The majority of metabolic energy in cells is generated by F₁F_o ATP synthase, a biological rotary motor that converts the proton motive force (pmf) to adenosine tri-phosphate (ATP) in both oxidative phosphorylation and photophosphorylation (Stewart et al., 2014; Walker, 2013). The enzyme consists of two reversible rotary motors, termed F₁ and F_o, coupled together by one central and one peripheral stalk, with the simplest subunit composition found in bacteria such as Escherichia coli (Figure 1—figure supplement 1). The F_o motor spans the membrane and converts the potential energy of the pmf into rotation of the central stalk that in turn drives conformational changes in the three catalytic sites of the α₃β₃ F₁ motor subunits to generate ATP. Moreover, this process is reversible so that, if the pmf drops below the threshold needed to power ATP synthesis, the motor has the ability to reverse and, in some bacterial species, operates primarily as a proton pump driven by ATP hydrolysis. Regulation of these ATPase/synthase activities is particularly important in times of cellular stress, primarily to prevent wasteful ATP consumption. However, different regulatory mechanisms are used by different F₁F_o subtypes (bacterial, chloroplastic, mitochondrial) and species (Sielaff et al., 2018; Morales-Rios et al., 2015; Stewart and Stock, 2012; Gledhill et al., 2007; Hahn et al., 2018; Stewart, 2014; Pullman and Monroy, 1963).

It has been postulated that bacterial ATP synthases are regulated by several mechanisms, with conformational changes of the ε subunit and catalytic nucleotide occupancies likely playing major roles (Sielaff et al., 2018; Hyndman et al., 1994; Feniouk et al., 2006). A long-standing question surrounds the role of the C-terminal domain of the ε subunit (εCTD). The εCTD is known to change conformation and block rotation of the central stalk in the F₁ motor from several bacteria, but this role is absent in the enzyme of mitochondria (Sielaff et al., 2018; Laget and Smith, 1979; Cingolani and Duncan, 2011; Sobti et al., 2016; Yagi et al., 2007; Shirakihara et al., 2015). In the active bacterial enzyme, the εCTD is proposed to adopt a condensed or ‘down’ conformation (Krah et al., 2017), and this state has been observed for isolated ε subunit from E. coli (Uhlin et al., 1997; Wilkens and Capaldi, 1998a) and Geobacillus stearothermophilus (or Bacillus PS3, hereafter termed PS3) (Yagi et al., 2007) as well as in isolated F₁ from Caldalkalibacillus thermarum (Ferguson et al., 2016). In autoinhibited states observed in crystal structures of F₁ from E. coli and PS3, the εCTDs are extended in similar ‘up’ conformations, so that a helix inserts into the F₁ central cavity and appears to block rotation of the complex by binding to both the central rotor subunit γ and several surrounding α and β subunits (Sielaff et al., 2018; Cingolani and Duncan, 2011; Mendoza-Hoffmann et al., 2018) (Figure 1—figure supplement 2). However, there is a key structural difference between these ‘up’ states: in E. coli, the εCTD consists of two helices (here termed εCTH1 and εCTH2; see Figure 1—figure supplement 2) separated by an extended loop, with each helix interacting with a different region of the γ subunit (Cingolani and Duncan, 2011; Rodgers and Wilce, 2000); whereas in PS3, εCTH1 and εCTH2 instead join to form one continuous helix (Shirakihara et al., 2015). In PS3 and a related Bacillus species, ATP is known to bind to the ε subunit and stabilise the ‘down’ conformation (Yagi et al., 2007; Krah et al., 2017; Kato-Yamada and Yoshida, 2003; Kato-Yamada, 2005; Imamura et al., 2009), and a clear mechanism of regulation by the εCTD and ATP levels has been proposed (Shirakihara et al., 2015). However, the physiological importance of the εCTD in ATP synthases of other bacteria, such as E. coli, has remained uncertain. Crosslinking studies have suggested that the εCTD of E. coli ATP synthase could act as a ratchet, preventing hydrolysis - but not synthesis - of ATP (Tsunoda et al., 2001). However, deletion studies performed in vivo, in which the E. coli εCTD was removed, showed minimal impact on cell growth or viability (Shah and Duncan, 2015), although deletion of just the five terminal residues results in decreased respiratory growth due to increased inhibition of ATP synthesis (Taniguchi et al., 2011). For the highly latent enzyme from C. thermarum, an early study indicated the εCTD is involved in inhibition (Keis et al., 2006), but thus far crystallographic studies of its F₁ complex have shown the εCTD in the down conformation irrespective of ATP binding to ε (Ferguson et al., 2016). Despite confusion over the possible importance of the εCTD for the bacteria noted above, one recent study demonstrated that the εCTD can impact the virulence of Streptococcus pneumoniae. F₁F_o is essential for viability of S. pneumoniae and functions in pH homeostasis and in its acid tolerance response (Ferrándiz and de la Campa, 2002; Cortes et al., 2015). In a mouse model of pneumococcal bacteraemia, a frameshift mutation that scrambled the sequence of the εCTD was found to increase pneumococcal survival in the spleen, most likely by reducing killing by splenic macrophages (Gerlini et al., 2014). Therefore, it is important to characterize more fully the bacterial εCTD and its different conformational states and interactions within bacterial ATP synthases.

