Tài liệu Báo cáo khoa học: A point mutation in the ATP synthase of Rhodobacter capsulatus results in differential contributions of DpH and Du in driving the ATP synthesis reaction pptx

A point mutation in the ATP synthase of
Rhodobacter capsulatus
results in differential contributions of DpH and Du in driving
the ATP synthesis reaction
Paola Turina and B. Andrea Melandri
Department of Biology, Laboratory of Biochemistry and Biophysics, University of Bologna, Italy
The interface between the c-subunit oligomer and the
a subunit i n the F
0
sector of the ATP synthase is believed to
form the core o f the rotating motor powered by the protonic
flow. Besides the essential cAsp61 and aArg210 residues
(Escherichia coli numbering), a few other residues at this
interface, although nonessential, show a h igh degree of
conservation, among these aGlu219. The homologous resi-
due aGlu210 in the ATP s ynthase o f t he photosynthetic
bacterium Rhodobacter capsulatus has been substituted by a
lysine. Inner membranes prepared from the mutant strain
showed approximately half of the ATP synthesis activity
when driven both by light and by a cid-base transitions. As
estimated with the ACMA assay, proton pumping rates in
the i nner membranes were also reduced to a similar extent in
the mutant. The most striking impairment of ATP synthesis
in the mutant, a decrease as low as 12 times as compared to
the wild-type, w as observed in the absence of a transmem-
brane e lectrical m embrane potential (Du)atlowtrans-
membrane pH difference ( DpH). Therefore, the mutation
seems t o affect both the mechanism responsible for coupling
F
1
with proton translocation by F

0
, and the mechanism
determining the relative contribution of DpH and Du in
driving ATP synthesis.
Keywords: ATP synthase; mutagenesis; Rhodobacter cap-
sulatus; DpH; Du.
Membrane-bound F
0
F
1
-ATPases (ATP synthases) catalyze
ATP synthesis in bacteria, chloroplasts and mitochondria at
the expenses of an electrochemical potential gradient of
protons (or Na
+
ions in so me species). The membrane-
embedded h ydrophobic F
0
sector is involved in proton
translocation across the membrane, and the hydrophilic F
1
sector contains the catalytic sites (reviewed in [1–3]). A
wealth of high resolution structural information f or the
soluble part has appeared since the first crystal structure of
the mitochondrial F
1
was reported in 1994 [4], paralleled by
an increasing amount of experimental evidence supporting a
rotational mechanism of catalysis (reviewed in [5]).
In the most investigated Escherichia c oli enzyme, F

1
consists of five types of subunits in stoichiometry a
3
b
3
cde
and F
0
consists of three types of subunits in stoichiometry
ab
2
c
9)12
.Thec subunit monomers span the membrane as a
hairpin of two a helices [6] and are arranged in a oligomer in
the form of a ring (see, for example, the crystallographic
evidence in [7]). Subunit a most likely consists of five
transmembrane helices [8–10], the fourth of which has been
shown by extensive cross-linking analysis to pack against
the second transmembrane segment of subunit c [11]. The
fourth and fifth transmembrane helices, residues 206–271,
house the most conserved regions of the subunit.
In view of the ATP-driven rotation of the c-and
e-subunit shaft within the a
3
b
3
subunit barrel in F
1
,itis

proposed that the c subunit ring in F
0
, which is connected to
the ce shaft [12–14], r otates against t he a subunit, which is
connected to the a
3
b
3
barrel through the b and d subunits
[15,16]. Experimental evidence consistent with this idea has
been presented [17–19].
A few mechanistic models for torque generation in F
0
have been proposed, which emphasize the role of electro-
static interactions [2 0–22] or the r ole of conf ormational
changes within the c subunit [23]. All models include a
central role for the essential carboxyl group of the c subunit
and for the essential Arg residue in the a subunit (cAsp61
and aArg210, respectively, in E. coli).
Besides the cAsp61/aArg210 couple in the middle of t he
membra ne, the remai ning a/c interface regions are believed
to form the a ccess pathways for protons. P robably lining the
acidic access pathway is residue aGlu219, based on cross-
linking data [11]. Several lines of evidence support a close
spatial and functional interaction between aG lu219 and
aHis245, including the f act that in the ATP synthases of
mitochondria and of photosynthetic bacteria the position of
these two amino acids in the primary sequence are inverted
[24], the fact that the E. coli double mutant aGlu219 fi
His/aHis245 fi Glu has an ATP synthase activity signifi-

cantly higher than that of either of the single mutation
strains [25], and their close position in the proposed
topological models [8–10]. Although t hese residues were
shown to be nonessential by extensive mutagenic analysis
Correspondence to B. A. Melandri, Laboratory of Biochemistry and
Biophysics, Department of Biology, University of Bologna, Via
Irnerio, 42, I-40126 Bologna, Italy. Fax: + 39 051 242576,
Tel.: + 39 051 2091293, E-mail:
Abbreviations: GTA, gene transfer agent; Bchl, bacteriochlorophyll;
ACMA, 9-amino-6-chloro-2-methoxyacridine; RC, photosynthetic
reaction center; Á
~
ll
H
þ
, transmembrane difference of electrochemical
potential of protons; Du, bulk-to-bulk transmembrane electrical
potential difference; Dw, surface electrical potential difference.
Enzyme: ATP synthase (EC 3.6.3.14).
Note: a website is available at />(Received 12 November 2001, revised 21 February 2002, accepted 21
February 2002)
Eur. J. Biochem. 269, 1984–1992 (2002) Ó FEBS 2002 doi:10.1046/j.1432-1033.2002.02843.x
[25], their impo rtant functional role is indicated both by
their high degree o f conservation a nd by the d eleterious
effects of their mutations on the E. coli ATP syn thase.
In this work, the photosynthetic bacterium Rhodobacter
capsulatus has been used as a convenient system for
genetic manipulation and for investigating catalytic prop-
erties of the ATP synthase, as a variety of functional
studies can be carried out in its well-coupled inner

membrane preparations (chromatophores). The subunit
composition of this ATP synthase is very similar to the
E. coli enzyme [26,27], except that the additional subunit
b¢, homologous to b, is found in F
0
,asitistypicalfor
photosynthetic organisms.
As in other photosynthetic bacteria, in Rb. capsulatus an
inverted mitochondrial-like arrangement for the residue pair
aGlu219/aHis245 is found. The corresponding residues in
the Rb. capsulatus are aHis173 and aGlu210 [27]. We have
introduced the single point mutation aGlu210 fi Lys and
have investigated detailed functional aspects of the ATP
synthase in native membranes. The corresponding mutation
in the E. coli enzyme, aGlu219 fi Lys, was examined i n
two previous studies [28,29], where it was shown to cause
reduced cell growth and reduced proton pumping by an
ATP hydrolytic activity similar to wild-type. The r esults of
the present work are consistent with the data obtained in
E. coli and reveal novel functional properties of the mutated
enzyme.
EXPERIMENTAL PROCEDURES
Bacterial strains, growth conditions, membrane
preparations
Rb. c apsulatus strain B100 is wild-type strain B10 cured of
phages. A s pontaneous rifampicin-resistant o f B100, J1,
was used in the GTA
1
procedures. Rb. capsulatus strains
were grown photoeterotrophically on a synthetic medium

