Glycolytic-to-oxidative fiber-type switch and mTOR signaling activation are early-onset features of SBMA muscle modified by high-fat diet

Animal care and experimental procedures were conducted in accordance with the Italian Institute of Technology and the University of Trento ethics committees and were approved by the Italian Ministry of Health. Generation and genotyping of knock-in AR21Q and AR113Q mice were previously described [106]. Mice were randomized and fed a standard diet (Mucedola 4RF21), purified normal chow diet (NCD, D12450B Research Diet), and high-fat diet (HFD, D12451 Research Diet) as indicated. The operator was blind for genotype and treatment. For rotarod analysis (Ugo Basile), mice received a weekly session, which included one trial followed by two test trials at 21 rpm speed for a maximum period of 600 s and the average of recordings was used. For rapamycin treatment, 154-day-old mice were injected intraperitoneally with rapamycin (4.0 mg/kg, Gold Biotechnology). For analysis of the rate of protein synthesis, animals were starved 30 min, injected with puromycin (0.040 μmol/g puromycin dissolved in 100 μl PBS), and sacrificed 30 min after injection. In vivo and ex vivo muscle force was measured as previously described [8].

Anonymized control and patient biopsy sample collection was approved by the ethics committee of the University of Padova (Italy). Written informed consent was obtained from each patient. All patients who underwent muscle biopsy were clinically affected and showed weakness and/or fasciculation and/or muscle atrophy. Myopathic changes together with neurogenic atrophy were observed in muscle biopsies.

Muscles collected immediately after euthanasia were flash-frozen in isopentane precooled in liquid nitrogen and embedded in optimal cutting temperature (OCT) compound (Tissue Tek, Sakura), and cross sections (10 µm thick) were cut with a cryostat (CM1850 UV, Leica Microsystems). Cryosections were processed for hematoxylin and eosin (H/E) and nicotinamide adenine dinucleotide (NADH) staining, as previously described [70]. For immunofluorescence analysis, muscle cryosections were incubated with M.O.M. IgG blocking solution (Vector), washed with phosphate-buffered saline (PBS) and incubated with a solution of PBS containing 0.5 % bovine serum albumin (BSA). Full list of antibodies used is provided in the Supplementary Information. Sections were mounted with aqueous mounting medium (Fluorescence Mounting Medium, Dako). Images were taken using an upright epifluorescence microscope (Zeiss Axio Imager M2) equipped with an X-Cite 120Q fluorescence light source and a Zeiss Mrm Color Camera. Multichannel images and mosaics were taken using Zeiss Axio Vision Software (V.4.8.2 SP3).

Muscles were mechanically pulverized and homogenized in lysis buffer [20 mM HEPES, 5 mM EGTA, 2 % sodium dodecyl sulfate (SDS)] and processed by SDS–polyacrylamide gels (SDS-PAGE) as previously described [70]. Full list of antibodies used is provided in the Supplementary Information. Signal intensities were quantified by ImageQuant LAS 4000 mini (GE Healthcare BioSciences). Lactate dehydrogenase (LDH) and citrate synthase activity was measured as previously described [42, 99]. For proteasome activity assay, muscles were lysed using ice-cold lysis buffer [50 mM HEPES, 5 mM EDTA, 150 mM NaCl, 2 mM ATP, 1 % Triton X-100], sonicated and centrifuged. Protein extracts were incubated in 25 mM HEPES, 0.5 mM EDTA, 0.05 % NP40, 0.001 % SDS, and 0.5 mM N-Succinyl-Leu-Leu-Val-Tyr-7-Amido-4-Methylcoumarin (Sigma). The absorption was recorded at 460 nm by excitation at 355 nm (Victor3-V luminometer, PerkinElmer).

