Oxidation of NADH and ROS production by respiratory complex I

BBABIO-47555; No. of pages: 9; 4C: 5 Biochimica et Biophysica Acta xxx (2015) xxx–xxx

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Oxidation of NADH and ROS production by respiratory complex I☆ Andrei D. Vinogradov ⁎, Vera G. Grivennikova Department of Biochemistry, School of Biology, Moscow State University, Moscow 119991

a r t i c l e

i n f o

Article history: Received 29 September 2015 Received in revised form 2 November 2015 Accepted 7 November 2015 Available online xxxx Keywords: NADH:quinone oxidoreductase Proton pumping Hydrogen peroxide Superoxide Bacterial plasma membranes Mitochondria

a b s t r a c t Kinetic characteristics of the proton-pumping NADH:quinone reductases (respiratory complexes I) are reviewed. Unsolved problems of the redox-linked proton translocation activities are outlined. The parameters of complex I-mediated superoxide/hydrogen peroxide generation are summarized, and the physiological significance of mitochondrial ROS production is discussed. This article is part of a Special Issue entitled Respiratory complex I, edited by Volkerzickermann and Ulrich Brandt. © 2015 Elsevier B.V. All rights reserved.

1. Introduction Respiratory complex I (proton-translocating NADH:quinone oxidoreductases, mitochondrial complex I, bacterial NDH-1)1 catalyzes a reaction of great metabolic significance: ð1Þ where Q stands for the natural membrane-bound quinone (ubiquinone with various number of isoprenoid units at position 6 of benzoquinone + ring) or menaquinone, H+ in and Hout stand for intramitochondrial (cytoplasmic) and extruded protons, respectively, and p.m.f. (protonmotive force) is the transmembrane gradient of electrochemical activity of H+ (Δμ~ Hþ ). Reaction (1) is only reversible when the oxidoreduction is tightly coupled to proton pumping activity. Thus, this reversible reaction only proceeds when the enzyme is bound to a membrane that is poorly permeable to protons. The enzyme detached from the membrane or

bound to a membrane freely permeable for protons catalyze reaction (1) irreversibly due to a significant thermodynamic gap between the standard redox potentials of NADH/NAD+ (− 320 mV) and QH2/Q (+60 mV). The membrane-bound and all solubilized or detergent-dispersed preparations of the enzyme catalyze the irreversible reaction: NADH þ 2Aox →NADþ þ 2Ared þ 2Hþ

ð2Þ

where Aox and Ared stand for artificial oxidized and reduced forms of the A red /A ox couple. Ferricyanide 3 − anion (Ferri) and/or hexaammineruthenium3 + cation (HAR) are routinely used in assays of the enzyme. Reaction (2) is not coupled to proton translocating activity. Membrane-bound and various solubilized preparations of complex I as well as their FMN-containing fragments catalyze the reversible NADH:APAD+ transhydrogenase reaction. Mitochondrial and bacterial complex I also catalyzes NADH oxidation by oxygen: NADH þ О2 þ Нþ →NADþ þ Н2 О2

ð3Þ

and/or

+

Abbreviations: APAD , 3-acetylpyridine adenine dinucleotide; Ferri, ferricyanide; FMN, flavin mononucleotide; HAR, hexaammineruthenium(III); Qn, 2,3-dimethoxy,5methyl,6-(n)isoprenyl,1,4-benzoquinone; ROS, reactive oxygen species; SMP, submitochondrial particles; O•2 , superoxide anion; SOD, superoxide dismutase; p.m.f., protonmotive force (transmembrane gradient of electrochemical activity of H+, Δμ~ Ηþ ). ☆ This article is part of a Special Issue entitled Respiratory complex I, edited by Volkerzickermann and Ulrich Brandt. ⁎ Corresponding author. E-mail address: [email protected] (A.D. Vinogradov). 1 Bacterial plasma membrane proton pumping NADH:quinone oxidoreductases (NDH-1) and corresponding mitochondrial enzymes are designated as complex I throughout the text for the sake of simplicity.

NADH þ 2О2 →NADþ þ 2O•2 þ Hþ

ð4Þ

the formation of so-called ROS (reactive oxygen species – hydrogen peroxide and superoxide) at the highest rate of about 0.2–0.3% of those depicted by Eqs. (1) and (2). Spectacular progress in visualization of atomic structures of the fragments [1,2] and the entire complex I [3–5] isolated from various sources have been achieved during the last decade. However, as important as the information provided by the molecular architecture of the enzyme

http://dx.doi.org/10.1016/j.bbabio.2015.11.004 0005-2728/© 2015 Elsevier B.V. All rights reserved.

