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Mitochondrial Dynamics and Mitophagy in Skeletal Muscle Health and Aging

Jean-Philippe Leduc-Gaudet, Sabah N. A. Hussain, Esther Barreiro and 1 more

International Journal of Molecular Sciences · 2021 · Review · CC BY

doi:10.3390/ijms22158179

Abstract

The maintenance of mitochondrial integrity is critical for muscle health. Mitochondria, indeed, play vital roles in a wide range of cellular processes, including energy supply, Ca2+ homeostasis, retrograde signaling, cell death, and many others. All mitochondria-containing cells, including skeletal muscle cells, dispose of several pathways to maintain mitochondrial health, including mitochondrial biogenesis, mitochondrial-derived vesicles, mitochondrial dynamics (fusion and fission process shaping mitochondrial morphology), and mitophagy—the process in charge of the removal of mitochondria though autophagy. The loss of skeletal muscle mass (atrophy) is a major health problem worldwide, especially in older people. Currently, there is no treatment to counteract the progressive decline in skeletal muscle mass and strength that occurs with aging, a process termed sarcopenia. There is increasing data, including our own, suggesting that accumulation of dysfunctional mitochondria contributes to the development of sarcopenia. Impairments in mitochondrial dynamics and mitophagy were recently proposed to contribute to sarcopenia. This review summarizes the current state of knowledge on the role played by mitochondrial dynamics and mitophagy in skeletal muscle health and in the development of sarcopenia. We also highlight recent studies showing that enhancing mitophagy in skeletal muscle is a promising therapeutic target to prevent or even treat skeletal muscle dysfunction in the elderly.

Full text

From PubMed Central, U.S. National Library of Medicine, as current on Oct 19, 2026. Paragraph numbers are ours, so an answer can point at one.

1. Introduction

Skeletal muscle, which accounts for over 40% of total body mass, is highly adaptable and of critical importance for general health. Indeed, it plays key roles in posture, mobility, thermogenesis, glucose homeostasis, and energy metabolism. The regulation of skeletal muscle mass is mainly controlled by a fine balance between protein synthesis and protein degradation. Skeletal muscle has a striking ability to adapt to various physiological and pathological conditions by changing its size, composition, and metabolic properties. Muscle atrophy, or the loss of muscle mass, is an important clinical problem that leads to negative health outcomes, especially during aging.

The progressive decline in muscle mass and strength that occurs with aging, a process known as sarcopenia, can have serious consequences on the lifestyle and functional status of the elderly. For instance, lower muscle mass and strength has been associated with functional impairment, falls, and physical disability. Sarcopenia is a strong predictor of hospitalization and is highly linked to short- and long-term mortality in older adults. Given the increasing population of older adults all over the world, it is not surprising that the age-related decline in muscle mass, strength, and function has become a major global health concern. Highlighting its serious health consequences, sarcopenia was recognized in 2016 as a muscle condition/disease and received its own ICD (international classification of diseases)-10-Clinical Modification code (M62.84).

Several biological mechanisms have been proposed to explain the development of sarcopenia. Among these mechanisms, strong experimental evidence indicates that the accumulation of mitochondrial dysfunction plays an important role in the muscle aging process. Indeed, many studies conducted in both rodents and humans have shown that aging is associated with a decline in mitochondrial bioenergetics. Several studies have also shown that the mitochondrial permeability transition pore (mPTP) becomes dysfunctional in old rodents and humans. Alterations in mPTP function are undoubtedly contributing to the aging-related loss of muscle mass and function, since mPTP opening can trigger several atrophy-regulating pathways. First, mPTP opening can result in mitochondrial reactive oxygen species (ROS) overproduction. ROS can, in turn, trigger the muscle atrophy program by activating the Fork head box O (FoxO) transcription factor family. Second, mPTP opening can result in mitochondrial DNA release and consequential activation of the NLRP3 (NOD-, LRR- and pyrin domain-containing protein 3) inflammasome, which was shown to increase the expression of ubiquitin ligases involved in muscle atrophy. Third, cytochrome C release secondary to mPTP opening was shown to increase proteasomal activity secondary to caspase 3 activation. Fourth, mPTP opening can lead to the release of pro-apoptotic factors such as endonuclease G, which was shown to accumulate in myonuclei of old individuals. Further strengthening the view that mitochondria play key roles in the aging-related loss of muscle mass and function, the most efficient interventions to attenuate sarcopenia are exercise training and calorie restriction, both well-known to improve mitochondrial health.

While the accumulation of mitochondrial dysfunction is now generally seen as a hallmark of muscle aging, the mechanisms underlying the progressive decline in mitochondrial function remain poorly understood. In this article, we critically review the available literature that has positioned impairments in mitochondrial dynamics and mitophagy as potential contributors. We also review recent evidence showing that targeting mitophagy might be a promising strategy to prevent or even treat sarcopenia.

2. The Importance of Mitochondria Dynamics for Skeletal Muscle Health

Initially considered as bean-shaped organelles, mitochondria are now known to display a complex architecture, especially in skeletal muscle cells. Mitochondria are dynamic organelles that are constantly undergoing fission and fusion so as to adapt their morphology to the cellular environment (Figure 1). Mitochondrial fusion is mainly regulated by the Mitofusin 1 & 2 (Mfn 1 & 2) and OPtic Atrophy 1 (Opa 1) proteins. Mitochondrial fission is mainly governed by Dynamin-Related Protein 1 (Drp1) (reviewed in). Several other proteins targeting the outer membrane, such as mitochondrial fission 1 protein (Fis1), mitochondrial fission factor (Mff), and mitochondrial dynamic protein 49 and 51 (MiD49, and MiD51), participate in the fragmentation of mitochondria. Figure 1 shows the known regulators involved in mitochondrial fusion and fission.

In the past decade, several studies have established the importance of mitochondrial dynamics in maintaining skeletal muscle and mitochondrial integrity and function. It was, for instance, reported that mice lacking both Mfn-1 and Mfn-2 in skeletal muscle display severe mitochondrial dysfunction, accumulation of mitochondrial DNA damage, and a severe deficit in growth. Mfn2 deletion in skeletal muscle was shown to result in oxidative stress and muscle atrophy in adult mice. Deletion of both Mfn1 and Mfn2 in adult skeletal muscle resulted in a major decrease in exercise performance, indicating that mitochondrial fusion is essential to muscle performance in mice.

Specific deletion of Opa1 in skeletal muscle leads to mitochondrial dysfunction, oxidative stress, endoplasmic reticulum (ER) stress, and inflammation. Opa1 deficiency also promotes the secretion of Fibroblast Growth Factor 21 (FGF21) from skeletal muscle, which leads to altered lipid homeostasis, inflammation, and the senescence of different tissues. Taken altogether, the available literature clearly indicates that mitochondrial fusion proteins are essential for the maintenance of mitochondrial and skeletal muscle health.

Several recent studies have highlighted the physiological importance of mitochondrial fission in the maintenance of skeletal muscle health. Muscle-specific Drp1 overexpression was shown to impair skeletal muscle growth in mice. Inhibition of mitochondrial fission by genetic silencing of Fis1 and Drp1 in skeletal muscle has been shown to protect against muscle wasting induced by starvation or by overexpression of the atrophying transcription factor FoxO3a. Muscle-specific Drp1 knockout in mice leads to a severe myopathic phenotype including muscle wasting, weakness, and signs of muscle degeneration and regeneration. The myopathic phenotype was evident about 2 months after muscle deletion of Drp1 and was also present at 6 months. The importance of Drp1 in regulating muscle function was further emphasized by our group, which revealed that muscle-selective Drp1 knockdown decreases mitochondrial respiration and increases markers of muscle regeneration, denervation, fibrosis, and oxidative stress. Interestingly, both Drp1 knockout and knockdown altered autophagy and mitophagy levels. As mentioned above, overexpression of Drp1 impaired skeletal muscle growth, whereas deletion or insufficient expression of Drp1 in skeletal muscle led to 40–50% atrophy, indicating that Drp1 is a key player in regulating skeletal muscle development and maintenance. Evidence from Drosophila also indicates that manipulating mitochondrial dynamics can impact myofibril development. Another recent study showed that loss of Fis1 leads to mitochondrial dysfunction, proteostasis impairment, muscle degeneration, and reduced lifespan in Fis1 mutant flies. Taken altogether, the findings discussed above clearly indicate that mitochondrial fission is essential for skeletal muscle health.

3. Mitochondrial Dynamics in the Skeletal Muscle Aging Process: Current State of Knowledge and Considerations for Future Research

As highlighted above, the available literature clearly positions mitochondrial dynamics as key processes in the maintenance of muscle and mitochondrial health. It is, therefore, not surprising that many studies have investigated whether impairment in mitochondrial dynamics could contribute to sarcopenia. However, and as will be reviewed below, the involvement of mitochondrial dynamics in the skeletal muscle aging process remains unclear as conflicting data exist in the field (Table 1).

In their elegant study, Sebastian et al. reported that mice lacking Mfn2 in skeletal muscle from birth display accelerated sarcopenia. They also reported that aged control (wild type, WT) mice displayed a reduction in Mfn2 content. However, the finding that Mfn2 content declines with aging is far from being universal. Indeed, Mfn2 content has been reported as decreased, unchanged, or even increased in aged rodent skeletal muscles. In humans, while Mfn2 expression was decreased in aged skeletal muscle in one study, others have reported either similar or even higher Mfn2 content in aged skeletal muscles. Smartly conflicting data also exist for Drp1 and Opa1.

In a recent study, Tezze et al. reported that Opa1 content normalized to complex II was lower in muscle from old sedentary individuals but not in old sportsmen. Interestingly, Opa1 content normalized to complex II content correlated with muscle fiber diameter and specific force in elderly individuals. Somewhat in line with this finding, Joseph et al. reported that Opa1 content was lower in the skeletal muscle of both low functioning and high functioning individuals. However, several studies did not detect any impact of aging on Opa1 content in the skeletal muscle of aged mice, rats, and humans. It is worth mentioning that Tezze et al. also reported that 18-month-old mice display lower levels of Opa1, but also showed that one week of exercise training was sufficient to restore Opa1 in aged skeletal muscles.

