Accessible Published by De Gruyter January 27, 2015

Thiol switches in mitochondria: operation and physiological relevance

Jan Riemer, Markus Schwarzländer, Marcus Conrad and Johannes M. Herrmann
From the journal Biological Chemistry

Abstract

Mitochondria are a major source of reactive oxygen species (ROS) in the cell, particularly of superoxide and hydrogen peroxide. A number of dedicated enzymes regulate the conversion and consumption of superoxide and hydrogen peroxide in the intermembrane space and the matrix of mitochondria. Nevertheless, hydrogen peroxide can also interact with many other mitochondrial enzymes, particularly those with reactive cysteine residues, modulating their reactivity in accordance with changes in redox conditions. In this review we will describe the general redox systems in mitochondria of animals, fungi and plants and discuss potential target proteins that were proposed to contain regulatory thiol switches.

Introduction – mitochondria contain two distinct redox networks

Mitochondria are essential organelles of eukaryotic cells. They produce not only the bulk of cellular energy in the form of ATP, but they also generate numerous important metabolites and cofactors, and they serve as critical signaling stations that, for example, integrate cellular signals to initiate apoptosis. In turn, mitochondria also communicate their metabolic and fitness state to the remainder of the cell to trigger cellular adaptation processes.

Mitochondria contain two distinct aqueous subcompartments, the intermembrane space (IMS) and the matrix. Both subcompartments differ strongly with respect to their biological activity, their protein composition (Herrmann and Riemer, 2010) as well as their redox properties. Firstly, whereas the IMS is connected to the cytosol via porins, which allow the free diffusion of molecules of up to 5 kDa (including GSH/GSSG, NADPH/NADP+ or hydrogen peroxide), the matrix is strictly separated from the IMS as the transport across the inner membrane is tightly controlled by substrate-specific carriers. Secondly, most cysteine residues of matrix proteins are believed to be reduced in the matrix, while many IMS proteins contain structural disulfide bonds which are introduced by the mitochondrial disulfide relay (also called MIA pathway) (Chacinska et al., 2004; Naoe et al., 2004; Allen et al., 2005; Mesecke et al., 2005; Bihlmaier et al., 2007; Banci et al., 2009; Kawano et al., 2009; Milenkovic et al., 2009; Bien et al., 2010; von der Malsburg et al., 2011; Fischer et al., 2013; Koch and Schmid, 2014).

The IMS may even be considered as two separate subcompartments because the peripheral IMS, which is adjacent to the outer membrane, is separated from the cristae space by cristae junctions (Frey and Mannella, 2000; Scorrano et al., 2002). Recently, a protein complex termed MICOS was identified and found to be located in the inner membrane and is critical for the separation of these two subcompartments as well as for mitochondrial functionality (Harner et al., 2011; Hoppins et al., 2011; von der Malsburg et al., 2011; Körner et al., 2012). Cristae junctions restrict (and perhaps even control) the diffusion of molecules into and out of the cristae. Hence, the small molecule environment of the cristae space – specifically with regards to its pH, glutathione redox potential and levels of reactive oxygen species (ROS) – likely differs from that of the peripheral IMS and is more strongly influenced by the respiratory chain, which is mainly localized in cristae membranes. There is good evidence that proteins of the IMS and the inner membrane are partitioned between the cristae and the peripheral inner membrane but the mechanisms by which segregation takes place are unknown (Vogel et al., 2006; Wurm and Jakobs, 2006; Stoldt et al., 2012; Daum et al., 2013).

The mitochondrial matrix is separated from the IMS by the inner membrane, which only allows a tightly regulated exchange of small molecules. Thus, the small molecule environment of the matrix is mainly controlled independently from the remainder of the cell. This is nicely illustrated for example by the regulation of GSH/GSSG ratio in the matrix (Hu et al., 2008; Kojer et al., 2012). The matrix GSH/GSSG ratio is similar to its cytosolic counterpart, but is subject to completely different regulatory machineries including mitochondria-specific ROS-generating pathways (see below) and a partly mitochondria-specific set of reducing enzymes, including factors for the reductive regeneration of NADPH and GSH. The differences between the mitochondrial compartments lead to differences in the generation, handling and perception of oxidizing and reductive influences.

In the following we will illustrate this notion by describing and discussing the mechanisms that drive the reversible operation of molecular cysteine switches. Molecular switch operating is driven by the balance of opposing influences (Figure 1). Oxidizing influences mainly stem from the production of ROS at different sites in mitochondria; reducing influences ultimately originate from NADPH, which is regenerated by different metabolic pathways. Both thiol switch drivers rely on enzymatic machineries for the modulation and the relay of signals.

Figure 1: H2O2-controlled thiol switches in mitochondria.Reactive oxygen species (ROS) can be produced at numerous sites in mitochondria, e.g. by flavine-containing enzymes of the TCA cycle or complexes of the respiratory chain. The main species produced is superoxide (O2·-) that rapidly dismutates – either enzymatically or non-enzymatically – to hydrogen peroxide (H2O2). H2O2 reacts with its targets, which include reactive protein thiols or dedicated redox enzymes such as glutathione peroxidases (Gpx) or peroxiredoxins (Prx). The reaction with Gpx and Prx either leads to the ‘detoxification’ of H2O2 or can result in the indirect oxidation of proteins with less reactive thiols. The localization of the ROS source and the competition between degrading pathways and H2O2 diffusion determines which targets are oxidized. Thiol oxidation in target proteins can have different consequences like changes in protein activity, localization and stability (visualized by the traffic lights). Importantly, oxidation is counteracted by reducing systems that depend on the availability of NADPH and the activity of reducing systems [thioredoxin-thioredoxin reductase (Trx, TRR) and glutaredoxin, glutathione, glutathione reductase (Grx, GSH, GR)]. The SH shown in the figure depicts thiols which, in vivo, might be in their protonated or deprotonated state.

Figure 1:

H2O2-controlled thiol switches in mitochondria.

Reactive oxygen species (ROS) can be produced at numerous sites in mitochondria, e.g. by flavine-containing enzymes of the TCA cycle or complexes of the respiratory chain. The main species produced is superoxide (O2·-) that rapidly dismutates – either enzymatically or non-enzymatically – to hydrogen peroxide (H2O2). H2O2 reacts with its targets, which include reactive protein thiols or dedicated redox enzymes such as glutathione peroxidases (Gpx) or peroxiredoxins (Prx). The reaction with Gpx and Prx either leads to the ‘detoxification’ of H2O2 or can result in the indirect oxidation of proteins with less reactive thiols. The localization of the ROS source and the competition between degrading pathways and H2O2 diffusion determines which targets are oxidized. Thiol oxidation in target proteins can have different consequences like changes in protein activity, localization and stability (visualized by the traffic lights). Importantly, oxidation is counteracted by reducing systems that depend on the availability of NADPH and the activity of reducing systems [thioredoxin-thioredoxin reductase (Trx, TRR) and glutaredoxin, glutathione, glutathione reductase (Grx, GSH, GR)]. The SH shown in the figure depicts thiols which, in vivo, might be in their protonated or deprotonated state.

Redox influences and redox enzymes in mitochondria

Oxidizing influences in mitochondria

Endogenous ROS have for a long time been considered as the unwanted but unavoidable downside of mitochondrial metabolism that, over time, cause deleterious oxidative stress in eukaryotic cells. However, this traditional concept has been challenged by many recent studies which demonstrated that ROS (in particular hydrogen peroxide) also serve as critical signaling molecules (Chandel et al., 2000; Albrecht et al., 2011; Zarse et al., 2012; Mouchiroud et al., 2013; Gladyshev, 2014; Sies, 2014; Yee et al., 2014).

ROS encompass a variety of different molecules including the superoxide radical (O2·-), hydrogen peroxide (H2O2) and the hydroxyl radical (·OH) that are derived from the partial reduction of molecular oxygen (O2). ROS can also result from the ‘detoxification’ of other ROS, as is the case during the dismutation of O2·- to H2O2 (and oxygen). H2O2 is a particularly suitable candidate to act as a signaling molecule, because of its relatively long half-life (Sies, 1993), relatively high concentration (nanomolar to low micromolar), and reasonable membrane permeability. In mitochondria channels in the outer membrane (porins) and the inner membrane, mitochondrial aquaporin-8 might considerably accelerate the exchange of hydrogen peroxide (Calamita et al., 2005; Marchissio et al., 2012). Perhaps most importantly, H2O2 can selectively oxidize specific cysteine residues in target proteins, making it an ideal candidate as a signaling molecule (Winterbourn and Hampton, 2008). H2O2 can oxidize thiols to sulfenic, sulfinic and sulfonic acid, of which the first two can be enzymatically converted back to thiols. Sulfenic acid can react with another thiol to a disulfide. In contrast, ·OH is highly reactive (reacting with diffusion-limited rate constants) and will indiscriminately oxidize biomolecules, rendering it unsuitable as a signaling molecule.

Mitochondrial pathways are major sources of cellular H2O2 (Figure 2). Different complexes of the respiratory chain release O2·- towards both the matrix and the IMS (Murphy, 2009; Bleier et al., 2014; Dröse et al., 2014). O2·- is rapidly converted to H2O2 by superoxide dismutases (copper-zinc SOD in the IMS and cytosol, manganese SOD in the matrix). The ∼10 000 fold increase in the rate of O2·- dismutation by SODs not only accelerates the production of H2O2, but also competes with the production of peroxinitrite (ONOO-) by the reaction of O2·- with NO·. While dedicated NO synthases have been reported in mammalian mitochondria (Giulivi et al., 1998; Nisoli et al., 2003), reports of a dedicated NO synthase in plant mitochondria have turned out to be incorrect (Gas et al., 2009); instead a likely source of mitochondrial NO is the mitochondrial respiratory chain by nitrite reduction (Gupta et al., 2011). Both H2O2 and ONOO- represent major electron sinks for the matrix thiol machinery and are likely to be main drivers behind thiol oxidation, both directly and indirectly.

Figure 2: Sources of mitochondrial reactive oxygen species.There are several sites in mitochondria that can, in principle, generate superoxide (O2·-). The rates of superoxide production can vary with the local oxygen tension, potentials and half-life of redox centers, metabolic flux topology and rates, and the accumulation of damage in proteins. The site of production is of central importance for the physiological impact as it determines which ROS-handling systems and which targets the respective ROS will encounter. Sites for superoxide production include the complexes of the respiratory chain, which release superoxide towards the IMS (complex III) or matrix (complexes I, II and III), respectively. Moreover, flavoproteins in the different compartments may release ROS as byproducts of their catalytic activities. Players that may act as ROS sources because of their cofactor biochemistry are marked in red. Purple arrows show examples for which the generation of ROS was experimentally shown. αKGDH, α-ketoglutarate dehydrogenase; ALR, augmenter of liver regeneration; AOX, alternative oxidase; CB5R, cytochrome b5 reductase; cytb2, cytochrome b2; DHOH, dihydroorotate dehydrogenase; GPDH, glycerol-3P-dehydrogenase; MAO, monoamine oxiadase; P450, cytochrome P450; PDH, pyruvate dehydrogenase

Figure 2:

Sources of mitochondrial reactive oxygen species.

There are several sites in mitochondria that can, in principle, generate superoxide (O2·-). The rates of superoxide production can vary with the local oxygen tension, potentials and half-life of redox centers, metabolic flux topology and rates, and the accumulation of damage in proteins. The site of production is of central importance for the physiological impact as it determines which ROS-handling systems and which targets the respective ROS will encounter. Sites for superoxide production include the complexes of the respiratory chain, which release superoxide towards the IMS (complex III) or matrix (complexes I, II and III), respectively. Moreover, flavoproteins in the different compartments may release ROS as byproducts of their catalytic activities. Players that may act as ROS sources because of their cofactor biochemistry are marked in red. Purple arrows show examples for which the generation of ROS was experimentally shown. αKGDH, α-ketoglutarate dehydrogenase; ALR, augmenter of liver regeneration; AOX, alternative oxidase; CB5R, cytochrome b5 reductase; cytb2, cytochrome b2; DHOH, dihydroorotate dehydrogenase; GPDH, glycerol-3P-dehydrogenase; MAO, monoamine oxiadase; P450, cytochrome P450; PDH, pyruvate dehydrogenase

H2O2 can also be produced by flavin- or ubiquinone-utilizing enzymes and by the reaction of iron-sulfur clusters with O2·-. The contribution of these different sources to mitochondrial H2O2 release in intact cells is unknown. The amounts of H2O2 produced and released by mitochondria may strongly fluctuate depending on the metabolic load (i.e. electron donors), the activity state of metabolic enzymes and the respiratory chain, the oxygen concentration, and the repertoire of redox enzymes. For instance, while the respiratory chain is commonly viewed as the main generator of O2·-, isolated mitochondria have recently been reported to produce O2·-/H2O2 at much higher rates from 2-oxoacid dehydrogenase complexes than from complex I under certain conditions (Quinlan et al., 2014; Goncalves et al., 2015).

