Structure and conformations of the bovine ATP synthase by ...

Structure and conformations of the bovine ATP synthase bysingle-particle electron cryomicroscopy

by

Anna Zhou

A thesis submitted in conformity with the requirementsfor the degree of Master of Science

Graduate Department of Medical BiophysicsUniversity of Toronto

© Copyright 2016 by Anna Zhou

Abstract

Structure and conformations of the bovine ATP synthase by single-particle electron

cryomicroscopy

Anna Zhou

Master of Science

Graduate Department of Medical Biophysics

University of Toronto

2016

Adenosine triphosphate (ATP), the energy currency of biology, is synthesized primarily

by the mitochondrial ATP synthase in most eukaryotes. Proton translocation in the

membrane-bound FO region leads to ATP synthesis in the catalytic F1 region through

rotation of a central rotor. The generation of rotation is proposed to occur through a

Brownian ratchet mechanism requiring two offset half-channels in the FO region. Here,

using single-particle electron cryomicroscopy (cryo-EM), seven distinct conformations

within three rotational states of the ATP synthase are presented at sub-nanometre

resolution. An average of the FO region of all seven maps provided a detailed view of the

proton-translocating a subunit, allowing an atomic model to be built using evolutionary

co-variance. The arrangement of subunits in the membrane region suggests two half-

channels for proton translocation. These channels, along with an observed oscillation

of the c8-ring in the FO region between substates in each rotational state, supports the

Brownian ratchet mechanism for the generation of rotor rotation and ATP synthesis.

ii

Acknowledgements

I would like to thank:

My supervisor, John Rubinstein, for his constant support and encouragement, and for

providing me with opportunities to learn and grow in my graduate studies and beyond.

My supervisory committee members, Mitsu Ikura and John Brumell, for their helpful

ideas and feedback.

My labmates, the Molecular Structure & Function community at the SickKids Research

Institute, and the students and faculty of the Department of Medical Biophysics for all of

the laughs, good food, and great conversations.

My family and friends, for always being there for me during my graduate school journey.

iii

Contents

List of Figures viii

List of Movies viii

List of Abbreviations viii

1 Introduction 1

1.1 The ATP synthase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1

1.1.1 Biological context . . . . . . . . . . . . . . . . . . . . . . . . . . . 1

1.1.2 Structure and composition . . . . . . . . . . . . . . . . . . . . . . 3

1.1.3 Rotary catalysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8

1.1.4 The IF1 inhibitor . . . . . . . . . . . . . . . . . . . . . . . . . . . 10

1.1.5 Brownian ratchet mechanism for proton translocation . . . . . . . 11

1.1.6 Anchoring of the peripheral stalk . . . . . . . . . . . . . . . . . . 14

1.1.7 Oligomerization and membrane curvature . . . . . . . . . . . . . . 15

1.2 Single particle electron cryomicroscopy (cryo-EM) . . . . . . . . . . . . . 16

1.2.1 Introduction to biological cryo-EM . . . . . . . . . . . . . . . . . 16

1.2.2 Microscopy theory . . . . . . . . . . . . . . . . . . . . . . . . . . 16

1.2.3 Protein structure determination with single particle cryo-EM . . . 18

1.2.4 Specimen preparation . . . . . . . . . . . . . . . . . . . . . . . . . 19

1.2.5 Image processing . . . . . . . . . . . . . . . . . . . . . . . . . . . 21

1.2.6 Near-atomic resolution with single particle cryo-EM . . . . . . . . 24

iv

1.2.7 Study of the structure of the ATP synthase using single particle

cryo-EM . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 26

1.3 Thesis objectives and overview . . . . . . . . . . . . . . . . . . . . . . . . 27

1.4 Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 28

2 Methods 29

2.1 Protein purification . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 29

2.2 Specimen preparation and microscopy . . . . . . . . . . . . . . . . . . . . 29

2.3 Image processing and 3D map refinement . . . . . . . . . . . . . . . . . . 30

2.4 Map analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 31

2.5 Building of atomic models . . . . . . . . . . . . . . . . . . . . . . . . . . 35

3 Results and Discussion 37

3.1 Three rotational states of the ATP synthase . . . . . . . . . . . . . . . . 37

3.2 Membrane curvature and a novel feature in the FO region . . . . . . . . . 44

3.3 The arrangement of subunits in the FO region . . . . . . . . . . . . . . . 47

3.4 An atomic model of the a subunit . . . . . . . . . . . . . . . . . . . . . . 51

3.5 A model for proton translocation . . . . . . . . . . . . . . . . . . . . . . 55

3.6 Conformational changes between substates . . . . . . . . . . . . . . . . . 60

3.7 Implications of flexibility in the ATP synthase . . . . . . . . . . . . . . . 63

4 Conclusions and future directions 66

4.1 Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 66

4.2 Strategies for higher resolution of the intact ATP synthase . . . . . . . . 67

4.3 Unanswered questions . . . . . . . . . . . . . . . . . . . . . . . . . . . . 69

4.3.1 The catalytic cycle of the ATP synthase . . . . . . . . . . . . . . 69

4.3.2 Dynamics of the intact ATP synthase . . . . . . . . . . . . . . . . 70

4.3.3 Structure of FO subunits . . . . . . . . . . . . . . . . . . . . . . . 71

v

Bibliography 72

Appendices 95

A Movie captions 96

vi

List of Figures

1.1 Structure of the mitochondrion . . . . . . . . . . . . . . . . . . . . . . . 2

1.2 The mitochondrial electron transport chain . . . . . . . . . . . . . . . . . 4

1.3 Structure of the mitochondrial ATP synthase . . . . . . . . . . . . . . . . 5

1.4 Binding change mechanism and rotary catalysis. . . . . . . . . . . . . . . 9

1.5 IF1 inhibitor protein bound to F1. . . . . . . . . . . . . . . . . . . . . . . 12

1.6 Brownian ratchet mechanism . . . . . . . . . . . . . . . . . . . . . . . . . 13

1.7 Single particle cryo-EM sample preparation . . . . . . . . . . . . . . . . . 20

2.1 Program for masking contamination in micrographs. . . . . . . . . . . . . 32

2.2 Particle image collection and processing. . . . . . . . . . . . . . . . . . . 33

3.1 Three main rotational states of the ATP synthase. . . . . . . . . . . . . . 38

3.2 Seven conformations of the ATP synthase. . . . . . . . . . . . . . . . . . 39

3.3 Fourier shell correlation (FSC) curves. . . . . . . . . . . . . . . . . . . . 40

3.4 Local resolution. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 41

3.5 Atomic models fit into maps. . . . . . . . . . . . . . . . . . . . . . . . . . 43

3.6 Features in the FO region. . . . . . . . . . . . . . . . . . . . . . . . . . . 46

3.7 Segmentation of the FO region. . . . . . . . . . . . . . . . . . . . . . . . 48

3.8 Evolutionary co-variance constraints. . . . . . . . . . . . . . . . . . . . . 53

3.9 Model of the a subunit. . . . . . . . . . . . . . . . . . . . . . . . . . . . . 54

3.10 Proposed proton translocation channels. . . . . . . . . . . . . . . . . . . 57

3.11 Disease-causing mutations in the a subunit . . . . . . . . . . . . . . . . . 59

3.12 Differences between substates. . . . . . . . . . . . . . . . . . . . . . . . . 62

vii

List of Movies

Movie 1. Conformational changes during the rotary cycle

Movie 2. Conformational changes between substates (Side view)

Movie 3. Conformational changes between substates (Top view)

Movie 4. Brownian ratchet mechanism

List of Symbols and Abbreviations

2D two–dimensional3D three–dimensionalA angstromADP adenosine diphosphateATP adenosine triphosphateCMC critical micelle-forming concentrationcryo-EM electron cryo-microscopyCTF contrast transfer functionDAPIT Diabetes-Associated Protein in Insulin-sensitive TissuesDDM dodecylmaltoside∆x translation along x axis∆y translation along y axisEMDB Electron Microscopy Data BankETC electron transport chainFSC Fourier shell correlationFT Fourier transformIF1 inhibitory factor 1IF11-60 ATP synthase inhibitor with residues 61 onwards deletedIMS intermembrane spacekDa kilodaltonLHON Leber hereditary optic neuropathyLS Leigh syndromeMC5DM1 Mitochondrial complex V deficiency, mitochondrial 1MIBSN Mitochondrial infantile bilateral striatal necrosisMLASA3 Myopathy, lactic acidosis, and sideroblastic anemia 3MPTP mitochondrial permeability transition pore

viii

MSA multiple sequence alignmentNADH nicotinamide adenine dinucleotideNARP Neuropathy, ataxia, and retinitis pigmentosaNMR nuclear magnetic resonanceOSCP oligomycin-sensitivity conferring proteinOTR orthogonal tilt reconstructionPDB Protein Data Bankϕ Euler angle describing rotation about the z” axisPi inorganic phosphatePSF point spread functionψ Euler angle describing rotation about the z axisRCT random conical tiltROTAN rotational analysisSNR signal-to-noise ratioθ Euler angle describing rotation about the -y’ axis

ix

Chapter 1

Introduction

1.1 The ATP synthase

1.1.1 Biological context

The mitochondrion is the organelle found in most eukaryotic cells where most adenosine

triphosphate (ATP), the energy currency of biology, is generated from the oxidation of

nutrients. Due to the relative cellular concentrations of ATP, adenosine diphosphate

(ADP) and inorganic phosphate (Pi), free energy is released when ATP is hydrolyzed

by the cell to ADP and Pi. ATP hydrolysis is therefore used to drive many cellular

processes. However, the production of ATP is energetically costly. Cells have evolved

proteins that work together to store the energy from metabolism in the chemical form of

ATP. In eukaryotes, these proteins make up the electron transport chain (ETC), a series of

metalloprotein complexes located in the inner membrane of mitochondria. In addition to

this inner membrane, which has highly folded cristae structures (Fig. 1.1), mitochondria

also have an outer membrane. The double membrane encloses the intermembrane space

(IMS) between its layers and surrounds the matrix.

During oxidative phosphorylation, substrates that were reduced during the breakdown

of nutrients (eg. carbohydrates and fatty acids), including nicotinamide adenine dinu-

1

Chapter 1. Introduction 2

Intermembrane space

Cristae

Inner membraneOuter membrane

Matrix

Figure 1.1: Structure of the mitochondrion. The mitochondrion is the organelle where ATP isproduced in most eukaryotes. It has a double membrane, which surrounds the matrix. Betweenthe membranes is the intermembrane space (IMS). The inner membrane forms highly-foldedstructures called cristae.


cleotide (NADH) and succinate, pass their electrons to the mitochondrial ETC protein

complexes (Fig. 1.2). Complex I (NADH dehydrogenase) receives electrons directly from

NADH. Complex II (succinate dehydrogenase) receives electrons directly from succinate.

Electrons are then transferred from both complex I and complex II to complex III (cy-

tochrome bc1 complex), then to cytochrome c, and finally complex IV (cytochrome c

oxidase). During these electron transfers, energy from the oxidation-reduction reactions

are used by complexes I, III and IV to pump protons across the inner mitochondrial

membrane from the matrix to the IMS. An electrochemical gradient or the proton motive

force (PMF)is established, which is required for ATP synthesis. The ETC culminates with

the reduction of molecular oxygen to water by complex IV. In the final step of oxidative

phosphorylation, the ATP synthase uses the PMF, or more specifically the energy from

the flow of protons down the proton gradient from the IMS to the matrix, to synthesize

ATP from ADP and Pi.

1.1.2 Structure and composition

The ATP synthase (Fig. 1.3) is a ∼550 kDa reversible molecular motor. It can be

divided into four functional parts: the soluble F1 (Fraction 1) region that catalyzes ATP

synthesis; the membrane-bound FO (Fraction binding Oligomycin) region through which

proton translocation occurs; the central rotor that couples proton translocation with ATP

synthesis; and the peripheral stalk that holds the F1 region stationary relative to FO. The

ATP synthase is oriented in the inner mitochondrial membrane with the F1 region in the

matrix.

The F1 region is made of subunits α3β3γδε [46, 1]. These subunits are conserved across

eukaryotes. The α and β subunits are nearly identical in their structures despite only

sharing ∼20% sequence identity, differing mostly in their C-terminal 40 residues. There

are six nucleotide binding sites in the α3β3 hexamer, three of which are catalytic sites and

three of which are non-catalytic. The nucleotide-binding site at the catalytic interface


¹⁄2 O2H2O

ADP + Pi ATPNADH NAD+ + H+

succinate fumarate

Complex I

Complex II

QComplex III Complex IV

cyt c

ATP synthase

Intermembrane space

MatrixH+H+

H+

H+

H+H+

H+ H+

H+

H+

H+

H+

H+

H+

H+

H+

H+

Figure 1.2: The mitochondrial electron transport chain. The electron transport chain (ETC) islocated in the inner mitochondrial membrane. Electron transfer from substrates reduced duringmetabolism to the metalloprotein complexes in the ETC is used to form an electrochemicalgradient across the inner membrane. The proton motive force (PMF) that is established isharnessed by the ATP synthase (blue) to generate ATP.


F1

FOafA6L e, g

b

OSCP

α β

γ

δε

d

F6

c8

Figure 1.3: Structure of the mitochondrial ATP synthase. The mitochondrial ATP synthaseis found in the inner membrane of mitochondria. It has a catalytic F1 region and a proton-translocating FO region, connected by a central rotor and peripheral stalk. Crystal structures ofthe bovine ATP synthase are shown (PDB IDs: 2WSS, 2XND, 2CLY [129, 166, 41]). There areno crystal structures for the membrane subunits except the c8-ring. Scale bars 25 A.


is formed predominantly by the β subunit, with the adjacent α subunit providing an

essential “arginine finger” (Arg373 in Escherichia coli and Saccharomyces cerevisiae).

The binding site at the non-catalytic interface is formed mainly by the α subunit. The

specific residues involved in catalysis and the binding of substrates were first determined

mostly from mutational studies of the E. coli enzyme [109] and later confirmed by crystal

structures of the F1 region from various organisms [105, 82, 80]. Crystal structures of the

F1 region also revealed that the non-catalytic sites are different from the catalytic sites in

that they lack an equivalent residue in an appropriate position to act as a catalytic base

(Glu188 in the β subunit is replaced by Gln208 in the α subunit). The non-catalytic sites

are unable to cycle between different conformations during the catalytic cycle (described

in section 1.1.3).

The membrane-bound FO region is made of subunits a, c8, e, g, f, A6L, Diabetes-

Associated Protein in Insulin-sensitive Tissues (DAPIT), 6.8 kDa proteolipid, and two

transmembrane α-helices from the b subunit [33]. The core membrane-bound subunits a,

b and c are found in all ATP synthases and are the only subunits necessary for proton

translocation in the minimal versions of the ATP synthase in bacteria [30, 59, 67]. The

additional mitochondrial ATP synthase subunits, A6L, f, e, g, DAPIT and 6.8 kDa

proteolipid, are each predicted to have one transmembrane α-helix [18]. The functions

of the essential A6L and f subunits are unknown [43, 96]. The non-essential e and g

subunits are predicted to be involved in oligomerization of the mitochondrial ATP synthase

(discussed in section 1.1.7) [7]. DAPIT and 6.8 kDa proteolipid are also non-essential

and are proposed to be involved in maintaining the populations of ATP synthase in

mitochondria [114, 51].

The F1 and FO regions contain parts of the central rotor and peripheral stalk, which

connect the two regions. The central rotor is made of subunits γδε in the F1 region [1] and

the c8-ring in the FO region. The γ subunit is bound to the c8-ring [166]. The peripheral

stalk that holds the catalytic F1 subunits stationary relative to the FO region is made of


subunits d, F6, oligomycin-sensitivity conferral protein (OSCP) and the soluble portion

of the b subunit [35, 34]. Despite its name, OSCP does not bind oligomycin, an ATP

synthase inhibitor. Oligomycin binds the FO region to inhibit proton translocation [155]

and requires OSCP. While the sequence identity of similar peripheral stalk proteins in

the bovine and yeast enzymes is low (∼14.5% between the bovine F6 subunit and the

similar h subunit in yeast), their functions appear to be conserved, as deletion of the h

subunit in yeast can be complemented with cDNA encoding the bovine F6 subunit [164].

The structure of the peripheral stalk of the eukaryotic ATP synthase is quite different

from the structure of the E. coli ATP synthase peripheral stalk, which is composed of

subunit δ (the bacterial equivalent to the mammalian OSCP subunit) and two copies of

the membrane-bound bacterial b subunit [45].

Crystal structures of the F1 region and peripheral stalk of the bovine ATP synthase

have been determined, providing important insight into the architecture of these protein

complexes [41, 129] (Fig. 1.3). There are structures of F1 with and without the c8-ring

[166, 129] and a structure of a portion of the peripheral stalk with segments of subunits b,

d and F6 [41]. The only high-resolution structures available for subunits in the FO region

are of the c-ring. c-rings with eight subunits are predicted to be conserved in all animals

[166]. Although the numbers of c subunits in the crystallized c-rings of different organisms

differ from 8 to 15 [166, 152, 124], all c subunits were found to have an outward-facing

acidic residue on helix 2 that is located near the middle of the membrane bilayer. This

residue is conserved and proposed to be involved in proton binding (section 1.1.5).

The difference in the number of c subunits means that different organisms may have

different bioenergetic costs of producing ATP. This cost is associated with the ratio of the

number of ATP molecules produced to the number of oxygen atoms reduced by the ETC,

termed the P/O ratio [166]. The P/O ratio depends, in part, on the number of c subunits

that translocate protons for each full turn of the rotor that results in the synthesis of three

ATP molecules. For the c-ring of the mammalian mitochondrial ATP synthase with eight


c subunits, theoretically, the maximum P/O ratio is 2.7 for electrons from NADH (that

enter the ETC at NADH dehydrogenase) and 1.6 for electrons from succinate (that enter

the ETC at succinate dehydrogenase) [166]. P/O ratios of 2.5 and 1.5, respectively, have

been observed [89, 64]. For yeast, which have a different ETC protein complex I that does

not translocate protons, and which have ten c subunits in the c-ring of its ATP synthase,

the P/O ratio is around 1.3 for both NADH and succinate [81]. The bioenergetic cost of

ATP production also depends on the [ATP]/[ADP] ratio in the cell.

1.1.3 Rotary catalysis

ATP is synthesized through a rotary catalytic mechanism. Proton translocation across

the FO region is driven by the transmembrane PMF, and occurs from the IMS to the

matrix at the interface between the c8-ring and the a subunit. The movement of protons

is coupled to rotation of the central rotor. Since the central rotor is asymmetric and

extends into the α3β3 hexamer [152], a sequence of concerted conformational changes is

induced in the three catalytic αβ sites as the rotor turns. These changes are described by

the binding change mechanism first hypothesized by Paul Boyer [22], and are illustrated

in Figure 1.4. Depending on the position of the rotor, each αβ catalytic site assumes one

of three conformations: loose, tight or open. The loose conformation binds ADP and

Pi, the tight conformation contains ATP, and the open conformation does not contain

nucleotide. During ATP synthesis, each site cycles through the three states, binding ADP

and Pi and releasing ATP, resulting in the production of three ATP molecules per 360◦

rotation of the rotor. The position of the peripheral stalk with respect to the catalytic

subunits is fixed, resulting in at least three distinct conformations of the ATP synthase.

