Mitochondrial respiratory chain complexes as sources

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to label SNO sites with deuterium-NEM (d5) [43,115] (Fig. 2C). The amount of nitrosothiols in complex I upon treatment with MitoSNO. [42] in normoxia and ...
Biochimica et Biophysica Acta 1844 (2014) 1344–1354

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Review

Mitochondrial respiratory chain complexes as sources and targets of thiol-based redox-regulation☆ Stefan Dröse a, Ulrich Brandt b,d,⁎, Ilka Wittig c,d a

Clinic of Anesthesiology, Intensive-Care Medicine and Pain Therapy, University Hospital Frankfurt, 60590 Frankfurt am Main, Germany Radboud University Medical Centre, Nijmegen Centre for Mitochondrial Disorders, Geert Grooteplein-Zuid 10, 6525 GA Nijmegen, The Netherlands Functional Proteomics, SFB 815 Core Unit, Faculty of Medicine, Johann Wolfgang Goethe University, 60590 Frankfurt am Main, Germany d Cluster of Excellence “Macromolecular Complexes”, Goethe-University, Frankfurt am Main, Germany b c

a r t i c l e

i n f o

Article history: Received 6 January 2014 Received in revised form 5 February 2014 Accepted 8 February 2014 Available online 19 February 2014 Keywords: Mitochondria Respiratory chain complex Reactive oxygen species (ROS) Active/deactive transition S-nitrosylation Redox proteomics

a b s t r a c t The respiratory chain of the inner mitochondrial membrane is a unique assembly of protein complexes that transfers the electrons of reducing equivalents extracted from foodstuff to molecular oxygen to generate a proton-motive force as the primary energy source for cellular ATP-synthesis. Recent evidence indicates that redox reactions are also involved in regulating mitochondrial function via redox-modification of specific cysteine-thiol groups in subunits of respiratory chain complexes. Vice versa the generation of reactive oxygen species (ROS) by respiratory chain complexes may have an impact on the mitochondrial redox balance through reversible and irreversible thiol-modification of specific target proteins involved in redox signaling, but also pathophysiological processes. Recent evidence indicates that thiol-based redox regulation of the respiratory chain activity and especially S-nitrosylation of complex I could be a strategy to prevent elevated ROS production, oxidative damage and tissue necrosis during ischemia–reperfusion injury. This review focuses on the thiol-based redox processes involving the respiratory chain as a source as well as a target, including a general overview on mitochondria as highly compartmentalized redox organelles and on methods to investigate the redox state of mitochondrial proteins. This article is part of a Special Issue entitled: Thiol-Based Redox Processes. © 2014 Elsevier B.V. All rights reserved.

1. Introduction Mitochondria are extraordinary organelles holding key positions in a number of fundamental cellular processes including ATP-synthesis, biosynthetic pathways, ion homeostasis, oxygen sensing and apoptosis. All of these pathways encompass redox-reactions as central elements.

Abbreviations: A/D-transition, ʻactive/deactive-transition’ of mitochondrial complex I; Gpx1 and 4, glutathione peroxidases 1 and 4; DIGE, difference gel electrophoresis; GELSILOX, gel-based stable isotope labeling of oxidized cysteines; GR, glutathione reductase; GSH, glutathione; Grx2, glutaredoxin-2; GSNO, S-nitrosoglutathione; GSSG, glutathione-disulfide; IAA, iodoacetic acid; IAM, iodoacetamide; ICAT, isotope-coded affinity tag; IEF, isoelectric focusing; IMS, intermembrane space; LC, liquid-chromatography; MS, mass spectrometry; NEM, N-ethyl maleimide; SDS, sodium dodecylsulfate; NNT, nicotinamide nucleotide transhydrogenase; Δp, proton-motive force; Prx 3, peroxiredoxin 3; RET, reverse electron transfer; RNS, reactive nitrogen species; ROS, reactive oxygen species; TCA cycle, tricarboxylic acid cycle; TCEP, tris(2-carboxyethyl)phosphine; TOM, translocases of the outer membrane; Trx2, thioredoxin 2; TrxR2, thioredoxin reductase 2; VDAC, voltage-dependent anion channel ☆ This article is part of a Special Issue entitled: Thiol-Based Redox Processes. ⁎ Corresponding author at: Nijmegen Centre for Mitochondrial Disorders (NCMD), Radboud University Nijmegen Medical Centre, Geert Grooteplein-Zuid 10, Route 772, 6525 GA Nijmegen, The Netherlands. Tel.: +31 24 36 67098. E-mail address: [email protected] (U. Brandt).

http://dx.doi.org/10.1016/j.bbapap.2014.02.006 1570-9639/© 2014 Elsevier B.V. All rights reserved.

In addition, mitochondria contain major cellular generators of reactive oxygen species (ROS) that include components of the respiratory chain and a number of other redox enzymes [1–4], as well as powerful antioxidative defense systems [5–8] making mitochondria also a central player in cellular redox homeostasis. Elevated mitochondrial ROS production has been associated with a number of pathophysiological processes [9] including neurodegenerative diseases like Morbus Alzheimer and Morbus Parkinson [10], cancer [11,12] and oxidative damage during ischemia–reperfusion injury [13,14]. In addition, recent findings and novel concepts imply mitochondrial ROS as regulatory agents in a number of signal transduction pathways [15–17]. Hence, the functions of mitochondrial ROS seem to be highly ambivalent, deleterious and disease-causing on one side, taking part in physiological redox-regulation on the other. To unravel this ‘ROS paradoxon’ one faces two major challenges concerning the biochemistry of the elusive agents involved: (1) the sources and underlying molecular mechanisms of mitochondrial ROS production and their control under different physiological and pathophysiological circumstances have to be elucidated and (2) the ROS targets under these conditions have to be identified. Since oxidation and reduction of thiol proteins are thought to be the major mechanisms by which reactive oxidants integrate into cellular signal transduction pathways [18,19], recent research has focused on thiol-based redox modifications [20]. The processes and mechanisms

