NMR-Based Metabolomic Analysis For The Effects Of α-Ketoglutarate Supplementation On C2C12 Myoblasts in Different Energy States Ⅱ

3. Discussion 

Myoblasts are the primary source of muscle repair and regeneration in damaged myofibers. The process of muscle regeneration involves the activation, proliferation, and differentiation of myoblasts, during which proliferation and differentiation occur simultaneously sometimes [22,23]. Concerning the mass and functions of muscle decline, supplementation of new myoblasts is a potential therapeutic strategy for repairing muscle injuries [24]. Therefore, it is vital to identify valid factors to promote the growth of myoblasts during the process of skeletal muscle regeneration, which may be beneficial to the development of ‘regenerative medicine’ for treating muscular diseases [25,26]. AKG has been recognized as a potential nutrition supplement for skeletal muscle. However, AKG-induced metabolic changes in myoblasts in different energy states remain to be systematically clarified. In the present work, we applied the conditions of normal culture and low-glucose culture to mimic the two states of normal energy and energy-defficiency, respectively, and performed the cellular metabolomic analysis to exploit the effects of AKG supplementation on metabolic profiles of aqueous extracts derived from the mouse myoblast cell line C2C12. Our work reveals signifificant beneficial effects of AKG supplementation for promoting the proliferation and differentiation of myoblasts, indicating the crucial role of AKG in enhancing the functions of skeletal muscle.

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3.1. AKG Promotes Proliferation and Differentiation of Myoblasts 

We found that AKG supplementation distinctly promoted the proliferation of myoblasts. In the two different energy states, myoblasts fused with each other to form myotubes, which became longer and thicker after AKG supplementation, indicating that AKG also promoted the myotube differentiation of myoblasts. Furthermore, under both normal culture and low-glucose culture conditions, the myoblasts exhibited enhanced expressions of MyOD1 acting as a key regulatory protein in the process of myoblast differentiation to form myotubes [27]. This result also verified the effect of AKG on promoting skeletal muscle differentiation. Even though previous studies have demonstrated that AKG can enable C2C12 myotubes to become thicker [10] or promote muscle hypertrophy [18], our study on C2C12 myoblasts provides a new perspective to mechanistically understanding the beneficial effects of AKG supplements for improving functions of skeletal muscle in the two states of normal energy and energy deficiency. 

3.2. AKG Improves Glutamate and Glutamine Metabolism 

Even though AKG supplementation dramatically raised intracellular AKG levels in both Nor-A and Low-A myoblasts, this work did not focus on AKG transport since it was not a rate-limiting step involved in the AKG metabolism [28]. As a product of catabolism related to proteins and other nitrogen-containing compounds, ammonia can damage cell functions [29]. As a precursor of glutamate and glutamine, AKG can react with ammonia to form glutamate, which further reacts with ammonia to form glutamine [30]. Glutamine can serve as a carbon source to generate energy and regulate the activity of signal transduction pathways, thereby promoting cell proliferation [31,32]. As shown in Table S2, AKG supplementation signifificantly up-regulated the glutamine level in myoblasts under normal culture conditions, although it did not signifificantly change the glutamate level. Additionally, myoblasts under low-glucose culture conditions showed a strengthened dependence on glutamine in the lack of carbon sources, and profoundly enhanced intracellular levels of glutamate and glutamine acting as characteristic metabolites between Low and Nor cells. In fact, in the absence of glutamine breakdown, increasing the intracellular AKG level would also activate the mTORC1-related pathway to promote cellular proliferation [11]. Signifificantly, Low-A cells displayed a declined level of glutamine, implying that AKG supplement reduced the demand for glutamine in skeletal muscle cells in the state of energy efficiency with a high glutamine level. These results reveal that AKG supplementation signifificantly improves glutamate and glutamine metabolism, thereby promoting the proliferation of myoblasts cultured in both normal and low-glucose media. 

