Transcription Factor EB Controls Metabolic Flexibility during Exercise

TFEB Regulates Mitochondrial Biogenesis in Muscle

To examine potential effects of TFEB in mitochondrial function, we analyzed mitochondrial morphology in muscles overexpressing or lacking TFEB. Electron microscopy (EM) analyses showed a striking increase of mitochondrial density and volume in TFEB-overexpressing muscles (Figures 1B, 1C, 1E, and 1F). Interestingly, mitochondrial density and size were normal in the TFEB KO muscles (Figures 1E and 1F). Consistent with the EM data, increase of mitochondrial DNA (mtDNA) was found in TFEB transgenic muscles, while no differences were observed in TFEB KO muscles (Figure 1G). However, while the cristae shape, matrix density, and outer membrane morphology were normal in TFEB-overexpressing muscles (Figure 1C), abnormalities were found in approximately 10% of the mitochondria from TFEB KO muscles (Figures 1D and 1H). An increase in the number of mitochondria was also observed in C2C12 muscle cells transfected with TFEB-GFP, as detected by immunofluorescence confocal and confirmed by EM analyses (Figures S2A and S2B). Quantitative real-time PCR also revealed an increase of mtDNA content in TFEB-overexpressing cells (Figure S2C).

Importantly, quantitative real-time PCR analysis revealed that TFEB overexpression in muscle and in C2C12 cells induces the expression of many genes involved in mitochondrial biogenesis and function, including the master gene of mitochondrial biogenesis, PGC1α, a known direct target of TFEB (Settembre et al., 2013) (Figures 2A and S2D). Moreover, another PGC-1 family member, PGC1β, was also upregulated by TFEB overexpression. Consistently, we found a significant induction of peroxisome proliferator-activated receptor α (PPARα), PPARβ/δ, and PPARγ in TFEB-overexpressing muscles. However, TFEB deletion did not affect the expression of PGC1α/β and PPAR genes, with the exception of PPARα, which was downregulated. In order to elucidate the possible mechanisms underlying the induction of mitochondrial biogenesis observed in TFEB-transfected muscles, we examined the expression of nuclear respiratory factors 1 and 2 (NRF1and NRF2). The mRNA levels of NRF2 were increased, as well as NRF downstream genes, including mitochondrial transcription factor A (TFAM). Chromatin immunoprecipitation (ChIP) experiments confirmed the direct recruitment of TFEB on NRF1 and NRF2 promoters (Figure 2B), but not on TFAM promoter (data not shown). Finally, overexpression of TFEB in skeletal muscle increased the expression of mitochondrial enzymes. Subunits of the four respiratory chain complexes and the ATP synthase, as well as genes encoding electron transport and tricarboxylic acid cycle proteins, were induced by TFEB overexpression and were reduced by TFEB deletion (Figure 2A). Importantly, immunoblotting analyses confirmed the increase of complex I (NDUFA9), complex II (SDHA), and complex IV (COX5a) proteins in TFEB-transfected muscles (Figures 2C and 2D). TFEB deletion did not alter NDUFA9 and COX5a expression but significantly reduced the level of SDHA protein (Figures 2C and 2D). To better characterize the involvement of TFEB in mitochondrial respiration, we analyzed the specific activities of enzymes involved in oxidative phosphorylation. Biochemical analysis of muscle samples infected with AAV2.1-TFEB compared to wild-type (WT) muscles showed increase of citrate synthase, mitochondrial respiratory chain complex I (CI), CII, CIII, and CIV activities (Figure 2E). Consistent with the western blot analyses, TFEB deletion led to decrease of CII activity, the complex that contains the SDHA flavoprotein, while the other respiratory complexes had normal activities (Figure 2E). The changes in respiratory chain activities were corroborated by histochemical analyses for COX and SDH activity in AAV2.1-TFEB-transfected and TFEB KO muscles. Indeed, only SDH activity was greatly reduced in the absence of TFEB, while both COX and SDH were increased in TFEB-overexpressing muscles (Figure 2F). To further investigate the role of TFEB in mitochondrial function, we generated an inducible muscle-specific transgenic mouse line. Acute activation of TFEB by tamoxifen treatment in adult mice recapitulated the phenotype of AAV2.1-TFEB overexpression on mitochondria biogenesis (Figure S3). Consistent with the increase of SDH and respiratory chain complex activity, mitochondrial respiration was significantly enhanced by TFEB expression in adult transgenic muscles (Figures S3A and S3B).

