Review Article | Open Access
Vincent J. Miller, Frederick A. Villamena, Jeff S. Volek, "Nutritional Ketosis and Mitohormesis: Potential Implications for Mitochondrial Function and Human Health", Journal of Nutrition and Metabolism, vol. 2018, Article ID 5157645, 27 pages, 2018. https://doi.org/10.1155/2018/5157645
Nutritional Ketosis and Mitohormesis: Potential Implications for Mitochondrial Function and Human Health
Impaired mitochondrial function often results in excessive production of reactive oxygen species (ROS) and is involved in the etiology of many chronic diseases, including cardiovascular disease, diabetes, neurodegenerative disorders, and cancer. Moderate levels of mitochondrial ROS, however, can protect against chronic disease by inducing upregulation of mitochondrial capacity and endogenous antioxidant defense. This phenomenon, referred to as mitohormesis, is induced through increased reliance on mitochondrial respiration, which can occur through diet or exercise. Nutritional ketosis is a safe and physiological metabolic state induced through a ketogenic diet low in carbohydrate and moderate in protein. Such a diet increases reliance on mitochondrial respiration and may, therefore, induce mitohormesis. Furthermore, the ketone β-hydroxybutyrate (BHB), which is elevated during nutritional ketosis to levels no greater than those resulting from fasting, acts as a signaling molecule in addition to its traditionally known role as an energy substrate. BHB signaling induces adaptations similar to mitohormesis, thereby expanding the potential benefit of nutritional ketosis beyond carbohydrate restriction. This review describes the evidence supporting enhancement of mitochondrial function and endogenous antioxidant defense in response to nutritional ketosis, as well as the potential mechanisms leading to these adaptations.
All cells of the human body require ATP as the fundamental energy source to support life. Because mitochondria produce the majority of ATP, impaired mitochondrial function is implicated in the majority of today’s most concerning chronic and degenerative health conditions including obesity, cardiovascular disease, cancer, diabetes, sarcopenia, and neurodegenerative diseases . Much of this association between mitochondrial function and disease can be attributed to excessive mitochondrial production of reactive oxygen species (ROS) .
Although mitochondrial ROS (mtROS) are generally considered harmful, which is certainly the case at high concentrations, modest levels stimulate necessary biological processes such as proliferation, differentiation, and immunity . Adaptations that enhance resistance to oxidative stress are also induced by mtROS , possibly decreasing net ROS production during basal metabolism. This adaptive response is called mitohormesis [4–6] and is a promising mechanism through which lifestyle interventions that enhance mitochondrial function may, in turn, enhance resistance to chronic and degenerative diseases.
By dramatically shifting energy metabolism towards ketogenesis and fatty acid oxidation, ketogenic diets are likely to have a profound effect on mitochondrial function. However, despite the rapidly growing amount of research on ketogenic diets and their effects on various disease states, only a small amount of this research has focused on mitochondrial function or oxidative stress. The well-established increase in fat oxidation induced by a ketogenic diet [7, 8] clearly indicates prominent connection with mitochondrial function and, in turn, oxidative stress and mitohormesis [5, 6, 9]. Therefore, the purpose of this review is to describe the current, but limited, understanding of how ketogenic diets may affect mitochondrial function and resistance to oxidative stress, particularly within the context of extending human healthspan.
2. Nutritional Ketosis
The use of lifestyle interventions to treat and prevent chronic disease is attractive because of their potential to lower medical costs and produce more robust and holistic improvements in health. Ketogenic diets have been studied sporadically for more than 100 years, but over the last 15 years, a growing number of researchers have contributed to what is now a critical mass of discoveries that link the process of keto-adaptation to a broad range of health benefits [10–33]. Early clinical research focused on the use of “extreme” versions of ketogenic diets to treat seizures, but recent research indicates that benefits related to the management of epilepsy, weight loss, metabolic syndrome, and type 2 diabetes can be achieved with an approach that is less restrictive in carbohydrate and protein, and therefore more satisfying, sustainable, and feasible for the general population. A “well-formulated” ketogenic diet is generally characterized by a total carbohydrate intake of less than 50 g/d and a moderate protein intake of approximately 1.5 g/d per kg of reference weight . This typically increases circulating β-hydroxybutyrate (BHB) and acetoacetate (ACA) from concentrations that are typically less than 0.3 mM into the range of nutritional ketosis, which for BHB, we define as 0.5–3 mM . This range is below the typical 5–10 mM range for BHB that occurs during prolonged fasting, and well below concentrations characteristic of ketoacidosis [34, 35]. From the perspective of meeting energy demands, the reduced carbohydrate and moderate protein intakes necessarily make ketogenic diets high in fat. Despite this contradiction with mainstream dietary guidelines, ketogenic diets may be beneficial for many health conditions, particularly the previously mentioned conditions related to mitochondrial impairment, which includes obesity [10, 11], diabetes [12–14], cardiovascular disease [15–17], cancer [15, 18–26], neurodegenerative diseases [19, 20, 27–30], and even aging [31–33, 36, 37].
In both the nutrition literature and public dietary guidelines, nonstarchy vegetables are one of the few dietary components nearly unanimously agreed upon as healthful. Given their health-supporting characteristics and low carbohydrate content, they should be a prominent component of any ketogenic diet. Beyond the primary features of a well-formulated ketogenic diet, such as macronutrient proportions, adequate mineral intake, and appropriate selection of fat sources, which have been discussed more thoroughly elsewhere [34, 35], inclusion of nonstarchy vegetables is an important consideration, given that reports in the literature of adverse effects resulting from ketogenic diets are often associated with extreme implementations that typically lack plant matter. In fact, for this reason, it has recently been recommended to increase the nonstarchy vegetable content of ketogenic diets used to treat epilepsy . Beyond adding variety to the diet, benefits of nonstarchy vegetables that may be particularly relevant to nutritional ketosis include the maintenance of adequate micronutrient status and the presence of prebiotic fiber as substrate for the gut microbiota. In addition to the importance of prebiotic fiber for basic health, the short-chain fatty acids produced by the gut microbiota from this dietary fiber support ketogenesis  and metabolic signaling related to mitochondrial function and antioxidant defense . Furthermore, nonstarchy vegetables are a source of the many micronutrients needed to support energy metabolism. As such, there is more to a ketogenic diet than simply restricting carbohydrate. Selection of a variety of nutrient-dense foods is therefore an important component of nutritional ketosis that should be given consideration in any clinical or academic implementation.
3. Formation of mtROS and Associated Antioxidant Defense
As with other sources of oxidative stress, mtROS can damage enzymes and cell membranes and subsequently facilitate the pathogenesis of chronic disease . Oxidative damage to mitochondrial DNA (mtDNA) is of particular concern because of its proximity to the electron transport chain (mtETC). Furthermore, compared to nuclear DNA, mtDNA is more prone to oxidative damage and is not repaired as effectively [42–45], although this has been debated based on more recent evidence [46–50]. Unrepaired mtDNA damage leads to mitochondrial dysfunction, which is implicated in the pathogenesis of a variety of chronic diseases  and associated with shorter lifespan . Therefore, while moderate levels of mtROS have important roles in signaling and health, protection against excessive levels is also important.
Although there are numerous sites of mtROS formation, the most prominent are those in the mtETC, where the superoxide radical (O2•−) is formed through reduction of O2 by leaked electrons, particularly at complexes I and III [41, 53–55]. Production of O2•− at complex I is particularly high during reverse electron transport (RET), which occurs when a high proton-motive force (Δp) develops across the inner mitochondrial membrane in conjunction with the pool of coenzyme Q (CoQ) being in a highly reduced state (i.e., mostly present as ubiquinol) as a result of electron transfer through complex II and electron transfer flavoprotein:ubiquinone oxidoreductase (ETF-QO) [56–62]. This dependence of mtROS production on Δp during RET is mainly influenced by proton gradient (ΔpH) .
Formation of O2•− at complexes I and III primarily occurs in the mitochondrial matrix, but some of the O2•− produced at complex III is produced in the intermembrane space . Within the matrix, O2•− is rapidly dismutated into hydrogen peroxide (H2O2) by manganese superoxide dismutase (SOD2) [41, 53]. Some O2•− may escape into the mitochondrial intermembrane space  and cytosol , where copper/zinc superoxide dismutase (SOD1) can dismutate it into H2O2 . The large majority of mitochondrial H2O2 is removed by peroxiredoxin (Prx) 3, followed by much smaller contributions from Prx5 and glutathione peroxidases (GPx) 1 and 4 . GPx also removes other peroxides, including lipid hydroperoxides . Catalase is another antioxidant enzyme capable of removing H2O2 but is primarily located in peroxisomes and is therefore unlikely to directly remove mitochondrial H2O2 [41, 66]. However, H2O2 can be transported out of mitochondria , and it is possible that the majority of mitochondrial H2O2 is removed in the cytosol. Since Prxs and GPxs rely on NADPH for recycling of their cofactors (thioredoxins and glutathione, resp.) , and since NADH is required for recycling of NADPH , activity of these enzymes would decrease availability of NADH for oxidative phosphorylation. Therefore, transport of H2O2 out of mitochondria for removal in the cytosol may be a more likely defense mechanism , implying a more important role of catalase and other antioxidant enzymes outside of mitochondria. Despite the lower reactivity of H2O2, it is still reactive and can oxidize metal ions, particularly iron, to form the hydroxyl radical (•OH), which readily damages DNA, lipids, and proteins . •OH is scavenged by metallothioneins I and II [69, 70] and glutatathione , indicating that these antioxidant proteins may be important defenses against byproducts of unaddressed mtROS. Other important antioxidant enzymes include glutamate-cysteine ligase (GCL), which is the rate-limiting step in glutathione synthesis, and glutathione reductase (GSR) and thioredoxin reductase (TRXR), which recycle glutathione and thioredoxin, respectively, to their reduced forms .
Increased reliance on mitochondrial respiration will increase the flow of electrons through the mtETC and, in turn, increase the potential for mtROS formation. Although oxidative stress is traditionally viewed as harmful, a modest increase in ROS is now established as a signaling stimulus that induces hormetic adaptation . In regard to mitohormesis and mtROS, such adaptation is largely centered around antioxidant defense [4–6], making mitohormesis an attractive target for the prevention and treatment of chronic disease.
Although mitohormesis has not been studied comprehensively in higher-level organisms, its occurrence is supported by compelling evidence in lower-level organisms. For example, inhibition of glycolysis in C. elegans increased fat oxidation (based on nematode triglyceride content) and mitochondrial O2 consumption, which was followed by increases in ROS production at day 2 and catalase activity at day 6 . The increase in catalase activity occurred in conjunction with increases in lifespan and resistance to the mitochondrial stressors sodium azide and paraquat. However, antioxidant treatment (N-acetylcysteine) decreased the elevation of ROS at day 2 and eliminated the resistance to sodium azide and paraquat treatments, indicating a requirement of ROS as a stimulus .
In a subsequent series of experiments, glucose metabolism in C. elegans was inhibited by knockdown of the insulin receptor, insulin-like growth factor 1 (IGF-1) receptor, and insulin receptor substrate 1 (IRS-1) . Consistent with the previous study , inhibition of glucose metabolism increased mitochondrial respiration concomitant with ROS-dependent increases in lifespan, stress resistance, and antioxidant enzyme activity. However, in this case, detection of ROS was mitochondria-specific, and repeated measures allowed for changes in antioxidant enzyme activities to be evaluated more closely in relation to the timing of changes in mtROS. Compared to controls, inhibition of glucose metabolism resulted in higher mitochondrial O2 consumption at 12 h, higher mtROS production at 24 h, and higher activities of SOD and catalase at 48 h, suggesting a dependence of antioxidant activity on mtROS and a dependence of mtROS on mitochondrial respiration. The most striking result is the lower mtROS at 120 h, indicating that the initial increase in mtROS and subsequent increase in antioxidant enzyme activity ultimately lowered net mtROS production to a level lower than controls, which is the proposed explanation for the more than twofold increase in lifespan. As with the previous study, this demonstration of mitohormesis is further supported by the changes in ROS production, antioxidant enzyme activity, and lifespan having been prevented with antioxidant treatment.
The occurrence of mitohormesis is further supported by the potential for mtROS to simultaneously induce bioenergetic and antioxidant adaptations through a single signaling mediator. As discussed later in this review, this mediator is the transcription factor peroxisome proliferator-activated receptor γ coactivator 1α (PGC-1α), and its role in simultaneously inducing bioenergetic and antioxidant adaptations has been demonstrated in several experimental models of mtROS production, including treatments with paraquat and H2O2. Paraquat is an herbicide that is reduced by the mtETC and subsequently initiates mtROS production by reacting with O2 to produce O2•− [74, 75], and H2O2 is a common form of mtROS. Treatment of a fibroblast cell line (10T1/2) with H2O2 has induced expression of both antioxidant enzymes (SOD1, SOD2, and GPx1) and proteins involved in mitochondrial oxidative phosphorylation, all in a manner largely dependent on PGC-1α . Demonstrating the hormetic benefit of this response in a variety of brain cells, overexpression of PGC-1α protected against cell death induced by H2O2 or paraquat treatment, and this occurred in conjunction with changes in gene expression similar to those observed with the 10T1/2 cells . Although the central role of PGC-1α in linking mitochondrial bioenergetics with antioxidant defense appears to not have been thoroughly investigated in vivo, some suggestive evidence does exist. In skeletal muscle of mice treated with paraquat, content of proteins involved in mitochondrial oxidative phosphorylation and uncoupling have been found to increase in conjunction with greater nuclear localization of PGC-1α . Traditional antioxidant proteins were not measured, but, as will be discussed later, the increase in uncoupling proteins can be regarded as an indication of enhanced antioxidant defense based on the potential of these proteins to decrease mtROS production.
Ketogenic and low-carbohydrate diets greatly increase reliance on fat oxidation [78–89], which would logically be expected to increase mitochondrial respiration and mtROS production and, in turn, induce mitohormesis. Furthermore, mtROS produced through RET appears to have particular relevance to hormetic adaptation, including increased lifespan . Nutritional ketosis is likely to increase RET by altering the FADH2 to NADH ratio. As the primary source of acetyl CoA shifts from glycolysis to β-oxidation and ketolysis, this ratio increases, more than doubling for β-oxidation of longer-chain fatty acids. Electrons from FADH2 reduce the CoQ pool through complex II and ETF-QO, thereby increasing RET [91, 92]. This induction of RET by alteration of substrate availability can also be influenced by configuration of mtETC complexes into supercomplexes . The greater potential for mtROS production through RET is consistent with evidence of mitochondria producing more H2O2 during oxidation of palmitoyl carnitine versus pyruvate [93, 94]. Furthermore, succinate is generated during ketolysis by succinyl-CoA:3-oxoacid CoA-transferase (SCOT), which also promotes RET by reducing the CoQ pool through complex II. Demonstrating the likely role of RET in mitohormesis, particularly within the context of nutritional ketosis, extension of lifespan in C. elegans through BHB treatment is dependent on both complex I function and expression of bioenergetic and antioxidant proteins .
A study on hippocampal mitochondrial function in rats more directly supports the induction of mitohormesis by a ketogenic diet. After the first day of the diet (Bio-Serv F3666), H2O2 production by isolated mitochondria was increased . After the third day, mitochondrial levels of oxidized glutathione (GSSG) and hippocampal levels of 4-hydroxy-2-nonenal (4-HNE) were also increased, further indicating an increase in oxidative stress. However, at completion of the first week, upregulation of antioxidant signaling occurred, indicated by increased nuclear content and transcriptional activity of nuclear factor erythroid-derived 2-like 2 (NFE2L2), which persisted through the remainder of the study. By the third week, mitochondrial H2O2 production decreased to below baseline . In the liver, content of reduced acetyl CoA, which is indicative of mitochondrial redox status, decreased after three days of the ketogenic diet, but increased relative to the control diet after three weeks, indicating an initial increase in oxidative stress followed by a decrease . This was in conjunction with changes in NFE2L2 nuclear content and transcriptional activity similar to those observed in the hippocampus. As with the previously described C. elegans experiments, the time course of these observations is a strong indication of mitohormesis, and the similarity in results between the liver and hippocampus suggests that a ketogenic diet can induce mitohormesis in a variety of tissues.
Several other rodent studies provide additional evidence of ketogenic diets upregulating antioxidant defense, but without enough data to convincingly attribute the results to mitohormesis. Content of SOD2 has increased in the livers of mice fed a ketogenic diet (% energy: 89 fat, <1 carbohydrate, and 10 protein), which occurred in conjunction with increased median lifespan and decreases in tumors and age-associated losses of physical and cognitive performance . In addition, activity of GCL and the protein content of its two subunits increased in the hippocampal homogenate of rats fed a ketogenic diet (Bio-Serv F3666) for 3 weeks . This was in conjunction with higher levels of reduced glutathione (GSH) and lower ROS production in hippocampal mitochondria. The ketogenic diet also increased resistance to mtDNA damage in hippocampal mitochondria exposed to H2O2 . Consistent with these results, total antioxidant capacity and activities of GPx and catalase were increased in hippocampal homogenate of rats fed a ketogenic diet (% energy: 86 fat, <1 carbohydrate, and 13 protein) for 8 weeks . Furthermore, in cortical homogenate of rats induced with traumatic brain injury, a ketogenic diet increased cytosolic and mitochondrial protein contents of NAD(P)H:quinone oxidoreductase 1 (NQO1) and SOD1, as well as mitochondrial protein content of SOD2, and also prevented mitochondrial oxidative damage (indicated by 4-HNE) .
Additional evidence, although disparate and primarily based on neuronal mitochondrial function related to epileptic seizures, further supports the potential for nutritional ketosis to induce mitohormesis . Much of this is based on signal transduction, antioxidant defense, and oxidative capacity, all of which will be discussed in proceeding sections.
5. Ketones as Antioxidants and Signaling Molecules
Although ketones may not induce mitohormesis directly, they do influence antioxidant defense (Figure 1). Furthermore, ketone metabolism is highly relevant to mitochondrial adaptation since the ketogenic and ketolytic enzymes needed to support nutritional ketosis are located in mitochondria.
BHB, in addition to being an important energy substrate, is also a signaling molecule [100–102]. Although not induced through mtROS, BHB inhibits class I and II histone deacetylases (HDACs) in a dose-dependent manner, resulting in greater histone acetylation regardless of whether BHB is elevated through fasting, caloric restriction, or infusion . This inhibition is associated with increased expression of forkhead box O (FOXO) 3a and metallothionein II and increased protein content of FOXO3a, SOD2, and catalase . Consistent with these changes, the kidneys of mice with elevated blood BHB concentrations (∼1.2 mM) were protected from paraquat-induced (50 mg/kg) oxidative damage to proteins and lipids, which was indicated by lower levels of protein carbonyls, 4-HNE, and lipid peroxides . Upregulation of antioxidant defense by BHB-induced HDAC inhibition also appears to be the mechanism through which exogenous BHB extends lifespan in C. elegans . The dependence of this response on FOXO3a, NFE2L2, and several bioenergetic signaling proteins that influence the activities of these two transcription factors  is indicative of the overlap between bioenergetics and antioxidant defense that is characteristic of mitohormesis.
Additional indications of exogenous BHB upregulating antioxidant defense have been observed, although without consideration of HDAC inhibition. In rats, injection of BHB has increased activities of SOD and catalase and prevented the increase in lipid peroxidation and decreases in SOD, catalase, and GSH induced by paraquat injection, all of which were observed in kidney homogenate . Furthermore, BHB also prevented the paraquat-induced decrease in nuclear NFE2L2, indicating involvement of antioxidant signaling . Similarly, BHB treatment has increased FOXO3a, SOD2, and catalase content in cardiomyocytes , indicating that BHB may also influence antioxidant defense in the heart. In this study, BHB also prevented the decrease of FOXO3a, SOD2, and catalase content that resulted from H2O2 treatment . Despite the amount of research that has been done on the antiseizure mechanisms of ketogenic diets, the influence of BHB on HDAC inhibition and related antioxidant defense appears to have not yet been investigated in brain tissue. However, BHB appears to inhibit HDAC2 in microvascular and neuronal brain cells , and BHB-induced HDAC inhibition is thought to have a role in the antiseizure effects of ketogenic diets .
In addition to BHB inducing upregulation of antioxidant defense, ketones have direct antioxidant capacity. BHB scavenges •OH, as does ACA, although to a lesser extent . The applicability of this antioxidant capacity has been investigated in vitro and in vivo in the context of hypoglycemia. In cultured hippocampal neurons, treatment with BHB or ACA decreased ROS during hypoglycemia induced through inhibition of glycolysis, and in hypoglycemic rats, infusion of BHB decreased hippocampal lipid peroxidation .
More specific to mitochondrial function, treatment with BHB + ACA (1 mM each) has decreased O2•− production in isolated rat neuronal mitochondria following glutamate exposure . This occurred in conjunction with decreased NADH levels, suggesting that ketones may additionally decrease mtROS production by enhancing electron transport along the mtETC after NADH oxidation and, in turn, decreasing mitochondrial Δp and associated O2•− production. The observed decrease in mitochondrial O2•− production occurred independently of glutathione , but in isolated and stunned hearts from guinea pigs, treatment with 5 mM ACA increased GSH and the NADPH/NADP+ ratio , suggesting that glutathione may be involved to some extent.
