From the Division of Biochemistry and Molecular Biology, Louisiana State University, Baton Rouge, Louisiana 70803
Received for publication, February 15, 2001, and in revised form, April 3, 2001
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Acetyl-CoA carboxylase catalyzes the first
committed step in fatty acid synthesis in all plants, animals, and
bacteria. The Escherichia coli form is a multimeric protein
complex consisting of three distinct and separate components: biotin
carboxylase, carboxyltransferase, and the biotin carboxyl carrier
protein. The biotin carboxylase component catalyzes the
ATP-dependent carboxylation of biotin using bicarbonate as
the carboxylate source and has a distinct architecture that is
characteristic of the ATP-grasp superfamily of enzymes. Included in
this superfamily are D-Ala D-Ala ligase,
glutathione synthetase, carbamyl phosphate synthetase, N5-carboxyaminoimidazole ribonucleotide
synthetase, and glycinamide ribonucleotide transformylase, all of which
have known three-dimensional structures and contain a number of highly
conserved residues between them. Four of these residues of biotin
carboxylase, Lys-116, Lys-159, His-209, and Glu-276, were
selected for site-directed mutagenesis studies based on their
structural homology with conserved residues of other ATP-grasp enzymes.
These mutants were subjected to kinetic analysis to characterize their
roles in substrate binding and catalysis. In all four mutants, the
Km value for ATP was significantly increased,
implicating these residues in the binding of ATP. This result is
consistent with the crystal structures of several other ATP-grasp
enzymes, which have shown specific interactions between the
corresponding homologous residues and cocrystallized ADP or nucleotide
analogs. In addition, the maximal velocity of the reaction was
significantly reduced (between 30- and 260-fold) in the 4 mutants
relative to wild type. The data suggest that the mutations have
misaligned the reactants for optimal catalysis.
Acetyl-CoA carboxylase catalyzes the first committed step in long
chain fatty acid synthesis, namely the formation of malonyl-CoA from
acetyl-CoA, MgATP, and bicarbonate. Found in all plants, animals, and
bacteria, this enzyme is biotin-dependent, with the following two-step reaction mechanism (1).
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
The Escherichia coli form of this enzyme consists of
three separable components. The biotin carboxylase component catalyzes the first half-reaction, which involves the phosphorylation of bicarbonate to form a carboxyphosphate intermediate, followed by the
transfer of the carboxyl group to the 1' nitrogen of biotin (2). The
carboxyltransferase component catalyzes the second half-reaction.
In vivo the biotin molecule is linked to the biotin carboxyl
carrier protein through an amide bond to a specific lysine residue.
Both biotin carboxylase and carboxyltransferase retain activity in the
absence of the other two components and will also use free biotin as a
substrate (3). The crystal structure of the biotin carboxylase
component has been solved and is the only three-dimensional structure
of a biotin-dependent carboxylase, making it the paradigm
for structure-function analysis of this class of enzymes (4).
Two years after the solution of the crystal structure, Artymiuk et al. (5) observed that biotin carboxylase had a strong structural homology to glutathione synthetase and D-Ala D-Ala ligase. Despite the remarkable similarity in the three-dimensional structures of biotin carboxylase, D-Ala D-Ala ligase, and glutathione synthetase, there is only an 11% primary sequence identity between the three enzymes (5). Although biotin carboxylase is metabolically unrelated to these two enzymes, all three enzymes are mechanistically homologous in that they catalyze the ATP-dependent ligation of a carboxylate-containing substrate to an amine-containing substrate via formation of an acylphosphate intermediate (5, 6). Structural similarity between the three enzymes includes a common three-domain architecture in which the flexible central domain extends away from the main body of the protein. The crystal structure of biotin carboxylase was originally determined in the absence of any ligands or substrate analogs (4), and its central domain (known as the B-domain) was in the "open" conformation, extending far out from the main body of the enzyme. In contrast, the structures of D-Ala D-Ala ligase and glutathione synthetase were solved in the presence of ADP and ATP, respectively, which revealed that the central domain forms a "lid" that clamps down over the active site upon nucleotide binding (7, 8). Using the structures of D-Ala D-Ala ligase and glutathione synthetase, Artymiuk et al. (5) identified several active-site residues of biotin carboxylase as potentially important for catalysis; among these were Lys-116, Lys-159, His-209, Lys-238, Glu-276, Glu-288 and Asn-290. Soon after the observations of Artymiuk et al. (5), the three-dimensional structure of carbamyl phosphate synthetase was reported and found to be homologous to biotin carboxylase, D-Ala D-Ala ligase, and glutathione synthetase (9). The structural and mechanistic similarity of all four enzymes suggested they were linked through evolution, and thus, they became the charter members of the ATP-grasp family of enzymes. The name "ATP-grasp" derives from the novel nucleotide binding fold observed in these enzymes. The ATP-grasp family of enzymes expanded even further to include several enzymes involved in purine biosynthesis based on a position-specific iterative BLAST sequence alignment (6, 10). The three-dimensional structures of two of these enzymes, N5-carboxyaminoimidazole ribonucleotide synthetase (11) and glycinamide ribonucleotide transformylase, (12) have been determined with nucleotides bound.
