The majority of adenosine triphosphate (ATP) powering cellular processes in eukaryotes is produced by the mitochondrial F1Fo ATP synthase. Here, we present the atomic models of the membrane Fo domain and the entire mammalian (ovine) F1Fo, determined by cryo-electron microscopy. Subunits in the membrane domain are arranged in the ‘proton translocation cluster’ attached to the c-ring and a more distant ‘hook apparatus’ holding subunit e. Unexpectedly, this subunit is anchored to a lipid ‘plug’ capping the c-ring. We present a detailed proton translocation pathway in mammalian Fo and key inter-monomer contacts in F1Fo multimers. Cryo-EM maps of F1Fo exposed to calcium reveal a retracted subunit e and a disassembled c-ring, suggesting permeability transition pore opening. We propose a model for the permeability transition pore opening, whereby subunit e pulls the lipid plug out of the c-ring. Our structure will allow the design of drugs for many emerging applications in medicine.
Subscribe to Journal
Get full journal access for 1 year
only $4.92 per issue
All prices are NET prices.
VAT will be added later in the checkout.
Tax calculation will be finalised during checkout.
Rent or Buy article
Get time limited or full article access on ReadCube.
All prices are NET prices.
Sazanov, L. A. A giant molecular proton pump: structure and mechanism of respiratory complex I. Nat. Rev. Mol. Cell Biol. 16, 375–388 (2015).
Wikström, M., Sharma, V., Kaila, V. R., Hosler, J. P. & Hummer, G. New perspectives on proton pumping in cellular respiration. Chem. Rev. 115, 2196–2221 (2015).
Walker, J. E. The ATP synthase: the understood, the uncertain and the unknown. Biochem. Soc. Trans. 41, 1–16 (2013).
Forgac, M. Vacuolar ATPases: rotary proton pumps in physiology and pathophysiology. Nat. Rev. Mol. Cell Biol. 8, 917–929 (2007).
Abrahams, J. P., Leslie, A. G., Lutter, R. & Walker, J. E. Structure at 2.8-Å resolution of F1-ATPase from bovine heart mitochondria. Nature 370, 621–628 (1994).
Schagger, H. & Pfeiffer, K. Supercomplexes in the respiratory chains of yeast and mammalian mitochondria. EMBO J. 19, 1777–1783 (2000).
Letts, J. A., Fiedorczuk, K. & Sazanov, L. A. The architecture of respiratory supercomplexes. Nature 537, 644–648 (2016).
Guo, R., Zong, S., Wu, M., Gu, J. & Yang, M. Architecture of human mitochondrial respiratory megacomplex I2III2IV2. Cell 170, 1247–1257 (2017).
Letts, J. A., Fiedorczuk, K., Degliesposti, G., Skehel, M. & Sazanov, L. A. Structures of respiratory supercomplex I + III2 reveal functional and conformational crosstalk. Mol. Cell 75, 1131–1146 (2019).
Davies, K. M., Blum, T. B. & Kuhlbrandt, W. Conserved in situ arrangement of complex I and III2 in mitochondrial respiratory chain supercomplexes of mammals, yeast and plants. Proc. Natl Acad. Sci. USA 115, 3024–3029 (2018).
Davies, K. M., Anselmi, C., Wittig, I., Faraldo-Gómez, J. D. & Kühlbrandt, W. Structure of the yeast F1Fo-ATP synthase dimer and its role in shaping the mitochondrial cristae. Proc. Natl Acad. Sci. USA 109, 13602–13607 (2012).
Giorgio, V. et al. Dimers of mitochondrial ATP synthase form the permeability transition pore. Proc. Natl Acad. Sci. USA 110, 5887–5892 (2013).
Carraro, M., Checchetto, V., Szabó, I. & Bernardi, P. F-ATP synthase and the permeability transition pore: fewer doubts, more certainties. FEBS Lett. 593, 1542–1553 (2019).
Mnatsakanyan, N., Beutner, G., Porter, G. A., Alavian, K. N. & Jonas, E. A. Physiological roles of the mitochondrial permeability transition pore. J. Bioenerg. Biomembr. 49, 13–25 (2017).