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that regulate such processes in mitochondria are largely unknown. However, it seems plausible that the respiratory chain should play a central role, since it comprises the major mitochondrial ROS generators complex I (NADH:ubiquinone oxidoreductase) [21–26], complex II (succinate:ubiquinone oxidoreductase) [27–29] and complex III (cytochrome bc1 complex; ubiquinol:cytochrome c oxidoreductase) [30–35]. On the other hand, specific cysteine thiols of respiratory chain complexes have been identified as targets of ROS and reactive nitrogen species (RNS) during oxidative stress [8,36–41]. Taken together these observations suggest a feed-back loop that uses reversible redox modifications of respiratory chain complexes to avoid irreversible oxidative damage caused by an elevated mitochondrial ROS production. A recent example is the reversible S-nitrosylation of complex I that is protective against myocardial ischemia/reperfusion damage [42,43]. This review intends to highlight different aspects of the intertwined relation between the respiratory chain and mitochondrial thiol-based redox processes: respiratory chain complexes as sources and targets of thiol-based redox-regulation, the influence of respiratory chain activity on the ‘general’ redox environment and redox-status of antioxidative defense systems. Relevant methods for the analysis of thiol-based redox processes in mitochondria will be discussed and the mitochondrial disulfide relay that facilitates the import of proteins into the intermembrane space including some respiratory chain complex subunits [44] will be briefly mentioned.

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2. Mitochondria are highly compartmentalized redox organelles It is evident from electron microscopy and especially cryo-electron tomography that mitochondria are highly compartmentalized organelles [45,46]. The mitochondrial matrix is surrounded by an outer membrane (OM) and an inner membrane (IM) separated by the intermembrane space (IMS; Fig. 1). The IM forms large invaginations into the matrix forming the so-called cristae that can develop into complex networks with different shapes [46]. The cristae are functionally separated from the inner boundary membrane by cristae junctions that limit the diffusion of IM proteins and IMS proteins [47]. The majority of respiratory chain complexes is localized in the cristae membranes which are shaped by highly organized respiratory chain supercomplexes [48] and rows of complex V (ATP synthase) oligomers [49–51]. The redox milieu – mainly determined by the redox status of glutathione – differs substantially between the mitochondrial compartments [52], thus placing the respiratory chain complexes at the boundary of two quite different redox environments: a reducing matrix and a relatively oxidizing IMS and cristae lumen [47] (Fig. 1). This ‘boundary effect’ has fundamental consequences for the distribution, reactivity and functionality of surface exposed cysteine thiols of mitochondrial proteins. As a result, the mitochondrial sub-compartments represent distinct reaction rooms that allow compartmentalized redox processes including redox signaling [53]. The different redox environments are governed by the distribution

Fig. 1. Mitochondrial redox compartmentalization. The redox environment of the mitochondrial compartments is influenced by the distribution of ROS sources (only complexes of the oxidative phosphorylation – indicated by Roman numerals I–V – are shown), components of the antioxidative defense (only GSH-related processes are shown) and their regenerating (reducing) systems. GSH is transported into the matrix by a not yet unambiguously identified transporter (GSH-T). In general, the intermembrane space (IMS) is more oxidizing than the cytosol and the matrix space (MS). It is not clear whether the redox environment of the cristae lumen (CRL) – separated by cristae junctions (CRJ) – differs from the peripheral IMS located between the inner boundary part of the inner membrane (IM) and the outer membrane (OM). Importantly, the regeneration of the GSH-pool in the IMS relies on activities of cytosolic enzymes and the cytosolic NADPH-pool, while the regeneration of the matrix GSH-pools and other components of the antioxidative defense (not shown) depends on activities of mitochondrial enzymes including nicotinamide nucleotide transhydrogenase (NNT) and the matrix NADPH-pool. The latter is eventually controlled by the combined activities all NADH-generating and -consuming processes (e.g. respiratory chain and TCA cycle, fueled by the substrates succinate (suc), fumarate (fum), malate (mal), oxaloacetate (oxal), citrate (cit), isocitrate (isocit), α-ketoglutarate (α-keto) and succinyl-CoA (suc-CoA)). Superoxide produced by complexes I and II is released into the matrix, while superoxide produced at the Qo-site of complex III is mainly released into the IMS. This primary ROS is converted into H2O2 by the activities of the superoxide dismutases SOD1 and SOD2, respectively. The relatively oxidizing environment in the IMS is important for the Mia40-Erv1 pathway that is also essential for the correct folding and assembly of respiratory chain complex subunits. For a detailed description of all aspects see text. TOM, translocases of the outer membrane; GR, glutathione reductase; Gpx1,4, glutathione peroxidases 1 and 4; FMN, flavin mononucleotide; FAD, flavin adenine dinucleotide; Q, ubiquinone; QH2, ubiquinol; IIQ, Q-binding site of complex II; Qo and Qi, ubiquinol oxidation and ubiquinone reduction centers of complex III; bH and bL, high potential and low potential cytochrome b, c1, cytochrome c1; CuA and CuB, copper A and copper B of complex IV.

