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. 2024 Sep 24;77:103371. doi: 10.1016/j.redox.2024.103371

Interplay of ROS, mitochondrial quality, and exercise in aging: Potential role of spatially discrete signaling

Siobhan M Craige a,⁎⁎, Rebecca K Mammel a, Niloufar Amiri a,b, Orion S Willoughby a, Joshua C Drake a,
PMCID: PMC11474192  PMID: 39357424

1. Introduction

In a young, healthy state, repetitive exposure to acute energetic stressors such as time-restricted feeding (e.g. caloric restriction and intermittent fasting) or exercise result in long-term benefits. This is due to multiple, overlapping stress response pathways that rapidly initiate signaling events to restore homeostasis including detection mechanisms, stress-specific transcriptional responses, and structural modifications to cellular organelles, such as mitochondria. The culmination of these coordinated stress responses are adaptations that limit homeostatic disturbance from future stressors of similar magnitude, otherwise known as stress resilience. However, with advancing age, a gradual decline in resilience occurs. This decline manifests as a reduction in the ability to regain homeostasis in response to stressors, potentially resulting in inadequate or even counter-productive stress responses. The decline in resilience ultimately narrows an organism's capacity to adapt to stressors, limiting what is known as the “homeodynamic space”- the range within which the body can maintain homeostasis despite challenges [1]. Nevertheless, repeated exposure to stressors related to energy metabolism such as the case with regular exercise, play a critical role in maintaining and even enhancing resilience, leading to healthspan extension [2].

While life-long exercise-mediated healthspan extension is an integrative phenomenon, skeletal muscle is particularly salient for healthspan extension. Skeletal muscle accounts for ∼40 % of body mass and, during periods of prolonged contraction such as during exercise, energy production can increase by as much as 100-fold [3,4]. Mitochondria meet a large portion of the energetic demand of skeletal muscle during exercise by oxidizing nutrient substrates to generate ATP. The loss of skeletal muscle with age, termed sarcopenia, is a common phenotype of aging and is causally linked to functional decline with age [5], as well as development of age-related disease [6,7] and increased mortality [8,9]. In humans, capacity for mitochondria to produce ATP declines with age [[10], [11], [12], [13]], which may be a harbinger for several age-related chronic conditions and diseases (e.g. sarcopenia [14,15], functional impairment [16], Alzheimer's Disease [17], diabetes [18], etc.).

In the 1950s Denham Harman proposed the free radical theory of aging [19] which posited reactive oxygen species (ROS) accumulation over time resulted in the biological consequences of aging. He later amended this theory to focus on mitochondrial ROS damage accumulation [20]. In skeletal muscle, ROS can be produced as natural byproducts of ATP production or enzymatically by multiple proteins. In response to muscle contraction, known ROS producers include the electron transport chain (ETC), NADPH oxidase 2 (NOX2), and NADPH oxidase 4 (NOX4). Additionally, ROS can originate from extracellular sources, such as other cell types like the endothelium, where both xanthine oxidase [21] and NOX4 [22] are important ROS producers. Superoxide can be reduced to hydrogen peroxide (H2O2) by extracellular superoxide dismutases (SOD). H2O2 is capable of crossing membranes or being imported by aquaporins. In skeletal muscle mitochondria, superoxide production occurs through the ETC, which can be converted to H2O2 by SOD2. Alternatively, mitochondrial H2O2 may be generated by NOX4 which could then be transported to the cytosol via mitochondrial aquaporins (see Powers et al. for an overview of skeletal muscle ROS-producers [23]). Nitric oxide (NO) also contributes to the skeletal muscle mitochondrial ROS environment as NO readily reacts with superoxide to form peroxynitrite, a highly reactive ROS [24]. NO is produced by three nitric oxide synthase (NOS) isoforms found in skeletal muscle: endothelial NOS (eNOS), neuronal NOS (nNOS), and inducible NOS (iNOS) [25]. In skeletal muscle, eNOS has been reported to colocalize with mitochondria [[25], [26], [27]], though this finding remains controversial. NOS activity in skeletal muscle is increased after acute exercise [28] and may influence mitochondrial responses to exercise.

While the nuanced production of skeletal muscle ROS in response to exercise is still an active area of research, studies have identified unique roles for different ROS producers at specific locations. For instance, cytosolic ROS production by NOX2 is crucial for glucose uptake [29], whereas NOX4 may play a more significant role in fatty acid oxidation and mitochondrial adaptation to exercise [22,30]. NOX4, when localized to the sarcoplasmic reticulum (SR), leads to oxygen-dependent ryanodine receptor-Ca(2+)-release channel (RyR1) oxidation and calcium release [31]; in dystrophic mice, NOX4-RyR1-induced Ca2+ leak can trigger NOX2 ROS production [32]. Both NOX2 and NOX4 are essential for mediating acute exercise transcriptional responses [22,33]. Although the details of skeletal muscle ROS production are still being uncovered with the development of new tools, the overall understanding is shaped by central paradigms. One such paradigm is that when ROS production in any location exceeds the capacity of antioxidants, accumulation in ROS can result in dysregulation of reduction/oxidation (redox) circuitry [34], resulting in system stress and macromolecular oxidative damage which may be particularly important in the context of mitochondrial health and aging [35].

In mitochondria in particular, ROS-mediated, post-translational modifications of proteins in the electron transport chain, TCA cycle, and/or β-oxidation can have detrimental consequences for cellular energy production [18]. Elevated ROS levels can impact other cellular pathways as well, disrupting intracellular communication [36], further impairing energy production and generating a deleterious feedback loop that results in even more ROS being produced. This negative cycle served as the underlying rationale behind Harman's free radical theory of aging. Harman's theory was the catalyst for research in later years that provided evidence for redox system dysregulation underlying age-related proteome modifications [37,38], particularly in skeletal muscle [39,40].

In the early 1980s, ROS were demonstrated to be produced in rat skeletal muscle [41] and later in human skeletal muscle in response to exercise via electron paramagnetic resonance [42]. Within the framework of Harman's free radical theory of aging, subsequent studies used antioxidants with the idea of improving exercise adaptations. However, utilizing antioxidants to reduce ROS have since been found to blunt adaptive mitochondrial responses to exercise [[43], [44], [45]], suggesting ROS are essential for adaptive signaling under energetically demanding conditions. This finding is supported by work in skeletal muscle cells demonstrating ROS, specifically H2O2, are essential for expression of multiple genes in vivo [[22], [46]] and in vitro [[47], [48], [49], [50]]. As evidenced by oxidatively sensitive reporters, exercise-induced ROS precedes and coincides with mitophagy in skeletal muscle [51], as well as mitochondrial fragmentation in C2C12 myoblasts [52], a murine derived cell line. Research in other cell types demonstrates the crucial role of mitochondrial H2O2 in facilitating mitophagy induced by energetic stress, suggesting mitochondrial ROS may serve as physiologic regulators for energetic stress conditions such as exercise [53]. The notion of blunting exercise-induced ROS mitigating adaptive responses to exercise aligns with the understanding that oxidized thiols (cysteine) can modify protein function, including protein phosphatases [54] and kinases [55], which are highly sensitive to rapid shifts in ROS levels. Such fine tailoring of intracellular communication via ROS suggests that localized ROS signaling [56,57] might influence health outcomes, with ROS-mediated mitohormesis [58] counteracting age-related functional regression. Here, we will discuss concepts of skeletal muscle adaptation to regular exercise that are pertinent to maintaining mitochondrial adaptive homeostasis with age, the role of ROS in those processes, and how understanding ROS as essential signaling molecules gives insight to the role of ROS in extending healthspan.

