Abstract

The γ-subunit of cyanobacterial and chloroplast ATP synthase, the rotary shaft of F1-ATPase, equips a specific insertion region that is only observed in photosynthetic organisms. This region plays a physiologically pivotal role in enzyme regulation, such as in ADP inhibition and redox response. Recently solved crystal structures of the γ-subunit of F1-ATPase from photosynthetic organisms revealed that the insertion region forms a β-hairpin structure, which is positioned along the central stalk. The structure–function relationship of this specific region was studied by constraining the expected conformational change in this region caused by the formation of a disulfide bond between Cys residues introduced on the central stalk and this β-hairpin structure. This fixation of the β-hairpin region in the α3β3γ complex affects both ADP inhibition and the binding of the ε-subunit to the complex, indicating the critical role that the β-hairpin region plays as a regulator of the enzyme. This role must be important for the maintenance of the intracellular ATP levels in photosynthetic organisms.

Introduction

FoF1-ATP synthase (FoF1) is ubiquitous in energy-transducing membranes, such as chloroplast and cyanobacterial thylakoid membranes, mitochondrial inner membranes, and bacterial plasma membranes. FoF1 synthesizes ATP from ADP and inorganic phosphate by using the electrochemical proton gradient formed across these membranes via the photosynthetic or respiratory electron transfer reaction [1,2]. When FoF1 catalyzes ATP hydrolysis, protons are transported in the opposite direction across membranes from that for ATP synthesis conditions. FoF1 consists of the membrane-embedded portion Fo and the water-soluble portion F1. Fo works as a proton translocation device and is composed of a, b, and c subunits, with a stoichiometry of a1b2c10–15 [39], whereas F1 is the catalytic core for ATP synthesis and hydrolysis, and is composed of five different subunits designated α–ε, with a stoichiometry of α3β3γ1δ1ε1 [10]. The minimum catalytically active complex for F1-ATPase is the α3β3γ complex [1114], and the catalytic sites reside on each of the three β-subunits at the interface with the α-subunits [15]. The rotary catalysis mechanism was first proposed by Boyer and co-workers based on detailed kinetic analyses of F1-ATPase [16]. Following the crystal structure analysis of F1-ATPase, which clearly indicated that the γ-subunit is a rotary shaft within the α3β3 hexamer [15], the counterclockwise continuous rotation of the γ-subunit during ATP hydrolysis was observed by single-molecule observation techniques [17,18].

Since ATP is an essential physiological energy resource for all living cells, the cellular ATP level must be controlled in response to changes in the environment, and futile ATP hydrolysis by FoF1 must be prevented. The most common regulatory mechanism of F1-ATPase is ADP-induced inhibition (ADP inhibition), irrespective of the source of the enzyme. MgADP produced by ATP hydrolysis remains at the catalytic site, and strongly inhibits ATP hydrolysis but not ATP synthesis [1923]. Another inhibitory mechanism, referred to as ε-inhibition, is attributed to the function of the intrinsic ε-subunit, which works as an inhibitor of ATP hydrolysis for bacterial and chloroplast enzymes. In the FoF1 complex, the ε-subunit is located at the bottom of the γ-subunit, and the C-terminal helix-turn-helix domain of the ε subunit of bacterial F1 shows a large conformational change from a retracted form to an extended form in response to external stimuli, such as a change in ATP level [2427]. This large conformational change is supposed to be the cause of the ε-inhibition [28,29]. Although the ε-subunit of cyanobacteria and chloroplasts appears to inhibit ATP hydrolysis more strongly than the bacterial ε-subunit [3032], the conformational change at the C-terminal helical region is unlikely to occur in the case of the cyanobacterial ε-subunit [33]. Single-molecule observations of ε-inhibition on rotation of the γ-subunit in cyanobacterial and bacterial F1-ATPase showed that the ε-subunit of cyanobacterial F1 completely stops the rotation [34], whereas the ε-subunit of bacterial F1 decreases the average rotation speed and increases the pause duration [35]. These inhibitions are important to avoid futile ATP hydrolysis and to ensure efficient ATP synthesis in vivo.

Because of the unique rotating shaft movement, the γ-subunit was expected to function in regulating ATP synthesis/hydrolysis. Cyanobacterial and chloroplast γ-subunits consist of an insertion region comprising 30–40 amino acids, in comparison with the γ-subunits of the bacterial and mitochondrial ATP synthases [36,37]. Although the insertion region of the chloroplast γ-subunit contains nine additional amino acids, including two redox-reactive cysteines [3840], the insertion region of cyanobacteria lacks this 9-amino-acid portion. In our previous study, we determined the crystal structure of the γ–ε complex of F1-ATPase from the thermophilic cyanobacterium Thermosynechococcus elongatus BP-1 at a resolution of 1.98 Å [41]. According to this structure, the insertion region forms a β-hairpin structure, which is positioned along the central stalk, and the top of the insertion region appears to interact with the DELSEED region of the β-subunit. This negatively charged region is considered to be important for the enzyme activity of FoF1 [4244]. Furthermore, higher levels of B-factors in the insertion region of the γ-subunit indicate the considerable flexibility of this region [41]. A similar structure is also observed in the γ-subunit of chloroplast FoF1 [45].

Based on these findings, we expected that the β-hairpin structure of the insertion region plays a regulatory role via certain movement in response to environmental stimuli, although the cyanobacterial γ-subunit does not contain the redox-regulation motif including two cysteines. We therefore prepared a mutant F1-ATPase of T. elongatus BP-1 to restrict the movement of this region by forming a disulfide bond between the introduced Cys residues on the β-hairpin structure and the central stalk of the γ-subunit. Our experiments restricting the β-hairpin structure clearly indicated that the insertion region is essential to control both ADP inhibition and ε-inhibition to prevent futile ATP hydrolysis.

