Abstract

Background: Glycolysis controls mTORC1 signaling and protein synthesis. In skeletal muscle, glucose metabolism increases with both exercise/contraction intensity and volume, and therefore, high-intensity muscle contraction (HiMC) such as resistance exercise facilitates glycolysis including glucose uptake and glycogen breakdown. However, it is unknown whether glycolysis regulates HiMC-induced mTORC1 activation and increase in protein synthesis.

Methods: To determine whether glycolysis regulates basal and HiMC-induced mTORC1 signaling and protein synthesis, we employed 2-deoxyglucose (2-DG) to inhibit glycolysis and isometrically contracted the gastrocnemius muscle of Sprague Dawley rats using percutaneous electrical stimulation.

Results: Inhibition of glycolysis by 2-DG inhibited basal phosphorylation of p70S6K and 4E-BP1 (downstream targets of mTORC1) and protein synthesis (all P < 0.05) independent of AMPK phosphorylation. AMPK phosphorylation was comparably increased after HiMC at 0 h post HiMC and returned to basal levels 6 h post HiMC in both vehicle- and 2-DG-treated groups. Glycolysis inhibition attenuated muscle contraction-induced phosphorylation of 4E-BP1 at 6 h post HiMC (P < 0.05) but not p70S6K phosphorylation and protein synthesis.

Conclusion: Although glycolysis is involved in basal but not HiMC-induced muscle protein synthesis, it regulates both basal and HiMC-induced mTORC1 signaling, and may play key roles in skeletal muscle adaptation to HiMC.

Keywords: Glycolytic metabolism, glucose, exercise, metabolomics

1. Introduction

Skeletal muscles comprise 4050% of the total body weight and are important not only for body movement but also for energy metabolism. In fact, skeletal muscle is responsible for approximately 80% of insulin-stimulated glucose uptake in the postprandial state [1, 2]. Thus, the maintenance of skeletal muscle mass is essential for an active and healthy life and perturbations in this homeostasis increase the risk of lifestyle-related diseases.

Skeletal muscle mass is mainly regulated by the balance between muscle protein synthesis and degradation, where the muscle protein balance is negative during atrophy and positive during hypertrophy. Compared with muscle protein degradation, protein synthesis is more sensitive and dynamically responds to physiological stimuli,including dietary intake and muscle contraction [3, 4], and is regulated by mechanistic target of rapamycin (mTOR) signaling [5].Muscle contraction, especially high-intensity muscle contraction (HiMC) such as during resistance exercise, is well known as a physiological stimulus for increasing mTOR, especially mTOR complex 1 (mTORC1) signaling and muscle protein synthesis [6-8]. However, although the mechanism underlying HiMC-induced muscle anabolism has been gradually elucidated, the mechanism upstream of HiMC-induced mTORC1 activation is still not completely understood [9- 13].

Muscle contraction and exercise intensity (e.g., % 1 repetition maximum (1RM) and % maximum voluntary contraction) are important variables for determining muscle anabolic response to resistance exercise [14] ; force-time integral/exercise volume (e.g., number of sets) is also a determining factor [15- 18]. Therefore, factors other than mechanical stress are involved in the regulation of HiMC-induced mTORC1 activation. Generally, insulin-mediated PI3K/Akt signaling activation and leucine-induced mTORC1 translocation to the lysosome are the major molecular mechanisms underlying mTORC1 activation [19, 20]. Although HiMC sensitizes skeletal muscle to both insulin and leucine during the recovery period [21, 22], which potentially facilitates mTORC1 activation, previous studies have reported that HiMC-induced mTORC1 activation occurs independently of PI3K/Akt signaling activation and mTORC1 translocation to the lysosome [8, 23-25], indicating that HiMC can facilitate mTORC1 signaling independently of insulin and leucine signaling.

