Ei-Ichiro Sunamura, Hiroki Konno, Mari Imashimizu-Kobayashi, Yasushi Sugano and Toru Hisabori∗
Chemical Resources Laboratory, Tokyo Institute of Technology, Nagatsuta 4259-R1-8, Midori-Ku, Yokohama, 226-8503 Japan
∗Corresponding author: E-mail, [email protected]; Fax, +81-45-924-5268
(Received April 15, 2010; Accepted April 21, 2010)
The FoF1-ATPase, which synthesizes ATP with a rotary motion,
is highly regulated in vivo in order to function efficiently,
although there remains a limited understanding of the
physiological significance of this regulation. Compared with
its bacterial and mitochondrial counterparts, the γ subunit
of cyanobacterial F1, which makes up the central shaft of
the motor enzyme, contains an additional inserted region.
Although deletion of this region results in the acceleration of
the rate of ATP hydrolysis, the functional significance of the
region has not yet been determined. By analysis of rotation,
we successfully determined that this region confers the
ability to shift frequently into an ADP inhibition state; this is
a highly conserved regulatory mechanism which prevents
ATP synthase from carrying out the reverse reaction. We
believe that the physiological significance of this increased
likelihood of shifting into the ADP inhibition state allows the
intracellular ATP levels to be maintained, which is especially
critical for photosynthetic organisms.
Keywords: ADP inhibition • ATP synthase • Cyanobacteria •
γ subunit • Regulation.
Abbreviations: F1, coupling factor 1; FCCP, carbonyl cyanide
p-(trifluoro-methoxy) phenylhydrazone; LDAO, lauryl
dimethylamine-N-oxide; PCA, pechloric acid; 1-methoxy PMS,
1-methoxy-5-methyl phenazinium methylsulfate; PMSF,
phenylmethylsulfonyl fluoride; rps, round per second; TF1, F1 of
thermophilic Bacillus PS3.
Introduction
FoF1-ATP synthase synthesizes ATP from ADP and inorganic
phosphate by being coupled with proton translocation across
the cytoplasmic membranes of bacteria, thylakoid membranes
of chloroplasts and inner membranes of mitochondria (Senior
1990, Boyer 1997, Yoshida et al. 2001). When the proton motive
force for ATP synthesis is insufficient, this enzyme potentially
Rapid Paper
Physiological Impact of Intrinsic ADP Inhibition of
Cyanobacterial FoF1 Conferred by the Inherent
Sequence Inserted into the γ Subunit
can hydrolyze ATP, and protons can be transported in the
opposite direction. The enzyme consists of the membraneembedded portion Fo and the water-soluble portion F1. Fo, the
proton translocation device, is composed of a, b and c subunits
with a stoichiometry of a1b2c10–15 (Stock et al. 1999, Seelert
et al. 2000, Jiang et al. 2001, Mitome et al. 2004, Meier et al.
2005). F1, the catalytic core for ATP synthesis and hydrolysis, is
composed of five subunits designated α–ε with a stoichiometry
of α3β3γ1δ1ε1 (Yoshida et al. 1979). The minimum composition,
which provides the basic functionality of F1-ATPase, is α3β3γ
(Kaibara et al. 1996, Du et al. 2001), and the catalytic sites
reside on each of the three β subunits at the interface with the
α subunits (Abrahams et al. 1994). The rotary catalysis mechanism was first proposed by P. D. Boyer and co-workers based on
detailed analysis of the kinetics of the ATPase reaction of F1
(Gresser et al. 1982). Following determination of the central
axis structure of the γ subunit in the α3β3 hexagon (Abrahams
et al. 1994), several groups tried to prove the rotation of the γ
subunit during ATP hydrolysis (Duncan et al. 1995, Sabbert
et al. 1996), and continuous rotation of the γ subunit coupled
with ATP hydrolysis was conclusively determined by single
molecule observation experiments (Noji et al. 1997, Hisabori
et al. 1999, Omote et al. 1999). After this, the discrete 120° step
rotation of γ per single molecule of ATP consumption and 80°
and 40° substeps within this 120° step were observed (Yasuda
et al. 1998, Yasuda et al. 2001). Recent thorough analyses provided the evidence that ADP release occurs at the 240° position
of γ rotated from the ATP binding position (Adachi et al. 2007,
Ariga et al. 2007).
Given the critical role of ATP synthesis as a key reaction
required for the maintenance of a number of metabolic pathways, the FoF1 complex must be subject to a number of
regulatory mechanisms which act optimally to accommodate
changes in environmental conditions. ADP-mediated inhibition of F1-ATPase (ADP inhibition) is a common regulatory
mechanism: the ATP hydrolysis reaction is inhibited by tight
binding of ADP-Mg to the catalytic site(s) (Minkov et al. 1979,
Plant Cell Physiol. 51(6): 855–865 (2010) doi:10.1093/pcp/pcq061, available online at www.pcp.oxfordjournals.org
© The Author 2010. Published by Oxford University Press on behalf of Japanese Society of Plant Physiologists.
All rights reserved. For permissions, please email: [email protected]
Plant Cell Physiol. 51(6): 855–865 (2010) doi:10.1093/pcp/pcq061 © The Author 2010.
855
E.-I. Sunamura et al.
Bar-Zvi and Shavit 1982, Vasilyeva et al. 1982, Zhou et al. 1988,
Digel et al. 1996, Boyer 1997). Detailed analyses of the rotational
behavior of F1 of thermophilic Bacillus PS3 (TF1) allowed this
inhibition to be assigned to the pause of rotation motion
at 80° within the 120° catalytic step (Hirono-Hara et al. 2001,
Hirono-Hara et al. 2005). The intrinsic inhibitory function of
the ε subunit (ε inhibition) is another mechanism which
regulates ATP hydrolysis, and is well characterized in bacteria
and in chloroplasts (Nelson et al. 1972, Richter et al. 1984,
Aggeler and Capaldi 1996, Kato et al. 1997, Nowak et al. 2002).
By using the single molecule observation technique, we recently
determined that ε inhibition stops rotation of the γ subunit
at a similar position to ADP-induced inhibition (Konno et al.
2006). There are several reports that neither ADP inhibition nor
ε inhibition affects ATP synthesis (Bald et al. 1998, Tsunoda
et al. 2001), and the physiological significance of these forms of
inhibition remains unknown.
Analysis of the crystal structure (Yagi et al. 2007) and
biochemical analysis (Kato-Yamada et al. 2000) have led to
the suggestion that binding of ATP to the ε subunit may result
in a conformational change of the C-terminal helices of the
TF1-ε subunit, and that mutual conformational change of
these C-terminal helices is dependent on the ATP concentration. The authors therefore concluded that the observed
conformational change reflects the dynamic attenuation of
activity against ATP concentrations in the cells, although the
evidence which shows the relevance of the ε inhibition for the
maintenance of the cellular ATP level is still missing.
