J Biol Chem, Vol. 274, Issue 46, 32667-32671, November 12, 1999
The Mechanism of ATP Hydrolysis at the Noncatalytic Sites of
the Transcription Termination Factor Rho*
Dong-Eun
Kim
and
Smita S.
Patel§
From the Department of Biochemistry, Ohio State University,
Columbus, Ohio 43210
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ABSTRACT |
Escherichia coli transcription
termination factor rho is a hexamer with three catalytic subunits that
turnover ATP at a fast rate and three noncatalytic subunits that
turnover ATP at a relatively slow rate. The mechanism of the ATPase
reaction at the noncatalytic sites was determined and was compared with
the ATPase mechanism at the catalytic sites. A sequential mechanism for
ATP binding or hydrolysis that was proposed for the catalytic sites was
not observed at the noncatalytic sites. Pre-steady-state pulse-chase experiments showed that three ATPs were tightly bound to the
noncatalytic sites and these were simultaneously hydrolyzed at a rate
of 1.8 s
1 at 18 °C. The apparent bimolecular rate
constant for ATP binding was determined as 5.4 × 105
M1 s1 in the presence of
poly(C) RNA. The ATP hydrolysis products dissociated from the
noncatalytic sites at 0.02 s1. The hydrolysis of ATP at
the noncatalytic sites was at least 130 times slower, and the overall
ATPase turnover was 1500 times slower than that at the catalytic sites.
These results from studies of the rho protein are likely to be general
to hexameric helicases. We propose that the ATPase activity at the
noncatalytic site is too slow to drive translocation of the protein on
the nucleic acid or to provide energy for nucleic acid unwinding.
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INTRODUCTION |
The Escherichia coli rho protein is a transcription
termination factor that is required for the release of certain nascent RNAs from the transcription complex (1, 2). Rho has various activities
such as RNA binding (3), RNA-dependent ATP hydrolysis (4,
5), and an ATPase-dependent helicase activity that unwinds RNA-DNA duplexes (6). The rho protein functions as a hexamer of
identical subunits (7) and shows structural and mechanistic similarities to hexameric DNA helicases such as E. coli DnaB
and the T7 gene 4 helicase (8, 9). The structure and mechanism of the
rho protein appear to be similar to the
3
3 F1-ATPase protein. The
tertiary structure of the RNA-binding domain residing at the N terminus
of rho is homologous to the structure of the N-terminal domain of the
F1-ATPase, despite there being no amino acid sequence
homology between them (10). The C-terminal domain of rho shows
significant amino acid homology to the -subunit of
F1-ATPase. The amino acid sequence homology between rho and the subunit is more striking than the amino acid sequence homology between the and subunits of the F1-ATPase protein
(11). In addition to the structural similarity, the ATPase mechanisms of both rho and the T7 gene 4 helicase show striking similarity to the
binding change mechanism of the F1-ATPase protein (9, 12).
Unlike the F1-ATPase protein, the rho protein is a
homohexamer. However, several studies suggest that the hexamer is
asymmetric with a C3/6 symmetry (13). This asymmetry
results in three high affinity and three low affinity ATP binding sites
(14, 15) and two classes of nucleic acid binding sites on the rho
hexamer (16, 17). Our previous studies showed that the ATPase turnover rate was different at the two classes of nucleotide binding sites on
the rho hexamer (18). In the presence of poly(C) RNA, the ATPase
turnover at the high affinity noncatalytic sites was 1500 times slower
than at the low affinity catalytic sites. In this paper, the mechanism
of the ATPase reaction at the noncatalytic sites was determined using
pre-steady-state kinetic approaches. This study allowed us to make a
comparison between the mechanisms of ATP hydrolysis at the noncatalytic
and the catalytic sites of the rho hexamer.
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MATERIALS AND METHODS |
Protein, Nucleotides, RNA Homopolymer, and Other
Reagents--
Purified rho protein was provided by Dr. Katsuya
Shigasada (Department of Genetics and Molecular Biology, Kyoto
University, Kyoto, Japan). The rho protein was overexpressed in
E. coli strain HB101 carrying the rho overexpression plasmid
pKS26 (19) and purified according to Finger and Richardson (20) with
slight modifications. The rho protein concentration was determined by UV absorption at 280 nm using an extinction coefficient of 0.325 (mg/ml)1 cm1 (7). The ATP and RNA
homopolymer, poly(C), was purchased from Amersham Pharmacia Biotech.
