The ATP synthase: the understood, the uncertain
and the unknown
John E. Walker1
MRC Mitochondrial Biology Unit, Wellcome Trust/MRC Building, Hills Road, Cambridge CB2 0XY, U.K.
Keilin Memorial Lecture
Delivered at the Biochemical Society
Annual Symposium held at Robinson
College, Cambridge, on 10 January
2012
Sir John Walker
Abstract
The ATP synthases are multiprotein complexes found in the
energy-transducing membranes of bacteria, chloroplasts and
mitochondria. They employ a transmembrane protonmotive
force, p, as a source of energy to drive a mechanical rotary
mechanism that leads to the chemical synthesis of ATP from ADP
and Pi . Their overall architecture, organization and mechanistic
principles are mostly well established, but other features are
less well understood. For example, ATP synthases from bacteria,
mitochondria and chloroplasts differ in the mechanisms of
regulation of their activity, and the molecular bases of these
different mechanisms and their physiological roles are only
just beginning to emerge. Another crucial feature lacking a
molecular description is how rotation driven by p is generated,
and how rotation transmits energy into the catalytic sites of
the enzyme to produce the stepping action during rotation.
One surprising and incompletely explained deduction based on
the symmetries of c-rings in the rotor of the enzyme is that the
amount of energy required by the ATP synthase to make an
ATP molecule does not have a universal value. ATP synthases
from multicellular organisms require the least energy, whereas
the energy required to make an ATP molecule in unicellular
organisms and chloroplasts is higher, and a range of values has
been calculated. Finally, evidence is growing for other roles of
Key words: adenosine triphosphate (ATP), ATP synthase, bacterium, chloroplast, energy cost,
mitochondrion, protonmotive force, rotor, stator.
Abbreviations used: p, protonmotive force; AMP-PNP, adenosine 5 -[β,γ -imido]triphosphate;
DAPIT, diabetes-associated protein in insulin-sensitive tissue; OSCP, oligomycin-sensitivityconferral protein.
1
email [email protected]
Biochem. Soc. Trans. (2013) 41, 1–16; doi:10.1042/BST20110773
ATP synthases in the inner membranes of mitochondria. Here
the enzymes form supermolecular complexes, possibly with
specific lipids, and these complexes probably contribute to, or
even determine, the formation of the cristae.
Keilin Memorial Lecture
Keilin Memorial Lecture
Introduction
“From the earliest times breathing and the warmth of the
body were recognized as the chief manifestations of life”. So
wrote David Keilin at the beginning of the first chapter of
his book The History of Cell Respiration and Cytochrome
[1]. This famous work, completed posthumously by his
daughter, Joan Keilin, traces the history of respiration from
the Roman physician, surgeon and philosopher, Galen of
Pergamon (AD 129–216) via Harvey, Lavoisier, Spallanzani,
Berzelius, Hoppe-Seyler and others through to Keilin’s own
great scientific work on the cytochromes and cytochrome
c oxidase. Keilin conducted his studies at the Molteno
Institute for Parasitological Research in Cambridge from
1925 until his death in 1963. Since then, a deep understanding
has emerged of how the cytochromes a, a3 , b, c1 and
c, which Keilin described, participate in electron-transfer
processes in mitochondria and bacteria, and, in a development
that Keilin could not have anticipated, how release of
cytochrome c from mitochondria triggers apoptosis [2]. This
knowledge about electron transfer has been integrated into
a wider comprehension of bioenergetics, stemming from
chemiosmosis, developed in the 1960s and 1970s by Peter
Mitchell [3], and Mitchell acknowledged his indebtedness to
Keilin’s work in his Nobel Lecture [4]. In his chemiosmotic
theory, Mitchell described how the redox energy derived
from oxidative metabolism is used to generate a proton
electrochemical gradient, μ̃H+ across the inner membranes
of mitochondria. He coined the phrase ‘protonmotive force’
(pmf or p), as a way of describing this gradient, and he
proposed that p drives the ATP synthase to make ATP,
the energy currency of the biological world, from ADP
and Pi . Since the 1980s, many of the details have emerged
of the atomic structures and molecular mechanisms of the
extraordinarily complicated multisubunit protein complexes
that carry out the key steps of respiration in mitochondria and
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Figure 1 Possible mechanisms that were considered to explain
the coupling of force to the synthesis of ATP
On the left, in a direct coupling mechanism, protons are carried through
the inner membranes of mitochondria into the catalytic region of
ATP synthase, where they participate directly in the chemistry of the
formation of ATP. On the right, in an indirect coupling mechanism,
the passage of protons through the membrane domain of ATP synthase
brings about intermediate effects that are then transmitted to the
catalytic sites of the enzyme and lead to synthesis of ATP. Today, we
know that the protonmotive force, p, is coupled indirectly to ATP
synthesis by a mechanical rotary mechanism.
bacteria [5–15]. Together, they provide a landmark in biology,
and they represent the culmination of the pioneering efforts
of Keilin and Mitchell.
The present article concerns the ATP synthase (also known
as the F-ATPase or F1 Fo -ATPase), which, although not
studied by Keilin himself, is one of the main beneficiaries of
the electron-transfer processes in mitochondria, bacteria and
chloroplasts, and is a key enzyme in energy conversion in
biology. Mitchell wrote extensively about the ATP synthase,
and he was interested especially in how p is coupled to ATP
synthesis. He favoured a direct coupling mechanism, where
protons are translocated through the membrane domain and
are directed into the catalytic domain of the enzyme of
the ATP synthase, where they participate directly in the
chemistry of ATP formation [3]. Other scientists supported
the indirect coupling method that we now know to operate
[16,17], but the mechanism of indirect coupling remained
obscure until the structure of F1 -ATPase was determined [18]
(Figure 1).
In the present article, I first summarize what is well
established and understood about the ATP synthase. I
go on to describe more recent and active research where
the interpretation of new findings has some attendant
uncertainty. I conclude with a summary of the main
outstanding problems in the field and how they might be
solved.
The understood
General features of ATP synthases
The ATP synthases are multisubunit enzyme complexes that
are integral components of energy-transducing membranes
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close relatives of the V-ATPases (vacuolar ATPases) found
in secretory membranes that use the energy from ATP
hydrolysis to generate ion gradients across those membranes,
and of the A-ATPases (archaeal ATPases) that synthesize
ATP in the archaea. Under aerobic conditions, the ATP
synthases make ATP from ADP and Pi using p, generated by
respiration or photosynthesis, as a source of energy. Instead
of using p as an energy intermediate, some bacteria, such as
Ilyobacter tartaricus, generate an Na + ion motive force and
couple it to ATP synthesis [20]. The structures and functions
of these Na + -dependent ATP synthases are broadly similar
to the ATP synthases that are driven by p. Many bacterial
ATP synthases can operate also in reverse under anaerobic
conditions, and hydrolyse ATP made by glycolysis, using
the energy that is released to translocate protons out of the
bacterial cytoplasm thereby generating p, which then drives
other essential cellular functions, such as chemotaxis, and
membrane transport steps that depend on p. In contrast,
the ATP hydrolytic activities of the ATP synthases from
mitochondria, chloroplasts and some bacteria are inhibited;
in vivo, these enzymes can only synthesize ATP although
their ATP hydrolase activities can be activated artificially
in vitro [21].