Conflicting data also surround the impact of aging on Drp1 content. Indeed, studies in rodents have reported increased or unchanged Drp1 content in aged skeletal muscles. The literature in humans also contains conflicting data, with some studies reporting a decreased or an unchanged Drp1 content in skeletal muscle of old individuals.

Clearly, no consensus on the role that mitochondrial dynamics play in the muscle aging process has been reached. Recently developed transgenic mouse lines with photoactivatable mitochondria might prove useful in addressing this issue. Defining whether impairment in mitochondrial dynamics is a likely contributor to the aging-related loss of muscle mass and strength in humans will require longitudinal studies. Such studies will need to pay particular attention to physical activity levels of participants, as the latter can impact the expression of proteins regulating mitochondrial dynamics. Further studies will also need to take into account the possibility that the impact of aging on mitochondrial dynamics and morphology might be fiber type-specific. Indeed, we recently reported in aged rats that the oxidative soleus muscle displayed signs of mitochondrial fragmentation, while the glycolytic white gastrocnemius displayed signs of mitochondrial elongation with aging. Other studies have also highlighted that mitochondrial morphology and dynamics differ across muscle fiber types. Importantly, a major gap in knowledge in the field that needs to be addressed urgently is whether sex differences exist in the role that mitochondrial dynamics play in the skeletal muscle aging process. Indeed, and to the best of our knowledge, no previous study has investigated whether the content of proteins regulating mitochondrial dynamics is differently affected in aged males vs. females.

Studies aiming at defining whether mitochondrial dynamics can be targeted to counteract sarcopenia are also needed. To date, the modulation (overexpression or deletion) of proteins regulating mitochondrial dynamics in mice has either shown no benefit to muscle cells or was proven deleterious to muscle health (Drp1 overexpression, Drp1 knockout, Drp1 knockdown, and Mfn1+Mfn2 knockout). Strengthening the need for such mechanistic studies, promoting Drp1-mediated mitochondrial fission in middle-aged flies was shown to extend lifespan and improve muscle mitochondrial function, likely by facilitating mitophagy (mitochondrial fission can, indeed, facilitate the removal of damaged/dysfunctional mitochondria through mitophagy). This study, therefore, suggested that Drp1 overexpression could be a potential strategy to attenuate the aging-related accumulation of mitochondrial dysfunction and counter sarcopenia. However, we recently reported that knocking down or overexpressing Drp1 late in life in mouse skeletal muscle (from 18 to 22 months of age) is detrimental to skeletal muscle mass and mitochondrial health. These recent findings, therefore, indicate that targeting Drp1 expression is unlikely to be a viable target to counter sarcopenia and indicate that mitochondrial fission must be maintained within a specific physiological range to preserve muscle and mitochondrial health. Whether enhancing/overexpressing mitochondrial fusion proteins, such as Mfn2 in muscles of aged WT mice, can attenuate the progression of sarcopenia has never been tested.

4. The Role of Autophagy in Skeletal Muscle Health and Aging

The word autophagy derives from two Greek words “auto”, meaning self, and “phagy”, meaning eating. Three different delivery systems have been described in mammals: macro-autophagy, chaperone-mediated autophagy, and micro-autophagy. In this review, the term autophagy is synonymous with macroautophagy, which is characterized by the formation of double-membrane vesicles known as autophagosomes that surround portions of cytoplasm, organelles, and protein aggregates that are subsequently delivered to lysosomes for degradation. The proper maintenance of this catabolic process is essential for survival and has important roles in various physiological functions. Under normal conditions, autophagy mainly prevents the accumulation of damaged organelles and misfolded proteins. In response to stress, such as starvation, autophagy acts mainly as a pro-survival mechanism, by providing metabolic substrates such as amino acids through the bulk degradation of cytoplasmic components; however, excessive self-degradation can lead to cell death and pathological phenotypes.

The formation of autophagosomes and the degradation of its cargo involve several steps including initiation, elongation, maturation, fusion, and degradation (Figure 2). These steps require energy and involve more than 30 autophagy-related (ATG) proteins in yeast, and many of these proteins are highly conserved in mammals.

The initiation and expansion of the phagophore depend on the activation of the ULK1 and PI3-K complexes and the synthesis of phosphatidylinositol 3-phosphate (PI3P). In addition, ATG9-containing vesicles also provide additional lipid/membrane sources for pre-autophagosomal structures. The elongation and the maturation of autophagosomes require the ATG12 conjugation system which includes ATG5, ATG7, ATG10, ATG12, and ATG16L1 proteins. The covalent conjugation ATG12 and ATG5 is accomplished by two enzymes ATG7 and ATG10. The E1-like enzyme ATG7 activates ATG12 via the formation of a thioester bond between the C-terminus of ATG12 and the cysteine507 of ATG7. Subsequently, activated ATG12 is transferred to the E2 enzyme ATG10, and is eventually conjugated to lys149 of ATG5. The conjugate ATG12-ATG5 subsequently forms a protein complex with ATG16L1 localized on the pre-autophagosomal structure. The ATG12-ATG5-ATG16L complex exhibits E3-like activity that facilitates the conjugation of LC3 to phosphatidylethanolamine (PE). LC3 exists in two forms: the free mature form referred to LC3-I present in the cytosol and the rapidly lipidated form (LC3-II), which is formed during autophagosome formation. LC3 is first processed by autophagin (ATG4) into LC3-I with the exposure of a glycine residue at the C-terminus. The second conjugation system ensures that the cytosolic form of LC3 (LC3-I) links to lipid-conjugated phosphatidylethanolamine (PE) to form LC3-PE (LC3-II). The LC3-II protein is rapidly degraded when the autophagosome fuses with the lysosome. It has also been postulated that WD Repeat Domain, Phosphoinositide Interacting 2 (WIPI2) could act upstream of Atg16L1 to recruit the ATG12-ATG5-ATG16L1 and LC3 proteins to the autophagosome membrane. Since LC3 localizes to the autophagosome membrane, monitoring the free mature form referred to LC3B-I present in the cytosol and the rapidly lipidated form (LCB3-II) by immunoblotting is currently the most widely used method to indirectly assess the levels of autophagosome formation. Importantly, an increase in LC3B-II protein content may not necessarily imply that autophagy is elevated, since insufficient autophagosome fusion with lysosomes or defective lysosome function may also cause higher levels of the LC3B-II protein, irrespective of the degree of autophagy. Currently, autophagic flux assays are the gold standard method for assessing autophagy in a given cell or tissue. In mouse skeletal muscle, autophagy flux has been accomplished by using inhibitors of autophagy, including for instance colchicine, leupeptin, and chloroquine. In addition, GFP-LC3 transgenic mice are also often used to evaluate autophagy in vivo. It is worth mentioning that monitoring autophagy in humans in vivo is impossible and that studies in humans are restricted to assessing indirect markers of autophagy activity. Extensive descriptions of autophagy monitoring procedures, as well as their advantages and limitations, have been reviewed in.

It is now generally accepted that autophagy contributes to mammalian cellular homoeostasis by removing damaged organelles under normal physiological conditions and even more under stress conditions, including nutrient and energy deprivation. For example, during acute starvation, autophagic production of amino acids in the liver is important for regulating hepatic gluconeogenesis. There is also evidence that autophagy plays important roles in lifespan and health span in mammals. In mice, muscle-specific deletion of ATG7 (a key regulator of autophagy) was shown to result in severe muscle atrophy, in the deterioration of neuromuscular junctions, and in a reduced lifespan. This severe phenotype is associated with significant mitochondrial abnormalities and multilamellar body structures in mice with muscle-selective inhibition of autophagy. Importantly, plasmid mediated-ATG7 overexpression for 14 days in aged skeletal muscles restored autophagy, ameliorated the integrity of neuromuscular junctions, and increased myofiber size. In humans, mutations in ATG7 cause neurodevelopmental disorders involving neurologic, muscular, and endocrine hypofunction. It was also reported that p62/SQSTM1 (sequestosome-1) accumulates in aged skeletal muscles, suggesting that a decline in autophagy may contribute to sarcopenia. In line with this view, several studies have observed an accumulation of lipofuscin particles in aged muscles from rodents and humans, suggesting that the autophagy-lysosomal function is defective. Furthermore, inactivation of autophagy regulating genes has also been associated with decreased lifespan in Caenorhabditis elegans, whereas promotion of autophagy in Drosophila extended lifespan. It has also been reported that muscle-specific overexpression of FoxO transcription factors in Drosophila decreased muscle aging and extended life span, at least in part by sustaining basal levels of autophagy. It is worth mentioning that FoxO transcription factors are now known to be major regulators of several autophagy-related proteins. Transgenic overexpression of Atg5 (key regulator of autophagy) in mice was shown to improve motor function and extend lifespan by 17%, which is most likely due to enhanced autophagy. There is evidence that aging-related alterations in autophagy likely lead to accumulation of damaged macromolecules and organelles during aging. However, it is important to mention here that the molecular machinery regulating autophagy is still only partly understood. Further studies will, therefore, be required to refine our understanding of the exact role that autophagy plays in the skeletal muscle aging process.

