Towards the molecular mechanism of respiratory complex I · core subunits of the best-studied complexes I; the nomenclature of Bos taurus complex I is used throughout the present

Biochem. J. (2010) 425, 327–339 (Printed in Great Britain) doi:10.1042/BJ20091382 327

REVIEW ARTICLETowards the molecular mechanism of respiratory complex IJudy HIRST1

Medical Research Council Mitochondrial Biology Unit, Wellcome Trust/MRC Building, Hills Road, Cambridge CB2 0XY, U.K.

Complex I (NADH:quinone oxidoreductase) is crucial torespiration in many aerobic organisms. In mitochondria, itoxidizes NADH (to regenerate NAD+ for the tricarboxylic acidcycle and fatty-acid oxidation), reduces ubiquinone (the electronsare ultimately used to reduce oxygen to water) and transportsprotons across the mitochondrial inner membrane (to produce andsustain the protonmotive force that supports ATP synthesisand transport processes). Complex I is also a major contributor toreactive oxygen species production in the cell. Understanding themechanisms of energy transduction and reactive oxygen speciesproduction by complex I is not only a significant intellectualchallenge, but also a prerequisite for understanding the rolesof complex I in disease, and for the development of effectivetherapies. One approach to defining a complicated reactionmechanism is to break it down into manageable parts that canbe tackled individually, before being recombined and integrated

to produce the complete picture. Thus energy transductionby complex I comprises NADH oxidation by a flavin mono-nucleotide, intramolecular electron transfer from the flavin tobound quinone along a chain of iron–sulfur clusters, quinonereduction and proton translocation. More simply, molecularoxygen is reduced by the flavin, to form the reactive oxygenspecies superoxide and hydrogen peroxide. The present reviewsummarizes and evaluates experimental data that pertain tothe reaction mechanisms of complex I, and describes and dis-cusses contemporary mechanistic hypotheses, proposals andmodels.

Key words: electron transport chain, enzyme mechanism,mitochondrion, NADH:quinone oxidoreductase, proton-coupledelectron transfer, proton translocation.

COMPLEX I IN THE MITOCHONDRIAL ELECTRON TRANSPORTCHAIN

Complex I (NADH:quinone oxidoreductase) is an entry point forelectrons into the respiratory electron transport chains of manyaerobic organisms. In mitochondria, the electron transport chaintransfers two electrons from NADH to oxygen; it extracts thepotential energy in three sequential steps, and uses it to transportprotons across the mitochondrial inner membrane, to produceand sustain the protonmotive force (�p) which supports thesynthesis of ATP and the transport of metabolites. Mitochondrialcomplex I oxidizes NADH, produced predominantly by thetricarboxylic acid cycle and the β-oxidation of fatty acids, toNAD+, to regenerate the NAD+ pool in the mitochondrial matrix(Figure 1). The two electrons from NADH oxidation reduceQ (ubiquinone) to QH2 (ubiquinol) in the inner membrane;the QH2 is subsequently reoxidized by complex III (QH2:cytochrome c oxidoreductase, cytochrome bc1). Finally, thesubstrates and cofactors of complex I have the lowest potentialsin the mitochondrial respiratory chain, so it is not surprisingthat complex I is increasingly recognized as a major contributorto ROS (reactive oxygen species) formation, with possiblephysiological and pathological effects [1].

THE COMPOSITION AND STRUCTURE OF MITOCHONDRIALCOMPLEX I

Complex I has an unusual L-shaped structure, with one arm inthe plane of the mitochondrial inner membrane, and one armprotruding into the matrix (Figure 1). The L-shaped structure,

observed by electron microscopy, is conserved in all knowncomplexes I, from mitochondria to bacteria [2,3], althoughthe protein composition of the eukaryotic and prokaryoticenzymes varies significantly. Common to all complexes I are14 ‘core subunits’ (comprising the ‘minimal enzyme’ of manyprokaryotes; [4]) that are considered sufficient for energytransduction. The core subunits comprise two distinct groups(see Table 1). The seven predominantly hydrophilic subunits areencoded in the nucleus and constitute the membrane-extrinsicarm. The seven highly hydrophobic subunits are dominated byTMHs (transmembrane helices); in eukaryotes they are encodedby the mitochondrial genome, and they constitute the membrane-intrinsic arm [5]. Eukaryotic complexes I contain ‘supernumerary’subunits also; they vary between species and augment bothenzyme domains [6], but they are very unlikely to participatedirectly in energy transduction and are not considered further inthe present review. Table 1 summarizes the nomenclatures for thecore subunits of the best-studied complexes I; the nomenclatureof Bos taurus complex I is used throughout the present review,regardless of the species referred to.

The structure of the hydrophilic domain of complex Ifrom Thermus thermophilus, reported to 3.3 Å (1 Å = 0.1 nm)resolution [8], is the first high-resolution structural model for anypart of complex I. It describes the seven hydrophilic core subunits,and one supernumerary subunit, Nqo15, which is particular to T.thermophilus and a few other organisms. The seven core subunitsform a bulbous Y-shaped domain; the lower part (the PSST,TYKY, 30 and 49 kDa subunits) comprises the interface withthe membrane domain, and in the upper part, two domains areformed by the 51 and 24 kDa subunits, and by the 75 kDa subunit

Abbreviations used: DQ, decylubiquinone; FMN, flavin mononucleotide; HAR, hexa-ammineruthenium III; Q, ubiquinone; QH2, ubiquinol; ROS, reactiveoxygen species; SQ, semiquinone; TMH, transmembrane helix.

1 email [email protected]

c© The Authors Journal compilation c© 2010 Biochemical Society

328 J. Hirst

Table 1 The nomenclature for the 14 core subunits of complex I, the conserved cofactors bound by the hydrophilic subunits and the predicted TMHs whichcomprise the ND subunits in B. taurus (predicted using ConPredII [7], and not verified experimentally)

Enzyme domain B. taurus Homo sapiens Y. lipolytica E. coli T. thermophilus Cofactors and TMHs

Hydrophilic arm (nuclear encoded in eukaryotes) 75 kDa NDUFS1 NUAM NuoG Nqo3 [2Fe–2S], 2 × [4Fe–4S]51 kDa NDUFV1 NUBM NuoF Nqo1 FMN, [4Fe–4S]49 kDa NDUFS2 NUCM NuoCD Nqo430 kDa NDUFS3 NUGM Nqo524 kDa NDUFV2 NUHN NuoE Nqo2 [2Fe–2S]PSST NDUFS7 NUIM NuoB Nqo6 [4Fe–4S]TYKY NDUFS8 NUKM NuoI Nqo9 2 × [4Fe–4S]

Hydrophobic arm (mitochondrial encoded in eukaryotes) ND1 ND1 NU1M NuoH Nqo8 8 TMHsND2 ND2 NU2M NuoN Nqo14 8 TMHsND3 ND3 NU3M NuoA Nqo7 3 TMHsND4 ND4 NU4M NuoM Nqo13 13 TMHsND5 ND5 NU5M NuoL Nqo12 16 TMHsND6 ND6 NU6M NuoK Nqo10 5 TMHsND4L ND4L NULM NuoJ Nqo11 3 TMHs

Figure 1 The reactions catalysed by complex I in the mitochondrion

NADH produced by the breakdown of carbohydrates, fats and proteins, is oxidized to NAD+, and two electrons are transferred, via an FMN and a chain of iron–sulfur clusters, to reduce Q to QH2.The redox reaction regenerates NAD+, provides electrons for the reduction of O2 to water, and provides the free energy for proton translocation across the inner membrane. Proton translocationcontributes to the �p which is used, for example, to synthesize ATP. The fully reduced flavin in complex I also reduces O2 to superoxide, a cause of cellular oxidative stress. ET chain, electrontransport chain; TCA cycle, tricarboxylic acid cycle.