In photosynthetic organisms, redox regulation of ATP
hydrolysis activity via the formation and reduction of a disulfide bond on the γ subunit is a well known regulatory mechanism (Hisabori et al. 2002). This mode of regulation has been
extensively studied in ATP synthase obtained from higher
plant chloroplasts (Mills et al. 1980, Nalin and McCarty 1984,
Samra et al. 2006), and those of Chlamydomonas reinhardtii
(Ross et al. 1995), and its role in preventing the futile ATP
hydrolysis in the dark is well understood, although there
remains limited evidence relating to the specific physiological
role of this regulatory system.
In contrast, the γ subunit of cyanobacterial thylakoid
membrane ATP synthase is much less clear, since the subunit
possesses the inserted region as in that from higher plant
chloroplasts, but lacks the nine amino acid sequence including
the two regulatory cysteines (Werner et al. 1990, Werner-Grune
et al. 1994). We recently reported that deletion of this
insertion drastically changed the ATP hydrolysis activity of
the α3β3γ complex from thermophilic cyanobacteria, and led
to a complex which is less sensitive to ε inhibition (Konno
et al. 2006).
In order to determine the significance of this inserted region
for the enzyme complex, we studied a mutant complex with
the subunit γ lacking this insertion, at both the single molecule
and whole cell levels. Our results clearly indicate that the
cyanobacterial ATP synthase equips special machinery to maintain the intracellular ATP level efficiently.
856
Results
The ATPase complex lacking the inserted region
on γ is less prone to ADP inhibition
In a previous study, we found that the α3β3γ complex of
Thermosynechoccus elongatus BP-1, which lacks the inserted
region (Leu198–Val222) on the γ subunit, showed a higher
ATP hydrolysis activity (Konno et al. 2006). To study the role of
this region in the enzyme, and to determine the cause of the
observed higher activity caused by the deletion in more detail,
a newly designed α3β3γ∆198–222 was constructed which is applicable to single molecule experiments. To this end, the corresponding sequence was deleted from the γ subunit of the
enzyme complex for rotation study, α3β3γG112C, A125C. Hereafter,
the original complex for rotation is referred to as α3β3γwild and
the complex lacking the inserted region on γ as α3β3γ∆198–222.
Following purification of these complexes which had been
expressed in Escherichia coli, the bound nucleotides on the
complexes were quantified. The α3β3γwild complex contained
1.6 ± 0.2 mol mol−1 ATP and 1.9 ± 0.1 mol mol−1 ADP, and
the α3β3γ∆198–222 complex contained 1.6 ± 0.4 mol mol−1 and
1.2 ± 0.5 mol mol−1, respectively (mean ± SD, n = 5).
The ATP hydrolysis activities of α3β3γwild and α3β3γ∆198–222
were then measured in the absence and presence of lauryl
dimethylamine-N-oxide (LDAO) (Fig. 1A). LDAO is known
to be effective in recovering F1-ATPase from ADP inhibition
(Dunn et al. 1990). In the absence of LDAO, α3β3γ∆198–222 demonstrated a ∼10-fold higher ATP hydrolysis activity compared
with α3β3γwild. The ATP hydrolysis activity of α3β3γwild was
activated about 11-fold by the addition of LDAO, while that of
α3β3γ∆198–222 was not activated. In a previous report (Konno
et al. 2006), the ATP hydrolysis activity of the deletion mutant
was activated 2.3-fold by the addition of LDAO. The apparent
discrepancy between these two studies is likely to be due to
the extent of suppression of the ATP hydrolysis activity of the
complex by ADP inhibition, probably due to the difference in
the amounts of the tightly bound ADP of these preparations.
In order to corroborate the difference in sensitivity to ADP
inhibition observed above, an additional effector of ADP inhibition, the phytotoxin tentoxin, was employed. Tentoxin is
a plant-specific toxin of fungal origin and is known to act as
a specific inhibitor for F1-ATPase in tentoxin-sensitive plant
chloroplasts (Avni et al. 1992). The effect of tentoxin on
cyanobacterial ATP synthase have been the subject of extensive
studies (Ohta et al. 1993, Meiss et al. 2008); low concentrations
(3–10 µM) of tentoxin inhibit ATP hydrolysis activity, whereas
this activity is stimulated by alleviation of ADP inhibition when
high concentrations (∼1 mM) of tentoxin are used. The ATP
hydrolysis activities of α3β3γwild and α3β3γ∆198–222 were therefore measured at various tentoxin concentrations (Fig. 1B).
Although the basal activities of these complexes were found
to be very different, both complexes were sensitive to tentoxin.
A maximum inhibition of 80% was achieved in α3β3γwild
at 5–10 µM tentoxin, while that of α3β3γ∆198–222 was ∼50%.
Plant Cell Physiol. 51(6): 855–865 (2010) doi:10.1093/pcp/pcq061 © The Author 2010.
Physiological impact of intrinsic ADP inhibition of cyanobacterial FoF1
ATP Hydrolysis Activity
(µmol Pi · min-1 · mg-1)
20
A
15
10
5
ATP Hydrolysis Activity
(µmol Pi · min-1 · mg-1)
0
LDAO
−
+
α3β3γwild
−
+
α3β3γ∆198-222
B
12
8
4
0
0
10-8
10-7 10-6 10-5
Tentoxin (M)
10-4
10-3
Fig. 1 The specific properties of the ATPase complex lacking the
inserted region on γ. (A) Effect of LDAO on ATP hydrolysis activities
of α3β3γwild and α3β3γ∆198–222. ATP hydrolysis activity was measured
using an ATP-regenerating system in the presence (black bar) and
absence (white bar) of 0.1% (w/v) LDAO at 25°C. For the assay, 2 nM
of complex was used. The ATP concentration in the assay mixture was
2 mM. The results of three independent experiments were averaged
(mean ± SD). (B) Effects of tentoxin on ATP hydrolysis activities of
α3β3γwild and α3β3γ∆198–222. ATP hydrolysis activity was measured using
an ATP-regenerating system following incubation with the indicated
concentrations of tentoxin. The line graphs of the activities of α3β3γwild
(filled circles) and α3β3γ∆198–222 (open circles) are shown. For the assay,
8.2 nM F1 was used. The activities were determined from a steady-state
slope. The samples were pre-incubated with the indicated tentoxin
concentrations at 25°C for at least 30 min, and tentoxin was also
included in the reaction buffer.
In contrast, the responses to higher concentrations of tentoxin
were very different. Although the activity of α3β3γwild increased
4-fold by addition of 1 mM tentoxin, α3β3γ∆198–222 showed
only partial relief from tentoxin inhibition at this concentration. The observed tentoxin sensitivity of α3β3γ∆198–222 was
similar to that of the tentoxin-sensitive TF1 mutant, whose
activity was not enhanced by 1 mM tentoxin (Pavlova
et al. 2004).