Recent studies have indicated that glucose metabolism, specifically glycolysis, controls mTORC1 signaling independently of insulin signaling [26, 27]. Glucose metabolism increases with both exercise/contraction intensity and volume [28, 29], and therefore, resistance exercise facilitates glycolysis including glucose uptake and glycogen breakdown [23, 30, 31].Furthermore, a recent study showed that an increase in muscle glycogen levels after pharmacological inhibition of glycogen phosphorylase facilitated HiMC-induced mTORC1 activation [32]. Thus, increased glucose utilization during resistance exercise may contribute to mTORC1 activation and promote protein synthesis. 2-Deoxyglucose (2-DG), a glucose analog, is incorporated by tissues in a similar manner to that of glucose. However, although incorporated 2-DG is converted by hexokinase (HK) to its phosphorylated form, it is not further metabolized, and accumulated
phosphorylated 2-DG acts as a competitive inhibitor of HK, inhibiting glycolysis [33, 34]. Interestingly, 2-DG inhibits mTORC1 signaling independently of AMP-activated protein kinase (AMPK), akinase activated during an energy deficit [35], suggesting that 2-DG-mediated inhibition of glycolysis attenuates HiMC-induced mTORC1 activation and increase in protein synthesis. Therefore, we investigated the effect of 2-DG-mediated inhibition of glycolysis on mTORC1 signaling and protein synthesis following HiMC. We found that while glycolysis plays a role in basal but not HiMC-induced muscle protein synthesis, it regulates both basal and HiMC-induced mTORC1 signaling.

2. Materials and Methods
2.1. Animals

All experimental procedures in this study were approved by the Ethics Committee for Animal Experiments of the Nagoya Institute of Technology. Male Sprague Dawley rats (10 weeks old)purchased from Japan SLC (Shizuoka, Japan) were housed for one week at 22-24°C under a 12 h light/dark cycle and with ad libitum access to food and water.

2.2.Experimental procedures

After overnight fasting, the right gastrocnemius muscle was contracted isometrically via percutaneous electrical stimulation (100 Hz; one or five sets often 3-s contractions, 7-s rest between contractions, 3-min rest between sets) as described previously under isoflurane anesthesia [36]. The left gastrocnemius muscle acted as the control. The torque production was measured during electrical stimulation as previously described [37]. 2-DG [500 mg/kg in phosphate buffered saline (PBS)] or vehicle was randomly injected intraperitoneally 1 h before exercise. Muscle samples were obtained immediately and 6 h after HiMC. The samples obtained immediately after one set of HiMC were used only for measuring muscle lactate levels. Tissues were frozen rapidly in liquid nitrogen and stored at -80。C until use.

2.3. Measurement of glycogen level

Powdered frozen muscle samples (10 mg) were homogenized in 500 μL buffer containing 30% KOH saturated with Na2 SO4. Samples were boiled at 95°C for 30 min, followed by addition of 600 μL 95% ethanol. After centrifugation at 840 × g for 30 min at 4 °C, the supernatant was discarded, and the pellet was dissolved in 600 μL ultra-pure water (glycogen solution). Then, 100 μL glycogen solution was mixed with equal volume of 5% phenol, followed by addition of 500 μL concentrated H2 SO4. The absorbance was measured at 490 nm.

2.4. Measurement of lactate level

Powdered frozen muscle samples (20 mg) were homogenized with 0.6 N HClO4 and centrifuged at 3000 × g for 5 min at 20°C. Hydrazine/glycine buffer (0.4 M hydrazine, 0.5 M glycine, pH 9.0) was combined with 40 mM β -NAD and 0.03 mL lactate dehydrogenase (5 mg protein/mL). NADH formation was measured at 340 nm.

2.5. Metabolomic analysis

Muscle samples were homogenized in 15 volumes of 50% acetonitrile, followed by the addition of equal volumes of 50% acetonitrile. After centrifugation at 2,300 × g for 5 min at 4 °C, the supernatants were ultrafiltered using an ultrafiltration tube (Ultrafree-MC PLHCC; Human Metabolome Technologies, Tsuruoka, Japan) and the filtrates were dried. The dried residues were redissolved in 50 mL ultrapure water and used for capillary electrophoresis-mass spectrometry (CE-MS). The metabolites were analyzed using CE-time of flight (TOF) MS (Agilent CE-TOFMS system) and CE-QqQMS (Agilent CE and 6460 Triple Quad LC/MS systems; Agilent Technologies, Santa Clara, CA). Cationic and anionic metabolites were analyzed using a fused-silica capillary (i.d. 50 μm × 80 cm) with cation buffer solution (p/n: H3301- 1001; Human Metabolome Technologies) and anionic buffer solution (p/n: H3302- 1023; Human Metabolome Technologies), respectively, as the electrolyte. CE-TOFMS and CE-QqQMS data were analyzed using automatic integration software MasterHand ver. 2.17.1.11 (Keio University, Japan) and MassHunter (Agilent Technologies),respectively.

