From CREST "Genetic Programming" Team 13, Teikyo
University Biotechnology Research Center 3F, Nogawa 907, Miyamae-ku,
Kawasaki 216-0001, Japan, § PRESTO, Chemical Resources
Laboratory, Tokyo Institute of Technology, Yokohama 226-8503, Japan,
** Tsukuba Research Laboratory, Hamamatsu Photonics KK,
Tokodai, Tsukuba 300-2635, Japan,
Chemical Resources
Laboratory, Tokyo Institute of Technology, Yokohama 226-8503, Japan,
and §§ Faculty of Science and Technology, Keio University,
Yokohama 223-8522, Japan
Received for publication, March 12, 2001, and in revised form, March 28, 2001
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
The binding change model for the
F1-ATPase predicts that its rotation is intimately
correlated with the changes in the affinities of the three catalytic
sites for nucleotides. If so, subtle differences in the nucleotide
structure may have pronounced effects on rotation. Here we show by
single-molecule imaging that purine nucleotides ATP, GTP, and ITP
support rotation but pyrimidine nucleotides UTP and CTP do not,
suggesting that the extra ring in purine is indispensable for proper
operation of this molecular motor. Although the three purine
nucleotides were bound to the enzyme at different rates, all showed
similar rotational characteristics: counterclockwise rotation, 120°
steps each driven by hydrolysis of one nucleotide molecule, occasional
back steps, rotary torque of ~40 piconewtons (pN)·nm, and
mechanical work done in a step of ~80 pN·nm. These latter
characteristics are likely to be determined by the rotational mechanism
built in the protein structure, which purine nucleotides can energize.
With ATP and GTP, rotation was observed even when the free energy of
hydrolysis was The FoF1 ATP synthase is an enzyme that
synthesizes ATP from ADP and inorganic phosphate (Pi) using
proton flow across a membrane (1). The Fo portion of the
enzyme resides in the membrane and mediates proton translocation. The
F1 portion, consisting of
For the coupling between the proton flow in Fo and the
chemical reaction (ATP synthesis/hydrolysis) in F1, Boyer
and Kohlbrenner (3) and Oosawa and Hayashi (4) independently suggested
a rotational catalysis model. The essence is that Fo is a
rotary motor (or turbine) driven by the proton flow, that
F1 is another rotary motor driven by ATP hydrolysis, and
that the two motors have a common rotary shaft, yet their genuine
rotary directions are opposite to each other's. Rotation of the shaft
in Fo's genuine direction, as occurs in cells, results in
the reverse rotation of the F1 motor and thus in ATP
synthesis in F1. When F1 gains control and
hydrolyzes ATP, protons are pumped through Fo in the reverse direction.
A crystal structure of F1 (5) strongly supported the
rotation model and has inspired many experiments, which, together, have
proved that the In Boyer's model, binding changes play the major role (1). On the
three If binding and release, rather than the cleavage of the terminal
phosphate, are indeed the source of mechanical torque production in the
F1 motor, the rotational characteristics are expected to differ among different NTPs. The free energy gain in overall
hydrolysis is similar among NTPs, but the operation of the
F1 motor, which can work at near 100% efficiency, may well
depend on delicate energy balances in various steps of hydrolysis. Here
we show that purine but not pyrimidine nucleotides support
F1 rotation, suggesting that the interaction between the
catalytic site and the additional ring in purines is critical to the
proper operation of this molecular machine.
Chemicals and Proteins--
Nucleotides (GTP, ITP, UTP, and CTP)
were purchased from Sigma. ATP and other enzymes were from Roche
Molecular Biochemicals. The purity of each nucleotide was assessed on
an anion exchange column. Nucleotides were applied on DEAE 2SW (Tosoh,
Japan) equilibrated with 50 mM sodium phosphate (pH 7.0)
and eluted with an isocratic flow of the same buffer. Because DEAE 2SW
has higher affinity for compounds with more negative charges,
nucleotides are eluted in the order of nucleoside mono-, di-, and
triphosphate. The elution profiles for UTP are shown in Fig.
1, a and b. Besides
the UDP peak at 13 ml (identical with the peak for commercial UDP),
several contaminant peaks (C1-C5) were resolved. The void peak at 4 ml is unlikely to be a nucleotide(s), because A260
(green) and A280 (red),
within an absorbance peak of a nucleotide, were extremely low compared
with A220 (blue). C1 at 7 ml was
probably UMP, because it was eluted earlier, and the relative
magnitudes of A220, A260 and A280 were the same as those for UTP and UDP.
C4 was judged as contaminating GTP, because its elution volume and the
relative magnitudes of A220,
A260, and A280 all
matched with those for GTP (Fig. 1c). The GTP contamination
amounted to 0.01% of UTP, as determined from the peak area of
A260. The other contaminants, C2, C3, and C5,
were of unknown origin; their signatures did not match with those of
ATP, GTP, ITP, dATP, or their diphosphates. C2, C3, and C5 in UTP
amounted to 0.03, 0.01, and 0.39%, respectively. Although the levels
of contaminants were low, they may nevertheless affect the rotation
assay, because the affinity of F1-ATPase for UTP is
extremely low (see "Results"). We therefore used the fraction at
the UTP peak (23 ml) for the rotation assay. CTP was also found to be
contaminated by unknown compounds (1.5%); thus, its peak fraction was
used for the rotation assay. ATP, GTP, and ITP contained less than 5%
contaminants, which were mostly their hydrolysis products; these were
used without purification.