In contrast to the crystallographic reports mentioned above with isolated ε subunits or soluble F₁, our previous cryo-EM study (Sobti et al., 2016) provided maps of the intact E. coli ATP synthase. These data were obtained using purified protein without the addition of exogenous nucleotide, thus likely representing a state autoinhibited by ε, here termed ATP synthase^AI. Here we have used similar methodologies to investigate the effect of ATP on the conformation of E. coli F₁F_o ATP synthase, by cryo-freezing and subsequently imaging the enzyme after a brief incubation with 10 mM MgATP. The maps obtained (termed ATP synthase^+ATP hereafter) indicate that, in the presence of MgATP, the εCTD became detached from the body of the enzyme, coupled with a large change in one of the three catalytic β subunits. Further analysis identified an intermediate in which the εCTD was in a ‘half-up’ conformation, suggesting that the enzyme can still rotate, even when the ε subunit is not in its condensed down position, as well as an εCTD ‘down’ conformation likely showing an active intermediate. Further inspection of our maps demonstrated peaks present in all six nucleotide-binding sites in the ATP synthase^+ATP enzyme (three catalytic β sites and three noncatalytic α sites), in contrast to the autoinhibited protein in which only four binding pockets (a single catalytic β site and three noncatalytic α sites) were occupied with nucleotide. Together with our previous work, these results reveal three distinct positions taken up by the εCTD in response to different catalytic nucleotide–bound states, providing further insight into the mechanism of ATP synthase regulation in E. coli and related species.

The structural information obtained here using E. coli F₁F_o ATP synthase incubated with MgATP likely reflects conformations uninhibited by the εCTD. Although the maps generated are of limited resolution, the position of the catalytic subunits and εCTD were clearly identifiable. Two different εCTD conformations were observed: a ‘down’ conformation, in which εCTH1 and εCTH2 are arranged next to one another and interacts with the N-terminal region of the ε subunit, and a ‘half-up’ conformation in which the εCTH1 lies against the γ subunit (Figure 2). Because the ATP and ADP concentrations present were comparable to those observed in intact cells (Bennett et al., 2009) it is likely that both these conformations are present in E. coli. Furthermore, analysis of the maps in light of the recent high-resolution structure of related S. oleracea enzyme has highlighted a unique C-terminal fold of the peripheral stalk b subunits in E. coli ATP synthase that derives from the homodimeric nature of this subunit in this system.

