RONS and Myokines in Skeletal Muscle Glucose Uptake: History
View Latest Version
Please note this is an old version of this entry, which may differ significantly from the current revision.
Contributor:
Paola Llanos
,
Jesus Palomero

The skeletal muscle is the largest organ in the body that performs different functions, including those related to the movement of the body such as stability, equilibrium, and locomotion; vital functions such as breathing; and those associated with the maintenance of metabolic homeostasis, in which the generation and expenditure of energy and heat production are critical. The adequate interplay of these functions leads to the maintenance of life in organisms. Glucose is essential in metabolism since it is one of the main substrates that produces ATP, the key molecule that transfers energy during chemical reactions in organisms. To produce ATP, glucose needs to be transported from the extracellular space into the cytosol of the cell. This process is called glucose uptake, and it is critical in skeletal muscle since it provides enough glucose to the cell to produce ATP and satisfy the high demand for energy of the skeletal muscle. Glucose uptake in skeletal muscle tissue is a process mainly regulated by insulin, which is a hormone synthesized in the pancreas and released into the blood stream, where it is transported until it binds to specific insulin receptors that are anchored at the plasma membrane of skeletal muscle cells.

  • ROS
  • RNS
  • nitric oxide
  • myokines
  • cytokines
  • glucose transport
  • GLUT4

1. Introduction

Skeletal muscle is mainly composed of fibers, which are postmitotic multinuclear cells formed from the fusion of single cells, known as myoblasts [1][2]. Muscle fibers continuously generate reactive oxygen and nitrogen species (RONS), and the production of RONS may be augmented in different situations, such as during contractile activity, inflammation, regeneration, and the aging of skeletal muscle [3]. When RONS are generated in excess and the intracellular antioxidant systems are unable to neutralize these highly reactive molecules, the skeletal muscle is exposed to oxidative distress and the RONS react with cellular macromolecules, such as lipids, proteins, carbohydrates, and nucleic acids, producing irreversible changes in those molecules and compromising the viability, integrity, and function of the cells [4]. However, when the level of intracellular RONS is moderate or relatively low, i.e., oxidative eustress, some of that RONS may act as signaling molecules, mediators or second messengers, which, through reversible redox reactions with specific residues of macromolecules, may modulate and regulate cellular signaling pathways that drive different cellular processes [4]. Over the last two decades, researchers in the field of redox biology have claimed and pointed out the importance of investigation and improving our knowledge regarding the function of RONS participating as signaling molecules in different pathways and how they are involved in the modulation, regulation, and control of cellular functions. In addition to the general physiological control of glucose uptake in skeletal muscle led by insulin, it has been proposed that RONS may be involved in the regulation of glucose uptake in skeletal muscle. However, it is uncertain whether this process might be independent or dependent of the effect of insulin [5][6][7]. Consequently, there is a need to investigate the role of RONS in the glucose uptake process in skeletal muscle. This knowledge may help us to find new targets for the treatment of the impairment of glucose uptake and insulin resistance manifested in type 2 diabetes, obesity, and aging.
Skeletal muscle is considered a secretory organ and liberates different cytokines, peptides, proteins, or hormones known as myokines that may have endocrine and auto/paracrine effects [8]. Myokines, such as myostatin, irisin and IL-6, among others, induce changes in the muscle itself, as well as in other organs and tissues [9]. These peptides, proteins or hormones regulate or alter the functionality and adaptation of muscle tissue, modifying its metabolism and other functions including hypertrophy or muscular atrophy, angiogenesis, and inflammatory processes [10]. Myokines may interfere with the prevention of obesity, metabolic syndrome, and type 2 diabetes. Consequently, there is an important focus in biomedical research to evaluate the potential of myokines as new therapeutic targets [9]. It appears that either the auto/paracrine effect of different myokines or the endocrine effect of different hormones, cytokines and factors that interact with skeletal muscle may affect glucose uptake and insulin resistance. However, the molecular signaling pathways and the physiological impact of these molecules remain largely unexplored.

2. Reactive Oxygen and Nitrogen Species in Skeletal Muscle

Around 1954, the first report was published that indicated that skeletal muscle produces free radical intermediates [11]. Twenty years later, two studies indicated that exercise was related to the generation of products from different reactions that occur between free radicals, referred to today as RONS, and cellular biomolecules [12][13]. These findings were consolidated in the 1980s when it was demonstrated that intense exercise produces free radicals in skeletal muscle [14]. Studies in the 1990s uncovered the fact that the contractile activity of skeletal muscle produced specific reactive species. Thus, it was demonstrated that superoxide was released from the diaphragm when this muscle contracted [15], nitric oxide was released from skeletal muscle [16], and hydroxyl radicals were generated in skeletal muscle during contractions [17]. The previous studies and other derived investigations have built upon this knowledge and created the current scenario that describes the generation of reactive oxygen and nitrogen species in skeletal muscle. For more detail, the reviews of [3][18][19][20] may be consulted.
Basically, the main ROS generated in skeletal muscle cells (and cells from other tissues) are free radical species, such as superoxide anion (O2·−) and hydroxyl radical (HO·), and others considered nonradical species, such as hydrogen peroxide (H2O2). All of these are considered reactive oxygen species (ROS) since they originate from a reduction of oxygen and the reactive atom is oxygen. In the case of reactive nitrogen species (RNS), which are reactive species generated from nitrogen, the most relevant are nitric oxide (NO·), which is a free radical species, and peroxynitrite anion (ONOO-), which is a nonradical species, although very reactive [19]. Superoxide and nitric oxide are the primary species generated in skeletal muscle fibers and lead to the formation of other secondary species such as hydrogen peroxide, hydroxyl radical and peroxynitrite, all of which may be generated in the intracellular and extracellular space, in the mitochondrial matrix, and in the mitochondrial intermembrane space. The contractile activity of skeletal muscle fibers evokes an increase in the intracellular generation of superoxide, hydrogen peroxide and nitric oxide [21][22][23][24]. In addition, contractile activity increases the production of superoxide, hydrogen peroxide, hydroxyl radicals and nitric oxide extracellularly in the muscle interstitial space [25][26]. Most of the studies indicated that the increase in ROS generation caused by contractions in skeletal muscle was attributed to the increase in mitochondrial respiration due to the high demand for energy production in the mitochondrial matrix by the electron transport chain and ATP synthase, which conduct the formation of ATP, which is the source of energy for contractile activity. Therefore, mitochondria were attributed as the main source of ROS production in skeletal muscle. However, later studies indicated a lower increase in ROS and discovered other nonmitochondrial sources for ROS generation [19]. Thus, NADPH oxidases were identified in the skeletal muscle plasma membrane [27], the sarcoplasmic reticulum [28], and T-tubules [29]. The product of NADPH oxidase activity is superoxide, and it is known from the later studies that NADPH oxidases contribute substantially to the production of superoxide and, potentially, hydrogen peroxide in skeletal muscle [30]. There are other sources of superoxide production that are less studied that may contribute to intracellular superoxide production in skeletal muscle, such as the enzyme phospholipase A2 [31]. In addition, the extracellular release of superoxide driven by a lipoxygenase was identified in skeletal muscle [32]. Another enzyme that may contribute to superoxide generation in the extracellular space of skeletal muscle is the xanthine oxidase located in the endothelial cells associated with skeletal muscle. It has been reported that this enzyme increases the superoxide concentration in the extracellular fluid of skeletal muscle during contractile activity [33].
