Structure of a mitochondrial ATP synthase with bound native cardiolipin

Abstract

The mitochondrial ATP synthase fuels eukaryotic cells with chemical energy. Here we report the cryo-EM structure of a divergent ATP synthase dimer from mitochondria of Euglena gracilis, a member of the phylum Euglenozoa that also includes human parasites. It features 29 different subunits, 8 of which are newly identified. The membrane region was determined to 2.8 Å resolution, enabling the identification of 37 associated lipids, including 25 cardiolipins, which provides insight into protein-lipid interactions and their functional roles. The rotor-stator interface comprises four membrane-embedded horizontal helices, including a distinct subunit a. The dimer interface is formed entirely by phylum-specific components, and a peripherally associated subcomplex contributes to the membrane curvature. The central and peripheral stalks directly interact with each other. Last, the ATPase inhibitory factor 1 (IF₁) binds in a mode that is different from human, but conserved in Trypanosomatids.

eLife digest

Every living thing uses the energy-rich molecule called adenosine triphosphate, or ATP, as fuel. It is the universal molecular currency for transferring energy. Cells trade it, mitochondria make it, and the energy extracted from it is used to drive chemical reactions, transport molecules across cell membranes, energize nerve impulses and contract muscles.

ATP synthase is the enzyme that makes ATP molecules. It is a multi-part complex that straddles the inner membrane of mitochondria, the energy factories in cells. The enzyme complex interacts with fatty molecules in the mitochondrial inner membrane, creating a curvature that is required to produce ATP more efficiently. The mitochondrial ATP synthase has been studied in many different organisms, including yeast, algae, plants, pigs, cows and humans. These studies show that most of these ATP synthases are similar to each other, but obtaining a high resolution structure has been a challenge.

Some single-cell organisms have unusual ATP synthases, which provide clues about how the enzyme evolved in pursuit of the most energy efficient arrangement. One such organism is the photosynthetic Euglena gracilis, which is closely related to the human parasites that cause sleeping sickness and Chagas disease.

Now, Mü̈hleip et al. have extracted ATP synthase from E. gracilis and reconstructed its structure using electron cryo-microscopy. The high resolution of this reconstruction allowed for the first time to examine the fatty molecules associated with ATP synthase, called cardiolipins. This is important, because cardiolipins are thought to modulate the rotating motor of the enzyme and affect how the complex sits in the membrane.

The analysis revealed that the ATP synthase in E. gracilis has 29 different protein subunits, 13 of which are only found in organisms of the same family. Some of the newly discovered subunits are glued together by fatty molecules and extend into the surrounding mitochondrial membrane. This distinctive structure suggests an adaptation which likely evolved independently in E. gracilis for efficiency.

These results represent an important advance in the field, and provide direct evidence for the functional roles of cardiolipin. This information will be used to reconstruct the evolution of this mighty molecule and to further study the roles of cardiolipin in energy conversion. Moreover, the analysis identified similarities between the ATP synthase in E. gracilis and human parasites, which could provide new therapeutic targets in disease-causing parasites.

Introduction

The mitochondrial ATP synthase is a membrane protein complex that generates most of the ATP in eukaryotic cells. The synthesis of ATP from ADP and inorganic phosphate proceeds via rotary catalysis, which uses the energy of the electrochemical gradient across the mitochondrial inner membrane. The translocation of protons through the membrane-bound F_o part, mediated by subunit a, drives the rotation of a membrane-embedded c-ring and the attached central stalk, which together form the rotor. The torque of the rotor against the stator subunits induces conformational changes in the (αβ)₃ headpiece, thereby triggering catalysis (Abrahams et al., 1994; Noji et al., 1997). The mitochondrial ATP synthase forms dimers, which in turn associate into dimer rows along the high-curvature membrane regions of the cristae (Davies et al., 2012; Paumard et al., 2002; Strauss et al., 2008). Loss of ATP synthase dimers results in aberrant cristae morphology, indicating that dimers are required for membrane bending and proper cristae formation in mitochondria (Davies et al., 2012; Paumard et al., 2002).

Previous biochemical and mass spectrometry analysis showed that a highly divergent ATP synthase with numerous new subunits is found in the phylum of Euglenozoa, belonging to the Excavata supergroup (Perez et al., 2014; Yadav et al., 2017; Zíková et al., 2009). A combination of transcriptome analysis and sequencing of nuclear and mitochondrial genomes showed the phylogenetic relationship between the protozoan Euglena gracilis and Kinetoplastids, which include human parasites such as Trypanosoma and Leishmania that cause sleeping sickness, Chagas disease, and Leishmaniasis (Dobáková et al., 2015; Ebenezer et al., 2019). However, while the reported recruitment of additional subunits in the Euglenozoan ATP synthase implies a distinct architecture, neither the complete composition, nor the functions or structures of the specific subunits have been reported. Previous attempts were limited by resolution (Mühleip et al., 2017) or restricted to the conserved F₁ subcomplex (Montgomery et al., 2018). In addition, lipids are known to be important to the structure and function of mitochondrial ATP synthases (Eble et al., 1990; Kühlbrandt, 2019; Laage et al., 2015; Srivastava et al., 2018). Proper cristae formation specifically depends on cardiolipin (CL) (Mileykovskaya and Dowhan, 2009), an anionic lipid that accounts for 20% of total lipids in the inner mitochondrial membrane (Calvayrac and Douce, 1970; Daum and Vance, 1997; Zinser and Daum, 1995). Cardiolipin is also essential for the activity of isolated mitochondrial ATP synthases (Laird et al., 1986; Pitotti et al., 1972; Santiago et al., 1973), and promotes the formation of dimer rows (Acehan et al., 2011). Although bound lipids have previously been observed in structures of rotary ATPases (Klusch et al., 2017; Murphy et al., 2019; Vasanthakumar et al., 2019), the atomic models of recently reported cryo-EM structures of mitochondrial ATP synthases do not include cardiolipin (Gu et al., 2019; Guo et al., 2017; Klusch et al., 2017; Murphy et al., 2019; Srivastava et al., 2018).

To reveal how the mitochondrial ATP synthase is modulated by lipids and organised into a dimer shaping discoid cristae in Euglenozoan mitochondria, we determined the structure of the ATP synthase dimer from E. gracilis. The atomic model of the entire 2-MDa complex contains 29 different subunits (eight newly identified) and provides a comprehensive description of a markedly distinct mitochondrial ATP synthase dimer, including a previously unseen binding mode of its natural inhibitor protein IF₁. The membrane region was determined to 2.8 Å resolution, enabling the identification of a structurally divergent subunit a and visualisation of 37 associated native lipids. Importantly, cardiolipin binding sites are found at the rotor-stator interface, dimer interface, and in a peripheral F_o cavity. These data provide insight into protein-lipid interactions in the mitochondrial ATP synthase and its evolution, suggesting functional roles of lipids in proton translocation, dimerization and stabilisation.

Results and discussion

Overall structure

The mitochondrial ATP synthase dimer was purified natively from E. gracilis and subjected to cryo-EM structure determination (Supplementary file 1). Symmetry expansion of the pseudo-C₂-symmetric dimer particles was used to classify individual F₁/c-ring monomers into rotational states 1, 2 and 3 (named according to bovine nomenclature; Zhou et al., 2015) and resolve structures at 3.0- to 3.9 Å resolution. Using masked refinement, maps of the membrane region, the rotor and the peripheral stalk tip were refined to 2.8 and 3.3 and 3.8 Å resolution, respectively, enabling the construction of atomic models (Figure 1—figure supplements 1–2; Video 1; Supplementary file 2). The model of the complete mitochondrial ATP synthase contains 29 different proteins, of which 14 are phylum-specific, displaying a distinct architecture, particularly in the membrane region. The interactions between the monomers are expanded, resulting in a 45° dimer angle, compared to ~100° in yeast and mammals (Figure 1A) (Davies et al., 2012; Gu et al., 2019; Hahn et al., 2016). The membrane-bound F_o region is composed of 22 proteins, of which 13 have no homologs in animals and fungi. The well-resolved membrane region allowed the identification of 37 native lipids, leading to the assignment of 25 cardiolipins, whereas the 12 remaining phophoslipids could not be unambiguously identified and were modelled as phosphatidic acid.

Figure 1. Structure of the E. gracilis ATP synthase dimer.

