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

Download asset Open asset

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
   2. Alastair G Stewart

   (2019) Electron Microscopy Data Bank

   ID EMD-20008. Autoinhibited E. coli ATP synthase state 3.

   http://www.ebi.ac.uk/pdbe/entry/emdb/EMD-20008
4. 1. Sobti M
   2. Stewart AG
   3. Sobti M
   4. Alastair G Stewart

   (2019) Electron Microscopy Data Bank

   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
   2. Stewart AG

   (2019) Electron Microscopy Data Bank

   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
   2. Stewart AG

   (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

1. 1. Avron M

   (1962)

   Light-dependent adenosine triphosphatase in chloroplasts

   The Journal of Biological Chemistry 237:2011–2017.

   • PubMed
   • Google Scholar
2. 1. Bennett BD
   2. Kimball EH
   3. Gao M
   4. Osterhout R
   5. Van Dien SJ
   6. Rabinowitz JD

   (2009) Absolute metabolite concentrations and implied enzyme active site occupancy in Escherichia coli

   Nature Chemical Biology 5:593–599.

   https://doi.org/10.1038/nchembio.186

   • PubMed
   • Google Scholar
3. 1. Chorev DS
   2. Baker LA
   3. Wu D
   4. Beilsten-Edmands V
   5. Rouse SL
   6. Zeev-Ben-Mordehai T
   7. Jiko C
   8. Samsudin F
   9. Gerle C
   10. Khalid S
   11. Stewart AG
   12. Matthews SJ
   13. Grünewald K
   14. Robinson CV

   (2018) Protein assemblies ejected directly from native membranes yield complexes for mass spectrometry

   Science 362:829–834.

   https://doi.org/10.1126/science.aau0976

   • PubMed
   • Google Scholar
4. 1. Cingolani G
   2. Duncan TM

   (2011) Structure of the ATP synthase catalytic complex (F₍₁)) from Escherichia coli in an autoinhibited conformation

   Nature Structural & Molecular Biology 18:701–707.

   https://doi.org/10.1038/nsmb.2058

   • PubMed
   • Google Scholar
5. 1. Cipriano DJ
   2. Dunn SD

   (2006) The role of the epsilon subunit in the Escherichia coli ATP synthase: The C-terminal domain is required for efficient energy coupling

   The Journal of Biological Chemistry 281:501–507.

   https://doi.org/10.1074/jbc.M509986200

   • PubMed
   • Google Scholar
6. 1. Cortes PR
   2. Piñas GE
   3. Cian MB
   4. Yandar N
   5. Echenique J

   (2015) Stress-triggered signaling affecting survival or suicide of streptococcus pneumoniae

   International Journal of Medical Microbiology 305:157–169.

   https://doi.org/10.1016/j.ijmm.2014.12.002

   • PubMed
   • Google Scholar
7. 1. Davies RB
   2. Smits C
   3. Wong ASW
   4. Stock D
   5. Christie M
   6. Sandin S
   7. Stewart AG

   (2017) Cryo-EM analysis of a domain antibody bound rotary ATPase complex

   Journal of Structural Biology 197:350–353.

   https://doi.org/10.1016/j.jsb.2017.01.002

   • PubMed
   • Google Scholar
8. 1. Del Rizzo PA
   2. Bi Y
   3. Dunn SD
   4. Shilton BH

   (2002) The "Second Stalk" of Escherichia coli ATP synthase: structure of the isolated dimerization domain

   Biochemistry 41:6875–6884.

   https://doi.org/10.1021/bi025736i

   • PubMed
   • Google Scholar
9. 1. Drobinskaya IY
   2. Kozlov IA
   3. Murataliev MB
   4. Vulfson EN

   (1985) Tightly bound adenosine diphosphate, which inhibits the activity of mitochondrial F₁-ATPase, is located at the catalytic site of the enzyme

   FEBS Letters 182:419–424.

   https://doi.org/10.1016/0014-5793(85)80346-X

   • PubMed
   • Google Scholar
10. 1. Duncan TM
    2. Bulygin VV
    3. Zhou Y
    4. Hutcheon ML
    5. Cross RL