containing malate as a carbon source [30]; kanamycin and
tetracycline were added at 25 and 2 lgÆmL
)1
, respectively.
Cultures were illuminated by two opposite panels e ach
carrying nine 100-W i ncandescent light bulbs; excessive
warming was prevented by water cooling. The mutant and
wild-type strain we re grown in p arallel and cells were
harvested at D
600
¼ 1.2–1.4. Intra-cytoplasmic mem-
branes (chromatophores) were prepared according to the
method described previously [30], r esuspended in 50 m
M
glycyl-glycine/NaOH, 5 m
M
MgCl
2
, pH 7.5, rapidly
frozen as small d roplets in liquid n itrogen, and stored a t
)80 °C.
Subunit a mutagenesis
The wild-type copy of subunit a was c arried by the pFo16
plasmid which contained the whole atp2 operon of
Rb. c apsulatus cloned into the pTZ19U plasmid [27]. The
aE210 fi K mutation was introduced into this plasmid by
using the QuickChange Site-Directed Mutagenesis K it
(Stratagene), based on linear PCR, using the following
mutagenic oligonucleotides: 5¢-CGCGATGTATGCGC
TC

AAGATCCTCGTGGCC-3¢ and 5¢-GGCCACGAGG
AT
CTTGAGCGCATACATCGCG-3¢. The mutation
introduced an additional restriction site for XhoII, therefore
its p resence was confirmed by restriction site analysis. The
mutated atp2 operon was then cloned into the broad-host-
range plasmid pRK415 [31] carrying the tetracycline
resistance, as described previously [27]. The new plasmid,
pKFo102, was subsequently introduced into Rb. capsulatus
B100 strain by triparental conjugation [32]. The wild-type
chromosomal copy of the atp2 operon was deleted by taking
advantage of the so-called GTA, bacteriophage-like parti-
cles produced by Rb. capsulatus [33] as described previously
[26]. GTA particles are produced by Rb. c apsulatus cells
which pack, randomly, pieces of DNA about 4.6-kb long,
either from the chromosome o r from resident p lasmids of
donor cells, and transfer them to acceptor cells, where they
are integrated into the chromosome by homologous
recombination. This results in a n exchange between the
incoming DNA and the corresponding chromosomal DNA
which is lost in the process. In this work, the GTA exchange
donor was the J1 strain carrying the p Fo39 plasmid. This
plasmid was a gift from R. Borghese in our department and
had been constructed by inserting the kanamycin resistence
cassette of Tn903 in place of the atpBEXF g enes (subunits
a, c, b¢, b) leaving only a C-terminal truncated atpI gene
(subunit i). Af ter GTA transfer, kanamycin-resistant colo-
nies appeared that could contain the kanamycin resistance
cassette (i.e. the F
0

deletion) either on the chromosomes or
on the pKFo102 plasmid. Restriction analysis of the
plasmid isolated from such colonies allowed the selection
of those carrying the F
0
deletion on the chromosomes. Two
mutated s trains were selected, w hich carried mutated
plasmids originated from two different PCR runs. A
pseudo-wild-type strain was constructed in parallel, which
carried the wild-type F
0
operon on the plasmid and the
deletion of the chromosomal F
0
operon. The cells used for
chromatophores preparations were routinely checked for
the presence of the mutation by XhoII restriction analysis of
the resident plasmid.
Western blot
The amount of ATP synthase in the membrane was
evaluated by quantitative Western blot on SD S/PAGE
isolated chromatophores protein, using a yeast anti-(b
subunit) antiserum (kindly provided by J. Velours, Bor-
deaux), the luminol assay for detection, and a purified ATP
synthase (isolated from Rb. capsulatus as described previ-
ously [34]) as a standard. The amounts of chromatophores
and standard protein in the different lanes of a single gel
were kept in the linear range of the luminol assay response.
Light-induced ATP synthesis
Light-driven ATP synthesis was carried out at 30 °Cin

the following buffer: 100 m
M
glycyl-glycine/NaOH,
pH 7.7 5, 50 m
M
KCl, 10 m
M
Mg-acetate, 0 .1% bovine
serum albumin, 5 m
M
P
i
,0.2m
M
succinic acid, 0.8 m
M
AMP, 10 l
M
Bchl. The chromatophores suspension was
illuminated from two opposite sides by two 100 W
incandescent bulbs. The reaction was started by addition
of 100 l
M
ADP. After s topping the r eaction with 7 %
perchloric acid, t he ATP c oncentration i n each s ample
was measured in a luminometer (LKB 1 250) with the
ATP-Monitoring Kit ( Bioorbit). The small amount of
ATP synthesized in the dark (due to the adenylate kinase
reaction) was subtracted. For the experiments reported in
Ó FEBS 2002 The aE210K mutation in Rb. capsulatus ATP synthase (Eur. J. Biochem. 269) 1985

Fig. 1, the cuvette contained 2 0 l
M
ADP, 6 0 l
M
lucif-
erine and 2–10 lgÆmL
)1
purified luciferase (32–160 ·
10
3
light units ÆmL
)1
) from Sigma (L9009) in the reaction
mixture described above, and the luminescence was
detected in real-time a t room temperature essentially as
described previously [35]. The assay mixture was illumi-
nated by a halogen lamp (160 WÆm
)2
light intensity,
filtered through 1 cm water and two layers of 8 8 A
Wratten fi lters) and different illumination times were
determined by an electronic shutter controlled by a
Uniblitz T132 Driver . The photomultiplier was shielded
against actinic light by a copper sulfate solution. The
amount of sy nthetized ATP w as evaluated by a dding
10–25 n
M
standard ATP.
ATP synthesis induced by acid-base transitions
Acid-base driven ATP synthesis was carried out similarly

as described in [ 36] by rapidly injecting an acidified
chromatophores suspension into a l uminometer cuvette
containing a basic solution and monitoring the ATP
concentration c ontinuously with luciferine/luciferase i n the
luminometer. The chromatophores were first resuspended
in 5 m
M
P
i
/NaOH, pH 7.3, 2 m
M
MgCl
2
,1m
M
AMP,
5 l
M
valinomycin, 10% sucr ose, and either 1 m
M
KCl
(+Du)or100m
M
KCl (–Du) and left incubating in this
medium  1 h at room temperature. This time was enough
to equilibrate the K
+
concentration across the membrane,
as judged by the d isappearance of the K
+