Rotational property of the α3β3γ∆198–222 complex
In order to determine the reason for the higher ATP hydrolysis
activity observed in α3β3γ∆198–222, the rotational behavior of
the complex was analyzed using single molecule observation
techniques. For this purpose, a polystyrene bead with a diameter
of approximately 340 nm was attached to the complex, and
rotation of the γ subunit in the complex observed for 10–20 min
by phase contrast microscopy. Fig. 2A and B shows the typical
time courses of rotation of the γ subunit in the presence of
20 µM ATP at room temperature. Initial observations showed
that the duration of rotation of γ in the α3β3γ∆198–222 complex
was longer than that in the α3β3γwild complex. This ATP concentration was too high to observe the stepwise rotation,
and we could not obtain a real rotation rate because of the
viscous friction of large beads even when we only focused
on the rotating period of each trace. However, the average
rotation rates of both complexes at the rotating period
were similar [α3β3γwild, 3.8 ± 1.1 rounds per second (rps);
and α3β3γ∆198–222, 4.5 ± 1.5 rps calculated from 100 continual
revolutions], implying that the difference in ATP hydrolysis
activity is mainly due to the difference of the pause duration
and rotation duration observed in the rotation profiles.
The pause duration was therefore analyzed (Fig. 2C, D) using
the pause length histogram, which was obtained from pauses
>1 s. Based on the binding rate constant for ATP, kon of
1.0 × 107 M−1 s−1, determined by dwell time analysis (Fig. 3),
ATP binding should take about 5 ms at 20 µM ATP. We therefore considered that pauses >1 s were not ATP binding dwell.
For curve fitting to the histograms obtained from both complexes, double exponential equations gave adequate results,
indicating that they contained at least two independent pauses.
Consequently, the results gave two independent lifetimes (τ),
the time constants of the marked events. The short lifetimes
(τsp) of α3β3γwild and α3β3γ∆198–222 were 8.6 and 8.2 s, respectively. These short-lived pause data are, however, not reliable
enough since the bin width of these histograms was 10 s. From
the analysis of the relevance between ADP inhibition and rotation of TF1 ATPase, Hirono-Hara et al. (2001) reported similar
results, though the short lifetimes (τsp) observed in this study
are marginally longer. As they reported, the origin of these short
pauses is not yet known. In contrast, the long pauses with longer
lifetimes (τlp) of α3β3γwild and α3β3γ∆198–222 were 124 and 62 s,
respectively (Fig. 2C, D). These long pauses can be considered
as the ADP-inhibited state, which occurs at the 80° position
(Hirono-Hara et al. 2001), when the ATP binding position was
set as 0° as in many previous reports (Yasuda et al. 1998, Yasuda
et al. 2001). The long pause position during rotation was then
determined, as shown in Fig. 4, by using a medium exchange
method from 20 µM to 200 nM ATP, enabling the determination of the ATP binding position. Consequently, the pause
position was determined as 73 ± 18° (n = 7) from four molecules,
quite close to the position for ADP inhibition. From these pause
duration analyses, the long lifetime of α3β3γ∆198–222 was found
to be half of that of α3β3γwild, suggesting that the mutant can
easily evade ADP inhibition.
Next, rotation durations which appeared between two
pauses were collected and analyzed. As shown in Fig. 2E
and F, the histograms obtained were clearly fitted to a single
exponential equation. The lifetime (τr) for α3β3γwild and that
for α3β3γ∆198–222 were 19 and 45 s, respectively. As these
Plant Cell Physiol. 51(6): 855–865 (2010) doi:10.1093/pcp/pcq061 © The Author 2010.
857
E.-I. Sunamura et al.
C
A
E
N = 157
B
τsp = 8.6 s
τlp = 124 s
D
N = 230
τsp = 8.2 s
τlp = 62 s
Time (s)
Pause Duration (s)
Number of Events
Number of Events
Revolutions
N = 174
τr = 19 s
F
N = 186
τr = 45 s
Rotation Duration (s)
Fig. 2 Rotation of the γ subunit in α3β3γwild and α3β3γ∆198–222. (A, B) The typical time courses of the rotation of α3β3γwild (A) and of α3β3γ∆198–222
(B) are shown. Many particles with duplex 340 nm diameter beads were recorded on video tapes and the rotations were analyzed using custom
software (Yasuda et al. 1998). Rotation observation was performed at 25°C in the presence of 20 µM ATP using an ATP-regenerating system.
(C, D) The histograms of pause duration are shown. Pauses >1 s were collected and analyzed. The histograms of α3β3γwild (C) and α3β3γ∆198–222 (D)
were constructed from data for 36 and 47 molecules, respectively. In the insets, the vertical axis is drawn with a finer scale, and the bars
for the data less than 20 s in C and those less than 30 s in D was omitted. The histograms are fitted with a double exponential equation,
Nsp × exp (−t/τsp) + Nlp × exp (−t/τlp). (E, F) The histograms of rotation duration between one pause (>1 s) and the next pause (>1 s) are shown.
The histograms of α3β3γwild (E) and α3β3γ∆198–222 (F) were constructed from data for 36 and 47 molecules, respectively. The histograms are fitted
with a single exponential equation, Nr × exp (−t/τr).
lifetimes (τr) obtained by the analysis would show the transition tendencies from active state to ADP-inhibited state,
we conclude that the α3β3γ∆198–222 complex is less prone to
drop into ADP inhibition and has the tendency to rotate
continuously.
Physiological significance of the inserted region on
the γ subunit of ATP synthase in cyanobacteria
Our major interest in the inserted region on the γ subunit of
cyanobacterial ATP synthase is the following: does the ability
to drop into ADP inhibition conferred by this region actually
have a physiological significance for cell viability or some other
cell metabolic process? In order to answer this question,
cyanobacterial cells lacking the inserted region on the γ subunit
were investigated under various physiological conditions. To
this end, we used Synechocystis sp. PCC6803, which has
a high efficiency of natural transformation, and transformants
are generated by simply supplying the foreign DNA to the
growth medium. The desired transformants were successfully
obtained by integration of a kanamycin resistance cassette
(KmR), and the inserted region of γ (Leu198–Val222) was
deleted by cultivation of the transformant cells in the presence
of kanamycin (for details, see the Materials and Methods).
Hereafter the transformant containing this deletion at γ is
referred to as the γ∆198–222 mutant. To examine the effect of
the introduced KmR itself on cell growth, a transformant containing only the KmR (wild + KmR) was used as a control.
The growth rates of the wild-type cells and the γ∆198–222
mutant cells were measured under continuous light
(30 µE m−2 s−1) conditions (Fig. 5A). Since no differences in
858
growth rates between the wild type and the mutant cells
were observed, 8 h light–16 h dark (8L/16D) cycle conditions
were then examined (Fig. 5B). In the dark, the growth of both
types of cells slowed down and immediately recovered in the
light. Under these conditions, again no remarkable differences
between the wild-type cells and the mutant cells were observed.
