Archives of Toxicology

, Volume 89, Issue 8, pp 1209–1226 | Cite as

Interactions between mitochondrial reactive oxygen species and cellular glucose metabolism

  • Dania C. Liemburg-Apers
  • Peter H. G. M. Willems
  • Werner J. H. KoopmanEmail author
  • Sander Grefte
Open Access
Review Article


Mitochondrial reactive oxygen species (ROS) production and detoxification are tightly balanced. Shifting this balance enables ROS to activate intracellular signaling and/or induce cellular damage and cell death. Increased mitochondrial ROS production is observed in a number of pathological conditions characterized by mitochondrial dysfunction. One important hallmark of these diseases is enhanced glycolytic activity and low or impaired oxidative phosphorylation. This suggests that ROS is involved in glycolysis (dys)regulation and vice versa. Here we focus on the bidirectional link between ROS and the regulation of glucose metabolism. To this end, we provide a basic introduction into mitochondrial energy metabolism, ROS generation and redox homeostasis. Next, we discuss the interactions between cellular glucose metabolism and ROS. ROS-stimulated cellular glucose uptake can stimulate both ROS production and scavenging. When glucose-stimulated ROS production, leading to further glucose uptake, is not adequately counterbalanced by (glucose-stimulated) ROS scavenging systems, a toxic cycle is triggered, ultimately leading to cell death. Here we inventoried the various cellular regulatory mechanisms and negative feedback loops that prevent this cycle from occurring. It is concluded that more insight in these processes is required to understand why they are (un)able to prevent excessive ROS production during various pathological conditions in humans.


Glucose GLUT1 GLUT4 Mitochondria ROS Oxidative stress 



Mitochondrial membrane potential




12-Hydroperoxyeicosatetraenoic acid


Apoptosis signal-regulating kinase 1


AMP-activated protein kinase


Adenine nucleotide translocator


Ataxia telangiectasia mutated


Complex I or NADH: ubiquinone oxidoreductase


Complex II or succinate:ubiquinone oxidoreductase


Complex III or ubiquinol:cytochrome-c oxidoreductase


Complex IV or cytochrome-c oxidase


Copper/zinc-dependent superoxide dismutase or SOD1


Complex V or F1Fo-ATP synthase


Carbohydrate response element-binding protein


Cyclophilin D


Electron transport chain


Dehydroascorbic acid


Dihydroorotate dehydrogenase


Reduced flavin adenine dinucleotide


Flavin mononucleotide




Glyceraldehyde-3-phosphate dehydrogenase


Gα-interacting protein-interacting protein, C-terminus


Glucose/glucose oxidase


Glucose transporter


Glutathione peroxidase






Oxidized glutathione


Hypoxia-inducible factor-1




Lactate dehydrogenase


Monoamine oxidases

MAPK p38

p38 mitogen-activated protein kinase


Mouse embryonic fibroblasts


Sn-glycerol-3-phosphate dehydrogenase


Mitochondrial inner membrane


Manganese-dependent superoxide dismutase or SOD2


Mitochondrial outer membrane


Inorganic phosphate carrier


Permeability transition pore


Reduced nicotinamide adenine dinucleotide


Nicotinamide nucleotide transhydrogenase


NAD(P)H oxidase


Oxygen-dependent degradation domains


2-Oxoglutarate dehydrogenase


Oxidative phosphorylation


Poly(ADP-ribose) polymerase


Pyruvate dehydrogenase




Inorganic phosphate


Phosphoinositide 3-kinase


Phosphatidylinositol-3-kinase-related protein kinase


Proton motive force


Peroxisome proliferator-activated receptor δ


Pentose phosphate pathway




Rieske iron–sulfur protein


Reactive oxygen species


Copper/zinc-dependent superoxide dismutase or CuZnSOD


Manganese-dependent superoxide dismutase or MnSOD


Specificity protein


Tricarboxylic acid




Thioredoxin-interacting protein


Uncoupling protein


Xanthine/xanthine oxidase


Mitochondria are among the prime ATP-generating organelles, which are necessary for cellular functioning (Koopman et al. 2012, 2013). To this end, glucose is oxidized to pyruvate in the cytosol by the glycolysis pathway. Next, pyruvate enters the mitochondria where it is converted into acetyl-coenzyme-A that is further oxidized within the tricarboxylic acid (TCA) cycle. Alternatively, acetyl-coenzyme-A can be generated by fatty acid breakdown during β-oxidation. Conversion of acetyl-coenzyme-A by the TCA cycle yields reduced nicotinamide adenine dinucleotide (NADH) and reduced flavin adenine dinucleotide (FADH2). These two molecules serve as electron donors for mitochondrial complex I (CI or NADH:ubiquinone oxidoreductase) and complex II (CII or succinate:ubiquinone oxidoreductase) of the electron transport chain (ETC). The electrons are subsequently transported by ubiquinone to complex III (CIII or ubiquinol:cytochrome-c oxidoreductase) and by cytochrome-c to complex IV (CIV or cytochrome-c oxidase) where they react with oxygen to form water. At CI, CIII and CIV protons are expelled from the mitochondrial matrix across the mitochondrial inner membrane (MIM). This results in establishment of an inward-directed proton motive force (PMF) that consists of a chemical (ΔpH) and electrical (Δψ) component (Mitchell 1961). Via complex V (CV or F1Fo-ATP synthase), protons are allowed to flow back into the matrix to fuel generation of ATP from ADP and inorganic phosphate (Pi). Together with the ETC, CV constitutes the mitochondrial oxidative phosphorylation (OXPHOS) system.

Mitochondrial ROS generation

Both as a consequence of normal electron transport and during mitochondrial dysfunction, electrons can escape from the ETC to induce formation of superoxide anions by one-electron reduction of oxygen. This means that, under certain conditions, mitochondria can substantially contribute to the generation of cellular reactive oxygen species (ROS; Adam-Vizi and Chinopoulos 2006; Murphy 2009). Interestingly, several proteins involved in glycolysis, mitochondrial electron transport, β-oxidation and the TCA cycle are also able to generate superoxide, hydrogen peroxide and/or other ROS. These include CI (Grivennikova and Vinogradov 2013; Murphy 2009; Treberg et al. 2011), CII (Quinlan et al. 2012a; Siebels and Drose 2013), CIII (Muller et al. 2004; Murphy 2009), dihydroorotate dehydrogenase (DHOH; Forman and Kennedy 1975; Orr et al. 2012), pyruvate dehydrogenase (PDH; Fisher-Wellman et al. 2013; Starkov et al. 2004), aconitase (Gardner 2002; Vasquez-Vivar et al. 2000), 2-oxoglutarate dehydrogenase (Odh, or α-ketoglutarate dehydrogenase; Bunik and Sievers 2002; Quinlan et al. 2014; Starkov et al. 2004; Tretter and Adam-Vizi 2004) and Sn-glycerol-3-phosphate dehydrogenase (mGPDH; Orr et al. 2012). In addition, various other mitochondrial proteins like monoamine oxidases (MAOs) and p66shc/cytochrome-c (Di Lisa et al. 2009; Giorgio et al. 2005; Hauptmann et al. 1996) are capable of ROS production. Regarding the ETC, CI and CIII are the most well characterized (Murphy 2009). In case of CI, superoxide production can occur at two sites: the flavin mononucleotide (FMN) site and the iron–sulfur cluster (Genova et al. 2001; Herrero and Barja 2000; Johnson et al. 2003; Kussmaul and Hirst 2006; Lambert and Brand 2004; Treberg et al. 2011). Alternatively, hydrogen peroxide might be directly formed at the FMN site (Grivennikova and Vinogradov 2013). In CIII, evidence was provided that superoxide is produced only at the quinol-oxidizing (QO) site (Kramer et al. 2004; Muller et al. 2003; Murphy 2009). Inhibitor studies suggested that superoxide and/or hydrogen peroxide can also be produced at the flavin site of CII (Quinlan et al. 2012a).

However, in these studies, the exact sites and magnitude of ROS production depend on the used OXPHOS substrates and inhibitors, respectively. In the absence of inhibitors, (native) ROS production appears to be much lower (Quinlan et al. 2012b, 2013). Since these studies use isolated mitochondria, the situation might also be different in intact cells and tissues.

Maintaining redox homeostasis

To prevent unintentional generation of redox signals and induction of oxidative stress, mitochondria possess powerful antioxidant systems. One of these consists of manganese-dependent superoxide dismutase (MnSOD or SOD2), an enzyme that is localized in the mitochondrial matrix and rapidly converts superoxide to hydrogen peroxide. This conversion is also catalyzed by the copper/zinc-dependent superoxide dismutase (Cu/ZnSOD or SOD1), which is localized in the cytosol, nucleus and mitochondrial intermembrane space (Murphy 2009; Tyler 1975; Weisiger and Fridovich 1973). In turn, hydrogen peroxide can be converted into water by the action of catalases that are mainly located in the peroxisomes and also in mitochondria (Salvi et al. 2007). However, within mitochondria, hydrogen peroxide is mainly removed by the action of glutathione peroxidase-1 (Gpx1; Cox et al. 2010; Esposito et al. 2000; Esworthy et al. 1997), peroxiredoxins 3 and 5 (Prx3 and Prx5) and the thioredoxin-2 (Trx2) system (Chae et al. 1999; Chang et al. 2004; Cox et al. 2010), which require glutathione (GSH). Oxidized GSH (GSSG) is recycled to GSH by the action of glutathione reductase. Similarly, oxidized Trx2 is recycled by Trx reductase. Both of these systems require NADPH (Arner 2009; Carlberg and Mannervik 1985), which is regenerated in the cytosol by glucose-6-phosphate-dehydrogenase (G6PDH) via the pentose phosphate pathway (PPP; Le Goffe et al. 2002). Alternatively, NADPH can be regenerated in the mitochondrial matrix by nicotinamide nucleotide transhydrogenase (NNT), which uses NADH and the PMF (Hatefi and Yamaguchi 1996; Rydstrom 2006; Yin et al. 2012).

ROS signaling

ROS have the ability to modulate the transcription and activity of enzymes, receptors and transporter (Mailloux et al. 2014; Martinez-Reyes and Cuezva 2014; Sena and Chandel 2012), for instance during adaptation to exercise (Gomez-Cabrera et al. 2005; Silveira et al. 2006). To fulfill a signaling function, ROS should be able to induce reversible protein modifications, thereby affecting its activity and/or function. Hydrogen peroxide is believed to be a main player in ROS signaling due to its physicochemical properties, which include a relatively low reactivity, long half-life and the ability to diffuse through membranes (Forman et al. 2010; Winterbourn and Hampton 2008). Mechanistically, hydrogen peroxide can oxidize thiol groups (–SH) on exposed cysteine residues in proteins, resulting in the formation of sulfenic acid (–SO, known as S-oxidation or sulfenylation) (Carballal et al. 2003; Charles et al. 2007; Seres et al. 1996). Subsequently, the sulfenic acid group can: (1) form inter- and intramolecular disulfide bonds with other thiol groups leading to altered protein structure or the formation of homo- and/or heterodimers (Brennan et al. 2004; Delaunay et al. 2002; Rehder and Borges 2010; Yang et al. 2007), (2) react with GSH (–SSG; thereby inducing S-glutathionylation of the protein; (Chen et al. 2007b; Hurd et al. 2008; McLain et al. 2013), or (3) react with amides to form a sulfenyl amide (Salmeen et al. 2003; Sivaramakrishnan et al. 2010). Although not discussed in this review, mitochondria are also exposed to various reactive nitrogen species (RNS), which can induce oxidative protein modifications (Beltran et al. 2000; Boveris et al. 2006; Hogg 2002; Rossig et al. 1999; Zaobornyj and Ghafourifar 2012). Interestingly, most of the above modifications are reversible (reviewed in detail elsewhere: Forman et al. 2010; Handy and Loscalzo 2012; Mailloux et al. 2014), i.e., disulfide bonds between thiol groups can be reduced by the Trx reductase system, while glutathionylated thiol groups can be reduced by glutaredoxin (Grx2) utilizing the GSH pool (Beer et al. 2004; Handy and Loscalzo 2012; Mailloux et al. 2014). Regarding ROS regulation of protein activity, several phosphatases contain thiol groups in their active site, which upon oxidation lead to a loss of dephosphorylation activity (Rhee et al. 2000; Tonks 2005). Also, proteins of the TCA cycle and OXPHOS are regulated by ROS (Mailloux et al. 2014). For example, S-glutathionylation has been shown to reduce the activity of CI (Hurd et al. 2008) and Odh (McLain et al. 2013).

Oxidative stress induction

When mitochondrial ROS production exceeds the capacity of the cell’s antioxidant systems or when the latter systems are less active, increased ROS levels can induce cell damage (oxidative stress). In principle, superoxide can react with protein iron–sulfur (Fe–S) clusters (Liochev and Fridovich 1994), which, in the presence of hydrogen peroxide, induce generation of hydroxyl radicals (Fenton reaction). The latter are highly reactive and can damage lipids, proteins and DNA (Martinez-Reyes and Cuezva 2014; Valko et al. 2006). Given the fact that iron is effectively sequestered (Kakhlon and Cabantchik 2002), the relevance of the Fenton reaction is not fully established in vivo. Superoxide can react with NO to form peroxynitrite (ONOO; Pacher et al. 2005). When rising too high, hydrogen peroxide can induce over-oxidization of cysteine residues from sulfenic acid (–SO) to sulfinic acid (–SO2H) and sulfonic acid (–SO3H). In case of CI, such over-oxidation is associated with irreversible deactivation of CI (Hurd et al. 2008; Mailloux et al. 2014). In general, if ROS levels exceed a certain threshold, they will impair OXPHOS complexes and further stimulate ROS production (Galloway and Yoon 2012). In the light of the above, it is not surprising that increased ROS levels, although not always oxidative stress, are observed during various pathological conditions. For example, primary fibroblasts derived either from CI deficient mice or patients show increased ROS levels, but no obvious signs of oxidative stress (Koopman et al. 2007; Valsecchi et al. 2013; Verkaart et al. 2007a, b). Increased ROS levels also have been observed in multiple types of cancer (e.g., prostate, colorectal, ovarian, pancreatic, breast, liver, bladder, melanoma, glioma), neurogenerative diseases (e.g., Alzheimer’s disease and Parkinson’s disease) and during insulin-resistance and diabetes (Afanas’ev 2011; Freeman et al. 2006; Kumar et al. 2008; Pi and Collins 2010; Sabens Liedhegner et al. 2012; Sanchez-Gomez et al. 2013). Below we will discuss the interplay between ROS levels, glucose uptake and metabolism in detail.

