A giant molecular proton pump: structure and mechanism of respiratory complex I.

REVIEWS

A giant molecular proton pump: structure and mechanism of respiratory complex I Leonid A. Sazanov

Abstract | The mitochondrial respiratory chain, also known as the electron transport chain (ETC), is crucial to life, and energy production in the form of ATP is the main mitochondrial function. Three proton-translocating enzymes of the ETC, namely complexes I, III and IV, generate proton motive force, which in turn drives ATP synthase (complex V). The atomic structures and basic mechanisms of most respiratory complexes have previously been established, with the exception of complex I, the largest complex in the ETC. Recently, the crystal structure of the entire complex I was solved using a bacterial enzyme. The structure provided novel insights into the core architecture of the complex, the electron transfer and proton translocation pathways, as well as the mechanism that couples these two processes. Chemiosmotic coupling A process that links the electron transport chain to ATP synthesis.

Midpoint redox potential (Em). A measure of the tendency of a chemical species to acquire electrons and thereby be reduced. The species with large positive potential have high affinity for electrons and vice versa. Em denotes the potential at which the compound is half oxidized and half reduced.

Institute of Science and Technology Austria, 3400 Klosterneuburg, Austria. e‑mail: [email protected] doi:10.1038/nrm3997 Published online 20 May 2015

Most eukaryotic cells contain mitochondria, the ‘power plants’ that are thought to be the remnants of an ancient endosymbiotic event 1. Mitochondrial respiratory enzymes represent more elaborate versions of their bacterial counterparts, and energy production in the form of ATP is the main mitochondrial function in addition to many other roles, such as signalling and cell death. Although ATP is consumed throughout the cell, it is primarily synthesized in the mitochondrial matrix by oxidative phosphorylation. Electrons harvested from the catabolic processes of glycolysis, fatty acid oxidation and the tricarboxylic acid (TCA) cycle enter the electron transport chain (ETC) on the inner mitochondrial membranes. Electron transfer through the ETC is coupled to proton translocation out of the mitochondrial matrix. Energy is transduced via chemiosmotic coupling 2, whereby the electrochemical gradient of protons (proton motive force) across the membrane drives F1FO-ATP synthase3,4. Most enzymes of the ETC are large multi-subunit protein assemblies (complexes I–IV) containing many redox cofactors, with complex I being the largest and most elaborate. This complexity has made it challenging to acquire a mechanistic understanding of the ETC. Moreover, even though the basic functional principles of most components of the ETC have been elucidated, the details are still being hotly debated. We now know that each complex in the chain functions by a unique mechanism and that there are no direct analogues with other enzymes.

The first structure of a component of the ETC to be determined was that of the F1-ATP synthase in 1993 (REF. 3), followed later by that of complex IV5,6, complex III7 and complex II8. Recently, the crystal structure of the entire complex I (from Thermus thermophilus) was solved9, providing many insights into its organization and mechanism. In this Review, I first provide an overview of the mitochondrial respiratory chain and the multiple proton-pumping enzymes involved. Second, I discuss the structural insights that have been gained from the crystal structure of the T. thermophilus complex I. Last, I review the most recent views on the electron transfer and proton translocation pathways and the possible mechanism that couples the two processes.

The mitochondrial respiratory chain The mammalian mitochondrial ETC includes protonpumping enzymes known as complex I (NADH– ubiquinone oxidoreductase), complex III (cytochrome bc1) and complex IV (cytochrome c oxidase) (FIG. 1). They contain multiple redox cofactors to facilitate intra-protein electron transfer, whereas electron transport between complexes is mediated by membrane-embedded ubiquinone and soluble cytochrome c, which are mobile carriers. Free energy is released at each step along the chain, as the redox potentials of electron donors and acceptors gradually increase. Complex I is the entry point for lowpotential (‘high-energy’) electrons from NADH (with a midpoint redox potential (Em) at pH 7 (Em,7) of –320 mV),

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REVIEWS Complex I NADH–ubiquinone oxidoreductase

Complex II Succinate–quinone oxidoreductase

Complex III Cytochrome bc1 complex

Complex IV Cytochrome c oxidase

Complex V F1FO-ATP synthase Mitochondrial matrix

NADH 6 H+

2 H+

NAD+ + H+ Succinate

4 H+

Fumarate + 2H+ ½O2

– Δψ

QH2

QH2

2QH2

QH2

Q

Q

2Q

Q

H2O

ATP ADP + Pi

Inner mitochondrial membrane

+ 4 H+

4 H+

Cytochrome c 2e–

2 H+

2.7 H+ IMS

Figure 1 | The electron transport chain. The mammalian mitochondrial electron transport chain (ETC) includes the proton-pumping enzymes complex I (NADH–ubiquinone oxidoreductase), complex III (cytochrome bc1) and complex IV Nature Reviews | Molecular Cell Biology (cytochrome c oxidase), which generate proton motive force that in turn drives F1FO-ATP synthase. Electron transport between complexes is mediated by membrane-embedded ubiquinone (Q) and soluble cytochrome c. Complex I is the entry point for electrons from NADH, which are used to reduce Q to ubiquinol (QH2). QH2 is subsequently used by complex III to reduce cytochrome c in the intermembrane space (IMS), and complex IV uses cytochrome c to reduce molecular oxygen, which is the ultimate electron acceptor. For each NADH molecule oxidized, 10 protons are translocated across the membrane from the matrix to the IMS. Complex II (succinate–quinone oxidoreductase) provides an additional entry point for electrons into the chain. The structure of each respiratory complex is presented: complex I from Thermus thermophilus (protein databank (PDB) identifier 4HEA)9, complex II from Sus scrofa (PDB identifier 1ZOY)8, complex III from Bos taurus (PDB identifier 1BGY)7 and complex IV from B. taurus (PDB identifier 1OCC)6. The structure of F1FO-ATP synthase was generated by merging crystal structures of subcomplexes from the B. taurus enzyme within an 18 Å resolution cryoelectron microscopy map89. The FO domain of ATP synthase has not been resolved in its entirety and therefore some subunits are not shown. ΔΨ, membrane potential. The PDB file for the ATP synthase was provided by J. E. Walker, and the ETC image was prepared by G. Minhas, Medical Research Council, Mitochondrial Biology Unit, Cambridge, UK.

which are used to reduce ubiquinone (Em,7 = +100 mV) to ubiquinol. Ubiquinol is subsequently used by complex III to reduce cytochrome c (Em,7 = +260 mV) in the intermembrane space (IMS), and complex IV uses cytochrome c to reduce molecular oxygen, the ultimate electron acceptor (Em,7 = +820 mV), to water (FIG. 1). The reactions catalysed by the complexes can be summarized as follows: Complex I: NADH + H+ + Q + 4H+in → NAD+ + QH2 + 4H+out

Complex III: QH2 + 2 cyt c 3+ + 2H+in → Q + 2 cyt c 2+ + 4H+out

Complex IV: O2 + 4 cyt c 2+ + 8H+in → 2 H2O + 4 cyt c 3+ + 4H+out

(in which Q denotes ubiquinone and QH 2 ubiquinol, cyt c denotes cytochrome c, and ‘in’ denotes the mitochondrial matrix and ‘out’ the IMS). Overall, for each NADH molecule oxidized, the combined action of these three complexes leads to the translocation of 10 protons across the membrane from the matrix to the IMS. Additional entry points into the chain for less ‘energetic’ electrons (~0 mV) are provided by complex II (succinate–quinone oxidoreductase) and other ubiquinone-reducing enzymes, such as electron transfer flavoprotein–ubiqionone oxidoreductase