Untargeted metabolomics experiments were performed as previously described [2]. For real-time analysis, total RNA was extracted with Trizol (Invitrogen), purified using RNeasy MinElute Cleanup Kit (QIAGEN), and reverse transcribed into cDNA using the iScript Reverse Transcription Supermix (Bio-Rad). Gene expression was measured by quantitative real-time PCR using 7900 HT Fast Real-Time PCR System (Applied Biosystems). The list of primers is provided in Supplementary Table 1. For microarray analysis, total RNA was purified using RNeasy MinElute Cleanup Kit (QIAGEN). High-quality RNA was used for labeling and array hybridization (Agilent Mouse GE 4X44 K slides). Slides were scanned on the Agilent DNA Microarray Scanner (G2505C) using the AgilentHD_GX_1Color Profile (scan area: 61 × 21.6 mm; scan resolution: 5 μm, dye channel set to 100 % Green PMT) of Agilent ScanControl software 8.1.3. Images were analyzed with Feature Extraction Software 10.7.3.1 (Agilent Technologies) using default parameters (protocol GE1_107_Sep09). Expression data were analyzed using the limma package from R and false discovery rate (FDR) control for statistical assessment of the microarray data (corrected P < 0.05 were considered significant). Gene set enrichment analysis (GSEA) was performed to identify sets of related genes altered in each experimental group (http://​www.​broad.​mit.​edu/​gsea). Heat maps were generated using MeV TM4 software V 4.9 (http://​www.​tm4.​org). Microarray data are available at the Gene Expression Omnibus database under accession number GSE68441.

Mitochondrial membrane potential was measured in isolated fibers from flexor digitorum brevis muscles. Mitochondrial membrane potential was measured by epifluorescence microscopy based on the accumulation of tetramethyl rhodamine methyl ester (TMRM) fluorescence, as previously described [82]. For mitochondrial complex activity, total muscle lysates were extracted and the enzymatic activities of the respiratory chain complexes I–IV were assayed as previously described [99].

All data are presented as mean ± standard error of the mean (SEM). Statistical differences of continuous data from two experimental groups were calculated using two-sample t tests. Comparisons of data from more than two groups were performed using a one-way ANOVA followed by a Fisher’s least significant difference post hoc test. Statistical comparisons of lifespan curves were performed using the log-rank and Gehan–Breslow–Wilcoxon tests. Statistical significance threshold was set at P < 0.05, unless otherwise indicated.

To elucidate the pathological processes underlying muscle atrophy in SBMA, we used SBMA knock-in mice that express AR with a pathogenic polyglutamine tract of 113 glutamine residues (AR113Q) and, as controls, AR21Q and wild-type littermates [72]. No signs of muscle atrophy were detected in AR21Q mice (Supplementary Fig. 1a), as previously described [107]. To determine whether expression of polyglutamine-expanded AR differentially affects muscles composed primarily of either type I slow-oxidative fibers (e.g., soleus) or type II fast-glycolytic fibers [e.g., quadriceps, gastrocnemius, tibialis anterior (TA)], we assessed the wet weight of these muscles as a function of disease progression. The mass of quadriceps, gastrocnemius, and TA was reduced by 13–35 % beginning at 60 or 90 days of age, whereas the mass of soleus did not change over the course of disease in AR113Q mice compared to age-matched control mice (Fig. 1a; Supplementary Fig. 1b). By H/E analysis, the mean myofiber cross-sectional area (CSA) of quadriceps, gastrocnemius, and TA muscles was decreased by 10–30 % in 90- and 180-day-old AR113Q mice, whereas that of soleus was increased (Supplementary Fig. 2a–c). Tetanic force production was decreased by 27 % in the gastrocnemius, but not soleus (Supplementary Fig. 1c, d), indicating that structural alterations are associated with functional deficits. Notably, the degree of atrophy correlated with the levels of expression of AR, suggesting a dose-dependent effect of polyglutamine-expanded AR in muscle (Fig. 1b). These results indicate that expression of AR113Q preferentially affects glycolytic muscles compared to oxidative muscles.

Analysis of muscle biopsies has revealed that the number of type I oxidative fibers is increased in the glycolytic muscles of SBMA patients, suggesting metabolic alterations [38]. By NADH staining, we observed a progressive shift from fast-glycolytic towards slow-oxidative fiber subtype in the quadriceps and gastrocnemius of AR113Q mice, which was detected as early as 40 days of age (Fig. 1c; Supplementary Fig. 3a, b). Staining of type IIa (fast-oxidative) and IIb (fast-glycolytic) fibers confirmed that the number of type IIb fibers was decreased, whereas that of type IIa was increased in the quadriceps of AR113Q mice. Notably, the mean CSA of oxidative fibers was decreased by 10 % in 40-day-old AR113Q mice and progressed to 16 % in 180-day-old mice, whereas that of glycolytic fibers was decreased by 10 % at 40 days of age and progressed to 40 % at 180 days of age in the quadriceps and gastrocnemius (Fig. 1d; Supplementary Fig. 4a, b). These results indicate an early-onset glycolytic-to-oxidative fiber-type switch with atrophy of glycolytic fibers exceeding that of oxidative fibers in the muscle of SBMA knock-in mice.