/valinomycin
diffusion potential (monitored by the carotenoid shift
signal) i nduced by the initial K
+
gradient [36]. The
chromatophores were then mixed with the acidic solution
[30 m
M
succinic acid/NaOH, pH 4.6–6.5, 2 m
M
MgCl
2
,
5m
M
P
i
,either1m
M
(+Du) or 100 m
M
KCl (–Du), 1 m
M
AMP, 5 l
M
valinomycin] and incubated at room tempera-
ture at variable t imes between 2 and 30 min, depending on
the pH of t he suspending buffer, prior t o injection of
100 lL i nto the basic solution. This latter contained
850 lL o f basic solution so that the final concentrations

after chromatophores addition would be 200 m
M
Tricine,
2m
M
MgCl
2
,5m
M
P
i
,either150m
M
KOH + 30 m
M
NaOH (+Du)or100m
M
KOH + 80 m
M
NaOH (–Du),
pH 8.65, 1 m
M
AMP, 100 l
M
ADP, 50 lLoftheATP-
Monitoring Kit and 5 lgÆmL
)1
of purified luciferase
(80 · 10
3

light units ÆmL
)1
). The final Bchl c oncentration
varied between 1 and 8 l
M
. The ATP concentration was
evaluated by adding 100–200 n
M
standard ATP in each
cuvette. The basic solution was thermostated so that the
ATP synthesis reaction took place a t 13 °C. The p H
measured after mixing the chroma tophores with the acidic
solution was taken as the internal pH, the pH measured
after mixing the acidified chromatophores with the basic
solution (8.55 ± 0.05) was taken as the external pH. Their
difference is the indicated DpH. F or the lowest DpH
differences the Ôacidic Õ solution contained 20 m
M
Tricine
instead of succinic acid. Assuming complete equilibration
of the K
+
during the 1 h preincubation (see above), the
value of t he K
+
/valinomycin diffusion potential deter-
mined by the K
+
transmembrane concentration difference
during the acid-base transition can be approximated, ( on

the basis of the Nernst equation or the Goldman equation
for monovalent ions). For T ¼ 12 °C, Du ¼ 124 mV or
Du ¼ 0 mV are obtained b ased on the Nernst equation,
Du ¼ 75 mV or Du ¼ 0.4 mV results by applying
the Goldman equation as described previously [36],
for [K
+
]
in
/[K
+
]
out
¼ 1/150 m
M
and [ K
+
]
in
/[K
+
]
out
¼
100/100 m
M
, respectively.
ACMA assay
ACMA fluorescence quenching assay was carried out in a
Jasco FP 500 spectrofluorimeter (wavelen gth 412 and

482 nm for excitation and emission, respectively) at 1 5 °C
in the following mixture: 20 m
M
Tricine/KOH, pH 8.0,
50 m
M
KCl, 0.5 m
M
MgCl
2
,0.2m
M
succinic acid , 5 l
M
Antimycin, 0.2 l
M
myxothiazol, 2 l
M
valinomycin, 1.5 l
M
ACMA, 20 l
M
Bchl, 400 l
M
ATP. The response of ACMA
to DpH was empirically calibrated using artificially induced
protonic g radients, established by HCl and NaOH a ddi-
tions, under similar temperature and buffer c onditions,
except for the presence of 20 m
M

succinic acid, as described
previously [37].
Fig. 1. ATP s ynthesis in c hromatophores i nduced by light. Light-
induced ATP synthesis in the absence (A) or in the presence of 2 l
M
valimomycin (B). The increase in luminescence associated with ATP
production follow ing a short period of illumination has been mon-
itored. The reactio n assays, describe d in the Experime ntal proce dures
section, contained luciferase ( 32–160 · 10
3
light unitsÆmL
)1
)and
luciferine (60 l
M
) as reported previously [35]. ADP and Bchl concen-
trations were 20 l
M
and 10 l
M
, respectively. The temperature was
28 °C. (d,m) wild-type and (s,n) mutant chromatophores. (B) The
data points h ave been fitted with a n arbitrary function. The d ashed
lines extrapolate the linear part of the curves. (C) The first derivatives
of the fi tting functions were calculated b etween 0 a nd 1000 ms
(extrapolating the fittin g function when necessary). The ratios of the
wild-type derivatives over the mutant, indicating the ratios of the ATP
synthesis rates, are plotted for the exp eriments in the absence (dashed
line) or in the presence (continu ous line) of valinomycin. As n o ATP
synthesis could be detected with added valinomycin for short illumin-

ation times, the rat io function was truncated for times £ 150 ms.
1986 P. Turina and B. A. Melandri (Eur. J. Biochem. 269) Ó FEBS 2002
ATP hydrolysis assays
ATP hydrolysis was measured routinely at 30 °Cinthe
following buffer: 20 m
M
Tricine/KOH, pH 8.0, 50 m
M
KCl, 2 m
M
MgCl
2
,0.2m
M
succinic acid, 20 l
M
Bchl. Th e
reaction was started by adding 1 m
M
ATP. After stopping
the reaction at different time s with 5% trichloroacetic acid,
the P
i
concentration was measured by molybdate colori-
metric assay as described previously [38]. For more sensitive
measurements, the re leased P
i
was measu red w ith t he
EnzCheck Phosphate Assay Kit (Molecular Probes) or with
the malachite green assay [39], both methods giving similar

results.
Other methods
Bchl concentration was measured in acetone/methanol
extract [40]. The protein concentration of c hromatophores
was determined using the BCA assay (Pierce). An aliquot
of the c hromatophores was e xtracted with acetone/meth-
anol (7 : 2 , v/v). After centrifugation, the p rotein pellet
was dissolved in 0.1
M
NaOH/1% SDS for determination
of protein concentration. The light-induced transmem-
brane electric potential difference was evaluated following
the electrochromic signal of endogenous carotenoid [41].
The c oncentration of photo-oxidizable reaction centre
(RC) and of total photo-oxidizable cytochrome (c
1
+ c
2
)
were measured as described previously [42], following
trains of closely spaced flashes. The amount of phosp-
holipid was determined by t he method described previ-
ously [43].
RESULTS
The single point mutation aE210K was introduced into the
atp2 operon o f Rb. capsulatus, c ontaining the F
0
genes,
cloned in an E. coli strain. The mutated operon was then
transferred into a broad-host-range vector and the resulting

plasmid introduced by conjugation into Rb. capsulatus
wild-type cells. Finally, a GTA transfer was allowe d to
take place , wh ich generated the deletion o f the chromoso-
mal atp2 operon by substitution with a kanamycin
resistance cassette. Therefore, the resulting strain carried
the deletion of the chromosomal atp2 operon and several
copies of a plasmid carrying the mutated atp2 operon. As a
control, a parallel at p2-deleted strain w as created, in which
the r esident p lasmid carried the w ild-type operon. This
pseudo-wild-type strain is referred to as wild-type i n the
following procedures.
Characterization of mutant chromatophores
Phototrophic g rowth of the aE210K mutant cells was
slower than the wild-type cells. Accordingly, the light-
induced ATP synthesis r ate catalyzed by the mutant
chromatophores was about 40% lower on a Bchl basis
than the rate c atalyzed by wild-type c hromatophores
(Table 1). T he same reduction res ulted also when the
ADP concentration was varied between 20 and 500 l
M
,
indicating that the mutation does not affect the apparent K
m
for ADP. In contrast, no significant difference could be
observed in t he ATP hydrolysis r ate. The concentration of
ATP synthase was estimated on a Bchl basis by quantitative
Western blot analysis and was found to be the same within
experimental error for both mutant and wild-type chro-
matophores. The specific activity of ATP s ynth esis was
8±2and13±3ATP/(F

