Taurine Administration Counteracts Aging-Associated Impingement Of Skeletal Muscle Regeneration By Reducing Inflammation And Oxidative Stress Part 2
Jun 12, 2023
3.4. Taurine Attenuates Oxidative Stress in TA Muscles of Aged Mice
Age-related sarcopenia is often associated with enhanced ROS production [5]. Taurine has been found at particularly high concentrations in tissues exposed to elevated levels of oxidants, suggesting a role in the attenuation of oxidative stress [40,60,61]. Thus, we evaluated whether the effect of taurine in skeletal muscle homeostasis of aged mice was correlated to the modulation of oxidative stress. To this purpose, we analyzed the levels of the Gp91phox protein, the catalytic subunit of the enzymatic complex nicotinamide adenine dinucleotide phosphate (NADPH) oxidase 2 (NOX2), responsible for the conversion of molecular oxygen to superoxide (O2 −) [62,63]. A significant increase in the Gp91phox protein level was observed in the muscles of the old mice compared to that present in the young mice (Figure 4A, B), highlighting an age-related generation of superoxide in older muscles. However, in the old mice treated with taurine, the expression of the Gp91phox protein returned to levels comparable to those of the young group. Another molecule involved in the maintenance of cellular levels of NADPH with both pro- and antioxidant activity is glucose-6-phosphate dehydrogenase (G6PD), whose altered level has been described as a consequence of NO signaling dysregulation [64,65]. We observed a significant increase in the G6PD protein in the TA muscles of the old mice compared to the young group, while the presence of high levels of taurine reduced G6PD to levels comparable to those of the young group (Figure 4A, C). These data suggest that taurine can counteract the deregulation of redox-related circuits, and consequently decreases NOX2-dependent ROS production.
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To confirm the role of taurine as an anti-oxidant molecule, using real-time PCR analysis, we further analyzed the expression levels of several antioxidant genes such as SOD1, CAT, and GPX1, known to undergo upregulation as a consequence of increased ROS production during aging [66]. As shown in Figure 4D–F, all the molecule expression levels were upregulated in the muscles of the old mice compared to those of the young group, but when the mice were treated with taurine, SOD1, CAT, and GPX1 expression was reduced, reaching a level comparable to what was found in the young mice-derived muscle extracts. Moreover, using Western blot analysis, we analyzed the levels of SOD in the different experimental groups (as indicated in the Figure), showing that the increased levels of SOD observed in the muscle extracts of the old mice were significantly reduced in the presence of taurine, confirming the results of the RT-PCR analysis (Figure 4G, H). These results demonstrate an important role played by taurine in the attenuation of the elevated level of oxidative stress that characterizes aged muscle. Consistently, taurine reduced ROS accumulation detected in the TA muscles of old mice (Supplemental Material) [67]. To investigate whether the abundance of ROS detected can induce oxidative modification of proteins, we performed an immunofluorescence analysis using 4-hydroxy-2-nonenal (4-HNE) adducts as markers of damage or alteration of muscle proteins due to oxidative stress [68]. Our results, shown in Figure 4I, J, demonstrated that 4-HNE expression was higher in the TA muscles of old mice, where slow MHC was also strongly downregulated; in the presence of taurine, 4-HNE expression was significantly decreased with a concomitant slow-MHC upregulation. These results are in agreement with our previous data (see Figure 3C, D) and strongly suggest taurine’s important role in the attenuation of ROS accumulation, preserving the slow-fiber phenotype during aging.

