Mitochondrial respiratory chain complexes as sources and targets of thiol-based redox-regulation

Biochimica et Biophysica Acta xxx (2014) xxx–xxx

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Review

Mitochondrial respiratory chain complexes as sources and targets ofthiol-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, Germanyb Radboud University Medical Centre, Nijmegen Centre for Mitochondrial Disorders, Geert Grooteplein-Zuid 10, 6525 GA Nijmegen, The Netherlandsc Functional Proteomics, SFB 815 Core Unit, Faculty of Medicine, Johann Wolfgang Goethe University, 60590 Frankfurt am Main, Germanyd Cluster of Excellence “Macromolecular Complexes”, Goethe-University, Frankfurt am Main, Germany

Abbreviations: A/D-transition, ʻactive/deactive-transitiGpx1 and 4, glutathione peroxidases 1 and 4; DIGE, dGELSILOX, gel-based stable isotope labeling of oxidizreductase; GSH, glutathione; Grx2, glutaredoxin-2; GSNglutathione-disulfide; IAA, iodoacetic acid; IAM, iodoacaffinity tag; IEF, isoelectric focusing; IMS, intermembrane spMS, mass spectrometry; NEM, N-ethyl maleimide; SDS, sotinamide nucleotide transhydrogenase; Δp, proton-motivRET, reverse electron transfer; RNS, reactive nitrogen specieTCA cycle, tricarboxylic acid cycle; TCEP, tris(2-carboxyethyof the outer membrane; Trx2, thioredoxin 2; TrxR2, thvoltage-dependent anion channel☆ This article is part of a Special Issue entitled: Thiol-Ba⁎ Corresponding author at: Nijmegen Centre for Mito

Radboud University Nijmegen Medical Centre, Geert Gr6525 GA Nijmegen, The Netherlands. Tel.: +31 24 36 670

E-mail address: [email protected] (U. Bran

http://dx.doi.org/10.1016/j.bbapap.2014.02.0061570-9639/© 2014 Elsevier B.V. All rights reserved.

Please cite this article as: S. Dröse, et al., MBiochim. Biophys. Acta (2014), http://dx.doi

a b s t r a c t
a r t i c l e i n f o
Article history:Received 6 January 2014Received in revised form 5 February 2014Accepted 8 February 2014Available online xxxx

Keywords:MitochondriaRespiratory chain complexReactive oxygen species (ROS)Active/deactive transitionS-nitrosylationRedox proteomics

The respiratory chain of the inner mitochondrial membrane is a unique assembly of protein complexes thattransfers the electrons of reducing equivalents extracted from foodstuff to molecular oxygen to generate aproton-motive force as the primary energy source for cellular ATP-synthesis. Recent evidence indicates thatredox reactions are also involved in regulating mitochondrial function via redox-modification of specificcysteine-thiol groups in subunits of respiratory chain complexes. Vice versa the generation of reactive oxygenspecies (ROS) by respiratory chain complexes may have an impact on the mitochondrial redox balance throughreversible and irreversible thiol-modification of specific target proteins involved in redox signaling, but alsopathophysiological processes. Recent evidence indicates that thiol-based redox regulation of the respiratorychain 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-basedredox processes involving the respiratory chain as a source as well as a target, including a general overviewon mitochondria as highly compartmentalized redox organelles and on methods to investigate the redox stateof 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 ina 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.

on’ of mitochondrial complex I;ifference gel electrophoresis;ed cysteines; GR, glutathioneO, S-nitrosoglutathione; GSSG,etamide; ICAT, isotope-codedace; LC, liquid-chromatography;dium dodecylsulfate; NNT, nico-e force; Prx 3, peroxiredoxin 3;s; ROS, reactive oxygen species;l)phosphine; TOM, translocasesioredoxin reductase 2; VDAC,

sed Redox Processes.chondrial Disorders (NCMD),ooteplein-Zuid 10, Route 772,98.dt).

itochondrial respiratory chai.org/10.1016/j.bbapap.2014.0

In addition, mitochondria contain major cellular generators of reac-tive oxygen species (ROS) that include components of the respiratorychain and a number of other redox enzymes [1–4], as well as power-ful antioxidative defense systems [5–8] making mitochondria also acentral player in cellular redox homeostasis. Elevated mitochondrialROS production has been associated with a number of pathophysiolog-ical processes [9] including neurodegenerative diseases like MorbusAlzheimer and Morbus Parkinson [10], cancer [11,12] and oxidativedamage during ischemia–reperfusion injury [13,14]. In addition, recentfindings and novel concepts imply mitochondrial ROS as regulatoryagents 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 physiologicalredox-regulation on the other. To unravel this ‘ROS paradoxon’ onefaces two major challenges concerning the biochemistry of the elusiveagents involved: (1) the sources and underlyingmolecular mechanismsofmitochondrial ROSproduction and their control under different phys-iological and pathophysiological circumstances have to be elucidatedand (2) the ROS targets under these conditions have to be identified.

Since oxidation and reduction of thiol proteins are thought to be themajor mechanisms by which reactive oxidants integrate into cellularsignal transduction pathways [18,19], recent research has focused onthiol-based redox modifications [20]. The processes and mechanisms

n complexes as sources and targets of thiol-based redox-regulation,2.006

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that regulate such processes in mitochondria are largely unknown.However, it seems plausible that the respiratory chain should playa central role, since it comprises the major mitochondrial ROS genera-tors complex I (NADH:ubiquinone oxidoreductase) [21–26], complexII (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 respiratorychain complexes have been identified as targets of ROS and reactive ni-trogen species (RNS) during oxidative stress [8,36–41]. Taken togetherthese observations suggest a feed-back loop that uses reversible redoxmodifications of respiratory chain complexes to avoid irreversible oxi-dative damage caused by an elevated mitochondrial ROS production.A recent example is the reversible S-nitrosylation of complex I that isprotective against myocardial ischemia/reperfusion damage [42,43].