    (1995) Rotation of subunits during catalysis by Escherichia coli F1-ATPase

    PNAS 92:10964–10968.

    https://doi.org/10.1073/pnas.92.24.10964

    • PubMed
    • Google Scholar
11. 1. Emsley P
    2. Lohkamp B
    3. Scott WG
    4. Cowtan K

    (2010) Features and development of coot

    Acta Crystallographica. Section D, Biological Crystallography 66:486–501.

    https://doi.org/10.1107/S0907444910007493

    • PubMed
    • Google Scholar
12. 1. Feniouk BA
    2. Suzuki T
    3. Yoshida M

    (2006) The role of subunit epsilon in the catalysis and regulation of FOF₁-ATP synthase

    Biochimica Et Biophysica Acta (BBA) - Bioenergetics 1757:326–338.

    https://doi.org/10.1016/j.bbabio.2006.03.022

    • PubMed
    • Google Scholar
13. 1. Feniouk BA
    2. Junge W

    (2005) Regulation of the F0F₁-ATP synthase: the conformation of subunit epsilon might be determined by directionality of subunit gamma rotation

    FEBS Letters 579:5114–5118.

    https://doi.org/10.1016/j.febslet.2005.08.030

    • PubMed
    • Google Scholar
14. 1. Ferguson SA
    2. Cook GM
    3. Montgomery MG
    4. Leslie AG
    5. Walker JE

    (2016) Regulation of the thermoalkaliphilic F1-ATPase from Caldalkalibacillus thermarum

    PNAS 113:10860–10865.

    https://doi.org/10.1073/pnas.1612035113

    • PubMed
    • Google Scholar
15. 1. Ferrándiz MJ
    2. de la Campa AG

    (2002) The membrane-associated F₍₀₎F₍₁₎ ATPase is essential for the viability of streptococcus pneumoniae

    FEMS Microbiology Letters 212:133–138.

    https://doi.org/10.1111/j.1574-6968.2002.tb11256.x

    • PubMed
    • Google Scholar
16. 1. Gerlini A
    2. Colomba L
    3. Furi L
    4. Braccini T
    5. Manso AS
    6. Pammolli A
    7. Wang B
    8. Vivi A
    9. Tassini M
    10. van Rooijen N
    11. Pozzi G
    12. Ricci S
    13. Andrew PW
    14. Koedel U
    15. Moxon ER
    16. Oggioni MR

    (2014) The role of host and microbial factors in the pathogenesis of pneumococcal bacteraemia arising from a single bacterial cell bottleneck

    PLOS Pathogens 10:e1004026.

    https://doi.org/10.1371/journal.ppat.1004026

    • PubMed
    • Google Scholar
17. 1. Gledhill JR
    2. Montgomery MG
    3. Leslie AG
    4. Walker JE

    (2007) How the regulatory protein, IF₍₁), inhibits F₍₁)-ATPase from bovine mitochondria

    PNAS 104:15671–15676.

    https://doi.org/10.1073/pnas.0707326104

    • PubMed
    • Google Scholar
18. 1. Goddard TD
    2. Huang CC
    3. Meng EC
    4. Pettersen EF
    5. Couch GS
    6. Morris JH
    7. Ferrin TE

    (2018) UCSF ChimeraX: meeting modern challenges in visualization and analysis

    Protein Science 27:14–25.

    https://doi.org/10.1002/pro.3235

    • PubMed
    • Google Scholar
19. 1. Guerrero KJ
    2. Xue ZX
    3. Boyer PD

    (1990)

    Active/inactive state transitions of the chloroplast F₁ ATPase are induced by a slow binding and release of ^Mg2+. Relationship to catalysis and control of F₁ ATPases

    The Journal of Biological Chemistry 265:16280–16287.