Overall structure of ATP synthases
Irrespective of their source, ATP synthases have related
structures, as summarized in Figure 2. They consist of two
major functional domains, a membrane-extrinsic F1 sector
and a membrane-intrinsic Fo sector joined together by central
and peripheral stalks. The F1 domain is the catalytic part of
the enzyme where ATP is formed from ADP and Pi . The Fo
domain contains a motor, which generates rotation using the
potential energy stored in p, as explained in Figure 3. The
rotational energy of the motor is transmitted to the catalytic
domain by the central stalk, which is attached directly to the
rotary motor. The peripheral stalk links the α 3 β 3 domain to
subunit a in the Fo domain, so that together they form the
integral stator of the enzyme.
The F1 domain of the ATP synthase is an assembly
of five globular proteins, α, β, γ , δ and ε, with the
stoichiometry α 3 β 3 γ 1 δ 1 ε1 [22]. The combined molecular
mass of these subunits is approximately 350 kDa. In the
α 3 β 3 γ 1 subassembly, which is common to all ATP synthases,
the three α- and the three β-subunits form an approximately
spherical α 3 β 3 structure approximately 100 Å (1 Å = 0.1 nm)
in diameter, with the six subunits arranged in alternation
around an elongated α-helical structure in the γ -subunit
[18]. This part of the γ -subunit is completely enveloped
in the α 3 β 3 domain and occupies its central axis. Thus the
α 3 β 3 structure resembles an orange made of six segments
arranged around the central pith stalk of the fruit. The
rest of the γ -subunit protrudes approximately 30 Å beyond
the α 3 β 3 domain, and forms part of a ‘foot’ that attaches the
α 3 β 3 γ 1 complex firmly to the Fo domain. This attachment is
augmented by another protein, known for historical reasons
as the ε-subunit in bacteria and chloroplasts, and as the
The ATP synthase: the understood, the uncertain and the unknown
Figure 2 Organization of protein subunits in ATP synthases
The bacterial and chloroplast ATP synthases are depicted on the left, and the more complex mitochondrial enzyme is shown
on the right. The upper part of each model contains the subunits in the F1 catalytic domain. One of the three α-subunits (red)
has been removed to expose the elongated α-helical structure in the γ -subunit (dark blue), which lies approximately along
the central axis of the spherical α 3 β 3 domain. The γ -subunit (and associated subunits) is in contact with the Fo membrane
domain, which contains the c-ring (brown) and the associated a-subunit (grey). The number of c-subunits in the c-ring
differs between species. The rotor of the enzyme consists of the ensemble of the c-ring and the γ -subunit (and associated
subunits). The pathway for protons through the Fo domain is in the vicinity of the interface between the c-ring and a-subunit.
The peripheral stalk is on the right of each model. In some bacterial enzymes, it consists of the δ-subunit (light blue) and
two identical b-subunits (pink). In other bacterial enzymes, and in the chloroplast enzyme, the two b-subunits are replaced
by single copies of homologous, but non-identical, subunits b and b’. The N-terminal domain of the δ-subunit binds to the
N-terminal region of one of the three α-subunits, and the b- and b’-subunits interact with the a-subunit via their N-terminal
transmembrane α-helices. In the mitochondrial enzyme, the peripheral stalk consists of single copies of subunits OSCP, b,
d and F6 . OSCP is the homologue of the bacterial δ-subunit. The sequences of mitochondrial subunits b, d and F6 are not
evidently related to those of the bacterial b and b’ subunits. Their structures also differ significantly, although they are both
dominated by α-helices, except for the C-terminal domain of OSCP (and presumably the bacterial and chloroplast δ-subunit),
which contain β-structures. The membrane domains of the mitochondrial enzyme contain a number of membrane subunits
with single transmembrane α-helices that are not found in bacteria and chloroplasts. These supernumerary subunits have
no known roles in the generation of ATP; subunits e, f, g, A6L and DAPIT and the 8 kDa proteolipid, which are found in the
complex when phospholipids are maintained during the purification of the complex, are shown.
δ-subunit in mitochondria. Despite their different names,
these ε- and δ-subunits are folded and probably bind in
a similar way, and their functions are probably related
also, although they are not identical. In mitochondria, the
attachment of the foot of the γ -subunit to the Fo domain
is augmented by a second subunit. Known, confusingly, as
the ε-subunit, this mitochondrial protein has no counterpart
in bacterial and chloroplast enzymes [23]. The γ -subunit,
and the associated ε-subunits in the bacterial and chloroplast
enzymes, and the associated δ- and ε-subunits in the
mitochondrial enzyme, are bound firmly to one end of
a hydrophobic cylindrical structure made of a ring of csubunits in the Fo membrane domain of the enzyme [24,25].
This cylinder, together with the γ -subunit and its associated
ε- and δ-subunits, forms the rotor of the enzyme.
How ATP is hydrolysed in the F1 domain
The understanding of the mechanism of how ATP is
hydrolysed in the catalytic F1 domain of the enzyme is based
on a series of high-resolution structures determined by X-ray
crystallography with the bovine [18,25–38] and yeast [24,39–
41] enzymes. Many of them represent either the ground
state in the catalytic cycle of ATP hydrolysis or synthesis
[24,25,30,33,35,37,39,42], or of the closely related ADPinhibited state [18,26–29,32,34,36,38,43]. One such structure
is summarized in Figure 4. In these structures, the three
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Figure 3 Model to explain the generation of rotation in the Fo
membrane domain of ATP synthases
The a-subunit (green) is shown in contact with the cylindrical ring of
c-subunits (blue). The C-terminal α-helix of each c-subunit contains
a carboxy group (in the side chain of an aspartate or glutamate
residue; dark blue circles) exposed on the external circumference of
the cylinder. From the bottom of the Figure, protons from the side
of the membrane distal from the F1 domain of the ATP synthase are
shown entering a half-channel and neutralizing a negatively charged
carboxy group in the interface between the a-subunit and the c-ring
[119]. Once neutralized, this residue moves by Brownian motion to the
more hydrophobic environment of the lipid bilayer, generating a rotation
in the direction indicated. This rotational step brings another negatively
charged carboxy group into the lower half-channel allowing another
proton to be consumed, generating another rotational substep. Thus, as
more negative charge is neutralized by protons, the protons are carried
around on the external surface the ring by the neutralized carboxy group,
until they reach a second site in contact with the a-subunit, where the
local environment reionizes the carboxy group, releasing the proton
through a second half-channel on the opposite of the membrane from
which it entered. The direction and magnitude of p ( − 180 mV inwards
in mitochondria) ensures that the rotation is unidirectional, and there is
no requirement for a ratchet mechanism. The number of protons required
to generate each 360◦ rotation of the c-ring corresponds to the number
of c-subunits that form the ring. The γ -subunit (and associated subunits)
are attached firmly to the ring, and each 360◦ rotation of this part of the
rotor provides the energy to generate three molecules of ATP from
the F1 domain.