(A) Atomic model of the complete E. gracilis ATP synthase dimer with both subcomplexes in rotational state-1. The 2-MDa dimeric F₁F_o-complex contains 29 different proteins. Dashed lines indicate C₂-symmetry axis and 45° dimer angle. (B) OSCP/F₁/c-ring subcomplex in rotational state-1, bound to its natural inhibitor protein IF₁ (cyan), remaining F_o transparent. (C) Density map showing the lumen-exposed F_o region. Detergent belt shown in yellow; c-ring β-barrel in dark grey, F_o subunits as in (A). (D) Close-up of the lumenal interface of ATPEG1 (blue) with the c-ring (grey). The interaction occurs mostly via hydrophobic interactions (blue and grey sticks). (E–G) Euglenozoa-specific F_o-subunits with known folds. (E) ATPTB1 in blue superposed with Mdm38 (PDB ID: 3SKQ) (Lupo et al., 2011), six conserved helices coloured yellow, rest grey. (F) ATPTB3 in blue superposed with a bacterial homoisocitrate dehydrogenase in orange (PDB ID: 4YB4)(Takahashi et al., 2016), adopts a Rossman-fold. (G) ATPEG5 in yellow is a structurally conserved ortholog of the cytochrome c oxidase subunit VIb superfamily; bovine subunit VIb in purple (PDB ID: 2Y69) (Kaila et al., 2011).

Figure 1—figure supplement 1. Cryo-EM data processing and classification scheme.

(A) Representative micrograph. Scale bar 500 nm. (B) Representative 2D classes. (C) Fourier Shell Correlation (FSC) plots of all seven maps shown in D, with resolutions calculated according to the 0.143-cutoff criterion. (D) Cryo-EM data processing scheme. Final maps include the membrane region (from all rotational states), F₁/c-ring maps from rotational states 1–3, the rotor and peripheral stalk tip (both from rotational state-1), as well as a consensus map of the ATP synthase dimer with both F₁/c-ring subcomplexes in rotational state-1. Rotational state-3 displayed partial occupancy of IF₁ and was separated into two classes with or without bound inhibitor using focused classification.

Figure 1—figure supplement 2. Local resolution estimation and model-map-correlations.

(A–G) Density maps of masked regions of the ATP synthase coloured according to local resolution (left panels, local resolution as highlighted in color bars in Å) with Model-Map-FSC curves of respective atomic model refined into one half map to give FSC_work (black) and FSC_test (red)(right panels). (A) Membrane region, (B) consensus map of complete ATPS synthase dimer with both monomers in rotational state-1. (C–E) F₁/c-ring maps of rotational states 1–3, (F) rotor in rotational state-1, (G) peripheral stalk tip of rotational state-1.

Video 1. Density map of E. gracilis ATP synthase dimer with regions corresponding to protein shown in grey and the detergent belt coloured gold.

Download video file ^{(17.9MB, mp4)}

Following reports of mitochondrial ATP synthases from various organisms, different names have previously been given to subunits performing the same role in F-type ATP synthases (Kühlbrandt, 2019). For the mitochondrial ATP synthase from E. gracilis, we adopt a nomenclature that is consistent with the conserved F_o subunits from yeast (subunits a-d, f, i/j, k, 8, OSCP) (Guo et al., 2017) and euglenozoa-specific subunits previously identified in Trypanosoma brucei (ATPTB1, 3, 4, 6, 12) (Perez et al., 2014; Zíková et al., 2009), whereas the additional subunits are named ATPEG1 to 8, according to their molecular weight (Supplementary file 3).

Identification of the transmembrane subunits

All known mitochondrial ATP synthases form dimers in the membrane through transmembrane F_o subunits. However, in E. gracilis, only subunit c and a set of phylum-specific subunits were identified (Perez et al., 2014; Yadav et al., 2017; Zíková et al., 2009), suggesting a divergent F_o composition. The core subunit a mediates proton translocation and contains the strictly conserved R176 (Saccharomyces cerevisiae numbering). Despite its functional importance, subunit a was identified neither in the E. gracilis genome project (Ebenezer et al., 2019), nor in the mitochondrial genome and transcriptome analysis (Dobáková et al., 2015). Our cryo-EM map allowed tracing of subunit a, for which we identified six structurally conserved membrane-embedded helices (H1_a to H6_a) directly from side-chain densities (Figure 2—figure supplement 1A to C). Using this information, we then found the matching sequence in the available genomic data and mapped it to a mitochondrial contig, which also contains subunit 8 and nad1 in a single open reading frame (Figure 2—figure supplement 1E). Thus, by combining the information from cryo-EM and sequencing, we report the most divergent subunit a found up to date.

In all previously reported ATP synthase structures, the horizontal helix H5_a bends around the c-ring, following its curvature, thereby contacting four of the ten subunits in the c-rings of yeast and algae (Figure 2B,D) (Allegretti et al., 2015; Guo et al., 2017; Hahn et al., 2016). By contrast, in the E. gracilis structure, the N-terminal part of H5_a is kinked towards the lumen, and therefore does not interact with the c-ring, and instead extends towards the lumenal membrane surface (Figure 2A,C). This structural rearrangement results in a smaller interface between subunits a and c, with only three c-subunits forming interactions, which also has implications for the formation of the matrix half-channel, as discussed below.

Figure 2. E. gracilis subunit a fold comparison.

Top view (upper panel) and side view (lower panel) of the E. gracilis (left) and S. cerevisiae (right) (Guo et al., 2017) subunit a (green) and c-ring (grey). Both structures contain the conserved H1-6_a, with E. gracilis displaying two helices (H4’_a and H4’’_a) in an extension segment and a C-terminal extension. Whereas the N-terminal region of H5_a (dark green) is kinked towards the c-ring in the yeast complex, it extends towards the lumen in the E. gracilis structure, thereby diminishing its interface with the c-ring. Unlike its yeast homolog, the N-terminus of E. gracilis subunit a is not involved in dimerisation, but contributes a strand to a β-sheet along the lumenal side of the detergent micelle.

Figure 2—figure supplement 1. Identification of subunits a and 8.

(A-D) Map quality and model fit of E.gracilis subunit a. H4’’_a, H5_a and H6_a (A), subunit a N-terminus (B), C-terminus (C), and subunit 8 N-terminus and transmembrane helix (D) are shown with map density as mesh. (E) Sequence of the translated E. gracilis genome contig containing the newly identified subunit a (green), subunit 8 (pink), as well as NADH dehydrogenase subunit 1 (nad1, blue) in a single predicted open reading frame (contig_684056 of PRJEB27422, E. gracilis Genome Project). An identical consensus contig was obtained by assembling sequence reads of the E. gracilis mitochondrial genome (PRJNA294935)(Dobáková et al., 2015), indicating that subunit a and subunit 8 are mitochondrially encoded.

In the yeast mitochondrial ATP synthase, subunit a interacts with transmembrane subunits b, f,  i/j, k, 8 and membrane-associated subunit d (Guo et al., 2017). Since no homologs were reported for any of these subunits in E. gracilis or T. brucei (Perez et al., 2014; Yadav et al., 2017; Zíková et al., 2009), we next investigated their putative location through superimposition of our F_o model with the yeast counterpart (Guo et al., 2017). Based on the matching position and topology of the transmembrane helices as well as conserved positions of interactions with subunit a, we identified all six associated subunits, which are structurally conserved, but display no significant sequence similarity to yeast counterparts (Figure 3). Subunits b,  f, i/j, k and 8 contain a single transmembrane helix associated with subunit a, whereas subunit d forms a clamp around subunit 8 at the base of the peripheral stalk, containing a structurally conserved two-helix motif at the matrix side of the membrane (Figure 3A). Finally, subunit k is bound peripherally to subunit a, as in yeast, however the H5_a kink results in a 15-Å displacement of the transmembrane helix of subunit k away from the c-ring, compared to its yeast counterpart (Figure 3B). These data show that despite sequence divergence, the assembly of the central F_o subunits around subunit a is architecturally conserved between Euglenozoa and Metazoa.

Figure 3. Conserved subunits of the F_o region.

(A) Side view of the conserved E. gracilis F_o subunits. Transmembrane helices with structural equivalents in yeast are labelled. (B) Top view of the superimposed conserved F_o subcomplexes from E. gracilis (coloured) and yeast (grey) PDB ID: 6B2Z (Guo et al., 2017). Although subunit k does not superimpose well, it occupies the same position relative to the H5_a.

The striking architectural divergence of the E. gracilis ATP synthase dimer is brought about by euglenozoa-specific subunits and extensions of the structurally conserved F_o subunits, which render them on average 2.5 times larger than in the yeast mitochondrial ATP synthase. Only subunit b is truncated. The extensions of the conserved F_o subunits are mostly involved in forming interactions with the euglenozoa-specific subunits, thus providing a platform for the observed increased molecular mass of the F_o (Figure 4—figure supplement 1A,C).

