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: sazanov@ist.ac.at
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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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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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
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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
14 TMH
(antiporter-like)
ND4L
ND4L
NuoK
Nqo11
3 TMH
ND5
ND5
NuoL
Nqo12
16 TMH
(antiporter-like)
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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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.
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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
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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.
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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
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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
Cytoplasm
His241
Lys385
Periplasm
Lys216
Lys235
Glu132 Glu377
Nqo12
Lys204
Nqo13
Glu123 Lys345
Glu163 Glu213
11_Glu67
11_Glu32
Glu130
Asp72
Lys216 Lys186 Glu112
10_Tyr59
E-channel
Nqo14
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
Nqo8
b
Cytoplasm
H+
His248
Lys405
H+
Glu140
Lys250 Lys219Glu137
Lys223
Lys392
Periplasm
MrpA
c
MrpD
Na+?
d
Na+?
Arg163
Lys216
Lys223 Glu140
Lys392
MrpA
MrpD
Glu132
Asp166
Glu377
Nqo12
Nqo13
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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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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.
NATURE REVIEWS | MOLECULAR CELL BIOLOGY
VOLUME 16 | JUNE 2015 | 385
© 2015 Macmillan Publishers Limited. All rights reserved
REVIEWS
NADH
FMN
Fe–S
cluster
e–
N2
H+
H+
H+
H+
Q–
Cytoplasm
HL
12 8
7
H+
Nqo12
5 12
H+
8
7
Nqo13
5
12 8
7
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,
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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.
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