Poly(C) RNA had a reported s20,w value of 7.1 in 0.015 M NaCl, 0.0015 M sodium
citrate buffer, pH 7.0, with an average length of 420 bases. Poly(C)
RNA concentration was determined by UV absorption at 269 nm using an
extinction coefficient of 6200 M1
cm1 for the cytosine base. These RNAs were dissolved in
TE buffer (40 mM Tris-HCl, pH 7.0, 0.5 mM EDTA)
and used without further purification. [-32P]ATP was
purchased from Amersham Pharmacia Biotech, and its purity was assessed
by polyethyleneimine-cellulose TLC and corrected for in all experiments.
The Three Syringe Rapid Chemical Quench-Flow
Experiments--
The kinetic experiments were conducted using a rapid
chemical quench-flow instrument (21) built by KinTek Corp. (State
College, PA) at a constant temperature of 18 or 10 °C. In the three
syringe experimental set-up, two delay times were used (KinTek RQF-3
software) as shown in Scheme I. The
pulse-chase kinetics were measured at fixed t1
and varying t2 (0.015-7 s). A 14-µl mixture
of rho protein (2.0 µM hexamer) and poly(C) RNA (2.68 µM) in 40 mM Tris-HCl (pH 7.8), 100 mM KCl, 10 mM MgCl2, 0.1 mM dithiothreitol, and 10% (v/v) glycerol was rapidly
mixed with a 14-µl mixture of ATP (100 µM) and
[-32P]ATP (0.03 µCi/µl) for
t1. After reacting the above for
t1, 31 µl of nonradioactive ATP (5 mM, final concentration after mixing and 10 mM
free MgCl2) was added from a third syringe in the
quench-flow instrument. The final unlabeled ATP concentration exceeded
the labeled ATP concentration by at least 200-fold. A total of 59-µl reactions were chased for varying times (t2) and
quenched with 120 µl of 0.5 M formic acid to stop the
reaction. Aliquots (1.0 µl) from each acid-quenched reaction at
varying time points were spotted on polyethyleneimine-cellulose TLC,
which was developed in 0.3 M potassium phosphate, pH 3.4. The resolved radioactive ATP and ADP were quantitated on a
PhosphorImager instrument (Molecular Dynamics). Product formation was
equal to the radioactivity corresponding to ADP divided by the total
radioactivity.
To estimate ATP hydrolyzed in the reaction time
(t1) prior to the ATP chase, 0.5 M
formic acid was added from the third syringe instead of the ATP chase.
This control experiment was repeated three times, and the values were
averaged and taken as a zero time point in the chase-time period (
t2). The efficiency of 0.5 M formic
acid as a quench was determined by loading the rho protein/RNA solution
into one sample loop, the reaction buffer into the other sample loop,
the quench solution into the third syringe line, and the
[-32P]ATP solution including the nonradioactive ATP
chase into the collecting microcentrifuge tube (21). In this
configuration, the enzyme solution was then rapidly mixed with the
quench solution for 5 ms prior to mixing with the ATP solution. Only
0.03% of ATP was hydrolyzed to ADP, which was the same as the level
observed in the absence of enzyme. This indicates that the quench
solution inactivated the enzyme within 5 ms.
Data Analysis--
The hydrolysis of radioactive ATP during the
chase-time, t2, was corrected as follows. A
correction factor, B, was calculated,
![B=[E]p×D×k<SUB><UP>cat</UP></SUB>×[<UP>ATP</UP>]p×D/[<UP>ATP</UP>]c](/content/vol274/issue46/fulltext/32667/img001.gif)
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(Eq. 1)
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where [E]p (1.0 µM) is the
concentration of rho hexamer before chase, D (28 µl/59
µl) is the dilution factor because of the addition of chase,
[ATP]p (50 µM) is the concentration of
radioactive ATP before chase, kcat is the ATPase
turnover number (30 and 8.8 s1 at 18 and 10 °C,
respectively), and [ATP]c (5.0 mM) is the
concentration of chase ATP. The value of B × t2 was subtracted from each data point in the
pulse-chase experiments. The corrected amounts of ADP formed during the
chase-time t2 were plotted and were fit to the
following single exponential equation with a y intercept using the Sigma Plot software version 5.0 (SPSS).