4. Discussion
In our previous studies, we demonstrated that taurine exerts a positive effect on myogenic differentiation and homeostasis in cell cultures [33]. Here, we investigated its effects in an in vivo experimental model. For this purpose, we used aged mice in which taurine was intraperitoneally injected every day for 5 weeks to assess the impact of taurine in the modulation of processes, such as regeneration, inflammation, and oxidative stress, which are known to be dysregulated during aging. We demonstrated that taurine accelerated the regenerative process of CTX-damaged TA muscles, preserving the architecture of skeletal muscle tissue. Indeed, 7 days after damage induction, in the presence of high levels of taurine, we observed a lower amount of inflammatory infiltration and fibrosis, and the regenerating fibers appeared larger compared to those of the vehicle-treated control muscles. This effect seems to be mediated by taurine-dependent stimulation of anabolic pathways, as demonstrated by the increased levels of phosphomTOR, rather than an effect on the modulation of catabolic processes; indeed, although the activation of other catabolic pathways cannot be excluded, ubiquitin ligase atrogin-1 is not significantly modulated by taurine. In general, skeletal muscle regeneration is guaranteed by the presence of satellite cells, whose number and activity significantly decrease during aging [69]. It has been demonstrated that the alteration of the immune response with aging, known as immunosenescence, is one of the main causes related to the hampered regenerative capability of skeletal muscle [70]. Indeed, immunosenescence promotes the development of a chronic low-grade inflammatory state, which may alter satellite cell proliferation and/or activity, thereby contributing to the impairment of the repairing capacity [69]. Thus, we verified whether the positive effect of taurine on skeletal muscle regeneration was mediated by the modulation of the inflammatory state. Here, we showed that the high number of macrophages present in old injured muscles was significantly decreased in the presence of taurine. This effect appeared to be mediated by NF-kB signaling since we showed that its elevated levels in CTX-injured muscles were decreased in the taurine-treated aged mice. These data are in agreement with what we have previously demonstrated in an in vitro experimental model [33] and are consistent with the role of taurine as an anti-inflammatory molecule exerting its effect, at least in part, through the inhibition of NF-kB activation [71]. In particular, it has been demonstrated that taurine may protect tissue damage from inflammation because its amino group can neutralize hypochlorous acid generated by inflammatory cells, downregulating the production of cytokines, and, finally, decreasing the immune response [72,73]. The chronic low-grade inflammatory state characterizing aged muscles may have a significant impact on the stimulation of catabolic pathways and mitochondrial dysfunctions, all contributing to the onset of sarcopenia [74]. In this context, the transcriptional co-activator PGC-1α appears to play a crucial role in skeletal muscle deterioration during aging. Indeed, it has been reported that PGC-1α plays a protective role in the inflammatory response, reducing proinflammatory cytokine production and exerting a regulatory mechanism for the expression of endogenous antioxidant proteins; moreover, it may improve muscle function, myofiber morphology, and integrity, suggesting its potential role in fiber repair and regeneration. Additionally, in cooperation with the MEF2C transcription factor, PGC-1α has been shown to regulate skeletal muscle fiber-type differentiation, promoting the switch from glycolytic fibers to more resistant oxidative ones [56,57]. Here, we showed that, in the absence of damage, no changes in PGC-1α levels and only a slight decrease in MEF2C levels were detected in TA muscle extracts of old mice compared to what was observed in young animals; however, their expression was significantly increased in the presence of taurine, reaching levels comparable to those found in the TA muscles of the young group. In addition, our results showed that taurine increases the levels of total MHC (MF20) and the slow-MHC and fast-MHC isoforms, suggesting its potential role in the metabolic shift of aged skeletal muscle fibers towards the oxidative, more resistant, phenotype [29]. These data reveal that the positive effect of taurine on skeletal muscle homeostasis of aged mice may be mediated by the stimulation of the PGC1-α/MEF2C pathway, favoring a possible metabolic shift of the myofibers towards the oxidative phenotype and preserving the more susceptible glycolytic fibers.

Taurine has been found at particularly high concentrations in tissues exposed to elevated levels of oxidants [40,75,76], and this prompted us to evaluate whether the observed positive effect of taurine on aged skeletal muscle homeostasis was related to the modulation of oxidative stress. A crucial mediator of ROS production in skeletal muscle tissue is the Gp91phox protein, which represents the catalytic subunit of the NOX2 complex and is also known to be overexpressed in dystrophic conditions [62,63,77–79]. Thus, we analyzed the Gp91phox protein in our experimental models, revealing that its level, while strongly upregulated in old mice compared to young ones, is significantly reduced in the presence of taurine. NOX2-dependent O2 − production is closely correlated with the availability of NADPH, although this substrate is also part of the antioxidant system contributing to the neutralization of ROS. In this context, one of the crucial enzymes involved in the maintenance of the cellular levels of NADPH is G6PD, which has pro- or antioxidant activity in skeletal muscle [65]. Here, we reported that the enhanced level of G6PD observed in old mice is significantly reduced in the presence of taurine, supporting the role of taurine as a potent modulator of NOX2-dependent ROS production in aged skeletal muscle. As a confirmation of this hypothesis, we showed that the accumulation of ROS in old muscle (see Supplemental Material) was strongly decreased by treatment with high doses of taurine. This effect was accompanied by the diminished formation of 4-HNE protein adducts, which are considered markers of lipid peroxidation and altered cellular redox homeostasis. We also showed that the endogenous antioxidant response in aged skeletal muscle is modulated in the presence of taurine, as revealed by the analysis of important anti-oxidant effectors such as SOD1, GPX1, and CAT. Indeed, the high levels of these molecules found in the TA muscle extracts of old mice were reduced upon taurine administration.