Experimental procedures

Materials

Pyruvate kinase, lactate dehydrogenase, and NADH were purchased from Roche Diagnostics (Basel, Switzerland). Other chemicals were of the highest grade commercially available.

Strains

Escherichia coli strains used were DH5α for cloning and BL21(DE3) uncΔ702 [Tcr, ATPase mutant, BL21(DE3) uncΔ702, asnA::Tn10] for expression of the α3β3γ complex of T. elongatus BP-1. The latter strain was a kind gift from Dr. C. S. Harwood (University of Iowa).

Construction of expression plasmids for the α3β3γ complex containing mutant γ-subunit

The expression plasmid for the α3β3γ complex of T. elongatus BP-1, pTR19FR, in which a deca-histidine tag was fused to the N-terminal of the β-subunit, was originally constructed in a previous study [34]. Using the plasmid as a template, expression plasmids for the complex containing the mutant γ-subunit were prepared. First, the γ-subunit-coding region, atpC, was amplified by PCR, and was cloned into the pGEM-T Easy Vector. Then, site-directed mutagenesis was performed by the overlap extension method to introduce the desired two Cys residues into the γ-subunit. The primers used for mutagenesis are shown in Supplemental Table S1. The vector containing mutated atpC was digested with EcoRI and NheI, and the mutated atpC fragment was fused to an EcoRI and NheI site of pTR19FR. Consequently, the expression plasmid for the α3β3γ containing the mutation at the insertion region of the γ-subunit was obtained.

Expression and purification of α3β3γ complex and ε-subunit

Expression and purification of the α3β3γ complex were performed as described previously [34] with some modifications. E. coli strain BL21 (DE3) unc 702, transformed with the desired plasmid, was cultured in 2xYT medium containing 100 µg/ml ampicillin and 0.2 mM isopropyl-β-d-thiogalactopyranoside (IPTG) at 37°C for 20 h. The expressed proteins were purified by Ni-affinity chromatography. The obtained protein complex was then subjected to size-exclusion column chromatography on a Superdex 200 Increase column 10/300 (GE Healthcare, Little Chalfont, U.K.), which was equilibrated with 50 mM HEPES–KOH, pH 8.0, 100 mM KCl, 0.1 mM MgCl2, and 0.1 mM ATP. The elution peak positioned at ∼11 ml at a flow rate of 0.5 ml/min was collected. The obtained complex protein was then flash-frozen in liquid nitrogen and stored at −80°C in 10% glycerol until use. Protein concentrations were determined using the Bradford method (Bio-Rad Protein Assay; Bio-Rad Laboratories, Inc., Hercules, U.S.A.) with bovine serum albumin as a standard.

The ε-subunit was expressed and purified as described elsewhere [34] with some modifications, and stored at −80°C in 10% glycerol until use.

Determination of the optimal oxidation and reduction conditions

Purified α3β3γ complex containing mutation in the γ-subunit was oxidized or reduced by various reagents (diamide, aldrithiol-2, and CuCl2 for oxidation, and DTT for reduction). The concentrations of the reagents used were 5, 50, and 500 µM for diamide and aldrithiol-2; 0.5, 5, and 50 mM for DTT; and 50 and 100 µM for CuCl2, for 30 and 60 min, at 25, 37, and 50°C. The reaction was then stopped by adding 5% (w/v, final concentration) trichloroacetic acid. After centrifugation, the supernatant was removed and the precipitate was washed away with 80 µl of acetone. The mixture was then centrifuged and air-dried after removing the supernatant. The precipitate was dissolved in AMS-labeling buffer (62.5 mM Tris–HCl, pH 6.8, 2% SDS, 7.5% glycerol, 0.01% BPB, and 2 mM AMS).

Measurement of ATP hydrolysis activity

ATP hydrolysis activities were measured as described previously [46] using an ATP-regenerating system (50 mM HEPES–KOH, pH 8.0, 100 mM KCl, 2 mM MgCl2, 2 mM phosphoenolpyruvate, 50 µg/ml pyruvate kinase/lactate dehydrogenase, 0.2 mM NADH, and 2 mM ATP) at 25°C. ATP hydrolysis was initiated by adding 2–20 µg of α3β3γ complexes to 1.2 ml of the assay buffer with or without LDAO (lauryldimethylamine-N-oxide) at a final concentration of 0.1% (v/v). The ATP hydrolysis rate after the addition of the enzyme was determined by monitoring the decrease in NADH absorption at 340 nm using a spectrophotometer V-550 (Jasco, Tokyo, Japan). The pH dependence of ATP hydrolysis activity was evaluated by the hydrazine sulfate reduction method [47]. To determine the extent of ε-inhibition, the ε-subunit was added into the ATP hydrolysis assay mixture. The ATP hydrolysis activity, the activation ratio by LDAO, and the extent of inhibition was then determined by the steady-state slope (Supplementary Figures S1–S3) [13].

Results and discussion

Preparation of locked mutant γ-subunit to restrict the movement of the β-hairpin structure of the insertion region

Based on the structure in the γ–ε complex of F1-ATPase obtained from T. elongatus BP-1 [41], we prepared mutant γ-subunits whose β-hairpin structure of the insertion region can be locked to the central stalk of the γ-subunit via the formation of a disulfide bond between the introduced Cys residues. Val32 and Ala50 on the N-terminal α-helix, and Ala201, Leu208, and Thr210 on the β-strand of the insertion region were selected as candidate residues for Cys substitution (Figure 1). Accordingly, we prepared three mutants by combination of these Cys substitutions (V32C/L208C, V32C/T210C, and A50C/A201C). Site-directed mutagenesis was performed as described in ‘Experimental Procedures,' and all mutants were successfully expressed in E. coli and accordingly purified.