![[<UP>ADP</UP>]<SUB>t2</SUB>=[<UP>ADP</UP>]<SUB>t1</SUB>+A(1−e<SUP><UP>−</UP>k×t2</SUP>)](/content/vol274/issue46/fulltext/32667/img002.gif)
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(Eq. 2)
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Where t1 and t2
are the two delay times (see Scheme I),
[ADP]t1 is the amount of ADP formed during the
reaction time (t1),
[ADP]t2 is the total amount of ADP formed, k is the rate of ATP hydrolysis during chase-time
t2, and A represents the amount of
ADP formed during chase-time t2.
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RESULTS AND DISCUSSION |
The two classes of ATP binding sites on the rho hexamer termed
catalytic and noncatalytic sites hydrolyze ATP at different rates. At
the catalytic sites, close to 30 ATPs are hydrolyzed/mole of rho
hexamer/second, and our previous studies have shown that the ATPase
turnover at the noncatalytic sites is about 1500 times slower (18). The
fast ATPase turnover at the catalytic sites precludes measurement of
the slow ATPase reaction at the noncatalytic sites. To dissect the
ATPase kinetics at the noncatalytic sites, we took advantage of the
tight binding of ATP at the noncatalytic sites and used pulse-chase
experiments to determine the kinetics of ATP binding and hydrolysis at
those sites.
Kinetics of ATP Hydrolysis at the Noncatalytic Sites of
Rho--
Scheme I shows the design of the pulse-chase experiments. A
complex of rho protein and poly(C) RNA was mixed with
[-32P]ATP for a fixed pulse-time
t1. During t1, the
radioactive ATP is presumed to bind both to the catalytic and the
noncatalytic sites and to undergo a few cycles of ATP hydrolysis at the
catalytic sites. To avoid extensive ATP hydrolysis at the catalytic
site, the pulse-time t1 was kept in the
millisecond range. After t1, the reaction was
chased with excess nonradioactive ATP. The chase-time t2 was varied from 15 ms to several seconds
before acid was added to stop the reaction. The free MgCl2
in the above reactions was always kept at 10 mM
concentration. During the chase-time t2, a
percentage of rho-bound radioactive ATP would dissociate, and a
percentage would hydrolyze to ADP and Pi dictated by the
respective rates of ATP dissociation and ATP hydrolysis on the protein.
The radioactive ATP that dissociates from rho can not rebind because it
will be diluted in the pool of excess nonradioactive ATP. From the
kinetics of radioactive ADP formation in the chase-time
t2, one can estimate both the number of ATP
molecules that remain bound and chased to product as well as their rate
of hydrolysis.
Fig. 1A shows the results of
such an experiment where pulse-time t1 was fixed
to 0.1 s, and the chase-time t2 was varied
from 0.015-7.0 s. There are two exponential phases in the pulse-chase time course. Because of the lack of time points shorter than 15 ms, the
fast exponential phase could not be defined. Hence, the data were fit
to a single exponential with a y intercept (Equation 2)
instead of the sum of two exponentials. This fit provided the rate
constant of the slow phase and the amplitudes of both the slow and the
fast phase. A total of four ATPs were chased to ADP during the
chase-period, indicating that at least four ATPs were bound in the
pulse-time of 0.1 s. Of the four ATPs, one was hydrolyzed at a
very fast rate estimated to be >150 s1 in the chase-time
t2 (assuming that the first time point of 15 ms
represented 3-4 half-lives), and the rest were hydrolyzed at a slow
rate equal to 1.9 s1.
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Fig. 1.
Pulse-chase kinetics of ATP hydrolysis.
A complex of rho (2.0 µM hexamer) and poly(C) RNA (2.68 µM) was mixed with [-32P]ATP (100 µM) for a fixed pulse period t1.