2. ROS and aging in adaptive homeostasis through exercise

The concept of mitohormesis proposes exposing mitochondria to multiple acute stressors, such as regular exercise, results in resilience [59]. Mitohormesis decreases with age, as well as in diabetes and insulin resistance, contributing to a decline of skeletal muscle mass and function [44,[59], [60], [61]]. Conceptually, it is reasonable to speculate that transient increases in ROS in response to acute exercise are rapidly detected within the local mitochondrial environment, contributing to the adaptations required for resilience. As aging is associated with a dampened acute stress response, potentially stemming from the perpetual generation of ROS in resting skeletal muscle (high background signal) [[62], [63], [64]], the production of ROS in response to acute exercise may be more difficult to recognize, alluding to a skewed signal-to-noise ratio. Thus far, efforts to counteract age-related decline through localized mitochondrial antioxidants have yielded inconsistent results [65,66]. However, some studies have demonstrated positive outcomes. For example, administration of elamipretide has been shown to improve muscle function in aging mice by targeting the inner mitochondrial membrane where it binds reversibly to cardiolipin and improves ATP production while reducing mitochondrial ROS levels [67]. An intriguing avenue for exploration would involve investigating whether exercise still causes a transient spike in mitochondrial ROS production even in the presence of elamipretide. However, the dynamics of mitochondrial H2O2 currently are poorly understood, and we are just now employing tools that allow for visualization of these dynamics [68,69]. Notably, several conceivable sources of localized mitochondrial ROS exist, including the ETC, p66shc [70], and NOX4 [71,72]. A recent study discovered ROS produced in specific mitochondrial microdomains caused a single cysteine oxidation in the complex I subunit NDUF-2.1 of C. elegans, which triggered a behavioral response to hypoxia [73], suggesting that the responses to energetic stress are significantly impacted by localized mitochondrial ROS production. Together, these data suggest that uncovering divergences in ROS production across microdomains may elucidate the variable outcomes witnessed with mitochondrial antioxidants.

3. Remodeling of the mitochondrial reticulum

In skeletal muscle, mitochondria form an intricate reticulum that extends along the length of the cell (i.e. myofiber) [[74], [75], [76], [77]] that functions as a syncytium with potential energy distributed across the reticulum [74,75,78]. Structural and functional integrity of the mitochondrial reticulum is maintained through synergistic processes that remodel the reticulum to maintain homeostasis. These processes include 1) dynamic reorganization of the reticulum, 2) recognition, removal, and degradation of damaged and/or dysfunctional regions (mitophagy), and 3) creation and incorporation of new proteins, lipids, and mtDNA (biogenesis), all of which function with a high degree of spatial specificity [[79], [80], [81]]. Mitochondrial remodeling processes are all sensitive to energetic stressors that produce ROS, such as exercise, to aid energetic production [74,76] and orchestrate adaptive responses to the mitochondrial reticulum that ensure sufficient energetic capacity during future events [51,79].

3.1. Mitochondrial dynamics

For mitochondria to respond and adapt to stressed environmental factors, they must be able to conform and maintain their shape, size, and distribution. This process requires cycles of fusion, the combining of mitochondrial membranes, and fission, the division of mitochondrial membranes creating a more fragmented mitochondrial reticulum [[82], [83], [84]]. The combination of those two processes have been coined mitochondrial dynamics. Similarly, like all other aspects of mitochondrial health, mitochondrial dynamics are impacted by exercise and ROS, while inefficient and unbalanced mitochondrial dynamics affect pathways that drive markers of aging. ROS play a crucial role in regulating various aspects of mitochondrial dynamics. For instance, in skeletal muscle cells, exposure to ROS leads to the fragmentation of the mitochondrial reticulum [52]. Additionally, investigations conducted on endothelial cells have highlighted the involvement of two specific proteins, Protein Disulfide Isomerase A1 (PDIA1) and Dynamin-Related Protein 1 (DRP1), both in vivo and in vitro [85,86]. Using a known regulator of mitochondrial fission, DRP1, new studies have emerged that proved the importance of mitochondrial dynamics in exercise performance and adaptations through the increased fused and tubular mitochondrial network post-exercise [87,88]. DRP1 can be activated by ROS, and ROS may be the molecular link between exercise and DRP1-mediated mitochondrial fission [89,90]. Specifically examining chronic vs. acute exercise effects on mitochondrial dynamics, Mesquits et al. showed mitochondrial fusion and fission markers, MFN1, MFN2, OPA1, and DRP1 increased at the mRNA level similar to others who investigated exercise type (high-impact vs. low-impact) on mitochondrial dynamics [91,92]. As previously highlighted, exercise impacts mitochondrial function and delays markers of aging. An increase in mitochondrial fragmentation is associated with aging [93]. A recent study investigating the relationship between mitochondrial fragmentation and sustained fitness with aging in C. elegans, found that age-associated increases in mitochondrial fragmentation could be delayed through exercise training in an AMPK-dependent manner. Therefore, manipulation of mitochondrial dynamics through the energetic sensor AMPK may be targets for therapies to aid in the aging-induced decrease in muscle function [93,94]. While mitochondrial dynamics are necessary for healthy mitochondrial function, few pathways regarding ROS and aging have been investigated, leaving room for further exploration. Recent work has aimed to addressed this gap as elevation in basal ROS levels in aged C. elegans is associated with hyperoxidized peroxiredoxins (PRDXs) and impaired mitochondrial remodeling responses to acute swim exercise [95], suggesting that the high background noise of elevated ROS with age impairs mechanisms responsible for acute fluctuations in ROS with exercise.

3.2. Enhanced mitophagy

Targeted degradation of damaged and/or dysfunctional regions of the mitochondrial reticulum through mitophagy maintains skeletal muscle health [96], in part, through limiting mitochondrial dysfunction and helping to maintain redox balance [97]. Mitochondrial ROS have been implicated in mitophagy initiation in contexts such as atrophy [98] and autophagosome recruitment in conditions of energetic stress [99,100]. Previous evidence in vitro suggests energetic stress via starvation increased mitochondrial H2O2, resulting in mitophagy mediated by oxidation and inactivation of ATG4 [53]; however, whether this is a universal energetic stress response is unknown. Experiments in skeletal muscle cells in vitro have demonstrated H2O2-induced mitochondrial fragmentation and mitophagy [52,101]. Mitophagy is initiated in response to exercise [51]; however, mitophagy induction in skeletal muscle is posited to be impaired with age [102], which would cause an increase in damaged mitochondria. Understanding how exercise-mediated mitophagy signaling changes with age will be necessary to maintain healthy mitochondria with aging. As a preceding and concomitant event to exercise-induced mitophagy in vivo, mitochondria are exposed to a bolus of oxidative stress in the hours following the cessation of exercise [51]. This observation raises the possibility that the mitochondrial H2O2 signal is a component of the mitophagy response. Several redox-sensitive proteins and transcription factors have been identified as regulators of mitophagy [103]. For example, the transcription factors nuclear factor erythroid 2-related factor 2 (NRF2), nuclear respiratory factor 1 (NRF1), and peroxisome proliferator-activated receptor gamma coactivator 1-alpha (PGC‐1α) are activated in response to ROS and can upregulate the expression of genes involved in mitophagy [104]. Recent findings illustrate that loss of PRDX-2, which may act as a relayer of oxidative equivalents to modify protein function, impairs mitophagy induction in C. elegans in response to acute swim exercise [95]. Further studies into relationships between H2O2 production and mitophagy induction in mammalian models are warranted.

Recent studies have highlighted that the specificity and localization of mitophagy may be regulated through spatial mitochondrial mechanisms in response to exercise, possibly through AMPK signaling [[105], [106], [107]]. New evidence suggests that distinct subcellular pools of the energetic sensor AMPK are required for modulating mitochondria to energetic stress [105,107] and, in skeletal muscle in particular, exercise-induced mitophagy (mitoAMPK) [106]. Within the mitochondrial microenvironment, AMPK and ROS are integral signaling molecules that may play a role in the localization of mitochondrial quality control mechanisms. Recently, downstream targets of AMPK have been investigated to elucidate mechanisms uncovering ULK1, DRP1, PARK2, and FUNDC1, all of which may play a role in driving exercise-induced mitophagy [51,87,108]. Investigating the upregulation and knockout of Parkin, a mitophagy marker, lends some insight as both mice and C. elegans require Parkin to adapt to stress and prevent aging-related impairments through skeletal muscle function and mass [109,110]. Together, these findings suggest that mitochondrial ROS can initiate mitophagy as part of a cellular quality control mechanism. Still, the relationship between mitophagy and ROS remains complex as the location, tissue type, and quantity of ROS produced can result in vastly different outcomes [92,111]. The mechanism(s) for which exercise-induced ROS and mitophagy result in beneficial adaptations and how aging alters these signaling pathways is unclear. Further research of these gaps in understanding has the potential to uncover new strategies for promoting healthy aging.