A considerable body of work has described the inhibition of F₁-ATPases by ADP, Mg²⁺ and εCTD (Sielaff et al., 2018; Hyndman et al., 1994; Laget and Smith, 1979; Cingolani and Duncan, 2011; Sobti et al., 2016; Richter, 2004; Zhou et al., 1988; Avron, 1962; Drobinskaya et al., 1985; Guerrero et al., 1990). Because the sample in the present study was imaged under conditions in which ADP accumulated at a linear rate (Figure 1—figure supplement 3), we propose that the maps represent structures of E. coli F₁F_o ATP synthase in which the autoinhibitory state of the εCTD is prevented. These results indicate that ATP induces the εCTD to exit from the central cavity of F₁ and so facilitate rotation of the enzyme. The contribution of the εCTD to regulate E. coli F₁F_o ATP synthase activity has previously been unclear. Our structural studies on E. coli F₁F_o ATP synthase support a mechanism whereby the εCTD is primed to act as a ‘safety lock” (Feniouk and Junge, 2005) or ‘ratchet” (Tsunoda et al., 2001; Cipriano and Dunn, 2006) to prevent ATP synthase from entering the ADP/Mg²⁺ inhibited state (Shah et al., 2013) even when ADP and Mg²⁺ concentrations are high (Figure 4). The ‘half-up’ state that we identified in our cryo-EM maps has been previously hypothesised (Cingolani and Duncan, 2011) and studies on the enzyme in the absence of the εCTH2 support the hypothesis that the εCTD ‘half-up’ conformation can act as a weak inhibitor, partially inhibiting without insertion into the F₁ motor (Nakanishi-Matsui et al., 2014). It is unlikely that the ‘half-up’ conformation we observe here is an artefact generated by the cysteine residue free construct used in this study, because in our previous work (Sobti et al., 2016) the same cysteine-free enzyme did not show the ‘half-up’ state. In addition, previous studies using γC87S mutant protein did not show any change in ε inhibition (Duncan et al., 1995), and in the structure of the native enzyme (Figure 1—figure supplement 10) loops in the γ subunit separate the cysteines in subunit γ and residues in the εCTH1, so that they are not in direct contact.

Figure 4

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Structural data from this and previous studies suggest that loading multiple catalytic sites with nucleotides disrupts access of the εCTD to the inhibitory ‘up’ state in E. coli (Figure 4). In a nucleotide-free solution, the F₁-ATPase adopts an autoinhibited state (seen in the crystal structure of isolated E. coli F₁-ATPase (Cingolani and Duncan, 2011) and the ATP synthase^AI cryo-EM map (Sobti et al., 2016); Figure 4a and b). The isolated F₁-ATPase shows one β subunit (β_DP) in a ‘half-closed’ state, with ADP bound, which interacts directly with the ‘up’ conformation of the εCTH2 (Figure 4a). In the ATP synthase^AI cryo-EM map, the same β subunit is unoccupied and has adopted an open conformation that no longer interacts with the εCTH2, which could facilitate a reversible mechanism whereby the helix is able to escape from its inserted position. When compared with isolated F₁, the reduced interactions with the ‘up’ state of subunit ε seen in the complete F₁F_o complex may explain why ε inhibition is significantly reduced upon rebinding of E. coli F₁ to F_o. In the ATP synthase^+ATP cryo-EM maps, all catalytic sites are at least partially occupied with nucleotide (Figure 1—figure supplement 7) and the εCTD is seen in two conformations that no longer contact the α and β subunits (Figure 4c and d). Because we only observed the εCTD in the ‘up’ conformation in the absence of ATP, these results indicate that nucleotide binding to the catalytic subunits, in either one or a combination of catalytic sites, is important for the release of the εCTD. Binding of ATP is unlikely to accelerate release of the εCTD from the ‘up’ state, as that would conflict with the known noncompetitive nature of ε inhibition with E. coli F₁ and F_oF₁. Rather, as the εCTH2 escapes from the central cavity of the ATP synthase^AI conformation (Figure 4b), rapid binding of ATP would induce a transition from the open (Figure 4b, ‘O’) to the closed conformation (Figure 4c/d, ‘C’), preventing re-entry of εCTH2 until the next catalytic dwell position. Finally, the ‘half-up’ conformation seen for the εCTD supports the prediction that the εCTH2 can remain mobile, anchored to γ by the εCTH1, so that it can re-enter the central cavity at a later step in the cycle (Cingolani and Duncan, 2011). It is also possible that this εCTH1 interaction alone may be able to modulate enzyme activity, as seen in a previous deletion study (Nakanishi-Matsui et al., 2014).

