Inner membranes prepared from the mutant strain showed approximately half of the ATP synthesis activity when driven both by light and by acid-base transitions.. ATP synthesis induced by
Trang 1A 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 in the F0sector of the ATP synthase is believed to
form the core of 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 high degree of
conservation, among these aGlu219 The homologous
resi-due aGlu210 in the ATP synthase of the 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 acid-base transitions As
estimated with the ACMA assay, proton pumping rates in
the inner 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, was observed in the absence of a transmem-brane electrical memtransmem-brane potential (Du) at low trans-membrane pH difference (DpH) Therefore, the mutation seems to affect both the mechanism responsible for coupling
F1 with proton translocation by F0, 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 F0F1-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 some species) The
membrane-embedded hydrophobic F0 sector is involved in proton
translocation across the membrane, and the hydrophilic F1
sector contains the catalytic sites (reviewed in [1–3]) A
wealth of high resolution structural information for the
soluble part has appeared since the first crystal structure of
the mitochondrial F1was 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 coli enzyme, F1
consists of five types of subunits in stoichiometry a3b3cde
and F0consists of three types of subunits in stoichiometry
ab2c9)12 The c 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 a3b3 subunit barrel in F1, it is proposed that the c subunit ring in F0, which is connected to the ce shaft [12–14], rotates against the a subunit, which is connected to the a3b3barrel 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 F0
have been proposed, which emphasize the role of electro-static interactions [20–22] or the role of conformational 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 the membrane, the remaining a/c interface regions are believed
to form the access pathways for protons Probably 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 aGlu219 and aHis245, including the fact 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 aGlu219fi His/aHis245fi 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 these 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: melandri@alma.unibo.it
Abbreviations: GTA, gene transfer agent; Bchl, bacteriochlorophyll;
ACMA, 9-amino-6-chloro-2-methoxyacridine; RC, photosynthetic
reaction center; ~ lHþ , 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 http://www.biologia.unibo.it/
(Received 12 November 2001, revised 21 February 2002, accepted 21
February 2002)
Trang 2[25], their important functional role is indicated both by
their high degree of conservation and by the deleterious
effects of their mutations on the E coli ATP synthase
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 colienzyme [26,27], except that the additional subunit
b¢, homologous to b, is found in F0, as it is typical for
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 aGlu210fi Lys and
have investigated detailed functional aspects of the ATP
synthase in native membranes The corresponding mutation
in the E coli enzyme, aGlu219fi Lys, was examined in
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 results of
the present work are consistent with the data obtained in
E coliand reveal novel functional properties of the mutated
enzyme
E X P E R I M E N T A L P R O C E D U R E S
Bacterial strains, growth conditions, membrane
preparations
Rb capsulatusstrain B100 is wild-type strain B10 cured of
phages A spontaneous rifampicin-resistant of B100, J1,
was used in the GTA1 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 each
carrying nine 100-W incandescent light bulbs; excessive
warming was prevented by water cooling The mutant and
wild-type strain were grown in parallel and cells were
harvested at D600 ¼ 1.2–1.4 Intra-cytoplasmic
mem-branes (chromatophores) were prepared according to the
method described previously [30], resuspended in 50 mM
glycyl-glycine/NaOH, 5 mM MgCl2, pH 7.5, rapidly
frozen as small droplets in liquid nitrogen, and stored at
)80 °C
Subunit a mutagenesis
The wild-type copy of subunit a was carried by the pFo16
plasmid which contained the whole atp2 operon of
Rb capsulatuscloned into the pTZ19U plasmid [27] The
aE210fi K mutation was introduced into this plasmid by
using the QuickChange Site-Directed Mutagenesis Kit
(Stratagene), based on linear PCR, using the following
mutagenic oligonucleotides: 5¢-CGCGATGTATGCGC
TCAAGATCCTCGTGGCC-3¢ and 5¢-GGCCACGAGG
ATCTTGAGCGCATACATCGCG-3¢ The mutation
introduced an additional restriction site for XhoII, therefore
its presence 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 capsulatus cells which pack, randomly, pieces of DNA about 4.6-kb long, either from the chromosome or from resident plasmids of donor cells, and transfer them to acceptor cells, where they are integrated into the chromosome by homologous recombination This results in an 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 pFo39 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 genes (subunits