    • PubMed
    • Google Scholar
20. Preprint

    1. Guo HS
    2. Rubinstein JL

    (2018) Structure of a bacterial ATP synthase

    bioRxiv.

    https://doi.org/10.7554/elife.43128

    • Google Scholar
21. 1. Hahn A
    2. Vonck J
    3. Mills DJ
    4. Meier T
    5. Kühlbrandt W

    (2018) Structure, mechanism, and regulation of the chloroplast ATP synthase

    Science 360:eaat4318.

    https://doi.org/10.1126/science.aat4318

    • PubMed
    • Google Scholar
22. 1. Huang H
    2. Yan Y
    3. Zuo Z
    4. Yang L
    5. Li B
    6. Song Y
    7. Liao L

    (2010) Determination of Adenosine phosphates in rat gastrocnemius at various postmortem intervals using high performance liquid chromatography

    Journal of Forensic Sciences 55:1362–1366.

    https://doi.org/10.1111/j.1556-4029.2010.01450.x

    • PubMed
    • Google Scholar
23. 1. Hyndman DJ
    2. Milgrom YM
    3. Bramhall EA
    4. Cross RL

    (1994) Nucleotide-binding sites on Escherichia coli F₁-ATPase. Specificity of noncatalytic sites and inhibition at catalytic sites by MgADP

    The Journal of Biological Chemistry 269:28871–28877.

    https://doi.org/10.1371/journal.pone.0107197

    • PubMed
    • Google Scholar
24. 1. Imamura H
    2. Nhat KP
    3. Togawa H
    4. Saito K
    5. Iino R
    6. Kato-Yamada Y
    7. Nagai T
    8. Noji H

    (2009) Visualization of ATP levels inside single living cells with fluorescence resonance energy transfer-based genetically encoded indicators

    PNAS 106:15651–15656.

    https://doi.org/10.1073/pnas.0904764106

    • PubMed
    • Google Scholar
25. 1. Ishmukhametov RR
    2. Galkin MA
    3. Vik SB

    (2005) Ultrafast purification and reconstitution of His-tagged cysteine-less Escherichia coli F1F_o ATP synthase

    Biochimica Et Biophysica Acta (BBA) - Bioenergetics 1706:110–116.

    https://doi.org/10.1016/j.bbabio.2004.09.012

    • PubMed
    • Google Scholar
26. 1. Ishmukhametov R
    2. Hornung T
    3. Spetzler D
    4. Frasch WD

    (2010) Direct observation of stepped proteolipid ring rotation in E. coli F₀F₁-ATP synthase

    The EMBO Journal 29:3911–3923.

    https://doi.org/10.1038/emboj.2010.259

    • PubMed
    • Google Scholar
27. 1. Kato-Yamada Y

    (2005) Isolated epsilon subunit of Bacillus subtilis F₁-ATPase binds ATP

    FEBS Letters 579:6875–6878.

    https://doi.org/10.1016/j.febslet.2005.11.036

    • PubMed
    • Google Scholar
28. 1. Kato-Yamada Y
    2. Yoshida M

    (2003) Isolated epsilon subunit of thermophilic F₁-ATPase binds ATP

    The Journal of Biological Chemistry 278:36013–36016.

    https://doi.org/10.1074/jbc.M306140200

    • PubMed
    • Google Scholar
29. 1. Keis S
    2. Stocker A
    3. Dimroth P
    4. Cook GM

    (2006) Inhibition of ATP hydrolysis by thermoalkaliphilic F1F_o-ATP synthase is controlled by the C terminus of the epsilon subunit

    Journal of Bacteriology 188:3796–3804.

    https://doi.org/10.1128/JB.00040-06

    • PubMed
    • Google Scholar
30. 1. Krah A
    2. Kato-Yamada Y
    3. Takada S

    (2017) The structural basis of a high affinity ATP binding ε subunit from a bacterial ATP synthase

    PLOS ONE 12:e0177907.

    https://doi.org/10.1371/journal.pone.0177907

    • PubMed
    • Google Scholar
31. 1. Laget PP
    2. Smith JB

    (1979) Inhibitory properties of endogenous subunit epsilon in the Escherichia coli F₁ ATPase

    Archives of Biochemistry and Biophysics 197:83–89.

    https://doi.org/10.1016/0003-9861(79)90222-4

    • PubMed
    • Google Scholar
32. 1. Lee LK
    2. Stewart AG
    3. Donohoe M
    4. Bernal RA
    5. Stock D