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Figure 2

TFEB Regulates Mitochondrial Function

(A) Quantitative real-time PCR of genes related to mitochondrial biogenesis and function in muscles of WT (blue bar), TFEB-overexpressing (red bar), and TFEB KO (green bar) mice. Data were normalized for GAPDH and expressed as fold induction relative to the WT. Data are expressed as mean ± SE, n = 3; ^∗p < 0.05, ^∗∗p < 0.01.

(B) ChIP analysis in muscles from TFEB-overexpressing and WT mice. The CLEAR elements of NRF1 and NRF2 promoters are depicted as red boxes. TSS (transcriptional start site) indicates the first codon. The histograms show the amount of immunoprecipitated DNA as detected by quantitative real-time PCR assay. Data represent mean ± SE of three independent experiments; ^∗p < 0.05, ^∗∗p < 0.01, ^∗∗∗p < 0.001.

(C) Western blots of mitochondrial respiratory chain proteins. Blot is representative of four independent experiments.

(D) Densitometric quantification of blots shown in (B). Data are shown as means ± SE (n = 4); ^∗p < 0.05.

(E) Mitochondrial respiratory chain activity in muscles of WT, AAV2.1-TFEB transfected, and TFEB KO mice. Data are expressed as a percentage relative to the WT. Data are expressed as mean ± SE, n = 4; ^∗p < 0.05, ^∗∗p < 0.01.

(F) H&E, succinate dehydrogenase (SDH), and cytochrome oxidase (COX) staining of AAV2.1-GFP, AAV2.1-TFEB transfected, and muscles from TFEB KO mice. The scale bars represent 100 μm. Magnification, 20×.

(G) ATP production rate per gram of tissue from TFEB-overexpressing and TFEB KO mice compared to controls. Data are shown as means ± SE (n = 5); ^∗∗p < 0.01, ^∗∗∗p < 0.001.

(H) Mitochondrial membrane potential of myofibers isolated from FDB muscles of WT and TFEB KO mice. Where indicated, 6 μM oligomycin or 4 μM FCCP were added. Each trace represents the tetramethylrhodamine methyl ester (TMRM) fluorescence of a single fiber. The fraction of myofibers with depolarizing mitochondria is indicated for each condition.

(I) Densitometric quantification of the carbonylated proteins extracted from WT and TFEB KO mice. Values are mean ± SEM (n = 4; ^∗p < 0.05). A representative immunoblot for carbonylated proteins is shown in Figure S3C.

We next assessed whether these TFEB-mediated changes of mitochondrial morphology and function had any impact on energy production. ATP levels were higher in TFEB-transfected muscles and lower in TFEB-deficient muscles compared to controls (Figure 2G). To understand the mechanisms underlying the significant decrease of ATP in TFEB KO muscles, we checked mitochondrial function in these animals. Fluorescent dyes, like tetramethylrhodamine methyl ester (TMRM), monitor the mitochondrial membrane potential (Δψm), the critical parameter that drives ATP production. Therefore, we checked the status of Δψm in isolated adult fibers of WT and TFEB KO muscles. As expected, in control mice oligomycin-dependent inhibition of ATP synthase did not alter Δψm (Figure 2H), and mitochondrial depolarization was achieved after membrane permeabilization by the protonophore carbonylcyanide-p-trifluoromethoxy phenylhydrazone (FCCP). Conversely, mitochondria of TFEB null fibers underwent a significant depolarization after oligomycin treatment (Figure 2G), suggesting that these fibers were at least in part relying on reverse activity of ATP synthase to preserve their membrane potential as a consequence of proton membrane leak. In addition, oxidative stress, revealed by protein carbonylation, was significantly higher in TFEB KO mice than controls (Figures 2I and S3C).