(see Figure 2). The structure as a whole encases the cofactorcohort of the hydrophilic arm; a cluster ‘chain’ runs between thesites of NADH oxidation and quinone reduction, in a commonconfiguration found in, for example, succinate dehydrogenaseand fumarate reductase, nitrate reductase and some formatedehydrogenases. The large domain of the 75 kDa subunit in T.thermophilus and a small number of other organisms, includingEscherichia coli, co-ordinates an additional [4Fe–4S] cluster, N7,which is not considered further in the present review; it is notconserved, and too distant (>20 Å) from the other clusters toparticipate in catalysis [9].

The conserved cofactors of complex I are a non-covalentlybound FMN (flavin mononucleotide) and eight iron–sulfurclusters, all bound by the hydrophilic domain (Table 1 andFigure 2). Prior to the structure, the clusters had been predictedcorrectly by sequence analyses, the overexpression of putativecluster-containing subunits, and EPR analyses (reviewed in[6,10]). No further permanently bound redox cofactors are known;

previous suggestions of a novel cofactor, perhaps a quinoid group,in the membrane domain, have not been substantiated by MS[11]. The confinement of all the cofactors to the hydrophilicdomain (Figures 1 and 2) is important in the formulation ofmechanistic hypotheses for complex I, and distinguishes it fromother proton-translocating redox enzymes, such as cytochromebc1 and cytochrome c oxidase. The NADH-binding site is locatedin the 51 kDa subunit of complex I, adjacent to the flavin (seebelow), but the location (and number) of the quinone-bindingsites remain unconfirmed; a putative binding site for the quinoneheadgroup is marked on Figure 2 (see below). Currently, thereis significant controversy about how to map the structure ofthe T. thermophilus hydrophilic domain on to low-resolutionelectron microscopy structures of the intact complex. Brandtand co-workers contend that the whole domain is ‘lifted away’from the membrane, placing cluster 4Fe[PS]/N2 35–60 Åaway from the membrane interface, and requiring quinone tomove substantially out of the membrane for catalysis [12,13]. In


Towards the molecular mechanism of respiratory complex I 329

Figure 2 The structure of the hydrophilic core subunits of complex I from T. thermophilus, and the chain of conserved cofactors [8]

The NADH and putative Q-binding sites (see text) are indicated. The subunits are named according to the nomenclature of B. taurus complex I. The conserved clusters (left, duplicate view butwithout the protein) are named according to their type (2Fe or 4Fe) and subunit, and then differentiated as all cysteine-ligated (C) or with one histidine ligand (H), or as clusters 1 or 2 (in TYKY). Thenon-conserved 4Fe cluster in the 75 kDa subunit has been omitted from the chain.

contrast, I (and others) contend that the hydrophilic domain ispositioned with 4Fe[PS]/N2 close to the membrane (see Figure 1,same orientation in Figure 2), so that the quinone headgroupcan approach 4Fe[PS]/N2, but remain in the membrane interface(reflecting its positions in succinate:ubiquinone oxidoreductase,cytochrome bc1 and the bacterial reaction centre). In support of thelatter model, a quinazoline inhibitor has been found to bind boththe N-terminal section of the 49 kDa subunit and ND1 [14], andcross-links have been formed by the PSST and 49 kDa subunitswith ND3 [15]. However, all of the evidence is circumstantialand the controversy will probably only be resolved when ahigh-resolution structural model of intact complex I becomesavailable.

The seven hydrophobic subunits of complex I comprise a largenumber of TMHs – the seven B. taurus subunits are predicted tocontain 56 of them (see Table 1), although the predictions have notbeen confirmed experimentally, and the subunit lengths arenot well conserved between species. For five of the ND subunits,fusion proteins (in Rhodobacter capsulatus [16] and Paracoccusdenitrificans [17]) have been used to infer the number of TMHs(and the protein orientations), and the results match the predictionsin Table 1. The arrangement of the ND subunits in the membranedomain has not been determined unambiguously, although ND4and ND5 have been located at the distal end by single-particleanalyses [18]. ND4, ND5 and ND2 are related to the Mrp familyof Na+/H+ antiporters [16], and so, despite the distance of ND4and ND5 from the hydrophilic arm, they are often suggested toparticipate in proton translocation (see below).

OVERVIEW OF COMPLEX I CATALYSIS

Under normal physiological conditions, mitochondrial complex Icatalyses the two-electron oxidation of NADH and reduction of Q.The redox reaction (eqn 1) is thermodynamically very favourable(�EpH7 =+ 0.4 V, although the physiologically relevant value of�E is determined also by the solution and the membrane con-ditions, the position of the Q-headgroup in the membrane [19],and by the NAD+/NADH and Q/QH2 ratios).

NAD+ + 2H+ + 2e− ↔ NADH + H+ EpH7 = −0.33 V [20]

Q + 2H+ + 2e− ↔ QH2 EpH7 = +0.07 V [19] (1)

For complex I in an energy-transducing membrane, the‘favourable’ redox reaction is opposed by ‘unfavourable’ protontranslocation, and the redox energy is transformed and conservedas �p. It is generally accepted that, in mitochondria at least,complex I translocates four protons for each two-electronoxidation of NADH. Therefore the �p supported by complex Iis thermodynamically limited to �pLim =�E/2, as the system isnever perfectly balanced some energy is always lost asheat (energy conservation is not completely efficient).As an estimation, in the mitochondrion �E ∼ 0.35 V so�pLim ∼ 0.175 V {pH 7.8, [NAD+] = 10[NADH] ([21] andreferences therein), and [Q] = [QH2] (value unknown)}. In vitro,if �p >�E/2, complex I catalyses the reverse reaction also: NAD+


330 J. Hirst

reduction and QH2 oxidation, driven by proton discharge acrossthe membrane [22]. Although the general events which comprisethe forward and reverse cycles are surely the reverse of oneanother (for example, NADH reduces the oxidized flavin, NAD+

oxidizes the reduced flavin), the two reactions are not necessarilymicroscopically the reverse of one another (the reaction is not atequilibrium so the forward and reverse cycles are not constrainedto identical pathways) [23]. A useful analogy is presented bya fixed-gear bicycle: rotate the pedals forward and the bikemoves forward, rotate the pedals backward and the bike movesbackward, so in a general sense the bicycle is reversible. However,on more detailed viewing, for example of the chain on the teethof the cogs, or the feet and ankles of the cyclist, it is obvious thatthe forward and backward motions are not the exact reverse ofeach other (a snapshot could easily reveal the direction of mo-tion). Similarly, in complex I, the structures (and lifetimes) oftransition states and reaction intermediates may vary according tothe direction of catalysis. The present review discusses complex Icatalysing the physiological ‘forward’ reaction; reverse catalysisby the mitochondrial enzyme has not been studied extensivelyand is probably non-physiological but, nevertheless, it may yieldimportant and complementary mechanistic information.