Further indicating that ketones influence mtROS production through alteration of electron transport, treatment of rat hippocampal slices with BHB + ACA (1 mM each) prevented the increase in mtROS and mitigated the decrease in ATP production that otherwise result from inhibition of mtETC complex I with rotenone . In mitochondria isolated from the brains of mice injected with BHB, although inhibition of complex I with rotenone and 1-methyl-4-phenylpyridinium increased rather than decreased mtROS production, the BHB treatment prevented the decrease in O2 consumption caused by inhibition of complex I, and this occurred independently of uncoupling . Consistent with the results from hippocampal brain slices, the BHB treatment also mitigated the decrease in ATP production caused by complex I inhibition . These effects were prevented by inhibition of complex II with 3-nitropropionic acid or malonate, indicating that BHB primarily influences mitochondrial respiration at complex II , which is consistent with ketolysis increasing formation of succinate and FADH2. However, in mutated cells prone to complex I disassembly and an associated severe decrease in complex I activity, treatment with BHB + ACA (5 mM each) increased both the assembly and activity of complex I , indicating that ketones somehow promote repair of complex I damage and may therefore influence mitochondrial respiration at more than one site.
Another aspect of mitochondrial function influenced by ketones is the mitochondrial permeability transition pore (mPTP). Prolonged opening of the mPTP is one of the mechanisms through which mtROS can induce cellular injury and promote disease . In neurons isolated from rat brain slices, treatment with BHB + ACA has decreased the mtROS production, mPTP opening, and cell death induced by H2O2 . This protective effect was duplicated with catalase, even in conjunction with diamide-induced opening of the mPTP, indicating that the protective effect of BHB and ACA is at least partly due to defense against ROS . In a mouse model of epilepsy, this decrease in mPTP opening was found to be induced exclusively by BHB, and in a manner dependent on the cyclophilin D subunit of the mPTP . BHB in combination with ACA also appears to promote opening of mitochondrial ATP-sensitive K+ (mtKATP) channels , which in heart mitochondria is known to protect against Ca+ overload  and dissipate membrane potential (ΔΨ) . Since high ΔΨ promotes mtROS production, dissipation of ΔΨ through mtKATP channels may partly explain the potential for ketones to decrease mtROS production. However, opening of mtKATP channels by pinacidil decreases mitochondrial ATP production , which is consistent with dissipation of ΔΨ and suggests a compromise between ATP and mtROS production.
Ketones may also be important, or even necessary, for the bioenergetic signaling associated with mitohormesis. As will be discussed later, peroxisome proliferator-activated receptor α (PPARα) is a nuclear receptor that is responsible for many of the bioenergetic adaptations associated with nutritional ketosis and mitohormesis . In mice, a ketogenic diet (% energy: 90 fat, 0 carbohydrate, and 10 protein) increased blood BHB concentration to 1-2 mM and upregulated expression of numerous PPARα targets in the liver . However, in mice fed a nonketogenic low-carbohydrate diet (% energy: 75 fat, 15 carbohydrate, and 10 protein), which did not raise blood concentration of BHB, the increased expression of PPARα targets did not occur , implying that induction of PPARα signaling by a ketogenic diet is dependent on ketones. This response may be, at least in part, a result of the epigenetic effects of BHB. In addition to HDAC inhibition, BHB also influences gene expression through β-hydroxybutyrylation of histone lysine residues . In the livers of mice subjected to prolonged fasting, this β-hydroxybutyrylation has been associated with upregulation of PPAR signaling, oxidative phosphorylation, fatty acid metabolism, the proteasome, and amino acid metabolism related to redox balance . Upregulation of these pathways is largely influenced by β-hydroxybutyrylation of the histone residue H3K9 , which is also involved in the upregulation of antioxidant defense through BHB-induced HDAC inhibition . This potential for BHB to influence expression of both mitochondrial and antioxidant genes through a common histone residue is further indication of the overlap between bioenergetics and antioxidant defense and suggests that if mitohormesis is indeed induced during nutritional ketosis, induction may be dependent on ketones and may therefore not occur during a low-carbohydrate diet that is not ketogenic.
In regard to the practicality of BHB signaling, many of the outcomes described above, including HDAC inhibition, were achieved with BHB concentrations within the range of 0.6–2 mM [37, 103, 105, 108, 109, 111, 112, 116], which is well within the physiological range of nutritional ketosis for humans and even suggests potential benefit at low to moderate levels.
6. Mitochondrial Uncoupling
As previously discussed, RET is a prominent source of mtROS and is dependent on a high Δp across the inner mitochondrial membrane. During ATP production, Δp is dissipated as H+ enters the mitochondrial matrix through ATP synthase. Mitochondrial uncoupling also dissipates Δp, but by allowing translocation of H+ into the matrix independent of ATP synthase. Uncoupling is therefore regarded as an antioxidant defense in that it can mitigate mtROS production [122–126]. In fact, only a small dissipation of ΔΨ or ΔpH (components of Δp) is needed for a large decrease in mtROS production [57–60, 127].
Mitochondrial uncoupling is primarily facilitated by uncoupling proteins (UCPs) and adenine nucleotide translocase (ANT) [124, 128, 129]. Although UCP1 is primarily expressed in brown adipose, UCP2 is expressed across a wide variety of tissues, and expression of UCP3 appears to be limited to skeletal muscle and the heart . Knockout of UCP2  or UCP3 [94, 132] increases mtROS production, and both proteins are inactivated through glutathionylation by GSH , further establishing their involvement in antioxidant defense. UCP2 and UCP3 may also be activated by products of lipid peroxidation induced by mtROS . However, the potential for UCP2 and UCP3 to reduce mtROS through uncoupling is not fully agreed upon;  UCPs may alternatively protect against oxidative damage merely by exporting lipid hydroperoxides . Furthermore, UCP3 is less abundant in type I and type IIa muscle fibers , which are more oxidative, and its expression and content are further decreased by endurance exercise training [135, 136], suggesting that UCP3 may not be a primary defense against mtROS.
Although the primary purpose of ANT is to exchange newly synthesized ATP in the mitochondrial matrix for cytosolic ADP , it shares a common feature with UCPs and other inner membrane proteins in that they translocate anions, including fatty acids. The uncoupling mechanism of ANT, along with UCP2 and UCP3, may be the exchange of protonated fatty acids from the mitochondrial intermembrane space for fatty acid anions in the matrix, thereby dissipating Δp [123, 137–139]. Inhibition studies indicate that ANT may contribute more to uncoupling than UCPs [140, 141].
Independent of nutritional ketosis, increased dietary fat intake increases expression of UCP2 and UCP3 in muscle , and fatty acids facilitate uncoupling through UCP2 [143, 144], UCP3 [94, 143, 144], and ANT . Given the high fat intake that is characteristic of a ketogenic diet, it is logical to expect nutritional ketosis to increase mitochondrial uncoupling.
Certain ionophores are capable of completely uncoupling mitochondria by transporting H+ across the inner membrane. Such ionophores are therefore commonly used to measure maximal mitochondrial respiration. In mice fed a ketogenic diet (Bio-Serv F3666, ∼6 : 1 ratio of fat to carbohydrate + protein) for 6 days, respiration of hippocampal mitochondria was fully uncoupled with the ionophore carbonyl cyanide 4-(trifluoromethoxy)phenylhydrazone (FCCP) . The ratio of respiration during oxidation of palmitic acid to maximally uncoupled respiration induced by FCCP was greater in response to the ketogenic diet, indicating increased uncoupling . Although this interpretation relies on the assumption that ATP production was not changed by diet, it is further supported by the higher levels of UCP2, UCP4, and UCP5 detected in mitochondria after the ketogenic diet. Furthermore, mtROS production was lower in the ketogenic diet group , supporting the role of uncoupling as an antioxidant defense. Although not based on direct measurement of mitochondrial function, in rats fed a ketogenic diet (% energy: 89.5 fat, 0.1 carbohydrate, and 10.4 protein), increased uncoupling in response to nutritional ketosis is further indicated by increases in fat oxidation and overall O2 consumption occurring in conjunction with decreases in CO2 production and energy expenditure . However, based on observations of greater palmitate-induced uncoupling (determined by measurement of ΔΨ) during state 4 respiration in rats fed a high-fat, low carbohydrate diet (% energy: 50 fat, 21 carbohydrate, and 29 protein)  that was likely too high in carbohydrate and protein to induce nutritional ketosis, it is possible that moderate carbohydrate restriction may increase mitochondrial uncoupling independently of ketones.
Several additional rodent studies have shown ketogenic diets to increase protein content of UCPs. However, since mitochondrial function was not measured in these studies, it is not known if uncoupling was affected by these changes in UCP content. In obese mice fed a ketogenic diet (0.4% of energy as carbohydrate), expression of UCP1 and UCP2 increased in adipose and the liver, respectively . Similarly, expression of UCP1 has increased in brown adipose of mice fed a low-carbohydrate diet (18.5% of energy) supplemented with ketone esters (6% w/v) . The increase in hepatic UCP2 expression during a ketogenic diet has been demonstrated by other studies as well [37, 150, 151]. Ketogenic diets also induce expression of UCP3 in skeletal muscle. In rats fed a ketogenic diet (% energy: 78.1 fat, 0 carbohydrate, and 21.9 protein) for 8 weeks, UCP3 expression increased in the soleus but not the extensor digitorum longus, which is consistent with the soleus containing mostly oxidative, type I muscle fibers . In humans, glycogen depleting exercise followed by two days of a low-carbohydrate diet (0.7 g/kg body mass) increased UCP3 expression in the vastus lateralis .
7. Oxidative Capacity
As the rate of oxidative phosphorylation approaches the capacity of the mtETC, Δp will increase and facilitate mtROS production . Higher oxidative capacity should therefore decrease the potential for mtROS production and subsequent oxidative damage. Furthermore, greater oxidative capacity may compensate for the resulting decrease in efficiency of ATP production associated with increased mitochondrial uncoupling. Since oxidative phosphorylation occurs exclusively in mitochondria, mitochondrial density is a key determinant of oxidative capacity .
As previously mentioned, numerous studies have demonstrated a profound increase in fat oxidation in response to ketogenic and low-carbohydrate diets. Some studies have even shown an increase in O2 consumption [148, 155–158]. However, fats contain fewer oxygen atoms per carbon than carbohydrates, thereby necessitating greater O2 intake to produce the same amount of energy . Furthermore, since β-oxidation and ketolysis produce a greater proportion of FADH2 to NADH, the resulting decrease in passage of electrons through complex I decreases potential for ATP production per unit of O2 consumption . Increased O2 consumption in response to a ketogenic diet may therefore merely be an effect of the differences in the metabolism and molecular structures of fat and carbohydrate rather than a true indication of increased capacity for oxidative phosphorylation. However, in rat hearts perfused with glucose, the addition of ketones has decreased O2 consumption . This discrepancy may be related to variations in mitochondrial uncoupling. Either way, several studies have shown ketogenic diets to increase mitochondrial content, and numerous studies have shown these diets to increase expression, content, or activity of mitochondrial proteins involved in oxidative phosphorylation and fat oxidation. Compared to O2 consumption alone, these findings provide more conclusive support for an increase in oxidative capacity in response to nutritional ketosis.
In rats fed a ketogenic diet (Bio-Serv F3666) for 22 days, mitochondrial density (determined by electron microscopy) in the hippocampus increased in conjunction with increased transcription of 39 of the 42 mitochondrial proteins analyzed . Similarly, mitochondrial content (mtDNA copy number) increased in skeletal muscle of mice fed a ketogenic diet (Research Diets D05052004; % energy: 89.5 fat, 0.1 carbohydrate, and 10.4 protein) for 10 months . Higher mtDNA copy number was also observed in skeletal muscle of rats fed a high-fat, low-carbohydrate diet (% energy: 60 fat, 20 carbohydrate, and 20 protein) for 4 weeks in conjunction with daily injections of heparin (0.5 U/g) to increase circulation of fatty acids . In humans, after just 3 days of a low-carbohydrate, high-fat diet (% energy: 50 fat, 34 carbohydrate, and 16 protein), fat oxidation significantly increased and 49% of the variance was explained by mtDNA content . Despite this, the content of mtDNA did not change significantly, but this was expected given the brief duration of the diet.
As will be discussed in the following sections, many of the signaling proteins involved in regulating antioxidant defense also regulate oxidative phosphorylation and fat oxidation. There is abundant evidence (Table 1) showing ketogenic and low-carbohydrate diets to increase expression, content, or activity of many targets of these signaling proteins, further indicating increased oxidative capacity. It is particularly striking that ketogenic or low-carbohydrate diets upregulate expression of proteins associated with each of the five mtETC complexes.
8. Bioenergetic Signal Transduction
Perturbations in bioenergetic homeostasis induce signal transduction that leads to upregulation of mitochondrial capacity and antioxidant defense. Three key enzymes involved in the sensing of these perturbations and the subsequent induction of signal transduction are AMP-activated protein kinase (AMPK) and silent mating type information regulation 2 homologues 1 and 3 (SIRT1 and SIRT3).
AMPK is activated through phosphorylation of the Thr172 residue of the AMPK α catalytic subunit [174–176], and this phosphorylation is largely regulated by molecules related to bioenergetic homeostasis including AMP, ADP, catecholamines, adiponectin, glycogen, and insulin. In general, AMPK is activated by energy deficit and induces signaling that upregulates energy production. AMP and ADP are direct byproducts of energy depletion while adiponectin and catecholamines serve as endocrine signals to increase energy production, often in response to energy depletion. In contrast, indications of energy surplus, such as glycogen and insulin, inhibit activation of AMPK. Nutritional ketosis increases the aforementioned factors that activate AMPK and decreases those that inhibit AMPK, suggesting that nutritional ketosis is similar to caloric restriction in inducing a signal of energy depletion.
AMP competes with ATP for binding to the γ regulatory subunit of AMPK [177, 178] and by doing so, greatly increases AMPK activity, but only in the presence of an upstream kinase such as liver kinase B1 (LKB1) . This binding of AMP to the γ subunit appears to promote AMPK activity through at least two mechanisms: facilitated phosphorylation of the α subunit [180–183] and inhibition of dephosphorylation by protein phosphatases 2Cα and 2Ac [179, 181, 183, 184]. ADP also binds to the γ subunit of AMPK to inhibit dephosphorylation [183, 185, 186] and possibly facilitate phosphorylation . This is important to the energy sensing sensitivity of AMPK based on the much higher physiological concentration of ADP compared to AMP . Data on changes in AMP and ADP levels in response to a ketogenic diet are lacking. However, the decreased availability of carbohydrate and increased mitochondrial uncoupling (previously described) during nutritional ketosis suggest a decline in ATP production, at least until compensatory adaptations occur. A decline in ATP implies a relative increase in AMP and ADP, which would facilitate AMPK phosphorylation and activation. In addition, ketogenic diets are commonly reported to have a satiating effect , which may further increase the AMP and ADP to ATP ratios through spontaneous caloric restriction.
Adiponectin increases AMPK activity in skeletal muscle [188, 189] and the liver  by promoting Thr172 phosphorylation, likely in response to an increase in the AMP to ATP ratio . Similarly, α-adrenergic signaling increases AMPK activity in skeletal  and cardiac muscle , and β-adrenergic signaling increases AMPK activity in adipose [192, 193], all through promotion of Thr172 phosphorylation. While activation through β-adrenergic signaling appears to involve the AMP to ATP ratio , α-adrenergic signaling appears to work independently of AMP and ATP . Increases in adiponectin have been observed during ketogenic or low-carbohydrate diets, although primarily in obese individuals [194–196]. BHB induces adiponectin secretion in adipocytes , indicating that the level of nutritional ketosis may be an important determinant of the extent to which ketogenic diets influence AMPK activity through adiponectin. In regard to catecholamines, epinephrine increases during fasting, and this appears to be dependent on carbohydrate restriction , implying that epinephrine is likely to be elevated during nutritional ketosis. Consistent with this, dietary carbohydrate restriction increases catecholamines at rest [155, 199] and in response to exercise [155, 199–202]. This may be, at least in part, a result of glycogen depletion [200, 203], suggesting both direct and indirect effects of glycogen on AMPK activity. The potential for nutritional ketosis to increase catecholamines is further supported by the dependency of the antiseizure effects of ketogenic diets on norepinephrine .
Glycogen influences AMPK activity by binding to a glycogen binding domain on the β regulatory subunit of AMPK [205, 206]. In human and rodent skeletal muscle, AMPK activity is lower when glycogen is bound to this domain [207, 208] and higher when muscle is depleted of glycogen [209–212]. In direct contrast to the effect of AMP and ADP, glycogen inhibits the phosphorylation of AMPK by upstream kinases such as LKB1 . Although muscle glycogen concentration has recently been demonstrated to be similar in ultra-endurance athletes regardless of a ketogenic or high-carbohydrate diet , concentrations generally decrease in response to dietary carbohydrate restriction [156, 166, 173, 214–221]. Furthermore, the long-term adaptations to nutritional ketosis that may enable some athletes to replenish glycogen at a normal rate may not apply to less physically active individuals.
Insulin inhibits AMPK activity by stimulating protein kinase B (PKB) to phosphorylate the Ser485 residue of the α subunit, thereby inhibiting phosphorylation at Thr172 . One of the most prominent features of nutritional ketosis is that, due to restricted carbohydrate intake, postprandial insulin is dramatically decreased. Furthermore, numerous studies have shown ketogenic or low-carbohydrate diets to decrease fasting insulin [155, 195, 223–225], particularly in the presence of metabolic dysregulation associated with hyperinsulinemia [84, 226–229].
Consistent with the mechanisms described above, changes in AMPK in response to a ketogenic or low-carbohydrate diet have been reported in several studies. In rodents, a ketogenic diet (Bio-Serv F3666) has increased AMPK activity in skeletal muscle  and AMPK phosphorylation in the liver , and a low-carbohydrate diet (18.5% of energy) supplemented with ketone esters (6% w/v) increased AMPK content in brown adipose . In humans, a nonketogenic low-carbohydrate diet (% energy: 50 fat, 30 carbohydrate, and 20 protein) has increased AMPK phosphorylation in skeletal muscle .
8.2. SIRT1 and SIRT3
The sirtuin isoforms SIRT1 [232, 233] and SIRT3 [234–236] are nicotinamide adenine dinucleotide- (NAD+) dependent deacetylases associated with longevity. Many reactions are regulated by the redox state of NAD+ and its phosphorylated form, NADP+. Among these reactions, a prominent role of reduced NADP+ (i.e., NADPH) is to support reductive biosynthesis and antioxidant defense, requiring the NADP+/NADPH ratio to be kept low . In contrast, the NAD+/NADH ratio is kept high to support energy metabolism , thereby linking sirtuin function to bioenergetic status . Although sirtuins are inhibited by high concentrations of NADH, their activity is influenced more by absolute NAD+ concentration than the NAD+/NADH ratio .
SIRT1 is present in the cytosol and nucleus , while SIRT3 is primarily located in mitochondria where it regulates bioenergetics and ROS production [239–241]. The sirtuins, particularly SIRT1, appear to participate in a feed-forward cycle of reciprocal activation with AMPK. In skeletal muscle, AMPK indirectly activates SIRT1 by increasing NAD+ through increased mitochondrial β-oxidation  and increased expression of nicotinamide phosphoribosyltransferase (NAMPT) , which is the rate-limiting enzyme in NAD+ synthesis . Completing the cycle, SIRT1 and SIRT3 can deacetylate and activate LKB1, thereby promoting further activation of AMPK. LKB1 is known to be activated by SIRT1 in adipose and liver  and by SIRT3 in cardiac muscle .
Based on the reciprocal activation described above, nutritional ketosis is likely to activate SIRT1 and SIRT3 indirectly through activation of AMPK. However, more direct activation of sirtuins by nutritional ketosis is possible. Since reduction of NAD+ to NADH occurs outside of mitochondria only during glycolysis, which is less active during nutritional ketosis, more cytosolic NAD+ remains oxidized, further facilitating activation of SIRT1 . In addition to the decrease in glucose availability during nutritional ketosis, glycolysis may be further inhibited through activation of pyruvate dehydrogenase kinase and subsequent inhibition of pyruvate dehydrogenase (PDH), which occurs in response to dietary carbohydrate restriction [248–251] or infusion of BHB, ACA, or fatty acids . Consistent with the relevance of these factors to nutritional ketosis, a ketogenic diet (% energy: 89 fat, <1 carbohydrate, and 10 protein) has decreased expression of PDH in mouse liver . More importantly, there is direct evidence of nutritional ketosis promoting an increase in NAD+ concentration. Treatment with BHB + ACA (1 mM each) has increased NADH oxidation in rat neocortical mitochondria , and a ketogenic diet (Bio-Serv F3666) has increased NAD+ concentration in rat hippocampus . There is also evidence of nutritional ketosis regulating sirtuin expression. A low-carbohydrate (20% of energy) diet combined with ketone esters (6% w/v) has increased SIRT1 protein content in brown adipose of mice , and a ketogenic diet (% energy: 90 fat, 0 carbohydrate, and 10 protein) has increased SIRT3 expression in mouse liver .
In addition to the downstream bioenergetic and antioxidant signaling induced by sirtuins, they directly facilitate ketogenesis and β-oxidation. SIRT1  and SIRT3  deacetylate 3-hydroxy-3-methylglutaryl CoA (HMG CoA) synthase, which is the rate-limiting enzyme for ketogenesis , resulting in increased levels of β-hydroxybutyrate . In addition, SIRT3 deacetylates and increases activity of long-chain acyl-CoA dehydrogenase (LCAD) , which participates in β-oxidation and therefore supports ketogenesis. SIRT3 has a similar influence on medium-chain acyl-CoA dehydrogenase (MCAD) as well . Since sirtuins facilitate ketogenesis, which then facilitates sirtuin activation, nutritional ketosis may promote, to some extent, a feed-forward cycle of sirtuin activity.