5. The Role of Mitophagy in Skeletal Muscle Health and Aging

Mitophagy is a specific form of autophagy where damaged mitochondria are selectively eliminated by the autophagy-lysosome system. In healthy skeletal muscles, damaged and depolarized mitochondria are selectively removed through mitophagy pathways (Figure 3). The most extensively studied mitophagy pathway involves the E3 ligase Parkin and PINK1 (Phosphatase and tensin homolog (PTEN)-induced kinase 1). Parkin, an E3 ubiquitin ligase encoded by the Park2 gene, is a 465 amino acid protein that translocates to depolarized mitochondria to initiate mitophagy. Loss of mitochondrial membrane potential (ΔΨm) by mitochondrial uncouplers blocks the import of PINK1 into the inner mitochondrial membrane (IMM). PINK1 then accumulates on the outer mitochondrial membrane (OMM), which, in turn, stimulates the recruitment of Parkin E3 ligase activity from the cytosol followed by removal of damaged mitochondria by autophagosomes. Once activated, Parkin has been reported to facilitate ubiquitination of numerous OMM proteins, including Mfn1, Mfn2, VDAC, Tom20 (translocase of the outer membrane 20), and Miro. These ubiquitinated proteins are then degraded by the proteasome and autophagy systems. In healthy mitochondria, cytosolic PINK1 protein is imported into the inner mitochondrial membrane and then degraded by mitochondrial proteases. Importantly, PINK1 was recently shown to be dispensable for mitochondrial recruitment of Parkin and activation of mitophagy in cardiomyocytes. It should be emphasized here that our current knowledge and understanding of Parkin-dependent mitophagy is largely based on cultured cell experiments in which non-physiological uncouplers such as CCCP were used to induce mitochondrial depolarization.

Mitophagy can also occur through Parkin-independent pathways. For instance, the selective elimination of damaged mitochondria can be achieved by p62/SQSTM1, Bnip3 (BCL2 Interacting Protein 3), and Bnip3L (BCL2 Interacting Protein 3 Like, also called NIX). Moreover, it has been shown that mitochondrial ubiquitin chains can interact with optineurin (Optn) and NDP52 (Nuclear dot protein 52 kD) via its LC3 interaction region (LIR) motif to remove dysfunctional mitochondria. A recent study also showed that deleting FUNDC1 (FUN14 Domain Containing 1) specifically in skeletal muscles in mice causes mitophagy impairment. As such, when assessing mitophagy, it important to keep in mind that various proteins, such as Bnip3, Bnip3L, NDP52, and p62/SQSTM1, can also trigger mitophagy in a Parkin-independent manner (Figure 3). It should also be mentioned that the translocation of cardiolipin from the inner to outer mitochondrial membrane is another event that can trigger mitophagy.

Only few studies have examined the impact of aging on mitophagy pathways in skeletal muscle. Although conflicting data exist, several studies reported that mitophagy-related proteins are decreased in the skeletal muscle of old rodents. Similarly, and while not a universal finding, Parkin content was reported lower in the skeletal muscle of old mice and rats. Importantly, the majority of studies that reported an increase in Parkin content in aged muscles also reported that mitophagy was impaired/insufficient in aged skeletal muscles. In humans, we reported that the ratio of Parkin to VDAC (voltage-dependent anion channel) is significantly reduced in atrophied muscles of old men compared to muscles of young men, suggesting that mitophagy might be impaired during muscle aging in humans. In line with this finding, it has been reported that inactive old women have significantly lower expression levels of Bnip3, Beclin 1, Atg7, and Park2 (the gene coding for Parkin), compared to a more active cohort. These studies suggest the presence of a strong relationship between decreased markers of mitophagy and the development of skeletal muscle atrophy during aging. Based on the available literature, it is possible that disrupted mitophagy and unbalanced mitochondrial dynamics might lead to the accumulation of dysfunctional mitochondria during skeletal muscle aging (Figure 4).

Several recent studies using loss- and gain-of-function approaches in vivo have provided mechanistic insights into the physiological function of Parkin in skeletal muscle health and aging. For instance, it has been reported that muscles from Parkin-deficient mice display lower mitophagy flux in response to acute exercise and endurance training. We and others have reported that muscles of Parkin-deficient mice display several aging-like features, including contractile dysfunction and impaired mitochondrial energetics. We also reported that the loss of Parkin in skeletal muscle triggers an increase in the sensitization of the mPTP. Taken together, these studies highlight the importance of Parkin in the maintenance of mitochondrial integrity in skeletal muscle. Using gain-of-function, we recently showed that Parkin overexpression increased muscle mass, fiber size, and mitochondrial enzyme activity in both young and old muscles. Furthermore, in old mice, we found that Parkin overexpression increased muscle strength, increased mitochondrial content, and protected against oxidative stress, fibrosis, and apoptosis. In line with these findings, Parkin overexpression in skeletal muscles of Drosophila led to higher mitochondrial citrate synthase enzymatic activity and attenuated the accumulation of protein aggregates, a marker of cellular aging, whereas Parkin loss-of-function in Drosophila resulted in a decreased lifespan and increased locomotor defects. We also recently showed that Parkin overexpression prevents sepsis-induced skeletal muscle atrophy, likely by improving mitochondrial quality and content. Taken altogether, these studies highlight the importance of proteins regulating mitophagy in skeletal muscle health and aging.

Importantly, mechanistic studies aiming at assessing the role played by proteins involved in Parkin-independent mitophagy in sarcopenia are currently lacking. Assessing whether modulating proteins regulating Parkin-independent mitophagy in aged mice can attenuate sarcopenia might open new therapeutic avenues.

6. Enhancing Mitophagy to Counteract Sarcopenia: Potential Applicability and Future Research

In the last years, several studies have provided the proof of principle that stimulating mitophagy can counteract sarcopenia. As discussed above, we have shown that overexpressing Parkin in aged skeletal muscles attenuates sarcopenia. Recently, Dr Auwerx’s research group reported that urolithin A supplementation, a natural dietary microflora-derived metabolite from ingested ellagitannins and ellagic acid, stimulates Parkin-mediated mitophagy and prolongs lifespan in C. elegans, and improves muscle function and running performance in old rodents while increasing Parkin expression. Administration of urolithin A in the first-in-human clinical trial has also been shown to be safe and to induce a molecular signature of mitochondrial health in the skeletal muscle of healthy sedentary elderly individuals. Another natural compound from tomatoes, tomatidine, has also been shown to attenuate disuse-induced muscle atrophy and improve skeletal muscle mass and function in mice, and to prolong lifespan and health span in C. elegans by stimulating mitophagy. It was also reported that lifespan extension in response to resveratrol supplementation in rats was associated with an activation of the Pink1/Parkin pathway. It is worth mentioning that two small molecules, T0466 and T0467, were recently shown to increase the mitochondrial translocation of Parkin in human dopaminergic neurons and myoblasts. Interestingly, both compounds were shown to reduce unfolded mitochondrial protein levels, presumably through enhanced PINK1-Parkin signaling. Recently, it was shown that injecting old mice with a miR-181a mimic (miR-181a being a microRNA known to enhance mitophagy in myoblasts) resulted in improved skeletal muscle force, increased myofiber size, and a trend for increased succinate dehydrogenase activity. Taken together, the available literature position mitophagy-modulating compounds as potential therapeutic tools to counteract skeletal muscle atrophy and weakness occurring with aging. Large-scale clinical trials are now needed to assess the efficacy of mitophagy-inducing compounds in fighting sarcopenia.

Interestingly, physical activity, which is well known to improve muscle and mitochondrial health in the elderly, was recently shown to stimulate mitophagy and increase Parkin expression in the skeletal muscles of rodents and humans. It was also shown that physically active older adults display higher Parkin in mitochondrial fractions prepared from muscle biopsies. It is, therefore, possible that the beneficial effects of physical activity on muscle and mitochondrial health in older individuals might, at least in part, involve an upregulation of mitophagy. Similarly, accumulating experimental evidence indicates that calorie restriction, one of the most efficient interventions to attenuate sarcopenia in rodents and to improve mitochondrial health, also stimulates mitophagy (see for a detailed review). Perhaps not surprisingly, evidence also indicates that fasting likely triggers mitophagy. Interestingly, time-restricted eating, which involves regular periods of fasting, has emerged in the last few years as an intervention with the potential to improve mitochondrial health and increase health span. It was even recently suggested that time-restricted eating might be an avenue to fight sarcopenia. Whether time-restricted eating can efficiently stimulate mitophagy in aged skeletal muscles and attenuate the aging-related decline in muscle mass and function represent an interesting research avenue that requires further study.

Recently, it was reported that long-term exposure to a ketogenic diet attenuated the aging-related decline of relative muscle mass and increased markers of mitochondrial content in aged skeletal muscles. Interestingly, the ketone body β-hydroxybutyrate, which is elevated during nutritional ketosis, was shown to induce mitophagy in young and aged myocytes and elicits favorable mitochondrial adaptations in myocytes. These findings combined are particularly interesting when considering the availability of recently developed ketone esters known to significantly increase circulating β-hydroxybutyrate in humans. However, whether a ketogenic diet or ketone supplements can efficiently stimulate mitophagy and attenuate sarcopenia in humans remains unknown.

7. Conclusions

In this review, we critically summarized the current state of knowledge on the role played by mitochondrial dynamics and mitophagy in skeletal muscle health and in the development of sarcopenia. Data from mechanistic studies have clearly highlighted that mitochondrial dynamics and mitophagy are critical to skeletal muscle and mitochondrial health. Whether impairment in mitochondrial dynamics contributes to sarcopenia still remains unknown, as the literature is filled with conflicting data. Studies aiming at modulating the expression of proteins regulating dynamics late in life should provide answers as to whether mitochondrial dynamics can be targeted to counter sarcopenia. While conflicting data also exist for mitophagy, a growing body of evidence indicates that mitophagy is impaired in aged skeletal muscles. Importantly, enhancing mitophagy through genetic or nutritional approaches improves skeletal muscle function in aged rodents and improves mitochondrial health in the elderly. Enhancing mitophagy in skeletal muscle, therefore, appears as a promising therapeutic target to prevent or even treat skeletal muscle dysfunction in the elderly.

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    Not in our library10.1089/rej.2017.1989

  19. 19.

    Liu Y., Bi X., Zhang Y., Wang Y., Ding W. Mitochondrial dysfunction/NLRP3 inflammasome axis contributes to angiotensin II-induced skeletal muscle wasting via PPAR-gamma. Lab. Investig. J. Tech. Methods Pathol. 2020;100:712-726.

