Branched-chain ketoacid overload inhibits insulin action in the muscle

Abstract

Branched-chain α-keto acids (BCKAs) are catabolites of branched-chain amino acids (BCAAs). Intracellular BCKAs are cleared by branched-chain ketoacid dehydrogenase (BCKDH), which is sensitive to inhibitory phosphorylation by BCKD kinase (BCKDK). Accumulation of BCKAs is an indicator of defective BCAA catabolism and has been correlated with glucose intolerance and cardiac dysfunction. However, it is unclear whether BCKAs directly alter insulin signaling and function in the skeletal and cardiac muscle cell. Furthermore, the role of excess fatty acids (FAs) in perturbing BCAA catabolism and BCKA availability merits investigation. By using immunoblotting and ultra-performance liquid chromatography MS/MS to analyze the hearts of fasted mice, we observed decreased BCAA-catabolizing enzyme expression and increased circulating BCKAs, but not BCAAs. In mice subjected to diet-induced obesity (DIO), we observed similar increases in circulating BCKAs with concomitant changes in BCAA-catabolizing enzyme expression only in the skeletal muscle. Effects of DIO were recapitulated by simulating lipotoxicity in skeletal muscle cells treated with saturated FA, palmitate. Exposure of muscle cells to high concentrations of BCKAs resulted in inhibition of insulin-induced AKT phosphorylation, decreased glucose uptake, and mitochondrial oxygen consumption. Altering intracellular clearance of BCKAs by genetic modulation of BCKDK and BCKDHA expression showed similar effects on AKT phosphorylation. BCKAs increased protein translation and mTORC1 activation. Pretreating cells with mTORC1 inhibitor rapamycin restored BCKA's effect on insulin-induced AKT phosphorylation. This study provides evidence for FA-mediated regulation of BCAA-catabolizing enzymes and BCKA content and highlights the biological role of BCKAs in regulating muscle insulin signaling and function.

Branched-chain amino acids (BCAAs) and their catabolites are bioactive molecules with a broad repertoire of metabolic actions in cellular health and disease (1–3). Leucine, isoleucine, and valine (BCAAs) are reversibly transaminated by branched-chain aminotransferase (BCAT) to yield branched-chain α-keto acids (BCKAs), specifically α-ketoisocaproate or ketoleucine (KIC), α-ketoisovalerate or ketovaline (KIV), and α-keto-beta-methylvalerate or ketoisoleucine (KMV). BCKAs are irreversibly decarboxylated by BCKA dehydrogenase (BCKDH), a rate-limiting step in BCAA catabolism. BCKDH activity is tightly regulated by BCKA dehydrogenase kinase (BCKDK)-mediated inhibitory phosphorylation (4–6) of BCKDH or BCKDH dephosphorylation and activation by protein phosphatase 2C (PP2Cm) (7, 8). Transcriptional regulation of the BCAA-catabolizing enzymes and BCAA catabolism is induced by Kruppel-like factor 15 (KLF15) (9), which also regulates glucose and fat oxidation (10–12). Emerging studies have shown that not only inborn mutations in BCAA enzymes, but also dysfunctional BCKA oxidation, is observed in metabolic disorders like cancer (13, 14), obesity and insulin resistance (15–17), ischemia (18), and diabetic cardiomyopathy (19, 20).

Skeletal and cardiac muscle insulin signaling and sensitivity are critical for metabolic homeostasis and organ function (21, 22). A strong association between plasma BCAAs, muscle BCAA catabolic gene expression, and insulin resistance (IR) is reported in several human and rodent models of obesity (1, 23). Whether defects in muscle BCAA-catabolizing enzyme expression and activity precede or follow obesity-induced IR remains unclear. Strikingly, in ob/ob and diet-induced obese (DIO) mice, BCKDK inhibition normalized BCAA catabolism and attenuated IR (23). Therefore, restoring BCAA catabolism and clearing BCAA metabolites in endocrine disorders present beneficial functional outcomes. Conversely, accumulation of BCAA catabolites likely causes maladaptive substrate metabolism and organ dysfunction. However, it remains unknown whether BCKAs directly impact muscle insulin signaling to influence substrate utilization and mitochondrial function. A recent report demonstrated that incubation of cardiomyocytes with excess BCKAs reduces glucose uptake (24) and suppresses mitochondrial respiration (25), questioning the effect of BCKA on insulin signaling and action.

In the current study, we demonstrated post-translational changes in BCAA-catabolizing enzyme expression and increased circulating BCKAs, but not BCAAs, in the fasted mouse heart. However, in the skeletal muscle but not the heart of mice subjected to DIO, circulating BCKA levels increased with a concomitant decrease in BCAA-catabolizing enzyme expression. The effects of DIO on BCAA-catabolizing enzyme expression and BCKA content were replicated in vitro by treating cells with saturated FA palmitate. Additionally, we observed that treatment with exogeneous BCKAs or increasing intracellular BCKA levels by modulating BCKDHA and BCKDK enzymes resulted in impaired insulin signaling and function in the cardiac and skeletal muscle. Last, we demonstrated that BCKAs can activate protein translation, and its effect on insulin signaling is likely attributable to mTOR signaling activation.

Results

Overnight fasting augments BCKDH phosphorylation and protein expression in the cardiac tissue but not in the gastrocnemius muscle

We examined whether acutely limiting nutrient intake (fasting) alters transcriptional or post-translational levels of BCAA catabolic enzymes in the cardiac and skeletal muscle (Fig. 1A). Male C57BL/6J mice subjected to overnight fasting exhibited a decline in body weight, serum glucose, and liver weight (Table S1) when compared with either ad libitum fed or refed mice. Ventricular weight to body weight ratio remained unchanged between groups (Table S1). Fasting increased serum BCKA levels (total and individual), whereas refeeding restored them to ad libitum fed levels (Fig. 1B). Unlike BCKAs, serum BCAAs levels remained unchanged following fasting but increased after refeeding (Fig. 1C). To confirm whether changes in circulating BCKAs were an outcome of altered BCAA catabolism, we examined protein markers of the BCAA catabolic pathway in the gastrocnemius muscle and heart. Inactivating phosphorylation of BCKDE1α at Ser-293 (suggestive of decreased BCKDHA activity) and total protein content of BCKDHA and BCKDK were unaltered in the gastrocnemius muscle (Fig. 1D). However, a marked increase in phosphorylation of BCKDE1α at Ser-293 was observed in the fasted hearts, which returned to fed levels upon 4 h of refeeding (Fig. 1E). Interestingly, BCKDHA protein levels were decreased upon fasting and were restored in the hearts of refed mice (Fig. 1E). In the heart, BCKDK protein levels remained unaltered across all the groups (Fig. 1E). Phosphorylated BCKDE1α Ser-293 levels in the heart (Fig. S1A) but not the gastrocnemius muscles (Fig. S1B) correlated with circulating BCKAs. Because nutritional changes are accompanied by changes in circulating insulin (decreased in fasting and increased in refeeding), we examined whether protein expression of BCAA-catabolizing enzymes corresponded with changes in AKT phosphorylation, an effector of insulin signaling. Immunoblot assessment of isoform-specific AKT phosphorylation revealed that AKT1 and AKT2, at Ser-473 and Ser-474, respectively, increased following refeeding in both gastrocnemius muscle (Fig. 1D) and heart (Fig. 1E). However, fasting decreased AKT2 Ser-474, but not AKT1 Ser-473, phosphorylation in both gastrocnemius muscle (Fig. 1D) and heart (Fig. 1E). Regardless of the tissue-specific changes in BCAA-catabolizing enzymes, AKT phosphorylation in both the heart (Fig. S1C) and gastrocnemius muscles (Fig. S1D) was negatively correlated with serum BCKAs. Notably, the changes in the BCAA catabolic protein expression and intracellular and circulating BCKAs were independent of mRNA expression changes of BCAA catabolic enzymes (Bckdha, Bckdk, Bcat2, Klf15, Oxct2a, Ivd2, Hmgcs1, and Mut) in both gastrocnemius and cardiac muscle (Fig. S1, E and F). Therefore, short-term physiological changes in insulin is associated with altered protein expression of BCAA-catabolizing enzymes and systemic BCKA content.

Figure 1.

Fasting induces post-translational changes in BCAA catabolic enzymes in the cardiac but not skeletal muscle. A, study design of C57BL/6J mice ad libitum fed or fasted for 16 h or refed for 4 h following fasting, n = 5 each group. B and C, analysis of serum BCKAs (B) and serum BCAAs (C) by UPLC MS/MS. Immunoblot and densitometric analysis of total and phosphorylated BCKDHA E1α Ser-293, total BCKDK, phosphorylated AKT1 Ser-473, total AKT1, phosphorylated AKT2 Ser-474, and total AKT2 in the gastrocnemius muscle (D) and heart (E). Statistical analysis was performed using two-way ANOVA followed by Tukey's multiple-comparison test. *, p < 0.05; **, p < 0.01; ****, p < 0.0001, as indicated. *, fed versus fasted; #, fasted versus refed; @, fasted versus refed. Data are presented as mean ± S.D.