3.3. Mitochondrial biogenesis

Mitochondrial biogenesis, the influx of newly synthesized proteins, lipids and mtDNA that results in expansion of the mitochondrial reticulum, is well described to be induced by exercise in skeletal muscle, with the first discovery made in 1967 by John Holloszy [112]. Transcriptionally, mitochondrial biogenesis depends on activation of PGC‐1α through overlapping upstream signals that are sensitive to exercise and ROS in skeletal muscle (e.g. p38 MAPK, AMPK, and p53) [79]. In humans, aging is associated with a reduction in PGC‐1α expression [94], which may suggest an age-related impairment in mitochondrial biogenesis. Over-expression of PGC‐1α in aged male mice maintains muscle mass, thus warding off the development of sarcopenia [113], suggesting increasing mitochondrial biogenesis has geroprotective effects in skeletal muscle. Direct measures of the synthesis of mitochondrial proteins (biogenesis) via isotope tracing studies have shown no differences in the synthesis of mitochondrial proteins between young and old humans and mice [114,115]. While these findings suggesting mitochondrial biogenesis, as a whole, may not be negatively affected by aging, others have found evidence for altered fidelity to translated proteins with age can negatively impact skeletal muscle health [116,117], as well as influence longevity [117,118] and age-related pathology [119]. Therefore, age-related changes to translational fidelity may cause functional detriments to mitochondrial quality in skeletal muscle. Knock-in mice expressing the ribosomal ambiguity mutation RPS9 D95 N produce more mitochondrial-ROS in skeletal muscle and show evidence of oxidative damage, such as lipid peroxidation, above that which occurs with aging [117]. Evidence suggests exercise-induced ROS are a necessary signal for PGC‐1α expression [120]. Therefore, it is possible that increased ROS-production with age alters the fidelity of synthesized proteins, contributing to loss of mitochondrial quality. Understanding how the response to acute exercise is altered due to poor translational fidelity in skeletal muscle and whether exercise training can mitigate this phenomenon will be an interesting area for future investigation.

3.4. Dysregulated intra- and intercellular signaling may contribute to decline in mitochondrial quality control

Evidence in recent years has elucidated that mitochondrial quality control processes are governed by localized, discrete spatial signaling mechanisms, particularly in skeletal muscle. These signals operate on two levels: intracellular (within myocytes), and intercellular (through neighboring cells). As described above, the redox environment within myofibers influences overall functions, including regulation of mitochondrial quality control. We and others have recently demonstrated AMPK's mitochondrial localization across multiple tissues, including skeletal muscle, where it resides on the outer mitochondrial membrane as mitoAMPK [106,107]. In skeletal muscle, mitoAMPK is activated at spatially distinct domains across the mitochondrial reticulum in response to prolonged contraction and specifically in response to mitochondrial complex I inhibition via metformin [106]. Additionally, inhibiting mitoAMPK activity effectively attenuated exercise-induced mitophagy in skeletal muscle [106], linking localized mitoAMPK activation to mitophagy during energetic stress.

While AMPK is conventionally recognized as sensing ADP and AMP levels [121,122] evidence suggests it is regulated by ROS as well [89,123,124]. Notably, ROS can activate AMPK even when cellular energy remains consistent [125,126]. One possibility that may govern local tagging of mitochondria for mitophagy is a coordinated response to energetic stress through the localized production of ROS by mitochondrial complexes or other proteins that produce ROS localized to the mitochondria, such as NOX4 [127,128]. Indeed, NOX4 contains an ATP binding motif (Walker-A) [72], potentially allowing it to sense low energy and increase ROS production under these conditions. The sensitivity of AMPK to energetic stress has been shown to decline with age in skeletal muscle [129], as well as the abundance of the various subunit isoforms [130]. However, the regulation or potential dysregulation of subcellular AMPK pools with aging, and possible connections to the age-related changes in mitochondrial quality control described above, remains unexplored.

While the redox environment of skeletal muscle is influence by localized ROS signaling within myofibers themselves, neighboring tissue/cell types are regulated as well. Recent advancements in single-cell sequencing and multi-omics approaches have begun to deconvolute the responses to exercise and aging on a cell type-specific level, offering promising insights into the distinct contributions of various cell types to skeletal muscle health and aging. Notably, an investigation spanning multiple tissues at the single-cell level revealed prominent anti-aging effects of exercise on the central nervous system and vasculature [131]. Remarkably, resetting the circadian clock through BMAL1 in only endothelial (vascular) cells closely mirrored exercise benefits, emphasizing the functional importance of the vasculature across tissues. Given the well-established connection between a functional vascular endothelium and skeletal muscle metabolism, it follows that capillary density correlates with muscle oxidative capacity, highlighting the microvasculature's role in determining mitochondrial metabolism [[132], [133], [134], [135]]. The close physical proximity between the vasculature and myofibers allows for significant intercellular crosstalk between the vascular endothelium and myofibers beyond simply blood supply. Indeed, endothelial cells comprise a substantial portion of muscle tissue [136] and the mitochondrial reticulum closely line the skeletal muscle microvasculature [[137], [138], [139]]. Thus, endothelial cells may be responsible for altering myofiber redox state, thereby impacting mitochondrial quality control. Recent findings have demonstrated ROS produced by NOX4 in endothelial cells are required for skeletal muscle mitochondrial metabolism in response to acute exercise [22]. NOX4 expression has been shown to be elevated in aorta with aging [140], and required for PGC‐1α expression [30]. Thus, it is feasible that age-related changes in endothelial NOX4 may underlie age-related declines in translational fidelity in skeletal muscle [116,117]. NOX4 primarily produces H2O2 [141,142], both NOX4 expression and H2O2 can drive endothelial expression of eNOS [[143], [144], [145], [146], [147]]. Deletion of eNOS significantly impairs muscle β-oxidation and mitochondrial biogenesis [[148], [149], [150], [151]]. Moreover, recent evidence demonstrates that deletion of eNOS in hepatocytes blunts exercise-induced increases in hepatic fatty acid oxidation and coincides with inhibition of the mitophagy activator ULK1. [152] Whether eNOS regulates exercise-induced mitophagy in skeletal muscle has not yet been tested. Together these data underscore the potential role of intercellular ROS communication on influencing myofiber redox state and mitochondrial quality control. While influence from other cell types, such as infiltrating immune cells and mesenchymal cells or contraction signaling from motor neurons across the neuromuscular junction likely influence myofiber redox state and mitochondrial quality control as well, these interactions remain to be elucidated. Understanding the mechanisms behind localized stress responses on myofiber redox state and mitochondrial quality control in aging could provide valuable insights into the development of interventions aimed at preserving skeletal muscle mitochondrial function and promoting healthy aging.

4. Conclusion

Aging leads to a progressive decline in homeostatic mechanisms, resulting in impaired responses to temporary physiological stressors, such as exercise. This decline compromises the capacity to handle future disturbances and impairs resilience. In a young, healthy state, skeletal muscle adapts to exercise through a well-coordinated series of intra- and intercellular stress signaling processes. Spatial regulation of mitochondrial quality control and the adaptive response to exercise is as emerging area of research in exercise metabolism that has implications for geroscience. Evidence suggest skeletal muscle ROS are essential signals for the adaptive responses of mitochondria and quality control mechanisms to exercise. However, the accumulation of ROS and ROS-mediated damage with age may reduce both the receiving and responding to adaptive signals resulting from a diminished signal-to-noise ratio (Fig. 1). The site-specific production of ROS in response to exercise and its significance in both intra- and intercellular contexts may significantly advance our understanding of how aging alters the adaptive response to exercise. Deconvoluting ROS regulation of mitochondrial quality control signaling with spatial resolution as it relates to aging could open new avenues towards promoting healthspan.

Fig. 1.

