A Unique Resting Position of the ATP-synthase from
Chloroplasts*
Christian
Mellwig and
Bettina
Böttcher
From the EMBL-Heidelberg, Meyerhofstrasse 1, Heidelberg 69117, Germany
Received for publication, December 17, 2002, and in revised form, March 3, 2003
 |
ABSTRACT |
The chloroplast ATP-synthase catalyzes ATP
synthesis coupled to transmembrane proton transport. The enzyme
consists of two parts, a membrane-embedded F0
part and an extrinsic F1 part, which are linked by two
connectors. One of these rotates during catalysis and the other remains
static. Although the atomic structures of various sub-complexes and
individual subunits have been reported, only limited structural
information on the complex, as a whole, is available. In particular,
information on the static connector is lacking. We contribute a
three-dimensional map at about 20-Å resolution, derived from electron
cryomicroscopy of enzymes embedded in vitrified buffer followed by
single particle image analysis. In the three-dimensional map both
connectors, between the F1 part and the
F0 part, are clearly visible. The static connector is tightly attached to an
subunit and faces the side of the
neighboring
subunit. The three-dimensional map provides a scaffold
for fitting in the known atomic structures of various subunits and
sub-complexes, and suggests that the oxidized, non-activated
ATP-synthase from chloroplasts adopts a unique resting position.
 |
INTRODUCTION |
F-type ATP-synthases are found in bacteria, chloroplasts, and
mitochondria. They catalyze ATP synthesis/hydrolysis coupled to
transmembrane proton transport. All ATP-synthases consist of two parts,
a membrane-embedded F0 part and a membrane extrinsic F1 part, which are connected by a thinner connecting region.
In the ATP-synthase from chloroplasts, the F1 part is
composed of five different subunits with the stoichiometry
(
)3

. The
and
subunits surround the
central
subunit, which, together with the
subunit, forms the
central stalk. The F0 part is involved in proton
translocation and consists of four different subunits I, II, III, and
IV. Subunit III is the major component, 14 copies of the subunit III
form a ring (1). A segment of this ring, together with subunit IV,
forms the proton channel (2). Subunits I and II have an amphiphilic
character, with a single transmembrane helix serving as a membrane
anchor. The homologous b subunits in Escherichia coli form a
dimer with inner-dimer contacts in the transmembrane N-terminal region
(3) as well as between amino acids 53 and 122 (4). The b-dimer connects
the F0 part to the F1 part by interacting with
the
subunit (5) at the top of F1 (6) and forms a
peripheral stalk.
According to a current functional model for F-type ATPases (2, 7), the
subunit III-ring in the membrane and the
and
subunit in the
stalk region form a rotor, which rotates during proton translocation.
This rotation induces conformational changes in the catalytic
nucleotide binding sites during ATP synthesis/hydrolysis. A second
static connection (stator) formed by subunits I and II, prevents
co-rotation of the (
)3 core complex. The orientation of the rotor determines the occupancy of the three catalytic nucleotide binding sites. There are three equivalent orientations of the central
rotor relative to the individual catalytic binding sites as shown by
micro videograms (8). Because rotational catalysis requires functional
equivalence of the catalytic binding sites, all three conformations
should occur with the same probability. In the ATP-synthases from
chloroplasts, this type of rotational multisite catalysis requires
activation of the enzyme by a transmembrane potential difference of
protons, 
H+ (9, 10). Up to now it is not
understood if the inactivate chloroplast ATP-synthase also adopts each
of the three conformations with the same probability or takes up a
unique resting conformation.