The sequence analysis studies identified several residues as being
strictly conserved throughout the entire ATP-grasp family of enzymes.
Not surprisingly, the conserved residues in biotin carboxylase were
Lys-116, Lys-159, His-209, Glu-276, Glu-288, and Asn-290. Site-directed
mutagenesis studies of Glu-288 and Asn-290 confirmed that these two
residues were indeed important for catalysis (13). In fact, mutation of
Glu-288 to lysine resulted in a completely inactive mutant (14).
Recently, Thoden et al. (14) determined the crystal
structure of the inactive mutant of biotin carboxylase, E288K,
cocrystallized with ATP. The structure showed that the B-domain of
biotin carboxylase does exhibit the characteristic "trap door"
closure in the presence of nucleotide, with some atoms moving by more
than 8 Å. As expected, comparison of the structure of the mutant
biotin carboxylase-ATP complex with the structures of the other enzymes
of the ATP-grasp superfamily revealed a significant degree of homology.
For example, Lys-116 of biotin carboxylase and the residue homologous
to Lys-116 in the other ATP-grasp enzymes were found to interact with
ATP. However, there were some notable differences between the structure
of biotin carboxylase and the structures of the other ATP-grasp
enzymes. Namely, the biotin carboxylase crystal structure suggested
that Lys-159, His-209, and Glu-276 did not interact with ATP, whereas the structures of the other ATP-grasp enzymes indicated that these residues did interact with ATP. Thus, the objective of this study is to
test the hypothesis that residues Lys-116, Lys-159, His-209, and
Glu-276 of biotin carboxylase are involved in binding ATP.
![]() |
MATERIALS AND METHODS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Chemicals and Enzymes-- Sodium bicarbonate labeled with 14C was from Amersham Pharmacia Biotech and had a specific activity of 0.1 mCi/mmol. His binding resin was from Novagen. Pyruvate kinase was from Roche Molecular Biochemicals. Restriction grade bovine thrombin was from Enzyme Research Laboratories. Primers were synthesized by Life Technologies, Inc. All other reagents were from Sigma or Aldrich. The growth and purification of wild type and mutant forms of biotin carboxylase were performed as previously described (13).
Site-directed Mutagenesis-- Site-directed mutagenesis of biotin carboxylase was carried out by the PCR method of overlap extension as previously described (13). The following mutants were constructed: H209A, E276Q, K159Q, K116Q, and K116A. The pairs of internal mutagenic primers used to make each site-directed mutant can be found in Table I. The entire gene of each mutant was sequenced to confirm that the desired mutation was made and that no other mutations were incorporated during polymerase chain reaction.
|
Enzymatic Assays-- The rate of ATP hydrolysis by biotin carboxylase in the absence or presence of biotin was measured spectrophotometrically by coupling the production of ADP to pyruvate kinase and lactate dehydrogenase and monitoring the oxidation of NADH at 340 nm. Each assay contained 0.5 mM phosphoenolpyruvate, 0.2 mM NADH, 10 units of pyruvate kinase, 18 units of lactate dehydrogenase, and 100 mM HEPES at pH 8. To ensure the formation of the MgATP complex, MgCl2 was included at concentrations at least twice that of the highest concentration of ATP in each assay. Since the Km for biotin is high (134 mM), the ionic strength of the reaction mixture was held constant with KCl when the initial velocities were measured as a function of biotin concentration.
For experiments in which bicarbonate was varied, all solutions (except for coupling enzymes, which were diluted into degassed buffers) were degassed to lower the level of endogenous bicarbonate (13) and stored in septum vials capped with rubber septa. All assay reactions were performed in a total volume of 1 ml and included the following components: 60 mM biotin, 0.5 mM phosphoenol pyruvate, 0.2 mM NADH, 21 units of pyruvate kinase, 35 units of lactate dehydrogenase, and 100 mM HEPES at pH 8.