Fig. 1

Age related changes in inter and intra-cellular redox homeostasis and the response to exercise. A. In young skeletal muscle, baseline ROS are low and redox homeostasis is maintained. Thus there are no deficiencies in energy production. B. In response to exercise in young skeletal muscle, ROS are transiently increased in multiple cell types, acting as signaling molecules to aid in mitochondrial quality control processes (e.g. mitophagy and mitochondrial dynamics) to maintain a healthy mitochondrial reticulum. C. With aging, baseline ROS are elevated, resulting in poor redox homeostasis and contributing to a more fragmented mitochondrial reticulum. D. The elevated baseline ROS in aging skeletal muscle may impair the ability of transient exercise-induced ROS to effectively signal for mitochondrial quality control processes (skewed signal-to-noise ratio).

Funding

This work was supported by National Institutes of Health grants (R00AG057825, R01AG080731) from the National Institute on Aging to J.C.D. and (K01AR073332, R03AR083493, R56AR083948) from the National Institute of Arthritis, Musculoskeletal and Skin Disease to S.M.C.

CRediT authorship contribution statement

Siobhan M. Craige: Writing – review & editing, Writing – original draft, Supervision, Funding acquisition, Conceptualization. Rebecca K. Mammel: Writing – review & editing, Writing – original draft. Niloufar Amiri: Writing – review & editing, Writing – original draft. Orion S. Willoughby: Writing – review & editing, Writing – original draft. Joshua C. Drake: Writing – review & editing, Writing – original draft, Supervision, Funding acquisition, Conceptualization.

Declaration of competing interest

Declarations of interest: none.

Contributor Information

Siobhan M. Craige, Email: craigesm@vt.edu.

Joshua C. Drake, Email: joshuacd@vt.edu.

Data availability

No data was used for the research described in the article.