Nitric oxide, the primary reactive nitrogen species (RNS), is generated enzymatically through nitric oxide synthase (NOS). NOS catalyzes the conversion of the amino acid L-arginine into L-citrulline, with cofactors NADPH and oxygen, and generates nitric oxide [19][20]. There are three NOS isoenzymes: neuronal (nNOS), endothelial (eNOS) and inducible (iNOS). nNOS is expressed in neurons and skeletal muscle, eNOS is expressed mainly in endothelial cells and in skeletal muscle with less expression, and iNOS appears to be associated with inflammatory processes that, in some circumstances, might affect the skeletal muscle [23]. Balon and Nadler were the first to identify the release of NO from skeletal muscle, which was augmented by contractile activity [16]. Other studies reported that skeletal muscle releases nitric oxide to the extracellular space, although they were uncertain which cells (i.e., skeletal muscle fibers, lymphocytes, fibroblasts, and endothelial cells) originate NO and the contribution they make to the total extracellular NO concentration [34][35]. Further work uncovered the release of NO specifically from skeletal muscle cells, such as myotubes [21][26]. In addition, it was indicated that the contractive activity of skeletal muscle produced an increase in the intracellular concentration of NO in a model of myotubes of a rat skeletal muscle origin [21]. Further studies, using the model of isolated single muscle fibers, demonstrated that contractile activity induced in fibers by electrical stimulation produced a rapid increase in the intracellular concentration of NO, which also rapidly decreased when contractile activity ceased [23]. Moreover, studies performed with a model of skeletal muscle fibers subjected to a protocol of passive elongation showed that the passive stretch applied to fibers isolated from old mice produced an increase in nitric oxide intracellular activity [36].
Hydrogen peroxide, hydroxyl radical and peroxynitrite are reactive species derived from the primary RONS, superoxide and nitric oxide, that play a relevant role in oxidative distress [37]. In skeletal muscle cells and other tissues, hydrogen peroxide is enzymatically generated in the cytosol, the mitochondrial matrix, and the extracellular space. Hydrogen peroxide is generated by the dismutation of superoxide in a reaction that is catalyzed by the enzyme superoxide dismutase (SOD). There are SOD isoenzymes that are expressed in different subcellular compartments. Manganese-SOD (Mn-SOD) is expressed in the mitochondrial matrix, copper/zinc-SOD (Cu/Zn-SOD) is expressed in the cytosol and mitochondrial intermembrane space, and there is an extracellular SOD (ec-SOD) that appears in the interstitial fluid in skeletal muscle [19][20]. Therefore, hydrogen peroxide is generated at different subcellular locations. When H2O2 concentration is high, it may appear oxidative distress in the cell. However, a lower H2O2 concentration leads to oxidative eustress, which is crucial for regulating and maintaining cellular processes through the modulation of different signaling pathways in the cell [4][37]. H2O2 may produce another secondary reactive species, such as hydroxyl radicals (HO·). This may occur when H2O2 is in the presence of divalent metals, mainly iron (Fe++), and H2O2 is decomposed to HO· by the Fenton reaction. The hydroxyl radical is a strong oxidant that is not diffusible through membranes and reacts with every close molecule. This produces alterations in macromolecules (lipids, proteins, and nucleic acids), which may disturb cellular homeostasis and other functions [19][20]. In other words, the hydroxyl radical mainly evokes oxidative distress. Peroxynitrite (ONOO-) is produced in a reaction between nitric oxide and superoxide [38]. This reaction is three times faster compared with the reaction of superoxide dismutation that produces hydrogen peroxide. Therefore, this is the primary reaction when both superoxide and hydrogen peroxide appear close together. Peroxynitrite shows a high oxidizing power and is very reactive; moreover, it may oxidize thiol groups of peptides, damage DNA and cause the nitration of proteins [19][20]. Hence, an abundance of peroxynitrite may potentially evoke oxidative distress.
Concomitant to RONS generation, cells are equipped with an antioxidant system that neutralizes the activity of reactive molecules. Therefore, there is a balance between the production and elimination of RONS to maintain the level of RONS at a steady state, facilitating homeostasis and cellular function. A moderate increase in the level of ROS is critical to trigger and modulate cellular signaling pathways that control several cellular functions. As mentioned above, this state is referred to as oxidative eustress [4]. However, when the level of RONS is elevated, if there is an excess of RONS being produced or the antioxidant system is unable to neutralize the RONS produced, then these reactive species interact with and oxidize other molecules (lipids, proteins, DNA, and carbohydrates), causing oxidative damage to them and possibly affecting their normal functions, which, ultimately, could compromise cellular homeostasis and viability. This state, already mentioned above, is currently referred to as oxidative distress in redox biology [4], in place of the classical term of oxidative stress.