The additional 13 euglenozoa-specific F_o subunits determine the architecture of the ATP synthase dimer, giving the F_o a markedly different overall shape, making it almost three times the size of its yeast counterpart (Figure 4—figure supplement 1). They contribute to the dimerization interface, the peripheral stalk and F_o periphery. The C-terminal helix of euglenozoa-specific ATPEG1 (H5_EG1) extends 50 Å from the membrane region into the lumen, where it interacts with the N-terminal extensions of subunit c, which together form a ten-stranded β-barrel (Figure 1C and D) protruding ~20 Å into the lumen. The N-terminal residues of subunit c (A24, I25) form a hydrophobic interface with hydrophobic residues (M148, M152, L155, I159, L163) of the amphipathic ATPEG1 helix. The position of the lumenal H5_EG1 on the c-ring β-barrel remains largely unchanged in all three rotational states, suggesting a mechanism of transient rotor-stator interaction during c-ring rotation. A similar lumenal interaction has previously been reported in the bovine ATP synthase, where subunit e extends from the membrane to contact the c-ring (Zhou et al., 2015). In the porcine ATP synthase tetramer, subunit e has been proposed to interact with the 6.8 kDa proteolipid, which has been suggested to reside inside the c-ring (Gu et al., 2019).

Other euglenozoa-specific F_o subunits contain structural domains that were shown to be functionally important in mitochondria (Figure 1E to G). ATPTB1 is a membrane-associated protein on the matrix side of the F_o periphery that adopts an Mdm38-like fold, which was shown to associate with yeast mitochondrial ribosomes at the inner mitochondrial membrane (Frazier et al., 2006). ATPTB3 is an isocitrate dehydrogenase ortholog that adopts a Rossman fold located at the tip of the peripheral stalk. ATPEG5 is a structurally conserved ortholog of the cytochrome c oxidase subunit VIb superfamily.

Dimer interface and associated lipids

The defining feature of mitochondrial ATP synthases is the formation of dimers in the crista membrane. In yeast, the two monomers are connected through F_o, and the dimer interface is formed on the lumenal side by the conserved F_o subunits a and i/j, as well as subunits k and e (Guo et al., 2017). Our atomic model of the E. gracilis mitochondrial ATP synthase shows that in contrast to yeast, subunits a, i/j, k do not contribute to the dimer interface, which instead is formed by species- and phylum-specific subunits and extensions of apparent homologs (Figure 4, Figure 4—figure supplement 1, Figure 4—figure supplement 2). The extensive dimer contacts are stacked across three layers: the matrix side, the transmembrane region and the lumenal side (Figure 4A and Figure 4—figure supplement 1A,C). On the matrix side, an extension of subunit d adopts an elaborated ferredoxin-like all-β fold that forms a dimer interface close to the symmetry axis (Figure 4A and Figure 4—figure supplement 1E,F). The curved H3 of ATPEG1 forms a homotypic dimerisation motif along the matrix surface of the membrane plane. ATPEG2 contributes to the dimer interface with both a transmembrane helix that interacts with ATPEG1 and its termini, which link the two F_o-parts, each extending into the rotor-stator interface of one monomer. On the lumenal surface, ATPEG1 interacts with the C-terminal extension of subunit f. The different F_o subunit composition and their involvement in dimer formation result in a 45° dimer angle, compared to ~100° in yeast (Figure 1A) (Guo et al., 2017). Thus, despite the presence of conserved F_o subunits and the contribution to their extensions to the dimer interface (Figure 4—figure supplement 2), the E. gracilis mitochondrial ATP synthase displays a fundamentally different dimer architecture, when compared to the yeast (Figure 4—figure supplement 1), mammalian and Polytomella ATP synthases (Gu et al., 2019; Murphy et al., 2019), suggesting that dimer formation evolved independently in different lineages.

Figure 4. The dimer interface and associated lipids.

(A and B) Views of the dimer interface along (A) and perpendicular (B) to the membrane plane. The dimerisation motifs (interacting subunits coloured) are stacked along the C₂-symmetry axis and formed by two copies of subunit d (red) and ATPEG1 (blue), which interacts with its symmetry-related copy, as well as ATPEG2 (green) and subunit f (yellow). Asterisks in (B) indicate positions of lipid-binding sites. (C to G) Close-ups of the lipid-binding sites indicated in (B). Bound lipids at the dimer interface identified as cardiolipin (CDL; C to E) or phospholipids modelled as phosphatidic acid (PL; F and G). Interacting residues (subunits coloured) include at least one arginine residue. Density shown as grey mesh.

Figure 4—figure supplement 1. Architecture and dimer interface comparison between the yeast and E. gracilis ATP synthase.

The E. gracilis (A,C) and yeast (B,D) F_o subcomplexes display different architectures. (A,C) The E. gracilis complex is held together by subunit d (red), subunit f (yellow), ATPEG-1 (light blue) and ATPEG2 (dark green). The subunit a/c-ring subcomplexes (light green and dark grey respectively) are offset along the along the F_o long axis. (B,D) The yeast ATP synthase dimer (PDB ID: 6B2Z) (Guo et al., 2017) is held together by subunit a (light green), subunit i/j, as well as subunit k (orange) and subunit g (dark blue, interaction not modelled). (E-F) Subunit d extension contributes to the dimer interface and peripheral stalk. (E) Subunit d (red) and subunit b (white) together form a ten-stranded all-β-fold at the dimer interface on the matrix side that contributes to the dimer interface. The smaller β-sheet (β2) is formed by subunit d, which adopts a ferredoxin-like fold containing a β-hairpin insertion (strands 4 an 5), which is part of the larger β-sheet (β1) that contains two β-strands (labelled A and B) of subunit b. (F) Top view of the dimer interface formed by subunit d, with interacting residues shown; green asterisk marks C₂-symmetry axis.

Figure 4—figure supplement 2. Species-specific subunits and extensions form the dimer interface.

The E. gracilis ATP synthase dimer side view (A) and bottom view (B). Phylum-specific subunits previously identified in T. brucei (ATPTB1, ATPTB3, ATPTB4, ATPTB6, ATPTB12, p18) shown in blue. Species-specific subunits with no detectable homologs outside euglenoids (ATPEG1-8) are shown in yellow. Conserved subunits with homologs or close structural equivalents in yeast are shown in grey, with species-specific extensions highlighted in red.

Figure 4—figure supplement 3. Bound lipids of the dimer interface.

Side view (A) and cut view (B) of the dimer interface with central cavity (grey rectangle in (A)) containing bound lipids (cardiolipin red, unidentified phospholipids light pink). Subunits involved in dimerization or lipid binding are highlighted. (B) View of ATP synthase monomer cut along a plane though the C₂-symmetry axis. Coloured rectangles highlight lipid binding sites at the dimer interface as shown in Figure 4C to G.

In addition to the described protein-protein interactions, we identified nine bound phospholipids occupying the dimer interface (Figure 4C to G, Figure 4—figure supplement 2). Five of them are cardiolipin molecules linking dimerising subunits close to the C₂-symmetry axis (Figure 4C to E). CDL11 links two horizontal helices of the two symmetry-related copies of ATPEG1, which extend along the matrix side of the membrane region (Figure 4E, Video 2). These protein-lipid interactions indicate a functionale role of cardiolipin in the stabilisation of the dimer contacts, which is consistent with its proposed role in mediating subunit interactions between the transmembrane helices in mitochondrial supercomplexes (Mileykovskaya and Dowhan, 2014; Wu et al., 2016).

Video 2. Atomic model of the E. gracilis ATP synthase dimer with bound lipids.

Download video file ^{(61.1MB, mp4)}

The cryo-EM map of the membrane region is shown as mesh. Closeup view of cardiolipin (CDL11) linking H1_EG1 and H3_EG1 of the two ATPEG1 copies at the matrix side of the membrane region.

Rotor-stator interface and proton path

Proton translocation occurs at the rotor-stator-interface, which is canonically formed in the membrane by horizontal helices H5_a, H6_a and the c-ring (Allegretti et al., 2015) (Figure 5—figure supplement 1B,D). In the E. gracilis ATP synthase, the essential R178 of H5_a is conserved, and interacts with Gln232 on H6_a. Intriguingly, we found two additional horizontal helices, contributed by ATPEG4 (H1_EG4) and subunit k (H1_k) (Figure 5A,C and Figure 5—figure supplement 1A). ATPEG4 is bound to subunit a in the membrane, and its amphipathic, horizontal helix (H1_EG4) is positioned close to the matrix side of the membrane, extending in parallel to H6_a at a distance of ~13 Å (Figure 5A), allowing it to interact with the same transmembrane helix of the c-ring as H5_a and H6_a (Figure 5—figure supplement 2A).

Figure 5. The rotor-stator interface is flanked by bound cardiolipin.