(ETF–QO), glycerol‑3‑phosphate dehydrogenase (GPDH) and dihydroorotate dehydrogenase. Although none of these proteins pumps protons, any ubiquinol produced then enters the ETC at complex III. Complex II also catalyses a key step in the TCA cycle, so that the rate of succinate-to-fumarate conversion is controlled by the ubiquinol/ubiquinone ratio in the membrane, providing a feedback mechanism between the TCA cycle and oxidative phosphorylation8. Another feedback mechanism, linked directly to the proton motive force, may be provided by nicotinamide nucleotide transhydro genase, which catalyses hydride transfer from NADH to NADP+ coupled to inward proton translocation10. Apart from providing reducing equivalents that are required to mitigate oxidative damage, this enzyme may also regulate TCA cycle activity at the level of NAD- and NADP‑linked isocitrate dehydrogenases11,12. The coupling between electron transfer and proton translocation may be direct (that is, involving chemical redox reaction intermediates that are protonated or de‑protonated, resulting in net proton translocation) or indirect (that is, involving long-range conformational changes). Complexes III and IV use direct coupling, which is mediated by membrane-embedded cofactors (haems and metal centres) (BOX 1). By contrast, ATP synthase does not contain any cofactors in the membrane,

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REVIEWS Box 1 | The mechanisms of respiratory complexes III and IV Mitchell83 proposed a Q‑cycle involving the quinone–quinol (Q–QH2) shuttle as a means for proton translocation across the membrane. This principle is realized in complex III (cytochrome bc1 complex): two turns of the cycle result in the release of four protons into the intermembrane space (IMS) and consumption of two protons from the matrix side47 (see the figure, part a). Electron transfer in the first step of the Q‑cycle is shown by solid arrows; dashed arrows indicate the same steps with a second ubiquinol. As ubiquinol is oxidized at the QO site, one electron is transferred along the high-potential chain to the 2Fe–2S centre on the Rieske protein, then to cytochrome c1 and finally onto the soluble carrier cytochrome c. The second electron is transferred along the low-potential chain to ubiquinone at the Qi site via two b‑type haems, leading to the formation of a ubisemiquinone radical (Q•–). The steps are repeated with a second ubiquinol; a second cytochrome c protein is reduced, and a further electron reduces ubisemiquinone to ubiquinol. The oxidation of two ubiquinol molecules at the QO site releases four protons into the IMS. Two protons are taken up from the matrix as ubiquinol at the Qi site is reduced. In complex IV (cytochrome c oxidase), four electrons delivered by cytochrome c are used in the catalytic cycle of the haem a3–CuB binuclear centre to reduce an oxygen molecule to two water molecules5,6 (see the figure, part b). CuA accepts electrons from cytochrome c one at a time (black arrows). The electrons are subsequently transferred to haem a, and on to the haem a3–CuB binuclear centre, where an oxygen molecule is bound. To reduce oxygen into water, four ‘chemical’ protons are taken from the matrix side (grey arrow). In addition, four protons are pumped across the membrane into the IMS (dashed arrows), so that in total eight protons are removed from the matrix. How exactly this coupling is achieved is still being debated; however, it is thought that delivery of each electron to the binuclear centre is accompanied by uptake of one substrate proton and translocation of one vectorial proton via a charge-compensation mechanism involving a key conserved Glu residue near the binuclear centre84.

a

b 2H+ 2H+

1e– Cytochrome c1

IMS

2Fe–2S 1e–

2QH2 2Q

QO site

4e– CuA 4e–

4e–

O2 CuB

1e– 1e–

Membrane

4H+

Cytochrome c

Haem a3

Haem bL Haem a

Q•– or Q2– QH2

1e–

Haem bH

2H2O

Qi site Complex III

Mitochondrial matrix

2H+

Complex IV 4H+

4H+

Nature Reviews Cell Biology and proton translocation back| Molecular into the matrix drives rotation of the ring of membrane-embedded subunits, resulting in conformational changes in the catalytic hydrophilic F1 domain3,4. It was shown early on that complex I also does not contain redox centres in the membrane domain and, in contrast to what occurs in complexes III and IV, electron transfer and proton translocation pathways are spatially separated. This, together with observed changes in crosslinking patterns upon reduction13–15 and the distal location of antiporter-like subunits16–19, led to the proposal that complex I might

function via conformational changes. Now that the structure of complex I has been solved, the details of the mechanism can be analysed.

The structure of complex I The overall architecture of complex I (TABLE 1) is described below, along with the description of its two main domains. Overall architecture. Bacterial complex I represents the minimal version of the enzyme, with 14 strictly conserved core subunits that are necessary and sufficient for function. Subunits are shared equally between the peripheral and membrane arms, which together form an L‑shaped molecule, as observed by single-particle electron microscopy (EM) for both bacterial and mitochondrial enzymes20–23. The peripheral arm comprises the NADH-oxidizing dehydrogenase module (N‑module), which provides electron input into the chain of Fe–S clusters, and the connecting Q‑module, which conducts electrons to the quinone-binding site. The membrane arm (also known as the membrane domain) comprises the proton-translocating P-module24 and subunit NuoH (in Escherichia coli; known as Nqo8 in Thermus spp.). NuoH is unrelated to other known proteins and so does not belong to any evolutionary module; it forms most of the interface to the peripheral arm (BOX 2). With the exception of this junction, to which quinone binds, the two arms of complex I are functionally and evolutionarily independent: the peripheral arm catalyses oxidation–reduction reactions, and the membrane arm catalyses proton transport. During the course of evolution, mitochondrial complex I has acquired ~30 supernumerary (or accessory) subunits in addition to the core subunits that are present in the bacterial enzyme25,26. This increases the total molecular weight of complex I by almost twofold, from ~550 kDa in bacteria to ~1 MDa in mitochondria. Except for the 42 kDa and 39 kDa subunits (bovine nomenclature), most supernumerary subunits are small (~10–20 kDa), and about 12 are predicted to contain a single transmembrane helix (TMH)27,28. As the core subunits coordinate all cofactors and are sufficient for function, the role of the supernumerary subunits is not clear. They are likely to assist in the assembly, regulation and stability of the complex, similarly to the supernumerary subunits in complex IV6. Structural information on mitochondrial complex I is currently limited, with no full atomic structures available. The electron density map of complex I from the fungus Yarrowia lipolytica was initially obtained at 6.3 Å resolution29. The fit of the T. thermophilus structure onto this density map confirmed strong preservation of the core structure during evolution30. The presence of additional electron density indicated that the supernumerary subunits form a shell around the core29,30. Recently, a cryo-EM map of bovine complex I at 5 Å resolution was published27, with about 14 supernumerary subunits identified on the basis of structural homology. In agreement with previous reports, core subunits were found



REVIEWS Table 1 | Nomenclature* of the core subunits of complex I Module

Bos taurus

Homo sapiens

Escherichia coli (Rhodobacter capsulatus)

Thermus thermophilus (Paracoccus denitrificans, Aquifex aeolicus)

Cofactors and comments‡

75 kDa

NDUFS1

NuoG

Nqo3

• N1b (2Fe[75]) • N4 (4Fe[75]C) • N5 (4Fe[75]H) • (N7)§

51 kDa

NDUFV1

NuoF

Nqo1

• FMN • N3 (4Fe[51])

24 kDa

NDUFV2

NuoE

Nqo2

N1a (2Fe[24])

49 kDa

NDUFS2

NuoD (NuoCD||)

Nqo4

No cofactor

30 kDa

NDUFS3

NuoC

Nqo5

No cofactor

TYKY

NDUFS8

NuoI

Nqo9

• N6a (4Fe[TY]1) • N6b (4Fe[TY]2)