a, c, b¢, b) leaving only a C-terminal truncated atpI gene (subunit i) After GTA transfer, kanamycin-resistant colo-nies appeared that could contain the kanamycin resistance cassette (i.e the F0deletion) 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 F0deletion on the chromosomes Two mutated strains were selected, which carried mutated plasmids originated from two different PCR runs A pseudo-wild-type strain was constructed in parallel, which carried the wild-type F0 operon on the plasmid and the deletion of the chromosomal F0operon 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 SDS/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°C in the following buffer: 100 mM glycyl-glycine/NaOH,
pH 7.75, 50 mM KCl, 10 mM Mg-acetate, 0.1% bovine serum albumin, 5 mM Pi, 0.2 mM succinic acid, 0.8 mM
AMP, 10 lM Bchl The chromatophores suspension was illuminated from two opposite sides by two 100 W incandescent bulbs The reaction was started by addition
of 100 lM ADP After stopping the reaction with 7% perchloric acid, the ATP concentration in each sample was measured in a luminometer (LKB 1250) 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
Trang 3Fig 1, the cuvette contained 20 lM ADP, 60 lM
lucif-erine and 2–10 lgÆmL)1 purified luciferase (32–160·
103light unitsÆmL)1) from Sigma (L9009) in the reaction
mixture described above, and the luminescence was
detected in real-time at 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 88 A
Wratten filters) 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 synthetized ATP was evaluated by adding
10–25 nM 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 luminometer cuvette containing a basic solution and monitoring the ATP concentration continuously with luciferine/luciferase in the luminometer The chromatophores were first resuspended
in 5 mM Pi/NaOH, pH 7.3, 2 mM MgCl2, 1 mM AMP,
5 lM valinomycin, 10% sucrose, and either 1 mM KCl (+Du) or 100 mM 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 disappearance of the K+/valinomycin diffusion potential (monitored by the carotenoid shift signal) induced by the initial K+ gradient [36] The chromatophores were then mixed with the acidic solution [30 mM succinic acid/NaOH, pH 4.6–6.5, 2 mM MgCl2,
5 mMPi, either 1 mM(+Du) or 100 mMKCl (–Du), 1 mM
AMP, 5 lMvalinomycin] and incubated at room tempera-ture at variable times between 2 and 30 min, depending on the pH of the suspending buffer, prior to injection of
100 lL into the basic solution This latter contained
850 lL of basic solution so that the final concentrations after chromatophores addition would be 200 mMTricine,
2 mM MgCl2, 5 mM Pi, either 150 mM KOH + 30 mM
NaOH (+Du) or 100 mMKOH + 80 mMNaOH (–Du),
pH 8.65, 1 mM AMP, 100 lM ADP, 50 lL of the ATP-Monitoring Kit and 5 lgÆmL)1 of purified luciferase (80· 103light unitsÆmL)1) The final Bchl concentration varied between 1 and 8 lM The ATP concentration was evaluated by adding 100–200 nM standard ATP in each cuvette The basic solution was thermostated so that the ATP synthesis reaction took place at 13°C The pH measured after mixing the chromatophores 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 For the lowest DpH differences the ÔacidicÕ solution contained 20 mM Tricine instead of succinic acid Assuming complete equilibration
of the K+ during the 1 h preincubation (see above), the value of the 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 based 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 mM and [K+]in/[K+]out ¼ 100/100 mM, respectively
ACMA assay ACMA fluorescence quenching assay was carried out in a Jasco FP 500 spectrofluorimeter (wavelength 412 and
482 nm for excitation and emission, respectively) at 15°C
in the following mixture: 20 mM Tricine/KOH, pH 8.0,
50 mM KCl, 0.5 mM MgCl2, 0.2 mM succinic acid, 5 lM
Antimycin, 0.2 lMmyxothiazol, 2 lMvalinomycin, 1.5 lM
ACMA, 20 lMBchl, 400 lMATP The response of ACMA
to DpH was empirically calibrated using artificially induced protonic gradients, established by HCl and NaOH addi-tions, under similar temperature and buffer condiaddi-tions, except for the presence of 20 mMsuccinic acid, as described previously [37]
Fig 1 ATP synthesis in chromatophores induced 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 following a short period of illumination has been
mon-itored The reaction assays, described in the Experimental procedures
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 have been fitted with an arbitrary function The dashed
lines extrapolate the linear part of the curves (C) The first derivatives
of the fitting functions were calculated between 0 and 1000 ms
(extrapolating the fitting 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 experiments in the absence (dashed
line) or in the presence (continuous line) of valinomycin As no ATP
synthesis could be detected with added valinomycin for short
illumin-ation times, the ratio function was truncated for times £ 150 ms.