Please cite this article as: A.D. Vinogradov, V.G. Grivennikova, Oxidation of NADH and ROS production by respiratory complex I, Biochim. Biophys. Acta (2015), http://dx.doi.org/10.1016/j.bbabio.2015.11.004

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A.D. Vinogradov, V.G. Grivennikova / Biochimica et Biophysica Acta xxx (2015) xxx–xxx

is, it serves only as excellent groundwork for hypotheses on possible catalytic mechanisms. We believe that studies on the catalytic activities still are and will be the powerful tool for understanding the mechanistic and, perhaps more physiologically important, regulatory properties of complex I. This short review discusses available information on some features of eukaryotic and bacterial complex I as they appear from studies of the reactions (1 and 2). We will also discuss reactions (3 and 4) and their possible physiological significance. Because little information is available on the catalytic properties of plant complex I, they are not discussed here; an interested reader is addressed to Refs. [6,7]. 2. Intramolecular electron transfer Complex I contains up to ten active redox components (FMN, 7–8 iron-sulfur clusters, and bound quinone(s)). The structures of complex I and fragments derived therefrom [1–4] unambiguously suggest the following sequence of intramolecular electron transfer on the way from NADH to the final quinone acceptor: NADH→FMN→N3ðN1aÞ→N1b→N4→N5→N6a→N6b→N2→Q

ð5Þ

During steady-state coupled or uncoupled NADH oxidation by bovine heart submitochondrial particles (SMP), all EPR-detectable ironsulfur centers of complex I are almost completely reduced [8,9]. Thus, the rate-limiting step of the overall NADH oxidase is reoxidation of the terminal electron-transferring component (N2, or specifically and tightly bound quinone) by bulk quinone. Only a very short description of the intramolecular electron transfer is needed for further discussion of the steady-state catalytic activity. Recently, the kinetics of intramolecular electron transfer in purified Escherichia coli complex I using ultrafast freeze-quenching techniques followed by EPR and optical spectroscopic analysis have been reported [10–12]. The results obtained by these groups and their interpretation significantly differ. Biphasic reduction of iron-sulfur clusters N1a and N2 (~ 50 μsec) followed by much slower reduction of N1b and N6b was interpreted by Verkhovskaya et al. so as to suggest that NAD+ dissociation from the nucleotide-binding site is the rate-limiting step in the overall reaction (rapid oxidation of the first NADH molecule results in reduction of N1a and N2, and further reduction of the enzyme proceeds significantly more slowly because the next NADH molecule can bind to the empty active site only after dissociation of NAD+) [10,11]. De Vries et al. interpreted their result as to suggest that rapid reduction of N2 results in six-fold decrease in electron tunneling rate between other iron-sulfur centers, thus synchronizing electron transfer with the proton pumping activity [12]. Whatever interpretation is correct, the enzyme turnover when it operates as a member of the respiratory chain (~hundreds per sec) is substantially slower than any intramolecular redox reaction. 3. Steady-state activity The accumulated data on NADH-dehydrogenating activities of membrane-bound complex I scattered in the literature are given in Table 1. Averaged NADH oxidase activity of fully activated2 uncoupled bovine heart SMP is about 1.5 μmol of NADH oxidized per min per mg of protein (Table 1), a value that is close to that seen at “saturating” concentrations of artificial quinone acceptors and other analogs or homologs of ubiquinone (Q1). Assuming a molar content of complex I in heart mitochondrial membranes of about 0.1 nmol per mg of protein [37], these numbers correspond to maximal steady-state enzyme turnover of ~ 250 s−1. Does this turnover reflect the maximal capacity 2 The mitochondrial enzyme exhibits complex kinetics in the quinone reductase activity due to slow transition between catalytically inert D- and active A-forms (operationally called the A/D-transition). This phenomenon reviewed in Refs [35,36] is beyond the scope of the present discussion.

of the enzyme, and how it can be regulated in situ? Natural ubiquinones are practically insoluble in aqueous media, and their analogs or homologs at “saturating” concentrations of about 100 μM are routinely used for assays of the membrane-bound or detergent-solubilized enzyme. The content of ubiquinone in heart inner mitochondrial membrane is about 5 nmol per mg of protein [37]. The bulk quinone is located in the lipid phase of the membrane. The content of phospholipids in the inner mitochondrial membrane (bovine heart) is about 0.4 mg per mg of protein [37]. Simple approximate calculations show that the concentration of ubiquinone in the lipid phase is as high as about 10 mM. Thus, in situ quinone reduction by complex I proceeds at concentrations of the substrate of about two orders of magnitude higher than that used under standard assay systems with added artificial quinone acceptors. The true concentration of “water-soluble” quinones in the membranes and their respective reactivity certainly depend on the lipid/water partition coefficient [38]. The sequence of events during the steady-state NADH:externally added quinone-acceptor reductase reaction catalyzed by particulate complex I is not clear. External quinone might either directly bind to the ubiquinone-specific site or oxidize tightly bound Q, or oxidize bulk natural reduced quinone swimming in the lipid phase. In studies on Q-pool behavior in the respiratory chain [39] recently confirmed by others [40], linear dependence of NADH oxidase activity of bovine heart SMP on the content (concentration) of oxidized ubiquinone have been demonstrated. If extrapolated to the state where ubiquinone is fully oxidized, the data would correspond to turnover number of complex I of about 500 s−1 (pH 7.4, 25 °C), a value that is substantially higher than those (averaged) measured at saturating concentrations of water-soluble quinones or in steady-state NADH oxidase assays (~250 s−1, see above). This is not surprising because during the steady state, fully uncoupled NADH oxidase operates at Qred/Qox ratio of about 1. Linear, not hyperbolic, dependence of the NADH oxidase rate on molar fraction of oxidized ubiquinone (Qox/(Qox + Qred)) has been demonstrated [39]. Such dependence is expected either if the total concentration of quinone is much lower than the apparent Km for Qox or if Qred competes with Qox for binding at the reactive site with similar affinity. The first possibility seems to be unlikely because of high concentration of ubiquinone in the membrane (see above). Kinetic competition between oxidized and reduced forms of ubiquinone (with similar affinity) seems the more plausible explanation and corroborates with well-established reversibility of reaction (1). Weak (competitive with oxidized Q1) inhibition of NADH:Q1 reductase activity catalyzed by highly purified bovine heart complex I by the product (Q1H2) have been reported [41]. Also, a competitive relation between oxidized and reduced quinone acceptor (Q2) was documented for respiratory complex II [42]. The linear dependence of NADH oxidase activity on the molar fraction of oxidized ubiquinone might shed some light on the mechanism of the respiratory control phenomenon. NADH oxidation in controlled state 4 can equally be slow either because p.m.f. inhibits proton translocating activity (see Eq. (1)) or because an increase in reduced ubiquinone concentration due to a decrease in Q-pool oxidation by further components of the respiratory chain (complex III and cytochrome c reductase), which are under the control of p.m.f. The relative contributions of “thermodynamic” (p.m.f.) and “kinetic” (Qred/Qox ratio) factors to the respiratory control phenomenon at the level of complex I remain to be established. 4. Artificial electron acceptors Ferricyanide (Ferri) [43,44] and hexaammineruthenium III (HAR) [21] are the most frequently used artificial electron acceptors for quantitation of complex I catalytic activity of preparations of different degree of resolution. The substrate (NADH) concentration–activity profile for Ferri reductase catalyzed by the membrane-bound or soluble, socalled high molecular mass preparations of complex I appears as a bell-shaped curve with a relatively sharp maximum. For any