ROS handling and reductive influences in mitochondria

ROS are subject to different enzymatic pathways that counteract or mediate (see below) their oxidizing influences (Figure 3). O2·- becomes rapidly dismutated by superoxide dismutases (SODs) in the IMS and the matrix (Sturtz et al., 2001). The product of this reaction – H2O2 – is subject to catalases, peroxiredoxins (Prx), glutathione peroxidases (GPx) and ascorbate peroxidases (APXs), which exist in distinct sets in the matrix and IMS. In the mitochondrial matrix, the contribution of catalase to H2O2 scavenging is limited (or even not existent) unless catalase is artificially overexpressed (Schriner et al., 2005). Prxs and GPxs become oxidized by the reduction of H2O2 and have to be reduced by the thioredoxin (Trx) and glutathione/glutaredoxin (Grx) systems, respectively (Grant, 2001; Trotter and Grant, 2005; Kumar et al., 2011; Toledano et al., 2013). Prxs and GPxs are very fast enzymes (e.g. the cytosolic Prx2 catalyzes with a rate of 2×107m-1s-1), and thus H2O2 reacts with them several orders of magnitudes more rapidly than with other protein thiols (Winterbourn and Hampton, 2008; Brandes et al., 2011). GPxs, and in particular GPx4 in mammals, also participate in the turnover of lipid hydroperoxides to their corresponding alcohols. Besides its documented reducing activity towards H2O2, human Prx5 is also known to reduce organic peroxides as well as peroxynitrite with second order rate constants of 106–107m-1s-1 whereas its reaction with hydrogen peroxide is lower (105m-1s-1) (Dubuisson et al., 2004; Knoops et al., 2011). The solely matrix-located Prx3 shows second order rate constants with H2O2 of approx. 107m-1s-1 (Parsonage et al., 2008; Cox et al., 2009).

Figure 3: Enzymatic and non-enzymatic systems for the conversion and degradation of mitochondrial reactive oxygen species.Superoxide (O2·-) as main ROS generated de novo is rapidly dismutated to H2O2, either non-enzymatically or by superoxide dimutases (SODs), which accelerate this reaction by a factor of about 10 000. H2O2 can leave the respective compartment by diffusing through membranes or in the case of the outer membrane by diffusing through porins/voltage-dependent anion channels (VDAC). This might play an important role during ROS signaling from mitochondria to the remainder of the cell. H2O2 can also undergo Fenton chemistry catalyzed by Fe2+ resulting in the generation of highly reactive hydroxyl radicals (·OH). However, the majority of H2O2 is presumably scavenged by multiple enzymatic and non-enzymatic systems, which convert H2O2 to water. These systems strongly differ between compartments and compete with each other and other proteins in the same compartment for reaction with H2O2. As electron donor these systems rely on NADPH, highlighting the importance of systems for NADPH regeneration in situ. APX, ascorbate peroxidase; αTCP, α-tocopherol; Ccp, cytochrome c peroxidase; CoQ10, coenzyme Q10; DHAR, dehydroascorbate reductase; Glr/GR, glutathione reductase; GPX/GPx/Gpx, glutathione peroxidase; GRX/Grx, glutaredoxin; GSH, glutathione; IDH/ICDH, isocitrate dehydrogenase; MDHAR, monodehydroascorbate reductase; NNT, nicotin nucleotid transhydrogenase; NTR/Trr/TXNRD, NADPH-thioredoxin reductase; PHGP, phospholipid hydroperoxide glutathione peroxidase; Por, porin; Prx/PRX, peroxiredoxin; Sod1/CSD1, copper-zinc superoxide dismutase; Sod2/MSD1, manganese superoxide dismutase; SRX, sulfiredoxin; TRX/Trx/TXN, thioredoxin; VDAC, voltage-dependent anion channel

Figure 3:

Enzymatic and non-enzymatic systems for the conversion and degradation of mitochondrial reactive oxygen species.

Superoxide (O2·-) as main ROS generated de novo is rapidly dismutated to H2O2, either non-enzymatically or by superoxide dimutases (SODs), which accelerate this reaction by a factor of about 10 000. H2O2 can leave the respective compartment by diffusing through membranes or in the case of the outer membrane by diffusing through porins/voltage-dependent anion channels (VDAC). This might play an important role during ROS signaling from mitochondria to the remainder of the cell. H2O2 can also undergo Fenton chemistry catalyzed by Fe2+ resulting in the generation of highly reactive hydroxyl radicals (·OH). However, the majority of H2O2 is presumably scavenged by multiple enzymatic and non-enzymatic systems, which convert H2O2 to water. These systems strongly differ between compartments and compete with each other and other proteins in the same compartment for reaction with H2O2. As electron donor these systems rely on NADPH, highlighting the importance of systems for NADPH regeneration in situ. APX, ascorbate peroxidase; αTCP, α-tocopherol; Ccp, cytochrome c peroxidase; CoQ10, coenzyme Q10; DHAR, dehydroascorbate reductase; Glr/GR, glutathione reductase; GPX/GPx/Gpx, glutathione peroxidase; GRX/Grx, glutaredoxin; GSH, glutathione; IDH/ICDH, isocitrate dehydrogenase; MDHAR, monodehydroascorbate reductase; NNT, nicotin nucleotid transhydrogenase; NTR/Trr/TXNRD, NADPH-thioredoxin reductase; PHGP, phospholipid hydroperoxide glutathione peroxidase; Por, porin; Prx/PRX, peroxiredoxin; Sod1/CSD1, copper-zinc superoxide dismutase; Sod2/MSD1, manganese superoxide dismutase; SRX, sulfiredoxin; TRX/Trx/TXN, thioredoxin; VDAC, voltage-dependent anion channel

In plants ascorbate has been well characterized as part of the ascorbate-glutathione cycle (Halliwell-Asada cycle). In the plant mitochondrial matrix this cycle encompasses the enzymes glutathione reductase 2 (GR2), monodehydroascorbate reductase (MDHAR), dehydroascorbate reductase (DHAR) and ascorbate peroxidase (APX) (Jimenez et al., 1997; Chew et al., 2003). DHAR draws electrons from GSH and MDHAR from NADPH to maintain a reduced ascorbate pool, which itself serves as electron donor for peroxide reduction via APX. Likewise electrons for Prx and GPx reduction are eventually provided by NADPH either through thioredoxins (Trx) and thioredoxin reductase (Trr), or glutaredoxins (Grx) and glutathione reductase (Glr). Despite the differences among the reducing systems in mitochondria of animals, fungi and plants, in all cases the rapid production of NADPH from different metabolic pathways is key for the efficient functioning of the reductive pathways (Figure 3). NADPH may be produced by different pathways including NADP-dependent isocitrate dehydrogenase, malic enzyme and methylenetetrahydrofolate dehydrogenase. In addition NADPH can be produced in the matrix harnessing the NADH pool, either through transhydrogenases or though NADH kinases, such as Pos5 in yeast. The individual contribution of each pathway under physiological conditions is not clear (Outten and Culotta, 2003; Fan et al., 2014).

While the repertoires of antioxidative enzymes of the mitochondrial matrix and the cytosol are largely known and characterized (Figure 3), the set of antioxidative enzymes of the IMS remains poorly defined. Recent proteomics studies of the yeast IMS identified a thioredoxin (Trx1), thioredoxin reductase (Trr1), the glutathione peroxidase Gpx3, cytochrome c peroxidase (Ccp1) and the copper/zinc SOD (Sod1) (Vögtle et al., 2012). In addition, we have identified Grx1 and Grx2 activity in the IMS of yeast cells (Kojer et al., 2014). In humans, Grx1, Sod1, TXNRD1 (Inarrea et al., 2007), GPx1 and GPx4 (Liang et al., 2009) were found to localize to the IMS (Pai et al., 2007). Even more incomplete is our knowledge on the plant IMS. The presence of a copper/zinc SOD (Cu/ZnSOD1, CSD1) has been suggested, but remains to be unambiguously shown (Huang et al., 2012).

Small redox molecules like alpha-tocopherol (vitamin E) and ubiquinone (coenzyme Q) support redox enzymes in their role in ROS depletion. These compounds are localized to the membranes and serve as scavengers of lipid peroxyl radicals. Therapeutic strategies using small molecules to reduce mitochondrial ROS levels are currently under intense development and testing (Smith et al., 2012).

In this context it is also important to note that H2O2 can (slowly) diffuse across membranes or (much faster) through porins/voltage-dependent anion channels (VDACs) in the outer membrane and presumably through aquaporins in the inner membrane. Hence, mitochondrial H2O2 has great potential to also influence reactions outside of mitochondria whereas the relevance for other mitochondria-generated ROS might be largely confined to the organelle.

Physiological role of ROS-handling enzymes in mammals

The importance of maintaining mitochondrial ROS at physiological levels has been demonstrated by specifically deleting selected mitochondrial redox enzymes in mice (Table 1). For instance, mice with targeted deficiencies in some members of the matrix-located thioredoxin-2/thioredoxin reductase-2/peroxiredoxin III axis and SOD2 develop severe phenotypes. The respective phenotypes include aberrations in embryonic brain, heart and blood cell development and neurodegeneration. In contrast, Grx2 deficient mice present more specific defects affecting lens epithelial cells (Wu et al., 2011), as well as cardiac tissue and skeletal muscle. These effects are probably a consequence of increased proton leakage and perturbed oxidative phosphorylation caused by dysregulation of glutathionylation and de-glutathionylation events in the electron transport chain complexes (Mailloux et al., 2013, 2014). For enzymes of the IMS, it appears to be more challenging to unequivocally assign distinct functions for redox enzymes in the control of ROS levels in the IMS as some of these, including SOD1, Grx1, TXNRD1, Gpx1 and Gpx4, are known to dually localize to the cytosol, IMS and other compartments of the cell (Table 1). Therefore, phenotypes and mechanisms obtained by reverse genetic studies in mice and cells need to be carefully interpreted as the contribution of the different compartments to a given phenotype can often not easily be attributed to these enzymes. This is, for instance, nicely illustrated for the short form of mammalian Gpx4 (also referred to as the ‘cytosolic’ form) that is present in the cytosol, nucleus and in the IMS, where it was found to be strongly associated with the outer leaflet of the inner membrane of mitochondria (Liang et al., 2009). As Gpx4 is efficiently reducing oxidized lipids in lipid bilayers, it is intriguing that cardiolipin, a phospholipid specific for the inner membrane of mitochondria, was found to be the first phospholipid to be oxidized in kidneys of inducible GPx4 knockout mice (Friedmann Angeli et al., 2014). The physiological role of Prdx6 is far from being clear as it fails to compensate for Gpx4 loss. This might be due to its dual function as Prdx6 does not directly act on oxidized esterified lipids and requires the release of fatty acid through its phospholipase A2 (PLA2) activity. Hence, Gpx4 and Prdx6 work via two different mechanisms as membrane repair via Prdx6 will cause a substantial loss of fatty acids from lipid molecules and thereby substantially impacts on membrane structure because of an increase of lysophospholipids (these are known to be strong detergents) in membranes. In contrast, Gpx4 acts directly on oxidized esterified fatty acids, thereby reducing them to the corresponding alcohols in membranes.

Table 1:

Summary of knockout mice for redox regulating enzymes with mitochondrial localization.

GeneLocalizationMajor pathophysiologic phenotype(s)References
SOD1Cytosol IMSIncreased susceptibility to ischemic and toxic insults; age-dependent skeletal muscle atrophy; glucose intolerance; altered bioenergetic function in miceHuang et al. (1997); Muller et al. (2006); Muscogiuri et al. (2013); Garratt et al. (2014)
SOD2MatrixNeurodegeneration and early postnatal death of mice; dilated cardiomyopathy; increased sensitivity of hemizygous mice to various stress-inducing agents and accelerated agingLi et al. (1995); Lebovitz et al. (1996); Flynn and Melov (2013)
Grx1Cytosol IMSImpaired NF-κB signaling; increased susceptibility to UVR-induced lens injury; attenuated actin polymerization and consequently impaired recruitment of neutrophils to inflammation sites and reduced bacterial handlingReynaert et al. (2006); Kronschläger et al. (2012); Sakai et al. (2012)
Grx2MatrixIncreased sensitivity to oxidative stress in primary mouse lens epithelial cells; increased levels of glutathionylated proteins and decreased oxidative phosphorylation in cardiac and skeletal muscleWu et al. (2011); Mailloux et al. (2013, 2014)
GPx1Cytosol IMSIncreased susceptibility to ischemic and toxic insults; increased atherosclerosis; increased insulin sensitivityCrack et al. (2001, 2006); Lewis et al. (2007); Wong et al. (2008); Loh et al. (2009)
GPx4Cytosol Nucleus Plasma membrane IMSEarly embryonic lethality; neurodegeneration in different brain regions; aberrant hair follicle development; increased thrombus formation in vitamin E-deprived endothelial-specific Gpx4 null mice; acute renal failureYant et al. (2003); Seiler et al. (2008); Wirth et al. (2010); Sengupta et al. (2013); Wortmann et al. (2013); Friedmann Angeli et al. (2014)
TXNRD1Cytosol IMSWidespread developmental retardation and embryonic death between E8.5 and E10.5; cerebellar hypoplasia; increased resistance of liver to acetaminophen poisoningJakupoglu et al. (2005); Soerensen et al. (2008); Patterson et al. (2013)
TXN2MatrixEmbryonic death at E10.5 and increased apoptosis rates in the developing brainNonn et al. (2003)
TXNRD2MatrixEmbryonic death hat E13.5 caused by impaired development of fetal blood cells and cardiac tissue; impaired cardiac function in response to transient ischemiaConrad et al. (2004); Horstkotte et al. (2011)
PrxIIIMatrixIncreased susceptibility to LPS-induced oxidative stress and decreased survival of macrophages; impaired synaptic plasticityLi et al. (2007, 2009)

Despite the myriad of reported phenotypes in enzymes contributing to mitochondrial ROS control (Table 1), it is still astonishing that almost nothing is known about the in vivo mechanisms of specific thiol switches in mitochondrial processes, including cell death signaling and bioenergetics that might be mediated by some of these enzymes in mammals.