Expression of BCAA catabolic enzymes is regulated differentially in the cardiac and skeletal muscles following DIO

During fasting, decreased insulin relieves inhibition of fatty acid oxidation in the muscle (26). Because muscle BCAA-catabolizing enzyme expression decreased in the presence of excess lipids, we postulated that FAs compete with BCAAs as muscle substrates by likely regulating expression and activity of BCAA-catabolizing enzymes to govern BCAA catabolism. We ascertained whether amino acid–metabolizing enzymes in the muscle and systemic BCKAs were temporally regulated in a model of hyperinsulinemia and progressive lipid excess (DIO). Male C57BL/6J mice were fed a high-fat high-sucrose (HFHS) diet, and blood and tissues were collected at 2, 4, 8, and 13 weeks after feeding to determine the BCAA catabolic enzyme expression in the gastrocnemius muscle and heart (Fig. 2A). Systemic BCKA levels were down-regulated only at 13 weeks after HFHS feeding (Fig. 2B). Circulating BCAA levels remained unaltered across chow and HFHS groups at all time points (Fig. S2A). To examine the effect of progressive lipid loading on BCAA-metabolizing enzymes, we compared the expression profile of genes involved in BCAA catabolism in the gastrocnemius muscles between 2 weeks and 13 weeks of HFHS feeding. mRNA expression of Mut and Ivd2, intermediary effector enzymes of the BCAA oxidation pathway, was reduced in the gastrocnemius muscle at 13-week compared with 2-week HFHS-fed mice (Fig. 2C), consistent with a prior report in mice fed a (60%) high-fat diet (27). Interestingly, Hmgcs1 levels were up-regulated in the gastrocnemius muscle in 13-week HFHS fed mice compared with 2-week HFHS fed mice (Fig. 2C), possibly as a compensatory mechanism to supply 3-hydroxy-3-methylglutaryl-CoA for Hmgcl to maintain CoA pools. In the heart, mRNA expression of the BCAA catabolic enzymes, except for KLF15, remained unaltered in a setting of DIO (Fig. 2D). Because the earliest changes in mRNA expression were not detected until 13 weeks, we used 4-week, and not 2-week, HFHS-fed mice as controls to analyze protein expression. Phosphorylation of BCKDE1α at Ser-293 in the gastrocnemius muscles was unchanged at 8 weeks but increased at 13 weeks in HFHS-fed mice compared with the 4-week HFHS-fed mice (Fig. 2E). Unlike the fasted state (acute lipid turnover), no changes in BCKDH complex phosphorylation were found in hearts from HFHS-fed mice (chronic lipid excess) compared with chow-fed (Fig. 2F). AKT1 Ser-473 phosphorylation trended to decline in the gastrocnemius muscles (Fig. 2E) of 13-week HFHS-fed mice but remained unchanged in the heart (Fig. 2F), signifying a likely reduction in insulin action with progressive nutrient overload in the skeletal muscle during obesity. Together, these data demonstrate that unlike acute fasting, chronic HFHS feeding down-regulates enzymes involved in BCAA catabolism in the gastrocnemius muscle but not the heart.

Figure 2.

Diet-induced obesity down-regulates BCAA-catabolizing enzymes in the skeletal muscle. A, study design of HFHS diet feeding of C57BL/6J mice at 2, 4, 8, and 13 weeks. B, serum BCKA measurements using UPLC MS/MS at 8 and 13 weeks after HFHS diet feeding (n = 5/group). C, quantification of Klf15, Mut, Ivd2, Acadm, and Hmgcs1 mRNA expression corrected to Rpl41/Cyclo (cyclophilin) reference genes in gastrocnemius muscle of 2- and 13-week HFHS-fed mice. D, quantification of Bckdha, Bckdk, Bcat2, Klf15, Ivd2, and Mut mRNA levels corrected to Rpl41/Rer1 reference genes in the heart of 4-, 8-, and 13-week HFHS-fed mice. E and F, immunoblot and densitometric analysis of total and phosphorylated BCKDHA E1α Ser-293, total and phosphorylated AKT1 Ser-473, and total and phosphorylated AKT2 Ser-474 in gastrocnemius muscle (E) and heart (F). Statistical analysis was performed using two-way ANOVA followed by Tukey's multiple-comparison test; *, p < 0.05; **, p < 0.01; ****, p < 0.0001 as indicated. *, within groups; #, between groups. Data are presented as mean ± S.D.

BCAA-catabolizing enzyme mRNA expression is decreased and intracellular BCKA levels are increased in C2C12 cells after palmitate exposure

Chronic lipid overload not only drives IR but also increases systemic and myocellular BCAAs (1, 9) and BCKAs (9). To simulate the gradual increase in FAs observed in obesity and IR ex vivo, C2C12 cells were treated with low (0.2 mm), intermediate (0.4 mm), and high (0.8 mm) palmitate for 16 h. Exposure to palmitate did not alter the expression of proximal BCAA-catabolizing genes (Bcat2, Bckdha) or the regulatory kinase, Bckdk (Fig. 3A). However, the intermediary (Acadsm, Ivd2) and distal (Hibch, Mut) genes were significantly down-regulated in the 0.8 mm palmitate-treated cells (Fig. 3A). Acadsm levels were also reduced in response to 0.4 mm palmitate treatment (Fig. 3A). As observed in HFHS-fed gastrocnemius muscles, Hmgcs1 was markedly up-regulated in all three palmitate-treated groups (Fig. 3A). Our data revealed that palmitate mimicked the effect of DIO on BCAA-catabolizing gene expression in vitro. To examine whether decreased mRNA expression of distal BCAA-oxidizing enzymes resulted in decreased BCAA catabolism, we measured intracellular BCKAs and BCAAs. Total and individual BCKAs were markedly elevated in 0.8 mm palmitate-treated cells (Fig. 3B), plausibly suggesting suppression of BCKA oxidation. This effect was not observed in cells treated with intermediate and low concentrations of palmitate (Fig. 3B). Intracellular BCAA levels remained unchanged across all treatment groups (Fig. 3C). Similar to that observed in physiological (fasting) and pathological (DIO) models of altered nutrient status, increases in BCKA levels corresponded with decreased phosphorylation of AKT1 and AKT2 at Ser-473 and Ser-474, respectively (Fig. 3, D and E), thereby calling into question the relationship between BCKAs and insulin signaling. Because lipid overload altered the expression of distinct BCAA catabolic enzymes and increased BCKA levels, at least in the skeletal muscle, we examined whether BCKAs influence muscle insulin signaling.

Figure 3.

Palmitate alters intracellular BCKA levels and transcript levels of distal BCAA catabolic genes in C2C12 cells. A, C2C12 cells preincubated with either 2% BSA or 2% BSA conjugated with 0.2, 0.4, or 0.8 mm palmitate for 16 h. Shown is quantification of proximal (Bcat2, Bckdh, and Bckdk), intermediary (Acadm and Ivd2), and distal (Hibch, Mut, and Hmgcs1) BCAA-catabolizing enzyme mRNA levels corrected to Rer1/Rpl7 reference gene levels. Statistical analysis was performed using two-way ANOVA followed by Tukey's multiple-comparison test: *, p < 0.05; **, p < 0.01; ****, p < 0.0001 as indicated. *, BSA versus 0.8 mm Pal; #, BSA versus 0.4 mm Pal; ^, BSA versus 0.2 mm Pal. B and C, intracellular BCKAs (B) and BCAAs (C) levels analyzed using UPLC MSMS. Statistical analysis was performed using one-way ANOVA followed by Tukey's multiple-comparison test: *, p < 0.05; **, p < 0.01; ****, p < 0.0001, as indicated. *, BSA versus 0.8 mm Pal. D, immunoblot and densitometric analysis of total and phosphorylated AKT1 Ser-473 and total and phosphorylated AKT2 Ser-474. Statistical analysis was performed using two-way ANOVA followed by Tukey's multiple-comparison test; *, p < 0.05; **, p < 0.01; ****, p < 0.0001, as indicated. *, BSA versus 0.8 mm Pal; #, BSA versus 0.4 mm Pal. E, linear regression of intracellular BCKAs correlated with the phosphorylated to total AKT1 Ser-473 protein expression. Statistical analysis was performed using one-way ANOVA followed by Tukey's multiple-comparison test: *, p < 0.05; **, p < 0.01; ****, p < 0.0001, as indicated. Data are presented as mean ± S.D.

BCKAs inhibit insulin signaling in muscle cells

BCKAs are emerging as clinically relevant biomarkers for IR (28, 29). BCKAs are not only generated intracellularly but also transported between tissues to support nitrogen balance, ketone body metabolism, and gluconeogenesis (30, 31). Acute (30-min) treatment of C2C12 cells with 0.05 mm 4-methyl 2-oxopentanoic acid sodium salt (KIC), sodium-3-methyl-2-oxobutyrate (KIV), 3-methyl-2-oxovaleric acid sodium salt (KMV), or a combination of all three (BCKAs) decreased insulin-induced phosphorylation of AKT1 at Ser-473 and AKT2 at Ser-474 (Fig. 4A). Moreover, treating C2C12 cells with pathological concentrations of KIC (1, 2, and 5 mm) also resulted in a concentration-dependent impairment of insulin-induced AKT1 phosphorylation at Ser-473 (Fig. S3A). Notably, insulin-mediated activating phosphorylation of insulin receptor substrate 1 (IRS1) at Tyr-612 was inhibited by KIC at a concentration of 5 mm but not 1 or 2 mm (Fig. S3A). KIC-mediated inhibition of AKT1 and IRS1 phosphorylation was observed at 15 min and persisted until 30 min of KIC treatment (Fig. S3B). However, phosphorylated AKT1 levels were restored to control levels at 60 min (Fig. S3B). Similar to C2C12 cells, the inhibitory effect of KIC on insulin-stimulated AKT1 phosphorylation was also recapitulated in the rat L6 muscle cell line (Fig. S3C). Because BCKAs inhibited insulin signaling per se, we next tested whether BCKAs modified palmitate-induced inhibition of insulin signaling. In L6 cells incubated with 0.4 mm palmitate, KIC exacerbated palmitate-mediated decreases in AKT1 Ser-473 phosphorylation and phosphorylation of IRS1 at Tyr-612 (Fig. S3D), whereas in C2C12 cells, KIC's effect on AKT and IRS phosphorylation was exacerbated by palmitate (Fig. 4B). To address whether other ketoacids also impact skeletal muscle insulin signaling in the presence of palmitate, we treated C2C12 cells with 5 mm KIV (Fig. 4C), KMV (Fig. 4D), and a combination of all three BCKAs (Fig. 5A) in the presence or absence of 0.4 mm palmitate. Similar to the effect of KIC, AKT1 Ser-473 phosphorylation was impaired by both KIV and KMV in the absence of palmitate (Fig. 3, B and C). Moreover, KIV's effects on AKT and IRS phosphorylation were also exacerbated by palmitate as seen with KIC in C2C12 cells (Fig. 4C). KMV did not exacerbate palmitate-induced impairment of insulin signaling (Fig. 4, C and D). A combination of all BCKAs significantly reduced AKT1 phosphorylation at Ser-473 but did not exacerbate palmitate-induced insulin signaling impairment in C2C12 cells (Fig. 5A). Immunoprecipitation studies revealed reduced activating tyrosine phosphorylation of IRS1 at residue 612 upon BCKA stimulation in C2C12 cells (Fig. 5B). Consistent with reduced AKT1 phosphorylation, insulin-induced AKT1 activity (Fig. 5C) and glucose uptake (Fig. 5D) were also decreased by KIC and KIV. KMV did not influence AKT1 activity and glucose uptake, perhaps reflecting its more modest ability to inhibit insulin signaling and function in C2C12 cells (Fig. 5D). Our data are consistent with a recent report that demonstrated that BCKAs inhibits insulin signaling in 3T3-L1 adipocytes (23). KIC inhibited insulin-mediated AKT1 Ser-473 phosphorylation in the absence of palmitate not only in skeletal muscle cell lines but also in primary cells, such as neonatal rat cardiomyocytes (NRCMs) (Fig. 5E) and adult rat cardiomyocytes (ARCMs) (Fig. 5F). KIC exacerbated palmitate-induced insulin signaling impairment in ARCMs, but not in NRCMs (Fig. 5, E and F). Similarly, treatment with KIV and KMV also decreased insulin-stimulated AKT1 Ser-473 phosphorylation in ARCMs (Fig. S4A–B). To deduce whether BCKAs induced inhibition of insulin signaling and glucose utilization by altering mitochondrial function, we examined extracellular oxygen consumption and ATP production in the adult mouse cardiomyocytes (AMCMs). BCKAs significantly reduced extracellular oxygen consumption in the AMCMs (Fig. 5G) in agreement with prior studies (9, 32). Treatment with BCKAs increased ATP production in the AMCMs (Fig. 5H), suggesting a plausible uncoupling of cellular respiration and ATP synthesis in response to BCKAs. These data suggest that BCKAs reprogram insulin signaling and insulin's effect on substrate uptake and mitochondrial function.