In addition, growth rates of these cells on an agar plate
under continuous dark conditions for 4 days were compared
(Supplementary Fig. S4). Even under such conditions that are
unfavorable for photosynthetic organisms, the γ∆198–222 mutant
and wild-type cells could survive. We therefore concluded that
ADP inhibition of the ATP synthase of cyanobacterial cells is
not critical to confer adaptability to light/dark conditions.
In order to explore further the potential physiological
relevance of the ADP inhibition of ATP hydrolysis activity of
ATP synthase, the dynamics of the cellular ATP level in intact
cells under light and dark conditions were measured (Fig. 6).
The cell suspension was directly denatured by addition of perchloic acid (PCA), and the amounts of ATP quantified by
a luciferin–luciferase assay after neutralization. Prior to conducting the experiments, cells were illuminated under normal
light conditions (30 µE m−2 s−1) for 20 min to build a sufficient
proton gradient for ATP synthesis. In the light, the ATP level of
the mutant cells was 70–80% of that of the wild-type cells.
When the light was turned off, the cellular ATP level decreased
sharply in both the wild-type and mutant cells. However, the
difference in the overall decrease of ATP was found to be
remarkable: amounts of ATP in the wild-type cells decreased to
50% of the original level whereas those in the γ∆198–222 mutant
cells decreased by ≤20%. When the cells were kept in the dark,
Plant Cell Physiol. 51(6): 855–865 (2010) doi:10.1093/pcp/pcq061 © The Author 2010.
P1
P1
Angle (deg)
P2
Number of events
Revolutions
Number of events
Physiological impact of intrinsic ADP inhibition of cyanobacterial FoF1
P2
Angle (deg)
Time (s)
Fig. 4 Stop angular position of long pauses (>10 s). After observation
for 10 min at 20 µM ATP, the solution was exchanged for 200 nM ATP
solution (arrows). The insets represent the histograms of angular
distribution of the beads. Black bars represent the angular distribution
from data at 200 nM ATP, and blue bars represent the angular
distribution of long pauses (indicated as ‘P1’ and ‘P2’) from the data
before buffer exchange.
Fig. 3 Rotation of the γ subunit in α3β3γwild and α3β3γ∆198–222 at an ATP
concentration of 200 nM. (A) Typical time courses of α3β3γwild (red)
and α3β3γ∆198–222 (blue) are shown. Insets represent the traces of
centroid of the beads. (B) The dwell time periods during rotation
observed in the trace shown in A were collected and the histograms
were prepared for α3β3γwild (red) and α3β3γ∆198–222 (blue). Each
histogram is constructed from four molecules and fitted with a single
exponential equation, Const. × exp (−kt).
the cellular ATP levels gradually recovered. Eventually, those
in the wild-type cells reached 80% of the original level within
15 min, whereas those in the γ∆198–222 mutant cells only recovered to 30–40%. When the cells were kept in the light again,
the ATP levels of both cells recovered to the original levels.
In order to determine whether the observed difference
in cellular ATP levels is attributable to the difference in ATP
synthesis activity of the thylakoid membranes, the ATP synthesis activities of the membranes of the wild-type and mutant
cells were measured using spheroplasts. Fig. 7 shows the time
courses of ATP synthesis by disrupted spheroplasts of the
wild-type and mutant cells. Compared with the earlier study
(Scholts et al. 1996), the observed activities were lower, probably due to the lower efficiency of lysozyme digestion used for
preparation. However, the efficiency of the digestion of the
mutant cells was not significantly different compared with
that of wild-type cells, when the efficiency of digestion was
evaluated by the amounts of phycocyanin released by this
treatment. The ATP synthesis activity was determined from
the linear portion of the measurement shown in Fig. 7, and
summarized in Table 1. These data suggest that the thylakoid
membranes of the mutant cells can catalyze light-driven ATP
synthesis at almost the same rate as the wild-type membranes.
Discussion
Phylogenetic analysis of the γ subunit clearly shows coevolution of the redox-sensitive chloroplast γ subunit with that
of cyanobacteria, though these two groups have a distinct
diversion point (Supplementary Fig. S1) as the cyanobacterial
γ subunit lacks a nine amino acid sequence containing two
regulatory cysteines (Supplementary Fig. S2). Since the phylogenetic distances of these two groups from other γ subunits of,
for example, E. coli or that of mitochondria are largely equivalent, it is difficult to provide a conclusive explanation as to the
Plant Cell Physiol. 51(6): 855–865 (2010) doi:10.1093/pcp/pcq061 © The Author 2010.
859
E.-I. Sunamura et al.
L
Abs730
Synthesized ATP
(µmoles · mg Chl-1)
A
Time (min)
Time (h)
Fig. 7 ATP synthesis of the spheroplasts obtained from wild-type and
γ∆198–222 mutant cells. Light-triggered ATP synthesis was measured
using the disrupted spheroplasts obtained from lysozyme-treated cells
(for details, see the Materials and Methods). Time courses of ATP
synthesis with wild-type (filled symbols) and mutant (open symbols)
cells are shown. Spheroplasts were illuminated with a light at 100
(squares) or 300 µE m−2 s−1 (circles). At 300 µE m−2 s−1, the effect of
carbonyl cyanide p-(trifluoro-methoxy) phenylhydrazone (FCCP) was
examined (triangles).
Abs730
B
Table 1 ATP synthesis activities of the spheroplasts of wild-type and
γ∆198–222 mutant cells
Time (h)
Spheroplasts
Amounts of Intracellular ATP
(nmoles · mg Chl-1)
Fig. 5 Growth of Synechocystis cells under continuous light and light/
dark conditions. (A) The growth rates of wild-type (filled circles),
γ∆198–222 mutant (open triangles) and wild-type + KmR (open squares)
cells were compared under continuous light conditions (30 µE m−2 s−1)
at 30°C. Two independent experiments were performed. (B) The
growth rates of wild-type (filled circles), γ∆198–222 mutant (open
triangles) and wild-type + KmR (open squares) cells were compared
under light/dark (8/16 h) conditions (30 µE m−2 s−1) at 30°C. Three
independent experiments were performed.
L
D
L
Time (min)
Fig. 6 Level of ATP in Synechocystis cells. The level of ATP in intact cells
of the wild-type (filled circles) and the γ∆198–222 mutant (open circles)
were compared. Late log phase cultures were put in an Erlenmeyer
flask and kept under light and dark conditions at 30°C. At the times
indicated, aliquots of 50 µl of the cell culture were taken and added to
10 µl of 12% (w/v) PCA. After neutralization, ATP in the supernatant
was quantified using a luciferin–luciferase assay. The results of three
independent experiments were averaged.