Position of Cys residues introduced for locking of the insertion region of the γ-subunit.

Figure 1.
Position of Cys residues introduced for locking of the insertion region of the γ-subunit.

Three mutants (V32C/L208C, V32C/T210C, and A50C/A201C) were prepared for locking the insertion region. (A) Side view of the cyanobacterial γ–ε complex structure (PDB ID: 5ZWL) superimposed on the whole F1-ATPase (α3β3γε complex) from Caldalkalibacillus thermarum (PDB ID: 5HKK). The γ-subunit, ε-subunit and insertion region are colored in green, sky blue, and orange, respectively. The β-, γ-, and ε-subunits of C. thermatum are colored in pink, blue, and red, respectively. Three α-subunits and βDP are removed for clarity. (B) Positions of introduced Cys residues for locking the insertion region of the γ-subunit are indicated. The introduced Cys residues are colored in purple. The figures were generated using UCSF Chimera (https://www.cgl.ucsf.edu/chimera/).

Figure 1.
Position of Cys residues introduced for locking of the insertion region of the γ-subunit.

Three mutants (V32C/L208C, V32C/T210C, and A50C/A201C) were prepared for locking the insertion region. (A) Side view of the cyanobacterial γ–ε complex structure (PDB ID: 5ZWL) superimposed on the whole F1-ATPase (α3β3γε complex) from Caldalkalibacillus thermarum (PDB ID: 5HKK). The γ-subunit, ε-subunit and insertion region are colored in green, sky blue, and orange, respectively. The β-, γ-, and ε-subunits of C. thermatum are colored in pink, blue, and red, respectively. Three α-subunits and βDP are removed for clarity. (B) Positions of introduced Cys residues for locking the insertion region of the γ-subunit are indicated. The introduced Cys residues are colored in purple. The figures were generated using UCSF Chimera (https://www.cgl.ucsf.edu/chimera/).

Determination of the optimal oxidation and reduction conditions

We examined the formation/cleavage conditions of the disulfide bond to lock and unlock the β-hairpin structure of the insertion region by changing the concentrations of oxidizing/reducing reagents, temperatures, and reaction periods. To evaluate the efficiency of disulfide bond formation, non-reducing SDS–PAGE analysis was performed. The locked γ-subunit showed a clear band shift on the gel as a consequence of the conformational constraints (Figure 2A,B). The oxidized and reduced γ-subunits were observed at 30 and 36 kDa, respectively. Similar band shifts caused by the intramolecular disulfide bond formation were also observed in our previous study [48]. Consequently, V32C/L208V, V32C/T210C, and A50C/A201C mutants were mostly oxidized (Figure 2A) by 50 µM diamide, and reduced by 50 mM DTT. It was confirmed that the oxidation and reduction treatments did not affect the ATP hydrolysis activity (Supplementary Figure S4A), the activation ratio by LDAO (Supplementary Figure S4B,C), pH dependence of the activity (Supplementary Figure S5) and the ε-inhibition (Supplementary Figure S6) of the wild-type complex. We therefore concluded that the change in the activity caused by oxidation or reduction in the mutants was simply originating from the locking or unlocking the insertion region. We then investigated the effect of locking of the insertion region on these mutants.

Non-reducing SDS–PAGE and western blotting analysis of unlocked and locked mutants.

Figure 2.
Non-reducing SDS–PAGE and western blotting analysis of unlocked and locked mutants.

(A) Untreated (U), reduced (R), and oxidized (O) α3β3γ complexes (5 µg/lane) were analyzed by 12% SDS–PAGE and stained by Coomassie Brilliant Blue R-250. V32C/L208V, V32C/T210C, and A50C/A201C mutants were mostly oxidized by 50 µM diamide for 30 min at 25°C, and reduced by 50 mM DTT for 30 min at 25°C. The oxidation and reduction conditions of purified wild-type, V32C/L208C, V32C/T210C, and A50C/A201C complexes were surveyed as described under ‘Experimental Procedures' and the most representative results are shown. (B) Western blotting of the untreated, unlocked, and locked γ-subunits (5 µg-α3β3γ/lane) in the mutant complexes. Electrophoresed proteins on 12% polyacrylamide gel were transferred to PVDF membrane, followed by immunoblotting with antibodies against the γ-subunit. The proteins were visualized by enhanced chemiluminescence reagent, and the signals were detected with a LAS-3000 mini luminescent image analyzer (Fujifilm, Tokyo, Japan).

Figure 2.
Non-reducing SDS–PAGE and western blotting analysis of unlocked and locked mutants.

(A) Untreated (U), reduced (R), and oxidized (O) α3β3γ complexes (5 µg/lane) were analyzed by 12% SDS–PAGE and stained by Coomassie Brilliant Blue R-250. V32C/L208V, V32C/T210C, and A50C/A201C mutants were mostly oxidized by 50 µM diamide for 30 min at 25°C, and reduced by 50 mM DTT for 30 min at 25°C. The oxidation and reduction conditions of purified wild-type, V32C/L208C, V32C/T210C, and A50C/A201C complexes were surveyed as described under ‘Experimental Procedures' and the most representative results are shown. (B) Western blotting of the untreated, unlocked, and locked γ-subunits (5 µg-α3β3γ/lane) in the mutant complexes. Electrophoresed proteins on 12% polyacrylamide gel were transferred to PVDF membrane, followed by immunoblotting with antibodies against the γ-subunit. The proteins were visualized by enhanced chemiluminescence reagent, and the signals were detected with a LAS-3000 mini luminescent image analyzer (Fujifilm, Tokyo, Japan).