2.6. Western blotting

Powdered frozen muscle samples were homogenized in 10 volumes of buffer containing 20 mM Tris-HCl (pH 7.5), 1% NP40, 1% sodium deoxycholate, 1 mM EDTA, 1 mM EGTA, 150 mM NaCl, and Halt TM protease and phosphatase inhibitor cocktail (Thermo Fisher Scientific, Waltham, MA) . After centrifugation at 10,000 × g for 10 min at 4°C, the supernatant was collected and the protein concentration of each sample was determined using a rapid protein assay kit (Wako, Osaka, Japan). The samples were diluted in 3X sample buffer and boiled at 95°C for 5 min. Then, using 10% or 15% sodium dodecyl sulfate-polyacrylamide gradient gels, equal amounts of protein were separated using electrophoresis and subsequently transferred to ClearTrans® SP polyvinylidene difluoride membranes (Wako). After transfer, the membranes were washed in Tris-buffered saline containing 0.1% Tween-20 (TBST) and blocked with 1% skim milk in TBST for 1 h at room temperature. The membranes were washed and incubated overnight at 4°C with primary antibodies. Antibodies against phospho-Akt (Thr308, cat. #13038), phospho-Akt (Ser473, cat. #9271), total-Akt (cat. #9272), phospho-p70S6K (Thr389, cat. #9205), phospho-p70S6K (Thr421/Ser424, cat. #9204), total-p70S6K (cat. #2708), phospho-rpS6 (Ser240/244, cat. #2215), total-rpS6 (cat. #2217), phospho-PRAS40 (Ser246, cat. #2640), total-PRAS40 (cat. #2610),phospho-4E-BP1 (Thr37/46, cat. #9459), total-4E-BP1 (cat. #9452), LC3 (cat. #2775), ubiquitin (cat. #3933), phospho-AMPK (cat. #2535), total-AMPK (cat. #2532), and phosphor-Raptor (Ser792, cat. #2083) were obtained from Cell Signaling Technology (Danvers, MA). Anti-p62 antibody (cat. #PM045) was obtained from MBL (Aichi, Japan). The membranes were washed again in TBST and incubated for 1 h at room temperature with the appropriate secondary antibody. The immunoblots were developed using chemiluminescent reagents and imaged using the C-DiGit Blot Scanner (LI-COR Biosciences, Lincoln, NE). The membranes were then stained with Coomassie Blue to verify equal loading in all lanes. Protein levels were semi-quantified via densitometry using Image Studio (LI-COR Biosciences).

2.7. Muscle protein synthesis

Muscle protein synthesis was measured using the in vivo SUnSET method [38]. Under anesthesia, 0.04 μmol puromycin/g body weight (Wako) diluted in a 0.02 M PBS stock solution was injected intraperitoneally. The gastrocnemius muscle was removed exactly 15 min after puromycin administration. Following homogenization as described above and centrifugation at 2000 × g for 3 min at 4°C, the supernatant was collected and processed for western blotting. A mouse monoclonal anti-puromycin antibody (cat. #MABE343; Millipore, Billerica, MA) was used to detect puromycin incorporation, which was evaluated as the sum of the intensity of all protein signals in the western blot.

2.8. Real-time (q) and reverse transcription PCR

Total RNA was extracted from 20 mg of powdered muscle using ISOGEN 2 (NIPPON Gene, Tokyo, Japan) according to manufacturer’s instructions, after which the concentration was measured using Synergy HT (BioTek, Winooski, VT). Next, 500 ng of total RNA was reverse transcribed into cDNA using PrimeScript™ RT Reagent Kit with gDNA Eraser (Takara Bio, Shiga, Japan) for qPCR. qPCR was performed using TB Green® Premix Ex Taq™ II (Takara Bio) and the appropriate primers. Gene expression levels were determined by the absolute quantification method and normalized to Gapdh mRNA expression. Primer sequences used in this study are listed in Table 1.