If we follow a pair of electrons from their entry to complex Ion a molecule of NADH, then catalysis comprises their transfer(as a pair) to the flavin, their transfer (one at a time) acrossthe chain of iron–sulfur clusters, and their reunion as quinoneis reduced to quinol. At an unknown point, but close to theirarrival on the bound quinone, the electrons, which so far haveretained most of their original potential energy, are harnessedto translocate protons across the membrane. Alternatively, anelectron may ‘escape’ by hijacking a molecule of O2. In thesections below, the mechanism of complex I is broken downinto separate steps (NADH oxidation and O2 reduction at theflavin site, intramolecular electron transfer, quinone reductionand proton translocation), although it is important to recognizethat the separate steps do not, and cannot, occur independently.

NADH OXIDATION AND THE REACTIONS OF THE FMN

In complex I, the primary function of the flavin is to oxidizeNADH. The interconversion of NADH and NAD+ is catalysedby many flavoproteins, and generally considered (but not proven)to proceed by hydride transfer, rather than by transfer of twoelectrons and one proton (thus avoiding the highly unstableNAD•). To transfer the hydride, the nicotinamide ring of NADHstacks above the flavin isoalloxazine ring system, juxtaposing thehydride donor (C-4 of the nicotinamide ring) and acceptor (N-5 of the flavin) in a defined orientation, separated by ∼3.5 Å[24]. Recently, structures of the hydrophilic domain of complexI from T. thermophilus with nucleotides bound [25] haverevealed the expected juxtaposition (see Figure 3), along withspecific interactions between the protein and the nucleotide(stacking interactions between three phenylalanine residues andthe adenine ring, a number of hydrophobic interactions, and hy-drogen bonds to the nicotinamide carboxyamide, two ribosemoieties and phosphates). However, the oxidation states of thebound nucleotides, flavin and clusters in these structures haveonly been inferred from the crystallization conditions, and fromspectroscopic data from different species of complex I. In thefuture, it will be crucial to elucidate structures in a variety of pre-cisely defined states, to fully understand the determinants of anystructural changes, and to address the structures of the reactivecomplexes.

Figure 3 The structure of the nucleotide-binding site in complex I from T.thermophilus [25]

The surface representations are for the protein (without the flavin or the nucleotide, grey),the flavin (blue) and the nucleotide (red). Residues in the 51 kDa subunit (T. thermophilusnumbering) which form hydrogen bonds (shorter than 3.5 A) are shown in yellow, phenylalanineresidues which form stacking interactions with the adenine ring are in cyan, and the conservedactive site glutamate residue is in green.

The flavin in complex I (at least in B. taurus) has anunusually low potential (−0.38 V at pH 7.8; [26]), close tothe potential of NAD+ (−0.35 V at pH 7.8; [20]). Therefore itcatalyses both NADH oxidation and NAD+ reduction, and thethermodynamic reversibility of NADH:flavin oxidoreduction hasbeen demonstrated electrochemically: the hydrophilic domainof B. taurus complex I, adsorbed to an electrode surface,establishes a Nernstian equilibrium between NAD+ and NADH[27]. Note that the ability of complex I to reduce NAD+ duringreverse electron transfer [22] does not necessarily require theinterconversion of NADH and NAD+ to be thermodynamicallyreversible (i.e. an infinitesimally small driving force, on eitherside of the equilibrium position, leads to net catalytic turnover;[28]). The implications of the electrochemical reversibilityare that NADH:flavin oxidoreduction is kinetically fast andthermodynamically efficient.

Catalytic NADH oxidation by complex I (followed readilyby NADH absorption at 340 nm) can be coupled to the reduc-tion of a variety of electron acceptors to evaluate different aspectsof the mechanism. For example, NADH:FeCN [ferricyanide,hexacyanoferrate(III)] [29] and NADH:APAD+ [21] oxidoreduc-tion involve only the flavin; rotenone- or piericidin A-sensitiveNADH:DQ (decylubiquinone) oxidoreduction comprises NADHoxidation, intramolecular electron transfer and quinone reduction,and, when complex I is incorporated in a vesicle, protontranslocation across the vesicular membrane. NADH:FeCNoxidoreduction is the fastest known reaction: for complexI from B. taurus it achieves several thousand NADH persecond [29]. In comparison, the highest rate of NADH:DQoxidoreduction from purified B. taurus complex I so far is∼100 s−1 [30]. Therefore NADH oxidation (comprising hereNADH binding, hydride transfer and NAD+ release) is fast(kcat

NADH/KmNADH > 1 × 107 M−1 · s−1 [21]) and not rate-limiting

in NADH:DQ oxidoreduction.FeCN [29], APAD+ [21] and hydrophilic quinones such as

Q-1 [31] and O2 [32], all react directly with the reduced flavin incomplex I: they are all inhibited by high NADH concentrations,and can be described by Ping Pong or Ping Pong Pong mechanisms(see Figure 4). So far, kinetic analyses based on these reactions



Figure 4 Kinetic analysis of a Ping Pong reaction catalysed by the flavin incomplex I

Left-hand panel: NADH oxidation by the flavin is described by a Michaelis–Menten reaction,to produce NAD+ and the reduced flavin. Similarly, APAD+ reduction produces APADH andreoxidizes the flavin. APAD+ binds only weakly to the oxidized flavin (K Ox

APAD+ is large) and soit does not inhibit turnover (top, right-hand panel). Conversely, NADH inhibits the reaction athigh concentration (bottom, right-hand panel) because it binds to the reduced flavin (K Red

NADH)and blocks the reaction of APAD+ . Data (right-hand panel) were modelled using the reactionscheme shown. Many different parameter combinations fit the data [21]; the values used herewere K m

NADH = 0.094 mM, k catNADH = 2700 s−1, K Red

NADH = 0.16 mM, K mAPAD+ = 0.32 mM,

k catAPAD+ = 340 s−1, K Ox

APAD+ = 5.0 M. Figure adapted with permission from [21]. c©2007American Chemical Society.

have provided estimated dissociation constants for NADH fromthe reduced flavin (20 μM [33] and 160–260 μM [21]) andNAD+ from the oxidized flavin (800 μM; [33]), but these valuesrefer to inhibited or product-bound states, and, due to theirkinetic complexity, even these simple one-site reactions haveso far eluded complete description. In addition, mutation of theconserved glutamate residue in the NADH-binding site of E.coli complex I has been used to suggest that its negative chargeis important for determining nucleotide-binding affinities [34],and three inhibitors are known to inhibit NADH oxidation bycomplex I directly. Diphenyleneiodonium forms a covalent adductwith the reduced flavin [35], and ADP-ribose (KOx

ADPR∼30 μMand KRed

ADPR∼500 μM [33]) and NADH-OH (KOxNADH−OH∼

0.3 nM and KRedNADH−OH∼7 nM [33]) are NADH analogues. Fi-

nally, HAR (hexa-ammineruthenium III) is often used to evaluatethe kinetic capability of the flavin site in complex I, becauseNADH:HAR oxidoreduction is relatively fast [36]. However,HAR reduction is not inhibited by high NADH concentrations,and its mechanism is not well understood (it has been proposedto occur by an ‘ordered’ mechanism): at present, HAR should notbe used to quantify changes in the rates of specific reactions atthe flavin, or to determine kinetic or thermodynamic constants.