This review intends to highlight different aspects of the intertwinedrelation between the respiratory chain and mitochondrial thiol-basedredox processes: respiratory chain complexes as sources and targets ofthiol-based redox-regulation, the influence of respiratory chain activityon the ‘general’ redox environment and redox-status of antioxidativedefense systems. Relevant methods for the analysis of thiol-basedredox processes in mitochondria will be discussed and the mitochon-drial disulfide relay that facilitates the import of proteins into the inter-membrane space including some respiratory chain complex subunits[44] will be briefly mentioned.

Fig. 1. Mitochondrial redox compartmentalization. The redox environment of the mitochondriaoxidative phosphorylation – indicated by Roman numerals I–V – are shown), components of t(reducing) systems. GSH is transported into the matrix by a not yet unambiguously identifiedthe cytosol and the matrix space (MS). It is not clear whether the redox environment of theIMS located between the inner boundary part of the inner membrane (IM) and the outer memof cytosolic enzymes and the cytosolic NADPH-pool, while the regeneration of thematrix GSH-pof mitochondrial enzymes including nicotinamide nucleotide transhydrogenase (NNT) and tNADH-generating and -consuming processes (e.g. respiratory chain and TCA cycle, fueled by(cit), isocitrate (isocit), α-ketoglutarate (α-keto) and succinyl-CoA (suc-CoA)). Superoxide ptheQo-site of complex III ismainly released into the IMS. This primary ROS is converted intoH2Ooxidizing environment in the IMS is important for theMia40-Erv1 pathway that is also essentiadescription of all aspects see text. TOM, translocases of the outermembrane; GR, glutathione redadenine dinucleotide; Q, ubiquinone; QH2, ubiquinol; IIQ, Q-binding site of complex II; Qo andpotential and low potential cytochrome b, c1, cytochrome c1; CuA and CuB, copper A and copp

Please cite this article as: S. Dröse, et al., Mitochondrial respiratory chaiBiochim. Biophys. Acta (2014), http://dx.doi.org/10.1016/j.bbapap.2014.0

2. Mitochondria are highly compartmentalized redox organelles

It is evident from electron microscopy and especially cryo-electrontomography thatmitochondria are highly compartmentalized organelles[45,46]. The mitochondrial matrix is surrounded by an outer membrane(OM) and an inner membrane (IM) separated by the intermembranespace (IMS; Fig. 1). The IM forms large invaginations into the matrixforming the so-called cristae that can develop into complex networkswith different shapes [46]. The cristae are functionally separated fromthe inner boundary membrane by cristae junctions that limit the diffu-sion of IM proteins and IMS proteins [47]. The majority of respiratorychain complexes is localized in the cristaemembranes which are shapedby highly organized respiratory chain supercomplexes [48] and rows ofcomplex V (ATP synthase) oligomers [49–51]. The redoxmilieu –mainlydetermined by the redox status of glutathione – differs substantiallybetween the mitochondrial compartments [52], thus placing the respi-ratory chain complexes at the boundary of two quite different redoxenvironments: a reducing matrix and a relatively oxidizing IMS andcristae lumen [47] (Fig. 1). This ‘boundary effect’ has fundamentalconsequences for the distribution, reactivity and functionality of surfaceexposed cysteine thiols of mitochondrial proteins. As a result, the mito-chondrial sub-compartments represent distinct reaction rooms thatallow compartmentalized redox processes including redox signaling[53]. The different redox environments are governed by the distribution

l compartments is influenced by the distribution of ROS sources (only complexes of thehe antioxidative defense (only GSH-related processes are shown) and their regeneratingtransporter (GSH-T). In general, the intermembrane space (IMS) is more oxidizing thancristae lumen (CRL) – separated by cristae junctions (CRJ) – differs from the peripheralbrane (OM). Importantly, the regeneration of the GSH-pool in the IMS relies on activitiesools and other components of the antioxidative defense (not shown) depends on activitieshe matrix NADPH-pool. The latter is eventually controlled by the combined activities allthe substrates succinate (suc), fumarate (fum), malate (mal), oxaloacetate (oxal), citrateroduced by complexes I and II is released into the matrix, while superoxide produced at2 by the activities of the superoxidedismutases SOD1and SOD2, respectively. The relativelyl for the correct folding and assembly of respiratory chain complex subunits. For a detaileductase; Gpx1,4, glutathione peroxidases 1 and 4; FMN, flavinmononucleotide; FAD, flavinQi, ubiquinol oxidation and ubiquinone reduction centers of complex III; bH and bL, higher B of complex IV.



3S. Dröse et al. / Biochimica et Biophysica Acta xxx (2014) xxx–xxx

of glutathione (GSH), antioxidant enzymes, ROS generators and theNADPH 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 antioxidativedefense systems/redox milieu in the matrix