Specific mitochondrial proteins are prone to undergo cysteine oxidation

Numerous in vitro studies have reported on the sensitivity of specific enzymes towards oxidants. The physiological relevance of these studies is often not clear as oxidative damage will arguably compromise the activity of any protein if sufficiently high levels of oxidants are applied. In the majority of cases studied, the inhibitory concentrations, for example of H2O2, are considerably larger than the concentrations found in vivo (Cocheme et al., 2011). Nonetheless, several recent proteomic analyses have reported oxidative (thiol) modifications on a number of mitochondrial enzymes, although in most cases it remains unclear which proportion of a given protein is modified and whether these modifications really influence mitochondrial pathways.

Redox regulation of the respiratory chain

The addition of H2O2 to cells was reported to preferentially damage (or modify) mitochondrial proteins (Garcia et al., 2010; Perluigi et al., 2010; Qin et al., 2011; Martinez-Acedo et al., 2012; Stauch et al., 2014). In several investigations H2O2 addition reduced respiratory activity and mitochondrial membrane potential. This might point at feedback control which limits electron flow in the respiratory chain under oxidative conditions in order to reduce the production of additional O2·- and, hence, H2O2. The way in which ROS interact with the respiratory chain is apparently highly complex and the numerous publications on the redox regulation of respiration have identified different enzymes of the electron transport chain as the primary responders to oxidation. A recent review provides a good summary of our current understanding of thiol-based redox regulation of the respiratory chain (Dröse et al., 2014).

Complex I (NADH ubiquinone oxidoreductase) has recently been found to feature a molecularly well-defined thiol redox switch that appears to be able to control respiration and superoxide production in vitro and in vivo (Chouchani et al., 2013). Cysteine residue 39 of the mammalian ND3 subunit of the complex, which is in proximity to the quinone binding site, is particularly susceptible to thiol modification under low oxygen conditions such as after ischemia in myocardial infarction and stroke (Galkin et al., 2008; Babot et al., 2014). Upon re-oxygenation (reperfusion) rapid reactivation of complex I is a major source of the deleterious ROS production in cardiomyocytes (Yellon and Hausenloy, 2007; Prime et al., 2009; Chouchani et al., 2014). Transient inactivation of complex I in these patients thus holds potential as a powerful therapeutic strategy. Hence, modification of this thiol in complex I (for example by nitrosylation) prevents reactivation of complex I and therefore protects against ischemia-reperfusion injury. The recently solved crystal structures of complex I (Hunte et al., 2010; Vinothkumar et al., 2014) will provide an excellent basis to unravel the molecular mechanisms by which this molecular switch on ND3 is controlled. It appears likely that this switch is also used under physiological conditions to prevent the burst of respiration upon rapid changes in the prevalent oxygen concentration of tissues. Conservation of the critical cysteine of ND3 across different eukaryotic kingdoms, such as animals and plants, is indicative of a fundamental and physiological role in regulating mitochondrial respiration beyond specific pathologies or stress conditions.

Redox regulation of mitochondrial enzymes with catalytic cysteine residues

A recent proteome-wide study identified oxidation-prone cysteine residues in mouse heart cells on the basis of their reactivity towards an iodoacetamide-containing compound (Weerapana et al., 2010). Interestingly, among the 50 most reactive cysteine residues in the cell, 19 are found in mitochondrial proteins. Proteins with such hyper-reactive cysteine residues include ‘professional’ redox enzymes such as thioredoxin Trx2 (also called Txn2) or the matrix superoxide dismutase Sod2. However, by far most mitochondrial proteins with hyper-reactive cysteine residues are metabolic enzymes, and the reactive cysteines are very often, but not always, critical residues in the reactive center. Examples of enzymes that may contain physiologically relevant thiol switches will be introduced in the following.

Aldehyde dehydrogenases

The aldehyde dehydrogenase Aldh2 was listed as the protein with the most reactive cysteine residue in mitochondria (Weerapana et al., 2010). Other aldehyde dehydrogenases were identified in the same screen, including Aldh5 and Aldh6. Aldehyde dehydrogenases, and in particular Aldh2, have been known to contain redox-sensitive thiol groups that are subject to oxidative inactivation (Loomes and Kitson, 1989; Moon et al., 2005; Wang et al., 2011) (Figure 4A). Recent studies on the reaction mechanism of aldehyde dehydrogenases indicated a transient covalent bond of the conserved reactive cysteine residue with the nicotinamide ring of NADP+ (Diaz-Sanchez et al., 2011; Tsybovsky et al., 2013).

Figure 4: Examples of proteins with potential thiol switches.(A) Structure of the active site of the human aldehyde dehydrogenase. The three adjacent cysteine residues are shown in green and purple, representing their reduced and oxidized states, respectively. They are close to the bound NADPH cofactors and it was suggested that one of them forms a covalent reaction intermediate with this cofactor during the reaction cycle. Oxidation of this cysteine would consequently abolish enzymatic activity. As the cysteine is deeply buried in the active center of the enzyme, it is likely that the oxidized form of the cysteine is stabilized because of it is partially inaccessible to reducing redox enzymes. Shown is the structure of chain A from PDB 1O02 (Perez-Miller and Hurley, 2003). (B) Potential thiol switch in plant alternative oxidase (AOX). AOX forms homodimers, which form an intermolecular disulfide bond in vitro (shown in green and purple). Reduction, which can be mediated by thioredoxins (TRX), opens the disulfide priming the dimer for activation by α-ketoacids (αKA). In the active form reduced coenzyme Q10 [CoQ10(red)] from the electron transport chain is oxidized [CoQ10(ox)] and its electrons are passed to oxygen, which is eventually reduced to water by the active sites. The structure model shown has been generated based on the crystal structure of Trypanosoma brucei AOX (PDB 3VV9) (Shiba et al., 2013) using the sequence of Arabidopsis thaliana AOX1a. The N-termini are highly flexible and T. brucei AOX does not contain the cysteine residue. This makes the depicted thiol-switch-dependant conformational change hypothetical.

Figure 4:

Examples of proteins with potential thiol switches.

(A) Structure of the active site of the human aldehyde dehydrogenase. The three adjacent cysteine residues are shown in green and purple, representing their reduced and oxidized states, respectively. They are close to the bound NADPH cofactors and it was suggested that one of them forms a covalent reaction intermediate with this cofactor during the reaction cycle. Oxidation of this cysteine would consequently abolish enzymatic activity. As the cysteine is deeply buried in the active center of the enzyme, it is likely that the oxidized form of the cysteine is stabilized because of it is partially inaccessible to reducing redox enzymes. Shown is the structure of chain A from PDB 1O02 (Perez-Miller and Hurley, 2003). (B) Potential thiol switch in plant alternative oxidase (AOX). AOX forms homodimers, which form an intermolecular disulfide bond in vitro (shown in green and purple). Reduction, which can be mediated by thioredoxins (TRX), opens the disulfide priming the dimer for activation by α-ketoacids (αKA). In the active form reduced coenzyme Q10 [CoQ10(red)] from the electron transport chain is oxidized [CoQ10(ox)] and its electrons are passed to oxygen, which is eventually reduced to water by the active sites. The structure model shown has been generated based on the crystal structure of Trypanosoma brucei AOX (PDB 3VV9) (Shiba et al., 2013) using the sequence of Arabidopsis thaliana AOX1a. The N-termini are highly flexible and T. brucei AOX does not contain the cysteine residue. This makes the depicted thiol-switch-dependant conformational change hypothetical.

Aldehyde dehydrogenases are critical for the detoxification of aldehydes, including those derived from lipid peroxidation. They are of high medical relevance as their inactivation causes alcohol-induced cell damage and cardiotoxicity (Chen et al., 2010). Polymorphisms in aldehyde dehydrogenases may explain why some alcoholism patients acquire certain organ-specific complications whereas others acquire different ones (Chao et al., 1997). Oxidative inactivation of Aldh2 is reversed by addition of dithiotreitol (DTT), which presumably resolves an inactivating disulfide in the active site (Wenzel et al., 2007). It is unclear whether the extreme redox-sensitivity of aldehyde dehydrogenases is the unavoidable consequence of their mode of action or whether it serves a purpose under physiological (or pathological) conditions to repress enzyme activity upon oxidative stress.

Mitochondrial thiolases

The mitochondrial matrix contains a number of thiolases (also known as acetyl-coenzyme A acetyltransferases) for the catabolic breakdown of fatty acids and other coenzyme A-activated metabolites. Cysteine residues of several mitochondrial thiolases such as acetoacetyl-CoA thiolase, ketoacyl-CoA thiolase and the 2-ketoacyl-CoA thiolase are among the most reactive protein thiols in mammalian cells (Weerapana et al., 2010). These enzymes are characterized by reactive cysteine residues in their active centers that form acyl-thioester reaction intermediates with their substrates (Modis and Wierenga, 1999; Kim and Battaile, 2002; Haapalainen et al., 2006). A number of recent studies suggest that oxidative modifications of these reactive cysteines are linked to diseases. For example the activity of the mitochondrial acetoacetyl-CoA thiolase was found to be reduced by 80% in colon cells of patients suffering from ulcerative colitis because of increased mitochondrial H2O2 levels (Santhanam et al., 2007). Reduced thiolase activity in ulcerative colitis was returned to normal by exposure to reductants (Santhanam et al., 2007). Redox-induced inactivation of the acetoacetyl-CoA thiolase was also reported for other cells under pathological conditions, such as in hepatocytes suffering from alcohol-mediated mitochondrial dysfunction or in cardiomyocytes of chronically diabetic rats (Grinblat et al., 1986; Moon et al., 2006).

Creatine kinase

Creatine kinases are expressed in many animal cells, particularly in those with high energy demand such as myocytes or neurons. The mitochondrial isoform of creatine kinase, which is localized in the intermembrane space, uses ATP to phosphorylate creatine. The resulting phosphocreatine diffuses into the cytosol where it is used by the two cytosolic isoforms of creatine kinase to phophorylate ADP. All three isoforms contain a highly conserved cysteine residue, which is important although not essential for activity (Okinaka et al., 1964). The cysteine residue of creatine kinase has a very low pKa of 5.6 so that it is predominantly present in the reactive thiolate anion form (Wang et al., 2001, 2006). This reactive cysteine was shown to be sensitive to H2O2 or glutathione disulfide and could be reactivated by the addition of DTT or glutathione (Suzuki et al., 1992; Thomas et al., 1994; Reddy et al., 2000; Wang et al., 2001). Overoxidation of creatine kinase was reported under pathological conditions for example in cancer or ischemic heart cells (Mekhfi et al., 1996; Choi et al., 2001). It is not known what percentage of the creatine kinase pool is oxidized in situ. However, as creatine kinase is rate-limiting for myocardial energetics (Gupta et al., 2013), oxidative inactivation may be very relevant in vivo.

Aconitase and homoaconitase

Aconitase catalyzes the conversion of citrate to isocitrate in the TCA cycle and contains an iron-sulfur cluster of the 4Fe-4S type in its active site. In contrast to 2Fe-2S clusters as found in ferredoxins, 4Fe-4S clusters are intrinsically labile and very sensitive towards oxidation. They also might lead to the generation of a hydrogen radical by a Fenton-like mechanism (Vasquez-Vivar et al., 2000). Numerous studies have reported the oxidative modification of aconitase and homoaconitase, which result in their inactivation during aging or in oxidative stress-related disorders (Bulteau et al., 2005). A number of recent articles indicated that the oxidative damage of aconitase might be of physiological relevance (Matasova and Popova, 2008; Lushchak et al., 2014; Stehling et al., 2014). In mammalian cells, there is a cytosolic isoform of aconitase which is also referred to as iron responsive protein that serves as transcription factor when it loses the 4Fe-4S cluster (Walden et al., 2006).