2.9. Statistical analysis

Data were analyzed using two-way analysis of variance (ANOVA) using JMP pro 15. Post-hoc analyses were performed using t-tests,with a Benjamini and Hochberg false discovery rate correction for multiple comparisons when appropriate. The level of significance was set at P < 0.05. There were no criteria for including or excluding animals and there were no exclusions.

3. Results
3.1. Peak torque and force-time integral

Peak torque did not differ between the two groups (Table 2) and 2-DG did not affect the force-time integral during contractions (Table 2). Therefore,equal contractions were observed for rats in both groups.

3.2. Muscle glycogen and metabolite levels

Resting muscle glycogen content was lower in the 2-DG group than in the vehicle group (r < 0.05; Fig. 1A). Muscle glycogen content decreased immediately post-HiMC (r < 0.05), although the change in muscle glycogen level in the 2-DG group was minor compared with that in the control group (r < 0.05; Fig. 1B). 2-DG decreased resting muscle lactate content (r < 0.05; Fig. 1C) and muscle lactate content increased immediately after one set of HiMC in both groups (r < 0.05); however, 2-DG attenuated the HiMC-induced increase in muscle lactate content (r < 0.05; Fig. 1D).These results indicate that 2-DG inhibits glycolysis during both rest and contractions.

We further analyzed muscle metabolite levels immediately after HiMC using metabolomics to better understand the effect of 2-DG on muscle metabolism (Table 3). Although the decrease in muscle glycogen levels was minor in the 2-DG group, glucose 1-phosphate levels decreased (r < 0.05) comparably in both groups. Generally, in the control group, HiMC decreased the levels of glycolytic metabolites (from glucose 6-phosphate to pyruvic acids), whereas the levels of tricarboxylic acid (TCA) cycle metabolites increased. In the 2-DG group, the levels of the above metabolites decreased/increased similarly, although the resting levels of some metabolites, including 2-phosphoglyceric acid, 3-phosphoglyceric acid, and malic acid, tended to be lower than those in the control groups. Moreover, the levels of most glucogenic amino acids decreased after 2-DG administration. HiMC did not alter malonyl CoA levels in the control group but led to a decrease in content (undetected in the contracted muscle) in the 2-DG group. HiMC reduced ATP levels similarly in both groups (r < 0.05); in contrast, HiMC increased ADP and AMP levels in the control group but had no effect on their levels in the 2-DG group. HiMC also increased inosine monophosphate (IMP) levels in both groups (r < 0.05).

3.3. Upstream and downstream factors of mTOR signaling

2-DG decreased the phosphorylation of p70S6K Thr389 (r < 0.05; main effect of 2DG; Fig. 2A, B), 4E-BP1 Thr37/46, and the γ form of 4E-BP1 at both 0 h and 6 h post HiMC except for 4E-BP1 Thr37/46 phosphorylation at 0 h post HiMC (r < 0.05; Fig. 2C, D). HiMC increased p70S6K Thr389 phosphorylation similarly across both groups at both time points (r < 0.05; main effect of HiMC) and decreased the phosphorylation of 4E-BP1 Thr37/46 (r < 0.05) but not that of the γ form of 4E-BP1 immediately after HiMC. 2-DG inhibited the HiMC-induced increase in 4E-BP1 and γ form of 4E-BP1 phosphorylation 6 h after HiMC.

AMPK Thr172 phosphorylation immediately increased post-HiMC similarly in both groups (r < 0.05; Fig. 3B). An increase in Raptor Ser792 phosphorylation, a downstream target of AMPK, was also observed immediately after HiMC (r < 0.05), with a comparable increase between the groups (Fig. 3C). 2-DG tended to increase the general phosphorylation levels of Raptor Ser792 (r = 0.06; main effect of 2-DG).However, neither 2-DG nor HiMC altered AMPK and Raptor phosphorylation 6 h after HiMC.