The redox micro-environments within different cellular compart-ments and organelles are mainly influenced by the GSH pool – ormore precisely by the ratio of GSH and the oxidized dimer GSSG – andother thiol-containing proteins like the thioredoxin/peroxiredoxin sys-tem and glutaredoxins [8,18,53,54,56]. GSH is not synthesized in mito-chondria, but has to be imported from the cytosol [57,58]. The redoxstate of the GSH/GSSG couple in mitochondria is about −280 mV to−330 mV as estimated with isolated organelles and tissue homoge-nates [59,60] and therefore more reduced than that in the cytoplasm(−260 mV to−200mV) [54]. It has to be noted that the investigationswith isolated mitochondria probably were dominated by the redox mi-lieu of the matrix, since recent studies with targeted redox-sensitivefluorescent proteins show that the IMS is significantly more oxidizingthan 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 wellas the GSH-dependent enzymes including glutathione peroxidases 1and 4 (Gpx1, Gpx4), glutathione reductase (GR) and glutaredoxin-2(Grx2) [8,53,54], rely on NADPH that serves as the common reductantfor the oxidized forms of these proteins and GSSG. In mitochondria,the proton-motive force (Δp)-dependent nicotinamide nucleotidetranshydrogenase (NNT) maintains the NADPH pool by utilizingNADH derived from the TCA cycle to reduce NADP+ to NADPH [62].Other mitochondrial NADPH regenerating enzymes comprise theNADP+-dependent isocitrate dehydrogenase and the malic enzyme[63–65]. Under normal conditions, these processes together keep themitochondrial NADPH pool in a more than 95% reduced state [8,54].About half of this is uncoupler sensitive and represents the NNT-dependent NADPH reduction since the reaction of NNT is driven byΔp [62,66]. Hence, NNT is influenced by the respiratory chain activityin two ways: (1) its activity depends critically on themembrane poten-tial and (2) via the activity of mitochondrial NADH-oxidase, i.e. theconcerted activities of complexes I, III and IV, this controls the level ofmitochondrial NADH needed as a substrate for the transhydrogenasereaction. In this regard it is important to note that ultimately theredox status of matrix GSH — and probably also of Prx3 and Grx2 isregulated by the availability of catabolites [55] oxidized in the TCAcycle to generate NADH. With isolated mitochondria it has beenshown that the efficiency of the antioxidative system is lower in thepresence of succinate as compared to NADH-generating substrates likemalate/glutamate [67,68]. In any case, the collective antioxidativedefense in the matrix is very effective in scavenging different ROS spe-cies. This holds especially for superoxide anions and H2O2, the primaryreactive species generated by the respiratory chain (see Section 2.3)[67–71]. Indeed, it has even been suggested that under ‘normal’ physio-logical conditions mitochondria are rather a sink for, than a majorsource for cellular ROS [6].

Recently, it has been proposed that surface exposed cysteine thiolsof mitochondrial proteins may play an important role against oxidativedamage [72]. Murphy and colleagues have estimated that within themitochondrial matrix the concentration of exposed protein thiols is60–90 mM [72] and therefore much higher than the GSH concentrationof 1–5 mM [8]. This is an interesting consideration, however the sheernumber of cysteine thiols might be not the most relevant parameter,since the direct reactivity of most cysteines with H2O2 or superoxideis very low [18,73]. Indeed, it has been shown that only few of thecysteines exposed on respiratory chain complexes do react with ROS


or thiol-reactive agents (see Section 5). Notably the non-catalyzed reac-tion of GSHwithH2O2 is rather inefficient [18], but is greatly acceleratedby glutathione peroxidases. Therefore, the combined scavenging activi-ties of GSH and mitochondrial glutathione peroxidases and especiallyPrx3 [7] should bemore important than the availability of surface cyste-ines on proteins in general. Notably, GSSG and oxidized Prx3 can be re-reduced promptly by the activity of GR and Trx2/TrxR2, respectively.Nevertheless, a transient glutathionylation of surface cysteines mightindeed be an important antioxidative defense, since this process couldprevent the formation of higher, irreversible oxidation states [8,72,74].In addition, the reactivity of a thiol group is critically dependent on itsactual pKa value and the compartment pH, since most physiologicaloxidants react only with the thiolate anion [18,75]. In this respect it isimportant to note that respiratory chain activity raises the matrix pHthereby facilitating deprotonation of thiols.

2.2. The oxidative redox environment in the IMS is essential for thiol basedprotein import

Recent investigations revealed that the mitochondrial intermem-brane space represents a unique and important cellular compartment(for an excellent review see [47]). Because the IMS is connected viathe porins (also called VDACs, voltage-dependent anion channels)with the cytosol, the physiological milieu of these two compartmentsis similar, but not identical [47]. The IMS is about 0.2–0.7 pH unitsmore acidic and more oxidizing than the cytosol [52,61]. It has beendemonstrated that the glutathione pools of IMS and cytosol are dynam-ically interconnected via the porins [76] and that GSSG is reduced toGSH by cytosolic glutathione reductase. This links the GSH-pool of theIMS to the cytosolic NADPH-pool. As mentioned above, the cristaelumen is functionally separated from the peripheral part of the inter-membrane space by cristae junctions and it is conceivable that thesetwo sub-compartments differ not only in their proteome, but also intheir ‘small molecule inventory’ (e.g. H+, ROS, GSH/GSSG, metabolites)[47].

The oxidizing environment in the IMS is prerequisite for theoxidation-driven Mia40-Erv1 pathway that induces the oxidation ofcysteine residues (i.e. the formation of disulfide bonds) of a definedgroup of proteins that enter this compartment via the TOM complex(for a recent review see [44]). The formation of intramolecular disulfidebonds triggers the folding of these proteins and keeps them in the IMS.The substrates of Mia40 are characterized by specific twin CX9C andtwin CX3C motifs [44,47]. Such cysteine-rich motifs are also present ina number of respiratory chain subunits and have probably a functionin the assembly and stability of the respective complexes. In additionto the complex III and IV subunits discussed by Herrmann and Riemer[44,47], mitochondrial complex I contains four subunits with cysteine-rich motifs, three of them exhibiting the canonical twin Cx9C pattern[77,78]. These subunits do not contain mitochondrial targeting se-quences and are probably located on the IMS-facing side of the mem-brane arm of complex I [78]. In vitro, glutathione plays an importantrole in substrate oxidation by Mia40 [79,80], since the presence of5–10 mM glutathione strongly increases Mia40-dependent proteinimport into the IMS [44]. Recently it was shown that –mediated by ex-change through the porins – the cytosolic glutathione pool influencesthe Mia40 redox state in vivo [76]. During catalysis, Mia40 is reducedand has to be re-oxidized by the sulfhydryloxidase Erv1. Erv1 finallytransfers the electrons via cytochrome c and complex IV of the respira-tory chain tomolecular oxygen [76]. This suggests that the redox state ofcytochrome c, which ismainly 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 mitochon-drial compartments is the production of ROS by distinct mitochondrial