Antonucci, S. et al. A novel class of cardioprotective small-molecule PTP inhibitors. Pharmacol. Res. 151, 104548 (2020).
Liu, X., Kim, C. N., Yang, J., Jemmerson, R. & Wang, X. Induction of apoptotic program in cell-free extracts: requirement for dATP and cytochrome c. Cell 86, 147–157 (1996).
Porter, G. A. Jr. & Beutner, G. Cyclophilin D, somehow a master regulator of mitochondrial function. Biomolecules 8, 176 (2018).
Bonora, M. et al. Mitochondrial permeability transition involves dissociation of F1FO ATP synthase dimers and C-ring conformation. EMBO Rep. 18, 1077–1089 (2017).
Giorgio, V. et al. Ca2+ binding to F-ATP synthase β subunit triggers the mitochondrial permeability transition. EMBO Rep. 18, 1065–1076 (2017).
Antoniel, M. et al. The unique histidine in OSCP subunit of F-ATP synthase mediates inhibition of the permeability transition pore by acidic pH. EMBO Rep. 19, 257–268 (2018).
Guo, L. et al. Arginine 107 of yeast ATP synthase subunit g mediates sensitivity of the mitochondrial permeability transition to phenylglyoxal. J. Biol. Chem. 293, 14632–14645 (2018).
Guo, L. et al. Arg-8 of yeast subunit e contributes to the stability of F-ATP synthase dimers and to the generation of the full-conductance mitochondrial megachannel. J. Biol. Chem. 294, 10987–10997 (2019).
Urbani, A. et al. Purified F-ATP synthase forms a Ca2+-dependent high-conductance channel matching the mitochondrial permeability transition pore. Nat. Commun. 10, 4341 (2019).
Mnatsakanyan, N. et al. A mitochondrial megachannel resides in monomeric F1FO ATP synthase. Nat. Commun. 10, 5823 (2019).
Bonora, M. et al. Role of the c subunit of the FO ATP synthase in mitochondrial permeability transition. Cell Cycle 12, 674–683 (2013).
He, J., Carroll, J., Ding, S., Fearnley, I. M. & Walker, J. E. Permeability transition in human mitochondria persists in the absence of peripheral stalk subunits of ATP synthase. Proc. Natl Acad. Sci. USA 114, 9086–9091 (2017).
He, J. et al. Persistence of the mitochondrial permeability transition in the absence of subunit c of human ATP synthase. Proc. Natl Acad. Sci. USA 114, 3409–3414 (2017).
Carroll, J., He, J., Ding, S., Fearnley, I. M. & Walker, J. E. Persistence of the permeability transition pore in human mitochondria devoid of an assembled ATP synthase. Proc. Natl Acad. Sci. USA 116, 12816–12821 (2019).
Alavian, K. N. et al. An uncoupling channel within the c-subunit ring of the F1FO ATP synthase is the mitochondrial permeability transition pore. Proc. Natl Acad. Sci. USA 111, 10580–10585 (2014).
Neginskaya, M. A. et al. ATP synthase C-subunit-deficient mitochondria have a small cyclosporine A-sensitive channel, but lack the permeability transition pore. Cell Rep. 26, 11–17 (2019).
Long, Z. & Sazanov, L. A. Structure and conformational plasticity of the intact thermus thermophilus V/A-type ATPase. Science https://doi.org/10.1126/science.aaw9144 (2019).
Guo, H., Suzuki, T. & Rubinstein, J. L. Structure of a bacterial ATP synthase. Elife 8, e43128 (2019).
Srivastava, A. P. High-resolution cryo-EM analysis of the yeast ATP synthase in a lipid membrane. Science 360, eaas9699 (2018).
Hahn, A., Vonck, J., Mills, D. J., Meier, T. & Kühlbrandt, W. Structure, mechanism and regulation of the chloroplast ATP synthase. Science 360, eaat4318 (2018).
Rees, D. M., Leslie, A. G. & Walker, J. E. The structure of the membrane extrinsic region of bovine ATP synthase. Proc. Natl Acad. Sci. USA 106, 21597–21601 (2009).