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of glutathione (GSH), antioxidant enzymes, ROS generators and the NADPH pool essential for the regeneration of the antioxidative systems [54]. The latter is influenced by the supply of respiratory substrates [55] and hence, the activity of the respiratory chain complexes. 2.1. The interplay between the respiratory chain and the antioxidative defense systems/redox milieu in the matrix The redox micro-environments within different cellular compartments and organelles are mainly influenced by the GSH pool – or more precisely by the ratio of GSH and the oxidized dimer GSSG – and other thiol-containing proteins like the thioredoxin/peroxiredoxin system and glutaredoxins [8,18,53,54,56]. GSH is not synthesized in mitochondria, but has to be imported from the cytosol [57,58]. The redox state of the GSH/GSSG couple in mitochondria is about − 280 mV to − 330 mV as estimated with isolated organelles and tissue homogenates [59,60] and therefore more reduced than that in the cytoplasm (−260 mV to −200 mV) [54]. It has to be noted that the investigations with isolated mitochondria probably were dominated by the redox milieu of the matrix, since recent studies with targeted redox-sensitive fluorescent proteins show that the IMS is significantly more oxidizing than both the cytosol and the matrix [52,61]. All components of the antioxidative defense, i.e. the thioredoxin/ peroxiredoxin system encompassing thioredoxin reductase 2 (TrxR2), thioredoxin 2 (Trx2) and peroxiredoxin 3 (Prx 3) [18,53,54] as well as the GSH-dependent enzymes including glutathione peroxidases 1 and 4 (Gpx1, Gpx4), glutathione reductase (GR) and glutaredoxin-2 (Grx2) [8,53,54], rely on NADPH that serves as the common reductant for the oxidized forms of these proteins and GSSG. In mitochondria, the proton-motive force (Δp)-dependent nicotinamide nucleotide transhydrogenase (NNT) maintains the NADPH pool by utilizing NADH derived from the TCA cycle to reduce NADP+ to NADPH [62]. Other mitochondrial NADPH regenerating enzymes comprise the NADP+-dependent isocitrate dehydrogenase and the malic enzyme [63–65]. Under normal conditions, these processes together keep the mitochondrial NADPH pool in a more than 95% reduced state [8,54]. About half of this is uncoupler sensitive and represents the NNTdependent NADPH reduction since the reaction of NNT is driven by Δp [62,66]. Hence, NNT is influenced by the respiratory chain activity in two ways: (1) its activity depends critically on the membrane potential and (2) via the activity of mitochondrial NADH-oxidase, i.e. the concerted activities of complexes I, III and IV, this controls the level of mitochondrial NADH needed as a substrate for the transhydrogenase reaction. In this regard it is important to note that ultimately the redox status of matrix GSH — and probably also of Prx3 and Grx2 is regulated by the availability of catabolites [55] oxidized in the TCA cycle to generate NADH. With isolated mitochondria it has been shown that the efficiency of the antioxidative system is lower in the presence of succinate as compared to NADH-generating substrates like malate/glutamate [67,68]. In any case, the collective antioxidative defense in the matrix is very effective in scavenging different ROS species. This holds especially for superoxide anions and H2O2, the primary reactive species generated by the respiratory chain (see Section 2.3) [67–71]. Indeed, it has even been suggested that under ‘normal’ physiological conditions mitochondria are rather a sink for, than a major source for cellular ROS [6]. Recently, it has been proposed that surface exposed cysteine thiols of mitochondrial proteins may play an important role against oxidative damage [72]. Murphy and colleagues have estimated that within the mitochondrial matrix the concentration of exposed protein thiols is 60–90 mM [72] and therefore much higher than the GSH concentration of 1–5 mM [8]. This is an interesting consideration, however the sheer number of cysteine thiols might be not the most relevant parameter, since the direct reactivity of most cysteines with H2O2 or superoxide is very low [18,73]. Indeed, it has been shown that only few of the cysteines exposed on respiratory chain complexes do react with ROS

or thiol-reactive agents (see Section 5). Notably the non-catalyzed reaction of GSH with H2O2 is rather inefficient [18], but is greatly accelerated by glutathione peroxidases. Therefore, the combined scavenging activities of GSH and mitochondrial glutathione peroxidases and especially Prx3 [7] should be more important than the availability of surface cysteines on proteins in general. Notably, GSSG and oxidized Prx3 can be rereduced promptly by the activity of GR and Trx2/TrxR2, respectively. Nevertheless, a transient glutathionylation of surface cysteines might indeed be an important antioxidative defense, since this process could prevent the formation of higher, irreversible oxidation states [8,72,74]. In addition, the reactivity of a thiol group is critically dependent on its actual pKa value and the compartment pH, since most physiological oxidants react only with the thiolate anion [18,75]. In this respect it is important to note that respiratory chain activity raises the matrix pH thereby facilitating deprotonation of thiols. 2.2. The oxidative redox environment in the IMS is essential for thiol based protein import Recent investigations revealed that the mitochondrial intermembrane space represents a unique and important cellular compartment (for an excellent review see [47]). Because the IMS is connected via the porins (also called VDACs, voltage-dependent anion channels) with the cytosol, the physiological milieu of these two compartments is similar, but not identical [47]. The IMS is about 0.2–0.7 pH units more acidic and more oxidizing than the cytosol [52,61]. It has been demonstrated that the glutathione pools of IMS and cytosol are dynamically interconnected via the porins [76] and that GSSG is reduced to GSH by cytosolic glutathione reductase. This links the GSH-pool of the IMS to the cytosolic NADPH-pool. As mentioned above, the cristae lumen is functionally separated from the peripheral part of the intermembrane space by cristae junctions and it is conceivable that these two sub-compartments differ not only in their proteome, but also in their ‘small molecule inventory’ (e.g. H+, ROS, GSH/GSSG, metabolites) [47]. The oxidizing environment in the IMS is prerequisite for the oxidation-driven Mia40-Erv1 pathway that induces the oxidation of cysteine residues (i.e. the formation of disulfide bonds) of a defined group of proteins that enter this compartment via the TOM complex (for a recent review see [44]). The formation of intramolecular disulfide bonds triggers the folding of these proteins and keeps them in the IMS. The substrates of Mia40 are characterized by specific twin CX9C and twin CX3C motifs [44,47]. Such cysteine-rich motifs are also present in a number of respiratory chain subunits and have probably a function in the assembly and stability of the respective complexes. In addition to the complex III and IV subunits discussed by Herrmann and Riemer [44,47], mitochondrial complex I contains four subunits with cysteinerich motifs, three of them exhibiting the canonical twin Cx9C pattern [77,78]. These subunits do not contain mitochondrial targeting sequences and are probably located on the IMS-facing side of the membrane arm of complex I [78]. In vitro, glutathione plays an important role in substrate oxidation by Mia40 [79,80], since the presence of 5–10 mM glutathione strongly increases Mia40-dependent protein import into the IMS [44]. Recently it was shown that – mediated by exchange through the porins – the cytosolic glutathione pool influences the Mia40 redox state in vivo [76]. During catalysis, Mia40 is reduced and has to be re-oxidized by the sulfhydryloxidase Erv1. Erv1 finally transfers the electrons via cytochrome c and complex IV of the respiratory chain to molecular oxygen [76]. This suggests that the redox state of cytochrome c, which is mainly determined by respiratory chain activity, may have some impact on the Mia40-Erv1 pathway. 2.3. Respiratory chain complexes as ROS sources An important factor determining the redox state of the mitochondrial compartments is the production of ROS by distinct mitochondrial