The structure of the soluble F1 region is well-characterized by crystal structures of the

region with and without substrate analogs and inhibitors [80, 21, 52, 56]. Crystallization

has been facilitated by the ease of dissociation of the F1 region and central rotor from the

FO region and peripheral stalk in vitro. A combination of this high-resolution structural


T

LO

O

TL

L

O

T

ATP hydrolysis

ATP synthesis

ADP

ADP

ADP

ATP ATP

ATP

Figure 1.4: Binding change mechanism and rotary catalysis. Cross-sections through the α3β3hexamer are illustrated with α subunits in red, β subunits in yellow, the central rotor in blue,and the peripheral stalk in green. Each catalytic site in the F1 region assumes either an open(O), tight (T) or loose (L) conformation, depending on the position of the asymmetric rotor.The sequence of conformational changes in the catalytic sites is reversed between ATP synthesisand ATP hydrolysis. Due to the fixed position of the peripheral stalk, the enzyme has at leastthree distinct conformations.


information [21, 105], computational studies [63, 116] and single molecule experiments

[112, 2, 6, 101, 161] has provided a detailed understanding of catalysis in the F1 region.

These experiments were done by observing ATP hydrolysis by the isolated F1 region,

which does not require the presence of a PMF across membranes to occur. During ATP

hydrolysis in the intact enzyme, the binding of ATP and release of ADP and Pi in the F1

region result in conformational changes in the catalytic sites that drive rotor rotation.

Although not confirmed experimentally, the order of the structural changes observed

during the hydrolytic cycle are assumed to be the opposite during ATP synthesis.

1.1.4 The IF1 inhibitor

Energy released through ATP hydrolysis by the ATP synthase can be coupled to the

maintenance of a proton gradient across the inner mitochondrial membrane. However,

wasteful ATP hydrolysis, as would occur due to an uncoupling agent that dissipates

the proton gradient, dissociation of F1 from FO, or production of free F1 during the

assembly of the ATP synthase [128] is detrimental to the cell because it compromises

oxidative phosphorylation. ATP hydrolysis during ischemia, or lack of oxygen in tissues,

during which the PMF collapses, leads to cell death even after reperfusion (termed

ischemia/reperfusion injury). In mitochondria, futile ATP hydrolysis is inhibited by

inhibitory factor 1 (IF1) [126]. The bovine IF1 is a naturally-occurring 84-residue protein

that binds the ATP synthase in conditions that promote ATP hydrolysis [138]. Its

C-terminal 49-81 residues are involved in dimerization, and its N terminus binds and

inhibits the ATP synthase. At pH >6.5, IF1 exists as an α-helical, autoinhibited tetramer

[28]. At pH <6.5, protonation of His49 leads to the dissociation of the tetramer into two

active dimers held together by a coiled coil [29]. The IF1 homodimer is able to inhibit

pairs of ATP synthases.

An IF1 mutant containing only residues 1-60 and lacking the dimerization region

(IF11-60) is a potent, monomeric inhibitor with a Ki of 30 nm−1 [17]. A crystal structure


of IF11-60 bound to the bovine F1 region in a ratio of 1:1 showed that IF1 binds at the

loose (ADP-containing) catalytic site [56]. Another crystal structure where the IF11-60

mutant is bound at all three catalytic sites provided insight into the mechanism through

which IF1 binds the ATP synthase to inhibit ATP hydrolysis [15] (Fig. 1.5). The N

terminus of IF1 is disordered in solution before it interacts with a catalytic αβ site in

the open conformation. The interaction is mediated through polar/charged interactions

that induce folding. ATP hydrolysis leads to the formation of additional IF1:F1 contacts.

Hydrolysis of a second ATP molecule results in the final inhibited form of IF1:F1, in which

IF1 is stabilized in an α-helix via numerous electrostatic and hydrophobic interactions in

the active site of the αβ heterodimer in the loose conformation.

1.1.5 Brownian ratchet mechanism for proton translocation

While the cooperative conformational changes in the F1 region that couple rotor rotation

to ATP synthesis are well understood, the lack of atomic models for almost all of the

subunits in the FO region means that the mechanism through which proton translocation

is coupled to rotor rotation is still unclear. Movement of protons in FO is believed to

be coupled to rotation of the central rotor by the Brownian ratchet model (Fig. 1.6)

[77, 78]. In this model, there are two offset aqueous half-channels near the a subunit/c-ring

interface, each allowing access to different sides of the membrane. A proton will enter one

half-channel in the direction of the proton gradient to protonate the negatively charged

Glu58 residue on one c subunit (bovine ATP synthase numbering). Once the Glu58

residue is protonated and neutralized, it can no longer interact favourably with the highly

conserved, positively charged Arg159 on the a subunit. Instead, the protonated Glu58

residue will partition into the hydrophobic lipid bilayer and the c-ring will rotate. While

one Glu58 residue becomes protonated, a previously protonated Glu58 residue on an

adjacent c subunit becomes aligned with the second half-channel. The Glu58 residue

becomes hydrated, and the proton is released. Since the deprotonated Glu58 residue is


O

T

L

O

T

L

Figure 1.5: IF1 inhibitor protein bound to F1. Structure of the F1 region of the ATP synthasewith IF11-60 bound at all three catalytic sites (PDB ID:4Z1M [16]), looking in the directionfrom the FO region towards the F1 region. IF11-60 is in purple, α subunits in red, β subunits inyellow and the central rotor in blue. The conformations of the catalytic pairs are indicated aseither open (O), loose (L) or tight (T). IF1 is in its final inhibitory conformation in the loosecatalytic site. This conformation is the most α-helical and has the most interactions with F1.Scale bar 25 A.


Arg

a subunit c-ring

-COO- -COOHproton

Figure 1.6: Brownian ratchet mechanism. The Brownian ratchet mechanism for coupling ofproton translocation in the FO region to rotor rotation requires two offset half-channels atthe interface of the a subunit (green) and c-ring (yellow). Each half-channel is only accessiblefrom one side of the membrane. Rotation is generated by the protonation and deprotonationof conserved acidic residues on the c-ring, as described in section 1.1.5. The path of protonsis indicated by black arrows. An acidic residue on a c subunit becomes protonated at onehalf-channel, and moves through the membrane bilayer before becoming deprotonated at theother half-channel. The essential and conserved Arg residue on the a subunit that interacts withdeprotonated acidic residues on the c-ring is indicated in blue.


now negatively charged, it cannot partition back into the lipid bilayer, and will interact

with the positively charged Arg159 residue. Glu58 residues protonated at one half-channel

travel through the lipid bilayer before reaching the second half-channel and releasing the

proton.

The random movement of particles due to collisions with the surrounding liquid or

gas molecules is called Brownian motion. Electrostatic and hydrophobic forces prevent an

unprotonated, charged acidic residue on a c subunit from entering the lipid bilayer due

to the large energetic penalty. Thus, Brownian oscillations of the central rotor relative

to the other FO subunits, induced by the small random motions of the water and lipid

molecules surrounding it, will only result in rotation by one c subunit upon the binding of

protons to the c-ring at an unprotonated conserved acidic residue. The direction of rotor

rotation when external forces such as ATP hydrolysis are not present is determined by

the electric field, and therefore the side of the membrane more likely to deliver a proton

to a half-channel to bind the c-ring. Thus, in the presence of the PMF established by

the ETC, net rotation is in the direction of ATP synthesis (clockwise when viewed from

the IMS). Structural information for the FO region is required to determine the exact

locations of the half-channels and to trace the path of proton translocation.

1.1.6 Anchoring of the peripheral stalk

While rotating within the catalytic α and β subunits, the central rotor applies a torque

to drive ATP-generating conformational changes. To counteract this force and keep

the catalytic subunits stationary relative to the FO region, which is necessary for the

efficient coupling of proton translocation to ATP synthesis, the peripheral stalk anchors

the α3β3 hexamer to the FO region. How and where the peripheral stalk may change

in conformation to accommodate the torque from rotor rotation has not been shown.

Identifying regions of flexibility in the peripheral stalk can elucidate how it contributes to

the efficient coupling of the F1 and FO motors during ATP synthesis.


Cross-linking has shown that the b subunit is in direct contact with both the a subunit

and c-ring in the E. coli ATP synthase [40, 76]. These three subunits (a, b and c)

are the only FO subunits necessary for proton translocation in bacterial ATP synthases

[30, 59, 67], suggesting that the transmembrane α-helices of the b subunit are involved in

the transport of protons. A previous 18 A map of the bovine ATP synthase [14] showed

the approximate location of where the b subunit enters the membrane region near the a

subunit. However, where the two transmembrane α-helices of the b subunit are located in

relation to other FO subunits and their interactions with other subunits remain unclear.

Determining the structure of the membrane-bound segment of the b subunit in the intact

ATP synthase will provide insight into how it anchors the peripheral stalk and contributes

to proton translocation.

1.1.7 Oligomerization and membrane curvature

F1 regions form characteristic rows of “lollipop” shapes on cristae that can be used to

identify ATP synthase molecules in electron micrographs of mitochondrial membranes

[119]. Rows of ATP synthase dimers were first observed with electron microscopy on

freeze-fractured and deep-etched Paramecium mitochondria [5]. Dimers were subsequently

observed to be formed by yeast, bovine, chloroplast and fungal ATP synthases [7, 39, 144].

Electron tomography of mitochondria and mitochondrial membranes showed that these

“dimer ribbons” are located at regions of high membrane curvature, suggesting that ATP

synthase dimers are responsible for creating and maintaining cristae [120, 153, 39]. More

recently, single particle cryo-EM [14] and electron tomography of two-dimensional (2D)

crystals [71] of the bovine ATP synthase showed that ATP synthase monomers are

sufficient for inducing high membrane curvature. From this observation, dimerization

was proposed to be driven by the resulting reduction in the membrane strain caused by

the presence of ATP synthase monomers [14]. Subunits e and g in the FO region are

necessary for dimer formation. Yeast strains that lacked these subunits had abnormal


cristae structure and did not have detectable ATP synthase dimers [7, 9]. However,

specific protein-protein interactions that mediate dimer and oligomer formation have not

been determined and the mechanism of dimerization is still unknown.

1.2 Single particle electron cryomicroscopy (cryo-EM)

1.2.1 Introduction to biological cryo-EM

Transmission electron microscopy (EM) is used to study biological specimens at cellular to

near-atomic scales. When applied to studying individual protein molecules, the technique

is referred to as single particle EM. One method of specimen preparation in single particle

EM involves fixing and staining the sample, which increases the resistance of the specimen

to radiation damage when exposed to an electron beam in the microscope, and also

increases electron scattering and image contrast. However, this method dehydrates and

flattens proteins, distorting their structure [62]. A method that preserves proteins in

native-like states has also been developed, called single particle cryo-EM. Using this

technique, frozen, hydrated specimens are prepared for electron microscopy. Samples

are applied to a layer of holey amorphous carbon (and more recently, gold [139], see

section 4.2), and flash frozen in a thin (<100 nm) layer of vitreous ice. Vitreous ice is a

non-crystalline type of ice that preserves protein structure for imaging. This technique

allows for the use buffers with diverse chemical components that provide native-like

conditions for the proteins of interest, including detergents for solubilizing membrane

proteins such as the ATP synthase.

1.2.2 Microscopy theory

Visualization of protein specimens in cryo-EM is possible due to the scattering of high-

energy electrons when they interact with biological matter. Contrast in images is generated

by two phenomena: amplitude contrast and phase contrast. Amplitude contrast derives


mainly from the scattering of electrons at high enough angles that they become excluded

from the objective aperture. The number of these lost electrons depends on a sample’s

molecular weight and specimen thickness [147]. Amplitude contrast is produced by

the difference in the number of scattering events that occur at each position in the

frozen specimen. However, few electrons are scattered by biological material because

it is composed primarily of light elements (hydrogen, carbon, oxygen and nitrogen).

Additionally, the density of protein is not much higher than that of the ice in which it is

embedded for cryo-EM [135, 44], resulting in only a small difference in scattering. Thus,

amplitude contrast contributes little to the contrast seen in cryo-EM images. Increase

in amplitude contrast can be achieved by using heavy metal stains, but as mentioned in

section 1.2.1, this has the disadvantage of damaging protein structure.

For cryo-EM images, the ability to see protein particles relies primarily on phase

contrast. Phase contrast in images arises from the change in phase of the scattered

electron beam before re-combining with the unscattered electron beam. Specifically, when

electrons interact with the specimen during cryo-EM, they undergo a phase shift of 90◦

[130] compared to electrons that travel through vacuum. When the microscope is in

focus, there is very little phase contrast because the difference in amplitude between the

unscattered beam and the unscattered beam combined with the scattered beam with

a 90◦ phase shift is small. To increase the contrast in cryo-EM images, the microscope

is over- or under-focused in practice. This “defocus” gives additional phase shifts to

scattered waves according to their scattering angle, and thereby increases the changes in

amplitude and image contrast upon combination of scattered and unscattered beams.

Microscope images are approximate 2D projections of macromolecules [55]. Image

formation involves the contrast transfer function (CTF), which describes the signal

contrast as a function of frequency. The specimen projection image is convoluted by

the microscope’s point spread function (PSF), and the CTF is the Fourier transform

of the PSF [55]. The effects of the CTF oscillations, which decrease in amplitude and


increase in frequency with increasing resolution, must be corrected computationally in

order to recover high resolution information. The CTF depends on the defocus used when

acquiring images. Because there are many points in a CTF where it is equal to zero and

there is no information for the image, a range of defocus values must be used in order

to collect information at all resolutions. It is simplest to correct for the CTF in Fourier

space, such as by multiplication of the Fourier transform of the image with a Weiner filter

[50].

1.2.3 Protein structure determination with single particle cryo-

EM

Single particle cryo-EM involves acquiring many images of individual proteins that have

been frozen in random orientations in vitreous ice. These images are processed com-

putationally to produce a three-dimensional (3D) density map. An advantage of using

cryo-EM for structure determination is that the treatment of images of individual proteins

means that different conformations that are present simultaneously in a sample can po-

tentially be separated into homogeneous datasets through image analysis. This separation

allows for more accurate structure determination for each conformation identified. Also,

single particle cryo-EM does not require protein crystals, which are difficult to obtain for

large macromolecular complexes that are dynamic, flexible and have unstable subunit

interactions. Crystallization is especially difficult for membrane proteins due to the added

complication of hydrophobic domains that need to be kept soluble in aqueous buffers.

Cryo-EM is a more straightforward technique for membrane proteins, because once they

have been purified, solubilised in detergent buffer, and flash-frozen in vitreous ice, they

can be imaged directly. Different detergents will exert different effects on proteins, and

the detergent that results in the highest quality images for each sample is determined

empirically. Another advantage of single particle cryo-EM in comparison to other structure

determination methods like X-ray crystallography is the requirement of relatively low


concentrations of protein (∼0.02 to 2 µM).

The signal in single particle cryo-EM improves with molecular weight, making the

technique well-suited for studying the structures of large macromolecular complexes.

Atomic structures of subcomplexes and subunits from other structural biology methods,

such as nuclear magnetic resonance (NMR) and crystallography, can be docked into

cryo-EM maps of large complexes to learn about their overall structure. The structures

of large molecules with high internal symmetry have been studied most effectively with

single particle cryo-EM, with icosahedral viruses first reaching near-atomic resolution

[70, 93, 170, 171]. It has been more difficult to use single particle cryo-EM to study

proteins less than ∼300 kDa in size. More recently, with the development of direct electron

detector technology, as well as improvements in image processing algorithms, structures

of smaller specimens lacking internal symmetry have also been determined to near-atomic

resolution (discussed in section 1.2.6).

1.2.4 Specimen preparation

Specimen preparation for single particle cryo-EM starts with the application of several

microlitres of the sample to a ∼10-50 nm thick layer of amorphous carbon supported

by a 3 mm copper/rhodium mesh grid (Fig. 1.7). The grids are glow-discharged to

increase their hydrophilicity. There are small holes of up to a few µm in diameter in the

carbon layer. After the sample has been applied and allowed to adsorb, the grid is blotted

to remove excess material, leaving a thin layer of sample across the holes. The grid is

then plunge-frozen in an ethane/propane mixture kept near liquid nitrogen temperature

(∼-196◦C). The specimen is frozen in microseconds, preventing water molecules from

crystallizing and damaging protein structure. Liquid nitrogen itself is not used to flash

freeze grids because it is close to its boiling point at ambient pressure, and heat transferred

from a grid would cause a layer of nitrogen gas to form around the specimen. The slowing

of heat transfer due to this protective gas layer would give enough time for water molecules


3 mm 60 μm 1 μm 1 μm

90°

protein particlesice

EM Grid Grid square Hole in carbon film Cross-section of hole in carbon film

Figure 1.7: Specimen preparation for single particle cryo-EM. A metal mesh grid supports athin layer of amorphous carbon with small holes in it. The sample is adsorbed to the grid, andproteins are flash frozen in a thin layer of ice in the holes.


in the sample to crystallize before freezing. This effect does not occur with short-chain

hydrocarbons at liquid nitrogen temperature because they are far from their boiling point.

A continuous, rather than holey, layer of carbon can also be used as a specimen support

in cryo-EM. The advantages of using a continuous carbon layer include the requirement

of lower protein concentrations and the ability to exchange buffers after protein has been

adsorbed to a grid. Disadvantages include the induction of preferred orientations of the

protein particles on the grid, as well as an increase in background noise in images from the

carbon itself. Having preferred orientations of particles on grids results in the collection

of more information for some views of the protein particle than others, and prevents

3D maps from being built with isotropic resolution. Extra background in the images

lowers the signal-to-noise ratio (SNR) and decreases the ability of programs to detect

and align particle images accurately. These effects can hinder the ability to determine

protein structure to high resolution (see section 1.2.5 for a detailed description of image

processing). Like the use of different detergents, the decision to use holey carbon or

continuous carbon support is dependent on the behaviour of the sample and is determined

empirically.

1.2.5 Image processing

Since cryo-EM images must be acquired with a low exposure of electrons to limit radiation

damage to specimens, they have low SNRs. Thus, thousands of particle images must

be averaged together to improve SNRs enough to access high resolution information.

Assuming Gaussian noise, the SNR of averages of particle images improves proportionally

with the square root of the number of images averaged. The SNR of averages is also

affected by the quality of images, including any background density caused by buffer

components or contamination, and by the ability to accurately separate the images into

homogeneous datasets before averaging.

To produce 2D averages, particle images are aligned by in-plane rotations and transla-


tions, and then classified into different subsets using multivariate statistical analysis. 2D

classification can be done with or without a reference [50]. All images in a class are then

averaged. 2D class averages that do not represent 2D projections of the specimen can be

discarded at this stage of image processing as they likely include other image features

and/or particle images that cannot be aligned well due to poor quality.

Following 2D classification, selected images are used to produce 3D maps. Since

the images acquired with an electron microscope are approximate 2D projections of

the protein [55], the Fourier projection theorem is used. This theorem states that the

Fourier transform of a 2D projection of an object is a central slice through the 3D

Fourier transform of the object [55]. The orientation of this central slice is related to the

orientation of the protein recorded in the image. By capturing 2D projections of a protein

in many random orientations, a 3D map of the protein can be computed. The resolution

of the 3D map depends, in part, on how well Fourier space is sampled, or, equivalently in

real space, how thoroughly the different views of the protein have been covered by the

acquired particle images.