THE REDUCTION OF MOLECULAR O2 BY THE REDUCED FLAVIN

There has been much debate about the site(s) and mechanism(s) ofO2 reduction by complex I, leading to the production of superoxideand/or H2O2. Essentially every possibility has been discussed[the reduced flavin and flavosemiquinone, NAD• radical, iron–sulfur clusters and SQ (semiquinone) intermediates]. Oxygenreduction by the reduced flavin [32] is now recognized asan important contributor that links superoxide production bycomplex I to the status of the mitochondrial NAD+/NADHpool, but a second site (either 4Fe[PS]/N2 or a SQ) may also

Figure 5 The mechanism of O2 reduction by the reduced flavin incomplex I

Top panel: schematic representation of how the rate of O2 reduction by complex I is determined(see text for details). Bottom left-hand panel: dependence of the rate of H2O2 productionon E SET (matching the expected potential dependence of the fully reduced flavin, not theflavosemiquinone), and showing the position of E 1/2. Bottom right-hand panel: dependence ofE 1/2 on pH, and comparison with values for the flavin determined by EPR and redox titrations[26]. Figure adapted with permission from [32]. c©2006 National Academy of Sciences.

contribute under some conditions [1]. A further possibility, the‘conveyor molecule’ mechanism, is that superoxide productionis increased by physiological or pharmacological agents that arereduced by the flavin, then reoxidized by O2 in redox-cyclingreactions; hydrophilic quinones have recently provided the firstwell-characterized (although non-physiological) example [31].The present review focuses on the direct reaction of O2 withthe reduced flavin in complex I.

In complex I from B. taurus, O2 reduction by the reducedflavin is much slower than NADH oxidation or NAD+ reduction(∼0.5 NADH · s−1 in atmospheric O2). Consequently, whencomplex I is incubated in a mixture of NADH and NAD+, anequilibrium is established between the oxidized and reducedflavins (the flavosemiquinone is unstable and present only inlow amounts). Because O2 reacts comparatively slowly with thereduced flavin, it does not perturb the equilibrium significantly,and the rate of reaction is the product of the reduced flavin and O2

concentrations, and the corresponding bimolecular rate constant[32] (see Figure 5). In reality, the equilibrium position is a functionof nucleotide binding also, because O2 reduction is blocked bybound nucleotides (it is inhibited by high NADH concentrations[32,37], and solvent access to the flavin is blocked in thenucleotide-bound T. thermophilus structures; [25]). However,knowledge of the nucleotide-binding constants is too limited fora quantitative evaluation at present, so in Figure 5 the rate of O2

reduction (H2O2 production) is plotted against the potential of theNAD+/NADH pool (expressed as ESET, from the Nernst equationfor NAD+ reduction). The midpoint potential (E1/2) of the curve isconsistent with the reduction potential of the flavin determined byEPR [26] (rather than with any of the SQ or cluster potentials, seebelow), and the variation of E1/2 with pH matches that expectedfor the flavin also (rather than matching a pH-independent clusterpotential). Finally, the sigmoidal shape shows clearly that, in therate-determining step, O2 reacts with the fully reduced flavin, notwith the flavosemiquinone.

In complex I from B. taurus, O2 reduction by the (two-electron) reduced flavin generates superoxide (a one-electronreduced species) rather than H2O2, indicating that the ‘second’electron is dissipated to the clusters, or that the nascent


332 J. Hirst

Table 2 The eight conserved iron–sulfur clusters in complex I

est., estimated; n.d., not detected

Subunit Cluster Ligation1 EPR [39] E pH7 (V)2 Comments on EPR signals

24 kDa 2Fe[24] C · X4 · C · X35 · C · X3 · C N1a est. -0.5 Not observed in intact B. taurus complex I3

51 kDa 4Fe[51] C · X2 · C · X2 · C · X39 · C N3 -0.25 Spin interactions with flavosemiquinone [26], E pH7 from redox titration [10,41].75 kDa 2Fe[75] C · X10 · C · X2 · C · X13 · C N1b est. -0.4 Reduced partially by NADH [42]3

4Fe[75]C C · X2 · C · X2 · C · X43 · C N5 -0.26 N5 signal fast relaxing, E pH7 from redox titration [10,41].4Fe[75]H H · X3 · C · X2 · C · X5 · C – n.d. Not observed by EPR

TYKY 4Fe[TY]1 C · X2 · C · X2 · C · X42 · C N4 -0.25 N4 may arise from 4Fe[TY]1 and/or 4Fe[TY]2: for simplicity it is attributed here to 4Fe[TY]1. EpH7 (N4) from redoxtitration [10,41].

4Fe[TY]2 C · X28 · C · X2 · C · X2 · C – n.d.PSST 4Fe[PS] C · C · X63 · C · X29 · C N2 -0.15 Spin interactions with ubisemiquinones [43]. E pH7 from redox titration [10,41].

1 The ligation motifs are for the clusters in B. taurus complex I; there are small differences between species, for example, in T. thermophilus extra residues in the 2Fe[75], 4Fe[TY]1 and 4Fe[TY]2motifs.

2 Cluster potentials are reported for B. taurus complex I.3 Literature reports of N1a in B. taurus complex I result from confusion between N1a and N1b; this confusion has also caused problems with defining the stoichiometry and potential of N1b.

superoxide dissociates before it can be further reduced [32].It has been suggested [8] that the distal 2Fe[24] cluster (seeFigure 2) transiently accepts the second electron (to quench theflavosemiquinone), a suggestion that is supported by a recent studyof E. coli complex I [38]. The distal cluster in E. coli complexI has a relatively high potential, so it is ‘already’ reduced at allof the values of ESET considered, and E. coli complex I producespredominantly H2O2. Although the mechanism provides a role forthe distal cluster, it is unclear why a ‘special’ cluster is required.In addition, multiple turnovers are not well accounted for, it isunclear whether the potential of even the mitochondrial clusteris appropriate, and why would mitochondrial complex I haveevolved to produce superoxide instead of H2O2? It is possiblethat the apparent link between 2Fe[24] potential and superoxidecompared with H2O2 production is simply coincidental, and that2Fe[24] is an evolutionary remnant, with no special role in thecontemporary enzyme.

INTRAMOLECULAR ELECTRON TRANSFERIn complex I, seven iron–sulfur clusters, one [2Fe–2S] cluster andsix [4Fe–4S] clusters, form a chain which connects the flavin withthe quinone-binding site, and the distal [2Fe–2S] cluster has noknown role in energy transduction (see Figure 2 and Table 2).Essentially all extant information about the properties of theclusters in complex I has been derived using EPR spectroscopy[10]: reduced 2Fe and 4Fe clusters are paramagnetic, so thedependence of signal intensity on potential defines the reductionpotential (see Table 2). When complex I from B. taurus is reducedby NADH, five EPR signals, from one reduced 2Fe cluster andfour reduced 4Fe clusters, are apparent (see Figure 6). Therehas been extensive discussion about how to assign the five EPRsignals to the eight clusters, based mostly on comparisons ofspectra from overexpressed subunits and enzyme fragments withthose from the isolated enzyme [10,39]. Note that in Figure 2and Table 2, the names of the clusters are based on the structure;the clusters are often named by their proposed EPR signals (N1a,N1b, N2, etc.), but the original assignment is not correct [39]and this practice continues to confuse. In addition, the EPRspectra vary between species (for example, NADH-reduced E.coli complex I exhibits N1a, but not N5 [10,40]); the variationsmay result from ‘insignificant’ differences in cluster environmentor reduction potential, or they may not. In any case, transferringreduction potentials and EPR signal assignments between speciesof complex I is not necessarily reliable.