Phosphorylation of Akt Thr308 and Ser473 increased immediately after HiMC in both groups (r < 0.05; Fig. 4B, C); 6 h after HiMC, Akt Thr308 phosphorylation (r < 0.05), but not that of Akt Ser473, was increased. 2-DG decreased the general phosphorylation of Akt Ser473 (r < 0.05; main effect of 2DG) at both time points. Although Akt phosphorylation increased immediately after HiMC, phosphorylation of PRAS40 Ser246 (r < 0.05), a downstream target of Akt, decreased 0 h post HiMC (Fig. 4D). In contrast, HiMC increased PRAS40 Ser246 phosphorylation 6 h after contraction (r < 0.05). 2-DG inhibited both basal and contraction-induced phosphorylation of PRAS40 Ser246 at both time points.

3.4.Protein synthesis

2-DG decreased basal muscle protein synthesis at both time points (r < 0.05; main effect of 2-DG; Fig. 5) while HiMC increased muscle protein synthesis 6 h after HiMC in the control group (r < 0.05). However, HiMC-mediated increase in muscle protein synthesis at 6 h post HiMC was not inhibited in the 2-DG group.

3.5. Protein degradation pathway

Although protein synthesis is considered a primary factor implicated in exercise-mediated muscle hypertrophy, protein degradation can also contribute to this process. Thus, we investigated the major protein degradation pathway. Neither 2-DG nor HiMC altered LC3 (LC3-I and II) expression (Fig. 6B, C). 2-DG decreased p62 expression (r < 0.05; main effect of 2-DG; Fig. 6D) but increased FoxO1, FoxO3,Atrogin- 1, and Murf- 1 transcription (r < 0.05; Fig. 6E–H); HiMC decreased these mRNA levels but only in the 2-DG group (r < 0.05). Moreover, 2-DG and HiMC did not alter the levels of ubiquitinated proteins (Fig. 6I), which are indicative of protein degradation.

3.6. PGC-1α and c-Myc

We further investigated the mRNA levels of PGC- 1α and c-Myc as markers of mitochondrial biogenesis. Both HiMC and 2-DG increased PGC- 1α and c-Myc mRNA levels 6 h after contraction (r < 0.05; Fig. 7A, B).

4. Discussion

Resistance exercise/high-intensity muscle contraction (HiMC) is known to stimulate mTORC1 and muscle protein synthesis. Currently, exercise volume/force-time integral, rather than contraction intensity, is considered to be an important variable in the regulation of mTORC1 and protein synthesis, indicating that not only mechanical stress, but also the metabolic aspect during and/or after muscle contraction affect mTORC1 activity and muscle protein synthesis. Recent studies indicated that glycolysis, which is robustly activated during HiMC, regulates mTORC1 and protein synthesis [27, 32]. However, no studies have directly investigated the role of glycolysis in the regulation of mTORC1 signaling and muscle protein synthesis. Therefore, in this study, we investigated the effect of 2-DG-mediated glycolysis inhibition on HiMC-induced mTORC1 activation and increase in protein synthesis. We found that glycolysis is a significant contributor to mTORC1 signaling and protein synthesis in skeletal muscle and that 2-DG inhibits basal mTORC1 signaling and muscle protein synthesis. Furthermore, 2-DG partially C25-140 price inhibited HiMC-induced mTORC1 signaling but not HiMC-induced muscle protein synthesis. Therefore, our results suggest that glycolysis plays an important role in hand infections both basal mTORC1 and muscle protein synthesis. Moreover, HiMC-mediated activation of glycolysis affects HiMC-induced mTORC1 signaling, although it does not significantly affect HiMC-induced muscle protein synthesis.

In this study, 2-DG administration inhibited basal mTORC1 signaling, as assessed by the phosphorylation levels of p70S6K and 4E-BP1. This inhibition occurred independent of AMPK signaling activation. Furthermore, while 2-DG did not alter the phosphorylation of Akt Thr308, a downstream factor of the insulin/PI3K signaling pathway, it decreased the phosphorylation of Akt Ser473, a downstream target of mTORC2. Therefore, in agreement with the results of previous studies on cells other than skeletal muscle [27, 39], glycolysis controls mTORC1 activity in skeletal muscle independently of AMPK and insulin/PI3K signaling.