TFEB Regulates Mitochondrial Biogenesis in Skeletal Muscle through a PGC1α- and PGC1β-Independent Mechanism

Both PGC1α and PGC1β are master regulators of mitochondrial biogenesis and oxidative metabolism. However, recent findings suggest the presence of an independent pathway that regulates mitochondrial biogenesis during exercise (Rowe et al., 2012). Therefore, we examined whether TFEB is the missing sensor of physical activity that coordinates the metabolic responses independently of PGC1α. First, we checked expression and localization of endogenous TFEB in PGC1α KO mice before and after exercise. TFEB was expressed at lower levels and more cytosolic in PGC1α KO mice when compared to controls (Figures 3A and 3B). Importantly, exercise restored a normal TFEB expression, induced TFEB nuclear translocation, and triggered upregulation of genes related to mitochondrial biogenesis (Figures 3C and S4A).

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Figure 3

Exercise Induces TFEB Expression and Nuclear Localization in PGC1α^−/− Muscle

(A) TFEB mRNA levels of PGC1α^−/− muscles infected with AAV2.9 control virus (light gray) or AAV2.9-TFEB (dark gray). Data were compared with TFEB mRNA level of WT mice (white). The levels of TFEB were measured in sedentary condition and post-exercise as indicated. Error bars represent mean ± SE for n = 3; ^∗p < 0.05, ^∗∗∗p < 0.001.

(B) TFEB immunohistochemical analysis of PGC1α^−/− muscles from sedentary and exercised mice. GCN muscle cryosections were immunostained by using anti-TFEB antibody. Control means endogenous TFEB in muscles infected with AAV2.9 control virus. TFEB means the transgene TFEB in muscles infected with AAV2.9-TFEB. Arrows indicate exercise-induced TFEB nuclear localization. The scale bars represent 50 μm.

(C) Analysis of genes related to mitochondrial biogenesis in PGC1α^−/− skeletal muscle infected with AAV2.9 control or with AAV2.9-TFEB virus. The mRNA levels were measured before and after acute exercise, as indicated. Data are shown as mean ± SE, n = 3; ^∗p < 0.05, ^∗∗p < 0.01, ^∗∗∗p < 0.001.

(D and E) Muscle fatigue and physical performance are improved by TFEB expression in PGC1α-deficient muscle.

(D) The mice were systemically injected with AAV2.9-TFEB or AAV2.9 control virus. After 3 weeks from the virus injection, mice were subjected to run until exhaustion, as described in the Experimental Procedures. Data are shown as mean ± SE, n = 6; ^∗p < 0.05.

(E) Soleus muscles were transected by intramuscular injection of AAV2.1-TFEB or AAV2.1 control virus, which resulted in 100% transfection efficacy. Index for fatigue means tetanic muscle force generated after 100 s of stimulation divided by the maximal tetanic tension produced during the fatigue protocol. Data are shown as mean ± SE, n = 4; ^∗p < 0.05, ^∗∗∗p < 0.001.

EM analysis of overexpressed TFEB in PGC1α KO mice revealed increased mitochondrial volume and density (Figure S4B). TFEB overexpression in PGC1α KO muscle also resulted in increased COX and SDH activity along with increased activities of CI, II, III, and IV (Figures S4C and S4D). The levels of PGC1α targets such as TFAM, NRF1, and NRF2 were also significantly upregulated by TFEB overexpression, even in the absence of PGC1α (Figure S5A). Importantly, systemic delivery of TFEB in muscle-specific PGC1α KO mice improved their exercise tolerance (Figure 3D). Indeed, TFEB expression was able to restore normal fatigue index when expressed in PGC1α KO muscle (Figure 3E). Altogether, these findings suggest that the induction of mitochondrial biogenesis in TFEB-overexpressing muscles does not depend on the presence of PGC1α.