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proteins. Main ROS generators have been localized in the respiratory chain [1,2,4], but also other oxidoreductases like the DLD (dihydrolipoamide dehydrogenase) component of pyruvate dehydrogenase and α-ketoglutarate dehydrogenase [81] and glycerol-3phosphate dehydrogenase [82,83] have been shown to produce ROS under certain conditions [3,6]. Within the respiratory chain, complexes I and III have been generally regarded as the main ROS producers [1–4]. However, recent investigations revealed that also mitochondrial complex II can be a significant ROS source, when the downstream respiratory chain is blocked and when the concentration of succinate and other dicarboxylates binding competitively to complex II is low [27–29]. Complex II can also modulate the ROS production of complex I and complex III, when succinate is the predominant substrate of the respiratory chain [84,85]. A fundamental factor affecting the mitochondrial redox environment is the directed release of ROS by the generators into different compartments (Fig. 1). Most respiratory chain complexes transfer one electron from reduced cofactors onto molecular oxygen which leads to the formation of superoxide as the primary ROS [1,4]. The charged, membrane-impermeable superoxide dismutates to hydrogen peroxide (H2O2) and water, a reaction that is greatly accelerated by the superoxide dismutases (SOD) present in the intermembrane space (SOD 1, Cu/Zn-SOD) and the matrix (SOD 2, Mn-SOD) (Fig. 1). The membrane-permeable H2O2 is generally regarded as the main ‘second messenger’ involved in thiol-modifications during redox signaling, since it has a longer lifetime due to its lower reactivity as compared to other ROS like superoxide or the hydroxyl radical. The latter ROS exhibit very short diffusion distances since they can react nonspecifically with different amino acid residues, lipids or nucleic acids [18,61,73,86]. Several studies indicate that complexes I and III release superoxide into different compartments [69,87,88]. Complex I produces ROS in two fundamentally different situations: (1) in the so called forward mode when the electrons are supplied by NADH and the downstream electron transfer is blocked either by an inhibitor like rotenone that binds to its Q-site or by inhibition of complex III or IV [22,24,25,84,89] and (2) during the so called reverse electron transfer (RET), when electrons are transferred from succinate by complex II or other dehydrogenases via ubiquinol to complex I, which requires a high membrane potential (Δ+ μ ) [21,23,90]. There is a general agreement that ROS are generated by electron transfer from FMNH2 in the forward mode, while there is an ongoing discussion whether this holds true also during RET [91] or whether instead electrons leak onto oxygen from a semiquinone in the Q-binding site of complex I [92]. In any case and in agreement with the experimental data [87,88], superoxide generated by complex I should be completely released into the matrix, since the structure of mitochondrial complex I revealed that both sites reside in the peripheral arm of complex I at some distance from the membrane plane [93]. In vitro, complex III produces ROS at the ubiquinol oxidation site (Qo site), if the ubiquinone reduction site is blocked by inhibitors like antimycin A [30–32,94,95]. While there is an ongoing controversial discussion about mechanistic details (whether superoxide is produced in a semi-forward mode from accumulated semiquinone or during a semi-reverse mode from the reduced cofactor heme bL [32,33,96]), structural considerations and experimental data indicate that superoxide from the Qo site is released primarily into the IMS or cristae lumen [69,87,88]. It is worthwhile to mention that especially ROS from this site have been linked to cellular redox signaling [15,33]. There is no direct experimental evidence pointing to the topology of ROS release from complex II. Yet, it seems likely that ROS produced at the flavin site (more precisely from the semi- or fully reduced FAD) of complex II [27,29] will be also released completely into the matrix [28]. However, mutagenesis studies in lower eukaryotes suggest that complex II can also produce ROS at its Q-binding site [97–99] making it impossible to draw any firm conclusions on the topology of ROS release by complex II.

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2.4. The mitochondrial redox compartments In summary, it can be concluded that the mitochondrial matrix and the IMS feature distinct redox environments that are controlled by the distribution of GSSG/GSH and thiol-containing proteins and that are influenced by the activity of the respective ROS sources (Fig. 1). The glutathione pool as the main redox buffer in the IMS relies on the activity of cytosolic enzymes and NADPH for the regeneration of GSH from GSSG. The main ROS source in the IMS is the Qo-site of complex III, but glycerol-3-phosphate dehydrogenase might also generate ROS in this compartment under certain conditions. The glutathione pool in the matrix is regulated by the activities of local NADP+-reducing enzymes and the matrix NADPH pool. NADPH is also essential for the reduction of the thioredoxin/peroxiredoxin system representing another, powerful ROS scavenging machinery. Main ROS sources in the matrix are complex I and probably also other NAD+-reducing dehydrogenases. 3. Mitochondrial redox proteomics 3.1. General aspects of thiol labeling for redox proteomics Reversible cysteine thiol oxidations/nitrosylations (inter- and intramolecular disulfides, S-gluathionylation, sulfenic acid, nitrosothiols) are among the most intensively studied oxidative and nitrosative modifications since they can modulate protein function [18,100]. High dynamics, instability and low abundance hamper direct identification of thiol modifications by mass spectrometry (MS). A long list of available thiol-reactive reagents provides a versatile tool box for detection, enrichment and quantification of such modifications by gel-based or gel-free redox proteomics (reviewed in [101]). In all cases, the redox status of a biological sample needs to be stabilized initially to obtain a snapshot of the steady-state oxidation levels at a given cellular or mitochondrial condition and to avoid artifacts by inadvertent oxidations during sample preparation [102]. This can be accomplished either by rapid protonation and precipitation of proteins with trichloroacetic acid [103] or by blocking free thiols with excess of the membrane permeable highly thiol-reactive reagent N-ethyl maleimide (NEM; Fig. 2A) [41,104–107]. In contrast to iodoacetamide (IAM) or iodoacetic acid (IAA) the reaction of NEM is more efficient at physiological pH enabling NEM-labeling of redox sensitive cysteines in their native conformation and physiological environment [104–106]. As recently described for mammalian complex I, this can be an advantage for targeted proteomics of thiol modifications of a specific protein or protein conformation in combination with functional studies [108]. Workflows for a comprehensive analysis of the cellular or mitochondrial redox proteome include a step completely denaturing the sample to unmask hidden thiols in native protein complexes. To achieve sufficient thiol-labeling under denaturing conditions a slightly alkaline pH is required to obtain the reactive thiolate form of the cysteines. At higher pH IAM and IAA are more thiol specific than NEM that exhibits some reactivity with other amino acid side chains [105]. Reversible thiol modifications (inter- and intramolecular disulfides, S-gluathionylation, S-nitrosylations, sulfenic acid) can be reduced by dithiothreitol or tris(2-carboxyethyl)phosphine (TCEP) following removal of excess thiol blocking reagent. Labeling with a different reagent then allows detection of these modifications (Fig. 2A). 3.2. Gel based methods Labeling with IAM or NEM coupled fluorescent dyes (Fluorescein, Cy3, Cy5) offers high sensitivity for detection of oxidized proteins separated by polyacrylamide gel electrophoresis (PAGE) in one or two dimensions. An example of this approach can be found in Galkin et al. [108], who applied fluorescein-NEM to identify the thiol that is accessible only after active/deactive transition of complex I using a 3D Blue