8.3. Direct Involvement of AMPK and Sirtuins in Redox Balance
Although the majority of links between energy sensing and antioxidant defense are manifested further downstream, there is some direct influence at the level of AMPK and sirtuins. AMPK is activated by oxidative stress [259, 260], likely through ATP depletion and a subsequent increase in the AMP to ATP ratio, or facilitation of tyrosine phosphorylation, which occurs independently of AMP and ATP concentrations . SIRT3 contributes more directly to antioxidant defense by deacetylating and activating SOD2 [261–263]. The overlapping effect of SIRT3 on antioxidant defense and bioenergetics is further supported by SIRT3 knockout increasing lipid peroxidation in conjunction with decreased O2 consumption in mouse skeletal muscle and also by SIRT3 knockdown increasing H2O2 production and decreasing O2 consumption in myoblasts .
9. Downstream Bioenergetic and Antioxidant Signaling
AMPK and sirtuins are the interface between the metabolic stimuli of nutritional ketosis and the downstream signaling that influences expression of proteins related to bioenergetics and antioxidant defense. Some of the primary downstream signaling molecules involved include PGC-1α, FOXO3a, nuclear respiratory factors 1 and 2 (NRF-1 and NRF-2), mitochondrial transcription factor A (TFAM), and NFE2L2.
The coordinated effects of AMPK, SIRT1, and SIRT3 are primarily mediated through PGC-1α, which is activated through phosphorylation by AMPK [242, 265] and deacetylation by SIRT1 [77, 242, 266–269]. SIRT3 also increases PGC-1α activity , possibly through cAMP response element binding protein (CREB) [271, 272], but the exact mechanism has not been elucidated. In addition to phosphorylating PGC-1α, activated AMPK also increases PGC-1α expression [260, 273–276]. Activation of β2-adrenergic receptors [277–280] and the adiponectin AdipoR1 receptor  also increase PGC-1α expression, independently of AMPK activation [278, 281]. PGC-1α activity is increased by oxidative stress [76, 77, 282–284], possibly through activation of AMPK [259, 260] or p38 mitogen-activated protein kinase (MAPK) [283, 284], or inhibition of glycogen synthase kinase 3β, which inhibits PGC-1α through phosphorylation [77, 283]. In contrast, insulin decreases PGC-1α activity through phosphorylation by PKB . Once activated, PGC-1α interacts with the PPAR family of nuclear receptors  and the FOXO family of transcription factors  to influence expression of a variety of bioenergetic and antioxidant proteins. PGC-1α most notably increases transcription of proteins involved in mitochondrial biogenesis and respiration [76, 242, 265, 267, 269, 274, 279, 282, 285, 288–293] but also increases transcription of antioxidant proteins including SOD1 , SOD2 [76, 282, 289, 292–294], catalase , GPx [76, 294], thioredoxins [282, 283, 292], TRXR [282, 292], Prx3 [282, 292], and Prx5 [282, 292], as well as the mitochondrial uncoupling proteins UCP2 [76, 265, 282, 288, 294], UCP3 [76, 265, 294], and ANT [76, 295].
PGC-1α coactivates all three known PPAR isoforms (PPARα, PPARδ, and PPARγ) . Although each isoform is expressed in a variety of tissues, PPARα is prominently expressed in the liver, PPARδ in skeletal muscle, the heart, and the pancreas, and PPARγ in adipose [286, 296]. PGC-1α was discovered and named based on its promotion of brown adipose differentiation through coactivation of PPARγ and subsequent induction of mitochondrial biogenesis and UCP1 expression . However, it is the PGC-1α coactivation of PPARα that is responsible for the upregulated transcription of many of the enzymes responsible for increased ketogenesis and fatty acid metabolism in response to a ketogenic diet . Consistent with the role of PGC-1α in inducing mitochondrial biogenesis, it also shifts skeletal muscle fiber composition towards type I [298, 299] and type IIa , which are more oxidative. AMPK also contributes to fiber type changes and is required for the transition of highly glycolytic, type IIb fibers to more oxidative, type IIa fibers . Although PGC-1α is primarily known for inducing transcription of nuclear DNA, it may also, in conjunction with SIRT1, induce expression of mtDNA .
PGC-1α is also influenced by p38 MAPK, which is well known for being involved in development  and adaptation  in skeletal muscle. PGC-1α is activated by p38 MAPK [283, 303] through phosphorylation , which prevents repression  by blocking interaction with the p160 myb binding protein . In addition, expression of PGC-1α is increased by p38 MAPK [305, 306], and the overlap in bioenergetic and antioxidant signaling is further indicated based on p38 MAPK activation by AMPK [307–309], oxidative stress [310–314], and β-adrenergic signaling [280, 315, 316].
Nutritional ketosis may facilitate PGC-1α activity through multiple mechanisms. Since PGC-1α is activated by AMPK and SIRT1, nutritional ketosis may initiate PGC-1α activity through these enzymes. As previously mentioned, catecholamines and adiponectin facilitate PGC-1α activity by promoting its expression, and insulin inhibits PGC-1α through downstream phosphorylation, all independent of AMPK. As previously discussed, a ketogenic diet may increase catecholamines and adiponectin and is well known to decrease insulin, indicating that nutritional ketosis may directly facilitate PGC-1α activity through these hormones. Supporting these potential mechanisms, a ketogenic or low-carbohydrate diet has increased expression, protein content, and activation of PGC-1α [149, 231, 317], as well as expression of its target PPARα [87, 148]. Furthermore, in skeletal muscle of mice following a ketogenic diet, the resulting increases in O2 consumption and expression of genes related to fat oxidation appear to be dependent on PGC-1α . Ketones likely contribute to this signaling as well based on the recent observation that the increased hepatic expression of PPARα targets induced by a ketogenic diet did not occur with a nonketogenic low-carbohydrate diet .
The FOXO family of transcription factors is highly conserved and promotes longevity and resistance to cellular stress. Although there are a variety of FOXO isoforms with varying tissue distribution [318–320], FOXO3a has been the most thoroughly studied in relation to energy sensing, mitochondrial function, and antioxidant defense. Similar to PGC-1α, FOXO3a is activated through phosphorylation by AMPK [321–323] and deacetylation by SIRT1 [324, 325] and SIRT3 [326–329], and its transcriptional activity is at least partly dependent on AMPK  and SIRT1 . In a variety of organisms, tissues, and cell types, FOXO3a increases mitochondrial biogenesis and expression of TFAM , but is more known for increasing expression of antioxidant and repair proteins, including SOD2 [287, 330, 331], catalase [287, 330, 332, 333], glutathione S-transferase (GST) , thioredoxins [287, 323], Prx3 [287, 334], Prx5 , and metallothioneins I and II , as well as UCP2 [287, 322] and the DNA repair enzyme growth arrest and DNA damage-inducible 45 (GADD45) [322, 324, 335, 336]. FOXO3a is also activated by oxidative stress [324, 331, 333], possibly in a SIRT1-dependent manner , and likely mediated through c-Jun N-terminal protein kinase (JNK), which allows FOXOs to translocate to the nucleus by promoting dissociation of 14-3-3 [337, 338]. Furthermore, FOXO3a and SIRT3 interact in mitochondria to induce mitochondrial gene expression in an AMPK-dependent manner . FOXO3a also induces expression of LKB1  and NAMPT , indicating a feed-forward cycle of activation with AMPK and sirtuins. Like PGC-1α, FOXO3a transcriptional activity is inhibited by insulin through PKB .
As with PGC-1α, nutritional ketosis may activate FOXO3a by increasing activity of AMPK and sirtuins or by decreasing insulin. Expression of FOXO3a is increased by fasting, caloric restriction, and BHB [103, 105], all of which are or can be components of a ketogenic diet. Furthermore, BHB treatment has extended lifespan in C. elegans in a manner dependent on FOXO3a , and a ketogenic diet (% energy: 89 fat, <1 carbohydrate, and 10 protein) has increased median lifespan and decreased tumors and age-associated losses of physical and cognitive performance, all in conjunction with increased hepatic content of FOXO3a .
9.3. NRF-1, NRF-2, and TFAM
NRF-1 and NRF-2 are transcription factors that increase expression of TFAM , which is required for full initiation of mtDNA transcription [343–345] and hence mitochondrial biogenesis. PGC-1α induces expression of NRF-1 and NRF-2 and facilitates TFAM expression by coactivating NRF-1 . Oxidative stress increases this signaling [346, 347] in conjunction with increased mitochondrial biogenesis . AMPK also contributes to mitochondrial biogenesis, but by inducing mitochondrial fission through phosphorylation of mitochondrial fission factor (MFF) , which is in addition to and independent of AMPK’s role in activating PGC-1α.
Nuclear factor erythroid-derived 2-like 2 (NFE2L2 or NRF2) is a transcription factor that has a prominent role in antioxidant signaling and also influences mitochondrial bioenergetics. The NFE2L2 abbreviation is used in this review to avoid confusion with nuclear respiratory factor 2, which despite being a different protein, has overlapping function with NFE2L2 and shares the same NRF2 abbreviation . Although the mechanisms of NFE2L2 signaling are not fully elucidated , oxidative stress has a clear role in interacting with cysteine residues of Kelch-like ECH-associated protein 1 (Keap1), which decreases proteasomal degradation of NFE2L2 and thereby allows entry of NFE2L2 into the nucleus to induce transcription [351–355]. Although the influence of PGC-1α on antioxidant enzyme expression is not dependent on NFE2L2 [76, 356], PGC-1α increases NFE2L2 expression , indicating that NFE2L2 activity is influenced by perturbations in both energy and redox homeostasis. NFE2L2 primarily increases expression of antioxidant enzymes, including SOD1 , SOD2 , catalase [358–361], GPx , NQO1 [354, 359–362], GCL [359–361], GST , GSR [359–361], and Prx1 , but also increases expression of proteins involved in mitochondrial biogenesis and bioenergetics including NRF-1, NRF-2, TFAM, cytochrome c oxidase, and citrate synthase .
In the previously described C. elegans experiments demonstrating mitohormesis, knockout of the NFE2L2 homologue SKN-1 attenuated the increases in antioxidant enzyme activity and lifespan , indicating that mitohormesis may, at least in part, be dependent on NFE2L2 signaling. Similarly, a ketogenic diet (Bio-Serv F3666) increased nuclear content of NFE2L2 and expression of its target NQO1 in the hippocampi of rats, all of which occurred after an initial increase in mtROS . This increase in NFE2L2 content appears to have mediated the subsequent decrease in mtROS to a level below baseline , thereby further indicating a likely role of NFE2L2 in the induction of mitohormesis during a ketogenic diet.
Additional evidence, although independent of mitohormesis, further supports the induction of NFE2L2 activity by nutritional ketosis. Succinate is a byproduct of ketolysis and is oxidized to fumarate by succinate dehydrogenase. Therefore, the increased presence of ketones and increased rate of ketolysis during nutritional ketosis are likely to increase fumarate, which can succinylate cysteine residues of proteins . In particular, fumarate can succinylate Keap1, thereby allowing NFE2L2 to enter the nucleus to induce transcription [364, 365]. In the retinas of rats injected with BHB, the nuclear content of NFE2L2 and the total homogenate content of SOD2 and GCL increased in conjunction with increased fumarate concentration . BHB injection also decreased retinal ROS production and degeneration following induction of ischemia, and this protection was dependent on NFE2L2 . These effects were observed at blood concentrations of BHB between 1 and 2 mM, which is consistent with nutritional ketosis.
10. Overlap between Bioenergetic and Antioxidant Signal Transduction
As described throughout the previous sections, there are many instances of codependencies and feed-forward loops in bioenergetic and antioxidant signal transduction, which supports the well-known potential for metabolic stimuli, such as diet or exercise, to have a profound physiological influence. Given the central role of mitochondria in oxidative phosphorylation and ROS production, the overlap between bioenergetic and antioxidant signaling is not surprising and is possibly an outcome of evolution favoring efficiency. PGC-1α is at the center of this overlapping and complex network of codependencies. The likely role of PGC-1α as a coactivator of FOXO3a indicates a possible dependence of FOXO3a transcriptional activity on PGC-1α , indicating FOXO3a as a central mediator as well. Furthermore, FOXO3a induces transcription of PGC-1α [287, 322, 367], and formation and antioxidant transcriptional activity of the PGC-1α-FOXO3a complex are partly dependent on interaction with SIRT1 . In muscle, expression of many of the bioenergetic and antioxidant proteins previously discussed is dependent on PGC-1α . Upstream, activation of PGC-1α is dependent on AMPK  and SIRT1 [242, 269] and partly dependent on SIRT3 . Furthermore, activation of SIRT1 is dependent on AMPK , which may also be the case for SIRT3. AMPK and PGC-1α are therefore two key factors, with critical supporting roles of the sirtuins, in the signal transduction linking bioenergetics to antioxidant defense. Further supporting the relevance of this linkage to nutritional ketosis, expression of AMPK, SIRT1, FOXO3a, and NFE2L2 is required for extension of lifespan in C. elegans by exogenous BHB , and expression of AMPK, p38 MAPK, and NFE2L2 is required for the extension of lifespan, also in C. elegans, by mitohormesis induced through inhibition of glucose metabolism . The induction of AMPK [259, 260], SIRT3 [263, 329], p38 MAPK [310–313], PGC-1α [76, 77, 260, 282, 283], FOXO3a [324, 331, 333], and NFE2L2 [358–360] activity by oxidative stress also makes this signaling highly relevant to mitohormesis [263, 282, 360], especially given that activation of these proteins has been shown to decrease mitochondrial or cellular ROS [76, 263, 282, 289, 323, 332, 334, 356, 359, 367]. Furthermore, mitochondrial biogenesis  and the activities of AMPK [259, 260], SIRT3 , p38 MAPK [311, 312], PGC-1α [76, 77, 260, 283], FOXO3a , and NFE2L2  are increased by H2O2, more specifically associating this signaling with mitohormesis. Given that AMPK and sirtuins are upstream of the majority of this signaling and that AMPK and sirtuin activities are stimulated by both bioenergetic and oxidative stressors, these stressors are likely the primary signals through which nutritional ketosis may induce the mitochondrial and antioxidant adaptations characteristic of mitohormesis (Figure 2).
11. Exercise as an Adjunct to Nutritional Ketosis
Although resting skeletal muscle is less metabolically active than the heart, kidneys, brain, or liver, it rivals even the brain in being the body’s most metabolically demanding tissue when considered relative to total tissue mass . Physical activity can greatly increase this demand, making exercise a practical and powerful way to induce bioenergetic adaptations.
In skeletal muscle, impaired mitochondrial function contributes to age-associated atrophy, impaired contraction, and insulin resistance . While exercise provides a direct stimulus for mitochondrial adaptation in muscle, with great potential to prevent or treat the aforementioned conditions, the global effects of exercise on bioenergetic homeostasis may lead to mitochondrial adaptations in other tissues as well. Based on this, exercise has the potential to influence any condition for which impairments in global energy metabolism or local mitochondrial function are a contributing factor, which is arguably the case for a majority of chronic diseases. Exercise is therefore an excellent adjunct to nutritional ketosis because it facilitates β-oxidation and ketogenesis by increasing energy demand and depleting glycogen storage, which is likely to augment the signaling induced by nutritional ketosis.
In skeletal muscle, oxidative capacity and mitochondrial content are related to fiber type. Compared to type II fibers, type I fibers have larger mitochondria  with greater oxidative enzyme content . While fiber type is plastic, particularly in response to endurance exercise, transformation from oxidative, slow-twitch fibers (type I) to glycolytic, fast-twitch fibers (type II) is unlikely to occur [372, 373]. Type II fibers, however, can shift in humans from highly glycolytic (type IIx) to more oxidative (type IIa) . Compared to type IIx fibers, type IIa fibers have greater citrate synthase activity, indicating greater mitochondrial content . The relevance of oxidative capacity and fiber type to oxidative stress has been demonstrated by greater mitochondrial respiration with less H2O2 production in permeabilized fibers from rat muscle consisting primarily of type I or IIa fibers versus type IIb fibers . Although muscle fiber-type transformation has been well characterized in response to exercise, this appears to not be the case for ketogenic diets. However, in rats, β-hydroxyacyl-CoA dehydrogenase (β-HAD) has been shown to increase most prominently in glycolytic, type IIb fibers following 4 weeks of a ketogenic diet (% energy: 70 fat, 6 carbohydrate, and 24 protein) , suggesting transition of these fibers towards type IIa fibers and, in turn, indicating potential for nutritional ketosis to promote a more oxidative muscle fiber composition.
Bioenergetic and oxidative stressors may be largely responsible for inducing many of the beneficial adaptations to exercise, and for this reason, exercise research provides much of the basis for mitohormesis [4–6]. As previously discussed, an increase in fat oxidation appears to be a prerequisite for increasing mtROS and, in turn, inducing mitohormesis. Given that ketogenic diets prominently increase fat oxidation during submaximal exercise [8, 88, 214–216, 218, 219, 376–381], the combination of the two interventions may induce mitohormetic adaptations to a greater extent. Furthermore, much of the signaling that is relevant to mitohormesis, and likely induced by nutritional ketosis, is also induced by exercise, further suggesting the possibility of an additive or even synergistic effect. Demonstrating this, exercise or muscle contraction increases activity, activation, or expression of AMPK [209–211, 275, 284, 382–386], SIRT1 [384–389], SIRT3 [272, 390, 391], NFE2L2 [358, 360, 392], p38 MAPK [284, 305, 313–315, 393–395], PGC-1α [275–279, 284, 305, 314, 385–389, 396–400], NRF-1 , and TFAM [358, 388, 389]. Exercise also increases expression or activity of antioxidant enzymes [313, 358, 360, 396, 397, 401], uncoupling proteins , and bioenergetic proteins involved in oxidative phosphorylation [396, 397, 400] and the citric acid cycle , all of which appear to be at least partly mediated by ROS-induced activity of p38 MAPK [284, 310, 313, 314], PGC-1α [284, 310, 397, 401], TFAM [310, 314, 358, 397], NRF-1 [310, 358, 397], NRF-2 [358, 360], and NFE2L2 .
In addition to increased mitochondrial demand and mtROS production, there are several other commonalities in the mechanisms through which exercise and nutritional ketosis induce adaptive signaling. Exercise-induced activation of AMPK is greater when the exercise is performed in a glycogen depleted state [209–211, 382, 383], and exercise-induced activation of p38 MAPK  and PGC-1α [277–279] occurs at least partly through β-adrenergic signaling. Although changes in NAD+ and NADH are difficult to measure and are complicated by conflicting results, exercise is also likely to increase sirtuin activation by increasing the NAD+ to NADH ratio .
In controlled studies on exercise-trained humans and animals, ketogenic diets have been shown to increase fat oxidation [8, 167] and expression or activity of carnitine palmitoyltransferase [167, 168] and β-HAD [168, 172], demonstrating that nutritional ketosis induces adaptation beyond exercise. Similarly, in a study comparing the independent and combined effects of exercise and a ketogenic diet on rats, the combination resulted in greater β-HAD and citrate synthase activities in skeletal muscle and higher maximal O2 consumption than either intervention alone, further indicating the potential for exercise to magnify adaptations induced by nutritional ketosis .
Among the chronic and degenerative diseases in which impaired mitochondrial function is a contributing factor, many respond favorably to lifestyle interventions focused on diet and exercise. The therapeutic potential of nutritional ketosis stands out in this regard. For example, in just the first 10 weeks of an ongoing clinical trial with hundreds of type 2 diabetics following a ketogenic diet, glycated hemoglobin (HbA1c) decreased to below the diagnostic threshold in more than a third of patients, and prescription medication was reduced or eliminated for more than half of patients . Convincing arguments for a ketogenic diet to be the default treatment for diabetes are a decade old  and have continued to gain support since then . Similar arguments are developing for obesity [10, 11], neurodegenerative diseases [19, 20, 27–30], cardiovascular disease [15–17], cancer [18–26], and even aging [31, 32]. Although the mechanisms through which a ketogenic diet may improve these conditions expand beyond mitochondrial function, the great extent to which nutritional ketosis increases reliance on mitochondrial metabolism strongly suggests that mitochondrial adaptation is a central factor.
The clinical relevance of nutritional ketosis to mitochondrial function is further indicated by promotion of ketogenic diets for treatment of mitochondrial disorders [19, 20, 26, 30, 247, 403]. The most prominent example is the study of mitochondrial adaptations as a mechanism for the well-known antiseizure effect of ketogenic diets [19, 29, 33, 162, 247, 403–405]. As previously discussed, the dramatic shift in energy metabolism and subsequent increase in circulating ketones induced by a ketogenic diet can enhance mitochondrial function and endogenous antioxidant defense. The primary mechanism behind these adaptations appears to be the increased demand for fat oxidation resulting from carbohydrate restriction. However, ketones themselves have important metabolic and signaling effects that enhance mitochondrial function and endogenous antioxidant defense, implying that a well-formulated ketogenic diet should have greater benefit than a nonketogenic low-carbohydrate diet. Regardless of the mechanism(s), the potential outcomes imply protection against chronic disease through improved mitochondrial function and, in turn, decreased potential for oxidative stress and subsequent pathology. However, further research is needed to better understand how nutritional ketosis influences mitochondrial function across different tissues and how these influences relate to human disease. Future research should also focus on further differentiation of the effects of carbohydrate restriction from the direct effects of ketones.
