Cholesterol and free fatty acids in serum also remained unaltered in GF mice compared to PF mice (fig. S2F). Further analysis of glucose uptake in skeletal muscles, using FDG-PET/MRI (fluorodeoxyglucose–positron emission tomography/magnetic resonance imaging) imaging, revealed no obvious differences in glucose uptake for the back and hindleg muscles of GF mice compared to PF mice (fig. S2, K and L). However, we observed an increased accumulation of glycogen in the quadriceps (fast glycolytic) muscles of GF mice compared to PF mice (P < 0.001; Fig. 3H), implying possible impaired utilization of glucose by the quadriceps muscle of GF mice. To further corroborate our findings, we assessed whether disruption of the gut microbiota–host environment using antibiotics had effects on skeletal muscle. We monitored muscle samples from PF mice exposed to low-dose penicillin to examine consequences of disruption of host metabolic functions mediated by antibiotics through alterations in microbial community composition (16, 17). Whereas low-dose penicillin increased FoxO3 gene expression and reduced MyoD gene expression in skeletal muscle of PF mice (P < 0.05; fig. S3A), the expression of Atrogin-1, Murf-1, Pgc1α, and Tfam genes remained largely unaffected under these experimental conditions (fig. S3, A and B). However, electron transport chain genes (CoxVa, CoxVIIb, and CytC) were down-regulated in PF mice treated with low-dose penicillin compared to untreated animals (P < 0.05; fig. S3C), indicating reduced oxidative metabolic capacity in skeletal muscle.

Altered metabolites in skeletal muscle, liver, and serum of GF mice To characterize the metabolic phenotype of GF mice in detail, we measured proton nuclear magnetic resonance (NMR) spectra at 600 MHz for the skeletal muscle, liver, and serum samples from GF, PF, and C-GF mice (Fig. 4). Cross-validated principal components analysis and orthogonal partial least squares analysis models revealed differences in 1H NMR metabolic profiles for the skeletal muscle, liver, and serum from GF mice compared to PF mice and GF mice compared to C-GF mice (fig. S4, A to C). Transplanting GF mice with gut microbiota of PF mice, as observed in C-GF mice normalized the metabolite profiles to those observed in PF mice. Fig. 4 Metabolite analysis of the muscle, liver, and serum from GF mice. (A to C) Shown is the average 1H NMR spectrum of hydrophilic phase after Folch extraction for 25 metabolites. The 1H NMR spectrum is shown for (A) gastrocnemius muscle from PF (n = 8), GF (n = 8), and C-GF (n = 10) mice; (B) liver tissue from PF (n = 7), GF (n = 8), and C-GF (n = 10) mice; (C) serum from PF (n = 8), GF (n = 8), and C-GF (n = 9) mice. 1, taurocholic acid; 2, bile acids; 3, low-density lipoprotein (LDL); 4, very-low-density lipoprotein (VLDL); 5, leucine; 6, 3-hydroxybutyrate; 7, alanine; 8, acetate; 9, glutamine; 10, glutamate; 11, pyruvate; 12, glutathione; 13, hypotaurine; 14, dimethylamine; 15, sarcosine; 16, trimethylamine; 17, dimethylglycine; 18, unknown (δ 3.11) (s); 19, choline; 20, glycerophosphorylcholine; 21, taurine; 22, betaine; 23, glycine; 24, unknown (δ 3.59) (d); and 25, unknown (δ 3.71) (s). Metabolites in red were found in higher concentrations in GF mice compared to C-GF or PF mice; metabolites in blue were found in lower concentrations in GF mice compared to C-GF or PF mice. s, singlet; d, doublet; a.u., arbitrary units. Our metabolite profiling identified large quantities of amino acids such as alanine and glycine in skeletal muscle of GF mice (Fig. 4A and Table 1). The expression of the gene encoding alanine transaminase (Alt) that results in transamination of alanine was also increased in muscle of GF mice (P < 0.01; fig. S5A). Typically, the amino acids alanine and glycine, and their carbon skeletons, enter different metabolic pathways to generate energy. However, we did not observe any change in adenosine 5′-triphosphate (ATP) concentrations in the skeletal muscle of GF mice compared to PF mice or C-GF mice (fig. S5B). We next tested whether higher alanine concentrations in the muscle of GF mice served as a source for hepatic gluconeogenesis through the glucose-alanine shuttle. Reduced alanine (Fig. 4B