In contrast to basal mTORC1 signaling, 2-DG did not inhibit HiMC-induced phosphorylation of p70S6K Thr389, although it inhibited HiMC-induced phosphorylation of 4E-BP1, indicating that HiMC-induced phosphorylation of p70S6K is regulated by a glycolysis-independent process. The reason for these opposing results regarding mTORC1 target protein phosphorylation is unclear. However, a previous study reported that HiMC differentially regulates the phosphorylation of p70S6K and 4E-BP1 [40, 41], and while p70S6K Thr 389 phosphorylation is rapamycin (an mTORC1 inhibitor)-sensitive, 4E-BP1 Thr37/46 is rapamycin-insensitive [5, 42]. Furthermore, PRAS40 competes with mTORC1 substrates, including p70S6K and 4E-BP1, for binding to Raptor [43, 44], although dephosphorylation of PRAS40 is associated with dephosphorylation of 4E-BP1 rather than p70S6K. Although why PRAS40 regulates 4E-BP1 rather than p70S6K is not clear, 2-DG inhibited both basal and HiMC-induced PRAS40 phosphorylation in this study. Therefore, glycolysis may regulate mTORC1/4E-BP1 signaling rather than mTORC1/p70S6K signaling via PRAS40.

A recent study identified dihydroxyacetone phosphate (DHAP) as a glycolytic metabolite that activates mTORC1 [27]. Hence, we hypothesized that HiMC increases DHAP levels while 2-DG inhibits this increase. However, the levels of DHAP and other glycolytic metabolites generally decreased immediately after HiMC, and 2-DG did not alter their levels. Therefore, our results indicate that glycolytic metabolites do not regulate mTORC1 in skeletal muscle. In contrast, while muscle lactate levels did not differ between non-contracted and contracted muscles immediately after five sets of HiMC, they were higher in the contracted muscle than in the non-contracted muscle immediately after one set of HiMC. Furthermore, muscle glycogen levels were extremely low after five sets of HiMC. Therefore, glycolysis was possibly more active with high levels of glycolytic metabolites in the contracted muscle during the early phase of HiMC (e.g., first/second set of HiMC), and thus it remains possible that glycolytic metabolites regulate mTORC1 in skeletal muscle.

A possible mechanism underlying mTORC1 inhibition by 2-DG is the 2-DG-induced decrease in amino acids including branched-chain amino acids (BCAA) in skeletal muscle. Indeed, leucine is known to activate mTORC1 in skeletal muscle [45]. However, HiMC did not alter BCAA concentrations in skeletal muscle, indicating that although basal mTORC1 inhibition by 2-DG is mediated, at least in part, by the decrease in BCAA, glycolysis-associated mTORC1 activation by HiMC is a BCAA-independent process.

AMPK is known to inhibit mTORC1 in some cells, including skeletal muscles [46], and pharmacological AMPK activation during HiMC inhibits mTORC1 activation [47]. In this study, while phosphorylation of AMPK Thr172 was comparable between the groups, 2-DG tended to facilitate HiMC-induced phosphorylation of Raptor Ser792, a downstream target of AMPK, immediately post-HiMC, indicating that 2-DG slightly activates AMPK. However, a recent study reported that the activity of AMPK, at least that of α2-AMPK, does not regulate HiMC-induced mTORC1 activation [32]. In particular, compared with wild-type mice, HiMC-induced phosphorylation of Raptor Ser792 is inhibited in dominant negative α2-AMPK kinase dead transgenic (KD-AMPK) mice; p70S6K and 4E-BP1 phosphorylation levels were unaltered in the KD-AMPK mice. Furthermore, although 2-DG tended to stimulate AMPK/Raptor signaling immediately after HiMC, it did not alter AMPK/Raptor signaling 3 h post HiMC. Therefore, it is unlikely that the 2-DG-induced slight activation of AMPK/Raptor signaling inhibited HiMC-induced mTORC1/4E-BP1 signaling in this study.