In order to construct a 3D map using the Fourier projection theorem, the projection

direction must be determined for each image so that transformations can be applied

to align the coordinate system of each particle image with an arbitrary, fixed reference

coordinate system. Transformations are described by three Euler angles (ϕ, θ, ψ) and two

in-plane translations along the x- and y-axes (∆x and ∆y). Since these transformations are

not known a priori, they must be first determined roughly, often using 2D class averages

with higher SNRs than the noisy individual particle images, before further refinement.

There are several methods for the determination of initial transformations:

� Rotational analysis (ROTAN) [12, 137] was designed for analysis of proteins that

often assume a particular type of orientation on cryo-EM grids. This can be due to

the existence of a long axis in the protein’s overall shape, thin ice, interactions with

the air/water interface, and/or preferential adsorption of one side of the protein to


carbon. For example, the long axis of the ATP synthase leads to mainly side views

in cryo-EM particle images. The change in position of asymmetric features for class

averages sharing a single rotation axis is used to determine the relative orientations

of the class averages.

� The common lines method, employing the Fourier projection theorem, requires

three images with different projection angles. Their Euler angles are determined by

the three common lines shared by the images. High SNRs are required in the images

because lines of single pixels are used in the analysis. The real space method using

the same approach is known as angular reconstitution [162]. This method does not

work for images of the ATP synthase because particles are primarily rotated around

one axis (the long axis of the enzyme), and the images lack the third orthogonal

line needed for unambiguous assignment of Euler angles [12].

� Tilt methods for initial 3D map building require the same specimen to be imaged

at different angles relative to the electron beam. Depending on the specific method

chosen, orthogonal tilt reconstruction (OTR) [92] or random conical tilt (RCT)

[127], pairs of images are either collected at ± 45◦, or 90◦ and ∼30◦, respectively.

Untilted images are first aligned and classified to determine the in-plane rotation

angle, ϕ. The orientations of tilted images corresponding to the identical projections

can then be determined based on the microscope tilt and in-plane rotations of their

corresponding untilted image.

� The random initial starts method makes use of 3D shapes such as spheres or

ellipsoids as starting references for projection matching, in which each individual

particle image is compared to a set of projections calculated from the reference.

Particle images are then assigned transformation parameters according to the

projection of the starting model that it most represents. In a similar (but not

random) method, a low-pass filtered map of a similar protein or the protein of


interest, when available, can be used as a reference.

Once rough estimates for the projection directions of images or class averages are

obtained, they are refined by several rounds of alignment and map building. In an iterative

process, maps built using refined alignments become the reference for the next round of

alignment. Alignment algorithms make use of projection matching, as described above.

This process is most easily and effectively implemented in Fourier space. Alignments can

be ranked using different algorithms, including amplitude-weighted phase residual [60]

and SNR-weighted correlation coefficient [151].

When alignments and the 3D map no longer change with additional rounds of refine-

ment, the resolution to which the map can be interpreted is determined. Most commonly,

the Fourier shell correlation (FSC) [50] is used to determine the resolution of maps in

cryo-EM. The FSC method for resolution determination involves calculating the normal-

ized cross-correlation coefficient at different spatial frequencies between two maps that

were processed using independent halves of the dataset. The highest resolution shell

that is statistically significant and above the noise level is the overall resolution of the

3D map. An FSC threshold of 0.143 is most frequently used to determine resolution,

and is equivalent to the figure-of-merit used in X-ray crystallography to indicate the

interpretability of crystal structures [133].

1.2.6 Near-atomic resolution with single particle cryo-EM

As previously mentioned, in order to limit radiation damage that causes structural

changes in proteins, low electron exposures must be used during imaging in cryo-EM.

The resulting low SNR of images means that it is often difficult to determine accurate

orientation parameters for particle images. In addition, CTF parameters such as defocus

and astigmatism may be computed incorrectly, resulting in blurring of particle images

when CTF correction is applied. Such errors during map building limit the resolutions

of the constructed maps. The nature of different samples leads to varying degrees of


difficulty in particle image alignment. It is often easier to reach high resolution with

complexes that are relatively larger, more rigid, and more homogeneous, and that have

symmetry and/or distinctive and easily aligned features. Sub-nanometre resolutions are

difficult to reach with complexes that are flexible, small, and heterogeneous.

Despite these challenges, near-atomic resolution structures from cryo-EM have become

increasingly feasible [84, 37, 10]. This is due in large part to the development of direct

electron detectors that are capable of capturing images with higher SNRs than was

possible previously, especially for high resolution information [104]. These detectors allow

image processing programs to more accurately align images for map building. They have

also contributed to the ability of programs to classify different protein conformations in

a single sample into homogeneous datasets for refinement [141, 95], which increases the

resolution of maps because protein features are not blurred by the averaging of many

different conformations in a single 3D class.

As the fast read rates of direct electron detectors allow for movies instead of single

images to be collected, the frames of these movies can be aligned before averaging [23] to

remove the blurring that would otherwise occur in a single image due to any specimen

movement during imaging (eg. due to stage drift or beam-induced motion). Image

processing programs have also been developed that are able to estimate and remove

local beam-induced motion of individual particle images [142, 136], which varies across

the imaged area and cannot be corrected by the whole-frame drift correction of movies.

Localized blurring in images is believed to occur due to radiation damage to the specimen

that causes it to distort during imaging [54], and electric charging of the sample that

causes deflection of electrons. Correction of these effects allows for recovery of high

resolution information from individual particle images.


1.2.7 Study of the structure of the ATP synthase using single

particle cryo-EM

The structure of the intact ATP synthase has been studied previously with single particle

cryo-EM. A 32 A map of the bovine ATP synthase showed the overall shape of the complex

[137], but did not provide information on the arrangement of subunits in the membrane

region. A structure of the S. cerevisiae ATP synthase, which loses subunits e and g when

extracted from mitochondria with detergent buffer, was then determined at 24 A [86].

Comparison of the two maps allowed localization of the e and g subunits in the FO region.

A 18 A map of the bovine ATP synthase [14] revealed the arrangement of subcomplexes

in the FO region, as well as where the peripheral stalk enters the FO region. However,

the arrangement, interactions, and secondary structure of individual membrane-bound

subunits remained unclear. Since different conformations of the enzyme could not be

separated when the 18 A map was built, unique rotational positions of the central rotor

and different conformational states of the αβ catalytic sites were not resolved.

More recently, the technological advances described in the previous section have

allowed for maps of rotary ATPases to be resolved to subnanometre resolutions using

single-particle cryo-EM. Three rotational states of the S. cerevisiae V-type ATPase were

resolved to 6.9-8.3 A [172], allowing for the analysis of flexibility in the enzyme. In these

maps, two highly tilted α-helices in the a subunit at its interface with the c-ring were also

revealed. A 6.2 A map of a dimer of the colourless green alga Polytomella sp. F-type ATP

synthase also showed two highly tilted α-helices in the proton-translocating a subunit

that are in contact with the c-ring [4]. For the mammalian mitochondrial ATP synthase,

different conformational states of the intact enzyme, as well as secondary structure in

the FO region, have not been shown to date. Thus, a clear understanding of both the

mechanism that couples proton translocation to rotation of the central rotor in the FO

region and the dynamic changes in conformation in the ATP synthase during its catalytic

cycle are lacking.


1.3 Thesis objectives and overview

Although the catalytic F1 region of the ATP synthase has been extensively studied and

the mechanism of ATP synthesis in the α3β3 hexamer is well understood, there is a lack

of high resolution structural information for the FO region in the context of the intact

ATP synthase. This means that how proton translocation in FO generates rotor rotation

remains unknown. In addition, regions of flexibility in the enzyme that are required for

efficient coupling of the mismatched three-step F1 and eight-step FO motors have not

been analyzed. Thus, the objectives for this thesis were:

� to better understand the coupling of proton translocation with rotation of the

central rotor by resolving the secondary structure and arrangement of subunits in

the FO region, and

� to investigate how the symmetry-mismatched F1 and FO regions are coupled effi-

ciently by identifying and analyzing different conformations of the ATP synthase

and determining regions of flexibility.

In this thesis, using single-particle cryo-EM, we present maps of the bovine mitochondrial

ATP synthase in seven distinct conformations. These maps were refined to 6.4-7.4 A

resolution, and show three rotational states with two to three substates within each

rotational state. The conformational changes that occur in the transitions between

substates reveal flexibility throughout the enzyme, particularly in the peripheral stalk,

that is important for efficient coupling of the symmetry-mismatched F1 and FO motors

during catalysis. By averaging the FO regions of all seven maps, α-helices of membrane-

bound subunits were resolved. We show that the mammalian F-type ATP synthase a

subunit, like the S. cerevisiae V-type ATPase and algal F-type ATP synthase a subunits

[172, 4], has two highly tilted α-helices that are in contact with the c8-ring. We built an

atomic model of the a subunit based on map density using constraints from evolutionary

co-variance analysis. This model has since been corroborated by a crystal structure of


the α-proteobacterium Paracoccus denitrificans ATP synthase [108]. The structure of the

a subunit in complex with two transmembrane α-helices of the b subunit suggests the

location of two half channels for proton translocation, supporting the Brownian ratchet

mechanism for the generation of rotation required for ATP synthesis.

This work has been published in eLife [173]. The EM maps of the seven conformations

are available in the Electron Microscopy Data Bank (EMDB) under accession codes

EMD-3164, EMD-3165, EMD-3166, EMD-3167, EMD-3168, EMD-3169 and EMD-3170.

The EM map of the average of the seven maps aligned at their FO regions is deposited

under the accession code EMDB-3181. The fitted atomic structures are available on the

Protein Data Bank (PDB) under PDB IDs 5ARA, 5ARE, 5ARH, 5ARI, 5FIJ, 5FIK and

5FIL.

1.4 Acknowledgements

Bovine ATP synthase was purified by John V. Bason and Martin G. Montgomery in the

laboratory of Prof. Sir John Walker (Medical Research Council’s Mitochondrial Biology

Unit, Cambridge, United Kingdom). Microscopy and map building was done by Alexis

Rohou in the laboratory of Prof. Nikolaus Grigorieff (Janelia Research Campus, Virginia,

United States). Analysis of evolutionary co-variance and generation of the atomic model

of the a subunit was done by Daniel Schep in the Rubinstein laboratory.

Chapter 2

Methods

2.1 Protein purification

Bovine ATP synthase was purified by John V. Bason and Martin G. Montgomery as

described previously [138]. Briefly, metal (Ni-NTA) affinity chromatography was used

to isolate ATP synthase in complex with residues 1-60 of heterologously expressed IF1

with a C-terminal His6-tag. After elution from the column, the complex was PEG-

precipitated and resolubilised in buffer containing 20 mM Tris-HCl (pH 7.2), 100 mM

NaCl, 10% (vol/vol) glycerol, 0.05% (wt/vol) dodecylmaltoside (DDM), 2 mM ATP and

0.02% (wt/vol) NaN3. Glycerol was removed from purified ATP synthase using 7000

MWCO Zeba Spin Desalting Columns (Thermo Scientific) according to the manufacturer’s

instructions before sample preparation for cryo-EM.

2.2 Specimen preparation and microscopy

Preparation of grids was done by Anna Zhou. Nanofabricated grids [99] were glow-

discharged for 2 minutes in air and loaded into a Vitrobot plunge freezing device (FEI

Company, Eindhoven, Netherlands) at 100% humidity and 4◦C. 2 µl of purified ATP

synthase at ∼8 mg/mL were applied onto the grids, followed by equilibration for 5 s,

29

Chapter 2. Methods 30

blotting for 27 s and freezing by plunging into a liquid ethane/propane (30/70% vol/vol)

mixture.

Microscopy was done by Alexis Rohou. Movies were recorded from three grids using

a Titan Krios (FEI) operating at 300 kV. An area of 2.5 µm diameter was illuminated

with a parallel beam at 3 e−/A2/s and using a 70 µm objective aperture. Micrographs

were recorded at 18000× nominal magnification on a K2 Summit direct detector device

(Gatan Inc.) used in super-resolution counting movie mode, with a calibrated physical

pixel size of 1.64 A and super-resolution pixel size of 0.82 A. An exposure of 8 e−/pixel/s

without a specimen was used for 20.1 s and 67 frames, resulting in a total exposure of the

specimen area of 60.3 e−/A2. Data was automatically collected using SerialEM [102].

2.3 Image processing and 3D map refinement

Movie processing, alignment and averaging were done by Alexis Rohou. Distortion in

each frame due to magnification anisotropy [172] measured previously using the program

mag distortion estimate [58] was corrected with the program mag distortion correct [58].

Fourier-space cropping was used to downsample frames to the physical pixel size of 1.64

A. To correct for whole-frame movement, frames were aligned using Unblur [58]. The

aligned sums were used to estimate CTF parameters with CTFFIND4 [132].

Automatic particle image picking was done by Anna Zhou. Imaging of the edges of

carbon holes occurred often in movies that were collected automatically using SerialEM.

Regions of carbon contamination were masked in the aligned averages of automatically

recorded movies with a novel program written by Anna Zhou, maskmicrograph.f90. This

program prevented the automatic particle-picking function in Relion [143] from picking

carbon and ice contamination features as particle images. The program first applies a

high-pass filter to the Fourier transform (FT) of an aligned average micrograph to remove

low frequency information (eg. ice gradients). It then uses a translating box to identify


where areas in the image have a greater variance and/or lower mean than user-set intensity

threshold values, and finally masks these areas by setting them to the mean value of the

micrograph (Fig. 2.1).

Anna Zhou used Relion [141, 143] to determine coordinates for 408,934 candidate

proteins in masked micrographs automatically (Fig. 2.2A). Local beam-induced motion in

particle images was corrected by Alexis Rohou using alignparts lmbfgs [136] (Fig. 2.2B).

195,233 candidate particle images were selected by Anna Zhou for further analysis from

5,825 micrographs after 2D classification with Relion [141] (Fig. 2.2C).

Map building was done by Alexis Rohou. 256 × 256 pixel filtered and aligned [58, 136,

13] particle images were down-sampled by a factor of 2 for determination of orientation

parameters. Using an 18 A map of the bovine ATP synthase filtered to 20 A resolution

as a reference [14], initial particle orientation parameters were determined with five

rounds of mode 3 in FREALIGN (systematic parameter search) [61]. Particles were

then classified into different 3D maps with a likelihood-based algorithm [95], alternating

between orientation parameter refinement and class occupancy refinement every three or

four rounds.

2.4 Map analysis

Map analysis was done by Anna Zhou. Segmentation of 3D maps into subunits and

subcomplexes of the ATP synthase was performed manually with UCSF Chimera [57, 121].

The Segger plugin [121] in UCSF Chimera was first used to segment the maps [121].

Segments generated by Segger were ungrouped to the regions initially produced by

the watershed method employed by Segger. Ungrouped segments were then combined

manually to generate the segments for individual FO subunits and subcomplexes according

to known and predicted structures [172, 4, 90, 113].

Atomic structures were flexibly fit into 3D maps using UCSF Chimera and NAMD as


A B C D

E

Figure 2.1: Program for masking contamination in micrographs. The effects of each step of themaskmicrograph.f90 program on a micrograph with carbon and ice contamination are shownhere. A) Original micrograph with carbon and ice contamination. B) The micrograph afterhigh-pass FT filtering. C) Identification of areas of carbon and ice contamination based onuser-set variance and mean intensity thresholds. D) Micrograph with areas identified to containcontamination masked with the mean intensity value of the micrograph. E) Comparison of theresults from Relion automatic particle picking [143] for the unmasked and masked micrographs,showing a decrease in the number of features picked from areas of carbon and ice contaminationfor the masked micrograph. Particle-like features identified by the Relion autopicking programare circled in blue. Scale bar 200 A.


A B

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Figure 2.2: Particle image collection and processing. After particle images were picked usingRelion [141], local beam-induced movement was corrected, and particle images were 2D classified.A) A representative micrograph with examples of bovine ATP synthase particle images circled inblue. Scale bar 200 A. B) Trajectories of particle images and other image features during movieacquisition determined by the alignparts lmbfgs algorithm [136]. Trajectories are exaggerated5-fold. C) Averages of 2D classes selected for 3D classification and map building. Scale bar 100A.


described in the Molecular Dynamics Flexible Fitting (MDFF ) tutorial [159, 160]. For

crystal structures of various subcomplexes of the ATP synthase [56, 129, 41, 166], different

regions of the cryo-EM maps corresponding to the subcomplexes were first manually

segmented in UCSF Chimera. The crystal structures were then rigidly docked into the

map segments using the Fit in Map function in UCSF Chimera. For flexible fitting, a map

segment was converted to an MDFF potential using the command mdff griddx in VMD

[69]. Using the VMD plugin AutoPSF, the crystal structure to be fit into the density

was prepared for NAMD by generating a PSF file with information about structure

connectivity and partial charges. A PDB file with scaling factors for each atom was then

prepared with mdff gridpdb. Secondary structure restraints, including conservation of

peptide bonds in their cis/trans configuration in the given structure and conservation of

the hand of chiral centres, were applied to preserve the secondary structure of the crystal

structure during flexible fitting. Two consecutive NAMD configuration files were then

prepared for MDFF simulations. The first used a small scaling factor (-gscale option)

of 0.3 for 50 ps, and the second used a higher scaling factor of 10 for 2 ps for energy

minimization. Higher scaling factor values meant stronger forces were used to fit the

crystal structure into the map density. After the MDFF simulation was run using NAMD,

the trajectory was viewed in VMD. The final structure of the trajectory, representing the

fitted atomic structure, was saved as a PDB file to be used in analysis of conformational

changes between the different states.

Conformational changes between substates were analyzed by linear interpolation in

UCSF Chimera (Moviemaker). To approximate and quantify the change in conformation

observed between substates, the rotation of the α3β3 hexamer in the F1 region was

measured. This was done by matching the α3β3 hexamer of the fitted structure of the

first substate of a transition to the α3β3 hexamer of the fitted structure of the second

substate of the transition using the Fit in Map function in UCSF Chimera. Using the

measure rotation command, the magnitude of rotation, as well as the axis of rotation


between the first and second substates were determined.

To increase the SNR in the FO region, the FO regions of the seven maps were aligned

in UCSF Chimera with Fit in Map and averaged in real space using volaver in the Situs

package [167]. Local resolution of maps was assessed using ResMap with maps that were

downsized by a factor of 2 [83].

2.5 Building of atomic models

An atomic model of the a subunit was built by Daniel Schep using the a subunit map

density and analysis of evolutionary co-variance constraints. Co-varying residues in the a

subunit were determined using EVcouplings [65] without the assumption of transmembrane

α-helices and using all other default settings. The top 90 co-varying residue pairs were

used to build a atomic model of the a subunit. For analysis of co-varying residues between

the a subunit and c subunit, GREMLIN [118] was used with Jackhmmr to generate

multiple sequence alignments using an E-value threshold of 1 × 10−10.