non-catalytic α-subunits and the three catalytic β-subunits of
the enzyme are arranged in alternation around an asymmetric
α-helical structure in the single γ -subunit. The α- and βsubunits are similar, each consisting of an N-terminal domain
with six β-strands, a central nucleotide-binding domain
made of both α-helices and β-strands, and an α-helical Cterminal domain containing six α-helices in β-subunits and
seven in α-subunits. Because the γ -subunit is asymmetrical,
F1 -ATPase is an inherently asymmetrical enzyme, and a
structure of yeast F1 -ATPase without bound nucleotides
demonstrated that the asymmetry is derived from the protein
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obliges each of the three catalytic β-subunits to adopt a
different conformation with different nucleotide-binding and
catalytic properties. Two of them have similar conformations,
but in structures, now recognized as the ADP-inhibited state
of the enzyme (see the section on Regulatory mechanisms),
one subunit, designated β DP , contains bound MgADP,
and the second subunit, β TP , has bound MgAMP-PNP
(adenosine 5 -[β,γ -imido]triphosphate) (AMP-PNP is a
non-hydrolysable analogue of ATP). However, in other
structures, which represent the ground state of the active
catalytic cycle, two molecules of MgAMP-PNP [24,25,35,39],
MgADP-BeF [33] or MgADP [30,33,37,42,43] are bound to
both β TP - and β DP -subunits in the same structure. Hence
the β DP - and β TP -subunits can bind either ADP or ATP.
In all of these structures, the third subunit, the β E -subunit,
adopts a radically different conformation, in which part of
the nucleotide-binding domain and the attached C-terminal
domain have hinged outwards together, in response to the
curvature of the γ -subunit. In these structures, this subunit
has no bound nucleotide, and so it is known as the ‘empty’ or
‘open’ state, designated β E . None of the α-subunits opens
in a similar way. All three of them remain closed, each
binding an Mg2 + ion and a nucleotide, and they remain bound
throughout the catalytic cycle. The nucleotides bound to αsubunits have no known direct role in the formation of ATP.
Other intermediate states in the catalytic cycle have been
resolved by X-ray crystallography, most notably a transition
state analogue structure [31], where the catalytic cycle has
been arrested before the release of MgADP from the β E subunit, and a pre-nucleotide-release state that demonstrates
that, following the hydrolysis of an ATP molecule, the Mg2 +
ion and Pi are released from the enzyme in an unknown
(possibly random) order before the nucleotide ADP [44]. The
same state has been captured in a structure of yeast F1 -ATPase
inhibited with the yeast inhibitor protein IF1 (G.C. Robinson,
J.V. Bason, M.G. Montgomery, I.M. Fearnley, D.M. Mueller,
A.G.W. Leslie and J.E. Walker, unpublished work).
In order to explain the interconversion of catalytic sites
through ‘tight’, ‘loose’ and ‘open’ states required by a
binding change mechanism of catalysis of ATP hydrolysis
by F1 -ATPase, it was proposed that the interconversion
of sites was effected by a mechanical rotation of the γ subunit, each 360◦ rotation taking each β-subunit through
the three states represented by the β TP -, β DP - and β E subunits, thereby hydrolysing three ATP molecules [18].
It was shown subsequently by biophysical experiments
conducted with single enzyme complexes that, during ATP
hydrolysis, in either a bacterial α 3 β 3 γ complex or the intact
F1 Fo -ATPase, the direction of rotation is counterclockwise
(as viewed from the membrane domain of the enzyme)
[45,46], and that the rotation proceeds in 120◦ steps which
were resolved subsequently into 90◦ and 30◦ substeps (or
80◦ and 40◦ substeps in other experiments) [47,48]. The
ground-state structure [35] probably represents the state of
the enzyme at the end of the complete 120◦ rotary step.
During ATP synthesis, the direction of rotation is reversed,
The ATP synthase: the understood, the uncertain and the unknown
Figure 4 Structure of the F1 catalytic domain of bovine ATP synthase in the ground state of the catalytic cycle
The subunits of the protein are depicted in ribbon representation. The α-, β-, γ -, δ- and ε-subunits are red, yellow, blue,
green and purple respectively. Top left: the complete F1 domain with three α- and three β-subunits and single copies of
γ -, δ- and ε-subunits [30]. In the intact ATP synthase, the rotor of the enzyme consists of the central stalk interacting via
its foot with loop regions between the N- and C-terminal α-helices of the eight c-subunits in the Fo membrane domain
[25]. In the top right, bottom left and bottom right diagrams, the three different conformations of the catalytic β-subunits
that are present in the structure in the top left panel are shown, together with the central stalk (γ -, δ- and ε-subunits)
and an α-subunit, for reference. Each β-subunit has three domains: the N-terminal domain (top) is made of β-sheets, the
nucleotide-binding domain (middle) is a mixed α-helix/β-sheet structure, and the C-terminal domain (bottom) is a bundle
of six α-helices. Each of the three β-subunits has been obliged by the asymmetry of the α-helical coiled-coil region of the
γ -subunit to adopt a different conformation, denoted β TP , β DP and β E , with different affinities for nucleotides, the β TP - and
β DP -subunits can each bind either ADP or ATP, and the β E -subunit is unable to bind any nucleotide. During the catalytic cycle
of ATP synthesis or hydrolysis, the rotation of the central stalk takes each β-subunit through each of the three conformations.
As the rotor turns, this action leads to the binding and entrapment of ADP and phosphate at the β E -site as it closes, ATP
formation at the β DP -site and ATP release from the β TP -site as it opens and converts back into the β E -site. Each 360◦ rotation
of the central stalk (clockwise as viewed from beneath) leads to the formation of an ATP molecule from each of the three
β-subunits.
and it is assumed that the order of the structural changes
accompanying ATP hydrolysis is reversed also.