The reaction was then chased with 5 mM (final concentration
after mixing) of nonradioactive ATP, and the chase-time
t2 was varied from 0.015 to 7.0 s. The
experiment was carried out in a rapid quench-flow instrument at
18 °C as shown in Scheme I. The filled circles ( ) in
both panels show the amount of ATP hydrolyzed in the pulse period
t1. The data were corrected for radioactive ATP
hydrolysis in the chase-time ("Materials and Methods"). The
left y axis shows the corrected amount of ATP
hydrolysis during the chase-time, which was obtained by subtracting the
amount of ATP hydrolyzed during pulse-time t1
and divided by [rho]. The right y axis shows
the uncorrected ATP hydrolysis. Insets in both panels show
the time-course up to 500 ms. A, the pulse-chase kinetics of
ATP hydrolysis with pulse-time t1 fixed at 100 ms. Computer fit of the kinetics to Equation 2 ("Materials and
Methods") represented by a solid line showed that 3.8 ± 0.5 ATPs were hydrolyzed to product during the 100 ms pulse-time.
One ATP (0.96 ± 0.23 ATPs/hexamer) was hydrolyzed at a fast rate,
and three ATPs (2.80 ± 0.31 ATPs/hexamer) were hydrolyzed at a
rate of 1.92 ± 0.60 s1. B, the
pulse-chase kinetics of ATP hydrolysis with pulse-time
t1 fixed at 30 ms. Under these conditions, a
total of 1.8 ATPs were hydrolyzed to product/rho hexamer. Less than one
ATP (0.51 ± 0.13 ATPs/hexamer) was hydrolyzed at a fast rate, and
1.3 ± 0.1 ATPs/hexamer were hydrolyzed at a rate of 2.31 ± 0.81 s1.
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The ATP that was hydrolyzed at a rate greater than the steady-state
ATPase turnover was most likely bound to one of the catalytic sites.
The remaining three ATPs must be bound to the noncatalytic sites,
because these were hydrolyzed at a rate 15 times slower than the
steady-state ATPase rate of 30 s1. Similar kinetics were
observed when the pulse-time t1 was changed to
30 ms (Fig. 1B). Only the total number of ATPs chased to
product was different. After a 30-ms pulse, a total of 1.8 radioactive ATPs/rho hexamer were chased to products. Of the 1.8 ATPs, less than 1 ATP (0.51 ATP) was hydrolyzed to ADP at a fast rate and about 1.3 ATPs
were hydrolyzed at a slow rate. These results indicate that only a
limited number of ATPs were bound to the noncatalytic sites during the
pulse-time of 30 ms. This result prompted us to carry out a systematic
study in which the pulse-chase kinetics were measured at various fixed
pulse-times to measure the apparent rate of ATP binding to the
noncatalytic sites.
Kinetics of ATP Binding to the Noncatalytic Sites of
Rho--
Examination of the pulse-chase kinetics at varying
pulse-times provided an estimate of the apparent bimolecular rate of
ATP binding to the noncatalytic sites. The chase kinetics were measured at various pulse-times t1 that ranged from 10 to
150 ms. Each chase-time course was analyzed to obtain the number of
ATPs bound to the noncatalytic and the catalytic sites and the rate of
ATP hydrolysis at the noncatalytic sites. These parameters were
obtained from computer fit of the data to a single exponential with a
y intercept (Equation 2 under "Materials and Methods").
As the pulse-time t1 was increased from 10 to
150 ms, the total number of ATPs chased to product increased to a
maximum of three to four. As shown in Fig.