    Not in our library10.1038/s41374-019-0355-1

  20. 20.

    Wang X.H., Zhang L., Mitch W.E., LeDoux J.M., Hu J., Du J. Caspase-3 cleaves specific 19 S proteasome subunits in skeletal muscle stimulating proteasome activity. J. Biol. Chem. 2010;285:21249-21257.

    Not in our library10.1074/jbc.m109.041707

  21. 21.

    Coggan A.R., Spina R.J., King D.S., Rogers M.A., Brown M., Nemeth P.M., Holloszy J.O. Skeletal muscle adaptations to endurance training in 60- to 70-yr-old men and women. J. Appl. Physiol. 1992;72:1780-1786.

    Not in our library10.1152/jappl.1992.72.5.1780

  22. 22.

    Song W., Kwak H.B., Lawler J.M. Exercise training attenuates age-induced changes in apoptotic signaling in rat skeletal muscle. Antioxid. Redox Signal. 2006;8:517-528.

    Not in our library10.1089/ars.2006.8.517

  23. 23.

    St-Jean-Pelletier F., Pion C.H., Leduc-Gaudet J.P., Sgarioto N., Zovilé I., Barbat-Artigas S., Reynaud O., Alkaterji F., Lemieux F.C., Grenon A. The impact of ageing, physical activity, and pre-frailty on skeletal muscle phenotype, mitochondrial content, and intramyocellular lipids in men. J. Cachexia Sarcopenia Muscle. 2016.

    Not in our library10.1002/jcsm.12139

  24. 24.

    Lozano-Montoya I., Correa-Pérez A., Abraha I., Soiza R.L., Cherubini A., O’Mahony D., Cruz-Jentoft A.J. Nonpharmacological interventions to treat physical frailty and sarcopenia in older patients: A systematic overview—The SENATOR Project ONTOP Series. Clin. Interv. Aging. 2017;12:721-740.

    Not in our library10.2147/cia.s132496

  25. 25.

    Baker D.J., Betik A.C., Krause D.J., Hepple R.T. No decline in skeletal muscle oxidative capacity with aging in long-term calorically restricted rats: Effects are independent of mitochondrial DNA integrity. J. Gerontol. Ser. A Biol. Sci. Med. Sci. 2006;61:675-684.

    Not in our library10.1093/gerona/61.7.675

  26. 26.

    Mayhew M., Renganathan M., Delbono O. Effectiveness of caloric restriction in preventing age-related changes in rat skeletal muscle. Biochem. Biophys. Res. Commun. 1998;251:95-99.

    Not in our library10.1006/bbrc.1998.9438

  27. 27.

    Hepple R.T., Baker D.J., McConkey M., Murynka T., Norris R. Caloric restriction protects mitochondrial function with aging in skeletal and cardiac muscles. Rejuvenation Res. 2006;9:219-222.

    Not in our library10.1089/rej.2006.9.219

  28. 28.

    Gouspillou G., Hepple R.T. Facts and controversies in our understanding of how caloric restriction impacts the mitochondrion. Exp. Gerontol. 2013;48:1075-1084.

    Not in our library10.1016/j.exger.2013.03.004

  29. 29.

    Ogata T., Yamasaki Y. Ultra-High-resolution scanning electron microscopy of mitochondria and sarcoplasmic reticulum arrangement in human red, white, and intermediate muscle fibers. Anat. Rec. 1997;248:214-223.

    Not in our library10.1002/(sici)1097-0185(199706)248:2<214::aid-ar8>3.0.co;2-s

  30. 30.

    Chan D.C. Fusion and fission: Interlinked processes critical for mitochondrial health. Annu. Rev. Genet. 2012;46:265-287.

    Not in our library10.1146/annurev-genet-110410-132529

  31. 31.

    Suen D.F., Norris K.L., Youle R.J. Mitochondrial dynamics and apoptosis. Genes Dev. 2008;22:1577-1590.

    Not in our library10.1101/gad.1658508

  32. 32.
    Mitochondrial dynamics: overview of molecular mechanisms · 2018
    No notice
  33. 33.

    Chen H., Vermulst M., Wang Y.E., Chomyn A., Prolla T.A., McCaffery J.M., Chan D.C. Mitochondrial fusion is required for mtDNA stability in skeletal muscle and tolerance of mtDNA mutations. Cell. 2010;141:280-289.

    Not in our library10.1016/j.cell.2010.02.026

  34. 34.

    Sebastian D., Sorianello E., Segales J., Irazoki A., Ruiz-Bonilla V., Sala D., Planet E., Berenguer-Llergo A., Munoz J.P., Sanchez-Feutrie M., et al. Mfn2 deficiency links age-related sarcopenia and impaired autophagy to activation of an adaptive mitophagy pathway. EMBO J. 2016;35:1677-1693.

    Not in our library10.15252/embj.201593084

  35. 35.

    Bell M.B., Bush Z., McGinnis G.R., Rowe G.C. Adult skeletal muscle deletion of Mitofusin 1 and 2 impedes exercise performance and training capacity. J. Appl. Physiol. 2019;126:341-353.

    Not in our library10.1152/japplphysiol.00719.2018

  36. 36.

    Tezze C., Romanello V., Desbats M.A., Fadini G.P., Albiero M., Favaro G., Ciciliot S., Soriano M.E., Morbidoni V., Cerqua C., et al. Age-Associated Loss of OPA1 in Muscle Impacts Muscle Mass, Metabolic Homeostasis, Systemic Inflammation, and Epithelial Senescence. Cell Metab. 2017;25:1374-1389.e6.

    Not in our library10.1016/j.cmet.2017.04.021

  37. 37.

    Pereira R.O., Tadinada S.M., Zasadny F.M., Oliveira K.J., Pires K.M.P., Olvera A., Jeffers J., Souvenir R., Mcglauflin R., Seei A., et al. OPA1 deficiency promotes secretion of FGF21 from muscle that prevents obesity and insulin resistance. EMBO J. 2017;36:2126-2145.

    Not in our library10.15252/embj.201696179

  38. 38.

    Rodríguez-Nuevo A., Díaz-Ramos A., Noguera E., Díaz-Sáez F., Duran X., Muñoz J.P., Romero M., Plana N., Sebastián D., Tezze C., et al. Mitochondrial DNA and TLR9 drive muscle inflammation upon Opa1 deficiency. EMBO J. 2018;37.

    Not in our library10.15252/embj.201796553

  39. 39.
    Muscle-specific Drp1 overexpression impairs skeletal muscle growth via translational attenuation · 2015
    No notice
  40. 40.

    Giovarelli M., Zecchini S., Martini E., Garrè M., Barozzi S., Ripolone M., Napoli L., Coazzoli M., Vantaggiato C., Roux-Biejat P., et al. Drp1 overexpression induces desmin disassembling and drives kinesin-1 activation promoting mitochondrial trafficking in skeletal muscle. Cell Death Differ. 2020;27:2383-2401.

    Not in our library10.1038/s41418-020-0510-7

  41. 41.

    Romanello V., Guadagnin E., Gomes L., Roder I., Sandri C., Petersen Y., Milan G., Masiero E., Del Piccolo P., Foretz M., et al. Mitochondrial fission and remodelling contributes to muscle atrophy. EMBO J. 2010;29:1774-1785.

    Not in our library10.1038/emboj.2010.60

  42. 42.
    DRP1-mediated mitochondrial shape controls calcium homeostasis and muscle mass · 2019
    No notice
  43. 43.

    Dulac M., Leduc-Gaudet J.P., Reynaud O., Ayoub M.B., Guerin A., Finkelchtein M., Hussain S.N., Gouspillou G. Drp1 knockdown induces severe muscle atrophy and remodelling, mitochondrial dysfunction, autophagy impairment and denervation. J. Physiol. 2020.

    Not in our library10.1113/jp279802

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    Avellaneda J., Rodier C., Daian F., Brouilly N., Rival T., Luis N.M., Schnorrer F. Myofibril and mitochondria morphogenesis are coordinated by a mechanical feedback mechanism in muscle. Nat. Commun. 2021;12:2091.

    Not in our library10.1038/s41467-021-22058-7

  45. 45.

    Lee T.T., Chen P.L., Su M.P., Li J.C., Chang Y.W., Liu R.W., Juan H.F., Yang J.M., Chan S.P., Tsai Y.C., et al. Loss of Fis1 impairs proteostasis during skeletal muscle aging in Drosophila. Aging Cell. 2021;20:e13379.

    Not in our library10.1111/acel.13379

  46. 46.
    Dysregulation of Mitochondrial Quality Control Processes Contribute to Sarcopenia in a Mouse Model of Premature Aging · 2013
    No notice
  47. 47.

    O’Leary M.F., Vainshtein A., Iqbal S., Ostojic O., Hood D.A. Adaptive plasticity of autophagic proteins to denervation in aging skeletal muscle. Am. J. Physiol. Cell Physiol. 2013;304:C422-C430.

    Not in our library10.1152/ajpcell.00240.2012

  48. 48.

    O’Connell K., Ohlendieck K. Proteomic DIGE analysis of the mitochondria-enriched fraction from aged rat skeletal muscle. Proteomics. 2009;9:5509-5524.

    Not in our library10.1002/pmic.200900472

  49. 49.

    Iqbal S., Ostojic O., Singh K., Joseph A.M., Hood D.A. Expression of mitochondrial fission and fusion regulatory proteins in skeletal muscle during chronic use and disuse. Muscle Nerve. 2013;48:963-970.

    Not in our library10.1002/mus.23838

  50. 50.

    Faitg J., Leduc-Gaudet J.P., Reynaud O., Ferland G., Gaudreau P., Gouspillou G. Effects of Aging and Caloric Restriction on Fiber Type Composition, Mitochondrial Morphology and Dynamics in Rat Oxidative and Glycolytic Muscles. Front. Physiol. 2019;10:420.