The combined high-resolution structural information
from the ground state [35], transition-state analogue [31]
and pre-nucleotide-release state [44] of the bovine F1 -ATPase
provides a detailed account of the mechanism of ATP binding
and hydrolysis, and release of the products of hydrolysis
(summarized in Figure 5). The upper part of the Figure shows
the interconversion of the β TP -, β DP - and β E -subunits with
the accompanying 120◦ rotary steps described above. The
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Figure 5 Structural description of the binding of substrates and the release of products during the catalytic cycle of ATP hydrolysis by
bovine F1 -ATPase
The structures are arranged to depict the events that occur cyclically in one of the three catalytic sites of the enzyme. In
the upper row, the conformations of the β E -, β TP - and β DP -subunits are taken from the ‘ground-state’ (GS) structure of
bovine F1 -ATPase [35], and are placed in the order described in [35], each conversion requiring a 120◦ step of the central
stalk of the enzyme. In the β DP -subunit, ATP is about to undergo hydrolysis. β TP -TS (transition state) depicts the active site
containing the transition analogue of ATP, ADP-AlF4 [31]. β E -TS represents the state immediately following the scission of
the γ -phosphate of ATP, before release of any of the products [44]. Its formation is postulated to require a 90◦ rotary step
[47]. F1 -PH represents the state of the catalytic site in β E -subunit of the structure of bovine F1 -ATPase determined with
crystals grown in the presence of phosphonate, a chelator of Mg2 + ions. The formation of this state from the transition-state
conformation is postulated to require a 30◦ rotary substep [47]. The Mg2 + ion, its hydrating water molecules and bound
nucleotide are shown in green, red and grey respectively. In β TP -GS, the Mg2 + ion is occluded.
lower part of the Figure shows intermediates formed during a
120◦ step. They are the transition-state intermediate, the state
following the scission of the γ -phosphate bond, but before
product release, and the state following the release of Pi and
the Mg2 + ion, where ADP is still bound to the enzyme. The
release of the Mg2 + ion involves the disruption of its six coordinating interactions (with four ordered water molecules,
the β-hydroxy group of the threonine residue βThr163 and
phosphate groups in the nucleotide) [44]. In the final step
(lower left), the ADP is released, and the β E conformation is
regenerated.
The main features of the mechanism of ATP synthase and
the synthesis of ATP are summarized in a series of movies
at http://www.mrc-mbu.cam.ac.uk/research/atp-synthase/
molecular-animations-atp-synthase. These movies demonstrate the rotary action of the enzyme driven by proton
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catalytic interfaces in the F1 domain, and after its formation,
ATP is shown leaving those interfaces.
The uncertain
Regulatory mechanisms
The different types of ATP synthases have various
mechanisms of inactivation and reactivation of ATP synthesis
under conditions of low and increasing available energy. In
mitochondria and chloroplasts, when p is low, MgADP
(without Pi ) remains bound to one of the three catalytic sites
of the enzyme forming an inactive complex. In chloroplasts,
it is thought that this inactive ADP-inhibited state is
stabilized during the hours of darkness by formation of an
The ATP synthase: the understood, the uncertain and the unknown
Figure 6 The bovine ATPase inhibitor protein, IF1 , and its interactions with bovine F1 -ATPase
(A) Structure of the dimeric bovine inhibitor from residues 19–84, with the identical monomers coloured blue and pink [52].
The coiled-coil region extends from residues 49 to 81. Residues 1–18 were not resolved. (B) Structure of bovine F1 -ATPase
inhibited by the monomeric bovine IF1 from residues 1 to 60. Residues 8–50 of IF1 (light blue) were resolved in this structure
[37]. (C) Binding site of bovine IF1 between the α-helical C-terminal domains of the α DP - and β DP -subunits (red and yellow
respectively). (D) Interaction of bovine IF1 with the antiparallel α-helical coiled-coiled region of the γ -subunit (dark blue)
in the central stalk region of F1 -ATPase. The interaction of the inhibitor involves a short α-helix (residues 14–18) and its
extended N-terminal region. In (B), part of the δ-subunit and the ε-subunit are coloured green and magenta respectively.
Scale bars in (A) and (B) are each 20 Å.
intramolecular disulfide linkage in the γ -subunit of the ATP
synthase. When daylight is restored, the enzyme is reactivated
by thioredoxin-regulated reduction of the disulfide and
energy-dependent dissociation of the MgADP [49,50].
Mitochondria contain an inhibitor protein known as IF1
[51]. In vitro, at a pH of ∼6.7 or below, this protein forms a
1:1 complex with either F1 -ATPase or with the intact enzyme
complex, the formation of this inhibited complex requiring
the hydrolysis of two ATP molecules. In the presence of p,
the direction of rotation of the rotor reverses, the bound
IF1 is released and ATP synthesis resumes. Hence it has
been assumed from these properties that the physiological
role of IF1 is to prevent the wasteful hydrolysis of ATP
in mitochondria under anaerobic conditions, although there
remains doubt that this is its true role in vivo.
The mode of inhibition of F1 -ATPase by IF1 has been
studied in detail (Figure 6). The active bovine protein is
a homodimer, where the C-terminal regions of the largely
α-helical proteins form an antiparallel coiled coil [52]
(Figure 6A). The protruding N-terminal regions provide the
inhibitory part of the protein, and each active dimer can bind
to two F1 -ATPase moieties [53]. At pH values of ∼8.0 and
higher, the dimers of bovine IF1 form dimers of dimers
and higher aggregates, occluding the inhibitory portion of the
protein and rendering the inhibitor inactive. Disaggregation
of these oligomers into dimers appears to be controlled by
the ionic state of a histidine residue, His49 , which is thought
to provide a pH-sensitive switch between inactive and active
states [54]. Removal of the capacity to dimerize by deletion of
the C-terminal region from residues 61–84 produces a potent
monomeric inhibitor that binds to a single F1 -ATPase moiety
[37]. In some species, for example Saccharomyces cerevisiae,
IF1 lacks the C-terminal dimerization region, and the protein
is naturally monomeric (G.C. Robinson, J.V. Bason, M.G.
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Montgomery, I.M. Fearnley, D.M. Mueller, A.G.W. Leslie
and J.E. Walker, unpublished work).
The structures of the monomeric inhibited bovine [37] and
yeast F1 –IF1 (G.C. Robinson, J.V. Bason, M.G. Montgomery,
I.M. Fearnley, D.M. Mueller, A.G.W. Leslie and J.E. Walker,
unpublished work) complexes show that IF1 is bound in
a complex binding site at a catalytic interface between the
α DP - and β DP -subunits (Figure 6B). In the bovine complex,
the inhibitor protein occupies a deep groove lined with αhelices in the C-terminal domains of the α DP - and β DP subunits (Figure 6C), and its N-terminal region interacts
with the coiled-coil region of the γ -subunit via a short
α-helix (Figure 6D), and extends into the central aqueous
cavity of F1 -ATPase. Yeast IF1 binds to yeast F1 -ATPase
in a similar manner. However, the yeast inhibitor arrests
the catalytic cycle at an earlier stage than in the bovine
complex, where the Mg2 + ion and Pi have been released
following hydrolysis of ATP, but ADP remains bound to the
β E -subunit (G.C. Robinson, J.V. Bason, M.G. Montgomery,
I.M. Fearnley, D.M. Mueller, A.G.W. Leslie and J.E. Walker,
unpublished work). In the bovine inhibited complex, the
ADP molecule has been released from the β E -subunit. The
residues in bovine IF1 that provide its binding energy
have been identified by systematic mutational analysis and
measurement of kinetic constants [55]. Most of the residues
are involved in hydrophobic interactions with the C-terminal
domain of the β DP - and β TP -subunits. Additional binding
energy is provided by an ionic interaction involving Glu30
of IF1 and Arg408 in the β DP -subunit. This information is
being used to construct more potent inhibitor proteins to
be used, for example, to probe the catalytic pathway of the
enzyme, and for investigation of the pathway of formation of
the inhibited F1 –IF1 complex.