2A, during a 10-150-ms
pulse-time, less than one ATP (0.60 ± 0.08) was hydrolyzed at a
very fast rate (within the fastest time point of 15 ms). The number of
ATPs that were more slowly hydrolyzed to product increased from one to
a maximum of three ATPs (2.8 ± 0.2) as the pulse-time
t1 was increased (Fig. 2B). This
increase in ATP site occupation at the noncatalytic sites was
exponential and fit to a rate constant of 27 s1. The rate
of slow ATP hydrolysis remained constant around an average value of 1.8 s1 (Fig. 2C). We assume that the fast
hydrolysis of ATP at one site occurs at the catalytic site, and the
slow hydrolysis of close to three ATPs occurs at the noncatalytic
sites. Assuming that ATP binding is the rate-limiting step at the
noncatalytic sites, an apparent bimolecular rate constant for ATP
binding to the noncatalytic site, kon, of
5.4 × 105 M1
s1 was calculated by dividing the observed exponential
rate by the concentration of ATP in the reaction (27 s1/50 µM).
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Fig. 2.
Pulse-chase ATPase kinetics as a function of
pulse-time t1. The experiment
described in Fig. 1 was carried out at various pulse-time
t1 (10-150 ms) at 18 °C. Each kinetic
experiment was analyzed as described in Fig. 1. A, shows the
t1 dependence of the number of ATPs hydrolyzed
to product at the catalytic sites (y intercept). An average
of 0.60 ± 0.08 ATP were hydrolyzed to product at the catalytic
sites (dashed line). B, shows the
t1 dependence of the ATP bound to the
noncatalytic sites. The site occupancy at the noncatalytic sites
increased in an exponential manner with a rate constant of 26.8 ± 5.6 s1 (solid line). The amplitude provided
the maximum number of ATPs hydrolyzed to product at the noncatalytic
sites as 2.80 ± 0.2. C, the t1
dependence of the rate of ATP hydrolysis at the noncatalytic sites. The
ATP hydrolysis rate at the noncatalytic sites remained constant around
1.84 ± 0.19 s1 (dashed line).
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The Rate of ATP Hydrolysis at the Catalytic Site of Rho--
In
the pulse-chase experiments described above, we found that the ATP
bound at the catalytic site was completely hydrolyzed to ADP in the
shortest chase-time of 15 ms. Thus, we could only estimate its rate of
hydrolysis at >150 s1. To obtain a more accurate value
for the rate of ATP hydrolysis at the catalytic site, we performed the
pulse-chase experiment both at a lower temperature of 10 °C and at a
higher concentration of rho. Fig. 3 shows
the results of the pulse-chase experiment in which the pulse-time was
fixed at 0.3 s and the chase-time was varied. The same kinetic
pattern for hydrolysis of ATP was observed at the lower temperature.
One ATP was hydrolyzed at a fast rate and two to three (2.2 ± 0.3 ATPs) were hydrolyzed at a slow rate. The ATP hydrolysis at the
catalytic site was still too fast to measure. Most of the ATP at the
catalytic site was hydrolyzed in the fastest time of 15 ms providing an
estimate of >150 s1 for the hydrolysis rate at 10 °C.
The ATP hydrolysis rate at the noncatalytic sites decreased to a value
of 1.12 s1, about 1.6 times slower than the rate of
hydrolysis at 18 °C. Assuming the same temperature dependence for
the catalytic site, we estimate that ATP hydrolysis at the catalytic
sites occurs at a rate >240 s1 at 18 °C.
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Fig. 3.
Pulse-chase kinetics of ATP hydrolysis at
10 °C. The experiment was carried out as described in Fig. 1.
The final concentration of rho in the experiment prior to the addition
of ATP chase was 2.0 µM hexamer, poly(C) RNA at 2.68 µM, [-32P]ATP at 100 µM;
the chase [ATP] was 5 mM; and free MgCl2 was
10 mM in the chase reaction. The pulse-time
t1 was fixed at 0.30 s, and the chase-time
t2 was varied from 0.015 to 7.0 s. Computer
fit of the data to Equation 2 showed that a total of 3.2 ± 0.44 ATPs were hydrolyzed to product/hexamer. One ATP (1.0 ± 0.1 ATPs/hexamer) was hydrolyzed at a fast rate, and 2.2 ± 0.30 ATPs/hexamer were hydrolyzed at a rate of 1.12 ± 0.46 s1. Inset shows the time course up to 600 ms
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A Minimal Mechanism For ATP Binding and Hydrolysis at the Catalytic
and Noncatalytic Sites of Rho--
The results presented in this paper
provide a kinetic pathway for the ATPase reaction at the noncatalytic
sites of the rho hexamer. Combined with the recently published model of
ATPase reaction at the catalytic site (12), it allows us to make a comparison between the ATPase reaction pathway at the catalytic versus the noncatalytic sites (Fig.