The NTPase Activity--
ATP, GTP, ITP, and UTP hydrolysis
activities of Rotation Assay--
Streptavidin-conjugated TF1 for
the rotation assay was prepared as described previously (9). Rotation
in the presence of ATP, GTP, or ITP was observed in the TF1
adsorbed on a Ni2+-NTA-modified polystyrene bead by
attaching an actin filament to the Proton Pump by Reconstituted
TFoF1--
Nucleotide-driven
proton-translocation into TFoF1 liposomes was
detected as the decrease in the pyranine fluorescence in an FP 777 fluorometer (JASCO, Japan) at 23 °C. TFoF1
liposomes (about 5 µg of TFoF1) containing
pyranine inside were preincubated at 23 °C in 1.5 ml of a reaction
mixture containing 10 mM MOPS-KOH (pH 7.0), 50 mM KCl, 2 mM nucleotide, 25 mM
K2SO4, 25 mM
Na2SO4, and 20 mM
p-xylene-bispyridinium bromide, which quenched the pyranine fluorescence outside the liposomes. Excitation and emission wavelengths were 460 and 510 nm, respectively. The reaction was started by the
addition of 4 mM MgCl2. After recording the
fluorescence change, 10 µM FCCP was added to recover the
initial fluorescence, which would ensure that the fluorescence decrease
was caused by the proton uptake into the liposomes.
Nucleotide-depleted
F1-ATPase--
F1-ATPase is known to be
inhibited by Mg-ADP tightly bound to a catalytic site (15, 16).
TF1 used in the previous study (10) contained ~0.3 mol of
tightly bound nucleotide/mol of enzyme; thus, the hydrolysis activity
might have been underestimated. Here, we employed an improved
purification protocol (see "Experimental Procedures") and obtained
a sample containing only 0.084 mol of bound nucleotide (0.037 mol of
ATP and 0.047 mol of ADP) per mol of enzyme as determined from the peak
heights in the reverse-phase chromatography (Fig.
2a). Before use, the protein
was further purified by size exclusion chromatography, because
TF1 tended to aggregate upon storage. For the final sample,
the ratio of A280 to
A260, a convenient measure of the purity, was
greater than 2.0 (Fig. 2b). As expected, the ATP hydrolysis
activity of this nucleotide-depleted enzyme was higher than that
reported previously (see below).
Hydrolysis Activities--
ATP, GTP, ITP, and UTP hydrolysis
activities of the nucleotide-depleted TF1 were determined
as the rate of oxidation of NADH using the ATP-, GTP-, ITP-, or
UTP-regenerating system. Hydrolysis of the purine nucleotides, ATP,
GTP, and ITP, gradually decelerated to a steady state as shown in Fig.
3, a and b. This
turnover-dependent inactivation is attributed to the Mg-ADP
(GDP, IDP) inhibition. The hydrolysis rates were therefore determined
during the initial 10 s after the injection of the enzyme into the
assay mixture (Fig. 3, a and b,
insets).
The rate of ATP hydrolysis did not obey simple Michaelis-Menten
kinetics and was fitted with two sets of Km and
Vmax: Vmaxa = 87.5 s
In contrast to the purine nucleotides, UTP hydrolysis activity
gradually increased with time (Fig. 3, a and b).
This cannot be ascribed to the slow response of the UTP-regenerating
system, because it could generate UTP from UDP much faster (0.5 s
As seen in Fig. 3c, the maximal hydrolysis rates for the
four nucleotides, ATP, GTP, ITP, and UTP, do not differ greatly. The
major difference among the four is in the apparent
Km values, UTP being the highest. In this regard, it
is possible that CTP is also hydrolyzed with a similar
Vmax but with a Km far above
10 mM. Another difference is that ATP hydrolysis
shows significant deviation from the simple Michaelis-Menten kinetics, which is not apparent for the other nucleotides. This may be correlated with the stronger tendency toward inhibition with ATP than with the
other nucleotides (Fig. 3, a and b). The
possibility that the other nucleotides also show deviation at much
higher concentrations (giving possibly the same
Vmax) cannot be dismissed.
GTP- and ITP-driven Rotation--
GTP and ITP supported the
rotation of the
Because the F1 motor has been shown to be a 120° stepper
(10), we compare 3 times the rotational rate of a short actin filament (0.5-1.3 µm) with the corresponding hydrolysis rate (Fig. 3). At low
nucleotide concentrations ([ATP] < 0.1 µM, [GTP] < 1 µM, [ITP] < 10 µM) where nucleotide
binding is rate-limiting, the two rates agreed with each other,
indicating that three molecules of purine nucleotide, whether ATP, GTP,
or ITP, drive a full turn. The agreement also shows that the rotation
in the presence of GTP or ITP was not due to possible ATP
contamination, which was far below 5%. At higher nucleotide
concentrations, the rotational rate saturated around 4 revolutions
s
As seen in Fig. 4a, back steps occurred occasionally in
GTP-driven rotation. ITP-driven rotation also showed occasional back steps (data not shown). The velocity of back steps was as fast as the
forward one and thus is unlikely to be of purely thermal origin (10). A
stochastic mistake in the order of substrate binding or product release
among the three catalytic sites could explain the back steps.