In complex I, assignment of the 2Fe signals is straightforward.The overexpressed 75 kDa subunit exhibits a 2Fe signal verysimilar to that from the intact enzyme (signal N1b) [39,44],whereas the overexpressed 24 kDa subunit exhibits a differentsignal (N1a) that is observed in the reduced flavoproteinsubcomplex (51 + 24 kDa), but not in intact B. taurus complexI (the cluster is not reduced by NADH) [42]. Therefore N1b isfrom 2Fe[75] and N1a is from 2Fe[24]. Signal N1a is observedin NADH-reduced E. coli complex I [40], as 2Fe[24] has a higherreduction potential in E. coli than in B. taurus [45]. The 4Fe signalsN2 and N3 can be assigned with confidence, to 4Fe[PS] and4Fe[51] respectively, since N2 interacts paramagnetically with aSQ intermediate [43], and N3 with the flavosemiquinone [26].The assignment of N4 and N5 is still disputed [39,46]. Here, weconsider that N4 is from one (or both) of 4Fe[TY]1 and 4Fe[TY]2,and N5 is from 4Fe[75]C [39]. Our assignment (see Table 2) isconsistent with all extant data, but has not been demonstrateddirectly yet; here, for simplicity, we take the assignments of N4to 4Fe[TY]1 and N5 to 4Fe[75]C as our model.

Combining the EPR signal assignment with data from redoxtitrations (see Table 2) [10,41] produces a reduction potential‘profile’ for electron transfer along the cofactor chain (seeFigure 7A). Cluster 4Fe[PS]/N2 has the highest potential. Clusters4Fe[51]/N3, 4Fe[TY]1/N4 and 4Fe[75]C/N5 are all considered‘isopotential’: in redox titrations they are all reduced at around−0.25 V (although the reduced-cluster stoichiometry has not beenestablished unambiguously). Cluster 2Fe[75]/N1b is enigmaticbecause it is reduced over an unusually wide potential range [42],but its potential is certainly below −0.25 V. Further reductionof B. taurus complex I requires very low potential donors: newsignals are observed at approx. −1 V [42], but they are notsimple rhombic signals and are probably due to interactionsbetween adjacent reduced clusters. Interestingly, signal N1a wasnot observed even at −1 V: the reduction of 2Fe[24]/N1amay be limited kinetically, because protein-film voltammetrystudies found that its reduction potential is above −0.5 V [45].In Figure 7(A), the estimated potentials for 2Fe[24]/N1a and2Fe[75]/N1b, and also for 4Fe[75]H and 4Fe[TY]2 (which arevery probably oxidized in NADH), include a consideration of theirpositions between clusters that are reduced at higher potentials,so that electrostatic interactions decrease their apparent reductionpotentials in redox titrations [47]. Finally, redox titrations relyon equilibrium potentials: the enzyme is in a ‘resting’ statethat may not be catalytically relevant, and equilibrium potentialsmay not reflect the free-energy changes during turnover. With



Figure 6 EPR spectra of complex I from B. taurus reduced by NADH

The spectra recorded at 12, 40 and 4 K (black) are compared with their simulated spectra(red). The inset for the 12 K spectrum (not on the x-axis scale) shows how the modelled 12K spectrum comprises N1b (grey), N2 (green), N3 (blue), N4 (magenta) and N5 (orange). Theg-values used were: N1b (gz,y,x = 2.024, 1.941, 1.926), N2 (gz,y,x = 2.055, 1.926, 1.925), N3(gz,y,x = 2.041, 1.927, 1.865), N4 (gz,y,x = 2.114, 1.930, 1.882) and N5 (gz,y,x = 2.064, 1.928,1.898). Figure adapted with permission from [42]. c©2008 American Chemical Society.

these points in mind, the alternating potential or ‘rollercoaster’profile [48] in Figure 7(A) is consistent with rapid electron transferalong the chain (single low-potential clusters impede electrontransfer much less than adjacent low-potential clusters) [49].In Figure 7(A), the longest distance for transfer (4Fe[75]H to4Fe[TY]1/N4), is compensated for by an increase in reductionpotential (kforw = 1.8 × 105 s−1 estimated by Dutton’s equation;[49]). The predicted rate-limiting step in forward electron transferis 4Fe[51]/N3 to 2Fe[75]/N1b (kforw = 7.1 × 104 s−1). Duringreverse-electron flow (from 4Fe[PS]/N2 to 4Fe[51]/N3) the rate-limiting step is predicted to be the longest distance transfer(4Fe[TY]1/N4 to 4Fe[75]H) (krev = 4.3 × 103 s−1; [49]). Recently,Verkhovskaya et al. [50] used freeze-quenching to show that thetwo highest potential clusters in E. coli complex I (4Fe[PS]/N2and 2Fe[24]/N1a) were reduced by NADH with an apparent timeconstant of ∼90 μs (assuming no electron transfer occurs inthe frozen sample), and that the second pair of reduced clusters(observed as signals N1b and N4, referred to as N6b) wereformed more slowly (∼1 ms). The results are consistent with fastintramolecular electron transfer from the flavin to 4Fe[PS]/N2(<90 μs, it is not clear whether the rate-limiting step is hydridetransfer, flavin reoxidation or intramolecular electron transfer)and with rate-limiting NAD+ dissociation (within the NADH to4Fe[PS]/N2 component) during turnover.

Figure 7 Electron transfer reactions in complex I

(A) Potential energy profile for the substrates and cofactors in complex I (based on data from B.taurus). The substrate potentials are two electron potentials, and two values for Q are included,for �p = 0 V and �p = 0.15 V. The substrate potentials and the two flavin potentials, FMN1(ox/semi) and FMN2 (semi/red) [26] are at pH∼7.8, and they include the coupled protonations.Cluster 2Fe[24]/N1a is not part of the chain. Values for 4Fe[51]/N3, 4Fe[75]C/N5, 4Fe[TY]1/N4and 4Fe[PS]/N2 are from [41]; values for the other clusters are estimates. Edge-to-edge distancesare indicated (in A). (B) Possible scheme for the transfer of two electrons to bound quinone,upon the oxidation of NADH (see text for details). Two electrons are highlighted in red for theiridentification.

It is likely that the cluster chain in complex I is already partiallyreduced during steady-state catalysis, so that a single electron pairdoes not traverse the complete chain during a single turnover.Figure 7(B) shows a model for rapid and efficient intramolecularelectron transfer in complex I. At −0.3 V (the potential ofthe mitochondrial NAD+/NADH pool), complex I contains fourelectrons, spaced alternately along the chain. NADH oxidationis reversible and fast (in comparison with Q reduction), and amixture of states is established, including ‘poised’ states such as{NADH:FMN} (NADH bound to oxidized flavin) [alternativepoised states are {NAD+:FMNH−}, {NADH:FMNH−} and{FMNH−}], as well as non-reactive states [{NAD+:FMN}and {FMN}]. Interestingly, if quinone binds to a non-reactivestate, then it is unlikely to be reduced, provided that sufficient�p is present (see Figure 7A). Consequently, turnover onlyoccurs when a poised enzyme binds Q, and NADH oxidation(or its equivalent) and Q reduction are enabled together. A SQintermediate is observed in the presence of �p (see below);even though NADH is a two-electron donor and SQ formationrequires only one electron, the flavin remains predominantlyoxidized because the cluster chain retains the extra electron,most likely on 2Fe[75]/N1b. NAD+ dissociation is followed byreversible nucleotide binding and reaction, as described above,and transfer of the second electron from the chain generatesthe quinol product, which then dissociates. Thus Figure 7(B)shows how the effective rate of electron transfer is increased (in


334 J. Hirst

analogy to the Grotthus mechanism of proton transfer) above thealready high rates predicted by Dutton’s model and implied by thedata of Verkhovskaya et al. (i.e. electron delivery to 4Fe[PS]/N2is very unlikely to exert any rate-limiting effect on catalysis).In addition, provided that the individual electron transfers arefast and reversible, the redox potential energy of the NADHis transferred efficiently to the bound quinone (in analogy toefficient energy transfer by Newton’s cradle) – this may be keyfor thermodynamic reversibility in catalysis by complex I.