4). Although the two pathways are shown
as independent ones, note that they occur simultaneously at the two
types of sites on the rho hexamer. Fig. 4A summarizes the
ATPase reaction at the noncatalytic sites. Three ATPs bind to the
noncatalytic sites of rho with an apparent bimolecular rate constant of
5.4 × 105 M1
s1. These ATPs are hydrolyzed at a rate of about 1.8 s1, and the products, ADP and/or Pi,
dissociate from the active site at a rate of 0.02 s1. The
dissociation rate was obtained from previous studies (18). In contrast,
hydrolysis of ATP has been proposed to be sequential at the catalytic
sites as shown in Fig. 4B (12). The rho-ATP complex at the
catalytic site is formed with an apparent bimolecular rate of 3 × 106 M1 s1,
estimated from the ratio
kcat/Km. Only one ATP is
hydrolyzed at a time at the catalytic site. This rate is estimated at
>240 s1 from this study as well as a published study on
the ATPase kinetics (12). In that report (12), a pre-steady-state burst
of ATP hydrolysis was observed, implying that a step following ATP
hydrolysis is a rate-limiting step. After the rate-limiting step, the
next catalytic subunit presumably goes through the same cycle of ATP binding and hydrolysis. When we compare the ATPase kinetics at the two
types of sites, it is clear that sequential hydrolysis of ATP does not
occur at the noncatalytic sites. The results in this paper show that
all three ATPs are hydrolyzed simultaneously at the noncatalytic sites
at a slow rate relative to the catalytic site (Fig. 2, B and
C). Hence, there does not appear to be any cooperativity in
ATP binding and hydrolysis at the noncatalytic sites. The rate of ATP
hydrolysis is at least 130 times slower (>240 s1/1.8
s1), and product release is 1500 times slower at the
noncatalytic sites compared with the corresponding steps at the
catalytic site, assuming that kcat is largely
determined by the product release step at the catalytic sites. This
indicates that the ATP and/or the ADP and Pi species are
bound to the noncatalytic sites, whereas multiple ATPase turnovers
occur at the catalytic sites.
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Fig. 4.
Comparison of the mechanism of ATP hydrolysis
at the noncatalytic and catalytic sites of rho. The
shaded subunits in the hexamer depict the catalytic sites on
the rho hexamer. They are shown alternating with the noncatalytic
subunits as seen in the F1-ATPase protein. The symbol
A denotes a tightly bound ATP; D,
ADP·Pi; and a, a loosely bound ATP that
readily exchanges with free ATP in the medium. A, the ATPase
kinetic pathway at the noncatalytic sites. B, the sequential
ATPase reaction at the catalytic sites. An estimate for the bimolecular
rate of ATP binding was obtained from
kcat/Km = 3.3 × 106 M1 s1
(12).
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Role of the Noncatalytic Sites--
Noncatalytic sites have been
identified thus far in three hexameric proteins including rho protein,
the T7 gene 4 helicase (8), and the F1-ATPase (22, 23). In
the 33 structure of
F1-ATPase, the noncatalytic sites reside primarily on the
subunits. The ATP binding site of the subunits lacks critical amino acids that are necessary for catalysis of ATP hydrolysis. Thus,
hydrolysis of ATP at the noncatalytic sites of F1-ATPase is
not easily detectable. Because rho is a homohexamer, the catalytic and
noncatalytic active sites must be induced either during hexamer formation or upon ATP and/or RNA binding. If the catalytic and noncatalytic active sites of rho contain the same amino acids, then the
observed difference in the ATPase rates at the two sites suggests that
the conformation of some of the critical amino acid residues that are
involved in ATP hydrolysis are different. This hypothesis will be
tested when the high resolution structure of rho hexamer in the
presence of ATP is available.
The noncatalytic sites were first identified in the
F1-ATPase protein. However, the exact role of the
noncatalytic sites in F1-ATPase is still unclear.