UTP Does Not Drive
If hydrolysis of each UTP molecule produces a 120° step as with
purine nucleotides, the rotational rate is expected to be 4.4 s Torque and Efficiency--
To compare torque and energy conversion
efficiency among three purine nucleotides, ATP, GTP, and ITP, that
supported
The torque of 40 pN·nm times the angular displacement of 2 Proton Pump--
Nucleotide-driven proton uptake by the
reconstituted TFoF1 liposome was measured, to
see whether In this paper, we have shown that the purine ring of a nucleotide
is indispensable for Our rotation assay has revealed that many of the rotational
characteristics are common to all three purine nucleotides. (i) The
rotary torque is constant around 40 pN·nm. (ii) The rotation consists
of discrete 120° steps. (iii) Each 120° step is driven by the
hydrolysis of one nucleotide molecule. (iv) Backward steps as fast as
the forward ones occur occasionally. The first three points imply that
the mechanical work done in a 120° step is about 80 pN·nm for all
three nucleotides, and thus the energy conversion efficiency can reach
~100% with all three purine nucleotides. As to the last statement,
we have shown in this study that the mechanical work of ~80 pN·nm
is done even when Chemical kinetics, in contrast, are different among the three purine
nucleotides. First, the hydrolysis curves in Fig. 3c are
shifted toward higher concentrations in the order of ATP, GTP, and ITP,
and the rotation curves follow the same trend. Apparently, this is due
to the difference in the rate of nucleotide binding, ATP being the
fastest and ITP the slowest. Analysis of step intervals at low
nucleotide concentrations supported this view; the rate constant was
2.5 × 107 M According to the binding change mechanism for ATP synthase (1), much of
the energy available from A second difference in the chemical kinetics is that the ATP hydrolysis
was described by two sets of Km and
Vmax, whereas one set sufficed for GTP and ITP.
The two sets for the case of ATP are usually ascribed to two modes of
catalysis: bisite and trisite, where one or two, or two or three,
respectively, of the three catalytic sites are alternately filled with
a nucleotide. The possibility, however, exists that the enzyme operates
in the bisite mode at all concentrations at or above micromolar and
that the apparent deviation from the simple Michaelis-Menten
kinetics is due to the Mg-ADP inhibition. Single Km
values for the hydrolysis of GTP and ITP, which are shown to be less
prone to inhibition, support this view, although two sets of
Km and Vmax with similar
Vmax/Km could also explain
the simple kinetics. Resolution calls for the determination of the number of nucleotides bound in the catalytic sites of uninhibited enzyme, which is not an easy experiment.
The pyrimidine nucleotides, CTP and UTP, were poor substrates for the
TF1 and TFoF1. CTP was not
hydrolyzed and, naturally, did not drive 80 pN·nm/molecule, indicating ~100% efficiency.
Reconstituted FoF1-ATPase actively translocated protons by hydrolyzing ATP, GTP, and ITP, but CTP and UTP were not even
hydrolyzed. Isolated F1 very slowly hydrolyzed UTP (but not
CTP), suggesting possible uncoupling from rotation.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
3
3
1
1
1 subunits, is external to the membrane and catalyzes ATP synthesis. The
ATP synthase is a completely reversible molecular machine in that ATP
hydrolysis in F1 can produce a reverse flow of protons through Fo. Isolated F1 only catalyzes ATP
hydrolysis and hence is called F1-ATPase. Its minimal,
stable subcomplex capable of ATP hydrolysis is
3
3
(2).
subunit is (part of) the common rotor shaft and
that
3
3 subunits, which surrounded
in
the crystal, are the stator in the F1 motor (6-8).
Single-molecule imaging of F1, in particular, has revealed
that the
subunit rotates in a unique direction consistent with the
crystal structure, that
makes discrete 120° steps, and that the
energy conversion efficiency of the F1 motor driven by ATP
hydrolysis can reach ~100% (9, 10). The precise mechanism of
rotation, however, is not yet clear.
subunits, each of which hosts a catalytic site, ATP and its
hydrolysis products, ADP and Pi, are in equilibrium. Thus,
in the absence of an external energy supply, the change in free energy
associated with ATP hydrolysis should manifest as the higher affinity
for ATP than for ADP and Pi. During ATP synthesis, the
mechanical energy supplied by the
rotation driven by F0
somehow decreases the affinity for ATP (and probably increases the
affinity for ADP and Pi), resulting in the appearance of
ATP in the medium. In the reverse reaction of ATP hydrolysis, the free
energy difference between the stronger ATP binding and weaker ADP/Pi binding drives the rotation of
. Boyer proposes
that these binding changes occur sequentially on the three catalytic
sites, synchronously with the
rotation. In the crystal structure of F1 (5), the three
subunits carried a different
nucleotide, an analog of ATP, ADP, and none, in support of the
sequential binding change mechanism.
EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
View larger version (36K):
[in a new window]
Fig. 1.
Contaminants in UTP. 775 nmol of UTP was
loaded on the anion exchange column equilibrated with 50 mM
sodium phosphate (pH 7.0) and eluted with an isocratic flow of the
equilibration buffer. a, elution profiles monitored as
A220 (blue),
A260 (green), and
A280 (red). UTP contained UDP at 3%.
b, expansions of profiles in a. Inset,
further expansion. Five contaminant peaks, C1-C5, were resolved.
c, elution profiles for combined purine nucleotides. 20 nmol
of ATP, 10 nmol of GTP, and 10 nmol of ITP were loaded together.