The ‘down’ conformation of the ε subunit has been observed for the isolated ε subunit of PS3 with ATP bound (Yagi et al., 2007) and on F₁ from C. thermarum (Ferguson et al., 2016). For ATP synthase^+ATP cryo-EM maps containing the εCTD in the ‘down’ conformation, close inspection showed very weak density, consistent with a fraction of these complexes showing nucleotide bound to the ε subunit. Docking of the C. thermarum (Ferguson et al., 2016) ε subunit with ATP bound, showed good correlation between the position occupied by the nucleotide and additional density seen in the ATP synthase^+ATP map (Figure 1—figure supplement 11). Partial occupancy would be consistent with the concentration of ATP employed in our study being slightly below the K_d of ~22 mM observed for ATP binding to isolated E. coli ε subunit (Yagi et al., 2007), and so because weak density attributable to nucleotide bound to the εCTD was only seen with the down conformation, it is possible that the ‘half-up’ conformation, with the associated probable disorder of εCTH2, might result when nucleotide is not bound to the ε subunit. Further work, for example, using a range of ATP concentrations, will be needed to resolve this point.

Although the movements of the εCTD are well defined by this study, the resolution of the reconstructions is limited compared to others (Guo and Rubinstein, 2018; Murphy et al., 2019). We believe this is likely due to inherent flexibility in the complex, possible varying subunit or lipid stoichiometry (Chorev et al., 2018), the detergent used, ice thickness, particle density and the lower accelerator voltage used during image acquisition (200 vs. 300 kV). Moreover, we only observed three rotational states whereas seven were seen in bovine mitochondrial ATP synthase (Zhou et al., 2015) and thirteen in Polytomella sp. mitochondrial ATP synthase (Murphy et al., 2019). We did employ methods similar to those used in these studies in an attempt to observe rotary sub-steps in the E. coli enzyme, but were unsuccessful. Although the complex should be undergoing rotation, due to the ATP hydrolysis that is occurring during freezing, single molecule studies have shown that in order for the complex to spend significant time in rotary sub-steps it had to be imaged under high drag, such as in 30% polyethylene glycol with a large object attached (Ishmukhametov et al., 2010). Indeed, the poor density seen at the membrane interface of the subunit b dimer, points towards these reconstructions containing multiple conformations, which may contribute to limiting the resolution obtained. It is possible that further work to obtain a much larger dataset, containing many more particles, may enable identification of more of the rotary sub-steps that are likely present in this sample.

Recent work on Polytomella sp. mitochondrial ATP synthase (Murphy et al., 2019) has described this ATP synthase in great detail, showing a flexible coupling between the F₁ and F_o motors mediated by a hinge region in OSCP (analogous to the δ subunit in E. coli ATP synthase). Comparison of the peripheral stalks in this study, by superposition of either the F_o stator or the N-terminal region of subunit δ (Figure 3—figure supplement 2), shows that the peripheral stalk likely bends and twists as the enzyme undergoes rotary catalysis, as suggested in previous studies (Hahn et al., 2018; Stewart et al., 2012). We do not observe movement in the delta/OSCP hinge, most likely due to our inability to resolve rotary substates in our maps.