Although various high resolution structures of sub-complexes from
different F-type ATPases are known (for example, bovine MF1
(
)3

(11); chloroplast CF1
(
)3
(12); E. coli EF1 (
)3
(13); yeast F1c10
(
)3
c10 (14)), none of these
sub-complexes include homologues of the smaller subunits that form the
stator in the chloroplast ATP-synthase. Because the small static
subunits are missing, which are asymmetrically attached to the
(
)3 core, the individual nucleotide binding sites are
only defined in respect to the rotor and not in respect to the stator,
which is insufficient to distinguish between the three possible
conformations. Information on this issue can be obtained by electron
microscopy and image reconstruction of a complete ATP-synthase. Up to
now, projection maps of the ATP-synthase from mitochondria (15),
chloroplasts (16), and E. coli (17), which show
rotor and stator in the connecting region, have been obtained. However,
two-dimensional projection maps alone, without prior knowledge of the
three-dimensional shape, are inadequate to discern between different
conformational states and projections of the same object in different
directions. Such a discrimination can only be done, if the
three-dimensional volumes are also known. Nevertheless,
three-dimensional maps at 30- to 35-Å resolution, of the
negatively stained E. coli (18) and chloroplast enzymes
(19), show the stator either weakly or not at all. Therefore,
three-dimensional maps are still insufficient to decide whether or not
the complex adopts different conformations. Here we present a
three-dimensional map of the ATP-synthase from chloroplasts embedded in
vitrified buffer, in which the stator and rotor are clearly visible and
only a single conformation, as expected for a unique resting position,
is observed.
 |
EXPERIMENTAL PROCEDURES |
Preparation of the Protein for Electron Microscopy--
The
ATP-synthase was purified according to a previous study (19), which
yields a complex where the regulatory disulfide bond of the
-subunit
is oxidized. The buffer was changed by passage through a Sephadex G-50
spin-column equilibrated in 0.1 mM dodecyl maltoside, 10 mM MgCl2, 10 mM KCl, 150 mM NaCl, 10 mM NaH2PO4, pH 7.2. Grids coated with a perforated carbon film were glow-discharged and used within 30 min. After adding 2 mM
AMP-PNP,1 2 µl of the
sample was applied to the pretreated grid. The grid was mounted in a
modified controlled environment freezing apparatus (20). After blotting
for 15 s with filter paper (Whatman No. 1), the samples were
plunged into liquid ethane.
Electron Microscopy and Image Processing--
The vitrified
samples were transferred with a Gatan 626 cryo-holder into a Philips
CM-200-FEG electron microscope. The microscope was operated
under low dose conditions at 200 kV accelerating voltage. Micrographs
were taken on Kodak SO-163 film, at a nominal magnification of ×50,000
with an underfocus of 3-5.1 µm. The micrographs were developed for
10 min in full-strength Kodak D-19 developer at room temperature.
For image processing, suitable micrographs were scanned with a Zeiss
SCAI scanner with a pixel size of 14 µm corresponding to 2.8 Å at
specimen level. The particle images were selected interactively and
were boxed off from the micrographs using the MRC image
processing programs (21). Further image processing was carried out
using the IMAGIC-5 software package (22) as described previously (23)
except for the following modifications. The phases of the particle
images were corrected for the contrast transfer function. The particle
images were band-pass filtered, including information between 1/16 and
1/110 Å
1.
The band-pass filtering and the contrast transfer function of the
microscope lead to an underestimation of the low resolution amplitudes,
causing dark fringes and an overestimation of holes in the complex. To
minimize this effect in the final three-dimensional map, the low
resolution amplitudes of the raw three-dimensional map
(Araw) were scaled relative to the amplitudes of
a binarized three-dimensional map (Abin), in
which gray values inside the particles were set to 1 and outside to 0. The threshold for binarizing was chosen so that the total number of
voxels set to 1 accounted for the expected molecular mass of the
ATP-synthase surrounded by the detergent micelle (~750 kDa). Raw map
and binarized map were Fourier-transformed. For each band of
spatial frequencies (R1
R
R2) the ratio of the average
amplitudes A was calculated,
|
(Eq. 1)
|
The ratio varied at low spatial frequencies R and
reached constant values for a small band of frequencies at about
RR = 1/45 Å
1. At frequencies below this
band, the ratio was used as a scaling factor and at frequencies above
this band the constant ratio in the band was used for scaling of the
amplitudes Araw in the raw map as
follows. For band < RR,
|
(Eq. 2)
|
and for band
RR,
|
(Eq. 3)
|
The scaled three-dimensional Fourier transform was transformed
to real space to give the final three-dimensional map (Fig. 5), which
showed the same gross features as the raw map but smaller holes and
reduced dark fringes, which facilitated fitting of the high resolution
structures. In this approach, the absolute threshold for binarization
was not too critical, however, the absolute size and shape of the
binarized reference map were crucial. Therefore, scaling the electron
microscopic reconstructions with atomic models of smaller
sub-complexes, or models lacking the detergent micelle, yields poorer
results than using the binarized reconstruction for scaling (data not shown).