Please cite this article as: A.D. Vinogradov, V.G. Grivennikova, Oxidation of NADH and ROS production by respiratory complex I, Biochim. Biophys. Acta (2015), http://dx.doi.org/10.1016/j.bbabio.2015.11.004

A.D. Vinogradov, V.G. Grivennikova / Biochimica et Biophysica Acta xxx (2015) xxx–xxx

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Table 1 Specific NADH-oxidizing activities of mitochondrial or plasma membrane respiratory complex I (representative values) a Activity (μmol/min per mg)

Eukaryotes Bos taurus Y. lipolytica N. crassa Prokaryotes E. coli P. denitrificans R. capsulatus

NADH oxidase

NADH:quinone reductase

NADH:ferricyanide reductase (Ferri 0.5 mM)

NADH:HAR reductase (HAR 2 mM)

NADH:APAD+ reductase

0.7–2.0 [13–17]

0.3–1.6 [13,18,19]

2.0–8.4 [20,21]

0.8 [24]

0.2 [25] 0.2–0.3 [27]

0.4 [25] 0.1–0.4 [27,28]

18.0b [22] 2.7–8.0 [20,21,23] 0.9–1.2 [25,26] 2.5 [28]

0.4 [29] 0.8 [14,15,33,34]

0.4–3.1 [29,30] 0.8 [34]

1.4–4.6 [29–31] 0.7 [20] 1.6–2.1c [33]

2.2d [32] 2.0 [20]

0.5 [34]

0.5 [34]

a

Specific activities reported in the literature even for the same type of preparations are inherently variable due to somewhat different assay conditions. This is particularly true for data where artificial electron acceptors were used. To the best of our knowledge, no study was reported where oxidase and acceptor reductase activities extrapolated to infinity concentrations of the latter have been compared. b Extrapolated to infinity HAR concentration. c 1 mM Ferri in the reaction mixture. d 0.35 mM HAR in the reaction mixture.

oxidoreductase reaction, the enzyme turnover number or Vmax is meaningful only if the activity determined at various concentration of an electron acceptor is extrapolated to its infinite concentration. Because of the evidently non-hyperbolic NADH concentration–activity profile, such extrapolation is somewhat ambiguous, and great precautions should be taken if complex I catalytic activity is to be quantitated using Ferri as the electron acceptor. The simplest mechanistic explanation for the bell-shaped dependence is that the reaction proceeds by a ping-pong mechanism, and NADH (at high concentrations) binds to the active site of the reduced enzyme, thus sterically protecting accessibility of the Ferri anion to FMN buried in a deep cleft. This interpretation hardly agrees with the kinetics of NADH-HAR reductase: it proceeds according to a ternary complex mechanism and shows no inhibition at high NADH concentration [20,21,45]. Thus, the kinetic behavior makes HAR a convenient acceptor for studies directed to quantitation of the enzyme turnover or its content (that is assumed to be proportional to Vmax) in various membrane or solubilized preparations. The specific NADH dehydrogenating activity of membrane-bound complex I in bovine heart SMP as extrapolated to Vmax is at least by an order of magnitude higher than their NADH oxidase corresponding to turnover of about 2500 s−1. This number determines the lower limit for the rate of product release from the enzyme active site: NAD+ dissociation is an inevitable step independent of the particular site where any electron acceptor interacts with the enzyme. Thus, at least for the mammalian enzyme, the half time of NAD+ release should be less than 0.3 msec. This value is substantially lower than that for the slow phase of N1b and N6b reduction (about 2 msec) in purified E. coli complex I interpreted as a characteristic time for NAD+ dissociation [10,11]. The NADH:HAR reductase activity of SMP and purified bovine heart complex I at low concentrations of the acceptor is strongly (up to 10-fold) stimulated by ATP and other purine tri- and diphosphonucleotides [45,46]. This effect was interpreted as evidence for an allosteric nucleotide-binding site in the mammalian enzyme [46] or, alternatively, as a change in the reaction pathways of the reaction [45] in the presence of ATP, ADP, and ADP-ribose, competitive inhibitors of the NADH/NAD+-binding site [47]. Kinetically, ATP acts as a competitive (with NADH) activator, thus decreasing the apparent Km for NADH [48]. Grivennikova et al. suggest that ATP bound at a putative allosteric site increases the midpoint potential of an enzyme component that donates electrons to HAR [46]. Remarkably, no activating effect of ATP was observed for prokaryotic Paracoccus denitrificans membranes, whereas it was seen for eukaryotic Yarrowia lipolytica SMP. This species-dependent activation by ATP hardly corroborates with reaction pathway change induced by nucleotides, the mechanism