The IF1 protein is a highly specific and effective inhibitor
of ATP hydrolysis by either F1 -ATPase or the intact ATP
synthase. This specificity has been put to practical use by
using residues 1–60 of bovine IF1 with a C-terminal affinity
tag to purify ATP synthase complexes from a wide range
of vertebrates and also from some invertebrates and fungal
species (M.J. Runswick, J.V. Bason, R. Chen, T. Charlesworth,
T. Walpole, M.G. Montgomery and J.E. Walker, unpublished
work). However, despite similarities between the subunits
of the F1 domains of mitochondrial and bacterial enzymes,
for reasons that are not understood, mitochondrial IF1 has
no effect on bacterial ATP synthases, and so this procedure
cannot be used for the affinity purification of the bacterial
enzymes.
Another potential use of the IF1 active fragment with a
C-terminal affinity tag is in the isolation of the so-called
ecto-ATPase, a version of the F-ATP synthase that has
been proposed to be associated with plasma membranes
(see, for example, [56]). Although individual subunits of the
enzyme have been detected by immunological means in these
membranes, and ATP hydrolysis activity has been shown to
be associated with the membranes, the presence of a fully
assembled enzyme complex capable of hydrolysing ATP has
not been demonstrated, and so the existence of such an ectopic
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of the subunit composition and enzyme properties of the
complex would help to resolve this issue.
The bacterial ATP synthase from Paracoccus denitrificans
and other Alphaproteobacteria have their own inhibitor
protein, known as the ξ -subunit, that is thought to
prevent ATP hydrolysis [57]. However, the protein is not
obviously closely related to mitochondrial IF1 and its mode
of action is not known. Other bacterial ATP synthases,
exemplified by the enzymes from Escherichia coli and from
the moderate thermophile Bacillus PS3 appear to be able both
to synthesize and to hydrolyse ATP, depending on whether
they are operating under aerobic or anaerobic conditions. An
inhibitory mechanism has been demonstrated in vitro for the
enzymes from E. coli [58] and Bacillus PS3 [59] involving
the ε-subunit which is bound to both to the γ -subunit
in the central stalk of the F1 -domain and to the c-ring in the
membrane domain of the enzyme (Figure 2). In a structure
of the isolated ε-subunit [60], the N-terminal domain of the
protein is folded into a ten-stranded β-sandwich, and the
C-terminal region consists of two side-by-side α-helices. In
the intact enzyme, the β-sandwich domain is involved in
binding the subunit to the γ -subunit and the c-ring, and the
α-helical C-terminal region has been proposed to adopt two
different conformations, ‘down’ and ‘up’ [61]. In the ‘down’
conformation, the two α-helices are held associated closely
with the β-sandwich by an ATP molecule bound (evidently
without an Mg2 + ion) between the two domains. In the
absence of bound ATP, the two α-helices are thought to
assume the ‘up’ position, where it has been proposed that
they could interact with the α 3 β 3 -complex and inhibit the
enzyme in the ATP hydrolysis direction. The ‘up’ position
has been captured in a structure of the intact F1 -domain
of the E. coli enzyme [62], but a coherent explanation of
the physiological context for such an inhibitory mechanism
is lacking. Other bacterial ATP synthases, exemplified by
the enzymes from Caldalkalibacillus thermarum [21] and
Mycobacterium tuberculosis [63], synthesize ATP in the usual
way under aerobic conditions, but they are not able to
hydrolyse ATP under physiological conditions. Their latent
ATP hydrolase activities can be activated only under nonphysiological conditions. Currently, there is no adequate
molecular explanation for these observations.
Generation of rotation
The cylinder of the rotor of ATP synthases is made of csubunits [24,25], simple membrane proteins folded into two
transmembrane α-helices, with the loop regions joining the
α-helices exposed on the same side of the membrane as the F1
domain, and with some of the loops at least in contact with,
and binding to, the foot of the central stalk. The N-terminal
α-helices of c-subunits form an inner ring in the core of the
cylinder, and the C-terminal α-helices provide its external
annulus. Most importantly for the generation of rotation of
the rotor by p, each c-subunit contains an essential acidic
amino acid residue (aspartate or glutamate depending on the
The ATP synthase: the understood, the uncertain and the unknown
species) found around the midpoint of the C-terminal α-helix
in the membrane domain of the enzyme [64].
During ATP synthesis, the rotation of the c-ring is driven
by p in a clockwise direction (as viewed from the membrane
domain towards the F1 domain), with an estimated rotary
speed of approximately 100–150 rev./s, depending on the
species. In the ATP synthases that hydrolyse ATP under
anaerobic conditions, the energy to drive rotation is now
provided by the hydrolysis of ATP, which drives the rotor in
a counterclockwise direction, and pumps protons across the
energy transducing membrane in the outward direction (away
from the F1 domain of the enzyme), thereby generating the
transmembrane p. The c-ring is in contact with, and rotates
against, the surface of another hydrophobic membrane
protein, known variously as subunit a (in bacteria), ATPase6 or A6 (in mitochondria) and subunit IV (in chloroplasts).
For simplicity, it will be referred to in the present paper as
subunit a. A model of its transmembrane topography has
been proposed [65]. Otherwise, little is known about the
detailed structure of this protein. It contains a basic amino
acid (Arg210 in the protein from E. coli) that participates in the
translocation of protons through the Fo membrane domain
of the enzyme [66]. This residue is strictly conserved in all
species that have been examined. It is likely that the a protein
provides a pathway (a channel or two half-channels) to allow
protons to access the negatively charged carboxy residues on
the C-terminal α-helices of c-subunits on the external surface
of the c-ring in the rotor, and two polar residues (an aspartate
and a glutamine residue) have been proposed to lie on this
pathway [67–69]. Following neutralization of the successive
carboxy groups on the surface of the c-ring, rotation of the
ring, and successive reionization of the carboxy groups,
the protons are released on the opposite side of the membrane
domain of the enzyme. One significant question that arises
is how is the stepping action of the γ -subunit generated
from the turning of the rotor? The most likely explanation
is that, before a 120◦ step occurs, energy is stored from the
turning rotor by an elastic element, and then released in a
quantum or packet (or two subpackets corresponding to the
substeps) in order to generate the stepping action. However,
the site of transient energy storage is not known, although it
could reside in the central stalk, or in the peripheral stalk, or
both.