To demonstrate the effect of the scaling approach, a model experiment
was performed: from the yeast F1c10 sub-complex
(Protein Data Bank 1QO1 (14)) a density map at 20-Å resolution was calculated (Fig. 1A). This map was convoluted with the
contrast transfer function as calculated for a defocus of 3000 nm using a Philips CM-200-FEG operating at 200 kV. The resulting map showed dark
fringes and holes at places where holes were not observed in the
original map. After correcting phases for the contrast transfer
function, the holes were observed at the correct positions (Fig.
1C). However, they were darker and larger than in the
original data. In addition, dark fringes surrounded the particle. Only little improvement was observed, when various reconstructions were
calculated using different defocus values and had phases corrected for
the contrast transfer functions were combined (Fig. 1D). We
assume, that the dark fringes and holes are predominantly effects of
uncorrected amplitudes at low spatial frequencies. In the "true"
reconstruction, we do not expect any negative values (dark
fringes and dark holes). This situation was
approximated by a binarized map (Fig. 1E), the threshold of
which was chosen so the total number of voxels set to 1 accounted for the expected molecular mass. The binarized map gave an
estimate for the average amplitudes of the low spatial frequencies of a
reconstruction without dark fringes. The map was used for scaling the
amplitudes of the low spatial frequencies of the combined
reconstruction (Fig. 1C) as described above. Indeed, scaling
reduced the dark fringes and decreased the size of the holes (Fig.
1F). Although the scaled map (Fig. 1F) was still
not identical to the starting map (Fig.
1A), it was more similar in
its overall appearance than the map (Fig. 1D), where only
phases were corrected for the contrast transfer function.

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Fig. 1.
Demonstration of the effect of amplitude
scaling on the final image. A, from the coordinates of
the yeast F1c10 sub-complex (14) (1QO1), a
density map at 20-Å resolution was calculated. A central slice
parallel to the long molecule axis is shown. B, the density
determined in A was modulated by a contrast transfer
function calculated for a nominal defocus of 3000 nm. C, the
phase information of the density shown in B was corrected
for the contrast transfer function. D, four maps similar to the one in
C, but calculated and phase-corrected for different defocus
values (3000, 3300, 3600, and 3900 nm) were combined. E, the
map shown in D was binarized with a suitable threshold,
describing a volume that would approximately fit the molecular mass of
the complex. F, the amplitude information of the binarized
map shown in E was used to scale the phase-corrected image
shown in D as described.
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|
To estimate the resolution of the three-dimensional map, two
independent maps of half of the class averages were calculated, and
their Fourier shell correlation was determined. The atomic models were
fitted manually to the three-dimensional map using the program O (24).
For the difference map, the fitted atomic models were filtered to the
resolution of the three-dimensional reconstruction. After binarizing
both maps with a threshold value of 0.3, a difference volume was calculated.
 |
RESULTS |
Electron Microscopy and Image Processing--
Fig.
2 shows a typical micrograph of the
ATP-synthase from chloroplasts embedded in a thin layer of vitrified
buffer. The protein formed a uniform monodisperse particle
distribution. For image processing about 10,000 particle images were
selected from 150 micrographs. The particle images were aligned,
classified according to their similarity, and averaged. The class
averages (Fig. 3A) showed
projections of particles with a larger and a smaller domain, which were
linked by one or two connectors. The larger domain corresponded to the
F1 part and the smaller domain to the F0 part,
shielded by the detergent micelle.