References

  • 1.Pomatto L.C.D., Davies K.J.A. The role of declining adaptive homeostasis in ageing. J. Physiol. 2017;595:7275–7309. doi: 10.1113/JP275072. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 2.Pomatto L.C.D., Sun P.Y., Davies K.J.A. To adapt or not to adapt: consequences of declining Adaptive Homeostasis and Proteostasis with age. Mech. Ageing Dev. 2019;177:80–87. doi: 10.1016/j.mad.2018.05.006. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 3.Janssen I., H S.B., Baumgartner R.N., Ross R. Estimation of skeletal muscle mass by bioelectrical impedance analysis. J. Appl. Physiol. 2000;89(1985):465–471. doi: 10.1152/jappl.2000.89.2.465. [DOI] [PubMed] [Google Scholar]
  • 4.Weibel E.R., Hoppeler H. Exercise-induced maximal metabolic rate scales with muscle aerobic capacity. J. Exp. Biol. 2005;208:1635–1644. doi: 10.1242/jeb.01548. [DOI] [PubMed] [Google Scholar]
  • 5.Cruz-Jentoft A.J., Landi F., Topinkova E., Michel J.P. Understanding sarcopenia as a geriatric syndrome. Curr. Opin. Clin. Nutr. Metab. Care. 2010;13:1–7. doi: 10.1097/MCO.0b013e328333c1c1. [DOI] [PubMed] [Google Scholar]
  • 6.Cleasby M.E., Jamieson P.M., Atherton P.J. Insulin resistance and sarcopenia: mechanistic links between common co-morbidities. J. Endocrinol. 2016;229:R67–R81. doi: 10.1530/JOE-15-0533. [DOI] [PubMed] [Google Scholar]
  • 7.Burns J.M.J., D K., Watts A., Swerdlow R.H., Brooks W.M. Reduced lean mass in early Alzheimer Disease and its association with brain atrophy. Arch. Neurol. 2010;67:428–433. doi: 10.1001/archneurol.2010.38. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 8.Metter E.J., T L.A., Schrager M., Conwit R. Skeletal muscle strength as a predictor of all-cause mortality in healthy men. J Gerontol A Biol Sci Med Sci. 2002;57:359–365. doi: 10.1093/gerona/57.10.b359. [DOI] [PubMed] [Google Scholar]
  • 9.Rantanen T., H T., Leveille S.G., Visser M., Foley D., Masaki K., Guralnik J.M. Muscle strength and body mass index as long-term predictors of mortality in initially healthy men. J Gerontol A Biol Sci Med Sci. 2000;55:168–173. doi: 10.1093/gerona/55.3.m168. [DOI] [PubMed] [Google Scholar]
  • 10.Trounce I., B E., Marzuki S. Decline in skeletal muscle mitochondrial respiratory chain function: possible factor in ageing. Lancet. 1989;333 doi: 10.1016/s0140-6736(89)92143-0. [DOI] [PubMed] [Google Scholar]
  • 11.Boffoli D., S S.C., Vergari R., Solarino G., Santacroce G., Papa S. Decline with age of the respiratory chain activity in human skeletal muscle. Biochim. Biophys. Acta. 1994;1226:73–82. doi: 10.1016/0925-4439(94)90061-2. [DOI] [PubMed] [Google Scholar]
  • 12.Short K.R., et al. Decline in skeletal muscle mitochondrial function with aging in humans. Proc. Natl. Acad. Sci. U. S. A. 2005;102:5618–5623. doi: 10.1073/pnas.0501559102. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 13.Porter C., et al. Mitochondrial respiratory capacity and coupling control decline with age in human skeletal muscle. Am. J. Physiol. Endocrinol. Metab. 2015;309:E224–E232. doi: 10.1152/ajpendo.00125.2015. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 14.Pugh T.D., et al. A shift in energy metabolism anticipates the onset of sarcopenia in rhesus monkeys. Aging Cell. 2013;12:672–681. doi: 10.1111/acel.12091. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 15.del Campo A., et al. Muscle function decline and mitochondria changes in middle age precede sarcopenia in mice. Aging. 2018;10:34–55. doi: 10.18632/aging.101358. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 16.Tian Q., et al. Muscle mitochondrial energetics predicts mobility decline in well-functioning older adults: the baltimore longitudinal study of aging. Aging Cell. 2022;21 doi: 10.1111/acel.13552. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 17.Tian Q., et al. Skeletal muscle mitochondrial function predicts cognitive impairment and is associated with biomarkers of Alzheimer's disease and neurodegeneration. Alzheimers Dement. 2023;19:4436–4445. doi: 10.1002/alz.13388. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 18.Jia D., Tian Z., Wang R. Exercise mitigates age-related metabolic diseases by improving mitochondrial dysfunction. Ageing Res. Rev. 2023;2023 doi: 10.1016/j.arr.2023.102087. [DOI] [PubMed] [Google Scholar]
  • 19.Harman D. Aging: a theory based on free radical and radiation chemistry. J. Gerontol. 1956;11:298–300. doi: 10.1093/geronj/11.3.298. [DOI] [PubMed] [Google Scholar]
  • 20.Harman D. The biologic clock: the mitochondria? J. Am. Geriatr. Soc. 1972;20:145–147. doi: 10.1111/j.1532-5415.1972.tb00787.x. [DOI] [PubMed] [Google Scholar]
  • 21.Hellsten Y., Frandsen U., Orthenblad N., Sjodin B., Richter E.A. Xanthine oxidase in human skeletal muscle following eccentric exercise: a role in inflammation. J. Physiol. 1997;498(Pt 1):239–248. doi: 10.1113/jphysiol.1997.sp021855. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 22.Specht K.S., et al. Nox4 mediates skeletal muscle metabolic responses to exercise. Mol. Metabol. 2021;45 doi: 10.1016/j.molmet.2020.101160. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 23.Powers S.K., Radak Z., Ji L.L., Jackson M. Reactive oxygen species promote endurance exercise-induced adaptations in skeletal muscles. J Sport Health Sci. 2014;13:780–792. doi: 10.1016/j.jshs.2024.05.001. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 24.Radi R. Peroxynitrite, a stealthy biological oxidant. J. Biol. Chem. 2013;288:26464–26472. doi: 10.1074/jbc.R113.472936. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 25.Kobzik L., Stringer B., Balligand J.L., Reid M.B., Stamler J.S. Endothelial type nitric oxide synthase in skeletal muscle fibers: mitochondrial relationships. Biochem. Biophys. Res. Commun. 1995;211:375–381. doi: 10.1006/bbrc.1995.1824. [DOI] [PubMed] [Google Scholar]
  • 26.Bates T.E., Loesch A., Burnstock G., Clark J.P. Mitochondrial nitric oxide synthase: a ubiquitous regulator of oxidative phosphorylation? Biochem. Biophys. Res. Commun. 1996;218:40–44. doi: 10.1006/bbrc.1996.0008. [DOI] [PubMed] [Google Scholar]
  • 27.Aguirre E., Lopez-Bernardo E., Cadenas S. Functional evidence for nitric oxide production by skeletal-muscle mitochondria from lipopolysaccharide-treated mice. Mitochondrion. 2012;12:126–131. doi: 10.1016/j.mito.2011.05.010. [DOI] [PubMed] [Google Scholar]
  • 28.Roberts C.K., Barnard R.J., Jasman A., Balon T.W. Acute exercise increases nitric oxide synthase activity in skeletal muscle. J. Appl. Physiol. 1999;277(1985):E390–E394. doi: 10.1152/ajpendo.1999.277.2.E390. [DOI] [PubMed] [Google Scholar]
  • 29.Henriquez-Olguin C., et al. Cytosolic ROS production by NADPH oxidase 2 regulates muscle glucose uptake during exercise. Nat. Commun. 2019;10:4623. doi: 10.1038/s41467-019-12523-9. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 30.Xirouchaki C., et al. Skeletal muscle NOX4 is required for adaptive responses that prevent insulin resistance. Sci. Adv. 2021;7:1–24. doi: 10.1126/sciadv.abl4988. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 31.Sun Q.A., et al. Oxygen-coupled redox regulation of the skeletal muscle ryanodine receptor-Ca2+ release channel by NADPH oxidase 4. Proc. Natl. Acad. Sci. U. S. A. 2011;108:16098–16103. doi: 10.1073/pnas.1109546108. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 32.Cully T.R., Rodney G.G. Nox4 - RyR1 - Nox2: regulators of micro-domain signaling in skeletal muscle. Redox Biol. 2020;36 doi: 10.1016/j.redox.2020.101557. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 33.Henriquez-Olguin C., et al. NOX2 inhibition impairs early muscle gene expression induced by a single exercise bout. Front. Physiol. 2016;7:282. doi: 10.3389/fphys.2016.00282. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 34.Jones D.P. Redefining oxidative stress. Antioxidants Redox Signal. 2006;8:1865–1879. doi: 10.1089/ars.2006.8.1865. [DOI] [PubMed] [Google Scholar]
  • 35.Sies H. Oxidative stress: introductory remarks. Oxidative Stress. 1985;1–8 doi: 10.1016/B978-0-12-642760-8.50005-3. [DOI] [Google Scholar]
  • 36.Bhatti J.S., Bhatti G.K., Reddy P.H. Mitochondrial dysfunction and oxidative stress in metabolic disorders - a step towards mitochondria based therapeutic strategies. Biochim. Biophys. Acta, Mol. Basis Dis. 2017;1863:1066–1077. doi: 10.1016/j.bbadis.2016.11.010. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 37.Jones D.P. Redox theory of aging. Redox Biol. 2015;5:71–79. doi: 10.1016/j.redox.2015.03.004. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 38.Castejon-Vega B., Cordero M.D., Sanz A. How the disruption of mitochondrial redox signalling contributes to ageing. Antioxidants. 2023;12 doi: 10.3390/antiox12040831. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 39.Powers S.K., Wiggs M.P., Duarte J.A., Zergeroglu A.M., Demirel H.A. Mitochondrial signaling contributes to disuse muscle atrophy. Am. J. Physiol. Endocrinol. Metab. 2012;303:E31–E39. doi: 10.1152/ajpendo.00609.2011. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 40.Gouspillou G., Hepple R.T. Editorial: mitochondria in skeletal muscle health, aging and diseases. Front. Physiol. 2016;7:446. doi: 10.3389/fphys.2016.00446. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 41.Davies K.J.A., Quintanilha A.T., Brooks G.A., Packer L. Free radicals and tissue damage produced by exercise. Biochem. Biophys. Res. Commun. 1982;107:1198–1205. doi: 10.1016/s0006-291x(82)80124-1. [DOI] [PubMed] [Google Scholar]