Studies on model organisms suggest that even under in vivo conditions considerable fractions of 4Fe-4S cluster-containing proteins carry cysteine modifications: in yeast 35% of homoaconitase was found to contain disulfides (Brandes et al., 2011). Similarly, 37% of aconitase was found to be thiolated in Escherichiacoli under unstressed conditions and this number increased to 68% after treatment of cultures with hypochlorous acid (Leichert et al., 2008).

Branched chain aminotransferase

The catalytic breakdown of branched chain amino acids (isoleucine, leucine, valine) is catalyzed by mitochondrial enzymes. In the first step of this pathway, the amino acids are converted by the mitochondrial isoform of branched chain aminotransferase (BCATm) to their respective α-ketoacids. In mammals, including humans, BCATm contains a cysteine pair (C315 and C318 in the human enzyme) that forms a disulfide bond under oxidizing conditions (Conway et al., 2002, 2003). C315 has a low pKa and is hence mainly present in a thiolate anion form that reacts efficiently with H2O2 to sulfenic acid and further to the disulfide (Conway et al., 2004). The same residue is also prone to nitrosylation by NO (Coles et al., 2009). Thiolation as well as nitrosylation thereby leads to the inactivation of the enzyme which is fully reversible upon reduction. Interestingly, this thiol switch is not present in closely related orthologs of nematodes, insects or bacteria, suggesting that it evolved in branch chain aminotransferases of mammals to allow redox-regulation of amino acid catabolism. A recent study proposed that BCATm can use its disulfide bond to oxidize other proteins in a reaction similar to that of the oxidoreductases Mia40 and protein disulfide isomerase and thus promote oxidative protein folding in the matrix (El Hindy et al., 2014). While this is an intriguing hypothesis, direct evidence for such a role under in vivo conditions in the mitochondrial matrix is still missing.

Alternative oxidase and other targets for thiol switching in plants

The respiratory chain of plant mitochondria contains an alternative oxidase (AOX) that passes electrons directly from ubiquinone to oxygen without pumping protons. AOX activity does not lead to ATP production (as no protons are pumped) and hence uncouples respiratory metabolism from energy conservation. The control of AOX activity is important to balance metabolic redox status and cellular energy charge, depending on nutrient status and environmental conditions of the plant. AOX is only fully active as a dimer in a reduced state, when it can be activated by α-keto acids, such as pyruvate (Millar et al., 1993) (Figure 4B). The requirement for reduction has been convincingly shown in vitro (Umbach and Siedow, 1993; Umbach et al., 1994, 2002; Rhoads et al., 1998) and can be overcome in mutants of the critical cysteine at position 78 (Cys-78). TRXs can catalyze the reduction in vitro (Gelhaye et al., 2004; Yoshida et al., 2013). As redox dependence of AOX activity has been predominantly studied in isolated mitochondria and protein thiols oxidize during mitochondrial isolation, the significance of this thiol switch under physiological conditions is still unclear. Overexpression of a Cys-78 mutant of AOX1a lacking the thiol switch in Arabidopsis does not show a phenotype as compared to over-expressors of the wild type form (although overexpression was not performed in the KO background meaning that a mix of wild type and mutant protein will be present)(Umbach et al., 2005). It has been argued that AOX may stably exist in its reduced state in vivo and that oxidation is merely an experimental artefact. However, in some plants [young soybean roots (Millar et al., 1998); wheat leaves (Bartoli et al., 2005)] an oxidized population of AOX is found that is reduced, i.e. activated, depending on developmental state and stress impact (drought). In other systems no oxidized AOX population has been found. The means of analyzing AOX thiol redox status in vivo require improvement to decide what physiological significance this thiol switching may have for plant AOX.

Two further candidates for matrix proteins with thiol switches of plants were recently discovered: citrate synthase (Schmidtmann et al., 2014) and NAD-dependent isocitrate dehydrogenase (Yoshida and Hisabori, 2014). Both can be reduced in a thioredoxin-mediated manner in vitro, and the induced thiol switch can strongly alter enzymatic activity, opening the possibility of redox dependent regulation. Yet, it remains unclear whether those thiol switches can regulate metabolic flux in vivo.

ROS and the redox state of cysteine residues with low reactivity – the relevance of enzymes for specificity and kinetics

How can H2O2 oxidize cysteine residues in a specific, localized and reversible manner? H2O2 reacts only slowly with most protein thiols (Winterbourn and Hampton, 2008) while catalytic thiols of dedicated detoxifying enzymes (catalase, GPx, Prx, APX) make exceptions from that rule (Forman et al., 2014). Despite the high reactivity and abundance of these H2O2-detoxifying enzymes, a diverse set of proteins containing cysteine residues of low reactivity towards H2O2 appear to be targets for H2O2-induced oxidation. Different models that are not mutually exclusive have been put forward to explain how this could be explained mechanistically (Figure 5). All of them emphasize the importance of indirect, enzyme-mediated transfer of redox status from oxidants or reductants onto target proteins.

Figure 5: Concepts for operation and maintenance of mitochondrial protein thiol switches based on cysteine residues with low reactivity towards hydrogen peroxide.The origins of redox perturbations that operate a mitochondrial protein thiol switch with low reactivity towards H2O2 are still not clearly resolved. As a basic requirement a thermodynamic drive to oxidize a reduced thiol switch or to reduce an oxidized one must exist, which is fulfilled in the mitochondrion by the presence of H2O2 and a sufficiently reducing NADPH pool, respectively. The balance between oxidizing and reducing pressures generates a functional steady-state of the thiol switches within a compartment. Different, distinct steady-states can be adopted by shifting the rate of oxidation and reduction, respectively. Changes can be controlled either kinetically [models (1), (3)] or thermodynamically [models (2), (4)], by modifying the oxidizing [models (1), (2)] or the reducing side [models (3), (4)]. This is illustrated by four models, which together provide a realistic concept for the operation and maintenance of protein thiol switches in vivo. See text for a detailed explanation of the individual models.

Figure 5:

Concepts for operation and maintenance of mitochondrial protein thiol switches based on cysteine residues with low reactivity towards hydrogen peroxide.

The origins of redox perturbations that operate a mitochondrial protein thiol switch with low reactivity towards H2O2 are still not clearly resolved. As a basic requirement a thermodynamic drive to oxidize a reduced thiol switch or to reduce an oxidized one must exist, which is fulfilled in the mitochondrion by the presence of H2O2 and a sufficiently reducing NADPH pool, respectively. The balance between oxidizing and reducing pressures generates a functional steady-state of the thiol switches within a compartment. Different, distinct steady-states can be adopted by shifting the rate of oxidation and reduction, respectively. Changes can be controlled either kinetically [models (1), (3)] or thermodynamically [models (2), (4)], by modifying the oxidizing [models (1), (2)] or the reducing side [models (3), (4)]. This is illustrated by four models, which together provide a realistic concept for the operation and maintenance of protein thiol switches in vivo. See text for a detailed explanation of the individual models.

  • The enzyme-mediated oxidation model Prx and GPx serve as mediators of H2O2 signaling. They react rapidly with H2O2 via their highly reactive thiols, which are oxidized in turn to subsequently transfer the oxidation to specific target molecules that otherwise have a low reactivity towards H2O2 directly. Given the dominating role of kinetic regulation in thiol redox biology, this type of regulation is likely to act as a major operator of physiological switches. One (non-mitochondrial) example for such a mode of action is the yeast transcription factor Yap1, which resides in the cytosol under non-stress conditions (Delaunay et al., 2002). Upon exposure of cells to H2O2, Yap1 receives a disulfide bond from GPx3, which has itself been oxidized by H2O2. This mediates re-localization of Yap1 to the nucleus to elicit its function in transcriptional regulation.

  • The floodgate model The generation of large amounts of H2O2 saturate (or inactivate by oxidation) the fast detoxifying pathways leaving slower-reacting thiol groups in target proteins time for reaction or allowing new functions for these ‘over-oxidized’ detoxifying enzymes (Wood et al., 2003; Kil et al., 2012). These more slowly reacting cysteine residues may either operate as thiol switches or fulfill a role as ‘redox buffer’ (Hansen et al., 2009; Brandes et al., 2011). In addition, micro-compartmentation and diffusion gradients may play a critical role: protein thiols with lower reactivity but localized in close proximity to the site of H2O2 generation may preferentially react with H2O2 when the more highly reactive H2O2 scavengers are localized far away. This model may, for instance, account for the mitochondrial IMS, which appears to contain only limited amounts of fast detoxifying systems (Kojer et al., 2014).

  • The oxidative influences are balanced and fine-tuned by reducing pathways, leading to a dynamic equilibrium between oxidizing and reducing inputs to set the duration and extent of cysteine residue oxidation as a result. The significance of the reductive pathways in the dynamic regulation of protein thiol switches are summarized in the following models:

  • The selective reduction model Despite sufficient thermodynamic drive for reduction, only specific cysteine residues are efficiently reduced by the Trx and Grx systems, because of steric, i.e. kinetic, constraints. This increases the extent and half-life of oxidation of non-target as compared to target cysteine residues, assuming equal rates of oxidation. A given cysteine residue may even switch between being a target and a non-target depending on structural re-organization of protein associations or complexes which alter the accessibility of the reduction machinery to the thiol switch.

  • The reductant exhaustion model Under conditions of overwhelming oxidation or upon insufficient flux through the metabolic pathways reducing NADP+, NADPH is depleted (oxidized, i.e. low NADPH/NADP+ ratio causing less reducing redox potential) and the thermodynamic drive from reduction decreases. This will result in increased amounts of oxidized thiols for prolonged times. Such an ‘NADPH depletion’ scenario is unlikely to be useful for physiological regulation as it is not specific to a given thiol switch; instead mitochondrial thiol switches will be oxidized in the order of their reactivity and if NADPH depletion persists long enough in the order of their midpoint potential starting from the most reducing couples (most negative potentials). Nevertheless more general responses may be triggered by exhaustion of reductant, for instance in mitochondrial pathology or in the early stages of mitochondria-mediated cell death.

Importantly, both models of oxidation (1+2) ultimately result in the oxidation of cysteine residues, even those with a low reactivity towards H2O2. Disulfides, sulfenic acids or glutathionylated cysteine residues that are formed in turn may serve in translating the H2O2 signal into a change in protein activity, localization or stability. The reductive pathways (3+4) balance oxidation generating a dynamic equilibrium in which the extent and duration of cysteine residue oxidation is shaped by all inputs (models 1, 2, 3, 4).

Reactive cysteine residues in the catalytic site of mitochondrial enzymes as critical mediators of H2O2-dependent regulation

A large number of (abundant) mitochondrial enzymes, such as the thiolases and dehydrogenases described above, contain highly reactive cysteine residues without being ‘professional’ H2O2 scavengers (see above). In these proteins the cysteines are often part of the active site and may be able to serve as acceptor sites even at low H2O2 concentrations (Figure 6). In contrast to peroxidases the oxidized cysteines in these proteins may be more stable as they are often buried in substrate-binding groves whereas peroxidases have evolved to interact efficiently with their specific electron acceptors. It is unknown which fraction of H2O2 reacts with these enzymes but there is good evidence that transient oxidation of these enzymes can regulate their activity. As the reactive cysteine residues typically contribute to catalytic activity their oxidation leads to inactivation. These enzymes can in turn sense ambient H2O2 levels directly and translate this information into a change in activity. It will be an exciting task to study the physiological relevance of such a thiol switch-mediated redox sensing in mitochondria in the future.

Figure 6: Enzymes with highly reactive cysteine residues might be direct targets of hydrogen peroxide.(A) H2O2 efficiently reacts with professional redox enzymes such as glutathione peroxidases or peroxiredoxins. (B) Protein thiols can also directly react with H2O2 resulting in higher oxidation states, such as sulfenic acid. However, their very low reactivity largely prevents this reaction, which is therefore presumably only relevant under highly oxidizing conditions or if mediated by other enzymes. (C) In contrast, enzymes with highly reactive cysteine residues, which are abundant in mitochondria, are good candidates to serve as direct targets for regulation by H2O2. Their reduction depends on glutaredoxins and thioredoxins, but the rates of their recovery under physiological conditions are not known. As reactive cysteine residues are typically crucial for catalytic activity, their oxidation leads to inactivation. Whether inactivation is just an unavoidable consequence of their reaction mechanism or serves a purpose for redox regulation remains to be explored.

Figure 6:

Enzymes with highly reactive cysteine residues might be direct targets of hydrogen peroxide.