proteins. Main ROS generators have been localized in the respira-tory chain [1,2,4], but also other oxidoreductases like the DLD(dihydrolipoamide dehydrogenase) component of pyruvate dehydro-genase and α-ketoglutarate dehydrogenase [81] and glycerol-3-phosphate dehydrogenase [82,83] have been shown to produce ROSunder certain conditions [3,6]. Within the respiratory chain, complexesI and III have been generally regarded as the main ROS producers [1–4].However, recent investigations revealed that also mitochondrial com-plex II can be a significant ROS source, when the downstream respirato-ry chain is blocked and when the concentration of succinate and otherdicarboxylates binding competitively to complex II is low [27–29].Complex II can alsomodulate the ROS production of complex I and com-plex III, when succinate is the predominant substrate of the respiratorychain [84,85].

A fundamental factor affecting the mitochondrial redox environ-ment is the directed release of ROS by the generators into differentcompartments (Fig. 1). Most respiratory chain complexes transferone electron from reduced cofactors onto molecular oxygen whichleads to the formation of superoxide as the primary ROS [1,4]. Thecharged, membrane-impermeable superoxide dismutates to hydrogenperoxide (H2O2) and water, a reaction that is greatly acceleratedby the superoxide dismutases (SOD) present in the intermembranespace (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 sig-naling, since it has a longer lifetime due to its lower reactivity as com-pared to other ROS like superoxide or the hydroxyl radical. The latterROS exhibit very short diffusion distances since they can react non-specifically with different amino acid residues, lipids or nucleic acids[18,61,73,86].

Several studies indicate that complexes I and III release superoxideinto different compartments [69,87,88]. Complex I produces ROS intwo fundamentally different situations: (1) in the so called forwardmode when the electrons are supplied by NADH and the downstreamelectron transfer is blocked either by an inhibitor like rotenone thatbinds 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 elec-trons are transferred from succinate by complex II or other dehydroge-nases via ubiquinol to complex I, which requires a high membranepotential (Δμ

+) [21,23,90]. There is a general agreement that ROS aregenerated by electron transfer from FMNH2 in the forward mode,while there is an ongoing discussion whether this holds true alsoduring RET [91] or whether instead electrons leak onto oxygen froma semiquinone in the Q-binding site of complex I [92]. In any caseand in agreement with the experimental data [87,88], superoxide gen-erated by complex I should be completely released into the matrix,since the structure of mitochondrial complex I revealed that bothsites reside in the peripheral arm of complex I at some distance fromthe membrane plane [93]. In vitro, complex III produces ROS at theubiquinol oxidation site (Qo site), if the ubiquinone reduction site isblocked by inhibitors like antimycin A [30–32,94,95]. While there isan ongoing controversial discussion aboutmechanistic details (whethersuperoxide is produced in a semi-forward mode from accumulatedsemiquinone or during a semi-reverse mode from the reduced co-factor heme bL [32,33,96]), structural considerations and experimentaldata indicate that superoxide from the Qo site is released primarilyinto the IMS or cristae lumen [69,87,88]. It is worthwhile to mentionthat especially ROS from this site have been linked to cellular redox sig-naling [15,33]. There is no direct experimental evidence pointing to thetopology of ROS release from complex II. Yet, it seems likely that ROSproduced at the flavin site (more precisely from the semi- or fully re-duced FAD) of complex II [27,29] will be also released completely intothematrix [28]. However, mutagenesis studies in lower eukaryotes sug-gest 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 ofROS release by complex II.


2.4. The mitochondrial redox compartments

In summary, it can be concluded that the mitochondrial matrix andthe IMS feature distinct redox environments that are controlled by thedistribution of GSSG/GSH and thiol-containing proteins and that are in-fluenced by the activity of the respective ROS sources (Fig. 1). The gluta-thione pool as the main redox buffer in the IMS relies on the activityof cytosolic enzymes and NADPH for the regeneration of GSH fromGSSG. The main ROS source in the IMS is the Qo-site of complex III, butglycerol-3-phosphate dehydrogenase might also generate ROS in thiscompartment under certain conditions. The glutathione pool in thema-trix is regulated by the activities of local NADP+-reducing enzymes andthematrix NADPH pool. NADPH is also essential for the reduction of thethioredoxin/peroxiredoxin system representing another, powerful ROSscavenging machinery. Main ROS sources in the matrix are complex Iand 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 intra-molecular disulfides, S-gluathionylation, sulfenic acid, nitrosothiols) areamong themost intensively studied oxidative and nitrosative modifica-tions since they canmodulate protein function [18,100]. High dynamics,instability and low abundance hamper direct identification of thiolmodifications by mass spectrometry (MS). A long list of availablethiol-reactive reagents provides a versatile tool box for detection,enrichment and quantification of such modifications by gel-based orgel-free redox proteomics (reviewed in [101]).