  • 42.Bailey D.M., et al. Electron paramagnetic spectroscopic evidence of exercise-induced free radical accumulation in human skeletal muscle. Free Radic. Res. 2007;41:182–190. doi: 10.1080/10715760601028867. [DOI] [PubMed] [Google Scholar]
  • 43.Gomez-Cabrera M.C., et al. Oral administration of vitamin C decreases muscle mitochondrial biogenesis and hampers training-induced adaptations in endurance performance. Am. J. Clin. Nutr. 2008;87:142–149. doi: 10.1093/ajcn/87.1.142. [DOI] [PubMed] [Google Scholar]
  • 44.Ristow M., et al. Antioxidants prevent health-promoting effects of physical exercise in humans. Proc Natl Acad Sci. 2009;106:8665–8670. doi: 10.1073/pnas.0903485106. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 45.Bruns D.R., et al. Differential effects of vitamin C or protandim on skeletal muscle adaptation to exercise. J. Appl. Physiol. 2018;125(1985):661–671. doi: 10.1152/japplphysiol.00277.2018. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 46.McArdle F., et al. Preconditioning of skeletal muscle against contraction-induced damage: the role of adaptations to oxidants in mice. J. Physiol. 2004;561:233–244. doi: 10.1113/jphysiol.2004.069914. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 47.Li Y., Chen Y., Li A.S., Reid M.B. Hydrogen peroxide stimulates ubiquitin-conjugating activity and expression of genes for specific E2 and E3 proteins in skeletal muscle myotubes. Am J Physiol Cell Physiol. 2003;285:C806–C812. doi: 10.1152/ajpcell.00129.2003. [DOI] [PubMed] [Google Scholar]
  • 48.Franco A.A., Odom R.S., Rando T.A. Regulation of antioxidant enzyme gene expression in response to oxidative stress and during differentiation of mouse skeletal muscle. Free Radic. Biol. Med. 1999;27:1122–1132. doi: 10.1016/s0891-5849(99)00166-5. [DOI] [PubMed] [Google Scholar]
  • 49.Irrcher I., Ljubicic V., Hood D.A. Interactions between ROS and AMP kinase activity in the regulation of PGC-1alpha transcription in skeletal muscle cells. Am J Physiol Cell Physiol. 2009;296:C116–C123. doi: 10.1152/ajpcell.00267.2007. [DOI] [PubMed] [Google Scholar]
  • 50.Silveira L.R., Pilegaard H., Kusuhara K., Curi R., Hellsten Y. The contraction induced increase in gene expression of peroxisome proliferator-activated receptor (PPAR)-gamma coactivator 1alpha (PGC-1alpha), mitochondrial uncoupling protein 3 (UCP3) and hexokinase II (HKII) in primary rat skeletal muscle cells is dependent on reactive oxygen species. Biochim. Biophys. Acta. 2006;1763:969–976. doi: 10.1016/j.bbamcr.2006.06.010. [DOI] [PubMed] [Google Scholar]
  • 51.Laker R.C., et al. Ampk phosphorylation of Ulk1 is required for targeting of mitochondria to lysosomes in exercise-induced mitophagy. Nat. Commun. 2017;8:548. doi: 10.1038/s41467-017-00520-9. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 52.Fan X., Hussien R., Brooks G.A. H2O2-induced mitochondrial fragmentation in C2C12 myocytes. Free Radic. Biol. Med. 2010;49:1646–1654. doi: 10.1016/j.freeradbiomed.2010.08.024. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 53.Scherz-Shouval R., et al. Reactive oxygen species are essential for autophagy and specifically regulate the activity of Atg4. EMBO J. 2007;26:1749–1760. doi: 10.1038/sj.emboj.7601623. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 54.Wright V.P., Reiser P.J., Clanton T.L. Redox modulation of global phosphatase activity and protein phosphorylation in intact skeletal muscle. J. Physiol. 2009;587:5767–5781. doi: 10.1113/jphysiol.2009.178285. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 55.Powers S.K., Duarte J., Kavazis A.N., Talbert E.E. Reactive oxygen species are signalling molecules for skeletal muscle adaptation. Exp. Physiol. 2010;95:1–9. doi: 10.1113/expphysiol.2009.050526. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 56.Martinez Guimera A., Welsh C.M., Proctor C.J., McArdle A., Shanley D.P. 'Molecular habituation' as a potential mechanism of gradual homeostatic loss with age. Mech. Ageing Dev. 2018;169:53–62. doi: 10.1016/j.mad.2017.11.010. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 57.Guimera A.M., Shanley D.P., Proctor C.J. Modelling the role of redox-related mechanisms in musculoskeletal ageing. Free Radic. Biol. Med. 2019;132:11–18. doi: 10.1016/j.freeradbiomed.2018.09.013. [DOI] [PubMed] [Google Scholar]
  • 58.Plecita-Hlavata L., et al. Potential of mitochondria-targeted antioxidants to prevent oxidative stress in pancreatic beta-cells. Oxid. Med. Cell. Longev. 2019;2019 doi: 10.1155/2019/1826303. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 59.Musci R.V., Hamilton K.L., Linden M.A. Exercise-induced mitohormesis for the maintenance of skeletal muscle and healthspan extension. Sports (Basel) 2019;7 doi: 10.3390/sports7070170. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 60.Sparks L.M., et al. Differences in mitochondrial coupling reveal a novel signature of mitohormesis in muscle of healthy individuals. J. Clin. Endocrinol. Metab. 2016;101:4994–5003. doi: 10.1210/jc.2016-2742. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 61.Miller C.J., et al. Disruption of Nrf2/ARE signaling impairs antioxidant mechanisms and promotes cell degradation pathways in aged skeletal muscle. Biochim. Biophys. Acta. 2012;1822:1038–1050. doi: 10.1016/j.bbadis.2012.02.007. [DOI] [PubMed] [Google Scholar]
  • 62.Vasilaki A., et al. Free radical generation by skeletal muscle of adult and old mice: effect of contractile activity. Aging Cell. 2006;5:109–117. doi: 10.1111/j.1474-9726.2006.00198.x. [DOI] [PubMed] [Google Scholar]
  • 63.Close G.L., Kayani A.C., Ashton T., McArdle A., Jackson M.J. Release of superoxide from skeletal muscle of adult and old mice: an experimental test of the reductive hotspot hypothesis. Aging Cell. 2007;6:189–195. doi: 10.1111/j.1474-9726.2007.00277.x. [DOI] [PubMed] [Google Scholar]
  • 64.Phaniendra A., Jestadi D.B., Periyasamy L. Free radicals: properties, sources, targets, and their implication in various diseases. Indian J. Clin. Biochem. 2015;30:11–26. doi: 10.1007/s12291-014-0446-0. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 65.Jang Y.C., Van Remmen H. The mitochondrial theory of aging: insight from transgenc and knockout mouse models. Exp. Gerontol. 2009;44:256–260. doi: 10.1016/j.exger.2008.12.006. [DOI] [PubMed] [Google Scholar]
  • 66.Jia Q., Sieburth D. Mitochondrial hydrogen peroxide positively regulates neuropeptide secretion during diet-induced activation of the oxidative stress response. Nat. Commun. 2021;12:2304. doi: 10.1038/s41467-021-22561-x. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 67.Campbell M.D., et al. Improving mitochondrial function with SS-31 reverses age-related redox stress and improves exercise tolerance in aged mice. Free Radic. Biol. Med. 2019;134:268–281. doi: 10.1016/j.freeradbiomed.2018.12.031. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 68.Hoehne M.N., et al. Spatial and temporal control of mitochondrial H(2) O(2) release in intact human cells. EMBO J. 2022;41 doi: 10.15252/embj.2021109169. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 69.Fernandez-Puente E., et al. Expression and functional analysis of the hydrogen peroxide biosensors HyPer and HyPer2 in C2C12 myoblasts/myotubes and single skeletal muscle fibres. Sci. Rep. 2020;10:871. doi: 10.1038/s41598-020-57821-1. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 70.Granatiero V., et al. Role of p66shc in skeletal muscle function. Sci. Rep. 2017;7:6283. doi: 10.1038/s41598-017-06363-0. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 71.Vendrov A.E., et al. NOX4 NADPH Oxidase-dependent mitochondrial oxidative stress in aging-associated cardiovascular disease. Antioxidants Redox Signal. 2015;23 doi: 10.1089/ars.2014.6221. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 72.Shanmugasundaram K., et al. NOX4 functions as a mitochondrial energetic sensor coupling cancer metabolic reprogramming to drug resistance. Nat. Commun. 2017;8:997. doi: 10.1038/s41467-017-01106-1. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 73.Onukwufor J.O., et al. A reversible mitochondrial complex I thiol switch mediates hypoxic avoidance behavior in C. elegans. Nat. Commun. 2022;13:2403. doi: 10.1038/s41467-022-30169-y. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 74.Glancy B., et al. Power grid protection of the muscle mitochondrial reticulum. Cell Rep. 2017;19:487–496. doi: 10.1016/j.celrep.2017.03.063. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 75.Glancy B., et al. Mitochondrial reticulum for cellular energy distribution in muscle. Nature. 2015;523:617–620. doi: 10.1038/nature14614. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 76.Bleck C.K.E., Kim Y., Willingham T.B., Glancy B. Subcellular connectomic analyses of energy networks in striated muscle. Nat. Commun. 2018;9:5111. doi: 10.1038/s41467-018-07676-y. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 77.Kirkwood S.P., M E.A., Brooks G.A. Mitochondrial reticulum in limb skeletal muscle. Am. J. Physiol. 1986;251:C395–C402. doi: 10.1152/ajpcell.1986.251.3.C395. [DOI] [PubMed] [Google Scholar]