3.3. AKG Promotes Antioxidant Capacity of Myoblasts under Low-Glucose Culture Condition 

Under normal culture conditions, AKG supplementation decreased cellular levels of methionine, glutathione, and glycine, implying that the glutathione metabolism was signifificantly down-regulated. As known, dietary supplementation with AKG can enhance the CAT and SOD expressions under oxidative stress [33]. We found that Nor-A cells did not show statistically signifificant differences in the CAT and SOD expressions and total antioxidant capacity from Nor cells, indicating that AKG supplementation did not signifificantly alter cellular oxidative stress. Compared to Nor cells, Low cells displayed a down-regulated CAT expression and a decreased total antioxidant capacity. Notably, AKG supplementation restored the total antioxidant capacity of myoblasts cultured in a low-glucose medium. Furthermore, Low-A cells displayed a signifificantly increased level of glycine relative to Low cells, indicating that AKG supplementation could favor the activation of cellular antioxidants in a state of energy deficiency, thereby protecting cells and maintaining cell growth and metabolism under adverse conditions. Additionally, it is known that taurine acts as an antioxidant [34,35]. In this study, Low cells exhibited a profoundly elevated intracellular level of taurine compared to Nor cells, which potentially enhanced the antioxidant ability of myoblasts under low-glucose culture conditions. Signifificantly, AKG supplementation further raised the intracellular level of taurine, and correspondingly enhanced the antioxidant ability of cells without sufficient energy supply. It has been reported that AKG has an antioxidative function and exhibits a vital role in scavenging ROS in organisms [36]. Here, we also demonstrated that AKG supplement signifificantly promotes the antioxidant capacity of myoblasts under low-glucose culture conditions. 

3.4. AKG Enhances Cellular Energy Status 

Acting as a source of energy, AKG can provide energy for cell processes and regulates cellular energy mechanisms. When cells were cultured in a normal medium with sufficient glucose, AKG supplementation increased intracellular levels of the upstream glucose and lactate, thereby potentially preserving the cell energy source. Compared with NorA cells, Nor-A cells showed a decreased level of phosphocreatine (PCr) and a basically unchanged level of creatine (Cr). PCr and Cr can be converted to each other through the reaction catalyzed by creatine kinase (CK). Note that both PCr and Cr are the two critical components of the CK/PCr system serving as the rapidly available source for ATP synthesis in skeletal muscle [37]. The decreased phosphocreatine level together with the increased ATP content in Nor-A cells, implied that the AKG supplement facilitated the conversion of PCr to Cr for ATP synthase in myoblasts in the state of normal energy.

On the other hand, Low cells displayed a decreased ATP content and a remarkably decreased PCr, as well as a dramatically increased Cr level compared with Nor cells, implying that more PCr was converted to Cr to meet energy demands in cells cultured in a low-glucose medium. Signifificantly, AKG supplement increased the ATP content by about one time in myoblasts in the state of energy deficiency as shown in Figure 7F. Unexpectedly, Low-A cells showed a higher ATP level than Nor cells, potentially owing to both an optimal phenotype of cells achieved at a moderate glucose level and AKG supplementation [38]. Nevertheless, the underlying molecular mechanisms should be explored in detail in the future. 

Low-A cells exhibited reduced phosphorylation of AMPK as indicated by the distinctly declined ratio of p-AMPK to AMPK relative to Low cells, indicating that AKG supplementation enhanced the antioxidant effect and improved the proliferation and metabolism of myoblasts in the state of energy efficiency. The underlying molecular mechanism also remains to be addressed in the future. Under normal culture conditions, AKG supplementation distinctly increased the intracellular ATP level, but did not signifificantly change the ratio of p-AMPK to AMPK, potentially due to the need to maintain a certain energy state in cells with sufficient energy supply. As is known, an up-regulated ATP level favors energy utilization and physiological activities of the organism such as cell proliferation and differentiation. However, it seems that several previous studies suggested apparently contradictory molecular mechanisms for further understanding the beneficial effects of AKG supplements. For example, one study suggested that AKG extends Drosophila lifespan by inhibiting mTOR and activating AMPK [39], while another study suggested that AKG activates mTOR signaling and promotes the ATP synthase in Lipopolysaccharide-challenged piglets [40]. Potentially, these differences in AKG-related mechanisms might be attributed to specifific strains and genetic backgrounds relevant to the detailed experiments. Further work must be conducted to clarify the molecular mechanisms underlying the beneficial effects of AKG in different energy states. 