Straight, ideal α-helices built in UCSF Chimera according to MEMSAT-SVM [113]

were manually arranged in the average a subunit density to satisfy co-variance constraints.

Loops to connect the α-helices were built using Modeller [47] in UCSF Chimera, and fit

into the a subunit map segment with MDFF. Rosetta [131] was used to model the loop

between residues 115 and 148 (loopmodel command [42]) with the quick ccd method, and

to idealize the bond lengths and angles in the output structure with the lowest energy out

of 100 output structures (idealize.jd2 command). The idealized model was then energy

minimized using UCSF Chimera.

The N terminus of the b subunit crystal structure [41] was extended by Anna Zhou

into the FO region by first building the two transmembrane α-helices in UCSF Chimera

based on transmembrane α-helix prediction from MEMSAT-SVM [113]. The α-helices

were docked rigidly into the b subunit map segment from the FO region, then connected


using Modeller. The extended b subunit structures were then fit into the maps using

MDFF as described in the previous section.

Calculations with Relion and MDFF were performed using the Scinet cluster [94] and

SickKids High Performance Computing Facility.

Chapter 3

Results and Discussion

3.1 Three rotational states of the ATP synthase

The resolution of the previous 18 A cryo-EM map of the bovine ATP synthase was limited

by the inability to separate different rotational positions of the central rotor within the

α3β3 catalytic hexamer, and did not allow for the unambiguous assignment of the three

αβ catalytic sites to different conformational states [14]. Here, three rotational states of

the ATP synthase in which the central rotor is rotated ∼120◦ relative to each other were

initially identified by 3D classification of 195,233 particle images (Fig. 3.1). The three

αβ catalytic pairs assume different conformations in accordance with rotor position in

these three states.

Further 3D classification identified two to three substates in each rotational state.

State 1 (43,039 particles) had two substates (States 1a and 1b), State 2 (48,053 particles)

had three substates (States 2a, 2b and 2c) and State 3 (46,257 particles) had two substates

(States 3a and 3b) (Fig. 3.2). The resolutions of the seven maps of the ATP synthase

were between 6.4 and 7.4 A (Fig. 3.3), with higher resolution in the F1 region compared

to the FO region (Fig. 3.4). The difference in resolution between the two regions was

expected due to the decrease in SNR in the FO region and blurring of the periphery of

37

Chapter 3. Results and Discussion 38

TightLoose Open

A

BState 1 State 2 State 3

Figure 3.1: Three main rotational states of the ATP synthase. A) Maps of states 1, 2 and 3. B)Map sections through the F1 region as indicated by the blue and purple arrows in (A). ∼120◦

turns of the two α-helices of the γ subunit in the central rotor (indicated by orange arrows) isseen in map sections boxed in blue, while the changes between loose, tight, and open statesof the three αβ catalytic sites is seen in map sections boxed in purple. The state of the αβcatalytic pair boxed in red is indicated. Scale bars 25 A.


State 1a6.7 Å20,104 particles (10.3%)

State 1b7.4 Å22,935 particles (11.7%)



State 2c7.4 Å18,899 particles (9.7%)



F1

FO

Figure 3.2: Seven maps of the ATP synthase. Each map shows the ATP synthase in a distinctconformation. All maps have the structural features discussed below, including a novel featureextending from the bent FO region. The bend is indicated by the dashed line. Percentagesare relative to the dataset of 195,233 particle images selected from 2D classification for 3Dclassification. Scale bar 25 A.


A

B

C

0.143

0

0.25

0.5

0.75

1

30 15 10 8 7 6 5 4

FSC

Resolution (Å)

state 1astate 1b

0.143

0

0.25

0.5

0.75

1

30 15 10 8 7 6 5 4

FSC

Resolution (Å)

state 2astate 2bstate 2c

0.143

0

0.25

0.5

0.75

1

30 15 10 8 7 6 5 4

FSC

Resolution (Å)

state 3astate 3b

Figure 3.3: Fourier shell correlation (FSC) curves for maps of each state. FSC curves for mapsof states 1a and 1b (A), states 2a, 2b and 2c (B), and states 3a and 3b (C).


4.0 4.8 5.6 6.4 7.2 8.0 8.8 9.6

detergent micellec8-ring

a subunit

Figure 3.4: Local resolution. Local resolution of State 1a as determined by Resmap [83] iscoloured according to the legend, showing higher resolution in the F1 region than in the FO

region. Scale bar 25 A.


membrane-bound proteins by the detergent micelle [87]. In addition, the extraction of the

ATP synthase from its native membrane environment may have decreased the stability of

the FO region, producing heterogeneity in this region that lowers resolution. The uneven

number of substates in each rotational state suggests that the conformations we were able

to refine to sub-nanometre resolution are a sampling of many conformations that may

exist in solution. Other conformations may not have been present in sufficiently large

populations in our dataset to be identified.

The ability to separate different rotational states and substates was possible in this

work due to the advances in direct electron detector technology and image processing

described in section 1.2.6. Because particle images were separated into near-homogeneous

datasets during image analysis, their orientations were refined more accurately, and the 3D

maps reached higher resolution than when image analysis incorrectly combines different

conformations into one class. The ability to separate distinct conformations present in

the sample also allowed for the analysis of dynamics of the ATP synthase, which will be

discussed further in section 3.5.

The particle images were distributed almost equally between the three rotational states,

unlike what was observed for the rotational states of the S. cerevisiae V-type ATPase

[172]. The near equal distribution of states may have been due to the IF11-60 inhibitor,

which would be expected to primarily stabilize the ATP synthase in three different states

depending on which of the αβ catalytic sites it binds to. Additionally, the presence of the

IF11-60 inhibitor may have limited the substates that were identified in the sample to

those that were most stable when it is bound. The addition of the IF11-60 inhibitor also

meant that the ATP synthase was not visualized during its catalytic cycle. Consequently,

the substates that we identified are not representative of catalytic intermediates of the

rotary cycle. They appear to be the result of Brownian motion based on the difference in

orientation of the c8-ring relative to the a subunit between substates (discussed further in

section 3.6), which suggests that they are conformations that are energetically similar or


State 1a State 1b

OSCP

β

αγεδ

c8-ring

F6

b

d

a

Figure 3.5: Atomic models of F1 and the peripheral stalk (PDB IDs 2WSS, 2CLY, 2XND[129, 41, 166]) and models of the a (green) and b (red-orange) subunits created in this thesiswere fit into maps of States 1a and 1b, showing the arrangement of all subunits except A6L, f, eand g in the intact ATP synthase. A region of unassigned density near the peripheral stalk isindicated by the black arrow. Scale bar 25 A.


equivalent.

At the resolutions of the maps, it was possible to fit crystal structures accurately

into map density for analysis of conformational changes (Fig. 3.5). Linear interpolation

between a combination of crystal structures fit into the maps of the three rotational states

(including the F1:peripheral stalk complex [129], peripheral stalk alone [41], F1:c-ring

complex [166], and atomic models of the a and b subunits built in this work and described

in sections 3.3 and 3.4) shows the large conformational changes in the central rotor and

catalytic α3β3 hexamer that occur during the full rotary cycle (Movie 1, see caption in

Appendix A). As the central rotor turns, concerted conformational changes are observed

in each of the three αβ catalytic sites. Each of the sites changes from what appear to

be the open (empty), to loose (ADP-binding), to tight (ATP-bound), then back to the

open conformation in the catalytic cycle. These changes are described by the binding

change mechanism as discussed in section 1.1.3. Large conformational changes in the

peripheral stalk can also be observed in this movie. Implications of this flexibility will

be discussed further in section 3.7. It is important to note that movies made by flexible

fitting of atomic structures into cryo-EM maps at sub-nanometre resolution followed by

linear interpolation may include small changes in conformation that may be artifacts of

the fitting. These changes can arise due to the low resolution in some areas of the map,

especially in the FO region, that allow for erroneous changes in crystal structures during

flexible fitting.

3.2 Membrane curvature and a novel feature in the

FO region

High membrane curvature of the inner mitochondrial membrane increases the surface area

that is available for oxidative phosphorylation and ATP synthesis. Electron tomography

of mitochondrial membranes showed that ATP synthases form rows of dimers that


induce high membrane curvature, producing characteristic cristae structures [153, 39,

38]. The dimerization interface is located in the FO region, between the peripheral

stalks of ATP synthase monomers. As was previously observed in the 18 A map and

membrane-reconstituted 2D crystals of the bovine ATP synthase [14, 71], as well as

electron tomograms of mitochondrial membranes [153, 39], the FO regions in our maps

have a bend in the portion farthest from the c8-ring where the dimerization interface

is expected to be located (Fig. 3.6A). This portion is thought to contain the e and g

subunits, since a corresponding density is not present in a map of the S. cerevisiae ATP

synthase lacking these subunits [14, 86]. Subunit f is expected to be located near subunits

e and g [18], and may also be located in this bent density.

Subunits e and g are associated with dimer formation and normal mitochondrial

morphology because their deletion was shown to result in the formation of balloon-like

cristae in yeast mitochondria [38]. The bent region of FO was measured in the 18 A map

to be ∼40◦ in relation to the long axis of the enzyme [14]. Since the e and g subunits

of dimerized ATP synthases have been biochemically shown to be in close contact [24]

and likely lying in the same plane, this measurement was in good agreement with the

80◦ angle measured between the long axes of dimerized ATP synthases in sub-tomogram

averages of intact bovine mitochondria [39] and further supported the role of subunits e

and g in dimerization.

Deletion of the e subunit [7], mutation of the GxxxG motif in the transmembrane

α-helix of either the e or g subunit [8, 27], or deletion of the first transmembrane α-helix

of the b subunit [146] results in the loss of the g subunit and abnormal mitochondrial

morphology. In contrast, deleting the g subunit or the first transmembrane α-helix of the b

subunit does not result in loss of the e subunit, but does result in abnormal morphological

phenotypes [7, 146]. These experiments suggested that the e and b subunits are involved

in anchoring the g subunit, and the g subunit is involved in maintaining normal cristae

structure. Subunit g is predicted to have three soluble α-helices in addition to its single


State 1a Average of FO region from all seven maps

A B C

Figure 3.6: Features in the FO region. A) The bent portion of FO region furthest from thec-ring is boxed in red. The novel feature extending from this portion and thought to be fromthe e subunit is indicated by the red arrows on the map and corresponding slices in (B). Slicesthrough the FO region of state 1a (B) and of the average FO region (C) show the increased SNRin the average, with better resolution of α-helices in the a subunit indicated by the blue arrows.A low density feature predicted to be from lipids or disordered protein is indicated by the whitearrows. Scale bars 25 A.


transmembrane α-helix [25, 75]. These amphipathic α-helices may embed themselves in

one membrane layer in a mechanism known to be used to induce membrane curvature by

proteins involved in vesicle trafficking [103], cell division [140, 145], and viral infection

[97, 134]. It was proposed in previous work [11] that the bending of the membrane by the

g subunit of monomeric ATP synthases would lead to strain in the membrane bilayer, and

the dimerization of ATP synthase would be favoured because it would reduce this strain.

In this model, dimerization would occur without the need for protein-protein interactions.

However, it remains unknown if protein-protein interactions mediate the formation of

ATP synthase dimers or higher order oligomers.

In this work, an extramembranous feature that has not been described previously for

maps of the ATP synthase [14, 4, 86, 137] is observed extending towards the c8-ring from

the bent FO density predicted to contain the e and g subunits (Fig. 3.6A). Previously,

a bridge-like feature was observed in bovine ATP synthase dimers in the same location

with negative-stain EM, and was proposed to be formed by the interaction between the

soluble, conserved coiled coil regions of the e subunits of two ATP synthase monomers

[48, 107]. While the novel feature in our maps extends from the density corresponding

to the expected location of the e subunit, its position in these maps does not suggest a

clear mechanism for dimerization. Thus, it can only be speculated that this feature is

involved in protein-protein interactions that form dimers, and perhaps also higher order

oligomers. The mechanism of dimerization is unclear and remains to be determined by

higher resolution crystal structures or EM maps of the dimerized complex.

3.3 The arrangement of subunits in the FO region

By aligning and averaging the FO region of the seven maps to increase the SNR of similar

features in the region (Fig. 3.6C), we were able to resolve secondary structure and

assign density to various membrane-bound subunits (Fig. 3.7). Assignment of density to


a subunitb subunit

A6L

e and g subunits

90˚ 90˚

Figure 3.7: Segmentation of the FO region. Segments for the a subunit (green), two transmem-brane α-helices from the b subunit (red), a transmembrane α-helix from the A6L subunit (blue),and the region predicted to contain the e and g dimerization subunits (orange) are shown. Thec8-ring is omitted for clarity. Scale bar 25 A.


individual subunits was done manually by first segmenting and removing the detergent

micelle in the maps, then identifying α-helices for each subunit based on biochemical

and structural data [4, 172, 90, 14, 88, 113]. This process was inexact because the DDM

micelle surrounding the FO region obscures the ends of transmembrane α-helices within

the detergent micelle. Also, α-helices in the dimerization region remained unresolved,

which meant that the secondary structure and locations of the e, g and f subunits could

not be determined. Despite these limitations, we were able to learn new information from

the segments presented in Figure 3.7.

The c8-ring was the easiest to segment because its density was mostly isolated from

that of the other FO subunits, except for a small contact point with the a subunit. This

limited contact between the c-ring and a subunit was also observed in the membrane-

bound regions of maps of other rotary ATPases [87, 14, 4]. Its importance is discussed

more in section 3.4. In the segment for the proton-translocating a subunit, we observed

two highly tilted α-helices that are in contact with the c8-ring (Fig. 3.7, green density),

as was also seen in the a subunits of the Polytomella sp. F-type ATP synthase, the S.

cerevisiae V-ATPase [4, 172], and most recently, the P. denitrificans ATP synthase [108].

This segment allowed us to predict and model the fold of the a subunit, which will be

described in the next section.

The average map of the FO region shows where the N terminus of the b subunit

enters the membrane region and forms two transmembrane α-helices in contact with the

a subunit, revealing exactly where the peripheral stalk is anchored in the FO region to

allow it to resist the torque from the rotor during catalysis (Fig. 3.7, red density). The

location of these b subunit α-helices shows that, in agreement with previous cross-linking

studies [40] and evolutionary co-variance analysis [66] (described in depth in the following

section) with the E. coli ATP synthase, it is in close proximity with the a and c subunits.

Density for two α-helices in the same position was also observed in the P. denitrificans

ATP synthase crystal structure [108] and was also attributed to the N termini of the b


and b’ subunits of its peripheral stalk.

The proximity of the b subunit to the a subunit/c-ring interface, along with studies

that have shown it is essential for proton translocation in bacterial ATP synthases, support

its involvement in the proton-translocating mechanism that generates rotor rotation in

FO. Comparison of the map density containing the a and b subunits with the maps of the

membrane-bound VO regions from yeast and T. thermophilus V-ATPases [86, 87] shows

that the arrangement of α-helices is conserved across rotary ATPases. As will be discussed

further in our model for proton translocation channels in section 3.5, this conservation

of structure also supports the role of the b subunit in forming half-channels for proton

translocation. Based on the segmented density, the two predicted transmembrane α-helices

from the b subunit were extended from the available crystal structure [41] in accordance

with transmembrane α-helix predictions [113] (Fig. 3.5A, red-orange subunit). The b

subunit is expected to have a sharp kink in its structure where it enters the FO region,

and where there are two of each of the helix-breaking residues, glycine and proline, in

its sequence. Indeed, a sharp bend in the peripheral stalk is observed in the 18 A map

[14] and in our maps. The predicted break in the α-helix was not modelled here since its

exact location is not known. High resolution structures of the intact enzyme will show

where it occurs.

An additional transmembrane α-helix that is in contact with the a subunit at a separate

interface from the b subunit is assigned to the A6L subunit due to its predicted proximity

to the peripheral stalk from cross-linking [90] (Fig. 3.7, blue density). In the bent FO

region, density for the e and g (and possibly f) subunits and the novel extramembranous

feature was segmented (Fig. 3.7, orange density). However, the resolution in this area did

not allow for a clear separation of protein from the detergent micelle due to their similar

densities (1.36 g/ml for protein [135] and 1.19 g/ml for DDM [158]), and the segment

likely contains some detergent density. Near the density from which the novel feature

extends, an additional low density feature is observed (Fig. 3.6B,C, white arrow). It is


unclear what this feature may be. It could arise from endogenous lipids remaining in

the complex after purification, or a disordered or flexible portion of a protein subunit.

Since the purification did not include the necessary phospholipids for the association of

DAPIT and 6.8 kDa proteolipid, these subunits are not expected to be present in the maps

described here [32, 31]. Structural studies with ATP synthase complexes that include

these subunits will be required to determine their locations relative to, and interactions

with, other FO subunits.

Higher resolution will be required to determine the structures of each of the subunits

in the FO region. Knowledge of the structures of individual subunits will show how

membrane curvature is generated by the FO region. It will also clarify the roles of the f

and A6L subunits, which have been suggested to be involved in organization and structural

support of the proton translocating subunits [175]. Notably, there was an additional

unassigned density in the seven maps that appeared to wrap around the peripheral stalk

just above the FO region and near where the b subunit enters the membrane (Fig. 3.5,

black arrow). This density may be from the d subunit, since the crystal structure of the

peripheral stalk [41] that was modelled here does not include the full subunit. It could

also be from either the f or A6L subunits that are predicted to be near the peripheral

stalk [90], as the apparent winding of the density around the base of the peripheral stalk

suggests that it could be involved in structural support and reinforcement. The origin of

this feature would be determined with improved resolution of the FO region.

3.4 An atomic model of the a subunit

An atomic model of the a subunit was built to fit into the segmented map density using

analysis of evolutionary co-variance. Evolutionary co-variance predicts that residues

that are in close proximity in a structure will evolve together in a way that maintains

energetically favourable interactions [65, 118]. Thus, sequences related by evolution can


be used to determine residue pairs that have evolved in concert, and that likely interact

in a protein structure. Co-variance constraints from this analysis can be used to orient

α-helices in a structural model by indicating which sides of the α-helices may have contacts

with each other. The use of this technique for the mitochondrial ATP synthase a subunit

was robust because thousands of related sequences (20,637) were used in the analysis. The

pattern of evolutionary couplings and the predicted fold of the bovine a subunit looked

similar to what was predicted for the E. coli a subunit [66] (not shown), indicating the

relevance of cross-linking and aqueous accessibility experiments done in the E. coli enzyme

[49] to the bovine enzyme. This analysis was informative for the proton translocation

model that is presented in section 3.5.

Using the top 90 co-variance constraints shown in Figure 3.8A, we could trace the

α-helical sequences unambiguously in the a subunit within the segmented density from

the average FO region map (Fig. 3.9A). Our model was further supported by evolutionary

co-variance analysis that predicted interactions between residues on the two highly tilted

α-helices of the a subunit expected to be in contact with the c8-ring [172, 4] and residues of

c subunit helix 2 that face outwards on the c8-ring (Fig. 3.8B). This a/c subunit covariance

analysis also supported our placement of the two α-helices in relation to the different

sides of the membrane, as described below. Our model violates 6% of the constraints

(defined here as having Cα-Cα distances greater than 15 A), which is within the expected

false discovery rate of the method [98]. Since the loops connecting the α-helices of the

model were built only to fit into the density as described in Materials and Methods, and

were not determined experimentally, they were not interpreted.