proposed by Birrell et al. [45]. Whether an allosteric ATP-binding site presumably located at some accessory subunit [49] of eukaryotic complexes has any physiological significance is an open question. Only the HAR reductase activity of eukaryotic complex I is affected by ATP. Recently, a striking observation relevant to HAR reductase activity was reported by Varghese et al. [50]. A point mutation (A341V) in the Y. lipolytica FMN-containing subunit eliminated NADH:HAR reductase activity, leaving all other activities including paraquat reductase unaffected [50]. This observation suggests a different reaction mechanism for paraquat and HAR reduction and demonstrates that our current knowledge on the reduction of the artificial electron acceptors is far from complete. 5. Proton pumping activity At present, the molecular mechanism of the redox-linked proton translocation at coupling Site 1 remains a black box. Numerous earlier Mitchellian direct transmembrane redox loop models should be discarded in light of the structural arrangement of multiple redox components within the enzyme structure. The ubiquinone reactive site (iron-sulfur cluster N2) is located approximately 30 Å from the coupling membrane plane [51,52]. The hydrophobic domain is composed of a number of subunits including three transmembrane homologs of bacterial Na+/H+ antiporters [53] (subunits L, M, and N according to E. coli nomenclature) [2]. A simple and attractive possibility is that these subunits operate as the proton-conductive channels required for the pumping mechanism. For any model of coupling, the stoichiometric coefficient, n in Eq. (1), should be defined. A consensus has been reached that 4 protons are translocated per molecule of NADH oxidized and ubiquinone reduced (2ē) [54,55,and references cited therein]. Here we will briefly discuss several points that might cast a new perspective on studies of the coupling mechanism. (i) The basic structure of Thermus thermophilus complex I is remarkably similar to that of the E. coli and eukaryotic enzymes. Menaquinone (E7m ~ −70 mV) serves as the electron acceptor in T. thermophilus membranes [56], whereas ubiquinones (E7m ~ +60 mV) are the acceptors in eukaryotic membranes. The thermodynamic restrictions hardly permit the T. thermophilus enzyme and ubiquinone-specific complexes I to operate by the same proton pumping mechanism. To the best of our knowledge, the value of n in Eq. (1) as applied to T. thermophilus is not known. In fact, among prokaryotes the n value was experimentally evaluated only for E. coli [57] and P. denitrificans [58,59]. Presumably, the same number of proton-conducting subunits is present in

Please cite this article as: A.D. Vinogradov, V.G. Grivennikova, Oxidation of NADH and ROS production by respiratory complex I, Biochim. Biophys. Acta (2015), http://dx.doi.org/10.1016/j.bbabio.2015.11.004

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rotenone (and other multiple rotenone-like inhibitors) or by transformation of the enzyme to its inactive D-form is not known. It could well be the initial one-electron reduction of the specifically tightly bound Q, or accepting the second electron to form reduced bound quinol, or dismutation of two semiquinones to form bound QH2, or electron exchange between bound QH2 and Q arriving from the bulk quinone pool. The key question of what the particular step(s) is(are) coupled with proton pumping remains to be answered.

prokaryotic and eukaryotic enzymes. The possibility cannot be excluded that variable stoichiometry exists in the reaction (Eq. 1), being controlled by some special regulatory mechanism that turns “on” one, two, or three proton-translocating subunits depending on particular physiological conditions (steady-state p.m.f.). It does not seem unlikely that if a channel-forming subunit is specialized for redox-linked proton translocation its functional state (conformation) would be voltage-dependent, as it is well established for a number of cation-specific transporters [60–62]. The structures of putative H+-translocating subunits N, L, and M are different, thus their “voltage sensitivity” (if it exists) might also be different, resulting in variable stoichiometry of the overall reaction (1) at different p.m.f. values. It should be emphasized that an attractive possibility of the direct participation of N, L, and M subunits in redox-linked energy coupling remains just a proposal having no strong experimental evidence, although “partial” uncoupling has been observed for Yarrowia complex I: variants in Yarrowia complex I deleted by two of the three subunits that are assumed to be candidates for proton pumping translocate protons with half of the stoichiometry observed for the intact parental enzyme [63]. (ii) Counter cation translocation is required to measure proton movement across the coupling membrane as a pH-change because the electrical component of the p.m.f. should be dissipated. Unexpectedly, valinomycin did not affect the initial rate or the maximal pH change if the proton translocation was initiated by NADH-external quinone oxidoreduction in coupled SMP [54]. Strong stimulation of proton translocation by valinomycin, as expected, was seen under the same experimental conditions if the overall NADH oxidase (all three coupling sites) was activated. Formally, these data can be interpreted as to suggest that, in contrast to complex III and cytochrome oxidase, complex I operates solely as the ΔpH generator. Work aimed to confirm or discard such a possibility is in progress in our laboratory. (iii) The NADH oxidase activity of bovine heart SMP is completely (more than 90%) inhibited by specific N2-ubiquinone junction site-directed inhibitors – rotenone and piericidin. When complex I activity is assayed with externally added ubiquinone homologs, a substantial rotenone-insensitive reaction is observed. The trivial explanation of this phenomenon is that water-soluble quinones accept electrons from some other than the natural ubiquinone reactive site, just as other artificial acceptors do (Ferri, HAR). However, studies on the proton pumping activity of bovine heart SMP do not agree with this simple interpretation [54]. Rotenone-inhibited particles catalyze the NADH:Q1 reductase reaction coupled with proton translocation with the same stoichiometry of 4 H+/2ē [54]. Moreover, the irreversibly stabilized D-form of complex I, which is phenomenologically equivalent to the rotenone-inhibited enzyme, also shows proton translocating activity with the same stoichiometry [64]. These unexpected observations are certainly relevant to the mechanism of redox coupled proton translocation. The particular step of the electron transfer from N2 to bulk ubiquinone blocked by