PSST

NDUFS7

NuoB

Nqo6

N2 (4Fe[PS])

–

ND1

ND1

NuoH

Nqo8

8–9 TMH

P-module

ND2

ND2

NuoN

Nqo14

14 TMH (antiporter-like)

ND3

ND3

NuoA

Nqo7

3 TMH

ND4

ND4

NuoM

Nqo13


ND4L

ND4L

NuoK

Nqo11

3 TMH

ND5

ND5

NuoL

Nqo12


ND6

ND6

NuoJ

Nqo10

5 TMH

Peripheral arm Dehydrogenase (N)-module

Connecting (Q)-module

Membrane arm

FMN, flavin mononucleotide; TMH, transmembrane helix. *Nuo nomenclature originates from NADH–ubiquinone oxidoreductase, Nqo nomenclature originates from NADH–quinone oxidoreductase, and ND nomenclature originates from NADH dehydrogenase. ‡Cofactors (FMN and Fe–S clusters) coordinated by each subunit are listed for the peripheral arm; comments apply to the membrane arm. The traditional nomenclature for Fe–S clusters (Nx, derived from initially described electron paramagnetic resonance (EPR) signatures38), as well as the nomenclature proposed recently51 on the basis of re‑assignment of EPR signals to structurally observed clusters, is shown. In the new nomenclature, clusters are named according to their nuclearity (2Fe or 4Fe), their subunit location (using the bovine nomenclature) and, when necessary, as ligated by four Cys (C) or three Cys and one His (H). §Cluster N7 is present only in some bacteria (for example, E. coli and T. thermophilus). ||Subunits NuoC (30 kDa) and NuoD (49 kDa) are fused in E. coli and some other bacteria.

to be similar to the bacterial enzyme. This study also showed that several small supernumerary subunits and the mammal-specific 42 kDa subunit (a member of the nucleoside kinase family) form an additional connection between the peripheral and membrane arms, possibly stabilizing this fragile area. The 39 kDa subunit, which contains tightly bound NADPH and is homologous to short-chain dehydrogenases31, was also found to localize near the junction. Subunit B16.6, which is identical to the apoptosis-inducing factor GRIM‑19 (REF. 26), was shown to form a long α‑helix that embraces the ‘heel’ of complex I. Other supernumerary subunits form a shell mostly around the membrane domain, with almost no extra protein mass around the N‑module of the peri pheral arm, possibly because it is the last to be added during assembly 27. More recently, the resolution of the structural characterization of Y. lipolytica complex I was improved to ~3.8 Å32. About 25% of the total complex was solved at the atomic level, including large parts of the core subunits but excluding the supernumerary subunits. As had been observed for bovine complex I, the core subunits were found to be structurally highly similar to

T. thermophilus, including conservation of key functional residues and features, such as the central hydrophilic axis in the membrane domain. Some of supernumerary subunits were preliminarily identified on the basis of assignments done for the bovine complex 27. Overall, in the membrane domain 18 extra TMHs were found to be distributed around the core subunits, thus extending the total number of TMHs to 82. Apart from all the remarkable similarities to the bacterial structures (and to their earlier interpretation), there are two notable differences: first, it has been suggested that the fourth proton translocation channel takes a different route compared with the bacterial structure9; second, a different conformation of several loops near the quinone-binding site was observed (see below). The first atomic structure of complex I was that of the peripheral arm of the enzyme from T. thermophilus, which was determined by X‑ray crystallography at 3.1 Å resolution33,34. Later, the crystal structure of the membrane arm from E. coli complex I was solved at 3.0 Å resolution35,36. Finally, the crystal structure of the entire complex from T. thermophilus was determined recently at 3.3 Å resolution and is still the only completely solved

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REVIEWS Box 2 | Evolutionary origins and nomenclature of complex I Complex I is a member of a family of membrane-bound oxidoreductases that is related to a class of membrane-bound [NiFe] hydrogenases; the latter couple substrate oxidation and hydrogen reduction to active proton transport85. Homology of subunits in related proteins suggests that complex I originated from the unification of pre-evolved subcomplexes (modules) with distinct functions45,86,87. The modules trace back to two unrelated protein families. Oxidoreductase activity was provided by soluble [NiFe] hydrogenases that gave rise to the NADH-oxidizing dehydrogenase module (N-module) and the connecting Q-module (combined already in NAD+-reducing hydrogenase)45,88, which together constitute the peripheral arm (see the figure part a; TABLE 1). Proton translocation activity was provided by Mrp cation/H+ antiporters, which are homologous to the P-module41 of the membrane arm. The combination of soluble hydrogenase and antiporter was likely to have resulted in the emergence of several known types of membrane-bound hydrogenases, which later evolved into complex I45. A non-canonical, but widely spread, ancestral complex I‑like enzyme comprising 11 subunits, which uses as yet unknown electron donors, seems to lack the N-module86 (not shown). The figure shows the structure of the entire Thermus thermophilus complex I (protein databank identifier 3M9S) and the evolutionary modules. Homology-based architectures of the NAD+-reducing [NiFe] hydrogenase and the Mrp antiporter (the 3D structures of which are not known) are also shown (see the figure, part b). The MrpBEFG subunits are unrelated to complex I.

a Complex I NADH

NAD+

b

NADH α

NAD+

Nqo1 and Nqo2

γ

N-module Nqo3 Nqo6, Nqo5, Nqo4 and Nqo9 H+

Q-module Cytoplasm

H+

H+

2H+

Nqo14, Nqo13 and Nqo12

MrpC

MrpD

Nqo8 Nqo7, Nqo10 and Nqo11

P-module

structure of the whole enzyme9. As the architecture and the sequences of the core subunits — including the key residues involved in the coordination of cofactors for electron transfer or proton translocation — are so well conserved, these structures provide the foundation for understanding the function and mechanism of the human enzyme, as well as the molecular basis for human pathologies associated with mutations in complex I9,34,35,37.

[NiFe] hydrogenases The class of hydrogenases with the most members. [NiFe] hydrogenases catalyse the reversible 2H+ + 2e− ↔ H2 reaction; their core comprises the large subunit hosting the Ni–Fe active site and the small subunit hosting the Fe–S clusters.

MrpA

MrpBEFG

Periplasm

A class of redox cofactors found in molybdenum- and tungsten-containing enzymes, such as nitrate reductase.

Na+

[NiFe] hydrogenase Q QH2

Molybdopterin

H2

δβ

H+

The peripheral arm. The T. thermophilus peripheral arm contains nine subunits: core Nqo1–6, Nqo9 and two additional subunits that are not part of the nqo operon: frataxin-like Nqo15 (REF. 34) and a possible chaperone, Nqo16 (REF. 9) (FIG. 2a). All known cofactors of complex I are found in the peripheral arm: the primary electron acceptor flavin mononucleotide (FMN, found in the distal tip of the domain) and 8–9 Fe–S clusters38. Seven of the clusters form a 95 Å-long redox chain connecting FMN to the quinone-binding site at the interface with the membrane domain (FIG. 2b). FMN is coordinated by subunit Nqo1 at the deep end of a solvent-exposed cavity that also contains the NADH-binding site33,34 (FIG. 2c). FMN is within 14 Å (the maximum distance for physiological electron transfer 39) of both Fe–S cluster N3 (coordinated by Nqo1) and off-path binuclear cluster N1a (coordinated by thioredoxin-like subunit Nqo2). The amino-terminal domain of subunit Nqo3, which is related to [FeFe] hydrogenases, contains the