Change in ATP hydrolysis activity and the LDAO-activation ratio by unlocking and locking the β-hairpin structure of the insertion region

First, we examined the ATP hydrolysis activity and the activation ratio by LDAO of unlocked and locked mutants at pH 7.0 (Figure 3, white bars) and pH 8.0 (Figure 3, gray bars). LDAO is a nonionic detergent that releases F1-ATPase from the intrinsic regulatory mechanism, ADP inhibition. Therefore, the activation ratio by LDAO is considered to be a good indicator of the extent of ADP inhibition of the enzyme. The activity of A50C/A201C was lower and those of V32C/L208C and V32C/T210C was higher than that of WT, and there was no obvious relationship between the lock of the β-hairpin structure of the insertion region and the change in activity of the mutants. Currently, we cannot explain the reason of the change in ATPase activity caused by these cysteine substitutions. The activation ratio by LDAO of locked mutants of A50C/A201C and V32C/T210C was lower than that of unlocked ones though the activation ratio for V32C/L208C was not affected.

ATP hydrolysis activity and LDAO sensitivity at pH 8 and pH 7 of the locked and unlocked mutants.

Figure 3.
ATP hydrolysis activity and LDAO sensitivity at pH 8 and pH 7 of the locked and unlocked mutants.

Gray and white bars represent the results of the assays at pH 8 and pH 7, respectively. The conditions of the insertion region in the mutant complexes are shown as: – (untreated), U (unlocked), and L (locked), respectively. (A and B) ATP hydrolysis activities were measured using an ATP-regenerating system in the absence (A) or presence (B) of LDAO (final concentration of 0.1%). The assay was performed at 25°C, and the activities were determined from the steady-state slope. The results of three experiments were averaged. Error bar represents SD. (C) The ratios of the activities in the presence versus those in the absence of LDAO were calculated from (A) and (B).

Figure 3.
ATP hydrolysis activity and LDAO sensitivity at pH 8 and pH 7 of the locked and unlocked mutants.

Gray and white bars represent the results of the assays at pH 8 and pH 7, respectively. The conditions of the insertion region in the mutant complexes are shown as: – (untreated), U (unlocked), and L (locked), respectively. (A and B) ATP hydrolysis activities were measured using an ATP-regenerating system in the absence (A) or presence (B) of LDAO (final concentration of 0.1%). The assay was performed at 25°C, and the activities were determined from the steady-state slope. The results of three experiments were averaged. Error bar represents SD. (C) The ratios of the activities in the presence versus those in the absence of LDAO were calculated from (A) and (B).

The influence of pH on the insertion region must be critical because pH change in chloroplasts appears to be a significant physiological parameter in photosynthetic organisms. Indeed, a drastic pH change is known to occur during light–dark transition, which must markedly affect the activities of various enzymes involved in photosynthesis [49,50]. Although fluctuation of intracellular pH in cyanobacteria has not yet been reported, that in stroma of chloroplasts where F1-ATPase protrudes from thylakoid membranes has been observed between pH 7 (dark) and pH 8 (light) [49,50]. Therefore, assuming that the fluctuation of intracellular pH in cyanobacteria is similar to that of the stroma in chloroplasts, we examined the ATPase activity of the mutants at pH 7.0 and pH 8.0 (Figure 3). The activities of WT and the mutants at pH 7.0 were lower than those at pH 8.0, irrespective of the lock or unlock of the β-hairpin structure of the insertion region. In addition, the LDAO-activation ratio of WT at pH 7.0 was threefold or higher than that at pH 8.0. These results suggest that the decrease in the activities corresponding with lowering pH observed in the WT and most of the mutants was probably caused by ADP inhibition. In contrast, the activation ratio by LDAO of locked A50C/A201C was not pH dependent, although the unlocked one demonstrated a tendency similar to that of the WT, implying that the bottom of the insertion region, where the mutation A50C/A201C was introduced, may be responsible for regulation regarding the pH dependence of the extent of ADP inhibition.

pH dependence of ADP inhibition

To thoroughly characterize the effect of the bottom of the insertion region on the pH dependence of ADP inhibition, we investigated the enzyme activity by a colorimetric method, which quantifies the liberated phosphate. Consequently, similar pH dependence of the activity and the activation ratio by LDAO were observed for WT and the A50C/A201C mutant (Figure 4) as in the results from the coupling assay (Figure 3). Unlocked A50C/A201C showed the same pH dependence as WT, but locked A50C/A201C did not. These results also support the idea that the pH dependence of ADP inhibition is attributable to the conformation of the bottom of the insertion region. In addition, at pH 8.2, the activity of unlocked A50C/A201C was about threefold higher than that of locked A50C/A201C, while the activation ratio by LDAO of unlocked A50C/A201C was less than twice of that of locked A50C/A201C. Therefore, it is unlikely that the decrease in the activity at high pH, caused by locking the β-hairpin structure of the insertion region, results only from ADP inhibition. There are many negatively charged amino acids around the mutation positions (Supplementary Figure S7). These amino acids might contribute to the pH dependence of the activity and ADP inhibition in response to a pH change. According to the structure of chloroplast FoF1-ATP synthase [45], the mutation positions of A50C/A201C are near the Cys residues, which play roles in the redox regulation of chloroplast F1. Considering these findings together, the redox regulation observed in the chloroplast ATPase is supposed to be due to ADP inhibition and the flexibility of the bottom of the insertion region must provide the required conformational change for this regulation.

pH dependence of ATP hydrolysis activity and LDAO-activation ratios of WT and A50C/A201C.

Figure 4.
pH dependence of ATP hydrolysis activity and LDAO-activation ratios of WT and A50C/A201C.

The insertion region in the A50C/A201C complex was unlocked (open circle), locked (open square), or untreated (open triangle). The activity of WT was measured as well (closed triangle). The results of three independent experiments were averaged. Error bar represents SD. (A and B) pH dependence of ATP hydrolysis activities of WT and A50C/A201C in the absence (A) and those in the presence (B) of LDAO (final concentration of 0.1%) were measured. (C) pH dependence of the LDAO-activation ratios of ATP hydrolysis activities of WT and A50C/A201C were calculated from (A) and (B). The results of three independent experiments were averaged. Error bar represents SD. Large error bars observed at pH 7.2 must be due to the small denominator values obtained for the activity in the absence of LDAO.