Inset, their expanded profiles. Contaminating ADP, GDP, and
IDP were eluted before ATP, GTP, and ITP. Peak C4 was identified as GTP
from the coincidence in the elution time and the relative magnitudes of
A220, A260, and
A280. The amounts of the contaminants in C1-C5
were 0.10, 0.03, 0.01, 0.01, and 0.39% of UTP, respectively.
3
3
subcomplex (
-His
tag/
S107C) of F1 derived from thermophilic
Baccillus strain PS3
(TF1)1 was
expressed in Escherichia coli as described previously (9) and purified as follows. The cell lysate containing the enzyme was
applied on a Ni2+-NTA Superflow column (Qiagen)
equilibrated with 50 mM imidazole (pH 7.0) and 100 mM NaCl. The column was washed with 100 mM
imidazole (pH 7.0) and 100 mM NaCl, and then the enzyme was
eluted with 500 mM imidazole (pH 7.0) and 100 mM NaCl. Ammonium sulfate was added to the fraction
containing the enzyme to a final concentration of 10% saturation and
the sample was applied to a butyl-Toyopearl column (Tosoh, Japan)
equilibrated with 500 mM imidazole (pH 7.0), 100 mM NaCl, and 10% saturated ammonium sulfate. The column
was washed with 10 column volumes of a solution containing 100 mM sodium phosphate (pH 7.0), 2 mM EDTA, and
10% saturated ammonium sulfate to remove endogenously bound
nucleotides. The enzyme was eluted with 50 mM Tris-Cl (pH
8.0) and 2 mM EDTA and stored as precipitant in 70%
saturated ammonium sulfate containing 2 mM dithiothreitol.
Before use, the enzyme was dissolved in 100 mM sodium
phosphate (pH 7.0) and 2 mM EDTA and passed through a size exclusion column (Superdex 200 HR 10/30; Amersham Pharmacia Biotech) equilibrated with 100 mM sodium phosphate (pH 7.0) and 2 mM EDTA. The amount of nucleotides remaining on the enzyme
was determined as described previously (11). Samples with less than 0.1 mol of nucleotide/mol of enzyme were used for the measurement of
hydrolysis activity. Whole TFoF1 complex was
reconstituted from the mutant TF1 (
-His tag/
S107C)
and authentic Fo from the thermophile and was incorporated
into liposomes as described previously (12).
3
3
were measured at
23 °C in a medium containing 50 mM MOPS-KOH (pH 7.0), 50 mM KCl, 2 mM MgCl2, and an ATP (or
GTP, ITP, or UTP)-regenerating system consisting of 2.5 mM
phosphoenolpyruvate, 100 µg/ml lactate dehydrogenase, 0.2 mM NADH, and pyruvate kinase at 200 µg/ml for ATP
hydrolysis, 500 µg/ml for GTP and ITP hydrolysis, or 750 µg/ml for
UTP hydrolysis. The reaction was initiated by the addition of the
enzyme to 1.3 ml of assay mixture, and hydrolysis was monitored as NADH
oxidation determined from the absorbance decrease at 340 nm. The CTP
hydrolysis activity was estimated from Pi released in the
same medium without a regenerating system during a 60-min incubation at
23 °C. Hydrolysis by the reconstituted TFoF1
was also measured as Pi released in 50 mM
MOPS-KOH (pH 7.0), 25 mM NaSO4, 25 mM K2SO4, 4 mM
MgCl2, and 2 mM indicated nucleotide during a
20-min incubation at 23 °C.
subunit (10). The assay mixture
contained 50 mM MOPS-KOH, (pH 7.0), 50 mM KCl,
2 mM MgCl2, 10 mg/ml bovine serum albumin, and
an indicated Mg-nucleotide. Fluorescent actin filaments at the bottom
of a flow chamber were observed with an inverted epifluorescence microscope (IX70; Olympus). Assays at less than 10 µM GTP
or ITP were made on the TF1 directly attached to coverslips
to which Ni2+-NTA had been bound covalently (13); the
probability of finding a rotating actin filament was higher than on the
Ni2+-NTA-modified polystyrene bead. Images were taken with
an intensified CCD camera (ICCD-350F; VideoScope) and recorded on 8-mm
videotapes. Rotation by UTP or CTP was observed through an aggregate of
biotinylated polystyrene beads with a diameter of 440 nm, which was
attached to
(14). Images of bead aggregates were obtained in the
bright field in the absence of the oxygen scavenger system and recorded with a CCD camera (CCD-300-RC; Dage-MTI). Possible nucleotide contamination from actin filaments and the oxygen scavenger system was
thus excluded.
RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
View larger version (20K):
[in a new window]
Fig. 2.
Nucleotide-depleted
3
3
subcomplex. a, assessment of nucleotides tightly
bound to the
3
3
subcomplex. 100 pmol
of the purified subcomplex was precipitated by perchloric acid and
subjected to reverse-phase chromatography with the monitor wavelength
of 260 nm (black line). Bound ATP and ADP gave
peaks at 188 and 208 s, respectively, with a height of 0.49 and
0.23 milliabsorbance units above the tail of the main peak. As
controls, 10 pmol of ADP gave a peak at 204 s with a height of
1.28 milliabsorbance units (blue line),
and 100 pmol of the commercial ADP showed a contaminant ATP peak at
183 s (red line). b, absorbance
spectrum of the nucleotide-depleted
3
3
subcomplex. The ratio of A280 (0.57) to
A260 (0.279) was 2.04.
View larger version (23K):
[in a new window]
Fig. 3.