The model of the a subunit consists of six α-helices, five of which are transmembrane

α-helices (Fig. 3.9B). Because detergent molecules in the buffer adhere to hydrophobic

regions of membrane-embedded portions of protein complexes, the detergent micelle

present in our maps around the FO region indicates the relative location of the membrane

bilayer. The N terminus of the a subunit is located in the IMS. α-helix #1 spans the


A

B

a subunit

a subunit

c subunit helix 2

MNENLFTSFITPVILGLPLVTLIVLFPSLLFPTSNRLVSNRFVTLQ

QW

MLQ

LVSKQM

MSIH

NSKG

QTWTLM

LMSLILFIGSTNLLGLLPHSFTPTTQLSMNLGMAIPLWAGAVITGFRNKTKASLAHFLPQGTPTPLIPMLV

IIETISL

FIQPM

ALAV

RLTA

NIT

AGH

LLIH

LIG

GAT

LALM

SIST

TTAL

ITFT

ILIL

LTILE

FAVA

MIQ

AYVFTLLVSLYLHDNT1

50

100

15020

0

226

MNENLFTSFITPVILGLPLVTLIVLFPSLLFPTSNRLVSNRFVTLQ

QW

MLQ

LVSKQM

MSIH

NSKGQTW

TLMLM

SLILFIGSTNLLGLLPHSFTPTTQLSMNLGMAIPLWAGAVITGFRNKTKASLAHFLPQGTPTPLIPMLVIIE

TISLFIQ

PMAL

AVRL

TAN

ITAG

HLL

IHLI

GG

ATLA

LMSI

STTT

ALIT

FTIL

ILLT

ILEFA

VAMIQ

AYVFTLLVSLYLHDNT

1

50

100

15020

0

226

Transmembrane prediction(MEMSAT-SVM)

LFSYAILG

FALSEA

MG

LFCL MV A

FLI LF

46

72

Highly conserved residues

Figure 3.8: Evolutionary co-variance constraints. A) The top 90 evolutionary co-varianceconstraints between residues in the a subunit (shown in a numbered circle) are indicated by blacklines. Transmembrane α-helices predicted in the a subunit by the MEMSAT-SVM algorithm[113] are indicated in green in the sequence, and by the dark blue rectangles in the circular bardiagram surrounding the a subunit sequence. A soluble α-helix is indicated by the light bluerectangle. B) The top six evolutionary co-variance constraints between residues in the a subunitand the outward-facing α-helix of the c subunit are indicated by black lines. Highly conservedresidues in the a subunit are in red. This figure was created by Daniel Schep and is published ineLife [173].


A B

C terminus

N terminus

Matrix

IMS

1

2 3

4

6

5

Figure 3.9: Model of the a subunit. A) The top 90 evolutionary co-variance constraints betweenresidues in the a subunit (green) are indicated by red lines. B) The model of the a subunit iscoloured from blue (N terminus) to red (C terminus). The α-helices in the model are numberedfrom 1 to 6. Arg159 on α-helix #5 is coloured blue. The approximate boundaries of the lipidbilayer are indicated by black lines. IMS, intermembrane space. Scale bar 25 A.


lipid bilayer perpendicular to the membrane plane. It is followed by α-helix #2, which

travels along the micelle surface. Moving in the direction towards the c8-ring, α-helix #2

is followed by a tilted hairpin consisting of α-helices #3 and #4. This hairpin does not

completely cross the lipid bilayer, as was observed for the Polytomella sp. F-type ATP

synthase [4]. The structure ends with a longer hairpin that is in contact with the c8-ring

consisting of α-helices #5 and #6, which is analogous to the highly tilted hairpin observed

in the S. cerevisiae V-type ATPase [172] and Polytomella sp. F-type ATP synthase [4] a

subunits. The C terminus is located in the matrix. The bundle of four α-helices created

by the two hairpins was also recently observed in a 4.0 A crystal structure of the P.

denitrificans ATP synthase [108], and the same sequence of α-helices was assigned. The

full a subunit was not resolved in this crystal structure and, as in our maps, did not show

side-chain density.

Our model places the conserved Arg159 that is essential for proton translocation in

the middle of α-helix #5 near the IMS side of the membrane, in contrast to the proposed

model of the Polytomella sp. ATP synthase a subunit [4]. It is near the predicted

location of the Arg159 residue that the a subunit has a small point of contact with the

c8-ring. This, as discussed further in section 3.7, is important to the function of the ATP

synthase. Firstly, the minimal contact between the a subunit and the c8-ring allows for

more water and lipid molecules to be inserted that can induce Brownian oscillations in the

c8-ring. Fewer protein-protein interactions can also mean there is less friction to hinder

the movement of the c8-ring, contributing to more smooth and rapid rotor rotation.

3.5 A model for proton translocation

The a subunit, b subunit and c-ring are the only subunits essential for proton translocation

in the minimal ATP synthase structures found in bacteria [30, 59, 67], suggesting that

these subunits may also be sufficient for forming the channels for proton translocation in


eukaryotic ATP synthases. However, the path of proton movement remains unclear. As

described in section 1.1.5, the Brownian ratchet mechanism for generation of rotation by

proton translocation is contingent on two offset half-channels in the FO region [77, 78].

Brownian fluctuations from water and lipid molecules in contact with the c-ring, and

possibly also from protein-protein interactions, are proposed to cause small, random

oscillations of the c-ring. Rotation of the c-ring by a full c subunit occurs upon binding

of a proton to a conserved acidic residue located near the centre of the membrane bilayer

when it is aligned with one aqueous half-channel. Protons are released to the other side of

the lipid bilayer through the other half-channel. From the Polytomella sp. ATP synthase

structure, it was proposed that the aqueous half-channel on the lumenal side (equivalent

to the IMS side in mitochondria) is formed in part by α-helices of the a subunit, while the

half-channel on the matrix side involves an interface between the a subunit and c-ring [4].

This is consistent with a previous model for the location of the half-channels proposed

based on cysteine mutation and ion accessibility studies for the E. coli ATP synthase a

and c subunits [49]. Here, we propose a model in which the transmembrane α-helices of

the b subunit also participate in proton translocation, in addition to the a subunit and

c-ring. This model is compelling because of the striking conservation of the architecture

of the bovine ATP synthase FO region with that of the S. cerevisiae V-ATPase VO region,

despite the lack of sequence similarity of the bovine ATP synthase a subunit to the

V-ATPase a subunit [172].

In this model (Fig. 3.10), the opening of the matrix half-channel is formed by the

matrix ends of a subunit α-helices #5 and #6 at the interface with the c8-ring, as was

proposed for the Polytomella sp. and E. coli ATP synthases [4, 49]. The opening of the

IMS half-channel is formed at the interface of the IMS ends of the b subunit transmembrane

α-helices and a subunit α-helices #5 and #6. These openings would channel protons

towards and away from where the acidic Glu58 residues on the c8-ring interact with the

conserved Arg159 residue on the a subunit, near the middle of the membrane bilayer.


H+ H+

Matrix

IMS

A B

half-channel

1half-channel

2

Figure 3.10: Proposed proton translocation channels. Arrows indicate a potential path of protontranslocation during ATP synthesis. The intermembrane space half-channel, consisting of thetwo transmembrane α-helices from the b subunit and the intermembrane space ends of a subunitα-helices #5 and 6, is in red. The matrix half-channel, consisting of the matrix ends of a subunitα-helices #5 and 6 at their interface with the c8-ring, is in purple. The conserved a subunitArg159 is indicated in blue, and the c8-ring is shown in a pink surface representation. Theapproximate boundaries of the lipid bilayer are indicated by black lines. IMS, intermembranespace. Scale bar 25 A.


Notably, human disease-causing mutations identified in the a subunit to date are all

located on α-helices #5 and #6 in our model, near the proposed half-channel openings

(Fig. 3.11). Conserved residues in the hairpin composed of α-helices #3 and #4, and

not just in α-helices #5 and #6, are also likely to participate in proton translocation

since the bundle of four α-helices is highly conserved in rotary ATPases. High-resolution

structures by cryo-EM or X-ray crystallography are ultimately required to determine the

specific residues that are involved in proton translocation.


Leu220Leu222Leu217

Leu156

Ser148

Thr192Arg159

Matrix

IMS

Figure 3.11: Disease-causing mutations in the a subunit. Mutations in the a subunit that havebeen identified in Neuropathy, ataxia, and retinitis pigmentosa (NARP), Leber hereditary opticneuropathy (LHON), Leigh syndrome (LS), Mitochondrial infantile bilateral striatal necrosis(MIBSN), Mitochondrial complex V deficiency, mitochondrial 1 (MC5DM1) and Myopathy,lactic acidosis, and sideroblastic anemia 3 (MLASA3) are coloured red on our atomic model ofthe a subunit [74, 26, 36, 85, 157]. The sequences of the human and bovine a subunits are highlyconserved at the C-terminal α-helices. The strictly conserved a subunit Arg159 is indicatedin blue. The approximate boundaries of the lipid bilayer are indicated by black lines. IMS,intermembrane space. Scale bar 25 A.


3.6 Conformational changes between substates

Using linear interpolation of the flexibly fitted crystal structures described in section 3.1,

the changes in conformation during transitions between substates of each rotational state

were analyzed to determine regions of flexibility in the ATP synthase. Conformational

changes were observed in nearly all of the subunits during the transitions between substates,

as was observed in transitions between the three rotational states of the S. cerevisiae

V-ATPase [172]. In the peripheral stalk specifically, there were two main regions of

flexibility: one close to the OSCP subunit near the C terminus of the b subunit, and one

close to the FO region. During the transition between states 1a and 1b (Movie 2, panel

A), and between states 2a and 2b (Movie 2, panel B), a bending of the peripheral stalk

is observed near the OSCP subunit, while between states 2b and 2c (Movie 2, panel C),

and 3a and 3b (Movie 2, panel D), a bending of the peripheral stalk is observed near the

FO region. Movie 3 shows the same transitions between the substates viewed from the F1

region towards the FO region. Here, again, it is seen that the transitions between states

1a and 1b (Movie 3, panel A) and states 2a and 2b (Movie 3, panel B) are similar, and

the transitions between states 2b and 2c (Movie 3, panel C) and states 3a and 3b (Movie

3, panel D) are similar.

From the previous 18 A map of the bovine ATP synthase [14], two contact points

between the peripheral stalk and the F1 region, other than the interaction of the OSCP

subunit with the N terminus of the α subunit farthest from the peripheral stalk, were

observed. One of these contact points was near the N terminus of the closest α subunit,

and the second was halfway along the interface of a non-catalytic αβ pair. It was found

that docking of a crystal structure with a truncated peripheral stalk [129] into the 18

A map required flexibility in the peripheral stalk somewhere between these two contact

points. This finding is consistent with the flexible region near the OSCP subunit observed

in our analysis of conformational changes between substates. In addition, the two regions

of flexibility that we identified correspond to the two poorly resolved regions in the


peripheral stalk of the P. denitrificans ATP synthase crystal structure [108], which further

confirmed that the regions are mobile.

The rotation of the α3β3 hexamer in the F1 region in relation to the rest of the

enzyme could be used to approximate and quantify the conformational changes observed

between substates (Fig. 3.12). The magnitude of the rotations ranged between 10◦ and

16◦. Although the axes of rotation for the α3β3 hexamer in the four transitions were

all different, the axes for the movements from state 1a to 1b and from state 2a to 2b

are similar, and the axes for the movements from state 2b to 2c and from state 3a to

3b are similar. The first two movements (Fig. 3.12A and B) have tilted axes that pass

through the central rotor and peripheral stalk, and the second two movements have axes

that are almost parallel to the long axis of the enzyme and near the central rotor (Fig.

3.12C and D). Differences in the axes of rotation of the α3β3 hexamer for what appear

to be overall similar movements in Movies 2 and 3 are expected because of the different

positions of the central rotor. In addition, the conformations captured in our maps are

likely a sampling of a continuum of conformations that the ATP synthase can assume in

solution, and they may not capture the full range of the observed motions, or all possible

kinds and combinations of motions.

Although only two points of flexibility in the peripheral stalk were obvious from the

analysis of our maps, this does not exclude the possibility of other regions of flexibility

that also contribute to smoothing catalysis. The points that we observed may have simply

been the most pronounced for the states that we were able to identify. The ability to

resolve more states may reveal other points of flexibility, and can also reveal a larger range

of motion than was seen in our analysis. Higher resolution for the current states and

other states will also mean that more subtle motions can be detected and flexibility can

be described more accurately. The use of linear interpolation between atomic structures

fit into cryo-EM maps will likely become more prevalent as a means to explore protein

dynamics as it becomes easier to separate different conformations of a protein within a


A B D

C

States 1a and 1b States 2a and 2b

States 2b and 2c

States 3a and 3b

Figure 3.12: Differences between substates. The differences between substates are shown byoverlaying their 3D maps, as well as their corresponding fitted atomic structures. The axis ofthe movement of the α3β3 hexamer used to approximate the overall movement is indicated as ablack rod. There is a rotation of 10◦ between states 1a (red) and 1b (green) (A), 11◦ betweenstates 2a (red) and 2b (green) (B), 12◦ between states 2b (red) and 2c (green) (C), and 16◦

between states 3a (red) and 3b (green) (D). Scale bar 25 A.


single sample by cryo-EM.

3.7 Implications of flexibility in the ATP synthase

Flexibility in the peripheral stalk may be involved in accommodating conformational

changes in the α and β subunits during the catalytic cycle to improve enzymatic efficiency.

In addition, it may be a site of transient energy storage while resisting torque generated

by the rotation of the rotor within the stationary F1 region. The observed flexibility in the

peripheral stalk of the mitochondrial ATP synthase is different from what was observed

for the E. coli ATP synthase, in which the peripheral stalk is composed of a homodimer

of b subunits [45], and also from what was observed for the T. thermophilus V/A-type

ATP synthase, in which the two peripheral stalks are each composed of two different

subunits that form a right-handed coiled coil around each other [91]. For the E. coli ATP

synthase, single molecule magnetic bead experiments showed that the peripheral stalk

was rigid in comparison to the central rotor [165], and for T. thermophilus, the agreement

of the crystal structure of the peripheral stalk [91] with the cryo-EM map of the intact

enzyme [87, 88] also suggested a rigid structure. Because the composition of the bovine

ATP synthase is very different from these organisms, with four distinct subunits that

interact with each other to form a complex structure, it is possible that flexibility in the

peripheral stalk is a feature that evolved in obligate aerobes for more efficient catalysis

compared to archael and bacterial enzymes. The analysis of flexible regions through linear

interpolation between fitted atomic models of substates was also extended to the rotor,

which appeared to be much more rigid than the peripheral stalk. Higher resolution and

biophysical experiments will be required to confirm this result.

In addition to the conformational changes observed in the peripheral stalk, the c8-

ring rotates slightly against the a subunit during transitions between substates of each

rotational state. Since this motion is smaller than the conformational changes observed


elsewhere in the enzyme during the transitions, it is unlikely that it indicates a disruption

of the interface of the c8-ring and the a subunit in the sample. Rather, it further illustrates

the inherent flexibility of the enzyme. In addition, the rotation of the c8-ring against

the a subunit between substates, even in sample conditions that limit rotary catalysis,

indicates a lack of rigid interaction between the c8-ring and a subunit that is consistent

with the Brownian ratchet model. In this model, as described in section 1.1.5, due to

Brownian motion, the c8-ring oscillates as a proton competes with the conserved a subunit

Arg159 residue to interact with a c subunit Glu58 residue. The c8-ring only rotates when

a proton binds a Glu58 residue. Movie 4 shows this rotational oscillation of the c8-ring

between states 2a and 2c, with the Glu58 residues on the c8-ring changing between the

proton-locked conformation [123] in the lipid bilayer to the open conformation [122, 156]

near the proposed locations of the two half-channels and the conserved Arg159 on the a

subunit.

While the c8-ring oscillates between substates, the α-helices of the γ subunit in the

central rotor do not change their positions within the α3β3 complex. This suggests the

presence of flexibility in the rotor that can be involved in smoothing its rotation against

the stationary catalytic region during ATP synthesis, which has also been proposed

previously by molecular simulations and mechanical modeling of the γ subunit [115].

From this compliance in the central rotor, and our observation of two points of flexibility

in the peripheral stalk during the transitions between substates (Movies 2 and 3), it

appears that both the central rotor and peripheral stalk may contribute to the elastic

coupling of the F1 and FO domains for smooth power transition. Due to the symmetry

mismatch between the three-step motor of the α3β3 complex in the F1 region and the

eight-step motor of the c8-ring in the FO region, such compliance and elasticity in the

structure of the ATP synthase may enhance enzymatic efficiency [172, 174, 115, 165].

In addition to dimerization, the subunits in the FO region have been shown to

facilitate the opening of the mitochondrial permeability transition pore (MPTP), an


inducible, non-specific pore located in the mitochondrial inner membrane [3, 53]. MPTP

opening leads to loss of membrane potential and cell death; it is thus implicated in

conditions with altered cell death, including muscular dystrophy, ischemic heart disease,

and neurodegenerative diseases [19]. The flexibility we observed in the peripheral stalk,

and throughout the structure of the ATP synthase, supports recently proposed models in

which the mitochondrial ATP synthase forms the MPTP. In one model, the dissociation of

the peripheral stalk from the FO region occurs upon binding of various MPTP modulators

to the ATP synthase. This dissociation results in uncoupling of the F1 region from the

c-ring, and allows the c-ring to expand and form the pore [73]. A second model proposes

that the MPTP is formed by ATP synthase dimers at the interface of the monomers.

The pore opens when Ca2+ binds to the catalytic sites in the α3β3 complex in place of

Mg2+ and causes conformational changes [19]. While the details of these specific models

remain to be tested, it has been shown that ATP synthase activity can be regulated

by the binding of allosteric modulators, as well as by post-translational modifications

[72, 150, 163, 19]. Thus, the flexibility we observed in the structure of the ATP synthase

may contribute to conformational changes that are involved in both the opening of the

MPTP and regulation of catalysis, in addition to smoothing the transition of power

between the mismatched F1 and FO motors.

Chapter 4

Conclusions and future directions

4.1 Conclusions

In this thesis, we presented cryo-EM maps of the intact bovine ATP synthase in seven

distinct conformations. A novel feature was shown to extend from the portion of the

FO region predicted to be involved in dimerization. Analysis of conformational changes

between the substates within each of the three rotational states indicated two main

regions of flexibility in the peripheral stalk that may contribute to the elastic coupling

of the F1 and FO motors during enzymatic activity. Additionally, an oscillation of the

c8-ring relative to the a subunit within each rotational state supports the Brownian

ratchet mechanism for generation of rotation in the FO region. Averaging the FO regions

from the seven maps revealed the locations of several membrane-bound subunits, as

well as the anchor point for the peripheral stalk in the FO region. Segmentation of the

proton-translocating a subunit from the average FO map allowed for the determination of

its fold when combined with analysis of evolutionary co-variance constraints. An atomic

model of the a subunit, along with the location of two transmembrane α-helices from the

b subunit, suggests two potential half-channels for proton translocation, also supporting

the Brownian ratchet mechanism of ATP synthesis.