Given that oxidative and glycolytic fibers primarily use oxidative phosphorylation and glycolysis to generate ATP, respectively, we hypothesized that the lipid profile of AR113Q mice was altered. Using untargeted high-resolution LC–MS/MS lipidomic and unsupervised principal component analyses, we identified major lipid alterations in muscle, but not serum, with 45 lipids significantly (P ≤ 0.001) enriched (fold change >1.5) and 22 lipids decreased (fold change <0.5) in AR113Q mice (Fig. 2a; Supplementary Fig. 5a; Supplementary Table 2). Lysophosphatidylcholines, lysophosphatidylethanolamines, ceramides, diglycerides, and polyunsaturated fatty acids were increased, whereas species enriched in polyunsaturated lipids and phosphatidylglycerol were decreased, suggesting increased lipid synthesis and turnover. To investigate the molecular pathways underlying the metabolic alterations detected in glycolytic SBMA muscles, we performed whole genome microarray analysis on quadriceps muscle. We identified 550 upregulated (fold change ≥1.5) genes and 286 downregulated (fold change ≤0.6) genes in the quadriceps of 180-day-old AR113Q mice compared to age-matched control mice (P < 0.05, false discovery rate <0.05) (Supplementary Fig. 5b; Supplementary Table 3). Consistent with the glycolytic-to-oxidative metabolic shift described above, the transcript levels of markers of oxidative fibers, such as troponin I1 (Tnni1), troponin C1 (Tnnc1), troponin T1 (Tnnt1), ankyrin repeat domain 2 (Ankrd2), myoglobin (Mb), myosin heavy chain 7b (Myh7b), were all upregulated, whereas those of glycolytic fibers were either decreased, such as 5′-AMP-activated protein kinase subunit gamma-3 (Prkag3), or remained unchanged, such as troponin I2 (TnnI2), myosin heavy chain 4 (Myh4), and parvalbumin (Pvalb) (Supplementary Table 4). Gene set enrichment analysis (GSEA) revealed that the top pathways enriched in genes differentially expressed in AR113Q and control mice were metabolic pathways (Fig. 2b; Supplementary Table 5). The gene set “starch/sucrose metabolism”, which includes glycolytic genes, was downregulated, whereas “peroxisome proliferator-activated receptor alpha (PPARα) signaling”, “glycerolipid and fatty acid metabolism”, and “tricarboxylic acid (TCA) cycle” were upregulated. Similar changes in metabolic pathways were also detected in transgenic mice overexpressing AR97Q (Supplementary Table 5) [57]. By real-time PCR analysis, we validated the upregulation of genes involved in lipid metabolism, such as fatty acid synthase (Fasn), which catalyzes the synthesis of palmitate from malonyl-coenzyme A and acetyl-coenzyme A in a process that requires NADPH, aldehyde dehydrogenase 9 subfamily A1 (Aldh9a1), which detoxifies from accumulation of aldehydes generated by lipid peroxidation through NAD(P)+-dependent oxidation to carboxylic acids, diacylglycerol O-acyltransferase 2 (Dgat2), which catalyzes the synthesis of triglycerides from diacylglycerol and acyl-coenzyme A, the mitochondrial protein cell death-inducing DNA fragmentation factor alpha subunit-like effector A (Cidea), which plays a role in thermogenesis and lipolysis, the sarcoplasmic reticulum SERCA Ca(2+)-ATPase A2 (Atp2a2), and Ppara, which is involved in the regulation of muscle metabolism (Fig. 2c). Importantly, the expression of these genes was not altered in soleus (Supplementary Fig. 6a). The expression and activity of key enzymes in the glycolytic pathway, such as hexokinase II (Hk II), phosphofructokinase 1 (Pfkm), 6-phosphofructo-2-kinase/fructose-2,6-biphosphatase 3 (Pfkb3), glyceraldehyde 3-phosphate dehydrogenase (Gapdh), phosphoglycerate kinase 1 (Pgk1), and lactate dehydrogenase (LDH), were downregulated in the quadriceps muscle of AR113Q mice, whereas the activity of citrate synthase (CS) was increased (Fig. 2d–f; Supplementary Fig. 5c, d). Alterations in glycolytic gene expression preceded those in lipid gene expression, as they were detected as early as 90 days of age in the quadriceps of AR113Q mice (Supplementary Fig. 7a, b). In soleus muscle, the transcript levels of Gapdh, and not the other glycolytic genes, were decreased by 40 % at 180 days of age (Supplementary Fig. 6b). Our microarray analysis also revealed that the transcript levels of SLC2A3, which encodes the glucose transporter 3 (GLUT3), were decreased by 60 %, suggesting altered glucose uptake, whereas those of SLC2A5, which encodes the solute carrier family 2 (facilitated glucose/fructose transporter), member 5 (GLUT5), was upregulated by 4.7-fold in the muscle of AR113Q mice, suggesting compensatory mechanisms for carbohydrate uptake (Supplementary Table 2). Fasting serum glucose and insulin levels were similar in AR113Q and control mice (Supplementary Fig. 8a), and intraperitoneal glucose and insulin tolerance tests showed normal glucose clearance and insulin sensitivity in AR113Q mice (Supplementary Fig. 8b, c). Moreover, no overt alterations in glycogen storage were detected in the quadriceps of 180-day-old AR113Q mice and SBMA patients (Supplementary Fig. 9). Taken together, these results indicate that glycolytic muscles in SBMA knock-in mice show enhanced lipid metabolism and gene expression programs, as well as impaired glycolysis.