5. Conclusions
Collectively, our results show that, in aged muscle, taurine administration counteracts aging impingement of skeletal muscle regeneration, attenuates low levels of chronic inflammation, and decreases high levels of oxidative stress. Although the molecular mechanisms underlying these effects have not been completely elucidated, our data demonstrate that taurine administration ameliorates the microenvironment that allows the maintenance of skeletal muscle homeostasis and counteracts the aging process.

Supplementary Materials: Representative micrographs of TA cross-sections showing ROS levels assessed using CM-H2DCFDA and (B) quantification of fluorescence intensity. Statistical analysis was performed using one-way ANOVA multiple comparisons, *** p < 0.001, n = 3 mice per group.
Author Contributions: Conceptualization, B.M.S.; methodology, A.B., S.S., E.L., and B.M.S.; data analysis, A.B., E.L., and D.F.; validation, B.M.S., G.D., and G.S.; writing—original draft preparation, B.M.S.; critical review of the manuscript, D.F., G.S., L.T. and G.D.; funding acquisition, B.M.S. All authors have read and agreed to the published version of the manuscript.
Funding: This work was supported by Progetto di ricerca di interesse di Ateneo-Linea D.3.2, Anno 2015, Università Cattolica del Sacro Cuore to BMS. Università Cattolica del Sacro Cuore contributed to the funding of this research project and its publication.
Institutional Review Board Statement: The animal study protocol was approved by the Italian Ministry of Health (Ministero della Salute) (n. 150/2017-PR on the date 13 December 2017).
Acknowledgments: The authors thank Maria Teresa Viscomi for providing the 4-HNE antibody and Filippo Biamonte, Gabriella Proietti, and Francesca Forte for their technical support.

References
1. Cruz-Jentoft, A.J.; Bahat, G.; Bauer, J.; Boirie, Y.; Bruyère, O.; Cederholm, T.; Cooper, C.; Landi, F.; Rolland, Y.; Sayer, A.A.; et al. Sarcopenia: Revised European consensus on definition and diagnosis. Age Ageing 2019, 48, 16–31. [CrossRef] [PubMed]
2. Alway, S.E.; Myers, M.J.; Mohamed, J.S. Regulation of Satellite Cell Function in Sarcopenia. Front. Aging Neurosci. 2014, 6, 246. [CrossRef] [PubMed]
3. Snijders, T.; Parise, G. Role of muscle stem cells in sarcopenia. Curr. Opin. Clin. Nutr. Metab. Care 2017, 20, 186–190. [CrossRef] [PubMed]
4. Dalle, S.; Rossmeislova, L.; Koppo, K. The role of inflammation in age-related sarcopenia. Front. Physiol. 2017, 8, 1045. [CrossRef]
5. Meng, S.J.; Yu, L.J. Oxidative stress, molecular inflammation, and sarcopenia. Int. J. Mol. Sci. 2010, 11, 1509–1526. [CrossRef]
6. Marzetti, E.; Calvani, R.; Cesari, M.; Buford, T.W.; Lorenzi, M.; Behnke, B.J.; Leeuwenburgh, C. Mitochondrial dysfunction and sarcopenia of aging: From signaling pathways to clinical trials. Int. J. Biochem. Cell Biol. 2013, 45, 2288–2301. [CrossRef]
7. Ferri, E.; Marzetti, E.; Calvani, R.; Picca, A.; Cesari, M.; Arosio, B. Role of Age-Related Mitochondrial Dysfunction in Sarcopenia. Int. J. Mol. Sci. 2020, 21, 5236. [CrossRef]