Conflicts of Interest
The authors declare that they have no conflicts of interest.
- D. Sebastian, M. Palacin, and A. Zorzano, “Mitochondrial dynamics: coupling mitochondrial fitness with healthy aging,” Trends in Molecular Medicine, vol. 23, no. 3, pp. 201–215, 2017.
- D. F. Dai, Y. A. Chiao, D. J. Marcinek, H. H. Szeto, and P. S. Rabinovitch, “Mitochondrial oxidative stress in aging and healthspan,” Longevity & Healthspan, vol. 3, no. 1, p. 6, 2014.
- R. B. Hamanaka and N. S. Chandel, “Mitochondrial reactive oxygen species regulate cellular signaling and dictate biological outcomes,” Trends in Biochemical Sciences, vol. 35, no. 9, pp. 505–513, 2010.
- P. C. Tapia, “Sublethal mitochondrial stress with an attendant stoichiometric augmentation of reactive oxygen species may precipitate many of the beneficial alterations in cellular physiology produced by caloric restriction, intermittent fasting, exercise and dietary phytonutrients: “Mitohormesis” for health and vitality,” Medical Hypotheses, vol. 66, no. 4, pp. 832–843, 2006.
- M. Ristow and K. Zarse, “How increased oxidative stress promotes longevity and metabolic health: the concept of mitochondrial hormesis (mitohormesis),” Experimental Gerontology, vol. 45, no. 6, pp. 410–418, 2010.
- M. Ristow and K. Schmeisser, “Mitohormesis: promoting health and lifespan by increased levels of Reactive Oxygen Species (ROS),” Dose Response, vol. 12, no. 2, pp. 288–341, 2014.
- J. S. Volek, M. L. Fernandez, R. D. Feinman, and S. D. Phinney, “Dietary carbohydrate restriction induces a unique metabolic state positively affecting atherogenic dyslipidemia, fatty acid partitioning, and metabolic syndrome,” Progress in Lipid Research, vol. 47, no. 5, pp. 307–318, 2008.
- J. S. Volek, D. J. Freidenreich, C. Saenz et al., “Metabolic characteristics of keto-adapted ultra-endurance runners,” Metabolism, vol. 65, no. 3, pp. 100–110, 2016.
- J. Milder and M. Patel, “Modulation of oxidative stress and mitochondrial function by the ketogenic diet,” Epilepsy Research, vol. 100, no. 3, pp. 295–303, 2012.
- A. Paoli, “Ketogenic diet for obesity: friend or foe?” International Journal of Environmental Research and Public Health, vol. 11, no. 2, pp. 2092–2107, 2014.
- N. B. Bueno, I. S. de Melo, S. L. de Oliveira, and T. da Rocha Ataide, “Very-low-carbohydrate ketogenic diet v. low-fat diet for long-term weight loss: a meta-analysis of randomised controlled trials,” British Journal of Nutrition, vol. 110, no. 7, pp. 1178–1187, 2013.
- A. L. McKenzie, S. J. Hallberg, B. C. Creighton et al., “A novel intervention including individualized nutritional recommendations reduces hemoglobin A1c level, medication use, and weight in type 2 diabetes,” JMIR Diabetes, vol. 2, no. 1, p. 14, 2017.
- R. D. Feinman and J. S. Volek, “Carbohydrate restriction as the default treatment for type 2 diabetes and metabolic syndrome,” Scandinavian Cardiovascular Journal, vol. 42, no. 4, pp. 256–263, 2008.
- R. D. Feinman, W. K. Pogozelski, A. Astrup et al., “Dietary carbohydrate restriction as the first approach in diabetes management: critical review and evidence base,” Nutrition, vol. 31, no. 1, pp. 1–13, 2015.
- A. Paoli, A. Rubini, J. S. Volek, and K. A. Grimaldi, “Beyond weight loss: a review of the therapeutic uses of very-low-carbohydrate (ketogenic) diets,” European Journal of Clinical Nutrition, vol. 67, no. 8, pp. 789–796, 2013.
- D. G. Cotter, R. C. Schugar, and P. A. Crawford, “Ketone body metabolism and cardiovascular disease,” American Journal of Physiology-Heart and Circulatory Physiology, vol. 304, no. 8, pp. H1060–H1076, 2013.
- C. Kosinski and F. R. Jornayvaz, “Effects of ketogenic diets on cardiovascular risk factors: evidence from animal and human studies,” Nutrients, vol. 9, no. 5, 2017.
- P. N. Hyde, M. B. Lustberg, V. J. Miller, R. A. LaFountain, and J. S. Volek, “Pleiotropic effects of nutritional ketosis: conceptual framework for ketoadaptation as a breast cancer therapy,” Cancer Treatment and Research Communications, vol. 12, no. 2017, pp. 32–39, 2017.
- A. F. Branco, A. Ferreira, R. F. Simoes et al., “Ketogenic diets: from cancer to mitochondrial diseases and beyond,” European Journal of Clinical Investigation, vol. 46, no. 3, pp. 285–298, 2016.
- C. E. Stafstrom and J. M. Rho, “The ketogenic diet as a treatment paradigm for diverse neurological disorders,” Frontiers in Pharmacology, vol. 3, p. 59, 2012.
- C. L. Oliveira, S. Mattingly, R. Schirrmacher, M. B. Sawyer, E. J. Fine, and C. M. Prado, “A Nutritional perspective of ketogenic diet in cancer: a narrative review,” Journal of the Academy of Nutrition and Dietetics, 2017, in press.
- B. A. Simone, C. E. Champ, A. L. Rosenberg et al., “Selectively starving cancer cells through dietary manipulation: methods and clinical implications,” Future Oncology, vol. 9, no. 7, pp. 959–976, 2013.
- T. N. Seyfried, J. Marsh, L. M. Shelton, L. C. Huysentruyt, and P. Mukherjee, “Is the restricted ketogenic diet a viable alternative to the standard of care for managing malignant brain cancer?” Epilepsy Research, vol. 100, no. 3, pp. 310–326, 2012.
- E. J. Fine and R. D. Feinman, “Insulin, carbohydrate restriction, metabolic syndrome and cancer,” Expert Review of Endocrinology & Metabolism, vol. 10, no. 1, pp. 15–24, 2014.
- B. G. Allen, S. K. Bhatia, C. M. Anderson et al., “Ketogenic diets as an adjuvant cancer therapy: history and potential mechanism,” Redox Biology, vol. 2, pp. 963–970, 2014.
- S. Vidali, S. Aminzadeh, B. Lambert et al., “Mitochondria: the ketogenic diet–a metabolism-based therapy,” International Journal of Biochemistry & Cell Biology, vol. 63, pp. 55–59, 2015.
- A. L. Hartman, “Neuroprotection in metabolism-based therapy,” Epilepsy Research, vol. 100, no. 3, pp. 286–294, 2012.
- M. Gasior, M. A. Rogawski, and A. L. Hartman, “Neuroprotective and disease-modifying effects of the ketogenic diet,” Behavioural Pharmacology, vol. 17, no. 5-6, pp. 431–439, 2006.
- M. Maalouf, J. M. Rho, and M. P. Mattson, “The neuroprotective properties of calorie restriction, the ketogenic diet, and ketone bodies,” Brain Research Reviews, vol. 59, no. 2, pp. 293–315, 2009.
- A. Paoli, A. Bianco, E. Damiani, and G. Bosco, “Ketogenic diet in neuromuscular and neurodegenerative diseases,” BioMed Research International, vol. 2014, Article ID 474296, 10 pages, 2014.
- C. L. Moreno and C. V. Mobbs, “Epigenetic mechanisms underlying lifespan and age-related effects of dietary restriction and the ketogenic diet,” Molecular and Cellular Endocrinology, vol. 455, pp. 33–40, 2017.
- C. Edwards, N. Copes, and P. C. Bradshaw, “D-β-hydroxybutyrate: an anti-aging ketone body,” Oncotarget, vol. 6, no. 6, pp. 3477-3478, 2015.
- R. L. Veech, P. C. Bradshaw, K. Clarke, W. Curtis, R. Pawlosky, and M. T. King, “Ketone bodies mimic the life span extending properties of caloric restriction,” IUBMB Life, vol. 69, no. 5, pp. 305–314, 2017.
- J. S. Volek and S. D. Phinney, The Art and Science of Low Carbohydrate Living, Beyond Obesity, LLC, Miami, FL, USA, 2011.
- J. S. Volek and S. D. Phinney, The Art and Science of Low Carbohydrate Performance, Beyond Obesity, LLC, Miami, FL, USA, 2012.
- M. N. Roberts, M. A. Wallace, A. A. Tomilov et al., “A ketogenic diet extends longevity and healthspan in adult mice,” Cell Metabolism, vol. 26, no. 3, pp. 539 e535–546 e535, 2017.
- J. C. Newman, A. J. Covarrubias, M. Zhao et al., “Ketogenic diet reduces midlife mortality and improves memory in aging mice,” Cell Metabolism, vol. 26, no. 3, pp. 547 e548–557 e548, 2017.
- A. W. C. Yuen, I. A. Walcutt, and J. W. Sander, “An acidosis-sparing ketogenic (ASK) diet to improve efficacy and reduce adverse effects in the treatment of refractory epilepsy,” Epilepsy & Behavior, vol. 74, pp. 15–21, 2017.
- P. A. Crawford, J. R. Crowley, N. Sambandam et al., “Regulation of myocardial ketone body metabolism by the gut microbiota during nutrient deprivation,” Proceedings of the National Academy of Sciences, vol. 106, no. 27, pp. 11276–11281, 2009.
- G. den Besten, K. van Eunen, A. K. Groen, K. Venema, D. J. Reijngoud, and B. M. Bakker, “The role of short-chain fatty acids in the interplay between diet, gut microbiota, and host energy metabolism,” Journal of Lipid Research, vol. 54, no. 9, pp. 2325–2340, 2013.
- B. Halliwell and J. M. C. Gutteridge, Free Radicals in Biology and Medicine, Oxford University Press, Oxford, UK, 5th edition, 2015.
- T. H. Zastawny, M. Dabrowska, T. Jaskolski et al., “Comparison of oxidative base damage in mitochondrial and nuclear DNA,” Free Radical Biology and Medicine, vol. 24, no. 5, pp. 722–725, 1998.
- F. M. Yakes and B. Van Houten, “Mitochondrial DNA damage is more extensive and persists longer than nuclear DNA damage in human cells following oxidative stress,” Proceedings of the National Academy of Sciences, vol. 94, no. 2, pp. 514–519, 1997.
- S. P. LeDoux, G. L. Wilson, E. J. Beecham, T. Stevnsner, K. Wassermann, and V. A. Bohr, “Repair of mitochondrial DNA after various types of DNA damage in Chinese hamster ovary cells,” Carcinogenesis, vol. 13, no. 11, pp. 1967–1973, 1992.
- S. W. Ballinger, B. Van Houten, G. F. Jin, C. A. Conklin, and B. F. Godley, “Hydrogen peroxide causes significant mitochondrial DNA damage in human RPE cells,” Experimental Eye Research, vol. 68, no. 6, pp. 765–772, 1999.
- I. N. Shokolenko, G. L. Wilson, and M. F. Alexeyev, “Aging: a mitochondrial DNA perspective, critical analysis and an update,” World Journal of Experimental Medicine, vol. 4, no. 4, pp. 46–57, 2014.
- K. Szczepanowska and A. Trifunovic, “Origins of mtDNA mutations in ageing,” Essays In Biochemistry, vol. 61, no. 3, pp. 325–337, 2017.
- M. Pinto and C. T. Moraes, “Mechanisms linking mtDNA damage and aging,” Free Radical Biology and Medicine, vol. 85, pp. 250–258, 2015.
- M. F. Alexeyev, “Is there more to aging than mitochondrial DNA and reactive oxygen species?” FEBS Journal, vol. 276, no. 20, pp. 5768–5787, 2009.
- T. E. S. Kauppila, J. H. K. Kauppila, and N. G. Larsson, “Mammalian mitochondria and aging: an update,” Cell Metabolism, vol. 25, no. 1, pp. 57–71, 2017.
- M. Muftuoglu, M. P. Mori, and N. C. de Souza-Pinto, “Formation and repair of oxidative damage in the mitochondrial DNA,” Mitochondrion, vol. 17, pp. 164–181, 2014.
- G. Barja and A. Herrero, “Oxidative damage to mitochondrial DNA is inversely related to maximum life span in the heart and brain of mammals,” FASEB Journal, vol. 14, no. 2, pp. 312–318, 2000.
- M. P. Murphy, “How mitochondria produce reactive oxygen species,” Biochemical Journal, vol. 417, no. 1, pp. 1–13, 2009.
- M. Rigoulet, E. D. Yoboue, and A. Devin, “Mitochondrial ROS generation and its regulation: mechanisms involved in H2O2 signaling,” Antioxidants & Redox Signaling, vol. 14, no. 3, pp. 459–468, 2011.
- V. Adam-Vizi and C. Chinopoulos, “Bioenergetics and the formation of mitochondrial reactive oxygen species,” Trends in Pharmacological Sciences, vol. 27, no. 12, pp. 639–645, 2006.
- A. Boveris and B. Chance, “The mitochondrial generation of hydrogen peroxide. General properties and effect of hyperbaric oxygen,” Biochemical Journal, vol. 134, no. 3, pp. 707–716, 1973.
- S. S. Korshunov, V. P. Skulachev, and A. A. Starkov, “High protonic potential actuates a mechanism of production of reactive oxygen species in mitochondria,” FEBS Letters, vol. 416, no. 1, pp. 15–18, 1997.
- S. Miwa, J. St-Pierre, L. Partridge, and M. D. Brand, “Superoxide and hydrogen peroxide production by Drosophila mitochondria,” Free Radical Biology and Medicine, vol. 35, no. 8, pp. 938–948, 2003.
- A. J. Lambert and M. D. Brand, “Superoxide production by NADH:ubiquinone oxidoreductase (complex I) depends on the pH gradient across the mitochondrial inner membrane,” Biochemical Journal, vol. 382, no. 2, pp. 511–517, 2004.
- T. V. Votyakova and I. J. Reynolds, “DeltaPsi(m)-Dependent and -independent production of reactive oxygen species by rat brain mitochondria,” Journal of Neurochemistry, vol. 79, no. 2, pp. 266–277, 2001.
- L. Tretter, K. Takacs, V. Hegedus, and V. Adam-Vizi, “Characteristics of α-glycerophosphate-evoked H2O2 generation in brain mitochondria,” Journal of Neurochemistry, vol. 100, no. 3, pp. 650–663, 2007.
- Y. Liu, G. Fiskum, and D. Schubert, “Generation of reactive oxygen species by the mitochondrial electron transport chain,” Journal of Neurochemistry, vol. 80, no. 5, pp. 780–787, 2002.
- M. D. Brand, “Mitochondrial generation of superoxide and hydrogen peroxide as the source of mitochondrial redox signaling,” Free Radical Biology and Medicine, vol. 100, pp. 14–31, 2016.
- D. Han, E. Williams, and E. Cadenas, “Mitochondrial respiratory chain-dependent generation of superoxide anion and its release into the intermembrane space,” Biochemical Journal, vol. 353, no. 2, pp. 411–416, 2001.
- D. Han, F. Antunes, R. Canali, D. Rettori, and E. Cadenas, “Voltage-dependent anion channels control the release of the superoxide anion from mitochondria to cytosol,” Journal of Biological Chemistry, vol. 278, no. 8, pp. 5557–5563, 2003.
- A. G. Cox, C. C. Winterbourn, and M. B. Hampton, “Mitochondrial peroxiredoxin involvement in antioxidant defence and redox signalling,” Biochemical Journal, vol. 425, no. 2, pp. 313–325, 2009.
- G. P. Bienert, J. K. Schjoerring, and T. P. Jahn, “Membrane transport of hydrogen peroxide,” Biochimica et Biophysica Acta, vol. 1758, no. 8, pp. 994–1003, 2006.
- J. Rydstrom, “Mitochondrial NADPH, transhydrogenase and disease,” Biochimica et Biophysica Acta, vol. 1757, no. 5-6, pp. 721–726, 2006.
- M. V. Kumari, M. Hiramatsu, and M. Ebadi, “Free radical scavenging actions of metallothionein isoforms I and II,” Free Radical Research, vol. 29, no. 2, pp. 93–101, 1998.
- P. J. Thornalley and M. Vasak, “Possible role for metallothionein in protection against radiation-induced oxidative stress. Kinetics and mechanism of its reaction with superoxide and hydroxyl radicals,” Biochimica et Biophysica Acta, vol. 827, no. 1, pp. 36–44, 1985.
- S. P. Mezyk, “Determination of the rate constant for the reaction of hydroxyl and oxide radicals with cysteine in aqueous solution,” Radiation Research, vol. 145, no. 1, pp. 102–106, 1996.
- T. J. Schulz, K. Zarse, A. Voigt, N. Urban, M. Birringer, and M. Ristow, “Glucose restriction extends Caenorhabditis elegans life span by inducing mitochondrial respiration and increasing oxidative stress,” Cell Metabolism, vol. 6, no. 4, pp. 280–293, 2007.
- K. Zarse, S. Schmeisser, M. Groth et al., “Impaired insulin/IGF1 signaling extends life span by promoting mitochondrial L-proline catabolism to induce a transient ROS signal,” Cell Metabolism, vol. 15, no. 4, pp. 451–465, 2012.
- H. M. Cocheme and M. P. Murphy, “Complex I is the major site of mitochondrial superoxide production by paraquat,” Journal of Biological Chemistry, vol. 283, no. 4, pp. 1786–1798, 2008.
- P. R. Castello, D. A. Drechsel, and M. Patel, “Mitochondria are a major source of paraquat-induced reactive oxygen species production in the brain,” Journal of Biological Chemistry, vol. 282, no. 19, pp. 14186–14193, 2007.
- J. St-Pierre, S. Drori, M. Uldry et al., “Suppression of reactive oxygen species and neurodegeneration by the PGC-1 transcriptional coactivators,” Cell, vol. 127, no. 2, pp. 397–408, 2006.
- R. M. Anderson, J. L. Barger, M. G. Edwards et al., “Dynamic regulation of PGC-1α localization and turnover implicates mitochondrial adaptation in calorie restriction and the stress response,” Aging Cell, vol. 7, no. 1, pp. 101–111, 2008.
- P. H. Bisschop, J. de Metz, M. T. Ackermans et al., “Dietary fat content alters insulin-mediated glucose metabolism in healthy men,” American Journal of Clinical Nutrition, vol. 73, no. 3, pp. 554–559, 2001.
- B. Ukropcova, O. Sereda, L. de Jonge et al., “Family history of diabetes links impaired substrate switching and reduced mitochondrial content in skeletal muscle,” Diabetes, vol. 56, no. 3, pp. 720–727, 2007.
- J. O. Hill, J. C. Peters, G. W. Reed, D. G. Schlundt, T. Sharp, and H. L. Greene, “Nutrient balance in humans: effects of diet composition,” American Journal of Clinical Nutrition, vol. 54, no. 1, pp. 10–17, 1991.
- P. Schrauwen, W. D. van Marken Lichtenbelt, W. H. Saris, and K. R. Westerterp, “Changes in fat oxidation in response to a high-fat diet,” American Journal of Clinical Nutrition, vol. 66, no. 2, pp. 276–282, 1997.
- J. M. Schwarz, R. A. Neese, S. Turner, D. Dare, and M. K. Hellerstein, “Short-term alterations in carbohydrate energy intake in humans. Striking effects on hepatic glucose production, de novo lipogenesis, lipolysis, and whole-body fuel selection,” Journal of Clinical Investigation, vol. 96, no. 6, pp. 2735–2743, 1995.
- B. Mittendorfer and L. S. Sidossis, “Mechanism for the increase in plasma triacylglycerol concentrations after consumption of short-term, high-carbohydrate diets,” American Journal of Clinical Nutrition, vol. 73, no. 5, pp. 892–899, 2001.
- G. Allick, P. H. Bisschop, M. T. Ackermans et al., “A low-carbohydrate/high-fat diet improves glucoregulation in type 2 diabetes mellitus by reducing postabsorptive glycogenolysis,” Journal of Clinical Endocrinology & Metabolism, vol. 89, no. 12, pp. 6193–6197, 2004.
- P. Schrauwen, W. D. van Marken Lichtenbelt, W. H. Saris, and K. R. Westerterp, “Role of glycogen-lowering exercise in the change of fat oxidation in response to a high-fat diet,” American Journal of Physiology-Endocrinology and Metabolism, vol. 273, no. 3, pp. E623–E629, 1997.
- P. Schrauwen, W. D. Lichtenbelt, W. H. Saris, and K. R. Westerterp, “Fat balance in obese subjects: role of glycogen stores,” American Journal of Physiology-Endocrinology and Metabolism, vol. 274, no. 6, pp. E1027–E1033, 1998.
- P. Garcia-Roves, J. M. Huss, D. H. Han et al., “Raising plasma fatty acid concentration induces increased biogenesis of mitochondria in skeletal muscle,” Proceedings of the National Academy of Sciences, vol. 104, no. 25, pp. 10709–10713, 2007.