In addition, we only performed metabolic profiling on aqueous metabolites extracted from C2C12 myoblasts. Analyzing the portion of hydrophobic metabolites in conjunction with that of aqueous metabolites would provide a much more comprehensive metabolic profile. Unfortunately, hydrophobic metabolites are usually related to poor NMR spectra with declined spectral resolution and decreased S/N ratio due to crowded, overlapped, or broadened peaks, especially peaks from lipid metabolites. It is difficult to accurately calculate integrals of most hydrophobic metabolites based on poor NMR spectra. Further efforts should be made in the future to improve the NMR spectra of hydrophobic metabolites.

4. Materials and Methods

4.1. Cell Culture

For obtaining C2C12 myoblasts, C2C12 cells were firstly cultured in normal DMEM to reach 50% confluence for 24 h, and then cultured in fresh normal DMEM with or without AKG supplementation (Nor-A GM, Nor GM), or in fresh low-glucose DMEM with or without AKG supplementation (Low-A GM, Low GM) for another 24 h (Figure S2A). The final concentration of AKG was 2 mm. Correspondingly, the obtained myoblasts were classifified into the following four groups: Nor-A, Nor, Low-A, and Low myoblasts. 

For obtaining C2C12 myotubes, C2C12 cells were firstly cultured in normal DM (the differentiation medium was replaced every two days) to form myotubes for 10 days, and then cultured in fresh normal DM with or without AKG supplementation (Nor-A DM, Nor DM), or in fresh low-glucose DM with or without AKG supplementation (Low-A DM, Low DM) for another 2 days (Figure S2B). Similarly, the obtained myotubes were also classified into the following four groups: Nor-A, Nor, Low-A, and Low myotubes. 

4.2. MTS Cell Proliferation Assay and Morphologies of Myoblasts and Myotubes

 C2C12 myoblast cells were seeded at a density of 5 × 103 cells per well in 96-well plates for 24 h by 100 µL of the medium. Then, the culture medium was replaced by 100µL of fresh medium, and the cells were incubated for another 24 h. Equivalent volumes of vehicle culture media were treated as controls. CellTiter 96 AQueous solution (MTS, Promega, Madison, WI, USA) was added to each well, and the absorbance of formazan at a wavelength of 490 nm on a microplate reader (BioTek, Winooski, VT, USA) after incubation in the dark for 3 h. In addition, C2C12 myoblasts or myotubes were washed three times using PBS to remove the dead cells. Thereafter, cell morphological images were taken randomly on a fluorescence microscope (Motic, Xiamen, China). 

4.3. Western Blotting

C2C12 myoblast cells were lysed in a RIPA buffer (Sangon Biotech, Shanghai, China) containing protease and phosphatase inhibitors, followed by brief sonication. Cell lysates were then loaded into sodium dodecyl sulfate-polyacrylamide gel, and, thereafter, transferred onto PVDF membranes (GE, Freiburg, Germany). Membranes were blocked with 5% non-fat milk and incubated with primary antibodies overnight at 4 ◦C with shaking. After incubation with the secondary antibody for 1 h at room temperature, the signal was visualized by the commercially enhanced chemiluminescence reagent (ECL, Beyotime, Shanghai, China). The used antibodies were as follows: GAPDH (Proteintech, Wuhan, China), MyoD1 (Santa Cruz Biotechnology, Dallas, TX, USA), CAT (Proteintech, Wuhan, China), SOD (Proteintech, Wuhan, China), p-AMPK (CST, Boston, MA, USA), AMPK (CST, Boston, MA, USA). 

4.4. Intracellular Metabolite Extraction and Samples Preparation 

Aqueous metabolites were extracted from C2C12 myoblast cells for NMR analyses according to the protocol described previously [41]. Cells were quickly rinsed thrice by cold phosphate-buffered saline (PBS, pH 7.4) to reduce the residual medium. The residual PBS was removed immediately by vacuum suction. Subsequently, metabolic activities of cells were aborted by methanol, and cells were scraped off by a cell scraper (Costar, Washington, DC, USA). Cells were then collected into a 15 mL centrifuge tube. Thereafter, methanol, chloroform, and water in the volume ratio of 4:4:2.85 were applied in a dual-phase extraction for extracting intracellular metabolites. Only the polar phase was lyophilized and subjected to metabolomic profiling. The aqueous cell extract powder was resolved in 550 µL of phosphate buffer [50 mm, pH 7.4, 100% D2O, 0.05 mm sodium 3-(trimethylsilyl) propionate-2,2,3,3-d4 (TSP)], vortexed and then centrifuged at 12,000× g for 15 min at 4 ◦C. The supernatants were transferred into 5 mm NMR tubes for NMR-based metabolomic analysis.