8. Watt, F.M.; Hogan, B.L.M. Out of Eden: Stem cells and their niches. Science 2000, 287, 1427–1430. [CrossRef]
9. Bodensteiner, J.B.; Engel, A.G. Intracellular calcium accumulation in Duchenne dystrophy and other myopathies: A study of 567,000 muscle fibers in 114 biopsies. Neurology 1978, 28, 439–446. [CrossRef]
10. Poenie, M.; Epel, D. Ultrastructural localization of intracellular calcium stores by a new cytochemical method. J. Histochem. Cytochem. 1987, 35, 939–956. [CrossRef]
11. Tidball, J.G. Inflammatory processes in muscle injury and repair. Am. J. Physiol. Regul. Integr. Comp. Physiol. 2005, 288, 345–353. [CrossRef] [PubMed]
12. Tedesco, F.S.; Dellavalle, A.; Diaz-Manera, J.; Messina, G.; Cossu, G. Repairing skeletal muscle: Regenerative potential of skeletal muscle stem cells. J. Clin. Investig. 2010, 120, 11. [CrossRef]
13. Karpati, G.; Carpenter, S.; Prescott, S. Small-caliber skeletal muscle fibers do not suffer necrosis in mdx mouse dystrophy. Muscle Nerve 1988, 11, 795–803. [CrossRef] [PubMed]
14. Verdijk, L.B.; Snijders, T.; Drost, M.; Delhaas, T.; Kadi, F.; Van Loon, L.J.C. Satellite cells in human skeletal muscle; from birth to old age. Age 2014, 36, 545–557. [CrossRef]
15. Snijders, T.; Verdijk, L.B.; Smeets, J.S.J.; McKay, B.R.; Senden, J.M.G.; Hartgens, F.; Parise, G.; Greenhaff, P.; van Loon, L.J.C. The skeletal muscle satellite cell response to a single bout of resistance-type exercise is delayed with aging in men. Age 2014, 36, 1–5. [CrossRef] [PubMed]
16. Cosgrove, B.D.; Gilbert, P.M.; Porpiglia, E.; Mourkioti, F.; Lee, S.P.; Corbel, S.Y.; Llewellyn, M.E.; Delp, S.L.; Blau, H.M. Rejuvenation of the muscle stem cell population restores strength to injured aged muscles. Nat. Med. 2014, 20, 255–264. [CrossRef]
17. Cuthbertson, D.; Smith, K.; Babraj, J.; Leese, G.; Waddell, T.; Atherton, P.; Wackerhage, H.; Taylor, P.M.; Rennie, M.J. Anabolic signaling deficits underlie amino acid resistance of wasting, aging muscle. FASEB J. 2005, 19, 1–22. [CrossRef]
18. Altenhöfer, S.; Radermacher, K.A.; Kleikers, P.W.M.; Wingler, K.; Schmidt, H.H.H.W. Evolution of NADPH Oxidase Inhibitors: Selectivity and Mechanisms for Target Engagement. Antioxid. Redox Signal. 2015, 23, 406–427. [CrossRef]
19. Bua, E.; Johnson, J.; Herbst, A.; Delong, B.; McKenzie, D.; Salamat, S.; Aiken, J.M. Mitochondrial DNA-deletion mutations accumulate intracellularly to detrimental levels in aged human skeletal muscle fibers. Am. J. Hum. Genet. 2006, 79, 469–480. [CrossRef]
20. Tanhauser, S.M.; Laipis, P.J. Multiple deletions are detectable in mitochondrial DNA of aging mice. J. Biol. Chem. 1995, 270, 24769–24775. [CrossRef]
21. Joseph, A.M.; Adhihetty, P.J.; Leeuwenburgh, C. Beneficial effects of exercise on age-related mitochondrial dysfunction and oxidative stress in skeletal muscle. J. Physiol. 2016, 594, 5105–5123. [CrossRef]
22. Handschin, C.; Rhee, J.; Lin, J.; Tarr, P.T.; Spiegelman, B.M. An autoregulatory loop controls peroxisome proliferator-activated receptor gamma coactivator 1alpha expression in muscle. Proc. Natl. Acad. Sci. USA 2003, 100, 7111–7116. [CrossRef] [PubMed]