Regulation of glucose uptake by reactive oxygen species

Stimulation of cellular glucose uptake is frequently observed during conditions of oxidative stress. Exogenous addition of hydrogen peroxide stimulates glucose uptake in skeletal muscle (Higaki et al. 2008; Jensen et al. 2008; Kim et al. 2006), C2C12 myoblasts, clone 9 liver cells and 3T3 fibroblasts (Prasad and Ismail-Beigi 1999). Upon electrical stimulation, endogenous ROS also induced an increase in glucose uptake in muscle cells (Merry et al. 2010; Pinheiro et al. 2010). Interestingly, in L6 myoblasts, inhibition of cellular glucose uptake was associated with increased ROS levels (Andrisse et al. 2014), perhaps suggesting a role for glucose in ROS scavenging. Cellular glucose uptake is mediated by glucose transporters (GLUTs), of which fourteen isoforms have been described with different kinetic properties and modes of regulation (Carruthers et al. 2009; Joost and Thorens 2001). Here we will primarily focus on GLUT1 and GLUT4, which are abundantly expressed in muscle cells and well studied. GLUT1 is expressed during all stages of embryonic development (Hogan et al. 1991). After birth, GLUT1 expression decreases, but most cell types still express low levels of GLUT1 to mediate basal glucose uptake. However, GLUT1 expression remains high in cells that primarily depend on glycolysis for ATP generation such as erythrocytes and tumor cells. In the latter, GLUT1 is frequently up-regulated (Baer et al. 1997; Brown and Wahl 1993; Nishioka et al. 1992), which is associated with poor survival in various malignant tumors (Szablewski 2013). GLUTs are expressed in a highly tissue-specific manner (Bell et al. 1990; Gould and Holman 1993). This, in combination with the fact that the different GLUTs display different functional characteristics, allows for a tissue-specific regulation of glucose uptake (Gould and Holman 1993). For example, GLUT4 expression is up-regulated in differentiating muscle cells (Mitsumoto and Klip 1992), which show increased levels of OXPHOS complexes and higher respiration rates (Mitsumoto and Klip 1992; Remels et al. 2010). This suggests that under certain conditions (i.e., differentiation), cells (co)express both GLUT1 and other GLUTs to facilitate increased glucose uptake to support increased cellular respiration. In fact, under basal conditions, the majority of GLUT4 in muscle cells is retained in intracellular vesicles that are derived from the trans-Golgi network. Retention of these vesicles requires the activity of the Rab GTPase activating proteins TBC1D1 and TBC1D4 (Eguez et al. 2005; Larance et al. 2005; Sano et al. 2003). Stimulation by insulin or exercise induces translocation of these vesicles to the plasma membrane. Although the full signaling cascade regulating vesicle translocation is still incompletely understood, Akt-mediated inhibition of TBC1D1/TBC1D4 plays an essential role (Eguez et al. 2005; Funaki et al. 2004; Kohn et al. 1998; Larance et al. 2005; Ng et al. 2008; Sano et al. 2003). It appears that following translocation to the plasma membrane, an additional activation step is required for stimulation of glucose uptake (Funaki et al. 2004; Somwar et al. 2002; Sweeney et al. 1999).

Transcriptional regulation of GLUT expression by ROS

Protein expression levels of human GLUT1 are controlled by a promoter region and several putative enhancer regions that contain binding sites for various transcription factors including specificity proteins (Sp1; Vinals et al. 1997) and hypoxia-inducible factor-1 (HIF-1; Ebert et al. 1995). Mild oxidative stress induced by either glucose/glucose oxidase (Glc/GO) or xanthine/xanthine oxidase (Xan/XO) has been shown to up-regulate GLUT1 expression by increasing the transcription rate and mRNA stability leading to increased GLUT1 protein and glucose transport activity (Kozlovsky et al. 1997). As far as we know, there is no experimental evidence demonstrating the involvement of Sp1 in ROS-induced stimulation of GLUT1 expression, and therefore, we here focus on the role of HIF-1.

HIF-1 consists of two subunits, HIF-1α and HIF-1β. Under normoxic conditions, prolines within the oxygen-dependent degradation domains (ODDs) of HIF-1α are hydroxylated by prolyl-4-hydroxylases (PHDs; Ivan et al. 2001). This hydroxylation acts as an ubiquitination signal leading to proteasomal degradation of HIF-1α. In the absence of oxygen, HIF-1α ubiquitinylation is inhibited allowing its interaction with HIF-1β to drive transcription of various target genes, including GLUT1 (Hayashi et al. 2004; Iyer et al. 1998; Ouiddir et al. 1999; Wood et al. 1998). During hypoxia, ROS levels increase and play an important role in HIF-1α stabilization (Brunelle et al. 2005; Chandel et al. 2000; Guzy et al. 2005; Mansfield et al. 2005; Sanjuan-Pla et al. 2005; Schroedl et al. 2002). Preventing ROS-mediated HIF-1α stabilization represses GLUT1 expression and glucose uptake in Lewis lung carcinoma, HT-29 colon, and T47D breast cancer cells (Jung et al. 2013). Upon mitochondrial DNA depletion (Chandel et al. 2000; Mansfield et al. 2005) and in mouse embryonic fibroblasts (MEFs) lacking cytochrome-c (Mansfield et al. 2005), hypoxia-induced HIF-1α stabilization is abrogated. This suggests that hypoxia-induced ROS are of mitochondrial origin. Knockout of the Rieske iron–sulfur protein (RISP) in mitochondrial CIII decreases ROS production during hypoxia and attenuates hypoxic stabilization of HIF-1α (Guzy et al. 2005). Therefore, RISP-mediated mitochondrial ROS production appears to be involved in HIF-1α stabilization during hypoxia. At the RISP site, electrons are transferred one-by-one from ubiquinol to cytochrome-c1. This one-electron donation generates a highly reactive ubisemiquinone, which can act as a source for superoxide generation. Over-expression of catalase (Chandel et al. 2000; Guzy et al. 2005) or GPx1 (Brunelle et al. 2005; Emerling et al. 2005) abolishes HIF-1α stabilization during hypoxia, whereas over-expression of SOD1 or SOD2 does not (Brunelle et al. 2005; Guzy et al. 2005). In addition, exogenous hydrogen peroxide is sufficient to stabilize HIF-1α under normoxic conditions (Chandel et al. 2000; Jung et al. 2008; Mansfield et al. 2005). This suggests that the stabilization of HIF-1α primarily involves hydrogen peroxide via inactivation of PHDs (Fig. 1a) and subsequent reduction of HIF-1α ubiquitinylation (Chandel et al. 1998; Guzy and Schumacker 2006). However, HIF-1α ubiquitinylation is incompletely blocked by exogenous or hypoxia-derived hydrogen peroxide (Guzy et al. 2005), suggesting the involvement of additional mechanisms.
Fig. 1

Interplay between ROS and glucose. a Glucose uptake can be regulated by: (1) altering the expression level of glucose transporters (GLUTs; blue), (2) stimulating translocation of GLUTs from internal vesicles to the plasma membrane and (3) changing the intrinsic activity of GLUTs at the plasma membrane. b Glycolytic conversion of glucose into pyruvate and subsequent pyruvate entry into the mitochondria (1) stimulates ROS production by hyperpolarizing the mitochondrial membrane potential (Δψ↑). Subsequently, ROS stimulate glucose uptake (see a), thereby triggering additional ROS production. Glucose flux through the pentose phosphate pathway (stimulated by AMPK and ATM) generates NADPH (2), which is an important cofactor in ROS scavenging. c Hyperpolarization of the mitochondrial membrane potential (Δψ↑) is prevented by: (1) GLUT1 internalization, (2) GLUT1 mRNA degradation, (3) reduction of pyruvate to lactate and subsequent secretion of lactate. A hyperpolarized mitochondrial membrane potential is diminished by: (4) transient uncoupling of the mitochondrial membrane potential (PTP, UCP) or enhancing oxidative phosphorylation efficiency by HK–CV interaction. Proteins that are activated by ROS are depicted in yellow (for details, see main text). 4-HDDE 4-hydroxydodecadienal, 12-HPETE 12-hydroperoxyeicosatetraenoic acid, Δψ mitochondrial membrane potential, ATM ataxia telangiectasia mutated, CV complex V, GIPC Gα-interacting protein-interacting protein, C-terminus, GLC glucose, Glut1 glucose transporter 1, HIF-1 hypoxia-inducible factor 1, HK hexokinase, LAC lactate, LDH lactate dehydrogenase, MCT monocarboxylate transporter, P-AMPK phosphorylated (activated) AMP-activated protein kinase, PHD prolyl hydroxylase domain, P-p38, phosphorylated (activated) p38 mitogen-activated protein kinase, PI3K phosphoinositide 3-kinase, PTP permeability transition pore, PYR pyruvate, ROS reactive oxygen species, TXNIP thioredoxin-interacting protein, UCP uncoupling protein (color figure online)

Alternatively, ROS-induced HIF-1α stabilization was proposed to be mediated by specific signaling routes. Phosphoinositide 3-kinase (PI3K) appears to be involved in HIF-1α stabilization by mitochondrial ROS in Hep3B cells (Fig. 1a; arrow 1) (Chandel et al. 2000). Compatible with this mechanisms, PI3K inhibition lowered GLUT1 transcription under hypoxic and normoxic conditions (Chen et al. 2001). In addition, both p38 mitogen-activated protein kinase (MAPK p38) and its upstream kinases MKK3/6 are activated by hypoxia-derived ROS and are essential in HIF-1α stabilization and the ensuing increase in GLUT1 mRNA levels (Emerling et al. 2005). Activation of MAPK p38 by ROS is mediated by apoptosis signal-regulating kinase 1 (ASK1), which is normally bound to Trx. Oxidation of Trx releases ASK1 leading to activation of its downstream targets MKK3, MKK4, MKK6 and MMK7 and subsequently JNK and MAPK p38 (Nagai et al. 2007). Also, AMP-activated protein kinase (AMPK) can be activated by ROS during hypoxia (Emerling et al. 2009). This protein plays a key role in energy metabolism and cellular adaptation to ROS (Wu et al. 2014). Various metabolic stress conditions can activate AMPK through phosphorylation of Thr172 by upstream kinases such as LKB1 (Hardie 2011). In addition, AMP-interaction stimulates AMPK activation by promoting AMPK phosphorylation, preventing AMPK dephosphorylation and inducing allosteric activation of AMPK (Davies et al. 1995; Hardie and Ashford 2014). Oxidative stress can inhibit the ETC, potentially leading to an increased AMP:ATP ratio (Hawley et al. 2010). However, under hypoxic conditions, AMPK is also activated by mitochondrial ROS in an LKB1-dependent and AMP:ATP-independent manner (Emerling et al. 2009; Mungai et al. 2011). In muscle cells, AMPK activation stimulated GLUT1 expression and twofold increased cellular glucose uptake (Fryer et al. 2002). Besides activation of HIF-1α, AMPK can up-regulate GLUT1 via degradation of thioredoxin-interacting protein (TXNIP) (Fig. 1a; arrow 1). TXNIP reduces the level of GLUT1 mRNA in the nucleus. TXNIP is degraded upon Ser308 phosphorylation by AMPK, leading to increased GLUT1 mRNA and protein levels (Wu et al. 2013).

Regulation of GLUT-vesicle translocation to the plasma membrane by ROS

The cellular capacity for glucose uptake is co-determined by the abundance of functional GLUTs at the plasma membrane. This means that glucose uptake can be regulated by GLUT trafficking between the cytosol and the plasma membrane. During exercise, muscle contraction is associated with increased ROS levels, believed to represent a fast adaptive response to an increase in energy demand (Chambers et al. 2009; Murrant et al. 1999; Wretman et al. 2001). In these cells, endogenous (i.e., those induced by contraction) and exogenous ROS stimulate glucose uptake via a mechanism involving activation of Akt and/or AMPK (Higaki et al. 2008; Sandstrom et al. 2006; Toyoda et al. 2004). ROS can activate Akt (Fernandes et al. 2011; Niwa et al. 2003), which probably involves inactivation of cysteine-based phosphatases (Okoh et al. 2011). ROS can activate AMPK by modification of cysteine residues on its α-catalytical subunit (Zmijewski et al. 2010). Once activated, Akt and AMPK phosphorylate TBC1D4 and TBC1D1, leading to GLUT4 translocation (Geraghty et al. 2007; Kramer et al. 2006; Taylor et al. 2008; Thong et al. 2007).

Translocation of GLUT1 to the plasma membrane (Fig. 1a; arrow 2) is regulated by Ataxia telangiectasia mutated (ATM; Andrisse et al. 2013). This protein is a member of the family of phosphatidylinositol-3-kinase-related protein kinases (PIKKs) and plays an important role in the response to DNA damage after which it becomes phosphorylated. However, ATM can also localize to the cytosol where it is activated by ROS and involved in cytosolic signaling (Alexander and Walker 2010). ATM is thiol-oxidized by ROS to form an active dimer of two covalently linked monomers (Guo et al. 2010). Activated ATM localizes near mitochondria and mitochondria are necessary for ROS-induced ATM activation (Morita et al. 2014). This suggests that ATM dimerization and activation are stimulated by mitochondrial ROS. Evidence was provided that ATM is activated by ROS during treatment with the chemotherapeutic agent doxorubicin (Kurz et al. 2004). In muscle, ATM activation by doxorubicin mediates targeting of GLUT1 to the cell surface by phosphorylation of GLUT1 at S490. This serine residue is part of a C-terminal PDZ motif, and phosphorylation of this residue induces interaction of GLUT1 with the PDZ-interacting protein, Gα-interacting protein and C-terminus (GIPC1) (Andrisse et al. 2013). GIPC1 interaction promotes GLUT1 trafficking to the cell surface and stimulates glucose uptake (Fig. 1a; arrow 2) (Wieman et al. 2009).

Regulation of GLUT1 activity at the plasma membrane by ROS

Regulation of glucose uptake can also occur directly at the plasma membrane. This type of regulation occurs on a relatively short-term time scale (i.e., within 1 h) and represents a relatively fast mechanism that allows the cells to cope with metabolic stress (Fig. 1a; arrow 3). The intrinsic activity of GLUTs can be modulated by conformational changes or posttranslational modifications that increase glucose affinity (Asano et al. 1991; Levine et al. 2002). In addition, the maximal rate of cellular glucose uptake (Vmax) can be modulated by activation or deactivation of GLUTs at the plasma membrane. A number of metabolic inhibitors and AMPK activators acutely stimulate glucose uptake without increasing the amount of GLUT molecules at the plasma membrane (Abbud et al. 2000; Barnes et al. 2002; Hamrahian et al. 1999; Shetty et al. 1993; Shi et al. 1995). It was proposed that this increase is due to the release of “masking proteins,” which display an inhibitory interaction with the GLUT cytoplasmic domain under basal conditions (Shi et al. 1995). Stomatin was proposed being a masking protein as it interacts with the C-terminus of GLUT1, and its over-expression reduces GLUT1 intrinsic activity (Rungaldier et al. 2013; Zhang et al. 2001). Another potential masking protein is TXNIP, which displays an inhibitory interaction with GLUT1 (Wu et al. 2013). AMPK-induced degradation of TXNIP (see previous section) would unmask GLUT1 leading to enhancement of GLUT1-mediated glucose influx (Fig. 1a; arrow 3). Although stimulation of GLUT intrinsic activity is often observed during metabolic stress, the role of ROS in this pathway is still incompletely understood. In a leukemic cell line, ROS generated by NAD(P)H oxidase (Nox) stimulated glucose uptake via Src-mediated phosphorylation of GLUT1 (Prata et al. 2008). On the other hand, antioxidants were ineffective in preventing GLUT1 activation during azide-induced CIV inhibition (Hamrahian et al. 1999). Taken together, various signaling pathways have been implied in the stimulation of glucose uptake as an adaptive response during oxidative stress (Fig. 1a). At a relatively slow timescale, ROS-induced signals stimulate glucose uptake via up-regulation of GLUT protein expression. Rapid stimulation of glucose uptake can occur at the level of GLUT translocation and regulation of GLUT intrinsic activity. Interestingly, several of the signaling proteins are ROS-sensitive (i.e., AMPK and TXNIP) and involved in both slow and fast responses. This suggests that ROS-induced stimulation of glucose uptake is part of a (adaptive) mechanism triggered by oxidative and/or metabolic stress.

Effects of increased glucose uptake on ROS production and scavenging

The above evidence supports the conclusion that increased ROS levels stimulate cellular glucose uptake both at slow and fast time scales. However, inhibition of GLUT1 activity in myoblasts was paralleled by increased ROS levels (Andrisse et al. 2014). This might indicate that glucose uptake also plays a role in regulating the balance between ROS production and scavenging, suggesting that glucose uptake must be tightly controlled to maintain cellular energy homeostasis and redox status.