(A) H2O2 efficiently reacts with professional redox enzymes such as glutathione peroxidases or peroxiredoxins. (B) Protein thiols can also directly react with H2O2 resulting in higher oxidation states, such as sulfenic acid. However, their very low reactivity largely prevents this reaction, which is therefore presumably only relevant under highly oxidizing conditions or if mediated by other enzymes. (C) In contrast, enzymes with highly reactive cysteine residues, which are abundant in mitochondria, are good candidates to serve as direct targets for regulation by H2O2. Their reduction depends on glutaredoxins and thioredoxins, but the rates of their recovery under physiological conditions are not known. As reactive cysteine residues are typically crucial for catalytic activity, their oxidation leads to inactivation. Whether inactivation is just an unavoidable consequence of their reaction mechanism or serves a purpose for redox regulation remains to be explored.


Corresponding authors: Jan Riemer, Cellular Biochemistry, University of Kaiserslautern, Erwin-Schrödinger-Str. 13, D-67663 Kaiserslautern, Germany, e-mail: ; Markus Schwarzländer, INRES – Chemical Signalling, University of Bonn, Friedrich-Ebert-Allee 144, D-53113 Bonn, Germany, e-mail: ; Marcus Conrad, Institute of Developmental Genetics, Helmholtz Center Munich, German Research Center for Environmental Health, Ingolstädter Landstraße 1, D-85764 Neuherberg, Germany, e-mail: ; and Johannes M. Herrmann, Cell Biology, University of Kaiserslautern, Erwin-Schrödinger-Str. 13, D-67633 Kaiserslautern, Germany, e-mail:

Acknowledgments

The authors thank Stephan Wagner and Michael W. Wöllhaf for help with the structural models in Figure 4, and Stefan Dröse and Bruce Morgan for critical reading and valuable comments on the manuscript. Work in the author’s laboratories is supported by the DFG in the framework of the Priority Program SPP1710, by the Emmy Noether Programme (SCHW 1719/1-1) and the Landesschwerpunkt BioComp.

References

Albrecht, S.C., Barata, A.G., Grosshans, J., Teleman, A.A., and Dick, T.P. (2011). In vivo mapping of hydrogen peroxide and oxidized glutathione reveals chemical and regional specificity of redox homeostasis. Cell Metab. 14, 819–829. Search in Google Scholar

Allen, S., Balabanidou, V., Sideris, D.P., Lisowsky, T., and Tokatlidis, K. (2005). Erv1 mediates the Mia40-dependent protein import pathway and provides a functional link to the respiratory chain by shuttling electrons to cytochrome c. J. Mol. Biol. 353, 937–944. Search in Google Scholar

Babot, M., Birch, A., Labarbuta, P., and Galkin, A. (2014). Characterisation of the active/de-active transition of mitochondrial complex I. Biochim. Biophys. Acta 1837, 1083–1092. Search in Google Scholar

Banci, L., Bertini, I., Cefaro, C., Ciofi-Baffoni, S., Gallo, A., Martinelli, M., Sideris, D.P., Katrakili, N., and Tokatlidis, K. (2009). Mia40 is an oxidoreductase that catalyzes oxidative protein folding in mitochondria. Nat. Struct. Mol. Biol. 16, 198–206. Search in Google Scholar

Bartoli, C.G., Gomez, F., Gergoff, G., Guiamet, J.J., and Puntarulo, S. (2005). Up-regulation of the mitochondrial alternative oxidase pathway enhances photosynthetic electron transport under drought conditions. J. Exp. Bot. 56, 1269–1276. Search in Google Scholar

Bien, M., Longen, S., Wagener, N., Chwalla, I., Herrmann, J.M., and Riemer, J. (2010). Mitochondrial disulfide bond formation is driven by intersubunit electron transfer in Erv1 and proof read by glutathione. Mol. Cell 37, 516–528. Search in Google Scholar

Bihlmaier, K., Mesecke, N., Terzyiska, N., Bien, M., Hell, K., and Herrmann, J.M. (2007). The disulfide relay system of mitochondria is connected to the respiratory chain. J. Cell Biol. 179, 389–395. Search in Google Scholar

Bleier, L., Wittig, I., Heide, H., Steger, M., Brandt, U., and Drose, S. (2014). Generator-specific targets of mitochondrial reactive oxygen species. Free Radic. Biol. Med. 78C, 1–10. Search in Google Scholar

Brandes, N., Reichmann, D., Tienson, H., Leichert, L.I., and Jakob, U. (2011). Using quantitative redox proteomics to dissect the yeast redoxome. J. Biol. Chem. 286, 41893–41903. Search in Google Scholar

Bulteau, A.L., Lundberg, K.C., Ikeda-Saito, M., Isaya, G., and Szweda, L.I. (2005). Reversible redox-dependent modulation of mitochondrial aconitase and proteolytic activity during in vivo cardiac ischemia/reperfusion. Proc. Natl. Acad. Sci. USA 102, 5987–5991. Search in Google Scholar

Calamita, G., Ferri, D., Gena, P., Liquori, G.E., Cavalier, A., Thomas, D., and Svelto, M. (2005). The inner mitochondrial membrane has aquaporin-8 water channels and is highly permeable to water. J. Biol. Chem. 280, 17149–17153. Search in Google Scholar

Chacinska, A., Pfannschmidt, S., Wiedemann, N., Kozjak, V., Sanjuan Szklarz, L.K., Schulze-Specking, A., Truscott, K.N., Guiard, B., Meisinger, C., and Pfanner, N. (2004). Essential role of Mia40 in import and assembly of mitochondrial intermembrane space proteins. EMBO J. 23, 3735–3746. Search in Google Scholar

Chandel, N.S., McClintock, D.S., Feliciano, C.E., Wood, T.M., Melendez, J.A., Rodriguez, A.M., and Schumacker, P.T. (2000). Reactive oxygen species generated at mitochondrial complex III stabilize hypoxia-inducible factor-1α during hypoxia: a mechanism of O2 sensing. J. Biol. Chem. 275, 25130–25138. Search in Google Scholar

Chao, Y.C., Young, T.H., Tang, H.S., and Hsu, C.T. (1997). Alcoholism and alcoholic organ damage and genetic polymorphisms of alcohol metabolizing enzymes in Chinese patients. Hepatology 25, 112–117. Search in Google Scholar

Chen, C.H., Sun, L., and Mochly-Rosen, D. (2010). Mitochondrial aldehyde dehydrogenase and cardiac diseases. Cardiovasc. Res. 88, 51–57. Search in Google Scholar

Chew, O., Whelan, J., and Millar, A.H. (2003). Molecular definition of the ascorbate-glutathione cycle in Arabidopsis mitochondria reveals dual targeting of antioxidant defenses in plants. J. Biol. Chem. 278, 46869–46877. Search in Google Scholar

Choi, H., Park, C.S., Kim, B.G., Cho, J.W., Park, J.B., Bae, Y.S., and Bae, D.S. (2001). Creatine kinase B is a target molecule of reactive oxygen species in cervical cancer. Mol. Cells 12, 412–417. Search in Google Scholar

Chouchani, E.T., Methner, C., Nadtochiy, S.M., Logan, A., Pell, V.R., Ding, S., James, A.M., Cocheme, H.M., Reinhold, J., Lilley, K.S., et al. (2013). Cardioprotection by S-nitrosation of a cysteine switch on mitochondrial complex I. Nat. Med. 19, 753–759. Search in Google Scholar

Chouchani, E.T., Pell, V.R., Gaude, E., Aksentijevic, D., Sundier, S.Y., Robb, E.L., Logan, A., Nadtochiy, S.M., Ord, E.N., Smith, A.C., et al. (2014). Ischaemic accumulation of succinate controls reperfusion injury through mitochondrial ROS. Nature 515, 431–435. Search in Google Scholar

Cocheme, H.M., Quin, C., McQuaker, S.J., Cabreiro, F., Logan, A., Prime, T.A., Abakumova, I., Patel, J.V., Fearnley, I.M., James, A.M., et al. (2011). Measurement of H2O2 within living Drosophila during aging using a ratiometric mass spectrometry probe targeted to the mitochondrial matrix. Cell Metab. 13, 340–350. Search in Google Scholar

Coles, S.J., Easton, P., Sharrod, H., Hutson, S.M., Hancock, J., Patel, V.B., and Conway, M.E. (2009). S-Nitrosoglutathione inactivation of the mitochondrial and cytosolic BCAT proteins: S-nitrosation and S-thiolation. Biochemistry 48, 645–656. Search in Google Scholar

Conrad, M., Jakupoglu, C., Moreno, S.G., Lippl, S., Banjac, A., Schneider, M., Beck, H., Hatzopoulos, A.K., Just, U., Sinowatz, F., et al. (2004). Essential role for mitochondrial thioredoxin reductase in hematopoiesis, heart development, and heart function. Mol. Cell. Biol. 24, 9414–9423. Search in Google Scholar

Conway, M.E., Yennawar, N., Wallin, R., Poole, L.B., and Hutson, S.M. (2002). Identification of a peroxide-sensitive redox switch at the CXXC motif in the human mitochondrial branched chain aminotransferase. Biochemistry 41, 9070–9078. Search in Google Scholar

Conway, M.E., Yennawar, N., Wallin, R., Poole, L.B., and Hutson, S.M. (2003). Human mitochondrial branched chain aminotransferase: structural basis for substrate specificity and role of redox active cysteines. Biochim. Biophys. Acta 1647, 61–65. Search in Google Scholar

Conway, M.E., Poole, L.B., and Hutson, S.M. (2004). Roles for cysteine residues in the regulatory CXXC motif of human mitochondrial branched chain aminotransferase enzyme. Biochemistry 43, 7356–7364. Search in Google Scholar

Cox, A.G., Peskin, A.V., Paton, L.N., Winterbourn, C.C., and Hampton, M.B. (2009). Redox potential and peroxide reactivity of human peroxiredoxin 3. Biochemistry 48, 6495–6501. Search in Google Scholar

Crack, P.J., Cimdins, K., Ali, U., Hertzog, P.J., and Iannello, R.C. (2006). Lack of glutathione peroxidase-1 exacerbates Aβ-mediated neurotoxicity in cortical neurons. J. Neural. Transm. 113, 645–657. Search in Google Scholar

Crack, P.J., Taylor, J.M., Flentjar, N.J., de Haan, J., Hertzog, P., Iannello, R.C., and Kola, I. (2001). Increased infarct size and exacerbated apoptosis in the glutathione peroxidase-1 (Gpx-1) knockout mouse brain in response to ischemia/reperfusion injury. J. Neurochem. 78, 1389–1399. Search in Google Scholar

Daum, B., Walter, A., Horst, A., Osiewacz, H.D., and Kuhlbrandt, W. (2013). Age-dependent dissociation of ATP synthase dimers and loss of inner-membrane cristae in mitochondria. Proc. Natl. Acad. Sci. USA 110, 15301–15306. Search in Google Scholar

Delaunay, A., Pflieger, D., Barrault, M.B., Vinh, J., and Toledano, M.B. (2002). A thiol peroxidase is an H2O2 receptor and redox-transducer in gene activation. Cell 111, 471–481. Search in Google Scholar

Diaz-Sanchez, A.G., Gonzalez-Segura, L., Rudino-Pinera, E., Lira-Rocha, A., Torres-Larios, A., and Munoz-Clares, R.A. (2011). Novel NADPH-cysteine covalent adduct found in the active site of an aldehyde dehydrogenase. Biochem. J. 439, 443–452. Search in Google Scholar

Dröse, S., Brandt, U., and Wittig, I. (2014). Mitochondrial respiratory chain complexes as sources and targets of thiol-based redox-regulation. Biochim. Biophys. Acta 1844, 1344–1354. Search in Google Scholar

Dubuisson, M., Vander Stricht, D., Clippe, A., Etienne, F., Nauser, T., Kissner, R., Koppenol, W.H., Rees, J.F., and Knoops, B. (2004). Human peroxiredoxin 5 is a peroxynitrite reductase. FEBS Lett. 571, 161–165. Search in Google Scholar

El Hindy, M., Hezwani, M., Corry, D., Hull, J., El Amraoui, F., Harris, M., Lee, C., Forshaw, T., Wilson, A., Mansbridge, A., et al. (2014). The branched-chain aminotransferase proteins: novel redox chaperones for protein disulfide isomerase-implications in Alzheimer’s disease. Antioxid. Redox. Signal. 20, 2497–2513. Search in Google Scholar

Fan, J., Ye, J., Kamphorst, J.J., Shlomi, T., Thompson, C.B., and Rabinowitz, J.D. (2014). Quantitative flux analysis reveals folate-dependent NADPH production. Nature 510, 298–302. Search in Google Scholar

Fischer, M., Horn, S., Belkacemi, A., Kojer, K., Petrungaro, C., Habich, M., Ali, M., Kuttner, V., Bien, M., Kauff, F., et al. (2013). Protein import and oxidative folding in the mitochondrial intermembrane space of intact mammalian cells. Mol. Biol. Cell. 24, 2160–2170. Search in Google Scholar

Flynn, J.M. and Melov, S. (2013). SOD2 in mitochondrial dysfunction and neurodegeneration. Free Radic. Biol. Med. 62, 4–12. Search in Google Scholar