- M. Vogt, A. Puntschart, H. Howald et al., “Effects of dietary fat on muscle substrates, metabolism, and performance in athletes,” Medicine & Science in Sports & Exercise, vol. 35, no. 6, pp. 952–960, 2003.
- A. Prince, Y. Zhang, C. Croniger, and M. Puchowicz, “Oxidative metabolism: glucose versus ketones,” Advances in Experimental Medicine and Biology, vol. 789, pp. 323–328, 2013.
- F. Scialo, D. J. Fernandez-Ayala, and A. Sanz, “Role of mitochondrial reverse electron transport in ROS signaling: potential roles in health and disease,” Frontiers in Physiology, vol. 8, p. 428, 2017.
- D. Speijer, “Oxygen radicals shaping evolution: why fatty acid catabolism leads to peroxisomes while neurons do without it: FADH(2)/NADH flux ratios determining mitochondrial radical formation were crucial for the eukaryotic invention of peroxisomes and catabolic tissue differentiation,” Bioessays, vol. 33, no. 2, pp. 88–94, 2011.
- D. Speijer, “Being right on Q: shaping eukaryotic evolution,” Biochemical Journal, vol. 473, no. 22, pp. 4103–4127, 2016.
- J. St-Pierre, J. A. Buckingham, S. J. Roebuck, and M. D. Brand, “Topology of superoxide production from different sites in the mitochondrial electron transport chain,” Journal of Biological Chemistry, vol. 277, no. 47, pp. 44784–44790, 2002.
- E. J. Anderson, H. Yamazaki, and P. D. Neufer, “Induction of endogenous uncoupling protein 3 suppresses mitochondrial oxidant emission during fatty acid-supported respiration,” Journal of Biological Chemistry, vol. 282, no. 43, pp. 31257–31266, 2007.
- C. Edwards, J. Canfield, N. Copes, M. Rehan, D. Lipps, and P. C. Bradshaw, “D-beta-hydroxybutyrate extends lifespan in C. elegans,” Aging, vol. 6, no. 8, pp. 621–644, 2014.
- J. B. Milder, L. P. Liang, and M. Patel, “Acute oxidative stress and systemic Nrf2 activation by the ketogenic diet,” Neurobiology of Disease, vol. 40, no. 1, pp. 238–244, 2010.
- S. G. Jarrett, J. B. Milder, L. P. Liang, and M. Patel, “The ketogenic diet increases mitochondrial glutathione levels,” Journal of Neurochemistry, vol. 106, no. 3, pp. 1044–1051, 2008.
- D. R. Ziegler, L. C. Ribeiro, M. Hagenn et al., “Ketogenic diet increases glutathione peroxidase activity in rat hippocampus,” Neurochemical Research, vol. 28, no. 12, pp. 1793–1797, 2003.
- T. Greco, T. C. Glenn, D. A. Hovda, and M. L. Prins, “Ketogenic diet decreases oxidative stress and improves mitochondrial respiratory complex activity,” Journal of Cerebral Blood Flow & Metabolism, vol. 36, no. 9, pp. 1603–1613, 2016.
- J. C. Newman and E. Verdin, “Ketone bodies as signaling metabolites,” Trends in Endocrinology & Metabolism, vol. 25, no. 1, pp. 42–52, 2014.
- J. C. Newman and E. Verdin, “β-hydroxybutyrate: much more than a metabolite,” Diabetes Research and Clinical Practice, vol. 106, no. 2, pp. 173–181, 2014.
- P. Rojas-Morales, E. Tapia, and J. Pedraza-Chaverri, “β-hydroxybutyrate: a signaling metabolite in starvation response?” Cell Signal, vol. 28, no. 8, pp. 917–923, 2016.
- T. Shimazu, M. D. Hirschey, J. Newman et al., “Suppression of oxidative stress by β-hydroxybutyrate, an endogenous histone deacetylase inhibitor,” Science, vol. 339, no. 6116, pp. 211–214, 2013.
- T. Wei, W. Tian, F. Liu, and G. Xie, “Protective effects of exogenous β-hydroxybutyrate on paraquat toxicity in rat kidney,” Biochemical and Biophysical Research Communications, vol. 447, no. 4, pp. 666–671, 2014.
- M. Nagao, R. Toh, Y. Irino et al., “β-Hydroxybutyrate elevation as a compensatory response against oxidative stress in cardiomyocytes,” Biochemical and Biophysical Research Communications, vol. 475, no. 4, pp. 322–328, 2016.
- K. Tanegashima, Y. Sato-Miyata, M. Funakoshi, Y. Nishito, T. Aigaki, and T. Hara, “Epigenetic regulation of the glucose transporter gene Slc2a1 by β-hydroxybutyrate underlies preferential glucose supply to the brain of fasted mice,” Genes Cells, vol. 22, no. 1, pp. 71–83, 2017.
- T. A. Simeone, K. A. Simeone, and J. M. Rho, “Ketone bodies as anti-seizure agents,” Neurochemical Research, vol. 42, no. 7, pp. 2011–2018, 2017.
- M. L. Haces, K. Hernandez-Fonseca, O. N. Medina-Campos, T. Montiel, J. Pedraza-Chaverri, and L. Massieu, “Antioxidant capacity contributes to protection of ketone bodies against oxidative damage induced during hypoglycemic conditions,” Experimental Neurology, vol. 211, no. 1, pp. 85–96, 2008.
- M. Maalouf, P. G. Sullivan, L. Davis, D. Y. Kim, and J. M. Rho, “Ketones inhibit mitochondrial production of reactive oxygen species production following glutamate excitotoxicity by increasing NADH oxidation,” Neuroscience, vol. 145, no. 1, pp. 256–264, 2007.
- J. E. Squires, J. Sun, J. L. Caffrey, D. Yoshishige, and R. T. Mallet, “Acetoacetate augments β-adrenergic inotropism of stunned myocardium by an antioxidant mechanism,” American Journal of Physiology-Heart and Circulatory Physiology, vol. 284, no. 4, pp. H1340–H1347, 2003.
- D. Y. Kim, J. Vallejo, and J. M. Rho, “Ketones prevent synaptic dysfunction induced by mitochondrial respiratory complex inhibitors,” Journal of Neurochemistry, vol. 114, no. 1, pp. 130–141, 2010.
- K. Tieu, C. Perier, C. Caspersen et al., “D-β-hydroxybutyrate rescues mitochondrial respiration and mitigates features of Parkinson disease,” Journal of Clinical Investigation, vol. 112, no. 6, pp. 892–901, 2003.
- S. Frey, G. Geffroy, V. Desquiret-Dumas et al., “The addition of ketone bodies alleviates mitochondrial dysfunction by restoring complex I assembly in a MELAS cellular model,” Biochimica et Biophysica Acta, vol. 1863, no. 1, pp. 284–291, 2017.
- D. B. Zorov, M. Juhaszova, and S. J. Sollott, “Mitochondrial reactive oxygen species (ROS) and ROS-induced ROS release,” Physiological Reviews, vol. 94, no. 3, pp. 909–950, 2014.
- D. Y. Kim, L. M. Davis, P. G. Sullivan et al., “Ketone bodies are protective against oxidative stress in neocortical neurons,” Journal of Neurochemistry, vol. 101, no. 5, pp. 1316–1326, 2007.
- D. Y. Kim, K. A. Simeone, T. A. Simeone et al., “Ketone bodies mediate antiseizure effects through mitochondrial permeability transition,” Annals of Neurology, vol. 78, no. 1, pp. 77–87, 2015.
- D. Y. Kim, M. G. Abdelwahab, S. H. Lee et al., “Ketones prevent oxidative impairment of hippocampal synaptic integrity through KATP channels,” PLoS One, vol. 10, no. 4, Article ID e0119316, 2015.
- J. A. Crestanello, N. M. Doliba, A. M. Babsky et al., “Opening of potassium channels protects mitochondrial function from calcium overload,” Journal of Surgical Research, vol. 94, no. 2, pp. 116–123, 2000.
- E. L. Holmuhamedov, S. Jovanovic, P. P. Dzeja, A. Jovanovic, and A. Terzic, “Mitochondrial ATP-sensitive K+ channels modulate cardiac mitochondrial function,” American Journal of Physiology-Heart and Circulatory Physiology, vol. 275, no. 5, pp. H1567–H1576, 1998.
- M. Grabacka, M. Pierzchalska, M. Dean, and K. Reiss, “Regulation of ketone body metabolism and the role of PPARα,” International Journal of Molecular Sciences, vol. 17, no. 12, 2016.
- Z. Xie, D. Zhang, D. Chung et al., “Metabolic regulation of gene expression by histone lysine β-hydroxybutyrylation,” Molecular Cell, vol. 62, no. 2, pp. 194–206, 2016.
- M. D. Brand, C. Affourtit, T. C. Esteves et al., “Mitochondrial superoxide: production, biological effects, and activation of uncoupling proteins,” Free Radical Biology and Medicine, vol. 37, no. 6, pp. 755–767, 2004.
- K. S. Echtay, “Mitochondrial uncoupling proteins–what is their physiological role?” Free Radical Biology and Medicine, vol. 43, no. 10, pp. 1351–1371, 2007.
- S. A. Mookerjee, A. S. Divakaruni, M. Jastroch, and M. D. Brand, “Mitochondrial uncoupling and lifespan,” Mechanisms of Ageing and Development, vol. 131, no. 7-8, pp. 463–472, 2010.
- R. J. Mailloux and M. E. Harper, “Uncoupling proteins and the control of mitochondrial reactive oxygen species production,” Free Radical Biology and Medicine, vol. 51, no. 6, pp. 1106–1115, 2011.
- V. P. Skulachev, “Role of uncoupled and non-coupled oxidations in maintenance of safely low levels of oxygen and its one-electron reductants,” Quarterly Reviews of Biophysics, vol. 29, no. 2, pp. 169–202, 1996.
- R. G. Hansford, B. A. Hogue, and V. Mildaziene, “Dependence of H2O2 formation by rat heart mitochondria on substrate availability and donor age,” Journal of Bioenergetics and Biomembranes, vol. 29, no. 1, pp. 89–95, 1997.
- A. S. Divakaruni and M. D. Brand, “The regulation and physiology of mitochondrial proton leak,” Physiology, vol. 26, no. 3, pp. 192–205, 2011.
- M. Klingenberg, “The ADP and ATP transport in mitochondria and its carrier,” Biochimica et Biophysica Acta, vol. 1778, no. 10, pp. 1978–2021, 2008.
- J. Nedergaard and B. Cannon, “The ‘novel’ ‘uncoupling’ proteins UCP2 and UCP3: what do they really do? Pros and cons for suggested functions,” Experimental Physiology, vol. 88, no. 1, pp. 65–84, 2003.
- S. Krauss, C. Y. Zhang, L. Scorrano et al., “Superoxide-mediated activation of uncoupling protein 2 causes pancreatic β cell dysfunction,” Journal of Clinical Investigation, vol. 112, no. 12, pp. 1831–1842, 2003.
- A. J. Vidal-Puig, D. Grujic, C. Y. Zhang et al., “Energy metabolism in uncoupling protein 3 gene knockout mice,” Journal of Biological Chemistry, vol. 275, no. 21, pp. 16258–16266, 2000.
- R. J. Mailloux, E. L. Seifert, F. Bouillaud, C. Aguer, S. Collins, and M. E. Harper, “Glutathionylation acts as a control switch for uncoupling proteins UCP2 and UCP3,” Journal of Biological Chemistry, vol. 286, no. 24, pp. 21865–21875, 2011.
- M. K. Hesselink, H. A. Keizer, L. B. Borghouts et al., “Protein expression of UCP3 differs between human type 1, type 2a, and type 2b fibers,” FASEB Journal, vol. 15, no. 6, pp. 1071–1073, 2001.
- A. P. Russell, E. Somm, M. Praz et al., “UCP3 protein regulation in human skeletal muscle fibre types I, IIa and IIx is dependent on exercise intensity,” Journal of Physiology, vol. 550, no. 3, pp. 855–861, 2003.
- A. P. Russell, G. Wadley, M. K. Hesselink et al., “UCP3 protein expression is lower in type I, IIa and IIx muscle fiber types of endurance-trained compared to untrained subjects,” Pflügers Archiv, vol. 445, no. 5, pp. 563–569, 2003.
- P. Jezek, H. Engstova, M. Zackova et al., “Fatty acid cycling mechanism and mitochondrial uncoupling proteins,” Biochimica et Biophysica Acta, vol. 1365, no. 1-2, pp. 319–327, 1998.
- T. C. Esteves and M. D. Brand, “The reactions catalysed by the mitochondrial uncoupling proteins UCP2 and UCP3,” Biochimica et Biophysica Acta, vol. 1709, no. 1, pp. 35–44, 2005.
- M. Klingenberg and K. S. Echtay, “Uncoupling proteins: the issues from a biochemist point of view,” Biochimica et Biophysica Acta, vol. 1504, no. 1, pp. 128–143, 2001.
- N. Parker, C. Affourtit, A. Vidal-Puig, and M. D. Brand, “Energization-dependent endogenous activation of proton conductance in skeletal muscle mitochondria,” Biochemical Journal, vol. 412, no. 1, pp. 131–139, 2008.
- L. S. Khailova, E. A. Prikhodko, V. I. Dedukhova, E. N. Mokhova, V. N. Popov, and V. P. Skulachev, “Participation of ATP/ADP antiporter in oleate- and oleate hydroperoxide-induced uncoupling suppressed by GDP and carboxyatractylate,” Biochimica et Biophysica Acta, vol. 1757, no. 9-10, pp. 1324–1329, 2006.
- D. W. Gong, Y. He, and M. L. Reitman, “Genomic organization and regulation by dietary fat of the uncoupling protein 3 and 2 genes,” Biochemical and Biophysical Research Communications, vol. 256, no. 1, pp. 27–32, 1999.
- K. S. Echtay, E. Winkler, K. Frischmuth, and M. Klingenberg, “Uncoupling proteins 2 and 3 are highly active H+ transporters and highly nucleotide sensitive when activated by coenzyme Q (ubiquinone),” Proceedings of the National Academy of Sciences, vol. 98, no. 4, pp. 1416–1421, 2001.
- M. Jaburek, M. Varecha, R. E. Gimeno et al., “Transport function and regulation of mitochondrial uncoupling proteins 2 and 3,” Journal of Biological Chemistry, vol. 274, no. 37, pp. 26003–26007, 1999.
- A. Andreyev, T. O. Bondareva, V. I. Dedukhova et al., “The ATP/ADP-antiporter is involved in the uncoupling effect of fatty acids on mitochondria,” European Journal of Biochemistry, vol. 182, no. 3, pp. 585–592, 1989.
- P. G. Sullivan, N. A. Rippy, K. Dorenbos, R. C. Concepcion, A. K. Agarwal, and J. M. Rho, “The ketogenic diet increases mitochondrial uncoupling protein levels and activity,” Annals of Neurology, vol. 55, no. 4, pp. 576–580, 2004.
- S. Iossa, M. P. Mollica, L. Lionetti, R. Crescenzo, M. Botta, and G. Liverini, “Skeletal muscle oxidative capacity in rats fed high-fat diet,” International Journal of Obesity, vol. 26, no. 1, pp. 65–72, 2002.
- T. Yamazaki, S. Okawa, and M. Takahashi, “The effects on weight loss and gene expression in adipose and hepatic tissues of very-low carbohydrate and low-fat isoenergetic diets in diet-induced obese mice,” Nutrition & Metabolism, vol. 13, p. 78, 2016.
- S. Srivastava, Y. Kashiwaya, M. T. King et al., “Mitochondrial biogenesis and increased uncoupling protein 1 in brown adipose tissue of mice fed a ketone ester diet,” FASEB Journal, vol. 26, no. 6, pp. 2351–2362, 2012.
- A. R. Kennedy, P. Pissios, H. Otu et al., “A high-fat, ketogenic diet induces a unique metabolic state in mice,” American Journal of Physiology-Endocrinology and Metabolism, vol. 292, no. 6, pp. E1724–1739, 2007.
- M. K. Badman, A. R. Kennedy, A. C. Adams, P. Pissios, and E. Maratos-Flier, “A very low carbohydrate ketogenic diet improves glucose tolerance in ob/ob mice independently of weight loss,” American Journal of Physiology-Endocrinology and Metabolism, vol. 297, no. 5, pp. E1197–1204, 2009.
- A. J. McAinch, J. S. Lee, C. R. Bruce, R. J. Tunstall, J. A. Hawley, and D. Cameron-Smith, “Dietary regulation of fat oxidative gene expression in different skeletal muscle fiber types,” Obesity Research, vol. 11, no. 12, pp. 1471–1479, 2003.
- M. J. Arkinstall, R. J. Tunstall, D. Cameron-Smith, and J. A. Hawley, “Regulation of metabolic genes in human skeletal muscle by short-term exercise and diet manipulation,” American Journal of Physiology-Endocrinology and Metabolism, vol. 287, no. 1, pp. E25–E31, 2004.
- K. Schwerzmann, H. Hoppeler, S. R. Kayar, and E. R. Weibel, “Oxidative capacity of muscle and mitochondria: correlation of physiological, biochemical, and morphometric characteristics,” Proceedings of the National Academy of Sciences, vol. 86, no. 5, pp. 1583–1587, 1989.
- J. Langfort, W. Pilis, R. Zarzeczny, K. Nazar, and H. Kaciuba-Uscilko, “Effect of low-carbohydrate-ketogenic diet on metabolic and hormonal responses to graded exercise in men,” Journal of Physiology and Pharmacology, vol. 47, no. 2, pp. 361–371, 1996.
- B. Simi, B. Sempore, M. H. Mayet, and R. J. Favier, “Additive effects of training and high-fat diet on energy metabolism during exercise,” Journal of Applied Physiology, vol. 71, no. 1, pp. 197–203, 1991.
- S. Schnyder, K. Svensson, B. Cardel, and C. Handschin, “Muscle PGC-1α is required for long term systemic and local adaptations to a ketogenic diet in mice,” American Journal of Physiology-Endocrinology and Metabolism, vol. 312, no. 5, pp. E437–E446, 2017.
- L. M. Burke, M. L. Ross, L. A. Garvican-Lewis et al., “Low carbohydrate, high fat diet impairs exercise economy and negates the performance benefit from intensified training in elite race walkers,” Journal of Physiology, vol. 595, no. 9, pp. 2785–2807, 2017.
- K. N. Frayn, “Calculation of substrate oxidation rates in vivo from gaseous exchange,” Journal of Applied Physiology, vol. 55, no. 2, pp. 628–634, 1983.
- X. M. Leverve, “Mitochondrial function and substrate availability,” Critical Care Medicine, vol. 35, no. 9, pp. S454–S460, 2007.
- K. Sato, Y. Kashiwaya, C. A. Keon et al., “Insulin, ketone bodies, and mitochondrial energy transduction,” FASEB Journal, vol. 9, no. 8, pp. 651–658, 1995.
- K. J. Bough, J. Wetherington, B. Hassel et al., “Mitochondrial biogenesis in the anticonvulsant mechanism of the ketogenic diet,” Annals of Neurology, vol. 60, no. 2, pp. 223–235, 2006.
- S. Ahola-Erkkila, C. J. Carroll, K. Peltola-Mjosund et al., “Ketogenic diet slows down mitochondrial myopathy progression in mice,” Human Molecular Genetics, vol. 19, no. 10, pp. 1974–1984, 2010.
- M. Balietti, B. Giorgetti, G. Di Stefano et al., “A ketogenic diet increases succinic dehydrogenase (SDH) activity and recovers age-related decrease in numeric density of SDH-positive mitochondria in cerebellar Purkinje cells of late-adult rats,” Micron, vol. 41, no. 2, pp. 143–148, 2010.
- P. M. Nemeth, B. W. Rosser, R. M. Choksi, B. J. Norris, and K. M. Baker, “Metabolic response to a high-fat diet in neonatal and adult rat muscle,” American Journal of Physiology-Cell Physiology, vol. 262, no. 2, pp. C282–C286, 1992.
- W. C. Miller, G. R. Bryce, and R. K. Conlee, “Adaptations to a high-fat diet that increase exercise endurance in male rats,” Journal of Applied Physiology, vol. 56, no. 1, pp. 78–83, 1984.
- J. H. Goedecke, C. Christie, G. Wilson et al., “Metabolic adaptations to a high-fat diet in endurance cyclists,” Metabolism, vol. 48, no. 12, pp. 1509–1517, 1999.
- B. Cheng, O. Karamizrak, T. D. Noakes, S. C. Dennis, and E. V. Lambert, “Time course of the effects of a high-fat diet and voluntary exercise on muscle enzyme activity in Long-Evans rats,” Physiology & Behavior, vol. 61, no. 5, pp. 701–705, 1997.
- S. Ahola, M. Auranen, P. Isohanni et al., “Modified Atkins diet induces subacute selective ragged-red-fiber lysis in mitochondrial myopathy patients,” EMBO Molecular Medicine, vol. 8, no. 11, pp. 1234–1247, 2016.
- J. R. Garbow, J. M. Doherty, R. C. Schugar et al., “Hepatic steatosis, inflammation, and ER stress in mice maintained long term on a very low-carbohydrate ketogenic diet,” American Journal of Physiology-Gastrointestinal and Liver Physiology, vol. 300, no. 6, pp. G956–G967, 2011.