23. Puigserver, P.; Spiegelman, B.M. Peroxisome proliferator-activated receptor-gamma coactivator 1 alpha (PGC-1 alpha): Transcriptional coactivator and metabolic regulator. Endocr. Rev. 2003, 24, 78–90. [CrossRef] [PubMed]
24. Knutti, D.; Kralli, A. PGC-1, a versatile coactivator. Trends Endocrinol. Metab. 2001, 12, 360–365. [CrossRef]
25. Russell, A. PGC-1alpha and exercise: Important partners in combating insulin resistance. Curr. Diabetes Rev. 2005, 1, 175–181. [CrossRef] [PubMed]
26. Kelly, D.P.; Scarpulla, R.C. Transcriptional regulatory circuits controlling mitochondrial biogenesis and function. Genes Dev. 2004, 18, 357–368. [CrossRef] [PubMed]
27. Lin, J.; Handschin, C.; Spiegelman, B.M. Metabolic control through the PGC-1 family of transcription coactivators. Cell Metab. 2005, 1, 361–370. [CrossRef]
28. Ventura-Clapier, R.; Garnier, A.; Veksler, V. Transcriptional control of mitochondrial biogenesis: The central role of PGC-1alpha. Cardiovasc. Res. 2008, 79, 208–217. [CrossRef]
29. Wang, Y.; Pessin, J.E. Mechanisms for fiber-type specificity of skeletal muscle atrophy. Curr. Opin. Clin. Nutr. Metab. Care 2013, 16, 243–250. [CrossRef]
30. Da Boit, M.; Sibson, R.; Meakin, J.R.; Aspden, R.M.; Thies, F.; Mangoni, A.A.; Gray, S.R. Sex differences in the response to resistance exercise training in older people. Physiol. Rep. 2016, 4, e12834. [CrossRef]
31. Bamman, M.M.; Hill, V.J.; Adams, G.R.; Haddad, F.; Wetzstein, C.J.; Gower, B.A.; Ahmed, A.; Hunter, G.R. Gender differences in resistance-training-induced myofiber hypertrophy among older adults. J. Gerontol. Ser. A Biol. Sci. Med. Sci. 2003, 58, 108–116. [CrossRef] [PubMed]
32. Safdar, A.; Hamadeh, M.J.; Kaczor, J.J.; Raha, S.; deBeer, J.; Tarnopolsky, M.A. Aberrant mitochondrial homeostasis in the skeletal muscle of sedentary older adults. PLoS ONE 2010, 5, e10778. [CrossRef] [PubMed]
33. Barbiera, A.; Sorrentino, S.; Lepore, E.; Carfì, A.; Sica, G.; Dobrowolny, G.; Scicchitano, B.M. Taurine Attenuates Catabolic Processes Related to the Onset of Sarcopenia. Int. J. Mol. Sci. 2020, 21, 8865. [CrossRef] [PubMed]
34. Rogeri, P.S.; Zanella, R.; Martins, G.L.; Garcia, M.D.A.; Leite, G.; Lugaresi, R.; Gasparini, S.O.; Sperandio, G.A.; Ferreira, L.H.B.; Souza-junior, T.P.; et al. Strategies to Prevent Sarcopenia in the Aging Process: Role of Protein Intake and Exercise. Nutrients 2021, 14, 52. [CrossRef]
35. Børsheim, E.; Bui, Q.U.T.; Tissier, S.; Kobayashi, H.; Ferrando, A.A.; Wolfe, R.R. Effect of amino acid supplementation on muscle mass, strength and physical function in elderly. Clin. Nutr. 2008, 27, 189–195. [CrossRef]
36. Dillon, E.L.; Sheffield-Moore, M.; Paddon-Jones, D.; Gilkison, C.; Sanford, A.P.; Casperson, S.L.; Jiang, J.; Chinkes, D.L.; Urban, R.J. Amino acid supplementation increases lean body mass, basal muscle protein synthesis, and insulin-like growth factor-I expression in older women. J. Clin. Endocrinol. Metab. 2009, 94, 1630–1637. [CrossRef]
37. Katsanos, C.S.; Kobayashi, H.; Sheffield-Moore, M.; Aarsland, A.; Wolfe, R.R. Aging is associated with diminished accretion of muscle proteins after the ingestion of a small bolus of essential amino acids. Am. J. Clin. Nutr. 2005, 82, 1065–1073. [CrossRef]