H+ H+ Mrp antiporter Nature Reviews | Molecular Cell Biology

Fe–S clusters N1b, N4 and N5 from the main redox chain, whereas its large carboxy-terminal domain, which is related to molybdopterin-containing enzymes, coordinates the Fe–S cluster N7. This cluster is too far from the main chain to participate in electron transfer and seems to be an evolutionary relic34,40 that is present only in some bacteria. The ferredoxin-like subunit Nqo9 coordinates the Fe–S clusters N6a and N6b, providing the link to the terminal Fe–S cluster N2. This cluster donates electrons to quinone and is coordinated by subunit Nqo6 at the interface with Nqo4, which are related to the small and large subunits of [NiFe] hydrogenases, respectively. The membrane arm. The membrane arm comprises 7 subunits: Nqo7, Nqo8 and Nqo10–14, which together contain 64 TMHs9,35,36 (BOX 2; FIG. 2a; TABLE 1). Subunits Nqo12, Nqo13 and Nqo14 are termed antiporter-like because they are homologous to each other and to the bacterial cation/H+ Mrp antiporter complex subunits MrpA and MrpD31,41, all of which contain 14 conserved TMHs each. Subunit Nqo12 contains a C‑terminal extension comprising two TMHs that are connected by an unexpected structural element, a 110 Å-long α‑helix (HL) that runs along the cytoplasmic membrane surface, linking the three antiporter-like subunits as a likely coupling element 9,35,36. Another element (βH) is formed from a series of connected β‑hairpins and helices on the opposite (periplasmic) side of the arm35.



REVIEWS In the antiporter-like subunits, 14 common helices can be subdivided into a highly conserved 10‑TMH core (comprising TM 4–13) and the less conserved TM1– TM3 and TM14. In the core, two sets of five helices (TM4–TM8 and TM9–TM13) are related to each other

by symmetry along a pseudo-twofold screw axis parallel to the length of the membrane arm. This is in contrast to known transporters, such as LeuT, in which the key domains are usually related by symmetry along a twofold axis running through the centre of the protein42. Nqo1 FMN

Nqo3 Nqo2 Nqo15

a

b

Nqo5

13.5 (12.3)

Nqo16 N2 Nqo4

Nqo9

N1a

8_AH1 Q

30 Å

Nqo14

Nqo12

Nqo13

N1b 13.9 (10.7) 24.2 (20.5)

N5

7_TM1 8_TM1

Nqo11 Nqo7 Nqo10

N6a

12.2 (9.4)

Decylubiquinone

N6b 14.2 (10.5)

d

c

N2

Tyr87 3.1 Glu97 C4

His38

Gly67 Nqo1 NADH

4.8

Phe70

N7

16.9 (14.0)

180 Å N2

14.2 (11.0)

N4 12.2 (8.5)

Nqo8

Periplasm

10.9 (7.6)

22.3 (19.4) N3

Nqo6

Cytoplasm

NADH

FMN

2.5 Asp139

11.9 (8.6)

Quinone Nqo4

Phe78

N5 FMN Glu185

Lys202 Phe205

Figure 2 | Structure of Thermus thermophilus complex I. a | The Thermus thermophilus complex I contains 14 strictly conserved core subunits (Nqo1– Nqo14), which are necessary and sufficient for function. The subunits are shared equally between the peripheral arm — comprising the NADH-oxidizing dehydrogenase module (N‑module; which provides electron input into the chain of Fe–S clusters) and the connecting Q‑module (which conducts electrons to the quinone-binding site) — and the membrane arm, which comprises the proton-translocating P‑module. The primary electron acceptor flavin mononucleotide (FMN) is shown as magenta spheres, and Fe–S clusters as red and orange spheres; the Fe–S cluster N2 is also indicated. The key helices (7_TM1, 8_TM1 and 8_AH1; in which the prefixes indicate the subunits) around the entrance into the quinone reaction chamber (indicated as Q) and approximate membrane position are also shown. b | Arrangement of redox centres is depicted. The main pathway

of electron transfer is indicated by solid arrows, and a diversion to cluster N1a by a dashed arrow. The distances between the| centres (given Å) were Nature Reviews Molecular CellinBiology calculated both centre-to‑centre and edge‑to‑edge (shown in parentheses). The positions of NADH33 and the quinone9 headgroup are based on experimental data. The entire ubiquinone tail was modelled into the quinone-binding cavity (protein databank (PDB) identifiers 4HEA and 3IAM). c | The NADH-binding site33, viewed from the solvent-exposed side, is shown. FMN and residues involved in NADH binding are shown as stick models, with carbons shown in yellow; the carbons of NADH are shown in pink (PDB identifier 3IAM). Potential interactions with Nqo1 residues are indicated by dashed lines. d | Bound decylubiquinone is shown with experimental electron density9. Nqo4 residues interacting with the headgroup are indicated. Distances for potential polar interactions (in Å) are indicated. Parts a and d from REF. 9, Nature Publishing Group.

380 | JUNE 2015 | VOLUME 16


REVIEWS The symmetry-related helices TM7 and TM12 are interrupted in the middle of the bilayer by an extended loop. Such helices are functionally important for ion transport, introducing flexibility and charge to the protein, which is embedded deep within the membrane42,43. The discontinuous helices are strategically located: the TM7 helix contacts the traverse α‑helix HL (see above), whereas TM12 contacts a key conserved Glu residue from TM5 of the neighbouring antiporter-like sub unit. In addition, the TM8 helix, found in the centre of subunits at the interface of symmetry-related domains, is partly unwound in the middle by a π-bulge44, which is usually found at functional sites in other proteins. An 11-TMH bundle of smaller subunits (Nqo7, Nqo10 and Nqo11) forms a connection between the antiporter-like subunits and Nqo8. Subunit Nqo8 is the most conserved subunit in the membrane domain. It is unique to the family of complex I‑related proteins and is involved in quinone binding at the junction between the peripheral and membrane arms; this suggests that Nqo8 is probably key to the coupling mechanism45. Surprisingly, the Nqo8 core TM2–TM6 helices were found to have the same fold as one of the five-TMH symmetry-related domains in the antiporter-like sub units9. In contrast to the rest of the membrane domain, all TMHs of this subunit are highly tilted relative to the membrane normal. Helices TM1, TM6 and amphipathic AH1, as well as TM1 from Nqo7, frame the entrance into the quinone-binding site (FIG. 2a).

Mechanism of complex I The mechanism of complex I is unique in that it must couple spatially separated electron transfer and proton translocation pathways, as discussed below.

π‑bulge (Also known as π–helix). A protein feature created by the insertion of a single additional amino acid into a pre-existing α‑helix, destabilizing secondary structure in potential functional sites.