- D. Cameron-Smith, L. M. Burke, D. J. Angus et al., “A short-term, high-fat diet up-regulates lipid metabolism and gene expression in human skeletal muscle,” American Journal of Clinical Nutrition, vol. 77, no. 2, pp. 313–318, 2003.
- J. W. Helge and B. Kiens, “Muscle enzyme activity in humans: role of substrate availability and training,” American Journal of Physiology-Regulatory, Integrative and Comparative Physiology, vol. 272, no. 5, pp. R1620–R1624, 1997.
- R. A. Lapachet, W. C. Miller, and D. A. Arnall, “Body fat and exercise endurance in trained rats adapted to a high-fat and/or high-carbohydrate diet,” Journal of Applied Physiology, vol. 80, no. 4, pp. 1173–1179, 1996.
- A. Woods, S. R. Johnstone, K. Dickerson et al., “LKB1 is the upstream kinase in the AMP-activated protein kinase cascade,” Current Biology, vol. 13, no. 22, pp. 2004–2008, 2003.
- R. J. Shaw, M. Kosmatka, N. Bardeesy et al., “The tumor suppressor LKB1 kinase directly activates AMP-activated kinase and regulates apoptosis in response to energy stress,” Proceedings of the National Academy of Sciences, vol. 101, no. 10, pp. 3329–3335, 2004.
- S. A. Hawley, J. Boudeau, J. L. Reid et al., “Complexes between the LKB1 tumor suppressor, STRAD alpha/beta and MO25 alpha/beta are upstream kinases in the AMP-activated protein kinase cascade,” Journal of Biology, vol. 2, no. 4, p. 28, 2003.
- J. W. Scott, S. A. Hawley, K. A. Green et al., “CBS domains form energy-sensing modules whose binding of adenosine ligands is disrupted by disease mutations,” Journal of Clinical Investigation, vol. 113, no. 2, pp. 274–284, 2004.
- B. Xiao, R. Heath, P. Saiu et al., “Structural basis for AMP binding to mammalian AMP-activated protein kinase,” Nature, vol. 449, no. 7161, pp. 496–500, 2007.
- M. Suter, U. Riek, R. Tuerk, U. Schlattner, T. Wallimann, and D. Neumann, “Dissecting the role of 5′-AMP for allosteric stimulation, activation, and deactivation of AMP-activated protein kinase,” Journal of Biological Chemistry, vol. 281, no. 43, pp. 32207–32216, 2006.
- J. W. Scott, F. A. Ross, J. K. Liu, and D. G. Hardie, “Regulation of AMP-activated protein kinase by a pseudosubstrate sequence on the gamma subunit,” EMBO Journal, vol. 26, no. 3, pp. 806–815, 2007.
- M. J. Sanders, P. O. Grondin, B. D. Hegarty, M. A. Snowden, and D. Carling, “Investigating the mechanism for AMP activation of the AMP-activated protein kinase cascade,” Biochemical Journal, vol. 403, no. 1, pp. 139–148, 2007.
- L. Chen, J. Wang, Y. Y. Zhang et al., “AMP-activated protein kinase undergoes nucleotide-dependent conformational changes,” Nature Structural & Molecular Biology, vol. 19, no. 7, pp. 716–718, 2012.
- G. J. Gowans, S. A. Hawley, F. A. Ross, and D. G. Hardie, “AMP is a true physiological regulator of AMP-activated protein kinase by both allosteric activation and enhancing net phosphorylation,” Cell Metabolism, vol. 18, no. 4, pp. 556–566, 2013.
- S. P. Davies, N. R. Helps, P. T. Cohen, and D. G. Hardie, “5′-AMP inhibits dephosphorylation, as well as promoting phosphorylation, of the AMP-activated protein kinase. Studies using bacterially expressed human protein phosphatase-2C alpha and native bovine protein phosphatase-2AC,” FEBS Letters, vol. 377, no. 3, pp. 421–425, 1995.
- J. S. Oakhill, R. Steel, Z. P. Chen et al., “AMPK is a direct adenylate charge-regulated protein kinase,” Science, vol. 332, no. 6036, pp. 1433–1435, 2011.
- B. Xiao, M. J. Sanders, E. Underwood et al., “Structure of mammalian AMPK and its regulation by ADP,” Nature, vol. 472, no. 7342, pp. 230–233, 2011.
- A. A. Gibson, R. V. Seimon, C. M. Lee et al., “Do ketogenic diets really suppress appetite? A systematic review and meta-analysis,” Obesity Reviews, vol. 16, no. 1, pp. 64–76, 2015.
- E. Tomas, T. S. Tsao, A. K. Saha et al., “Enhanced muscle fat oxidation and glucose transport by ACRP30 globular domain: acetyl-CoA carboxylase inhibition and AMP-activated protein kinase activation,” Proceedings of the National Academy of Sciences, vol. 99, no. 25, pp. 16309–16313, 2002.
- T. Yamauchi, J. Kamon, Y. Minokoshi et al., “Adiponectin stimulates glucose utilization and fatty-acid oxidation by activating AMP-activated protein kinase,” Nature Medicine, vol. 8, no. 11, pp. 1288–1295, 2002.
- Y. Minokoshi, Y. B. Kim, O. D. Peroni et al., “Leptin stimulates fatty-acid oxidation by activating AMP-activated protein kinase,” Nature, vol. 415, no. 6869, pp. 339–343, 2002.
- M. Xu, Y. T. Zhao, Y. Song et al., “alpha1-adrenergic receptors activate AMP-activated protein kinase in rat hearts,” Sheng Li Xue Bao, vol. 59, no. 2, pp. 175–182, 2007.
- H. J. Koh, M. F. Hirshman, H. He et al., “Adrenaline is a critical mediator of acute exercise-induced AMP-activated protein kinase activation in adipocytes,” Biochemical Journal, vol. 403, no. 3, pp. 473–481, 2007.
- S. K. Moule and R. M. Denton, “The activation of p38 MAPK by the beta-adrenergic agonist isoproterenol in rat epididymal fat cells,” FEBS Letters, vol. 439, no. 3, pp. 287–290, 1998.
- M. R. Ruth, A. M. Port, M. Shah et al., “Consuming a hypocaloric high fat low carbohydrate diet for 12 weeks lowers C-reactive protein, and raises serum adiponectin and high density lipoprotein-cholesterol in obese subjects,” Metabolism, vol. 62, no. 12, pp. 1779–1787, 2013.
- I. Partsalaki, A. Karvela, and B. E. Spiliotis, “Metabolic impact of a ketogenic diet compared to a hypocaloric diet in obese children and adolescents,” Journal of Pediatric Endocrinology and Metabolism, vol. 25, no. 7-8, pp. 697–704, 2012.
- T. Hu, L. Yao, K. Reynolds et al., “The effects of a low-carbohydrate diet vs. a low-fat diet on novel cardiovascular risk factors: a randomized controlled trial,” Nutrients, vol. 7, no. 9, pp. 7978–7994, 2015.
- E. P. Plaisance, M. Lukasova, S. Offermanns, Y. Zhang, G. Cao, and R. L. Judd, “Niacin stimulates adiponectin secretion through the GPR109A receptor,” American Journal of Physiology-Endocrinology and Metabolism, vol. 296, no. 3, pp. E549–E558, 2009.
- S. Klein and R. R. Wolfe, “Carbohydrate restriction regulates the adaptive response to fasting,” American Journal of Physiology-Endocrinology and Metabolism, vol. 262, no. 5, pp. E631–E636, 1992.
- J. Langfort, D. Czarnowski, M. Zendzian-Piotrowska, R. Zarzeczny, and J. Gorski, “Short-term low-carbohydrate diet dissociates lactate and ammonia thresholds in men,” Journal of Strength and Conditioning Research, vol. 18, no. 2, pp. 260–265, 2004.
- A. Steensberg, G. van Hall, C. Keller et al., “Muscle glycogen content and glucose uptake during exercise in humans: influence of prior exercise and dietary manipulation,” Journal of Physiology, vol. 541, no. 1, pp. 273–281, 2002.
- H. Sasaki, N. Hotta, and T. Ishiko, “Comparison of sympatho-adrenal activity during endurance exercise performed under high- and low-carbohydrate diet conditions,” Journal of Sports Medicine and Physical Fitness, vol. 31, no. 3, pp. 407–412, 1991.
- B. Fagerberg, O. Andersson, U. Nilsson, T. Hedner, B. Isaksson, and P. Bjorntorp, “Weight-reducing diets: role of carbohydrates on sympathetic nervous activity and hypotensive response,” International Journal of Obesity, vol. 8, no. 3, pp. 237–243, 1984.
- T. Mikulski, A. Ziemba, and K. Nazar, “Influence of body carbohydrate store modification on catecholamine and lactate responses to graded exercise in sedentary and physically active subjects,” Journal of Physiology and Pharmacology, vol. 59, no. 3, pp. 603–616, 2008.
- P. Szot, D. Weinshenker, J. M. Rho, T. W. Storey, and P. A. Schwartzkroin, “Norepinephrine is required for the anticonvulsant effect of the ketogenic diet,” Developmental Brain Research, vol. 129, no. 2, pp. 211–214, 2001.
- E. R. Hudson, D. A. Pan, J. James et al., “A novel domain in AMP-activated protein kinase causes glycogen storage bodies similar to those seen in hereditary cardiac arrhythmias,” Current Biology, vol. 13, no. 10, pp. 861–866, 2003.
- G. Polekhina, A. Gupta, B. J. Michell et al., “AMPK β subunit targets metabolic stress sensing to glycogen,” Current Biology, vol. 13, no. 10, pp. 867–871, 2003.
- J. F. Wojtaszewski, C. MacDonald, J. N. Nielsen et al., “Regulation of 5′AMP-activated protein kinase activity and substrate utilization in exercising human skeletal muscle,” American Journal of Physiology-Endocrinology and Metabolism, vol. 284, no. 4, pp. E813–E822, 2003.
- G. R. Steinberg, M. J. Watt, S. L. McGee et al., “Reduced glycogen availability is associated with increased AMPKα2 activity, nuclear AMPKα2 protein abundance, and GLUT4 mRNA expression in contracting human skeletal muscle,” Applied Physiology, Nutrition, and Metabolism, vol. 31, no. 3, pp. 302–312, 2006.
- M. J. Watt, G. R. Steinberg, S. Chan, A. Garnham, B. E. Kemp, and M. A. Febbraio, “Beta-adrenergic stimulation of skeletal muscle HSL can be overridden by AMPK signaling,” FASEB Journal, vol. 18, no. 12, pp. 1445-1446, 2004.
- Z. P. Chen, T. J. Stephens, S. Murthy et al., “Effect of exercise intensity on skeletal muscle AMPK signaling in humans,” Diabetes, vol. 52, no. 9, pp. 2205–2212, 2003.
- G. D. Wadley, R. S. Lee-Young, B. J. Canny et al., “Effect of exercise intensity and hypoxia on skeletal muscle AMPK signaling and substrate metabolism in humans,” American Journal of Physiology-Endocrinology and Metabolism, vol. 290, no. 4, pp. E694–E702, 2006.
- W. Derave, H. Ai, J. Ihlemann et al., “Dissociation of AMP-activated protein kinase activation and glucose transport in contracting slow-twitch muscle,” Diabetes, vol. 49, no. 8, pp. 1281–1287, 2000.
- A. McBride, S. Ghilagaber, A. Nikolaev, and D. G. Hardie, “The glycogen-binding domain on the AMPK beta subunit allows the kinase to act as a glycogen sensor,” Cell Metabolism, vol. 9, no. 1, pp. 23–34, 2009.
- S. D. Phinney, B. R. Bistrian, W. J. Evans, E. Gervino, and G. L. Blackburn, “The human metabolic response to chronic ketosis without caloric restriction: preservation of submaximal exercise capability with reduced carbohydrate oxidation,” Metabolism, vol. 32, no. 8, pp. 769–776, 1983.
- S. D. Phinney, E. S. Horton, E. A. Sims, J. S. Hanson, E. Danforth, and B. M. LaGrange, “Capacity for moderate exercise in obese subjects after adaptation to a hypocaloric, ketogenic diet,” Journal of Clinical Investigation, vol. 66, no. 5, pp. 1152–1161, 1980.
- L. M. Burke, D. J. Angus, G. R. Cox et al., “Effect of fat adaptation and carbohydrate restoration on metabolism and performance during prolonged cycling,” Journal of Applied Physiology, vol. 89, no. 6, pp. 2413–2421, 2000.
- M. P. Harber, S. Schenk, A. L. Barkan, and J. F. Horowitz, “Alterations in carbohydrate metabolism in response to short-term dietary carbohydrate restriction,” American Journal of Physiology-Endocrinology and Metabolism, vol. 289, no. 2, pp. E306–E312, 2005.
- T. W. Zderic, C. J. Davidson, S. Schenk, L. O. Byerley, and E. F. Coyle, “High-fat diet elevates resting intramuscular triglyceride concentration and whole body lipolysis during exercise,” American Journal of Physiology-Endocrinology and Metabolism, vol. 286, no. 2, pp. E217–E225, 2004.
- E. V. Lambert, D. P. Speechly, S. C. Dennis, and T. D. Noakes, “Enhanced endurance in trained cyclists during moderate intensity exercise following 2 weeks adaptation to a high fat diet,” European Journal of Applied Physiology and Occupational Physiology, vol. 69, no. 4, pp. 287–293, 1994.
- E. Hultman and J. Bergstrom, “Muscle glycogen synthesis in relation to diet studied in normal subjects,” Acta Medica Scandinavica, vol. 182, no. 1, pp. 109–117, 1967.
- J. D. Symons and I. Jacobs, “High-intensity exercise performance is not impaired by low intramuscular glycogen,” Medicine & Science in Sports & Exercise, vol. 21, no. 5, pp. 550–557, 1989.
- S. Horman, D. Vertommen, R. Heath et al., “Insulin antagonizes ischemia-induced Thr172 phosphorylation of AMP-activated protein kinase alpha-subunits in heart via hierarchical phosphorylation of Ser485/491,” Journal of Biological Chemistry, vol. 281, no. 9, pp. 5335–5340, 2006.
- M. J. Sharman, A. L. Gomez, W. J. Kraemer, and J. S. Volek, “Very low-carbohydrate and low-fat diets affect fasting lipids and postprandial lipemia differently in overweight men,” Journal of Nutrition, vol. 134, no. 4, pp. 880–885, 2004.
- M. Noakes, P. R. Foster, J. B. Keogh, A. P. James, J. C. Mamo, and P. M. Clifton, “Comparison of isocaloric very low carbohydrate/high saturated fat and high carbohydrate/low saturated fat diets on body composition and cardiovascular risk,” Nutrition & Metabolism, vol. 3, p. 7, 2006.
- T. L. Hernandez, J. P. Sutherland, P. Wolfe et al., “Lack of suppression of circulating free fatty acids and hypercholesterolemia during weight loss on a high-fat, low-carbohydrate diet,” American Journal of Clinical Nutrition, vol. 91, no. 3, pp. 578–585, 2010.
- S. Kodama, K. Saito, S. Tanaka et al., “Influence of fat and carbohydrate proportions on the metabolic profile in patients with type 2 diabetes: a meta-analysis,” Diabetes Care, vol. 32, no. 5, pp. 959–965, 2009.
- J. S. Volek, S. D. Phinney, C. E. Forsythe et al., “Carbohydrate restriction has a more favorable impact on the metabolic syndrome than a low fat diet,” Lipids, vol. 44, no. 4, pp. 297–309, 2009.
- E. C. Westman, W. S. Yancy, J. C. Mavropoulos, M. Marquart, and J. R. McDuffie, “The effect of a low-carbohydrate, ketogenic diet versus a low-glycemic index diet on glycemic control in type 2 diabetes mellitus,” Nutrition & Metabolism, vol. 5, no. 1, p. 36, 2008.
- J. C. Mavropoulos, W. S. Yancy, J. Hepburn, and E. C. Westman, “The effects of a low-carbohydrate, ketogenic diet on the polycystic ovary syndrome: a pilot study,” Nutrition & Metabolism, vol. 2, no. 1, p. 35, 2005.
- S. S. McDaniel, N. R. Rensing, L. L. Thio, K. A. Yamada, and M. Wong, “The ketogenic diet inhibits the mammalian target of rapamycin (mTOR) pathway,” Epilepsia, vol. 52, no. 3, pp. e7–e11, 2011.
- B. Draznin, C. Wang, R. Adochio, J. W. Leitner, and M. A. Cornier, “Effect of dietary macronutrient composition on AMPK and SIRT1 expression and activity in human skeletal muscle,” Hormone and Metabolic Research, vol. 44, no. 9, pp. 650–655, 2012.
- S. Imai, C. M. Armstrong, M. Kaeberlein, and L. Guarente, “Transcriptional silencing and longevity protein Sir2 is an NAD-dependent histone deacetylase,” Nature, vol. 403, no. 6771, pp. 795–800, 2000.
- J. Landry, A. Sutton, S. T. Tafrov et al., “The silencing protein SIR2 and its homologs are NAD-dependent protein deacetylases,” Proceedings of the National Academy of Sciences, vol. 97, no. 11, pp. 5807–5811, 2000.
- P. Onyango, I. Celic, J. M. McCaffery, J. D. Boeke, and A. P. Feinberg, “SIRT3, a human SIR2 homologue, is an NAD-dependent deacetylase localized to mitochondria,” Proceedings of the National Academy of Sciences, vol. 99, no. 21, pp. 13653–13658, 2002.
- M. B. Scher, A. Vaquero, and D. Reinberg, “SirT3 is a nuclear NAD+-dependent histone deacetylase that translocates to the mitochondria upon cellular stress,” Genes & Development, vol. 21, no. 8, pp. 920–928, 2007.
- D. B. Lombard, F. W. Alt, H. L. Cheng et al., “Mammalian Sir2 homolog SIRT3 regulates global mitochondrial lysine acetylation,” Molecular and Cellular Biology, vol. 27, no. 24, pp. 8807–8814, 2007.
- W. Ying, “NAD+/NADH and NADP+/NADPH in cellular functions and cell death: regulation and biological consequences,” Antioxidants & Redox Signaling, vol. 10, no. 2, pp. 179–206, 2008.
- K. A. Anderson, A. S. Madsen, C. A. Olsen, and M. D. Hirschey, “Metabolic control by sirtuins and other enzymes that sense NAD+, NADH, or their ratio,” Biochimica et Biophysica Acta, vol. 1858, no. 12, pp. 991–998, 2017.
- R. Nogueiras, K. M. Habegger, N. Chaudhary et al., “Sirtuin 1 and sirtuin 3: physiological modulators of metabolism,” Physiological Reviews, vol. 92, no. 3, pp. 1479–1514, 2012.
- A. S. Bause and M. C. Haigis, “SIRT3 regulation of mitochondrial oxidative stress,” Experimental Gerontology, vol. 48, no. 7, pp. 634–639, 2013.
- A. Giralt and F. Villarroya, “SIRT3, a pivotal actor in mitochondrial functions: metabolism, cell death and aging,” Biochemical Journal, vol. 444, no. 1, pp. 1–10, 2012.
- C. Canto, Z. Gerhart-Hines, J. N. Feige et al., “AMPK regulates energy expenditure by modulating NAD+ metabolism and SIRT1 activity,” Nature, vol. 458, no. 7241, pp. 1056–1060, 2009.
- M. Fulco, Y. Cen, P. Zhao et al., “Glucose restriction inhibits skeletal myoblast differentiation by activating SIRT1 through AMPK-mediated regulation of Nampt,” Developmental Cell, vol. 14, no. 5, pp. 661–673, 2008.
- J. R. Revollo, A. A. Grimm, and S. Imai, “The NAD biosynthesis pathway mediated by nicotinamide phosphoribosyltransferase regulates Sir2 activity in mammalian cells,” Journal of Biological Chemistry, vol. 279, no. 49, pp. 50754–50763, 2004.
- F. Lan, J. M. Cacicedo, N. Ruderman, and Y. Ido, “SIRT1 modulation of the acetylation status, cytosolic localization, and activity of LKB1. Possible role in AMP-activated protein kinase activation,” Journal of Biological Chemistry, vol. 283, no. 41, pp. 27628–27635, 2008.
- V. B. Pillai, N. R. Sundaresan, G. Kim et al., “Exogenous NAD blocks cardiac hypertrophic response via activation of the SIRT3-LKB1-AMP-activated kinase pathway,” Journal of Biological Chemistry, vol. 285, no. 5, pp. 3133–3144, 2010.
- D. C. Wallace, W. Fan, and V. Procaccio, “Mitochondrial energetics and therapeutics,” Annual Review of Pathology: Mechanisms of Disease, vol. 5, no. 1, pp. 297–348, 2010.
- C. T. Putman, L. L. Spriet, E. Hultman et al., “Pyruvate dehydrogenase activity and acetyl group accumulation during exercise after different diets,” American Journal of Physiology-Endocrinology and Metabolism, vol. 265, no. 5, pp. E752–E760, 1993.
- S. J. Peters, T. A. St Amand, R. A. Howlett, G. J. Heigenhauser, and L. L. Spriet, “Human skeletal muscle pyruvate dehydrogenase kinase activity increases after a low-carbohydrate diet,” American Journal of Physiology-Endocrinology and Metabolism, vol. 275, no. 6, pp. E980–E986, 1998.