and Table 1) along with low expression of the gene encoding glucose-6-phosphatase (G6Pase), a gluconeogenic marker, was observed in the liver of GF mice compared to PF mice (P < 0.05; fig. S5C). Table 1 Differences in metabolite concentrations between GF, PF, and C-GF mice. s, singlet; d, doublet; dd, doublet of doublets; t, triplet; q, quartet; m, multiplet. View this table: A large amount of glycine was observed in skeletal muscle of GF mice compared to PF mice (Fig. 4A and Table 1). Apart from a possible increase in protein degradation of muscle tissue, glycine can be derived from intermediates of glycolysis (18). However, no changes in expression of genes encoding key enzymes of this intermediate pathway, such as phosphoglycerate dehydrogenase (Phgdh) and serine hydroxymethyltransferases (Shmt1 and Shmt2) were observed in the muscle of GF mice (fig. S5D). Glycine can also be generated from choline via betaine, dimethylglycine, and sarcosine. Reduced quantities of choline and dimethylglycine along with higher amounts of glycine were found in the serum of GF mice compared to PF mice (Fig. 4C and Table 1). Intermediates in the choline metabolic pathway, glycerophosphocholine and betaine, were also reduced in the liver of GF mice (Fig. 4B and Table 1). However, no changes in expression of the genes encoding the enzymes involved in these intermediate steps: Choline dehydrogenase (Chdh), betaine homocysteine S-methyltransferase (Bhmt), Bhmt2, and sarcosine dehydrogenase (Sardh) were observed in the skeletal muscle of GF mice when compared to PF mice (fig. S5E). The metabolite profile of the liver from GF mice was characterized by high taurine, hypotaurine, and tauro-conjugated bile acids (taurocholic acid), showing that bile acid metabolism was affected in GF mice compared to PF mice. We also observed increased dimethylamine, a product of choline degradation, in the liver of GF mice compared to PF mice (Fig. 4B and Table 1). Furthermore, glycerophosphocholine, betaine (trimethylglycine), and oxidized glutathione were also present in lower amounts in the liver of GF mice, indicating potential perturbation of the cysteine metabolic pathway although the amount of sarcosine remained high. In addition, glutamine, alanine, leucine, and valine along with glutamate were reduced in the liver of GF mice compared to PF mice. Pyruvate was lower in the liver of GF mice compared to PF mice. Expression profiling of genes encoding enzymes involved in mitochondrial oxidative and glucose metabolism revealed reduced expression of several genes involved in these pathways, indicating metabolic dysfunction in the liver of GF mice compared to PF mice (P < 0.01; fig. S5, F and G). The amount of glycine was high in serum and muscle of GF mice compared to PF or C-GF mice (Fig. 4C and Table 1). However, the amounts of choline, trimethylamine, and dimethylglycine were lower in serum of GF mice compared to PF or C-GF mice. In addition, acetate, pyruvate, and 3-hydroxybutyrate were lower in serum of GF mice. We also observed differences in serum lipid profiles, with high amounts of very-low-density lipoproteins and low amounts of low-density lipoproteins in GF mice compared to PF mice (Fig. 4), in agreement with an earlier report (19). Overall, the metabolite data indicated disrupted energy homeostasis in GF mice compared to PF mice, with a marked perturbation of the amino acid metabolic pathway (Fig. 5). Fig. 5 Multicompartment metabolic reaction network. Metabolites are connected on the basis of the shortest paths of reactions that are mediated by enzymes encoded in the Mus musculus genome or on the basis of nonenzymatic reactions from the Kyoto Encyclopedia of Genes and Genomes (KEGG) database. Metabolites in orange were found in higher concentrations in GF mice compared to PF or C-GF mice. Conversely, metabolites in blue were found in lower concentrations in GF mice compared to PF and C-GF mice. Alanine was higher in the skeletal muscle (gastrocnemius) and lower in the liver. The background shading indicates the three different subnetworks for gastrocnemius muscle (purple), liver (green), and serum (pink). Overlap exemplifies similarity between affected metabolic compartments.