1
F
0
Æs) for mutant and wild-type,
respectively, whereas the specific ATP hydrolysis activities
were 2.0 ± 0.7 and 2.3 ± 1.0 ATP/(F
1
F
0
Æs). These values
are from a single preparation of chromatophores but are
representative of several preparations, obtained from strains
carrying mutated plasmids originating from two different
PCR runs.
The lower ATP synthesis rate of the mutant was not
due to a lower efficiency of the electron transport chain or
higher permeability of the membrane as the ATP
synthesis rate induced by acid-base transitions was
similarly reduced (see below and Fig. 3A). The possibility
of a higher percentage of open, and therefore uncoupled,
membrane fragments o r of r ight-side-out vesicles in the
mutant was also r uled out by measuring t he extent of
flash-induced carotenoid shift and the extent of cyto-
chrome c accessible photo-o xidizable RC, which were
comparable to those of w ild-type chromatophores (not
shown).
The mutant and wild-type preparations were further
characterized as to their RC, phospholipid and protein
content ( Table 1 ). The ATP synthase/Bchl ratio, the size of
the a ntenna (Bchl/RC) and the phospholipid content were

very similar in both strains. The most striking difference was
found in the protein content, which w as approximately
1.6-fold lower in the mutant chromatophores on a B chl
basis. It is possible that this difference affects the adsorption
of ACMA to the m embrane and therefore t he probe
response to DpH (see below).
Table 1. Catalytic activities and composition of chromatophores from
wild-type and mutant cells. All values reported are from a s ingle
chromatophores preparation but are representative of several different
preparations.
Wild-type Mutant aE210 fi K
Photophosphorylation rate
a
74 ± 4 44 ± 3
(mM ATP/M Bchl/s)
ATP Hydrolysis Rate
b
13 ± 3 11 ± 2
(mM P
i
/M Bchl/s)
Bchl/ATP Synthase Ratio
c
178 ± 35 180 ± 31
(moles/mole)
Protein/Bchl 48.4 29.7
(mg/lmole)
Bchl/RC 71 89
(moles/mole)
phospholipids/Bchl 13 ± 1.8 12 ± 2.0

(moles/mole)
a
The ATP synthesized at each illumination time (n ¼ 5) was
measured after denaturation in a luminometer with the ATP
Monitoring Kit as described in the Experimental procedures.
b
The
P
i
released at each time point (n ¼ 5) was analyzed after dena-
turation by the colorimetric assay described previously [38].
c
After
SDS/PAGE and transfer onto nitrocellulose paper of chromato-
phores and known amounts of isolated Rb. capsulatus ATP
synthase, the unknown amount of ATP synthase was evaluated by
detecting the b subunit with an anti-(b subunit) Ig in a linear range
of response.
Ó FEBS 2002 The aE210K mutation in Rb. capsulatus ATP synthase (Eur. J. Biochem. 269) 1987
Light-induced ATP synthesis at low D
~
ll
H
þ
The ATP synthesis rates reported in Table 1 were measured
under light close to saturation at times ranging from 2 to
10 s. Under these conditions, a high steady-state D
~
ll
H

þ
was
obtained, which resulted in a linear increase of the ATP
yields with illumination time. In order to investigate the
activity of the mutated enzyme at lower D
~
ll
H
þ
values, shorter
illumination times were chosen (from 100 ms to 2 s ). The
assays were also supplemented with the ionophore valino-
mycin, which largely prevents the onset of the electrical
component of D
~
ll
H
þ
, thus further reducing its total extent
during short illumination times. Due to the low ATP yields
expected in these experiments, the luciferine/luciferase ATP
detection system was added into t he assay cuvette at high
concentration (32–160 · 10
3
light units ÆmL
)1
of luciferase),
so that the ATP-induced luminescence was directly detected.
Figure 1A shows the ATP yields obtained from m utant
and wild-type chromatophores as a function of illumination

time in the absence of valinomycin. A linear relationship
was observed under these conditions, and the ATP yield by
the mutant chromatophores was  50% of that by the wild-
type at every time point. W hen the assay c ontained
valinomycin, the data reported in Fig. 1 B were obtained.
In this case, the ATP yield of both wild-type and mutant as
a function of illumination time presented a lag phase before
becoming linear. A similar lag phase had a lready been
observed in wild-type c hromatophores [35] and c an be
interpreted as being due to a lag phase in the onset of D
~
ll
H
þ
when its e lectrical c omponent is being dissipated b y
valinomycin.
Strikingly, this lag phase was much more pronounced in
thecaseofthemutant.Thelargedifferencerelativetothe
wild-type can be best appreciated by taking the derivative of
the fi tting f unctions of the mutant and wild-type data a nd
plotting their ratio, as in Fig. 1C. This derivative represents
the r ate o f ATP synthesis at each time point. The twofold
difference between the mutant and wild-type observed in the
presence of a Du is approached o nly for illumination times
longer than 1 s, whereas between 200 and 500 ms the ATP
synthesis rate catalyzed by the mutant is much decreased, up
to 12-fold lower with respect to the wild-type. Clearly, the
effect of the mutation is much larger in the absence of a
significant Du and at low D
~

ll
H
þ
values.
ATP synthesis induced by acid-base transitions
The functioning of the mutated enzyme was also studied by
using the technique of acid-base transitions. This technique
allows one to control the extent of DpH across vesicle
Fig. 2. ATP synthesis driven by acid-base transitions. Chromatophores
were preincubated in resuspending and acidic media as described in the
Experimental procedures, and injected into the basic medium as
indicated by the arrow. The ATP synthesis following chromatophores
injection was mo nitored continuously with t he luciferine/luciferase
ATP Monitoring Kit in a luminometer. The high signal-to-noise ratio
was obtained due to ad ded purifie d luciferase (80 · 10
3
light
unitsÆmL
)1
). The internal and external pH’s were 4.96 and 8.54,
respectively, and the [K
+
]
out
and [K
+
]
in
were 150 m
M

and 1 m
M
,
respectively, thus inducing a K
+
/valinomycin diffus ion potential
(+Du). Bchl concentration in t he luminometer cuvette was 1.1 l
M
.
The reaction temperature was 1 3 °C. Oligomycin was added during
the preincubation time at a concentratio n of 20 lgÆmL
)1
and was
present at the same concentration in the luminometer cuvette.
Fig. 3. Rates of ATP synthesis induced by acid-base transitions in the
presence and in t he absence of a diffusion potential. (A) Data are
obtained from measurements similar to those reported in Fig. 2. The
compositio n of the preincubation and reaction mixtures are detailed in
the Experimental procedures. Bchl concentration varied between 1 and
8 l
M
. The reaction temperature was 13 °C. The ade nylate kinase rate
has been subtracted . Data are averages of 2–3 measurements. (d,m)
wild-type and ( s,n) mutant chromatophores. Rates w ere measured
both in the p re sence (d,s) and in the ab sence (m,n)ofadiffusion
potential (Du), i.e. with [K
+
]
out
and [K