66

Chapter 4. Conclusions and future directions 67

Several novel components were presented. A novel extramembranous feature in the

FO region was identified that is postulated to be involved in dimerization of the ATP

synthase. How the b subunit anchors the peripheral stalk in the membrane region and

how it may contribute to proton translocation was revealed. The first model for the fold

of the proton-translocating a subunit of a rotary ATPase was produced by combining

cryo-EM with analysis of evolutionary co-variance. Dynamics of the intact mitochondrial

ATP synthase were explored using cryo-EM through the separation of several distinct

conformations during image processing. This new information contributes to the overall

understanding of the function of the ATP synthase, from ATP synthesis to maintenance

of mitochondrial morphology.

4.2 Strategies for higher resolution of the intact ATP

synthase

An atomic or near-atomic model for the intact mitochondrial ATP synthase by cryo-EM

remains elusive. Several avenues through which this may be achieved in the near future

will now be discussed.

Map resolution depends on image quality, which determines the ability to recover

high resolution information and the accuracy of alignment of particle images during map

refinement. When imaging membrane proteins in detergent buffer, there is often low SNR

in particle images because protein-detergent complexes are most stable, and aggregation

is avoided at detergent concentrations close to the critical micelle-forming concentration

(CMC). At this concentration, detergent molecules produce high background noise in

microscope images. The use of detergent also decreases the ability to control ice thickness

due to the reduction in surface tension of the buffer, further lowering the contrast of

images with thick ice. There is often a thin layer of buffer in the middle of carbon


holes that pushes protein into areas of thicker buffer at the edges of the holes. Because

the reduction in surface tension results in such uneven particle distribution, it can also

be difficult to image uniformly distributed and isolated protein particles in detergent

buffer. The quality of images of membrane proteins can be improved by using detergent

alternatives such as amphipols, which are amphipathic polymers that are able to stabilize

membrane proteins in solution but do not have the same detrimental effects to image

quality as detergent [125]. Detergents with lower CMCs can also be used that allow for

the removal of almost all free detergent molecules from the sample, thus mitigating the

effects of background noise and lowered surface tension. Other strategies to improve

particle image quality include the use of gold grids that reduce the crinkling of the carbon

support layer and the beam-induced motion of particles in ice [139], and per-particle CTF

estimation and correction, a method that is currently being explored in our laboratory.

While the IF1 inhibitor in our sample appeared to stabilize the three rotational states

of the ATP synthase, many conformational changes could still occur. The flexibility of

the ATP synthase structure that was observed in this work likely limited the resolution

of our maps. In order to improve the resolution achievable by single particle cryo-EM,

the sample can be altered in ways that fix the ATP synthase in one or a few distinct

states, rather than allowing for a continuum of conformations. Reduction in the number

of conformations present simultaneously in the sample can be achieved with cross-linking,

mutagenesis, or the addition of various substrate analogues or inhibitors that lock the

enzyme in a small number of states.

In an opposite approach, to determine the structure of many different conformations

in a heterogeneous sample at high resolution, the collection of large datasets will be

required. This will become more feasible and efficient with the ongoing development of

microscopes that streamline the cryo-EM pipeline with automated specimen-loading and

data collection capabilities. The maskmicrograph program that was written in this thesis

for masking carbon and ice contamination prior to automatic particle image selection


may no longer be required when there are improved automated data collection schemes

that are able to avoid imaging carbon.

4.3 Unanswered questions

Several unanswered questions about the ATP synthase will now be discussed, which may

be addressed by using the strategies for reaching near-atomic resolution discussed above.

4.3.1 The catalytic cycle of the ATP synthase

Various models for the catalytic cycle in the F1 region currently exist based on studies

of the hydrolytic pathway (with assumption of its reversibility during ATP synthesis).

While three 120◦ steps in the full 360◦ cycle have been resolved in both the bacterial

and human enzymes [169, 154], different sizes and numbers of substeps between the

catalytic dwells that correspond to substrate release and product binding have been

proposed to make up each of the 120◦ incremental steps. These differences exist because

of the difficulty of capturing intermediate catalytic states of the enzyme by crystallization,

and of correlating available crystal structures to the states observed in single molecule

experiments [112, 2, 6, 101, 161]. The substeps during ATP hydrolysis for the human ATP

synthase (and likely also the bovine ATP synthase based on its high sequence identity and

crystal structure [1]) have been observed to be different from the eubacterial and bacterial

enzymes. In the human enzyme, three dwell positions were identified: 0◦ for ATP binding,

65◦ for Pi release and 90◦ for ATP hydrolysis [154]. In contrast, for the E. coli, Bacillus

PS3 and yeast enzymes, there were two dwells identified: 0◦ for ATP binding, and 80-90◦

for ATP hydrolysis [112, 169, 168, 111, 2, 149, 110, 117, 20, 148, 100]. The order of

phosphate/ADP release during ATP hydrolysis also appears to differ between human

and bacterial enzymes [16, 154, 79]. Furthermore, different types of rotary ATPases

have different catalytic cycles. For example, substeps were not observed within the 120◦


incremental steps of the Na+-pumping V-ATPase of Enterococcus hirae [106].

With the ability to automatically acquire large cryo-EM datasets and to accurately

align and classify particle images with high SNR from direct electron detectors, it will be

possible to resolve all catalytic intermediate states of the ATP synthase at high resolution

and reveal the occupancy of the αβ catalytic sites at each catalytic dwell. Unlike the

sample used in this thesis, which included an inhibitor that limited the conformations

that were present and also meant that our maps did not represent catalytically relevant

states, experiments to visualize catalytic intermediates will need to be done with active

enzymes. The application of this approach to the study of ATP synthases from different

organisms and to various types of rotary ATPases will allow for a thorough comparison of

their catalytic cycles.

4.3.2 Dynamics of the intact ATP synthase

Another area that remains to be explored in depth is the dynamics of the ATP synthase

during its catalytic cycle. Although our work suggested that both the rotor and peripheral

stalk are flexible and contribute to the smooth transition of power between the F1 and

FO regions, biophysical experiments and molecular simulations with the bovine enzyme

can be used to confirm this. Dynamics can also be probed in more detail if many

conformations of the uninhibited ATP synthase are resolved at high resolution, and

linear interpolation between fitted atomic structures is used to reveal precisely where the

regions of flexibility in the active ATP synthase are located. For this type of analysis,

large cryo-EM datasets will be required to resolve the different conformations in the

sample at sub-nanometre resolution. A large number of particle images means that

classification and separation of the dataset into homogeneous populations for refinement

will not result in a resolution-limiting reduction in the number of particle images for each

conformation. Dynamics can then be explored with confidence that the resulting maps

represent meaningful conformations.


4.3.3 Structure of FO subunits

At the resolution of our maps, it is still unknown which residues in the a, b and c subunits

are involved in proton translocation. Elucidating this would require structures of the FO

region at high enough resolution to observe bound water molecules that may indicate the

path of proton translocation [68]. In addition, although we were able to assign density

in the average FO region to different subunits (a, b and A6L), side chains and specific

interactions between subunits were not resolved. Improved algorithms and/or detectors in

cryo-EM may make it possible to determine the structures of individual membrane-bound

subunits or subcomplexes of subunits to near-atomic or atomic resolution in the near

future. Alternatively, X-ray crystallography or NMR can be employed. The resolution

of the maps from this thesis would allow atomic structures to be fit unambiguously

into a complete mosaic structure of the FO region. In addition to revealing the proton

translocation channels, high resolution information for the FO region will help elucidate

the yet unknown functions of several membrane-bound subunits, and provide insight

into how each of these proteins may support both the structure and function of the

proton-translocating subunits. Structures of the e and g subunits will also reveal how the

FO region induces membrane curvature and the mechanism of dimerization. These will

be important advancements in completing the structural picture of the ATP synthase.

Bibliography

[1] Abrahams, J. P., Leslie, A. G., Lutter, R., & Walker, J. E. Structure at 2.8 A

resolution of F1-ATPase from bovine heart mitochondria. Nature, 370(6491):621–628,

1994.

[2] Adachi, K., Oiwa, K., Nishizaka, T., Furuike, S., Noji, H., Itoh, H., Yoshida, M.,

& Kinosita, K. Coupling of Rotation and Catalysis in F1-ATPase Revealed by

Single-Molecule Imaging and Manipulation. Cell, 130(2):309–321, 2007.

[3] Alavian, K. N., Beutner, G., Lazrove, E., Sacchetti, S., Park, H. A., Licznerski,

P., Li, H., Nabili, P., Hockensmith, K., Graham, M., Porter Jr, G. A., & Jonas,

E. A. An uncoupling channel within the c-subunit ring of the F1FO ATP synthase

is the mitochondrial permeability transition pore. Proc. Natl. Acad. Sci. U. S. A.,

111(29):10580–10585, 2014.

[4] Allegretti, M., Klusch, N., Mills, D. J., Vonck, J., Kuhlbrandt, W., & Davies, K. M.

Horizontal membrane-intrinsic alpha-helices in the stator a-subunit of an F-type

ATP synthase. Nature, 521(7551):237–240, 2015.

[5] Allen, R. D., Schroeder, C. C., & Fok, A. K. An investigation of mitochondrial inner

membranes by rapid-freeze deep-etch techniques. J. Cell Biol., 108(6):2233–2240,

1989.

72

Bibliography 73

[6] Ariga, T., Muneyuki, E., & Yoshida, M. F1-ATPase rotates by an asymmetric,

sequential mechanism using all three catalytic subunits. Nat. Struct. Mol. Biol.,

14(9):841–846, 2007.

[7] Arnold, I., Pfeiffer, K., Neupert, W., Stuart, R. A., & Schagger, H. Yeast mitochon-

drial F1F0-ATP synthase exists as a dimer: identification of three dimer-specific

subunits. EMBO J., 17(24):7170–7178, 1998.

[8] Arselin, G., Giraud, M. F., Dautant, A., Vaillier, J., Brethes, D., Coulary-Salin,

B., Schaeffer, J., & Velours, J. The GxxxG motif of the transmembrane domain of

subunit e is involved in the dimerization/oligomerization of the yeast ATP synthase

complex in the mitochondrial membrane. Eur. J. Biochem., 270(8):1875–1884, 2003.

[9] Arselin, G., Vaillier, J., Salin, B., Schaeffer, J., Giraud, M. F., Dautant, A., Brethes,

D., & Velours, J. The modulation in subunits e and g amounts of yeast ATP synthase

modifies mitochondrial cristae morphology. J. Biol. Chem., 279(39):40392–40399,

2004.

[10] Bai, X. C., Yan, C., Yang, G., Lu, P., Ma, D., Sun, L., Zhou, R., Scheres, S. H., &

Shi, Y. An atomic structure of human gamma-secretase. Nature, 525(7568):212–217,

2015.

[11] Baker, L. A. The structure of the bovine mitochondrial ATP synthase by single

particle electron cryomicroscopy. Ph.D. thesis, University of Toronto, 2012.

[12] Baker, L. A. & Rubinstein, J. L. Angle determination for side views in single

particle electron microscopy. J. Struct. Biol., 162(2):260–270, 2008.

[13] Baker, L. A., Smith, E. A., Bueler, S. A., & Rubinstein, J. L. The resolution depen-

dence of optimal exposures in liquid nitrogen temperature electron cryomicroscopy

of catalase crystals. J. Struct. Biol., 169(3):431–437, 2010.

Bibliography 74

[14] Baker, L. A., Watt, I. N., Runswick, M. J., Walker, J. E., & Rubinstein, J. L. Ar-

rangement of subunits in intact mammalian mitochondrial ATP synthase determined

by cryo-EM. Proc. Natl. Acad. Sci. U. S. A., 109(29):11675–11680, 2012.

[15] Bason, J. V., Montgomery, M. G., Leslie, A. G. W., & Walker, J. E. Pathway

of binding of the intrinsically disordered mitochondrial inhibitor protein to F1-

ATPase. Proc. Natl. Acad. Sci. U. S. A., 111(31):11305–11310, 2014. URL http:

//www.ncbi.nlm.nih.gov/pubmed/25049402.

[16] Bason, J. V., Montgomery, M. G., Leslie, A. G. W., & Walker, J. E. How release of

phosphate from mammalian F ¡sub¿1¡/sub¿ -ATPase generates a rotary substep.

Proc. Natl. Acad. Sci., 112(19):201506465, 2015. URL http://www.pnas.org/

lookup/doi/10.1073/pnas.1506465112.

[17] Bason, J. V., Runswick, M. J., Fearnley, I. M., & Walker, J. E. Binding of the

inhibitor protein IF1 to bovine F 1-ATPase. J. Mol. Biol., 406(3):443–453, 2011.

URL http://dx.doi.org/10.1016/j.jmb.2010.12.025.

[18] Belogrudov, G. I., Tomich, J. M., & Hatefi, Y. Membrane topography and near-

neighbor relationships of the mitochondrial ATP synthase subunits e, f, and g. J.

Biol. Chem., 271(34):20340–20345, 1996.

[19] Bernardi, P., Di Lisa, F., Fogolari, F., & Lippe, G. From ATP to PTP and Back: A

Dual Function for the Mitochondrial ATP Synthase. Circ. Res., 116(11):1850–1862,

2015.

[20] Bilyard, T., Nakanishi-Matsui, M., Steel, B. C., Pilizota, T., Nord, A. L., Hosokawa,

H., Futai, M., & Berry, R. M. High-resolution single-molecule characterization of the

enzymatic states in Escherichia coli F1-ATPase. Philos. Trans. R. Soc. Lond. B. Biol.

Sci., 368(1611):20120023, 2013. URL http://rstb.royalsocietypublishing.

org/content/368/1611/20120023.abstract.

Bibliography 75

[21] Bowler, M. W., Montgomery, M. G., Leslie, A. G. W., & Walker, J. E. Ground

state structure of F1-ATPase from bovine heart mitochondria at 1.9 ?? resolution.

J. Biol. Chem., 282(19):14238–14242, 2007.

[22] Boyer, P. D. The ATP synthase–a splendid molecular machine. Annu. Rev. Biochem.,

66:717–749, 1997.

[23] Brilot, A. F., Chen, J. Z., Cheng, A., Pan, J., Harrison, S. C., Potter, C. S.,

Carragher, B., Henderson, R., & Grigorieff, N. Beam-induced motion of vitrified

specimen on holey carbon film. J. Struct. Biol., 177(3):630–637, 2012.

[24] Brunner, S., Everard-Gigot, V., & Stuart, R. A. Su e of the yeast F1F0-ATP

synthase forms homodimers. J. Biol. Chem., 277(50):48484–48489, 2002.

[25] Buchan, D. W. a., Minneci, F., Nugent, T. C. O., Bryson, K., & Jones, D. T.

Scalable web services for the PSIPRED Protein Analysis Workbench. Nucleic Acids

Res., 41(Web Server issue):349–357, 2013.

[26] Burrage, L. C., Tang, S., Wang, J., Donti, T. R., Walkiewicz, M., Luchak, J. M.,

Chen, L. C., Schmitt, E. S., Niu, Z., Erana, R., Hunter, J. V., Graham, B. H.,

Wong, L. J., & Scaglia, F. Mitochondrial myopathy, lactic acidosis, and sideroblastic

anemia (MLASA) plus associated with a novel de novo mutation (m.8969G¿A) in

the mitochondrial encoded ATP6 gene. Mol. Genet. Metab., 113(3):207–212, 2014.

URL http://dx.doi.org/10.1016/j.ymgme.2014.06.004.

[27] Bustos, D. M. & Velours, J. The modification of the conserved GXXXG motif of the

membrane-spanning segment of subunit g destabilizes the supramolecular species of

yeast ATP synthase. J. Biol. Chem., 280(32):29004–29010, 2005.

[28] Cabezon, E., Butler, P. J. G., Runswick, M. J., & Walker, J. E. Modulation of the

oligomerization state of the bovine F1-ATPase inhibitor protein, IF1, by pH. J.

Biol. Chem., 275(33):25460–25464, 2000.

Bibliography 76

[29] Cabezon, E., Runswick, M. J., Leslie, A. G., & Walker, J. E. The structure

of bovine IF(1), the regulatory subunit of mitochondrial F-ATPase. EMBO J.,

20(24):6990–6996, 2001.

[30] Cain, B. D. & Simoni, R. D. Impaired proton conductivity resulting from mutations

in the a subunit of F1F0 ATPase in Escherichia coli. J. Biol. Chem., 261(22):10043–

10050, 1986.

[31] Carroll, J., Fearnley, I. M., Wang, Q., & Walker, J. E. Measurement of the molecular

masses of hydrophilic and hydrophobic subunits of ATP synthase and complex I in

a single experiment. Anal. Biochem., 395(2):249–255, 2009.

[32] Chen, R., Runswick, M. J., Carroll, J., Fearnley, I. M., & Walker, J. E. Association

of two proteolipids of unknown function with ATP synthase from bovine heart

mitochondria. FEBS Lett., 581(17):3145–3148, 2007.

[33] Collinson, I. R., Runswick, M. J., Buchanan, S. K., Fearnley, I. M., Skehel, J. M.,

van Raaij, M. J., Griffiths, D. E., & Walker, J. E. Fo membrane domain of ATP

synthase from bovine heart mitochondria: purification, subunit composition, and

reconstitution with F1-ATPase. Biochemistry, 33(25):7971–7978, 1994.

[34] Collinson, I. R., Skehel, J. M., Fearnley, I. M., Runswick, M. J., & Walker, J. E.

The F1F0-ATPase complex from bovine heart mitochondria: the molar ratio of

the subunits in the stalk region linking the F1 and F0 domains. Biochemistry,

35(38):12640–12646, 1996.

[35] Collinson, I. R., van Raaij, M. J., Runswick, M. J., Fearnley, I. M., Skehel, J. M.,

Orriss, G. L., Miroux, B., & Walker, J. E. ATP synthase from bovine heart

mitochondria. In vitro assembly of a stalk complex in the presence of F1-ATPase

and in its absence. J. Mol. Biol., 242(4):408–421, 1994.

Bibliography 77

[36] Craig, K., Elliott, H. R., Keers, S. M., Lambert, C., Pyle, A., Graves, T. D., Wood-

ward, C., Sweeney, M. G., Davis, M. B., Hanna, M. G., & Chinnery, P. F. Episodic

ataxia and hemiplegia caused by the 8993T-¿C mitochondrial DNA mutation. J

Med Genet, 44(12):797–799, 2007.

[37] Cronin, N., Yang, J., Zhang, Z., Kulkarni, K., Chang, L., Yamano, H., & Barford, D.

Atomic-resolution structures of the APC/C subunits Apc4 and the Apc5 N-terminal

domain. J. Mol. Biol., 2015.

[38] Davies, K. M., Anselmi, C., Wittig, I., Faraldo-Gomez, J. D., & Kuhlbrandt,

W. Structure of the yeast F1Fo-ATP synthase dimer and its role in shaping the

mitochondrial cristae. Proc. Natl. Acad. Sci. U. S. A., 109(34):13602–13607, 2012.

[39] Davies, K. M., Strauss, M., Daum, B., Kief, J. H., Osiewacz, H. D., Rycovska,

A., Zickermann, V., & Kuhlbrandt, W. Macromolecular organization of ATP

synthase and complex I in whole mitochondria. Proc. Natl. Acad. Sci. U. S. A.,

108(34):14121–14126, 2011.