6. Purified complex I Only limited data on catalytic activities of purified complex I are available, although several properties of the enzymes isolated from mammalian [65–67] and yeast [26,68] mitochondria, or plasma membranes of E. coli [30–32], P. denitrificans [33], and thermophilic bacteria (T. thermophilus [69], Aquifex aeolicus [70], Rhodothermus marinus [71]) have been described. A requirement for added phospholipids for the rotenone-sensitive activity of purified bovine heart complex I was demonstrated many years ago [72]. More recently, the same phenomenon was demonstrated for the detergent solubilized purified enzymes from Bos taurus [66], Y. lipolytica [73], and E. coli [32]. The turnover numbers of purified complex I are significantly lower than for complex I-catalyzed NADH oxidase activity of the parent membranes. This is explained either by an inadequate assay using externally added “water-soluble” quinones (see above) or by the presence of some, yet unidentified specific factor(s) required for full catalytic activity in the natural membranes. 7. Complex I-mediated ROS production Membrane-bound or purified complex I from all species studied so far catalyze reactions (3) and/or (4). Quantitative and even qualitative appraisals of the contribution of complex I to overall ROS production by intact mitochondria are extremely difficult due to the presence of multiple intramitochondrial enzymes involved in the formation (for example, dihydrolipoyl dehydrogenase) and utilization of superoxide (superoxide dismutases) and hydrogen peroxide (thioredoxins, glutathione peroxidase, catalase). Here we discuss only the data obtained for systems where ROS production can be unambiguously attributed to the membrane-bound or purified complex I. NADH or succinate oxidation by tightly coupled bovine heart SMP is accompanied by superoxide production at the rate of about 1 nmol/min per mg of protein corresponding to about 0.2–0.3% of the total oxygen consumption. The contribution of particular respiratory complexes to the total superoxide production can be quantitatively evaluated as shown in Table 2. The highest production, which is inhibited by rotenone and uncouplers, is seen during coupled succinate oxidation, thus suggesting that about 70% of one-electron oxygen reduction proceeds via energy-dependent reverse electron transfer (reversal of reaction (1)). The residual rotenone-insensitive generation corresponds to the combined activities of complexes II and III. It was somehow surprising that the rate of generation during more rapid coupled NADH oxidation is about three-fold

Table 2 Oxidase activities and superoxide generation by bovine heart coupled SMP (pH 8.0, 30 °C)a Substrate-donor

1. Succinate (10 mM) + rotenone 2. Succinate (10 mM) + NADH (1 mM) 3. NADH (1 mM) 4. NADH (50 μM) + rotenone a

Oxidase activity, μmol/min per mg

Generation of O•2 , nmol/min per mg

– uncoupler (state 4)

+ uncoupler (state 3)

– uncoupler (state 4)

+ uncoupler (state 3)

0.3 0.3 0.7 0.6 0.6 b0.002

0.9 – – 1.9 1.8 b0.002

1.0 0.3 0.3 0.4 1.0 1.4

0.1 – – 0.1 0.9 –

Adapted from Ref. [17,74].

Please cite this article as: A.D. Vinogradov, V.G. Grivennikova, Oxidation of NADH and ROS production by respiratory complex I, Biochim. Biophys. Acta (2015), http://dx.doi.org/10.1016/j.bbabio.2015.11.004