38. Paddon-Jones, D.; Sheffield-Moore, M.; Zhang, X.J.; Volpi, E.; Wolf, S.E.; Aarsland, A.; Ferrando, A.A.; Wolfe, R.R. Amino acid ingestion improves muscle protein synthesis in the young and elderly. Am. J. Physiol. Endocrinol. Metab. 2004, 286, E321–E328. [CrossRef]
39. Tieland, M.; van de Rest, O.; Dirks, M.L.; van der Zwaluw, N.; Mensink, M.; van Loon, L.J.C.; de Groot, L.C.P.G.M. Protein Supplementation Improves Physical Performance in Frail Elderly People: A Randomized, Double-Blind, Placebo-Controlled Trial. J. Am. Med. Dir. Assoc. 2012, 13, 720–726. [CrossRef]
40. Scicchitano, B.M.; Sica, G. The Beneficial Effects of Taurine to Counteract Sarcopenia. Curr. Protein Pept. Sci. 2018, 19, 673–680. [CrossRef]
41. Schaffer, S.W.; Azuma, J.; Mozaffari, M. Role of antioxidant activity of taurine in diabetes. Can. J. Physiol. Pharmacol. 2009, 87, 91–99. [CrossRef] [PubMed]
42. Pierno, S.; De Luca, A.; Camerino, C.; Huxtable, R.J.; Camerino, D.C. Chronic Administration of Taurine to Aged Rats Improves the Electrical and Contractile Properties of Skeletal Muscle Fibers. J. Pharmacol. Exp. Ther. 1998, 286, 1183–1190. [PubMed]
43. Gomez, R.; Caletti, G.; Arbo, B.D.; Hoefel, A.L.; Schneider, R.; Hansen, A.W.; Pulcinelli, R.R.; Freese, L.; Bandiera, S.; Kucharski, L.C.; et al. Acute intraperitoneal administration of taurine decreases glycemia and reduces food intake in type 1 diabetic rats. Biomed. Pharmacother. 2018, 103, 1028–1034. [CrossRef] [PubMed]
44. Luo, H.; Geng, C.J.; Miao, S.M.; Wang, L.H.; Li, Q. Taurine attenuates the damage of lupus nephritis mouse via inactivation of the NF-κB pathway. Ann. Palliat. Med. 2021, 10, 137–147. [CrossRef] [PubMed]
45. Aïnad-Tabet, S.; Grar, H.; Haddi, A.; Negaoui, H.; Guermat, A.; Kheroua, O.; Saïdi, D. Taurine administration prevents the intestine from the damage induced by beta-lactoglobulin sensitization in a murine model of food allergy. Allergol. Immunopathol. 2019, 47, 214–220. [CrossRef]
46. Caletti, G.; Herrmann, A.P.; Pulcinelli, R.R.; Steffens, L.; Morás, A.M.; Vianna, P.; Chies, J.A.B.; Moura, D.J.; Barros, H.M.T.; Gomez, R. Taurine counteracts the neurotoxic effects of streptozotocin-induced diabetes in rats. Amino Acids 2018, 50, 95–104. [CrossRef]
47. Caletti, G.; Olguins, D.B.; Pedrollo, E.F.; Barros, H.M.T.; Gomez, R. Antidepressant effect of taurine in diabetic rats. Amino Acids 2012, 43, 1525–1533. [CrossRef]
48. Costa, A.; Toschi, A.; Murfuni, I.; Pelosi, L.; Sica, G.; Adamo, S.; Scicchitano, B.M. Local overexpression of V1a-vasopressin receptor enhances regeneration in tumor necrosis factor-induced muscle atrophy. Biomed. Res. Int. 2014, 2014, 235426. [CrossRef]
49. Guardiola, O.; Andolfi, G.; Tirone, M.; Iavarone, F.; Brunelli, S.; Minchiotti, G. Induction of Acute Skeletal Muscle Regeneration by Cardiotoxin Injection. J. Vis. Exp. 2017, 119, 54515. [CrossRef]
50. Fischer, A.H.; Jacobson, K.A.; Rose, J.; Zeller, R. Hematoxylin and eosin staining of tissue and cell sections. CSH Protoc. 2008, 2008, pdb.prot4986. [CrossRef]