The electron transfer pathway. The electron donor NADH binds to its binding pocket in the Nqo1 subunit of the peripheral arm (FIG. 2c) in an extended conformation, enabling effective hydride transfer to FMN33. Analysis of the distances between redox centres (FIG. 2c) suggests that the overall electron transfer pathway comprises the following: NADH→FMN→N3 →N1b→N4→N5→N6a→N6b→N2→Q. The first step (FMN reduction) and last step (quinone reduction) in the chain involve the transfer of two electrons, whereas Fe–S clusters transfer one electron at a time. With turnover rates of about 200 s−1, each catalytic cycle would take ~5 ms46, which is much slower than both the calculated47 and the measured48 electron transfer rates from NADH to N2 of about 100 μs. As most complex I Fe–S clusters are reduced under physiological steady-state conditions49, it is thought that the overall rate of electron transfer is limited by quinone binding and release48. N3, N4 and N6a are equipotential at about –250 mV, whereas N2 has the highest potential (–100 mV to –150 mV), as can be expected for the terminal cluster in the redox chain38,50. By contrast, the intermediate clusters N1b, N5 and N6b have lower potentials, in part owing to electrostatic interactions with reduced clusters nearby, which results in alternating high and low potentials, or a ‘roller-coaster’ redox profile along the chain50,

as is common in redox enzymes. Such an arrangement optimizes the rate of electron transfer along the chain and may help to achieve efficient energy conversion51. Clusters N5 and N2 are unusual and may not be simple ‘stepping stones’ in the redox chain. 4Fe–4S cluster N5 is coordinated by three Cys residues and a His residue, instead of the usual four Cys residues. It is separated from the next cluster (N6a) by 14 Å, the longest distance in the chain34, and has a very low potential. Thus, it represents a major bottleneck in the pathway and is most likely to control the overall rate of electron transfer. The cluster is surrounded by charged and polar residues despite being buried deep in the protein. It is possible that the unusual coordination and environment of cluster N5 help it to sense the redox state of downstream clusters and to control electron transfer accordingly. The terminal 4Fe–4S cluster N2 is also unusually coordinated, with two of the four Cys residues being consecutive (Nqo6 residues 45 and 46). This results in unfavourable geometry, so that in the reduced state one of the Cys residues disconnects from the cluster 33. The midpoint potential of N2 is pH dependent, indicating that its reduction is coupled to proton binding 38, possibly to one of the disconnected Cys residues. Such a change in coordination of the 4Fe–4S cluster following reduction is linked to modest, but significant, conformational shifts of several helices nearby33. This represents a novel direct connection between the redox state of the cluster and protein conformation, which may facilitate conformational coupling (see below). Coordination of a 4Fe–4S cluster by consecutive Cys residues is rare, with only one other example known (adenosine 5ʹ‑phosphosulfate reductase52). As such coordination is fully conserved in complex I across species, the conformational flexibility and/or unusual redox properties of the cluster must be essential for coupling. Similarly, the off-pathway 2Fe–2S cluster N1a is fully conserved, which suggests that it has functional significance, possibly preventing the excessive production of reactive oxygen species (ROS)34,37. In addition, the bifurcation of electron transfer from FMN to the main chain and to N1a might ensure a long (millisecond) lifetime of a state in which N2 is reduced but there is no second electron available to complete ubiquinone reduction48. Such a state might be required to initiate a conformational change to prime the proton pump mechanism. The electron transfer path ends in the quinone-binding site formed between subunits Nqo4, Nqo6, Nqo7 and Nqo8 (FIG. 2d). This site is extremely unusual, as it is long, enough to accommodate nearly an entire quinone molecule (including most of the tail) and shielded from the solvent. The quinone headgroup binds in the deep end of a cavity, about 15 Å out from the membrane surface. This is in contrast to other membrane proteins, the quinonebinding sites of which are usually open and accommodate only the quinone headgroup9. Surprisingly, the quinonebinding site is lined mostly by hydrophilic residues, which may guide the quinone headgroup deep into the cavity. Importantly, owing to tight protein packing near the bound headgroup, quinone can be protonated only by the coordinating residues (invariant Tyr87 and His38 from Nqo4) and not by solvent water molecules. The charged



REVIEWS species (either Q2− or charged residues nearby) can exist in the chamber because it is relatively hydrophilic and distal from the membrane. Thus, if the protein controls quinone protonation, the charge can be used to drive conformational changes, and only after they are completed will the quinone be protonated and released. Proton translocation channels. Each symmetry-related set of five helices in the antiporter-like subunits forms an apparent half-channel for proton translocation, with TM4–TM8 comprising the cytoplasmic half and TM9– TM13 the periplasmic half 35 (FIG. 3a). The half-channels are formed by conserved polar residues and polar cavities containing water molecules, some of which were identified by crystallography 35. Any proton-translocating channel should have a residue (or another chemical group) with a regulated pKa (that is, a changeable affinity for a proton) and a ‘gate’ (that is, a conformational switch between proton input and proton output) to achieve vectorial proton transport. The Lys residues in the centre of each half channel are key; they are conserved in proton channels across species, essential for activity and located on the breaks in symmetry-related TM7 and TM12 (thus termed LysTM7 and LysTM12). The only exception is a conserved Glu residue replacing Lys in TM12 of subunit Nqo13. Thus, five of six key residues in proton channels in complex I are Lys residues. This is highly unusual, as proton pumping normally involves carboxylates (Asp or Glu), with a positively charged residue such as Arg modulating the pKa of the key carboxylate53. In the antiporter-like subunits of complex I, these roles seem to be reversed: the conserved essential Glu residues from TM5 (GluTM5) can modulate the pKa of nearby LysTM7 from the same subunit and that of LysTM12 from the neighbouring subunit 35. This modulation can occur in an alternating manner (that is, LysTM7 will be de‑protonated when neighbouring LysTM12 is protonated) during the catalytic cycle, consistent with the directionality of the proton pump. In addition, the negatively charged C termini of the first halves of the broken TM12 helices can interact electrostatically with LysTM12 (or GluTM12). One known example of a protein in which a Lys residue participates in proton translocation is the amino acid transporter ApcT from Methanocaldococcus jannaschii, in which protonation of a key Lys residue is suggested to be linked to conformational changes owing to interactions with a broken TMH54. The reasons for the reversal of LysTM12 (which is conserved in Mrp) to GluTM12 (which is conserved in Nqo13 of complex I) may lie in the evolutionary origins of complex I from antiporters. Homology modelling of the two main subunits of Mrp antiporters (MrpA, which is homologous to Nqo12, and MrpD, which is homologous to Nqo13 and Nqo14) suggests that the proteins share similar proton translocation pathways through these subunits (FIG. 3b). Mrp complexes catalyse active efflux of Na+ in electrogenic exchange for H+ entering the cell (that is, in the opposite direction to the normal complex I reaction), with a likely stoichiometry of 2 H+ per 1 Na+ (REF. 55). Therefore, it is probable that Na+ is transported

at the MrpA–MrpD interface, driven by conformational changes analogous to those in complex I. Despite the overall homology of MrpA to Nqo12, the MrpA–MrpD interface is more similar to the Nqo13–Nqo14 interface, with GluTM5 from MrpA facing LysTM12 in MrpD (FIG. 3c). Because of its position at the subunit interface in the vicinity of key LysTM7 from MrpA and LysTM12 from MrpD, the conserved GluTM5 residue is likely to be involved in binding Na+. Similarly, in Na+/H+ antiporters from the NhaA family (which are not related to the Mrp family), conserved Asp residues, which are located near the breaks in TMHs, are proposed to be involved in Na+ binding 56,57. Additional coordination of Na+ can be provided by exposed backbone carbonyls from the break in TM12, as is the case in Na+-coupled transporters58. Mutations of either GluTM5 or key Lys residues abolish Mrp antiporter activity 55. Electrostatic interactions between GluTM5 and LysTM7 (and LysTM12) can lead to coupling of Na+ translocation at the interface of the subunits to H+ translocation via the interiors of MrpA and MrpD; this explains the key role of Lys in the Mrp family. If evolutionary ancestors of complex I included Mrp-like antiporters45, the key role of Lys residues in proton translocation would be preserved. In the case of the Nqo12–Nqo13 interface, it seems that the additional conserved Arg163 residue in Nqo12 replaced Na+ (FIG. 3d), similarly to Na+ being replaced by Arg in CaiT from E. coli or by Lys in ApcT, which made these transporters Na+ independent 54,59. The conversion of LysTM12 in MrpD to GluTM12 in Nqo13 would then have been necessary to preserve complementary electrostatic interactions between subunits; that is, the GluTM5–Arg pair interacts with GluTM12 rather than GluTM5 interacting directly with LysTM12. Furthermore, the presence of Arg163 and the additional conserved Asp166 in TM6 of Nqo12 would lead to shorter distances between charged residues (FIG. 3c,d; see Supplementary information S1 (figure)), resulting in stronger interactions and potentially tighter coupling in this distal part of the complex. Subunits Nqo13 and Nqo14 probably emerged from MrpD gene duplication, and their interface resembles that of MrpA–MrpD, as no additional Arg is present and LysTM12 is not replaced by Glu. It is therefore possible that a residual antiporter activity is restored at this interface in thermally deactivated46 bovine complex I60, which has lost its tight coupling to oxidoreductase activity. Despite some reports to the contrary 61, there is no clear experimental evidence that native, intact complex I from any species functions as an antiporter in vivo. An additional fourth proton channel consists of two connected half-channels, with the cytoplasmic half formed by the 5-TMH antiporter-like part of Nqo8 and the periplasmic part formed by the small subunits Nqo7, Nqo10 and Nqo11 (REF. 9). A large number of charged residues of Nqo8 are embedded in the membrane, which is unusual for any membrane protein and especially for a protein that does not translocate any large or highly charged substrates. Some of these charged residues form the central part of the channel, including an Asp residue and three interacting Glu residues (the channel is thus