Figure 4.

BCKAs impair insulin signaling in C2C12 cells. A, differentiated C2C12 cells were treated with either 0.05 mm KIC, KMV, or KIV, a combination of all three BCKAs, or vehicle control for 30 min, followed by 100 nm insulin for 15 min. Data show immunoblot and densitometric analyses of total and phosphorylated AKT1 Ser-473 and total and phosphorylated AKT2 Ser-474. Statistical analysis was performed using one-way ANOVA followed by Tukey's multiple-comparison test: *, p < 0.05; **, p < 0.01; ****, p < 0.0001, as indicated. a, CON versus VEH + INS; b, VEH + INS versus KIC + INS; c, VEH + INS versus KMV + INS; d, VEH + INS versus KIV + INS; e, VEH + INS versus BCKA + INS. B–D, differentiated C2C12 cells were preincubated with either 2% BSA or 2% BSA conjugated with 0.4 mm palmitate for 16 h, followed by either 5 mm KIC (a), KIV (b), or KMV (c) treatment for 30 min. Myotubes were subjected to 100 nm insulin for 15 min in the presence or absence of individual BCKAs. Shown are immunoblot and densitometric analyses of total and phosphorylated AKT Ser-473 and total and phosphorylated IRS Tyr-612. Statistical analysis was performed using two-way ANOVA followed by Tukey's multiple-comparison test: *, p < 0.05; **, p < 0.01; ****, p < 0.0001, as indicated. Only statistically significant groups have been denoted on each graph. a, VEH versus BSA + INS; b, PA versus PA + INS; c, BSA + INS versus PA + INS; d, BSA + INS versus BSA + KIC/KMV/KIV + INS; e, BSA + KIC/KIV/KMV versus BSA + KIC/KMV/KIV + INS; f, BSA + KIC/KMV/KIV + INS versus PA + KIC/KMV/KIV + INS; g, PA + INS versus PA + KIC/KMV/KIV + INS. Data are presented as mean ± S.D.

Figure 5.

BCKAs impair insulin signaling and function in the muscle. A, differentiated C2C12 myotubes were preincubated with either 2% BSA or 2% BSA conjugated with 0.4 mm palmitate for 16 h, followed by 5 mm total BCKAs for 30 min. Myotubes were subjected to 100 nm insulin for 15 min in the presence or absence of BCKAs. Data show immunoblot and densitometric analyses of total and phosphorylated AKT Ser-473 levels and IRS Tyr-612. Statistical analysis was performed using two-way ANOVA followed by Tukey's multiple-comparison test: *, p <0.05; **, p < 0.01; ****, p <0.0001, as indicated. Only statistically significant groups have been denoted on each graph. a, VEH versus BSA + INS; d, PA versus PA + INS; e, BSA + INS versus PA + INS; c, BSA + INS versus BSA + BCKA + INS; b, BSA + BCKA versus BSA + BCKA + INS. B, C2C12 myotubes were treated with 5 mm total BCKAs for 30 min, followed by 100 nm insulin for 15 min. Cell lysates were then immunoprecipitated (IP) with IRS1 antibody and analyzed by immunoblotting (IB) using anti-pIRS Tyr-612 antibody (top). The bottom panels show the respective protein levels of pIRS Tyr-612, IRS1, actin, and total protein loading. C, quantification of insulin-induced AKT activity in C2C12 myotubes following 100 nm insulin stimulation for 15 min in the presence or absence of 5 mm KIC, KMV, and KIV treated for 30 min. Non-insulin–, non-BCKA–treated cells served as controls, n = 3. Statistical analysis was performed using Student's t test, *, p < 0.05; *, −INS versus +INS; @, VEH + INS versus KIC + INS; ^, VEH + INS versus KIV + INS. D, 2-DG uptake measured in C2C12 myotubes treated with 5 mm individual and total BCKAs for 30 min, followed by 1 μm insulin stimulation for 20 min. Statistical analysis was performed using one-way ANOVA, followed by Tukey's multiple-comparison test: *, p < 0.05; **, p < 0.01; ****, p < 0.0001, as indicated. *, VEH versus +INS; @, VEH + INS versus KIC + INS; ^, VEH + INS versus KIV + INS; #, VEH + INS versus BCKA + INS. E and F, isolated rat NRCMs (E) and ARCMs (F) were preincubated with either 2% BSA or 2% BSA conjugated with 0.4 mm palmitate for 16 h, followed by 5 mm KIC for 30 min. Data show immunoblot and densitometric analyses of total and phosphorylated AKT1 Ser-473 levels in cardiomyocytes subjected to either 200 nm (NRCMs) or 100 nm (ARCMs) insulin stimulation for 20 min in the presence or absence of KIC. Statistical analysis was performed using two-way ANOVA followed by Tukey's multiple-comparison test: *, p < 0.05; **, p < 0.01; ****, p < 0.0001, as indicated. Only statistically significant groups have been denoted on each graph. a, VEH versus BSA + INS; b, PA versus PA + INS; c, BSA + INS versus PA + INS; d, BSA + INS versus BSA + KIC + INS; e, BSA + KIC versus BSA + KIC + INS; f, BSA + KIC + INS versus PA + KIC + INS; g, PA + INS versus PA + KIC + INS. G and H, mitochondrial function analyzed in AMCMs treated with 5 mm BCKAs by measuring extracellular oxygen consumption (G) and ATP production (H). Statistical analysis for oxygen consumption was performed using one-way ANOVA followed by Tukey's multiple-comparison test: *, p < 0.05; **, p < 0.01; ****, p < 0.0001 as indicated; *, VEH versus KIC; @, VEH versus KMV; ^, VEH versus KIV; #, VEH versus BCKA, n = 3; Statistical analysis was performed using Student's t test, *, p < 0.05. Data are presented as mean ± S.D.

Altered clearance of BCKAs by genetic and pharmacological modulation of BCAA-catabolizing enzymes influences muscle insulin signaling

We next examined whether BCKA-mediated impairment in insulin signaling can be precipitated by increasing endogenous BCKA accumulation as opposed to exposure to exogenous BCKAs. Enzymatic activity of BCKDH determines the catabolic fate of BCKAs (1). BCKDH activity is inhibited by BCKDK-mediated phosphorylation, an effect that is reversed by PPM1K-mediated BCKDH dephosphorylation (Fig. 6A). We hypothesized that targeting BCAA-catabolizing enzymes will modulate oxidation and clearance of intracellular BCKAs, influencing BCKA's effect on insulin signaling. C2C12 myotubes were transduced with adenoviruses overexpressing BCKDK and BCKDHA, the key subunit of BCKDH complex (Fig. S5A). Intracellular BCKAs were significantly increased in BCKDK-overexpressing cells and modestly decreased in BCKDHA-overexpressing cells (Fig. 6B), signifying that altering BCAA metabolic enzyme levels changes cellular BCKA content. We next examined whether this change in intracellular BCKA levels precipitates alterations in insulin signaling. Consistent with increased accumulation of intracellular BCKAs, BCKDK overexpression decreased insulin-mediated phosphorylation of AKT1 at Ser-473 and AKT2 at Ser-474 (Fig. 6C). AKT1 phosphorylation at both the Ser-473 and Ser-474 residues of AKT2 (Fig. 6D) was increased in myotubes overexpressing BCKDHA in the presence of insulin. Silencing BCKDHA in C2C12 myoblasts (Fig. S5B) impaired AKT1 Ser-473 phosphorylation (Fig. 6E) and phosphorylated AKT content, as measured by sandwich ELISA analysis (Fig. S5C). Improved insulin signaling induced by BCKDHA overexpression resulted in increased pyruvate dehydrogenase (PDH) activity (Fig. S5D). However, BCKDHA silencing did not impact PDH activity (Fig. S5D). Similar effects on AKT1 Ser-473 phosphorylation were observed in NRCMs with either overexpression or knockdown of BCKDHA (Fig. 6F).

Figure 6.