Comparison of the hydrolysis and rotational
rates for different nucleotides. a, time courses of
hydrolysis monitored as NADH oxidation through the regeneration system.
Blue, 2 mM ATP; green, 1 mM GTP; orange, 1 mM ITP;
red, 3 mM UTP. Inset, enlarged view
of the initial portion. The thick parts of the
lines show the portion in which the hydrolysis rate shown in
c was estimated. b, time courses of hydrolysis at
lower nucleotide concentrations. Blue, 600 nM
ATP; green, 3 µM GTP; orange, 10 µM ITP; red, 100 µM UTP.
c, comparison between hydrolysis and rotational rates.
Closed circles, hydrolysis rates determined as in
a and b. Open circles,
rotational rates for 0.5-1.2-µm actin filaments; 3 times the
measured rates in revolutions s 1 are plotted
for comparison with the hydrolysis rates. Error
bars, S.D. Blue, ATP; green, GTP;
orange, ITP; red, UTP. Rotational rates for ATP
are from Ref. 10. Solid lines, except the one for
ATP hydrolysis, show fit with V = Vmax[NTP]/([NTP] + Km),
where Vmax and Km are 187 s
1 and 64 µM for GTP
hydrolysis, 157 s
1 and 202 µM
for ITP hydrolysis, 256.8 s
1 and 5.5 mM for UTP hydrolysis, 3.82 revolutions
s
1 and 0.69 µM for ATP-driven
rotation, 3.28 revolutions s
1 and 3.43 µM for GTP-driven rotation, and 3.95 revolutions
s
1 and 13.3 µM for ITP-driven
rotation. The rates of ATP hydrolysis were fitted with
V = (VmaxaKmb[NTP] + Vmaxb[NTP]2)/([NTP]2 + Kmb[NTP] + Kma
Kmb), where
Vmaxa = 87.5 s
1, Vmaxb = 313 s
1, Kma = 5.25 µM, and Kmb= 429 µM.
1, Vmaxb = 313 s
1, Kma = 5.25 µM, and Kmb = 429 µM (Fig. 3c). The maximum rate of ATP
hydrolysis, Vmaxb, of 313 s
1 is higher than the previous value (10) of
177 s
1, presumably because of the thorough
removal of the bound nucleotide and aggregated enzyme. An additional
factor is the increase of the amount of pyruvate kinase, which may
limit the overall reaction rate, to 0.2 mg/ml, compared with 0.05 mg/ml
in the previous study. The rates for GTP and ITP hydrolysis could be
accounted for by simple Michaelis-Menten kinetics;
Vmax and Km were 187 s
1 and 64 µM, respectively, for
GTP and 157 s
1 and 202 µM for ITP.
1) than the acceleration (0.017 s
1 at 1 mM UTP). Slow binding of
UTP to noncatalytic sites could explain the acceleration, but evidence
is absent. Because the activation took a longer time at lower UTP
concentrations, fully activated rate of UTP hydrolysis was estimated
from the slope between 150 and 200 s at >300 µM,
between 250 and 300 s at 100 µM, and between 1000 and 1200 s at 30 µM. As seen in Fig. 3c, hydrolysis of UTP was much slower than that of the purine nucleotides. To confirm that the measured hydrolysis rate was that of UTP and not of
contaminants, we also measured Pi released in the medium. Although the rate of Pi release was around half the
hydrolysis rate determined from NADH oxidation, presumably because of
the absence of the UTP-regenerating system, TF1 produced
Pi equivalent to >40% of UTP in 5 min. Thus,
TF1 did hydrolyze UTP, not contaminating other
nucleoside-triphosphates, which amounted to <0.5% (see
"Experimental Procedures"). CTP, in contrast, was not hydrolyzed;
the rate of Pi release at 2 mM CTP measured
over 60 min was indistinguishable from the rate without the enzyme
(<0.01 s-1).
subunit in TF1. The rotation, observed
through the motion of a fluorescent actin filament attached to the
subunit, was counterclockwise without exception, as in the case of
ATP-driven rotation. Rotation assays at less than 10 µM
GTP or 10 µM ITP were made on the enzyme directly attached to a coverslip that had been covalently modified with Ni2+-NTA (13). Compared with the previous method (10) of
attaching the enzyme on Ni2+-NTA-coated beads, the direct
attachment resulted in a higher percentage of finding rotating
filaments at low nucleotide concentrations.
1. This is simply due to the hydrodynamic friction
against the rotating actin filament (10). At low nucleotide
concentrations, the rotation was resolved into discrete 120° steps
(Fig. 4, a and b).
The histograms of dwell times between steps were fitted with a single
exponential decay with the rate of 2.9 s
1 for
1 µM GTP and 11.1 s
1 for 10 µM ITP (Fig. 4, c and d), implying
that a first order reaction, binding of GTP (ITP), triggers each 120°
step. These rate constants agree with the hydrolysis rates (Fig.
3c), confirming again that each 120° step is driven by the
hydrolysis of one molecule of GTP or ITP.
View larger version (47K):
[in a new window]
Fig. 4.
Stepping rotation at low nucleotide
concentrations. a and b, time courses of
rotation at 1 µM GTP (actin length 0.95 µm) and at 10 µM ITP (actin length 1.01 µm). Insets,
traces of the centroid of the actin filament. c and
d, histograms of dwell times between steps. Rotation
records in a and b were analyzed and are shown in
c and d. Lines indicate an exponential
fit: constant × exp( kt), where k is the
rate for nucleotide binding (2.9 s
1 for 1 µM GTP and 11.1 s
1 for 10 µM ITP).