    (2010) The structure of the peripheral stalk of thermus thermophilus H+-ATPase/synthase

    Nature Structural & Molecular Biology 17:373–378.

    https://doi.org/10.1038/nsmb.1761

    • PubMed
    • Google Scholar
33. 1. Mendoza-Hoffmann F
    2. Zarco-Zavala M
    3. Ortega R
    4. García-Trejo JJ

    (2018) Control of rotation of the F1FO-ATP synthase nanomotor by an inhibitory α-helix from unfolded ε or intrinsically disordered ζ and IF1 proteins

    Journal of Bioenergetics and Biomembranes 50:403–424.

    https://doi.org/10.1007/s10863-018-9773-9

    • Google Scholar
34. 1. Micheli V
    2. Simmonds HA
    3. Bari M
    4. Pompucci G

    (1993) HPLC determination of oxidized and reduced pyridine coenzymes in human erythrocytes

    Clinica Chimica Acta 220:1–17.

    https://doi.org/10.1016/0009-8981(93)90002-L

    • PubMed
    • Google Scholar
35. 1. Morales-Rios E
    2. Montgomery MG
    3. Leslie AG
    4. Walker JE

    (2015) Structure of ATP synthase from Paracoccus denitrificans determined by X-ray crystallography at 4.0 Å resolution

    PNAS 112:13231–13236.

    https://doi.org/10.1073/pnas.1517542112

    • PubMed
    • Google Scholar
36. Preprint

    1. Murphy BJK
    2. Langer JD
    3. Mills DJ
    4. Yildiz Ö
    5. Kühlbrandt W

    (2019) Rotary substates of mitochondrial ATP synthase reveal the basis of flexible F1-Fo coupling

    bioRxiv.

    https://doi.org/10.1101/543108

    • Google Scholar
37. 1. Nakanishi-Matsui M
    2. Sekiya M
    3. Yano S
    4. Futai M

    (2014) Inhibition of F1-ATPase rotational catalysis by the carboxyl-terminal domain of the ϵ subunit

    The Journal of Biological Chemistry 289:30822–30831.

    https://doi.org/10.1074/jbc.M114.578872

    • PubMed
    • Google Scholar
38. 1. Nakano M
    2. Imamura H
    3. Toei M
    4. Tamakoshi M
    5. Yoshida M
    6. Yokoyama K

    (2008) ATP hydrolysis and synthesis of a rotary motor V-ATPase from thermus thermophilus

    Journal of Biological Chemistry 283:20789–20796.

    https://doi.org/10.1074/jbc.M801276200

    • PubMed
    • Google Scholar
39. 1. Pullman ME
    2. Monroy GC

    (1963)

    A naturally occurring inhibitor of mitochondrial adenosine triphosphatase

    The Journal of Biological Chemistry 238:3762–3769.

    • PubMed
    • Google Scholar
40. 1. Punjani A
    2. Rubinstein JL
    3. Fleet DJ
    4. Brubaker MA

    (2017) cryoSPARC: algorithms for rapid unsupervised cryo-EM structure determination

    Nature Methods 14:290–296.

    https://doi.org/10.1038/nmeth.4169

    • PubMed
    • Google Scholar
41. 1. Richter ML

    (2004) Gamma-epsilon interactions regulate the chloroplast ATP synthase

    Photosynthesis Research 79:319–329.

    https://doi.org/10.1023/B:PRES.0000017157.08098.36

    • PubMed
    • Google Scholar
42. 1. Rodgers AJ
    2. Wilce MC

    (2000) Structure of the γ–ɛ complex of ATP synthase

    Nature Structural & Molecular Biology 7:1051–1054.

    https://doi.org/10.1038/80975

    • Google Scholar
43. 1. Scheres SH

    (2012) RELION: implementation of a bayesian approach to cryo-EM structure determination

    Journal of Structural Biology 180:519–530.

    https://doi.org/10.1016/j.jsb.2012.09.006

    • PubMed
    • Google Scholar
44. 1. Shah NB
    2. Hutcheon ML
    3. Haarer BK
    4. Duncan TM