51. Franceschi, C.; Bonafè, M.; Valensin, S.; Olivieri, F.; De Luca, M.; Ottaviani, E.; De Benedictis, G. Inflamm-aging. An evolutionary perspective on immunosenescence. Ann. N. Y. Acad. Sci. 2000, 908, 244–254. [CrossRef]
52. Saito, Y.; Chikenji, T.S. Diverse Roles of Cellular Senescence in Skeletal Muscle Inflammation, Regeneration, and Therapeutics. Front. Pharmacol. 2021, 12, 1–13. [CrossRef] [PubMed]
53. Bakkar, N.; Guttridge, D.C. NF-κB signaling: A tale of two pathways in skeletal myogenesis. Physiol. Rev. 2010, 90, 495–511. [CrossRef] [PubMed]
54. Kumar, A.; Takada, Y.; Boriek, A.M.; Aggarwal, B.B. Nuclear factor-κB: Its role in health and disease. J. Mol. Med. 2004, 82, 434–448. [CrossRef] [PubMed]
55. Thoma, A.; Lightfoot, A.P. Nf-kb and inflammatory cytokine signaling: Role in skeletal muscle atrophy. In Advances in Experimental Medicine and Biology; Springer: Berlin/Heidelberg, Germany, 2018; Volume 1088, pp. 267–279.
56. Lin, J.; Wu, H.; Tarr, P.T.; Zhang, C.Y.; Wu, Z.; Boss, O.; Michael, L.F.; Puigserver, P.; Isotani, E.; Olson, E.N.; et al. Transcriptional co-activator PGC-1α drives the formation of slow-twitch muscle fibers. Nature 2002, 418, 797–801. [CrossRef] [PubMed]
57. Sandri, M.; Lin, J.; Handschin, C.; Yang, W.; Arany, Z.P.; Lecker, S.H.; Goldberg, A.L.; Spiegelman, B.M. PGC-1α protects skeletal muscle from atrophy by suppressing FoxO3 action and atrophy-specific gene transcription. Proc. Natl. Acad. Sci. USA 2006, 103, 16260–16265. [CrossRef]
58. Calabria, E.; Ciciliot, S.; Moretti, I.; Garcia, M.; Picard, A.; Dyar, K.A.; Pallafacchina, G.; Tothova, J.; Schiaffino, S.; Murgia, M. NFAT isoforms control activity-dependent muscle fiber type specification. Proc. Natl. Acad. Sci. USA 2009, 106, 13335–13340. [CrossRef]
59. Wilkinson, D.J.; Piasecki, M.; Atherton, P.J. The age-related loss of skeletal muscle mass and function: Measurement and physiology of muscle fiber atrophy and muscle fiber loss in humans. Ageing Res. Rev. 2018, 47, 123–132. [CrossRef]
60. Marcinkiewicz, J.; Kontny, E. Taurine and inflammatory diseases. Amino Acids 2014, 46, 7. [CrossRef]
61. Oliveira, M.W.S.; Minotto, J.B.; de Oliveira, M.R.; Zanotto-Filho, A.; Behr, G.A.; Rocha, R.F.; Moreira, J.C.F.; Klamt, F. Scavenging and antioxidant potential of physiological taurine concentrations against different reactive oxygen/nitrogen species. Pharmacol. Rep. 2010, 62, 185–193. [CrossRef]
62. Ferreira, L.F.; Laitano, O. Regulation of NADPH oxidases in skeletal muscle. Free Radic. Biol. Med. 2016, 98, 18–28. [CrossRef] [PubMed]
63. Whitehead, N.P.; Yeung, E.W.; Froehner, S.C.; Allen, D.G. Skeletal muscle NADPH oxidase is increased and triggers stretch-induced damage in the mdx mouse. PLoS ONE 2010, 5, e15354. [CrossRef] [PubMed]
64. Stanton, R.C. Glucose-6-phosphate dehydrogenase, NADPH, and cell survival. IUBMB Life 2012, 64, 362–369. [CrossRef] [PubMed]