The role of glucose in ROS scavenging

Glucose entry into the PPP is protective against hydrogen peroxide-induced cytotoxicity (Le Goffe et al. 2002). Mechanistically, this protection is probably due to an increased PPP flux, leading to a higher NADPH/NADP+ ratio and GSH level. Compatible with this hypothesis, skin fibroblasts derived from patients with MERFF (myoclonic epilepsy with ragged red fibers) displayed GLUT1 up-regulation and increased NADPH and GSH levels (Wu and Wei 2012). Preventing this NADPH increase induced ROS over-production and cell death (Wu and Wei 2012). Glucose entry into the PPP and subsequently increased levels of NADPH is stimulated by ATM-induced activation of G6PDH (Fig. 1b; arrow 2), which is the first and rate determining enzyme of the PPP (Cosentino et al. 2011). Moreover, GLUT1 contributes to ROS scavenging by mediating the transport of dehydroascorbic acid (DHA), the oxidized form of vitamin C, which is recycled back to vitamin C inside the cell (Rumsey et al. 1997). Vitamin C is a potent antioxidant, which can prevent oxidative cell death (De Rosa et al. 2010; Guaiquil et al. 2001). Experimental evidence suggests that GLUT1 also co-localizes with mitochondria to facilitate mitochondrial uptake of DHA and quench ROS induced by mitochondrial uncoupling (Kc et al. 2005). In summary, enhanced PPP glucose entry and GLUT-mediated antioxidant uptake appear to be relevant for cellular ROS removal.

The role of glucose in stimulation of ROS production

A high glycolytic flux was associated with increased ROS levels (Talior et al. 2003; Zhou et al. 2005), whereas inhibition of mitochondrial pyruvate uptake lowered ROS levels (Nishikawa et al. 2000; Yu et al. 2006). Increased glycolytic flux is generally associated with and increased TCA cycle flux (Ishihara et al. 1996). The latter results in accumulation of OXPHOS substrates and elevated NADH/NAD+ and FADH2/FAD ratios (Ido 2007; Ying 2008). A high level of mitochondrial NADH leads to a fully reduced FMN site in CI, potentially stimulating superoxide production (Kussmaul and Hirst 2006). Similarly, a more reduced ETC might stimulate superoxide formation by CIII (Turrens et al. 1985). Reduced ubiquinone, in combination with a highly negative (hyperpolarized) Δψ, favors reverse electron transfer from CII to CI, which also stimulates superoxide production (Batandier et al. 2006; Murphy 2009). Hyperglycemic conditions induced mitochondrial fragmentation in clone 9 liver cells, H9c2 cardiomyoblasts, and smooth muscle cells, which was strictly required to allow increased ROS production (Yu et al. 2011). However, induction of mitochondrial fragmentation by over-expression of the fission-promoting protein Drp1 (Dynamin-related protein 1), did not stimulate ROS levels in HeLa cells, suggesting that increased ROS levels are not a de facto consequence of mitochondrial fragmentation (Distelmaier et al. 2012). It was proposed that fragmented mitochondria might produce more ROS due to a bigger relative membrane surface (allowing better uptake of metabolic substrates) and ensuing Δψ hyperpolarization (Yu et al. 2006). High extracellular glucose levels stimulate TXNIP expression (Stoltzman et al. 2008), which can aggravate oxidative stress by binding Trx via disulfide bridges and thereby inhibiting its reducing potential (Hwang et al. 2014; Kaimul et al. 2007; Li et al. 2015; Nishiyama et al. 1999; Schulze et al. 2004). Taken together, the current experimental evidence suggests that increased glucose uptake and glycolytic conversion to pyruvate can increase ROS levels.

Cellular and metabolic adaptation to increased ROS levels

Integrating the above mechanisms, it is conceivable that certain (pathological) conditions favor activation of a self-amplifying cycle of glucose uptake and glucose-stimulated ROS production, ultimately leading to cell death. Glucose-stimulated ROS production might be counterbalanced by the combined action of endogenous antioxidant systems (Sect. 1.2) and glucose-stimulated increase in ROS scavenging (Sect. 3.1). If these systems are insufficient, cells might also prevent glucose-induced oxidative stress by other means. As discussed in Sect. 3.2, a very high glycolytic flux eventually results in increased ETC electron input, a highly negative (hyperpolarized) Δψ, and a (probably) more reduced ETC. These phenomena all favor mitochondrial ROS production by the ETC. As discussed in the following sections, several mechanisms have been described that reduce ETC-mediated ROS generation. The latter generally are associated with a less reduced state of CI and CIII, likely associated with reduced electron leak and ROS generation, and include lowering the amount of electrons fed into the ETC and induction of (partial) Δψ depolarization.

Reducing the ETC electron input

In principle, ETC electron input can be reduced by lowering cellular glucose uptake. A high extracellular glucose concentration triggers a rapid reduction in GLUT1 levels at the plasma membrane, without affecting their total cellular levels (Greco-Perotto et al. 1992; Sasson et al. 1997). Such a reduction can be induced by inhibition of GLUT translocation to the plasma membrane, stimulation of GLUT internalization or both (Fig. 1c; process 1). On a slower time scale, down-regulation of GLUT expression (Fig. 1c; process 2) also lowers the total and plasma membrane levels of GLUT (Riahi et al. 2010; Totary-Jain et al. 2005). Regulation of these two processes is mediated by various signaling pathways. First, a high rate of glucose uptake induces the buildup of glycolytic intermediates, which activate the transcription factor carbohydrate response element-binding protein (chREBP) to drive the expression of a number of target genes, including TXNIP, that negatively regulate glycolysis (Stoltzman et al. 2008). TXNIP over-expression inhibits glucose uptake, while TXNIP knockdown stimulates glucose uptake (Parikh et al. 2007). TXNIP inhibits glucose uptake by interacting with GLUT1 and possibly inducing GLUT1 internalization through clathrin-coated pits (Fig. 1c; process 1). In addition, TXNIP reduces glucose uptake by suppressing GLUT1 mRNA levels (Fig. 1c; process 2; Wu et al. 2013). A TXNIP mutant unable to bind Trx still inhibited glucose uptake (Parikh et al. 2007; Patwari et al. 2009), suggesting that TXNIP-induced down-regulation of glucose uptake does not exclusively depend on Trx (and possibly redox status). Besides having a stimulatory effect on glucose uptake, ROS also exert a negative feedback on glucose uptake. For instance, exogenous application of ROS induced GLUT1 internalization and reduced glucose uptake (Fig. 1c; process 1) in retinal endothelial cells (Fernandes et al. 2004, 2011). Similarly, a prolonged exposure to exogenous ROS triggered Akt inactivation and GLUT1 internalization (Fernandes et al. 2011). In contrast, a short exposure to exogenous ROS stimulated Akt activation and translocation of GLUT1 to the plasma membrane (Fernandes et al. 2011). The above suggests a mechanism in which ROS first transiently stimulates glucose uptake (perhaps to increase NADPH levels) and subsequently down-regulates glucose uptake to prevent that too much electrons are fed into the ETC. However, internalization of GLUT1 does not always occur efficiently and high glucose-induced ROS production is prolonged (Cohen et al. 2007; Rosa et al. 2009). The exact reason for the absence of GLUT1 reduction at the plasma membrane is not well understood. It is possible that regulatory proteins such as Akt are bypassed. It may also be a consequence of irreversible damage (induced by ROS) of regulatory proteins, such as those comprising the proteasome machinery. In muscle, increased ROS levels during enhanced glucose uptake reduce the level of GLUT1 mRNA by increasing the expression of 12-lipoxygenase. This enzyme converts arachidonic acid to 12-hydroperoxyeicosatetraenoic acid (12-HPETE) (Alpert et al. 2002) and ROS-induced oxidation of the latter leads to formation of 4-hydroxydodecadienal (4-HDDE). This molecule activates the peroxisome proliferator-activated receptor δ (PPARδ) to drive the expression of calreticulin (Riahi et al. 2010). On its turn, calreticulin interacts with a cis-acting element in the 3′UTR of GLUT1 mRNA, making it susceptible to degradation (Totary-Jain et al. 2005) (Fig. 1c; process 2).

Inhibition of lactate dehydrogenase (LDH) enhanced oxidative stress and cell death in tumor cells (Le et al. 2010), whereas stimulating pyruvate-to-lactate conversion reduced oxidative stress (Brand 1997). This suggests that ETC electron input is reduced by stimulating conversion of pyruvate into lactate and secretion of the latter by the cell into the extracellular environment (Fig. 1c; process 3). Another mechanism to reduced ETC electron input is shown by the fact that ROS can stimulate glutathionylation or sulfenation, and thereby inactivation, of several TCA cycle enzymes including PDH, KGDHC, aconitase, isocitrate dehydrogenase, and CII (Bulteau et al. 2005; Chen et al. 2007b; Kil and Park 2005; McLain et al. 2011, 2013; Yan et al. 2013). Although ROS-induced inhibition of the TCA cycle results in a decline of NADH and in turn diminish the mitochondrial electron feed and hence reduced state of CI, it may also be detrimental to cells (Tretter and Adam-Vizi 2000). Increased ROS levels can also reduce the glycolytic flux by activation of poly(ADP-ribose) polymerase (PARP), leading to subsequent inactivation of glyceraldehyde-3-phosphate dehydrogenase (GAPDH) (Du et al. 2003; Nishikawa et al. 2000). Although this mechanism might be useful to lower ETC electron input, it also makes glucose enter the polyol pathway, which consumes NADPH and generates NADH (Giacco and Brownlee 2010). This is unfavorable since NADPH depletion and increased NADH/NAD+ ratios are associated with increased ROS production and/or levels.

Depolarization of Δψ

The magnitude of Δψ is mainly determined by the gradient of protons across the MIM, but also other charged ions and small molecules (e.g., ATP3−, ADP4−) can play a role. Therefore, every mechanism that reduces proton influx or increases proton efflux into the MIM, when not counterbalanced, stimulates (partial) Δψ depolarization. In this sense, Δψ depolarization can be induced by ETC inhibition, CV stimulation (increased coupling efficiency) and/or increased trans-MIM proton leak (uncoupling). The latter is mediated by mitochondrial uncoupling proteins (UCPs) 1-3 (Fig. 1c; process 4, UCP). In mitochondrial fractions from various tissues, GDP-induced inhibition of UCP1 and UCP2 was associated with Δψ hyperpolarization and increased production of hydrogen peroxide (Negre-Salvayre et al. 1997). Similarly, in muscle mitochondria and intact muscle fibers inhibition of UCP3 by GDP-stimulated superoxide and hydrogen peroxide production, respectively (Anderson et al. 2007; Talbot et al. 2004). Also genetic intervention studies revealed that absence of either UCP2 or UCP3 resulted in higher ROS production and oxidative stress in mitochondria and cells (Anderson et al. 2007; Arsenijevic et al. 2000; Brand et al. 2002; Lee et al. 2009; McLeod et al. 2005; Seifert et al. 2008; Toime and Brand 2010; Valsecchi et al. 2013). In line with these results, UCP2 or UCP3 over-expression lowered mitochondrial ROS production (Lee et al. 2005; Teshima et al. 2003; Valsecchi et al. 2013). Surprisingly, ROS can activate both UCP2 and UCP3 via deglutathionylation rather than glutathionylation, although the latter is normally stimulated by high ROS levels (Mailloux et al. 2011). In this sense, high levels of ROS activate UCPs and thereby constituting a negative feedback loop that lowers mitochondrial ROS production presumably via mitochondrial uncoupling and ensuing Δψ depolarization. In addition to the role of UCPs in controlling mitochondrial ROS production via Δψ depolarization, also a role for these proteins in metabolism has been proposed (Bouillaud 2009; Huppertz et al. 2001). This suggests that UCP-mediated metabolic changes might (co)determine mitochondrial ROS generation.

Fast Δψ depolarization can also be induced by opening of the mitochondrial permeability transition pore (PTP) (Fig. 1; process 4, PTP). Although the mitochondrial matrix protein Cyclophilin D (CypD) is a well-characterized regulator of PTP opening, the exact molecular composition of this channel remains elusive. Genetic intervention studies revealed that the adenine nucleotide translocase (ANT) and the mitochondrial phosphate carrier (PiC) are not core components but also regulators of PTP opening (Gutierrez-Aguilar et al. 2014; Kokoszka et al. 2004; Kwong et al. 2014; Varanyuwatana and Halestrap 2012). More recently, a central role for mitochondrial CV has been proposed (Alavian et al. 2014; Bonora et al. 2013; Giorgio et al. 2013). However, it may well be that an interaction of the CV with ANT and PiC is the mechanism of PTP formation (Halestrap 2014). Most of the above proteins have been shown to be prone to ROS-induced modification. The ANT contains several thiol residues that are sensitive to redox modification and involved in regulating PTP opening (Costantini et al. 2000; McStay et al. 2002; Queiroga et al. 2010). Upon diamide-induced oxidative stress, Cys160 cross-links to Cys257 thereby locking the ANT in its “c-conformation,” which sensitizes the PTP to elevated calcium levels within the mitochondrial matrix (McStay et al. 2002). Carbon monoxide is able to induce glutathionylation of ANT and decreasing PTP opening, while in turn diamide is able to deglutathionylate ANT (Martinez-Reyes and Cuezva 2014). Cys203 of CypD is another target for oxidative modification, shown to be involved in ROS-induced PTP opening and cell death. Upon Cys203 mutation, PTP opening is reduced to a level similar to that in CypD-negative cells (Linard et al. 2009; Nguyen et al. 2011). The α-subunit of the CV can also be glutathionylated leading to a decrease in CV activity (Garcia et al. 2010). Whether this decrease reduces PTP opening still needs to be elucidated. Although PTP opening might reduce the mitochondrial ROS levels by lowering ROS production and/or allow the release of mitochondrial ROS into the cytosol, this phenomenon has also been associated with induction of superoxide “flashes” (Wang et al. 2008; Zhang et al. 2013). However, the existence and chemical nature of these flashes is currently debated as the biosensor used for superoxide detection also appears to be pH-sensitive (Muller 2009; Schwarzlander et al. 2012, 2014). More importantly, sustained PTP opening triggers apoptosis and has been linked to pathophysiology and cell death (Brenner and Moulin 2012; Ichas and Mazat 1998). This suggests that only short reversible PTP openings, associated with reversible Δψ depolarizations (Blanchet et al. 2014), are suited to reduce mitochondrial ROS levels and preserve cell viability.

Other mechanisms that reduce ROS

Increased coupling efficiency of the OXPHOS system (i.e., between ETC electron transport and CV-mediated ATP production) has also been described as a mechanism to depolarize Δψ and lower ROS production (Fig. 1; process 4, HK/CV) (Starkov and Fiskum 2003). Recruitment of hexokinase (HK) to the outer surface of the mitochondrial outer membrane (MOM) was associated with decreased hydrogen peroxide production (Sun et al. 2008). A mechanism was proposed in which HK uses OXPHOS-derived ATP to metabolize glucose and form ADP. The latter is exchanged with ATP across the MIM by the ANT and used to fuel ATP production by CV. In principle, such a mechanism would increase coupled respiration and thereby reduce ETC electron leak and ROS production. However, glucose-6-phosphate accumulation is stimulated under conditions of increased glycolytic flux. This accumulation might inhibit HK activity and thereby increase mitochondrial ROS production (da-Silva et al. 2004). As increased HK-mediated ADP cycling enhances ETC electron transport, this mechanism should also result in a less reduced state of the FMN site of CI and thereby might decrease mitochondrial ROS formation. The latter could also be achieved by lowering NADH production (Sect. 4.1) or by decreasing OXPHOS activity. In case of CII and CV, their glutathionylation has been associated with reduced activity (Chen et al. 2007b, 2008; Garcia et al. 2010). CI also contains subunits that are sensitive to oxidative modification. These include the 51-kDa (NDUFV1) subunit (Cys187, Cys206, Cys425) and 75-kDa (NDUFS1) subunit (Cys367 Cys531 Cys704 Cys226 Cys727), which can be glutathionylated and thereby diminish the activity of CI (Beer et al. 2004; Chen et al. 2007a; Hurd et al. 2008; Kang et al. 2012). NDUFV1 and NDUFS1 glutathionylation is reversible and protects CI from further oxidative damage such as sulfenylation and thereby irreversible deactivation (Hurd et al. 2008). Compatible with inhibitor studies (Koopman et al. 2010), CI inhibition by glutathionylation was linked to increased superoxide production (Taylor et al. 2003). This suggests that CI inactivation is not necessarily associated with reduced mitochondrial ROS production. Taken together, the above suggests that oxidative stress triggered by increased glucose uptake could trigger adaptive responses to reduce mitochondrial ROS production via reducing ETC electron input, depolarization of Δψ or increasing coupled respiration.