Figure 4.
pH dependence of ATP hydrolysis activity and LDAO-activation ratios of WT and A50C/A201C.

The insertion region in the A50C/A201C complex was unlocked (open circle), locked (open square), or untreated (open triangle). The activity of WT was measured as well (closed triangle). The results of three independent experiments were averaged. Error bar represents SD. (A and B) pH dependence of ATP hydrolysis activities of WT and A50C/A201C in the absence (A) and those in the presence (B) of LDAO (final concentration of 0.1%) were measured. (C) pH dependence of the LDAO-activation ratios of ATP hydrolysis activities of WT and A50C/A201C were calculated from (A) and (B). The results of three independent experiments were averaged. Error bar represents SD. Large error bars observed at pH 7.2 must be due to the small denominator values obtained for the activity in the absence of LDAO.

Effect of locking the insertion region on ε-inhibition

ε-inhibition is conferred by the intrinsic function of the ε-subunit. We expected that the binding of the ε-subunit may affect the conformation around the insertion region because the ε-subunit is located adjacent to the γ-subunit in the structure and a β-strand of the γ-subunit forms a β-sheet structure with the ε-subunit [41]. As shown in Figure 5, the extent of the ε-inhibition of locked mutants (open square), especially V32C/T210C, which is locked at the top of the insertion region, was weaker than that of unlocked mutants (open circle), indicating that the locking of the insertion region has a negative effect on ε-inhibition.

The effects of locking the insertion region on ε inhibition.

Figure 5.
The effects of locking the insertion region on ε inhibition.

The extents of the inhibition of the ATP hydrolysis activities of 17.7 nM A50C/A201C (A), 4.7 nM V32C/T210C (B), and 4.7 nM V32C/L208C (C) mutants in untreated (open triangle), unlocked (open circle), and locked (open square) forms were measured in the presence of various concentrations of the ε-subunit. The results of 17.7 nM WT are shown with closed triangle as a control.

Figure 5.
The effects of locking the insertion region on ε inhibition.

The extents of the inhibition of the ATP hydrolysis activities of 17.7 nM A50C/A201C (A), 4.7 nM V32C/T210C (B), and 4.7 nM V32C/L208C (C) mutants in untreated (open triangle), unlocked (open circle), and locked (open square) forms were measured in the presence of various concentrations of the ε-subunit. The results of 17.7 nM WT are shown with closed triangle as a control.

Relative conformational change between the insertion region and the central stalk of the γ-subunit caused by the ε-subunit

The locking of the β-hairpin structure of the insertion region had a negative effect on ε-inhibition (Figure 5). In other words, the locking may interfere with the conformational change of the γ-subunit caused by the binding of the ε-subunit. Therefore, we investigated the effect of ε-binding on the disulfide bond formation ability. The disulfide bond formation efficiency must reflect the change of the distance between two introduced Cys residues on the γ-subunit. The ε-subunit itself should not chemically affect the oxidation or reduction because the ε-subunit has no Cys residue. When oxidation or reduction treatment of α3β3γ was carried out in the absence of the ε-subunit, the locked γ-subunit was obtained at a rate of more than 80% by oxidation (Figure 6A and Supplementary Figure S8). In contrast, the locked γ-subunit was hardly obtained by oxidation, except for A50C/A201C, in the presence of the ε-subunit (Figure 6B and Supplementary Figure S8). The disulfide bond formation of V32C/T210C was strongly suppressed by the addition of the ε-subunit. These results indicate that binding of the ε-subunit to the α3β3γ complex induced the relative conformational change between the α-helix of the central stalk and the β-hairpin structure of the insertion region of the γ-subunit, especially at the top of the insertion region. A similar strong effect of the ε-subunit on the insertion region was also observed when the ε-inhibition was evaluated (Figure 5).

The effects of the binding of the ε subunit to α3β3γ on disulfide bond for locking the insertion region.

Figure 6.
The effects of the binding of the ε subunit to α3β3γ on disulfide bond for locking the insertion region.

(A and B) The ratios of the disulfide bond formation on the locked mutant complex by the oxidation treatment in the absence (A) and the presence (B) of the ε subunit were calculated when the complex was incubated with 0, 5, 50 and 500 µM diamide. The ratios of the disulfide bond formation were calculated from the band intensities of the unlocked and locked γ-subunits shown in Supplemental Figure S8. The band intensities were quantified using Image J software. The results of three experiments were averaged. Error bar represents SD.

Figure 6.
The effects of the binding of the ε subunit to α3β3γ on disulfide bond for locking the insertion region.

(A and B) The ratios of the disulfide bond formation on the locked mutant complex by the oxidation treatment in the absence (A) and the presence (B) of the ε subunit were calculated when the complex was incubated with 0, 5, 50 and 500 µM diamide. The ratios of the disulfide bond formation were calculated from the band intensities of the unlocked and locked γ-subunits shown in Supplemental Figure S8. The band intensities were quantified using Image J software. The results of three experiments were averaged. Error bar represents SD.

Considering these findings together, the binding of the ε-subunit induces the conformational change of the γ-subunit, which plays a significant role in ε-inhibition as well. Because all the previously reported structures of the ε-subunits of photosynthetic organisms show a retracted conformation [33,41,45] and are clearly different from the bacterial one, which can reach the DELSEED region of the β-subunit, the roles of the C-terminal α-helical regions of the chloroplast ε-subunit and the bacterial one must be very different from each other in the complex. The β-hairpin structure of the insertion region of the γ-subunit of photosynthetic organisms may therefore accomplish an alternative role of the C-terminal α-helical region of the bacterial ε-subunit in enzyme regulation.