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Fig. 2. Methods for detection and quantification of redox modifications. A, Strategies for cysteine labeling of reversible redox/nitrosative modifications in proteins (R−) from tissues, intact cells or isolated mitochondria. The first step of sample preparation includes lysis and stabilization of redox state by labeling free thiols (R-S−) of cysteine residues with the first thiol probe (black ★). Reversible redox modifications S-sulfenylation (R-SOH), S-glutathionylation (R-S-SG), mixed disulfides (R-S-S-R′) and S-nitrosylation (R-SNO) are reduced by dithiothreitol or tris(2-carboxyethyl)phosphine (TCEP). Alternatively S-nitrosylations are selectively reduced by ascorbate and copper(II) [116,117]. The secondary thiol labels (blue or brown★) mark reversible oxidized cysteine residues of proteins. Irreversible redox modifications e.g. sulfinic acid (R-SO2H) or sulfonic acid (R-SO3H) cannot be reduced and labeled by thiol probes. B, to quantify redox modifications the first label is used to block free thiols and reversibly oxidized thiols are labeled differentially to compare two or more conditions. This can be achieved in a redox or SNO-DIGE approach (left) by using thiol reactive reagents coupled to fluorescent dyes Cy3 (green) or Cy5 (red) [113] or by using ICAT and quantitative mass spectrometry (right) [41,128]. C, following another strategy, the redox state of a cysteine thiol containing peptide can be analyzed by using probes containing light and heavy isotope coded thiol reactive reagents and quantitative mass spectrometry. This can be achieved by using light NEM (d0) as the label for free thiols (black) and heavy deuterium NEM (d5) as label for reversible oxidized modifications (gray) [43,115]. OxICAT uses the isotope-coded affinity tags (ICAT) to enrich light (brown) and heavy (orange) labeled peptides by biotin affinity columns and enable analysis of redox state in very complex protein mixtures [124]. For a detailed description see text. red, reduced cysteine peptide; ox, oxidized cysteine peptide; m/z, mass-to-charge ratio; IEF, isoelectric focusing.

Native/double SDS separation followed by identification of the modified thiol by MS (see Section 5.1). Classical 2D IEF/SDS gels separate complex mixtures of proteins according to their isoelectric point in the first dimensional isoelectric focusing (IEF) and according to their molecular mass in the second dimension (SDS-PAGE) to provide a detailed 2D-map of the proteome under study [109]. If according to the approach described above oxidized thiols are reduced following NEM treatment and are subsequently labeled by a fluorescent dye, the laser scanner will detect only reversibly oxidized proteins [110]. By introducing redox-DIGE, Hurd et al. [111] adapted the difference gel electrophoresis (2D-DIGE, [112]) approach for the detection of proteins differentially oxidized by mitochondrial ROS [111]. After blocking free thiols by NEM, reversibly oxidized proteins of control and redox-challenged mitochondria are reduced, then differentially alkylated with Cy3- or Cy5-maleimides and finally separated in the same 2D gel to identify ROS specific targets (Fig. 2B). Differentially labeled protein spots are then identified by MS. The use of internal standards like in 2D-DIGE [112] could allow quantitative comparison of redox-modifications under different conditions of ROS production to identify generator-specific protein targets in mitochondria. SNO-DIGE is a variant of redox-DIGE specifically designed to quantify S-nitrosylations [113]. This method uses ascorbate and copper(II) to allow for mild and specific reduction of SNO-sites that are subsequently labeled differentially with Cy3-NEM and Cy5-NEM. The introduction of

internal SNO standards enables quantification and comparison of more than two conditions within one SNO-DIGE experiment [114]. The redox- and SNO-DIGE methods give a comprehensive map of differentially oxidized and nitrosylated proteins, although MS identification of cysteines modified with fluorescent dyes is rarely achieved. 3.3. Quantitative redox proteomics by mass spectrometry In purely liquid-chromatography (LC)–MS based redox proteomics better detectable thiol labels are used. Cysteine containing peptides alkylated with NEM, IAM or IAA can be identified readily by LC–MS and have been used to distinguish between reduced and oxidized thiols in a qualitative manner (reviewed in [100]). In order to identify generator specific thiols for redox signaling quantitative redox proteomic approaches using stable isotope labeling are required. Differential cysteine labeling with stable isotopes produces chemically identical light and heavy peptides that elute at the same retention time in chromatographic separations, exhibit the same ionization properties, but can be discriminated and quantified based on a mass shift of neighboring heavy and light precursor ion peaks. Depending on the experimental design the heavy/light ratios reflect the redox state or the differential oxidation state under two conditions of a thiol containing peptide. Tandem MS/MS is used for identification of the peptides and the modified cysteines. A targeted approach to identify thiol modifications in