+
]
in
equal to 1 and 150 m
M
,
respectively, or 100 and 100 m
M
, respectively. The data points have
been fitted with arbitrary functions. (B) An enlarged view of (A),
showing only the data obtained in the absence of Du.(C)Theratioof
the best-fitting functions and of the data points of wild-type over
mutant are plotted for data in the presence (d)andabsence(m)ofDu.
1988 P. Turina and B. A. Melandri (Eur. J. Biochem. 269) Ó FEBS 2002
membranes and to independently superimpose an electrical
diffusion potential Du, by varying the K
+
gradient across
the vesicle membrane in the presence of valinomycin. In the
present work, the [K
+
]
in
and [K
+
]
out
for + Du was 1 and
150 m
M

, respectively, and the [K
+
]
in
and [ K
+
]
out
for –Du
was 100 and 100 m
M
, respectively. The DpH values were
varied up to 3.8 units by decreasing the internal pH at
constant external pH. Rb. c apsulatus chromatophores had
been shown to catalyze, by this technique, rates of ATP
synthesis comparable to those obtained by illumination [36].
At [K
+
]
in
¼ [K
+
]
out
¼ 100 m
M
(–Du condition), the
absence of any significant diffusion potential due to other
ionic species, e.g. monoionic succinate [44] was e xcluded
observing the electrochromic spectral shift of carotenoids.

We therefore conclude that in chromatophores of Rb. c ap-
sulatus, under these conditions, DpH represents the only
driving force for ATP synthesis.
When carried out at room temperature, the linear phase
of the D
~
ll
H
þ
-induced ATP synthesis decays within a few
hundred milliseconds [36] , due to the r elatively h igh
permeability o f t he chromatophores membrane and to the
high H
+
flow through the ATP synthase, thus requiring for
its measurement a quench-flow apparatus. In the present
work, the du ration of this linear phase was increased to a
few seconds by decreasing the reaction temperature to
13 °C, and the transition was carried out manually by
rapidly injecting the acidic chromatophores suspension into
the luminometer cuvette containing the b asic solution and
the luciferine/luciferase detection kit (plus additional
80 · 10
3
light units ÆmL
)1
of luciferase). This method has
already been applied at room temperature for measuring the
ATP synthesis activity of ATP synthases incorporated into
liposomes [45,46].

Figure 2 shows some of the traces obtained by this
method. The initial high rates of ATP synthesis decay within
a few seconds mainly due to the decay of the D
~
ll
H
þ
-induced
by the acid-base transitio n. The slow increase of ATP
concentration seen thereafter was uncoupler- (not shown)
and oligomycin-insensitive, and can be attributed to the
adenylate kinase a ctivity present in chromatophores pre-
parations [36]. Such a rate has been routinely subtracted
from the initial ATP synthesis rate, which is uncoupler- and
oligomycin-sensitive and can be attributed to the ATP
synthase. This initial rate has been measured as a function of
DpH in the presence and absence of Du for both mutant and
wild-type chromatophores preparations. The resulting data
are shown in Fig. 3A. In the presence of a Du,theATP
synthesis rates of the m utants wer e about twofold lower
with respect to the wild-type at every DpH tested. Figure 3B
is an enlarged view of Fig. 3A showing only the r ates
obtained in the absence of a Du. In this case, the impairment
of the ATP synthesis rates in th e mutant was higher than
twofold, especially in the low DpH range.
Again, this latter comparison can be best appreciated by
plotting the ratio of the fi tting f unctions of wild-type over
mutant, either in the presence or in the absence of a Du,as
shown in Fig. 3 C. In the presence of a Du, the difference
between the two strains was twofold over the entire DpH

range tested. In the absence of a Du, the ATP synthesis rate
of the mutant was up to eightfold lower with respect to the
wild-type for DpH values r anging between 2 .4 and 3.3,
whereas a twofold factor was approached at the h ighest
DpH tested. The trace –Du in Fig. 3C can b e d irectly
compared to the +valinomycin trace in Fig. 1C, because, in
the absence of a Du,theDpH is low in the short illumination
times (200–500 ms), and it increases at longer times.
Efficiency of proton pumping as estimated
with the ACMA assay
The proton-pumping activities of the mutant and wild-type
chromatophores were compared first by measuring the
ATP-induced fluorescence quenching of ACMA. For better
comparison of the in itial rates of quenching, t he ATP
hydrolysis rates were slowed down by decreasing the
reaction temperature to 15 °C. The ACMA fluorescence
quenching as a function of time after addition of ATP is
shown i n Fig. 4A f or chromatophores of both strains. N o
significant difference between the two chromatophores
preparations could be detected. However, when the ACMA
quenching was calibrated as a function of known DpH’s
generated during acid-base transitions, as described previ-
ously [37], the r esponse turned out to be sign ifican tly
different for the mutant and wild-type c hromatophores
(Fig. 4 B). This different respon se was systematically found
in different experimental sessions and i n different chro-
matophores preparations. Therefore, after converting the
fluorescence quenching values into DpH values according to
this calibration procedure, the initial rate of DpH formation
resulted to be higher in the wild-type w ith respect to the

mutant chromatophores by a factor of about 1.8, as shown
in Fig. 4C. The rates of ATP hydrolysis measured under the
same conditions of the ACMA quenching did not differ
significantly (see Fig. 4D).
Under t he assumption that the buffering capacity and
inner volume of chromatophores were the same for both
preparations, these data can be taken to indicate that the
mutated enzyme was less efficient than the wild-type in
ATP-driven proto n tr anslocation ( ATP slipping). The
reason for this d ifferent response of ACMA to calibration
is presently unclear, but it could be related to the only gross
structural difference found between wild-type and mutant
chromatophores, i.e. the different protein/Bchl ratio (see
Table 1), which might affect the adsorption equilibria
between the free and membrane-bound probe [37].
DISCUSSION
The main finding of this work is t hat a single point mutation
can a ffect the ATP synthesis rate o f an ATP synthase
differently according to whether a Du is present or not. To
our knowledge, no previous reports of a similar effect exist
in the literature on ATP synthases .
Two different lines of evidence reported in this work
support this conclusion. In the light-driven ATP synthesis
measurements (Fig. 1), the Du was c ollapsed by the
ionophore valinomycin and a significantly higher impair-
ment in the catalysis by the mutated enzyme (up to 12-fold)
was seen during the first 1000 ms of illumination, i.e. when a
DpH was building up but had not yet reached its maximum,
stationary value. In the ATP synthesis induced by acid-base
transitions ( Fig. 3 ), a Du in the presence o f the K