860
ATP synthesis [µmol ATP min−1 (mg Chl)−1]
100 µE m−2 s−1
300 µE m−2 s−1
Wild type
0.19 ± 0.03
0.66 ± 0.05
γ∆198–222 mutant
0.19 ± 0.04
0.61 ± 0.09
ATP synthesis activity was determined using linear regressions by comparing ATP
synthesis in the absence of FCCP with that in the presence of FCCP. The results of
three different membrane preparations were averaged (mean ± SD).
evolutionary origin of the γ subunit containing the inserted
region. It is likely, however, that a particular γ subunit type must
have evolved in photosynthetic organisms to confer the necessary regulation of their activities under light–dark conditions
when solar energy was harnessed for ATP synthesis.
In order to determine the origin of this inserted sequence on
the cyanobacterial γ subunit, 198LETADDEIFRLTTRGSHLEVN
REK222V, a BLAST search was carried out using the sequence of
the inserted region of the T. elongatus BP-1 (Supplementary
Fig. S2). However, only sequences showing similarity within
the γ subunit of ATP synthase of other origin were obtained,
suggesting that the inserted sequence originally evolved in the
γ subunit itself. Since no crystal or solution structures of the
γ subunit of cyanobacterial and chloroplast ATP synthase have
been reported to date, the interaction of this inserted region
with other parts of the complex remains unknown.
Since, as mentioned previously, the α3β3γ∆198–222 deletion
mutant complex lacked one bound ADP after purification,
the affinity for ADP at a binding site of this complex must be
lower than that of the α3β3γwild complex. This difference in
Plant Cell Physiol. 51(6): 855–865 (2010) doi:10.1093/pcp/pcq061 © The Author 2010.
Physiological impact of intrinsic ADP inhibition of cyanobacterial FoF1
affinity for ADP must be reflected in the extent of ADP inhibition, and must cause the observed difference in ATP hydrolysis
activity in the absence and presence of the detergent or tentoxin (see Fig. 1A and B), though the reason why the change of
the γ subunit affects the ADP binding affinity remains unknown.
Consistent with our results, an increase in the ATP hydrolysis
activity by the deletion of the entire inserted region of the
γ subunit of the chloroplast ATP synthase has already been
reported by Samra et al. (2006).
Observation of single molecule behavior of F1-ATPase
has now become a powerful tool to reveal a change in the
molecular motion caused by small phenomena occurring at
a catalytic site or a non-catalytic site of this large enzyme complex. Accordingly, a difference in rotational motion between
α3β3γwild and α3β3γ∆198–222 was observed (Fig. 2A, B). From the
analysis of the duration (Figs. 2C–F), we concluded that the
lower ATP hydrolysis activity of the α3β3γwild complex was due
to the frequent shift into the ADP inhibition state, and the
difficulties of recovery from this inhibition. In contrast, the
pause duration of the mutant complex was shorter than that
of the wild-type complex, and the rotation duration of the
mutant complex was longer than that of the wild type, implying that the mutant complex is less susceptible to the ADP
inhibition state.
For photosynthetic organisms, ATP formation in the light is
the first and most important energy conversion process.
Regulation of ATP synthase is therefore critical to drive the
whole photosynthetic system efficiently. Since inhibition of the
reverse reaction, ATP hydrolysis, appears to be the simplest
regulatory system for this enzyme, the mutant strain lacking
the inserted region in the γ subunit was prepared, and it was
expected that this would possess a higher ATP hydrolysis
activity in vivo. As shown in Fig. 5, however, the mutant strain
did not show any remarkable differences in cell growth rate
under normal light conditions and 8L/16D conditions.
The change in the amount of ATP in the cells was therefore
examined when the cells were transferred from light to dark
conditions (Fig. 6). In the light, the amount of ATP in the
mutant cells was slightly lower than that observed in wild-type
cells. This difference was not due to the difference in the
amounts of the ATP synthase in thylakoid membranes, since
similar amounts of membrane-localized β subunits were
detected in both cells (Supplementary Fig. S5). In addition,
when the ATP synthesis activities of the spheroplasts obtained
from wild-type and mutant cells were measured, they were
largely equivalent (Fig. 7 and Table 1). Hence, we concluded
that the lower ATP level of the mutant cells in the light is due
to the higher ATP hydrolysis activity of the ATP synthase complex in the mutant cells, since the ATP level must be determined
by the balance between ATP synthesis and hydrolysis reactions.
By turning off the light, the amount of ATP in wild-type cells
immediately dropped to half of the original level, and that in
the mutant cells reached 20%, suggesting that the intracellular
ATP was vigorously hydrolyzed by the mutant ATP synthase.
Thereafter it recovered to 70–80% of the original level within
15 min in the wild-type cells, although only 30–40% recovery
was observed in the mutant cells. In cyanobacteria, ATP synthesis is promoted by photophosphorylation, oxidative phosphorylation, glycolysis and transphosphorylation reactions, and
ATP hydrolysis by the reverse reaction of ATP synthase
and other ATP-hydrolyzing enzymes. When cells were transferred from light to dark conditions, the cellular ATP level
decreased suddenly and then recovered gradually. One of the
possibilities for this gradual recovery is the slow down of the
ATP hydrolysis reaction induced by ADP inhibition of ATP
synthase, although oxidative phosphorylation is constantly
active irrespective of light/dark conditions. This may explain
why the recovery level was lower in the case of the mutant cells
(see Fig. 6), as the ATP synthase in the mutant cells must
rarely have shifted into the ADP inhibition state. The major part
of the recovery was attributed to the oxidative phosphorylation (Bottomley and Stewart 1976) as mentioned, since the
recovery of ATP in the dark was almost diminished by addition
of KCN to the culture, which can block oxidative phosphorylation (Supplementary Fig. S6).
Werner-Grune et al. (1994) previously introduced the
regulatory region of the γ subunit of spinach chloroplast ATP
synthase into the F1-ATPase in Synechocystis cells to confer
a redox regulation property to this enzyme. However, this
mutation did not confer any advantage in terms of cell viability
under photosynthetic conditions. In their case, the reason that
the mutant did not show any specific characteristics is likely to
have been that the cyanobacterial cells were already compensated by the inserted region for the photosynthetic conditions,
by using strong ADP inhibition as shown in our study.
In the case of eukaryotic cells including photosynthetic
plants, the intracellular AMP/ATP ratio is known to be an
important signal which controls various metabolic pathways
(Hoppe et al. 2009). In contrast, catabolic pathways are more
affected than anabolic pathways (growth rates) when the
intracellular ADP/ATP ratio is controlled by the expression of
ATPase in E. coli (Koebmann et al. 2002). Although we could
not find any physiological disadvantage in mutant cells caused
by a lower intracellular ATP level in the dark, our study is the
first report to show successfully the physiological significance of
the regulatory system of ATP synthase under photosynthetic
conditions in vivo.
Materials and Methods
Materials
Biotin-PEAC5-maleimide and 1-methoxy-5-methyl phenazinium
methylsulfate (1-methoxy PMS) were purchased from Dojindo
(Kumamoto, Japan). Tentoxin, ATP, ADP, diadenosine pentaphosphate, phosphoenolpyruvate and bovine serum albumin
(BSA) were obtained from Sigma (St Louis, MO, USA). Pyruvate
kinase, lactate dehydrogenase and NADH were purchased from
Roche Diagnostics (Basel, Switzerland). Other chemicals were
of the highest grade commercially available.