65. Cacchiarelli, D.; Martone, J.; Girardi, E.; Cesana, M.; Incitti, T.; Morlando, M.; Nicoletti, C.; Santini, T.; Sthandier, O.; Barberi, L.; et al. MicroRNAs Involved in Molecular Circuitries Relevant for the Duchenne Muscular Dystrophy Pathogenesis Are Controlled by the Dystrophin/nNOS Pathway. Cell Metab. 2010, 12, 341–351. [CrossRef]
66. Kozakowska, M.; Pietraszek-Gremplewicz, K.; Jozkowicz, A.; Dulak, J. The role of oxidative stress in skeletal muscle injury and regeneration: Focus on antioxidant enzymes. J. Muscle Res. Cell Motil. 2015, 36, 377–393. [CrossRef] [PubMed]
67. King, N.; McGivan, J.D.; Griffiths, E.J.; Halestrap, A.P.; Suleiman, M.S. Glutamate loading protects freshly isolated and perfused adult cardiomyocytes against intracellular ROS generation. J. Mol. Cell. Cardiol. 2003, 35, 975–984. [CrossRef]
68. Eckl, P.M.; Ortner, A.; Esterbauer, H. Genotoxic properties of 4-hydroxyalkenals and analogous aldehydes. Mutat. Res. 1993, 290, 183–192. [CrossRef]
69. Domingues-Faria, C.; Vasson, M.P.; Goncalves-Mendes, N.; Boirie, Y.; Walrand, S. Skeletal muscle regeneration and impact of aging and nutrition. Ageing Res. Rev. 2016, 26, 22–36. [CrossRef]
70. Shaw, A.C.; Goldstein, D.R.; Montgomery, R.R. Age-dependent dysregulation of innate immunity. Nat. Rev. Immunol. 2013, 13, 875–887. [CrossRef]
71. Barua, M.; Liu, Y.; Quinn, M.R. Taurine Chloramine Inhibits Inducible Nitric Oxide Synthase and TNF-α Gene Expression in Activated Alveolar Macrophages: Decreased NF-κB Activation and IκB Kinase Activity. J. Immunol. 2001, 167, 2275–2281. [CrossRef]
72. Kim, C.; Jang, J.S.; Cho, M.R.; Agarawal, S.R.; Cha, Y.N. Taurine chloramine induces heme oxygenase-1 expression via Nrf2 activation in murine macrophages. Int. Immunopharmacol. 2010, 10, 440–446. [CrossRef] [PubMed]
73. Schuller-Levis, G.B.; Park, E. Taurine: New implications for an old amino acid. FEMS Microbiol. Lett. 2003, 226, 195–202. [CrossRef]
74. Ji, L.L.; Kang, C. Role of PGC-1α in sarcopenia: Etiology and potential intervention—A mini-review. Gerontology 2015, 61, 139–148. [CrossRef] [PubMed]
75. Green, T.R.; Fellman, J.H.; Eicher, A.L.; Pratt, K.L. Antioxidant role and subcellular location of hypotaurine and taurine in human neutrophils. Biochim. Biophys. Acta BBA-Gen. Subj. 1991, 1073, 91–97. [CrossRef]
76. Schaffer, S.W.; Ju Jong, C.; Kc, R.; Azuma, J. Physiological roles of taurine in heart and muscle. J. Biomed. Sci. 2010, 17, 1–8. [CrossRef]
77. Kim, J.H.; Kwak, H.B.; Thompson, L.V.; Lawler, J.M. Contribution of oxidative stress to pathology in diaphragm and limb muscles with Duchenne muscular dystrophy. J. Muscle Res. Cell Motil. 2013, 34, 1–13. [CrossRef]
78. Pelosi, L.; Forcina, L.; Nicoletti, C.; Scicchitano, B.M.; Musarò, A. Increased Circulating Levels of Interleukin-6 Induce Perturbation in Redox-Regulated Signaling Cascades in Muscle of Dystrophic Mice. Oxid. Med. Cell. Longev. 2017, 2017, 1987218. [CrossRef]
79. Petrillo, S.; Pelosi, L.; Piemonte, F.; Travaglini, L.; Forcina, L.; Catteruccia, M.; Petrini, S.; Verardo, M.; D’Amico, A.; Musaró, A.; et al. Oxidative stress in Duchenne muscular dystrophy: Focus on the NRF2 redox pathway. Hum. Mol. Genet. 2017, 26, 2781–2790. [CrossRef]
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