In previous studies on E. coli ATP synthase, the enzyme has been shown to become inactive four hours after purification, possibly due to aggregation (Ishmukhametov et al., 2010). In this study, the enzyme was purified rapidly (see Materials and methods) and EM grids were made shortly after purification to ensure active enzyme was being imaged. The images obtained showing free monodisperse protein with little evidence of aggregation. Furthermore, the enzyme was purified in digitonin rather than the mixture of phosphatidylcholine, octyl glucopyranoside, sodium deoxycholate and sodium cholate used previously (Ishmukhametov et al., 2010; Ishmukhametov et al., 2005). Digitonin was selected because enzyme purified in the original mixture of lipids and detergents was not stable on the size exclusion chromatography (SEC) employed as a final purification step prior to grid freezing. Because the enzyme had been purified in a different detergent and further purified using SEC, we assessed the activity of the digitonin solubilized enzyme using ATP regeneration assays after 0, 1, 2, 3, 4 and 8 hr to compare with data taken on enzyme purified in previous studies (Ishmukhametov et al., 2010) (Figure 1—figure supplement 12). Importantly the enzyme showed little change in activity over 8 hr, suggesting that E. coli ATP synthase purified with digitonin and SEC may be less prone to inactivation or aggregation.

The present study has exploited the ability to manipulate sample conditions immediately before freezing to obtain images of E. coli ATP synthase at a time when ADP formation from ATP proceeds linearly, and hence likely unhindered, to identify the possible conformational changes that occur to remove the εCTD from the autoinhibitory position. This outcome was achieved by optimizing experimental design, considering particle concentration, as well as mixing and blotting time. Interestingly, employing similar methods on the related bacterial A/V type ATPase from Thermus thermophilus, showed that no conformational changes occur in the catalytic or related F subunits upon addition of the ATP transition state mimic fluoroaluminate (Davies et al., 2017). This could be due to different nucleotide affinity, operating temperature or indeed function of the complexes (Nakano et al., 2008).

In summary, this study presents structural data of E. coli ATP synthase imaged in the presence of ~9.75 mM ATP and ~0.3 mM ADP, comparable to the concentrations present in the intact bacterium. Compared to our previous work on the autoinhibited enzyme in the absence of ATP, the cryo-EM reconstructions presented here show that a single β subunit and the C-terminal domain of subunit ε change conformation and position substantially upon addition of ATP. Together, our studies provide strong support for the hypothesis that the ε subunit acts as an emergency brake to minimize wasteful hydrolysis of cellular ATP in situations where the ATP-to-ADP ratio is low. As the εCTD has recently been shown to impact pathogenesis of at least one bacterial species (Gerlini et al., 2014), the different conformations of the εCTD observed in these studies may have potential to guide future development of drugs that could target this bacteria-specific inhibitory mechanism.

ATP was purchased in the form of adenosine 5-triphosphate disodium salt hydrate (Sigma A7699).

Cryo-EM maps have been deposited to the EMDB under accession numbers EMD-9345, EMD-9346, EMD-9348, EMD-20006, EMD-20007 and EMD-20008.

1. 1. Meghna Sobti
   2. Alastair G Stewart

   (2019) Electron Microscopy Data Bank

   ID EMD-20006. Autoinhibited E. coli ATP synthase state 1.

   http://www.ebi.ac.uk/pdbe/entry/emdb/EMD-20006
2. 1. Meghna Sobti
   2. Alastair G Stewart

   (2019) Electron Microscopy Data Bank

   ID EMD-20007. Autoinhibited E. coli ATP synthase state 2.

   http://www.ebi.ac.uk/pdbe/entry/emdb/EMD-20007
3. 1. Meghna Sobti
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   ID EMD-20008. Autoinhibited E. coli ATP synthase state 3.

   http://www.ebi.ac.uk/pdbe/entry/emdb/EMD-20008
4. 1. Sobti M
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   ID EMD-9348. E. coli ATP synthase after incubation with ATP - State C.

   http://www.ebi.ac.uk/pdbe/entry/emdb/EMD-9348
5. 1. Sobti M
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   ID EMD-9345. E. coli ATP synthase after incubation with ATP - State A.