[40] Deleon-Rangel, J., Ishmukhametov, R. R., Jiang, W., Fillingame, R. H., & Vik, S. B.

Interactions between subunits a and b in the rotary ATP synthase as determined

by cross-linking. FEBS Lett., 587(7):892–897, 2013. URL http://dx.doi.org/10.

1016/j.febslet.2013.02.012.

[41] Dickson, V. K., Silvester, J. A., Fearnley, I. M., Leslie, A. G., & Walker, J. E.

On the structure of the stator of the mitochondrial ATP synthase. EMBO J.,

25(12):2911–2918, 2006.

[42] DiMaio, F., Tyka, M. D., Baker, M. L., Chiu, W., & Baker, D. Refinement of

protein structures into low-resolution density maps using rosetta. J. Mol. Biol.,

392(1):181–190, 2009.

Bibliography 78

[43] Dimmer, K. S., Fritz, S., Fuchs, F., Messerschmitt, M., Weinbach, N., Neupert,

W., & Westermann, B. Genetic basis of mitochondrial function and morphology in

Saccharomyces cerevisiae. Mol. Biol. Cell, 13(3):847–853, 2002.

[44] Dubochet, J., Adrian, M., Chang, J. J., Homo, J. C., Lepault, J., McDowall, A. W.,

& Schultz, P. Cryo-electron microscopy of vitrified specimens. Q. Rev. Biophys.,

21(2):129–228, 1988.

[45] Dunn, S. D. & Chandler, J. Characterization of a b2delta complex from Escherichia

coli ATP synthase. J. Biol. Chem., 273(15):8646–8651, 1998.

[46] Dunn, S. D. & Heppel, L. A. Properties and functions of the subunits of the

Escherichia coli coupling factor ATPase. Arch. Biochem. Biophys., 210(2):421–436,

1981.

[47] Eswar, N., Webb, B., Marti-Renom, M. A., Madhusudhan, M. S., Eramian, D.,

Shen, M. Y., Pieper, U., & Sali, A. Comparative protein structure modeling using

Modeller. Curr. Protoc. Bioinforma. / Ed. board, Andreas D.Baxevanis ...[et al.],

Chapter 5:Unit 5.6, 2006.

[48] Everard-Gigot, V., Dunn, C. D., Dolan, B. M., Brunner, S., Jensen, R. E., & Stuart,

R. A. Functional analysis of subunit e of the F1Fo-ATP synthase of the yeast

Saccharomyces cerevisiae: importance of the N-terminal membrane anchor region.

Eukaryot. Cell, 4(2):346–355, 2005.

[49] Fillingame, R. H. & Steed, P. R. Half channels mediating H+ transport and

the mechanism of gating in the Fo sector of Escherichia coli F1Fo ATP synthase.

Biochim. Biophys. Acta - Bioenerg., 1837(7):1063–1068, 2014. URL http://dx.

doi.org/10.1016/j.bbabio.2014.03.005.

[50] Frank, J. Three-dimensional electron microscopy of macromolecular assemblies.

Oxford University Press, Toronto, 2 edition, 2006.

Bibliography 79

[51] Fujikawa, M., Ohsakaya, S., Sugawara, K., & Yoshida, M. Population of ATP

synthase molecules in mitochondria is limited by available 6.8-kDa proteolipid

protein (MLQ). Genes Cells, 19(2):153–160, 2014.

[52] Gibbons, C., Montgomery, M. G., Leslie, A. G., & Walker, J. E. The structure

of the central stalk in bovine F(1)-ATPase at 2.4 A resolution. Nat. Struct. Biol.,

7(11):1055–1061, 2000.

[53] Giorgio, V., von Stockum, S., Antoniel, M., Fabbro, A., Fogolari, F., Forte, M.,

Glick, G. D., Petronilli, V., Zoratti, M., Szabo, I., Lippe, G., & Bernardi, P. Dimers

of mitochondrial ATP synthase form the permeability transition pore. Proc. Natl.

Acad. Sci. U. S. A., 110(15):5887–5892, 2013.

[54] Glaeser, R. M. Retrospective: Radiation damage and its associated ”Information

Limitations”. J. Struct. Biol., 163(3):271–276, 2008.

[55] Glaeser, R. M., Downing, K. H., DeRosier, D. J., Chiu, W., & Frank, J. Electron

Crystallography of Biological Macromolecules. Oxford University Press, Toronto,

2007.

[56] Gledhill, J. R., Montgomery, M. G., Leslie, A. G., & Walker, J. E. How the

regulatory protein, IF(1), inhibits F(1)-ATPase from bovine mitochondria. Proc.

Natl. Acad. Sci. U. S. A., 104(40):15671–15676, 2007.

[57] Goddard, T. D., Huang, C. C., & Ferrin, T. E. Visualizing density maps with UCSF

Chimera. J. Struct. Biol., 157(1):281–287, 2007.

[58] Grant, T. & Grigorieff, N. Measuring the optimal exposure for single particle

cryo-EM using a 2.6 A reconstruction of rotavirus VP6. Elife, 4:10.7554/eLife.06980,

2015.

Bibliography 80

[59] Greie, J. C., Heitkamp, T., & Altendorf, K. The transmembrane domain of subunit

b of the Escherichia coli F1F(O) ATP synthase is sufficient for H(+)-translocating

activity together with subunits a and c. Eur. J. Biochem., 271(14):3036–3042, 2004.

[60] Grigorieff, N. Three-dimensional structure of bovine NADH:ubiquinone oxidoreduc-

tase (complex I) at 22 A in ice. J. Mol. Biol., 277(5):1033–1046, 1998.

[61] Grigorieff, N. FREALIGN: high-resolution refinement of single particle structures.

J. Struct. Biol., 157(1):117–125, 2007.

[62] Harris, J. R. & Horne, R. W. Negative staining: A brief assessment of current

technical benefits, limitations and future possibilities. Micron, 25(1):5–13, 1994.

[63] Hayashi, S., Ueno, H., Shaikh, A. R., Umemura, M., Kamiya, M., Ito, Y., Ikeguchi,

M., Komoriya, Y., Iino, R., & Noji, H. Molecular mechanism of ATP hydrolysis in

F1-ATPase revealed by molecular simulations and single-molecule observations. J.

Am. Chem. Soc., 134(20):8447–8454, 2012.

[64] Hinkle, P. C. P/O ratios of mitochondrial oxidative phosphorylation. Biochim.

Biophys. Acta - Bioenerg., 1706(1-2):1–11, 2005.

[65] Hopf, T. A., Colwell, L. J., Sheridan, R., Rost, B., Sander, C., & Marks, D. S.

Three-dimensional structures of membrane proteins from genomic sequencing. Cell,

149(7):1607–1621, 2012.

[66] Hopf, T. a., Scharfe, C. P. I., Rodrigues, J. P. G. L. M., Green, A. G., Kohlbacher,

O., Sander, C., Bonvin, A. M. J. J., & Marks, D. S. Sequence co-evolution

gives 3D contacts and structures of protein complexes. Elife, 3:1–45, 2014. URL

http://www.ncbi.nlm.nih.gov/pubmed/25255213.

Bibliography 81

[67] Hoppe, J., Schairer, H. U., Friedl, P., & Sebald, W. An Asp-Asn substitution

in the proteolipid subunit of the ATP-synthase from Escherichia coli leads to a

non-functional proton channel. FEBS Lett., 145(1):21–29, 1982.

[68] Hosler, J. P., Ferguson-Miller, S., & Mills, D. a. Energy transduction: proton

transfer through the respiratory complexes. Annu. Rev. Biochem., 75:165–187, 2006.

[69] Humphrey, W., Dalke, A., & Schulten, K. VMD: Visual molecular dynamics. J.

Mol. Graph., 14(1):33–38, 1996.

[70] Jiang, W., Baker, M. L., Jakana, J., Weigele, P. R., King, J., & Chiu, W. Backbone

structure of the infectious epsilon15 virus capsid revealed by electron cryomicroscopy.

Nature, 451(7182):1130–1134, 2008.

[71] Jiko, C., Davies, K. M., Shinzawa-Itoh, K., Tani, K., Maeda, S., Mills, D. J.,

Tsukihara, T., Fujiyoshi, Y., Kuhlbrandt, W., & Gerle, C. Bovine F1Fo ATP

synthase monomers bend the lipid bilayer in 2D membrane crystals. Elife, 4:e06119,

2015.

[72] Johnson, K. M., Cleary, J., Fierke, C. A., Opipari Jr, A. W., & Glick, G. D.

Mechanistic basis for therapeutic targeting of the mitochondrial F1F0-ATPase. ACS

Chem. Biol., 1(5):304–308, 2006.

[73] Jonas, E. A., Porter Jr, G. A., Beutner, G., Mnatsakanyan, N., & Alavian, K. N. Cell

death disguised: The mitochondrial permeability transition pore as the c-subunit of

the FF ATP synthase. Pharmacol. Res., 2015.

[74] Jonckheere, A. I., Smeitink, J. A. M., & Rodenburg, R. J. T. Mitochondrial ATP

synthase: architecture, function and pathology. J. Inherit. Metab. Dis., 35(2):211–

225, 2012. URL http://link.springer.com/10.1007/s10545-011-9382-9.

Bibliography 82

[75] Jones, D. T. Protein secondary structure prediction based on position-specific

scoring matrices. J. Mol. Biol., 292(2):195–202, 1999. URL http://dx.doi.org/

10.1006/jmbi.1999.3091.

[76] Jones, P. C., Hermolin, J., Jiang, W., & Fillingame, R. H. Insights into the rotary

catalytic mechanism of F0F1 ATP synthase from the cross-linking of subunits b

and c in the Escherichia coli enzyme. J. Biol. Chem., 275(40):31340–31346, 2000.

[77] Junge, W., Lill, H., & Engelbrecht, S. ATP synthase: an electrochemical transducer

with rotatory mechanics. Trends Biochem. Sci., 22(11):420–423, 1997.

[78] Junge, W. & Nelson, N. Structural biology. Nature’s rotary electromotors. Science,

308(5722):642–644, 2005.

[79] Junge, W., Sielaff, H., & Engelbrecht, S. Torque generation and elastic power

transmission in the rotary F(O)F(1)-ATPase. Nature, 459(7245):364–370, 2009.

[80] Kabaleeswaran, V., Puri, N., Walker, J. E., Leslie, A. G. W., & Mueller, D. M.

Novel features of the rotary catalytic mechanism revealed in the structure of yeast

F1 ATPase. EMBO J., 25(22):5433–5442, 2006.

[81] Kerscher, S. J. Diversity and origin of alternative NADH:ubiquinone oxidoreductases.

Biochim. Biophys. Acta - Bioenerg., 1459(2-3):274–283, 2000.

[82] Komoriya, Y., Ariga, T., Iino, R., Imamura, H., Okuno, D., & Noji, H. Principal

role of the arginine finger in rotary catalysis of F 1-ATPase. J. Biol. Chem.,

287(18):15134–15142, 2012.

[83] Kucukelbir, A., Sigworth, F. J., & Tagare, H. D. Quantifying the local resolution of

cryo-EM density maps. Nat. Methods, 11(1):63–65, 2014.

[84] Kuhlbrandt, W. Biochemistry. The resolution revolution. Science, 343(6178):1443–

1444, 2014.

Bibliography 83

[85] Lamminen, T., Majander, A., Juvonen, V., Wikstrom, M., Aula, P., Nikoskelainen,

E., & Savontous, M. L. A mitochondrial mutation at nt 9101 in the ATP synthase

6 gene associated with deficient oxidative phosphorylation in a family with Leber

hereditary optic neuroretinopathy. Am. J. Hum. Genet., 56(5):1238–1240, 1995.

URL http://www.ncbi.nlm.nih.gov/pmc/articles/PMC1801467/.

[86] Lau, W. C., Baker, L. A., & Rubinstein, J. L. Cryo-EM structure of the yeast ATP

synthase. J. Mol. Biol., 382(5):1256–1264, 2008.

[87] Lau, W. C. Y. & Rubinstein, J. L. Structure of intact Thermus thermophilus

V-ATPase by cryo-EM reveals organization of the membrane-bound V(O) motor.

Proc. Natl. Acad. Sci. U. S. A., 107(4):1367–1372, 2010.

[88] Lau, W. C. Y. & Rubinstein, J. L. Subnanometre-resolution structure of the intact

Thermus thermophilus H+-driven ATP synthase. Nature, 481(7380):214–218, 2011.

URL http://dx.doi.org/10.1038/nature10699.

[89] Lee, C. P., Gu, Q., Xiong, Y., Mitchell, R. a., & Ernster, L. P/O ratios reassessed:

mitochondrial P/O ratios consistently exceed 1.5 with succinate and 2.5 with

NAD-linked substrates. FASEB J., 10(2):345–350, 1996.

[90] Lee, J., Ding, S., Walpole, T. B., Holding, A. N., Montgomery, M. G., Fearnley, I. M.,

& Walker, J. E. Organization of Subunits in the Membrane Domain of the Bovine

F-ATPase Revealed by Covalent Cross-linking. J. Biol. Chem., 290(21):13308–13320,

2015.

[91] Lee, L. K., Stewart, A. G., Donohoe, M., Bernal, R. A., & Stock, D. The structure

of the peripheral stalk of Thermus thermophilus H+-ATPase/synthase. Nat. Struct.

Mol. Biol., 17(3):373–378, 2010.

Bibliography 84

[92] Leschziner, A. E. & Nogales, E. The orthogonal tilt reconstruction method: An

approach to generating single-class volumes with no missing cone for ab initio

reconstruction of asymmetric particles. J. Struct. Biol., 153(3):284–299, 2006.

[93] Liu, H., Jin, L., Koh, S. B. S., Atanasov, I., Schein, S., Wu, L., & Zhou, Z. H.

Atomic structure of human adenovirus by cryo-EM reveals interactions among

protein networks. Science, 329(5995):1038–1043, 2010.

[94] Loken, C., Gruner, D., Groer, L., Peltier, R., Bunn, N., Craig, M., Henriques, T.,

Dempsey, J., Yu, C.-H., Chen, J., Dursi, L. J., Chong, J., Northrup, S., Pinto, J.,

Knecht, N., & Zon, R. V. Scinet: Lessons learned from building a power-efficient top-

20 system and data centre. Journal of Physics: Conference Series, 256(1):012026,

2010. URL http://stacks.iop.org/1742-6596/256/i=1/a=012026.

[95] Lyumkis, D., Brilot, A. F., Theobald, D. L., & Grigorieff, N. Likelihood-based

classification of cryo-EM images using FREALIGN. J. Struct. Biol., 183(3):377–388,

2013.

[96] Macreadie, I. G., Novitski, C. E., Maxwell, R. J., John, U., Ooi, B. G., McMullen,

G. L., Lukins, H. B., Linnane, A. W., & Nagley, P. Biogenesis of mitochondria:

the mitochondrial gene (aap1) coding for mitochondrial ATPase subunit 8 in

Saccharomyces cerevisiae. Nucleic Acids Res., 11(13):4435–4451, 1983.

[97] Maier, O., Galan, D., Wodrich, H., & W., C. M. An N-terminal Domain of

Adenovirus Protein VI Fragments Membranes By Inducing Positive Membrane

Curvature. Virology, 402(1):11–19, 2011.

[98] Marks, D. S., Colwell, L. J., Sheridan, R., Hopf, T. A., Pagnani, A., Zecchina, R.,

& Sander, C. Protein 3D structure computed from evolutionary sequence variation.

PLoS One, 6(12):e28766, 2011.

Bibliography 85

[99] Marr, C. R., Benlekbir, S., & Rubinstein, J. L. Fabrication of carbon films with

approximately 500nm holes for cryo-EM with a direct detector device. J. Struct.

Biol., 185(1):42–47, 2014.

[100] Martin, J. L., Ishmukhametov, R., Hornung, T., Ahmad, Z., & Frasch,

W. D. Anatomy of F1-ATPase powered rotation. Proc. Natl. Acad. Sci. U.

S. A., 111(10):3715–20, 2014. URL http://www.pubmedcentral.nih.gov/

articlerender.fcgi?artid=3956197{\&}tool=pmcentrez{\&}rendertype=

abstract.

[101] Masaike, T., Koyama-Horibe, F., Oiwa, K., Yoshida, M., & Nishizaka, T. Coop-

erative three-step motions in catalytic subunits of F(1)-ATPase correlate with 80

degrees and 40 degrees substep rotations. Nat. Struct. Mol. Biol., 15(12):1326–1333,

2008.

[102] Mastronarde, D. N. Automated electron microscope tomography using robust

prediction of specimen movements. J. Struct. Biol., 152(1):36–51, 2005.

[103] Masuda, M., Takeda, S., Sone, M., Ohki, T., Mori, H., Kamioka, Y., & Mochizuki,

N. Endophilin BAR domain drives membrane curvature by two newly identified

structure-based mechanisms. EMBO J., 25(12):2889–2897, 2006.

[104] McMullan, G., Chen, S., Henderson, R., & Faruqi, a. R. Detective quantum efficiency

of electron area detectors in electron microscopy. Ultramicroscopy, 109(9):1126–1143,

2009.

[105] Menz, R. I., Walker, J. E., & Leslie, A. G. Structure of bovine mitochondrial

F(1)-ATPase with nucleotide bound to all three catalytic sites: implications for the

mechanism of rotary catalysis. Cell, 106(3):331–341, 2001.

[106] Minagawa, Y., Ueno, H., Hara, M., Ishizuka-Katsura, Y., Ohsawa, N., Terada, T.,

Shirouzu, M., Yokoyama, S., Yamato, I., Muneyuki, E., Noji, H., Murata, T., &

Bibliography 86

Iino, R. Basic properties of rotary dynamics of the molecular motor enterococcus

hirae V1-ATPase. J. Biol. Chem., 288(45):32700–32707, 2013.

[107] Minauro-Sanmiguel, F., Wilkens, S., & Garcia, J. J. Structure of dimeric mito-

chondrial ATP synthase: novel F0 bridging features and the structural basis of

mitochondrial cristae biogenesis. Proc. Natl. Acad. Sci. U. S. A., 102(35):12356–

12358, 2005.

[108] Morales-Rios, E., Montgomery, M. G., Leslie, A. G., & Walker, J. E. Structure of atp

synthase from paracoccus denitrificans determined by x-ray crystallography at 4.0 a

resolution. Proceedings of the National Academy of Sciences, 112(43):13231–13236,

2015.

[109] Nakamoto, R. K., Shin, K., Iwamoto, A., Omote, H., Maeda, M., & Futai, M.

Escherichia coli F0F1-ATPase. Residues involved in catalysis and coupling. Ann. N.

Y. Acad. Sci., 671:335–343; discussion 343–344, 1992.

[110] Nakanishi-Matsui, M., Kashiwagi, S., Hosokawa, H., Cipriano, D. J., Dunn,

S. D., Wada, Y., & Futai, M. Stochastic high-speed rotation of Escherichia coli

ATP synthase F1 sector: the epsilon subunit-sensitive rotation. J. Biol. Chem.,

281(7):4126–31, 2006. URL http://www.jbc.org/content/281/7/4126.short$\

backslash$nhttp://www.ncbi.nlm.nih.gov/pubmed/16352612.