To determine whether PGC1β may compensate for the lack of PGC1α, we measured the effect of TFEB overexpression on mitochondrial biogenesis in cells that were silenced for PGC1α and PGC1β. Importantly, inhibition of both PGC1 factors did not prevent or reduce TFEB-mediated induction of genes related to mitochondrial biogenesis and mitochondrial respiratory chain activity (Figure S5B).

TFEB Controls Energy Balance in Skeletal Muscle during Exercise

Physical activity has a major impact on glucose homeostasis and mitochondrial biogenesis and function. Therefore, we checked whether acute exhausting versus mild and chronic exercise regiments are equally able to activate TFEB. As a readout of TFEB activation, we monitored its nuclear localization. While acute exhausting contraction led to nuclear translocation of TFEB (Figure 4A), mild exercise did not (Figure 4B). However, 7 weeks of training with progressive increase of intensity without reaching exhaustion induced a massive TFEB nuclear translocation with concomitant cytosolic depletion (Figure 4B). Therefore, intensity and duration of training are critical factors that affect TFEB nuclear translocation and transcriptional regulation of genes related to mitochondrial biogenesis and function (Figure 4C).

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Figure 4

Exercise Induces TFEB Nuclear Translocation

(A) GCN tissue sections of TFEB KO and WT mice were immunostained with anti-TFEB antibody and counter-stained with hematotoxylin as indicated. Arrows indicate nuclear localization of endogenous TFEB in WT mice after an acute bout of exercise. The scale bars represent 50 μm.

(B) Endogenous TFEB was immunostained in muscle cryosections of sedentary, 4 days mild exercised, and 7 weeks intense trained WT mice. Arrows indicate TFEB nuclear localization.

(C) Expression analysis of genes related to mitochondrial biogenesis and mitochondrial respiratory chain in WT skeletal muscle before and after acute exercise. Data are shown as mean ± SE, n = 3; ^∗p < 0.05, ^∗∗p < 0.01.

To determine the physiological consequences of TFEB translocation during physical activity, we examined exercise performance in both TFEB KO and inducible muscle-specific TFEB transgenic mice. High-intensity exercise revealed significant training intolerance of TFEB KO mice compared to controls (Figure 5A). Conversely, acute muscle-specific activation of TFEB enhanced physical performance (Figure 5A). To better understand the exercise intolerance of the TFEB KO mice, we examined energy expenditure during treadmill running. While WT mice maintained constant levels of energy expenditure during physical activity, the TFEB KO mice displayed a drop after 15 min of physical exercise (Figure 5B). Metabolic analyses revealed that in basal condition, TFEB KO mice have a higher respiratory exchange rate (RER) than controls (Figure 5C). These data suggest that TFEB KO mice depend on glucose oxidation more than controls. In addition, while WT mice maintained a relatively constant RER during running period, TFEB KO mice showed a drop in RER after 20 min (Figure 5C). This decrease indicates a shift in substrate usage from glucose to fat metabolism. Finally, we measured glucose and fatty acid levels in muscle and blood from TFEB KO and TFEB transgenic mice before and after exercise. TFEB KO mice showed lower blood glucose levels compared to WT mice in basal condition (Figure 5D). Exercise caused a 50% reduction of blood glucose in TFEB KO, TFEB transgenic, and control mice (Figure 5D). Insulin levels mirrored the changes of blood glucose, as they were reduced in basal condition in TFEB KO mice and dropped after exercise in the different genotypes (Figure 5E).

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Figure 5

TFEB Controls Energy Balance in Skeletal Muscle

(A) High-intensity exhaustive exercise. To determine exercise capacity, mice were run on a treadmill. During high-intensity exercise, transgenic mice (red) ran more, while TFEB KO mice (gray) ran half as much as WT mice (white). Data are shown as mean ± SE, n = 10; ^∗∗p < 0.01.