The cellular antioxidant system is constituted of both an enzymatic and nonenzymatic system. The main antioxidant enzymes are superoxide dismutase (SOD), which catalyzes the conversion of superoxide anions into hydrogen peroxide; glutathione peroxidase (GPx), which neutralizes hydrogen peroxide to convert it to water and appears at different subcellular locations, particularly the mitochondria and cytosol; and catalase (CAT), which transforms hydrogen peroxide to water. In addition, there are accessory proteins that protect cells against oxidation, such as peroxiredoxin (PRX), glutaredoxin (GRX), and thioredoxin reductase (TRX). For more details, other comprehensive reviews in the field, i.e., [19], may be consulted. The nonenzymatic antioxidant system of the cell involves compounds such as glutathione, vitamins C and E, carotenoids, polyphenols, alpha-lipoic acid, coenzyme Q10, uric acid and bilirubin. These compounds are involved in thiol–disulfide exchange reactions and play a role in maintaining the redox balance in the cell [3][19]. Glutathione (GSH) is the most abundant nonenzymatic antioxidant in the cell, and it plays an important role due to its high reductive power. Glutathione reacts with different RONS, transferring protons and neutralizing these reactive species. Furthermore, glutathione is the substrate for the antioxidant enzyme GPx, and participates in the recycling of other antioxidant enzymes, such as TRX and GRX. Moreover, glutathione leads to the reduction of other antioxidants such as vitamins E and C [19]. Glutathione is critical in contractile activity and in the adaptation of skeletal muscle to exercise. This has been an important matter of investigation in the field of redox biology. It is out of the scope here, but readers may refer to other comprehensive key publications that describe the crucial role of glutathione in the redox biology of skeletal muscle. i.e., [39][40][41].
To recapitulate, RONS are generated in skeletal muscle at rest and during contractile activity. A high level of RONS might cause the excessive oxidation of macromolecules, which leads to cellular damage and dysfunction. However, a moderate RONS level is essential to modulate redox signaling processes that regulate cellular homeostasis and adaptation. The term RONS refers to several specific species. Some of them are mainly prone to induce oxidative distress, such as hydroxyl radicals and peroxynitrite. However, other RONS appear to be responsible for oxidative eustress and, currently, they are considered to be signaling molecules. These include nitric oxide and hydrogen peroxide, and even superoxide, which is the precursor to hydrogen peroxide. Over the last two decades, the focus of research in this area has been centered on nitric oxide, in addition to other important roles of NO in physiology and pharmacology, and especially on hydrogen peroxide. Different studies indicate that hydrogen peroxide has a crucial role in redox signaling pathways that regulate cellular homeostasis and other functions. Consequently, there is great interest in the field of redox biology and the knowledge continues to grow, although there is still much to discover.

3. Myokines Involved in Glucose Uptake in Skeletal Muscle

Skeletal muscle is receiving increasing attention as an endocrine organ due to its release of peptides or proteins, referred to as myokines, that may influence the metabolism of virtually every organ in the body, including the muscle itself. Myokines present a new paradigm for understanding how muscles communicate with other organs [42][43]. However, special attention has also been given to the auto/paracrine effects of myokines within skeletal muscle affecting muscle functions [44]. This view, originally addressed over three decades ago, suggests that proteins and other peptides produced, expressed, and released by muscle fibers after exercise exert autocrine, paracrine, or endocrine effects through actions on their receptors [45]. However, a myokine may be secreted independently of muscle contractions [46]. It is worthy of note that myokines may represent potential therapeutic targets to combat obesity and associated metabolic disorders such as insulin resistance and T2D. Myokines produced by muscles during contraction may improve insulin sensitivity and glucose oxidation via autocrine actions [47]. A secretome-based analysis of the human myocyte culture medium has revealed more than 600 myokines to date [48]. However, most of these myokines are still insufficiently characterized. In addition, the signaling pathways of certain myokines are altered in the skeletal muscle of patients with T2D [49]. Taken together, this suggests that myokine secretion is an important factor contributing to the development of muscle metabolic defects in T2D. However, whether these myokines and their effects influence both glucose uptake and GLUT4 translocation remains largely unexplored. Thus, an update on the myokines that have been identified in association with glucose uptake and insulin resistance is presented.
Fibroblast growth factor 21 (FGF21) has emerged as a promising therapeutic agent for the treatment of obesity and T2D [50]. FGF21 is a protein preferentially expressed in the liver, but it has also been described as a myokine since its expression and secretion are regulated by insulin and Akt activation [51]. This peptide hormone is secreted by several organs and can act on multiple tissues to regulate energy homeostasis [52][53]. FGF21 has been proposed as a novel metabolic regulator given its ability to normalize glucose and lipid metabolism and prevent the development of obesity and diabetes [50]. Recently, it has been reported that FGF21 regulates glucose uptake through a mechanism mediated by GLUT4 and that is dependent on atypical PKC-ζ- in skeletal muscle [54]. FGF21 gene therapy in animals receiving a long-term high-fat-diet feeding or in ob/ob mice showed marked reductions in body weight, adipose tissue hypertrophy and inflammation, hepatic steatosis inflammation and fibrosis, and insulin resistance due to the higher expression of FGF21 [55].
Irisin is a myokine that is secreted after exercise and is associated with increased energy expenditure because of its ability to stimulate the browning of white adipose tissue [56]. In skeletal muscle, it has been proposed that irisin stimulates glucose uptake after the activation of AMPK in L6 myotubes [57]. Decreased irisin secretion contributes to muscle insulin resistance, which was observed in high-fat-diet mice when the insulin action was significantly inhibited [58][59]. In addition, irisin reverses insulin resistance in C2C12 muscle cells via the p38-MAPK-PGC-1α pathway and enhances mitochondrial function [60]. Irisin improves fatty acid oxidation and glucose utilization in T2D by regulating the AMPK signaling pathway [61].
The identification of IL-6 as a myokine has created much interest around its acting as a metabolic regulator molecule [62]. However, the elevation of systemic IL-6, often in obesity and metabolic syndrome, and the role of IL-6 in metabolic disease remains controversial [62]. After exercise, IL-6 plasma levels rise because of the increased local production in muscle, and this increase may enhance substrate metabolism and whole-body glucose homeostasis [63][64][65]. Acute IL-6 exposure increases glucose metabolism in resting human skeletal muscle without changing insulin-stimulated glucose transport and insulin signaling [66]. On the contrary, it has been reported that IL-6 administration increases insulin sensitivity in vitro in muscle via the AMP-activated protein kinase [47]. Interestingly, IL-6 induces lipolysis and free fatty acid release from adipocytes and skeletal muscle [67]. In addition, low-grade systemic inflammation is one of the earliest and main pathological events that might lead to the development of insulin resistance [68]. Therefore, IL-6 may exert both pro- and anti-inflammatory effects and may even promote muscle anabolism or catabolism depending on the target structure, the predominant cytokine environment, and the mode of release [69].