Trang 4ATP hydrolysis assays
ATP hydrolysis was measured routinely at 30°C in the
following buffer: 20 mM Tricine/KOH, pH 8.0, 50 mM
KCl, 2 mMMgCl2, 0.2 mMsuccinic acid, 20 lMBchl The
reaction was started by adding 1 mMATP After stopping
the reaction at different times with 5% trichloroacetic acid,
the Pi concentration was measured by molybdate
colori-metric assay as described previously [38] For more sensitive
measurements, the released Pi was measured with the
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 chromatophores
was determined using the BCA assay (Pierce) An aliquot
of the chromatophores was extracted with
acetone/meth-anol (7 : 2, v/v) After centrifugation, the protein pellet
was dissolved in 0.1MNaOH/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 concentration of photo-oxidizable reaction centre
(RC) and of total photo-oxidizable cytochrome (c1+ c2)
were measured as described previously [42], following
trains of closely spaced flashes The amount of
phosp-holipid was determined by the method described
previ-ously [43]
R E S U L T S
The single point mutation aE210K was introduced into the
atp2 operon of Rb capsulatus, containing the F0 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 allowed to
take place, which generated the deletion of 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 atp2-deleted strain was created, in which
the resident plasmid carried the wild-type operon This
pseudo-wild-type strain is referred to as wild-type in the
following procedures
Characterization of mutant chromatophores
Phototrophic growth of the aE210K mutant cells was
slower than the wild-type cells Accordingly, the
light-induced ATP synthesis rate catalyzed by the mutant
chromatophores was about 40% lower on a Bchl basis
than the rate catalyzed by wild-type chromatophores
(Table 1) The same reduction resulted also when the
ADP concentration was varied between 20 and 500 lM,
indicating that the mutation does not affect the apparent Km
for ADP In contrast, no significant difference could be
observed in the ATP hydrolysis rate 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 synthesis was
8 ± 2 and 13 ± 3 ATP/(F1F0Æs) for mutant and wild-type, respectively, whereas the specific ATP hydrolysis activities were 2.0 ± 0.7 and 2.3 ± 1.0 ATP/(F1F0Æs) 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 or of right-side-out vesicles in the mutant was also ruled out by measuring the extent of flash-induced carotenoid shift and the extent of cyto-chrome c accessible photo-oxidizable RC, which were comparable to those of wild-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 antenna (Bchl/RC) and the phospholipid content were very similar in both strains The most striking difference was found in the protein content, which was approximately 1.6-fold lower in the mutant chromatophores on a Bchl basis It is possible that this difference affects the adsorption
of ACMA to the membrane and therefore the 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 single 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 Rateb 13 ± 3 11 ± 2 (mM P i /M Bchl/s)
Bchl/ATP Synthase Ratio c 178 ± 35 180 ± 31 (moles/mole)
(mg/lmole)
(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.bThe
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.
Trang 5Light-induced ATP synthesis at low D~lHþ
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~lHþ 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~lHþvalues, 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~lHþ, 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 the assay cuvette at high
concentration (32–160· 103light unitsÆmL)1of luciferase),
so that the ATP-induced luminescence was directly detected
Figure 1A shows the ATP yields obtained from mutant
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 When the assay contained
valinomycin, the data reported in Fig 1B 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 already been
observed in wild-type chromatophores [35] and can be
interpreted as being due to a lag phase in the onset of D~l þ
when its electrical component is being dissipated by valinomycin
Strikingly, this lag phase was much more pronounced in the case of the mutant The large difference relative to the wild-type can be best appreciated by taking the derivative of the fitting functions of the mutant and wild-type data and plotting their ratio, as in Fig 1C This derivative represents the rate of ATP synthesis at each time point The twofold difference between the mutant and wild-type observed in the presence of a Du is approached only 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~lHþ 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 monitored continuously with the luciferine/luciferase
ATP Monitoring Kit in a luminometer The high signal-to-noise ratio
was obtained due to added purified 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 diffusion potential
(+Du) Bchl concentration in the luminometer cuvette was 1.1 l M
The reaction temperature was 13 °C Oligomycin was added during
the preincubation time at a concentration of 20 lgÆmL)1and 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 the absence of a diffusion potential (A) Data are obtained from measurements similar to those reported in Fig 2 The composition 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 adenylate kinase rate has been subtracted Data are averages of 2–3 measurements (d,m) wild-type and (s,n) mutant chromatophores Rates were measured both in the presence (d,s) and in the absence (m,n) of a diffusion 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) The ratio of the best-fitting functions and of the data points of wild-type over mutant are plotted for data in the presence (d) and absence (m) of Du.