The peripheral stalk
The peripheral stalk is an elongated and largely α-helical
structure that extends from the Fo membrane domain to
the topmost extremity of the F1 -domain, where it binds
to the N-terminal region of an α-subunit [38,70–72]. The
structure of the membrane-extrinsic region of the peripheral
stalk of the enzyme from bovine mitochondria has been
studied in detail. It is a complex of single copies of subunits
OSCP (oligomycin-sensitivity-conferral protein), b’ (subunit
b lacking its two transmembrane α-helices), d and F6 [73].
OSCP has two domains. Its α-helical N-terminal domain
provides the binding site for the N-terminal region of one
of three α-subunits, which extends above the α 3 β 3 domain
[38,71,72,74], and its C-terminal domain interacts with the
C-terminal region of the b-subunit and with F6 [38,75,76].
The region between the two domains appears to be flexible
and could provide an ‘elbow’ in an otherwise seemingly rather
rigid structure. The rest of the peripheral stalk is made of αhelices in the b, d and F6 subunits, which lie roughly parallel
to each other, giving the impression of a stiff structure, rather
than resembling a flexible rope [77]. The bacterial δ-subunit
is homologous with OSCP, and the structures of their Nterminal domains are very similar [78,79]. The sequences
of mitochondrial and bacterial b-subunits are not evidently
related [80], although the two bacterial b-subunits (or single
copies of related b and b’-subunits) are thought to form an αhelical coiled coil [81,82]. However, the detailed structures of
the C-terminal domain of the bacterial δ-subunit and of the band b’-subunits with the exception of their membrane-bound
N-terminal α-helices [83], and how they interact, remain to
be established.
The coupling of p to ATP synthesis depends upon an
integral peripheral stalk, and the peripheral stalk has been
proposed to perform a number of roles. They include the
clamping of subunit a to the c-ring [84], transient storage of
elastic energy during the 120◦ rotary steps of the enzyme [85]
and helping the α 3 β 3 domain to resist the rotational torque
of the rotor [38]. The possible presence of a flexible elbow
in the peripheral stalk argues against the first role, and the
only biophysical experiments that have addressed the issue
of where energy is stored transiently during the 120◦ steps
suggest that only the central stalk participates in this part of
the enzyme’s mechanism [86]. Therefore the only remaining
role appears to be that of resisting the rotational torque of the
rotor.
Although the location of the elastic element responsible
for transient energy storage is currently not known for
certain, the suggestion that it is most likely to reside
in the α-helical coiled-coil region of the γ -subunit is
supported by a molecular dynamics simulation based on
the structure of bovine F1 -ATPase [87]. This simulation
suggested that the coiled-coil region of the γ -subunit
undergoes structural changes during a 120◦ step. However,
in possible contradiction of this observation, pairwise
comparisons of all of the known structures of F1 -ATPase have
shown that the α-helical coiled-coil region of the γ -subunit
behaves as a rigid body in all of these structures, but there
are points of flexion above and below the coiled coil (M.G.
Montgomery, A.G.W. Leslie and J.E. Walker, unpublished
work). Thus the issue of how and exactly where transient
energy is stored remains unresolved.
Bioenergetic cost of making an ATP molecule
Knowledge of the main architectural features of the ATP
synthases and their functions allows a fundamental deduction
to be made about energy transduction in the organisms
in which these enzymes are found, namely the number of
protons that are required to be translocated through the
energy-transducing membrane for each ATP molecule that
is synthesized by the ATP synthase from ADP and Pi . This
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parameter is the energetic cost to the organism of making
each ATP molecule. Since each 360◦ rotation of the rotor
of the ATP synthase produces three ATP molecules from
the F1 domain of the enzyme, and each 360◦ rotation of the
c-ring requires the translocation of the same number of
protons as there are c-subunits in the ring, the bioenergetic
cost to the ATP synthase is the number of c-subunits in
the c-ring divided by 3 [24]. The c8 -rings in the bovine
enzyme [25] are the smallest c-rings that have been hitherto
observed, and it is likely that they occur in all vertebrate ATP
synthases, where, in examples from each class, the sequence
of the c-subunit is conserved almost completely. They are
probably found in the enzymes from invertebrates also,
where key residues that allow c8 -rings to form and to function
are conserved in all known sequences of c-subunits. It has
been estimated that there are approximately 50 000 species
of vertebrates on Earth today and approximately 2 million
species of invertebrates. The associated bioenergetic cost to
the ATP synthases in these multicellular organisms is 8/3 or
2.7 protons/ATP. By the same logic, in unicellular organisms
including fungi and bacteria (and also chloroplasts from green
plants), where ring sizes of 10, 11, 13, 14 and 15 have been
observed [24,88–91] (Figure 7), the bioenergetic cost to the
ATP synthase will be 3.3, 3.7, 4.3 and 5.0 protons/ATP
respectively. In mitochondria, ATP is produced in the matrix
of the organelle, and it is made available as a source of
cellular energy by exchanging it for external ADP. Pi is also
transported actively back into the matrix of mitochondria.
The combination of the electrogenic exchange of an internal
ATP for an external ADP and the non-electrogenic symport
of Pi and a proton adds one proton to the total required
to provide ATP to the cellular cytoplasm, and so the cost to
mitochondria containing an ATP synthase with a c8 -ring
will be 3.7 protons. In mammalian mitochondria, for each
two electrons transferred to oxygen or succinate, ten or
six protons respectively are translocated out of the matrix.
Therefore the number of moles of ADP phosphorylated
to ATP per two electrons transferred to oxygen, known
as the P/O ratio [92,93], will be 10:3.7 and 6:3.7 or 2.7
and 1.6 respectively for NADH and succinate, similar to
the experimental values of 2.5 and 1.5. P/O ratios have
been less studied in other species, and there is uncertainty
about the proton stoichiometries of their electron-transfer
complexes. Nonetheless, similar considerations apply to their
ATP synthases, except that in bacteria there is no requirement
to translocate ATP, and ADP and Pi across the energytransducing membrane.
The unknown
The issues
Just as the boundaries are blurred between what is understood
(we may ask ‘at what level?’) and what is uncertain, so
are the boundaries between the uncertain and the unknown
indistinct. Thus, in this final section of the present paper,
some of the outstanding features of the ATP synthase that
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already in foregoing sections. In this section, the emphasis is
on what remains to be established and how those goals might
be reached.