4.5. NMR Measurements and Data Preprocessing

Phase correction, baseline correction, and resonance alignment were carried out for all 1D NMR spectra using the MestReNova 9.0 software (Mestrelab Research S.L., Santiago de Compostela, Spain). For further multivariate statistical analysis, 1D 1H spectral region of 9.5–0.6 ppm was segmented into bins with a width of 0.01 ppm. The water region of 4.9–4.7 ppm was excluded to eliminate distortion from the residual water resonance in all 1D spectra. Peak integrals of the segments were normalized by the sum of peak integrals to compensate for potential variations in the concentrations of samples. The sum of the peak integrals was set to 100 for each spectrum. The normalized integrals were used to represent the relative levels of assigned metabolites. For pairwise comparisons of metabolite levels between the groups, singlet or nonoverlapped resonances in each NMR spectrum were selected for computing metabolite integrals. 

4.6. Resonance Assignments of Metabolites 

Two AKG resonance regions of 2.495–2.530 ppm and 2.996–3.019 ppm were excluded to compress metabolic differences between the groups. NMR resonances of metabolites were assigned using a combination of the Chenomx NMR Suite (version 8.6, Chenomx Inc., Edmonton, AB, Canada) and the Human Metabolome Database (HMDB, ca/ accessed on 6 January 2021). In addition, 2D 1H-13C HSQC and 1H-1H TOCSY spectra were used to confirm the assigned metabolites. 

4.7. Metabolomic Analysis 

Multivariate statistical analysis was performed on 1D 1H-NMR spectral data of C2C12 cell extracts by using the SIMCA-P software (version 12.0.1, Umetrics, Umea, Sweden). Pareto scaling was applied to the normalized NMR spectral data to enhance the significance of low-level metabolites without noise enlargement. Then, principal component analysis (PCA) was conducted to examine grouping trends and reveal metabolic differences. Moreover, orthogonal partial least-squares discriminant analysis (OPLS-DA) was conducted to check grouping trends and improve group separation. Cross-validation was performed to measure the robustness of the OPLS-DA model with a response permutation test (200 times). The reliability of the OPLS-DA model was raised as the R2 and Q2 approached 1. Important metabolites were identified with VIP > 1 from the OPLS-DA model. 

Univariate data analysis was conducted on the relative levels of the assigned metabolites between the groups, which were calculated based on the integrals of the metabolites relative to the sum of metabolite integrals. We quantitatively compared the relative levels of the metabolites between the groups using a two-tailed Student’s t-test with the Graphpad Prism software (version 6.0, La Jolla, San Diego, CA, USA). Data were expressed as the mean ± SD. Metabolites with p < 0.05 were identified to be differential metabolites. Characteristic metabolites were determined by a combination of the differential metabolites and important metabolites described above.

The metabolic pathway analysis was performed on the MetaboAnalyst 5.0 webserver (https://www.metaboanalyst.ca accessed on 6 January 2021), using a combination of metabolite sets enrichment analysis (p < 0.05) and pathway topological analysis (pathway impact value > 0.2). Signifificantly altered metabolic pathways were identified for pairwise comparisons of Nor-A vs. Nor, Low vs. Nor, Low-A vs. Low based on the relative levels of the assigned metabolites. 

4.8. Measurement of Cellular Total Antioxidant Capacity (T-AOC)

4.9. Measurement of Intracellular ATP Content

After 2.4 mL of lysis buffer was added to each dish of cells, lysates were collected into an Eppendorf tube. Eppendorf tubes were spun in the centrifuge for 5 min at 12,000× g.The ATP content was measured by the luciferase method using the ATP assay kit according to the manufacturer’s instructions. The protein concentration in each sample was detected by the BCA protein assay (Beyotime, Shanghai, China) to calculate the ATP concentration per mg protein. 

5. Conclusions

Sample Availability: Samples of the NMR experiments are available from the authors. 

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