    Not in our library10.3389/fphys.2019.00420

  51. 51.

    Koltai E., Hart N., Taylor A.W., Goto S., Ngo J.K., Davies K.J., Radak Z. Age-Associated declines in mitochondrial biogenesis and protein quality control factors are minimized by exercise training. Am. J. Physiol. Regul. Integr. Comp. Physiol. 2012;303:R127-R134.

    Not in our library10.1152/ajpregu.00337.2011

  52. 52.
    Mitochondrial morphology is altered in atrophied skeletal muscle of aged mice · 2015
    No notice
  53. 53.

    Capitanio D., Vasso M., De Palma S., Fania C., Torretta E., Cammarata F.P., Magnaghi V., Procacci P., Gelfi C. Specific protein changes contribute to the differential muscle mass loss during ageing. Proteomics. 2016;16:645-656.

    Not in our library10.1002/pmic.201500395

  54. 54.

    Yeo D., Kang C., Gomez-Cabrera M.C., Vina J., Ji L.L. Intensified mitophagy in skeletal muscle with aging is downregulated by PGC-1alpha overexpression in vivo. Free Radic. Biol. Med. 2019;130:361-368.

    Not in our library10.1016/j.freeradbiomed.2018.10.456

  55. 55.

    Zhao L., Zou X., Feng Z., Luo C., Liu J., Li H., Chang L., Wang H., Li Y., Long J., et al. Evidence for association of mitochondrial metabolism alteration with lipid accumulation in aging rats. Exp. Gerontol. 2014;56:3-12.

    Not in our library10.1016/j.exger.2014.02.001

  56. 56.

    Konopka A.R., Suer M.K., Wolff C.A., Harber M.P. Markers of Human Skeletal Muscle Mitochondrial Biogenesis and Quality Control: Effects of Age and Aerobic Exercise Training. J. Gerontol. Ser. A. 2013;69:371-378.

    Not in our library10.1093/gerona/glt107

  57. 57.

    Balan E., Schwalm C., Naslain D., Nielens H., Francaux M., Deldicque L. Regular Endurance Exercise Promotes Fission, Mitophagy, and Oxidative Phosphorylation in Human Skeletal Muscle Independently of Age. Front. Physiol. 2019;10:1088.

    Not in our library10.3389/fphys.2019.01088

  58. 58.

    Distefano G., Standley R.A., Dubé J.J., Carnero E.A., Ritov V.B., Stefanovic-Racic M., Toledo F.G., Piva S.R., Goodpaster B.H., Coen P.M. Chronological Age Does not Influence Ex-vivo Mitochondrial Respiration and Quality Control in Skeletal Muscle. J. Gerontol. Ser. A Biol. Sci. Med. Sci. 2017;72:535-542.

    Not in our library10.1093/gerona/glw102

  59. 59.

    Joseph A.M., Adhihetty P.J., Buford T.W., Wohlgemuth S.E., Lees H.A., Nguyen L.M., Aranda J.M., Sandesara B.D., Pahor M., Manini T.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.

    Not in our library10.1111/j.1474-9726.2012.00844.x

  60. 60.

    Crane J.D., Devries M.C., Safdar A., Hamadeh M.J., Tarnopolsky M.A. The effect of aging on human skeletal muscle mitochondrial and intramyocellular lipid ultrastructure. J. Gerontol. Ser. A Biol. Sci. Med. Sci. 2010;65:119-128.

    Not in our library10.1093/gerona/glp179

  61. 61.

    Wyckelsma V.L., Levinger I., McKenna M.J., Formosa L.E., Ryan M.T., Petersen A.C., Anderson M.J., Murphy R.M. Preservation of skeletal muscle mitochondrial content in older adults: Relationship between mitochondria, fibre type and high-intensity exercise training. J. Physiol. 2017;595:3345-3359.

    Not in our library10.1113/jp273950

  62. 62.

    Pham A.H., McCaffery J.M., Chan D.C. Mouse lines with photo-activatable mitochondria to study mitochondrial dynamics. Genesis. 2012;50:833-843.

    Not in our library10.1002/dvg.22050

  63. 63.

    Perry C.G., Lally J., Holloway G.P., Heigenhauser G.J., Bonen A., Spriet L.L. Repeated transient mRNA bursts precede increases in transcriptional and mitochondrial proteins during training in human skeletal muscle. J. Physiol. 2010;588:4795-4810.

    Not in our library10.1113/jphysiol.2010.199448

  64. 64.

    Granata C., Oliveira R.S.F., Little J.P., Renner K., Bishop D.J. Training intensity modulates changes in PGC-1α and p53 protein content and mitochondrial respiration, but not markers of mitochondrial content in human skeletal muscle. FASEB J. 2016;30:959-970.

    Not in our library10.1096/fj.15-276907

  65. 65.

    Meinild Lundby A.K., Jacobs R.A., Gehrig S., de Leur J., Hauser M., Bonne T.C., Flück D., Dandanell S., Kirk N., Kaech A., et al. Exercise training increases skeletal muscle mitochondrial volume density by enlargement of existing mitochondria and not de novo biogenesis. Acta Physiol. 2018;222.

    Not in our library10.1111/apha.12905

  66. 66.

    Mishra P., Varuzhanyan G., Pham A.H., Chan D.C. Mitochondrial Dynamics Is a Distinguishing Feature of Skeletal Muscle Fiber Types and Regulates Organellar Compartmentalization. Cell Metab. 2015;22:1033-1044.

    Not in our library10.1016/j.cmet.2015.09.027

  67. 67.

    Dahl R., Larsen S., Dohlmann T.L., Qvortrup K., Helge J.W., Dela F., Prats C. Three-Dimensional reconstruction of the human skeletal muscle mitochondrial network as a tool to assess mitochondrial content and structural organization. Acta Physiol. 2015;213:145-155.

    Not in our library10.1111/apha.12289

  68. 68.

    Lally J.S.V., Herbst E.A.F., Matravadia S., Maher A.C., Perry C.G.R., Ventura-Clapier R., Holloway G.P. Over-Expressing Mitofusin-2 in Healthy Mature Mammalian Skeletal Muscle Does Not Alter Mitochondrial Bioenergetics. PLoS ONE. 2013;8:e55660.

    Not in our library10.1371/journal.pone.0055660

  69. 69.
    Promoting Drp1-mediated mitochondrial fission in midlife prolongs healthy lifespan of Drosophila melanogaster · 2017
    No notice
  70. 70.

    Twig G., Elorza A., Molina A.J., Mohamed H., Wikstrom J.D., Walzer G., Stiles L., Haigh S.E., Katz S., Las G., et al. Fission and selective fusion govern mitochondrial segregation and elimination by autophagy. EMBO J. 2008;27:433-446.

    Not in our library10.1038/sj.emboj.7601963

  71. 71.

    Dulac M., Leduc-Gaudet J.P., Cefis M., Ayoub M.B., Reynaud O., Shams A., Moamer A., Nery Ferreira M.F., Hussain S.N., Gouspillou G. Regulation of muscle and mitochondrial health by the mitochondrial fission protein Drp1 in aged mice. J. Physiol. 2021.

    Not in our library10.1113/jp281752

  72. 72.

    Levine B., Mizushima N., Virgin H.W. Autophagy in immunity and inflammation. Nature. 2011;469:323-335.

    Not in our library10.1038/nature09782

  73. 73.

    Yorimitsu T., Klionsky D.J. Autophagy: Molecular machinery for self-eating. Cell Death Differ. 2005;12(Suppl. 2):1542-1552.

    Not in our library10.1038/sj.cdd.4401765

  74. 74.

    Rabinowitz J.D., White E. Autophagy and metabolism. Science. 2010;330:1344-1348.

    Not in our library10.1126/science.1193497

  75. 75.

    Levine B., Kroemer G. Autophagy in the pathogenesis of disease. Cell. 2008;132:27-42.

    Not in our library10.1016/j.cell.2007.12.018

  76. 76.

    Yang Z., Klionsky D.J. Eaten alive: A history of macroautophagy. Nat. Cell Biol. 2010;12:814-822.

    Not in our library10.1038/ncb0910-814

  77. 77.

    Tsukada M., Ohsumi Y. Isolation and characterization of autophagy-defective mutants of Saccharomyces cerevisiae. FEBS Lett. 1993;333:169-174.

    Not in our library10.1016/0014-5793(93)80398-e

  78. 78.

    Young A.R., Chan E.Y., Hu X.W., Kochl R., Crawshaw S.G., High S., Hailey D.W., Lippincott-Schwartz J., Tooze S.A. Starvation and ULK1-dependent cycling of mammalian Atg9 between the TGN and endosomes. J. Cell Sci. 2006;119:3888-3900.

    Not in our library10.1242/jcs.03172

  79. 79.

    Yamada T., Carson A.R., Caniggia I., Umebayashi K., Yoshimori T., Nakabayashi K., Scherer S.W. Endothelial nitric-oxide synthase antisense (NOS3AS) gene encodes an autophagy-related protein (APG9-like2) highly expressed in trophoblast. J. Biol. Chem. 2005;280:18283-18290.

    Not in our library10.1074/jbc.m413957200

  80. 80.

    Orsi A., Razi M., Dooley H.C., Robinson D., Weston A.E., Collinson L.M., Tooze S.A. Dynamic and transient interactions of Atg9 with autophagosomes, but not membrane integration, are required for autophagy. Mol. Biol. Cell. 2012;23:1860-1873.

    Not in our library10.1091/mbc.e11-09-0746

  81. 81.

    Mizushima N., Noda T., Yoshimori T., Tanaka Y., Ishii T., George M.D., Klionsky D.J., Ohsumi M., Ohsumi Y. A protein conjugation system essential for autophagy. Nature. 1998;395:395-398.

    Not in our library10.1038/26506

  82. 82.