Apelin is a peptide secreted from various tissues that has been classically characterized as an adipokine. Moreover, it has been described as a myokine that improves glucose metabolism and shows antidiabetic properties [70][71]. Apelin knockout mice are insulin resistant, a condition that can be reversed after apelin treatment [72]. It has been reported that both short- and long-term apelin treatments improve insulin sensitivity in obese and insulin-resistant mice, mainly due to the increase in glucose uptake in skeletal muscle that was observed in [72][73]. Using in vivo and in vitro pharmacological and genetic approaches, the involvement of eNOS, AMP-activated protein kinase, and Akt was reported in apelin-stimulated glucose uptake in the soleus muscle [73]. In C2C12 myotubes, apelin increased glucose uptake and Akt phosphorylation [72]. Apelin expression is induced by exercise signaling pathways and secreted in vitro in human primary myotubes [74]. However, it has recently been reported that exercise-induced insulin sensitization occurs independently of plasma apelin changes [75]. Interestingly, apelin treatment increases complete fatty acid oxidation, mitochondrial oxidative capacity, and biogenesis in the muscle of insulin-resistant mice. This suggests that the improvement in the insulin sensitivity triggered by apelin might be secondary to the decrease in adiposity, and it might be due to a direct action in skeletal muscle [76].
Myostatin is a myokine of the TGF-β superfamily expressed in both embryonic and adult skeletal muscle that regulates muscle mass and function, producing muscle atrophy [77]. Serum myostatin is upregulated in obesity and correlates with insulin resistance in humans [78]. The overexpression of myostatin in mice causes insulin resistance [79] whereas anti-myostatin antibodies prevent obesity [80] by stimulating fatty acid oxidation and increasing energy expenditure [81]. In addition, it has been reported that myostatin inhibits glucose uptake via the suppression of insulin-dependent and -independent signaling pathways in skeletal muscle [82]. Myostatin inhibits low basal glucose uptake, insulin-induced IRS-1 tyrosine (Tyr495) phosphorylation, and both the expression and activation of PI3K, along with diminishing Akt phosphorylation, which leads to a reduction in insulin-induced GLUT4 membrane translocation and glucose uptake [82]. In addition, this myokine decreases AMPK activity, which is accompanied by reduced GLUT4 gene expression and glucose uptake [82]. Moreover, it has been reported that myostatin regulates glucose metabolism via the AMPK pathway by promoting glucose consumption and glucose uptake, increasing glycolysis, and inhibiting glycogen synthesis in skeletal muscle cells [83].
Musclin is a factor secreted by skeletal muscle and is a potent regulator of glucose metabolism [84]. In humans, an increase in the circulating musclin level has been reported in diagnosed T2D patients [85] and those with metabolic syndrome [86]. In the latter, a positive correlation was found between both altered insulinemia and glycemia and a body composition profile with high visceral fat and lean mass [86]. The musclin-induced impairment of insulin-stimulated glucose uptake in skeletal muscle is related to Akt inhibition and PPARγ/LXRα in mice [87] and causes endoplasmic reticulum stress in rats [88]. Recently, it was found that a reduction in muscle-derived musclin production through chronic resistance exercise was involved in improving insulin resistance in rats with T2D [89]. Exercise training improves lipid metabolism and insulin sensitivity by upregulating GLUT4 and downregulating musclin in skeletal muscle [90].
Brain-derived neurotrophic factor (BDNF) is a member of the nerve growth factor family that is generated mainly by the brain [91], but is also secreted by skeletal muscle in response to contractions, and enhances fat oxidation via the activation of the AMP-activated protein kinase [92]. During short-duration aerobic exercise, immediately after a short-duration high-intensity exercise to exhaustion, there is a transient augmentation of serum BDNF concentration in humans [93]. Low levels of BDNF accompanying impaired glucose metabolism in T2D patients have been reported [94]. The repetitive subcutaneous or intracerebroventricular administration of BDNF ameliorates glucose metabolism by enhancing the glucose utilization in muscle in db/db mice [95]. Interestingly, a peripheral BDNF treatment promotes GLUT4 protein expression as well as hypophagia in skeletal muscle [96].

This entry is adapted from the peer-reviewed paper 10.3390/cells11244008

References

  1. Dumont, N.A.; Bentzinger, C.F.; Sincennes, M.C.; Rudnicki, M.A. Satellite Cells and Skeletal Muscle Regeneration. Compr. Physiol. 2015, 5, 1027–1059.
  2. Gillies, A.R.; Lieber, R.L. Structure and function of the skeletal muscle extracellular matrix. Muscle Nerve 2011, 44, 318–331.
  3. Palomero, J.; Jackson, M. Redox regulation in skeletal muscle during contractile activity and aging 1. J. Anim. Sci. 2010, 88, 1307–1313.
  4. Sies, H.; Jones, D.P. Reactive oxygen species (ROS) as pleiotropic physiological signalling agents. Nat. Rev. Mol. Cell Biol. 2020, 21, 363–383.
  5. Sylow, L.; Møller, L.L.V.; Kleinert, M.; Richter, E.A.; Jensen, T.E. Rac1-a novel regulator of contraction-stimulated glucose uptake in skeletal muscle. Exp. Physiol. 2014, 99, 1574–1580.
  6. Henríquez-Olguin, C.; Knudsen, J.R.; Raun, S.H.; Li, Z.; Dalbram, E.; Treebak, J.T.; Sylow, L.; Holmdahl, R.; Richter, E.A.; Jaimovich, E.; et al. Cytosolic ROS production by NADPH oxidase 2 regulates muscle glucose uptake during exercise. Nat. Commun. 2019, 10, 4623.
  7. Merry, T.L.; McConell, G.K. Skeletal muscle glucose uptake during exercise: A focus on reactive oxygen species and nitric oxide signaling. IUBMB Life 2009, 61, 479–484.
  8. Pedersen, B.K. Muscle as a Secretory Organ. Compr. Physiol. 2013, 3, 1337–1362.
  9. Parkin, J.M.; Carey, M.F.; Zhao, S.; Febbraio, M.A. Effect of ambient temperature on human skeletal muscle metabolism during fatiguing submaximal exercise. J. Appl. Physiol. 1999, 86, 902–908.
  10. Hoffmann, C.; Weigert, C. Skeletal Muscle as an Endocrine Organ: The Role of Myokines in Exercise Adaptations. Cold Spring Harb. Perspect. Med. 2017, 7, a029793.