Modulating intracellular BCKAs by modifying BCAA catabolic enzyme expression alters skeletal and cardiac muscle insulin signaling. A, brief schematic of enzymes regulating BCKA catabolism and BT2 action. B, UPLC mass spectrometric analysis of intracellular BCKA levels in C2C12 myotubes transduced with Ad-CMV-GFP-hBCKDK/HA (MOI 150) or Ad-CMV-mCherry-hBCKDHA/FLAG (MOI 150) and their appropriate controls. The graph represents mean ± S.D., n = 3. Statistical analysis was performed using Student's t test, *, p < 0.05. Data show immunoblot and densitometric analyses of total and phosphorylated AKT1 Ser-473 and total and phosphorylated AKT2 Ser-474 in C2C12 myotubes transduced with Ad-CMV-GFP or Ad-CMV-GFP-hBCKDK/HA (C), Ad-CMV-mCherry or Ad-CMV-mCherry-hBCKDHA/FLAG (D), or Ad-CMV-shGFP or Ad-CMV-shGFP-r/mBCKDHA (MOI 200) (E) for 48 h, followed by incubation with 100 nm insulin for 15 min. F, immunoblot and densitometric analyses of total and phosphorylated AKT1 Ser-473 levels in NRCMs transduced with either Ad-CMV-mCherry or Ad-CMV-mCherry-hBCKDHA/FLAG (MOI 200) or Ad-CMV-shGFP or Ad-CMV-shGFP-r/mBCKDHA (MOI 200) for 24 h, followed by 24-h serum starvation and incubation with 200 nm insulin for 20 min. Statistical analysis was performed using two-way ANOVA followed by Tukey's multiple-comparison test: *, p < 0.05; **, p < 0.01; ****, p < 0.0001, as indicated; *, −INS versus +INS groups; #, within +INS groups across different viral infection groups. Data are presented as mean ± S.D.

In addition to genetic manipulation of BCAA-catabolizing enzymes, we also employed pharmacological inhibition of BCKDK as an alternative approach to modulate intracellular BCKA levels. 3,6-Dichlorobenzothiophene-2-carboxylic acid (BT2) is a selective inhibitor of BCKDK (Fig. 6A), which increases BCKDH activity (20, 23) by inhibiting BCKDH phosphorylation. A 20-h exposure of C2C12 myotubes to BT2 modestly reduced intracellular BCKAs (Fig. S6A), albeit at high concentrations, signifying increased BCKDH activity, consistent with previous reports (20, 23). Moreover, BT2 markedly increased Klf15 and moderately increased Bcat2 mRNA levels (Fig. 7A), suggesting increased BCKA flux toward oxidation. BT2 also decreased BCKDE1α Ser-293–inactivating phosphorylation in a concentration-dependent manner (Fig. 7B). Moreover, BT2 increased insulin-mediated AKT phosphorylation in C2C12 cells (Fig. 7C) and phosphorylated AKT content, as measured by sandwich ELISA (Fig. S6B). Similar insulin-dependent effects of BT2 on AKT1 phosphorylation were also observed in NRCMs (Fig. 7D). We next questioned whether BT2 could rescue insulin signaling impairment in BCKDHA-depleted cells by compensatory action on other subunits of the BCKDH supercomplex. In BCKDHA-silenced C2C12 cells, BT2 treatment could not restore insulin-induced AKT phosphorylation (Fig. 7E). We observed that BT2 augments glucose transporter 1 (Glut1) but not Glut4 mRNA levels. (Fig. 7F). Similar to the effects of BT2, silencing BCKDK in C2C12 cells also increased Glut1, but not Glut4, and Klf15 mRNA expression (Fig. 7G). Silencing BCKDK also increased BCKDHA mRNA levels (Fig. 7H). Additionally, BT2 trended to increase insulin-induced PDH activity compared with vehicle-treated cells (Fig. S6B), supporting the view that BT2, by BCKDK inhibition, potentiates insulin's effect to activate PDH and stimulate muscle glucose oxidation (33–35).

Figure 7.

Pharmacological inhibition of BCKDK by BT2 improves muscle insulin signaling. A, qPCR analysis of BCAA catabolic genes Bckdha, Bckdk, Bcat2, Hadh (hydroxyacyl-coenzyme A dehydrogenase), Hibch, and Klf15 corrected to Rer1/Rpl7 reference gene levels in C2C12 myotubes treated with 500 and 750 μm BT2 for 20 h. B, Immunoblot analysis and densitometric quantification of total and phosphorylated BCKDH subunit E1 at Ser-293 in C2C12 myotubes treated with 160, 250, 320, 500, and 750 μm BT2 for 20 h. C, Immunoblotting and densitometric quantification of phosphorylated AKT1 Ser-473 and total AKT1 in differentiated C2C12 cells pretreated with 320, 500, and 750 μm BT2 for 20 h, followed by incubation with 100 nm insulin for 15 min. D, NRCMs were pretreated with 500 μm BT2 for 20 h, followed by 200 nm insulin incubation for 20 min. Data show immunoblot and densitometric analyses of phosphorylated BCKDE1α Ser-293 and total and phosphorylated AKT Ser-473. Statistical analysis was performed using two-way ANOVA followed by Tukey's multiple-comparison test: *, p < 0.05; **, p < 0.01; ****, p < 0.0001, as indicated; *, VEH versus +INS groups; #, VEH + INS versus BT2 + INS groups. E, immunoblot analysis and densitometric quantification of phosphorylated AKT1 Ser-473, AKT2 Ser-474, and total AKT1 and AKT2 in C2C12 myoblasts transduced with Ad-CMV-shGFP or Ad-CMV-shGFP-r/mBCKDHA (MOI 200) for 48 h, followed by incubation with 500 μm BT2 for 20 h and stimulation with 100 nm insulin for 15 min. Statistical analysis was performed using two-way ANOVA followed by Tukey's multiple-comparison test: *, p < 0.05; **, p < 0.01; ****, p < 0.0001, as indicated; *, −VEH versus +INS groups; #, shGFP + INS versus with shBCKDHA + INS group. F, qPCR analysis of Glut1 and Glut4 mRNA levels corrected to Rer1/Rpl7 reference gene levels in C2C12 myotubes treated with or without 500 μm BT2 for 20 h. G, qPCR analysis of BCAA catabolic genes Bckdha, Bckdk, and Klf15 and the glucose transporters Glut1 and Glut 4 corrected to Rer1/Rpl7 reference gene levels in C2C12 myotubes transduced with Ad-CMV-shGFP or Ad-CMV-shGFP-rBCKDK (MOI = 200) for 72 h. Statistical analysis was performed using one-way ANOVA followed by Tukey's multiple-comparison test: *, p < 0.05; **, p < 0.01; ****, p < 0.0001, as indicated. Data are presented as mean ± S.D.

Protein phosphatase 2A (PP2A) silencing rescues ketoleucine induced inhibition of insulin-mediated AKT 2 but not AKT1 phosphorylation

Because acute BCKA treatment impaired insulin signaling, we questioned whether BCKAs target upstream modulators of AKT phosphorylation. PP2A dephosphorylates and inactivates AKT (36), whereas insulin inhibits PP2A activity (37, 38), preventing AKT dephosphorylation and increasing AKT activity (38, 39). We examined whether KIC alters PP2A activity to modulate insulin signaling (Fig. 8A). Although KIC by itself had no effect on PP2A activity (data not shown), it prevented the inhibitory effect of insulin on PP2A activity, whereas pretreatment with the PP2A inhibitor, okadaic acid (OA), decreased KIC's effect (Fig. 8B). We next examined whether changes in PP2A activity corresponded with AKT phosphorylation. OA increased phosphorylation of both AKT isoforms, with a more significant effect observed for AKT1 Ser-473 (Fig. 8C). Ketoleucine exhibited a more significant inhibitory effect on AKT2 Ser-474 than AKT Ser-473 phosphorylation (Fig. 8C). Pretreatment with OA partially rescued AKT1 Ser-473 phosphorylation and significantly increased AKT2 Ser-474 phosphorylation (Fig. 8C). Interestingly, pretreatment of cells with KIC, followed by OA and insulin treatment, augmented AKT phosphorylation more than OA alone (data not shown). Given the off-target effects of OA, we sought to silence PP2A-c in C2C12 cells subjected to KIC and insulin treatment. PP2A silencing resulted in a significant increase in insulin-induced AKT1 and AKT2 phosphorylation (Fig. 8D). Treatment with KIC in PP2A-depleted cells restored insulin-induced AKT2 Ser-474 but not AKT1 Ser-473 phosphorylation (Fig. 8D). These data show that PP2A inactivation is sufficient to blunt KIC-induced inhibition of insulin-mediated AKT phosphorylation in an isoform-specific manner.

Figure 8.

Silencing PP2A partially rescues BCKA-induced insulin signaling impairment. A, outline of the treatment scheme used to test the effect of the PP2A inhibitor, OA, on KIC-mediated insulin signaling. B and C, PP2A activity expressed as percentage of mean control (B) and immunoblot and densitometric analyses of total to phosphorylated AKT1 Ser-473 and total to phosphorylated AKT2 Ser-474 in differentiated C2C12 myotubes pretreated with 250 nm OA for 30 min, followed by 5 mm KIC for 30 min and 100 nm insulin stimulation for 15 min (C). No insulin-treated cells were employed as controls. Other relevant groups are provided as an illustration in A. Statistical analysis was performed using one-way ANOVA followed by Tukey's multiple-comparison test: *, p < 0.05; **, p < 0.01; ****, p < 0.0001, as indicated; *, comparison with CON; #, comparison with +INS; ^, comparison of +KL + INS versus OA + KL + INS; @, +KL + INS versus OA + INS; +, comparison of +INS versus INS + OA + KIC; $, INS + OA versus INS + OA + KIC. D, C2C12 myotubes transfected with 20 nm siPP2A-c for 48 h, followed by 5 mm KIC treatment for 30 min and 100 nm insulin for 15 min. Data show immunoblot and densitometric analyses of total to phosphorylated AKT1 Ser-473 and total to phosphorylated AKT2 Ser-474. Statistical analysis was performed using two-way ANOVA followed by Tukey's multiple-comparison test: *, p < 0.05; **, p < 0.01; ****, p < 0.0001. Data are presented as mean ± S.D.