    Mizushima N., Sugita H., Yoshimori T., Ohsumi Y. A new protein conjugation system in human. The counterpart of the yeast Apg12p conjugation system essential for autophagy. J. Biol. Chem. 1998;273:33889-33892.

    Not in our library10.1074/jbc.273.51.33889

  83. 83.

    Mizushima N., Noda T., Ohsumi Y. Apg16p is required for the function of the Apg12p-Apg5p conjugate in the yeast autophagy pathway. EMBO J. 1999;18:3888-3896.

    Not in our library10.1093/emboj/18.14.3888

  84. 84.

    Mizushima N., Kuma A., Kobayashi Y., Yamamoto A., Matsubae M., Takao T., Natsume T., Ohsumi Y., Yoshimori T. Mouse Apg16L, a novel WD-repeat protein, targets to the autophagic isolation membrane with the Apg12-Apg5 conjugate. J. Cell Sci. 2003;116:1679-1688.

    Not in our library10.1242/jcs.00381

  85. 85.

    Tanida I., Mizushima N., Kiyooka M., Ohsumi M., Ueno T., Ohsumi Y., Kominami E. Apg7p/Cvt2p: A novel protein-activating enzyme essential for autophagy. Mol. Biol. Cell. 1999;10:1367-1379.

    Not in our library10.1091/mbc.10.5.1367

  86. 86.

    Shintani T., Mizushima N., Ogawa Y., Matsuura A., Noda T., Ohsumi Y. Apg10p, a novel protein-conjugating enzyme essential for autophagy in yeast. EMBO J. 1999;18:5234-5241.

    Not in our library10.1093/emboj/18.19.5234

  87. 87.

    Kuma A., Mizushima N., Ishihara N., Ohsumi Y. Formation of the approximately 350-kDa Apg12-Apg5.Apg16 multimeric complex, mediated by Apg16 oligomerization, is essential for autophagy in yeast. J. Biol. Chem. 2002;277:18619-18625.

    Not in our library10.1074/jbc.m111889200

  88. 88.

    Kabeya Y., Mizushima N., Ueno T., Yamamoto A., Kirisako T., Noda T., Kominami E., Ohsumi Y., Yoshimori T. LC3, a mammalian homologue of yeast Apg8p, is localized in autophagosome membranes after processing. EMBO J. 2000;19:5720-5728.

    Not in our library10.1093/emboj/19.21.5720

  89. 89.

    Kabeya Y., Mizushima N., Yamamoto A., Oshitani-Okamoto S., Ohsumi Y., Yoshimori T. LC3, GABARAP and GATE16 localize to autophagosomal membrane depending on form-II formation. J. Cell Sci. 2004;117:2805-2812.

    Not in our library10.1242/jcs.01131

  90. 90.
    WIPI2 Links LC3 Conjugation with PI3P, Autophagosome Formation, and Pathogen Clearance by Recruiting Atg12–5-16L1 · 2014
    No notice
  91. 91.

    Klionsky D.J., Abdel-Aziz A.K., Abdelfatah S., Abdellatif M., Abdoli A., Abel S., Abeliovich H., Abildgaard M.H., Abudu Y.P., Acevedo-Arozena A., et al. Guidelines for the use and interpretation of assays for monitoring autophagy (4th edition). Autophagy. 2021;1-382.

    Not in our library10.1080/15548627.2020.1797280

  92. 92.

    Mofarrahi M., Guo Y., Haspel J.A., Choi A.M., Davis E.C., Gouspillou G., Hepple R.T., Godin R., Burelle Y., Hussain S.N. Autophagic flux and oxidative capacity of skeletal muscles during acute starvation. Autophagy. 2013;9:1604-1620.

    Not in our library10.4161/auto.25955

  93. 93.

    Ju J.-S., Varadhachary A.S., Miller S.E., Weihl C.C. Quantitation of “autophagic flux” in mature skeletal muscle. Autophagy. 2010;6:929-935.

    Not in our library10.4161/auto.6.7.12785

  94. 94.

    Romanello V., Scalabrin M., Albiero M., Blaauw B., Scorrano L., Sandri M. Inhibition of the Fission Machinery Mitigates OPA1 Impairment in Adult Skeletal Muscles. Cells. 2019;8:597.

    Not in our library10.3390/cells8060597

  95. 95.

    Huang D.D., Yan X.L., Fan S.D., Chen X.Y., Yan J.Y., Dong Q.T., Chen W.Z., Liu N.X., Chen X.L., Yu Z. Nrf2 deficiency promotes the increasing trend of autophagy during aging in skeletal muscle: A potential mechanism for the development of sarcopenia. Aging. 2020;12:5977-5991.

    Not in our library10.18632/aging.102990

  96. 96.

    Oost L.J., Kustermann M., Armani A., Blaauw B., Romanello V. Fibroblast growth factor 21 controls mitophagy and muscle mass. J. Cachexia Sarcopenia Muscle. 2019;10:630-642.

    Not in our library10.1002/jcsm.12409

  97. 97.

    De Palma C., Morisi F., Cheli S., Pambianco S., Cappello V., Vezzoli M., Rovere-Querini P., Moggio M., Ripolone M., Francolini M., et al. Autophagy as a new therapeutic target in Duchenne muscular dystrophy. Cell Death Dis. 2012;3:e418.

    Not in our library10.1038/cddis.2012.159

  98. 98.

    Mizushima N. Methods for monitoring autophagy using GFP-LC3 transgenic mice. Methods Enzymol. 2009;452:13-23.

    Not in our library10.1016/s0076-6879(08)03602-1

  99. 99.

    Moulis M., Vindis C. Methods for Measuring Autophagy in Mice. Cells. 2017;6:14.

    Not in our library10.3390/cells6020014

  100. 100.

    Martinet W., Roth L., De Meyer G.R.Y. Standard Immunohistochemical Assays to Assess Autophagy in Mammalian Tissue. Cells. 2017;6:17.

    Not in our library10.3390/cells6030017

  101. 101.

    Schworer C.M., Mortimore G.E. Glucagon-induced autophagy and proteolysis in rat liver: Mediation by selective deprivation of intracellular amino acids. Proc. Natl. Acad. Sci. USA. 1979;76:3169-3173.

    Not in our library10.1073/pnas.76.7.3169

  102. 102.

    Carnio S., LoVerso F., Baraibar M.A., Longa E., Khan M.M., Maffei M., Reischl M., Canepari M., Loefler S., Kern H., et al. Autophagy impairment in muscle induces neuromuscular junction degeneration and precocious aging. Cell Rep. 2014;8:1509-1521.

    Not in our library10.1016/j.celrep.2014.07.061

  103. 103.

    Fernandez A.F., Sebti S., Wei Y., Zou Z., Shi M., McMillan K.L., He C., Ting T., Liu Y., Chiang W.C., et al. Disruption of the beclin 1-BCL2 autophagy regulatory complex promotes longevity in mice. Nature. 2018;558:136-140.

    Not in our library10.1038/s41586-018-0162-7

  104. 104.

    Madeo F., Zimmermann A., Maiuri M.C., Kroemer G. Essential role for autophagy in life span extension. J. Clin. Investig. 2015;125:85-93.

    Not in our library10.1172/jci73946

  105. 105.

    Melendez A., Talloczy Z., Seaman M., Eskelinen E.L., Hall D.H., Levine B. Autophagy genes are essential for dauer development and life-span extension in C. elegans. Science. 2003;301:1387-1391.

    Not in our library10.1126/science.1087782

  106. 106.

    Masiero E., Agatea L., Mammucari C., Blaauw B., Loro E., Komatsu M., Metzger D., Reggiani C., Schiaffino S., Sandri M. Autophagy is required to maintain muscle mass. Cell Metab. 2009;10:507-515.

    Not in our library10.1016/j.cmet.2009.10.008

  107. 107.
    Atg5flox-Derived Autophagy-Deficient Model of Pompe Disease: Does It Tell the Whole Story? · 2017
    Not checked
  108. 108.
    Mitochondrial dysfunction and oxidative stress mediate the physiological impairment induced by the disruption of autophagy · 2009
    No notice
  109. 109.

    Collier J.J., Guissart C., Olahova M., Sasorith S., Piron-Prunier F., Suomi F., Zhang D., Martinez-Lopez N., Leboucq N., Bahr A., et al. Developmental Consequences of Defective ATG7-Mediated Autophagy in Humans. N. Engl. J. Med. 2021;384:2406-2417.

    Not in our library10.1056/nejmoa1915722

  110. 110.

    Sakuma K., Kinoshita M., Ito Y., Aizawa M., Aoi W., Yamaguchi A. p62/SQSTM1 but not LC3 is accumulated in sarcopenic muscle of mice. J. Cachexia Sarcopenia Muscle. 2016;7:204-212.

    Not in our library10.1002/jcsm.12045

  111. 111.

    Hütter E., Skovbro M., Lener B., Prats C., Rabøl R., Dela F., Jansen-Dürr P. Oxidative stress and mitochondrial impairment can be separated from lipofuscin accumulation in aged human skeletal muscle. Aging Cell. 2007;6:245-256.

    Not in our library10.1111/j.1474-9726.2007.00282.x

  112. 112.

    Renault V., Rolland E., Thornell L.-E., Mouly V., Butler-Browne G. Distribution of satellite cells in the human vastus lateralis muscle during aging. Exp. Gerontol. 2002;37:1513-1514.

    Not in our library10.1016/s0531-5565(02)00095-5

  113. 113.

    Toth M.L., Sigmond T., Borsos E., Barna J., Erdelyi P., Takacs-Vellai K., Orosz L., Kovacs A.L., Csikos G., Sass M., et al. Longevity pathways converge on autophagy genes to regulate life span in Caenorhabditis elegans. Autophagy. 2008;4:330-338.

    Not in our library10.4161/auto.5618

  114. 114.