Forman, H.J., Ursini, F., and Maiorino, M. (2014). An overview of mechanisms of redox signaling. J. Mol. Cell. Cardiol. 73, 2–9. Search in Google Scholar

Frey, T.G. and Mannella, C.A. (2000). The internal structure of mitochondria. Trends Biochem. Sci 25, 319–324. Search in Google Scholar

Friedmann Angeli, J.P., Schneider, M., Proneth, B., Tyurina, Y.Y., Tyurin, V.A., Hammond, V.J., Herbach, N., Aichler, M., Walch, A., Eggenhofer, E., et al. (2014). Inactivation of the ferroptosis regulator Gpx4 triggers acute renal failure in mice. Nat. Cell Biol. 16, 1180–1191. Search in Google Scholar

Galkin, A., Meyer, B., Wittig, I., Karas, M., Schagger, H., Vinogradov, A., and Brandt, U. (2008). Identification of the mitochondrial ND3 subunit as a structural component involved in the active/deactive enzyme transition of respiratory complex I. J. Biol. Chem. 283, 20907–20913. Search in Google Scholar

Garcia, J., Han, D., Sancheti, H., Yap, L.P., Kaplowitz, N., and Cadenas, E. (2010). Regulation of mitochondrial glutathione redox status and protein glutathionylation by respiratory substrates. J. Biol. Chem. 285, 39646–39654. Search in Google Scholar

Garratt, M., Pichaud, N., Glaros, E.N., Kee, A.J., and Brooks, R.C. (2014). Superoxide dismutase deficiency impairs olfactory sexual signaling and alters bioenergetic function in mice. Proc. Natl. Acad. Sci. USA 111, 8119–8124. Search in Google Scholar

Gas, E., Flores-Perez, U., Sauret-Gueto, S., and Rodriguez-Concepcion, M. (2009). Hunting for plant nitric oxide synthase provides new evidence of a central role for plastids in nitric oxide metabolism. Plant Cell 21, 18–23. Search in Google Scholar

Gelhaye, E., Rouhier, N., Gerard, J., Jolivet, Y., Gualberto, J., Navrot, N., Ohlsson, P.I., Wingsle, G., Hirasawa, M., Knaff, D.B., et al. (2004). A specific form of thioredoxin h occurs in plant mitochondria and regulates the alternative oxidase. Proc. Natl. Acad. Sci. USA 101, 14545–14550. Search in Google Scholar

Giulivi, C., Poderoso, J.J., and Boveris, A. (1998). Production of nitric oxide by mitochondria. J. Biol. Chem. 273, 11038–11043. Search in Google Scholar

Gladyshev, V.N. (2014). The free radical theory of aging is dead. Long live the damage theory! Antioxid. Redox. Signal. 20, 727–731. Search in Google Scholar

Goncalves, R.L., Quinlan, C.L., Perevoshchikova, I.V., Hey-Mogensen, M., and Brand, M.D. (2015). Sites of superoxide and hydrogen peroxide production by muscle mitochondria assessed ex vivo under conditions mimicking rest and exercise. J. Biol. Chem. 290, 209–227. Search in Google Scholar

Grant, C.M. (2001). Role of the glutathione/glutaredoxin and thioredoxin systems in yeast growth and response to stress conditions. Mol. Microbiol. 39, 533–541. Search in Google Scholar

Grinblat, L., Pacheco Bolanos, L.F. and Stoppani, A.O. (1986). Decreased rate of ketone-body oxidation and decreased activity of D-3-hydroxybutyrate dehydrogenase and succinyl-CoA:3-oxo-acid CoA-transferase in heart mitochondria of diabetic rats. Biochem. J. 240, 49–56. Search in Google Scholar

Gupta, K.J., Igamberdiev, A.U., Manjunatha, G., Segu, S., Moran, J.F., Neelawarne, B., Bauwe, H., and Kaiser, W.M. (2011). The emerging roles of nitric oxide (NO) in plant mitochondria. Plant Sci. 181, 520–526. Search in Google Scholar

Gupta, A., Rohlfsen, C., Leppo, M.K., Chacko, V.P., Wang, Y., Steenbergen, C., and Weiss, R.G. (2013). Creatine kinase-overexpression improves myocardial energetics, contractile dysfunction and survival in murine doxorubicin cardiotoxicity. PLoS One 8, e74675. Search in Google Scholar

Haapalainen, A.M., Merilainen, G., and Wierenga, R.K. (2006). The thiolase superfamily: condensing enzymes with diverse reaction specificities. Trends Biochem. Sci. 31, 64–71. Search in Google Scholar

Hansen, R.E., Roth, D., and Winther, J.R. (2009). Quantifying the global cellular thiol-disulfide status. Proc. Natl. Acad. Sci. USA 106, 422–427. Search in Google Scholar

Harner, M., Korner, C., Walther, D., Mokranjac, D., Kaesmacher, J., Welsch, U., Griffith, J., Mann, M., Reggiori, F., and Neupert, W. (2011). The mitochondrial contact site complex, a determinant of mitochondrial architecture. EMBO J. 30, 4356–4370. Search in Google Scholar

Herrmann, J.M. and Riemer, J. (2010). The intermembrane space of mitochondria. Antioxid. Redox. Signal. 13, 1341–1358. Search in Google Scholar

Hoppins, S., Collins, S.R., Cassidy-Stone, A., Hummel, E., Devay, R.M., Lackner, L.L., Westermann, B., Schuldiner, M., Weissman, J.S., and Nunnari, J. (2011). A mitochondrial-focused genetic interaction map reveals a scaffold-like complex required for inner membrane organization in mitochondria. J. Cell Biol. 195, 323–340. Search in Google Scholar

Horstkotte, J., Perisic, T., Schneider, M., Lange, P., Schroeder, M., Kiermayer, C., Hinkel, R., Ziegler, T., Mandal, P.K., David, R., et al. (2011). Mitochondrial thioredoxin reductase is essential for early postischemic myocardial protection. Circulation 124, 2892-U2327. Search in Google Scholar

Hu, J., Dong, L., and Outten, C.E. (2008). The redox environment in the mitochondrial intermembrane space is maintained separately from the cytosol and matrix. J. Biol. Chem. 283, 29126–29134. Search in Google Scholar

Huang, T.T., Yasunami, M., Carlson, E.J., Gillespie, A.M., Reaume, A.G., Hoffman, E.K., Chan, P.H., Scott, R.W., and Epstein, C.J. (1997). Superoxide-mediated cytotoxicity in superoxide dismutase-deficient fetal fibroblasts. Arch. Biochem. Biophys. 344, 424–432. Search in Google Scholar

Huang, C.H., Kuo, W.Y., Weiss, C., and Jinn, T.L. (2012). Copper chaperone-dependent and -independent activation of three copper-zinc superoxide dismutase homologs localized in different cellular compartments in Arabidopsis. Plant Physiol. 158, 737–746. Search in Google Scholar

Hunte, C., Zickermann, V., and Brandt, U. (2010). Functional modules and structural basis of conformational coupling in mitochondrial complex I. Science 329, 448–451. Search in Google Scholar

Inarrea, P., Moini, H., Han, D., Rettori, D., Aguilo, I., Alava, M.A., Iturralde, M., and Cadenas, E. (2007). Mitochondrial respiratory chain and thioredoxin reductase regulate intermembrane Cu,Zn-superoxide dismutase activity: implications for mitochondrial energy metabolism and apoptosis. Biochem. J. 405, 173–179. Search in Google Scholar

Jakupoglu, C., Przemeck, G.K.H., Schneider, M., Moreno, S.G., Mayr, N., Hatzopoulos, A.K., de Angelis, M.H., Wurst, W., Bornkamm, G.W., Brielmeier, M. et al. (2005). Cytoplasmic thioredoxin reductase is essential for embryogenesis but dispensable for cardiac development. Mol. Cell. Biol. 25, 1980–1988. Search in Google Scholar

Jimenez, A., Hernandez, J.A., Del Rio, L.A., and Sevilla, F. (1997). Evidence for the presence of the ascorbate-glutathione cycle in mitochondria and peroxisomes of pea leaves. Plant Physiol. 114, 275–284. Search in Google Scholar

Kawano, S., Yamano, K., Naoe, M., Momose, T., Terao, K., Nishikawa, S., Watanabe, N., and Endo, T. (2009). Structural basis of yeast Tim40/Mia40 as an oxidative translocator in the mitochondrial intermembrane space. Proc. Natl. Acad. Sci. USA 106, 14403–14407. Search in Google Scholar

Kil, I.S., Lee, S.K., Ryu, K.W., Woo, H.A., Hu, M.C., Bae, S.H., and Rhee, S.G. (2012). Feedback control of adrenal steroidogenesis via H2O2-dependent, reversible inactivation of peroxiredoxin III in mitochondria. Mol. Cell 46, 584–594. Search in Google Scholar

Kim, J.J. and Battaile, K.P. (2002). Burning fat: the structural basis of fatty acid beta-oxidation. Curr. Opin. Struct. Biol. 12, 721–728. Search in Google Scholar

Knoops, B., Goemaere, J., Van der Eecken, V., and Declercq, J.P. (2011). Peroxiredoxin 5: structure, mechanism, and function of the mammalian atypical 2-Cys peroxiredoxin. Antioxid. Redox. Signal. 15, 817–829. Search in Google Scholar

Koch, J.R. and Schmid, F.X. (2014). Mia40 targets cysteines in a hydrophobic environment to direct oxidative protein folding in the mitochondria. Nat. Commun. 5, 3041. Search in Google Scholar

Kojer, K., Bien, M., Gangel, H., Morgan, B., Dick, T.P., and Riemer, J. (2012). Glutathione redox potential in the mitochondrial intermembrane space is linked to the cytosol and impacts the Mia40 redox state. EMBO J. 31, 3169–3182. Search in Google Scholar

Kojer, K., Peleh, V., Calabrese, G., Herrmann, J.M., and Riemer, J. (2015). Kinetic control by limiting glutaredoxin amounts enables thiol oxidation in the reducing mitochondrial intermembrane space. Mol. Biol. Cell. 26, 195–204. Search in Google Scholar

Körner, C., Barrera, M., Dukanovic, J., Eydt, K., Harner, M., Rabl, R., Vogel, F., Rapaport, D., Neupert, W., and Reichert, A.S. (2012). The C-terminal domain of Fcj1 is required for formation of crista junctions and interacts with the TOB/SAM complex in mitochondria. Mol. Biol. Cell. 23, 2143–2155. Search in Google Scholar

Kronschläger, M., Galichanin, K., Ekstrom, J., Lou, M.F., and Söderberg, P.G. (2012). Protective effect of the thioltransferase gene on in vivo UVR-300 nm-induced cataract. Invest. Ophthalmol. Vis. Sci. 53, 248–252. Search in Google Scholar

Kumar, C., Igbaria, A., D’Autreaux, B., Planson, A.G., Junot, C., Godat, E., Bachhawat, A.K., Delaunay-Moisan, A. and Toledano, M.B. (2011). Glutathione revisited: a vital function in iron metabolism and ancillary role in thiol-redox control. EMBO J. 30, 2044–2056. Search in Google Scholar

Lebovitz, R.M., Zhang, H., Vogel, H., Cartwright, J., Jr., Dionne, L., Lu, N., Huang, S., and Matzuk, M.M. (1996). Neurodegeneration, myocardial injury, and perinatal death in mitochondrial superoxide dismutase-deficient mice. Proc. Natl. Acad. Sci. USA 93, 9782–9787. Search in Google Scholar

Leichert, L.I., Gehrke, F., Gudiseva, H.V., Blackwell, T., Ilbert, M., Walker, A.K., Strahler, J.R., Andrews, P.C., and Jakob, U. (2008). Quantifying changes in the thiol redox proteome upon oxidative stress in vivo. Proc. Natl. Acad. Sci. USA 105, 8197–8202. Search in Google Scholar

Lewis, P., Stefanovic, N., Pete, J., Calkin, A.C., Giunti, S., Thallas-Bonke, V., Jandeleit-Dahm, K.A., Allen, T.J., Kola, I., Cooper, M.E. et al. (2007). Lack of the antioxidant enzyme glutathione peroxidase-1 accelerates atherosclerosis in diabetic apolipoprotein E-deficient mice. Circulation 115, 2178–2187. Search in Google Scholar

Li, Y., Huang, T.T., Carlson, E.J., Melov, S., Ursell, P.C., Olson, J.L., Noble, L.J., Yoshimura, M.P., Berger, C., Chan, P.H., et al. (1995). Dilated cardiomyopathy and neonatal lethality in mutant mice lacking manganese superoxide dismutase. Nat. Genet. 11, 376–381. Search in Google Scholar

Li, L., Shoji, W., Takano, H., Nishimura, N., Aoki, Y., Takahashi, R., Goto, S., Kaifu, T., Takai, T., and Obinata, M. (2007). Increased susceptibility of MER5 (peroxiredoxin III) knockout mice to LPS-induced oxidative stress. Biochem. Biophys. Res. Commun. 355, 715–721. Search in Google Scholar