Abbreviations

     
  • ADP inhibition

    ADP-induced inhibition

  •  
  • FoF1

    FoF1-ATP synthase

  •  
  • IPTG

    isopropyl-β-d-thiogalactopyranoside

  •  
  • LDAO

    lauryldimethylamine-N-oxide

Author Contribution

K.A. and T.H. conceived the study, and K.A. and K.K. performed the experiments. K.W. and T.H. supervised the research. K.A., K.I., K.K., S.M., K.W., and T.H. discussed the data. K.A. and T.H. wrote the paper, and K.K. and K.W. commented on the manuscript.

Funding

This study was supported by JSPS KAKENHI (MEXT-KAKENHI) [Grant Number 16H06556 to T.H.] and by Dynamic Alliance for Open Innovation Bridging Human, Environment and Materials.

Acknowledgements

We thank the Biomaterials Analysis Division, Tokyo Institute of Technology for supporting DNA sequencing analysis.

Competing Interests

The Authors declare that there are no competing interests associated with the manuscript.

References

References
1
Boyer
,
P.D.
(
1997
)
The ATP synthase—a splendid molecular machine
.
Annu. Rev. Biochem.
66
,
717
749
2
Yoshida
,
M.
,
Muneyuki
,
E.
and
Hisabori
,
T.
(
2001
)
ATP synthase—a marvellous rotary engine of the cell
.
Nat. Rev. Mol. Cell Biol.
2
,
669
677
3
Senior
,
A.E.
(
1990
)
The proton-translocating ATPase of Escherichia coli
.
Annu. Rev. Biophys. Biophys. Chem.
19
,
7
41
4
Stock
,
D.
,
Gibbons
,
C.
,
Arechaga
,
I.
,
Leslie
,
A.G.
and
Walker
,
J.E.
(
2000
)
The rotary mechanism of ATP synthase
.
Curr. Opin. Struct. Biol.
10
,
672
679
5
Seelert
,
H.
,
Poetsch
,
A.
,
Dencher
,
N.A.
,
Engel
,
A.
,
Stahlberg
,
H.
and
Muller
,
D.J.
(
2000
)
Structural biology. Proton-powered turbine of a plant motor
.
Nature
405
,
418
419
6
Jiang
,
W.
,
Hermolin
,
J.
and
Fillingame
,
R.H.
(
2001
)
The preferred stoichiometry of c subunits in the rotary motor sector of Escherichia coli ATP synthase is 10
.
Proc. Natl Acad. Sci. U.S.A.
98
,
4966
4971
7
Mitome
,
N.
,
Suzuki
,
T.
,
Hayashi
,
S.
and
Yoshida
,
M.
(
2004
)
Thermophilic ATP synthase has a decamer c-ring: indication of noninteger 10:3 H+/ATP ratio and permissive elastic coupling
.
Proc. Natl Acad. Sci. U.S.A.
101
,
12159
12164
8
Meier
,
T.
,
Polzer
,
P.
,
Diederichs
,
K.
,
Welte
,
W.
and
Dimroth
,
P.
(
2005
)
Structure of the rotor ring of F-Type Na+-ATPase from Ilyobacter tartaricus
.
Science
308
,
659
662
9
Watt
,
I.N.
,
Montgomery
,
M.G.
,
Runswick
,
M.J.
,
Leslie
,
A.G.
and
Walker
,
J.E.
(
2010
)
Bioenergetic cost of making an adenosine triphosphate molecule in animal mitochondria
.
Proc. Natl Acad. Sci. U.S.A.
107
,
16823
16827
10
Yoshida
,
M.
,
Sone
,
N.
,
Hirata
,
H.
,
Kagawa
,
Y.
and
Ui
,
N.
(
1979
)
Subunit structure of adenosine triphosphatase. Comparison of the structure in thermophilic bacterium PS3 with those in mitochondria, chloroplasts, and Escherichia coli
.
J. Biol. Chem.
254
,
9525
9533
PMID:
[PubMed]
11
Matsui
,
T.
and
Yoshida
,
M.
(
1995
)
Expression of the wild-type and the Cys-/Trp-less α3β3γ complex of thermophilic F1-ATPase in Escherichia coli
.
Biochim. Biophys. Acta
1231
,
139
146
12
Kaibara
,
C.
,
Matsui
,
T.
,
Hisabori
,
T.
and
Yoshida
,
M.
(
1996
)
Structural asymmetry of F1-ATPase caused by the gamma subunit generates a high affinity nucleotide binding site
.
J. Biol. Chem.
271
,
2433
2438
13
Hisabori
,
T.
,
Kato
,
Y.
,
Motohashi
,
K.
,
Kroth-Pancic
,
P.
,
Strotmann
,
H.
and
Amano
,
T.
(
1997
)
The regulatory functions of the gamma and epsilon subunits from chloroplast CF1 are transferred to the core complex, alpha3beta3, from thermophilic bacterial F1
.
Eur. J. Biochem.
247
,
1158
1165
14
Du
,
Z.
,
Tucker
,
W.C.
,
Richter
,
M.L.
and
Gromet-Elhanan
,
Z.
(
2001
)
Assembled F1-(αβ) and Hybrid F13β3γ-ATPases from Rhodospirillum rubrum α, wild type or mutant β, and chloroplast γ subunits. Demonstration of Mg2+ versus Ca2+-induced differences in catalytic site structure and function
.
J. Biol. Chem.
276
,
11517
11523
15
Abrahams
,
J.
,
Leslie
,
A.
,
Lutter