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lower than the succinate-supported reaction. Moreover, the addition of NADH (1 mM) to a sample where complex I produces superoxide via succinate-supported reverse electron transfer inhibits the reaction. The superoxide production increases and reaches the level of the succinate-supported rate at low concentration of NADH. The dependences of superoxide production and ferricyanide reductase activity of complex I on NADH concentration are almost the same: both appear as bell-shaped curves with a maximum at about 50 μM [17,75]. The simplest interpretation of bell-shaped dependence is that oxygen reduction proceeds via a ping-pong mechanism, and the accessibility of a one-electron donating site for oxygen is restricted in the reduced enzyme–NADH complexes. Other possible interpretations, such as the presence of two NADH/NAD+-binding sites in mammalian complex I, have been discussed [17]. Superoxide production at low (optimal) concentration of NADH is activated by rotenone. At very low NADH concentrations, purified bovine heart complex I produces ROS mostly (90%) as superoxide [18], whereas purified E. coli enzyme generates ROS as hydrogen peroxide [76]. Recently, membrane-bound complex I (SMP) as well as the purified enzyme were shown to generate both hydrogen peroxide and superoxide [75,77]. The partitioning between the products depends on NADH concentration as shown in Fig. 1. The superoxide production reaches a maximum at 10–50 μM NADH and gradually decreases in the millimolar range of the substrate [17,75]. The apparent KNADH as determined from the linear double-reciprocal m plot for the ascending part of the superoxide production titration curve is as low as about 0.5 μM [75]. As noted above (see 4. Artificial electron acceptors section), the apparent Km value for the substratedonor in the reaction catalyzed by any oxidoreductase depends on the redox potential gap between the primary electron acceptor (FMN for complex I) and the component that reacts with an acceptor. The very low KNADH for superoxide production [18,75] indicates that a compom nent immediately reacting with oxygen has a substantially higher redox potential than FMN. The iron-sulfur cluster N2 might serve as a one-electron donor for oxygen reduction as it has been proposed by Genova et al. [78]. This cluster is located close to the ubiquinonebinding site in a funnel-like area at the distance of 25–30 Å from the membrane plane [51,52]. All other iron-sulfur centers are well insulated by the protein environment. A direct approach to exclude cluster N2 as the site of one-electron oxygen reduction was used by Galkin and Brandt [25]. They found that a variant form of Y. lipolytica isolated complex I (R141M) that showed no EPR-detectable center N2 produced superoxide with the same rate (96%) as did the enzyme from the wild strain. They normalized their data to NADH:HAR reductase activities

5

of the wild and mutated strains. Normalization of their data to the “natural” activities shows 60% quinone reductase. This agrees with data reported previously by the same group [79] and originally interpreted as to suggest that the mutants lacking N2 were still capable of ubiquinone reduction at near normal rates, a possibility that seems hardly probable. Rather, the R141M mutation changes the spectral and redox properties of N2, as later demonstrated by Zwicker et al. for the H226M mutant, showing 80 mV negative shift of N2 midpoint potential, absence of its pH-dependence, and still being capable of redox-linked proton translocation with unaltered stoichiometry [80]. We believe that exclusion of iron-sulfur cluster N2 as a possible site of superoxide production should wait for more experimental verification. Hydrogen peroxide formation depends on NADH concentration quite differently. At low substrate concentrations (up to 3 μM), no production is seen (Fig. 1), and the rate reaches a constant value at higher (up to millimolar) range of NADH concentration, where its relative contribution to the overall ROS production is about 60% [75]. Complex I-catalyzed ROS production is inhibited by μmolar NAD+ concentrations [17]. Because of the very low activity of complex I in ROS generation as compared with major oxidase or NADH:artificial acceptor reductase reactions, it is safe to assume that all redox components of the enzyme are in equilibrium with the NAD+/NADH couple during the steady-state reaction (the contribution of the kinetic term to the apparent Km is negligible). Thus, the dependence of ROS production on NAD+/NADH ratio is indicative of the midpoint redox potential of the component(s) reacting with oxygen. The NAD+/NADH ratios reported in the literature for half-maximal ROS production by complex I are greatly variable (from 0.01 up to 7.0) [8,18,81–83]. Titration of complex I by the NAD+/NADH couple is not a true redox titration because the relative binding affinities of the reduced and oxidized enzyme to NAD+ and NADH [84] can significantly contribute to the apparent (NAD+/NADH)0,5 ratio required for ROS production. This ratio is dependent on the total pool nucleotide concentration. For example, it decreases from 0.2 to 0.05 for total ROS production by SMP when total NAD+ plus NADH concentration increases from 50 to 500 μM [83]. At optimal nucleotide concentration (50 μM), the rate of Н2О2 production fits the Nernst equation for a two-electron reaction with midpoint redox potential of − 350 mV ((NAD+/NADH)0.5 = 0.13) [75], a value close to the midpoint potential of FMN in complex I (− 370 mV at рН 8.0) [85]. The same dependence for superoxide production does not fit the Nernst equation, and half-maximal activity is observed at a significantly higher NAD+/NADH ratio (0.33) [75]. Superoxide production does proceed even when the nucleotide pool is

Fig. 1. Partitioning between superoxide and hydrogen peroxide as the products of complex I-mediated ROS production. Red and blue bars are superoxide and hydrogen peroxide, respectively. The lines on left diagram are drawn to emphasize complex kinetics of the NADH concentration dependence. Adapted from Ref. [75].