Zhou, A. et al. Structure and conformational states of the bovine mitochondrial ATP synthase by cryo-EM. Elife 4, e10180 (2015).
Gu, J. et al. Cryo-EM structure of the mammalian ATP synthase tetramer bound with inhibitory protein IF1. Science 364, 1068–1075 (2019).
Guo, H., Bueler, S. A. & Rubinstein, J. L. Atomic model for the dimeric FO region of mitochondrial ATP synthase. Science 358, 936–940 (2017).
Horvath, S. E. & Daum, G. Lipids of mitochondria. Prog. Lipid Res. 52, 590–614 (2013).
Meier, T., Matthey, U., Henzen, F., Dimroth, P. & Müller, D. J. The central plug in the reconstituted undecameric c cylinder of a bacterial ATP synthase consists of phospholipids. FEBS Lett. 505, 353–356 (2001).
Muhleip, A., McComas, S. E. & Amunts, A. Structure of a mitochondrial ATP synthase with bound native cardiolipin. Elife 8, e51179 (2019).
Allegretti, M. et al. Horizontal membrane-intrinsic α-helices in the stator a-subunit of an F-type ATP synthase. Nature 521, 237–240 (2015).
Mazhab-Jafari, M. T. et al. Atomic model for the membrane-embedded VO motor of a eukaryotic V-ATPase. Nature 539, 118–122 (2016).
Roh, S. H. et al. The 3.5-Å cryoEM structure of nanodisc-reconstituted yeast vacuolar ATPase Vo proton channel. Mol. Cell 69, 993–1004 (2018).
Zhang, L. & Hermans, J. Hydrophilicity of cavities in proteins. Proteins 24, 433–438 (1996).
Davies, K. M. et al. Macromolecular organization of ATP synthase and complex I in whole mitochondria. Proc. Natl Acad. Sci. USA 108, 14121–14126 (2011).
Murphy, B. J. et al. Rotary substates of mitochondrial ATP synthase reveal the basis of flexible F1-Fo coupling. Science 364, eaaw9128 (2019).
He, J. et al. Assembly of the membrane domain of ATP synthase in human mitochondria. Proc. Natl Acad. Sci. USA 115, 2988–2993 (2018).
Jiko, C. et al. Bovine F1Fo ATP synthase monomers bend the lipid bilayer in 2D membrane crystals. Elife 4, e06119 (2015).
Gerle, C. On the structural possibility of pore-forming mitochondrial FoF1 ATP synthase. Biochim. Biophys. Acta 1857, 1191–1196 (2016).
Pierce, B. G. et al. ZDOCK server: interactive docking prediction of protein–protein complexes and symmetric multimers. Bioinformatics 30, 1771–1773 (2014).
Papageorgiou, S., Melandri, A. B. & Solaini, G. Relevance of divalent cations to ATP-driven proton pumping in beef heart mitochondrial F0F1-ATPase. J. Bioenerg. Biomembr. 30, 533–541 (1998).
Tucker, W. C. et al. Observation of calcium-dependent unidirectional rotational motion in recombinant photosynthetic F1-ATPase molecules. J. Biol. Chem. 279, 47415–47418 (2004).
Ader, N. R. et al. Molecular and topological reorganizations in mitochondrial architecture interplay during Bax-mediated steps of apoptosis. Elife 8, e40712 (2019).
Carraro, M. et al. High-conductance channel formation in yeast mitochondria is mediated by F-ATP synthase e and g subunits. Cell. Physiol. Biochem. 50, 1840–1855 (2018).
Nesci, S., Trombetti, F., Algieri, C. & Pagliarani, A. A therapeutic role for the F1FO-ATP synthase. SLAS Discov. 24, 893–903 (2019).
Smith, A. L. Preparation, properties and conditions for assay of mitochondria: slaughterhouse material, small-scale. Methods Enzymol. 10, 81–86 (1967).
Letts, J. A., Degliesposti, G., Fiedorczuk, K., Skehel, M. & Sazanov, L. A. Purification of ovine respiratory complex I results in a highly active and stable preparation. J. Biol. Chem. 291, 24657–24675 (2016).