+
-
transporter valinomycin was either imposed by a [K
+
]
gradient across the membrane, or prevented by imposing
equally high K
+
concentrations in both intra- and extra-
vesicular compartments. In the latter case, the A TP
Ó FEBS 2002 The aE210K mutation in Rb. capsulatus ATP synthase (Eur. J. Biochem. 269) 1989
synthesis r ate was m uch lower (up to eightfold) in the
mutant with respect to the wild-type in the DpH range
between 2.4 and 3.3.
The fact that this higher impairment is seen only within a
limited pH
in
range strongly suggests that a step in the ATP
synthase functioning, which is rate-limiting at th e low H
+
in
concentrations, gradually ceases to b e r ate-limiting f or
increasing H
+
in
concentrations. It is this rate-limiting step,
apparently, which is strongly affected in the mutant. What
could be rate-limiting at t he lowest H
+
in

concentrations is
most likely the rate of H
+
binding to some of the H
+
binding sites in the H
+
translocation pathway, ultimately
cAsp61. Models of the F
0
rotation driven by the
protonmotive force [20,21] suggests a periplasmic (acidic)
and a cytoplasmic (basic) half-channel, placed between
subunit a and the c-subunit oligomer, connecting t he bulk
aqueous compartments to the cAsp61 in the middle of the
membrane. I n these models, an e lectrostatic con straint
implies that c arboxyl groups on the r ing are always
protonated ( electroneutral) when facing the lipid phase,
while they can be unprotonated and charged when facing
the a subunit. The t wo half-channels are spatially offset, so
that this electrostatic constraint imposes a directionality to
the c-oligomer rotational fluctuations. A ccording to the
model p roposed by Oster and co workers [ 21], the rate
constant k
in
for proton hopping into the channel from the
periplasmic s ide d epends upon [H
+
]
in

and upon the
potential drop Dw due to surface charges, i.e. k
in
a10
–pHin
exp(Dw/RT). As a Glu fi Lys substitution is expected to
change the e lectrostatic profile o f t he nearby region, a
possible explanation for the data presented is that residue
a210 (a219 in E. co li) belongs to the periplasmic half-
channel and that one of i ts roles is to create a favorable
electrostatic profile f or incoming protons. This role is
drastically altered when a lysine side-chain substitutes the
carboxylate.
Within this framework, the results obtained in the
presence of a Du indicate that different steps of the
reaction cycle b ecome rate-limiting under these conditions.
According to the cited model [21], the main effects of a Du
in speeding up the enzymatic rate are an increased rate o f
H
+
releaseatthecytoplasmicsideandadecreasedrateof
H
+
release at the periplasmic side. Particularly this last
effect can presumably compensate for the low bulk H
+
in
concentration b y effectively i ncreasing [H
+
] w ithin the

periplasmic half-channel. Under these conditions, i.e. in the
presence of a high bulk-to-bulk Du, the overall reaction
rate would be relatively less sensitive to the extent of the
surface p otential drop Dw at the entrance of this half-
channel.
Fig. 4. Proton pumping and ATP hydrolysis. The composition o f the reaction mixtures are detailed in the Experimental procedures. ATP and Bchl
concentrations were 400 and 20 l
M
, respectively. The t emperature was 15 °C. Photosynthetic electron transfer was inhibited by the addition of
myxothiazol and antimycin to avoid any proton pumping due to the measuring light. The onset of a membrane potential was prevented b y the
addition of 2 l
M
valinomycin. (d) wild-t ype and (m) mutant chromatophores. (A) ACMA fluorescence quenching as a function of time aft er
addition of ATP. The original recorder trac es have been digitalized and data po ints have been fitted with arbitrary exponential functions. (B)
Calibration of the ACMA fluorescence response to transmembrane pH differe nces. Th e ACMA fl uorescenc e que nching h as be en measure d after
imposing various DpH jumps. The reaction m ixture was identical to that in Fig. 4A, except that 20 m
M
succina te was added to i ncr ease the
buffering power at acidic pH and no ATP was added. The curves are the best-fit of data to the function described previously [37]:
DpH ¼ (A Æ Q%) / (B ) Q%) Æ exp[Q%/(B ) Q%) + C Æ Q%]. The values obtained for the empirical fitting parameters were A ¼ 17.8,
B ¼ 118 .8, C ¼ )0.046 and A ¼ 10.9, B ¼ 13.6, C ¼ )0.086 for wild-type and mutant, respectively. (C) The fluorescence quenching values
of (A) have been converted to DpH values according to the best-fit functions of (B). (D) P
i
concentration as a function of time measured after
addition of ATP to wild-type or mutant chromatophores, under experimental conditions identical to those of the fluorescence measurements of (A).
After trichloroacetic acid denaturation, th e released P
i
was analyzed with the EnzCh eck Phosphate Assay. The rates of AT P hydrolysis resulted
4.7 m
M

ATP/(M BchlÆs) for both wild-type and the E210K mutant.
1990 P. Turina and B. A. Melandri (Eur. J. Biochem. 269) Ó FEBS 2002
A s econd major effect of the aE210K mutation is the
generally lower ATP synthesis f ound regardless of D
~
ll
H
þ
extent and composition, which might also explain t he lower
growth rate, as found here and in E. coli strains carrying
the homologous aE219K mut ation [28,29]. This general
impairment of ATP synthesis amounted here ap proxi-
mately to a factor of 2. This is seen for light-driven ATP
synthesis, both for long and short illumination times
(Table 1 and Fig. 1A, respectively), a s well as f or ATP
synthesis driven b y acid-base transitions, both in the
presence of a Du andinitsabsenceathighH
+
in
concentrations (Fig. 3C).
Several hypotheses can be put forward t o e xplain this
impairment. One possibility is that the mutation perturbs
the electrostatic profile sensed b y t he unproto nated Asp61
during i ts r adial translocation, thereby i ncreasing t he
probability of protons leaking through F
0
without perform-
ing any work [21]. This perturbation would also reduce the
H
+

pumping efficie ncy, in line with t he conclusions
suggested by our (Fig. 4) and other [28,29] data. Evidence
for ATP slipping caused by the aE210K mutation, if
confirmed, w ould b e a n i ndependent element for assessing
theroleofGlu210intheprotonpathwaywithinF
0
.
Alternatively, it can be t hat the mutation has an even
longer-range effect, affecting t he conformation at the
interface between the c-ring and the ce shaft. The
conformational modification of t his interface could also
lead to a similar phenotype of decreased coupling efficiency
both in ATP synthesis and in ATP hydrolysis direction. As a
direct connection between subunit a and F
1
has not yet been
seen, s uch l ong-distance e ffect would p resumably o ccur
through a change of the c-ring conformation. This hypoth-
esis cannot be excluded especially in view of the results
recently obtained by Cain and coworkers [47], which showed
that structural changes resulting in the E. coli aR2 10 fi I,
aA217 fi R, a nd a G218 fi D mutants are propagated to
the e subunit, affecting its trypsin digestion and cross-linking
pattern. In addition, a change in the e conformation was
previously seen, a gain by trypsin digestion and cross-linking,
following dicyclohexylcarbodiimide modification of cAsp61
[48], and a movement in the cytoplasmic loop of the isolated
c subunit, which is in contact with the e subunit in the whole
enzyme [12,14], had been detected by NMR spectroscopy on
protonation of cAsp 61 [23].