QUINONE REDUCTION

Quinone reduction and proton translocation by complex I areless well understood than NADH oxidation and intramolecularelectron transfer; they both involve the enzyme’s membranedomain, for which there is currently no high-resolutionstructural model and only limited mechanistic information. Thechallenges for mechanistic studies are the lack of cofactors (fewspectroscopic opportunities), the lack of independent catalyticactivity and the experimental difficulties associated with thehighly hydrophobic quinone substrate (typically Q-10), and withcontrolling and quantifying an electrochemical potential acrossthe enzyme in a membrane. At present, extant data pertains mostlyto identification of the quinone-binding site (or sites), and toobservation and characterization of SQ intermediates.

A putative binding site for the quinone headgroup was identifiedin the structure of the hydrophilic domain of complex I from T.thermophilus, at the interface of the 49 kDa and PSST subunits;close to 4Fe[PS]/N2 and the membrane interface (see Figures 2and 8) [8]. The site matches that proposed previously on the basisof mutations to the 49 kDa subunit in R. capsulatus [51], and site-directed mutants in the 49 kDa and PSST subunits of complexI from Yarrowia lipolytica have now been used to characterizeit further [52]. A number of mutations affected NADH:DQoxidoreduction and inhibitor binding [52–55], consistent withthe proposed location of the binding site (see Figure 8).However, there are additional considerations. First, the struct-urally characterized hydrophilic domain is cleaved from thehydrophobic domain, with the bottom of the putative Q-bindingsite and helices H1 and H2 in the cleavage plane, and there aresignificant stretches of adjacent sequence which are not definedstructurally (in particular, the N-terminus of the 49 kDa subunit,and in PSST). Secondly, the decreased activities of the mutants donot necessarily mean that quinone binding or reduction is affected(alternatively, proton transfer, redox-coupled protonation or thesecondary structure may be disrupted). Indeed, mutations of tworesidues in PSST which are distant from the putative bindingsite also decreased the activity significantly (see Figure 8) [54].Nevertheless, residues in and around the putative quinone-bindingsite certainly do influence catalysis, showing that this region ofthe enzyme is important functionally [52].

Many diverse hydrophobic compounds inhibit complexI [57] and, because they inhibit NADH:quinone (but notNADH:ferricyanide) oxidoreduction they are termed ‘Q-siteinhibitors’. Some of them, notably piericidin A, clearly resemblethe quinone substrate, others, notably the acetogenins, display noapparent similarity. Q-site inhibitors may compete directly withquinone binding, but they may also bind elsewhere and trap theenzyme in a state in which the Q-site is occluded, or they mayinhibit a different step in the catalytic cycle (for example, protontranslocation). Radiolabelling experiments have localized boundinhibitors to several subunits, including ND1, ND5, PSST andthe 49 kDa subunit (see [14] and references therein). Importantly,recent observations that a quinazoline inhibitor binds both to ND1

Figure 8 Comparison of the structures of oxidized and reduced (nucleotide-bound) structures of the hydrophilic domain of T. thermophilus complex I

Oxidized structure: white [8]; reduced structure: wheat [25]. Conformational changes uponreduction, identified by Berrisford and Sazanov [25], have been highlighted in green (helicesH1 and H2 on the right, four helix bundle in 49 kDa subunit on left). The lower panel is rotatedby 90◦ relative to the top panel, and is a view from the membrane. Residues which have beenmutated in the 49 kDa and PSST subunits of Y. lipolytica complex I, and which cause at least75 % loss in activity without abolishing the N2 EPR signal, are in red. In Y. lipolytica PSST, theresidues are Val88 (=T. thermophilus Ile48) [52], Asp99 (not resolved in structure) [54], Asp115

(=Asp76) [54], and Asp136 (=Glu97) [54]. In Y. lipolytica 49 kDa, the residues are His91 (notresolved in structure) [55], Ala94 (=Thr37) [52], His95 (=His38) [52,55], Val97 (=Val40) [52],Leu98 (=Leu41) [52], Arg99 (=Arg42) [52], Asp143 (=Asp86) [53], Tyr144 (=Tyr87) [52,53],Tyr145 (=Leu88) [52], Met188 (=Val131) [52], Ser192 (=Thr135) [52], Lys407 (=Arg350) [52],Leu459 (=Pro402) [52], Phe461 (=Met404) [52] and Glu463 (=Asp406) [52,53]. Mutations ofthree additional residues {Y. lipolytica 49 kDa Arg141 (=Arg84) [53,55], Val460 (=Val403) [53]and Arg466 (=Arg409) [55]} caused substantial loss of activity but abolished N2. The putativeredox-Bohr group, Y. lipolytica 49 kDa His226 (=His169) [56], is shown in blue. �PSST and�49kDa indicate the stretches of sequence (26 and 17 residues in T. thermophilus respectively)which are not resolved structurally.

and to the N-terminal section of the 49 kDa subunit underline thefunctional significance of this region of the enzyme [14].

SQ species have been detected during steady-state catalysisby complex I in tightly coupled submitochondrial particles,and differentiated by their relaxation rates and responses tomembrane potential, pH and inhibitors [43,58,59]. SQNf is thefastest relaxing, and it is sensitive to uncouplers (which dissipate�p), and to rotenone and piericidin A. SQNf is proposed to interact,by both spin–spin and dipolar interactions, with 4Fe[PS]/N2; it isconsidered to be ∼12 Å from the cluster, with the vector between



Figure 9 Proton translocation mechanisms based on the oxidation and reduction of quinone species

(A) Mitchell’s original Q-cycle mechanism applied to complex I (left-hand side) and a second enzyme that reoxidizes QH2 to Q at the positive side of the membrane (right-hand panel). (B) Complex I(left-hand side) catalyses the ‘reductant-induced oxidation’ mechanism [67]. (C) Complex I catalyses by redox cycling between a SQ and a QH2, switching between different sides of the membrane,coupled to the separate reduction of a substrate quinone. (D) The redox-cycling SQ and QH2 are retained in the same site, and proton access is switched between different sides of the membrane[69]. See the text for details.

the two species close to the membrane plane, so that SQNf is ∼5 Åcloser to the membrane [43,58]. The proposed position of SQNf isconsistent with the putative quinone-binding site described above.SQNs is insensitive to uncouplers and slowly relaxing (thus it isconsidered distant from the clusters), and it has been characterizedthermodynamically [59]. At pH 7.8, the reduction potentials ofSQNs are EQ/SQ = −0.045 V and ESQ/QH2 =−0.063 V, so SQNs

is thermodynamically stable; because EQ/QH2 = −0.054 V (lowerthan for free quinone), QNs is bound more tightly to complex I thanQNsH2. There is no proposed location for SQNs, although ND4 inE. coli complex I is a possibility because it was labelled by anazido-ubiquinone derivative [60] and is probably located at thedistal end of the membrane arm. Finally, a third species, SQNx,which is also slowly relaxing, was described previously, but isnow not considered to originate in complex I [59].