Summary and conclusion

ROS are produced as a consequence of normal mitochondrial energy metabolism. When transiently and/or moderately increased, ROS can activate signaling pathways involved in cellular adaptation to various types of (metabolic) stress. One of these pathways is the stimulation of glucose uptake. When ROS levels are too high and/or remain increased during a prolonged period of time, a vicious circle of ROS-stimulated glucose uptake and glucose-stimulated ROS production can be triggered. This pathological cycle can be broken by restoring mitochondrial ROS production to normal levels. We presented three major mechanisms that, in principle, can lower mitochondrial ROS production: (1) reducing glucose uptake, (2) increasing lactate secretion and (3) depolarization of Δψ. Unfortunately, these mechanisms have also been associated with increases in ROS and/or appear to be not effective in all experimental models. Undesirable side effects include reduced NADPH production during reduced glucose uptake, a high rate of lactate secretion potentially inducing lactic acidosis and induction of mitochondrial dysfunction and apoptosis by (high-magnitude) and/or prolonged Δψ depolarization. We conclude that cellular glucose metabolism and mitochondrial ROS production are coupled by various signaling mechanisms, which need to be controlled by the cell to avoid oxidative stress. A more detailed understanding of how these pathways interact with mitochondrial ROS production, endogenous antioxidant systems and mitochondrial/cellular function is required to explain why oxidative stress induction still appears to contribute to pathology induction in humans (e.g., diabetes, cancer, mitochondrial dysfunction).



This work was supported by the CSBR (Centres for Systems Biology Research) initiative from ZonMW (Grant: #CSBR09/013 V).

Conflict of interest

The authors declare that they have no conflict of interest.