    Aparicio R., Rana A., Walker D.W. Upregulation of the Autophagy Adaptor p62/SQSTM1 Prolongs Health and Lifespan in Middle-Aged Drosophila. Cell Rep. 2019;28:1029-1040.e5.

    Not in our library10.1016/j.celrep.2019.06.070

  115. 115.

    Demontis F., Perrimon N. FOXO/4E-BP signaling in Drosophila muscles regulates organism-wide proteostasis during aging. Cell. 2010;143:813-825.

    Not in our library10.1016/j.cell.2010.10.007

  116. 116.

    Mammucari C., Milan G., Romanello V., Masiero E., Rudolf R., Del Piccolo P., Burden S.J., Di Lisi R., Sandri C., Zhao J., et al. FoxO3 controls autophagy in skeletal muscle in vivo. Cell Metab. 2007;6:458-471.

    Not in our library10.1016/j.cmet.2007.11.001

  117. 117.

    Zhao J., Brault J.J., Schild A., Cao P., Sandri M., Schiaffino S., Lecker S.H., Goldberg A.L. FoxO3 coordinately activates protein degradation by the autophagic/lysosomal and proteasomal pathways in atrophying muscle cells. Cell Metab. 2007;6:472-483.

    Not in our library10.1016/j.cmet.2007.11.004

  118. 118.
    Regulation of autophagy and the ubiquitin–proteasome system by the FoxO transcriptional network during muscle atrophy · 2015
    No notice
  119. 119.

    Pyo J.O., Yoo S.M., Ahn H.H., Nah J., Hong S.H., Kam T.I., Jung S., Jung Y.K. Overexpression of Atg5 in mice activates autophagy and extends lifespan. Nat. Commun. 2013;4:2300.

    Not in our library10.1038/ncomms3300

  120. 120.

    Garcia-Prat L., Martinez-Vicente M., Perdiguero E., Ortet L., Rodriguez-Ubreva J., Rebollo E., Ruiz-Bonilla V., Gutarra S., Ballestar E., Serrano A.L., et al. Autophagy maintains stemness by preventing senescence. Nature. 2016;529:37-42.

    Not in our library10.1038/nature16187

  121. 121.

    Narendra D., Tanaka A., Suen D.-F., Youle R.J. Parkin is recruited selectively to impaired mitochondria and promotes their autophagy. J. Cell Biol. 2008;183:795-803.

    Not in our library10.1083/jcb.200809125

  122. 122.
    PINK1 Is Selectively Stabilized on Impaired Mitochondria to Activate Parkin · 2010
    No notice
  123. 123.

    Kane L.A., Lazarou M., Fogel A.I., Li Y., Yamano K., Sarraf S.A., Banerjee S., Youle R.J. PINK1 phosphorylates ubiquitin to activate Parkin E3 ubiquitin ligase activity. J. Cell Biol. 2014;205:143-153.

    Not in our library10.1083/jcb.201402104

  124. 124.

    Vives-Bauza C., Zhou C., Huang Y., Cui M., de Vries R.L., Kim J., May J., Tocilescu M.A., Liu W., Ko H.S., et al. PINK1-dependent recruitment of Parkin to mitochondria in mitophagy. Proc. Natl. Acad. Sci. USA. 2010;107:378-383.

    Not in our library10.1073/pnas.0911187107

  125. 125.

    Matsuda N., Sato S., Shiba K., Okatsu K., Saisho K., Gautier C.A., Sou Y.-S., Saiki S., Kawajiri S., Sato F. PINK1 stabilized by mitochondrial depolarization recruits Parkin to damaged mitochondria and activates latent Parkin for mitophagy. J. Cell Biol. 2010;189:211-221.

    Not in our library10.1083/jcb.200910140

  126. 126.

    Glauser L., Sonnay S., Stafa K., Moore D.J. Parkin promotes the ubiquitination and degradation of the mitochondrial fusion factor mitofusin 1. J. Neurochem. 2011;118:636-645.

    Not in our library10.1111/j.1471-4159.2011.07318.x

  127. 127.
    The Mitochondrial Fusion-Promoting Factor Mitofusin Is a Substrate of the PINK1/Parkin Pathway · 2010
    No notice
  128. 128.

    Ziviani E., Tao R.N., Whitworth A.J. Drosophila parkin requires PINK1 for mitochondrial translocation and ubiquitinates mitofusin. Proc. Natl. Acad. Sci. USA. 2010;107:5018-5023.

    Not in our library10.1073/pnas.0913485107

  129. 129.

    Chen Y., Dorn G.W. PINK1-phosphorylated mitofusin 2 is a Parkin receptor for culling damaged mitochondria. Science. 2013;340:471-475.

    Not in our library10.1126/science.1231031

  130. 130.

    Geisler S., Holmström K.M., Skujat D., Fiesel F.C., Rothfuss O.C., Kahle P.J., Springer W. PINK1/Parkin-mediated mitophagy is dependent on VDAC1 and p62/SQSTM1. Nat. Cell Biol. 2010;12:119-131.

    Not in our library10.1038/ncb2012

  131. 131.
    Parkinson's Disease–Associated Kinase PINK1 Regulates Miro Protein Level and Axonal Transport of Mitochondria · 2012
    No notice
  132. 132.

    Deas E., Plun-Favreau H., Gandhi S., Desmond H., Kjaer S., Loh S.H., Renton A.E., Harvey R.J., Whitworth A.J., Martins L.M. PINK1 cleavage at position A103 by the mitochondrial protease PARL. Hum. Mol. Genet. 2011;20:867-879.

    Not in our library10.1093/hmg/ddq526

  133. 133.

    Greene A.W., Grenier K., Aguileta M.A., Muise S., Farazifard R., Haque M.E., McBride H.M., Park D.S., Fon E.A. Mitochondrial processing peptidase regulates PINK1 processing, import and Parkin recruitment. EMBO Rep. 2012;13:378-385.

    Not in our library10.1038/embor.2012.14

  134. 134.

    Jin S.M., Lazarou M., Wang C., Kane L.A., Narendra D.P., Youle R.J. Mitochondrial membrane potential regulates PINK1 import and proteolytic destabilization by PARL. J. Cell Biol. 2010;191:933-942.

    Not in our library10.1083/jcb.201008084

  135. 135.

    Kubli D.A., Cortez M.Q., Moyzis A.G., Najor R.H., Lee Y., Gustafsson A.B. PINK1 Is Dispensable for Mitochondrial Recruitment of Parkin and Activation of Mitophagy in Cardiac Myocytes. PLoS ONE. 2015;10:e0130707.

    Not in our library10.1371/journal.pone.0130707

  136. 136.

    Zhang J., Ney P.A. Role of BNIP3 and NIX in cell death, autophagy, and mitophagy. Cell Death Differ. 2009;16:939-946.

    Not in our library10.1038/cdd.2009.16

  137. 137.

    Bellot G., Garcia-Medina R., Gounon P., Chiche J., Roux D., Pouysségur J., Mazure N.M. Hypoxia-Induced Autophagy Is Mediated through Hypoxia-Inducible Factor Induction of BNIP3 and BNIP3L via Their BH3 Domains. Mol. Cell. Biol. 2009;29:2570-2581.

    Not in our library10.1128/mcb.00166-09

  138. 138.

    Hanna R.A., Quinsay M.N., Orogo A.M., Giang K., Rikka S., Gustafsson A.B. Microtubule-associated protein 1 light chain 3 (LC3) interacts with Bnip3 protein to selectively remove endoplasmic reticulum and mitochondria via autophagy. J. Biol. Chem. 2012;287:19094-19104.

    Not in our library10.1074/jbc.m111.322933

  139. 139.

    Rikka S., Quinsay M.N., Thomas R.L., Kubli D.A., Zhang X., Murphy A.N., Gustafsson A.B. Bnip3 impairs mitochondrial bioenergetics and stimulates mitochondrial turnover. Cell Death Differ. 2011;18:721-731.

    Not in our library10.1038/cdd.2010.146

  140. 140.

    Novak I., Kirkin V., McEwan D.G., Zhang J., Wild P., Rozenknop A., Rogov V., Lohr F., Popovic D., Occhipinti A., et al. Nix is a selective autophagy receptor for mitochondrial clearance. EMBO Rep. 2010;11:45-51.

    Not in our library10.1038/embor.2009.256

  141. 141.

    Wong Y.C., Holzbaur E.L. Optineurin is an autophagy receptor for damaged mitochondria in parkin-mediated mitophagy that is disrupted by an ALS-linked mutation. Proc. Natl. Acad. Sci. USA. 2014;111:E4439-E4448.

    Not in our library10.1073/pnas.1405752111

  142. 142.

    Heo J.M., Ordureau A., Paulo J.A., Rinehart J., Harper J.W. The PINK1-PARKIN Mitochondrial Ubiquitylation Pathway Drives a Program of OPTN/NDP52 Recruitment and TBK1 Activation to Promote Mitophagy. Mol. Cell. 2015;60:7-20.

    Not in our library10.1016/j.molcel.2015.08.016

  143. 143.

    Fu T., Xu Z., Liu L., Guo Q., Wu H., Liang X., Zhou D., Xiao L., Liu L., Liu Y., et al. Mitophagy Directs Muscle-Adipose Crosstalk to Alleviate Dietary Obesity. Cell Rep. 2018;23:1357-1372.

    Not in our library10.1016/j.celrep.2018.03.127

  144. 144.

    Chu C.T., Ji J., Dagda R.K., Jiang J.F., Tyurina Y.Y., Kapralov A.A., Tyurin V.A., Yanamala N., Shrivastava I.H., Mohammadyani D., et al. Cardiolipin externalization to the outer mitochondrial membrane acts as an elimination signal for mitophagy in neuronal cells. Nat. Cell Biol. 2013;15:1197-1205.

    Not in our library10.1038/ncb2837

  145. 145.

    Goljanek-Whysall K., Soriano-Arroquia A., McCormick R., Chinda C., McDonagh B. miR-181a regulates p62/SQSTM1, parkin, and protein DJ-1 promoting mitochondrial dynamics in skeletal muscle aging. Aging Cell. 2020;19:e13140.