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Fig. 2.
Micrograph of the ATP-synthase embedded in
vitrified buffer. Particle images usually selected for image
processing are marked by black circles. The scale
bar indicates 50 nm.
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Fig. 3.
A, class-averages; B, surface
representations of the three-dimensional map; and C,
calculated projections of the three-dimensional map in the same
directions as determined for the class averages in A. Views
are rotated along the long axis of the complex in steps of ~60°. In
the surface representation individual elements possibly corresponding
to the subunits are labeled 1, 3, and
5 and elements corresponding to the subunits are marked
2, 4, and 6. The scale bar
corresponds to 100 Å.
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|
The class averages were combined into a three-dimensional map by exact
filtered back-projection reconstruction (25) (Fig. 3B). All
class averages could be matched by projections of the three-dimensional
map (Fig. 3C). The Euler angles of the class averages
covered most of the asymmetric unit (Fig.
4), leaving only small areas unaccounted
for. To estimate the overall resolution, we calculated the Fourier
shell correlation between two maps, each representing half of the data.
The correlation dropped to 0.5 at (1/21) Å
1 and cut the
three-times noise correlation curve at (1/18) Å
1,
indicating an overall resolution of about 20 Å.

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Fig. 4.
Angular distribution of the orientations of
the class averages. A, front hemisphere. B,
back hemisphere. The first angle is the angle between the north
pole and the latitude line of the determined orientation for a
particular view. The second angle defines the longitude of the
orientation. The different orientations of the class averages are
represented by crosses.
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|
The F1 Part--
In our three-dimensional map the
F1 part was formed by six elongated elements arranged
roughly hexagonally around a central rod (see map in Fig.
5B, labeled 1-6).
These elements could also be recognized in the surface
representations (Fig. 3B) and most likely correspond to the
and
subunits. To decide which of the elements were
and
which were
subunits, we compared this region of the map to the
atomic structures of different F1 sub-complexes (PS3-(
)3 (26); chloroplasts-(
)3
(12); mitochondrial-(
)3
(27)). Although these
sub-complexes had different nucleotide occupancies and varied
significantly in the region nearest to the connecting region, the
subunits always extended further from the center than the
subunits.
We observed a similar pattern in our map (Fig. 5B,
2), where the elements 1, 3, and
5 extended further from the center than 2,
4, and 6. For this reason, we conclude that the
subunits form elements 1, 3, and 5 and that the
subunits are represented by elements 2,
4, and 6. The 
-dimers in our map were not
related by strict 3-fold symmetry. Therefore, we fitted the asymmetric
(
)3 sub-complex of the mitochondrial F1
(PDB 1E79 (11)), rather than the structure of the symmetric (
)3 sub-complex of the F1, from
chloroplasts (PDB 1FX0 (12)), into our three-dimensional map (Fig.
6). The match between the mitochondrial
F1 sub-complex and our map was best when the
subunit with the empty nucleotide binding site was superimposed to the elongated element 6 in our map. We propose that the rod in
the center of the six elongated elements (Fig. 5A,
elements 6 and 7, and 5B,
element 2) correspond to the two long helices of the
subunit in the atomic structure of the mitochondrial F1
part.

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Fig. 5.
Sections through the final three-dimensional
map. A, sections perpendicular to the supposed plane of
membrane. The sections are spaced by 1.1 nm and are 2.8-Å thick.
B, some selected sections parallel to the supposed plane of
membrane. Their approximate positions are indicated in A,
panel 9. The scale bar represents 100 Å.
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Fig. 6.