Muscle homeostasis is maintained by a fine balance between protein synthesis and degradation, two processes that simultaneously occur in muscle [100]. We hypothesized that the metabolic changes detected in SBMA muscle alter protein turnover. To assess the rate of new protein synthesis, we used in vivo surface sensing of translation (SUnSET) [92]. We intraperitoneally injected AR113Q and control mice with the antibiotic puromycin, and we analyzed the rate of new protein synthesis by Western blotting using an anti-puromycin antibody. The rate of new protein synthesis was increased by 35 and 73 % in the quadriceps muscle of 90- and 270-day-old AR113Q mice compared to age-matched control mice (Fig. 3a; Supplementary Fig. 10a); this increase was overt even upon 4 h of fasting (Supplementary Fig. 10b). We assessed protein degradation by analyzing ubiquitination of total protein lysates from quadriceps muscle of AR113Q and control mice. The amount of ubiquitinated proteins was increased by 1.33- and 1.84-fold in the quadriceps of 90- and 270-day-old AR113Q mice compared to control mice, respectively (Fig. 3b; Supplementary Fig. 10c). We observed increases in the expression of the E3 ubiquitin ligases, specific of muscle atrophy and regulated by transcription (Smart) and muscle ubiquitin ligase of the SCF complex in atrophy-1 (Musa1) (Fig. 3c) [56, 88], as well as increased proteasome activity starting at 90 days of age in the quadriceps muscle of AR113Q mice, further indicating enhanced protein degradation in the muscle of SBMA mice (Fig. 3d; Supplementary Fig. 10d). Notably, the net amount of protein content in the quadriceps and gastrocnemius muscles of AR113Q mice was reduced at 90 and 270 days of age (Fig. 3e; Supplementary Fig. 10e). These alterations were not detected in soleus (Supplementary Fig. 11a, b). These results show that protein turnover (protein synthesis and breakdown) is increased in SBMA glycolytic, but not oxidative muscles, and that the increased rate of protein degradation is not compensated by the increased rate of protein synthesis.