Nevertheless several studies indicate that the noncatalytic sites are
essential for optimal ATPase activity. It has been suggested that the
fully functional noncatalytic sites in F1-ATPase are
necessary for maintaining cooperative catalysis at the catalytic sites.
When wild-type subunits in F1-ATPase hexamer were
replaced with mutant subunits defective in ATP binding, the ATP
hydrolysis at the catalytic sites was inhibited in a
time-dependent manner (24). This time-dependent
inhibition was shown to be because of tight binding of inhibitory
Mg-ADP at the catalytic sites. These results suggest that the fully
functional noncatalytic sites in F1-ATPase are required for
continuous ATPase turnover at the catalytic sites.
Two experiments with rho suggest that the noncatalytic sites in rho are
essential. When two to three subunits of the wild-type rho hexamer were
exchanged for an inactive mutant protein, a loss in the ATPase activity
of rho was observed (25). Some of these substituted subunits could be
the noncatalytic subunits, but it can be argued also that this
inhibition was because of the loss in the function of the catalytic
sites. Inactivation of the rho ATPase was also observed when one rho
subunit was covalently modified with 8-azido-ATP (26). Assuming that
the catalytic and noncatalytic sites have an equal chance of being
covalently modified, the results argue that modification of the
noncatalytic sites inhibits ATP hydrolysis at the catalytic sites.
Is the hydrolysis of ATP at the noncatalytic sites coupled to movement
on the RNA? To address this question, we have calculated the coupling
constants (nt1 moved/ATP
hydrolyzed) for the catalytic and noncatalytic sites. Based on these
numbers, we conclude that the conformational changes resulting from the
hydrolysis of ATP at the noncatalytic sites cannot be directly coupled
to movement. Walstrom et al. (27, 28) have reported that rho
translocates along the RNA at a rate of ~20 nt/s and consumes about
1-2 ATP when translocating about 1 nucleotide of RNA. This provides a
coupling constant of about 0.5-1 nt/ATP hydrolyzed for the catalytic
sites. Similarly, Brennan et al. (29) have reported a
coupling constant close to 0.06 nt/ATP hydrolyzed. The ATP turnover at
the noncatalytic sites of rho occurs at 0.02 s1 (18). To
calculate the coupling constant for the noncatalytic sites, we have
corrected the above translocation rate (decreased 4-fold) because that
number was measured at 37 °C and the turnover number of 0.02 s1 at the noncatalytic sites was measured at 18 °C.
According to this rough calculation, if the hydrolysis at the
noncatalytic sites were directly coupled to movement of rho along the
RNA, then the rho protein will move close to 250 nt/ATP hydrolyzed. We
believe that this is unlikely because rho binds only about 60-70 nt of
RNA (30). We therefore propose that the energy from ATP hydrolysis at
the noncatalytic sites alone is not utilized directly for translocation
or for nucleic acid unwinding. At this time, the role of ATP binding or
its hydrolysis at the noncatalytic sites is unclear. The ATP at the
noncatalytic sites may play a regulatory role analogous to that
proposed for the noncatalytic sites of F1-ATPase; that is,
these sites mediate cooperative catalysis of ATP at the catalytic sites
and/or ensure continuous ATP hydrolysis at the catalytic sites.
Alternatively, the noncatalytic subunits may participate in
interactions with the nucleic acid during active ATP hydrolysis at the
catalytic site and thus facilitate processive translocation of the
protein on the nucleic acid.
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ACKNOWLEDGEMENT |
We thank Dr. Katsuya Shigesada (Kyoto
University, Japan) for the gift of purified rho protein and for
critical discussion of some of these studies.
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FOOTNOTES |
*
This research was supported by the National Institutes of
Health Grant GM55310.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.
Current address: Dept. of Biochemistry, Robert Wood Johnson
Medical School, 675 Hoes Ln., Piscataway, NJ 08854.
§
To whom correspondence should be addressed. Tel.: 732-235-3372;
Fax: 732-235-4783; E-mail: patelss@umdnj.edu.
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ABBREVIATIONS |
The abbreviation used is:
nt(s), nucleotide(s).
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