  11. Commoner, B.; Townsend, J.; Pake, G.E. Free Radicals in Biological Materials. Nature 1954, 174, 689–691.
  12. Dillard, C.J.; Litov, R.E.; Savin, W.M.; Dumelin, E.E.; Tappel, A.L. Effects of exercise, vitamin E, and ozone on pulmonary function and lipid peroxidation. J. Appl. Physiol. 1978, 45, 927–932.
  13. Brady, P.S.; Brady, L.J.; Ullrey, D.E. Selenium, Vitamin E and the Response to Swimming Stress in the Rat. J. Nutr. 1979, 109, 1103–1109.
  14. Davies, K.J.; Quintanilha, A.T.; Brooks, G.A.; Packer, L. Free radicals and tissue damage produced by exercise. Biochem. Biophys. Res. Commun. 1982, 107, 1198–1205.
  15. Reid, M.B.; Shoji, T.; Moody, M.R.; Entman, M.L. Reactive oxygen in skeletal muscle. II. Extracellular release of free radicals. J. Appl. Physiol. 1992, 73, 1805–1809.
  16. Balon, T.W.; Nadler, J.L. Nitric oxide release is present from incubated skeletal muscle preparations. J. Appl. Physiol. 1994, 77, 2519–2521.
  17. O’Neill, C.A.; Stebbins, C.L.; Bonigut, S.; Halliwell, B.; Longhurst, J.C. Production of hydroxyl radicals in contracting skeletal muscle of cats. J. Appl. Physiol. 1996, 81, 1197–1206.
  18. Jackson, M.J.; Pye, D.; Palomero, J. The production of reactive oxygen and nitrogen species by skeletal muscle. J. Appl. Physiol. 2007, 102, 1664–1670.
  19. Powers, S.K.; Jackson, M.J. Exercise-Induced Oxidative Stress: Cellular Mechanisms and Impact on Muscle Force Production. Physiol. Rev. 2008, 88, 1243–1276.
  20. Jackson, M.J. Control of Reactive Oxygen Species Production in Contracting Skeletal Muscle. Antioxid. Redox Signal. 2011, 15, 2477–2486.
  21. Silveira, L.R.; Pereira-Da-Silva, L.; Juel, C.; Hellsten, Y. Formation of hydrogen peroxide and nitric oxide in rat skeletal muscle cells during contractions. Free Radic. Biol. Med. 2003, 35, 455–464.
  22. Palomero, J.; Pye, D.; Kabayo, T.; Spiller, D.G.; Jackson, M.J. In Situ Detection and Measurement of Intracellular Reactive Oxygen Species in Single Isolated Mature Skeletal Muscle Fibers by Real Time Fluorescence Microscopy. Antioxid. Redox Signal. 2008, 10, 1463–1474.
  23. Pye, D.; Palomero, J.; Kabayo, T.; Jackson, M.J. Real-time measurement of nitric oxide in single mature mouse skeletal muscle fibres during contractions. J. Physiol. 2007, 581, 309–318.
  24. Sakellariou, G.K.; Pye, D.; Vasilaki, A.; Zibrik, L.; Palomero, J.; Kabayo, T.; McArdle, F.; Van Remmen, H.; Richardson, A.; Tidball, J.G.; et al. Role of superoxide-nitric oxide interactions in the accelerated age-related loss of muscle mass in mice lacking Cu, Zn superoxide dismutase. Aging Cell 2011, 10, 749–760.
  25. McArdle, A.; Pattwell, D.; Vasilaki, A.; Griffiths, R.D.; Jackson, M.J. Contractile activity-induced oxidative stress: Cellular origin and adaptive responses. Am. J. Physiol. Physiol. 2001, 280, C621–C627.
  26. Patwell, D.M.; McArdle, A.; Morgan, J.E.; Patridge, T.A.; Jackson, M. Release of reactive oxygen and nitrogen species from contracting skeletal muscle cells. Free Radic. Biol. Med. 2004, 37, 1064–1072.
  27. Javesghani, D.; Magder, S.A.; Barreiro, E.; Quinn, M.T.; Hussain, S.N.A. Molecular Characterization of a Superoxide-Generating NAD(P)H Oxidase in the Ventilatory Muscles. Am. J. Respir. Crit. Care Med. 2002, 165, 412–418.
  28. Xia, R.; Webb, J.A.; Gnall, L.L.M.; Cutler, K.; Abramson, J.J. Skeletal muscle sarcoplasmic reticulum contains a NADH-dependent oxidase that generates superoxide. Am. J. Physiol. Physiol. 2003, 285, C215–C221.
  29. Espinosa, A.; Leiva, A.; Peña, M.; Müller, M.; Debandi, A.; Hidalgo, C.; Carrasco, M.A.; Jaimovich, E. Myotube depolarization generates reactive oxygen species through NAD(P)H oxidase; ROS-elicited Ca2+ stimulates ERK, CREB, early genes. J. Cell. Physiol. 2006, 209, 379–388.
  30. Sakellariou, G.K.; Vasilaki, A.; Palomero, J.; Kayani, A.; Zibrik, L.; McArdle, A.; Jackson, M. Studies of Mitochondrial and Nonmitochondrial Sources Implicate Nicotinamide Adenine Dinucleotide Phosphate Oxidase(s) in the Increased Skeletal Muscle Superoxide Generation That Occurs During Contractile Activity. Antioxid. Redox Signal. 2013, 18, 603–621.
  31. Gong, M.C.; Arbogast, S.; Guo, Z.; Mathenia, J.; Su, W.; Reid, M.B. Calcium-independent phospholipase A2modulates cytosolic oxidant activity and contractile function in murine skeletal muscle cells. J. Appl. Physiol. 2006, 100, 399–405.
  32. Zuo, L.; Christofi, F.L.; Wright, V.P.; Bao, S.; Clanton, T.L. Lipoxygenase-dependent superoxide release in skeletal muscle. J. Appl. Physiol. 2004, 97, 661–668.
  33. Gomez-Cabrera, M.C.; Close, G.L.; Kayani, A.; McArdle, A.; Viña, J.; Jackson, M.J. Effect of xanthine oxidase-generated extracellular superoxide on skeletal muscle force generation. Am. J. Physiol. Integr. Comp. Physiol. 2010, 298, R2–R8.