Plant Cell Physiol. 51(6): 855–865 (2010) doi:10.1093/pcp/pcq061 © The Author 2010.
861
E.-I. Sunamura et al.
Strains
Escherichia coli strains used were DH5α for cloning and BL21
(DE3) unc∆702 [Tcr, ATPase mutant, BL21 (DE3) unc∆702,
asnA::Tn10] (Joshi et al. 1989, Nichols and Harwood 1997)
for expression of the T. elongatus α3β3γ complex. The latter
strain was a kind gift from Dr. Harwood (University of Iowa).
The glucose-tolerant strain of the unicellular cyanobacterium
Synechocystis sp. PCC6803 (Williams 1988) was used for analysis
of the γ∆198–222 mutant strain, which lacks the inserted region
on the γ subunit of F1.
Expression plasmid and protein preparation
An expression plasmid pTR19FRs, which was originally constructed for single molecule experiments with the α3β3γ complex of T. elongatus BP-1 in a previous study (Konno et al. 2006),
was used for expression of α3β3γG112C, A125C, and the complex
obtained was referred to as α3β3γwild in this article. For deletion
of the inserted region on the γ subunit corresponding to amino
acids from Leu198 to Val222, a 409 bp DNA fragment containing the deletion region was excised with SacI and NheI from the
coding region of the γ subunit and ligated into plasmid
pTR19FRs. The plasmid obtained was named pTR19FRsd and
used for expression of α3β3γ∆198–222, which lacks the inserted
region on the γ subunit. Expression and purification of the
complexes were performed as described (Konno et al. 2006).
The obtained complexes were labeled with biotin-PEAC5maleimide for the single molecule experiments as described
(Konno et al. 2006).
Assay of ATP hydrolysis activity
ATP hydrolysis activity was measured as described (Meiss et al.
2008). The reaction was initiated by adding 10 µl of the enzyme
solution into the assay solution (1.2 ml). To investigate the effect
of LDAO on the ATP hydrolysis activity, LDAO [final concentration of 0.1% (w/v)] was added to the assay mixture before addition of the enzyme. For the tentoxin assay, the enzyme, which
was incubated with tentoxin in advance, was added into the
assay mixture. Tentoxin pre-treatment was performed for at least
30 min with the tentoxin concentrations indicated.
Rotation assay
The rotation assay was performed as described (Meiss et al.
2008) with some modifications. Streptavidin-coated beads
with a diameter of 340 nm were used. Rotation was initiated by
addition of 60 µl of assay buffer (50 mM HEPES-KOH, pH 8.0,
100 mM KCl, 0.5 mM MgCl2, 20 µM ATP, 100 µg ml−1 pyruvate
kinase and 2 mM phosphoenolpyruvate) and the movement of
the beads observed using phase contrast microscopy and
video-recorded for further analysis.
Deletion of the inserted region sequence from the
atpC gene (sll1327) in Synechocystis sp. PCC6803
Deletion of the sequence for the inserted region from the
atpC gene (sll1327) was performed using the homologous
862
recombination technique with the Tn5-derived kanamycin
resistance cassette. For this purpose, the DNA fragment of
879 bp containing the downstream region of atpC and slr1411
was amplified by PCR using primers, atpC_F1 and slr1411_
BamHI_R, which contain BamHI sites (the sequences of the
primers used are shown in Supplementary Table S1). The PCR
product was cloned into pGEM-T Easy vector (Promega,
Madison, WI, USA) and then a Tn5-derived kanamycin resistance cassette (KmR) was inserted into the EcoO109I site
according to the manufacturer’s instructions. The DNA fragment of 711 bp containing the upstream region of the atpC
was amplified by PCR using primers atpC_F2 and atpC_R1, and
was cloned into pGEM-T Easy vector. For deletion of the
sequence for the inserted region from atpC, two primers,
atpC_del_F and atpC_del_R, which anneal to the sequence
on both sides of the inserted seqence of atpC, were designed.
After PCR amplification, the DNA fragment obtained was selfligated and then sequenced (Applied Biosystems, Foster City,
CA, USA) to confirm the deletion. The resulting plasmids were
digested with BamHI, and the 2,105 bp BamHI fragment containing the downstream region of atpC, KmR and slr1411 was
ligated into the atpC on the other plasmid.
A diagram of the resulting plasmid is shown in Supplementary Fig. S3. The plasmids were then transformed into
Synechocystis and selected on a BG11 plate containing 20 µg ml−1
kanamycin. Complete segregation and deletion mutation
were confirmed by PCR of total transformant DNA with
a primer set atpC_F1 and slr1411_BamHI_R, and atpC_check_F
and atpC_check_R, respectively (Supplementary Table S1).
Growth conditions of Synechocystis cells
For the liquid culture, Synechocystis cells were grown in
liquid BG11 medium (Stanier et al. 1971) containing 10 mM
HEPES-KOH (pH 7.4) bubbled with air containing 1% (v/v) CO2
at 30°C under continuous light. Alternatively, cells were
grown on 1.5% (w/v) agar (Agar BA-10, High Quality, INA,
Japan) plates containing BG11 and 0.3% (w/v) sodium
thiosulfate.
Quantification of intracellular ATP
The amount of ATP in intact cells was determined according
to Ohta et al. (1993). A 40 ml aliquot of cultures at an absorbance of 1.6–2.0 at 730 nm was placed in an Erlenmeyer flask
and kept under light (30 µE m−2 s−1) at 30°C with shaking
(96 min−1). After 30 min illumination, the light was turned off
and the cultures were incubated in the dark for 32 min. The
sample was then returned to the light. At the time indicated in
Fig. 6, 50 µl aliquots of cell culture were withdrawn and added
to 10 µl of 12% (w/v) PCA. A 50 µl aliquot of the supernatant
was neutralized with 125 µl of 2 M Tris-acetate (pH 7.7), and
ATP was quantified by a luciferin–luciferase assay using the
ATP bioluminescence assay kit CLS II (Roche Diagnostics,
Basel, Switzerland). The concentration of Chl was determined
according to Grimme and Boardman (1972).
Plant Cell Physiol. 51(6): 855–865 (2010) doi:10.1093/pcp/pcq061 © The Author 2010.
Physiological impact of intrinsic ADP inhibition of cyanobacterial FoF1
Preparation of spheroplasts
Spheroplasts were prepared according to Scholts et al. (1996)
with some modifications. Cells at an absorbance of 1.1–1.4 at
730 nm were harvested and incubated in 20 ml of mannitol
medium [500 mM mannitol, 10 mM Tricine-KOH, pH 7.8,
10 mM MgCl2, 5 mM NaH2PO4, 2.5 mM K2HPO4, 100 µM
phenylmethylsulfonyl fluoride (PMSF) and 1 mM 6-amino-ncaproic acid] with 0.2% lysozyme for 1.5 h at 35°C. The efficiency of lysozyme digestion was evaluated by phycocyanin
release upon osmotic shock caused by the addition of distilled
water. After centrifugation (3,000 × g for 5 min at 4°C), the
spheroplasts were suspended in mannitol medium at a Chl
concentration of 0.2–0.3 mg ml−1, and stored at 0°C.