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complex I followed the principle of differential labeling with light NEM (d0) to block free thiols and mild reduction by ascorbate and copper(II) to label SNO sites with deuterium-NEM (d5) [43,115] (Fig. 2C). The amount of nitrosothiols in complex I upon treatment with MitoSNO [42] in normoxia and ischemia of intact mouse hearts was estimated by quantitative MS [43]. Comprehensive redox proteomic approaches use thiol reactive reagents with biotin affinity tags. In 2001 Jaffrey et al. introduced the biotin-switch assay to assess the S-nitrosylome [116]. This method is based on the conversion of SNO sites to biotin labeled thiols followed by affinity chromatography and protein identification [116,117]. The biotin-switch approach was later adapted to enrich other thiol modifications e.g. mixed redox modifications and sulfenic acid [118–120]. A very elegant way to quantify the thiol redox proteome by an MS based approach uses isotope-coded affinity tag (ICAT) technology. The ICAT reagent features an IAM-moiety for thiol labeling, an isotopic tag with a mass difference of 9 Da between the light 12C and heavy 13C form for quantitative MS and a cleavable biotin tag for affinity enrichment of cysteine containing peptides [121] (Fig. 2C). The ICAT technology has been adapted for redox proteomics of complex protein mixtures employing at least three strategies. The Cohen group used ICAT reagents to label free thiols of control and redox-treated samples and compared both experimental conditions by quantitative MS [122,123]. The OxICAT method introduced by Leichert et al. [124] monitors the oxidation state of a thiol by labeling free thiols with light, and oxidized cysteines with heavy ICAT [124–127]. A third strategy blocks free thiols with NEM and uses light and heavy ICAT to compare two experimental conditions [41,128] (Fig. 2B). The latter approach was used in a comprehensive redox proteomics study to identify and quantify changes in thiol oxidation during ischemia and reperfusion in Langendorff perfused isolated mouse hearts and many redox modified mitochondrial targets were identified [41]. All approaches to identify and quantify thiol modifications show advantages and limitations. The advantage of the LC–MS based redox ICAT methods is beyond any doubt efficient enrichment, identification and quantification of reduced and oxidized cysteine sites in a proteome or sub-proteome. However these methods do not allow for parallel estimation of the overall protein abundance and identification of irreversible thiol oxidations. As much as protein oxidation, changes in protein abundance due to protein degradation or protein translocation to another cellular compartment could be a consequence of redox signaling or oxidative stress. The estimation of the redox state of a peptide by OxICAT is to some extent independent of protein abundance, but cellular conditions that shift from redox signaling to oxidative stress cannot be monitored since over-oxidized cysteines escape detection. To overcome this limitation the recently reported method GELSILOX (GEL-based Stable Isotope Labeling of OXidized cysteines) combines quantification of redox modification and protein abundance by using standard thiol labels (NEM, IAM) and 16O/18O isotopic labeling to distinguish between experimental condition and protein abundance [129]. By direct LC–MS identification and quantification of sulfinic or sulfonic acid containing peptides, the GELSILOX approach could be easily expanded to also detect these irreversible redox modifications. Many high throughput methods to quantify redox modification are now available. However the major challenge in redox proteomics remains freezing the redox state during the first step of sample preparation to avoid inadvertent oxidations and artificial results. It is difficult in particular to find the right conditions for redox proteomics in cell culture experiments, since cell lines are adapted to atmospheric oxygen pressure, but the physiological processes underlying this adaptation are unknown. In order to determine the physiological relevance of thiol modifications for protein function, protein complex assembly or conformation, and protein translocation or degradation many additional targeted experiments are required in-vivo and in-vitro. To this end the recently developed complexome profiling approach [130] that allows studying

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protein abundance and protein complex remodeling could be useful to study the functional consequences of redox signaling and stress.

4. Targets of mitochondrial ROS A number of in vitro and in vivo investigations have addressed the question which mitochondrial proteins contain redox-sensitive thiols, i.e. are oxidized, glutathionylated or S-nitrosylated upon induction of more or less defined oxidative stressors. However, only few studies used a physiological stimulus or relied at least on endogenous mitochondrial ROS sources. Often oxidative stress was applied by the addition of external hydrogen peroxide, which might be generally appropriate to identify redox-sensitive mitochondrial proteins, but does not consider redox compartmentalization or discriminate between distinct ROS sources (see Section 2). Respiratory chain complexes that have been identified as targets of (mitochondrial) ROS are discussed in Section 5. In their fundamental investigation using Redox-DIGE (see Section 3) Hurd et al. [111] detected a number of proteins in isolated rat heart mitochondria that were modified by externally added H2O2, ROS generated by respiratory chain complexes I and III (induced by RET and antimycin A, respectively) and externally added RNS (i.e. S-nitroso-N-acetyl-DL-penicillamine, SNAP). In general, only few proteins were modified under all conditions and ROS production by antimycin or RET led to the oxidation of only three and six proteins, respectively [111]. Among the identified proteins were VDAC1, mitochondrial creatine kinase and proteins of mitochondrial fatty acid and pyruvate metabolism. Redox-modification of VDAC1 and enzymes involved in lipid metabolism and oxidative phosphorylation (see Section 5) have been identified in a study by Kumar et al. who investigated the thiol proteome in isolated mouse hearts following ischemia/ reperfusion applied with a Langendorff perfusion system [41]. In this study, also two TCA cycle enzymes (malate dehydrogenase and α-ketoglutarate dehydrogenase) came up as ROS-targets. Redoxsensitivity of α-ketoglutarate-dehydrogenase, i.e. the glutathionylation of the lipoic acid of subunit E2, has been detected also in an in vitro study using rat heart mitochondria [131–133]. A remarkable finding of the study of Kumar et al. [41] is that the oxidation of proteinthiols in hearts subjected to 20 min global ischemia was maximal after 5 min of reperfusion and was almost completely reverted after 30 min. This indicates that these modifications are largely reversible. Since the observed redox-modifications coincide with the oxidative burst that occurs upon reperfusion, complex I in RET mode seems to be the main source for these ROS [4,28]. A completely different approach was followed by Dick and coworkers, who used in situ kinetic trapping to detect proteins that interact with mitochondrial thioredoxin 2 (Trx2) [134]. In HEK293 cells, they expressed an inducible mutated form of Trx2 that was still capable of attacking disulfide bridges of target proteins, but deficient in resolving the formed mixed disulfide intermediates. In combination with a streptavidin tag, this allowed to selectively pull out Trx2 target proteins and identify them subsequently by mass spectrometry. Oxidative stress was mainly applied by external H2O2 and only one target (methionyltRNA synthase) was confirmed after induction of endogenous ROS by the combined application of rotenone and antimycin A [134]. This investigation suggests that the mitochondrial protein biosynthesis machinery is a major target of ROS. Other targets were chaperones, proteins of the amino acid metabolism and ATP/ADP translocase 2 that has been implicated also in mitochondrial permeability transition. ROS are also potent inductors of the enigmatic mitochondrial permeability transition pore [135,136] that contributes to different forms of apoptotic and necrotic cell death and leads to permeabilization of the mitochondrial membranes [137]. Irrespective to the recently proposed complex V dimers [138], it is still uncertain, which role other proposed mPTP-proteins like cyclophilin D, ATP/ADP translocase, the phosphate carrier and VDAC might play. Oxidation of cysteine has