For experiments in which the concentration of magnesium was varied, the N-terminal His tag on biotin carboxylase was removed by thrombin cleavage to eliminate the possibility of magnesium binding to the His tag. A ratio of two units of thrombin per unit of biotin carboxylase was used (13).
The rate of ATP synthesis from MgADP and carbamyl phosphate was determined spectrophotometrically. The formation of ATP was coupled to hexokinase and glucose-6-phosphate dehydrogenase, with the production of NADPH monitored at 340 nm. Each assay contained 0.5 mM glucose, 0.4 mM NADP, 2.5 units of hexokinase, 2.5 units of glucose-6-phosphate dehydrogenase, 100 mM KCl, and 100 mM HEPES at pH 8. To ensure the formation of the MgADP complex, MgCl2 was included and held at concentrations at least twice that of the highest concentration of ADP in the assay.
Initial velocities were measured using a Uvikon 810 (Kontron Instruments) spectrophotometer interfaced to a PC equipped with a data acquisition program. The temperature of the reactions was maintained at 25 °C by a circulating water bath. All reactions were initiated by the addition of enzyme. Kinetic parameters were calculated per active site using a molecular mass of 50,000 daltons for the biotin carboxylase monomer (biotin carboxylase exists as a homodimer).
To determine if there was a stoichiometric production of ADP and carboxybiotin, the amount of carboxybiotin produced by biotin carboxylase was determined using a 14C fixation assay and compared with the production of ADP as previously described (13). The reaction mixtures contained 20 mM ATP, 70 mM bicarbonate, 100 mM biotin, 50 mM MgCl2, and 100 mM HEPES at pH 8 in a total volume of 0.5 ml.
Data Analysis--
The Km and
Vmax parameters were determined by nonlinear
regression analysis of the velocity versus [substrate]
data to the Michaelis-Menten equation using the program Enzfitter.
![]() |
RESULTS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Bicarbonate-dependent ATPase Reaction-- In the absence of biotin, biotin carboxylase from E. coli catalyzes a bicarbonate-dependent ATP hydrolysis (Reaction 1) as follows.
![]() |
![]() |
|
The Km for bicarbonate for each of the mutants was determined at fixed, nonsaturating levels of ATP and biotin. Since it was not possible to achieve saturation with biotin or with ATP, the Km values for bicarbonate are apparent Km values. The apparent Km values for wild type biotin carboxylase and the four mutants are shown in Table II. The mutations did not significantly affect the apparent Km for any mutant except that of H209A, where the Km was 15 times greater than that of wild type.
Biotin-dependent ATPase Reaction--
In the presence
of biotin, biotin carboxylase from E. coli catalyzes the
phosphorylation of bicarbonate by ATP to form a carboxyphosphate intermediate. The carboxyl group is then transferred from
carboxyphosphate to biotin to form carboxybiotin. When ATP hydrolysis
activity was examined in the presence of biotin, the
Km values for biotin for all mutants were not
significantly different compared with the wild type enzyme (Table
III). The Km values
for biotin are apparent because it was not possible to saturate with ATP. The largest change in Km was exhibited by H209A but was less than 10-fold. However, Vmax for
three of the four mutants decreased significantly between 30- and 200- fold. The one exception is K116Q, for which a Km
value could not be obtained; the activity of this mutant did not
increase with the addition of biotin over a range of 1-300
mM. To further investigate the role of this lysine residue
in the biotin-dependent ATPase reaction, a K116A mutant was
constructed, and the initial velocity as a function of biotin
concentration was measured. The Km for biotin for
the K116A mutant was 147 ± 12 mM, whereas the maximal velocity was 1.03 ± 0.04 min1. The
effect of this mutation was similar to that of the other three in that
the Vmax rather than the Km
was significantly altered with respect to wild type. These data suggest
that the lack of stimulation of ATP hydrolysis by biotin for K116Q
reflects a function of the mutant glutamine residue rather than of the role of the native lysine residue.