Fit of the atomic structures of various known
subunits or sub-complexes (colored) to the observed
density (gray). The following units were
fitted: mitochondrial ( )3  ( , light
blue; , dark blue;   , yellow, PDB
1E79 (11)), N-terminal domain of E. coli subunit
(green, PDB 1ABV (28)), ring of 14 copies of E. coli c-subunit (red, PDB 1C99 (36)). For better
comprehensibility, representative slices are shown perpendicular
to the supposed plane of membrane (A) and parallel to the
supposed plane of the membrane (B-D).
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Fitting the
and
subunits and the two long helices of the
subunit to the map, as described above, accounted for most of the
observed density in the F1 part. In the F1 part
only a peanut-shaped density at the top and an elongated density,
running parallel to the
subunit corresponding to element 1, was
unaccounted. The peanut-shaped feature at the top of F1
part consisted of two domains (Fig. 5B, 1). One
of the domains was located in the center of a crown-like region at the
top of the
and
subunits, formed by the
-barrels, as seen in
the x-ray structure (PDB 1E79 (11)). The other domain created a bridge
to the
subunit in element 1. We assume that most of the
peanut-shaped feature was occupied by subunit
, because its central
domain coincided with the area that is recognized by a monoclonal
antibody against the C-terminal region of subunit
in E. coli (6). Consequently, in our three-dimensional map the
N-terminal domain of subunit
must occupy the remaining peripheral
domain into which the structure of the N-terminal domain of E. coli
subunit (28) fitted nicely (Fig. 6).
The Connecting Region--
The F1 and F0
part were joined by two connecting elements, a thin peripheral
connector and one that was larger, and more centrally localized. The
latter was formed by subunits
and
. In the past, two different
conformations have been observed for the central connection. One was
derived from crystals of a
'
sub-complex of the E. coli enzyme (29) and the other from a mitochondrial F1
(11), and it has been argued that both conformations occur during
catalysis (13). The conformations vary in the orientation of the
subunit (or related
subunit in mitochondria F1)
relative to the
subunit with the
subunit rotating 81° and
undergoing a net translation of 23 Å. We fitted both types of central
connections into our map (not shown) and found the conformation
observed in the mitochondrial F1 was the best match (Fig.
6).
In contrast to an earlier reconstruction of the negatively stained
ATP-synthase from E. coli, where two peripheral connectors have been identified (18), only one peripheral connection was observed.
Whether or not the two connections observed in the ATP-synthase of
E. coli are a specialty of the bacterial enzyme or some
artifact probably caused by the staining procedure remains open.
In the chloroplast ATP-synthase the thinner, peripheral connection was
attached at the periphery of the
subunit of the F1 part
(Fig. 3B, element 1). The connection was wider at
the sites of interaction with the F0 and F1
part and thinner between these points. It remains uncertain if this
slimming is a genuine feature of the connection or only appears in our
model due to increased flexibility in this area. Whichever the case, we
think that this peripheral link is the stator, formed by subunits I and
II, in the ATP-synthase from chloroplasts. For the related b-subunits of E. coli, secondary structure determination indicates a
helix content of 80% (30), suggesting that the observed stator might contain only a little more than two extended helices, which is also
supported by x-ray analysis (31). In the F1 part, the
stator does not account for an individual feature and is presumably
tightly attached to the
subunit (element 1), pointing
toward the
subunit (element 6).
To follow the path of the stator through the F1 structure,
we calculated the difference between the volume occupied by the manually fitted x-ray structures and the volume occupied by our three-dimensional map (Fig.
7A). In the F1
part and in the connecting region, the fitted x-ray structures did not
account for subunits I and II, the C-terminal domain of the
subunit, and the disordered N-terminal residues of the
and
subunits. According to the difference, the stator contacted the
subunit (Fig. 5B, element 1), involved in forming
the tight catalytic nucleotide binding site in the center, at the
"bottom." Then the stator lined the side of the
subunit, which
faced the
subunit that forms part of the empty catalytic nucleotide
binding site (Fig. 5B, element 6). At the
top of the F1 part, the stator ended with a contact to the
subunit at the uppermost part of the N-terminal domain. The
observed position of the stator is in agreement with earlier cross-linking data for the stator-forming b and
subunits in E. coli (32, 33).