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been shown to occur in the ‘classical’ mPTP component cyclophilin D [139,140]. Furthermore, it was recently shown that redox-modifications, i.e. glutathionylation, regulate the activities of mitochondrial uncoupling proteins [141–143]. 5. Thiol-based redox modification of respiratory chain complexes The reversible redox modification of protein-thiols is an important response to changes in the cellular redox environment. Since mitochondria are central to oxidative stress and redox signaling, S-glutathionylation and other thiol-modifications of mitochondrial proteins are of particular interest [8,72,74]. Especially thiol modifications of respiratory chain complexes have been detected in a number of studies. This initiated a number of in vitro studies and prompted a discussion on physiological functions and consequences that is still ongoing. 5.1. Complex I and the active/deactive transition Mitochondrial complex I has been identified in a number of studies as a target of oxidative thiol-modifications. Oxidation of cysteinethiols or S-glutathionylation of the iron–sulfur cluster containing subunits PSST (Ndufs7) and 51-kDa (Ndufv1) and the accessory subunit NUJM (Ndufa11) was observed in Langendorff perfused isolated mouse hearts during ischemia/reperfusion [41]. The 51-kDa subunit (Ndufv1) was found modified in the post-ischemic myocardium of rats [40]. Furthermore, reversible thiol oxidation of the 75-kDa subunit (Ndufs1) has been demonstrated in a study using kinetic trapping of Trx2-interacting proteins in HEK 293T cells [134]. It should be noted that the 51-kDa subunit carries FMN, the primary superoxide source of complex I, and resides in close proximity to the 75-kDa subunit. On the other hand, cardioprotective S-nitrosylation of complex I subunits by different NO-generating compounds (GSNO, MitoSNO1) has been detected in mouse and rat heart models [43,144]. The implications of these observations are discussed in detail further below. The effect of S-glutathionylation, S-nitrosylation and the formation of intramolecular disulfides on complex I activity has been investigated in a number of in vitro studies using purified complex I or ʻmitochondrial membranes’ (i.e. submitochondrial particles (SMP) mainly derived from bovine heart mitochondria) [38,40,42,145–150]. A general outcome of these studies is that oxidative thiol-modifications of complex I cysteines result in a general reduction of catalytic activity and that the modifications are relatively specific for distinct cysteines located in the 51 kDa- and 75 kDa-subunits of complex I. Only prolonged exposure of complex I to oxidative stress resulted in unspecific labeling of additional cysteines and eventually in the loss of iron–sulfur clusters. While initial experiments showed that S-glutathionylation results in an increased ROS-generation by complex I [145], further studies could not confirm this finding [38,148]. It has been proposed that oxidative modification of thiol groups in complex I and other proteins, either by S-glutathionylation or S-nitrosylation, is a protective mechanism to prevent higher oxidation of the thiyl-radical or sulfenic acid that is formed during the primary event [8].

Control of the catalytic activity of mitochondrial complex I by the socalled active/deactive (A/D) transition involves a specific and well defined cysteine switch mechanism. Kotlyar and Vinogradov were the first to observe the A/D transition in submitochondrial particles from bovine heart [151]. Mitochondrial complex I converts into an inactive state, the ‘deactive’ or D-form, when incubated at 37 °C in the absence of substrates. This form slowly turns back into the ‘active’ or A-form in the presence of NADH (or NADPH) and ubiquinone (Fig. 3). Later it was shown by Maklashina et al. that the A/D transition is a characteristic of complex I from vertebrates [152]. In the lower eukaryotes Yarrowia lipolytica and Neurospora crassa A/D transition of complex is also observed, but deactivation occurs much faster and at lower temperatures. As also shown by Vinogradov and coworkers, divalent cations like Ca2+ [153] and covalent modification of a single cysteine [154] can prevent reactivation of deactive complex I. Remarkably, the ion binding site and the cysteine are only accessible in the D-form of the enzyme rendering the A-form resistant to this type of inhibition (Fig. 3). This indicates that the A/D transition goes along with a significant conformational change. Anoxia/reperfusion studies by Maklashina et al. using Langendorff hearts gave the first indications that the A/D transition is more than an in vitro phenomenon [155]. Galkin and Moncada showed induction of the D-form of complex I during prolonged hypoxia and suggested that this could make the cysteine exposed only in this state accessible for nitrosation. Since under these conditions simultaneous generation of ROS by the respiratory chain complexes and of NO by NO-synthases could lead to the formation of significant amounts of the endogenous nitrosating agent peroxynitrite, this could result in permanent deactivation of the enzyme [156,157]. Murphy and coworkers recently showed marked cardioprotective effects in mice associated with the S-nitrosation of the cysteine switch of the A/D transition [42]. Taken together these results demonstrate that the A/D transition and the associated cysteine switch are operational in vivo. However, the physiological role of this regulatory mechanism remains obscure and it will be important to understand the molecular mechanism and control of the A/D transition in detail. The single cysteine that is only exposed in the deactive form of complex I was identified by site specific labeling and mass spectrometry as described in Section 3.2 [108]. The highly conserved residue was found in the hydrophilic loop connecting the first two transmembrane helices of subunit ND3 of complex I, one of the central subunits of complex I encoded by mitochondrial DNA. This subunit is part of the proximal domain of the membrane arm of complex I [158]. In the loop carrying the cysteine controlling the A/D transition, pathogenic mutations in humans causing mitochondrial disorders were reported at three positions (Fig. 4). The X-ray structure of complex I [158] reveals that loop 1 of subunit ND3 resides next to a β-sheet of the 49-kDa subunit of the peripheral arm (Fig. 4). It was shown by site-directed mutagenesis in the yeast genetic model Y. lipolytica [159] that this β-sheet is part of the ubiquinone and inhibitor binding pocket of complex I. Two histidines essential for catalytic activity are located within the loop connecting the first and second strands of the β-sheet [160,161]. Therefore, they seem to be part of the entry path for the ubiquinone head group into the catalytic site. This puts the A/D cysteine switch of complex I into an ideal position to control activity (Fig. 4). The analysis of conformation specific crosslinks suggests

Fig. 3. The active/deactive transition of mitochondrial complex I. If the active (A) form of complex I is left idle in the absence of substrate, it converts into the apparently inactive state, the deactive (D) form, that can be reactivated by adding substrates to induce catalytic turnover. Reactivation of the D-form to the A-form can be prevented by the binding of divalent cations (Me2+) or covalent modification (R) of a single cysteine by oxidation, nitrosylation or any kind of SH-reagent like N-ethylmaleimide of iodoacetamide.