Rotation--
With commercial UTP at 4 mM, we occasionally observed slow rotation (e.g.
1.4 revolutions s
1). We suspected that this
might have been due to contaminants, because ATP at a concentration as
low as 300 nM, for example, could account for the observed
speed. Indeed, even CTP, which the
3
3
does not hydrolyze, produced rotation at 0.3 revolutions s
1 at 2 mM. We therefore purified
UTP and CTP (see "Experimental Procedures") and examined rotation
at 300 µM, the highest concentration available after the
column purification. To avoid possible nucleotide contamination from
other sources, we attached an aggregate of biotinylated polystyrene
beads to the
subunit in place of an actin filament and observed the
beads in bright field in the absence of the oxygen scavenger system. No
rotating beads were found in six assays with UTP in which 5543 beads
only fluctuated around a fixed point. Observation in each assay
continued for more than 30 min, much longer than the time needed to
activate UTP hydrolysis at 300 µM (~5 min). Six assays
with CTP also gave negative results.
1 at 300 µM UTP. We therefore
carried out parallel assays with 600 nM ATP where the
rotational rate would be 1.5 s
1. 40 rotating
beads were found out of 4299 fluctuating beads (five assays). We also
made an "unpredicted test" in which we observed beads without
knowing the identity of the nucleotide. No rotation was found with 300 µM UTP, and rotating beads were readily found in the
presence of 600 nM ATP. We conclude that UTP cannot support rotation of the
subunit carrying a bead aggregate, at least not at
the rate expected from the hydrolysis rate. One possibility is that UTP
hydrolysis, unlike the hydrolysis of purine nucleotides, is completely
decoupled from
rotation, even at no load. The other possibility is
that UTP, with a different base structure, cannot supply sufficient
power to drive the beads through 120°; in this case, the rate of UTP
hydrolysis by the loaded TF1 would also be very low.
rotation, the rotational velocities of actin filaments
attached to the
subunit on Ni2+-NTA coated-beads were
examined. Fig. 5 shows time courses of rotation for actin filaments with length around 2 µm (Fig.
5a) and 3-4 µm (Fig. 5b). The average
rotational velocity was determined for each curve over a portion
containing at least five consecutive revolutions without noticeably
unnatural intermissions (e.g. the last part of the
brown curve in Fig. 5b was omitted).
The results are summarized in Fig. 5c. Most data for
GTP-driven (green) and ITP-driven rotation
(orange) are on or below
the line representing the constant torque of 40 pN·nm, as
is also the case for ATP-driven rotation (blue). We think
that higher velocity values are more reliable, because any obstructions
against rotation would reduce the velocity, and we conclude that
TF1 exerts a constant rotary torque of about 40 pN·nm
regardless of the differences among the structures of purine rings.
(The average torque calculated over all points in Fig. 5c is
32 ± 11 pN·nm for ATP, 35 ± 11 pN·nm for GTP, and 32 ± 10 pN·nm for ITP; these average values, however, are not meaningful,
because the data in Fig. 5c have been selected for high
torque values.)
View larger version (16K):
[in a new window]
Fig. 5.
Estimation of torque in GTP- and ITP-driven
rotations. a, time courses of the rotation of an actin
filament with length around 2 µm. Blue, 2 mM
ATP; green, 2 mM GTP; orange, 2 mM ITP; pink, 2 mM ATP, 10 µM ADP, and 100 mM Pi;
gold, 2 mM GTP, 10 µM GDP, and 100 mM Pi. b, time courses of the
rotation with actin length of 3-4 µm. See a for color
coding. c, rotational rate versus the length of
the actin filament. Blue circles, 2 mM Mg-ATP; green circles, 2 mM Mg-GTP; orange circles, 2 mM Mg-ITP; pink squares, 2 mM ATP, 10 µM ADP, and 100 mM
Pi ( GATP =
80
pN·nm/molecule); gold diamonds, 2 mM GTP, 10 µM GDP, and 100 mM
Pi (
GGTP =
80
pN·nm/molecule). Large symbols indicate the
data in a and b. Solid
line, calculated rotational rate under an assumed constant
torque of 40 pN·nm; dashed lines, constant
torque at 80 and 20 pN·nm.
/3
radians (equal to 120°), ~80 pN·nm, is the mechanical work done in a 120° step. In the previous study (10), it was shown that TF1 does this much work even when the free energy available
from ATP hydrolysis,
GATP, was reduced to
90 pN·nm/molecule. Here, we examined the torque and work in the
presence of 2 mM ATP (or GTP), 10 µM ADP
(GDP), and 100 mM inorganic phosphate. Under these conditions,
GATP is calculated as
80
pN·nm/molecule from
GATP =
G0 + kBT·ln[ADP]·[Pi]/[ATP],
where
G0 =
50 pN·nm/molecule (17),
kBT = 4.1 pN·nm/molecule is
thermal energy at room temperature, and
GGTP should be
of a similar value. As seen in Fig. 5c, the experimental
points at
GATP =
80 pN·nm/molecule (purple) and
GGTP =
80
pN·nm/molecule (brown) are indistinguishable from other
points and also fall on or below the line for the constant torque
of 40 pN·nm or constant work of 80 pN·nm (the torque averaged over
plotted points is 35 ± 10 pN·nm for
GATP =
80 and 32 ± 9 pN·nm for
GGTP =
80 pN·nm/molecule) This
strengthens our contention that the efficiency of the energy conversion
in this motor, from the hydrolysis (of purine nucleotides) into the
mechanical work, can reach ~100%.
rotation is mechanically coupled to the proton pump.