PROTON TRANSFER HYPOTHESES

First, the question of whether some complexes I transport Na+,instead of protons, across energy-transducing membranes hasobvious implications for the mechanism of charge transfer.Steuber’s group co-reconstituted their preparation of complexI from Klebsiella pneumoniae with Na+-translocating ATPsynthase from Ilyobacter tartaricus and demonstrated bothNADH-supported ATP synthesis and ATP-supported NAD+

reduction [61]. It has since been claimed that Steuber’s resultsstem from the presence of an unrelated Na+-translocating enzyme(Na-NQR) [62], but this claim has not yet been either provenor disproven. Steuber’s group have proposed that the complexesI from E. coli and Y. lipolytica translocate Na+ also [63,64],but in both cases the proposal has been attacked vigorously,using well-characterized and highly pure enzymes [65,66]. ANa+-translocating prokaryotic complex I would be reasonable,as some bacteria rely on Na+-motive forces to support, forexample, ATP synthesis and the flagellar motor, but a Na+-translocating mitochondrial complex I seems extremely unlikelyand this proposal is not generally considered viable. Thereforeonly proton translocation is considered for the rest of the presentreview.

There are several classes of energy-transduction mechanismswhich should be considered for complex I. First, in a direct-coupling mechanism (exemplified, in the respiratory chain, bycytochrome c oxidase), proton translocation is initiated at thesame active site as the electron transfer event that drives it.Secondly, in an indirect-coupling mechanism (exemplified, in therespiratory chain, by ATP synthase), a significant conformationalchange is used to transfer the potential energy of the drivingreaction to a remote site, where proton translocation is initiated.Thirdly, in Q-cycle mechanisms (exemplified, in the respiratorychain, by cytochrome bc1), quinol is used as a mobile electron–proton carrier, to transport protons across the membrane dielectric.Although the three classes appear clearly distinguishable, inreality proposed mechanisms for complex I often fall betweenclasses, or include components from more than one class. First,we consider possible relationships between complex I catalysisand quinone reduction.

Figure 9(A) depicts how Mitchell’s original Q-cyclemechanism applies to complex I: Q is reduced at the negative(matrix) side of the membrane by two electrons (from NADH)and two protons to form QH2. A second enzyme reoxidizes theQH2 at the positive side of the membrane, releasing two pro-tons (to the intermembrane space). The system transports twoprotons and two electrons across the membrane (complex I doesnot ‘pump’ any protons) and it is conserved as a component ofthe following three mechanisms. Figure 9(B) depicts complex Icatalysing by the ‘reductant-induced oxidation’ mechanism, thatmirrors the mechanism of cytochrome bc1 [67]. Q is reduced, at thenegative side, by one electron from NADH and one electron froma QH2 (first half-cycle) or a SQ (second half-cycle) bound at thepositive side. Mechanism B predicts that each NADH oxidationresults in the oxidation of one QH2 to Q (positive side), and thereduction of two Q to QH2 (negative side), but when complex Iwas provided with NADH and two distinguishable quinone/quinolspecies, QAH2 and QB, no evidence for catalytic QA formation wasobserved [68]. The lack of cofactors in the membrane domain, andthe proton-to-electron stoichiometry, present further obstacles tomechanism B, although it cannot be discounted unambiguously.Mechanism B transports four protons and two electrons across the


336 J. Hirst

membrane; complex I pumps two protons per NADH. Figure 9(C)shows a modified version of mechanism B: at the negative sidea SQ is reduced to QH2 by one electron and two protons, andboth the SQ and QH2 are retained; the QH2 is shuttled acrossthe membrane and reoxidized at the positive side, by transfer of asingle electron to a ‘substrate quinone’ and release of two protons.The SQ then returns to the negative side to repeat the half-cycleand fully reduce the substrate quinone. Mechanism C transportssix protons and two electrons across the membrane; complex Ipumps four protons per NADH and so mechanism C accounts forthe experimentally observed stoichiometry. However, it requiresan intramolecular shuttle for SQ and QH2, and it still requires ele-ctron transfer across the membrane. Figure 9(D) presents a similarmechanism, but now the SQ and QH2 intermediates are retainedin the same site. Instead, proton access is switched between thenegative and positive sides: SQ is always protonated from thenegative side, QH2 is always deprotonated to the positive side.Mechanism D also accounts for the experimentally observedstoichiometry of complex I, it does not require electron transferacross the membrane, and it comprises two SQ species, consistentwith extant data. Mechanism D was described recently byOhnishi and Salerno [69], and is related to a previous mechanismsuggested by Dutton et al. [67]. However, it requires an unknowngating mechanism to impose directionality on the proton transferevents.

A second possible direct-coupling site in complex I is4Fe[PS]/N2, because its reduction is coupled to proton uptake.The coupled-protonation has been proposed to occur on a nearbyhistidine residue: in Y. lipolytica, the 49 kDa subunit H226Mmutant retained its ability to transport protons, but the reductionpotential of 4Fe[PS]/N2 became independent of pH (over the mea-sured range, pH 6–8) [56]: these results suggest that theredox-coupled protonation of 4Fe[PS]/N2 is not important inenergy transduction. Alternatively, Friedrich and co-workers[70] proposed that two tyrosine residues are protonated upon4Fe[PS]/N2 reduction, and that a glutamate residue is protonatedupon 4Fe[PS]/N2 oxidation, but these observations could notbe reproduced by Rich and co-workers [71]. Taken together,these observations do not form a strong case for a redox-linkedprotonation of 4Fe[PS]/N2 as the basis for proton translocationby complex I.

Recently, Berrisford and Sazanov [25] described a highlyspeculative redox-linked proton transfer mechanism for complexI, based on apparent changes in the electron density, and thusthe ligation, of 4Fe[PS]/N2 in three states of the enzyme. Theysuggest that, although the oxidized cluster (State I) is ligatedby four cysteine residues (Cys45, Cys46, Cys111 and Cys140 inthe T. thermophilus PSST subunit), the reduced cluster is onlyligated by three. Furthermore, when ‘cluster N6b’ (4Fe[TY]2) isoxidized, Cys45 is dissociated and protonated (State II), but when4Fe[TY]2 is reduced, Cys46 is dissociated and protonated (StateIII). The changes in electron density are certainly intriguing, but itis difficult to confidently assign the oxidation states of 4Fe[PS]/N2and 4Fe[TY]2 in the crystals: typically, 4Fe[PS]/N2 is the highestpotential cluster in complex, but its potential in T. thermophilushas not been defined unambiguously and may be lower than usual[41,72], and the oxidation state of 4Fe[TY]2 has been inferredusing EPR signal N4 (although N4 may originate from 4Fe[TY]1,and is either not observed or atypical in T. thermophilus) [41,72].However, the mechanism begins with the clusters oxidized(compare with Figure 7B), in State I. NADH is oxidized and4Fe[PS]/N2 is reduced (State I→II), and then 4Fe[TY]2 isreduced also (State II→III). One electron is transferred to boundquinone, and 4Fe[TY]2 is reoxidized (State III→II), and thenthe second electron is transferred and 4Fe[PS]/N2 is reoxidized

(State II→I). Each state change results in the dissociation andprotonation, or deprotonation and re-association, of one the cys-teine ligands of 4Fe[PS]/N2, but, incredibly, none of the ligationchanges are suggested to be involved directly in proton pumping!Instead, Berrisford and Sazanov [25] propose that all fourprotons are ‘pumped’ during quinol formation, one protonbeing transferred (from a protonated residue ‘loaded’ by Cys45

deprotonation) to the periplasm by an unknown conformationalchange, and three protons being pumped in subunits ND2,ND4 and ND5 by further unknown conformational changes (seebelow). The conformational changes are coupled to 4Fe[PS]/N2and 4Fe[TY]2 reduction, and they originate in helices H1 andH2, and in a four-helix bundle in the 49 kDa subunit respectively,but the observed changes are small (see Figure 8). In addition,there are no well-defined examples of [4Fe–4S] clusters withcysteine ligands that dissociate and protonate upon reduction,and the suggested mechanism relies heavily on the pH-dependentreduction potential of 4Fe[PS]/N2 (it is difficult to reconcilewith the proton-pumping activity of Y. lipolytica H226M; [56]).However, further work is required to better define the pKOx andpHRed values of 4Fe[PS]/N2. Currently, the best-defined valuesare from Y. lipolytica (pKOx = 5.7, pHRed = 7.3) [56], but theyprovide a �pK value of less than 2; although the values are fromequilibrium redox titrations, the separation is not sufficient for astrongly coupled proton pump.