- S. J. Peters, R. A. Harris, P. Wu, T. L. Pehleman, G. J. Heigenhauser, and L. L. Spriet, “Human skeletal muscle PDH kinase activity and isoform expression during a 3-day high-fat/low-carbohydrate diet,” Journal of Physiology-Endocrinology and Metabolism, vol. 281, no. 6, pp. E1151–E1158, 2001.
- T. L. Pehleman, S. J. Peters, G. J. Heigenhauser, and L. L. Spriet, “Enzymatic regulation of glucose disposal in human skeletal muscle after a high-fat, low-carbohydrate diet,” Journal of Applied Physiology, vol. 98, no. 1, pp. 100–107, 2005.
- M. S. Olson, S. C. Dennis, M. S. DeBuysere, and A. Padma, “The regulation of pyruvate dehydrogenase in the isolated perfused rat heart,” Journal of Biological Chemistry, vol. 253, no. 20, pp. 7369–7375, 1978.
- M. Elamin, D. N. Ruskin, S. A. Masino, and P. Sacchetti, “Ketone-based metabolic therapy: is increased NAD+ a primary mechanism?” Frontiers in Molecular Neuroscience, vol. 10, p. 377, 2017.
- M. D. Hirschey, T. Shimazu, J. A. Capra, K. S. Pollard, and E. Verdin, “SIRT1 and SIRT3 deacetylate homologous substrates: AceCS1,2 and HMGCS1,2,” Aging, vol. 3, no. 6, pp. 635–642, 2011.
- T. Shimazu, M. D. Hirschey, L. Hua et al., “SIRT3 deacetylates mitochondrial 3-hydroxy-3-methylglutaryl CoA synthase 2 and regulates ketone body production,” Cell Metabolism, vol. 12, no. 6, pp. 654–661, 2010.
- F. G. Hegardt, “Mitochondrial 3-hydroxy-3-methylglutaryl-CoA synthase: a control enzyme in ketogenesis,” Biochemical Journal, vol. 338, no. 3, pp. 569–582, 1999.
- M. D. Hirschey, T. Shimazu, E. Goetzman et al., “SIRT3 regulates mitochondrial fatty-acid oxidation by reversible enzyme deacetylation,” Nature, vol. 464, no. 7285, pp. 121–125, 2010.
- S. S. Bharathi, Y. Zhang, A. W. Mohsen et al., “Sirtuin 3 (SIRT3) protein regulates long-chain acyl-CoA dehydrogenase by deacetylating conserved lysines near the active site,” Journal of Biological Chemistry, vol. 288, no. 47, pp. 33837–33847, 2013.
- S. L. Choi, S. J. Kim, K. T. Lee et al., “The regulation of AMP-activated protein kinase by H2O2,” Biochemical and Biophysical Research Communications, vol. 287, no. 1, pp. 92–97, 2001.
- I. Irrcher, V. Ljubicic, and D. A. Hood, “Interactions between ROS and AMP kinase activity in the regulation of PGC-1α transcription in skeletal muscle cells,” American Journal of Physiology-Cell Physiology, vol. 296, no. 1, pp. C116–C123, 2009.
- X. Qiu, K. Brown, M. D. Hirschey, E. Verdin, and D. Chen, “Calorie restriction reduces oxidative stress by SIRT3-mediated SOD2 activation,” Cell Metabolism, vol. 12, no. 6, pp. 662–667, 2010.
- R. Tao, M. C. Coleman, J. D. Pennington et al., “Sirt3-mediated deacetylation of evolutionarily conserved lysine 122 regulates MnSOD activity in response to stress,” Molecular Cell, vol. 40, no. 6, pp. 893–904, 2010.
- Y. Chen, J. Zhang, Y. Lin et al., “Tumour suppressor SIRT3 deacetylates and activates manganese superoxide dismutase to scavenge ROS,” EMBO Reports, vol. 12, no. 6, pp. 534–541, 2011.
- E. Jing, B. Emanuelli, M. D. Hirschey et al., “Sirtuin-3 (Sirt3) regulates skeletal muscle metabolism and insulin signaling via altered mitochondrial oxidation and reactive oxygen species production,” Proceedings of the National Academy of Sciences, vol. 108, no. 35, pp. 14608–14613, 2011.
- S. Jager, C. Handschin, J. St-Pierre, and B. M. Spiegelman, “AMP-activated protein kinase (AMPK) action in skeletal muscle via direct phosphorylation of PGC-1α,” Proceedings of the National Academy of Sciences, vol. 104, no. 29, pp. 12017–12022, 2007.
- S. Nemoto, M. M. Fergusson, and T. Finkel, “SIRT1 functionally interacts with the metabolic regulator and transcriptional coactivator PGC-1α,” Journal of Biological Chemistry, vol. 280, no. 16, pp. 16456–16460, 2005.
- Z. Gerhart-Hines, J. T. Rodgers, O. Bare et al., “Metabolic control of muscle mitochondrial function and fatty acid oxidation through SIRT1/PGC-1α,” EMBO Journal, vol. 26, no. 7, pp. 1913–1923, 2007.
- J. T. Rodgers, C. Lerin, W. Haas, S. P. Gygi, B. M. Spiegelman, and P. Puigserver, “Nutrient control of glucose homeostasis through a complex of PGC-1α and SIRT1,” Nature, vol. 434, no. 7029, pp. 113–118, 2005.
- M. Lagouge, C. Argmann, Z. Gerhart-Hines et al., “Resveratrol improves mitochondrial function and protects against metabolic disease by activating SIRT1 and PGC-1α,” Cell, vol. 127, no. 6, pp. 1109–1122, 2006.
- X. Kong, R. Wang, Y. Xue et al., “Sirtuin 3, a new target of PGC-1α, plays an important role in the suppression of ROS and mitochondrial biogenesis,” PLoS One, vol. 5, no. 7, Article ID e11707, 2010.
- T. Shi, F. Wang, E. Stieren, and Q. Tong, “SIRT3, a mitochondrial sirtuin deacetylase, regulates mitochondrial function and thermogenesis in brown adipocytes,” Journal of Biological Chemistry, vol. 280, no. 14, pp. 13560–13567, 2005.
- O. M. Palacios, J. J. Carmona, S. Michan et al., “Diet and exercise signals regulate SIRT3 and activate AMPK and PGC-1α in skeletal muscle,” Aging, vol. 1, no. 9, pp. 771–783, 2009.
- I. Irrcher, V. Ljubicic, A. F. Kirwan, and D. A. Hood, “AMP-activated protein kinase-regulated activation of the PGC-1α promoter in skeletal muscle cells,” PLoS One, vol. 3, no. 10, Article ID e3614, 2008.
- W. J. Lee, M. Kim, H. S. Park et al., “AMPK activation increases fatty acid oxidation in skeletal muscle by activating PPARα and PGC-1,” Biochemical and Biophysical Research Communications, vol. 340, no. 1, pp. 291–295, 2006.
- S. Terada, M. Goto, M. Kato, K. Kawanaka, T. Shimokawa, and I. Tabata, “Effects of low-intensity prolonged exercise on PGC-1 mRNA expression in rat epitrochlearis muscle,” Biochemical and Biophysical Research Communications, vol. 296, no. 2, pp. 350–354, 2002.
- K. S. Rockl, M. F. Hirshman, J. Brandauer, N. Fujii, L. A. Witters, and L. J. Goodyear, “Skeletal muscle adaptation to exercise training: AMP-activated protein kinase mediates muscle fiber type shift,” Diabetes, vol. 56, no. 8, pp. 2062–2069, 2007.
- S. Miura, K. Kawanaka, Y. Kai et al., “An increase in murine skeletal muscle peroxisome proliferator-activated receptor-γ coactivator-1α (PGC-1α) mRNA in response to exercise is mediated by beta-adrenergic receptor activation,” Endocrinology, vol. 148, no. 7, pp. 3441–3448, 2007.
- M. Tadaishi, S. Miura, Y. Kai et al., “Effect of exercise intensity and AICAR on isoform-specific expressions of murine skeletal muscle PGC-1α mRNA: a role of β2-adrenergic receptor activation,” American Journal of Physiology-Endocrinology and Metabolism, vol. 300, no. 2, pp. E341–E349, 2011.
- S. Miura, Y. Kai, Y. Kamei, and O. Ezaki, “Isoform-specific increases in murine skeletal muscle peroxisome proliferator-activated receptor-γ coactivator-1α (PGC-1α) mRNA in response to β2-adrenergic receptor activation and exercise,” Endocrinology, vol. 149, no. 9, pp. 4527–4533, 2008.
- B. C. Frier, Z. Wan, D. B. Williams, A. L. Stefanson, and D. C. Wright, “Epinephrine and AICAR-induced PGC-1α mRNA expression is intact in skeletal muscle from rats fed a high-fat diet,” American Journal of Physiology-Cell Physiology, vol. 302, no. 12, pp. C1772–C1779, 2012.
- M. Iwabu, T. Yamauchi, M. Okada-Iwabu et al., “Adiponectin and AdipoR1 regulate PGC-1α and mitochondria by Ca2+ and AMPK/SIRT1,” Nature, vol. 464, no. 7293, pp. 1313–1319, 2010.
- I. Valle, A. Alvarez-Barrientos, E. Arza, S. Lamas, and M. Monsalve, “PGC-1α regulates the mitochondrial antioxidant defense system in vascular endothelial cells,” Cardiovascular Research, vol. 66, no. 3, pp. 562–573, 2005.
- B. L. Olson, M. B. Hock, S. Ekholm-Reed et al., “SCFCdc4 acts antagonistically to the PGC-1α transcriptional coactivator by targeting it for ubiquitin-mediated proteolysis,” Genes & Development, vol. 22, no. 2, pp. 252–264, 2008.
- Y. Zhang, G. Uguccioni, V. Ljubicic et al., “Multiple signaling pathways regulate contractile activity-mediated PGC-1α gene expression and activity in skeletal muscle cells,” Physiological Reports, vol. 2, no. 5, p. e12008, 2014.
- X. Li, B. Monks, Q. Ge, and M. J. Birnbaum, “Akt/PKB regulates hepatic metabolism by directly inhibiting PGC-1α transcription coactivator,” Nature, vol. 447, no. 7147, pp. 1012–1016, 2007.
- Y. X. Wang, “PPARs: diverse regulators in energy metabolism and metabolic diseases,” Cell Research, vol. 20, no. 2, pp. 124–137, 2010.
- Y. Olmos, I. Valle, S. Borniquel et al., “Mutual dependence of Foxo3a and PGC-1α in the induction of oxidative stress genes,” Journal of Biological Chemistry, vol. 284, no. 21, pp. 14476–14484, 2009.
- Z. Wu, P. Puigserver, U. Andersson et al., “Mechanisms controlling mitochondrial biogenesis and respiration through the thermogenic coactivator PGC-1,” Cell, vol. 98, no. 1, pp. 115–124, 1999.
- D. Kukidome, T. Nishikawa, K. Sonoda et al., “Activation of AMP-activated protein kinase reduces hyperglycemia-induced mitochondrial reactive oxygen species production and promotes mitochondrial biogenesis in human umbilical vein endothelial cells,” Diabetes, vol. 55, no. 1, pp. 120–127, 2006.
- R. B. Vega, J. M. Huss, and D. P. Kelly, “The coactivator PGC-1 cooperates with peroxisome proliferator-activated receptor alpha in transcriptional control of nuclear genes encoding mitochondrial fatty acid oxidation enzymes,” Molecular and Cellular Biology, vol. 20, no. 5, pp. 1868–1876, 2000.
- P. Wareski, A. Vaarmann, V. Choubey et al., “PGC-1α and PGC-1β regulate mitochondrial density in neurons,” Journal of Biological Chemistry, vol. 284, no. 32, pp. 21379–21385, 2009.
- Z. Lu, X. Xu, X. Hu et al., “PGC-1α regulates expression of myocardial mitochondrial antioxidants and myocardial oxidative stress after chronic systolic overload,” Antioxidants & Redox Signaling, vol. 13, no. 7, pp. 1011–1022, 2010.
- T. Geng, P. Li, M. Okutsu et al., “PGC-1α plays a functional role in exercise-induced mitochondrial biogenesis and angiogenesis but not fiber-type transformation in mouse skeletal muscle,” American Journal of Physiology-Cell Physiology, vol. 298, no. 3, pp. C572–C579, 2010.
- J. St-Pierre, J. Lin, S. Krauss et al., “Bioenergetic analysis of peroxisome proliferator-activated receptor gamma coactivators 1α and 1β (PGC-1α and PGC-1β) in muscle cells,” Journal of Biological Chemistry, vol. 278, no. 29, pp. 26597–26603, 2003.
- A. Gavalda-Navarro, J. A. Villena, A. Planavila, O. Vinas, and T. Mampel, “Expression of adenine nucleotide translocase (ANT) isoform genes is controlled by PGC-1α through different transcription factors,” Journal of Cellular Physiology, vol. 229, no. 12, pp. 2126–2136, 2014.
- L. Poulsen, M. Siersbaek, and S. Mandrup, “PPARs: fatty acid sensors controlling metabolism,” Seminars in Cell & Developmental Biology, vol. 23, no. 6, pp. 631–639, 2012.
- P. Puigserver, Z. Wu, C. W. Park, R. Graves, M. Wright, and B. M. Spiegelman, “A cold-inducible coactivator of nuclear receptors linked to adaptive thermogenesis,” Cell, vol. 92, no. 6, pp. 829–839, 1998.
- J. Lin, H. Wu, P. T. Tarr et al., “Transcriptional co-activator PGC-1α drives the formation of slow-twitch muscle fibres,” Nature, vol. 418, no. 6899, pp. 797–801, 2002.
- C. Handschin, S. Chin, P. Li et al., “Skeletal muscle fiber-type switching, exercise intolerance, and myopathy in PGC-1α muscle-specific knock-out animals,” Journal of Biological Chemistry, vol. 282, no. 41, pp. 30014–30021, 2007.
- K. Aquilano, P. Vigilanza, S. Baldelli, B. Pagliei, G. Rotilio, and M. R. Ciriolo, “Peroxisome proliferator-activated receptor gamma co-activator 1α (PGC-1α) and sirtuin 1 (SIRT1) reside in mitochondria: possible direct function in mitochondrial biogenesis,” Journal of Biological Chemistry, vol. 285, no. 28, pp. 21590–21599, 2010.
- A. Keren, Y. Tamir, and E. Bengal, “The p38 MAPK signaling pathway: a major regulator of skeletal muscle development,” Molecular and Cellular Endocrinology, vol. 252, no. 1-2, pp. 224–230, 2006.
- H. F. Kramer and L. J. Goodyear, “Exercise, MAPK, and NF-κB signaling in skeletal muscle,” Journal of Applied Physiology, vol. 103, no. 1, pp. 388–395, 2007.
- D. Knutti, D. Kressler, and A. Kralli, “Regulation of the transcriptional coactivator PGC-1 via MAPK-sensitive interaction with a repressor,” Proceedings of the National Academy of Sciences, vol. 98, no. 17, pp. 9713–9718, 2001.
- M. Fan, J. Rhee, J. St-Pierre et al., “Suppression of mitochondrial respiration through recruitment of p160 myb binding protein to PGC-1α: modulation by p38 MAPK,” Genes & Development, vol. 18, no. 3, pp. 278–289, 2004.
- T. Akimoto, S. C. Pohnert, P. Li et al., “Exercise stimulates Pgc-1α transcription in skeletal muscle through activation of the p38 MAPK pathway,” Journal of Biological Chemistry, vol. 280, no. 20, pp. 19587–19593, 2005.
- A. R. Pogozelski, T. Geng, P. Li et al., “p38γ mitogen-activated protein kinase is a key regulator in skeletal muscle metabolic adaptation in mice,” PLoS ONE, vol. 4, no. 11, Article ID e7934, 2009.
- X. Xi, J. Han, and J. Z. Zhang, “Stimulation of glucose transport by AMP-activated protein kinase via activation of p38 mitogen-activated protein kinase,” Journal of Biological Chemistry, vol. 276, no. 44, pp. 41029–41034, 2001.
- A. Pelletier, E. Joly, M. Prentki, and L. Coderre, “Adenosine 5′-monophosphate-activated protein kinase and p38 mitogen-activated protein kinase participate in the stimulation of glucose uptake by dinitrophenol in adult cardiomyocytes,” Endocrinology, vol. 146, no. 5, pp. 2285–2294, 2005.
- K. Lemieux, D. Konrad, A. Klip, and A. Marette, “The AMP-activated protein kinase activator AICAR does not induce GLUT4 translocation to transverse tubules but stimulates glucose uptake and p38 mitogen-activated protein kinases alpha and beta in skeletal muscle,” FASEB Journal, vol. 17, no. 12, pp. 1658–1665, 2003.
- C. Kang, K. M. O’Moore, J. R. Dickman, and L. L. Ji, “Exercise activation of muscle peroxisome proliferator-activated receptor-gamma coactivator-1c signaling is redox sensitive,” Free Radical Biology and Medicine, vol. 47, no. 10, pp. 1394–1400, 2009.
- S. Kurata, “Selective activation of p38 MAPK cascade and mitotic arrest caused by low level oxidative stress,” Journal of Biological Chemistry, vol. 275, no. 31, pp. 23413–23416, 2000.
- J. Huot, F. Houle, F. Marceau, and J. Landry, “Oxidative stress-induced actin reorganization mediated by the p38 mitogen-activated protein kinase/heat shock protein 27 pathway in vascular endothelial cells,” Circulation Research, vol. 80, no. 3, pp. 383–392, 1997.
- M. C. Gomez-Cabrera, C. Borras, F. V. Pallardo, J. Sastre, L. L. Ji, and J. Vina, “Decreasing xanthine oxidase-mediated oxidative stress prevents useful cellular adaptations to exercise in rats,” Journal of Physiology, vol. 567, no. 1, pp. 113–120, 2005.
- G. D. Wadley, M. A. Nicolas, D. S. Hiam, and G. K. McConell, “Xanthine oxidase inhibition attenuates skeletal muscle signaling following acute exercise but does not impair mitochondrial adaptations to endurance training,” American Journal of Physiology-Endocrinology and Metabolism, vol. 304, no. 8, pp. E853–E862, 2013.
- A. R. Gosmanov, N. C. Nordtvedt, R. Brown, and D. B. Thomason, “Exercise effects on muscle beta-adrenergic signaling for MAPK-dependent NKCC activity are rapid and persistent,” Journal of Applied Physiology, vol. 93, no. 4, pp. 1457–1465, 2002.
- A. R. Gosmanov and D. B. Thomason, “Insulin and isoproterenol differentially regulate mitogen-activated protein kinase-dependent Na+-K+-2Cl− cotransporter activity in skeletal muscle,” Diabetes, vol. 51, no. 3, pp. 615–623, 2002.
- F. R. Jornayvaz, M. J. Jurczak, H. Y. Lee et al., “A high-fat, ketogenic diet causes hepatic insulin resistance in mice, despite increasing energy expenditure and preventing weight gain,” American Journal of Physiology-Endocrinology and Metabolism, vol. 299, no. 5, pp. E808–E815, 2010.
- T. Furuyama, T. Nakazawa, I. Nakano, and N. Mori, “Identification of the differential distribution patterns of mRNAs and consensus binding sequences for mouse DAF-16 homologues,” Biochemical Journal, vol. 349, no. 2, pp. 629–634, 2000.
- M. J. Anderson, C. S. Viars, S. Czekay, W. K. Cavenee, and K. C. Arden, “Cloning and characterization of three human forkhead genes that comprise an FKHR-like gene subfamily,” Genomics, vol. 47, no. 2, pp. 187–199, 1998.
- W. H. Biggs, W. K. Cavenee, and K. C. Arden, “Identification and characterization of members of the FKHR (FOX O) subclass of winged-helix transcription factors in the mouse,” Mammalian Genome, vol. 12, no. 6, pp. 416–425, 2001.
- A. M. Sanchez, A. Csibi, A. Raibon et al., “AMPK promotes skeletal muscle autophagy through activation of forkhead FoxO3a and interaction with Ulk1,” Journal of Cellular Biochemistry, vol. 113, no. 2, pp. 695–710, 2012.
- E. L. Greer, P. R. Oskoui, M. R. Banko et al., “The energy sensor AMP-activated protein kinase directly regulates the mammalian FOXO3 transcription factor,” Journal of Biological Chemistry, vol. 282, no. 41, pp. 30107–30119, 2007.
- X. N. Li, J. Song, L. Zhang et al., “Activation of the AMPK-FOXO3 pathway reduces fatty acid-induced increase in intracellular reactive oxygen species by upregulating thioredoxin,” Diabetes, vol. 58, no. 10, pp. 2246–2257, 2009.
- A. Brunet, L. B. Sweeney, J. F. Sturgill et al., “Stress-dependent regulation of FOXO transcription factors by the SIRT1 deacetylase,” Science, vol. 303, no. 5666, pp. 2011–2015, 2004.
- Y. Olmos, F. J. Sanchez-Gomez, B. Wild et al., “SirT1 regulation of antioxidant genes is dependent on the formation of a FoxO3a/PGC-1α complex,” Antioxid Redox Signal, vol. 19, no. 13, pp. 1507–1521, 2013.
- N. R. Sundaresan, M. Gupta, G. Kim, S. B. Rajamohan, A. Isbatan, and M. P. Gupta, “Sirt3 blocks the cardiac hypertrophic response by augmenting Foxo3a-dependent antioxidant defense mechanisms in mice,” Journal of Clinical Investigation, vol. 119, no. 9, pp. 2758–2771, 2009.