[111] Nishizaka, T., Oiwa, K., Noji, H., Kimura, S., Muneyuki, E., Yoshida, M., &

Kinosita, K. Chemomechanical coupling in F1-ATPase revealed by simultaneous

observation of nucleotide kinetics and rotation. Nat. Struct. Mol. Biol., 11(2):142–

148, 2004.

[112] Noji, H., Yasuda, R., Yoshida, M., & Kinosita Jr, K. Direct observation of the

rotation of F1-ATPase. Nature, 386(6622):299–302, 1997.

Bibliography 87

[113] Nugent, T. & Jones, D. T. Transmembrane protein topology prediction using

support vector machines. BMC Bioinformatics, 10:159, 2009.

[114] Ohsakaya, S., Fujikawa, M., Hisabori, T., & Yoshida, M. Knockdown of DAPIT

(diabetes-associated protein in insulin-sensitive tissue) results in loss of ATP synthase

in mitochondria. J. Biol. Chem., 286(23):20292–20296, 2011.

[115] Okazaki, K. I. & Hummer, G. Elasticity, friction, and pathway of gamma-subunit

rotation in FoF1-ATP synthase. Proc. Natl. Acad. Sci. U. S. A., 2015.

[116] Okazaki, K. I. & Takada, S. Structural comparison of F1-ATPase: Interplay among

enzyme structures, catalysis, and rotations. Structure, 19(4):588–598, 2011.

[117] Omote, H., Sambonmatsu, N., Saito, K., Sambongi, Y., Iwamoto-Kihara, a.,

Yanagida, T., Wada, Y., & Futai, M. The gamma-subunit rotation and torque

generation in F1-ATPase from wild-type or uncoupled mutant Escherichia coli. Proc.

Natl. Acad. Sci. U. S. A., 96(14):7780–7784, 1999.

[118] Ovchinnikov, S., Kamisetty, H., & Baker, D. Robust and accurate prediction of

residue-residue interactions across protein interfaces using evolutionary information.

Elife, 3:e02030, 2014.

[119] Parsons, D. F. Mitochondrial Structure: Two Types of Subunits on Negatively

Stained Mitochondrial Membranes. Science, 140(3570):985–987, 1963.

[120] Paumard, P., Vaillier, J., Coulary, B., Schaeffer, J., Soubannier, V., Mueller, D. M.,

Brethes, D., Di Rago, J. P., & Velours, J. The ATP synthase is involved in generating

mitochondrial cristae morphology. EMBO J., 21(3):221–230, 2002.

[121] Pintilie, G. D., Zhang, J., Goddard, T. D., Chiu, W., & Gossard, D. C. Quantitative

analysis of cryo-EM density map segmentation by watershed and scale-space filtering,

Bibliography 88

and fitting of structures by alignment to regions. J. Struct. Biol., 170(3):427–438,

2010.

[122] Pogoryelov, D., Krah, A., Langer, J. D., Yildiz, O., Faraldo-Gomez, J. D., & Meier,

T. Microscopic rotary mechanism of ion translocation in the F(o) complex of ATP

synthases. Nat. Chem. Biol., 6(12):891–899, 2010.

[123] Pogoryelov, D., Yildiz, O., Faraldo-Gomez, J. D., & Meier, T. High-resolution

structure of the rotor ring of a proton-dependent ATP synthase. Nat. Struct. Mol.

Biol., 16(10):1068–1073, 2009.

[124] Pogoryelov, D., Yu, J., Meier, T., Vonck, J., Dimroth, P., & Muller, D. J. The c15

ring of the Spirulina platensis F-ATP synthase: F1/F0 symmetry mismatch is not

obligatory. EMBO Rep., 6(11):1040–1044, 2005.

[125] Popot, J.-L. Amphipols, nanodiscs, and fluorinated surfactants: three noncon-

ventional approaches to studying membrane proteins in aqueous solutions. Annu.

Rev. Biochem., 79:737–75, 2010. URL http://www.ncbi.nlm.nih.gov/pubmed/

20307193.

[126] Pullman, M. E. & Monroy, G. C. A Naturally Occurring Inhibitor of Mitochondrial

Adenosine Triphosphatase. J. Biol. Chem., 238:3762–3769, 1963.

[127] Radermacher, M., Wagenknecht, T., Verschoor, A., & Frank, J. Three-dimensional

reconstruction from a single-exposure, random conical tilt series applied to the 50S

ribosomal subunit of Escherichia coli. J. Microsc., 146(Pt 2):113–136, 1987.

[128] Rak, M., Gokova, S., & Tzagoloff, A. Modular assembly of yeast mitochondrial

ATP synthase. EMBO J., 30(5):920–930, 2011.

Bibliography 89

[129] Rees, D. M., Leslie, A. G., & Walker, J. E. The structure of the membrane extrinsic

region of bovine ATP synthase. Proc. Natl. Acad. Sci. U. S. A., 106(51):21597–21601,

2009.

[130] Reimer, L. & Kohl, H. Transmission electron microscopy - Physics of image

formation. Springer Series in Optical Sciences, New York, 5 edition, 2008.

[131] Rohl, C. A., Strauss, C. E., Misura, K. M., & Baker, D. Protein structure prediction

using Rosetta. Methods Enzymol., 383:66–93, 2004.

[132] Rohou, A. & Grigorieff, N. Ctffind4: Fast and accurate defocus estimation from

electron micrographs. Journal of structural biology, 2015. LR: 20150824; CI: Copy-

right (c) 2015; JID: 9011206; OTO: NOTNLM; 2015/06/13 [received]; 2015/08/11

[revised]; 2015/08/12 [accepted]; aheadofprint.

[133] Rosenthal, P. B. & Henderson, R. Optimal determination of particle orientation,

absolute hand, and contrast loss in single-particle electron cryomicroscopy. J. Mol.

Biol., 333(4):721–745, 2003.

[134] Rossman, J. S., Jing, X., Leser, G. P., & Lamb, R. a. Influenza Virus M2 Protein

Mediates ESCRT-Independent Membrane Scission. Cell, 142(6):902–913, 2010. URL

http://dx.doi.org/10.1016/j.cell.2010.08.029.

[135] Rubinstein, J. L. Structural analysis of membrane protein complexes by single

particle electron microscopy. Methods, 41(4):409–416, 2007.

[136] Rubinstein, J. L. & Brubaker, M. A. Alignment of cryo-EM movies of individual

particles by optimization of image translations. J. Struct. Biol., 2015.

[137] Rubinstein, J. L., Walker, J. E., & Henderson, R. Structure of the mitochondrial

ATP synthase by electron cryomicroscopy. EMBO J., 22(23):6182–6192, 2003.

Bibliography 90

[138] Runswick, M. J., Bason, J. V., Montgomery, M. G., Robinson, G. C., Fearnley, I. M.,

& Walker, J. E. The affinity purification and characterization of ATP synthase

complexes from mitochondria. Open Biol., 3(2):120160, 2013.

[139] Russo, C. J. & Passmore, L. A. Electron microscopy: Ultrastable gold substrates

for electron cryomicroscopy. Science, 346(6215):1377–1380, 2014.

[140] Salje, J., van den Ent, F., de Boer, P., & Lowe, J. Direct Membrane Binding by

Bacterial Actin MreB. Mol. Cell, 43(3):478–487, 2011.

[141] Scheres, S. H. RELION: implementation of a Bayesian approach to cryo-EM

structure determination. J. Struct. Biol., 180(3):519–530, 2012.

[142] Scheres, S. H. Beam-induced motion correction for sub-megadalton cryo-EM

particles. Elife, 3:e03665, 2014. URL http://elifesciences.org/lookup/doi/

10.7554/eLife.03665$\backslash$nhttp://www.ncbi.nlm.nih.gov/pubmed/

25122622.

[143] Scheres, S. H. Semi-automated selection of cryo-EM particles in RELION-1.3. J.

Struct. Biol., 189(2):114–122, 2015.

[144] Seelert, H. & Dencher, N. A. ATP synthase superassemblies in animals and plants:

Two or more are better, 2011.

[145] Shih, Y.-L., Huang, K.-F., Lai, H.-M., Liao, J.-H., Lee, C.-S., Chang, C.-M., Mak, H.-

M., Hsieh, C.-W., & Lin, C.-C. The N-terminal amphipathic helix of the topological

specificity factor MinE is associated with shaping membrane curvature. PLoS One,

6(6):e21425, 2011. URL http://www.pubmedcentral.nih.gov/articlerender.

fcgi?artid=3124506{\&}tool=pmcentrez{\&}rendertype=abstract.

[146] Soubannier, V., Vaillier, J., Paumard, P., Coulary, B., Schaeffer, J., & Velours, J.

In the absence of the first membrane-spanning segment of subunit 4(b), the yeast

Bibliography 91

ATP synthase is functional but does not dimerize or oligomerize. J. Biol. Chem.,

277(12):10739–10745, 2002.

[147] Spence, J. High-resolution electron microscopy. Oxford University Press, Toronto, 3

edition, 2003.

[148] Spetzler, D., York, J., Daniel, D., Fromme, R., Lowry, D., & Frasch, W.

Microsecond time scale rotation measurements of single F1-ATPase molecules.

Biochemistry, 45(10):3117–3124, 2006. URL http://eutils.ncbi.nlm.nih.gov/

entrez/eutils/elink.fcgi?dbfrom=pubmed{\&}id=16519506{\&}retmode=

ref{\&}cmd=prlinks$\backslash$npapers2://publication/doi/10.1021/

bi052363n$\backslash$nhttp://www.ncbi.nlm.nih.gov/pubmed/16519506.

[149] Steel, B. C., Nord, A. L., Wang, Y., Pagadala, V., Mueller, D. M., & Berry, R. M.

Comparison between single-molecule and X-ray crystallography data on yeast F1-

ATPase. Sci. Rep., 5:8773, 2015. URL http://www.ncbi.nlm.nih.gov/pubmed/

25753753.

[150] Stelzer, A. C., Frazee, R. W., Van Huis, C., Cleary, J., Opipari Jr, A. W., Glick,

G. D., & Al-Hashimi, H. M. NMR studies of an immunomodulatory benzodiazepine

binding to its molecular target on the mitochondrial F(1)F(0)-ATPase. Biopolymers,

93(1):85–92, 2010.

[151] Stewart, A. & Grigorieff, N. Noise bias in the refinement of structures derived from

single particles. Ultramicroscopy, 102(1):67–84, 2004.

[152] Stock, D., Leslie, A. G., & Walker, J. E. Molecular architecture of the rotary motor

in ATP synthase. Science, 286(5445):1700–1705, 1999.

[153] Strauss, M., Hofhaus, G., Schroder, R. R., & Kuhlbrandt, W. Dimer ribbons of ATP

synthase shape the inner mitochondrial membrane. EMBO J., 27(7):1154–1160,

2008.

Bibliography 92

[154] Suzuki, T., Tanaka, K., Wakabayashi, C., Saita, E.-i., & Yoshida, M. Chemo-

mechanical coupling of human mitochondrial F1-ATPase motor. Nat.

Chem. Biol., 10(11):930–936, 2014. URL http://dx.doi.org/10.1038/

nchembio.1635$\backslash$n10.1038/nchembio.1635$\backslash$nhttp:

//www.nature.com/nchembio/journal/v10/n11/abs/nchembio.1635.html{\#

}supplementary-information.

[155] Symersky, J., Osowski, D., Walters, D. E., & Mueller, D. M. Oligomycin

frames a common drug-binding site in the ATP synthase. Proc. Natl. Acad. Sci.

U. S. A., 109(35):13961–5, 2012. URL http://www.pubmedcentral.nih.gov/

articlerender.fcgi?artid=3435195{\&}tool=pmcentrez{\&}rendertype=

abstract.

[156] Symersky, J., Pagadala, V., Osowski, D., Krah, A., Meier, T., Faraldo-Gomez,

J. D., & Mueller, D. M. Structure of the c(10) ring of the yeast mitochondrial ATP

synthase in the open conformation. Nat. Struct. Mol. Biol., 19(5):485–91, S1, 2012.

[157] Thyagarajan, D., Shanske, S., Vazquez-Memije, M., De Vivo, D., & DiMauro, S. A

novel mitochondrial ATPase 6 point mutation in familial bilateral striatal necrosis.

Ann. Neurol., 38(3):468–72, 1995. URL http://www.ncbi.nlm.nih.gov/pubmed/

7668837.

[158] Timmins, P., Leonhard, M., Weltzien, H., Wacker, T., & Welte, W. A physical

characterization of some detergents of potential use for membrane protein crys-

tallization. FEBS Lett., 238(2):361–368, 1988. URL http://www.sciencedirect.

com/science/article/pii/0014579388805131.

[159] Trabuco, L. G., Villa, E., Mitra, K., Frank, J., & Schulten, K. Flexible fitting

of atomic structures into electron microscopy maps using molecular dynamics.

Structure, 16(5):673–683, 2008.

Bibliography 93

[160] Trabuco, L. G., Villa, E., Schreiner, E., Harrison, C. B., & Schulten, K. Molecular

dynamics flexible fitting: a practical guide to combine cryo-electron microscopy and

X-ray crystallography. Methods, 49(2):174–180, 2009.

[161] Uchihashi, T., Iino, R., Ando, T., & Noji, H. High-speed atomic force microscopy

reveals rotary catalysis of rotorless F-ATPase. Science, 333(6043):755–758, 2011.

[162] Van Heel, M. Angular reconstitution: a posteriori assignment of projection directions

for 3D reconstruction. Ultramicroscopy, 21(2):111–123, 1987.

[163] Vassilopoulos, A., Pennington, J. D., Andresson, T., Rees, D. M., Bosley, A. D.,

Fearnley, I. M., Ham, A., Flynn, C. R., Hill, S., Rose, K. L., Kim, H. S., Deng,

C. X., Walker, J. E., & Gius, D. SIRT3 deacetylates ATP synthase F1 complex

proteins in response to nutrient- and exercise-induced stress. Antioxid. Redox Signal.,

21(4):551–564, 2014.

[164] Velours, J., Vaillier, J., Paumard, P., Soubannier, V., Lai-Zhang, J., & Mueller,

D. M. Bovine Coupling Factor 6, with Just 14.5% Shared Identity, Replaces Subunit

h in the Yeast ATP Synthase. J. Biol. Chem., 276(11):8602–8607, 2001.

[165] Wachter, A., Bi, Y., Dunn, S. D., Cain, B. D., Sielaff, H., Wintermann, F., En-

gelbrecht, S., & Junge, W. Two rotary motors in F-ATP synthase are elastically

coupled by a flexible rotor and a stiff stator stalk. Proc. Natl. Acad. Sci. U. S. A.,

108(10):3924–3929, 2011.

[166] Watt, I. N., Montgomery, M. G., Runswick, M. J., Leslie, A. G., & Walker,

J. E. Bioenergetic cost of making an adenosine triphosphate molecule in animal

mitochondria. Proc. Natl. Acad. Sci. U. S. A., 107(39):16823–16827, 2010.

[167] Wriggers, W. Using Situs for the integration of multi-resolution structures. Biophys.

Rev., 2(1):21–27, 2010.

Bibliography 94

[168] Yasuda, R., Noji, H., Kinosita, K., & Yoshida, M. F1-ATPase is a highly efficient

molecular motor that rotates with discrete 120??steps. Cell, 93(7):1117–1124, 1998.

[169] Yasuda, R., Noji, H., Yoshida, M., Kinosita, K., & Itoh, H. Resolution of dis-

tinct rotational substeps by submillisecond kinetic analysis of F1-ATPase. Nature,

410(6831):898–904, 2001.

[170] Yu, X., Jin, L., & Zhou, Z. H. 3.88 A structure of cytoplasmic polyhedrosis virus

by cryo-electron microscopy. Nature, 453(7193):415–419, 2008.

[171] Zhang, X., Jin, L., Fang, Q., Hui, W. H., & Zhou, Z. H. 3.3 A Cryo-EM structure of a

nonenveloped virus reveals a priming mechanism for cell entry. Cell, 141(3):472–482,

2010.

[172] Zhao, J., Benlekbir, S., & Rubinstein, J. L. Electron cryomicroscopy observation of

rotational states in a eukaryotic V-ATPase. Nature, 521(7551):241–245, 2015.

[173] Zhou, A., Rohou, A., Schep, D. G., Bason, J. V., Montgomery, M. G., Walker,

J. E., Grigorieff, N., & Rubinstein, J. L. Structure and conformational states of the

bovine mitochondrial atp synthase by cryo-em. eLife, 2015.

[174] Zhou, M., Politis, A., Davies, R. B., Liko, I., Wu, K. J., Stewart, A. G., Stock,

D., & Robinson, C. V. Ion mobility-mass spectrometry of a rotary ATPase reveals

ATP-induced reduction in conformational flexibility. Nat. Chem., 6(3):208–215,

2014.

[175] Zickermann, V., Angerer, H., Ding, M. G., Nubel, E., & Brandt, U. Small single

transmembrane domain (STMD) proteins organize the hydrophobic subunits of

large membrane protein complexes. FEBS Lett., 584(12):2516–2525, 2010. URL

http://dx.doi.org/10.1016/j.febslet.2010.04.021.

Appendices

95

Appendix A

Movie captions

The movies in this thesis have been published in eLife [173].

Movie 1. Conformational changes during the rotary cycle. The transition by linear

interpolation between states 1a, 2a and 3a is shown, with pauses at each state. Large

conformational changes can be seen, including the turning of the c-ring (pink) against

the a subunit (green), the rotation of the rotor (blue, cyan and purple) within the α3β3

complex (red and yellow), and the conformational changes of each catalytic αβ pair. View

in loop.

Movie 2. Conformational changes between substates (Side view). A) The transition

by linear interpolation between states 1a and 1b, showing a bending of the peripheral

stalk near the OSCP (blue) and F6 subunit (magenta), around the C terminus of the b

subunit (red). B) The transition between states 2a and 2b, also showing a bending of the

peripheral stalk near the OSCP subunit. C) The transition between states 2b and 2c,

showing a bending of the peripheral stalk near the FO region. D) The transition between

states 3a and 3b, also showing a bending of the peripheral stalk near the FO region. Scale

bar 25 A. View in loop.

96

Appendix A. Movie captions 97

Movie 3. Conformational changes between substates (Top view). The same transitions as

in Movie 2, viewed from the F1 region towards the FO region. Scale bar 25 A. View in loop.

Movie 4. Brownian ratchet mechanism. The a subunit (green), b subunit (red) and

c8-ring (pink) are viewed in the direction of the matrix towards the intermembrane space.

Arg159 is shown as a blue sphere on the a subunit. The movement of the c8-ring between

substates (states 2a and 2c) is shown here, with the side chains of the Glu58 residues on

the c subunits represented as spheres. Glu58 on a c subunit changes from a proton-locked

conformation while in the lipid bilayer to an open conformation near Arg159, where it is

deprotonated. Upon re-protonation, it assumes the proton-locked position and can enter

the lipid bilayer again. The c8-ring oscillation is consistent with the Brownian ratchet

mechanism for coupling of proton translocation to rotor rotation. Scale bar 25 A. View

in loop.

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