Finally, it should be considered that the ATP synthase of
Rb. c apsulatus had been shown t o undergo significant
D
~
ll
H
þ
-induced activation phenomena [49], whose possible
interplay w ith the catalytic activity is not yet c lear. Very
similar activation phenomena have recently been seen in the
E. coli enzyme [50]. S trikingly, in both cases, the Du
component was driving this activation significantly more
than the DpH, suggesting t he presence o f an allosteric Du
sensor in the transmembrane portion of F
0
. It cannot be
ruled out that the aGlu210 is part of this activation switch,
whose perturbation could lead both to a decrease in the
functional efficiency of the enzyme population and to a
different sensitivity towards DpH and Du, respectively, as
observed in the present work.
ACKNOWLEDGEMENTS
This work has been supported by t he grant PRIN/01, Processi
Ossidoriduttivi e Trasduzione di Energia in Membrane Procariotiche ed
Eucariotiche, from the Italian M inistery for E ducation of University
and Research (MIUR). We are grateful to G. Venturoli for many
discussions and to F. Federici and D. Giovannini for excellent help in
the experiments.
REFERENCES
1. Weber, J. & Senior, A .E. (1997) Catalytic mechanism of F

1
-
ATPase. Biochim. Biophys. Acta 1319, 19–58.
2. Boyer, P.D. (1998) Catalytic site forms and controls in ATP
synthase catalysis. Biochim. Biophys. Acta 1365, 3–9.
3. Fillingame, R.H., Jiang, W. & Dmitr iev, O.Y. (2000) Coupling
H
+
transport to rotary catalysis in F-type ATP synthases: struc-
ture and organization of the transmembrane rotary motor. J. Exp.
Biol. 203, 9–17.
4. Abrahams, J.P., Leslie, A.G., Lutter, R. & Walker, J.E. (1994)
Structure at 2.8 A
˚
resolution of F
1
-ATPasefrombovineheart
mitochondria. Nature 370, 621–628.
5. Yoshida, M., Muneyuki, E. & Hisabori, T. (2001) ATP synthase –
a marvellous rotary engine of the cell. Nat. Rev. Mol. Cell. Biol. 2,
669–677.
6. Girvin, M.E., Rastogi, V.K., Abildgaard, F., Markley, J.L. &
Fillingame, R.H. (1998) Solution structure of the transmembrane
H
+
-transporting subunit c of the F
1
F
0
ATP synthase. Biochem-

istry 37, 8817–8824.
7. Stock, D., Leslie, A.G. & Walker, J.E. ( 1999) Molecular archi-
tecture of the rotary motor in ATP synthase. Science 286, 1700–
1705.
8. Valiyaveetil, F.I. & Fillingame, R.H. (1998) Transmembrane
topography of subunit a in th e Escherichia c oli F
1
F
0
ATP
synthase. J. Biol. Chem. 273, 16241–16247.
9. Long, J.C., Wang, S. & Vik, S.B. (1998) Membrane topology
of subu nit a of the F
1
F
0
ATP synthase as d etermined b y
labeling of uniq ue cysteine re sidues. J. Biol. Che m. 273, 16235–
16240.
10. Wada, T., Long, J.C., Zhang, D. & Vik, S.B. (1999) A novel
labeling approach supports the five-transmembrane model of
subunit a of the Escherichia coli ATP synthase. J. Biol. Chem. 274,
17353–17357.
11. Jiang, W. & Fillingame, R.H. (1998) I nteracting helical faces of
subunits a and c in the F
1
F
0
ATP syn thase of Escherichia coli
defined by disulfide cross-linking. Proc. Natl Acad. Sci. USA 95,

6607–6612.
12. Zhang, Y. & Fillingame, R.H. (1995) Changing the ion binding
specificity of the Escherichia coli H
+
-transporting AT P s ynthase
by directed mutagenesis of subunit c. J. Biol. Chem. 27 0, 24609–
24614.
13. Watts, S.D., Tang, C. & Capaldi, R.A. (1996) The stalk region of
the Escherichia coli ATP s ynthase. Tyrosine 205 of the gamma
subunit is in the interface between the F
1
and F
0
parts and can
interact with bot h the epsilon and c oligome r. J. Biol. Chem. 271,
28341–28347.
14. Hermolin, J., Dmitriev, O.Y., Zhang, Y. & Fillingame, R.H.
(1999) Defining the domain of binding of F
1
subunit ep silon with
the polar loop of F
0
subunit c in the Escherichia coli ATP synthase.
J. Biol. Chem. 274, 17011–17016.
15. Dunn, S.D., McLachlin, D.T. & Revington, M. (2000) The second
stalk of Escherichia coli ATP synthase. Biochim. Biophys. Acta
1458, 356–363.
16. Capaldi, R.A., Schulenberg, B., Murray, J. & Aggeler, R. (2000)
Cross-linking and electron microscopy studies of the structure and
functioning of the Escherichia coli ATP synthase. J. Exp. Biol. 203,

29–33.
17. Sambongi, Y., Iko, Y., Tanabe, M., Omote, H., Iwamoto-Kihara,
A., Ueda, I., Y anagida, T., Wada, Y. & Futai, M. (1999)
Mechanical rotation of the c subunit oligomer in ATP s ynthase
(F
0
F
1
): direct observation. Science 286, 1722–1724.
Ó FEBS 2002 The aE210K mutation in Rb. capsulatus ATP synthase (Eur. J. Biochem. 269) 1991
18. Pa
¨
nke, O., Gumbiowski, K., Junge, W. & Engelbrecht, S. (2000)
F-ATPase: spec ific observation of the rotating c subunit oligomer
of EF
0
EF
1
. FEBS Lett. 472, 34–38.
19. Tsunoda, S.P., Aggeler, R., Yoshida, M. & Capaldi, R.A. (2001)
Rotation of the c subunit oligomer in fully functional F
1
F
0
ATP
synthase. Proc. Natl Acad. Sci. USA 98, 898–902.
20. Junge, W., Lill, H. & Engelbrecht, S. (1997) A TP s ynthase: an
electrochemical transducer with rotatory mechanics. Trends Bio-
chem. Sci. 22, 420–423.
21. Elston, T., Wang, H. & Oster, G. (1998) Energy transduction in