Completion of the structure of ATP synthase
The current mosaic high-resolution structural model of
mitochondrial ATP synthase lacks the subunits in the
membrane domain, especially subunit a (Figure 9), which
is required for a molecular understanding of proton
translocation and the generation of rotation, but also of
the membrane domain of subunit b and the so-called
supernumerary subunits e, f, g [94,95], A6L [96], DAPIT
(diabetes-associated protein in insulin-sensitive tissue) and
6.8 proteolipid [97]. The way that the two transmembrane
α-helices of subunit b contact subunit a, and possibly the
c-ring, could also be important factors in understanding
the enzyme’s mechanism, and the possible roles of the
supernumerary subunits are discussed below. At least two
approaches are possible for deriving a complete structure of
the ATP synthase. One is to continue to determine structures
of individual subunits, or of subcomplexes that contain them,
and thereby to add to, and to complete, the mosaic structure.
The subcomplex approach is preferable to pursuing structures
of individual subunits, since some of the information about
how subunits interact with each other in the intact ATP
synthase is retained, and, for example, knowing exactly
how the a-protein and the c-ring interact could be crucial
for understanding proton translocation. Another approach
is to solve structures of complete complexes. Clearly, this
approach would be advantageous as information about
intersubunit contacts would be retained, but, so far, intact
ATP synthase complexes have resisted attempts to induce
them to form well-ordered three-dimensional crystals that
are suitable for X-ray analysis. Some of the possible reasons
for this are becoming apparent. One possible reason is
that the intact ATP synthase is inherently unstable in the
contact region between the a-subunit and the c-ring, and
the removal of native lipids from the membrane domain
and their replacement by detergents during purification adds
to this instability. In other words, in purified ATP synthase
complexes, the ATP synthetic and hydrolytic activities of
the enzyme tend to become uncoupled from p. We have
therefore put considerable effort into defining conditions of
purification where this contact remains intact and the enzyme
remains fully coupled to p. This objective has been achieved,
and it remains to be seen whether appropriate crystals of the
intact enzyme can be formed. However, a second possible
reason for the formation of well-ordered crystals of the
intact complex being difficult is that the isolated ATP synthase
complex may be a mixture of unresolved structural isomers,
differing in the mode of association of the peripheral stalk
with the asymmetrical α 3 β 3 -domain of the enzyme. This
issue has been discussed previously in the context of the
structure of the F1 -domain with the peripheral stalk attached,
where the peripheral stalk was aligned with one specific
non-catalytic interface between the α DP - and β TP -subunits
The ATP synthase: the understood, the uncertain and the unknown
Figure 7 Structures of c-rings in the rotor of ATP synthases
(A) Cross-sections, viewed from the F1 -domain of the intact enzyme, of the inner and outer rings of N-terminal and C-terminal
α-helices made from the c-subunits in ATP synthases from various sources. The rings are taken from the following species:
c8 , bovine mitochondria [25]; c10 , mitochondria from Saccharomyces cerevisiae [24]; c11 , Ilyobacter tartaricus [88]; c13 ,
Caldalkalibacillus thermarum [89]; c14 , spinach chloroplast [90]; c15 , Spirulina platensis [91]. (B) Side view of the c8 -ring
from bovine mitochondria shown in solid representation. The inner and outer rings of N- and C-terminal α-helices in the
bovine c-subunit are yellow and blue respectively. The red and pink regions are trimethyl-lysine and glutamic acid residues
respectively. The former is proposed to facilitate the binding of cardiolipin molecules (one such molecule is shown in
association with a trimethyl-lysine residue); the latter is involved in proton translocation through the inner membrane of
mitochondria and in the generation of rotation of the ring. Scale bars in both (A) and (B) are 50 Å.
[38]. Another possible approach is to solve the structure
of the intact complex by electron cryomicroscopy of single
particles (single intact ATP synthase complexes) embedded
in ice. This approach was used to produce models of the
mitochondrial ATP synthase complex at 30, 24 and 17 Å
resolution [98–100], and recently to produce a structure of the
A-type ATP synthase complex from Thermus thermophilus
at 9.7 Å resolution [101]. This model of the A-type enzyme
indicates that the rotor ring of 12 L-subunits and the I-protein
(equivalent to the c-and a-subunits respectively in the F-type
ATP synthases) have a surprisingly small contact area. The Atype I-subunit and the F-type a-subunit are rather distantly
related, and only the region around the essential arginine
residue (Arg215 in the E. coli a-subunit) in the C-terminal
regions of the protein is obviously conserved. Currently, one
of the technical issues that limits the resolution of structures
determined by electron cryomicroscopy is the ability to
align the images of the single particles accurately. In this
respect, the A-type ATP synthase is more favourable than
the F-type enzyme as the two peripheral stalks of the Atype enzyme facilitate the alignment process. However, other
technical issues originating from the electron microscope
itself contribute to the current limitations of the resolution
of this technique.
Reconciliation of structural and rotational
experiments
It is unlikely that a detailed structure of intact ATP synthase
on its own will be sufficient to explain the generation of
rotation and the transmission of energy into the catalytic
sites of the enzyme, including providing a description of
the generation of the stepping action of the central stalk
in the F1 -domain of the integral enzyme complex. It is
likely that a number of structures with the enzyme halted
at various points in the catalytic cycle as well as biophysical
single-‘molecule’ experiments will be required, and also
biochemical and mutational experiments conducted in the
light of the structural and biophysical information. One of
the current difficulties in the ATP synthase field is that it
is apparently difficult to reconcile structural data relating to
the catalytic sites of the enzyme with the observations of
dwells in the rotational cycle. This problem applies especially
to the positions adopted by the foot of the central stalk in
various structures of F1 -ATPase, where it has been noted
several times that the position could be influenced by crystal
lattice contacts, and therefore that the observed positions
should be interpreted cautiously [25,33,36]. It has also been
proposed that the conformations of catalytic subunits in these
structures are likewise influenced by crystal lattice contacts,
and therefore that the observed conformations and nucleotide
occupancies are ‘irrelevant’ in understanding catalysis by the
enzyme [48,102–104]. Therefore we have investigated these
issues systematically by comparing in a pairwise manner
all of the 50 or more available crystal structures with each
other (M.G. Montgomery, A.G.W. Leslie and J.E. Walker,
unpublished work). The main conclusions from this study
are that, with the possible exception of yeast F1 -ATPase
inhibited with the yeast inhibitor protein, IF1 , the structures
of the catalytic sites are not influenced by lattice contacts,
and therefore that the observed structures describe the cycle
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Figure 8 Arrangement of ATP synthase complexes in the inner membranes of mitochondria
Left: molecular structure of the ATP synthase from bovine mitochondria. This overall structure is a mosaic of high-resolution
molecular structures of F1 -ATPase [30], F1 -ATPase with the stator attached [38], the stator alone [77] and the F1 –c8 -ring
subcomplex [25], all determined by X-ray crystallography. They have been assembled within a mask of the low-resolution
structure of the intact ATP synthase complex determined by cryoelectron microscopy [98]. The green region is the residual
region of the membrane domain of the enzyme where no high-resolution structures have been determined to date. This
region contains subunit a, the membrane domain of subunit b (two transmembrane α-helices), and subunits ATPase-8, e, f,
g, DAPIT and 6.8 kDa proteolipid (each with a single transmembrane α-helix). Right (A–D): dimeric ATP synthase complexes
[109].