(A) View from the c-ring towards the membrane-embedded stator subunits. H5_a and H6_a are augmented by the tilted, amphipathic H1_EG4 (brown). Cardiolipin molecules flanking subunit a are shown in red (tails of acyl chains are mostly disordered and shown only for illustration). Proton half-channels on the lumen and matrix side are shown in orange and red respectively. Remaining subunits not shown for clarity. The conserved R178 and the H186 at the lumen channel exit are shown with interacting residues. (B) Entrance of the lumenal channel (orange) is lined by the termini of subunit f, ATPEG4, subunit 8, ATPEG3, as well as a lumenal segment of subunit i/j. Inside the F_o, the lumen channel is confined by transmembrane helices of subunits f and b. β-DDM occupying the exit of the lumen channel shown in orange with density map as mesh, c-ring in grey. Arrows indicate proposed path of proton flow. (C) Polar and protonatable residues between R178 and the matrix-side half channel (red mesh). Subunit k contributes a horizontal helix (H1_k) to the rotor-stator interface.

Figure 5—figure supplement 1. Comparison between E. gracilis and S. cerevisiae rotor-stator interfaces.

The E. gracilis ATP synthase (A,C) displays a larger rotor-stator interface than S. cerevisiae (B,D). (B) In yeast, the interactions (red) with the c-ring are mainly formed by the horizontal H5_a and H6_a in the membrane (green). (A) In E. gracilis, four horizontal helices (H5_a, H6_a, H1_EG4, H1_k) are found at the rotor-stator interface, with the H5_EG1 forming a lumenal c-ring interaction. (C,D) Interactions between the F_o stator (grey) and the c-ring (white transparent) are highlighted in red.

Figure 5—figure supplement 2. Native phospholipids of the rotor-stator interface.

Associated native phospholipids of the E. gracilis rotor-stator interface. (A) Top view of the rotor-stator interface. The offset half-channels are separated by the central R178 (lumenal channel with β-DDM shown in orange; internal end of matrix channel shown in red). The half channels are flanked on either side by a cardiolipin (shown in red). Species-specific structural elements constituting the rotor-stator interface near the matrix channel enclosed by dashed blue line. (F-H) Cardiolipins (B,C) and unidentified phospholipid (D) of the rotor-stator interface (map density shown in grey mesh).

The E. gracilis ATP synthase structure suggests a mechanism of proton translocation via two offset proton half channels, similar to those previously described in other F-type ATP synthases (Allegretti et al., 2015; Guo et al., 2017; Hahn et al., 2016). Protons enter the membrane region via the lumenal half-channel, which we traced as an internal cavity of the atomic model. The entrance of the lumenal half-channel is lined by the C-termini of subunit f and ATPEG4, extension of subunit i/j and the N-terminal regions of ATPEG3 and subunit 8, of which only the latter is structurally conserved in yeast (Figure 5B). Inside the F_o region, the lumenal half-channel is lined by the conserved transmembrane helices of subunits f and b and extends between H5_a, H6_a, as previously suggested (Guo et al., 2017). At the exit of the lumenal half-channel, we identified a β-DDM detergent molecule, with its hydrophilic maltose head group protruding into the channel and the acyl chain extending between H5_a and H6_a, confirming both the hydrophilic environment and accessibility of the membrane-intrinsic lumen channel (Figure 5B).

Proton translocation to the rotor-stator interface results in protonation of the conserved glutamate residue (E86 in E. gracilis) in the middle of the c-ring and subsequent counter-clockwise rotation of the c-ring when viewed from the F₁ to F_o (Noji et al., 1997). This proton transfer has been suggested to be ultimately mediated by a glutamate of H6_a (E223 in S. cerevisiae), which is paired with H185 of H6_a. This residue pair facing the lumenal half-channel is conserved in yeast, mammals and Polytomella (Gu et al., 2019; Guo et al., 2017; Klusch et al., 2017) but surprisingly not in E. gracilis (V225 and S187, respectively; Figure 5A). The absence of an acidic residue from the exit of the lumenal proton half-channel indicates that it is not strictly required for proton transfer to the c-ring in mitochondrial ATP synthases. Instead, this function appears to be compensated in our structure by H186 of H5_a, which extends towards the c-ring and interacts with Y228 of H6_a, thus forming an alternative residue pair at the lumenal channel exit (Figure 5B).

After almost a full rotation of the c-ring, the glutamate residue (E86) is deprotonated by R178 of subunit a. The translocated proton is then released into the matrix half-channel. In other ATP synthases, H5_a bends around the c-ring, thereby determining the channel path (Guo et al., 2017). Due to the unusually kinked H5_a, E. gracilis ATP synthase lacks the interaction of the N-terminal segment of H5_a following the c-ring curvature (Figure 2A). Instead, its functional role in forming the release channel is replaced by the N-termini of ATPEG4, ATPEG6 and ATPEG8 and subunit k which forms a fourth horizontal helix in the membrane that extends towards the c-ring (Figure 5C, Figure 5—figure supplement 2A). Thus, the reduced interface between subunits a and c is compensated by species-specific structural elements forming the matrix half channel (Figure 5C). Taken together with the horizontal H1_EG4 and the lumenal H5_EG1, the E. gracilis ATP synthase displays and increased number of c-ring interactions compared to its yeast counterpart. As a consequence, the E. gracilis rotor-stator interface displays larger buried surface area of ~830 Ų compared to ~480 Ų in yeast (Figure 5—figure supplement 1C and D).

Adjacent to the proton-half channels, the E. gracilis ATP synthase structure reveals two bound cardiolipins (CDL1, CDL2) flanking either side of the two horizontal helices H5_a and H6_a (Figure 5A and Figure 5—figure supplement 2A to C). The head groups of both lipids are bound around the middle of the membrane plane, with their acyl chains extending towards the rotor-stator interface. CDL1 is coordinated by R49 of ATPEG4, holding it near the loop connecting H5_a and H6_a. Near the matrix half-channel, CDL2 is coordinated collectively by R160 of H5_a, R50 of ATPEG6 and R22 of subunit k (Figure 5A; Figure 5—figure supplement 2B to D). Together, these two bound cardiolipins enclose the two horizontal membrane helices H5_a and H6_a, possibly acting to seal the F_o against proton leakage by recruiting a high density of acyl chains, as well as separating lipid and aqueous environments in the vicinity of the two half-channels. Thus, in addition to previous studies suggesting transient interactions of the metazoan c-ring rotor with cardiolipin (Duncan et al., 2016), the E. gracilis ATP synthase structure shows that some of the cardiolipin is bound specifically to the stator, indicating a potential functional role in proton translocation.

Peripheral F_o subcomplex and lipid cavity

A cluster of phylum-specific subunits is located at the F_o periphery, away from the dimer interface. Here, seven tightly associated subunits, ATPTB1, 6, 12 and ATPEG3, 5, 6, 8 form a subcomplex, connected to the conserved core in both, the lumen and matrix (Figure 6A,B). On the lumenal side, it is flanked by the terminal extensions of subunit a and subunit k (Figure 6—figure supplement 1A,B). The N-terminal ß-strand of subunit a, which is involved in the dimerization in yeast, shares a ß-sheet with ATPEG3, whereas the C-terminal of subunit k extending along the membrane plane interacts with six subunits of the subcomplex (all except ATPTB1). On the matrix side, ATPTB1 anchors the subcomplex to the conserved core through multiple contact sites. While the two moieties are extensively associated with each other outside the membrane, they are separated by more than 20-Å gap inside the membrane. Thus, a protein-enclosed membrane cavity is formed. In the cavity, we identified six bound cardiolipins (Figure 6B). To assess the lipid-binding capacity of the cavity, we performed coarse-grained molecular dynamics simulations of the entire mitochondrial ATP synthase dimer embedded in a phospholipid membrane containing 20% cardiolipin (Figure 6—figure supplement 2). The simulations indicate that the cavity is filled with a bilayer lipid array in which the lipid molecules can freely diffuse in or out of within the membrane (Video 3). Starting with random initial placement of phospholipids, the average residence time of cardiolipin was ~2.5 times higher than that of other lipid types included in the simulations (Figure 6—figure supplement 2E). The cardiolipin binding is induced by positively charged residues of subunits a, i/j, k, ATPTB1, ATPTB6, ATPEG3 extending into the cavity. To probe the dynamics of lipid binding, we calculated the probabilities of entering and remaining in the cavity. The lipid exhibiting the highest probability, especially over longer time intervals, is cardiolipin (Figure 6—figure supplement 2C,D), which is consistent with the assignment in the cryo-EM density maps (Figure 6B).

Figure 6. Peripheral F_o subcomplex and the peripheral stalk.