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Fig. 7.
Slices through a difference map between the
volume derived from electron microscopic data and the volume occupied
by the fitted atomic structures (Fig. 6), which was reduced to the same
resolution as the electron microscopic map. For comparison, both
maps were set to one inside the volume and to zero outside the volume.
The slices are gray where the map and the fitted structure
occupied the same volume. Additional density of the fitted x-ray
structures is white, and additional density in our map is
black. A, slices from the center of the complex
parallel to the long axis; B, slices perpendicular to the
long axis, which are equivalent to those in Fig. 5B but
rotated in plane to match those in A. The scale
bar corresponds to 100 Å.
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The F0 Part--
The F0 part had an
elliptical cross-section in the plane of the membrane (Fig.
5B, 4). An outer oval belt, probably formed by
detergent, surrounded an inner circular ring of similar diameter, as
the subunit III complex observed by atomic force microscopy (1).
Therefore, we conclude that the subunit III complex forms the inner
ring. To get some impression of its space requirements in the context
of our map, we generated a ring of 14 copies of the related E. coli c-subunit. There are two structures available from NMR
measurements that vary in the deprotonation of Asp-61, which in
E. coli has a pKa value of 7.1 (34).
Accordingly, we generated a ring of the protonated (35) and the
deprotonated (36) conformation of the related c-subunit (36) using
MolMol (37) and placed both in the observed density. The manual
placement was guided by the hole in the center of the F0
part in our map, which we superimposed on the central channel of the
modeled ring. A belt of unaccounted density surrounded both modeled
rings. This belt was smaller due to the modeled ring of the
deprotonated conformation. However, at the level of observed detail,
both fits were somewhat arbitrary. In Fig. 6 the modeled ring of the
deprotonated form is shown, because at pH 7.2 this conformation should
be adopted by the majority subunits. The ring was sealed from one side
by the central stalk and from the other side by a plug. The nature of
the plug was unclear, but it was probably formed by either detergent or
by tightly bound lipids. The latter was observed in two-dimensional
crystals of the related c complex of I. tartaricus (38). When the volume occupied by the modeled ring was
subtracted from the observed density, a belt with a bulge was left at
the side where the stator emerged. We conclude that the membrane
domains of subunits I, II, and IV formed this bulge.
 |
DISCUSSION |
ATP-synthases work with a rotational mechanism, which requires the
catalytic nucleotide binding sites to be functionally equivalent. Accordingly, there should be three equivalent conformations in which
the catalytic nucleotide binding sites have the same overall occupancy,
but vary in the occupancy of the individual binding sites as depicted
in Fig. 8. In all three conformations,
the core of (
)3
III14 should have
the same structural organization. Only the presence of the small
peripheral subunits (I, II, IV, and
) provides the means to
distinguish between the three conformations. In the three
conformations, the small peripheral subunits also have to adapt to
spatially different environments, because the invariable core does not
have a strict 3-fold symmetry. In our samples, we have the small static
subunits present, and therefore, we are in principle, able to discern
between the three conformations. Nevertheless, at the given resolution,
our data does not show any indication of the presence of multiple
conformational states. All observed projections can be matched by
projections calculated from the same three-dimensional map, which is
one indication that a conformationally homogeneous particle population
was explored (19). We can also exclude the possibility that particle
images representing different conformations were accidentally combined in the three-dimensional map for the following reason: if particles with the described variation in conformations would have been averaged
in our three-dimensional map, we would either expect the occurrence of
three weak peripheral connectors or of a central connector with a
3-fold symmetry-axis approximately parallel to the long molecule axis
and/or with a fuzzy outline. However, we observe only a single
peripheral connector and a central stalk with a well-defined asymmetric
shape. This stalk accommodates the atomic model of the mitochondrial
central stalk in only one orientation. The two alternative orientations
of the central stalk, which could be expected for the other two
equivalent conformations (Fig. 8), do not match the observed density
(not shown). Therefore, we conclude that the majority of the isolated,
inactive ATP-synthases adopts a single unique resting position
(equivalent to the one depicted in the center of Fig. 8). At present we
do not know if this is a specialty of the ATP-synthase from
chloroplasts, which requires activation by 
H+
for rotational multisite catalysis, or whether it is a genuine feature
for all ATP-synthases.