Li, L., Kaifu, T., Obinata, M., and Takai, T. (2009). Peroxiredoxin III-deficiency sensitizes macrophages to oxidative stress. J. Biochem. 145, 425–427. Search in Google Scholar

Liang, H., Yoo, S.E., Na, R., Walter, C.A., Richardson, A., and Ran, Q. (2009). Short form glutathione peroxidase 4 is the essential isoform required for survival and somatic mitochondrial functions. J. Biol. Chem. 284, 30836–30844. Search in Google Scholar

Loh, K., Deng, H., Fukushima, A., Cai, X., Boivin, B., Galic, S., Bruce, C., Shields, B.J., Skiba, B., Ooms, L.M., et al. (2009). Reactive oxygen species enhance insulin sensitivity. Cell Metab. 10, 260–272. Search in Google Scholar

Loomes, K.M. and Kitson, T.M. (1989). Reaction between sheep liver mitochondrial aldehyde dehydrogenase and various thiol-modifying reagents. Biochem. J. 261, 281–284. Search in Google Scholar

Lushchak, O.V., Piroddi, M., Galli, F., and Lushchak, V.I. (2014). Aconitase post-translational modification as a key in linkage between Krebs cycle, iron homeostasis, redox signaling, and metabolism of reactive oxygen species. Redox. Rep. 19, 8–15. Search in Google Scholar

Mailloux, R.J., Xuan, J.Y., Beauchamp, B., Jui, L.D., Lou, M., and Harper, M.E. (2013). Glutaredoxin-2 is required to control proton leak through uncoupling protein-3. J. Biol. Chem. 288, 8365–8379. Search in Google Scholar

Mailloux, R.J., Xuan, J.Y., McBride, S., Maharsy, W., Thorn, S., Holterman, C.E., Kennedy, C.R.J., Rippstein, P., deKemp, R., da Silva, J., et al. (2014). Glutaredoxin-2 is required to control oxidative phosphorylation in cardiac muscle by mediating deglutathionylation reactions. J. Biol. Chem. 289, 14812–14828. Search in Google Scholar

Marchissio, M.J., Frances, D.E., Carnovale, C.E. and Marinelli, R.A. (2012). Mitochondrial aquaporin-8 knockdown in human hepatoma HepG2 cells causes ROS-induced mitochondrial depolarization and loss of viability. Toxicol. Appl. Pharmacol. 264, 246–254. Search in Google Scholar

Martinez-Acedo, P., Nunez, E., Gomez, F.J., Moreno, M., Ramos, E., Izquierdo-Alvarez, A., Miro-Casas, E., Mesa, R., Rodriguez, P., Martinez-Ruiz, A., et al. (2012). A novel strategy for global analysis of the dynamic thiol redox proteome. Mol. Cell Proteomics 11, 800–813. Search in Google Scholar

Matasova, L.V. and Popova, T.N. (2008). Aconitate hydratase of mammals under oxidative stress. Biochemistry (Mosc) 73, 957–964. Search in Google Scholar

Mekhfi, H., Veksler, V., Mateo, P., Maupoil, V., Rochette, L., and Ventura-Clapier, R. (1996). Creatine kinase is the main target of reactive oxygen species in cardiac myofibrils. Circ. Res. 78, 1016–1027. Search in Google Scholar

Mesecke, N., Terziyska, N., Kozany, C., Baumann, F., Neupert, W., Hell, K., and Herrmann, J.M. (2005). A disulfide relay system in the intermembrane space of mitochondria that mediates protein import. Cell 121, 1059–1069. Search in Google Scholar

Milenkovic, D., Ramming, T., Muller, J.M., Wenz, L.S., Gebert, N., Schulze-Specking, A., Stojanovski, D., Rospert, S., and Chacinska, A. (2009). Identification of the signal directing Tim9 and Tim10 into the intermembrane space of mitochondria. Mol. Biol. Cell. 20, 2530–2539. Search in Google Scholar

Millar, A.H., Wiskich, J.T., Whelan, J., and Day, D.A. (1993). Organic acid activation of the alternative oxidase of plant mitochondria. FEBS Lett. 329, 259–262. Search in Google Scholar

Millar, A.H., Atkin, O.K., Ian Menz, R., Henry, B., Farquhar, G., and Day, D.A. (1998). Analysis of respiratory chain regulation in roots of soybean seedlings. Plant Physiol. 117, 1083–1093. Search in Google Scholar

Modis, Y. and Wierenga, R.K. (1999). A biosynthetic thiolase in complex with a reaction intermediate: the crystal structure provides new insights into the catalytic mechanism. Structure 7, 1279–1290. Search in Google Scholar

Moon, K.H., Kim, B.J., and Song, B.J. (2005). Inhibition of mitochondrial aldehyde dehydrogenase by nitric oxide-mediated S-nitrosylation. FEBS Lett. 579, 6115–6120. Search in Google Scholar

Moon, K.H., Hood, B.L., Kim, B.J., Hardwick, J.P., Conrads, T.P., Veenstra, T.D., and Song, B.J. (2006). Inactivation of oxidized and S-nitrosylated mitochondrial proteins in alcoholic fatty liver of rats. Hepatology 44, 1218–1230. Search in Google Scholar

Mouchiroud, L., Houtkooper, R.H., Moullan, N., Katsyuba, E., Ryu, D., Canto, C., Mottis, A., Jo, Y.S., Viswanathan, M., Schoonjans, K., et al. (2013). The NAD+/Sirtuin pathway modulates longevity through activation of mitochondrial UPR and FOXO signaling. Cell 154, 430–441. Search in Google Scholar

Muller, F.L., Song, W., Liu, Y., Chaudhuri, A., Pieke-Dahl, S., Strong, R., Huang, T.T., Epstein, C.J., Roberts, L.J., 2nd, Csete, M., et al. (2006). Absence of CuZn superoxide dismutase leads to elevated oxidative stress and acceleration of age-dependent skeletal muscle atrophy. Free Radic. Biol. Med. 40, 1993–2004. Search in Google Scholar

Murphy, M.P. (2009). How mitochondria produce reactive oxygen species. Biochem. J. 417, 1–13. Search in Google Scholar

Muscogiuri, G., Salmon, A.B., Aguayo-Mazzucato, C., Li, M., Balas, B., Guardado-Mendoza, R., Giaccari, A., Reddick, R.L., Reyna, S.M., Weir, G., et al. (2013). Genetic disruption of SOD1 gene causes glucose intolerance and impairs β-cell function. Diabetes 62, 4201–4207. Search in Google Scholar

Naoe, M., Ohwa, Y., Ishikawa, D., Ohshima, C., Nishikawa, S., Yamamoto, H., and Endo, T. (2004). Identification of Tim40 that mediates protein sorting to the mitochondrial intermembrane space. J. Biol. Chem. 279, 47815–47821. Search in Google Scholar

Nisoli, E., Clementi, E., Paolucci, C., Cozzi, V., Tonello, C., Sciorati, C., Bracale, R., Valerio, A., Francolini, M., Moncada, S., et al. (2003). Mitochondrial biogenesis in mammals: the role of endogenous nitric oxide. Science 299, 896–899. Search in Google Scholar

Nonn, L., Williams, R.R., Erickson, R.P., and Powis, G. (2003). The absence of mitochondrial thioredoxin 2 causes massive apoptosis, exencephaly, and early embryonic lethality in homozygous mice. Mol. Cell. Biol. 23, 916–922. Search in Google Scholar

Okinaka, S., Sugita, H., Momoi, H., Toyokura, Y., Watanabe, T., Ebashi, F., and Ebashi, S. (1964). Cysteine-stimulated serum creatine kinase in health and disease. J. Lab. Clin. Med. 64, 299–305. Search in Google Scholar

Outten, C.E. and Culotta, V.C. (2003). A novel NADH kinase is the mitochondrial source of NADPH in Saccharomyces cerevisiae. EMBO J. 22, 2015–2024. Search in Google Scholar

Pai, H.V., Starke, D.W., Lesnefsky, E.J., Hoppel, C.L., and Mieyal, J.J. (2007). What is the functional significance of the unique location of glutaredoxin 1 (GRx1) in the intermembrane space of mitochondria? Antioxid. Redox. Signal. 9, 2027–2033. Search in Google Scholar

Parsonage, D., Karplus, P.A., and Poole, L.B. (2008). Substrate specificity and redox potential of AhpC, a bacterial peroxiredoxin. Proc. Natl. Acad. Sci. USA 105, 8209–8214. Search in Google Scholar

Patterson, A.D., Carlson, B.A., Li, F., Bonzo, J.A., Yoo, M.H., Krausz, K.W., Conrad, M., Chen, C., Gonzalez, F.J., and Hatfield, D.L. (2013). Disruption of thioredoxin reductase 1 protects mice from acute acetaminophen-induced hepatotoxicity through enhanced NRF2 activity. Chem. Res. Toxicol. 26, 1088–1096. Search in Google Scholar

Perez-Miller, S.J. and Hurley, T.D. (2003). Coenzyme isomerization is integral to catalysis in aldehyde dehydrogenase. Biochemistry 42, 7100–7109. Search in Google Scholar

Perluigi, M., Di Domenico, F., Giorgi, A., Schinina, M.E., Coccia, R., Cini, C., Bellia, F., Cambria, M.T., Cornelius, C., Butterfield, D.A. et al. (2010). Redox proteomics in aging rat brain: involvement of mitochondrial reduced glutathione status and mitochondrial protein oxidation in the aging process. J. Neurosci. Res. 88, 3498–3507. Search in Google Scholar

Prime, T.A., Blaikie, F.H., Evans, C., Nadtochiy, S.M., James, A.M., Dahm, C.C., Vitturi, D.A., Patel, R.P., Hiley, C.R., Abakumova, I., et al. (2009). A mitochondria-targeted S-nitrosothiol modulates respiration, nitrosates thiols, and protects against ischemia-reperfusion injury. Proc. Natl. Acad. Sci. USA 106, 10764–10769. Search in Google Scholar

Qin, G., Liu, J., Cao, B., Li, B., and Tian, S. (2011). Hydrogen peroxide acts on sensitive mitochondrial proteins to induce death of a fungal pathogen revealed by proteomic analysis. PLoS One 6, e21945. Search in Google Scholar

Quinlan, C.L., Goncalves, R.L., Hey-Mogensen, M., Yadava, N., Bunik, V.I., and Brand, M.D. (2014). The 2-oxoacid dehydrogenase complexes in mitochondria can produce superoxide/hydrogen peroxide at much higher rates than complex I. J. Biol. Chem. 289, 8312–8325. Search in Google Scholar

Reddy, S., Jones, A.D., Cross, C.E., Wong, P.S., and Van Der Vliet, A. (2000). Inactivation of creatine kinase by S-glutathionylation of the active-site cysteine residue. Biochem. J. 347, 821–827. Search in Google Scholar

Reynaert, N.L., van der Vliet, A., Guala, A.S., McGovern, T., Hristova, M., Pantano, C., Heintz, N.H., Heim, J., Ho, Y.S., Matthews, D.E., et al. (2006). Dynamic redox control of NF-κB through glutaredoxin-regulated S-glutathionylation of inhibitory κB kinase β. Proc. Natl. Acad. Sci. USA 103, 13086–13091. Search in Google Scholar

Rhoads, D.M., Umbach, A.L., Sweet, C.R., Lennon, A.M., Rauch, G.S., and Siedow, J.N. (1998). Regulation of the cyanide-resistant alternative oxidase of plant mitochondria. Identification of the cysteine residue involved in α-keto acid stimulation and intersubunit disulfide bond formation. J. Biol. Chem. 273, 30750–30756. Search in Google Scholar

Sakai, J., Li, J., Subramanian, K.K., Mondal, S., Bajrami, B., Hattori, H., Jia, Y., Dickinson, B.C., Zhong, J., Ye, K., et al. (2012). Reactive oxygen species-induced actin glutathionylation controls actin dynamics in neutrophils. Immunity 37, 1037–1049. Search in Google Scholar

Santhanam, S., Venkatraman, A., and Ramakrishna, B.S. (2007). Impairment of mitochondrial acetoacetyl CoA thiolase activity in the colonic mucosa of patients with ulcerative colitis. Gut 56, 1543–1549. Search in Google Scholar

Schmidtmann, E., Konig, A.C., Orwat, A., Leister, D., Hartl, M. and Finkemeier, I. (2014). Redox regulation of Arabidopsis mitochondrial citrate synthase. Mol. Plant 7, 156–169. Search in Google Scholar

Schriner, S.E., Linford, N.J., Martin, G.M., Treuting, P., Ogburn, C.E., Emond, M., Coskun, P.E., Ladiges, W., Wolf, N., Van Remmen, H., et al. (2005). Extension of murine life span by overexpression of catalase targeted to mitochondria. Science 308, 1909–1911. Search in Google Scholar