In all cases, the redox status of a biological sample needs to be stabi-lized initially to obtain a snapshot of the steady-state oxidation levelsat a given cellular or mitochondrial condition and to avoid artifacts byinadvertent oxidations during sample preparation [102]. This can beaccomplished either by rapid protonation and precipitation of proteinswith trichloroacetic acid [103] or by blocking free thiols with excessof the membrane permeable highly thiol-reactive reagent N-ethylmaleimide (NEM; Fig. 2A) [41,104–107]. In contrast to iodoacetamide(IAM) or iodoacetic acid (IAA) the reaction of NEM is more efficient atphysiological pH enabling NEM-labeling of redox sensitive cysteines intheir native conformation and physiological environment [104–106].As recently described for mammalian complex I, this can be an advan-tage for targeted proteomics of thiol modifications of a specific proteinor protein conformation in combination with functional studies [108].Workflows for a comprehensive analysis of the cellular or mitochon-drial redox proteome include a step completely denaturing the sampleto unmask hidden thiols in native protein complexes. To achieve suffi-cient thiol-labeling under denaturing conditions a slightly alkaline pHis required to obtain the reactive thiolate formof the cysteines. At higherpH IAM and IAA are more thiol specific than NEM that exhibits somereactivity with other amino acid side chains [105]. Reversible thiolmodifications (inter- and intramolecular disulfides, S-gluathionylation,S-nitrosylations, sulfenic acid) can be reduced by dithiothreitol ortris(2-carboxyethyl)phosphine (TCEP) following removal of excessthiol blocking reagent. Labeling with a different reagent then allowsdetection 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 sepa-rated by polyacrylamide gel electrophoresis (PAGE) in one or twodimensions. An example of this approach can be found in Galkin et al.[108], who applied fluorescein-NEM to identify the thiol that is accessi-ble only after active/deactive transition of complex I using a 3D Blue



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, intactcells 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 thiolprobe (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 dithio-threitol or tris(2-carboxyethyl)phosphine (TCEP). Alternatively S-nitrosylations are selectively reduced by ascorbate and copper(II) [116,117]. The secondary thiol labels (blue orbrown★) 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 labeledby 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 quanti-tative 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 andheavy 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 deuteriumNEM (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 peptidesby 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, oxidizedcysteine peptide; m/z, mass-to-charge ratio; IEF, isoelectric focusing.


Native/double SDS separation followed by identification of themodifiedthiol by MS (see Section 5.1).

Classical 2D IEF/SDS gels separate complex mixtures of proteins ac-cording to their isoelectric point in the first dimensional isoelectricfocusing (IEF) and according to their molecular mass in the seconddimension (SDS-PAGE) to provide a detailed 2D-map of the proteomeunder study [109]. If according to the approach described above oxi-dized thiols are reduced followingNEM treatment and are subsequentlylabeled by a fluorescent dye, the laser scannerwill detect only reversiblyoxidized proteins [110]. By introducing redox-DIGE, Hurd et al. [111]adapted the difference gel electrophoresis (2D-DIGE, [112]) approachfor the detection of proteins differentially oxidized by mitochondrialROS [111]. After blocking free thiols by NEM, reversibly oxidizedproteins of control and redox-challenged mitochondria are reduced,then differentially alkylated with Cy3- or Cy5-maleimides and finallyseparated in the same 2D gel to identify ROS specific targets (Fig. 2B).Differentially labeled protein spots are then identified by MS. The useof internal standards like in 2D-DIGE [112] could allow quantitativecomparison of redox-modifications under different conditions of ROSproduction to identify generator-specific protein targets in mitochon-dria. SNO-DIGE is a variant of redox-DIGE specifically designed to quan-tify S-nitrosylations [113]. This method uses ascorbate and copper(II) toallow for mild and specific reduction of SNO-sites that are subsequentlylabeled differentially with Cy3-NEM and Cy5-NEM. The introduction of


internal SNO standards enables quantification and comparison of morethan two conditions within one SNO-DIGE experiment [114]. Theredox- and SNO-DIGE methods give a comprehensive map of differen-tially oxidized and nitrosylated proteins, although MS identification ofcysteines modified with fluorescent dyes is rarely achieved.

3.3. Quantitative redox proteomics by mass spectrometry

In purely liquid-chromatography (LC)–MS based redox proteomicsbetter detectable thiol labels are used. Cysteine containing peptidesalkylated with NEM, IAM or IAA can be identified readily by LC–MSand have been used to distinguish between reduced and oxidized thiolsin a qualitative manner (reviewed in [100]). In order to identify gener-ator specific thiols for redox signaling quantitative redox proteomic ap-proaches using stable isotope labeling are required. Differential cysteinelabeling with stable isotopes produces chemically identical light andheavy peptides that elute at the same retention time in chromato-graphic separations, exhibit the same ionization properties, but canbe discriminated and quantified based on a mass shift of neighboringheavy and light precursor ion peaks. Depending on the experimentaldesign the heavy/light ratios reflect the redox state or the differentialoxidation state under two conditions of a thiol containing peptide.TandemMS/MS is used for identification of the peptides and the modi-fied cysteines. A targeted approach to identify thiol modifications in




complex I followed the principle of differential labeling with light NEM(d0) to block free thiols andmild reduction by ascorbate and copper(II)to label SNO sites with deuterium-NEM (d5) [43,115] (Fig. 2C). Theamount of nitrosothiols in complex I upon treatment with MitoSNO[42] in normoxia and ischemia of intact mouse hearts was estimatedby quantitative MS [43]. Comprehensive redox proteomic approachesuse thiol reactive reagents with biotin affinity tags. In 2001 Jaffreyet al. introduced the biotin-switch assay to assess the S-nitrosylome[116]. This method is based on the conversion of SNO sites to biotinlabeled thiols followed by affinity chromatography and protein identifi-cation [116,117]. The biotin-switch approach was later adapted toenrich other thiol modifications e.g. mixed redox modifications andsulfenic acid [118–120].