Trang 6membranes 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+]inand [K+]outfor +Du was 1 and
150 mM, respectively, and the [K+]inand [K+]outfor –Du
was 100 and 100 mM, respectively The DpH values were
varied up to 3.8 units by decreasing the internal pH at
constant external pH Rb capsulatus 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 mM (–Du condition), the
absence of any significant diffusion potential due to other
ionic species, e.g monoionic succinate [44] was excluded
observing the electrochromic spectral shift of carotenoids
We therefore conclude that in chromatophores of Rb
cap-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~lHþ-induced ATP synthesis decays within a few
hundred milliseconds [36], due to the relatively high
permeability of the 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 duration 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 basic solution and
the luciferine/luciferase detection kit (plus additional
80· 103light unitsÆmL)1of 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~lHþ-induced
by the acid-base transition The slow increase of ATP
concentration seen thereafter was uncoupler- (not shown)
and oligomycin-insensitive, and can be attributed to the
adenylate kinase activity 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, the ATP
synthesis rates of the mutants were 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 rates
obtained in the absence of a Du In this case, the impairment
of the ATP synthesis rates in the 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 fitting functions of wild-type over
mutant, either in the presence or in the absence of a Du, as
shown in Fig 3C 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 ranging between 2.4 and 3.3,
whereas a twofold factor was approached at the highest
DpH tested The trace –Du in Fig 3C can be directly
compared to the +valinomycin trace in Fig 1C, because, in the absence of a Du, the DpH 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 initial rates of quenching, the 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 in Fig 4A for chromatophores of both strains No 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 response turned out to be significantly different for the mutant and wild-type chromatophores (Fig 4B) This different response was systematically found
in different experimental sessions and in 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 with 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 the 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 proton translocation (ATP slipping) The reason for this different 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]
D I S C U S S I O N
The main finding of this work is that a single point mutation can affect the ATP synthesis rate of 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 collapsed 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 of 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 ATP
Trang 7synthesis rate was much 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 pHinrange strongly suggests that a step in the ATP
synthase functioning, which is rate-limiting at the low H+in
concentrations, gradually ceases to be rate-limiting for
increasing H+inconcentrations It is this rate-limiting step,
apparently, which is strongly affected in the mutant What
could be rate-limiting at the 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 F0 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 the bulk
aqueous compartments to the cAsp61 in the middle of the
membrane In these models, an electrostatic constraint
implies that carboxyl groups on the ring are always
protonated (electroneutral) when facing the lipid phase,
while they can be unprotonated and charged when facing
the a subunit The two half-channels are spatially offset, so
that this electrostatic constraint imposes a directionality to
the c-oligomer rotational fluctuations According to the
model proposed by Oster and coworkers [21], the rate
constant kinfor proton hopping into the channel from the periplasmic side depends upon [H+]in and upon the potential drop Dw due to surface charges, i.e kina10–pHin
exp(Dw/RT) As a Glufi Lys substitution is expected to change the electrostatic profile of the nearby region, a possible explanation for the data presented is that residue a210 (a219 in E coli) belongs to the periplasmic half-channel and that one of its roles is to create a favorable electrostatic profile for 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 become 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 of
H+release at the cytoplasmic side and a decreased rate of
H+ release at the periplasmic side Particularly this last effect can presumably compensate for the low bulk H+in
concentration by effectively increasing [H+] within 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 potential drop Dw at the entrance of this half-channel
Fig 4 Proton pumping and ATP hydrolysis The composition of the reaction mixtures are detailed in the Experimental procedures ATP and Bchl concentrations were 400 and 20 l M , respectively The temperature 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 by the addition of 2 l M valinomycin (d) wild-type and (m) mutant chromatophores (A) ACMA fluorescence quenching as a function of time after addition of ATP The original recorder traces have been digitalized and data points have been fitted with arbitrary exponential functions (B) Calibration of the ACMA fluorescence response to transmembrane pH differences The ACMA fluorescence quenching has been measured after imposing various DpH jumps The reaction mixture was identical to that in Fig 4A, except that 20 m M succinate was added to increase 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, the released P i was analyzed with the EnzCheck Phosphate Assay The rates of ATP hydrolysis resulted 4.7 m M ATP/(M BchlÆs) for both wild-type and the E210K mutant.