ATP synthase. Nature 391, 510–514.
22. Vik, S.B., Long, J.C., Wada, T. & Zhang, D. (2000) A model for
the structure of subunit a of the Escherichia coli ATP synthase and
its role in proton translocation. Biochim. Biophys. Acta 1458,457–
466.
23. Rastogi, V.K. & Girvin, M.E. (1999) Structural changes linked to
proton translocation by subunit c of the ATP synthase. Nature
402, 263–268.
24. Hartzog, P.E. & Cain, B.D. (1994) Second-site suppressor muta-
tions at glycine 218 and histidine 245 in the alpha subunit of F
1
F
0
ATP synthase in Escherichia coli. J. Biol. Chem. 269, 32313–32317.
25. Cain, B.D. & Simoni, R.D. (1988) Interaction between Glu-219
and His-245 within the a subunit of F
1
F
0
-ATPase in Escherichia
coli. J. Biol. Chem. 263, 6606–6612.
26. Borghese, R., Crimi, M., Fava, L. & Melandri, B.A. (1998) The
ATP s ynthase atpHAGDC (F
1
) operon from Rhodobacter cap-
sulatus. J. Bacteriol. 180, 416–421.
27. Borghese, R., Turina, P., Lambertini, L. & Melandri, B.A. (1998)
The atpIBEXF operon coding for the F
0
sector of the ATP syn-

thase from the purple nonsulfur photosynthetic bacterium Rho-
dobacter capsulatus. Arch. Microbiol. 170 , 385–388.
28. Valiyaveetil, F.I. & Fillingame, R.H. (1997) On the role of Arg-210
and Glu-219 of subunit a in proton translocation by the Escher-
ichia coli F
0
F
1
-ATP synthase. J. Biol. Chem. 272, 32635–32641.
29. Hatch, L.P., Cox, G.B. & Howitt, S.M. (1998) Glutamate residues
at positions 219 and 252 in the a-subun it of the Escherichia coli
ATP synthase are not functionally equivalent. Biochim. Biophys.
Acta 1363, 217–223.
30. Baccarini-Melandri, A. & Melandri, B.A. (1971) Partial resolution
of phosphorylating system of Rps. Capsulata. Methods Enzymol.
23, 556–561.
31. Keen, N.T., Tamaki, S., Kobayashi, D. & Trollinger, D. (1988)
Improved broad-host-range plasmids for DNA cloning in gram-
negative bacteria. Gene 70, 191–197.
32. Sharak Genthner, B.R. & Wall, J.D. (1985) Ammonium uptake in
Rhodopseudomonas capsulata. Arch. Microbiol. 141, 219–224.
33. Yen, H.C. & Marrs, B. (1976) Map of genes for carotenoid and
bacteriochlorop hyll biosynthesis in Rhodopseudomonas capsulata.
J. Bacteriol. 126, 619–629.
34. Gabellini, N., Gao, Z., Eckerskorn, C., Lottspeich, F. &
Oesterhelt, D. (1988) Purification of the H
+
-ATPase from Rho-
dobacter capsulatus, identification of the F
1

F
0
components and
reconstitution of the active enzyme. Biochim. Biophys. Acta 934,
227–234.
35. Melandri, B.A., De Santis, A., Venturoli, G. & Baccarini-
Melandri, A. (1978) The rates of onset of photophosphorylation
and of the protonic electrochemical potential difference in chro-
matophores. FEBS Lett. 95, 130–134.
36. Turina, P., Melandri, B.A. & Gra
¨
ber, P. (1991) ATP synthesis in
chromatophores driven by artificially induced ion gradients. Eur.
J. Biochem. 196, 225–229.
37. Casadio, R. (1991) Measurements of transmembrane pH differ-
ences of low extents in bacterial chromatophores Eur. Biophys. J.
19, 189–201.
38. Taussky, M. & Schorr, J. (1953) A m icrocolorimetric method for
the determination of inorganic phosphate. J. Biol. Chem. 202,675–
686.
39. Lanzetta, P.A., Alvarez, L.J., Reinach, P.S. & Candia, O.A. (1979)
An improved assay for nanom ole amounts of inorganic phos-
phate. Anal. Biochem. 100, 95–97.
40. Clayton, R.K. (1963) Toward the isolation of a photoche mical
reaction center in Rhodopseudomonas sphaeroides. Bioc him. Bio-
phys. Acta 75, 312–323.
41. Jackson, B.J. & Nicholls, D.G. (1986) Methods for the determ ina-
tion of membrane p otential in bioenergetic s ystems. Methods
Enzymol. 127, 557–577.
42. Barz, W.P., Francia, F., Venturoli, G., Melandri, B.A., Vermeglio,

A. & Oesterhelt, D. (1995) Role of PufX protein in photosynthetic
growth of Rhodobacter sphaeroides. 1. PufX is required for efficient
light-driven electron transfer and p hotophosphorylation u nder
anaerobic conditions. Biochemistry 34, 15235–15247.
43. Petitou, M., Tuy, F. & Rosenfeld, C. (1978) A simplified proce-
dure for organic phosphorus dete rmination fro m phospho lipids.
Anal. Biochem. 91, 350–353.
44. Kaim, G. & Dimroth, P. (1999) ATP synthesis by F-type ATP
synthase is obligatorily dependent on the transmembrane voltage.
EMBO J. 18, 4118–4127.
45. Slooten, L. & V andenbranden, S. (1989) ATP-synthesis by
proteoliposomes inc orporat ing Rhodospirillum rubrum F
0
F
1
as
measured with firefly luc iferase: dependence on delta psi and delta
pH. Biochim. Biophys. Acta 976, 150–160.
46. Fischer, S ., Etzold, C., T u rina, P., Deckers-Hebestreit, G.,
Altendorf, K. & Gra
¨
ber, P. (1994) ATP synthesis catalyzed by the
ATP synthase of Escherichia coli reconstituted into liposomes.
Eur. J. Biochem. 225, 167–172.
47. Gardner, J.L. & Cain, B.D. (1999) Amino acid substitutions in the
a subunit affect the epsilon subunit of F
1
F
0
ATP synthase from

Escherichia coli. Arch. Biochem. Biophys. 361, 302–308.
48. Mendel-Hartvig, J. & Capaldi, R.A. (1991) Nucleotide-dependent
and dicyclohexylcarbodiimide-sensitive conformational changes in
the epsilon subunit of Escherichia coli ATP synthase. Biochemistry
30, 10987–10991.
49. Turina, P., Rumberg, B., Melandri, B.A. & Gra
¨
ber, P. (1992)
Activation of the H
+
-ATP syn thase in the ph otosynthetic bacter-
ium Rhodobacter capsulatus. J. Biol. Chem. 267, 11057–11063.
50. Fischer, S., Gra
¨
ber, P. & Turina, P. (2000) The activity of the ATP
synthase from Escherichia coli is regulated by the transmembrane
proton motive force. J. Biol. Chem. 275, 30157–30162.
1992 P. Turina and B. A. Melandri (Eur. J. Biochem. 269) Ó FEBS 2002