  • 78.Ghosh S., et al. Insights on the impact of mitochondrial organisation on bioenergetics in high-resolution computational models of cardiac cell architecture. PLoS Comput. Biol. 2018;14 doi: 10.1371/journal.pcbi.1006640. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 79.Drake J.C., Wilson R.J., Yan Z. Molecular mechanisms for mitochondrial adaptation to exercise training in skeletal muscle. FASEB J. 2016;30:13–22. doi: 10.1096/fj.15-276337. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 80.Nichenko A.S., Specht K.S., Craige S.M., Drake J.C. Sensing local energetics to acutely regulate mitophagy in skeletal muscle. Front. Cell Dev. Biol. 2022;10 doi: 10.3389/fcell.2022.987317. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 81.Drake J.C., Yan Z. Precision remodeling: how exercise improves mitochondrial quality in myofibers. Current Opinion in Physiology. 2019;10:96–101. doi: 10.1016/j.cophys.2019.05.005. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 82.Tilokani L., Nagashima S., Paupe V., Prudent J. Mitochondrial dynamics: overview of molecular mechanisms. Essays Biochem. 2018;62:341–360. doi: 10.1042/EBC20170104. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 83.Yapa N.M.B., Lisnyak V., Reljic B., Ryan M.T. Mitochondrial dynamics in health and disease. FEBS Lett. 2021;595:1184–1204. doi: 10.1002/1873-3468.14077. [DOI] [PubMed] [Google Scholar]
  • 84.Busquets-Cortes C., et al. Training enhances immune cells mitochondrial biosynthesis, fission, fusion, and their antioxidant capabilities synergistically with dietary docosahexaenoic supplementation. Oxid. Med. Cell. Longev. 2016;2016 doi: 10.1155/2016/8950384. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 85.Jendrach M., Mai S., Pohl S., Voth M., Bereiter-Hahn J. Short- and long-term alterations of mitochondrial morphology, dynamics and mtDNA after transient oxidative stress. Mitochondrion. 2008;8 doi: 10.1016/j.mito.2008.06.001. [DOI] [PubMed] [Google Scholar]
  • 86.Kim Y.M., et al. Redox regulation of mitochondrial fission protein drp1 by protein Disulfide Isomerase limits endothelial senescence. Cell Rep. 2018;23:3565–3578. doi: 10.1016/j.celrep.2018.05.054. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 87.Moore T.M., et al. The impact of exercise on mitochondrial dynamics and the role of Drp1 in exercise performance and training adaptations in skeletal muscle. Mol. Metabol. 2019;21:51–67. doi: 10.1016/j.molmet.2018.11.012. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 88.Axelrod C.L., Fealy C.E., Mulya A., Kirwan J.P. Exercise training remodels human skeletal muscle mitochondrial fission and fusion machinery towards a pro-elongation phenotype. Acta Physiol. 2019;225 doi: 10.1111/apha.13216. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 89.Trewin A.J., Berry B.J., Wojtovich A.P. Exercise and mitochondrial dynamics: keeping in shape with ROS and AMPK. Antioxidants. 2018;7 doi: 10.3390/antiox7010007. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 90.Cid-Castro C., Moran J. Differential ROS-mediated phosphorylation of Drp1 in mitochondrial fragmentation induced by distinct cell death conditions in cerebellar granule neurons. Oxid. Med. Cell. Longev. 2021;2021 doi: 10.1155/2021/8832863. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 91.Fiorenza M., et al. Metabolic stress-dependent regulation of the mitochondrial biogenic molecular response to high-intensity exercise in human skeletal muscle. J. Physiol. 2018;596:2823–2840. doi: 10.1113/JP275972. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 92.Mesquita P.H.C., et al. Acute and chronic effects of resistance training on skeletal muscle markers of mitochondrial remodeling in older adults. Physiol Rep. 2020;8 doi: 10.14814/phy2.14526. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 93.Campos J.C., et al. Exercise preserves physical fitness during aging through AMPK and mitochondrial dynamics. Proc. Natl. Acad. Sci. U. S. A. 2023;120 doi: 10.1073/pnas.2204750120. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 94.Joseph A.M., et al. The impact of aging on mitochondrial function and biogenesis pathways in skeletal muscle of sedentary high- and low-functioning elderly individuals. Aging Cell. 2012;11:801–809. doi: 10.1111/j.1474-9726.2012.00844.x. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 95.Xia Q., et al. Peroxiredoxin 2 regulates DAF-16/FOXO mediated mitochondrial remodelling in response to exercise that is disrupted in ageing. Mol. Metabol. 2024;88 doi: 10.1016/j.molmet.2024.102003. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 96.Masiero E., et al. Autophagy is required to maintain muscle mass. Cell Metabol. 2009;10:507–515. doi: 10.1016/j.cmet.2009.10.008. [DOI] [PubMed] [Google Scholar]
  • 97.Bin-Umer M.A., McLaughlin J.E., Butterly M.S., McCormick S., Tumer N.E. Elimination of damaged mitochondria through mitophagy reduces mitochondrial oxidative stress and increases tolerance to trichothecenes. Proc. Natl. Acad. Sci. U. S. A. 2014;111:11798–11803. doi: 10.1073/pnas.1403145111. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 98.Yamashita S.I., et al. Mitophagy reporter mouse analysis reveals increased mitophagy activity in disuse-induced muscle atrophy. J. Cell. Physiol. 2021;236:7612–7624. doi: 10.1002/jcp.30404. [DOI] [PubMed] [Google Scholar]
  • 99.Sciarretta S., Volpe M., Sadoshima J. NOX4 regulates autophagy during energy deprivation. Autophagy. 2014;10:699–701. doi: 10.4161/auto.27955. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 100.Sobhakumari A., et al. NOX4 mediates cytoprotective autophagy induced by the EGFR inhibitor erlotinib in head and neck cancer cells. Toxicol. Appl. Pharmacol. 2013;272:736–745. doi: 10.1016/j.taap.2013.07.013. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 101.Iqbal S., Hood D.A. Oxidative stress-induced mitochondrial fragmentation and movement in skeletal muscle myoblasts. Am J Physiol Cell Physiol. 2014;306:C1176–C1183. doi: 10.1152/ajpcell.00017.2014. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 102.Drake J.C., Yan Z. Mitophagy in maintaining skeletal muscle mitochondrial proteostasis and metabolic health with ageing. J. Physiol. 2017;595:6391–6399. doi: 10.1113/JP274337. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 103.Di Malta C., Cinque L., Settembre C. Transcriptional regulation of autophagy: mechanisms and diseases. Front. Cell Dev. Biol. 2019;7:114. doi: 10.3389/fcell.2019.00114. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 104.Liu L., et al. Mitophagy receptor FUNDC1 is regulated by PGC-1alpha/NRF1 to fine tune mitochondrial homeostasis. EMBO Rep. 2021;22 doi: 10.15252/embr.202050629. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 105.Schmitt D.L., et al. Spatial regulation of AMPK signaling revealed by a sensitive kinase activity reporter. Nat. Commun. 2022;13 doi: 10.1038/s41467-022-31190-x. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 106.Drake J.C., et al. Mitochondria-localized AMPK responds to local energetics and contributes to exercise and energetic stress-induced mitophagy. Proc. Natl. Acad. Sci. U. S. A. 2021;118:1–10. doi: 10.1073/pnas.2025932118. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 107.Zong Y., et al. Hierarchical activation of compartmentalized pools of AMPK depends on severity of nutrient or energy stress. Cell Res. 2019;29:460–473. doi: 10.1038/s41422-019-0163-6. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 108.Ma L., et al. Exercise protects aged mice against coronary endothelial senescence via FUNDC1-dependent mitophagy. Redox Biol. 2023;62 doi: 10.1016/j.redox.2023.102693. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 109.Gouspillou G., et al. Protective role of Parkin in skeletal muscle contractile and mitochondrial function. J. Physiol. 2018;596:2565–2579. doi: 10.1113/JP275604. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 110.Leduc-Gaudet J.P., Reynaud O., Hussain S.N., Gouspillou G. Parkin overexpression protects from ageing-related loss of muscle mass and strength. J. Physiol. 2019;597:1975–1991. doi: 10.1113/JP277157. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 111.Sakellariou G.K., et al. Mitochondrial ROS regulate oxidative damage and mitophagy but not age-related muscle fiber atrophy. Sci. Rep. 2016;6 doi: 10.1038/srep33944. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 112.Holloszy J.O. Biochemical adaptations in muscle. Effects of exercise on mitochondrial oxygen uptake and respiratory enzyme activity in skeletal muscle. J. Biol. Chem. 1967;242:2278–2282. [PubMed] [Google Scholar]