(A) Euglenozoa-specific subunits form a peripheral F_o subcomplex. Density of the F_o with proteins of the peripheral region coloured, c-ring model shown in grey, outline of the detergent belt (yellow dashed lines) with 2 nm offset towards the lumen indicated as determined by the density (transparent gold) (B) Atomic model of the F_o periphery, cavity lipids are shown in magenta. (C to E) Attachment of the peripheral stalk to F₁. (C) E. gracilis ATP synthase with proteins constituting the peripheral stalk coloured. (D) Side view of the peripheral stalk tip and F₁ (white, crown domain light grey). C-terminal helix of OSCP (light green) extending towards the membrane attaches OSCP to the rest of the peripheral stalk via subunit d (red). (E) The N-terminal extension of OSCP (yellow) interacts with the C-terminal extension of the rotor subunit γ (conserved region purple, extension dark blue).

Figure 6—figure supplement 1. The peripheral F_o subcomplex and inverted topology structural motif.

Side view (A) and front view (B) of the F_o region. The peripheral subcomplex (pink) and the conserved F_o-subunits (white) are joined together by ATPTB1 on the matrix side and by subunits a and k in the lumen and separated by the lipid-filled F_o cavity (B). (C) Hairpin helices of ATPTB1 and ATPEG3 extend into the membrane region from opposite sides in the membrane, where they interact, forming an inverted topology motif. (D) Inverted topology motif of the prokaryotic aspartate transporter Glt_Ph consisting of two hairpin loops (HP1 and HP2, PDB ID: 1XFH) (Yernool et al., 2004).

Figure 6—figure supplement 2. Coarse-grained molecular dynamics simulations of the E. gracilis ATP synthase dimer.

(A) The 45°-angle of the E. gracilis ATP synthase dimer (grey) generates significant membrane curvature (acyl chains in light blue; phosphate groups in orange, ethanolamine groups in red, choline groups in green), resulting in a 7 nm local displacement of the membrane from the bilayer plane (marked with bar). (B) Radial distribution function g (r) of the center of mass of head groups from different lipid types. The first two maxima indicate well-defined coordination shells around the transmembrane region. Phosphatidic acid (PA) and cardiolipin (CDL) have a higher density in the annulus of the transmembrane region than phosphatidyl choline (PC) and phosphatidylethanolamine (PE). (C) View of F_o cavity with orange sphere indicating region used for lipid binding analysis. (D) Probabilities of entering and staying bound in the cavity shown for each lipid type. (E) Mean residence time of each lipid type in the cavity. Error bars indicating the 90% confidence interval obtained from bootstrapping. Nonparametric significance tests were performed for each lipid type’s residence time against cardiolipin: * p-value<0.05, ** p-value<1×10⁻⁵. (F) Side view of F_o cavity with lipids forming a bilayer-like array.

Figure 6—figure supplement 3. Interactions of OSCP extension with F₁ and the peripheral stalk.

(A, B) Peripheral stalk architectures of S. cerevisiae (A) (Guo et al., 2017) and E. gracilis (B) viewed from the F₁ (not shown) towards the peripheral stalk. The two structures have subunits b, d, 8 and OSCP in common. Unlike in S. cerevisiae, subunit f does not contribute to the peripheral stalk in E. gracilis. Subunit F6 is not found in E. gracilis. Red arrowheads indicate different heights of OSCP attachment to subunit b and subunit d respectively. (C) The C-terminal extension of E. gracilis OSCP (red rectangle) extends in between the peripheral stalk (dark grey) and F₁ (light grey). (D) Close-up of OSCP/subunit d interaction in the E. gracilis peripheral stalk. The indicated residues are conserved in Euglenozoa. (E) The E. gracilis OSCP contains conserved structural elements (light green)(Srivastava et al., 2018) with the N-terminal domain consisting of helices 1,2,5 and 6 and the C-terminal domain consisting of a four-stranded β-sheet and helices 7 and 8 (dihydrolipicolinate reductase domain 2-like fold). The N-terminal extension (light yellow) interacts with subunit γ (blue) of the rotor, whereas the C-terminal extension (orange) is anchored to the rest of the peripheral stalk. (F-H) Multiple sequence alignments of the OSCP C-terminal region (F), with the Euglenozoa-specific extension highlighted red; OSCP N-terminal region (G) with the N-terminal helix found in the E. gracilis OSCP highlighted yellow; red rectangles indicate predicted cleavage sites of mitochondrial processing peptidase. (H) Subunit γ C-terminal region, with euglenozoa-specific extension highlighted red; Bos tarus (Bt), Homo sapiens (Hs), Caenorhabditis elegans (Ce), Saccharomyces cerevisiae (Sc), the trypanosomatids Trypanosoma brucei (Tb), Leishmania major (Lm), Endotrypanum monterogeii (Em), Crithidia fasciculata (Cf), Leptomonas pyrrhocoris (Lp), Blechomonas ayalai (Ba), Phytomonas sp. (Ps) and the euglenoid Euglena gracilis (Eg).

Video 3. Coarse-grained molecular dynamics simulation showing the diffusion of lipids into and out of the peripheral F_o cavity (orange sphere, as described in Figure 6—figure supplement 2C) within the lipid bilayer over a period of 4 µs.

Download video file ^{(25.7MB, mp4)}

A sliced view of the membrane bilayer is initially shown for reference, but later removed to allow viewing of the binding and unbinding phospholipids. Cardiolipin is indicated with purple, phosphatidic acid with yellow, phophatidylethanolamine with red, phosphatidylcholine with cyan acyl chains respectively. Lipids considered to be bound in the beginning or end of the simulation are visualized, demonstrating that cardiolipin replaces other lipids in the cavity during the simulation.

The membrane region around the identified F_o cavity is displaced ~20 Å towards the lumen, thus contributing to its curvature (Figure 6A and Video 1). The displacement appears to be induced by two helical hairpin motifs of ATPTB1 and ATPEG3 extending into the membrane from opposite sides. This structural feature occludes the lipid-filled cavity in the membrane (Figure 6—figure supplement 1A,B). The architecture resembles that of an inverted-topology structural repeat found in excitatory amino acid transporters (Figure 6—figure supplement 1C,D), where similar helical hairpins play the key role in the transport mechanism and ion binding (Crisman et al., 2009). Within the membrane, the distances between the hairpin loops of a glutamate transporter homolog (Yernool et al., 2004) and ATPTB1/ATPEG3 are remarkably similar, within a 3.0–4.3 Å range. However, unlike in the intra-protein helical hairpin repeats, the membrane-spanning ATPTB1-ATPEG3 motif is formed by two separate subunits, and involves hydrophobic interactions (Y100_TB1, W55_EG3), which would restrict conformational variation.

Peripheral stalk

The peripheral stalk connects the static membrane-embedded part to the catalytic subunits in the F₁, which in Euglenozoa is elaborated by three copies of subunit p18, each bound to subunit α (Gahura et al., 2018; Montgomery et al., 2018). The peripheral stalk in yeasts and mammals is built around a ~ 150 Å-long helix of subunit b, extending from the membrane to the functionally important and conserved oligomycin-sensitivity conferring protein (OSCP) on top of the F₁ crown (Rees et al., 2009). By contrast, subunit b is significantly shorter in E. gracilis, extending only ~65 Å into the matrix, and does not interact with OSCP. This is compensated by extensions of OSCP and subunit d mediating the attachment of the peripheral stalk, which is augmented by two species-specific soluble subunits ATPTB3 and ATPTB4 (Figure 6C,D; Figure 6—figure supplement 3). The additional elements significantly increase the size of the peripheral stalk compared to yeast. The C-terminal OSCP extension contains a 13-residue proline-rich region, followed by 26 residues arranged in two helices (D240-L246, V250-A265), which contact the large matrix extension of subunit d. This interaction is further clamped by ATPTB3 and ATPTB4 of the peripheral stalk tip and p18 on F₁ (Figure 6D). Thus, the E. gracilis peripheral stalk displays a different mode of interaction with the OSCP compared to all other reported ATP synthase structures.