BCKAs up-regulate mTORC1 and protein translation signaling in skeletal muscle cells

BCAAs and BCKAs augment mTOR signaling (40–43) in different cell types (44, 45). We examined whether the effect of BCKAs on inhibiting the insulin signaling pathway was associated with increased mTOR signaling (Fig. S7A), which is known to negatively regulate insulin signaling (46). Phosphorylation of mTORC1 at Ser-2448 and its downstream target P70S6K at Thr-389 was significantly increased by BCKDHA silencing in C2C12 cells (Fig. 9A). Translation initiation is regulated by mTOR-dependent phosphorylation and activation of eukaryotic initiation factor 4G (eIF4G), at Ser-1108 (47). We observed increased eIF4G Ser-1108 phosphorylation in C2C12 cells depleted of BCKDHA (Fig. 9A). We next assessed the effect of individual BCKAs on mTOR signaling and protein translation. Increased phosphorylated mTOR, P70S6K, and eIF4G were observed with KIC and KMV but not KIV treatment (Fig. 9B). Increased mTORC1 signaling inhibits eukaryotic elongation factor 2 (eEF2) kinase (eEF2K) and promotes dephosphorylation and activation of eEF2 (48). Consistent with mTORC1 activation, KIC and KMV, but not KIV, down-regulated eEF2 phosphorylation at Thr-56 (Fig. 9B). To ascertain whether the increased mTOR signaling cascade corresponded with increased protein translation, we measured puromycin incorporation in nascent proteins in response to BCKAs. Puromycin incorporation in both C2C12 (Fig. 9C) and AMCMs (Fig. S7B) was increased by BCKAs. Together, our findings are in agreement with a recent study in adipocytes (23) and show that acute incubation with BCKAs (KIC and KMV) or altering endogenous BCKAs by silencing BCKDHA up-regulates mTORC1 activity and protein translation signaling. To decipher whether BCKA-induced insulin signaling impairment is mediated by mTORC1, we treated BCKDHA-depleted cells with the mTORC1 inhibitor, rapamycin (Fig. 10A). Rapamycin restored phosphorylated AKT1 and AKT2 levels in BCKDHA-silenced cells (Fig. 10A). Pretreatment with rapamycin also rescued BCKA's (0.05 and 5 mm) inhibition of insulin-mediated phosphorylation at AKT2 Ser-474 residue and modestly at AKT1 Ser-473 (Fig. 10B). Further, we observed more pronounced effects of rapamycin in the 0.05 mm BCKA group (Fig. 10B). Together, our findings suggest that BCKAs are sensitive to rapamycin, thereby highlighting the role of mTORC1 in BCKA-mediated inhibition of insulin signaling.

Figure 9.

BCKAs activate mTORC1 and downstream protein translation in skeletal muscle. A, Immunoblot analysis and densitometric quantification of total and phosphorylated mTORC1 Ser-2448, p70S6K Thr-389, and eIFG Ser-1108 in C2C12 myotubes transduced with Ad-CMV-shGFP or Ad-CMV-shGFP-r/mBCKDHA for 48 h, followed by stimulation with 100 nm insulin for 15 min. *, CON versus +INS groups; #, shGFP + INS versus shBCKDHA + INS groups. B, C2C12 myotubes treated with 5 mm KIC, KMV, or KIV or appropriate vehicle control for 45 min. Shown are immunoblot analysis and densitometric quantification of total and phosphorylated mTORC1 Ser-2448, p70S6K Thr-389, eIFG Ser-1108, and eEF2 Thr-56. Statistical analysis was performed using two-way ANOVA followed by Tukey's multiple-comparison test: *, p < 0.05; **, p < 0.01; ****, p < 0.0001, as indicated. *, CON versus KIC; #, CON versus KMV. C, C2C12 myotubes pretreated with 0.05 mm total BCKAs for 30 min, followed by incubation with 1 μm puromycin for 30 min. Immunoblot and densitometric analysis of puromycin incorporation, n = 3; Statistical analysis was performed using Student's t test, *, p < 0.05. Data are presented as mean ± S.D.

Figure 10.

Rapamycin rescues BCKA-mediated impairment of insulin signaling in C2C12 cells. A, C2C12 myotubes transduced with Ad-CMV-shGFP or Ad-CMV-shGFP-r/mBCKDHA (MOI = 200) treated with 100 nm rapamycin for 1 h, followed by 100 nm insulin stimulation for 15 min. Statistical analysis was performed using two-way ANOVA followed by Tukey's multiple-comparison test. *, p < 0.05; **, p < 0.01; ****, p <0.0001, as indicated. *, comparison between CON versus +INS groups; #, comparison of +INS versus INS + RAP group. B, C2C12 myotubes pretreated with 100 nm rapamycin for 1 h, followed by treatment with 0.05 mm or 5 mm total BCKAs for 30 min and 100 nm insulin for 15 min. D–E, Immunoblot and densitometric analyses of total to phosphorylated AKT1 Ser-473 and AKT2 Ser-474 and total to phosphorylated S6. Statistical analysis was performed using two-way ANOVA followed by Tukey's multiple-comparison test. *, p < 0.05; **, p < 0.01; ****, p < 0.0001, as indicated. *, comparison of CON versus +INS groups; #, VEH + INS versus BCKAs + INS groups; @, BCKA + INS versus BCKA + INS + RAP groups. C, graphical abstract. In in vivo and in vitro models of increased fatty acid availability, BCAA catabolic enzyme expression was suppressed in the heart (physiological fasting) in the gastrocnemius muscles (DIO) and in C2C12 cells treated with palmitate (a), resulting in increased accumulation of BCKAs either in circulation or intracellularly (b). Exogenous supply of BCKAs results in impairment of insulin-induced AKT phosphorylation by decreasing IRS1 phosphorylation (c). Altered accumulation of intracellular BCKAs by adenoviral overexpression of BCKDK or silencing BCKDH (d) or genetic and pharmacological activation of BCKDHA (e and f) impaired (d) or sensitized (e and f) skeletal and cardiac muscle cells to insulin-induced AKT phosphorylation at both isoforms. Effects of BCKAs on insulin signaling are mediated in an isoform-specific manner by PP2A, an upstream modulator of AKT (g). BCKAs' influence on insulin signaling can also be attributed to the activation of mTORC1 and increased protein translation machinery. Images were created using BioRender. Data are presented as mean ± S.D.

Discussion

Prior studies by utilizing genetic (1, 23, 49) and diet-induced rodent models of obesity and IR have either supported (1, 15, 50, 51) or challenged (1, 52–54) the paradigm that increased circulating BCAAs are a hallmark of IR, T2D, and obesity. The inconsistencies across different studies relating circulating BCAA levels to murine obesity, IR and metabolism are partly attributable to differential dietary intake, tissue uptake, and nutritional status. Coordinate and intertissue regulation of BCAA-metabolizing enzymes also contributes to the reported inconsistencies because there are contradictory findings on the impact of IR on BCAA catabolic enzyme expression in the skeletal muscle (23, 27, 55). Remarkably, elevations in circulating BCKAs correlated better with the severity of IR and T2D pathogenesis (28, 29). Elevated BCKAs is an outcome of impaired action of BCAA-catabolizing enzymes, BCKDHA (56, 57), PPM1K (58–60), and KLF15, the transcriptional activator of BCAA metabolism (1, 9). Altered enzyme expression within tissues capable of BCAA uptake and/or BCAA oxidation leads to systemic alterations in BCKA content. Significant amounts of BCKAs are generated within the skeletal muscle for interorgan transport, specifically to the liver and kidney (61). Moreover, it is plausible that loss of BCAA-catabolizing enzyme function can overwhelm skeletal muscle BCKA concentration and perturb downstream metabolism, such as insulin signaling and glucose disposal within the skeletal muscle (22). We investigated whether increasing BCKAs or targeting BCAA catabolic enzymes exerts effects on insulin signaling and function to govern muscle metabolism.

In our study, lipotoxicity decreased BCAA-catabolizing enzyme expression during DIO or palmitate exposure in the skeletal muscle. In contrast, physiological fasting post-translationally modified BCAA-metabolizing enzymes in the heart. Changes in BCAA-catabolizing enzyme expression altered circulating and intracellular BCKA content, signifying the physiological importance of BCAA catabolism. Indeed, rodents with obesity and IR show decreased expression of BCATm, PPM1K, and BCKDHA in adipose tissue (59, 62–64). Our findings are also in agreement with a prior study demonstrating decreased expression of BCAA oxidative genes in human and rodent muscles with IR (27). Evidently, excess lipids down-regulate BCAA-catabolizing enzymes, elevate intracellular BCKAs, and induce incomplete oxidation of BCKA. Because sequential oxidation of BCKAs contributes to cellular CoA pools, we postulated that limiting the oxidation of intracellular BCKAs triggers a likely reduction in BCAA-derived CoA content. Notably, in both in vivo and in vitro models of lipid overload, we observed a compensatory increase in Hmgcs1 mRNA levels, which supplies 3-hydroxy-3-methylglutaryl-CoA for Hmgcl, which replenishes CoA pools, in a feed-forward loop. Not only gene expression but inhibitory phosphorylation of BCKDH, a surrogate for BCKDH inactivity, was increased in the gastrocnemius muscle of DIO mice at 13 weeks, supporting prior studies demonstrating loss of BCKDH activity and BCAA oxidation in adipose, liver, and heart (56, 57). Notably, studies contradicting our findings show unaltered skeletal muscle BCAA catabolic genes in DIO mice (19), in leptin-deficient ob/ob mice (23), and in IR women with normoglycemia (55). Moreover, in a prior study, inhibitory phosphorylation of BCKDHA in the gastrocnemius muscles is unchanged in mice fed a 60% high-fat diet for 10 weeks (19). We attribute this inconsistency to difference in diet composition, duration postfeeding, and also the nutritional state of the animal before euthanasia. Our study reports the temporal effect of FA on BCAA catabolic enzyme expression and BCKA content in the muscle, both via HFHS feeding in vivo and exposure to palmitate in vitro. Our data are consistent with previous reports linking lipid metabolism to remodeling of muscle BCAA catabolism (2, 27, 65).