  1. Abbud W, Habinowski S, Zhang JZ et al (2000) Stimulation of AMP-activated protein kinase (AMPK) is associated with enhancement of Glut1-mediated glucose transport. Arch Biochem Biophys 380(2):347–352. doi: 10.1006/abbi.2000.1935 PubMedGoogle Scholar
  2. Adam-Vizi V, Chinopoulos C (2006) Bioenergetics and the formation of mitochondrial reactive oxygen species. Trends Pharmacol Sci 27(12):639–645. doi: 10.1016/ PubMedGoogle Scholar
  3. Afanas’ev I (2011) Reactive oxygen species signaling in cancer: comparison with aging. Aging Dis 2(3):219–230PubMedCentralPubMedGoogle Scholar
  4. Alavian KN, Beutner G, Lazrove E et al (2014) An uncoupling channel within the c-subunit ring of the F1FO ATP synthase is the mitochondrial permeability transition pore. Proc Natl Acad Sci USA 111(29):10580–10585. doi: 10.1073/pnas.1401591111 PubMedCentralPubMedGoogle Scholar
  5. Alexander A, Walker CL (2010) Differential localization of ATM is correlated with activation of distinct downstream signaling pathways. Cell Cycle 9(18):3685–3686PubMedCentralPubMedGoogle Scholar
  6. Alpert E, Gruzman A, Totary H, Kaiser N, Reich R, Sasson S (2002) A natural protective mechanism against hyperglycaemia in vascular endothelial and smooth-muscle cells: role of glucose and 12-hydroxyeicosatetraenoic acid. Biochem J 362(Pt 2):413–422PubMedCentralPubMedGoogle Scholar
  7. Anderson EJ, Yamazaki H, Neufer PD (2007) Induction of endogenous uncoupling protein 3 suppresses mitochondrial oxidant emission during fatty acid-supported respiration. J Biol Chem 282(43):31257–31266. doi: 10.1074/jbc.M706129200 PubMedGoogle Scholar
  8. Andrisse S, Patel GD, Chen JE et al (2013) ATM and GLUT1-S490 phosphorylation regulate GLUT1 mediated transport in skeletal muscle. PLoS One 8(6):e66027. doi: 10.1371/journal.pone.0066027 PubMedCentralPubMedGoogle Scholar
  9. Andrisse S, Koehler RM, Chen JE et al (2014) Role of GLUT1 in regulation of reactive oxygen species. Redox Biol 2:764–771. doi: 10.1016/j.redox.2014.03.004 PubMedCentralPubMedGoogle Scholar
  10. Arner ES (2009) Focus on mammalian thioredoxin reductases–important selenoproteins with versatile functions. Biochim Biophys Acta 1790(6):495–526. doi: 10.1016/j.bbagen.2009.01.014 PubMedGoogle Scholar
  11. Arsenijevic D, Onuma H, Pecqueur C et al (2000) Disruption of the uncoupling protein-2 gene in mice reveals a role in immunity and reactive oxygen species production. Nat Genet 26(4):435–439. doi: 10.1038/82565 PubMedGoogle Scholar
  12. Asano T, Katagiri H, Takata K et al (1991) The role of N-glycosylation of GLUT1 for glucose transport activity. J Biol Chem 266(36):24632–24636PubMedGoogle Scholar
  13. Baer SC, Casaubon L, Younes M (1997) Expression of the human erythrocyte glucose transporter Glut1 in cutaneous neoplasia. J Am Acad Dermatol 37(4):575–577PubMedGoogle Scholar
  14. Barnes K, Ingram JC, Porras OH et al (2002) Activation of GLUT1 by metabolic and osmotic stress: potential involvement of AMP-activated protein kinase (AMPK). J Cell Sci 115(Pt 11):2433–2442PubMedGoogle Scholar
  15. Batandier C, Guigas B, Detaille D et al (2006) The ROS production induced by a reverse-electron flux at respiratory-chain complex 1 is hampered by metformin. J Bioenerg Biomembr 38(1):33–42. doi: 10.1007/s10863-006-9003-8 PubMedGoogle Scholar
  16. Beer SM, Taylor ER, Brown SE et al (2004) Glutaredoxin 2 catalyzes the reversible oxidation and glutathionylation of mitochondrial membrane thiol proteins: implications for mitochondrial redox regulation and antioxidant defense. J Biol Chem 279(46):47939–47951. doi: 10.1074/jbc.M408011200 PubMedGoogle Scholar
  17. Bell GI, Kayano T, Buse JB, Burant CF, Takeda J, Lin D, Fukumoto H, Seino S (1990) Molecular biology of mammalian glucose transporters. Diabetes Care 13(3):198–208PubMedGoogle Scholar
  18. Beltran B, Orsi A, Clementi E, Moncada S (2000) Oxidative stress and S-nitrosylation of proteins in cells. Br J Pharmacol 129(5):953–960. doi: 10.1038/sj.bjp.0703147 PubMedCentralPubMedGoogle Scholar
  19. Blanchet L, Grefte S, Smeitink JA, Willems PH, Koopman WJ (2014) Photo-induction and automated quantification of reversible mitochondrial permeability transition pore opening in primary mouse myotubes. PLoS One 9(11):e114090. doi: 10.1371/journal.pone.0114090 PubMedCentralPubMedGoogle Scholar
  20. Bonora M, Bononi A, De Marchi E et al (2013) Role of the c subunit of the FO ATP synthase in mitochondrial permeability transition. Cell Cycle 12(4):674–683. doi: 10.4161/cc.23599 PubMedCentralPubMedGoogle Scholar
  21. Bouillaud F (2009) UCP2, not a physiologically relevant uncoupler but a glucose sparing switch impacting ROS production and glucose sensing. Biochim Biophys Acta 1787(5):377–383. doi: 10.1016/j.bbabio.2009.01.003 PubMedGoogle Scholar
  22. Boveris A, Valdez LB, Zaobornyj T, Bustamante J (2006) Mitochondrial metabolic states regulate nitric oxide and hydrogen peroxide diffusion to the cytosol. Biochim Biophys Acta 1757(5–6):535–542. doi: 10.1016/j.bbabio.2006.02.010 PubMedGoogle Scholar
  23. Brand K (1997) Aerobic glycolysis by proliferating cells: protection against oxidative stress at the expense of energy yield. J Bioenerg Biomembr 29(4):355–364PubMedGoogle Scholar
  24. Brand MD, Pamplona R, Portero-Otin M et al (2002) Oxidative damage and phospholipid fatty acyl composition in skeletal muscle mitochondria from mice underexpressing or overexpressing uncoupling protein 3. Biochem J 368(Pt 2):597–603. doi: 10.1042/BJ20021077 PubMedCentralPubMedGoogle Scholar
  25. Brennan JP, Wait R, Begum S, Bell JR, Dunn MJ, Eaton P (2004) Detection and mapping of widespread intermolecular protein disulfide formation during cardiac oxidative stress using proteomics with diagonal electrophoresis. J Biol Chem 279(40):41352–41360. doi: 10.1074/jbc.M403827200 PubMedGoogle Scholar
  26. Brenner C, Moulin M (2012) Physiological roles of the permeability transition pore. Circ Res 111(9):1237–1247. doi: 10.1161/CIRCRESAHA.112.265942 PubMedGoogle Scholar
  27. Brown RS, Wahl RL (1993) Overexpression of Glut-1 glucose transporter in human breast cancer. An immunohistochemical study. Cancer 72(10):2979–2985PubMedGoogle Scholar
  28. Brunelle JK, Bell EL, Quesada NM et al (2005) Oxygen sensing requires mitochondrial ROS but not oxidative phosphorylation. Cell Metab 1(6):409–414. doi: 10.1016/j.cmet.2005.05.002 PubMedGoogle Scholar
  29. Bulteau AL, Lundberg KC, Ikeda-Saito M, Isaya G, Szweda LI (2005) Reversible redox-dependent modulation of mitochondrial aconitase and proteolytic activity during in vivo cardiac ischemia/reperfusion. Proc Natl Acad Sci USA 102(17):5987–5991. doi: 10.1073/pnas.0501519102 PubMedCentralPubMedGoogle Scholar
  30. Bunik VI, Sievers C (2002) Inactivation of the 2-oxo acid dehydrogenase complexes upon generation of intrinsic radical species. Eur J Biochem/FEBS 269(20):5004–5015Google Scholar
  31. Carballal S, Radi R, Kirk MC, Barnes S, Freeman BA, Alvarez B (2003) Sulfenic acid formation in human serum albumin by hydrogen peroxide and peroxynitrite. Biochemistry 42(33):9906–9914. doi: 10.1021/bi027434m PubMedGoogle Scholar
  32. Carlberg I, Mannervik B (1985) Glutathione reductase. Methods Enzymol 113:484–490PubMedGoogle Scholar
  33. Carruthers A, DeZutter J, Ganguly A, Devaskar SU (2009) Will the original glucose transporter isoform please stand up! Am J Physiol Endocrinol Metab 297(4):E836–E848. doi: 10.1152/ajpendo.00496.2009 PubMedCentralPubMedGoogle Scholar
  34. Chae HZ, Kim HJ, Kang SW, Rhee SG (1999) Characterization of three isoforms of mammalian peroxiredoxin that reduce peroxides in the presence of thioredoxin. Diabetes Res Clin Pract 45(2–3):101–112PubMedGoogle Scholar
  35. Chambers MA, Moylan JS, Smith JD, Goodyear LJ, Reid MB (2009) Stretch-stimulated glucose uptake in skeletal muscle is mediated by reactive oxygen species and p38 MAP-kinase. J Physiol 587(Pt 13):3363–3373. doi: 10.1113/jphysiol.2008.165639 PubMedCentralPubMedGoogle Scholar
  36. Chandel NS, Maltepe E, Goldwasser E, Mathieu CE, Simon MC, Schumacker PT (1998) Mitochondrial reactive oxygen species trigger hypoxia-induced transcription. Proc Natl Acad Sci USA 95(20):11715–11720PubMedCentralPubMedGoogle Scholar
  37. Chandel NS, McClintock DS, Feliciano CE et al (2000) Reactive oxygen species generated at mitochondrial complex III stabilize hypoxia-inducible factor-1alpha during hypoxia: a mechanism of O2 sensing. J Biol Chem 275(33):25130–25138. doi: 10.1074/jbc.M001914200 PubMedGoogle Scholar
  38. Chang TS, Cho CS, Park S, Yu S, Kang SW, Rhee SG (2004) Peroxiredoxin III, a mitochondrion-specific peroxidase, regulates apoptotic signaling by mitochondria. J Biol Chem 279(40):41975–41984. doi: 10.1074/jbc.M407707200 PubMedGoogle Scholar
  39. Charles RL, Schroder E, May G et al (2007) Protein sulfenation as a redox sensor: proteomics studies using a novel biotinylated dimedone analogue. Mol Cell Proteomics 6(9):1473–1484. doi: 10.1074/mcp.M700065-MCP200 PubMedGoogle Scholar
  40. Chen C, Pore N, Behrooz A, Ismail-Beigi F, Maity A (2001) Regulation of glut1 mRNA by hypoxia-inducible factor-1. Interaction between H-ras and hypoxia. J Biol Chem 276(12):9519–9525. doi: 10.1074/jbc.M010144200 PubMedGoogle Scholar
  41. Chen CL, Zhang L, Yeh A et al (2007a) Site-specific S-glutathiolation of mitochondrial NADH ubiquinone reductase. Biochemistry 46(19):5754–5765. doi: 10.1021/bi602580c PubMedCentralPubMedGoogle Scholar
  42. Chen YR, Chen CL, Pfeiffer DR, Zweier JL (2007b) Mitochondrial complex II in the post-ischemic heart: oxidative injury and the role of protein S-glutathionylation. J Biol Chem 282(45):32640–32654. doi: 10.1074/jbc.M702294200 PubMedGoogle Scholar
  43. Chen CL, Chen J, Rawale S et al (2008) Protein tyrosine nitration of the flavin subunit is associated with oxidative modification of mitochondrial complex II in the post-ischemic myocardium. J Biol Chem 283(41):27991–28003. doi: 10.1074/jbc.M802691200 PubMedCentralPubMedGoogle Scholar
  44. Cohen G, Riahi Y, Alpert E, Gruzman A, Sasson S (2007) The roles of hyperglycaemia and oxidative stress in the rise and collapse of the natural protective mechanism against vascular endothelial cell dysfunction in diabetes. Arch Physiol Biochem 113(4–5):259–267. doi: 10.1080/13813450701783513 PubMedGoogle Scholar
  45. Cosentino C, Grieco D, Costanzo V (2011) ATM activates the pentose phosphate pathway promoting anti-oxidant defence and DNA repair. EMBO J 30(3):546–555. doi: 10.1038/emboj.2010.330 PubMedCentralPubMedGoogle Scholar
  46. Costantini P, Belzacq AS, Vieira HL et al (2000) Oxidation of a critical thiol residue of the adenine nucleotide translocator enforces Bcl-2-independent permeability transition pore opening and apoptosis. Oncogene 19(2):307–314. doi: 10.1038/sj.onc.1203299 PubMedGoogle Scholar
  47. Cox AG, Winterbourn CC, Hampton MB (2010) Mitochondrial peroxiredoxin involvement in antioxidant defence and redox signalling. Biochem J 425(2):313–325. doi: 10.1042/BJ20091541 Google Scholar
  48. da-Silva WS, Gomez-Puyou A, de Gomez-Puyou MT et al (2004) Mitochondrial bound hexokinase activity as a preventive antioxidant defense: steady-state ADP formation as a regulatory mechanism of membrane potential and reactive oxygen species generation in mitochondria. J Biol Chem 279(38):39846–39855. doi: 10.1074/jbc.M403835200 PubMedGoogle Scholar
  49. Davies SP, Helps NR, Cohen PT, Hardie DG (1995) 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 Lett 377(3):421–425. doi: 10.1016/0014-5793(95)01368-7 PubMedGoogle Scholar
  50. De Rosa S, Cirillo P, Paglia A, Sasso L, Di Palma V, Chiariello M (2010) Reactive oxygen species and antioxidants in the pathophysiology of cardiovascular disease: does the actual knowledge justify a clinical approach? Curr Vasc Pharmacol 8(2):259–275PubMedGoogle Scholar
  51. Delaunay A, Pflieger D, Barrault MB, Vinh J, Toledano MB (2002) A thiol peroxidase is an H2O2 receptor and redox-transducer in gene activation. Cell 111(4):471–481PubMedGoogle Scholar
  52. Di Lisa F, Kaludercic N, Carpi A, Menabo R, Giorgio M (2009) Mitochondrial pathways for ROS formation and myocardial injury: the relevance of p66(Shc) and monoamine oxidase. Basic Res Cardiol 104(2):131–139. doi: 10.1007/s00395-009-0008-4 PubMedGoogle Scholar
  53. Distelmaier F, Valsecchi F, Forkink M et al (2012) Trolox-sensitive reactive oxygen species regulate mitochondrial morphology, oxidative phosphorylation and cytosolic calcium handling in healthy cells. Antioxid Redox Signal 17(12):1657–1669. doi: 10.1089/ars.2011.4294 PubMedCentralPubMedGoogle Scholar
  54. Du X, Matsumura T, Edelstein D et al (2003) Inhibition of GAPDH activity by poly(ADP-ribose) polymerase activates three major pathways of hyperglycemic damage in endothelial cells. J Clin Invest 112(7):1049–1057. doi: 10.1172/JCI18127 PubMedCentralPubMedGoogle Scholar
  55. Ebert BL, Firth JD, Ratcliffe PJ (1995) Hypoxia and mitochondrial inhibitors regulate expression of glucose transporter-1 via distinct Cis-acting sequences. J Biol Chem 270(49):29083–29089PubMedGoogle Scholar
  56. Eguez L, Lee A, Chavez JA et al (2005) Full intracellular retention of GLUT4 requires AS160 Rab GTPase activating protein. Cell Metab 2(4):263–272. doi: 10.1016/j.cmet.2005.09.005 PubMedGoogle Scholar
  57. Emerling BM, Platanias LC, Black E, Nebreda AR, Davis RJ, Chandel NS (2005) Mitochondrial reactive oxygen species activation of p38 mitogen-activated protein kinase is required for hypoxia signaling. Mol Cell Biol 25(12):4853–4862. doi: 10.1128/MCB.25.12.4853-4862.2005 PubMedCentralPubMedGoogle Scholar
  58. Emerling BM, Weinberg F, Snyder C et al (2009) Hypoxic activation of AMPK is dependent on mitochondrial ROS but independent of an increase in AMP/ATP ratio. Free Radic Biol Med 46(10):1386–1391. doi: 10.1016/j.freeradbiomed.2009.02.019 PubMedCentralPubMedGoogle Scholar
  59. Esposito LA, Kokoszka JE, Waymire KG, Cottrell B, MacGregor GR, Wallace DC (2000) Mitochondrial oxidative stress in mice lacking the glutathione peroxidase-1 gene. Free Radic Biol Med 28(5):754–766PubMedCentralPubMedGoogle Scholar
  60. Esworthy RS, Ho YS, Chu FF (1997) The Gpx1 gene encodes mitochondrial glutathione peroxidase in the mouse liver. Arch Biochem Biophys 340(1):59–63. doi: 10.1006/abbi.1997.9901 PubMedGoogle Scholar
  61. Fernandes R, Carvalho AL, Kumagai A et al (2004) Downregulation of retinal GLUT1 in diabetes by ubiquitinylation. Mol Vis 10:618–628PubMedGoogle Scholar
  62. Fernandes R, Hosoya K, Pereira P (2011) Reactive oxygen species downregulate glucose transport system in retinal endothelial cells. Am J Physiol Cell Physiol 300(4):C927–C936. doi: 10.1152/ajpcell.00140.2010 PubMedGoogle Scholar
  63. Fisher-Wellman KH, Gilliam LA, Lin CT, Cathey BL, Lark DS, Neufer PD (2013) Mitochondrial glutathione depletion reveals a novel role for the pyruvate dehydrogenase complex as a key H2O2-emitting source under conditions of nutrient overload. Free Radic Biol Med 65:1201–1208. doi: 10.1016/j.freeradbiomed.2013.09.008 PubMedCentralPubMedGoogle Scholar
  64. Forman HJ, Kennedy J (1975) Superoxide production and electron transport in mitochondrial oxidation of dihydroorotic acid. J Biol Chem 250(11):4322–4326PubMedGoogle Scholar
  65. Forman HJ, Maiorino M, Ursini F (2010) Signaling functions of reactive oxygen species. Biochemistry 49(5):835–842. doi: 10.1021/bi9020378 PubMedCentralPubMedGoogle Scholar
  66. Freeman H, Shimomura K, Cox RD, Ashcroft FM (2006) Nicotinamide nucleotide transhydrogenase: a link between insulin secretion, glucose metabolism and oxidative stress. Biochem Soc Trans 34(Pt 5):806–810. doi: 10.1042/BST0340806 PubMedGoogle Scholar