  • 113.Yang S., et al. Functional effects of muscle PGC-1alpha in aged animals. Skelet Muscle. 2020;10:14. doi: 10.1186/s13395-020-00231-8. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 114.Robinson M.M., et al. Enhanced protein translation underlies improved metabolic and physical adaptations to different exercise training modes in young and old humans. Cell Metabol. 2017;25:581–592. doi: 10.1016/j.cmet.2017.02.009. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 115.Miller B.F., Robinson M.M., Bruss M.D., Hellerstein M., Hamilton K.L. A comprehensive assessment of mitochondrial protein synthesis and cellular proliferation with age and caloric restriction. Aging Cell. 2012;11:150–161. doi: 10.1111/j.1474-9726.2011.00769.x. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 116.Moore J., et al. Random errors in protein synthesis activate an age-dependent program of muscle atrophy in mice. Commun. Biol. 2021;4:703. doi: 10.1038/s42003-021-02204-z. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 117.Shcherbakov D., et al. Premature aging in mice with error-prone protein synthesis. Sci. Adv. 2022;8 doi: 10.1126/sciadv.abl9051. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 118.Ke Z., et al. Translation fidelity coevolves with longevity. Aging Cell. 2017;16:988–993. doi: 10.1111/acel.12628. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 119.Brilkova M., et al. Error-prone protein synthesis recapitulates early symptoms of Alzheimer disease in aging mice. Cell Rep. 2022;40 doi: 10.1016/j.celrep.2022.111433. [DOI] [PubMed] [Google Scholar]
  • 120.Kang C., O'Moore K.M., Dickman J.R., Ji L.L. Exercise activation of muscle peroxisome proliferator-activated receptor-gamma coactivator-1alpha signaling is redox sensitive. Free Radic. Biol. Med. 2009;47:1394–1400. doi: 10.1016/j.freeradbiomed.2009.08.007. [DOI] [PubMed] [Google Scholar]
  • 121.Xiao B., et al. Structural basis for AMP binding to mammalian AMP-activated protein kinase. Nature. 2007;449:496–500. doi: 10.1038/nature06161. [DOI] [PubMed] [Google Scholar]
  • 122.Xiao B., et al. Structure of mammalian AMPK and its regulation by ADP. Nature. 2011;472:230–233. doi: 10.1038/nature09932. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 123.Morales-Alamo D., Calbet J.A.L. AMPK signaling in skeletal muscle during exercise: role of reactive oxygen and nitrogen species. Free Radic. Biol. Med. 2016;98:68–77. doi: 10.1016/j.freeradbiomed.2016.01.012. [DOI] [PubMed] [Google Scholar]
  • 124.Shao D., et al. A redox-dependent mechanism for regulation of AMPK activation by Thioredoxin1 during energy starvation. Cell Metabol. 2014;19:232–245. doi: 10.1016/j.cmet.2013.12.013. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 125.Zmijewski J.W., et al. Exposure to hydrogen peroxide induces oxidation and activation of AMP-activated protein kinase. J. Biol. Chem. 2010;285:33154–33164. doi: 10.1074/jbc.M110.143685. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 126.Emerling B.M., et al. Hypoxic activation of AMPK is dependent on mitochondrial ROS but independent of an increase in AMP/ATP ratio. Free Radic. Biol. Med. 2009;46:1386–1391. doi: 10.1016/j.freeradbiomed.2009.02.019. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 127.Block K., Gorin Y., Abboud H.E. Subcellular localization of Nox4 and regulation in diabetes. Proc. Natl. Acad. Sci. U. S. A. 2009;106:14385–14390. doi: 10.1073/pnas.0906805106. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 128.Sakellariou G.K., et al. 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. Antioxidants Redox Signal. 2013;18:603–621. doi: 10.1089/ars.2012.4623. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 129.Reznick R.M., et al. Aging-associated reductions in AMP-activated protein kinase activity and mitochondrial biogenesis. Cell Metabol. 2007;5:151–156. doi: 10.1016/j.cmet.2007.01.008. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 130.Hardman S.E., Hall D.E., Cabrera A.J., Hancock C.R., Thomson D.M. The effects of age and muscle contraction on AMPK activity and heterotrimer composition. Exp. Gerontol. 2014;55:120–128. doi: 10.1016/j.exger.2014.04.007. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 131.Sun S., et al. A single-cell transcriptomic atlas of exercise-induced anti-inflammatory and geroprotective effects across the body. Innovation. 2023;4 doi: 10.1016/j.xinn.2023.100380. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 132.Hoppeler H., Hudlicka O., Uhlmann E. Relationship between mitochondria and oxygen consumption in isolated cat muscles. J. Physiol. 1987;385:661–675. doi: 10.1113/jphysiol.1987.sp016513. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 133.Kayar S.R., Hoppeler H., Mermod L., Weibel E.R. Mitochondrial size and shape in equine skeletal muscle: a three-dimensional reconstruction study. Anat. Rec. 1988;222:333–339. doi: 10.1002/ar.1092220405. [DOI] [PubMed] [Google Scholar]
  • 134.Kusters Y.H., Barrett E.J. Muscle microvasculature's structural and functional specializations facilitate muscle metabolism. Am. J. Physiol. Endocrinol. Metab. 2016;310:E379–E387. doi: 10.1152/ajpendo.00443.2015. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 135.Larsson L., et al. Sarcopenia: aging-related loss of muscle mass and function. Physiol. Rev. 2019;99:427–511. doi: 10.1152/physrev.00061.2017. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 136.Lovric A., et al. Single-cell sequencing deconvolutes cellular responses to exercise in human skeletal muscle. Commun. Biol. 2022;5:1121. doi: 10.1038/s42003-022-04088-z. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 137.Glancy B., et al. In vivo microscopy reveals extensive embedding of capillaries within the sarcolemma of skeletal muscle fibers. Microcirculation. 2014;21:131–147. doi: 10.1111/micc.12098. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 138.Willingham T.B., Ajayi P.T., Glancy B. Subcellular specialization of mitochondrial form and function in skeletal muscle cells. Front. Cell Dev. Biol. 2021;9 doi: 10.3389/fcell.2021.757305. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 139.Kim Y., Lindberg E., Bleck C.K.E., Glancy B. Endothelial cell nanotube insertions into cardiac and skeletal myocytes during coordinated tissue development. Cardiovasc. Res. 2020;116:260–261. doi: 10.1093/cvr/cvz285. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 140.Canugovi C., et al. Increased mitochondrial NADPH oxidase 4 (NOX4) expression in aging is a causative factor in aortic stiffening. Redox Biol. 2019;26 doi: 10.1016/j.redox.2019.101288. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 141.Takac I., et al. The E-loop is involved in hydrogen peroxide formation by the NADPH oxidase Nox4. J. Biol. Chem. 2011;286:13304–13313. doi: 10.1074/jbc.M110.192138. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 142.Nisimoto Y., Diebold B.A., Cosentino-Gomes D., Lambeth J.D. Nox4: a hydrogen peroxide-generating oxygen sensor. Biochemistry. 2014;53:5111–5120. doi: 10.1021/bi500331y. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 143.Craige S.M., et al. NADPH oxidase 4 promotes endothelial angiogenesis through endothelial nitric oxide synthase activation. Circulation. 2011;124:731–740. doi: 10.1161/CIRCULATIONAHA.111.030775. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 144.Schroder K., et al. Nox4 is a protective reactive oxygen species generating vascular NADPH oxidase. Circ. Res. 2012;110:1217–1225. doi: 10.1161/CIRCRESAHA.112.267054. [DOI] [PubMed] [Google Scholar]
  • 145.Thomas S.R., Chen K., Keaney J.F., Jr. Hydrogen peroxide activates endothelial nitric-oxide synthase through coordinated phosphorylation and dephosphorylation via a phosphoinositide 3-kinase-dependent signaling pathway. J. Biol. Chem. 2002;277:6017–6024. doi: 10.1074/jbc.M109107200. [DOI] [PubMed] [Google Scholar]
  • 146.Cai H., et al. Akt-dependent phosphorylation of serine 1179 and mitogen-activated protein kinase kinase/extracellular signal-regulated kinase 1/2 cooperatively mediate activation of the endothelial nitric-oxide synthase by hydrogen peroxide. Mol. Pharmacol. 2003;63:325–331. doi: 10.1124/mol.63.2.325. [DOI] [PubMed] [Google Scholar]
  • 147.Drummond G.R., Cai H., Davis M.E., Ramasamy S., Harrison D.G. Transcriptional and posttranscriptional regulation of endothelial nitric oxide synthase expression by hydrogen peroxide. Circ. Res. 2000;86:347–354. doi: 10.1161/01.res.86.3.347. [DOI] [PubMed] [Google Scholar]
  • 148.Momken I., et al. Endothelial nitric oxide synthase (NOS) deficiency affects energy metabolism pattern in murine oxidative skeletal muscle. Biochem. J. 2002;368:341–347. doi: 10.1042/BJ20020591. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 149.Le Gouill E., et al. Endothelial nitric oxide synthase (eNOS) knockout mice have defective mitochondrial beta-oxidation. Diabetes. 2007;56:2690–2696. doi: 10.2337/db06-1228. [DOI] [PubMed] [Google Scholar]
  • 150.Tengan C.H., Rodrigues G.S., Godinho R.O. Nitric oxide in skeletal muscle: role on mitochondrial biogenesis and function. Int. J. Mol. Sci. 2012;13:17160–17184. doi: 10.3390/ijms131217160. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 151.Lee-Young R.S., et al. Endothelial nitric oxide synthase is central to skeletal muscle metabolic regulation and enzymatic signaling during exercise in vivo. Am. J. Physiol. Regul. Integr. Comp. Physiol. 2010;298:R1399–R1408. doi: 10.1152/ajpregu.00004.2010. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 152.Cunningham R.P., et al. Hepatocyte-specific eNOS deletion impairs exercise-induced adaptations in hepatic mitochondrial function and autophagy. Obesity. 2022;30:1066–1078. doi: 10.1002/oby.23414. [DOI] [PMC free article] [PubMed] [Google Scholar]

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