Scorrano, L., Ashiya, M., Buttle, K., Weiler, S., Oakes, S.A., Mannella, C.A., and Korsmeyer, S.J. (2002). A distinct pathway remodels mitochondrial cristae and mobilizes cytochrome c during apoptosis. Dev. Cell 2, 55–67. Search in Google Scholar

Seiler, A., Schneider, M., Forster, H., Roth, S., Wirth, E.K., Culmsee, C., Plesnila, N., Kremmer, E., Radmark, O., Wurst, W., et al. (2008). Glutathione peroxidase 4 senses and translates oxidative stress into 12/15-lipoxygenase dependent- and AIF-mediated cell death. Cell Metab. 8, 237–248. Search in Google Scholar

Sengupta, A., Lichti, U.F., Carlson, B.A., Cataisson, C., Ryscavage, A.O., Mikulec, C., Conrad, M., Fischer, S.M., Hatfield, D.L., and Yuspa, S.H. (2013). Targeted disruption of glutathione peroxidase 4 in mouse skin epithelial cells impairs postnatal hair follicle morphogenesis that is partially rescued through inhibition of COX-2. J. Invest. Dermatol. 133, 1731–1741. Search in Google Scholar

Shiba, T., Kido, Y., Sakamoto, K., Inaoka, D.K., Tsuge, C., Tatsumi, R., Takahashi, G., Balogun, E.O., Nara, T., Aoki, T., et al. (2013). Structure of the trypanosome cyanide-insensitive alternative oxidase. Proc. Natl. Acad. Sci. USA 110, 4580–4585. Search in Google Scholar

Sies, H. (1993). Strategies of antioxidant defense. Eur. J. Biochem. 215, 213–219. Search in Google Scholar

Sies, H. (2014). Role of metabolic H2O2 generation: redox signaling and oxidative stress. J. Biol. Chem. 289, 8735–8741. Search in Google Scholar

Smith, R.A., Hartley, R.C., Cocheme, H.M., and Murphy, M.P. (2012). Mitochondrial pharmacology. Trends Pharmacol. Sci. 33, 341–352. Search in Google Scholar

Soerensen, J., Jakupoglu, C., Beck, H., Forster, H., Schmidt, J., Schmahl, W., Schweizer, U., Conrad, M., and Brielmeier, M. (2008). The role of thioredoxin reductases in brain development. PLoS One 3, e1813. Search in Google Scholar

Stauch, K.L., Purnell, P.R., and Fox, H.S. (2014). Quantitative proteomics of synaptic and nonsynaptic mitochondria: insights for synaptic mitochondrial vulnerability. J. Proteome Res. 13, 2620–2636. Search in Google Scholar

Stehling, O., Wilbrecht, C., and Lill, R. (2014). Mitochondrial iron-sulfur protein biogenesis and human disease. Biochimie 100, 61–77. Search in Google Scholar

Stoldt, S., Wenzel, D., Hildenbeutel, M., Wurm, C.A., Herrmann, J.M., and Jakobs, S. (2012). The inner-mitochondrial distribution of Oxa1 depends on the growth conditions and on the availability of substrates. Mol. Biol. Cell. 23, 2292–2301. Search in Google Scholar

Sturtz, L.A., Diekert, K., Jensen, L.T., Lill, R., and Culotta, V.C. (2001). A fraction of yeast Cu,Zn-superoxide dismutase and its metallochaperone, CCS, localize to the intermembrane space of mitochondria. A physiological role for SOD1 in guarding against mitochondrial oxidative damage. J. Biol. Chem. 276, 38084–38089. Search in Google Scholar

Suzuki, Y.J., Edmondson, J.D., and Ford, G.D. (1992). Inactivation of rabbit muscle creatine kinase by hydrogen peroxide. Free Radic. Res. Commun. 16, 131–136. Search in Google Scholar

Thomas, C., Carr, A.C., and Winterbourn, C.C. (1994). Free radical inactivation of rabbit muscle creatinine kinase: catalysis by physiological and hydrolyzed ICRF-187 (ICRF-198) iron chelates. Free Radic. Res. 21, 387–397. Search in Google Scholar

Toledano, M.B., Delaunay-Moisan, A., Outten, C.E., and Igbaria, A. (2013). Functions and cellular compartmentation of the thioredoxin and glutathione pathways in yeast. Antioxid. Redox. Signal. 18, 1699–1711. Search in Google Scholar

Trotter, E.W. and Grant, C.M. (2005). Overlapping roles of the cytoplasmic and mitochondrial redox regulatory systems in the yeast Saccharomyces cerevisiae. Eukaryot. Cell 4, 392–400. Search in Google Scholar

Tsybovsky, Y., Malakhau, Y., Strickland, K.C., and Krupenko, S.A. (2013). The mechanism of discrimination between oxidized and reduced coenzyme in the aldehyde dehydrogenase domain of Aldh1l1. Chem. Biol. Interact. 202, 62–69. Search in Google Scholar

Umbach, A.L. and Siedow, J.N. (1993). Covalent and noncovalent dimers of the cyanide-resistant alternative oxidase protein in higher plant mitochondria and their relationship to enzyme activity. Plant Physiol. 103, 845–854. Search in Google Scholar

Umbach, A.L., Wiskich, J.T., and Siedow, J.N. (1994). Regulation of alternative oxidase kinetics by pyruvate and intermolecular disulfide bond redox status in soybean seedling mitochondria. FEBS Lett. 348, 181–184. Search in Google Scholar

Umbach, A.L., Gonzalez-Meler, M.A., Sweet, C.R., and Siedow, J.N. (2002). Activation of the plant mitochondrial alternative oxidase: insights from site-directed mutagenesis. Biochim. Biophys. Acta 1554, 118–128. Search in Google Scholar

Umbach, A.L., Fiorani, F., and Siedow, J.N. (2005). Characterization of transformed Arabidopsis with altered alternative oxidase levels and analysis of effects on reactive oxygen species in tissue. Plant Physiol. 139, 1806–1820. Search in Google Scholar

Vasquez-Vivar, J., Kalyanaraman, B., and Kennedy, M.C. (2000). Mitochondrial aconitase is a source of hydroxyl radical. An electron spin resonance investigation. J. Biol. Chem. 275, 14064–14069. Search in Google Scholar

Vinothkumar, K.R., Zhu, J., and Hirst, J. (2014). Architecture of mammalian respiratory complex I. Nature 515, 80–84. Search in Google Scholar

Vogel, F., Bornhovd, C., Neupert, W., and Reichert, A.S. (2006). Dynamic subcompartmentalization of the mitochondrial inner membrane. J. Cell Biol. 175, 237–247. Search in Google Scholar

Vögtle, F.N., Burkhart, J.M., Rao, S., Gerbeth, C., Hinrichs, J., Martinou, J.C., Chacinska, A., Sickmann, A., Zahedi, R.P., and Meisinger, C. (2012). Intermembrane space proteome of yeast mitochondria. Mol. Cell Proteomics 11, 1840–1852. Search in Google Scholar

von der Malsburg, K., Muller, J.M., Bohnert, M., Oeljeklaus, S., Kwiatkowska, P., Becker, T., Loniewska-Lwowska, A., Wiese, S., Rao, S., Milenkovic, D., et al. (2011). Dual role of mitofilin in mitochondrial membrane organization and protein biogenesis. Dev. Cell 21, 694–707. Search in Google Scholar

Walden, W.E., Selezneva, A.I., Dupuy, J., Volbeda, A., Fontecilla-Camps, J.C., Theil, E.C., and Volz, K. (2006). Structure of dual function iron regulatory protein 1 complexed with ferritin IRE-RNA. Science 314, 1903–1908. Search in Google Scholar

Wang, P.F., McLeish, M.J., Kneen, M.M., Lee, G., and Kenyon, G.L. (2001). An unusually low pK(a) for Cys282 in the active site of human muscle creatine kinase. Biochemistry 40, 11698–11705. Search in Google Scholar

Wang, P.F., Flynn, A.J., Naor, M.M., Jensen, J.H., Cui, G., Merz, K.M., Jr., Kenyon, G.L., and McLeish, M.J. (2006). Exploring the role of the active site cysteine in human muscle creatine kinase. Biochemistry 45, 11464–11472. Search in Google Scholar

Wang, J., Wang, H., Hao, P., Xue, L., Wei, S., Zhang, Y., and Chen, Y. (2011). Inhibition of aldehyde dehydrogenase 2 by oxidative stress is associated with cardiac dysfunction in diabetic rats. Mol. Med. 17, 172–179. Search in Google Scholar

Weerapana, E., Wang, C., Simon, G.M., Richter, F., Khare, S., Dillon, M.B., Bachovchin, D.A., Mowen, K., Baker, D., and Cravatt, B.F. (2010). Quantitative reactivity profiling predicts functional cysteines in proteomes. Nature 468, 790–795. Search in Google Scholar

Wenzel, P., Hink, U., Oelze, M., Schuppan, S., Schaeuble, K., Schildknecht, S., Ho, K.K., Weiner, H., Bachschmid, M., Munzel, T. et al. (2007). Role of reduced lipoic acid in the redox regulation of mitochondrial aldehyde dehydrogenase (ALDH-2) activity. Implications for mitochondrial oxidative stress and nitrate tolerance. J. Biol. Chem. 282, 792–799. Search in Google Scholar

Winterbourn, C.C. and Hampton, M.B. (2008). Thiol chemistry and specificity in redox signaling. Free Radic. Biol. Med. 45, 549–561. Search in Google Scholar

Wirth, E.K., Conrad, M., Winterer, J., Wozny, C., Carlson, B.A., Roth, S., Schmitz, D., Bornkamm, G.W., Coppola, V., Tessarollo, L., et al. (2010). Neuronal selenoprotein expression is required for interneuron development and prevents seizures and neurodegeneration. FASEB J. 24, 844–852. Search in Google Scholar

Wong, C.H., Bozinovski, S., Hertzog, P.J., Hickey, M.J., and Crack, P.J. (2008). Absence of glutathione peroxidase-1 exacerbates cerebral ischemia-reperfusion injury by reducing post-ischemic microvascular perfusion. J. Neurochem. 107, 241–252. Search in Google Scholar

Wood, Z.A., Poole, L.B., and Karplus, P.A. (2003). Peroxiredoxin evolution and the regulation of hydrogen peroxide signaling. Science 300, 650–653. Search in Google Scholar

Wortmann, M., Schneider, M., Pircher, J., Hellfritsch, J., Aichler, M., Vegi, N., Kolle, P., Kuhlencordt, P., Walch, A., Pohl, U., et al. (2013). Combined deficiency in glutathione peroxidase 4 and vitamin E causes multiorgan thrombus formation and early death in mice. Circ. Res. 113, 408–417. Search in Google Scholar

Wu, H., Lin, L., Giblin, F., Ho, Y.S., and Lou, M.F. (2011). Glutaredoxin 2 knockout increases sensitivity to oxidative stress in mouse lens epithelial cells. Free Radic. Biol. Med. 51, 2108–2117. Search in Google Scholar

Wurm, C.A. and Jakobs, S. (2006). Differential protein distributions define two sub-compartments of the mitochondrial inner membrane in yeast. FEBS Lett. 580, 5628–5634. Search in Google Scholar

Yant, L.J., Ran, Q.T., Rao, L., Van Remmen, H., Shibatani, T., Belter, J.G., Motta, L., Richardson, A., and Prolla, T.A. (2003). The selenoprotein GPX4 is essential for mouse development and protects from radiation and oxidative damage insults. Free Radic. Biol. Med. 34, 496–502. Search in Google Scholar

Yee, C., Yang, W., and Hekimi, S. (2014). The intrinsic apoptosis pathway mediates the pro-longevity response to mitochondrial ROS in C. elegans. Cell 157, 897–909. Search in Google Scholar

Yellon, D.M. and Hausenloy, D.J. (2007). Myocardial reperfusion injury. N. Engl. J. Med. 357, 1121–1135. Search in Google Scholar

Yoshida, K. and Hisabori, T. (2014). Mitochondrial isocitrate dehydrogenase is inactivated upon oxidation and reactivated by thioredoxin-dependent reduction in Arabidopsis. Front. Environm. Sci. 2, Article 38, 1–7. Search in Google Scholar

Yoshida, K., Noguchi, K., Motohashi, K., and Hisabori, T. (2013). Systematic exploration of thioredoxin target proteins in plant mitochondria. Plant Cell. Physiol. 54, 875–892. Search in Google Scholar

Zarse, K., Schmeisser, S., Groth, M., Priebe, S., Beuster, G., Kuhlow, D., Guthke, R., Platzer, M., Kahn, C.R., and Ristow, M. (2012). Impaired insulin/IGF1 signaling extends life span by promoting mitochondrial L-proline catabolism to induce a transient ROS signal. Cell Metab. 15, 451–465. Search in Google Scholar

Received: 2014-12-2
Accepted: 2015-1-19
Published Online: 2015-1-27
Published in Print: 2015-5-1

©2015 by De Gruyter