So, what is the evidence for a significant conformational changeduring complex I catalysis? First, two different large changesin structure have been observed by electron microscopy: the‘horseshoe’ conformation [73] and an ‘expansion’ over the wholestructure upon enzyme reduction [74]. Both observations are nowconsidered artefacts [74,75]. Furthermore, electron microscopyof the oxidized and reduced E. coli complexes I in ice (ratherthan negative stain) did not reveal any significant conformationalchange [75]. Secondly, several reports have described changes inthe cross-linking pattern, particularly between the subunits of thehydrophilic arm of complex I, upon nucleotide binding and/orreduction, and they have been taken to represent conformationalchanges, or changes in the relative positions of the subunits(see, for example [74,76]). However, there are only minimaldifferences between the structures of the oxidized and reducedstates of the hydrophilic arm of T. thermophilus complex I [25](see Figure 8). Fourier-transform IR spectroscopy studies alsosuggested that oxidized and NADH-reduced complex I do notsignificantly differ structurally (the observed differences werefully consistent with the oxidation and reduction of a set ofsimple FeS proteins) [71]. Thirdly, and finally, ND4 and ND5 arelocated at the distal end of the membrane arm [18], and theyare proposed to be important in proton translocation, becausethey are homologous with Na+/H+ antiporters [16], complex Iis inhibited by several amiloride derivatives (known antiporterinhibitors [77]), and a photoaffinity analogue of the inhibitorfenpyroximate has been located on ND5 [78] (although thisresult has been challenged recently [14]). Coupling a redoxreaction in the hydrophilic arm to proton translocation at the distalend of the hydrophobic arm would indeed require an unusualmechanism of long-range energy transfer. Such a mechanismcould be based on a ‘conformational change’. Alternatively,Verkhovskaya and co-workers [79] suggested an ‘electrostatictransmission’ mechanism, mediated by conserved charges in themembrane domain. Whether such a mechanism could drive threeprotons across the membrane from ∼100 Å away, remains to beestablished. Finally, a number of studies have mutated conservedresidues in the membrane subunits of bacterial complexes I, andobserved changes in Vmax and Km(Q) (see [79,80] and referencestherein), but the data are difficult to interpret in the absence of



high-resolution structural information. The residues identifiedmay comprise proton channels or parts of the energy-couplingmechanism, or they may simply exert secondary effects on theenzyme structure or stability.

CONCLUSIONS AND PERSPECTIVES

For many years, complex I was the poor relation of therespiratory chain enzymes, the refractory ‘black box’ whichresisted structural or mechanistic characterization, the enzymewith little new to stir the imagination. Complex I, indeed, isan enzyme more challenging than most: difficult to purify andcrystallize, lacking good spectroscopic handles, and comprisingan unwieldy gamut of subunits and cofactors: but, finally, complexI is coming to heel. The first structural model for the hydrophilicdomain of the enzyme has provided a solid starting point formechanistic proposals which integrate structure with functionaland spectroscopic data, and a further leap forward is to beanticipated when the structure of the hydrophobic domain, orof an intact complex I, is finally determined. However, structureis only part of the story (‘a picture of a racehorse does not tellyou how it runs’) and solving the mechanism of a complicatedredox enzyme such as complex I will certainly require innovativeand advanced biophysical and biochemical approaches. Indeed,this is an exciting time for complex I – not only a challengingenzyme full of opportunities, but also an enzyme which may playan important role in many human diseases.

The present review has aimed to provide a balanced overviewof experimental evidence pertaining to the mechanism ofcomplex I, and a discussion of contemporary mechanistichypotheses, proposals and models. Finally, current knowledgeof the mechanism of complex I is summarized as follows. (i)NADH is oxidized by the FMN in a hydride transfer reaction,with the nicotinamide ring juxtaposed on the isoalloxazinering system in a configuration common to many flavoenzymes.NADH:flavin oxidoreduction is fast (several thousand per second)and thermodynamically reversible; NAD+ dissociation may berate-limiting in NADH oxidation. (ii) The reduced flavin reactswith O2 in a slow, bimolecular reaction, to generate ROS. Thepopulation of reduced flavin is determined by the concentrationsof NADH and NAD+ (they set the potential of the flavin andblock O2 access by binding to the active site). ROS productionmay be accelerated by conveyor molecules that induce redox-cycling reactions. (iii) Complex I contains eight iron–sulfurclusters. Seven of them form a chain between the flavin andthe quinone-binding site. One of them, the distal cluster, is onthe opposite side of the flavin. The ‘distal’ cluster has beensuggested to act as a temporary electron storage device, tominimize the lifetime of the flavosemiquinone; the distal clusterhas no apparent role in energy transduction. (iv) The seven clustersof the chain are spaced less than 14 Å apart, consistent withfast electron transfer along the chain. Five reduced clusters havebeen observed by EPR in mitochondrial complex I, and theirreduction potentials measured. Controversy remains about howto assign the spectroscopic data to the structurally defined clu-sters, but, in the model most consistent with extant data, thecluster arrangement produces an alternating or ‘roller-coaster’free-energy profile. The chain is pre-loaded with four electronsduring turnover, increasing the apparent rate of electron transferand aiding efficient energy conservation. (v) NADH oxidation andintramolecular electron transfer are both fast and reversible, andmuch faster than quinone reduction. Proton translocation is mostprobably coupled to quinone binding, quinol release or quinonereduction by the terminal iron–sulfur cluster. A possible quinone-

binding site has been identified near to the terminal cluster, and SQintermediates have been observed by EPR. (vi) The mechanismsof quinone reduction and proton translocation are not known;they are the subject of much speculation, but little data. Proton-coupled reactions at the terminal iron–sulfur cluster, quinoneredox-cycling mechanisms and conformational changes are allpossibilities that are currently under discussion.

FUNDING

Work in my laboratory is funded by the Medical Research Council.

REFERENCES

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

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79 Euro, L., Belevich, G., Verkhovsky, M. I., Wikstrom, M. and Verkhovskaya, M. (2008)Conserved lysine residues of the membrane subunit NuoM are involved in energyconversion by the proton-pumping NADH:ubiquinone oxidoreductase (complex I).Biochim Biophys Acta 1777, 1166–1172

80 Kao, M.-C., Nakamaru-Ogiso, E., Matsuno-Yagi, A. and Yagi, T. (2005) Characterizationof the membrane domain NuoK (ND4L) NADH-quinone oxidoreductase from Escherichiacoli. Biochemistry 44, 9545–9554

Received 3 September 2009; accepted 30 September 2009Published on the Internet 23 December 2009, doi:10.1042/BJ20091382


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