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REVIEWS named E‑channel, from one-letter amino acid code E used to represent Glu), whereas others form a funnel-like connection to the quinone-binding site (FIG. 3a). TM5 from Nqo8 (which contains the conserved Glu213) and TM3 from Nqo10 (which contains the functionally important Tyr59 (REF. 35) and is surrounded on all sides by conserved Glu and Asp residues) are key discontinuous helices that are likely to be involved in proton translocation. Glu130 and the conserved Glu163 (FIG. 3a) are particularly close to each other and may share a proton, as is common for buried pairs of acidic residues62. Therefore, it seems that,

in the E‑channel, the more usual carboxylates, rather than Lys residues, have a key role in proton translocation. It was recently proposed that in Y. lipolytica proton translocation takes a different path in the cytoplasmic (matrix) half of the fourth channel (rather than following the E-channel): at the interface of subunits ND2 (Nqo14) and ND4L (Nqo11)32. Although a similar possibility was suggested when the structure of the E. coli complex I membrane domain was first solved35, it later became clear that this interface is considerably disrupted in the isolated membrane domain and is much more tightly closed in

a

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Glu163 Glu213 11_Glu67 11_Glu32 Glu130 Asp72 Lys216 Lys186 Glu112 10_Tyr59

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Figure 3 | Proton translocation channels. a | Two sets of five symmetry-related helices in the antiporter-like subunits Nqo12, Nqo13 and Nqo14 each form an apparent half-channel for proton translocation, with TM4–TM8 comprising the cytoplasmic half and TM9–TM13 the periplasmic half (not shown). Polar residues lining the channels are shown as stick models with carbons shown in dark blue for the first (amino-terminal) half-channel, in green for the second (carboxy-terminal) half-channel and in orange for connecting residues. Key residues for proton translocation in antiporter-like subunits — that is, GluTM5 (132, 123 and 112) and LysTM7 (216, 204 and 186) from the first half-channel, Lys or HisTM8 (241, 235 and 216) from the connection and Lys or GluTM12 (385, 377 and 345) from the second half-channel — are indicated. Residues with similar roles in the E‑channel are also indicated (Glu–Asp quartet comprises Glu213, Glu163 and Glu130 from Nqo8, and Asp72 from Nqo7; 11_Glu67, 11_Glu32, 10_Tyr59 are also important for proton translocation). The quinone-binding cavity is shown in brown, with the modelled ubiquinone molecule shown in cyan and residues connecting the cavity to the E‑channel shown in magenta. Previously suggested proton translocation pathways are indicated by grey arrows, and additional proposed paths (new entry sites and inter-subunit transfer) by black arrows. b | Evolutionary links between Mrp antiporter subunits and complex I are shown. Homology model of subunits MrpA and MrpD from the Bacillus subtilis Mrp antiporter — built with MODELLER 9v7 (REF. 90) using Escherichia coli complex I subunits NuoL and NuoM, respectively, as templates35 — suggests very similar proton translocation pathways. Polar residues lining the putative proton translocation pathways (grey arrows) are shown as stick models, with carbons shown in dark blue for the N‑terminal half-channel, in green for the C‑terminal half-channel and in orange for connecting residues. Key residues, GluTM5 (140 and 137) and LysTM7 (223 and 219) from the first half-channel, Lys or HisTM8 (248 and 250) from the connection and LysTM12 (405 and 392) from the second half-channel, are labelled. Possible Na+ translocation pathway is indicated by a black arrow. c | Key charged residues at the interface of subunits MrpA–MrpD are shown. Putative Na+ may be coordinated by GluTM5 from MrpA (Glu140). d | Key charged residues at the interface of subunits Nqo12 and Nqo13, with additional Arg163 and Asp166 residues, are shown.

Quinone

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b Cytoplasm

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H+

Glu140 Lys250 Lys219Glu137

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REVIEWS

Grotthuss-type mechanism A proton-hopping mechanism, whereby protons travel through networks of water molecules and protonatable side chains via the formation and cleavage of hydrogen bonds.

the entire complex 9. The path between ND2–ND4L to the matrix is in fact blocked by large hydrophobic residues both in T. thermophilus and in Y. lipolytica, which makes the proposal for proton translocation pathway being located here unlikely. By contrast, the E‑channel is packed with residues that can be protonated and is also more likely to be the site of proton translocation from evolutionary and internal symmetry considerations owing to its similarity to antiporter-like subunits. The residues constituting the E‑channel form part of a notable continuous hydrophilic axis of charged and polar residues that are surrounded by many water molecules (evidence for this has been obtained through both modelling and crystallography 35). The axis is located in the middle of the membrane and spans the entire length of the membrane domain of complex I, from the quinonebinding site to the tip of subunit Nqo12 (REF. 9) (see Supplementary information S1 (figure)). The residues along the axis are found either in the half-channels or in the areas that connect them, and most are located in or near the breaks in discontinuous helices (that is TM7, TM8 and TM12 in the antiporter-like subunits, and 8_TM5, 10_TM3; in which the prefixes indicate the subunits), enabling flexibility along this polar axis. This flexibility is probably key to the mechanism, as proton translocation is likely to be facilitated by conformational changes within the axis rather than just being electro statically driven63 owing to large distances between some of the charged residues along the axis (see Supplementary information S1 (figure)). Pure electrostatic coupling as in the ‘wave-spring’ model63 would require a very precise design to achieve proton gating, which seems incompatible with the inherent protein flexibility. Overall, the presence of four putative channels in complex I suggests that each of them translocates one proton across the membrane per catalytic cycle, which is consistent with known stoichiometry. A detailed analysis of putative proton translocation pathways, taking into account all protonatable residues and modelled internal water molecules (see Supplementary information S2 (figure)), indicates that the prediction of only two symmetry-related half-channels in antiporter-like subunits is probably a simplification. Distances longer than a normal hydrogen bond were included in the modelled pathways if there were no obstacles between the two stepping stones in a Grotthuss-type mechanism, allowing for side-chain movements and possible further water molecules. Additional proton input pathways into the central parts of subunits then seem possible: one from the cytoplasm roughly along central TM8 and another as a ‘side entry’ from the interface between subunits, through GluTM5 (FIG. 3a). Multiple input pathways would enable the effective capture of protons, which are present in low concentrations in the cytoplasm (which has a high pH). Conversely, an exit pathway into the periplasm seems to be possible only around TM12, as discussed above, and even that is identifiable only in some subunits. This would be consistent with the necessity for the protein to tightly control ejection of protons against the gradient into the low-pH periplasm. A similar organization is apparent in the E‑channel, which has a porous cytoplasmic half and