  34. Hirschfield, W.; Moody, M.R.; O’Brien, W.E.; Gregg, A.R.; Bryan, R.M.; Reid, M.B. Nitric oxide release and contractile properties of skeletal muscles from mice deficient in type III NOS. Am. J. Physiol. Integr. Comp. Physiol. 2000, 278, R95–R100.
  35. Kobzik, L.; Reid, M.B.; Bredt, D.S.; Stamler, J.S. Nitric oxide in skeletal muscle. Nature 1994, 372, 546–548.
  36. Palomero, J.; Pye, D.; Kabayo, T.; Jackson, M.J. Effect of passive stretch on intracellular nitric oxide and superoxide activities in single skeletal muscle fibres: Influence of ageing. Free. Radic. Res. 2012, 46, 30–40.
  37. Sies, H.; Belousov, V.V.; Chandel, N.S.; Davies, M.J.; Jones, D.P.; Mann, G.E.; Murphy, M.P.; Yamamoto, M.; Winterbourn, C. Defining roles of specific reactive oxygen species (ROS) in cell biology and physiology. Nat. Rev. Mol. Cell Biol. 2022, 23, 499–515.
  38. Szabó, C.; Ischiropoulos, H.; Radi, R. Peroxynitrite: Biochemistry, pathophysiology and development of therapeutics. Nat. Rev. Drug Discov. 2007, 6, 662–680.
  39. Ji, L.L.; Fu, R.; Mitchell, E.W. Glutathione and antioxidant enzymes in skeletal muscle: Effects of fiber type and exercise intensity. J. Appl. Physiol. 1992, 73, 1854–1859.
  40. Leeuwenburgh, C.; Hollander, J.; Leichtweis, S.; Griffiths, M.; Gore, M.; Ji, L.L. Adaptations of glutathione antioxidant system to endurance training are tissue and muscle fiber specific. Am. J. Physiol. Integr. Comp. Physiol. 1997, 272, R363–R369.
  41. Powers, S.K.; Ji, L.L.; Leeuwenburgh, C. Exercise training-induced alterations in skeletal muscle antioxidant capacity: A brief review. Med. Sci. Sport. Exerc. 1999, 31, 987–997.
  42. Pedersen, B.K.; Febbraio, M.A. Muscles, exercise and obesity: Skeletal muscle as a secretory organ. Nat. Rev. Endocrinol. 2012, 8, 457–465.
  43. Lightfoot, A.; Cooper, R.G. The role of myokines in muscle health and disease. Curr. Opin. Rheumatol. 2016, 28, 661–666.
  44. Eckel, J. Myokines in metabolic homeostasis and diabetes. Diabetologia 2019, 62, 1523–1528.
  45. Giudice, J.; Taylor, J.M. Muscle as a paracrine and endocrine organ. Curr. Opin. Pharmacol. 2017, 34, 49–55.
  46. Aguer, C.; Loro, E.; Di Raimondo, D. Editorial: The Role of the Muscle Secretome in Health and Disease. Front. Physiol. 2020, 11, 1101.
  47. Carey, A.L.; Steinberg, G.R.; Macaulay, S.L.; Thomas, W.G.; Holmes, A.G.; Ramm, G.; Prelovsek, O.; Hohnen-Behrens, C.; Watt, M.J.; James, D.E.; et al. Interleukin-6 Increases Insulin-Stimulated Glucose Disposal in Humans and Glucose Uptake and Fatty Acid Oxidation In Vitro via AMP-Activated Protein Kinase. Diabetes 2006, 55, 2688–2697.
  48. Goörgens, S.W.; Eckardt, K.; Jensen, J.; Drevon, C.A.; Eckel, J. Exercise and Regulation of Adipokine and Myokine Production. Prog. Mol. Biol. Transl. Sci. 2015, 135, 313–336.
  49. Balakrishnan, R.; Thurmond, D.C. Mechanisms by Which Skeletal Muscle Myokines Ameliorate Insulin Resistance. Int. J. Mol. Sci. 2022, 23, 4636.
  50. Kharitonenkov, A.; Shiyanova, T.L.; Koester, A.; Ford, A.M.; Micanovic, R.; Galbreath, E.J.; Sandusky, G.E.; Hammond, L.J.; Moyers, J.S.; Owens, R.A.; et al. FGF-21 as a novel metabolic regulator. J. Clin. Investig. 2005, 115, 1627–1635.
  51. Izumiya, Y.; Bina, H.A.; Ouchi, N.; Akasaki, Y.; Kharitonenkov, A.; Walsh, K. FGF21 is an Akt-regulated myokine. FEBS Lett. 2008, 582, 3805–3810.
  52. Von Holstein-Rathlou, S.; BonDurant, L.D.; Peltekian, L.; Naber, M.C.; Yin, T.C.; Claflin, K.E.; Urizar, A.I.; Madsen, A.N.; Ratner, C.; Holst, B.; et al. FGF21 Mediates Endocrine Control of Simple Sugar Intake and Sweet Taste Preference by the Liver. Cell Metab. 2016, 23, 335–343.
  53. Potthoff, M.J.; Kliewer, S.A.; Mangelsdorf, D.J. Endocrine fibroblast growth factors 15/19 and 21: From feast to famine. Genes Dev. 2012, 26, 312–324.
  54. Rosales-Soto, G.; Diaz-Vegas, A.; Casas, M.; Contreras-Ferrat, A.; Jaimovich, E. Fibroblast growth factor-21 potentiates glucose transport in skeletal muscle fibers. J. Mol. Endocrinol. 2020, 65, 85–95.
  55. Jimenez, V.; Jambrina, C.; Casana, E.; Sacristan, V.; Muñoz, S.; Darriba, S.; Rodó, J.; Mallol, C.; Garcia, M.; León, X.; et al. FGF21 gene therapy as treatment for obesity and insulin resistance. EMBO Mol. Med. 2018, 10, e8791.
  56. Mahajan, R.D.; Patra, S.K. Irisin, a Novel Myokine Responsible for Exercise Induced Browning of White Adipose Tissue. Indian J. Clin. Biochem. 2013, 28, 102–103.
  57. Lee, H.J.; Lee, J.O.; Kim, N.; Kim, J.K.; Kim, H.I.; Lee, Y.W.; Kim, S.J.; Choi, J.-I.; Oh, Y.; Kim, J.H.; et al. Irisin, a novel myokine, regulates glucose uptake in skeletal muscle cells via AMPK. Mol. Endocrinol. 2015, 29, 873–881.