Runswick, M. J. et al. The affinity purification and characterization of ATP synthase complexes from mitochondria. Open Biol. 3, 120160 (2013).
Scheres, S. H. RELION: implementation of a Bayesian approach to cryo-EM structure determination. J. Struct. Biol. 180, 519–530 (2012).
Zheng, S. Q. et al. MotionCor2: anisotropic correction of beam-induced motion for improved cryo-electron microscopy. Nat. Methods 14, 331–332 (2017).
Rohou, A. & Grigorieff, N. CTFFIND4: fast and accurate defocus estimation from electron micrographs. J. Struct. Biol. 192, 216–221 (2015).
Hahn, A. et al. Structure of a complete ATP synthase dimer reveals the molecular basis of inner mitochondrial membrane morphology. Mol. Cell 63, 445–456 (2016).
Bepler, T. et al. Positive-unlabeled convolutional neural networks for particle picking in cryo-electron micrographs. Nat. Methods 16, 1153–1160 (2019).
Kelley, L. A., Mezulis, S., Yates, C. M., Wass, M. N. & Sternberg, M. J. The Phyre2 web portal for protein modeling, prediction and analysis. Nat. Protoc. 10, 845–858 (2015).
Emsley, P. & Cowtan, K. Coot: model-building tools for molecular graphics. Acta Crystallogr. D Biol. Crystallogr. 60, 2126–2132 (2004).
Adams, P. D. et al. PHENIX: a comprehensive Python-based system for macromolecular structure solution. Acta Crystallogr. D Biol. Crystallogr. 66, 213–221 (2010).
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, 249–255 (2009).
Larkin, M. A. et al. Clustal W and Clustal X version 2.0. Bioinformatics 23, 2947–2948 (2007).
Ashkenazy, H., Erez, E., Martz, E., Pupko, T. & Ben-Tal, N. ConSurf 2010: calculating evolutionary conservation in sequence and structure of proteins and nucleic acids. Nucleic Acids Res. 38, W529–W533 (2010).
Pravda, L. et al. MOLEonline: a web-based tool for analyzing channels, tunnels and pores (2018 update). Nucleic Acids Res. 46, W368–W373 (2018).
Williams, C. J. et al. MolProbity: more and better reference data for improved all-atom structure validation. Protein Sci. 27, 293–315 (2018).
Barad, B. A. et al. EMRinger: side chain-directed model and map validation for 3D cryo-electron microscopy. Nat. Methods 12, 943–946 (2015).
Pettersen, E. F. et al. UCSF chimera—a visualization system for exploratory research and analysis. J. Comput. Chem. 25, 1605–1612 (2004).
Krissinel, E. & Henrick, K. Inference of macromolecular assemblies from crystalline state. J. Mol. Biol. 372, 774–797 (2007).
We thank J. Novacek from CEITEC (Brno, Czech Republic) for assistance with collecting the FEI Krios dataset and iNEXT for providing access to CEITEC. We thank the IST Austria EM facility for access and assistance with collecting the FEI Glacios dataset. Data processing was performed at the IST high-performance computing cluster. This work has been supported by iNEXT EM HEDC (proposal 4506), funded by the Horizon 2020 Programme of the European Commission.
The authors declare no competing interests.
Peer review information Peer reviewer reports are available. Inês Chen was the primary editor on this article and managed its editorial process and peer review in collaboration with the rest of the editorial team.
Publisher’s note Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
Extended Data Fig. 1 Purification of mammalian ATP synthase and processing chart of the State 1a F1Fo maps.
a, The last step of purification (preparation in LMNG, no divalent cations added) was sucrose gradient with fractions collected using Biocomp Instruments Gradient Station allowing A280 absorbance recording. In the elution profile M indicates monomeric peak used for collection of data on Krios TEM leading to F1Fo monomer structure, D is a dimeric peak and fractions of larger MW were used for collection of data on Glacios TEM leading to multimer structures. The associated Native and SDS-PAGE gels are shown on the right. SDS-PAGE shows expected subunit composition of the mammalian ATP synthase. b, The ‘monomer’ dataset example micrograph and 2D classes examples. c, The ‘multimer’ dataset example micrograph and 2D classes examples. d, The processing strategy for the ‘monomer’ dataset leading to the Fo consensus class map and to the composite state 1a map as described in the Methods section. e, State 1a map from the consensus refinement with F1Fo mask (at 3.8 Å overall resolution, local resolution-filtered in Relion due to lower resolution in the Fo domain) is shown alongside and overlayed with the composite 1a state F1Fo map, to illustrate their global similaritity.