   http://www.ebi.ac.uk/pdbe/entry/emdb/EMD-9345
6. 1. Sobti M
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   (2019) Electron Microscopy Data Bank

   ID EMD-9346. E. coli ATP synthase after incubation with ATP - State B.

   http://www.ebi.ac.uk/pdbe/entry/emdb/EMD-9346

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Author details

1. Meghna Sobti

   1. Molecular, Structural and Computational Biology Division, The Victor Chang Cardiac Research Institute, Darlinghurst, Australia
   2. St Vincent’s Clinical School, Faculty of Medicine, UNSW Sydney, Sydney, Australia

   Contribution

   Conceptualization, Formal analysis, Investigation, Methodology, Writing—original draft, Writing—review and editing

   Competing interests

   No competing interests declared

2. Robert Ishmukhametov

   Department of Physics, Clarendon Laboratory, University of Oxford, Oxford, United Kingdom

   Contribution

   Conceptualization, Methodology, Writing—original draft, Writing—review and editing

   Competing interests

   No competing interests declared

3. James C Bouwer

   Molecular Horizons, The University of Wollongong, Wollongong, Australia

   Contribution

   Investigation, Methodology, Writing—review and editing

   Competing interests

   No competing interests declared

4. Anita Ayer

   1. St Vincent’s Clinical School, Faculty of Medicine, UNSW Sydney, Sydney, Australia
   2. Vascular Biology Division, Victor Chang Cardiac Research Institute, Darlinghurst, Australia

   Contribution

   Formal analysis, Investigation, Methodology, Writing—review and editing

   Competing interests

   No competing interests declared

5. Cacang Suarna

   Vascular Biology Division, Victor Chang Cardiac Research Institute, Darlinghurst, Australia

   Contribution

   Formal analysis, Investigation, Methodology

   Competing interests

   No competing interests declared

6. Nicola J Smith

   1. St Vincent’s Clinical School, Faculty of Medicine, UNSW Sydney, Sydney, Australia
   2. Molecular Cardiology and Biophysics Division, Victor Chang Cardiac Research Institute, Darlinghurst, Australia

   Contribution

   Resources, Funding acquisition, Writing—review and editing

   Competing interests

   No competing interests declared

7. Mary Christie

   1. Molecular, Structural and Computational Biology Division, The Victor Chang Cardiac Research Institute, Darlinghurst, Australia
   2. St Vincent’s Clinical School, Faculty of Medicine, UNSW Sydney, Sydney, Australia

   Present address

   School of Life and Environmental Sciences, University of Sydney, Sydney, Australia

   Contribution

   Resources, Funding acquisition, Writing—original draft, Writing—review and editing

   Competing interests

   No competing interests declared

8. Roland Stocker

   1. St Vincent’s Clinical School, Faculty of Medicine, UNSW Sydney, Sydney, Australia
   2. Vascular Biology Division, Victor Chang Cardiac Research Institute, Darlinghurst, Australia

   Contribution

   Resources, Funding acquisition, Investigation, Methodology, Writing—review and editing

   Competing interests

   No competing interests declared

9. Thomas M Duncan

   Department of Biochemistry & Molecular Biology, SUNY Upstate Medical University, Syracuse, NY, United States

   Contribution

   Writing—original draft, Writing—review and editing

   Competing interests

   No competing interests declared

10. Alastair G Stewart

    1. Molecular, Structural and Computational Biology Division, The Victor Chang Cardiac Research Institute, Darlinghurst, Australia
    2. St Vincent’s Clinical School, Faculty of Medicine, UNSW Sydney, Sydney, Australia

    Contribution

    Conceptualization, Resources, Funding acquisition, Investigation, Methodology, Writing—original draft, Project administration, Writing—review and editing

    For correspondence

    a.stewart@victorchang.edu.au

    Competing interests

    No competing interests declared

    "This ORCID iD identifies the author of this article:" 0000-0002-2070-6030

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