Liposomes were incubated with 2 mM nucleotide until the
base line became stable, and then 4 mM MgCl2 was added to start the reaction (Fig. 6).
After 20 min, 10 µM FCCP was added to collapse the pH
gradients so as to ensure that the fluorescence decrease indicated the
proton uptake. Fig. 6 clearly shows that the purine nucleotides ATP,
GTP, and ITP drove the formation of pH gradient but UTP and CTP did
not; the latter two gave signals that were indistinguishable from the
control in the absence of a nucleotide. The initial rates of proton
uptake by GTP and ITP in 1 min after starting the reaction were 30 and 27% of the ATP-driven uptake, respectively. The rate of nucleotide hydrolysis by TFoF1, determined from the amount
of inorganic phosphate released in 20 min, was 7.2 ± 1.4 s
1 for ATP, 3.4 ± 0.5 s
1 for GTP, and 2.7 ± 0.5 s
1 for ITP. The three nucleotides show the
same order in proton pumping and hydrolysis activities. The agreement
suggests that GTP and ITP hydrolysis are coupled to proton pumping as
efficiently as with ATP. Pyridine nucleotides were not good substrates
for hydrolysis by TFoF1; the hydrolysis
activity was not detected by the phosphate measurement (<0.12
s
1 for 2 mM CTP and <0.09
s
1 for 2 mM UTP).
View larger version (23K):
[in a new window]
Fig. 6.
Proton pump activity of
TFoF1 reconstituted into liposomes.
Translocation of protons into TFoF1 liposomes
was monitored as the decrease in pyranine fluorescence. The reaction
mixture contained 2 mM ATP (blue
line), 2 mM GTP (green
line), 2 mM ITP (orange
line), 2 mM CTP (black
line), 2 mM UTP (red
line), or no nucleotide (dashed
black line). The reaction was started by the
addition of 4 mM MgCl2. At 1200 s, 10 µM FCCP was added to collapse the pH gradients.
DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
rotation and for proton pumping in the
FoF1-ATPase. Both rotation and proton pumping
were supported by the purine nucleotides, ATP, GTP, and ITP, but not by
the pyrimidine nucleotides, CTP and UTP. Our results are consistent
with the report by Perlin et al. (18), where GTP and ITP
were shown to drive proton pumping in the
FoF1-ATPase, and support the idea that
nucleotide hydrolysis is coupled to proton pumping through mechanical
rotation of the
subunit.
GATP or
GGTP is reduced to
80 pN·nm/molecule. We
did not ascertain a high efficiency in ITP-driven rotation, but Sorgato
et al. (19) have reported that the energy from ITP
hydrolysis is converted into a membrane potential by submitochondrial
particles as efficiently as ATP hydrolysis. Thus, the efficiency of
ITP-driven rotation is expected also to be close to 100%. The
similarities among the three nucleotides suggest that the mechanical
characteristics of the rotation such as the stepping, torque, and work
per step are inherent in the (structure of the) F1
motor. Purine nucleotides can trigger and let proceed the stepping
mechanism in which the step angle, torque, and work are preset, whereas
pyrimidine nucleotides cannot.
1
s
1 for ATP (10), 2.9 × 106
M
1 s
1 for GTP, and 1.1 × 106 M
1 s
1 for ITP.
rotation during synthesis is used to
release a tightly bound ATP into the medium. Conversely, during
hydrolysis, binding of ATP powers the reverse rotation of
. If the
above difference in the rate constant of nucleotide binding reflects a
similar difference in the affinity for the nucleotide, then the energy
provided by ITP binding will be the smallest and might have been
insufficient to power the stepping mechanism. Because all three purine
nucleotides supported rotation, it seems either that the affinity for
ITP is high enough (a 20-fold difference in the binding constant is
equivalent to a free energy difference of only 12 pN·nm/molecule), or
that the rate of nucleotide release decreases in the order of ATP to
ITP and makes the affinities more or less the same. In this regard, it
is possible that UTP, of which the hydrolysis curve in Fig.
3c is further shifted toward the right, cannot confer
sufficient binding energy to the stepping mechanism.
rotation or proton pump.
UTP was hydrolyzed by TF1, yet UTP did not support rotation
and pumping. Ca-ATP has also been reported to be an uncoupling
substrate for FoF1, in that Ca-ATP was
hydrolyzed without pumping protons (20). The kinetics of Ca-ATP
hydrolysis, however, was similar to that of Mg-ATP hydrolysis. Thus,
the nature of uncoupling seems different between the two cases, Ca-ATP
and (Mg-)UTP. Interestingly, the reconstituted
TFoF1 did not hydrolyze UTP, while
TF1 (the
3
3
subcomplex)
did. This finding is consistent with the report by Yokoyama et
al. (21) that the substrate specificity of this enzyme is higher
when it contains a fuller complement of subunits. The UTP hydrolysis
activity of the
3
3
subcomplex may be
ascribed to some flexibility in and around the catalytic site that
would be suppressed in FoF1; such hydrolysis
may proceed without a major structural change of the
subunit and
thus without rotation of the
subunit.