Extended Data Fig. 2 Processing chart of different rotational states of the monomer and of dimers from the ‘monomer’ Krios dataset.
The strategy is described in the Methods section.
Extended Data Fig. 3 Different oligomeric states from the ‘multimer’ Glacios dataset and resolution estimates of State 1a structure determination from the ‘monomer’ Krios dataset.
a, Side-view of the tetramer from the porcine study37 with indicated angles (measured using rotational axes going through the c-ring and F1 of each fitted monomer) and distances (between the IMS side centers of the c-rings) between the protomers. b, Different oligomeric states of ovine F1Fo from the ‘multimer’ dataset (LMNG, no divalent cations) with angles and distances indicated. In the ‘up-down’ dimers the Fo-Fo contacts were tight, with direct interactions between subunits DAPIT and 6.8pl from each monomer (leftmost) or between the c-rings and DAPIT subunits (second from the left). c, Processing chart of different oligomeric states from the ‘multimer’ dataset. d, Gold-standard Fourier shell correlation (FSC) curves for the overall and F1 / Fo-focused refinements of state 1a F1Fo, as well as corresponding local resolution maps estimated with Relion 3.0 implementation. Note that the resolution scale bars differ. e, Gold-standard Fourier shell correlation (FSC) curve for the Fo-focused refinement of the consensus Fo class from all rotational states, as well as corresponding local resolution map. f, The map versus model FSC curves for the consensus Fo and state 1a F1Fo models.
Representative densities of subunits α (a), β (b), OSCP (c), γ (d), catalytic ADP (o), and non-catalytic ADP and Mg2+ (p) in the F1-focus refined state 1a map. Densities for ATP8 (e), a (f), c (g), DAPIT (h), e (i), f (j), g (k), and IMS-side lipid (n) are from the Fo consensus class map. Matrix-side lipid density (m) is from the Fo-focus refined 1a state map. In panel (l) the arginine checkpoint is shown from the matrix side (arginine is marked with * here and in panel f). Note the absence of density between E58c and R159a indicating the absence of salt bridge.
a, A scheme of 360° catalytic cycle with dwell angles, catalytic events and nucleotide occupancies indicated, discussed in the Supplementary Note 1. Different conformations of the three catalytic β subunits (gray, salmon and yellow) are color-coded in the circumference as green for βDP, magenta for βTP and blue for βE. Note that during one 120° rotation between the two ATP binding dwells, starting from top and moving anti-clockwise, the ATP binding, Pi release and ATP hydrolysis generate 65°, 25° and 30° rotations respectively, while ADP is released before Pi. b, Scheme 1 for MgADP inhibition, discussed in the Supplementary Note 1. c, Scheme 2 for MgADP inhibition. d, Different rotational states of ovine F1Fo and their distribution (% of particles in the class), with the degree of subunit γ rotation shown. Subunits α and β of in-between states (1_2, 2_3 and 3_1) are transparent to indicate an ambiguous fit of some of these subunits into the density. The degree of subunit γ rotation from in-between to the main states is roughly consistent with angles between dwells in a.