The OSCP N-terminal domain contains a structurally conserved bundle of four α-helices, followed by the N-terminal extension with a terminal helix. The N-terminal helix of OSCP connects to the central stalk through an interaction with the C-terminal extension of subunit γ of the central stalk (Figure 6E). We observe this contact in all three rotational states, which provides the first observation of an interaction between the peripheral and central stalk in the ATP synthase (Figure 7—figure supplement 1A to C). This raises the question how central stalk rotation proceeds despite the observed anchoring. Studies in Escherichia coli F₁-ATPase showed that artificial crosslinking of rotor subunit γ with stator subunit α near the crown domain does not impair the hydrolysis activity nor the full rotation of subunit γ (Gumbiowski et al., 2001; Hilbers et al., 2013). Therefore, the interaction observed between the E. gracilis OSCP and subunit γ is consistent with rotational catalysis. However, a permanent interaction would require rotation around a pivot point, presumably the short, extended linker region (K296-G298) in subunit γ, which is less well resolved in the cryo-EM maps. The flexibility of the central stalk has previously been suggested to play an important role in the energy transmission in the ATP synthase (Guo et al., 2019; Murphy et al., 2019; Wächter et al., 2011). Thus, the anchoring of subunit γ may affect the energy required for transition between the individual rotational substeps. The extensions of OSCP and subunit γ are conserved in Kinetoplastida (Figure 6—figure supplement 3F to H), suggesting that the observed interaction is a structural feature of euglenozoan ATP synthases. Together, the matrix-exposed subunit γ/OSCP interactions and lumenal ATPEG1/c-ring contacts (Figure 1D) pin the F₁/c-ring-complex to the stator, likely providing additional stability.

Unusual binding mode of IF₁

IF₁ is a natural inhibitor protein of the F₁-subcomplex (αβ)₃γδε that interacts with subunits α, β and γ upon pH increase (Cabezon et al., 2000; Cabezón et al., 2001). Conventionally, it binds to F₁ via the C-terminal helix that protrudes into the β_DP-α_DP interface, followed by a short N-terminal helix, which extends into the central F₁ cavity and interacts with the α-helical coiled coil of subunit γ of the central stalk to stall its rotation (Figure 7B) (Cabezón et al., 2003; Gledhill et al., 2007). Mechanistically, the central stalk rotation of the mammalian mitochondrial F₁-ATPase is specifically inhibited by the N-terminal of IF₁ that enters through the open catalytic β_E-α_E interface, and the subsequent 120°-rotation of the central stalk is coupled to a deeper insertion of the α-helix towards subunit γ (Bason et al., 2014). Importantly, the minimal inhibitory region of IF₁ (A14-K47, mammalian numbering) has been shown to include the N-terminal helix (van Raaij et al., 1996).

Figure 7. Unusual binding mode of IF₁.

(A and B) Comparison of IF₁ binding mode between E. gracilis (rotational state-1) and Bos taurus (PDB ID: 2V7Q) (Gledhill et al., 2007). In the E. gracilis structure, both termini are located outside the F₁. (C) Superposition of the IF₁-inhibited E. gracilis F₁ and the bovine counterpart. Both structures share the conserved C-terminal helix but differ in the structure of the N-termini. (D) Close-up of the IF₁ binding site. The C-terminal helix of the E. gracilis IF₁ contains the conserved EERY, followed by the helix-breaking P46. In the Euglenozoa-specific N-terminal region W37 and E39 interact with α_DP, whereas D34 interacts with β_TP. (E) Multiple sequence alignment of IF₁ from different species; Bos tarus (Bt), Homo sapiens (Hs), Caenorhabditis elegans (Ce), Saccharomyces cerevisiae (Sc), the trypanosomatids Trypanosoma brucei (Tb), Leishmania major (Lm), Endotrypanum monterogeii (Em), Crithidia fasciculata (Cf), Leptomonas pyrrhocoris (Lp), Blechomonas ayalai (Ba), Phytomonas sp. (Ps) and the euglenoid Euglena gracilis (Eg). Euglenozoan species names shown in red. Residues are shaded in blue according to conservation. Residues conserved within Euglenozoa are highlighted red. (F,G) Bridging density in ATP synthase dimer. (F) E. gracilis ATP synthase dimer with both monomers in rotational state-1; map shown at low threshold. (G) From the C-terminal helix of IF₁ a continuous density (cyan) extends towards the C₂-symmetry axis, contacting the all-β fold formed by subunits d and b, thereby bridging the two F₁ subcomplexes.

Figure 7—figure supplement 1. Three main rotational states with bound nucleotides and IF₁.

(A-C) F₁/c-ring/OSCP subcomplexes in the rotational states 1 (A), 2 (B), and 3 (C), related by a ~ 120°-rotation of the rotor. (G-F) Nucleotide binding sites of the β_DP containing ADP (G), the β_TP containing ATP (H) and the α_TP containing ATP and Triton-X100 (green, I). In both the β_DP and β_TP, the arginine finger (αR388) extends towards the nucleotide, the former being in contrast to the IF₁-inhibited bovine F₁, in which the arginine finger points away from the nucleotide (Gledhill et al., 2007).

The E. gracilis F₁/cring maps of rotational states 1 and 2 showed strong density for IF₁, but weak density in rotational state 3. Subsequent focused classification of the IF₁ binding site suggested complete inhibitor binding for rotational states 1 and 2, whereas rotational state 3 displayed a mixed occupancy, with a minority (34%) of particles containing IF₁ and a majority (66%) not containing the inhibitor (Figure 1—figure supplement 1D). Unlike the bovine ATP synthase, which displays a nearly equal particle distribution among the three rotational states (Zhou et al., 2015), rotational state-3 (20%) is less populated than rotational states 1 (44%) and 2 (36%). Rather than reflecting dwell times during rotary catalysis, the distribution of rotational states is likely dictated by accessibility of the respective catalytic interfaces to IF₁ binding, which for rotational state-3 may be sterically hindered by the proximity to the peripheral stalk.

In all three rotational states, IF₁ is bound to the β_DP-α_DP interface adopting a similar conformation (Figure 7—figure supplement 1A,B,D,E, Figure 1—figure supplement 1D). While E. gracilis IF₁ shares the conserved C-terminal helix at the β_DP-α_DP interface, including an EERY consensus motif, it does not contain the N-terminal helix. Instead, a Euglenozoa-specific proline residue (P46) follows the EERY motif and produces a break in the C-terminal helix of IF₁, which causes the N-terminal region to extend in the opposite direction compared to bovine IF₁. This results in interactions with the C-terminal domain of the α_DP-subunit via a number of residues that are conserved in Euglenozoa (Figure 7D,E). As a consequence, the E. gracilis IF₁, extends around the C-terminal domain of the α_DP-subunit and exits the F₁ through the β_TP-α_DP interface in close proximity of the γ-protrusion of the central stalk (Figure 7A). Consequently, the N-terminus of the E. gracilis IF₁ does not contact the central α-helical coiled coil of the central stalk, indicating a different binding mode compared to the mammalian inhibitor. Importantly, the residues that anchor the extended N-terminal region of the E. gracilis IF₁ to the α_DP-subunit are conserved in Trypanosomatids, indicating that the newly identified binding mode for the inhibition of the mitochondrial ATP synthase is synthase is structurally conserved in the related group of parasites (Figure 7—figure supplement 1E).

The C-terminal region of both the bovine and T. brucei IF₁ has been characterised as a homo-oligomerisation domain, mediating the assembly of IF₁ into a dimer, which is the inhibitory form in both organisms (Cabezón et al., 2001; Gahura et al., 2018). Recently, mammalian IF₁ was shown to bridge neighboring mitochondrial ATP synthase dimers into a tetramer (Gu et al., 2019). Our unsharpened map of the ATP synthase dimer with both monomers in rotational state-1 (IF₁pointing towards the C₂-axis), shows a continuous density from the C-terminus of IF₁. This density extends towards the dimer interface and contacts the all-β-fold formed by subunits b and d (Figure 7—figure supplement 1F,G), suggesting that also in Euglenozoa both F₁ complexes in the ATP synthase dimer may be inhibited by a dimeric IF₁. However, in contrast to the mammalian ATP synthase, the specific binding mode of the Euglenozoan IF₁ assembly links two monomers within the dimer, rather than between neighbouring dimers.

Conclusions

This work describes the composition and structure of a divergent mitochondrial ATP synthase of Euglenozoa with native lipids. The distinct subunit a, which had not been previously identified by genomic sequencing, is assigned directly from the density map, and markedly deviates from the structures in all the previously characterized ATP synthases. The newly found elements are involved in proton transfer, including functional substitution of the otherwise mitochondrially conserved glutamate of H6a with a histidine residue of H5a. The proton release channel is lined by species-specific subunits that together with four horizontal helices contribute to an almost double-sized contact area between the rotor and stator in the membrane. In addition, a direct interaction between the peripheral and central stalks is reported for the first time in the ATP synthase. Taken together, it implies a different energetic landscape for transition between the rotational substeps. This exemplifies the extreme structural divergence of the mechanistic components of ATP synthase.