    (2013) F1-ATPase of Escherichia coli: the ε- inhibited state forms after ATP hydrolysis, is distinct from the ADP-inhibited state, and responds dynamically to catalytic site ligands

    The Journal of Biological Chemistry 288:9383–9395.

    https://doi.org/10.1074/jbc.M113.451583

    • PubMed
    • Google Scholar
45. 1. Shah NB
    2. Duncan TM

    (2015) Aerobic growth of Escherichia coli Is Reduced, and ATP Synthesis Is selectively inhibited when five c-terminal residues are deleted from the ϵ subunit of ATP synthase

    Journal of Biological Chemistry 290:21032–21041.

    https://doi.org/10.1074/jbc.M115.665059

    • PubMed
    • Google Scholar
46. 1. Shirakihara Y
    2. Shiratori A
    3. Tanikawa H
    4. Nakasako M
    5. Yoshida M
    6. Suzuki T

    (2015) Structure of a thermophilic F₁ -ATPase inhibited by an ε-subunit: deeper insight into the ε-inhibition mechanism

    FEBS Journal 282:2895–2913.

    https://doi.org/10.1111/febs.13329

    • Google Scholar
47. 1. Sielaff H
    2. Duncan TM
    3. Börsch M

    (2018) The regulatory subunit ε in Escherichia coli FOF1-ATP synthase

    Biochimica Et Biophysica Acta (BBA) - Bioenergetics 1859:775–788.

    https://doi.org/10.1016/j.bbabio.2018.06.013

    • Google Scholar
48. 1. Sielaff H
    2. Yanagisawa S
    3. Frasch WD
    4. Junge W
    5. Börsch M

    (2019) Structural asymmetry and kinetic limping of single rotary F-ATP synthases

    Molecules 24:504.

    https://doi.org/10.3390/molecules24030504

    • PubMed
    • Google Scholar
49. 1. Sobti M
    2. Smits C
    3. Wong AS
    4. Ishmukhametov R
    5. Stock D
    6. Sandin S
    7. Stewart AG

    (2016) Cryo-EM structures of the autoinhibited E. coli ATP synthase in three rotational states

    eLife 5:e21598.

    https://doi.org/10.7554/eLife.21598

    • PubMed
    • Google Scholar
50. 1. Stewart AG
    2. Lee LK
    3. Donohoe M
    4. Chaston JJ
    5. Stock D

    (2012) The dynamic stator stalk of rotary ATPases

    Nature Communications 3:687.

    https://doi.org/10.1038/ncomms1693

    • PubMed
    • Google Scholar
51. 1. Stewart AG
    2. Laming EM
    3. Sobti M
    4. Stock D

    (2014) Rotary ATPases--dynamic molecular machines

    Current Opinion in Structural Biology 25:40–48.

    https://doi.org/10.1016/j.sbi.2013.11.013

    • PubMed
    • Google Scholar
52. 1. Stewart AG

    (2014) The molecular V Brake

    Journal of Molecular Biology 426:273–274.

    https://doi.org/10.1016/j.jmb.2013.10.003

    • PubMed
    • Google Scholar
53. 1. Stewart AG
    2. Stock D

    (2012) Priming a molecular motor for disassembly

    Structure 20:1799–1800.

    https://doi.org/10.1016/j.str.2012.10.003

    • PubMed
    • Google Scholar
54. 1. Taniguchi N
    2. Suzuki T
    3. Berney M
    4. Yoshida M
    5. Cook GM

    (2011) The regulatory C-terminal domain of subunit ε of F₀F₁ ATP synthase is dispensable for growth and survival of Escherichia coli

    Journal of Bacteriology 193:2046–2052.

    https://doi.org/10.1128/JB.01422-10

    • PubMed
    • Google Scholar
55. 1. Tsunoda SP
    2. Rodgers AJ
    3. Aggeler R
    4. Wilce MC
    5. Yoshida M
    6. Capaldi RA

    (2001) Large conformational changes of the epsilon subunit in the bacterial F1F₀ ATP synthase provide a ratchet action to regulate this rotary motor enzyme

    PNAS 98:6560–6564.