|
Since the ATPase assay measured the production of ADP in the presence and absence of biotin, the question still remained as to whether carboxybiotin was being produced by the mutant enzymes. In other words, is there a 1:1 stoichiometry for the formation of ADP and carboxybiotin or is the hydrolysis of ATP uncoupled from the formation of carboxybiotin? If the 1:1 ratio were altered, this would suggest that the mutations had affected the carboxyl transfer step. The ratio of carboxybiotin to ADP produced during the ATPase reaction for wild type and the four mutants was determined (Table IV). All four mutants produced a ratio of carboxybiotin to ADP that was nearly 1:1. These results indicated that the mutations did not prevent the production of carboxybiotin, and therefore, the carboxyl transfer step had not been uncoupled from the hydrolysis of ATP.
|
ATP Synthesis Reaction-- Biotin carboxylase from E. coli has been shown to catalyze the transfer of the phosphoryl group of carbamyl phosphate to ADP to form ATP and carbamate as follows.
![]() |
![]() |
The kinetic parameters for the ATP synthesis reaction were determined in the absence of biotin (Table V). The mutations did not have a significant effect on the Km for either carbamyl phosphate or ADP. However, a modest decrease in Vmax of 4-fold or less was observed.
|
To test the ability of biotin to stimulate the phosphoryl transfer reaction of the wild type and four mutants of biotin carboxylase, initial velocities were measured at a saturating concentration of ADP and carbamyl phosphate, both in the absence and presence of 60 mM biotin (Table VI). The degree of stimulation of the ATP synthesis activity by biotin was decreased 10-fold by the E276Q mutation, whereas the H209A mutant showed no significant decrease in stimulation. Both K116Q and K159Q showed a decrease of ~2.5-fold compared with wild type (Table VI).
|
Magnesium Assay--
Biotin carboxylase requires two equivalents
of magnesium for activity. One equivalent is complexed to ATP, whereas
the role of the other equivalent is unknown. The effect of the four
mutations on the ability of magnesium to stimulate the
biotin-dependent ATPase activity of wild type and mutant
biotin carboxylase was evaluated by measuring initial velocity as a
function of [MgCl2]. All four mutants exhibited a
dependence on MgCl2 similar to that of wild type. This
suggests that these mutations did not affect the affinity of the enzyme
for magnesium (Fig. 1).
|
![]() |
DISCUSSION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
The objective of this study was to test the hypothesis that four
residues of biotin carboxylase, Lys-116, Lys-159, Glu-276, and His-209,
were involved in binding ATP. Each of the corresponding site-directed
mutants displayed an elevated Km value for ATP
relative to the wild type value. This suggests that all four conserved
active-site residues bind ATP. For the K116Q mutant, the increased
Km value for ATP was consistent with the three-dimensional structure of biotin carboxylase bound to ATP as well
as with the three-dimensional structures of other ATP-grasp enzymes
bound to ADP or AMPPNP.1 The
crystal structure of biotin carboxylase complexed with ATP revealed an
electrostatic interaction between the -amino group of Lys-116 and
the
-phosphoryl oxygen of ATP (14). As shown in Table
VII, the residues homologous to Lys-116
in enzymes of the ATP-grasp family also interacted with the oxygens of
the
or
phosphates, as determined by crystallography. Moreover,
mutation of the homologous residue in the carboxyphosphate domain of
carbamyl phosphate synthetase resulted in a 5-fold increase in the
Km for ATP (17).
|
Although all the available crystallographic and kinetic data implicate
Lys-116 in binding an or
phosphate oxygen, roles for the other
three residues are not as well defined. Site-directed mutagenesis data
for biotin carboxylase and the crystal structures of other ATP-grasp
enzymes suggest differing roles for Lys-159, His-209, and Glu-276.
First, the
-amino group of Lys-159 was not implicated in ATP binding
based on the crystal structure of biotin carboxylase complexed with
ATP. However, the residues homologous to Lys-159 in other ATP-grasp
enzymes have been shown to interact with the nucleotide (Table VII),
which supports the 90-fold increase in Km for ATP in
the K159Q mutant of biotin carboxylase. Site-directed mutagenesis of
the homologous residues of carbamyl phosphate synthetase
(carboxyphosphate domain) and D-Ala D-Ala ligase revealed a Km for ATP that was 31- and
50-fold higher than the wild type value, respectively (17, 18).
Together these data implicate Lys-159 in binding to ATP.