The signature feature of mitochondrial ATP synthase dimers is the induction of membrane curvature that leads to cristae formation. In E. gracilis it is induced at the dimer interface, which is formed by species- or phylum-specific subunits and extensions of conserved subunits. Additionally, a membrane subcomplex is peripherally associated with conserved subunits, thereby contributing to a local displacement of the membrane towards the lumen. This suggests that interactions shaping the inner mitochondrial membrane are likely to have evolved independently. At the dimer interface, bound cardiolipins are specifically coordinated by different subunits, and the cavity formed by the newly identified subcomplex is predominantly occupied by cardiolipins. Therefore, protein-lipid interactions also play an important role in membrane bending. These observations extend our understanding of the structural mechanisms leading to a membrane curvature.

Last, the newly described binding mode of the inhibitor IF₁ to the ATP synthase is different from human, but appears to be conserved in Trypanosomatids, providing a potential therapeutic target.

Materials and methods

Cell culture and mitochondria isolation

Euglena gracilis was obtained from Lebendkulturen Helbig and grown in Hutner’s modified media (Buetow and Padilla, 1963) in 2 l Erlenmeyer flasks (14 × 700 ml culture) at 25°C and 120 rpm in the dark. Cells were harvested in late-logarithmic phase by centrifugation at 1200 xg, 10 min, 4°C. The pelleted cells were transferred into a cold mortar and lysed by grinding them with 500-µm glass beads in buffer A (50 mM HEPES pH 7.5, 210 mM mannitol, 70 mM sucrose, 1 mM DTT, 1 mM EGTA, 5 mM EDTA) for 5 min. The lysate was centrifuged at 1500 xg, 10 min, 4°C and the supernatant was spun again at 20,000 xg, 20 min, 4°C. The resulting crude mitochondrial pellet was further purified on a discontinuous sucrose gradient in buffer B (20 mM HEPES-KOH pH 7.5, 2 mM EDTA, 15/23/32/60% sucrose) by centrifugation (103,745 xg, 1 hr, 4°C) in an SW28 rotor (Beckman Coulter) and enriched mitochondria were collected from the 32–60% (w/v) interface.

Isolation of E. gracilis ATP synthase dimers

Approximately 10 mg mitochondria were lysed in a total volume of 90 ml buffer C (25 mM HEPES/KOH pH 7.5, 25 mM KCl, 15 mM MgOAc₂, 1.7% Triton-X100, 2 mM DTT, one tablet EDTA-free Protease Inhibitor Cocktail) for 2 hr at 4°C and the lysate was cleared by centrifugation at 30,000 xg, 20 min, 4°C. The supernatant was layered on a sucrose cushion in buffer D (1 M sucrose, 25 mM HEPES/KOH pH 7.5, 25 mM KCl, 15 mM MgOAc₂, 1% Triton-X100, 2 mM DTT) and centrifuged 158,420 xg, 3 hr, 4°C in a Ti45 rotor (Beckman Coulter). The resulting pellet was resuspended in 200 µl buffer E (25 mM HEPES/KOH pH 7.5, 25 mM KCl, 15 mM MgOAc₂, 2 mM DTT, 0.05% β-DDM) and gel filtrated over a Superose 6 Increase 3.2/300 column (GE Healthcare) in buffer E. Fractions corresponding to ATP synthase dimers were pooled and concentrated to 25 µl in a vivaspin500 filter (100 kDa MWCO).

Electron cryo-microscopy and data processing

3 µl sample (~5 mg/ml) were applied to glow-discharged Quantifoil R1.2/1.3 Cu grids and vitrified by plunge-freezing into liquid ethane after blotting for 3 s. Samples were imaged on a Titan Krios operated at 300 kV at a magnification of 130 kx (1.05 Å/pixel) with a Quantum K2 camera (slit width 20 eV) at an exposure rate of 4 electrons/pixel/s with a 10-s exposure and 25 frames using the EPU software. A total of 9045 collected movies were motion-corrected and exposure-weighted using MotionCor2 (Zheng et al., 2017) and contrast transfer function (CTF) estimation was performed using Gctf (Zhang, 2016). All subsequent image processing was performed in RELION-3 (Zivanov et al., 2018). Bad images were removed manually by inspection in real and Fourier space. Initial rounds of Gaussian-based particle picking, classification and refinement were used to generate picking references from the data. Reference-based particle picking was performed using Gautomatch (developed by Dr Kai Zhang, MRC Laboratory of Molecular Biology, Cambridge, UK, http://www.mrc-lmb.cam.ac.uk/kzhang/Gautomatch) to pick 555,269 particles, which were subjected to reference-free two-dimensional (2D) classification, resulting in 540,669 particles for three-dimensional (3D) classification, from which 2 classes containing 171,033 particles were selected for a consensus 3D-refinement, applying C₂-symmetry. The resulting pre-aligned particles were C₂-symmetry expanded and a mask including one asymmetric unit was used for signal subtraction on the particles. Subsequent local-angular-search 3D-classification into three classes yielded three maps corresponding to the three main rotational states (Figure 1—figure supplement 1D). The class corresponding to rotational state-3 could be improved by sub-classification. Masked refinements of the F₁/c-ring regions yielded three final maps of rotational state-1 (150,744 particles), state-2 (122,085 particles) and state-3 (43,232 particles) at 3.04 Å, 3.14 Å and 3.92 Å resolution, respectively (Figure 1—figure supplement 2C to E). Rotational states 1 and 2 show strong density for IF₁, which is weak in rotational state-3. Focussed classification of the IF₁ binding site for rotational states 1 and 2 did not yield a class lacking IF₁, indicating complete occupancy. By contrast, rotational state-3 displayed a mixed occupancy, with a minority (34%) of particles containing IF₁ (refined to 4.2 Å resolution) and a majority (66%) not containing the inhibitor (refined to 4.1 Å resolution; Figure 1—figure supplement 1D). Masked refinements of the rotor (central stalk and c-ring) and peripheral stalk tip region in rotational state-1 yielded maps at 3.3 Å and 3.82 Å, respectively (Figure 1—figure supplement 2F,G). For the refinement of these maps, signal-subtracted particles were used for refinement, whereas original particles were used for final reconstruction. Masked refinement of combined particles from all three rotational states, yielded a 2.82 Å map of the static membrane region (Figure 1—figure supplement 2A). Finally, a consensus 4.32 Å resolution dimer map was generated by combining particles in which both F₁/c-ring subcomplexes are in rotational state-1 (Figure 1—figure supplement 2B). All final maps were generated from CTF-refined particles. All resolution estimates are according to Fourier shell correlations (FSC) that were calculated from independently refined half-maps using the 0.143-criterion with correction for the effect of the applied masks (Figure 1—figure supplement 1C).

Atomic model building and refinement

Atomic model building was performed in Coot (Emsley and Cowtan, 2004). F_o subunits were built de novo and identified from the density map as previously reported E. gracilis ATP synthase subunits (Perez et al., 2014; Yadav et al., 2017) or newly identified subunits using the E. gracilis transcriptome or genome (NCBI accession code: PRJEB27422). OSCP/F₁/c-ring models were built using a homology model (Waterhouse et al., 2018) of the yeast F₁/c₁₀-ring (PDB ID: 3ZRY) (Giraud et al., 2012), whereas OSCP, IF₁and p18 were built de novo. Real-space refinement of atomic models was performed in PHENIX (Afonine et al., 2018) using secondary structure restraints. Cardiolipins resolved in the cryo-EM map were identified by their unique structure containing two phosphatidyl groups linked by a central glycerol bridge (Video 2). Bound lipids other than cardiolipin could not be unambiguously identified from the head group of their phosphatidyl density and were thus modelled as phosphatidic acid. Acyl tails of lipids were truncated according to map density. The atomic model of the rotor in rotational state-1 was rigid-body fitted into the local-resolution-filtered F₁/c-ring maps of all three rotational states and the combined F₁/c-ring models were subsequently refined with reference restraints on the c-ring. To generate a composite model of the complete ATP synthase dimer, the atomic models of the membrane region, the OSCP/F₁/c-ring in rotational-state one and the peripheral stalk tip were combined after rigid-body fitting into the consensus map of the dimer with both rotors in rotational state-1 and refined in PHENIX using reference restraints. Model statistics were calculated using MolProbity (Chen et al., 2010). To evaluate potential overfitting of the atomic model during refinement, the atomic coordinates of the refined model were randomly displaced by shifts of up to 0.5 Å using ‘Shake’ in the CCPEM suite (Burnley et al., 2017). The shaken model was Real-space refined using PHENIX against one half map that had been reference-sharpened using Refmac (Brown et al., 2015). Subsequently, FSC_work and FSC_test between the model and the two unfiltered half-maps, were calculated as described (Brown et al., 2015) (Figure 1—figure supplement 2).