A very elegant way to quantify the thiol redox proteome by an MSbased approach uses isotope-coded affinity tag (ICAT) technology. TheICAT reagent features an IAM-moiety for thiol labeling, an isotopic tagwith a mass difference of 9 Da between the light 12C and heavy 13Cform for quantitative MS and a cleavable biotin tag for affinity enrich-ment of cysteine containing peptides [121] (Fig. 2C). The ICAT technol-ogy has been adapted for redox proteomics of complexproteinmixturesemploying at least three strategies. The Cohen group used ICAT reagentsto label free thiols of control and redox-treated samples and comparedboth experimental conditions by quantitativeMS [122,123]. TheOxICATmethod introduced by Leichert et al. [124] monitors the oxidation stateof a thiol by labeling free thiols with light, and oxidized cysteineswith heavy ICAT [124–127]. A third strategy blocks free thiols withNEM and uses light and heavy ICAT to compare two experimental con-ditions [41,128] (Fig. 2B). The latter approach was used in a compre-hensive redox proteomics study to identify and quantify changes inthiol oxidation during ischemia and reperfusion in Langendorff per-fused isolated mouse hearts and many redox modified mitochondrialtargets were identified [41]. All approaches to identify and quantifythiol modifications show advantages and limitations. The advantageof the LC–MS based redox ICAT methods is beyond any doubt efficientenrichment, identification and quantification of reduced and oxidizedcysteine sites in a proteome or sub-proteome. However these methodsdo not allow for parallel estimation of the overall protein abundanceand identification of irreversible thiol oxidations. As much as proteinoxidation, changes in protein abundance due to protein degradationor protein translocation to another cellular compartment could be aconsequence of redox signaling or oxidative stress. The estimation ofthe redox state of a peptide by OxICAT is to some extent independentof protein abundance, but cellular conditions that shift from redoxsignaling to oxidative stress cannot be monitored since over-oxidizedcysteines escape detection. To overcome this limitation the recentlyreported method GELSILOX (GEL-based Stable Isotope Labeling ofOXidized cysteines) combines quantification of redox modificationand protein abundance by using standard thiol labels (NEM, IAM)and 16O/18O isotopic labeling to distinguish between experimentalcondition and protein abundance [129]. By direct LC–MS identificationand quantification of sulfinic or sulfonic acid containing peptides,the GELSILOX approach could be easily expanded to also detect theseirreversible redox modifications.

Many high throughput methods to quantify redox modification arenow available. However the major challenge in redox proteomics re-mains freezing the redox state during the first step of sample prepara-tion to avoid inadvertent oxidations and artificial results. It is difficultin particular to find the right conditions for redox proteomics in cell cul-ture experiments, since cell lines are adapted to atmospheric oxygenpressure, but the physiological processes underlying this adaptationare unknown.

In order to determine the physiological relevance of thiol modifica-tions for protein function, protein complex assembly or conformation,and protein translocation or degradation many additional targetedexperiments are required in-vivo and in-vitro. To this end the recentlydeveloped complexome profiling approach [130] that allows studying


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

4. Targets of mitochondrial ROS

A number of in vitro and in vivo investigations have addressed thequestion which mitochondrial proteins contain redox-sensitive thiols,i.e. are oxidized, glutathionylated or S-nitrosylated upon induction ofmore or less defined oxidative stressors. However, only few studiesused a physiological stimulus or relied at least on endogenous mito-chondrial ROS sources. Often oxidative stress was applied by theaddition of external hydrogen peroxide, which might be generally ap-propriate to identify redox-sensitive mitochondrial proteins, but doesnot consider redox compartmentalization or discriminate between dis-tinct ROS sources (see Section 2). Respiratory chain complexes thathave been identified as targets of (mitochondrial) ROS are discussedin Section 5. In their fundamental investigation using Redox-DIGE(see Section 3) Hurd et al. [111] detected a number of proteins inisolated rat heart mitochondria that were modified by externallyadded H2O2, ROS generated by respiratory chain complexes I and III(induced by RET and antimycin A, respectively) and externally addedRNS (i.e. S-nitroso-N-acetyl-DL-penicillamine, SNAP). In general, onlyfew proteins were modified under all conditions and ROS productionby antimycin or RET led to the oxidation of only three and six proteins,respectively [111]. Among the identified proteins were VDAC1, mito-chondrial creatine kinase and proteins of mitochondrial fatty acidand pyruvate metabolism. Redox-modification of VDAC1 and enzymesinvolved in lipid metabolism and oxidative phosphorylation (seeSection 5) have been identified in a study by Kumar et al. who investi-gated the thiol proteome in isolated mouse hearts following ischemia/reperfusion applied with a Langendorff perfusion system [41]. Inthis study, also two TCA cycle enzymes (malate dehydrogenase andα-ketoglutarate dehydrogenase) came up as ROS-targets. Redox-sensitivity of α-ketoglutarate-dehydrogenase, i.e. the glutathionylationof the lipoic acid of subunit E2, has been detected also in an in vitrostudy using rat heart mitochondria [131–133]. A remarkable findingof the study of Kumar et al. [41] is that the oxidation of protein-thiols in hearts subjected to 20 min global ischemia was maximalafter 5 min of reperfusion and was almost completely reverted after30 min. This indicates that these modifications are largely reversible.Since the observed redox-modifications coincide with the oxidativeburst that occurs upon reperfusion, complex I in RET mode seems tobe the main source for these ROS [4,28].

A completely different approach was followed by Dick and co-workers, who used in situ kinetic trapping to detect proteins that inter-actwithmitochondrial thioredoxin 2 (Trx2) [134]. In HEK293 cells, theyexpressed an inducible mutated form of Trx2 that was still capable ofattacking disulfide bridges of target proteins, but deficient in resolvingthe formed mixed disulfide intermediates. In combination with astreptavidin tag, this allowed to selectively pull out Trx2 target proteinsand identify them subsequently by mass spectrometry. Oxidative stresswas mainly applied by external H2O2 and only one target (methionyl-tRNA synthase) was confirmed after induction of endogenous ROSby the combined application of rotenone and antimycin A [134]. Thisinvestigation suggests that the mitochondrial protein biosynthesismachinery is a major target of ROS. Other targets were chaperones,proteins of the amino acid metabolism and ATP/ADP translocase 2 thathas been implicated also in mitochondrial permeability transition.