a less clear connection to the periplasm. It is therefore likely that the central hydrophilic axis of complex I is usually poised for action, fully loaded with protons captured from the cytoplasm and re‑distributed between subunits. The Nqo12–Nqo13 interface is especially well adapted for inter-subunit proton transfer owing to the presence of the additional Arg163 and Asp166 residues in Nqo12 and the surrounding water molecules. Once during the catalytic cycle, the conformation of the membrane arm may be changed so that one proton is ejected into the periplasm from each of the four channels. These considerations are consistent with conservation patterns: the cytoplasmic surface of the membrane domain contains many conserved charged residues, which may be used for proton capture, whereas the periplasmic surface is essentially devoid of conserved exposed residues, except for small areas near the TM12 helices, where protons may be ejected (see Supplementary information S3 (figure)). Furthermore, the most detrimental mutations of key residues map to LysTM12 and GluTM12 residues64, which is consistent with the tight control of proton transfer near the exit points into the periplasm. Recent molecular dynamic studies65 are also consistent with the idea that the central axis is extensively connected by water networks to the cytoplasm but not to the periplasm. Coupling mechanism. Currently, the key question in complex I research is how exactly electron transfer and proton transfer are coupled, as these processes are separated by up to hundreds of angstroms. The redox reaction steps at which energy is released — partly following cluster N2 reduction but mostly during quinone reduction37 — must be taken into account. Time-resolved electron paramagnetic resonance (EPR) experiments with E. coli complex I revealed that cluster N2 reduction is fast but not followed by the appearance of a semiquinone radical, which suggests that the electron potential of the bound quinone–semiquinone pair is low and that most of the energy is released in a single step upon delivery of the second electron to quinone48. Recently, it was suggested that the electron potential of the bound quinone–quinol pair is also low (below –300 mV), and most redox energy is therefore released only following the protonation of quinol66. This would exclude any direct role of cluster N2 in driving conformational changes but contradicts the conservation of the seemingly unfavourable coordination of cluster N2 by tandem Cys residues. Thus, it remains to be determined whether the phenomenon of low potential of bound quinone is conserved. For example, one molecule of quinone remains bound to complex I throughout purification in E. coli 66, but not in T. thermophilus9, which suggests possible differences in potential. Models involving one or two ‘strokes’ per catalytic cycle have been discussed. A stroke in this context means a conformational change leading to proton translocation. The most widely accepted model currently is probably the ‘one-stroke one-site’ model, which proposes that all four protons are translocated at once, driven by the redox chemistry of one bound quinone molecule, which takes into account the known redox potentials of quinone

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REVIEWS reduction intermediates and the reversibility of the overall complex I reaction16,45. An alternative two-stroke67 or ‘two-state stabilization change’ model suggests that two sequential one-electron quinone reduction steps induce the conformational changes that result in the trans location of two protons per stroke. However, this model seems to be unlikely, as it assumes that one of the antiporter-like subunits does not translocate protons, which contradicts functional and structural data68. The idea of a second functional quinone-binding site69, which has been suggested to be located in antiporter-like subunit Nqo14 (ND2), is also not in agreement with structural and mutagenesis data64 indicating that all three antiporterlike subunits have similar roles in proton translocation. It is important to note that even though a single stroke involves a single large drop in the energy, this stroke is effectively divided into four parallel steps in four proton translocation channels, which is consistent with the general principles of bioenergetics, where large energy drops are usually broken into smaller intermediate steps. In this way, complex I differs from complex IV (cytochrome c oxidase), in which a large energy drop is divided into four consecutive, rather than parallel, steps (BOX 1). In the new publication on Y. lipolytica complex I32, the authors’ aforementioned ‘two-stroke’ model is suggested as an option. Thus, the mechanism is rather similar to the one proposed for the bacterial enzyme, as ‘charge stabilization’ means conformational changes that are driven by negatively charged quinone. A key role for the polar axis, discussed previously 9, was also noted in this study, although the authors suggested a more prominent role for electrostatic coupling, as proposed in the wave-spring model63, with local conformational changes also playing a part. The structure of the Y. lipolytica enzyme revealed a notable difference between the mitochondrial and bacterial structure around the quinone-binding site: the position of the conserved Nqo4 loop that contains His38 is different in the mitochondrial enzyme compared with the bacterial one, displacing bound quinone-like inhibitors further away from cluster N2. It was suggested that this may be due to the fact that the isolated mitochondrial enzyme was in its deactive (D) state, which is known to be conformationally different from the active (A) state on the basis of studies on D→A transition70. However, the same loop does not seem to show a similar shift in the bovine enzyme27, which should also be in a deactive state (of note, the structure of the bovine enzyme was analysed at lower resolution; the loop is visible but not well defined). Different conformations of two nearby loops in the ND1 and ND3 subunits were also noted32, although these are not well resolved in the current electron density. It is possible that the observed alternative conformation of the Nqo4 loop resembles part of the catalytic cycle, as quinone is moved away from cluster N2. However, this hypothesis needs to be verified by studying different redox states of the enzyme from the same species. When considering the mechanism it is remarkable that quinone would enter the cavity about 200 times per second, travel all the way to the vicinity of cluster N2, and then be reduced and protonated, moving out of the cavity as a quinol. Molecular dynamic studies are required to

determine the energetics of such unusual movements. It is possible that the opening up of the narrow entry point into the quinone-binding site forms part of the overall conformational cycle, enabling the bulky quinone headgroup to get in and out of the cavity. This is consistent with the fact that mutations in the key charged residues in the proton channels, even in the most distal LysTM12 of Nqo12 (REF. 71), completely abolish oxidoreductase activity, highlighting how tightly coupled this conformational machine is. Electron transfer from N2 initiates a cascade of conformational changes in the E-channel, then propagating to the antiporters. The architecture of the E-channel subunit Nqo8 suggests that it is flexible because its TMHs are highly tilted and it contains a large number of polar residues that are located in the membrane. The quinonebinding site is linked to the E‑channel by a hydrophilic funnel that consists of charged residues, culminating with a Glu–Asp quartet approaching the break in the highly conserved 10_TM3 (FIG. 3a), which is a ‘hot spot’ for human disease-linked mutations. The negatively charged ubiquinol (or charged residues nearby that control its protonation) can interact electrostatically with these negatively charged residues and drive conformational changes in the E‑channel. Cluster N2 could also contribute, as helices from Nqo4 and Nqo6 that directly contact Nqo8 move following N2 reduction33. Conformational changes in the E‑channel can be transmitted to the nearest antiporter-like subunit Nqo14 through interactions of key charged residues: Glu32 and Glu67 in Nqo11 and GluTM5 in Nqo14. In turn, the Nqo14–Nqo13 and Nqo13–Nqo12 pairs of subunits interact directly through contacts between the conserved Pro residue from the break in TM12 from one subunit and GluTM5 from the neighbouring subunit. As a result, key charged residues would be protonated and de‑protonated, and access to the cytoplasm and periplasm gated, as required for the pumping cycle (FIG. 4). It is energetically expensive to fold and assemble a protein with such an extensive polar axis in the middle of the membrane. The flexibility of the axis, together with the potential electrostatic and mechanical interactions of the residues forming this axis, suggests that the polar axis has a key role in driving conformational changes as they propagate from the E‑channel to the tip of the membrane domain. The distal subunit Nqo12 is the most conserved of the antiporter-like subunits, which indicates that a more precise design is needed to maintain coupling in areas that are most separated from the quinone-binding site. The traverse helix HL and the βH motif clearly play a part in keeping the membrane domain together 72,73. They probably also help in the coordination of conformational changes: helix HL can coordinate movements of each TM7 in the antiporter-like subunits, and on the opposite side of the membrane domain the interactions of the βH element with TM8 and TM12 in each antiporter can facilitate the coordination of conformational changes35. However, mutational analyses have not been conclusive so far 73,74, and the mechanistic role of helix HL remains to be defined.