- K. M. Jacobs, J. D. Pennington, K. S. Bisht et al., “SIRT3 interacts with the daf-16 homolog FOXO3a in the mitochondria, as well as increases FOXO3a dependent gene expression,” International Journal of Biological Sciences, vol. 4, no. 5, pp. 291–299, 2008.
- H. S. Kim, K. Patel, K. Muldoon-Jacobs et al., “SIRT3 is a mitochondria-localized tumor suppressor required for maintenance of mitochondrial integrity and metabolism during stress,” Cancer Cell, vol. 17, no. 1, pp. 41–52, 2010.
- A. H. Tseng, S. S. Shieh, and D. L. Wang, “SIRT3 deacetylates FOXO3 to protect mitochondria against oxidative damage,” Free Radical Biology and Medicine, vol. 63, pp. 222–234, 2013.
- K. Miyamoto, K. Y. Araki, K. Naka et al., “Foxo3a is essential for maintenance of the hematopoietic stem cell pool,” Cell Stem Cell, vol. 1, no. 1, pp. 101–112, 2007.
- G. J. Kops, T. B. Dansen, P. E. Polderman et al., “Forkhead transcription factor FOXO3a protects quiescent cells from oxidative stress,” Nature, vol. 419, no. 6904, pp. 316–321, 2002.
- W. Q. Tan, K. Wang, D. Y. Lv, and P. F. Li, “Foxo3a inhibits cardiomyocyte hypertrophy through transactivating catalase,” Journal of Biological Chemistry, vol. 283, no. 44, pp. 29730–29739, 2008.
- S. Nemoto and T. Finkel, “Redox regulation of forkhead proteins through a p66shc-dependent signaling pathway,” Science, vol. 295, no. 5564, pp. 2450–2452, 2002.
- C. B. Chiribau, L. Cheng, I. C. Cucoranu, Y. S. Yu, R. E. Clempus, and D. Sorescu, “FOXO3A regulates peroxiredoxin III expression in human cardiac fibroblasts,” Journal of Biological Chemistry, vol. 283, no. 13, pp. 8211–8217, 2008.
- Y. Furukawa-Hibi, K. Yoshida-Araki, T. Ohta, K. Ikeda, and N. Motoyama, “FOXO forkhead transcription factors induce G2-M checkpoint in response to oxidative stress,” Journal of Biological Chemistry, vol. 277, no. 30, pp. 26729–26732, 2002.
- H. Tran, A. Brunet, J. M. Grenier et al., “DNA repair pathway stimulated by the forkhead transcription factor FOXO3a through the Gadd45 protein,” Science, vol. 296, no. 5567, pp. 530–534, 2002.
- Q. Weng, Z. Liu, B. Li, K. Liu, W. Wu, and H. Liu, “Oxidative stress induces mouse follicular granulosa cells apoptosis via JNK/FoxO1 pathway,” PLoS One, vol. 11, no. 12, Article ID e0167869, 2016.
- Y. Furukawa-Hibi, Y. Kobayashi, C. Chen, and N. Motoyama, “FOXO transcription factors in cell-cycle regulation and the response to oxidative stress,” Antioxidants & Redox Signaling, vol. 7, no. 5-6, pp. 752–760, 2005.
- A. Peserico, F. Chiacchiera, V. Grossi et al., “A novel AMPK-dependent FoxO3A-SIRT3 intramitochondrial complex sensing glucose levels,” Cellular and Molecular Life Sciences, vol. 70, no. 11, pp. 2015–2029, 2013.
- N. Lutzner, J. De-Castro Arce, and F. Rosl, “Gene expression of the tumour suppressor LKB1 is mediated by Sp1, NF-Y and FOXO transcription factors,” PLoS One, vol. 7, no. 3, Article ID e32590, 2012.
- R. Tao, D. Wei, H. Gao, Y. Liu, R. A. DePinho, and X. C. Dong, “Hepatic FoxOs regulate lipid metabolism via modulation of expression of the nicotinamide phosphoribosyltransferase gene,” Journal of Biological Chemistry, vol. 286, no. 16, pp. 14681–14690, 2011.
- J. V. Virbasius and R. C. Scarpulla, “Activation of the human mitochondrial transcription factor A gene by nuclear respiratory factors: a potential regulatory link between nuclear and mitochondrial gene expression in organelle biogenesis,” Proceedings of the National Academy of Sciences, vol. 91, no. 4, pp. 1309–1313, 1994.
- R. P. Fisher and D. A. Clayton, “Purification and characterization of human mitochondrial transcription factor 1,” Molecular and Cellular Biology, vol. 8, no. 8, pp. 3496–3509, 1988.
- T. E. Shutt, M. F. Lodeiro, J. Cotney, C. E. Cameron, and G. S. Shadel, “Core human mitochondrial transcription apparatus is a regulated two-component system in vitro,” Proceedings of the National Academy of Sciences, vol. 107, no. 27, pp. 12133–12138, 2010.
- M. Falkenberg, M. Gaspari, A. Rantanen, A. Trifunovic, N. G. Larsson, and C. M. Gustafsson, “Mitochondrial transcription factors B1 and B2 activate transcription of human mtDNA,” Nature Genetics, vol. 31, no. 3, pp. 289–294, 2002.
- H. C. Lee, P. H. Yin, C. W. Chi, and Y. H. Wei, “Increase in mitochondrial mass in human fibroblasts under oxidative stress and during replicative cell senescence,” Journal of Biomedical Science, vol. 9, no. 6, pp. 517–526, 2002.
- S. Miranda, R. Foncea, J. Guerrero, and F. Leighton, “Oxidative stress and upregulation of mitochondrial biogenesis genes in mitochondrial DNA-depleted HeLa cells,” Biochemical and Biophysical Research Communications, vol. 258, no. 1, pp. 44–49, 1999.
- E. Q. Toyama, S. Herzig, J. Courchet et al., “Metabolism. AMP-activated protein kinase mediates mitochondrial fission in response to energy stress,” Science, vol. 351, no. 6270, pp. 275–281, 2016.
- S. Baldelli, K. Aquilano, and M. R. Ciriolo, “Punctum on two different transcription factors regulated by PGC-1α: nuclear factor erythroid-derived 2-like 2 and nuclear respiratory factor 2,” Biochimica et Biophysica Acta, vol. 1830, no. 8, pp. 4137–4146, 2013.
- H. K. Bryan, A. Olayanju, C. E. Goldring, and B. K. Park, “The Nrf2 cell defence pathway: Keap1-dependent and -independent mechanisms of regulation,” Biochemical Pharmacology, vol. 85, no. 6, pp. 705–717, 2013.
- K. Itoh, N. Wakabayashi, Y. Katoh et al., “Keap1 represses nuclear activation of antioxidant responsive elements by Nrf2 through binding to the amino-terminal Neh2 domain,” Genes & Development, vol. 13, no. 1, pp. 76–86, 1999.
- K. Itoh, N. Wakabayashi, Y. Katoh, T. Ishii, T. O’Connor, and M. Yamamoto, “Keap1 regulates both cytoplasmic-nuclear shuttling and degradation of Nrf2 in response to electrophiles,” Genes to Cells, vol. 8, no. 4, pp. 379–391, 2003.
- D. D. Zhang and M. Hannink, “Distinct cysteine residues in Keap1 are required for Keap1-dependent ubiquitination of Nrf2 and for stabilization of Nrf2 by chemopreventive agents and oxidative stress,” Molecular and Cellular Biology, vol. 23, no. 22, pp. 8137–8151, 2003.
- M. McMahon, K. Itoh, M. Yamamoto, and J. D. Hayes, “Keap1-dependent proteasomal degradation of transcription factor Nrf2 contributes to the negative regulation of antioxidant response element-driven gene expression,” Journal of Biological Chemistry, vol. 278, no. 24, pp. 21592–21600, 2003.
- A. Kobayashi, M. I. Kang, Y. Watai et al., “Oxidative and electrophilic stresses activate Nrf2 through inhibition of ubiquitination activity of Keap1,” Molecular and Cellular Biology, vol. 26, no. 1, pp. 221–229, 2006.
- S. L. Colombo and S. Moncada, “AMPKα1 regulates the antioxidant status of vascular endothelial cells,” Biochemical Journal, vol. 421, no. 2, pp. 163–169, 2009.
- K. Aquilano, S. Baldelli, B. Pagliei, S. M. Cannata, G. Rotilio, and M. R. Ciriolo, “p53 orchestrates the PGC-1α-mediated antioxidant response upon mild redox and metabolic imbalance,” Antioxidants & Redox Signaling, vol. 18, no. 4, pp. 386–399, 2013.
- T. L. Merry and M. Ristow, “Nuclear factor erythroid-derived 2-like 2 (NFE2L2, Nrf2) mediates exercise-induced mitochondrial biogenesis and the anti-oxidant response in mice,” Journal of Physiology, vol. 594, no. 18, pp. 5195–5207, 2016.
- S. B. Shelar, M. Narasimhan, G. Shanmugam et al., “Disruption of nuclear factor (erythroid-derived-2)-like 2 antioxidant signaling: a mechanism for impaired activation of stem cells and delayed regeneration of skeletal muscle,” FASEB Journal, vol. 30, no. 5, pp. 1865–1879, 2016.
- V. R. Muthusamy, S. Kannan, K. Sadhaasivam et al., “Acute exercise stress activates Nrf2/ARE signaling and promotes antioxidant mechanisms in the myocardium,” Free Radical Biology and Medicine, vol. 52, no. 2, pp. 366–376, 2012.
- C. J. Miller, S. S. Gounder, S. Kannan et al., “Disruption of Nrf2/ARE signaling impairs antioxidant mechanisms and promotes cell degradation pathways in aged skeletal muscle,” Biochimica et Biophysica Acta, vol. 1822, no. 6, pp. 1038–1050, 2012.
- K. Itoh, T. Chiba, S. Takahashi et al., “An Nrf2/small Maf heterodimer mediates the induction of phase II detoxifying enzyme genes through antioxidant response elements,” Biochemical and Biophysical Research Communications, vol. 236, no. 2, pp. 313–322, 1997.
- N. L. Alderson, Y. Wang, M. Blatnik et al., “S-(2-Succinyl)cysteine: a novel chemical modification of tissue proteins by a Krebs cycle intermediate,” Archives of Biochemistry and Biophysics, vol. 450, no. 1, pp. 1–8, 2006.
- H. Ashrafian, G. Czibik, M. Bellahcene et al., “Fumarate is cardioprotective via activation of the Nrf2 antioxidant pathway,” Cell Metabolism, vol. 15, no. 3, pp. 361–371, 2012.
- J. Adam, E. Hatipoglu, L. O’Flaherty et al., “Renal cyst formation in Fh1-deficient mice is independent of the Hif/Phd pathway: roles for fumarate in KEAP1 succination and Nrf2 signaling,” Cancer Cell, vol. 20, no. 4, pp. 524–537, 2011.
- Y. Izuta, T. Imada, R. Hisamura et al., “Ketone body 3-hydroxybutyrate mimics calorie restriction via the Nrf2 activator, fumarate, in the retina,” Aging Cell, 2017, in press.
- S. Borniquel, N. Garcia-Quintans, I. Valle et al., “Inactivation of Foxo3a and subsequent downregulation of PGC-1α mediate nitric oxide-induced endothelial cell migration,” Molecular and Cellular Biology, vol. 30, no. 16, pp. 4035–4044, 2010.
- S. Fourquet, R. Guerois, D. Biard, and M. B. Toledano, “Activation of NRF2 by nitrosative agents and H2O2 involves KEAP1 disulfide formation,” Journal of Biological Chemistry, vol. 285, no. 11, pp. 8463–8471, 2010.
- Z. Wang, Z. Ying, A. Bosy-Westphal et al., “Specific metabolic rates of major organs and tissues across adulthood: evaluation by mechanistic model of resting energy expenditure,” American Journal of Clinical Nutrition, vol. 92, no. 6, pp. 1369–1377, 2010.
- T. Ogata and Y. Yamasaki, “Ultra-high-resolution scanning electron microscopy of mitochondria and sarcoplasmic reticulum arrangement in human red, white, and intermediate muscle fibers,” Anatomical Record, vol. 248, no. 2, pp. 214–223, 1997.
- M. R. Jackman and W. T. Willis, “Characteristics of mitochondria isolated from type I and type IIb skeletal muscle,” American Journal of Physiology-Cell Physiology, vol. 270, no. 2, pp. C673–C678, 1996.
- S. Schiaffino and C. Reggiani, “Fiber types in mammalian skeletal muscles,” Physiological Reviews, vol. 91, no. 4, pp. 1447–1531, 2011.
- Z. Yan, M. Okutsu, Y. N. Akhtar, and V. A. Lira, “Regulation of exercise-induced fiber type transformation, mitochondrial biogenesis, and angiogenesis in skeletal muscle,” Journal of Applied Physiology, vol. 110, no. 1, pp. 264–274, 2011.
- M. D. Delp and C. Duan, “Composition and size of type I, IIA, IID/X, and IIB fibers and citrate synthase activity of rat muscle,” Journal of Applied Physiology, vol. 80, no. 1, pp. 261–270, 1996.
- E. J. Anderson and P. D. Neufer, “Type II skeletal myofibers possess unique properties that potentiate mitochondrial H2O2 generation,” American Journal of Physiology-Cell Physiology, vol. 290, no. 3, pp. C844–C851, 2006.
- G. D. Brinkworth, M. Noakes, P. M. Clifton, and J. D. Buckley, “Effects of a low carbohydrate weight loss diet on exercise capacity and tolerance in obese subjects,” Obesity, vol. 17, no. 10, pp. 1916–1923, 2009.
- J. Fleming, M. J. Sharman, N. G. Avery et al., “Endurance capacity and high-intensity exercise performance responses to a high fat diet,” International Journal of Sport Nutrition and Exercise Metabolism, vol. 13, no. 4, pp. 466–478, 2003.
- N. A. Johnson, S. R. Stannard, K. Mehalski et al., “Intramyocellular triacylglycerol in prolonged cycling with high- and low-carbohydrate availability,” Journal of Applied Physiology, vol. 94, no. 4, pp. 1365–1372, 2003.
- J. W. Helge, B. Wulff, and B. Kiens, “Impact of a fat-rich diet on endurance in man: role of the dietary period,” Medicine & Science in Sports & Exercise, vol. 30, no. 3, pp. 456–461, 1998.
- D. S. Rowlands and W. G. Hopkins, “Effects of high-fat and high-carbohydrate diets on metabolism and performance in cycling,” Metabolism, vol. 51, no. 6, pp. 678–690, 2002.
- N. K. Stepto, A. L. Carey, H. M. Staudacher, N. K. Cummings, L. M. Burke, and J. A. Hawley, “Effect of short-term fat adaptation on high-intensity training,” Medicine & Science in Sports & Exercise, vol. 34, no. 3, pp. 449–455, 2002.
- N. Fujii, T. Hayashi, M. F. Hirshman et al., “Exercise induces isoform-specific increase in 5′AMP-activated protein kinase activity in human skeletal muscle,” Biochemical and Biophysical Research Communications, vol. 273, no. 3, pp. 1150–1155, 2000.
- G. K. McConell, R. S. Lee-Young, Z. P. Chen et al., “Short-term exercise training in humans reduces AMPK signalling during prolonged exercise independent of muscle glycogen,” Journal of Physiology, vol. 568, no. 2, pp. 665–676, 2005.
- B. Guerra, A. Guadalupe-Grau, T. Fuentes et al., “SIRT1, AMP-activated protein kinase phosphorylation and downstream kinases in response to a single bout of sprint exercise: influence of glucose ingestion,” European Journal of Applied Physiology, vol. 109, no. 4, pp. 731–743, 2010.
- L. Li, C. Muhlfeld, B. Niemann et al., “Mitochondrial biogenesis and PGC-1α deacetylation by chronic treadmill exercise: differential response in cardiac and skeletal muscle,” Basic Research in Cardiology, vol. 106, no. 6, pp. 1221–1234, 2011.
- L. Li, R. Pan, R. Li et al., “Mitochondrial biogenesis and peroxisome proliferator-activated receptor-gamma coactivator-1α (PGC-1α) deacetylation by physical activity: intact adipocytokine signaling is required,” Diabetes, vol. 60, no. 1, pp. 157–167, 2011.
- C. L. Dumke, J. Mark Davis, E. Angela Murphy et al., “Successive bouts of cycling stimulates genes associated with mitochondrial biogenesis,” European Journal of Applied Physiology, vol. 107, no. 4, pp. 419–427, 2009.
- J. P. Little, A. Safdar, G. P. Wilkin, M. A. Tarnopolsky, and M. J. Gibala, “A practical model of low-volume high-intensity interval training induces mitochondrial biogenesis in human skeletal muscle: potential mechanisms,” Journal of Physiology, vol. 588, no. 6, pp. 1011–1022, 2010.
- V. Ljubicic, A. M. Joseph, P. J. Adhihetty et al., “Molecular basis for an attenuated mitochondrial adaptive plasticity in aged skeletal muscle,” Aging, vol. 1, no. 9, pp. 818–830, 2009.
- B. J. Gurd, G. P. Holloway, Y. Yoshida, and A. Bonen, “In mammalian muscle, SIRT3 is present in mitochondria and not in the nucleus; and SIRT3 is upregulated by chronic muscle contraction in an adenosine monophosphate-activated protein kinase-independent manner,” Metabolism, vol. 61, no. 5, pp. 733–741, 2012.
- F. Hokari, E. Kawasaki, A. Sakai, K. Koshinaka, K. Sakuma, and K. Kawanaka, “Muscle contractile activity regulates SIRT3 protein expression in rat skeletal muscles,” Journal of Applied Physiology, vol. 109, no. 2, pp. 332–340, 2010.
- T. Li, S. He, S. Liu, Z. Kong, J. Wang, and Y. Zhang, “Effects of different exercise durations on Keap1-Nrf2-ARE pathway activation in mouse skeletal muscle,” Free Radical Research, vol. 49, no. 10, pp. 1269–1274, 2015.
- U. Widegren, X. J. Jiang, A. Krook et al., “Divergent effects of exercise on metabolic and mitogenic signaling pathways in human skeletal muscle,” FASEB Journal, vol. 12, no. 13, pp. 1379–1389, 1998.
- R. Somwar, M. Perreault, S. Kapur et al., “Activation of p38 mitogen-activated protein kinase alpha and beta by insulin and contraction in rat skeletal muscle: potential role in the stimulation of glucose transport,” Diabetes, vol. 49, no. 11, pp. 1794–1800, 2000.
- J. W. Ryder, R. Fahlman, H. Wallberg-Henriksson et al., “Effect of contraction on mitogen-activated protein kinase signal transduction in skeletal muscle. Involvement of the mitogen- and stress-activated protein kinase 1,” Journal of Biological Chemistry, vol. 275, no. 2, pp. 1457–1462, 2000.
- N. A. Strobel, J. M. Peake, A. Matsumoto et al., “Antioxidant supplementation reduces skeletal muscle mitochondrial biogenesis,” Medicine & Science in Sports & Exercise, vol. 43, no. 6, pp. 1017–1024, 2011.
- M. C. Gomez-Cabrera, E. Domenech, M. Romagnoli et al., “Oral administration of vitamin C decreases muscle mitochondrial biogenesis and hampers training-induced adaptations in endurance performance,” American Journal of Clinical Nutrition, vol. 87, no. 1, pp. 142–149, 2008.
- K. Baar, A. R. Wende, T. E. Jones et al., “Adaptations of skeletal muscle to exercise: rapid increase in the transcriptional coactivator PGC-1,” FASEB Journal, vol. 16, no. 14, pp. 1879–1886, 2002.
- H. Pilegaard, B. Saltin, and P. D. Neufer, “Exercise induces transient transcriptional activation of the PGC-1α gene in human skeletal muscle,” Journal of Physiology, vol. 546, no. 3, pp. 851–858, 2003.
- G. Paulsen, K. T. Cumming, G. Holden et al., “Vitamin C and E supplementation hampers cellular adaptation to endurance training in humans: a double-blind, randomised, controlled trial,” Journal of Physiology, vol. 592, no. 8, pp. 1887–1901, 2014.
- M. Ristow, K. Zarse, A. Oberbach et al., “Antioxidants prevent health-promoting effects of physical exercise in humans,” Proceedings of the National Academy of Sciences, vol. 106, no. 21, pp. 8665–8670, 2009.
- A. T. White and S. Schenk, “NAD+/NADH and skeletal muscle mitochondrial adaptations to exercise,” American Journal of Physiology-Endocrinology and Metabolism, vol. 303, no. 3, pp. E308–E321, 2012.
- E. Paleologou, N. Ismayilova, and M. Kinali, “Use of the ketogenic diet to treat intractable epilepsy in mitochondrial disorders,” Journal of Clinical Medicine, vol. 6, no. 6, p. 56, 2017.
- J. M. Rho, “How does the ketogenic diet induce anti-seizure effects?” Neuroscience Letters, vol. 637, pp. 4–10, 2017.
- N. N. Danial, A. L. Hartman, C. E. Stafstrom, and L. L. Thio, “How does the ketogenic diet work? Four potential mechanisms,” Journal of Child Neurology, vol. 28, no. 8, pp. 1027–1033, 2013.
Copyright © 2018 Vincent J. Miller et al. This is an open access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.