,
R.
and
Walker
,
J.
(
1994
)
Structure at 2.8 Å resolution of F1-ATPase from bovine heart mitochondria
.
Nature
370
,
621
628
16
Grubmeyer
,
C.
,
Cross
,
R.L.
and
Penefsky
,
H.S.
(
1982
)
Mechanism of ATP hydrolysis by beef heart mitochondrial ATPase. Rate constants for elementary steps in catalysis at a single site
.
J. Biol. Chem.
257
,
12092
12100
PMID:
[PubMed]
17
Noji
,
H.
,
Yasuda
,
R.
,
Yoshida
,
M.
and
Kinosita
, Jr,
K.
(
1997
)
Direct observation of the rotation of F1-ATPase
.
Nature
386
,
299
302
18
Yasuda
,
R.
,
Noji
,
H.
,
Kinosita
, Jr,
K.
,
Motojima
,
F.
and
Yoshida
,
M.
(
1997
)
Rotation of the γ subunit in F1-ATPase; evidence that ATP synthase is a rotary motor enzyme
.
J. Bioenerg. Biomembr.
29
,
207
209
19
Minkov
,
I.B.
,
Fitin
,
A.F.
,
Vasilyeva
,
E.A.
and
Vinogradov
,
A.D.
(
1979
)
Mg2+-induced ADP-dependent inhibition of the ATPase activity of beef heart mitochondrial coupling factor F1
.
Biochem. Biophys. Res. Commun.
89
,
1300
1306
20
Dunham
,
K.R.
and
Selman
,
B.R.
(
1981
)
Regulation of spinach chloroplast coupling factor 1 ATPase activity
.
J. Biol. Chem.
256
,
212
218
PMID:
[PubMed]
21
Vasilyeva
,
E.A.
,
Minkov
,
I.B.
,
Fitin
,
A.F.
and
Vinogradov
,
A.D.
(
1982
)
Kinetic mechanism of mitochondrial adenosine triphosphatase. ADP-specific inhibition as revealed by the steady-state kinetics
.
Biochem. J.
202
,
9
14
22
Yoshida
,
M.
and
Allison
,
W.S.
(
1983
)
Modulation by ADP and Mg2+ of the inactivation of the F1-ATPase from the thermophilic bacterium, PS3, with dicyclohexylcarbodiimide
.
J. Biol. Chem.
258
,
14407
14412
PMID:
[PubMed]
23
Bald
,
D.
,
Amano
,
T.
,
Muneyuki
,
E.
,
Pitard
,
B.
,
Rigaud
,
J.L.
,
Kruip
,
J.
et al.  (
1998
)
ATP synthesis by F0F1-ATP synthase independent of noncatalytic nucleotide binding sites and insensitive to azide inhibition
.
J. Biol. Chem.
273
,
865
870
24
Rodgers
,
A.J.
and
Wilce
,
M.C.
(
2000
)
Structure of the γ(ε complex of ATP synthase
.
Nat. Struct. Biol.
7
,
1051
1054
25
Gibbons
,
C.
,
Montgomery
,
M.G.
,
Leslie
,
A.G.
and
Walker
,
J.E.
(
2000
)
The structure of the central stalk in bovine F1-ATPase at 2.4 Å resolution
.
Nat Struct Biol.
7
,
1055
1061
26
Tsunoda
,
S.P.
,
Rodgers
,
A.J.
,
Aggeler
,
R.
,
Wilce
,
M.C.
,
Yoshida
,
M.
and
Capaldi
,
R.A.
(
2001
)
Large conformational changes of the ε subunit in the bacterial F1F0 ATP synthase provide a ratchet action to regulate this rotary motor enzyme
.
Proc. Natl Acad. Sci. U.S.A.
98
,
6560
6564
27
Cingolani
,
G.
and
Duncan
,
T.M.
(
2011
)
Structure of the ATP synthase catalytic complex (F1) from Escherichia coli in an autoinhibited conformation
.
Nat. Struct. Mol. Biol.
18
,
701
707
28
Hara
,
K.Y.
,
Kato-Yamada
,
Y.
,
Kikuchi
,
Y.
,
Hisabori
,
T.
and
Yoshida
,
M.
(
2001
)
The role of the βDELSEED motif of F1-ATPase: propagation of the inhibitory effect of the ε subunit
.
J. Biol. Chem.
276
,
23969
23973
29
Kato-Yamada
,
Y.
,
Bald
,
D.
,
Koike
,
M.
,
Motohashi
,
K.
,
Hisabori
,
T.
and
Yoshida
,
M.
(
1999
)
ε subunit, an endogenous inhibitor of bacterial F1-ATPase, also inhibits F0F1-ATPase
.
J. Biol. Chem.
274
,
33991
33994
30
Richter
,
M.L.
,
Patrie
,
W.J.
and
McCarty
,
R.E.
(
1984
)
Preparation of the ε subunit and ε subunit-deficient chloroplast coupling factor 1 in reconstitutively active forms
.
J. Biol. Chem.
259
,
7371
7373
PMID:
[PubMed]
31
Weber
,
J.
,
Dunn
,
S.D.
and
Senior
,
A.E.
(
1999
)
Effect of the ε-subunit on nucleotide binding to Escherichia coli F1-ATPase catalytic sites
.
J. Biol. Chem.
274
,
19124
19128
32
Keis
,
S.
,
Stocker
,
A.
,
Dimroth
,
P.
and
Cook
,
G.M.
(
2006
)
Inhibition of ATP hydrolysis by thermoalkaliphilic F1Fo-ATP synthase is controlled by the C terminus of the ε subunit
.
J. Bacteriol.
188
,
3796
3804
33
Yagi
,
H.
,
Konno
,
H.
,
Murakami-Fuse
,
T.
,
Isu
,
A.
,
Oroguchi
,
T.
,
Akutsu
,
H.
et al.  (
2010
)
Structural and functional analysis of the intrinsic inhibitor subunit ε of F1-ATPase from photosynthetic organisms