  58. Yano, N.; Zhang, L.; Wei, D.; Dubielecka, P.M.; Wei, L.; Zhuang, S.; Zhu, P.; Qin, G.; Liu, P.Y.; Chin, Y.E.; et al. Irisin counteracts high glucose and fatty acid-induced cytotoxicity by preserving the AMPK-insulin receptor signaling axis in C2C12 myoblasts. Am. J. Physiol. Metab. 2020, 318, E791–E805.
  59. Yang, Z.; Chen, X.; Chen, Y.; Zhao, Q. Decreased irisin secretion contributes to muscle insulin resistance in high-fat diet mice. Int. J. Clin. Exp. Pathol. 2015, 8, 6490–6497.
  60. Ye, X.; Shen, Y.; Ni, C.; Ye, J.; Xin, Y.; Zhang, W.; Ren, Y. Irisin reverses insulin resistance in C2C12 cells via the p38-MAPK-PGC-1α pathway. Peptides 2019, 119, 170120.
  61. Xin, C.; Liu, J.; Zhang, J.D.; Zhu, D.; Wang, H.; Xiong, L.; Lee, Y.; Ye, J.; Lian, K.; Xu, C.; et al. Irisin improves fatty acid oxidation and glucose utilization in type 2 diabetes by regulating the AMPK signaling pathway. Int. J. Obes. 2016, 40, 443–451.
  62. Kistner, T.M.; Pedersen, B.K.; Lieberman, D.E. Interleukin 6 as an energy allocator in muscle tissue. Nat. Metab. 2022, 4, 170–179.
  63. Pedersen, B.K.; Åkerström, T.C.; Nielsen, A.R.; Fischer, C.P. Role of myokines in exercise and metabolism. J. Appl. Physiol. 2007, 103, 1093–1098.
  64. Steensberg, A.; Keller, C.; Starkie, R.L.; Osada, T.; Febbraio, M.A.; Pedersen, B.K. IL-6 and TNF-α expression in, and release from, contracting human skeletal muscle. Am. J. Physiol. Metab. 2002, 283, E1272–E1278.
  65. Febbraio, M.A.; Hiscock, N.; Sacchetti, M.; Fischer, C.P.; Pedersen, B.K. Interleukin-6 Is a Novel Factor Mediating Glucose Homeostasis During Skeletal Muscle Contraction. Diabetes 2004, 53, 1643–1648.
  66. Glund, S.; Deshmukh, A.; Long, Y.C.; Moller, T.; Koistinen, H.A.; Caidahl, K.; Zierath, J.R.; Krook, A. Interleukin-6 Directly Increases Glucose Metabolism in Resting Human Skeletal Muscle. Diabetes 2007, 56, 1630–1637.
  67. Knudsen, J.G.; Gudiksen, A.; Bertholdt, L.; Overby, P.; Villesen, I.; Schwartz, C.L.; Pilegaard, H. Skeletal muscle IL-6 regulates muscle substrate utilization and adipose tissue metabolism during recovery from an acute bout of exercise. PLoS ONE 2017, 12, e0189301.
  68. Shoelson, S.E.; Lee, J.; Goldfine, A.B. Inflammation and insulin resistance. J. Clin. Investig. 2006, 116, 1793–1801.
  69. Nelke, C.; Dziewas, R.; Minnerup, J.; Meuth, S.G.; Ruck, T. Skeletal muscle as potential central link between sarcopenia and immune senescence. eBioMedicine 2019, 49, 381–388.
  70. Chaves-Almagro, C.; Castan-Laurell, I.; Dray, C.; Knauf, C.; Valet, P.; Masri, B. Apelin receptors: From signaling to antidiabetic strategy. Eur. J. Pharmacol. 2015, 763, 149–159.
  71. Castan-Laurell, I.; Dray, C.; Knauf, C.; Kunduzova, O.; Valet, P. Apelin, a promising target for type 2 diabetes treatment? Trends Endocrinol. Metab. 2012, 23, 234–241.
  72. Yue, P.; Jin, H.; Aillaud, M.; Deng, A.C.; Azuma, J.; Asagami, T.; Kundu, R.K.; Reaven, G.M.; Quertermous, T.; Tsao, P.S. Apelin is necessary for the maintenance of insulin sensitivity. Am. J. Physiol. Endocrinol. Metab. 2010, 298, E59–E67.
  73. Dray, C.; Knauf, C.; Daviaud, D.; Waget, A.; Boucher, J.; Buléon, M.; Cani, P.D.; Attané, C.; Guigné, C.; Carpéné, C.; et al. Apelin Stimulates Glucose Utilization in Normal and Obese Insulin-Resistant Mice. Cell Metab. 2008, 8, 437–445.
  74. Besse-Patin, A.; Montastier, E.; Vinel, C.; Castan-Laurell, I.; Louche, K.; Dray, C.; Daviaud, D.; Mir, L.; Marques, M.-A.; Thalamas, C.; et al. Effect of endurance training on skeletal muscle myokine expression in obese men: Identification of apelin as a novel myokine. Int. J. Obes. 2013, 38, 707–713.
  75. Waller, J.D.; McNeill, E.H.; Zhong, F.; Vervaecke, L.S.; Goldfarb, A.H. Plasma Apelin Unchanged with Acute Exercise Insulin Sensitization. J. Sport. Sci. Med. 2019, 18, 537–543.
  76. Attané, C.; Foussal, C.; Le Gonidec, S.; Benani, A.; Daviaud, D.; Wanecq, E.; Guzmán-Ruiz, R.; Dray, C.; Bezaire, V.; Rancoule, C.; et al. Apelin Treatment Increases Complete Fatty Acid Oxidation, Mitochondrial Oxidative Capacity, and Biogenesis in Muscle of Insulin-Resistant Mice. Diabetes 2012, 61, 310–320.
  77. Rodgers, B.D.; Ward, C.W. Myostatin/Activin Receptor Ligands in Muscle and the Development Status of Attenuating Drugs. Endocr. Rev. 2021, 43, 329–365.
  78. Amor, M.; Itariu, B.K.; Moreno-Viedma, V.; Keindl, M.; Jürets, A.; Prager, G.; Langer, F.; Grablowitz, V.; Zeyda, M.; Stulnig, T.M. Serum Myostatin is Upregulated in Obesity and Correlates with Insulin Resistance in Humans. Exp. Clin. Endocrinol. Diabetes 2019, 127, 550–556.