Altered expression of genes encoding NMJ proteins in GF mice Given that we observed low quantities of choline in the serum of GF mice (Fig. 4C and Table 1) and that choline is a substrate for the derivation of the neurotransmitter acetylcholine and is essential for membrane integrity (20), we screened an array of molecules that affect NMJ development and function including acetylcholine receptors (21). We observed reduced expression of genes encoding different acetylcholine receptor subunits (Chrna1, Chrnb, Chrne, and Chrnd) in tibialis anterior muscle of GF mice compared to PF mice (P < 0.05; Fig. 6A), suggesting a potential impairment of acetylcholine receptor assembly. We therefore monitored the expression of genes associated with formation, maturation, and maintenance of NMJs, including Rapsyn, a 43-kDa receptor-associated synaptic protein (22) and the low-density lipoprotein receptor–related protein 4 (Lrp4), both reported to be important for the development and assembly of acetylcholine receptors in NMJs (23). Expression of both Rapsyn and Lrp4 genes was reduced in GF mice, and this change was reversed when GF mice were transplanted with the gut microbiota of PF mice (C-GF) (P < 0.05; Fig. 6B). Lrp4 associates with muscle-specific kinase (MuSK) to form a receptor complex necessary for agrin to bind (24). Whereas we noted elevated expression of the MuSK gene, no change in expression of the Agrn gene encoding agrin was observed in muscle of GF mice compared to PF mice (P < 0.05; Fig. 6B). The reduced expression of the gene encoding troponin in skeletal muscle of GF mice compared to PF mice and C-GF mice suggested possible impairment of myofiber contractility (P < 0.01; Fig. 6C). We therefore evaluated muscle strength in GF mice. GF mice displayed reduced muscle strength compared to PF mice when examined in a weights test (P < 0.05; Fig. 6D). GF mice also exhibited reduced locomotor and rearing activity compared to PF mice (P < 0.01 and P < 0.001; Fig. 6, E and F, respectively) and C-GF mice (P < 0.05 and P < 0.05; Fig. 6, E and F, respectively). Fig. 6 Differential expression of mouse neuromuscular junction proteins between GF and PF mice. (A) Shown are changes in expression of genes encoding acetylcholine receptor subunits (Chrn) in the TA muscle of GF, PF, and C-GF mice. Genes include Chrna1 (PF, n = 7; GF, n = 7; C-GF, n = 9), Chrnb (PF, n = 7; GF, n = 7; C-GF, n = 8), Chrnd (PF, n = 7; GF, n = 7; C-GF, n = 7), and Chrne (PF, n = 7; GF, n = 7; C-GF, n = 7). (B) Shown are changes in expression in TA muscle of PF, GF, and C-GF mice of genes encoding the receptor-associated protein of the synapse (Rapsyn; PF, n = 7; GF, n = 7; C-GF, n = 9), low-density lipoprotein receptor–related protein 4 (Lrp4), and Agrin (Agrn) (PF, n = 7; GF, n = 7; C-GF, n = 9). (C) Shown are changes in expression of the gene encoding fast-twitch troponin (Tnn) in TA muscle of PF (n = 6), GF (n = 7), and C-GF (n = 9) mice. (D) Analysis of hindlimb grip strength using the weights test in PF, GF, and C-GF mice (n = 6 per group). (E to F) Shown is the spontaneous activity of GF, PF, and C-GF mice in the open-field test measured by cumulative distance traveled (E) and cumulative vertical activities (F). PF (n = 8), GF (n = 9), and C-GF (n = 8) mice were monitored over a 2-hour period. All data are expressed as means ± SEM. Data were analyzed using ANOVA followed by Tukey’s post hoc test and were considered statistically significant at *P < 0.05, **P < 0.01, and ***P < 0.001 between indicated groups.