REVIEWS NADH FMN Fe–S cluster

e–

N2 H+

H+

H+

H+

Q– Cytoplasm

HL

12 8

7

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5 3 5 Periplasm βH H+ Nqo14 Nqo11 H+ Nqo10 Nqo8

Figure 4 | Proposed coupling mechanism of complex I. Key helices and residues of complex I are depicted schematically. Upon electron transfer from the Fe–S cluster N2, negatively charged quinone (or charged residues nearby) initiates a cascade of conformational changes, propagating from the E‑channel (at Nqo8, Nqo10 and Nqo11) to the antiporters via the central axis (indicatedNature by greyReviews arrows) comprising and | Molecular charged Cell Biology polar residues that are located around flexible breaks in key transmembrane helices (TMHs). Cluster N2‑driven shifts (dashed arrows) of Nqo4 and Nqo6 helices33 (not shown) are likely to assist overall conformational changes. Helix HL and the βH element help to coordinate conformational changes by linking discontinuous TMHs between the antiporters. Key charged residues can be protonated from the cytoplasm through several possible pathways, including inter-subunit transfer (indicated by black arrows) (FIG. 3). Following the reduction of quinone and completion of conformational changes, Lys or GluTM12 in the antiporters and Glu32 from Nqo11 in the E‑channel each eject a proton into the periplasm. TMHs are numbered and key charged residues (that is, GluTM5, LysTM7, Lys or HisTM8 and Lys or GluTM12 from Nqo12–Nqo14, as well as Glu67 and Glu32 from Nqo11, which interacts with Tyr59 from Nqo10, Glu213 from Nqo8 and some residues from the connection to the quinone cavity) are indicated by red circles for Glu, blue circles for Lys or His, and white circle for Tyr. FMN, flavin mononucleotide. Figure from REF. 9, Nature Publishing Group.

The key role of the quinone redox cycle in driving conformational changes is consistent with the reversibility of the overall reaction. Complex I functions close to an equilibrium in vivo and, under conditions of high proton motive force and in the presence of a highly reduced ubiquinone pool, the reaction can be driven in the reverse direction so that ubiquinol reduces NAD+. In this case, conformational changes driven by high proton motive force can result in high affinity for ubiquinol and in a low redox potential of bound ubiquinol45 so that electron transfer can proceed in reverse towards FMN.

Conclusion The structure and mechanism of complex I discussed in this Review provide an explanation of how this intricate machinery evolved from smaller building blocks, 1. Margulis, L. Origin of Eukaryotic Cells (Yale University Press, 1970). 2. Mitchell, P. Coupling of phosphorylation to electron and hydrogen transfer by a chemi-osmotic type of mechanism. Nature 191, 144–148 (1961). 3. Abrahams, J. P., Leslie, A. G., Lutter, R. & Walker, J. E. Structure at 2.8 Å resolution of F1‑ATPase from bovine heart mitochondria. Nature 370, 621–628 (1994).

4.

5.

achieving remarkable effectiveness in energy conversion. Redox processes drive proton translocation through eight half-channels, which work together and are connected via coupling elements (the central polar axis and the HL and βH elements). A unique quinone-binding site, shielded from the solvent, is perfectly suited to use redox energy to drive conformational changes. To fully resolve the mechanism that couples electron transfer and proton translocation, we will need to answer the following questions. Which particular intermediates in the ubiquinone redox cycle are coupled to which particular step of proton translocation? Which coupling elements actually move during catalysis? How exactly is proton release into the periplasm achieved? Are all or only some of the proton pathways discussed here functional? These questions can be answered by solving and comparing structures of complex I in different redox states obtained with X‑ray crystallography and cryo‑EM. The latest cryo‑EM methods may produce high-resolution maps and have an added advantage of the enzyme being in solution and free of crystal contacts (which may limit conformational freedom of movement). Advanced molecular dynamic studies will help in the detailed analysis of all the possible pathways and interactions, and structural studies can be complemented by sitedirected mutagenesis to verify the predictions. Another major future goal is solving the full atomic structure of the more elaborate mitochondrial complex. This will help to clarify the role of supernumerary subunits and will be of obvious importance for medical applications. Owing to its central role in bioenergetics and metabolism, defects in complex I result in a plethora of human diseases. Most of these are neurodegenerative disorders, probably because neuronal tissue is highly dependent on energy production by mitochondria, and so even mild decreases in complex I activity can result in pathology — for example, Leber hereditary optic neuropathy (LHON)37,75. Complex I is also a major source of ROS in mitochondria76, which can lead to cellular damage such as mitochondrial DNA damage, protein denaturation and/or lipid peroxidation, all of which have been implicated in the pathology of Parkinson disease77 and ageing 78. All the mutations in the 14 core subunits can be mapped on a homology model (based on the bacterial structure) of the human enzyme, providing a structural basis for the understanding of the pathologies, as discussed previously 9,35. The three most common mitochondrial DNA mutations result in about 90% of LHON cases79,80 (see Supplementary information S4 (figure)). The emerging role of complex I and ROS in cancer and apoptosis81,82, as well as in ageing, still remains to be elucidated.

Watt, I. N., Montgomery, M. G., Runswick, M. J., Leslie, A. G. & Walker, J. E. Bioenergetic cost of making an adenosine triphosphate molecule in animal mitochondria. Proc. Natl Acad. Sci. USA 107, 16823–16827 (2010). Iwata, S., Ostermeier, C., Ludwig, B. & Michel, H. Structure at 2.8 Å resolution of cytochrome c oxidase from Paracoccus denitrificans. Nature 376, 660–669 (1995).

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6. Tsukihara, T. et al. The whole structure of the 13‑subunit oxidized cytochrome c oxidase at 2.8 Å. Science 272, 1136–1144 (1996). 7. Iwata, S. et al. Complete structure of the 11‑subunit bovine mitochondrial cytochrome bc1 complex. Science 281, 64–71 (1998). 8. Sun, F. et al. Crystal structure of mitochondrial respiratory membrane protein complex II. Cell 121, 1043–1057 (2005).


REVIEWS 9.

10. 11.

12.

13.

14.

15.

16.

17.

18.

19.

20.

21. 22.

23.

24. 25. 26.

27.

28.

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Acknowledgements

The work in the author’s laboratory in the Medical Research Council Mitochondrial Biology Unit (Cambridge, UK) was funded by the UK Medical Research Council. Additional fund‑ ing was provided by the Royal Society and the European Molecular Biology Organization (EMBO).

Competing interests statement

The author declares no competing interests.

SUPPLEMENTARY INFORMATION See online article: S1 (figure) | S2 (figure) | S3 (figure) | S4 (figure) ALL LINKS ARE ACTIVE IN THE ONLINE PDF


The mechanism of coupling between electron transfer and proton translocation in respiratory complex I.

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