Please cite this article as: A.D. Vinogradov, V.G. Grivennikova, Oxidation of NADH and ROS production by respiratory complex I, Biochim. Biophys. Acta (2015), http://dx.doi.org/10.1016/j.bbabio.2015.11.004

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90% oxidized, i.e. at much more positive potential than that of the FMNH−/FMN couple. The dependence of superoxide production by purified complex I on the NAD+/NADH ratio measured by Kussmaul and Hirst fits the theoretical curve of the two-electron titration of the FMNH−/FMN couple with midpoint potential of −360 mV [18]. A single amino acid replacement (E95, a glutamate residue located near FMN) by glutamine in E. coli complex I results in a 15-fold increase in the rate of NADH-dependent hydrogen peroxide production [86]. The authors proposed the existence of closed and open states of the nucleotide-binding site in the FMN-containing subunit with different reactivity (accessibility) of flavin to oxygen, a model where the E95 residue keeps the site in the closed state. This observation seems to be relevant to the well-known stimulatory effects of guanidine on the ferricyanide reductase [87] and superoxide producing activities [75] of mammalian complex I. The electrostatic interaction between the large guanidinium cation and the glutamate anion might stabilize the open state. Interestingly, this negatively charged/neutral residue replacement was accompanied by about 5-fold decrease in HAR reductase activity [86]. The well-known “burst” of ROS upon anaerobic–aerobic state transition, such as organ reperfusion after ischemia, is a phenomenon that might be relevant to the unusual hysteretic kinetics of complex I. If no oxidized ubiquinone is available, the enzyme is transformed to the socalled deactivated state where electron transfer from N2 to ubiquinone is blocked (see Refs. [35,88] for reviews). The active (A) state-todeactivated state (D) transition has been detected for isolated complex I [89], SMP [13], intact heart mitochondria [90], and in ex vivo studies of perfused hearts [91]. In terms of catalytic activities, deactivation of complex I is equivalent to its inhibition by rotenone, which is known to increase ROS production. The back transformation of the D- to Aform is a slow process that is inhibited by free fatty acids and divalent metal cations [92–94]. Taken together, these data suggest the following scenario of the normal state → ischemia → reperfusion transition. Negligible complex I-mediated ROS production under the initial normal state occurs. It stops when no oxygen is available, and complex I become deactivated. A sudden increase in oxygen to the normal level would result in a burst of ROS because the deactivated enzyme will be directly oxidized by oxygen, not by ubiquinone. Increased ROS production is expected for the time needed for the slow D-to-A transformation and restoration of the normal ubiquinone reductase activity. When discussing forward and reverse electron transfer in complex I-mediated ROS production, a note should be made concerning the use of succinate as the respiratory substrate. The respiratory chain of coupled mitochondria or SMP oxidizing externally added succinate cannot be considered as a model of any physiologically conceivable situation. Succinate, an intermediate of the Krebs cycle, is produced and utilized in the mitochondrial matrix, providing one fifth of the reducing equivalents during complete oxidation of pyruvate. No other quantitatively significant cytoplasmic sources of succinate exists in aerobic metabolic pathways (α-oxoglutarate-dependent proline hydroxylation [95] and succinic semialdehyde transformation [96] are minor contributors to the total respiratory activity of mammalian tissues). This by no means excludes reverse electron transfer as a pathway for ROS production under some pathophysiological conditions, such as anoxia, where a significant amount of intramitochondrial succinate is accumulated [97]. Also, ubiquinol, the actual substrate for the reversal, is produced in several mitochondrial metabolic pathways such as fatty acid β-oxidation or oxidation of α-glycerophosphate. As discussed above, ROS-producing activity of complex I depends on the matrix redox potential, i.e. NAD+/NADH ratio [83]. The total amount of pyridine nucleotides in heart mitochondria can be approximated as 4–7 nmol per mg of protein [98–100], these values corresponding to 4–7 millimolar concentration. It can thus be assumed that the nucleotide-binding sites of complex I are always saturated. However, the concentrations of free NAD+/NADH nucleotides are not known. The physiologically relevant NAD+/NADH ratio in liver mitochondria

was approximated as about 8 [101]. In isolated cardiomyocytes respiring on 10 mM glucose [102] or in heart perfused by 10 mM glucose, this ratio is closed to 1 [103]. Thus, the apparent redox potential in the matrix is significantly higher than that conventionally used in the model experiments on ROS production. It appears that in vivo complex I-mediated ROS production is many-fold lower than measured under experimentally “optimal” conditions. In recent years, the term “p.m.f.(or energy)-dependent ROSproduction” is frequently used in the literature, thus implicitly assuming that the membrane energization and deenergization increases and decreases ROS production, respectively. We believe that this terminology can be misinterpreted by a reader who is only superficially familiar with mitochondrial bioenergetics. The production of ROS depends on the amount (concentration) of reactive sites accessible to oxygen. The steady-state NADH/NAD+ ratio, indeed, is decreased when respiration is activated by ADP (small drop of р.m.f.) or by uncoupler (complete dissipation of р.m.f.). However, it seems important to emphasize that р.m.f. itself does not affect the generation rate; it influences redox state of the oxygen reactive sites. No correlation between membrane energization and ROS production exists; for example, rotenone stimulates mitochondrial ROS production, whereas it completely deenergizes membranes. It might appear that this note is a matter of semantics; however, the expression “energy-dependent ROS generation” (by complex I) is misleading. 8. Notes on physiological and pathophysiological significance of complex I and other mitochondrial enzyme-mediated ROS production Mitochondria contain a number of enzymes capable of ROS generation. Their relative contributions along with complex I to the level of intracellular hydrogen peroxide are expected to be species- and tissue-dependent. To the best of our knowledge, only one study published more than 40 years ago was designed for very approximate quantitation of the relative contribution of various enzyme systems to H2O2 production in rat liver [104]. According to that report, about 15% of intracellular hydrogen peroxide is produced by mitochondria. The widespread opinion that mitochondria are the major source of ROS, apparently because mitochondria are indeed the major consumers of oxygen, is exemplified by citing D. Harman who wrote, “Mitochondria would be expected to be particularly subject to free radical induced change as over 90% of oxygen utilized by mammals takes place in them” [105]. This and other similar statements circulating in the literature are somehow misleading. Mitochondrial respiration inevitably decreases the local concentration of oxygen in the vicinity of the centers potentially capable of ROS production, which results in a decrease in the generation rate. The increase in atmospheric oxygen in the history of the Earth has led to great improvement in the energetics of life. On the other hand, it also necessitates protection against undesired deteriorating oxidative reactions. One strategy to solve the problem is to protect potentially reactive centers (flavins and/or iron-sulfur clusters) by the specific arrangement of the protein structure, including attachment of additional subunits that have no other function than to avoid their oxygen reactivity. Numerous experimental data show that almost all flavoand iron-sulfur proteins including those knowingly not operating functionally as oxidases do react with oxygen, thus producing either superoxide or hydrogen peroxide. In other words, their specific protection is not perfect. This is compensated by the widespread presence of SODs, peroxidases, and catalase. Does mitochondrial ROS production have any physiological function? Numerous reports have recently appeared in the literature where signaling cascade reactions with the participation of hydrogen peroxide are suggested and discussed (see for example review [106]). Note should be made that signaling molecules are defined, in contrast to metabolites, as those that bind to a receptor and induce its structural change without their chemical transformation. The structural