(B and C) Energy expenditure (B) and RER (C) were determined during exercise in TFEB KO and WT mice. The mice ran at a fixed speed of 10 m/min and an incline of 20°. The figure shows the mean RER measured at peak oxygen consumption. Data were transformed by Blom’s method to obtain both normally distributed data and normally distributed residual. Two-way ANOVA was used for comparison of RERs in the two groups during the entire resting period, whereas t test was used for comparison of individual time points, n = 8.

(D–F) Blood and muscle metabolites before and after high-intensity exercise in WT (white), TFEB transgenic (red), and TFEB KO mice (gray). Data are shown as mean ± SE, n = 8; ^∗p < 0.05, ^∗∗p < 0.01, ^∗∗∗p < 0.001.

(G) Enzymatic quantification of muscle glycogen before and after high-intensity exercise in WT (white), TFEB transgenic (red), and TFEB KO mice (gray). Data are shown as mean ± SE, n = 8; ^∗∗p < 0.01.

(H) Periodic acid-Schiff (PAS) staining of cryosections from AAV2.1-GFP, AAV2.1-TFEB transfected, and TFEB KO mice. Inserts show PAS staining after glycogen breakdown. The scale bars represent 100 μm.

(I and J) Quantitative analysis of serum (I) and muscle (J) NEFA before and after high-intensity exercise in TFEB KO (gray) and WT mice (white). Data are shown as mean ± SE, n = 8; ^∗p < 0.05, ^∗∗p < 0.01.

(K) Enzymatic quantification of B-hydroxybutyrate before and after high-intensity exercise in TFEB KO (gray box) and WT mice (white box). Data are shown as mean ± SE, n = 8; ^∗p < 0.05.

(L) TFEB ablation results in a decreased rate of insulin-stimulated GIR. EU clamps were used to assess whole-body insulin sensitivity by determining the GIR required to maintain euglycemia in WT (white) and TFEB KO (gray) mice. Data represent the means ± SE from five to six individual mice per group; ^∗p < 0.05.

(M) TFEB deletion results in a reduction of insulin-stimulated glucose uptake in muscle. Insulin-stimulated glucose uptake into muscle tissues was determined by 2-deoxy-d-[1-¹⁴C]glucose injection during the last 35 min of insulin infusion during the EU clamp. These data represent the means ± SE of five to six mice per group; ^∗p < 0.05. WT, white; TFEB KO, gray.

(N) The amount of glucose conversion to glycogen in GCN muscles from WT (white) and TFEB KO (gray) mice was determined during the EU clamp by infusion of [3-³H]glucose. Data represent the means ± SE from five to six mice per group; ^∗p < 0.05.

(O and P) TFEB KO mice do not show any significant difference in glucose homeostasis of liver or adipose tissue. (O) Glucose uptake into adipose tissue (EPI) and (P) insulin suppression of hepatic glucose output (HGP, liver) were determined during the EU clamp by infusion of 2-deoxy-d-[1-¹⁴C]glucose or [3-³H]glucose infusion into WT mice (white) and TFEB KO (gray) mice. These data represent the means ± SE from five to six mice per group.