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Fig. 8.
Schematic representations of the
three-functionally equivalent conformations of the ATP-synthase.
The ATP-synthase is depicted from the top, perpendicular to
the plane of membrane. The conformationally invariant core is marked by
a black outline and consists of subunits
(white), subunits (medium gray), the subunit
III ring (light gray), and the subunit
(black). The subunits forming the catalytic binding sites
are labeled according to the nomenclature introduced by Abrahams
et al. (27) (E for empty, D for ADP,
and T for ATP or ADP). This naming convention does not
necessarily reflect different occupancies in the catalytic binding
sites but is also used in other structures (40) where the identity of
the sites is determined by the orientation of the subunit. At the
periphery of F1 the stator (outlined by the
dashed lines) is formed by subunits , I, II, and IV and
adapts to the spatially different environments of the three
conformations. The putative, unique resting position is equivalent to
the conformation shown in the center.
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|
A prerequisite for a unique resting position would be functional
none-equivalent catalytic nucleotide binding sites. Somehow, the
ATP-synthase must "know" in which position to stop when catalysis halts, due to inactivation by an insufficient

H+. This could be achieved by a change of the
binding constants for nucleotides in one of the three catalytic
nucleotide binding sites. We think that in the ATP-synthase from
chloroplasts, the change in binding constants might be realized by the
stator, which is anchored in the membrane on one hand, where it could
sense changes in the 
H+, and on the other hand
is also tightly attached to one of the
subunits where it could
influence the properties of the adjacent nucleotide binding sites.
Therefore, the stator could "communicate" changes in

H+ to the binding sites by dislocating the
subunit slightly and thus changing the binding constant of the adjacent
nucleotide binding sites.
According to our fit, in the resting position, the stator is attached
to the tight catalytic nucleotide binding site (Fig. 8,
center), which carries an ADP in the
dicyclohexylcarbodiimide (DCCD) inhibited mitochondrial
F1 sub-complex (11). An ADP on a tight catalytic nucleotide
binding site, which cannot be removed, is also found in the purified
ATP-synthase from chloroplasts (39). It is conceivable that this
peculiar property of one of the binding sites is caused by the stator,
which in the resting position could force the adjacent site in a closed
conformation and thus trapping the bound ADP. Because rotational
catalysis requires functional equivalence of the catalytic nucleotide
binding sites, we speculate that activation by

H+, shifts the stator, which in turn relocates
the attached
subunit rendering the binding sites functionally equivalent and thus enabling multisite catalysis again.
 |
ACKNOWLEDGEMENTS |
We thank David Venzke for excellent technical
assistance, Remco Sprangers for help with MolMol, and Peter
Gräber and Susanne Fischer for valuable discussions.
 |
FOOTNOTES |
*
This work was supported by the Deutsche
Forschungsgemeinschaft (Grant BO 1150/3).The costs of publication of this
article were defrayed in part by the
payment of page charges. The article must therefore be hereby marked
"advertisement" in
accordance with 18 U.S.C. Section
1734 solely to indicate this fact.
To whom correspondence should be addressed. Tel.: 49-6221-387-304;
Fax: 49-6221-387-306; E-mail: boettcher@embl-heidelberg.de.
Published, JBC Papers in Press, March 6, 2003, DOI 10.1074/jbc.M212852200
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ABBREVIATIONS |
The abbreviations used are:
AMP-PNP, adenosine
5'-(
,
-imino)triphosphate;
PDB, Protein Data Bank.
 |
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Copyright © 2003 by The American Society for Biochemistry and Molecular Biology, Inc.