Please cite this article as: A.D. Vinogradov, V.G. Grivennikova, Oxidation of NADH and ROS production by respiratory complex I, Biochim. Biophys. Acta (2015), http://dx.doi.org/10.1016/j.bbabio.2015.11.004

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change of a receptor then leads to activation (or inhibition) of some metabolic pathways. Many hormones, second messengers (c-AMP, Са2 +), and protein factors are true signaling molecules. If expanded (unjustified in our opinion), the meaning of “signaling molecule” could be applied to any metabolite. In most schemes, the mechanism of hydrogen peroxide “signaling” is postulated as a trivial nonenzymatic oxidation of deprotonated sulfhydryl groups of the “target” that results in their catalytic (or further signaling) activities. It appears that if a regulatory signaling cascade does exist, all its steps must be enzymatically catalyzed, as is evident for the classic cascade regulation of glycogen metabolism. Another aspect worth brief discussion relevant to the physiological significance of mitochondrial ROS production is its dependence on oxygen concentration. In contrast to cytochrome oxidase, a reaction which has an apparent Km for oxygen significantly lower than the physiologically conceivable concentration (about 5-fold less than that in air-saturated aqueous solutions at normal pressure [107]), the specific activity of “major” respiratory chain-linked ROS generator, complex I [74,108,109], is simply proportional to oxygen concentration. Hyperbolic or other complex kinetics of complex I-mediated ROS production might be expected if it does have physiological function. On the other hand, simple proportionality of ROS formation to oxygen concentration might serve as an ideal oxygen sensing mechanism. The enzymes utilizing superoxide and hydrogen peroxide are frequently considered as an “antioxidant defense system”, whereas generation of ROS is considered as an evolutionarily unfavorable leakage reaction (except for NAD(P)H oxidases, which perform a clearly defined function). An alternative view is that hydrogen peroxide is a normal metabolic intermediate that fulfills some important not yet clear function(s). If a “leakage” hypothesis and “antioxidant defense system” are to be accepted, many more or less experimentally justified efforts to improve the defense (antioxidant drugs, particularly those mitochondrially targeted [110,111]) are certainly warranted. On the other hand, if the alternative view is correct, the use of so-called antioxidant drugs could result in unfavorable consequences. When discussing physiological and pathophysiological “oxidative stress”, one should keep in mind metabolic effects of the “reductive stress” that can be induced by the antioxidants [112]. Obvious expected consequences of “overreduction” are: 1) abnormal anabolic activity including malignant growth; 2) an increase in fatty acid and fat biosynthesis; 3) disorder of amino acid transport systems (participation of glutathione in the γ-glutamate cycle [113]); 4) a decrease of normal insulin level due to an increase in the insulin:glutathione transhydrogenase reaction [114,115]. Last, not the least consequence of reductive stress is a possible paradoxical activation of ROS production. Conflict of Interest We declare no conflict of interest. Acknowledgements We are indebted to Dr. J. Hirst for providing us data (Ref. [50]) before publication. We are grateful to anonymous Reviewer for his (her) kind linguistic corrections of our manuscript. The experimental works done in our laboratory were supported by Russian Foundation for Fundamental Research Grant 14-04-00279 to A.D.V. and Grant 15-04-02957 to V.G.G. References [1] L.A. Sazanov, P. Hinchliffe, Structure of the hydrophilic domain of respiratory complex I from Thermus thermophilus, Science 311 (2006) 1430–1436. [2] R.G. Efremov, R. Baradaran, L.A. Sazanov, The architecture of respiratory complex I, Nature 465 (2010) 441–445. [3] C. Hunte, V. Zickermann, U. Brandt, Functional modules and structural basis of conformational coupling in mitochondrial complex I, Science 329 (2010) 448–451.

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Please cite this article as: A.D. Vinogradov, V.G. Grivennikova, Oxidation of NADH and ROS production by respiratory complex I, Biochim. Biophys. Acta (2015), http://dx.doi.org/10.1016/j.bbabio.2015.11.004