TFEB KO mice are hypoglycemic, contain dysfunctional mitochondria, and produce less ATP. Thus, we reasoned that they use anaerobic glycolysis to produce energy. Consistent with this hypothesis, we found higher levels of lactate in the blood of TFEB KO mice before and after exercise compared to controls (Figure 5F). Conversely, lactate of transgenic mice was already lower than controls in resting condition and did not increase after exercise (Figure 5F). Therefore, TFEB transgenic mice better utilize glucose for energy production. To further confirm this finding, we measured glycogen levels in muscle. Glycogen levels were remarkably lower in TFEB KO mice and higher in TFEB transgenic in basal condition compared to WT. Enzymatic quantification showed that glycogen content was 3-fold less after TFEB ablation (Figure 5G), while it was 10-fold higher after TFEB overexpression compared to controls (Figure 5G). This was confirmed by periodic acid-Schiff (PAS) staining (Figure 5H). Exercise led to glycogen consumption in both TFEB KO muscles and controls (Figure 5G). The lower glycogen content detected in sedentary TFEB KO muscles explains the decrease of RER observed after a 15 min exercise (Figure 5C). Since glycogen is rapidly depleted in TFEB KO mice, the additional need for energy during exhausting exercise requires a switch from glycolysis to fatty acid oxidation. Consistently, while blood free fatty acid concentrations were reduced after exercise in both TFEB KO mice and controls (Figure 5I) and blood levels of non-esterified fatty acids (NEFAs) did not differ between genotypes (Figure 5I), their muscle content dramatically decreased after exercise only in TFEB KO mice (Figure 5J). Importantly, ketones did not differ between TFEB KO and controls (Figure 5K). These findings confirm a change in metabolic flexibility in the absence of TFEB that forced muscle cells to use lipids for ATP production. The exhaustion of the lipid fuel in KO mice results in inability to maintain the same exercise intensity of controls.

TFEB Controls Metabolic Flexibility and Energy Balance Independently of Autophagy

We and others have found that autophagy is important for mitochondrial quality control and is activated by exercise to clear dysfunctional mitochondria (Lo Verso et al., 2014). Thus, we checked whether TFEB controls autophagy in adult skeletal muscles. Surprisingly, TFEB activation was not sufficient to enhance autophagy flux and TFEB deletion did not impair autophagy flux in the presence or absence of nutrients (Figures S6A and S6B). Moreover, activation of TFEB did not induce protein breakdown and muscle loss. In fact, most of the atrophy-related genes belonging to the ubiquitin proteasome and autophagy-lysosome systems were not induced by TFEB expression (Figure S6C). Similarly, mitophagy genes were not upregulated. Since protein degradation is not affected by TFEB activation, we checked whether genes related to protein synthesis were modulated by TFEB. However, when we checked a cross-sectional area, we found a shift toward smaller size in transgenic mice, suggesting that protein synthesis was not induced. This decrease in fiber size is due to a metabolic shift because oxidative fibers are smaller than glycolytic skeletal muscle fibers (Figure S7A). Furthermore, we did not find any significant difference in myosin distribution between transgenic and control muscles (Figure S7B). Therefore, TFEB controls myofiber metabolism, but not myosin content/type, independently of autophagy or proteostasis.

TFEB Controls Glucose Homeostasis and Insulin Sensitivity Independently of PGC1α

Because muscles from TFEB KO mice contain lower glycogen levels than controls, we reasoned that they may have abnormal regulation of glucose homeostasis. Thus, we performed euglycemic-hyperinsulinemic (EU) clamps and observed that the glucose infusion rate (GIR) that is required to maintain a constant glycemia during insulin treatment was significantly reduced in TFEB KO compared to control mice (Figure 5L). The reduction of GIR was consequent to a decrease in skeletal muscle glucose uptake (Figure 5M), which caused a decrease of glycogen synthesis (Figure 5N). Although there was an apparent small decrease in adipose tissue glucose uptake, this was not statistically significant (Figure 5O). Moreover, insulin was equally effective in suppressing hepatic glucose output during the clamp experiment (Figure 5P). Together, these data demonstrate that TFEB deficiency in skeletal muscle results in peripheral insulin resistance-reduced glucose uptake and decreased glycogen content.

These findings are consistent with the transcriptomic signature of TFEB-overexpressing and TFEB KO muscles. To further confirm these findings, we monitored the expression levels of glucose homeostasis-related genes in muscles injected with AAV-TFEB and in muscles of TFEB transgenic mice and found a significant increase in the expression of GLUT1 and GLUT4, the GTPase involved in GLUT4 translocation (TBC1D1), and the rate-limiting enzymes of glycolysis (hexokinase 1 and 2) (Figure 6A). Immunoblotting analyses confirmed increased protein levels of GLUT1 in TFEB-overexpressing muscles (Figure 6B). The induction of genes involved in glucose uptake was also coupled with an increase of transcript and protein of GYS (Figures 6C and 6D).