.
Biochem. J.
425
,
85
94
34
Konno
,
H.
,
Murakami-Fuse
,
T.
,
Fujii
,
F.
,
Koyama
,
F.
,
Ueoka-Nakanishi
,
H.
,
Pack
,
C.G.
et al.  (
2006
)
The regulator of the F1 motor: inhibition of rotation of cyanobacterial F1-ATPase by the ε subunit
.
EMBO J.
25
,
4596
4604
35
Sekiya
,
M.
,
Hosokawa
,
H.
,
Nakanishi-Matsui
,
M.
,
Al-Shawi
,
M.K.
,
Nakamoto
,
R.K.
and
Futai
,
M.
(
2010
)
Single molecule behavior of inhibited and active states of Escherichia coli ATP synthase F1 rotation
.
J. Biol. Chem.
285
,
42058
42067
36
Hisabori
,
T.
,
Ueoka-Nakanishi
,
H.
,
Konno
,
H.
and
Koyama
,
F.
(
2003
)
Molecular evolution of the modulator of chloroplast ATP synthase: origin of the conformational change dependent regulation
.
FEBS Lett.
545
,
71
75
37
Werner-Grune
,
S.
,
Gunkel
,
D.
,
Schumann
,
J.
and
Strotmann
,
H.
(
1994
)
Insertion of a ‘chloroplast-like’ regulatory segment responsible for thiol modulation into γ-subunit of F0F1-ATPase of the cyanobacterium Synechocystis 6803 by mutagenesis of atpC
.
Mol. Gen. Genet.
244
,
144
150
38
Miki
,
J.
,
Maeda
,
M.
,
Mukohata
,
Y.
and
Futai
,
M.
(
1988
)
The γ-subunit of ATP synthase from spinach chloroplasts. Primary structure deduced from the cloned cDNA sequence
.
FEBS Lett.
232
,
221
226
39
Arana
,
J.L.
and
Vallejos
,
R.H.
(
1982
)
Involvement of sulfhydryl groups in the activation mechanism of the ATPase activity of chloroplast coupling factor 1
.
J. Biol. Chem.
257
,
1125
1127
PMID:
[PubMed]
40
Nalin
,
C.M.
and
McCarty
,
R.E.
(
1984
)
Role of a disulfide bond in the γ subunit in activation of the ATPase of chloroplast coupling factor 1
.
J. Biol. Chem.
259
,
7275
7280
PMID:
[PubMed]
41
Murakami
,
S.
,
Kondo
,
K.
,
Katayama
,
S.
,
Hara
,
S.
,
Sunamura
,
E.I.
,
Yamashita
,
E.
et al.  (
2018
)
Structure of the γ-ε complex of cyanobacterial F1-ATPase reveals a suppression mechanism of the γ subunit on ATP hydrolysis in phototrophs
.
Biochem. J.
475
,
2925
2939
42
Kagawa
,
Y.
and
Hamamoto
,
T.
(
1996
)
The energy transmission in ATP synthase: from the γ-c rotor to the α3β3 oligomer fixed by OSCP-b stator via the β DELSEED sequence
.
J. Bioenerg. Biomembr.
28
,
421
431
43
Ketchum
,
C.J.
,
Al-Shawi
,
M.K.
and
Nakamoto
,
R.K.
(
1998
)
Intergenic suppression of the γM23K uncoupling mutation in F0F1 ATP synthase by βGlu-381 substitutions: the role of the β380DELSEED386 segment in energy coupling
.
Biochem. J.
330
,
707
712
44
Hara
,
K.Y.
,
Noji
,
H.
,
Bald
,
D.
,
Yasuda
,
R.
,
Kinosita
, Jr,
K.
and
Yoshida
,
M.
(
2000
)
The role of the DELSEED motif of the β subunit in rotation of F1-ATPase
.
J. Biol. Chem.
275
,
14260
14263
45
Hahn
,
A.
,
Vonck
,
J.
,
Mills
,
D.J.
,
Meier
,
T.
and
Kuhlbrandt
,
W.
(
2018
)
Structure, mechanism, and regulation of the chloroplast ATP synthase
.
Science
360
,
eaat4318
46
Sunamura
,
E.
,
Konno
,
H.
,
Imashimizu-Kobayashi
,
M.
,
Sugano
,
Y.
and
Hisabori
,
T.
(
2010
)
Physiological impact of intrinsic ADP inhibition of cyanobacterial F0F1 conferred by the inherent sequence inserted into the γ subunit
.
Plant Cell Physiol.
51
,
855
865
47
Sakurai
,
H.
,
Shinohara
,
K.
,
Hisabori
,
T.
and
Shinohara
,
K.
(
1981
)
Enhancement of adenosine triphosphatase activity of purified chloroplast coupling factor 1 in aqueous organic solvent
.
J. Biochem.
90
,
95
102
48
Sunamura
,
E.
,
Konno
,
H.
,
Imashimizu
,
M.
,
Mochimaru
,
M.
and
Hisabori
,
T.
(
2012
)
A conformational change of the γ subunit indirectly regulates the activity of cyanobacterial F1-ATPase
.
J. Biol. Chem.
287
,
38695
38704
49
Heldt
,
W.H.
,
Werdan
,
K.
,
Milovancev
,
M.
and
Geller
,
G.
(
1973
)
Alkalization of the chloroplast stroma caused by light-dependent proton flux into the thylakoid space
.
Biochim. Biophys. Acta
314
,
224
241
50
Robinson
,
S.P.
(
1985
)
The involvement of stromal ATP in maintaining the pH gradient across the chloroplast envelope in the light
.
Biochim. Biophys. Acta, Bioenergetics
806
,
187
194