We and others have demonstrated that despite unchanged plasma BCAAs, systemic and tissue-specific BCKAs were elevated in obese animals (23) as well as in patients with obesity undergoing cardiac surgery (66). BCKAs are reported to alter carbohydrate metabolism in multiple insulin-responsive tissues (32, 67). BCKAs inhibit glucose uptake in cardiomyocytes (24) and skeletal muscle (68), and reducing BCKA levels by cardiac-specific BCATm deletion increases basal and insulin-induced glucose oxidation and ATP production (67). Moreover, in the liver and cultured hepatic cells, BCKAs up-regulate the glycogenolytic enzyme, glycogen phosphorylase (32). However, it was unclear whether BCKAs' effects on metabolism involved changes in insulin signaling. Moreover, a recent study demonstrated that BCAA restriction did not affect cardiac insulin signaling in Zucker fatty rats (69), indicating that downstream metabolites of BCAAs, such as BCKAs, are likely modifiers of insulin signaling. Interestingly, despite elevated levels of BCAAs, the BCAT2 knockout mouse unexpectedly developed resistance to diet-induced obesity and presented with increased glucose disposal, improved glucose tolerance, and enhanced insulin sensitivity (70), likely attributable to reduced BCKA levels. We demonstrated that supraphysiological and pathological levels (71) of exogenously supplied BCKAs impaired insulin signaling in the skeletal and cardiac muscle cells. Our data are consistent with a recent study in cultured 3T3-L1 adipocytes showing that BCKAs directly impair insulin signaling (23). In agreement with previous studies (24, 68), we also observed that KIC, KIV, and a combination of BCKAs suppress glucose uptake. Moreover, BCKA-mediated impairment of insulin-induced AKT phosphorylation was reversed by inhibiting PP2A. Whereas okadaic acid reversed KIC's effects on both of the AKT isoforms, PP2A-c depletion resulted in a selective restoration of phosphorylated AKT2 Ser-474 in KIC-treated cells. Because okadaic acid is also a potent inhibitor of PP4, PP5, and PP1, we cannot exclude the role of these phosphatases in altering KIC effects on AKT.

Prior studies have examined the direct effects of BCAAs and BCKAs but not the consequences of disrupted BCAA catabolic enzyme expression for cellular metabolism and insulin function. This study, for the first time, demonstrates that altering intracellular BCKA levels by overexpressing or silencing BCKDHA or BCKDK impacts insulin signaling both in the skeletal and cardiac muscle cells. Adenoviral silencing of BCKDHA or overexpression of BCKDK impaired AKT phosphorylation. Conversely, overexpression of BCKDHA or incubation with the BCKDK inhibitor, BT2, marginally decreased intracellular BCKA levels, which sensitized cardiac and skeletal muscle cells to insulin signaling. Our findings support the correlation between intracellular BCKA levels and insulin-mediated AKT phosphorylation in the C2C12 myotubes as well as neonatal and adult cardiomyocytes. Our ex vivo findings are consistent with the in vivo study wherein BT2 treatment in ob/ob mice improved insulin-induced AKT phosphorylation in the skeletal muscle but not in the liver or white adipose tissue (23). We theorize that overexpressing BCKDHA or exposure to BT2 will augment intracellular BCKA clearance and insulin signaling, when employed in a preexisting condition of metabolic stress. Interestingly, in our study, we find robust correlation of isoform-specific AKT changes, with BCKDHA expression levels. Previous studies reported that AKT2 Ser-474 phosphorylation is critical for maximal activation of AKT and insulin action (72). Indeed, a mixture of BCAAs/BCKAs attenuates AKT2 signaling in the hepatocytes in a mTORC1/C2-dependent manner (73). Nevertheless, our data indicate that lowering intracellular BCKA levels augments insulin signaling in the muscle. Even a modest reduction in intracellular BCKA levels by either BT2 treatment or BCKDHA overexpression sensitized insulin action, as reflected by increased PDH activity. Indeed, hearts with PPM1K deletion (18) and plasmodium with loss of BCKDH function (74) exhibited reduced PDH activity, leading to impaired glucose oxidation (75). In our study, increased PDH activity in BT2-treated C2C12 cells corresponded with the up-regulation of Glut1 mRNA expression, also observed upon BCKDK knockdown. BCKA levels are down-regulated in Glut1-overexpressing cardiomyocytes (76), suggesting that BCKA generation and utilization are intricately coupled to glucose metabolism via its effects on insulin signaling.

Muscle insulin signaling is inhibited by mTOR, P70S6K, and ribosomal S6, proteins activated by BCAAs (77). A key function of nutrient sensor mTOR is to maintain the available amino acid pool by regulating protein translation. Indeed, during DIO, glucolipotoxicity exacerbates IR with concomitant mTOR activation in the heart and skeletal muscle (78, 79). We demonstrate that the BCKDHA silencing and exposure to BCKAs activated mTORC1 signaling, with KIC and KMV exhibiting pronounced effects. Our data are in agreement with prior studies in cultured adipocytes (23) showing increased mTORC1 activity and defective insulin pathway in response to BCKA exposure (80). Pretreatment with the mTORC1 inhibitor, rapamycin, was able to rescue insulin signaling impairment induced by both BCKAs and BCKDHA depletion. mTOR activation increases protein translation and synthesis (81). A concerted action of eEF2, eEF2K, and eIF4G in conjunction with mTORC1 activation governs translation signaling (47, 48). In C2C12 and AMCMs, either depleting BCKDHA or treating with BCKAs up-regulated mTORC1 and protein translation signaling, as supported by puromycin incorporation studies. Excessive mitogenic signaling, including translation, can be an energetically expensive process compromising cellular survival. Indeed, cardiomyocytes incubated with BCKAs acutely down-regulated AKT signaling, triggering cell death (82). Supporting this paradigm, we and others demonstrate that BCKAs suppress mitochondrial oxygen consumption in AMCMs and cardiac (9), neuronal (31), and hepatic cells (32). We propose intracellular BCKAs as an anabolic sensor facilitating mTOR activation and also acting as an endogenous inhibitor of insulin signaling.

Taken together, we report that muscle BCAA-catabolizing enzyme expression and BCKA content are modulated by acute and chronic changes in insulin during physiological fasting and diet-induced obesity, as well as exposure to increasing concentrations of FAs. In an environment of chronic lipid overload down-regulation of BCAA catabolic enzyme expression accumulates BCKAs, causing IR. Cardiac and skeletal muscle cells depleted of BCKDHA or treated with exogenous BCKAs displayed defective insulin signaling with concomitant activation of mTORC1 and protein translation. BCKAs impaired glucose uptake and suppressed mitochondrial respiration. Additional studies are warranted to clarify whether the metabolic effects of BCKAs are a cause or effect of changes in insulin signaling, mitochondrial function, or protein translation. Moreover, augmenting BCKA clearance modestly by overexpressing BCKDHA or pharmacological targeting of BCKDK enhanced insulin signaling and glucose utilization. Our findings indicate that BCKAs can independently impair insulin signaling by modulating upstream effectors of AKT or through the activation of mTORC1 (Fig. 10C). Our study signifies that preferential cellular lipid metabolism in either physiological fasting or lipotoxicity involves two distinct processes: 1) down-regulation of BCAA-catabolizing enzymes and BCAA oxidation resulting in elevated BCKAs and 2) inhibition of insulin action and glucose utilization by BCKAs. This study advances the knowledge of the molecular nexus of BCAA metabolism and signaling with cellular insulin action.

Materials and methods

Cell lines and culture conditions

C2C12 cells were cultured in DMEM with 10% fetal bovine serum (FBS; Thermo Fisher Scientific). Differentiation was induced with 0.2% FBS for 4–6 days after cells reached 80% confluence. L6 cells were grown and maintained in α-minimal essential medium (Corning) containing 10% FBS. Differentiation was induced with 2% FBS for 4–6 days. Cells were grown to 70–80% confluence. All the cell lines were maintained at 37 °C in a humidified atmosphere of 5% CO₂.

NRCMs were isolated from 2-day-old Sprague–Dawley rat pups as described previously (79). Briefly, hearts were excised, and the ventricles were minced into small pieces and digested in several steps using collagenase-type 2 (2% (w/v); Worthington), DNase (0.5% (w/v); Worthington), and trypsin (2% (w/v); Worthington) with gentle stirring to dissociate heart pieces into single cells. Following digestion, nonmuscle cells and fibroblasts were eliminated using differential plating of the cell suspension for 2 h. Supernatant from differential plating, containing cardiomyocytes, was collected and suspended in DMEM/F-12 Ham's (Sigma) growth medium (containing 10% FBS, 10 μmol of cytosine-β-d-arabinofuranoside (ARAC; Sigma), insulin–transferrin–selenium (Corning), 1× antimycotic/antibiotic solution (Sigma)) and plated in primaria cell culture plates (Corning). The following day, cells were washed and cultured in serum-free DMEM no glucose medium (Gibco) (containing 10 μmol of ARAC, 50 mg/μl gentamycin, 1% penicillin–streptomycin, and 10 mm glucose) for 24 h.

ARCMs were isolated from adult male Sprague–Dawley rat hearts as described previously (79). Briefly, the Langendorff method was used for retrograde perfusion of the isolated heart with Tyrode buffer (containing 1.49 mm KCl, 0.33 mm KH₂PO₄, 11.69 mm NaCl, 12.51 mm taurine, 1.206 mm MgSO₄, 4.766 mm HEPES, 3.604 mm dextrose, and 0.396 mm l-carnitine at pH 7.4) in 5% CO₂, 95% O₂ at 37 °C. Following perfusion, the heart was digested with collagenase (60 mg of collagenase, 25 μm CaCl₂ in 75 ml of Tyrode buffer) for 30 min in a recirculating mode. The digested heart was excised, and the ventricular pieces were finely minced into a homogeneous solution. Incremental physiological concentrations of calcium (200 μm, 500 μm, and 1 mm) were used to make the ventricular myocytes calcium-tolerant, in the presence of 5% CO₂, 95% O₂. Dead cells were separated from the viable cardiomyocytes by gravity settlement. Cell pellet containing viable cardiomyocytes was resuspended in plating medium (containing medium 199 (Sigma), 26.2 mm NaHCO₃, 25 mm HEPES, 1.24 mm l-carnitine, 137 μm streptomycin (Sigma), 280.6 μm penicillin (Sigma), 10 mm taurine (Sigma), and 1% BSA Fraction V (Roche Applied Science) at pH 7.4) and seeded at a density of 50–75 × 10³ cells/plate on laminin (Roche Applied Science)-coated plates. Plating medium was replaced with fresh medium after 4 h, and cells were treated accordingly.