Data analysis and visualisation

The lumenal half-channel was traced as a void in the F_o-model, using HOLLOW (Ho and Gruswitz, 2008). The matrix half channel was visualised as a void (inverted contrast) in the density map of the membrane region. Images were rendered using PyMOL 2 (Schrödinger, LLC), Chimera (Pettersen et al., 2004) or ChimeraX (Goddard et al., 2018). Sequence alignments were performed using MAFFT (Katoh et al., 2002). Prediction of cleavage sites of the mitochondrial matrix protease was performed using MitoFates (Fukasawa et al., 2015). To assess the presence of the newly identified subunit a and subunit 8 in the E. gracilis mitochondrial genome, a BLAST search of the DNA sequence (experiment PRJNA294935; Dobáková et al., 2015) was performed, which showed that the query is entirely covered by individual sequencing reads. The respective sequences were subsequently used to assemble a consensus contig using CAP3 (Huang and Madan, 1999) (Figure 2—figure supplement 1E).

Molecular dynamics simulations

The consensus ATP synthase dimer model was placed in a planar bilayer (45% palmitoyl-oleoyl-phosphatidylcholine (POPC), 30% palmitoyl-oleoyl-phosphatidylethanolamine (POPE), 20% cardiolipin (CDL), 5% palmitoyl-oleoyl-phosphatidic acid (POPA); corresponding to 3609, 3011, 1604, and 400 molecules, respectively) using insane (Wassenaar et al., 2015), surrounded by an aqueous 150 mM NaCl solution. Three replicas with identical lipid types and quantities were generated, differing only in the initial lipid placement. The protein model was converted to a coarse-grained MARTINI representation using the martinize script (de Jong et al., 2013). Parameters for the lipids POPC, POPE, POPA, and CDL were used from the MARTINI force field for lipids, and parameters for the coarse-grained protein, water and ions were used from the MARTINI 2.0 forcefield (Marrink et al., 2007; Monticelli et al., 2008). The initial system box size was 50 × 50×35 nm with a total number of ~775,000 beads. Equilibration of the system was carried out by an energy minimization using the steep integrator for 10 ps total, followed by seven steps during which positional restraints on the protein and lipid beads were gradually lowered, while increasing the time step of the simulation, until a total of 91 ns equilibration time. The Berendsen semi-isotropic pressure coupling was used during equilibration (Berendsen et al., 1984). During production, three replicas of 4 μs each were simulated in 20-fs time steps using periodic boundary conditions. The protein bead scaffold was restrained utilizing the EleNeDyn forcefield (Periole et al., 2009) with a force constant of 500 kJ/mol/nm², and all beads were held with a positional restraint constant of 10 kJ/mol/nm². The temperature of the system was maintained at 298 K using the velocity-rescaling thermostat (Bussi et al., 2007) with temperature coupling separately for protein, lipids, and solvent. The pressure was maintained at 1 bar, using the Parrinello-Rahman barostat and semi-isotropic coupling (Parrinello and Rahman, 1981). Electrostatics were calculated with a dielectric constant of 15.0 and a cutoff of 1.1 nm was applied for short-range interactions. Molecular dynamics simulations were performed with GROMACS 2016.1 (Abraham et al., 2015).

Simulation data was visualised using VMD or the python package matplotlib (Humphrey et al., 1996; Hunter, 2007). The script cg_bonds (available on cgmartini.nl) was used to visualize the coarse-grained beads as whole molecules. The radial distribution function, as well as the distances between the F_o peripheral cavity and the lipid molecules were determined using GROMACS tools rdf and pairdist (Abraham et al., 2015), respectively. Residence time in the pocket was smoothed to avoid intermittent exit of lipids from the F_o peripheral cavity. A lipid was considered to begin a binding event if any of its beads reached within a 1.8 nm distance from the center of mass of the cavity, and was considered to be bound until leaving a buffer zone of 3.5 nm distance from the cavity center of mass. Error bars in Figure 6—figure supplement 1E represent a confidence interval of 90%, obtained after bootstrapping of the original distribution. The nonparametric Mann-Whitney U test was used to determine if the residence times of each lipid type originated from the same distribution as cardiolipin residence times (Mann and Whitney, 1947).

Over different time spans (τ), the probability of different lipid types (j) to enter, remain in, or exit the binding site was calculated. For each lipid type, the number of times (N_j) any bead of each lipid molecule of this type entered the binding site (N_j(o_k → i_k+τ)), or remained in the binding site (N_j(i_k → i_k+τ)) where k is the frame index in the simulation, was calculated. This quantity was normalised over the total number of possible events, including the other two possible binding events, a lipid molecule exiting the binding site (N_j(i_k → o_k+τ)), and a lipid molecule remaining outside the binding site (N_j(o_k → o_k+τ)), thus yielding a probability. Time spans τ of 1 ns to 3000 ns were considered. For example, the probability of entering the pocket (N_j(o_k → i_k+τ)) is expressed as:

$$\rho (o\to i{)}_{\tau ,j}=\frac{{\sum }_{j=1}^{Nlipidtypes}{\sum }_{k=1}^{Nframes}\left(N_j\right(o_k\to i_{k+\tau }\left)\right)}{{\sum }_{j=1}^{Nlipidtypes}{\sum }_{k=1}^{Nframes}\left(N_j\right(o_k\to i_{k+\tau })+(N_j(i_k\to i_{k+\tau })+\left(N_j\right(i_k\to o_{k+\tau })+(N_j(o_k\to o_{k+\tau }))}$$

Funding Statement

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Funding Information

This paper was supported by the following grants:

• Stiftelsen för Strategisk Forskning FFL15:0325 to Alexey Amunts.

• Ragnar Söderbergs stiftelse M44/16 to Alexey Amunts.

• Vetenskapsrådet NT_2015-04107 to Alexey Amunts.

• Cancerfonden 2017/1041 to Alexey Amunts.

• H2020 European Research Council ERC-2018-StG-805230 to Alexey Amunts.

• Knut och Alice Wallenbergs Stiftelse 2018.0080 to Alexey Amunts.

• European Molecular Biology Organization ALTF 260-2017 to Alexander Mühleip.

• European Molecular Biology Organization Young Investigator Program to Alexey Amunts.

Data availability

All data generated or analyzed during this study are included in this Article and the Supplementary Information. The cryo-EM maps have been deposited in the Electron Microscopy Data Bank with accession codes EMD-10467, EMD-10468, EMD-10469, EMD-10470, EMD 10471, EMD-10472, EMD-10473, EMD-10474, EMD-10475. The atomic models have been deposited in the Protein Data Bank under accession codes 6TDU, 6TDV, 6TDW, 6TDX, 6TDY, 6TDZ, 6TE0.

The following datasets were generated:

Mühleip A, McComas SE, Amunts A. 2019. Atomic Models. Protein Data Bank. 6TDU

Mühleip A, McComas SE, Amunts A. 2019. Atomic Models. Protein Data Bank. 6TDV

Mühleip A, McComas SE, Amunts A. 2019. Atomic Models. Protein Data Bank. 6TDW

Mühleip A, McComas SE, Amunts A. 2019. Atomic Models. Protein Data Bank. 6TDX

Mühleip A, McComas SE, Amunts A. 2019. Atomic Models. Protein Data Bank. 6TDY

Mühleip A, McComas SE, Amunts A. 2019. Atomic Models. Protein Data Bank. 6TDZ

Mühleip A, McComas SE, Amunts A. 2019. Atomic Models. Protein Data Bank. 6TE0

Mühleip A, McComas SE, Amunts A. 2019. Cryo-EM maps. Electron Microscopy Data Bank. EMD-10467

Mühleip A, McComas SE, Amunts A. 2019. Cryo-EM maps. Electron Microscopy Data Bank. EMD-10468

Mühleip A, McComas SE, Amunts A. 2019. Cryo-EM maps. Electron Microscopy Data Bank. EMD-10469

Mühleip A, McComas SE, Amunts A. 2019. Cryo-EM maps. Electron Microscopy Data Bank. EMD-10470

Mühleip A, McComas SE, Amunts A. 2019. Cryo-EM maps. Electron Microscopy Data Bank. EMD 10471

Mühleip A, McComas SE, Amunts A. 2019. Cryo-EM maps. Electron Microscopy Data Bank. EMD-10472

Mühleip A, McComas SE, Amunts A. 2019. Cryo-EM maps. Electron Microscopy Data Bank. EMD-10473

Mühleip A, McComas SE, Amunts A. 2019. Cryo-EM maps. Electron Microscopy Data Bank. EMD-10474

Mühleip A, McComas SE, Amunts A. 2019. Cryo-EM maps. Electron Microscopy Data Bank. EMD-10475