Extended Data Fig. 6 Comparison of different species, sequence conservation in F1Fo and surface properties of Fo.
a, Gradual flattening of the joint end of the key helix hairpin in subunit a of different ATPases. Stuctures are aligned by the c subunit that directly interacts with the arginine checkpoint in subunit a. Subunits DAPIT/k of the mitochondrial enzymes are also shown to illustrate their roles in bending helices towards the c-ring. b, Complementary surface charges of the c-ring (bottom) and the interacting surface of the central stalk (top) are shown on the right. Solvent-accessible surface is shown colored red for negative, white for neutral and blue for positive surface charges. State 1a model is shown on the left for orientation. c, Comparison of the Fo interactions in the mammalian (top) and yeast (bottom) dimers. Note the extensive contacts of subunits a and i(6.8pl) in yeast (circled). d, Mammalian ATP synthase model (in two views rotated for clarity) colored according to residue conservation from blue (most conserved) to red (least conserved). Most conserved areas are indicated. e, Side views of the dimeric Fo region with the approximate membrane position indicated according to exposed hydrophobic areas. (Left) Solvent-accessible surface charge distribution. (Right) Protein surface colored according to the Eisenberg hydrophobicity scale from white (hydrophobic) to red (hydrophilic). From both representations it is clear that the membrane has to be highly curved to accommodate a dimer (or even a monomer).
a, b, Lipid occupancies. a, Fo density of the consensus class from all the rotational states combined (3.8 Å resolution) shows clear density of the IMS-side lipid (ISL) while the matrix-side (MSL) lipid density (circled area) is blurred/averaged due to MSL rotation with the c-ring (matrix side is on top). b, In the Fo density of state 1a only (4.2 Å resolution) the MSL is better resolved due to its fixed position in one state. Density for ISL at the bottom is visible in both cases since ISL is attached to subunit e and does not rotate with the ring. c, Surface charge distribution shows that the c-ring cavity is highly hydrophobic, with lipids inside shown as spheres. The ring of V16c, indicated with green sticks, serves as a bottleneck in the 310 helix section of subunit c. d, e, Cryo-EM density straight from Fo-focused auto-refinement (no post-processing, that is B-factor is not applied) for the consensus Fo class (d) and for state 1a only (e), showing that subunit e remains helical till the C-terminus, where it is connected (arrow) to the density in the center of the c-ring. f, g, CryoEM analysis of the enzyme purified in the presence of Ca2+ (or Mg2+) ions. f, Typical 2D classes from Mg and Ca datasets. g, Slices through the Fo domain density (between two yellow lines in the Overview) viewed in the projection along the vertical axis. In the Mg monomer, the fitted model is shown as a cartoon in magenta, with yellow arrow indicating c-ring and blue arrow subunit a. At similar resolution in the Mg monomer the c-ring is clearly seen as a circular density, while it is considerably distorted in Step0 and Step1 classes and absent in Step2 and Step3. All maps of Mg and Ca monomers were pre-aligned, so the slices show identical orientation.
Extended Data Fig. 8 Purification and cryoEM data processing of ATP synthase in the presence of Mg2+ or Ca2+.
These preparations were in digitonin and started from the addition of divalent cations (5 mM of either Mg or Ca) to the intact mitochondria. a-b, The last step of purification was sucrose gradient: for the Mg dataset collected fractions included mostly tetramers (a) while for the Ca dataset mostly monomeric fractions were included (b), although these species would overlap in elution. The associated Native gels are shown below, with SDS-PAGE of selected fractions on the left. SDS-PAGE shows expected subunit composition of the mammalian ATP synthase, with α/β subunits band marked. Both tetramer and monomer fractions are dominated by ATP synthase, although some amounts of CIII and CIV co-elute with monomers. These contaminants are separated out during cryo-EM data processing. Examples of micrographs and 2D classes are shown in (c) for the Mg dataset and in (d) for the Ca dataset. Processing chart for the Mg dataset is in (e) and for the Ca dataset in (f), with details in Methods.
About this article
Cite this article
Pinke, G., Zhou, L. & Sazanov, L.A. Cryo-EM structure of the entire mammalian F-type ATP synthase. Nat Struct Mol Biol 27, 1077–1085 (2020). https://doi.org/10.1038/s41594-020-0503-8
Redox Biology (2021)
Molecular and Supramolecular Structure of the Mitochondrial Oxidative Phosphorylation System: Implications for Pathology
Nature Communications (2021)
FEBS Letters (2021)
The inhibition of gadolinium ion (Gd3+) on the mitochondrial F1FO-ATPase is linked to the modulation of the mitochondrial permeability transition pore
International Journal of Biological Macromolecules (2021)