AMCMs were isolated from male C57BL/6 mice as described previously (79, 83). Briefly, the isolated hearts were perfused retrogradely by the noncirculating Langendorff method using the perfusion buffer (113 mm NaCl, 4.7 mm KCl, 0.6 mm KH₂PO₄, 0.6 mm Na₂HPO₄, 1.2 mm MgSO₄·7H2O, 12 mm NaHCO₃, 10 mm KHCO₃, 10 mm HEPES, 30 mm taurine, 10 mm butanedione monoxime, and 5.5 mm glucose, pH 7.4). The isolated ventricular myocytes were exposed to increasing concentrations of calcium (100, 400, and 900 μm) to render myocytes calcium-tolerant. Cardiomyocytes were plated on laminin-coated plates at a density of 40–60 × 10³ cells/plate and incubated at 37 °C. The medium was changed to fresh cardiomyocyte culture medium (minimum essential medium containing 0.1% BSA, 10 mm butanedione monoxime, 100 units/ml penicillin, 2 mm glutamine, and 2 mm ATP) after 3 h of plating.

C2C12 and L6 cells were incubated with serum-free DMEM low glucose, leucine-free medium (D9443, Sigma); NRCMs were incubated with serum-free DMEM no glucose medium; and ARCMs were incubated with serum-free medium 199 for 16 h before treating with 4-methyl 2-oxopentanoic acid sodium salt (KIC, W387101, Sigma), sodium-3-methyl-2-oxobutyrate (KIV, 198994, Sigma), 3-methyl-2-oxovaleric acid sodium salt (KMV, 198978, Sigma), or a combination of all three (BCKAs). BCKAs were prepared in 1:6 HCl/water solution. All the experiments were conducted with either 0.05 or 5 mm BCKAs (either individual or a mixture) for 30 min unless mentioned otherwise. Insulin resistance was induced in C2C12 cells, L6 cells, and cardiomyocytes by incubating them in their respective media containing 2% (w/v) fatty acid–free BSA (Roche Applied Science) and 0.4 mm sodium palmitate (Sigma) for 16 h. Myotubes were incubated with 2% fatty acid–free BSA in the absence of palmitate to mimic an insulin-sensitive state. To examine insulin signaling, cells were incubated with 100 nm insulin (C2C12, L6, ARCMs) or 200 nm insulin (NRCMs) or PBS for 15 min (C2C12, L6) or 20 min (NRCMs, ARCMs). For the BT2 experiments, C2C12 and NRCMs were pretreated for 20 h with the desired concentration of BT2 (Sigma and Matrix Scientific). BT2 from Sigma was dissolved in cremaphore solution, generously gifted by Ayappan Subbiah (Sevengenes Inc.). Cells were washed once and harvested in ice-cold PBS, followed by centrifugation at 18,000 × g for 10 min at 4 °C. Cell pellets were snap-frozen in liquid nitrogen and stored at −80 °C until further use.

Plasmids, transfections, and adenovirus

Adenoviral vectors expressing BCKDK/HA (ADV-202031, MOI 200), BCKDHA/FLAG (ADV-202026, MOI 200), BCKDHA shRNA (shADV-253870, MOI 200), and BCKDK shRNA (shADV-202031, MOI 250) were obtained from Vector Biolabs. The control vectors used were Ad-GFP, Ad-mCherry (catalog nos. 1060, 1767, and 1122, respectively), and scrambled shRNA GFP, obtained from Vector Biolabs. Cells were transfected with either pooled siRNA against mouse PP2A-Cα (sc-36302, Santa Cruz Biotechnology, Inc.) or siRNA negative control (4390844, Thermo Fisher Scientific) using Lipofectamine RNAiMAX (Invitrogen) according to the manufacturer's instructions with a concentration of 15 nm siRNA per plate.

Animal studies

C57BL6J mice were procured from the Jackson Laboratory (Bar Harbor, ME, USA). 10-Week-old male C57BL/6J mice were divided into three groups and either fed ad libitum or fasted for 16 h or refed for 4 h following fasting (n = 5 for each group). Body weight and blood glucose were measured before euthanasia, and liver and ventricle weight were measured after dissection. Gastrocnemius muscles and heart tissue were snap-frozen in liquid nitrogen and stored at −80 °C until further processing.

8-Week-old C57BL6J male mice were fed either chow or an HFHS diet and euthanized following overnight fast at 2, 4, 8, and 13 weeks (n = 5 for each group). Diet composition details are included in Table S2. Serum was collected after centrifuging the blood at 5000 rpm for 5 min. Gastrocnemius muscle and heart tissue were snap-frozen in liquid nitrogen and stored at −80 °C until further processing. All protocols involving mice were approved by the Dalhousie University Institutional Animal Care and Use Committee.

Tissue and cell lysate processing and immunoblotting

Frozen hearts and gastrocnemius muscle compartment from mice were powdered and homogenized using a tissue homogenizer (Omni TH, Omni International) in ice-cold lysis buffer (containing 20 mm Tris-HCl, pH 7.4, 5 mm EDTA, 10 mm Na₄P₂O₇ (Calbiochem), 100 mm NaF, 1% Nonidet P-40, 2 mm Na₃VO₄, protease inhibitor (10 μl/ml; Sigma), and phosphatase inhibitor (10 μl/ml; Calbiochem)). Homogenate was centrifuged at 1200 × g for 30 min, and the supernatant was collected for determining protein concentrations. Cell pellets were sonicated in ice-cold lysis buffer and centrifuged at 16,000 × g for 15 min. Protein concentrations of the cell and tissue lysates were determined using the BCA protein assay kit (Pierce, Thermo Fisher Scientific). Lysates were subjected to SDS-PAGE, and proteins were transferred onto a nitrocellulose membrane (Bio-Rad). Proteins were visualized using a reversible protein stain (Memcode, Pierce, Thermo Fisher Scientific), and membranes were incubated with the primary antibodies listed in Table S3. Immunoblots were developed using the Western Lightning Plus-ECL enhanced chemiluminescence substrate (PerkinElmer Life Sciences). Densitometric analysis was performed using Image laboratory software (Bio-Rad), and the quantifications were normalized by total protein loading using GraphPad software (Clarivate).

Immunoprecipitation

Immunoprecipitation of IRS1 was performed as described previously (84). Briefly, cells were harvested in radioimmunoprecipitation assay buffer (10 mmol/liter Tris-HCl (pH 8.0), 1 mmol/liter EDTA, 0.5 mmol/liter EGTA, 1% Triton X-100, 0.1% SDS, 140 mmol/liter NaCl with protease inhibitor, and phosphatase/inhibitor mixture). Then 350 μg of protein in a volume of 500 μl was precleared with 100 μl of 50% protein A–agarose bead slurry (ab193256, Abcam) for 1 h at 4 °C. This was followed by incubation with 4 μg of IRS1 antibody and 100 μl of 50% protein A–agarose bead slurry at 4 °C overnight. Immune complexes were pelleted and washed three times with radioimmunoprecipitation assay buffer and warmed at 95 °C for 5 min with 2× SDS sample buffer. Immunoprecipitated proteins were analyzed by Western blotting as mentioned above and probed for phosphorylated IRS Tyr-612 antibody.

SunSET method

Protein synthesis was measured in vitro by the SunSET assay as described previously (85, 86). Briefly, C2C12 myotubes or AMCMs were pretreated with 0.05 mm BCKAs for 30 min, followed by treatment with 1 μm puromycin dihydrochloride (P8833, Sigma) for 30 min prior to harvesting. Puromycin incorporation was detected by Western blotting using the monoclonal puromycin antibody. The primary antibody concentrations used for C2C12 and AMCMs were 1:5000 and 1:1000, respectively. Mouse secondary antibody concentrations used for C2C12 and AMCMs were 1:10,000 and 1:2500, respectively.

qPCR analysis

mRNA levels of BCAA-catabolizing enzymes and Glut-related gene expression were determined in tissues and cells using qPCR by employing validated optimal reference gene pairs as described previously (83). Primer information of the target and reference genes are provided in Table S4. Powdered tissue and cell pellets were homogenized in Ribozol (Amresco). RNA was isolated as per the manufacturer's instructions, and RNA quality and quantity were examined using a QIAxcel Advanced System (Qiagen). cDNA was synthesized from 1 μg of RNA using qScript cDNA supermix (Quanta Biosciences), and cDNA samples were stored at −30 °C until further use. qPCR analysis was performed in 96-well plates using PerfeCTa SYBR Green Supermix Low ROX (Thermo Fisher Scientific) and a ViiA7 real-time PCR machine (Thermo Fisher Scientific).

AKT activity assay

AKT activity was measured using the KinaseSTAR AKT activity assay kit (K435, Biovision) as per the manufacturer's instructions. Briefly, C2C12 cells were plated at a density of 2 × 10⁶ and treated either with 100 nm insulin alone for 15 min or with 5 mm KIC, KMV, and KIV for 30 min, followed by 100 nm insulin for 15 min. The cells were lysed using cold kinase extraction buffer and pelleted at 13,000 rpm for 10 min at 4 °C. One part of the cell lysate was used for assaying protein concentration. 200 μg of the cell lysate was incubated with 2 μl of AKT antibody, and AKT was immunoprecipitated using Protein A–Sepharose beads. A kinase assay was performed by incubating the AKT protein–bound beads with 2 μl of GSK3α/ATP mixture at 30 °C for 4 h. The beads were spun down, and supernatant was collected and boiled in 2× SDS-PAGE loading buffer separated by 12% SDS-PAGE and analyzed by Western blotting against phosphorylated GSK3α Ser-21.