Assay of ATP synthesis
For the assay, the spheroplast preparation was treated with
osmotic shock by the addition of reaction medium (10 mM
Tricine-NaOH, pH 8.0, 10 mM NaCl, 5 mM MgCl2, 50 µM
1-methoxy PMS, 2 mM Na-phosphate, 50 µM diadenosine penta
phosphate and 100 µM PMSF). Spheroplasts containing
10–15 µg of Chl were incubated in the reaction medium for
1 min, and the solution was pre-illuminated for 2 min at a light
intensity of 100 or 300 µE m−2 s−1. The assay was then initiated
by adding final concentrations of 2 mM ADP at 30°C. This ADP
was treated with hexokinase and 2-deoxy-D-glucose to reduce
the ATP content in advance. At the times indicated in Fig. 7,
50 µl of the reaction medium was taken out and mixed with
10 µl of 12% (w/v) PCA. After centrifugation, the supernatants
were diluted 10-fold, and 50 µl of the solution neutralized by
adding 125 µl of 1 M Tris-acetate (pH 7.5). The amount of ATP
in the neutralized solution was then quantified by a luciferin–
luciferase assay. Sensitivity of ATP synthesis to an uncoupler,
FCCP (50 µM) was also examined.
Supplementary data
Supplementary data are available at PCP online.
Funding
This work was supported by the Ministry of Education, Culture,
Sports, Science and Technology of Japan [Grants-in-Aid for
Scientific Research on Priority Areas (No. 18074002 to T. H.)].
Acknowledgments
We thank T. Murakami-Fuse and Y. S. Kim for technical
assistance, and M. Yoshida and M. Tsumuraya for discussions.
References
Abrahams, J., Leslie, A., Lutter, R. and Walker, J.E. (1994) Structure at
2.8 Å resolution of F1-ATPase from bovine heart mitochondria.
Nature 370: 621–628.
Adachi, K., Oiwa, K., Nishizaka, T., Furuike, S., Noji, H., Itoh, H., et al.
(2007) Coupling of rotation and catalysis in F1-ATPase revealed by
single-molecule imaging and manipulation. Cell 130: 309–321.
Aggeler, R. and Capaldi, R.A. (1996) Nucleotide-dependent movement
of the ε subunit between α and β subunits in the Escherichia coli
F1F0-type ATPase. J. Biol. Chem. 271: 13888–13891.
Ariga, T., Muneyuki, E. and Yoshida, M. (2007) F1-ATPase rotates by an
asymmetric, sequential mechanism using all three catalytic subunits.
Nat. Struct. Mol. Biol. 14: 841–846.
Avni, A., Anderson, J.D., Holland, N., Rochaix, J.D., Gromet-Elhanan, Z.
and Edelman, M. (1992) Tentoxin sensitivity of chloroplasts
determined by codon 83 of β subunit of proton-ATPase. Science
257: 1245–1247.
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.
Bar-Zvi, D. and Shavit, N. (1982) Modulation of the chloroplast ATPase
by tight ADP binding. Effect of uncouplers and ATP. J. Bioenerg.
Biomembr. 14: 467–478.
Bottomley, P.J. and Stewart, W.D. (1976) ATP pools and transients in the
blue-green alga, Anabaena cylindrica. Arch Microbiol. 108: 249–258.
Boyer, P.D. (1997) The ATP synthase—a splendid molecular machine.
Annu. Rev. Biochem. 66: 717–749.
Digel, J.G., Kishinevsky, A., Ong, A.M. and McCarty, R.E. (1996)
Differences between two tight ADP binding sites of the chloroplast
coupling factor 1 and their effects on ATPase activity. J. Biol. Chem.
271: 19976–19982.
Du, Z., Tucker, W.C., Richter, M.L. and Gromet-Elhanan, Z. (2001)
Assembled F1-αβ and hybrid F1-α3β3γ-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.
Duncan, T.M., Bulygin, V.V., Zhou, Y., Hutcheon, M.L. and Cross, R.L.
(1995) Rotation of subunits during catalysis by Escherichia coli
F1-ATPase. Proc. Natl Acad. Sci. USA 92: 10964–10968.
Dunn, S.D., Tozer, R.G. and Zadorozny, V.D. (1990) Activation of
Escherichia coli F1-ATPase by lauryldimethylamine oxide and
ethylene glycol: relationship of ATPase activity to the interaction of
the ε and β subunits. Biochemistry 29: 4335–4340.
Gresser, M.J., Myers, J.A. and Boyer, P.D. (1982) Catalytic site
cooperativity of beef heart mitochondrial F1 adenosine
triphosphatase. Correlations of initial velocity, bound intermediate,
and oxygen exchange measurements with an alternating three-site
model. J. Biol. Chem. 257: 12030–12038.
Grimme, L.H. and Boardman, N.K. (1972) Photochemical activities of
a particle fraction P1 obtained from the green alga Chlorella fusca.
Biochem. Biophys. Res. Commun. 49: 1617–1623.
Hirono-Hara, Y., Ishizuka, K., Kinosita, K., Jr., Yoshida, M. and Noji, H.
(2005) Activation of pausing F1 motor by external force. Proc. Natl
Acad. Sci. USA 102: 4288–4293.
Hirono-Hara, Y., Noji, H., Nishiura, M., Muneyuki, E., Hara, K.Y., Yasuda, R.,
et al. (2001) Pause and rotation of F1-ATPase during catalysis. Proc.
Natl Acad. Sci. USA 98: 13649–13654.
Hisabori, T., Kondoh, A. and Yoshida, M. (1999) The γ subunit in
chloroplast F1-ATPase can rotate in a unidirectional and counterclockwise manner. FEBS Lett. 463: 35–38.
Hisabori, T., Konno, H., Ichimura, H., Strotmann, H. and Bald, D. (2002)
Molecular devices of chloroplast F1-ATP synthase for the regulation.
Biochim. Biophys. Acta 1555: 140–146.
Plant Cell Physiol. 51(6): 855–865 (2010) doi:10.1093/pcp/pcq061 © The Author 2010.
863
E.-I. Sunamura et al.
Hoppe, S., Bierhoff, H., Cado, I., Weber, A., Tiebe, M., Grummt, I., et al.
(2009) AMP-activated protein kinase adapts rRNA synthesis to
cellular energy supply. Proc. Natl Acad. Sci. USA 106: 17781–17786.
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. USA 98: 4966–4971.