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Figure 6

TFEB Controls Genes Related to Glucose Metabolism

(A and C) Quantitative real-time PCR of glucose uptake and glycogen biosynthesis-related genes in GCN muscles infected with AAV2.1-TFEB (black bar) and in TFEB transgenic muscles (red bar) compared with WT (white bar) muscles. Data were normalized for GAPDH and expressed as fold induction relative to the WT. Data are shown as mean ± SE, n = 3; ^∗p < 0.05, ^∗∗p < 0.01.

(B) Western blot of GLUT1 and densitometric quantification. Values are normalized for GAPDH; ^∗p < 0.05.

(D) Western blot analysis of total and phosphorylated GYS from extracts of GCN. Representative blot images are shown (left panel). Densitometric quantification is depicted on the right panel. Data are shown as mean ± SE, n = 3; ^∗p < 0.05.

(E–G) TFEB regulates glycogen synthesis independently of PGC1α.

(E) Quantitative real-time PCR of genes related to glucose uptake and glycogen biosynthesis in PGC1α^−/− and WT mice that were infected with AAV2.1-GFP (white) or AAV2.1-TFEB (black). Data are shown as mean ± SE, n = 3; ^∗p < 0.05, ^∗∗p < 0.01.

(F) PAS staining of cryosections from PGC1α^−/− muscles that were transfected by AAV2.1-GFP or AAV2.1-TFEB. The scale bars represent 100 μm.

(G) Enzymatic quantification of basal glycogen levels in PGC1α^−/− and WT muscle, infected with AAV2.1-GFP (white) or AAV2.1-TFEB (black).

GYS activity is negatively regulated by phosphorylation of the C-terminal region by GYS kinases (GSK3s) (Jensen and Lai, 2009). However, the phospho-GYS levels were unchanged in AAV-TFEB muscles, and therefore, the pGYS/GYS ratio was dramatically decreased in TFEB-overexpressing muscles when compared to controls (Figure 6D). Furthermore, expression analysis of genes related to glucose metabolism did not reveal any significant difference between PGC1α KO and controls after TFEB overexpression (Figure 6E). Finally, EM showed an accumulation of glycogen in PGC1α KO mice that were infected by AAV2.1-TFEB (Figure S4B). Quantitative and qualitative analyses of glycogen showed that TFEB induced a higher glycogen accumulation in PGC1α KO muscle than in WT (Figures 6F and 6G). These data indicate that TFEB is able to control muscle glycogen content in a PGC1α-independent manner.

Real-Time PCR

Quantitative real-time PCR analyses were performed on a LigthCycler 480 II (Roche). For detailed information on preparation and gene expression analysis, see the Supplemental Experimental Procedures.

Generation of Muscle-Specific TFEB KO and Inducible Muscle-Specific Transgenic Mice and In Vivo Experimental Procedures

Detailed information in the Supplemental Experimental Procedures.

Western Blot Analysis and Antibodies

Total homogenates were prepared in RIPA buffer (50 mM Tris HCl [pH 8], 150 mM NaCl, 1% NP-40, 0.5% sodium deoxycholate, and 0.1% SDS) with the addition of a protease inhibitor cocktail (Roche). Protein concentration was determined by the Lowry method. Aliquots, 50 μg each, were run through an SDS-PAGE and electroblotted onto a PVDF membrane, which was then matched with different antibodies. For the antibodies used in western blot analysis, see the Supplemental Experimental Procedures.

Biochemical Analysis of Mitochondrial Respiratory Chain Complex

Muscle samples stored in liquid nitrogen were homogenized in 10 mM phosphate buffer (pH 7.4), and the spectrophotometric activity of cI, cII, cIII, and cIV, as well as citrate synthase (CS), was measured as described (Bugiani et al., 2004). Detailed information in the Supplemental Experimental Procedures.