ROS are also potent inductors of the enigmatic mitochondrial per-meability transition pore [135,136] that contributes to different formsof apoptotic and necrotic cell death and leads to permeabilization ofthe mitochondrial membranes [137]. Irrespective to the recently pro-posed complex V dimers [138], it is still uncertain, which role other pro-posed mPTP-proteins like cyclophilin D, ATP/ADP translocase, thephosphate carrier and VDAC might play. Oxidation of cysteine has




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 uncouplingproteins [141–143].

5. Thiol-based redox modification of respiratory chain complexes

The reversible redox modification of protein-thiols is an impor-tant response to changes in the cellular redox environment. Sincemitochondria are central to oxidative stress and redox signaling,S-glutathionylation and other thiol-modifications of mitochondrialproteins are of particular interest [8,72,74]. Especially thiol modifica-tions of respiratory chain complexes have been detected in a numberof studies. This initiated a number of in vitro studies and prompted adiscussion on physiological functions and consequences that is stillongoing.

5.1. Complex I and the active/deactive transition

Mitochondrial complex I has been identified in a number of studiesas a target of oxidative thiol-modifications. Oxidation of cysteine-thiols or S-glutathionylation of the iron–sulfur cluster containingsubunits PSST (Ndufs7) and 51-kDa (Ndufv1) and the accessory subunitNUJM (Ndufa11) was observed in Langendorff perfused isolated mousehearts during ischemia/reperfusion [41]. The 51-kDa subunit (Ndufv1)was found modified in the post-ischemic myocardium of rats [40]. Fur-thermore, reversible thiol oxidation of the 75-kDa subunit (Ndufs1) hasbeen demonstrated in a study using kinetic trapping of Trx2-interactingproteins in HEK 293T cells [134]. It should be noted that the 51-kDasubunit carries FMN, the primary superoxide source of complex I, andresides in close proximity to the 75-kDa subunit. On the other hand,cardioprotective S-nitrosylation of complex I subunits by differentNO-generating compounds (GSNO, MitoSNO1) has been detected inmouse and rat heart models [43,144]. The implications of these obser-vations are discussed in detail further below.

The effect of S-glutathionylation, S-nitrosylation and the formationof intramolecular disulfides on complex I activity has been investigatedin a number of in vitro studies using purified complex I or ʻmitochon-drial membranes’ (i.e. submitochondrial particles (SMP)mainly derivedfrom bovine heart mitochondria) [38,40,42,145–150]. A general out-come of these studies is that oxidative thiol-modifications of complexI cysteines result in a general reduction of catalytic activity and thatthe modifications are relatively specific for distinct cysteines located inthe 51 kDa- and 75 kDa-subunits of complex I. Only prolonged exposureof complex I to oxidative stress resulted in unspecific labeling of addi-tional cysteines and eventually in the loss of iron–sulfur clusters.While initial experiments showed that S-glutathionylation results inan increased ROS-generation by complex I [145], further studies couldnot confirm this finding [38,148]. It has been proposed that oxidativemodification of thiol groups in complex I and other proteins, either byS-glutathionylation or S-nitrosylation, is a protectivemechanism to pre-vent higher oxidation of the thiyl-radical or sulfenic acid that is formedduring the primary event [8].

Fig. 3. The active/deactive transition of mitochondrial complex I. If the active (A) form of complexdeactive (D) form, that can be reactivated by adding substrates to induce catalytic turnover. Re(Me2+) or covalent modification (R) of a single cysteine by oxidation, nitrosylation or any kind


Control of the catalytic activity ofmitochondrial complex I by the so-called active/deactive (A/D) transition involves a specific and welldefined cysteine switch mechanism. Kotlyar and Vinogradov were thefirst to observe the A/D transition in submitochondrial particles frombovine heart [151]. Mitochondrial complex I converts into an inactivestate, the ‘deactive’ or D-form, when incubated at 37 °C in the absenceof substrates. This form slowly turns back into the ‘active’ or A-form inthe presence of NADH (or NADPH) and ubiquinone (Fig. 3). Later itwas shown byMaklashina et al. that theA/D transition is a characteristicof complex I from vertebrates [152]. In the lower eukaryotes Yarrowialipolytica and Neurospora crassa A/D transition of complex is also ob-served, 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 preventreactivation of deactive complex I. Remarkably, the ion binding siteand the cysteine are only accessible in the D-form of the enzymerendering the A-form resistant to this type of inhibition (Fig. 3). Thisindicates that the A/D transition goes along with a significant confor-mational change.

Anoxia/reperfusion studies by Maklashina et al. using Langendorffhearts gave the first indications that the A/D transition is more thanan in vitro phenomenon [155]. Galkin and Moncada showed inductionof the D-form of complex I during prolonged hypoxia and suggestedthat this could make the cysteine exposed only in this state accessiblefor nitrosation. Since under these conditions simultaneous generationof ROS by the respiratory chain complexes and of NO by NO-synthasescould lead to the formation of significant amounts of the endogenousnitrosating agent peroxynitrite, this could result in permanent deacti-vation of the enzyme [156,157]. Murphy and coworkers recentlyshowed marked cardioprotective effects in mice associated with theS-nitrosation of the cysteine switch of the A/D transition [42]. Takentogether these results demonstrate that the A/D transition and the as-sociated cysteine switch are operational in vivo. However, the physio-logical role of this regulatory mechanism remains obscure and it willbe important to understand the molecular mechanism and control ofthe A/D transition in detail.