Joshi, A.K., Ahmed, S. and Ferro-Luzzi Ames, G. (1989) Energy coupling
in bacterial periplasmic transport systems. Studies in intact
Escherichia coli cells. J. Biol. Chem. 264: 2126–2133.
Kaibara, C., Matsui, T., Hisabori, T. and Yoshida, M. (1996) Structural
asymmetry of F1-ATPase caused by the γ subunit generates a high
affinity nucleotide binding site. J. Biol. Chem. 271: 2433–2438.
Kato-Yamada, Y., Yoshida, M. and Hisabori, T. (2000) Movement of the
helical domain of the ε subunit is required for the activation of
thermophilic F1-ATPase. J. Biol. Chem. 275: 35746–35750.
Kato, Y., Matsui, T., Tanaka, N., Muneyuki, E., Hisabori, T. and
Yoshida, M. (1997) Thermophilic F1-ATPase is activated without
dissociation of an endogenous inhibitor, ε subunit. J. Biol. Chem. 272:
24906–24912.
Koebmann, B.J., Westerhoff, H.V., Snoep, J.L., Nilsson, D. and Jensen, P.R.
(2002) The glycolytic flux in Escherichia coli is controlled by the
demand for ATP. J. Bacteriol. 184: 3909–3916.
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.
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.
Meiss, E., Konno, H., Groth, G. and Hisabori, T. (2008) Molecular
processes of inhibition and stimulation of ATP synthase caused by
the phytotoxin tentoxin. J. Biol. Chem. 283: 24594–24599.
Mills, J.D., Mitchell, P. and Schurmann, P. (1980) Modulation of
coupling factor ATPase activity in intact chloroplasts, the role of the
thioredoxin system. FEBS Lett. 112: 173–177.
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.
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. USA
101: 12159–12164.
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.
Nelson, N., Nelson, H. and Racker, E. (1972) Partial resolution of
the enzymes catalyzing photophosphorylation. XII. Purification
and properties of an inhibitor isolated from chloroplast coupling
factor 1. J. Biol. Chem. 247: 7657–7662.
Nichols, N.N. and Harwood, C.S. (1997) PcaK, a high-affinity permease for
the aromatic compounds 4-hydroxybenzoate and protocatechuate
from Pseudomonas putida. J. Bacteriol. 179: 5056–5061.
Noji, H., Yasuda, R., Yoshida, M. and Kinosita, K., Jr. (1997) Direct
observation of the rotation of F1-ATPase. Nature 386: 299–302.
Nowak, K.F., Tabidze, V. and McCarty, R.E. (2002) The C-terminal
domain of the ε subunit of the chloroplast ATP synthase is not
required for ATP synthesis. Biochemistry 41: 15130–15134.
Ohta, Y., Yoshioka, T., Mochimaru, M., Hisabori, T. and Sakurai, H.
(1993) Tentoxin inhibits both photophosphorylation in thylakoids
864
and the ATPase activity of isolated coupling factor F1 from the
cyanobacterium Anacystis nidulans. Plant Cell Physiol. 34: 523–529.
Omote, H., Sambonmatsu, N., Saito, K., Sambongi, Y., Iwamoto-Kihara, A.,
Yanagida, et al. (1999) The γ-subunit rotation and torque generation
in F1-ATPase from wild-type or uncoupled mutant Escherichia coli.
Proc. Natl Acad. Sci. USA 96: 7780–7784.
Pavlova, P., Shimabukuro, K., Hisabori, T., Groth, G., Lill, H. and Bald, D.
(2004) Complete inhibition and partial re-activation of single
F1-ATPase molecules by tentoxin: new properties of the re-activated
enzyme. J. Biol. Chem. 279: 9685–9688.
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.
Ross, S.A., Zhang, M.X. and Selman, B.R. (1995) Role of the
Chlamydomonas reinhardtii coupling factor 1 γ-subunit cysteine bridge
in the regulation of ATP synthase. J. Biol. Chem. 270: 9813–9818.
Sabbert, D., Engelbrecht, S. and Junge, W. (1996) Intersubunit rotation
in active F-ATPase. Nature 381: 623–625.
Samra, H.S., Gao, F., He, F., Hoang, E., Chen, Z., Gegenheimer, P.A., et al.
(2006) Structural analysis of the regulatory dithiol-containing
domain of the chloroplast ATP synthase γ subunit. J. Biol. Chem. 281:
31041–31049.
Scholts, M.J.C., Aardewijn, P. and Walraven, H.S.V. (1996) Membrane
vesicles from Synechocystis 6803 showing proton and electron
transport and high ATP synthase activities. Photosynth. Res. 47:
301–305.
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.
Senior, A.E. (1990) The proton-translocating ATPase of Escherichia coli.
Annu. Rev. Biophys. Biophys. Chem. 19: 7–41.
Stanier, R.Y., Kunisawa, R., Mandel, M. and Cohen-Bazire, G. (1971)
Purification and properties of unicellular blue-green algae (order
Chroococcales). Bacteriol. Rev. 35: 171–205.
Stock, D., Leslie, A.G. and Walker, J.E. (1999) Molecular architecture of
the rotary motor in ATP synthase. Science 286: 1700–1705.
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. USA 98:
6560–6564.
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.
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.
Werner, S., Schumann, J. and Strotmann, H. (1990) The primary
structure of the γ-subunit of the ATPase from Synechocystis 6803.
FEBS Lett. 261: 204–208.
Williams, J. (1988) Construction of specific mutations in photosystem
II photosynthetic reaction center by genetic engineering methods
in Synechocystis 6803. Methods Enzymol. 167: 766–778.
Yagi, H., Kajiwara, N., Tanaka, H., Tsukihara, T., Kato-Yamada, Y.,
Yoshida, M., et al. (2007) Structures of the thermophilic F1-ATPase
ε subunit suggesting ATP-regulated arm motion of its C-terminal
domain in F1. Proc. Natl Acad. Sci. USA 104: 11233–11238.
Plant Cell Physiol. 51(6): 855–865 (2010) doi:10.1093/pcp/pcq061 © The Author 2010.
Physiological impact of intrinsic ADP inhibition of cyanobacterial FoF1
Yasuda, R., Noji, H., Kinosita, K., Jr. and Yoshida, M. (1998) F1-ATPase is
a highly efficient molecular motor that rotates with discrete 120
degree steps. Cell 93: 1117–1124.
Yasuda, R., Noji, H., Yoshida, M., Kinosita, K., Jr. and Itoh, H. (2001)
Resolution of distinct rotational substeps by submillisecond kinetic
analysis of F1-ATPase. Nature 410: 898–904.
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.
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.
Zhou, J.M., Xue, Z.X., Du, Z.Y., Melese, T. and Boyer, P.D. (1988)
Relationship of tightly bound ADP and ATP to control and
catalysis by chloroplast ATP synthase. Biochemistry 27:
5129–5135.
Plant Cell Physiol. 51(6): 855–865 (2010) doi:10.1093/pcp/pcq061 © The Author 2010.
865