  67. Fryer LG, Foufelle F, Barnes K, Baldwin SA, Woods A, Carling D (2002) Characterization of the role of the AMP-activated protein kinase in the stimulation of glucose transport in skeletal muscle cells. Biochem J 363(Pt 1):167–174PubMedCentralPubMedGoogle Scholar
  68. Funaki M, Randhawa P, Janmey PA (2004) Separation of insulin signaling into distinct GLUT4 translocation and activation steps. Mol Cell Biol 24(17):7567–7577. doi: 10.1128/MCB.24.17.7567-7577.2004 PubMedCentralPubMedGoogle Scholar
  69. Galloway CA, Yoon Y (2012) Perspectives on: SGP symposium on mitochondrial physiology and medicine: what comes first, misshape or dysfunction? The view from metabolic excess. J Gen Physiol 139(6):455–463. doi: 10.1085/jgp.201210771 PubMedCentralPubMedGoogle Scholar
  70. Garcia J, Han D, Sancheti H, Yap LP, Kaplowitz N, Cadenas E (2010) Regulation of mitochondrial glutathione redox status and protein glutathionylation by respiratory substrates. J Biol Chem 285(51):39646–39654. doi: 10.1074/jbc.M110.164160 PubMedCentralPubMedGoogle Scholar
  71. Gardner PR (2002) Aconitase: sensitive target and measure of superoxide. Methods Enzymol 349:9–23PubMedGoogle Scholar
  72. Genova ML, Ventura B, Giuliano G et al (2001) The site of production of superoxide radical in mitochondrial Complex I is not a bound ubisemiquinone but presumably iron-sulfur cluster N2. FEBS Lett 505(3):364–368PubMedGoogle Scholar
  73. Geraghty KM, Chen S, Harthill JE et al (2007) Regulation of multisite phosphorylation and 14-3-3 binding of AS160 in response to IGF-1, EGF, PMA and AICAR. Biochem J 407(2):231–241. doi: 10.1042/BJ20070649 PubMedCentralPubMedGoogle Scholar
  74. Giacco F, Brownlee M (2010) Oxidative stress and diabetic complications. Circ Res 107(9):1058–1070. doi: 10.1161/CIRCRESAHA.110.223545 PubMedCentralPubMedGoogle Scholar
  75. Giorgio M, Migliaccio E, Orsini F et al (2005) Electron transfer between cytochrome c and p66Shc generates reactive oxygen species that trigger mitochondrial apoptosis. Cell 122(2):221–233. doi: 10.1016/j.cell.2005.05.011 PubMedGoogle Scholar
  76. Giorgio V, von Stockum S, Antoniel M et al (2013) Dimers of mitochondrial ATP synthase form the permeability transition pore. Proc Natl Acad Sci USA 110(15):5887–5892. doi: 10.1073/pnas.1217823110 PubMedCentralPubMedGoogle Scholar
  77. Gomez-Cabrera MC, Borras C, Pallardo FV, Sastre J, Ji LL, Vina J (2005) Decreasing xanthine oxidase-mediated oxidative stress prevents useful cellular adaptations to exercise in rats. J Physiol 567(Pt 1):113–120. doi: 10.1113/jphysiol.2004.080564 PubMedCentralPubMedGoogle Scholar
  78. Gould GW, Holman GD (1993) The glucose transporter family: structure, function and tissue-specific expression. Biochem J 295(Pt 2):329–341PubMedCentralPubMedGoogle Scholar
  79. Greco-Perotto R, Wertheimer E, Jeanrenaud B, Cerasi E, Sasson S (1992) Glucose regulates its transport in L8 myocytes by modulating cellular trafficking of the transporter GLUT-1. Biochem J 286(Pt 1):157–163PubMedCentralPubMedGoogle Scholar
  80. Grivennikova VG, Vinogradov AD (2013) Partitioning of superoxide and hydrogen peroxide production by mitochondrial respiratory complex I. Biochim Biophys Acta 1827(3):446–454. doi: 10.1016/j.bbabio.2013.01.002 PubMedGoogle Scholar
  81. Guaiquil VH, Vera JC, Golde DW (2001) Mechanism of vitamin C inhibition of cell death induced by oxidative stress in glutathione-depleted HL-60 cells. J Biol Chem 276(44):40955–40961. doi: 10.1074/jbc.M106878200 PubMedGoogle Scholar
  82. Guo Z, Kozlov S, Lavin MF, Person MD, Paull TT (2010) ATM activation by oxidative stress. Science 330(6003):517–521. doi: 10.1126/science.1192912 PubMedGoogle Scholar
  83. Gutierrez-Aguilar M, Douglas DL, Gibson AK, Domeier TL, Molkentin JD, Baines CP (2014) Genetic manipulation of the cardiac mitochondrial phosphate carrier does not affect permeability transition. J Mol Cell Cardiol 72:316–325. doi: 10.1016/j.yjmcc.2014.04.008 PubMedCentralPubMedGoogle Scholar
  84. Guzy RD, Schumacker PT (2006) Oxygen sensing by mitochondria at complex III: the paradox of increased reactive oxygen species during hypoxia. Exp Physiol 91(5):807–819. doi: 10.1113/expphysiol.2006.033506 PubMedGoogle Scholar
  85. Guzy RD, Hoyos B, Robin E et al (2005) Mitochondrial complex III is required for hypoxia-induced ROS production and cellular oxygen sensing. Cell Metab 1(6):401–408. doi: 10.1016/j.cmet.2005.05.001 PubMedGoogle Scholar
  86. Halestrap AP (2014) The c ring of the F1Fo ATP synthase FORMS the mitochondrial permeability transition pore: a critical appraisal. Front Oncol 4:234. doi: 10.3389/fonc.2014.00234 PubMedCentralPubMedGoogle Scholar
  87. Hamrahian AH, Zhang JZ, Elkhairi FS, Prasad R, Ismail-Beigi F (1999) Activation of Glut1 glucose transporter in response to inhibition of oxidative phosphorylation. Arch Biochem Biophys 368(2):375–379. doi: 10.1006/abbi.1999.1320 PubMedGoogle Scholar
  88. Handy DE, Loscalzo J (2012) Redox regulation of mitochondrial function. Antioxid Redox Signal 16(11):1323–1367. doi: 10.1089/ars.2011.4123 PubMedCentralPubMedGoogle Scholar
  89. Hardie DG (2011) Energy sensing by the AMP-activated protein kinase and its effects on muscle metabolism. Proc Nutr Soc 70(1):92–99. doi: 10.1017/S0029665110003915 PubMedGoogle Scholar
  90. Hardie DG, Ashford ML (2014) AMPK: regulating energy balance at the cellular and whole body levels. Physiology 29(2):99–107. doi: 10.1152/physiol.00050.2013 PubMedCentralPubMedGoogle Scholar
  91. Hatefi Y, Yamaguchi M (1996) Nicotinamide nucleotide transhydrogenase: a model for utilization of substrate binding energy for proton translocation. FASEB J 10(4):444–452PubMedGoogle Scholar
  92. Hauptmann N, Grimsby J, Shih JC, Cadenas E (1996) The metabolism of tyramine by monoamine oxidase A/B causes oxidative damage to mitochondrial DNA. Arch Biochem Biophys 335(2):295–304. doi: 10.1006/abbi.1996.0510 PubMedGoogle Scholar
  93. Hawley SA, Ross FA, Chevtzoff C et al (2010) Use of cells expressing gamma subunit variants to identify diverse mechanisms of AMPK activation. Cell Metab 11(6):554–565. doi: 10.1016/j.cmet.2010.04.001 PubMedCentralPubMedGoogle Scholar
  94. Hayashi M, Sakata M, Takeda T et al (2004) Induction of glucose transporter 1 expression through hypoxia-inducible factor 1alpha under hypoxic conditions in trophoblast-derived cells. J Endocrinol 183(1):145–154. doi: 10.1677/joe.1.05599 PubMedGoogle Scholar
  95. Herrero A, Barja G (2000) Localization of the site of oxygen radical generation inside the complex I of heart and nonsynaptic brain mammalian mitochondria. J Bioenerg Biomembr 32(6):609–615PubMedGoogle Scholar
  96. Higaki Y, Mikami T, Fujii N et al (2008) Oxidative stress stimulates skeletal muscle glucose uptake through a phosphatidylinositol 3-kinase-dependent pathway. Am J Physiol Endocrinol Metab 294(5):E889–E897. doi: 10.1152/ajpendo.00150.2007 PubMedCentralPubMedGoogle Scholar
  97. Hogan A, Heyner S, Charron MJ et al (1991) Glucose transporter gene expression in early mouse embryos. Development 113(1):363–372PubMedGoogle Scholar
  98. Hogg N (2002) The biochemistry and physiology of S-nitrosothiols. Annu Rev Pharmacol Toxicol 42:585–600. doi: 10.1146/annurev.pharmtox.42.092501.104328 PubMedGoogle Scholar
  99. Huppertz C, Fischer BM, Kim YB et al (2001) Uncoupling protein 3 (UCP3) stimulates glucose uptake in muscle cells through a phosphoinositide 3-kinase-dependent mechanism. J Biol Chem 276(16):12520–12529. doi: 10.1074/jbc.M011708200 PubMedGoogle Scholar
  100. Hurd TR, Requejo R, Filipovska A et al (2008) Complex I within oxidatively stressed bovine heart mitochondria is glutathionylated on Cys-531 and Cys-704 of the 75-kDa subunit: potential role of CYS residues in decreasing oxidative damage. J Biol Chem 283(36):24801–24815. doi: 10.1074/jbc.M803432200 PubMedCentralPubMedGoogle Scholar
  101. Hwang J, Suh HW, Jeon YH et al (2014) The structural basis for the negative regulation of thioredoxin by thioredoxin-interacting protein. Nat Commun 5:2958. doi: 10.1038/ncomms3958 PubMedCentralPubMedGoogle Scholar
  102. Ichas F, Mazat JP (1998) From calcium signaling to cell death: two conformations for the mitochondrial permeability transition pore. Switching from low- to high-conductance state. Biochim Biophys Acta 1366(1–2):33–50PubMedGoogle Scholar
  103. Ido Y (2007) Pyridine nucleotide redox abnormalities in diabetes. Antioxid Redox Signal 9(7):931–942. doi: 10.1089/ars.2007.1630 PubMedGoogle Scholar
  104. Ishihara H, Nakazaki M, Kanegae Y et al (1996) Effect of mitochondrial and/or cytosolic glycerol 3-phosphate dehydrogenase overexpression on glucose-stimulated insulin secretion from MIN6 and HIT cells. Diabetes 45(9):1238–1244PubMedGoogle Scholar
  105. Ivan M, Kondo K, Yang H et al (2001) HIFalpha targeted for VHL-mediated destruction by proline hydroxylation: implications for O2 sensing. Science 292(5516):464–468. doi: 10.1126/science.1059817 PubMedGoogle Scholar
  106. Iyer NV, Kotch LE, Agani F et al (1998) Cellular and developmental control of O2 homeostasis by hypoxia-inducible factor 1 alpha. Genes Dev 12(2):149–162PubMedCentralPubMedGoogle Scholar
  107. Jensen TE, Schjerling P, Viollet B, Wojtaszewski JF, Richter EA (2008) AMPK alpha1 activation is required for stimulation of glucose uptake by twitch contraction, but not by H2O2, in mouse skeletal muscle. PLoS One 3(5):e2102. doi: 10.1371/journal.pone.0002102 PubMedCentralPubMedGoogle Scholar
  108. Johnson JE Jr, Choksi K, Widger WR (2003) NADH-Ubiquinone oxidoreductase: substrate-dependent oxygen turnover to superoxide anion as a function of flavin mononucleotide. Mitochondrion 3(2):97–110. doi: 10.1016/S1567-7249(03)00084-9 PubMedGoogle Scholar
  109. Joost HG, Thorens B (2001) The extended GLUT-family of sugar/polyol transport facilitators: nomenclature, sequence characteristics, and potential function of its novel members (review). Mol Membr Biol 18(4):247–256PubMedGoogle Scholar
  110. Jung SN, Yang WK, Kim J et al (2008) Reactive oxygen species stabilize hypoxia-inducible factor-1 alpha protein and stimulate transcriptional activity via AMP-activated protein kinase in DU145 human prostate cancer cells. Carcinogenesis 29(4):713–721. doi: 10.1093/carcin/bgn032 PubMedGoogle Scholar
  111. Jung KH, Lee JH, Thien Quach CH et al (2013) Resveratrol suppresses cancer cell glucose uptake by targeting reactive oxygen species-mediated hypoxia-inducible factor-1alpha activation. J Nucl Med 54(12):2161–2167. doi: 10.2967/jnumed.112.115436 PubMedGoogle Scholar
  112. Kaimul AM, Nakamura H, Masutani H, Yodoi J (2007) Thioredoxin and thioredoxin-binding protein-2 in cancer and metabolic syndrome. Free Radic Biol Med 43(6):861–868. doi: 10.1016/j.freeradbiomed.2007.05.032 PubMedGoogle Scholar
  113. Kakhlon O, Cabantchik ZI (2002) The labile iron pool: characterization, measurement, and participation in cellular processes(1). Free Radic Biol Med 33(8):1037–1046PubMedGoogle Scholar
  114. Kang PT, Zhang L, Chen CL, Chen J, Green KB, Chen YR (2012) Protein thiyl radical mediates S-glutathionylation of complex I. Free Radic Biol Med 53(4):962–973. doi: 10.1016/j.freeradbiomed.2012.05.025 PubMedCentralPubMedGoogle Scholar
  115. Kc S, Carcamo JM, Golde DW (2005) Vitamin C enters mitochondria via facilitative glucose transporter 1 (Glut1) and confers mitochondrial protection against oxidative injury. FASEB J 19(12):1657–1667. doi: 10.1096/fj.05-4107com PubMedGoogle Scholar
  116. Kil IS, Park JW (2005) Regulation of mitochondrial NADP+-dependent isocitrate dehydrogenase activity by glutathionylation. J Biol Chem 280(11):10846–10854. doi: 10.1074/jbc.M411306200 PubMedGoogle Scholar
  117. Kim JS, Saengsirisuwan V, Sloniger JA, Teachey MK, Henriksen EJ (2006) Oxidant stress and skeletal muscle glucose transport: roles of insulin signaling and p38 MAPK. Free Radic Biol Med 41(5):818–824. doi: 10.1016/j.freeradbiomed.2006.05.031 PubMedGoogle Scholar
  118. Kohn AD, Barthel A, Kovacina KS et al (1998) Construction and characterization of a conditionally active version of the serine/threonine kinase Akt. J Biol Chem 273(19):11937–11943PubMedGoogle Scholar
  119. Kokoszka JE, Waymire KG, Levy SE et al (2004) The ADP/ATP translocator is not essential for the mitochondrial permeability transition pore. Nature 427(6973):461–465. doi: 10.1038/nature02229 PubMedCentralPubMedGoogle Scholar
  120. Koopman WJ, Verkaart S, Visch HJ et al (2007) Human NADH: ubiquinone oxidoreductase deficiency: radical changes in mitochondrial morphology? Am J Physiol Cell Physiol 293(1):C22–C29. doi: 10.1152/ajpcell.00194.2006 PubMedGoogle Scholar
  121. Koopman WJ, Nijtmans LG, Dieteren CE et al (2010) Mammalian mitochondrial complex I: biogenesis, regulation, and reactive oxygen species generation. Antioxid Redox Signal 12(12):1431–1470. doi: 10.1089/ars.2009.2743 PubMedGoogle Scholar
  122. Koopman WJ, Willems PH, Smeitink JA (2012) Monogenic mitochondrial disorders. N Engl J Med 366(12):1132–1141. doi: 10.1056/NEJMra1012478 PubMedGoogle Scholar
  123. Koopman WJ, Distelmaier F, Smeitink JA, Willems PH (2013) OXPHOS mutations and neurodegeneration. EMBO J 32(1):9–29. doi: 10.1038/emboj.2012.300 PubMedCentralPubMedGoogle Scholar
  124. Kozlovsky N, Rudich A, Potashnik R, Ebina Y, Murakami T, Bashan N (1997) Transcriptional activation of the Glut1 gene in response to oxidative stress in L6 myotubes. J Biol Chem 272(52):33367–33372PubMedGoogle Scholar
  125. Kramer DM, Roberts AG, Muller F, Cape J, Bowman MK (2004) Q-cycle bypass reactions at the Qo site of the cytochrome bc1 (and related) complexes. Methods Enzymol 382:21–45. doi: 10.1016/S0076-6879(04)82002-0 PubMedGoogle Scholar
  126. Kramer HF, Witczak CA, Fujii N et al (2006) Distinct signals regulate AS160 phosphorylation in response to insulin, AICAR, and contraction in mouse skeletal muscle. Diabetes 55(7):2067–2076. doi: 10.2337/db06-0150 PubMedGoogle Scholar
  127. Kumar B, Koul S, Khandrika L, Meacham RB, Koul HK (2008) Oxidative stress is inherent in prostate cancer cells and is required for aggressive phenotype. Cancer Res 68(6):1777–1785. doi: 10.1158/0008-5472.CAN-07-5259 PubMedGoogle Scholar
  128. Kurz EU, Douglas P, Lees-Miller SP (2004) Doxorubicin activates ATM-dependent phosphorylation of multiple downstream targets in part through the generation of reactive oxygen species. J Biol Chem 279(51):53272–53281. doi: 10.1074/jbc.M406879200 PubMedGoogle Scholar
  129. Kussmaul L, Hirst J (2006) The mechanism of superoxide production by NADH: ubiquinone oxidoreductase (complex I) from bovine heart mitochondria. Proc Natl Acad Sci USA 103(20):7607–7612. doi: 10.1073/pnas.0510977103 PubMedCentralPubMedGoogle Scholar
  130. Kwong JQ, Davis J, Baines CP et al (2014) Genetic deletion of the mitochondrial phosphate carrier desensitizes the mitochondrial permeability transition pore and causes cardiomyopathy. Cell Death Differ 21(8):1209–1217. doi: 10.1038/cdd.2014.36 PubMedGoogle Scholar
  131. Lambert AJ, Brand MD (2004) Inhibitors of the quinone-binding site allow rapid superoxide production from mitochondrial NADH: ubiquinone oxidoreductase (complex I). J Biol Chem 279(38):39414–39420. doi: 10.1074/jbc.M406576200 PubMedGoogle Scholar
  132. Larance M, Ramm G, Stockli J et al (2005) Characterization of the role of the Rab GTPase-activating protein AS160 in insulin-regulated GLUT4 trafficking. J Biol Chem 280(45):37803–37813. doi: 10.1074/jbc.M503897200 PubMedGoogle Scholar
  133. Le Goffe C, Vallette G, Charrier L et al (2002) Metabolic control of resistance of human epithelial cells to H2O2 and NO stresses. Biochem J 364(Pt 2):349–359. doi: 10.1042/BJ20011856 PubMedCentralPubMedGoogle Scholar