A key regulator of protein turnover in muscle is mTOR, a sensor of cellular metabolism and a regulator of glycolysis and lipid biosynthesis [30]. mTOR stimulates protein synthesis through phosphorylation of the eukaryotic translation initiation factor 4E binding protein 1 (4EBP1) and S6 kinase 1 (S6K1), which in turn phosphorylates the ribosomal protein S6. mTOR signaling was increased in the quadriceps, but not spinal cord, of 90- and 270-day-old AR113Q male, and not female, mice (Fig. 3f; Supplementary Fig. 12a). S6K1 phosphorylation was increased also in gastrocnemius and TA, which show atrophy, but not soleus, which is spared in this mouse model of SBMA (Supplementary Fig. 12b); the increase in mTOR signaling preceded atrophy, as it was detected as early as 40 days of age (Supplementary Fig. 12c). Enhanced mTOR signaling was not associated with increased expression of AR in male AR113Q mice compared to control mice (Supplementary Fig. 12a). Importantly, we did not detect alterations in the phosphorylation levels of Akt and its downstream effectors glycogen synthase kinase 3β (GSK3β) and forkhead box O transcription factors (FOXOs), suggesting that mTOR activation is independent of Akt. To assess whether enhanced mTOR signaling is responsible for the increased rate of new protein synthesis in SBMA muscle, we administered rapamycin (4 mg/kg/day) to AR113Q and control mice via intraperitoneal injections every day for 20 days. Rapamycin treatment reduced S6 phosphorylation (Fig. 3g) and decreased the rate of new protein synthesis by 28 and 32 % in control and AR113Q mice, respectively (Fig. 3h). Importantly, rapamycin decreased Akt phosphorylation in muscle, the body weight of AR113Q mice starting from 16 days of treatment, without altering food intake, and testis size in both AR113Q and control mice (Supplementary Fig. 13a–d). Rapamycin did not modify skeletal muscle mass (Supplementary Fig. 13e). Taken together, these results indicate that mTOR signaling is enhanced prior to atrophy, and it increases the rate of new protein synthesis in SBMA glycolytic muscles.

Denervation may cause metabolic alterations in muscle associated with atrophy and loss of fast-glycolytic fibers [64]. Therefore, we investigated whether the metabolic changes detected in quadriceps result from denervation in SBMA knock-in mice (Fig. 4a). The transcript levels of two denervation markers, acetylcholine receptor subunit gamma (Chrng) and muscle-specific kinase (Musk), were upregulated by 90 days of age, but not at 40 days of age, when the metabolic changes were started to be detected. Moreover, these denervation markers were not upregulated in the soleus of 180-day-old AR113Q mice (Supplementary Fig. 14a). These results indicate that the metabolic alterations that occur in SBMA glycolytic muscles precede denervation and are not initiated, rather maybe contributed at later stages of disease, by pathogenic processes occurring in motor neurons.

mTOR controls mitochondrial gene expression through modulation of PGC1α (Ppargc1a) function [20]. PGC1α is a master regulator of muscle metabolism, and when overexpressed it shifts muscle metabolism towards oxidative phosphorylation [47]. PGC1α expression correlated with disease progression in the quadriceps, but not soleus of AR113Q mice (Fig. 4b; Supplementary Fig. 14b). In quadriceps, PGC1α induction showed a trend for significance (P = 0.08) at 40 days of age, suggesting that its dysregulation occurs very early in disease pathogenesis, with dysregulation increasing with disease progression. Importantly, rapamycin treatment decreased PGC1α transcript levels, indicating that in SBMA muscle PGC1α expression is regulated by mTOR (Fig. 4c). PGC1α promotes mitochondrial biogenesis [104]. Markers of mitochondria fission and fusion, such as dynamin-related protein 1 (Dnm1 l, Drp1) and opticatrophy gene 1 (Opa1), were upregulated in the quadriceps, but not soleus of AR113Q mice (Fig. 4b; Supplementary Fig. 14b). Moreover, the activity of mitochondrial complexes was increased in the quadriceps of AR113Q mice (Fig. 4d; Supplementary Fig. 15a). Mitochondrial abnormalities have been reported in SBMA [79]. We measured mitochondrial membrane potential in myofibers isolated from the fast-glycolytic flexor digitorum brevis muscle upon treatment with the F₁F₀-ATPase blocker, oligomycin (Fig. 4e; Supplementary Fig. 15b). We found that the number of fibers with mitochondria depolarized by oligomycin was significantly (P < 0.05) increased by 40 % in 90- and 180-day-old AR113Q mice. These data indicate that despite the increased beta oxidative metabolism, mitochondria are dysfunctional in SBMA muscle and are a potential source of oxidative stress.