    Not in our library10.1111/acel.13140

  146. 146.

    Carter H.N., Kim Y., Erlich A.T., Zarrin-Khat D., Hood D.A. Autophagy and mitophagy flux in young and aged skeletal muscle following chronic contractile activity. J. Physiol. 2018;596:3567-3584.

    Not in our library10.1113/jp275998

  147. 147.

    Chen C.C.W., Erlich A.T., Crilly M.J., Hood D.A. Parkin is required for exercise-induced mitophagy in muscle: Impact of aging. Am. J. Physiol. Endocrinol. Metab. 2018;315:E404-E415.

    Not in our library10.1152/ajpendo.00391.2017

  148. 148.

    McMullen C.A., Ferry A.L., Gamboa J.L., Andrade F.H., Dupont-Versteegden E.E. Age-Related changes of cell death pathways in rat extraocular muscle. Exp. Gerontol. 2009;44:420-425.

    Not in our library10.1016/j.exger.2009.03.006

  149. 149.

    Gaugler M., Brown A., Merrell E., DiSanto-Rose M., Rathmacher J.A., Reynolds T.H.T. PKB signaling and atrogene expression in skeletal muscle of aged mice. J. Appl. Physiol. 2011;111:192-199.

    Not in our library10.1152/japplphysiol.00175.2011

  150. 150.

    Drummond M.J., Addison O., Brunker L., Hopkins P.N., McClain D.A., LaStayo P.C., Marcus R.L. Downregulation of E3 ubiquitin ligases and mitophagy-related genes in skeletal muscle of physically inactive, frail older women: A cross-sectional comparison. J. Gerontol. Ser. A Biol. Sci. Med. Sci. 2014;69:1040-1048.

    Not in our library10.1093/gerona/glu004

  151. 151.

    Russ D.W., Wills A.M., Boyd I.M., Krause J. Weakness, SR function and stress in gastrocnemius muscles of aged male rats. Exp. Gerontol. 2014;50:40-44.

    Not in our library10.1016/j.exger.2013.11.018

  152. 152.

    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.

    Not in our library10.1113/jp277157

  153. 153.

    Chen C.C.W., Erlich A.T., Hood D.A. Role of Parkin and endurance training on mitochondrial turnover in skeletal muscle. Skelet Muscle. 2018;8:10.

    Not in our library10.1186/s13395-018-0157-y

  154. 154.

    Gouspillou G., Godin R., Piquereau J., Picard M., Mofarrahi M., Mathew J., Purves-Smith F.M., Sgarioto N., Hepple R.T., Burelle Y., et al. Protective role of Parkin in skeletal muscle contractile and mitochondrial function. J. Physiol. 2018;596:2565-2579.

    Not in our library10.1113/jp275604

  155. 155.

    Peker N., Donipadi V., Sharma M., McFarlane C., Kambadur R. Loss of Parkin impairs mitochondrial function and leads to muscle atrophy. Am. J. Physiol. Cell Physiol. 2018;315:C164-C185.

    Not in our library10.1152/ajpcell.00064.2017

  156. 156.

    Rana A., Rera M., Walker D.W. Parkin overexpression during aging reduces proteotoxicity, alters mitochondrial dynamics, and extends lifespan. Proc. Natl. Acad. Sci. USA. 2013;110:8638-8643.

    Not in our library10.1073/pnas.1216197110

  157. 157.

    Greene J.C., Whitworth A.J., Kuo I., Andrews L.A., Feany M.B., Pallanck L.J. Mitochondrial pathology and apoptotic muscle degeneration in Drosophila parkin mutants. Proc. Natl. Acad. Sci. USA. 2003;100:4078-4083.

    Not in our library10.1073/pnas.0737556100

  158. 158.

    Pesah Y., Pham T., Burgess H., Middlebrooks B., Verstreken P., Zhou Y., Harding M., Bellen H., Mardon G. Drosophila parkin mutants have decreased mass and cell size and increased sensitivity to oxygen radical stress. Development. 2004;131:2183-2194.

    Not in our library10.1242/dev.01095

  159. 159.

    Leduc-Gaudet J.P., Mayaki D., Reynaud O., Broering F.E., Chaffer T.J., Hussain S.N.A., Gouspillou G. Parkin Overexpression Attenuates Sepsis-Induced Muscle Wasting. Cells. 2020;9:1454.

    Not in our library10.3390/cells9061454

  160. 160.

    Ryu D., Mouchiroud L., Andreux P.A., Katsyuba E., Moullan N., Nicolet-Dit-Felix A.A., Williams E.G., Jha P., Lo Sasso G., Huzard D., et al. Urolithin A induces mitophagy and prolongs lifespan in C. elegans and increases muscle function in rodents. Nat. Med. 2016;22:879-888.

    Not in our library10.1038/nm.4132

  161. 161.

    Andreux P.A., Blanco-Bose W., Ryu D., Burdet F., Ibberson M., Aebischer P., Auwerx J., Singh A., Rinsch C. The mitophagy activator urolithin A is safe and induces a molecular signature of improved mitochondrial and cellular health in humans. Nat. Metab. 2019;1:595-603.

    Not in our library10.1038/s42255-019-0073-4

  162. 162.
    Systems-based Discovery of Tomatidine as a Natural Small Molecule Inhibitor of Skeletal Muscle Atrophy · 2014
    No notice
  163. 163.
    Tomatidine enhances lifespan and healthspan in C. elegans through mitophagy induction via the SKN-1/Nrf2 pathway · 2017
    No notice
  164. 164.
    Antiaging Properties of a Grape-Derived Antioxidant Are Regulated by Mitochondrial Balance of Fusion and Fission Leading to Mitophagy Triggered by a Signaling Network of Sirt1-Sirt3-Foxo3-PINK1-PARKIN · 2014
    Retracted
  165. 165.

    Shiba-Fukushima K., Inoshita T., Sano O., Iwata H., Ishikawa K.-I., Okano H., Akamatsu W., Imai Y., Hattori N. A Cell-Based High-Throughput Screening Identified Two Compounds that Enhance PINK1-Parkin Signaling. iScience. 2020;23:101048.

    Not in our library10.1016/j.isci.2020.101048

  166. 166.

    Cartee G.D., Hepple R.T., Bamman M.M., Zierath J.R. Exercise Promotes Healthy Aging of Skeletal Muscle. Cell Metab. 2016;23:1034-1047.

    Not in our library10.1016/j.cmet.2016.05.007

  167. 167.
    Ampk phosphorylation of Ulk1 is required for targeting of mitochondria to lysosomes in exercise-induced mitophagy · 2017
    No notice
  168. 168.

    Brandt N., Gunnarsson T.P., Bangsbo J., Pilegaard H. Exercise and exercise training-induced increase in autophagy markers in human skeletal muscle. Physiol. Rep. 2018;6:e13651.

    Not in our library10.14814/phy2.13651

  169. 169.

    Mehrabani S., Bagherniya M., Askari G., Read M.I., Sahebkar A. The effect of fasting or calorie restriction on mitophagy induction: A literature review. J. Cachexia Sarcopenia Muscle. 2020;11:1447-1458.

    Not in our library10.1002/jcsm.12611

  170. 170.

    Longo V.D., Panda S. Fasting, Circadian Rhythms, and Time-Restricted Feeding in Healthy Lifespan. Cell Metab. 2016;23:1048-1059.

    Not in our library10.1016/j.cmet.2016.06.001

  171. 171.

    Mattson M.P., Longo V.D., Harvie M. Impact of intermittent fasting on health and disease processes. Ageing Res. Rev. 2017;39:46-58.

    Not in our library10.1016/j.arr.2016.10.005

  172. 172.

    Tinsley G.M., Paoli A. Time-Restricted eating and age-related muscle loss. Aging. 2019;11:8741-8742.

    Not in our library10.18632/aging.102384

  173. 173.

    Wallace M.A., Aguirre N.W., Marcotte G.R., Marshall A.G., Baehr L.M., Hughes D.C., Hamilton K.L., Roberts M.N., Lopez-Dominguez J.A., Miller B.F., et al. The ketogenic diet preserves skeletal muscle with aging in mice. Aging Cell. 2021;20:e13322.

    Not in our library10.1111/acel.13322

  174. 174.

    Zhou Z., Hagopian K., Lopez-Dominguez J.A., Kim K., Jasoliya M., Roberts M.N., Cortopassi G.A., Showalter M.R., Roberts B.S., Gonzalez-Reyes J.A., et al. A ketogenic diet impacts markers of mitochondrial mass in a tissue specific manner in aged mice. Aging. 2021;13:7914-7930.

    Not in our library10.18632/aging.202834

  175. 175.

    Thai P.N., Seidlmayer L.K., Miller C., Ferrero M., Dorn G.W., Schaefer S., Bers D.M., Dedkova E.N. Mitochondrial Quality Control in Aging and Heart Failure: Influence of Ketone Bodies and Mitofusin-Stabilizing Peptides. Front. Physiol. 2019;10:382.

    Not in our library10.3389/fphys.2019.00382

  176. 176.

    Parker B.A., Walton C.M., Carr S.T., Andrus J.L., Cheung E.C.K., Duplisea M.J., Wilson E.K., Draney C., Lathen D.R., Kenner K.B., et al. beta-Hydroxybutyrate Elicits Favorable Mitochondrial Changes in Skeletal Muscle. Int. J. Mol. Sci. 2018;19:2247.

    Not in our library10.3390/ijms19082247

  177. 177.

    Soto-Mota A., Vansant H., Evans R.D., Clarke K. Safety and tolerability of sustained exogenous ketosis using ketone monoester drinks for 28 days in healthy adults. Regul. Toxicol. Pharmacol. RTP. 2019;109:104506.

    Not in our library10.1016/j.yrtph.2019.104506

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