Similar to mTORC1 signaling, basal muscle protein synthesis was inhibited by 2-DG administration. It is known that mTORC1 regulates basal muscle protein synthesis [12, 36, 48]; thus, our results suggest that glycolysis regulates basal muscle protein synthesis via mTORC1. A decrease in ATP levels is considered a mechanism of inhibition of mTORC1 and protein synthesis by 2-DG [35, 49]. However, a recent study in C2C12 cells reported that decreased ATP levels do not influence AMPK phosphorylation and muscle protein synthesis [50]. Moreover, in this study, 2-DG did not affect ATP levels, which instead decreased during HiMC. Therefore, these results suggest that ATP levels are not a major regulator of glycolysis-associated protein synthesis in skeletal muscle.

In contrast to the basal regulation of muscle protein synthesis, HiMC-induced muscle protein synthesis occurs through a rapamycin-sensitive mTOR/mTORC1-independent mechanism [5, 9, 12, 36, 48, 51]. In this study, glycolysis inhibition via 2-DG administration attenuated HiMC-induced rapamycin-insensitive mTORC1/4E-BP1 signaling but not muscle protein synthesis. Therefore, our results indicate that glycolysis-associated regulation of rapamycin-insensitive mTORC1/4E-BP1 signaling by HiMC does not control muscle protein synthesis, and that neither HiMC-induced rapamycin-sensitive nor insensitive mTORC1 activation play a role in the HiMC-induced regulation of muscle protein synthesis. In addition, we previously reported that mTORC2 inhibition through muscle-specific rictor knockout did not affect HiMC-induced muscle protein synthesis [52]; instead, the ATP-competitive mTOR kinase inhibitor AZD8055, which inhibits both mTORC1 and mTORC2, prevented the protein synthesis induced by HiMC [5]. Therefore, our results suggest that neither mTORC1 nor mTORC2 is involved in the regulation of HiMC-induced muscle protein synthesis but that mTOR activity per se and/or an unknown mTOR complex regulate HiMC-induced muscle protein synthesis.

In addition, we observed that 2-DG-mediated glycolysis inhibition upregulated atrogenes. Given that ubiquitinated protein levels did not change in this study, acute glycolytic inhibition may not significantly affect muscle protein degradation. Indeed, glycolytic and glycogen Biokinetic model handling enzyme levels are low in aged fast-type muscle [53], and therefore chronic inhibition of glycolysis may affect muscle atrophy. Interestingly, the atrogenes upregulated by 2-DG were not completely, but still significantly, downregulated by HiMC, indicating that glycolytic activity is important for atrogene regulation.

Finally, we investigated the mRNA levels of mitochondrial biogenesis markers, PGC- 1α and c-Myc. Both have been shown to increase during HiMC independently of rapamycin-sensitive mTOR
/mTORC1 signaling activation [36]. Here, 2-DG and HiMC synergistically increased PGC- 1α and c-Myc mRNA levels, indicating that glycolysis inhibition stimulates aerobic energy metabolism as a compensatory mechanism. However, it is of note that the role of c-Myc in skeletal muscle is currently not well understood,although it is believed to stimulate mitochondrial biogenesis [54]. In addition to being metabolic regulators, PGC- 1α and c-Myc are also protein synthesis regulators [55, 56]. Therefore, PGC- 1α and c-Myc may also facilitate muscle protein synthesis to compensate for the decrease in muscle protein synthesis after glycolysis inhibition.

5. Conclusions

2-DG-induced inhibition of glycolysis resulted in decreased basal mTORC1 signaling and muscle protein synthesis. Moreover, it attenuated HiMC-induced mTORC1 signaling to a certain extent, although it did not alter the response of muscle protein synthesis to HiMC. These findings indicate that although glycolysis has a minor role in the regulation of HiMC-induced muscle protein synthesis, it regulates both basal and HiMC-induced mTORC1 signaling. Given that skeletal muscle mTORC1 signaling regulates various cellular functions and processes including fiber type composition, cell size, force production, and energy metabolism [57-60], glycolysis during muscle contractions may play key roles in skeletal muscle adaptation to HiMC. Future studies should investigate the mechanisms how glycolysis regulates HiMC-induced mTORC1, the potential glycolysis-mTORC1 connection in humans, and its practical implications, such as whether higher glucose availability (e.g., high muscle glycogen) augments the resistance exercise-induced muscle hypertrophy.