Second, the crystal structure of biotin carboxylase complexed with ATP showed that the imidazole group of His-209 is about 4 Å from the hydroxyl groups of the ribose of ATP, whereas the crystal structures of other ATP-grasp enzymes showed that the residue homologous to His-209 hydrogen bonded to the 2' and 3' hydroxyl groups of the ribose (Table VII). As with K159Q, the Km for ATP in the H209A mutant of biotin carboxylase was elevated 70-fold relative to wild type. Again, the structural data from other ATP-grasp enzymes and the site-directed mutagenesis results of biotin carboxylase strongly suggest that His-209 interacts with ATP, presumably with the hydroxyl groups of ribose.
Third, when the three-dimensional structure of biotin carboxylase with
ATP was modeled with bound biotin, the ureido ring of biotin was
located near the side chain of Glu-276 (14). However, three-dimensional
structures of other ATP-grasp enzymes showed that the residue
homologous to Glu-276 coordinated to either magnesium or manganese and
that the divalent cation in each structure coordinated to the
phosphoryl oxygens of the phosphate group (Table VII). Site-directed mutagenesis of Glu-276 in biotin carboxylase indicated no
change in the Km for biotin but a nearly 40-fold
increase in the Km for ATP. This result supports the
hypothesis that Glu-276 coordinates to a magnesium ion in biotin
carboxylase and that mutation of Glu-276 distorts the binding of
magnesium and in turn the binding of ATP. It should be noted that
although the mutation of Glu-276 altered the binding of magnesium ion
to the residue, it did not appear to change the affinity of magnesium for the enzyme (Fig. 1). Additionally, when the homologous residue in
the carboxyphosphate domain of carbamyl phosphate synthetase was
mutated to alanine, the Km for ATP for the resulting mutant was 19-fold higher than wild type carbamyl phosphate synthetase (17). Again, the mutagenesis data for biotin carboxylase were consistent with the structures and mutagenesis data for the other ATP-grasp enzymes and inconsistent with the biotin carboxylase structure complexed with ATP.
The discrepancies described above can be explained by noting that the biotin carboxylase used in the crystal structure complexed with ATP contained a mutation of a critical residue (E288K). Although this mutation crippled the ability of the enzyme to hydrolyze ATP, it allowed crystals to be grown in the presence of ATP. The structural data from the other ATP-grasp enzymes in Table VII were obtained from wild type protein. Furthermore, the crystal structure of biotin carboxylase did not contain a divalent cation coordinated to ATP. Thus, the crystal structure of the E288K mutant form of biotin carboxylase complexed with ATP may not be a completely accurate description for the binding of ATP to this enzyme, which would explain the disagreement between the mutagenesis data and the crystal structure. Moreover, the evidence that ATP was bound incorrectly in the E288K mutant of biotin carboxylase may explain, at least in part, the lack of activity for the E288K mutant and provide insight into why the maximal velocities of the biotin-dependent ATPase activity for the four mutants characterized in this paper were significantly decreased.
Although the increase in the Michaelis constants of the four mutants of biotin carboxylase strongly suggests that these residues are involved in binding ATP, the concomitant decrease in their maximal velocities indicates that these residues also play a role in catalysis. If these four residues were solely involved in binding ATP, then the corresponding mutants should have the same maximal velocity as wild type biotin carboxylase. This brings us to the question of what role these residues could play in catalysis that would be consistent with their role in binding ATP. Any discussion of the catalytic roles of these residues must begin with the observation that biotin carboxylase exhibits the phenomenon of substrate-induced synergism with respect to biotin (13). That is, in the absence of biotin, the enzyme will cleave ATP into ADP and Pi in a bicarbonate-dependent manner, albeit at a very slow rate. However, in the presence of biotin, the rate of ATP hydrolysis increases 1100-fold. Thus, the hydrolysis of ATP is synergistic with the binding of biotin. A possible explanation for substrate-induced synergism in biotin carboxylase that is consistent with the data presented in this paper is that in the absence of biotin ATP binds to the enzyme in a large number of nonproductive modes, which is manifested as a low maximal velocity. However, upon the binding of biotin to biotin carboxylase, the number of nonproductive binding modes of ATP is reduced, allowing for a more reactive alignment of the substrates. This phenomenon is manifested as a dramatic increase in the maximal velocity. Recently, a more sophisticated version of this concept has been proposed (19, 20). In the current theory, the enzyme molecule pre-organizes the active site to allow the substrates to form near attack conformers. A near attack conformer refers to the juxtaposition of the substrates in the ground state such that they closely resemble the transition state. The effect of mutating any of the four residues in this study would be a shift in the active site geometry, which would possibly reduce the number of near attack conformers. This is manifested as a significantly reduced maximal velocity for the mutant enzyme. Biotin cannot properly cause the alignment of ATP for the reaction because of greater conformational flexibility of ATP in the active site due to the mutation. This concept of greater conformational flexibility of ATP is supported by both the increase in Km for ATP in each of the mutant enzymes and the presumably incorrect binding of ATP in the active site of the inactive E288K mutant. Recall that none of the mutants affected the carboxyl transfer from carboxyphosphate to biotin, yet the Vmax values were significantly decreased. Thus, the fact that these four ATP binding residues are conserved throughout the ATP-grasp superfamily of enzymes further attests to the notion that binding interactions and correct positioning of the substrates appear to play the dominant role in catalysis by biotin carboxylase.