Morphological Analysis

Histochemical and ultrastructural analyses were performed as described (Sciacco and Bonilla, 1996). Detailed information in the Supplemental Experimental Procedures.

Isolation of Skeletal Myofibers and Measurements of Mitochondrial Membrane Potential

Muscle fibers were isolated from FDB muscle and mitochondrial membrane potential measured by epifluorescence microscopy on the basis of the accumulation of TMRM fluorescence, as previously described (Lo Verso et al., 2014). Fibers were considered as depolarizing when they lost more than 10% of the initial value of TMRM fluorescence. Imaging was performed with a Zeiss Axiovert 100 TV inverted microscope equipped with a 12-bit digital cooled charge-coupled device camera (Micromax, Princeton Instruments). The results were analyzed with MetaFluor imaging software (Universal Imaging).

Acute Exercise, Training, and Fatigue Experiments

For acute and chronic exercise studies, 16-week-old mice performed concentric exercise on a treadmill (Biological Instruments, LE 8710 Panlab Technology 2B) with a 10° incline, according to the protocol of exercise previously described (Medina et al., 2015). Total running distance was recorded for each mouse. Detailed information in the Supplemental Experimental Procedures.

Immunohistochemistry

Detailed information in the Supplemental Experimental Procedures.

Electron Microscopy

Detailed information in the Supplemental Experimental Procedures.

Plasma Chemistry Analysis

Blood was collected from the orbital plexus under isoflurane (Vedco) anesthesia. Plasma was frozen in aliquots at −20°C or used immediately after collection. Specific enzymatic kits were used for determination of serum NEFAs (Wako) and lactate (Abcam). Plasma glucose was monitored by a glucometer. Insulin was measured by ELISA (Mercodia).

Tissue Metabolite Quantification

Detailed information in the Supplemental Experimental Procedures.

Whole-Body Indirect Calorimetry

Metabolic measurements were performed using an Oxymax indirect calorimetry system (Columbus Instruments) (Zong et al., 2011). Detailed information in the Supplemental Experimental Procedures.

In Vivo Assessment of Insulin Action and Glucose Metabolism

Four days before the experiment, the mice were anesthetized and an indwelling catheter was introduced into the left internal jugular vein. The mice were fully recovered from the surgery before the in vivo experiments, as reflected by their reaching preoperative weight. After an overnight fast, EU clamps were conducted in conscious mice as previously described (Ayala et al., 2006, Zong et al., 2012). Detailed information in the Supplemental Experimental Procedures.

ChIP Assays

We performed ChIP assays on adult skeletal muscle overexpressing TFEB3XFlag, and on WT muscle as control, by using the ChIP assay kit (Upstate) according to Milan et al. (2015). For immunoprecipitation, we used anti-FLAG antibody (F7425, Sigma-Aldrich). Oligonucleotide primers for amplification of a TFEB binding site on the GLUT1, GLUT4, nNOS, NRF1, NRF2, and TFAM promoters are listed in Table S7.

Microarray Data Analysis

Detailed information in the Supplemental Experimental Procedures.

We thank N. Brunetti, D. Medina, and C. Settembre for suggestions. We are grateful to E. Nusco and TIGEM Bioinformatic and HCS Facilities for technical support, in particular D. Carrella and S. Montefusco. We acknowledge support from the Italian Telethon Foundation to A.B. (TCR09003) and M.S. (TCP04009), the European Research Council (ERC) to A.B. (250154-CLEAR) and M.S. (282310-MyoPHAGY), the Italian Ministry of Education (MiUR) (PRIN 2010/2011) to M.S., CARIPARO and Foundation Leducq to M.S., the Pennsylvania University and Beyond Batten Disease Foundation to A.B., the Core Grant MRC-MBU and ERC FP7-322424 to M.Z., GR-2010-2306-756 and the Italian Ministry of Health to C.V., and the National Science Foundation of China (H0315: 81370531) to H.Z.