of catalytic events in the active sites of the enzyme. The
second conclusion is that in many structures of F1 -ATPase,
the ‘foot’ regions of the central stalk are displaced by lattice
contacts. A wide range of positions has been observed
with rotations in the range − 15◦ to + 32◦ relative to a
specific ground-state structure. These rotations cannot be
correlated with observations of rotation made in biophysical
experiments. This conclusion is not surprising. In the intact
ATPase or in F1 -ATPase actively hydrolysing ATP, or in a
biophysical rotation experiment, the central stalk experiences
a rotational torque, and energy is stored transiently, possibly
in the coiled-coil region of the γ -subunit in the central
stalk. Subsequently, this energy is released in a quantized
manner to give the observed stepping action of the rotary
motor [47,86]. In crystallization experiments involving F1 ATPase, the enzyme is inactive, the elastic element is relaxed,
and the foot can adopt any one of a wide range of positions,
as observed, that are strongly influenced by crystal lattice
contacts.
Thus it appears that, in order to understand fully
the mechanism of ATP synthase, it may be necessary
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the presence of an imposed p. It is not immediately
obvious how this requirement could be fulfilled either by
X-ray crystallography or by electron cryomicroscopy of
single-enzyme complexes. In principle, one possible way
would be to obtain the structural information in situ in
mitochondrial membrane vesicles, or in vesicles containing
the reconstituted enzyme complexes, but this approach will
await the improvement of current imaging techniques.
In the meantime, much can be done to improve the
correlation of structural data of F1 -ATPases with rotational
experiments. At present, rotational experiments have been
carried out almost entirely on bacterial enzymes, where the
mutations required for building the appropriate experimental
constructs can be introduced easily. In contrast, almost all
of the structural data on F1 -ATPases with which correlation with rotational observations is being attempted, are
derived from bovine and, increasingly, yeast mitochondrial
complexes. Therefore it would probably be beneficial to
obtain more structural data with higher resolution than
at present on bacterial complexes, and also to carry out
The ATP synthase: the understood, the uncertain and the unknown
rotational experiments on yeast complexes. We are attempting
to contribute to both approaches.
Roles of ATP synthase beyond oxidative
phosphorylation
Over the years, evidence has accumulated in bacteria and
mitochondria that respiratory complexes, including the ATP
synthase complex, are organized into super- and supramolecular complexes [105,106]. Similar observations have
been made about photosynthetic complexes in the thylakoid
membranes of chloroplasts [106,107]. In the inner membranes
of mitochondria, dimers of the ATP synthase are formed
by the association of their e-subunits in their membrane
domains [108]. These dimers form ribbons, or linear rows
[109–111] (Figure 8), which are localized at the region of
highest curvature along the edges of the cristae [112]. It is well
known that cardiolipin (diphosphatidylglycerol) is essential
for the activity of a coupled ATP synthase complex [113].
Cardiolipin molecules have been proposed to bind in the
vicinity of a fully trimethylated lysine residue at position
43 [114] of the c-subunit [25]. This trimethylated lysine
residue is located in the headgroup region of the bilayer
(Figure 7B), and its presence would hamper the binding
of the headgroups of phospholipids, providing sites for
cardiolipin to bind preferentially. The ADP–ATP translocase,
another abundant component of the inner membranes of
mitochondria, also contains a trimethyl-lysine residue, which
is also in the headgroup region of the membrane, and
would be likely to have a similar preference for binding
cardiolipin molecules. The proposal that these trimethyllysine residues mark cardiolipin-binding sites is borne out by
molecular dynamics simulations (A. Duncan, A.J. Robinson
and J.E. Walker, unpublished work), and the trimethylated
lysine residue in subunit c is conserved in all vertebrate
and invertebrate ATP synthase complexes that have been
examined, but not in fungi or bacteria (T. Walpole, D. Palmer,
S. Ding, K. Jayawardena, I.M. Fearnley and J.E. Walker,
unpublished work). The cardiolipin molecules bound to the
c8 -rings of mitochondrial ATP synthase may stabilize the c8 rings to help them survive the rotational torque to which they
are subjected. They may also help to lubricate the rotation
of the ring in the membrane environment. Cardiolipin has
been detected also in association with subunit a [115]. It is
known that approximately 75 % of cardiolipin in bovine heart
mitochondria is associated with the matrix leaflet of the inner
membrane [116,117], and the association of cardiolipin with
subunit c and the ADP–ATP translocase would contribute
to this local concentration. If those cardiolipin molecules
associated with ATP synthase complexes were approximately
cone-shaped [118], they would tend to give the cristae
a negative curvature, the opposite to what is observed.
Therefore the cardiolipin molecules associated with ATP
synthase are more likely to be flattened to the surfaces of
the proteins in the membrane sector of the enzyme, and their
role is likely to be specific to the action of the ATP synthase
itself.
Concluding remarks
The present article follows closely the contents of the Keilin
Memorial Lecture delivered in Cambridge on 10 January
2012. It is intended to be an account of the contributions
of my colleagues and I towards understanding the ATP
synthase. Therefore emphasis has been placed especially on
structural studies and, to some extent, on earlier biochemical
work, where we identified the subunits of ATP synthase
complexes, determined their sequences and the organization
especially of the ‘Fo ’ subunits, some of which we recognized
by reconstitution as constituting the hitherto unknown
peripheral stalk of the enzyme. In the present paper, these
contributions have been related to biophysical studies on the
rotational mechanism of the enzyme carried out largely by
Japanese colleagues. Notwithstanding the current difficulties
of their interpretation in a structural context mentioned
above, these experiments have complemented our work
exquisitely.
The present paper is not intended to be a comprehensive
review of the ATP synthase field. Thus I have not attempted
to describe and acknowledge the many other important
biochemical and genetic contributions that have come from
many other colleagues around the world. I hope to have the
opportunity to do so elsewhere.
Acknowledgements
I am grateful to all of my many colleagues who have contributed
to this work over the last 30 years or more. Mostly, they are too
numerous to mention individually, but some of their contributions
are described in references that accompany this article. However,
I do thank especially Michael Runswick and Martin Montgomery
for their contributions to many aspects of our research, and Andrew
Leslie, FRS, for his contributions to the X-ray crystallographic aspects.
I am also grateful to Martin Montgomery for his help in preparing
the present article.
Funding
Throughout this research, I have been in receipt of Medical Research
Council intramural funding. Additional financial support was provided
by the Human Frontiers of Science Project, the European Molecular
Biology Organization, and the European Union, via the collaborative
projects EMEP (European Membrane Protein Consortium) and EDICT
(European Drug Initiative in Channels and Transporters).
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Received 2 November 2012
doi:10.1042/BST20110773