The single cysteine that is only exposed in the deactive form of com-plex I was identified by site specific labeling and mass spectrometryas described in Section 3.2 [108]. The highly conserved residue wasfound in the hydrophilic loop connecting the first two transmembranehelices of subunit ND3 of complex I, one of the central subunits of com-plex I encoded bymitochondrial DNA. This subunit is part of the proximaldomain of themembrane arm of complex I [158]. In the loop carrying thecysteine controlling the A/D transition, pathogenic mutations in humanscausing mitochondrial disorders were reported at three positions(Fig. 4). The X-ray structure of complex I [158] reveals that loop 1 of sub-unit ND3 resides next to aβ-sheet of the 49-kDa subunit of the peripheralarm (Fig. 4). It was shown by site-directed mutagenesis in the yeast ge-netic model Y. lipolytica [159] that this β-sheet is part of the ubiquinoneand inhibitor binding pocket of complex I. Twohistidines essential for cat-alytic activity are located within the loop connecting the first and secondstrands of the β-sheet [160,161]. Therefore, they seem to be part of theentry path for the ubiquinone head group into the catalytic site. Thisputs the A/D cysteine switch of complex I into an ideal position to controlactivity (Fig. 4). The analysis of conformation specific crosslinks suggests

I is left idle in the absence of substrate, it converts into the apparently inactive state, theactivation of the D-form to the A-form can be prevented by the binding of divalent cationsof SH-reagent like N-ethylmaleimide of iodoacetamide.



Fig. 4. The loop of the ND3 subunit involved in the A/D transition of complex I. Structuralmodel of the loop connecting transmembrane helices 1 and 2 of the mitochondriallyencoded subunit ND3 and surrounding central subunits. The model was constructedusing the Pymol software package from the structural coordinates of bacterial complex Ifrom Thermus thermophilus (PDB ID: 4HEA) [158]. The highlighted residues were changedto the human variant where necessary using the Pymol mutagenesis wizard and thenumbering of the human proteins is indicated. The ND3 subunit is shown in yellow andthe 49-kDa subunit in blue. The cysteine involved in the A/D transition (magenta), twoconserved acid residues in its vicinity (yellow) and three residues changed in mito-chondrial disease (green) are shown in stick representation. The N-terminal-sheet ofthe 49-kDa subunit is highlighted in dark blue. The terminal iron–sulfur cluster N2 isdepicted in space-fill representation.


that also the accessory 39-kDa subunit takes part in the structural chang-es associated with the A/D transition of mitochondrial complex I [162].The 39-kDa subunit is of particular interest since it is homologous toshort chain dehydrogenases and contains a NADPH binding site of un-known function [163–165].

Since the ROS production by mitochondrial complex I is directlylinked to and controlled by its catalytic activity, studying the controlof the A/D transition by the cysteine switch in subunit ND3 in detailwill be prerequisite to investigate and understand its physiologicalrole in the context of the redox signaling networks of eukaryoticcells.

5.2. Complex II

Also the FAD-containing 70-kDa subunit (SdhA) of complex II hasbeen identified as a target of redox modification following ischemia/reperfusion in mouse heart [41]. Since mechanistic studies haveshown that ROS are generated at the flavin site of complex II [27,29]when the downstream respiratory chain is blocked, cysteines thatare in close proximity to FAD located in the 70-kDa subunit are likelythe primary targets of these ROS. In studies with isolated complex IIfrom bovine heart mitochondria and an in vivo rat heart ischemia/reperfusion model it was observed that de-glutathionylation of the70-kDa subunit (SdhA) of complex II results in a decrease of its cata-lytic activity [36]. S-glutathionylation of the 70-kDa subunit (SdhA) inpurified complex II enhanced electron transfer activity and decreasedthe production of superoxide [36]. Superoxide production by complexII also induces self-inactivation by the formation of a CII-derivedthiyl-radical. This implies that S-glutathionylation of complex IIunder ischemic conditions may have a protective effect by preservingelectron transfer activity and preventing a vicious circle of ROS-induced self-inactivation [36]. In follow-up studies, it was shown that


the S-glutathionylation protects from oxidative damage by tyrosinenitration [166,167].

5.3. Complexes III, IV and V

Redox-modification of respiratory chain complexes III, IV and Vhas been also observed in some in vitro studies. However, systematic in-vestigations on functional consequences are largely missing. A redoxmodificationwithin the core I subunit of complex III has been identifiedin isolated rat heart mitochondria after ROS induction at the Qo site byantimycin A [111] and S-nitrosylation of this subunit was induced byMitoSNO [113]. Cysteine thiols of Core II were found to be oxidizedafter ischemia/reperfusion in Langendorff-perfused mouse hearts [41].Subunit Va of complex IV (cytochrome c oxidase) was glutathionylatedin response to diamide treatment of human T-cells [168] and subunit Vbwas glutathionylated in rat hepatocytes exposed to the redox cyclermenadione [169]. Glutathionylation of the α1-subunit of complex Vwas 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 mousehearts [144] and has been identified as a thioredoxin-2 target in H2O2

treated HEK293 cells [134].

5.4. Concluding remarks

Recent years have brought a much better understanding of themechanisms by which the respiratory chain complexes may generateROS and how the antioxidative defense systems can modulate theredox environment in the different mitochondrial compartments.However, although numerous oxidative modifications of mitochondrialproteins have been reported, so far the evidence supporting a physio-logical relevance of suchmodifications is scarce. Indeed, even thewidelyaccepted notion that the respiratory chain is a major source for mito-chondrial ROS in vivo is still being challenged [170]. The application ofthe redox-proteomic approaches to analyze oxidative modifications atthe level of individual proteins as summarized above will be requiredto elucidate the importance of mitochondrial ROS production and itsconsequences in living cells. Control of the A/D transition by specificmodification of a single cysteine is a first example of how such studiescan shed light on mechanisms of redox regulation by oxidative proteinmodification.

Acknowledgements

The author's work was supported by the DeutscheForschungsgemeinschaft (SFB815 “Redox Regulation: Generator systemsand functional consequences”, projects A02 and Z01). The authorsthank Erik Bonke for drawing Fig. 1.

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Mitochondrial respiratory chain complexes as sources and targets of thiol-based redox-regulation

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