Trang 8A second major effect of the aE210K mutation is the
generally lower ATP synthesis found regardless of D~lHþ
extent and composition, which might also explain the lower
growth rate, as found here and in E coli strains carrying
the homologous aE219K mutation [28,29] This general
impairment of ATP synthesis amounted here
approxi-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), as well as for ATP
synthesis driven by acid-base transitions, both in the
presence of a Du and in its absence at high H+in
concentrations (Fig 3C)
Several hypotheses can be put forward to explain this
impairment One possibility is that the mutation perturbs
the electrostatic profile sensed by the unprotonated Asp61
during its radial translocation, thereby increasing the
probability of protons leaking through F0without
perform-ing any work [21] This perturbation would also reduce the
H+ pumping efficiency, in line with the conclusions
suggested by our (Fig 4) and other [28,29] data Evidence
for ATP slipping caused by the aE210K mutation, if
confirmed, would be an independent element for assessing
the role of Glu210 in the proton pathway within F0
Alternatively, it can be that the mutation has an even
longer-range effect, affecting the conformation at the
interface between the c-ring and the ce shaft The
conformational modification of this 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 F1has not yet been
seen, such long-distance effect would presumably occur
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 aR210fi I,
aA217fi R, and aG218 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, again 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 capsulatus had been shown to undergo significant
D~lHþ-induced activation phenomena [49], whose possible
interplay with the catalytic activity is not yet clear Very
similar activation phenomena have recently been seen in the
E coli enzyme [50] Strikingly, in both cases, the Du
component was driving this activation significantly more
than the DpH, suggesting the presence of an allosteric Du
sensor in the transmembrane portion of F0 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
A C K N O W L E D G E M E N T S
This work has been supported by the grant PRIN/01, Processi
Ossidoriduttivi e Trasduzione di Energia in Membrane Procariotiche ed
Eucariotiche, from the Italian Ministery for Education 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.
R E F E R E N C E S
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 & Dmitriev, 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 -ATPase from bovine heart 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 the Escherichia coli 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 subunit a of the F 1 F 0 ATP synthase as determined by labeling of unique cysteine residues J Biol Chem 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) Interacting helical faces of subunits a and c in the F 1 F 0 ATP synthase 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 ATP synthase
by directed mutagenesis of subunit c J Biol Chem 270, 24609– 24614.
13 Watts, S.D., Tang, C & Capaldi, R.A (1996) The stalk region of the Escherichia coli ATP synthase Tyrosine 205 of the gamma subunit is in the interface between the F 1 and F 0 parts and can interact with both the epsilon and c oligomer 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 epsilon 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., Yanagida, T., Wada, Y & Futai, M (1999) Mechanical rotation of the c subunit oligomer in ATP synthase (F 0 F 1 ): direct observation Science 286, 1722–1724.
Trang 918 Pa¨nke, O., Gumbiowski, K., Junge, W & Engelbrecht, S (2000)
F-ATPase: specific 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) ATP synthase: 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 synthase 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-subunit 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
bacteriochlorophyll 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 microcolorimetric 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 nanomole amounts of inorganic phos-phate Anal Biochem 100, 95–97.
40 Clayton, R.K (1963) Toward the isolation of a photochemical reaction center in Rhodopseudomonas sphaeroides Biochim Bio-phys Acta 75, 312–323.
41 Jackson, B.J & Nicholls, D.G (1986) Methods for the determina-tion of membrane potential in bioenergetic systems 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 photophosphorylation under anaerobic conditions Biochemistry 34, 15235–15247.
43 Petitou, M., Tuy, F & Rosenfeld, C (1978) A simplified proce-dure for organic phosphorus determination from phospholipids 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 & Vandenbranden, S (1989) ATP-synthesis by proteoliposomes incorporating Rhodospirillum rubrum F 0 F 1 as measured with firefly luciferase: dependence on delta psi and delta
pH Biochim Biophys Acta 976, 150–160.
46 Fischer, S., Etzold, C., Turina, 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 synthase in the photosynthetic 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.