![]() |
ACKNOWLEDGEMENTS |
---|
We thank Y. Harada, T. Nishizaka, K. Adachi, T. Hisabori, and E. Muneyuki for technical assistance and helpful discussions and H. Umezawa for laboratory management.
![]() |
FOOTNOTES |
---|
* The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
¶ Present address: Dept. of Structural Biology, Free University of Amsterdam, De Boelelaan 1087, 1081, Amsterdam, Netherlands.
Present address: Cold Spring Harbor Laboratory, 1 Bungtwon
Rd., Cold Spring Harbor, NY 11724.
¶¶ To whom correspondence should be addressed. Tel.: 81 44 750 1710; Fax: 81 44 750 1712.
Published, JBC Papers in Press, March 28, 2001, DOI 10.1074/jbc.M102200200
![]() |
ABBREVIATIONS |
---|
The abbreviations used are:
TF1, the 3
3
subcomplex
(
-His tag/
S107C) of F1 derived from thermophilic
Baccillus strain PS3;
NTA, nitrilotriacetic acid;
FCCP, carbonyl cyanide p-(trifluoromethoxy) phenylhydrazone;
TFoF1, FoF1-ATPase
derived from thermophilic Baccillus strain PS3;
MOPS, 4-morpholinepropanesulfonic acid;
pN, piconewton(s).
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1. | Boyer, P. D. (2000) Biochim. Biophys. Acta 1458, 252-262[Medline] [Order article via Infotrieve] |
2. | Matsui, T., and Yoshida, M. (1995) Biochim. Biophys. Acta 1231, 139-146[Medline] [Order article via Infotrieve] |
3. | Boyer, P. D., and Kohlbrenner, W. E. (1981) in Energy Coupling in Photosynthesis (Selman, B. R. , and Selman-Reimer, S., eds) , pp. 231-240, Elsevier, Amsterdam |
4. | Oosawa, F., and Hayashi, S. (1986) Adv. Biophys. 22, 151-183[Medline] [Order article via Infotrieve] |
5. | Abrahams, J. P., Leslie, A. G., Lutter, R., and Walker, J. E. (1994) Nature 370, 621-628[CrossRef][Medline] [Order article via Infotrieve] |
6. |
Aggeler, R.,
Haughton, M. A.,
and Capaldi, R. A.
(1995)
J. Biol. Chem.
270,
9185-9191 |
7. | Duncan, T. M., Bulygin, V. V., Zhou, Y., Hutcheon, M. L., and Cross, R. L. (1995) Proc. Natl. Acad. Sci. U. S. A. 92, 10964-10968[Abstract] |
8. | Sabbert, D., Engelbrecht, S., and Junge, W. (1996) Nature 381, 623-625[CrossRef][Medline] [Order article via Infotrieve] |
9. | Noji, H., Yasuda, R., Yoshida, M., and Kinosita, K., Jr. (1997) Nature 386, 299-302[CrossRef][Medline] [Order article via Infotrieve] |
10. | Yasuda, R., Noji, H., Kinosita, K., Jr., and Yoshida, M. (1998) Cell 93, 1117-1124[Medline] [Order article via Infotrieve] |
11. |
Tsunoda, S. P.,
Muneyuki, E.,
Amano, T.,
Yoshida, M.,
and Noji, H.
(1999)
J. Biol. Chem.
274,
5701-5706 |
12. |
Bald, D.,
Amano, T.,
Muneyuki, E.,
Pitard, B.,
Rigaud, J. L.,
Kruip, J.,
Hisabori, T.,
Yoshida, M.,
and Shibata, M.
(1998)
J. Biol. Chem.
273,
865-870 |
13. |
Adachi, K.,
Yasuda, R.,
Noji, H.,
Itoh, H.,
Harada, Y.,
Yoshida, M.,
and Kinosita, K., Jr.
(2000)
Proc. Natl. Acad. Sci. U. S. A.
97,
7243-7247 |
14. | Yasuda, R., Noji, H., Yoshida, M., Kinosita, K., Jr., and Itoh, H. (2001) Nature 410, 898-904[CrossRef][Medline] [Order article via Infotrieve] |
15. | Jault, J. M., Matsui, T., Jault, F. M., Kaibara, C., Muneyuki, E., Yoshida, M., Kagawa, Y., and Allison, W. S. (1995) Biochemistry 34, 16412-16418[Medline] [Order article via Infotrieve] |
16. |
Matsui, T.,
Muneyuki, E.,
Honda, M.,
Allison, W. S.,
Dou, C.,
and Yoshida, M.
(1997)
J. Biol. Chem.
272,
8215-8221 |
17. | Stryer, L. (1995) Biochemistry , 4th Ed. , pp. 443-462, Freeman, New York |
18. | Perlin, D. S., Latchney, L. R., Wise, J. G., and Senior, A. E. (1984) Biochemistry 23, 4998-5003[Medline] [Order article via Infotrieve] |
19. | Sorgato, M. C., Galiazzo, F., Valente, M., Cavallini, L., and Ferguson, S. J. (1982) Biochim. Biophys. Acta 681, 319-322[Medline] [Order article via Infotrieve] |
20. | Papageorgiou, S., Melandri, A. B., and Solaini, G. (1998) J. Bioenerg. Biomembr. 30, 533-541[CrossRef][Medline] [Order article via Infotrieve] |
21. |
Yokoyama, K.,
Hisabori, T.,
and Yoshida, M.
(1989)
J. Biol. Chem.
264,
21837-21841 |