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the S-glutathionylation protects from oxidative damage by tyrosine nitration [166,167]. 5.3. Complexes III, IV and V Redox-modification of respiratory chain complexes III, IV and V has been also observed in some in vitro studies. However, systematic investigations on functional consequences are largely missing. A redox modification within the core I subunit of complex III has been identified in isolated rat heart mitochondria after ROS induction at the Qo site by antimycin A [111] and S-nitrosylation of this subunit was induced by MitoSNO [113]. Cysteine thiols of Core II were found to be oxidized after ischemia/reperfusion in Langendorff-perfused mouse hearts [41]. Subunit Va of complex IV (cytochrome c oxidase) was glutathionylated in response to diamide treatment of human T-cells [168] and subunit Vb was glutathionylated in rat hepatocytes exposed to the redox cycler menadione [169]. Glutathionylation of the α1-subunit of complex V was observed in H2O2 challenged isolated brain and liver mitochondria [55]. The α1-subunit is also oxidized/S-nitrosylated after ischemia/ reperfusion [41] and GSNO treatment of Langendorff-perfused mouse hearts [144] and has been identified as a thioredoxin-2 target in H2O2 treated HEK293 cells [134]. Fig. 4. The loop of the ND3 subunit involved in the A/D transition of complex I. Structural model of the loop connecting transmembrane helices 1 and 2 of the mitochondrially encoded subunit ND3 and surrounding central subunits. The model was constructed using the Pymol software package from the structural coordinates of bacterial complex I from Thermus thermophilus (PDB ID: 4HEA) [158]. The highlighted residues were changed to the human variant where necessary using the Pymol mutagenesis wizard and the numbering of the human proteins is indicated. The ND3 subunit is shown in yellow and the 49-kDa subunit in blue. The cysteine involved in the A/D transition (magenta), two conserved acid residues in its vicinity (yellow) and three residues changed in mitochondrial disease (green) are shown in stick representation. The N-terminal-sheet of the 49-kDa subunit is highlighted in dark blue. The terminal iron–sulfur cluster N2 is depicted in space-fill representation.

that also the accessory 39-kDa subunit takes part in the structural changes associated with the A/D transition of mitochondrial complex I [162]. The 39-kDa subunit is of particular interest since it is homologous to short chain dehydrogenases and contains a NADPH binding site of unknown function [163–165]. Since the ROS production by mitochondrial complex I is directly linked to and controlled by its catalytic activity, studying the control of the A/D transition by the cysteine switch in subunit ND3 in detail will be prerequisite to investigate and understand its physiological role in the context of the redox signaling networks of eukaryotic cells. 5.2. Complex II Also the FAD-containing 70-kDa subunit (SdhA) of complex II has been identified as a target of redox modification following ischemia/ reperfusion in mouse heart [41]. Since mechanistic studies have shown that ROS are generated at the flavin site of complex II [27,29] when the downstream respiratory chain is blocked, cysteines that are in close proximity to FAD located in the 70-kDa subunit are likely the primary targets of these ROS. In studies with isolated complex II from bovine heart mitochondria and an in vivo rat heart ischemia/ reperfusion model it was observed that de-glutathionylation of the 70-kDa subunit (SdhA) of complex II results in a decrease of its catalytic activity [36]. S-glutathionylation of the 70-kDa subunit (SdhA) in purified complex II enhanced electron transfer activity and decreased the production of superoxide [36]. Superoxide production by complex II also induces self-inactivation by the formation of a CII-derived thiyl-radical. This implies that S-glutathionylation of complex II under ischemic conditions may have a protective effect by preserving electron transfer activity and preventing a vicious circle of ROSinduced self-inactivation [36]. In follow-up studies, it was shown that

5.4. Concluding remarks Recent years have brought a much better understanding of the mechanisms by which the respiratory chain complexes may generate ROS and how the antioxidative defense systems can modulate the redox environment in the different mitochondrial compartments. However, although numerous oxidative modifications of mitochondrial proteins have been reported, so far the evidence supporting a physiological relevance of such modifications is scarce. Indeed, even the widely accepted notion that the respiratory chain is a major source for mitochondrial ROS in vivo is still being challenged [170]. The application of the redox-proteomic approaches to analyze oxidative modifications at the level of individual proteins as summarized above will be required to elucidate the importance of mitochondrial ROS production and its consequences in living cells. Control of the A/D transition by specific modification of a single cysteine is a first example of how such studies can shed light on mechanisms of redox regulation by oxidative protein modification. Acknowledgements The author's work was supported by the Deutsche Forschungsgemeinschaft (SFB815 “Redox Regulation: Generator systems and functional consequences”, projects A02 and Z01). The authors thank Erik Bonke for drawing Fig. 1. References [1] M.P. Murphy, How mitochondria produce reactive oxygen species, Biochem. J. 417 (2009) 1–13. [2] A.J. Kowaltowski, N.C. Souza-Pinto, R.F. Castilho, A.E. Vercesi, Mitochondria and reactive oxygen species, Free Radic. Biol. Med. 47 (2009) 333–343. [3] M.D. Brand, The sites and topology of mitochondrial superoxide production, Exp. Gerontol. 45 (2010) 466–472. [4] S. Dröse, U. Brandt, Molecular mechanisms of superoxide production by the mitochondrial respiratory chain, Adv. Exp. Med. Biol. 748 (2012) 145–169. [5] A.I. Andreyev, Y.E. Kushnareva, A.A. Starkov, Mitochondrial metabolism of reactive oxygen species, Biochemistry (Mosc) 70 (2005) 200–214. [6] A.A. Starkov, The role of mitochondria in reactive oxygen species metabolism and signaling, Ann. N. Y. Acad. Sci. 1147 (2008) 37–52. [7] A.G. Cox, C.C. Winterbourn, M.B. Hampton, Mitochondrial peroxiredoxin involvement in antioxidant defence and redox signalling, Biochem. J. 425 (2010) 313–325. [8] M.P. Murphy, Mitochondrial thiols in antioxidant protection and redox signaling: distinct roles for glutathionylation and other thiol modifications, Antioxid. Redox Signal. 16 (2012) 476–495. [9] T.R. Figueira, M.H. Barros, A.A. Camargo, R.F. Castilho, J.C.B. Ferreira, A.J. Kowaltowski, F.E. Sluse, N.C. Souza-Pinto, A.E. Vercesi, Mitochondria as a source

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