  79. Allen, D.L.; Hittel, D.S.; McPherron, A. Expression and Function of Myostatin in Obesity, Diabetes, and Exercise Adaptation. Med. Sci. Sport. Exerc. 2011, 43, 1828–1835.
  80. Zhang, C.; McFarlane, C.; Lokireddy, S.; Masuda, S.; Ge, X.; Gluckman, P.D.; Sharma, M.; Kambadur, R. Inhibition of myostatin protects against diet-induced obesity by enhancing fatty acid oxidation and promoting a brown adipose phenotype in mice. Diabetologia 2011, 55, 183–193.
  81. Dong, J.; Dong, Y.; Chen, F.; Mitch, W.E.; Zhang, L. Inhibition of myostatin in mice improves insulin sensitivity via irisin-mediated cross talk between muscle and adipose tissues. Int. J. Obes. 2016, 40, 434–442.
  82. Liu, X.-H.; Bauman, W.A.; Cardozo, C.P. Myostatin inhibits glucose uptake via suppression of insulin-dependent and -independent signaling pathways in myoblasts. Physiol. Rep. 2018, 6, e13837.
  83. Chen, Y.; Ye, J.; Cao, L.; Zhang, Y.; Xia, W.; Zhu, D. Myostatin regulates glucose metabolism via the AMP-activated protein kinase pathway in skeletal muscle cells. Int. J. Biochem. Cell Biol. 2010, 42, 2072–2081.
  84. Nishizawa, H.; Matsuda, M.; Yamada, Y.; Kawai, K.; Suzuki, E.; Makishima, M.; Kitamura, T.; Shimomura, I. Musclin, a Novel Skeletal Muscle-derived Secretory Factor. J. Biol. Chem. 2004, 279, 19391–19395.
  85. Chen, W.-J.; Liu, Y.; Sui, Y.-B.; Zhang, B.; Zhang, X.-H.; Yin, X.-H. Increased circulating levels of musclin in newly diagnosed type 2 diabetic patients. Diabetes Vasc. Dis. Res. 2017, 14, 116–121.
  86. Sánchez, Y.L.; Yepes-Calderón, M.; Valbuena, L.; Milán, A.F.; Trillos-Almanza, M.C.; Granados, S.; Peña, M.; Estrada-Castrillón, M.; Aristizábal, J.C.; Narvez-Sanchez, R.; et al. Musclin Is Related to Insulin Resistance and Body Composition, but Not to Body Mass Index or Cardiorespiratory Capacity in Adults. Endocrinol. Metab. 2021, 36, 1055–1068.
  87. Liu, Y.; Huo, X.; Pang, X.; Zong, Z.; Meng, X.; Liu, G. Musclin Inhibits Insulin Activation of Akt/Protein Kinase B in Rat Skeletal Muscle. J. Int. Med. Res. 2008, 36, 496–504.
  88. Chen, W.-J.; Liu, Y.; Sui, Y.-B.; Yang, H.-T.; Chang, J.-R.; Tang, C.-S.; Qi, Y.-F.; Zhang, J.; Yin, X.-H. Positive association between musclin and insulin resistance in obesity: Evidence of a human study and an animal experiment. Nutr. Metab. 2017, 14, 46.
  89. Shimomura, M.; Horii, N.; Fujie, S.; Inoue, K.; Hasegawa, N.; Iemitsu, K.; Uchida, M.; Iemitsu, M. Decreased muscle-derived musclin by chronic resistance exercise is associated with improved insulin resistance in rats with type 2 diabetes. Physiol. Rep. 2021, 9, e14823.
  90. Yu, J.; Zheng, J.; Liu, X.; Feng, Z.; Zhang, X.; Cao, L.; Zhou, Z. Exercise improved lipid metabolism and insulin sensitivity in rats fed a high-fat diet by regulating glucose transporter 4 (GLUT4) and musclin expression. Braz. J. Med. Biol. Res. 2016, 49, e5129.
  91. Miranda, M.; Morici, J.F.; Zanoni, M.B.; Bekinschtein, P. Brain-Derived Neurotrophic Factor: A Key Molecule for Memory in the Healthy and the Pathological Brain. Front. Cell. Neurosci. 2019, 13, 363.
  92. Matthews, V.B.; Åström, M.-B.; Chan, S.; Bruce, C.; Krabbe, K.S.; Prelovsek, O.; Åkerström, T.; Yfanti, C.; Broholm, C.; Mortensen, O.H.; et al. Brain-derived neurotrophic factor is produced by skeletal muscle cells in response to contraction and enhances fat oxidation via activation of AMP-activated protein kinase. Diabetologia 2009, 52, 1409–1418.
  93. Vega, S.R.; Strüder, H.K.; Wahrmann, B.V.; Schmidt, A.; Bloch, W.; Hollmann, W. Acute BDNF and cortisol response to low intensity exercise and following ramp incremental exercise to exhaustion in humans. Brain Res. 2006, 1121, 59–65.
  94. Krabbe, K.S.; Nielsen, A.R.; Krogh-Madsen, R.; Plomgaard, P.; Rasmussen, P.; Erikstrup, C.; Fischer, C.; Lindegaard, B.; Petersen, A.M.W.; Taudorf, S.; et al. Brain-derived neurotrophic factor (BDNF) and type 2 diabetes. Diabetologia 2007, 50, 431–438.
  95. Yamanaka, M.; Tsuchida, A.; Nakagawa, T.; Nonomura, T.; Ono-Kishino, M.; Sugaru, E.; Noguchi, H.; Taiji, M. Brain-derived neurotrophic factor enhances glucose utilization in peripheral tissues of diabetic mice. Diabetes Obes. Metab. 2007, 9, 59–64.
  96. Suwa, M.; Yamamoto, K.-I.; Nakano, H.; Sasaki, H.; Radak, Z.; Kumagai, S. Brain-Derived Neurotrophic Factor Treatment Increases the Skeletal Muscle Glucose Transporter 4 Protein Expression in Mice. Physiol. Res. 2010, 59, 619–623.
More
© Text is available under the terms and conditions of the Creative Commons Attribution (CC BY) license; additional terms may apply. By using this site, you agree to the Terms and Conditions and Privacy Policy.
This entry is offline, you can click here to edit this entry!
Video Production Service
Feedback