    https://doi.org/10.1073/pnas.111128098

    • PubMed
    • Google Scholar
56. 1. Uhlin U
    2. Cox GB
    3. Guss JM

    (1997) Crystal structure of the epsilon subunit of the proton-translocating ATP synthase from Escherichia coli

    Structure 5:1219–1230.

    https://doi.org/10.1016/S0969-2126(97)00272-4

    • PubMed
    • Google Scholar
57. 1. Walker JE

    (2013) The ATP synthase: the understood, the uncertain and the unknown

    Biochemical Society transactions 41:1–16.

    https://doi.org/10.1042/BST20110773

    • PubMed
    • Google Scholar
58. 1. Walker JE
    2. Dickson VK

    (2006) The peripheral stalk of the mitochondrial ATP synthase

    Biochimica Et Biophysica Acta (BBA) - Bioenergetics 1757:286–296.

    https://doi.org/10.1016/j.bbabio.2006.01.001

    • PubMed
    • Google Scholar
59. 1. Wilkens S
    2. Capaldi RA

    (1998a) Solution structure of the epsilon subunit of the F₁-ATPase from Escherichia coli and interactions of this subunit with beta subunits in the complex

    The Journal of Biological Chemistry 273:26645–26651.

    https://doi.org/10.2210/pdb1bsh/pdb

    • PubMed
    • Google Scholar
60. 1. Wilkens S
    2. Capaldi RA

    (1998b) Electron microscopic evidence of two stalks linking the F₁ and F₀ parts of the Escherichia coli ATP synthase

    Biochimica Et Biophysica Acta (BBA) - Bioenergetics 1365:93–97.

    https://doi.org/10.1016/S0005-2728(98)00048-6

    • PubMed
    • Google Scholar
61. 1. Yagi H
    2. Kajiwara N
    3. Tanaka H
    4. Tsukihara T
    5. Kato-Yamada Y
    6. Yoshida M
    7. Akutsu H

    (2007) Structures of the thermophilic F₁-ATPase epsilon subunit suggesting ATP-regulated arm motion of its C-terminal domain in F₁

    PNAS 104:11233–11238.

    https://doi.org/10.1073/pnas.0701045104

    • PubMed
    • Google Scholar
62. 1. Yaginuma H
    2. Kawai S
    3. Tabata KV
    4. Tomiyama K
    5. Kakizuka A
    6. Komatsuzaki T
    7. Noji H
    8. Imamura H

    (2014) Diversity in ATP concentrations in a single bacterial cell population revealed by quantitative single-cell imaging

    Scientific Reports 4:6522.

    https://doi.org/10.1038/srep06522

    • PubMed
    • Google Scholar
63. 1. Zhang K

    (2016) Gctf: real-time CTF determination and correction

    Journal of Structural Biology 193:1–12.

    https://doi.org/10.1016/j.jsb.2015.11.003

    • PubMed
    • Google Scholar
64. 1. Zheng SQ
    2. Palovcak E
    3. Armache JP
    4. Verba KA
    5. Cheng Y
    6. Agard DA

    (2017) MotionCor2: anisotropic correction of beam-induced motion for improved cryo-electron microscopy

    Nature Methods 14:331–332.

    https://doi.org/10.1038/nmeth.4193

    • PubMed
    • Google Scholar
65. 1. Zhou JM
    2. Xue ZX
    3. Du ZY
    4. Melese T
    5. Boyer PD

    (1988) Relationship of tightly bound ADP and ATP to control and catalysis by chloroplast ATP synthase

    Biochemistry 27:5129–5135.

    https://doi.org/10.1021/bi00414a027

    • PubMed
    • Google Scholar
66. 1. Zhou A
    2. Rohou A
    3. Schep DG
    4. Bason JV
    5. Montgomery MG
    6. Walker JE
    7. Grigorieff N
    8. Rubinstein JL

    (2015) Structure and conformational states of the bovine mitochondrial ATP synthase by cryo-EM

    eLife 4:e10180.

    https://doi.org/10.7554/eLife.10180

    • PubMed
    • Google Scholar

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

• 2,993

  views

• 415

  downloads

• 51

  citations

Views, downloads and citations are aggregated across all versions of this paper published by eLife.