The question is now how does the binding of biotin to the enzyme position the substrates into a more reactive conformation. A conformational change (i.e. induced fit) in biotin carboxylase upon biotin binding could explain the large increase in rate for ATP hydrolysis. However, the major conformational change in biotin carboxylase occurs upon the binding of ATP (14), which binds to the enzyme first before bicarbonate and biotin (21). A crystal structure of biotin carboxylase with only biotin bound showed no difference in conformation compared with the unliganded structure of the enzyme (4). The lack of a large conformational change in biotin carboxylase upon binding biotin is consistent with the high Km for biotin (134 mM). Using this value as an apparent binding constant for biotin to calculate the binding energy, a relatively low value of 1.2 kcal/mol is obtained. The low binding energy of biotin to the enzyme is not suggestive of a large conformational change.
How then is biotin able to stimulate the rate of ATP hydrolysis if not
via a conformational change? Perhaps biotin only promotes very small
changes in the enzyme that result in the alignment of substrates for
catalysis. To this end, recent studies on hydrogen tunneling in
dehydrogenases have found a correlation between protein dynamics and
enzymatic activity (22, 23). Particularly intriguing is the case of
isocitrate dehydrogenase, to which the binding of its substrate,
isocitrate, induces shifts of less than an angstrom in the amino acid
side chains of the active site. These seemingly insignificant changes
in conformation are in fact, related to rate increases of many orders
of magnitude (24, 25). Thus, it may be that very subtle dynamic
behavior of biotin carboxylase is enough to generate the large increase
in the rate of ATP hydrolysis upon the binding of biotin. Further
studies will be required to determine this aspect of the mechanism. In
summary, the four active-site residues of biotin carboxylase, Lys-116,
Lys-159, His-209, and Glu-276, were shown to be involved in binding
ATP. Furthermore, these four residues have also been found to be
involved in catalysis, and their role in catalysis is to orient ATP in
a conformation that allows for optimal catalysis. Finally, the results
also suggest that the crystal structure of the mutant biotin
carboxylase, E288K complexed with ATP, may not be a completely accurate
depiction for the binding of ATP to the wild type form of biotin carboxylase.
![]() |
FOOTNOTES |
---|
* This research was supported in part by National Institutes of Health Grant GM51261 (to G. L. W.).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.
Supported by a Howard Hughes Medical Institute grant through the
Undergraduate Biological Sciences Education Program to Louisiana State University.
§ To whom correspondence should be addressed: Dept. of Biological Sciences, Rm. 508 Life Sciences Bldg., Louisiana State University, Baton Rouge, LA 70803. Tel.: 225-388-5209; Fax: 225-388-4638; E-mail: gwaldro@lsu.edu.
Published, JBC Papers in Press, May 9, 2001, DOI 10.1074/jbc.M101472200
![]() |
ABBREVIATIONS |
---|
The abbreviation used is:
AMPPNP, adenosine
5'-(,
-imino)- triphosphate.
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1. | Wakil, S. J., Stopps, J. K., and Joshi, V. C. (1983) Annu. Rev. Biochem. 52, 537-579[CrossRef][Medline] [Order article via Infotrieve] |
2. | Knowles, J. R. (1989) Annu. Rev. Biochem. 58, 195-221[CrossRef][Medline] [Order article via Infotrieve] |
3. |
Guchhait, R. B.,
Polakis, S. E.,
Dimroth, P.,
Stoll, E.,
Moss, J.,
and Lane, D. M.