Mitochondrial depolarization and accumulation of reactive oxygen species represent a signal for autophagy activation [90]. The autophagy markers microtubule-associated protein 1A/1B-light chain 3 (LC3) and sequestosome 1 (Sqstm1, p62) are upregulated in the muscle of AR113Q mice, indicating enhanced autophagy [16, 83]. The transcript levels of these autophagy markers were increased in the quadriceps, but not soleus, of 180-day-old AR113Q mice (Fig. 4f; Supplementary Fig. 14c). Upregulation of these autophagy markers in quadriceps was preceded by transcriptional induction of TfEB, the master regulator of autophagy gene expression [87]. To monitor autophagy flux, we measured the soluble mature form of LC3, namely LC3I, the membrane-bound form, LC3II, which accumulates upon autophagosome formation, and p62, which accumulates upon inhibition of autophagy flux (Fig. 4g; Supplementary Fig. 16a–c). Both LC3I and II increased at late stages of disease (180 and 270 days of age), and the ratio LC3II/LC3I did not change over the progression of disease, whereas p62 significantly accumulated in 270-day-old mice, suggesting defects in autophagy flux at very late stages of disease. These data indicate that metabolic changes in SBMA muscle precede TFEB upregulation and autophagy induction.

mTOR and PGC1α can be inhibited by feeding mice an HFD [6, 97]. Therefore, we administered AR113Q and control mice either an HFD or the NCD (Supplementary Fig. 17). Treatment was started at 40 days of age. The HFD reduced the changes in gene expression detected in SBMA muscle (Supplementary Fig. 18a, Tables 3–5), including Ppargc1a (PGC1α), genes involved in lipid metabolism, and TfEB (Fig. 5a), it reduced mitochondrial membrane depolarization by 50 % without altering the activity of mitochondrial complexes (Fig. 4d, e), it increased LC3 lipidation (Fig. 4g), and it restored the activation of mTOR signaling to normal levels (Fig. 5b). The HFD did not affect the levels of expression of polyglutamine-expanded AR (Supplementary Fig. 18b). The HFD increased the wet weight of quadriceps (Fig. 5c) but not gastrocnemius (Supplementary Fig. 19a), and reduced the number of slow-oxidative fibers by 20 % in both quadriceps and gastrocnemius muscles of SBMA mice (Fig. 5d; Supplementary Fig. 19b, c), it increased the mean CSA of oxidative fibers in the quadriceps and gastrocnemius by 11 and 7 %, respectively, whereas it did not affect the CSA of glycolytic fibers (Supplementary Fig. 20a, b). In CTR mice, the HFD caused glucose intolerance and insulin resistance (Supplementary Fig. 8b, c), signs of metabolic syndrome that normal mice develop upon exposure to HFD [35]. However, the HFD did not elicit signs of metabolic syndrome in AR113Q mice. The HFD increased fat deposition by 2.9- and 3.9-fold in AR113Q and CTR mice, respectively (Fig. 5e). Control mice fed the HFD exhibited a progressive increase in body weight (P = 0.00001) from 70 days of age compared to control mice fed the NCD (Fig. 5f; Supplementary Fig. 21a, b). AR113Q mice also showed a significant (P = 0.00001) increase in body weight starting from 84 days of age and were indistinguishable from control mice fed the NCD. The body weight of control and AR113Q mice fed the HFD versus NCD was increased by 1.84- and 1.49-fold, respectively, indicating that AR113Q are partially resistant to diet-induced obesity, despite similar food intake and locomotor activity (Supplementary Fig. 21c, d). AR113Q mice showed reduced performance on an accelerating rotarod (Supplementary Fig. 21e). The performance of control mice fed the HFD was significantly decreased, as expected, whereas that of AR113Q mice was restored to normal. The HFD also increased the force of gastrocnemius (Supplementary Fig. 21f). Importantly, the HFD extended the median life span of AR113Q mice from 161 days to 322 days (χ2LR = 3.955, P = 0.047; Gehan–Breslow: P = 0.046) (Fig. 5g). These results show that the HFD attenuates disease manifestations in knock-in SBMA mice.

We next assessed whether metabolic alterations also occur in the muscle of SBMA patients. The number of NADH-positive fibers was increased by 1.7-fold in the muscle of SBMA patients compared to normal subjects (Fig. 6a). The expression of the key glycolytic genes, HkII, Pfkfb3, and Gapdh, was significantly decreased in the glycolytic muscles of SBMA patients, suggesting that glycolysis is impaired in the muscle of SBMA patients (Fig. 6b). Finally, the phosphorylation of the mTOR downstream target S6K1 was augmented in the muscle of SBMA patients compared to control subjects, indicating that mTOR signaling is enhanced in SBMA patients (Fig. 6c). These results show metabolic alterations in the muscle of SBMA patients, thereby supporting the biological relevance of our findings in pathogenesis.