  134. Le A, Cooper CR, Gouw AM et al (2010) Inhibition of lactate dehydrogenase A induces oxidative stress and inhibits tumor progression. Proc Natl Acad Sci USA 107(5):2037–2042. doi: 10.1073/pnas.0914433107 PubMedCentralPubMedGoogle Scholar
  135. Lee KU, Lee IK, Han J et al (2005) Effects of recombinant adenovirus-mediated uncoupling protein 2 overexpression on endothelial function and apoptosis. Circ Res 96(11):1200–1207. doi: 10.1161/01.RES.0000170075.73039.5b PubMedGoogle Scholar
  136. Lee SC, Robson-Doucette CA, Wheeler MB (2009) Uncoupling protein 2 regulates reactive oxygen species formation in islets and influences susceptibility to diabetogenic action of streptozotocin. J Endocrinol 203(1):33–43. doi: 10.1677/JOE-09-0117 PubMedGoogle Scholar
  137. Levine KB, Cloherty EK, Hamill S, Carruthers A (2002) Molecular determinants of sugar transport regulation by ATP. Biochemistry 41(42):12629–12638PubMedGoogle Scholar
  138. Li L, Zhu K, Liu Y et al (2015) Targeting thioredoxin-1 with siRNA exacerbates oxidative stress injury after cerebral ischemia/reperfusion in rats. Neuroscience 284:815–823. doi: 10.1016/j.neuroscience.2014.10.066 PubMedGoogle Scholar
  139. Linard D, Kandlbinder A, Degand H, Morsomme P, Dietz KJ, Knoops B (2009) Redox characterization of human cyclophilin D: identification of a new mammalian mitochondrial redox sensor? Arch Biochem Biophys 491(1–2):39–45. doi: 10.1016/ PubMedGoogle Scholar
  140. Liochev SI, Fridovich I (1994) The role of O2.– in the production of HO.: in vitro and in vivo. Free Radic Biol Med 16(1):29–33PubMedGoogle Scholar
  141. Mailloux RJ, Seifert EL, Bouillaud F, Aguer C, Collins S, Harper ME (2011) Glutathionylation acts as a control switch for uncoupling proteins UCP2 and UCP3. J Biol Chem 286(24):21865–21875. doi: 10.1074/jbc.M111.240242 PubMedCentralPubMedGoogle Scholar
  142. Mailloux RJ, Jin X, Willmore WG (2014) Redox regulation of mitochondrial function with emphasis on cysteine oxidation reactions. Redox Biol 2:123–139. doi: 10.1016/j.redox.2013.12.011 PubMedCentralPubMedGoogle Scholar
  143. Mansfield KD, Guzy RD, Pan Y et al (2005) Mitochondrial dysfunction resulting from loss of cytochrome c impairs cellular oxygen sensing and hypoxic HIF-alpha activation. Cell Metab 1(6):393–399. doi: 10.1016/j.cmet.2005.05.003 PubMedCentralPubMedGoogle Scholar
  144. Martinez-Reyes I, Cuezva JM (2014) The H(+)-ATP synthase: a gate to ROS-mediated cell death or cell survival. Biochim Biophys Acta 1837(7):1099–1112. doi: 10.1016/j.bbabio.2014.03.010 PubMedGoogle Scholar
  145. McLain AL, Szweda PA, Szweda LI (2011) α-Ketoglutarate dehydrogenase: a mitochondrial redox sensor. Free Radical Res 45(1):29–36. doi: 10.3109/10715762.2010.534163 Google Scholar
  146. McLain AL, Cormier PJ, Kinter M, Szweda LI (2013) Glutathionylation of alpha-ketoglutarate dehydrogenase: the chemical nature and relative susceptibility of the cofactor lipoic acid to modification. Free Radic Biol Med 61:161–169. doi: 10.1016/j.freeradbiomed.2013.03.020 PubMedGoogle Scholar
  147. McLeod CJ, Aziz A, Hoyt RF Jr, McCoy JP Jr, Sack MN (2005) Uncoupling proteins 2 and 3 function in concert to augment tolerance to cardiac ischemia. J Biol Chem 280(39):33470–33476. doi: 10.1074/jbc.M505258200 PubMedGoogle Scholar
  148. McStay GP, Clarke SJ, Halestrap AP (2002) Role of critical thiol groups on the matrix surface of the adenine nucleotide translocase in the mechanism of the mitochondrial permeability transition pore. Biochem J 367(Pt 2):541–548. doi: 10.1042/BJ20011672 PubMedCentralPubMedGoogle Scholar
  149. Merry TL, Steinberg GR, Lynch GS, McConell GK (2010) Skeletal muscle glucose uptake during contraction is regulated by nitric oxide and ROS independently of AMPK. Am J Physiol Endocrinol Metab 298(3):E577–E585. doi: 10.1152/ajpendo.00239.2009 PubMedGoogle Scholar
  150. Mitchell P (1961) Coupling of phosphorylation to electron and hydrogen transfer by a chemi-osmotic type of mechanism. Nature 191:144–148PubMedGoogle Scholar
  151. Mitsumoto Y, Klip A (1992) Development regulation of the subcellular distribution and glycosylation of GLUT1 and GLUT4 glucose transporters during myogenesis of L6 muscle cells. J Biol Chem 267(7):4957–4962PubMedGoogle Scholar
  152. Morita A, Tanimoto K, Murakami T, Morinaga T, Hosoi Y (2014) Mitochondria are required for ATM activation by extranuclear oxidative stress in cultured human hepatoblastoma cell line Hep G2 cells. Biochem Biophys Res Commun 443(4):1286–1290. doi: 10.1016/j.bbrc.2013.12.139 PubMedGoogle Scholar
  153. Muller FL (2009) A critical evaluation of cpYFP as a probe for superoxide. Free Radic Biol Med 47(12):1779–1780. doi: 10.1016/j.freeradbiomed.2009.09.019 PubMedGoogle Scholar
  154. Muller FL, Roberts AG, Bowman MK, Kramer DM (2003) Architecture of the Qo site of the cytochrome bc1 complex probed by superoxide production. Biochemistry 42(21):6493–6499. doi: 10.1021/bi0342160 PubMedGoogle Scholar
  155. Muller FL, Liu Y, Van Remmen H (2004) Complex III releases superoxide to both sides of the inner mitochondrial membrane. J Biol Chem 279(47):49064–49073. doi: 10.1074/jbc.M407715200 PubMedGoogle Scholar
  156. Mungai PT, Waypa GB, Jairaman A et al (2011) Hypoxia triggers AMPK activation through reactive oxygen species-mediated activation of calcium release-activated calcium channels. Mol Cell Biol 31(17):3531–3545. doi: 10.1128/MCB.05124-11 PubMedCentralPubMedGoogle Scholar
  157. Murphy MP (2009) How mitochondria produce reactive oxygen species. Biochem J 417(1):1–13. doi: 10.1042/BJ20081386 PubMedCentralPubMedGoogle Scholar
  158. Murrant CL, Andrade FH, Reid MB (1999) Exogenous reactive oxygen and nitric oxide alter intracellular oxidant status of skeletal muscle fibres. Acta Physiol Scand 166(2):111–121. doi: 10.1046/j.1365-201x.1999.00555.x PubMedGoogle Scholar
  159. Negre-Salvayre A, Hirtz C, Carrera G et al (1997) A role for uncoupling protein-2 as a regulator of mitochondrial hydrogen peroxide generation. FASEB J 11(10):809–815PubMedGoogle Scholar
  160. Ng Y, Ramm G, Lopez JA, James DE (2008) Rapid activation of Akt2 is sufficient to stimulate GLUT4 translocation in 3T3-L1 adipocytes. Cell Metab 7(4):348–356. doi: 10.1016/j.cmet.2008.02.008 PubMedGoogle Scholar
  161. Nguyen TT, Stevens MV, Kohr M, Steenbergen C, Sack MN, Murphy E (2011) Cysteine 203 of cyclophilin D is critical for cyclophilin D activation of the mitochondrial permeability transition pore. J Biol Chem 286(46):40184–40192. doi: 10.1074/jbc.M111.243469 PubMedCentralPubMedGoogle Scholar
  162. Nishikawa T, Edelstein D, Du XL et al (2000) Normalizing mitochondrial superoxide production blocks three pathways of hyperglycaemic damage. Nature 404(6779):787–790. doi: 10.1038/35008121 PubMedGoogle Scholar
  163. Nishioka T, Oda Y, Seino Y et al (1992) Distribution of the glucose transporters in human brain tumors. Cancer Res 52(14):3972–3979PubMedGoogle Scholar
  164. Nishiyama A, Matsui M, Iwata S et al (1999) Identification of thioredoxin-binding protein-2/vitamin D(3) up-regulated protein 1 as a negative regulator of thioredoxin function and expression. J Biol Chem 274(31):21645–21650PubMedGoogle Scholar
  165. Niwa K, Inanami O, Yamamori T, Ohta T, Hamasu T, Kuwabara M (2003) Redox regulation of PI3K/Akt and p53 in bovine aortic endothelial cells exposed to hydrogen peroxide. Antioxid Redox Signal 5(6):713–722. doi: 10.1089/152308603770380016 PubMedGoogle Scholar
  166. Okoh V, Deoraj A, Roy D (2011) Estrogen-induced reactive oxygen species-mediated signalings contribute to breast cancer. Biochim Biophys Acta 1815(1):115–133. doi: 10.1016/j.bbcan.2010.10.005 PubMedGoogle Scholar
  167. Orr AL, Quinlan CL, Perevoshchikova IV, Brand MD (2012) A refined analysis of superoxide production by mitochondrial sn-glycerol 3-phosphate dehydrogenase. J Biol Chem 287(51):42921–42935. doi: 10.1074/jbc.M112.397828 PubMedCentralPubMedGoogle Scholar
  168. Ouiddir A, Planes C, Fernandes I, VanHesse A, Clerici C (1999) Hypoxia upregulates activity and expression of the glucose transporter GLUT1 in alveolar epithelial cells. Am J Respir Cell Mol Biol 21(6):710–718. doi: 10.1165/ajrcmb.21.6.3751 PubMedGoogle Scholar
  169. Pacher P, Schulz R, Liaudet L, Szabo C (2005) Nitrosative stress and pharmacological modulation of heart failure. Trends Pharmacol Sci 26(6):302–310. doi: 10.1016/ PubMedCentralPubMedGoogle Scholar
  170. Parikh H, Carlsson E, Chutkow WA et al (2007) TXNIP regulates peripheral glucose metabolism in humans. PLoS Med 4(5):e158. doi: 10.1371/journal.pmed.0040158 PubMedCentralPubMedGoogle Scholar
  171. Patwari P, Chutkow WA, Cummings K et al (2009) Thioredoxin-independent regulation of metabolism by the alpha-arrestin proteins. J Biol Chem 284(37):24996–25003. doi: 10.1074/jbc.M109.018093 PubMedCentralPubMedGoogle Scholar
  172. Pi J, Collins S (2010) Reactive oxygen species and uncoupling protein 2 in pancreatic beta-cell function. Diabetes Obes Metab 12(Suppl 2):141–148. doi: 10.1111/j.1463-1326.2010.01269.x PubMedGoogle Scholar
  173. Pinheiro CH, Silveira LR, Nachbar RT, Vitzel KF, Curi R (2010) Regulation of glycolysis and expression of glucose metabolism-related genes by reactive oxygen species in contracting skeletal muscle cells. Free Radic Biol Med 48(7):953–960. doi: 10.1016/j.freeradbiomed.2010.01.016 PubMedGoogle Scholar
  174. Prasad RK, Ismail-Beigi F (1999) Mechanism of stimulation of glucose transport by H2O2: role of phospholipase C. Arch Biochem Biophys 362(1):113–122. doi: 10.1006/abbi.1998.1026 PubMedGoogle Scholar
  175. Prata C, Maraldi T, Fiorentini D, Zambonin L, Hakim G, Landi L (2008) Nox-generated ROS modulate glucose uptake in a leukaemic cell line. Free Radical Res 42(5):405–414. doi: 10.1080/10715760802047344 Google Scholar
  176. Queiroga CS, Almeida AS, Martel C, Brenner C, Alves PM, Vieira HL (2010) Glutathionylation of adenine nucleotide translocase induced by carbon monoxide prevents mitochondrial membrane permeabilization and apoptosis. J Biol Chem 285(22):17077–17088. doi: 10.1074/jbc.M109.065052 PubMedCentralPubMedGoogle Scholar
  177. Quinlan CL, Orr AL, Perevoshchikova IV, Treberg JR, Ackrell BA, Brand MD (2012a) Mitochondrial complex II can generate reactive oxygen species at high rates in both the forward and reverse reactions. J Biol Chem 287(32):27255–27264. doi: 10.1074/jbc.M112.374629 PubMedCentralPubMedGoogle Scholar
  178. Quinlan CL, Treberg JR, Perevoshchikova IV, Orr AL, Brand MD (2012b) Native rates of superoxide production from multiple sites in isolated mitochondria measured using endogenous reporters. Free Radic Biol Med 53(9):1807–1817. doi: 10.1016/j.freeradbiomed.2012.08.015 PubMedCentralPubMedGoogle Scholar
  179. Quinlan CL, Perevoshchikova IV, Hey-Mogensen M, Orr AL, Brand MD (2013) Sites of reactive oxygen species generation by mitochondria oxidizing different substrates. Redox Biol 1(1):304–312. doi: 10.1016/j.redox.2013.04.005 PubMedCentralPubMedGoogle Scholar
  180. Quinlan CL, Goncalves RL, Hey-Mogensen M, Yadava N, Bunik VI, Brand MD (2014) The 2-oxoacid dehydrogenase complexes in mitochondria can produce superoxide/hydrogen peroxide at much higher rates than complex I. J Biol Chem 289(12):8312–8325. doi: 10.1074/jbc.M113.545301 PubMedCentralPubMedGoogle Scholar
  181. Rehder DS, Borges CR (2010) Cysteine sulfenic acid as an intermediate in disulfide bond formation and nonenzymatic protein folding. Biochemistry 49(35):7748–7755. doi: 10.1021/bi1008694 PubMedCentralPubMedGoogle Scholar
  182. Remels AH, Langen RC, Schrauwen P, Schaart G, Schols AM, Gosker HR (2010) Regulation of mitochondrial biogenesis during myogenesis. Mol Cell Endocrinol 315(1–2):113–120. doi: 10.1016/j.mce.2009.09.029 PubMedGoogle Scholar
  183. Rhee SG, Bae YS, Lee SR, Kwon J (2000) Hydrogen peroxide: a key messenger that modulates protein phosphorylation through cysteine oxidation. Sci STKE 2000(53):pe1. doi: 10.1126/stke.2000.53.pe1 PubMedGoogle Scholar
  184. Riahi Y, Sin-Malia Y, Cohen G et al (2010) The natural protective mechanism against hyperglycemia in vascular endothelial cells: roles of the lipid peroxidation product 4-hydroxydodecadienal and peroxisome proliferator-activated receptor delta. Diabetes 59(4):808–818. doi: 10.2337/db09-1207 PubMedCentralPubMedGoogle Scholar
  185. Rosa SC, Goncalves J, Judas F, Mobasheri A, Lopes C, Mendes AF (2009) Impaired glucose transporter-1 degradation and increased glucose transport and oxidative stress in response to high glucose in chondrocytes from osteoarthritic versus normal human cartilage. Arthritis Res Ther 11(3):R80. doi: 10.1186/ar2713 PubMedCentralPubMedGoogle Scholar
  186. Rossig L, Fichtlscherer B, Breitschopf K et al (1999) Nitric oxide inhibits caspase-3 by S-nitrosation in vivo. J Biol Chem 274(11):6823–6826PubMedGoogle Scholar
  187. Rumsey SC, Kwon O, Xu GW, Burant CF, Simpson I, Levine M (1997) Glucose transporter isoforms GLUT1 and GLUT3 transport dehydroascorbic acid. J Biol Chem 272(30):18982–18989PubMedGoogle Scholar
  188. Rungaldier S, Oberwagner W, Salzer U, Csaszar E, Prohaska R (2013) Stomatin interacts with GLUT1/SLC2A1, band 3/SLC4A1, and aquaporin-1 in human erythrocyte membrane domains. Biochim Biophys Acta 1828(3):956–966. doi: 10.1016/j.bbamem.2012.11.030 PubMedCentralPubMedGoogle Scholar
  189. Rydstrom J (2006) Mitochondrial NADPH, transhydrogenase and disease. Biochim Biophys Acta 1757(5–6):721–726. doi: 10.1016/j.bbabio.2006.03.010 PubMedGoogle Scholar
  190. Sabens Liedhegner EA, Gao XH, Mieyal JJ (2012) Mechanisms of altered redox regulation in neurodegenerative diseases–focus on S–glutathionylation. Antioxid Redox Signal 16(6):543–566. doi: 10.1089/ars.2011.4119 PubMedCentralPubMedGoogle Scholar
  191. Salmeen A, Andersen JN, Myers MP et al (2003) Redox regulation of protein tyrosine phosphatase 1B involves a sulphenyl-amide intermediate. Nature 423(6941):769–773. doi: 10.1038/nature01680 PubMedGoogle Scholar
  192. Salvi M, Battaglia V, Brunati AM et al (2007) Catalase takes part in rat liver mitochondria oxidative stress defense. J Biol Chem 282(33):24407–24415. doi: 10.1074/jbc.M701589200 PubMedGoogle Scholar
  193. Sanchez-Gomez FJ, Espinosa-Diez C, Dubey M, Dikshit M, Lamas S (2013) S-glutathionylation: relevance in diabetes and potential role as a biomarker. Biol Chem 394(10):1263–1280. doi: 10.1515/hsz-2013-0150 PubMedGoogle Scholar
  194. Sandstrom ME, Zhang SJ, Bruton J et al (2006) Role of reactive oxygen species in contraction-mediated glucose transport in mouse skeletal muscle. J Physiol 575(Pt 1):251–262. doi: 10.1113/jphysiol.2006.110601 PubMedCentralPubMedGoogle Scholar
  195. Sanjuan-Pla A, Cervera AM, Apostolova N et al (2005) A targeted antioxidant reveals the importance of mitochondrial reactive oxygen species in the hypoxic signaling of HIF-1alpha. FEBS Lett 579(12):2669–2674. doi: 10.1016/j.febslet.2005.03.088 PubMedGoogle Scholar
  196. Sano H, Kane S, Sano E et al (2003) Insulin-stimulated phosphorylation of a Rab GTPase-activating protein regulates GLUT4 translocation. J Biol Chem 278(17):14599–14602. doi: 10.1074/jbc.C300063200 PubMedGoogle Scholar
  197. Sasson S, Kaiser N, Dan-Goor M et al (1997) Substrate autoregulation of glucose transport: hexose 6-phosphate mediates the cellular distribution of glucose transporters. Diabetologia 40(1):30–39. doi: 10.1007/s001250050639 PubMedGoogle Scholar
  198. Schroedl C, McClintock DS, Budinger GR, Chandel NS (2002) Hypoxic but not anoxic stabilization of HIF-1alpha requires mitochondrial reactive oxygen species. Am J Physiol Lung Cell Mol Physiol 283(5):L922–L931. doi: 10.1152/ajplung.00014.2002 PubMedGoogle Scholar