(1974)
J. Biol. Chem.
249,
6633-6645 |
4. | Waldrop, G. L., Rayment, I., and Holden, H. (1994) Biochemistry 33, 10249-10256[Medline] [Order article via Infotrieve] |
5. | Artymiuk, P. J., Poirrette, A. R., Rice, D. W., and Willett, P. (1996) Nat. Struct. Biol. 3, 128-132[Medline] [Order article via Infotrieve] |
6. |
Galperin, M. Y.,
and Koonin, E. V.
(1997)
Protein Sci.
6,
2639-2643 |
7. | Fan, C., Moews, P. C., Walsh, C. T., and Knox, J. R. (1994) Science 266, 439-443[Medline] [Order article via Infotrieve] |
8. | Yamaguchi, H., Kato, H., Hata, Y., Nishioka, T., Kimura, A., Oda, J., and Katsube, Y. (1993) J. Mol. Biol. 229, 1083-1100[CrossRef][Medline] [Order article via Infotrieve] |
9. | Thoden, J. B., Holden, H. M., Wesenberg, G., Raushel, F. M., and Rayment, I. (1997) Biochemistry 36, 6305-6316[CrossRef][Medline] [Order article via Infotrieve] |
10. | Kanai, S., and Toh, H. (1999) J. Mol. Evol. 48, 482-492[Medline] [Order article via Infotrieve] |
11. | Thoden, J. B., Kappock, T. J., Stubbe, J., and Holden, H. M. (1999) Biochemistry 38, 15480-15492[CrossRef][Medline] [Order article via Infotrieve] |
12. | Thoden, J. B., Firestine, S., Nixon, A., Benkovic, S. J., and Holden, H. M. (2000) Biochemistry 39, 8791-8802[CrossRef][Medline] [Order article via Infotrieve] |
13. | Blanchard, C. Z., Lee, Y. M., Frantom, P. A., and Waldrop, G. L. (1999) Biochemistry 38, 3393-3400[CrossRef][Medline] [Order article via Infotrieve] |
14. |
Thoden, J. B.,
Blanchard, C. Z.,
Holden, H. M.,
and Waldrop, G. L.
(2000)
J. Biol. Chem.
275,
16183-16190 |
15. | Ogita, T., and Knowles, J. R. (1988) Biochemistry 27, 8028-8033[Medline] [Order article via Infotrieve] |
16. |
Polakis, S. E.,
Guchhait, R. B.,
Zwergel, E. E.,
Lane, M. D.,
and Cooper, T. G.
(1974)
J. Biol. Chem.
249,
6657-6667 |
17. | Stapleton, M. A., Javid-Majd, F., Harmon, M. F., Hanks, B. A., Grahmann, J. L., Mullins, L. S., and Raushel, F. M. (1996) Biochemistry 35, 14352-14361[CrossRef][Medline] [Order article via Infotrieve] |
18. | Shi, Y., and Walsh, C. T. (1995) Biochemistry 34, 2768-2776[Medline] [Order article via Infotrieve] |
19. | Bruice, T. C., and Lightstone, F. C. (1999) Acc. Chem. Res. 32, 127-136[CrossRef] |
20. | Bruice, T. C., and Benkovic, S. J. (2000) Biochemistry 39, 6267-6274[CrossRef][Medline] [Order article via Infotrieve] |
21. | Blanchard, C. Z., Amspacher, D., Strongin, R., and Waldrop, G. L. (1999) Biochem. Biophys. Res. Commun. 266, 466-471[CrossRef][Medline] [Order article via Infotrieve] |
22. | Sutcliffe, M. J., and Scrutton, N. S. (2000) Trends Biochem. Sci. 25, 405-408[CrossRef][Medline] [Order article via Infotrieve] |
23. | Kohen, A., and Klinman, J. H. (2000) J. Am. Chem. Soc. 122, 10738-10739[CrossRef] |
24. | Koshland, D. E. (1998) Nat. Med. 4, 1112-1114[CrossRef][Medline] [Order article via Infotrieve] |
25. | Koshland, D. E. (1994) Angew. Chem. Int. Ed. Engl. 33, 2375-2378[CrossRef] |
26. | Levert, K. L., Lloyd, R. B., and Waldrop, G. L. (2000) Biochemistry 39, 4122-4128[CrossRef][Medline] [Order article via Infotrieve] |
27. | Thoden, J. B., Wesenberg, G., Raushel, F. M., and Holden, H. M. (1999) Biochemistry 38, 2347-2357[CrossRef][Medline] [Order article via Infotrieve] |