Sandwich ELISA

AKT phosphorylated at Ser-473 was measured by sandwich ELISA using the DuoSet IC ELISA (DYC887B-2, R&D Systems) as per the manufacturer's protocol. Briefly, C2C12 cells were transduced with either Ad-CMV-shGFP or Ad-CMV-GFP-rm-shBCKDHA (MOI 200) for 48 h or treated with DMSO or 500 μm BT2 for 20 h, followed by 100 nm insulin treatment for 15 min. Cells were then solubilized in lysis buffer (1 mm EDTA, 0.5% Triton X-100, 5 mm NaF, 6 m urea, 1 mm activated Na₃VO₄, 2.5 mm Na₄P₂O₇, 10 μg/ml leupeptin, 10 μg/ml pepstatin, 100 μm phenylmethylsulfonyl fluoride, 3.0 μg/ml aprotinin in PBS, pH 7.2–7.4) at a concentration of 1 × 10⁷ cells/ml and centrifuged at 2000 × g for 5 min. One day prior to the assay, a 96-well high binding polystyrene plate (Greiner Bio-One) was coated with 6 μg/ml capture antibody (phosphorylated AKT Ser-473) diluted in 1× PBS overnight. For the assay, the supernatant and a seven-point standard curve (100–10,000 pg/ml) was diluted 6-fold with diluent 8 (1 mm EDTA, 0.5% Triton X-100, 5 mm NaF in PBS, pH 7.2–7.4) and further diluted in diluent 3 (1 mm EDTA, 0.5% Triton X-100, 5 mm NaF, 1 m urea in PBS, pH 7.2–7.4). On the day of the assay, the plate was washed three times with the wash buffer (0.05% Tween 20 in PBS, pH 7.2–7.4) and blocked at RT for 2 h with blocking buffer (1% BSA, 0.05% NaN₃ in PBS, pH 7.2–7.4). 100 μl of the sample or standard was added per well and incubated at RT for 2 h, followed by aspiration and washing. The samples and standards were then incubated with detection antibody (phosphorylated AKT Ser-473) conjugated with Streptavidin-HRP A at RT for 2 h. Colorimetric reagents were then added, and absorbance was measured immediately at 540 nm. The assay was normalized with protein concentration.

PP2A immunoprecipitation phosphatase assay

C2C12 cells were either treated with 100 nm insulin for 15 min or pretreated with 250 nm okadaic acid for 45 min, followed by 100 nm insulin for 15 min, or with 5 mm KIC for 30 min, followed by 100 nm insulin for 15 min, or with 250 nm okadaic acid for 45 min, followed by 5 mm KIC and 100 nm insulin for 15 min. Cells were lysed with lysis buffer (20 mm imidazole-HCl, 2 mm EDTA, 2 mm EGTA, pH 7.0, with 10 mg/ml each of aprotinin, leupeptin, pepstatin, 1 mm benzamidine, and 1 mm phenylmethylsulfonyl fluoride), and 300 μg of lysate was used to measure PP2A activity. Tissue was homogenized on ice and centrifuged at 12,000 rpm for 10 min at 4 °C. Cells were sonicated before centrifuging similarly. The supernatants were used to assay PP2A phosphatase activity by a standard kit (17-313, EMD Millipore) according to the manufacturer's instructions. The intensity of the color reaction was measured at 650 nm on a Bio-Rad microplate spectrophotometer.

Glucose uptake

2-Deoxyglucose (2-DG) uptake was measured in C2C12 cells using a colorimetric assay kit (ab136955, Abcam) as described previously (87). Briefly, C2C12 cells were plated in a 96-well plate and differentiated for 5 days. Myotubes were serum-starved for 12 h, followed by treatment with 5 mm KIC, KIV, KMV, BCKAs, or their vehicle controls for 30 min. Cells were incubated with glucose-free Krebs-Ringer-phosphate-HEPES buffer (20 mm HEPES, 5 mm KH₂PO₄, 1 mm MgSO₄, 1 mm CaCl₂, 136 mm NaCl, 4.7 mm KCl, pH 7.4) containing 2% BSA for 40 min. Cells were then incubated with or without 1 μm insulin for 20 min, followed by 10 μl of 10 mm 2-DG for 20 min. After adequate washes with 1× PBS, cells were harvested, and 2-DG absorbance was measured at 412 nm in the kinetic mode.

PDH activity assay

C2C12 myotubes were transduced with Ad-mCherry-hBCKDHA (MOI 120) or Ad-GFP-sh-r/mBCKDHA (MOI 200) and their respective controls for 48 h or treated with 500 μm BT2 or DMSO for 20 h, and PDH activity was measured using the PDH activity assay kit (K679, BioVision). Briefly, 1 × 10⁶ cells/condition were lysed with 100 μl of ice-cold PDH assay buffer and incubated on ice, followed by centrifugation at 10,000 rpm for 5 min. 50 μl of the supernatant was used and adjusted with 50 μl of PDH assay buffer, and the reaction was started with 50 μl of the reaction mixture (PDH assay buffer, PDH developer, and PDH substrate), and absorbance was measured immediately at 450 nm in a kinetic mode for 60 min at 37 °C using the Synergy plate reader. An NADH standard curve was used to calculate PDH activity in the samples in milliunits/ml.

Oxygen consumption

Oxygen consumption was measured using the Extracellular Oxygen Consumption Assay Kit (ab197243, Abcam) following the manufacturer's protocol. Briefly, AMCMs were seeded in a 96-well plate at a density of 2000 cells/well and treated with either 0.05 mm BCKAs or 5 mm BCKAs or their appropriate vehicle controls for 30 min. Following the treatment, the cells were incubated with an oxygen consumption reagent and covered with mineral oil. Quenching of the fluorescent probe by oxygen was measured by time-resolved fluorescence at excitation/emission of 360/630 in a Synergy H1 Hybrid reader.

ATP production

ATP levels were measured using the ATP determination kit (A22066, Invitrogen) as described previously (88). Briefly, AMCMs treated with 5 mm BCKAs or vehicle control were collected in the cell lysis buffer supplemented with protease and phosphatase inhibitors, sonicated, and centrifuged at 16,000 × g for 20 min at 4 °C. The lysate was used to determine the ATP content in terms of luminescence using the Synergy H1 Hybrid reader (BioTek). The assay was normalized to the amount of protein.

BCKA and BCAA measurements

Serum BCKA extraction

20 μl of the sample and 120 μl of internal standard (4 μg/ml in H₂O) containing leucine-d₃ (CDN Isotopes), 40 μl of MilliQ water, 60 μl of 4 m perchloric acid (VWR) were combined and vortexed. Proteins were precipitated in two sequential steps, followed by centrifugation at 13,000 rpm for 15 min at 4 °C. Supernatants collected from both steps were combined for measuring BCKAs.

Intracellular BCKA extraction

500,000 cells, 120 μl of internal standard (4 μg/ml in H₂O containing leucine-d₃ (CDN Isotopes, D-1973) and 0.8 ng/μl in H₂O containing sodium-2-keto-3-methyl-d₃-butyrate-3,4,4,4-d₄ (KIVd7; CDN Isotopes), 120 μl of 6 m perchloric acid (VWR) were combined and homogenized with a tissue homogenizer. Proteins were precipitated in two sequential steps, followed by centrifugation at 16,500 × g for 15 min at 4 °C. Supernatant collected from both steps was combined and split into two portions for measuring BCKAs. Samples were derivatized according to previously established protocols (89, 90).

BCKA derivatization and quantification

150 μl of extract and 500 μl of 25 mm OPD in 2 m HCl (made from o-phenylenediamine, 98%; VWR) were combined. The mixture was vortexed and then incubated at 80 °C for 20 min, followed by incubation on ice for 10 min. The derivatized extract was centrifuged at 500 × g for 15 min, and the supernatant was transferred to a tube containing 0.08 g of sodium sulfate (VWR) and 500 μl of ethyl acetate (VWR), following which they were centrifuged at 500 × g at RT for 15 min. This step was repeated twice, and the supernatants collected were vacuum-centrifuged at 30 °C for 45 min Samples were then reconstituted in 64 μl of 200 mm ammonium acetate (made from ammonium acetate, 98%; VWR) and transferred to amber glass UPLC vials (Waters). BCKAs were quantified with a Waters Acquity UPLC, Xevo-μTandem Mass Spectrometer and an Acquity UPLC BEH C18 (1.7 μm, 2.1 mm × 50 mm; Waters) and ACQBEHC18 VanGuard (130 Å, 1.7 μm, 2.1 × 5 mm; Waters) using multiple-reaction monitoring and internal standard calibration according to established protocol (66).

BCAA derivatization and quantification

10 μl of reconstituted extract was transferred to an autosampler vial and was combined with 70 μl of borate buffer (Waters) from the Waters AccQ-Tag derivatization kit (target pH: 8–10) and vortexed. 20 μl of AccQ-Tag Derivatization Agent (Waters, 186003836) was added, vortexed, and left standing for 1 min. Samples were derivatized (55 °C, 10 min) and vortexed. Derivatized samples were quantified with a Waters Acquity UPLC, Xevo-μ Tandem Mass Spectrometer and an AccQ-Tag Ultra RP Column 130 Å, 1.7-μm, 2.1-mm, 100-mm column using multiple-reaction monitoring and internal standard calibration (66).

Statistical analysis

Data are expressed as mean ± S.D. unless otherwise indicated. Statistical and Spearman's correlation analyses were conducted using Prism software (GraphPad, La Jolla, CA, USA). Comparisons between multiple groups were performed using two-way analysis of variance followed by a Tukey post hoc test or one-way ANOVA followed by a Tukey post hoc test, as appropriate. Linear regression statistical analysis was performed to assess significant correlations for normally distributed variables. p values of <0.05 were considered statistically significant.

Data availability

All data and materials used in the current study are available from the corresponding author upon request.