  199. Schulze PC, Yoshioka J, Takahashi T, He Z, King GL, Lee RT (2004) Hyperglycemia promotes oxidative stress through inhibition of thioredoxin function by thioredoxin-interacting protein. J Biol Chem 279(29):30369–30374. doi: 10.1074/jbc.M400549200 PubMedGoogle Scholar
  200. Schwarzlander M, Murphy MP, Duchen MR et al (2012) Mitochondrial ‘flashes’: a radical concept repHined. Trends Cell Biol 22(10):503–508. doi: 10.1016/j.tcb.2012.07.007 PubMedGoogle Scholar
  201. Schwarzlander M, Wagner S, Ermakova YG et al (2014) The ‘mitoflash’ probe cpYFP does not respond to superoxide. Nature 514(7523):E12–E14. doi: 10.1038/nature13858 PubMedGoogle Scholar
  202. Seifert EL, Bezaire V, Estey C, Harper ME (2008) Essential role for uncoupling protein-3 in mitochondrial adaptation to fasting but not in fatty acid oxidation or fatty acid anion export. J Biol Chem 283(37):25124–25131. doi: 10.1074/jbc.M803871200 PubMedGoogle Scholar
  203. Sena LA, Chandel NS (2012) Physiological roles of mitochondrial reactive oxygen species. Mol Cell 48(2):158–167. doi: 10.1016/j.molcel.2012.09.025 PubMedCentralPubMedGoogle Scholar
  204. Seres T, Ravichandran V, Moriguchi T, Rokutan K, Thomas JA, Johnston RB Jr (1996) Protein S-thiolation and dethiolation during the respiratory burst in human monocytes. A reversible post-translational modification with potential for buffering the effects of oxidant stress. J Immunol 156(5):1973–1980PubMedGoogle Scholar
  205. Shetty M, Loeb JN, Vikstrom K, Ismail-Beigi F (1993) Rapid activation of GLUT-1 glucose transporter following inhibition of oxidative phosphorylation in clone 9 cells. J Biol Chem 268(23):17225–17232PubMedGoogle Scholar
  206. Shi X, Bai S, Ford AC et al (1995) Stable inducible expression of a functional rat liver organic anion transport protein in HeLa cells. J Biol Chem 270(43):25591–25595PubMedGoogle Scholar
  207. Siebels I, Drose S (2013) Q-site inhibitor induced ROS production of mitochondrial complex II is attenuated by TCA cycle dicarboxylates. Biochim Biophys Acta 1827(10):1156–1164. doi: 10.1016/j.bbabio.2013.06.005 PubMedGoogle Scholar
  208. Silveira LR, Pilegaard H, Kusuhara K, Curi R, Hellsten Y (2006) The contraction induced increase in gene expression of peroxisome proliferator-activated receptor (PPAR)-gamma coactivator 1alpha (PGC-1alpha), mitochondrial uncoupling protein 3 (UCP3) and hexokinase II (HKII) in primary rat skeletal muscle cells is dependent on reactive oxygen species. Biochim Biophys Acta 1763(9):969–976. doi: 10.1016/j.bbamcr.2006.06.010 PubMedGoogle Scholar
  209. Sivaramakrishnan S, Cummings AH, Gates KS (2010) Protection of a single-cysteine redox switch from oxidative destruction: on the functional role of sulfenyl amide formation in the redox-regulated enzyme PTP1B. Bioorg Med Chem Lett 20(2):444–447. doi: 10.1016/j.bmcl.2009.12.001 PubMedCentralPubMedGoogle Scholar
  210. Somwar R, Koterski S, Sweeney G et al (2002) A dominant-negative p38 MAPK mutant and novel selective inhibitors of p38 MAPK reduce insulin-stimulated glucose uptake in 3T3-L1 adipocytes without affecting GLUT4 translocation. J Biol Chem 277(52):50386–50395. doi: 10.1074/jbc.M205277200 PubMedGoogle Scholar
  211. Starkov AA, Fiskum G (2003) Regulation of brain mitochondrial H2O2 production by membrane potential and NAD(P)H redox state. J Neurochem 86(5):1101–1107PubMedGoogle Scholar
  212. Starkov AA, Fiskum G, Chinopoulos C et al (2004) Mitochondrial alpha-ketoglutarate dehydrogenase complex generates reactive oxygen species. J Neurosci 24(36):7779–7788. doi: 10.1523/JNEUROSCI.1899-04.2004 PubMedGoogle Scholar
  213. Stoltzman CA, Peterson CW, Breen KT, Muoio DM, Billin AN, Ayer DE (2008) Glucose sensing by MondoA: Mlx complexes: a role for hexokinases and direct regulation of thioredoxin-interacting protein expression. Proc Natl Acad Sci USA 105(19):6912–6917. doi: 10.1073/pnas.0712199105 PubMedCentralPubMedGoogle Scholar
  214. Sun L, Shukair S, Naik TJ, Moazed F, Ardehali H (2008) Glucose phosphorylation and mitochondrial binding are required for the protective effects of hexokinases I and II. Mol Cell Biol 28(3):1007–1017. doi: 10.1128/MCB.00224-07 PubMedCentralPubMedGoogle Scholar
  215. Sweeney G, Somwar R, Ramlal T, Volchuk A, Ueyama A, Klip A (1999) An inhibitor of p38 mitogen-activated protein kinase prevents insulin-stimulated glucose transport but not glucose transporter translocation in 3T3-L1 adipocytes and L6 myotubes. J Biol Chem 274(15):10071–10078PubMedGoogle Scholar
  216. Szablewski L (2013) Expression of glucose transporters in cancers. Biochim Biophys Acta 1835(2):164–169. doi: 10.1016/j.bbcan.2012.12.004 PubMedGoogle Scholar
  217. Talbot DA, Lambert AJ, Brand MD (2004) Production of endogenous matrix superoxide from mitochondrial complex I leads to activation of uncoupling protein 3. FEBS Lett 556(1–3):111–115PubMedGoogle Scholar
  218. Talior I, Yarkoni M, Bashan N, Eldar-Finkelman H (2003) Increased glucose uptake promotes oxidative stress and PKC-delta activation in adipocytes of obese, insulin-resistant mice. Am J Physiol Endocrinol Metab 285(2):E295–E302. doi: 10.1152/ajpendo.00044.2003 PubMedGoogle Scholar
  219. Taylor ER, Hurrell F, Shannon RJ, Lin TK, Hirst J, Murphy MP (2003) Reversible glutathionylation of complex I increases mitochondrial superoxide formation. J Biol Chem 278(22):19603–19610. doi: 10.1074/jbc.M209359200 PubMedGoogle Scholar
  220. Taylor EB, An D, Kramer HF et al (2008) Discovery of TBC1D1 as an insulin-, AICAR-, and contraction-stimulated signaling nexus in mouse skeletal muscle. J Biol Chem 283(15):9787–9796. doi: 10.1074/jbc.M708839200 PubMedCentralPubMedGoogle Scholar
  221. Teshima Y, Akao M, Jones SP, Marban E (2003) Uncoupling protein-2 overexpression inhibits mitochondrial death pathway in cardiomyocytes. Circ Res 93(3):192–200. doi: 10.1161/01.RES.0000085581.60197.4D PubMedGoogle Scholar
  222. Thong FS, Bilan PJ, Klip A (2007) The Rab GTPase-activating protein AS160 integrates Akt, protein kinase C, and AMP-activated protein kinase signals regulating GLUT4 traffic. Diabetes 56(2):414–423. doi: 10.2337/db06-0900 PubMedGoogle Scholar
  223. Toime LJ, Brand MD (2010) Uncoupling protein-3 lowers reactive oxygen species production in isolated mitochondria. Free Radic Biol Med 49(4):606–611. doi: 10.1016/j.freeradbiomed.2010.05.010 PubMedCentralPubMedGoogle Scholar
  224. Tonks NK (2005) Redox redux: revisiting PTPs and the control of cell signaling. Cell 121(5):667–670. doi: 10.1016/j.cell.2005.05.016 PubMedGoogle Scholar
  225. Totary-Jain H, Naveh-Many T, Riahi Y, Kaiser N, Eckel J, Sasson S (2005) Calreticulin destabilizes glucose transporter-1 mRNA in vascular endothelial and smooth muscle cells under high-glucose conditions. Circ Res 97(10):1001–1008. doi: 10.1161/01.RES.0000189260.46084.e5 PubMedGoogle Scholar
  226. Toyoda T, Hayashi T, Miyamoto L et al (2004) Possible involvement of the alpha1 isoform of 5′AMP-activated protein kinase in oxidative stress-stimulated glucose transport in skeletal muscle. Am J Physiol Endocrinol Metab 287(1):E166–E173. doi: 10.1152/ajpendo.00487.2003 PubMedGoogle Scholar
  227. Treberg JR, Quinlan CL, Brand MD (2011) Evidence for two sites of superoxide production by mitochondrial NADH-ubiquinone oxidoreductase (complex I). J Biol Chem 286(31):27103–27110. doi: 10.1074/jbc.M111.252502 PubMedCentralPubMedGoogle Scholar
  228. Tretter L, Adam-Vizi V (2000) Inhibition of Krebs cycle enzymes by hydrogen peroxide: a key role of [alpha]-ketoglutarate dehydrogenase in limiting NADH production under oxidative stress. J Neurosci 20(24):8972–8979PubMedGoogle Scholar
  229. Tretter L, Adam-Vizi V (2004) Generation of reactive oxygen species in the reaction catalyzed by alpha-ketoglutarate dehydrogenase. J Neurosci 24(36):7771–7778. doi: 10.1523/JNEUROSCI.1842-04.2004 PubMedGoogle Scholar
  230. Turrens JF, Alexandre A, Lehninger AL (1985) Ubisemiquinone is the electron donor for superoxide formation by complex III of heart mitochondria. Arch Biochem Biophys 237(2):408–414PubMedGoogle Scholar
  231. Tyler DD (1975) Polarographic assay and intracellular distribution of superoxide dismutase in rat liver. Biochem J 147(3):493–504PubMedCentralPubMedGoogle Scholar
  232. Valko M, Rhodes CJ, Moncol J, Izakovic M, Mazur M (2006) Free radicals, metals and antioxidants in oxidative stress-induced cancer. Chem Biol Interact 160(1):1–40. doi: 10.1016/j.cbi.2005.12.009 PubMedGoogle Scholar
  233. Valsecchi F, Grefte S, Roestenberg P et al (2013) Primary fibroblasts of NDUFS4(−/−) mice display increased ROS levels and aberrant mitochondrial morphology. Mitochondrion 13(5):436–443. doi: 10.1016/j.mito.2012.12.001 PubMedGoogle Scholar
  234. Varanyuwatana P, Halestrap AP (2012) The roles of phosphate and the phosphate carrier in the mitochondrial permeability transition pore. Mitochondrion 12(1):120–125. doi: 10.1016/j.mito.2011.04.006 PubMedCentralPubMedGoogle Scholar
  235. Vasquez-Vivar J, Kalyanaraman B, Kennedy MC (2000) Mitochondrial aconitase is a source of hydroxyl radical. An electron spin resonance investigation. J Biol Chem 275(19):14064–14069PubMedGoogle Scholar
  236. Verkaart S, Koopman WJ, Cheek J et al (2007a) Mitochondrial and cytosolic thiol redox state are not detectably altered in isolated human NADH: ubiquinone oxidoreductase deficiency. Biochim Biophys Acta 1772(9):1041–1051. doi: 10.1016/j.bbadis.2007.05.004 PubMedGoogle Scholar
  237. Verkaart S, Koopman WJ, van Emst-de Vries SE et al (2007b) Superoxide production is inversely related to complex I activity in inherited complex I deficiency. Biochim Biophys Acta 1772(3):373–381. doi: 10.1016/j.bbadis.2006.12.009 PubMedGoogle Scholar
  238. Vinals F, Fandos C, Santalucia T et al (1997) Myogenesis and MyoD down-regulate Sp1. A mechanism for the repression of GLUT1 during muscle cell differentiation. J Biol Chem 272(20):12913–12921PubMedGoogle Scholar
  239. Wang W, Fang H, Groom L et al (2008) Superoxide flashes in single mitochondria. Cell 134(2):279–290. doi: 10.1016/j.cell.2008.06.017 PubMedCentralPubMedGoogle Scholar
  240. Weisiger RA, Fridovich I (1973) Mitochondrial superoxide simutase. Site of synthesis and intramitochondrial localization. J Biol Chem 248(13):4793–4796PubMedGoogle Scholar
  241. Wieman HL, Horn SR, Jacobs SR, Altman BJ, Kornbluth S, Rathmell JC (2009) An essential role for the Glut1 PDZ-binding motif in growth factor regulation of Glut1 degradation and trafficking. Biochem J 418(2):345–367. doi: 10.1042/BJ20081422 PubMedCentralPubMedGoogle Scholar
  242. Winterbourn CC, Hampton MB (2008) Thiol chemistry and specificity in redox signaling. Free Radic Biol Med 45(5):549–561. doi: 10.1016/j.freeradbiomed.2008.05.004 PubMedGoogle Scholar
  243. Wood SM, Wiesener MS, Yeates KM et al (1998) Selection and analysis of a mutant cell line defective in the hypoxia-inducible factor-1 alpha-subunit (HIF-1alpha). Characterization of hif-1alpha-dependent and -independent hypoxia-inducible gene expression. J Biol Chem 273(14):8360–8368PubMedGoogle Scholar
  244. Wretman C, Lionikas A, Widegren U, Lannergren J, Westerblad H, Henriksson J (2001) Effects of concentric and eccentric contractions on phosphorylation of MAPK(erk1/2) and MAPK(p38) in isolated rat skeletal muscle. J Physiol 535(Pt 1):155–164PubMedCentralPubMedGoogle Scholar
  245. Wu SB, Wei YH (2012) AMPK-mediated increase of glycolysis as an adaptive response to oxidative stress in human cells: implication of the cell survival in mitochondrial diseases. Biochim Biophys Acta 1822(2):233–247. doi: 10.1016/j.bbadis.2011.09.014 PubMedGoogle Scholar
  246. Wu N, Zheng B, Shaywitz A et al (2013) AMPK-dependent degradation of TXNIP upon energy stress leads to enhanced glucose uptake via GLUT1. Mol Cell 49(6):1167–1175. doi: 10.1016/j.molcel.2013.01.035 PubMedCentralPubMedGoogle Scholar
  247. Wu SB, Wu YT, Wu TP, Wei YH (2014) Role of AMPK-mediated adaptive responses in human cells with mitochondrial dysfunction to oxidative stress. Biochim Biophys Acta 1840(4):1331–1344. doi: 10.1016/j.bbagen.2013.10.034 PubMedGoogle Scholar
  248. Yan LJ, Sumien N, Thangthaeng N, Forster MJ (2013) Reversible inactivation of dihydrolipoamide dehydrogenase by mitochondrial hydrogen peroxide. Free Radical Res 47(2):123–133. doi: 10.3109/10715762.2012.752078 Google Scholar
  249. Yang Y, Song Y, Loscalzo J (2007) Regulation of the protein disulfide proteome by mitochondria in mammalian cells. Proc Natl Acad Sci USA 104(26):10813–10817. doi: 10.1073/pnas.0702027104 PubMedCentralPubMedGoogle Scholar
  250. Yin F, Sancheti H, Cadenas E (2012) Silencing of nicotinamide nucleotide transhydrogenase impairs cellular redox homeostasis and energy metabolism in PC12 cells. Biochim Biophys Acta 1817(3):401–409. doi: 10.1016/j.bbabio.2011.12.004 PubMedCentralPubMedGoogle Scholar
  251. Ying W (2008) NAD+/NADH and NADP+/NADPH in cellular functions and cell death: regulation and biological consequences. Antioxid Redox Signal 10(2):179–206. doi: 10.1089/ars.2007.1672 PubMedGoogle Scholar
  252. Yu T, Robotham JL, Yoon Y (2006) Increased production of reactive oxygen species in hyperglycemic conditions requires dynamic change of mitochondrial morphology. Proc Natl Acad Sci USA 103(8):2653–2658. doi: 10.1073/pnas.0511154103 PubMedCentralPubMedGoogle Scholar
  253. Yu T, Jhun BS, Yoon Y (2011) High-glucose stimulation increases reactive oxygen species production through the calcium and mitogen-activated protein kinase-mediated activation of mitochondrial fission. Antioxid Redox Signal 14(3):425–437. doi: 10.1089/ars.2010.3284 PubMedCentralPubMedGoogle Scholar
  254. Zaobornyj T, Ghafourifar P (2012) Strategic localization of heart mitochondrial NOS: a review of the evidence. Am J Physiol Heart Circ Physiol 303(11):H1283–H1293. doi: 10.1152/ajpheart.00674.2011 PubMedGoogle Scholar
  255. Zhang JZ, Abbud W, Prohaska R, Ismail-Beigi F (2001) Overexpression of stomatin depresses GLUT-1 glucose transporter activity. Am J Physiol Cell Physiol 280(5):C1277–C1283PubMedGoogle Scholar
  256. Zhang X, Huang Z, Hou T et al (2013) Superoxide constitutes a major signal of mitochondrial superoxide flash. Life Sci 93(4):178–186. doi: 10.1016/j.lfs.2013.06.012 PubMedGoogle Scholar
  257. Zhou J, Deo BK, Hosoya K et al (2005) Increased JNK phosphorylation and oxidative stress in response to increased glucose flux through increased GLUT1 expression in rat retinal endothelial cells. Invest Ophthalmol Vis Sci 46(9):3403–3410. doi: 10.1167/iovs.04-1064 PubMedGoogle Scholar
  258. Zmijewski JW, Banerjee S, Bae H, Friggeri A, Lazarowski ER, Abraham E (2010) Exposure to hydrogen peroxide induces oxidation and activation of AMP-activated protein kinase. J Biol Chem 285(43):33154–33164. doi: 10.1074/jbc.M110.143685 PubMedCentralPubMedGoogle Scholar

Copyright information

© The Author(s) 2015

Open AccessThis article is distributed under the terms of the Creative Commons Attribution 4.0 International License (, which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made.

Authors and Affiliations

  • Dania C. Liemburg-Apers
    • 1
  • Peter H. G. M. Willems
    • 1
  • Werner J. H. Koopman
    • 1
    Email author
  • Sander Grefte
    • 1
    • 2
  1. 1.Department of Biochemistry (286), Radboud Institute for Molecular Life Sciences (RIMLS)Radboud University Medical Center (RUMC)NijmegenThe Netherlands
  2. 2.Department of Human and Animal PhysiologyWageningen UniversityWageningenThe Netherlands

Personalised recommendations