From the Department of Biochemistry and Biophysics,
Texas A & M University, College Station, Texas 77843-2128, § Graduate School of Biomedical Sciences, Texas A & M
University System Health Science Center,
College Station, Texas 77843-1114, and ¶ Center for Structural
Biology, Institute of Biosciences and Technology,
Houston, Texas 77030-3303
Received for publication, November 27, 2002, and in revised form, December 26, 2002
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() |
---|
The N-1-(5'-phosphoribosyl)-ATP
transferase catalyzes the first step of the histidine biosynthetic
pathway and is regulated by a feedback mechanism by the product
histidine. The crystal structures of the
N-1-(5'-phosphoribosyl)-ATP transferase from Mycobacterium tuberculosis in complex with inhibitor
histidine and AMP has been determined to 1.8 Å resolution and without
ligands to 2.7 Å resolution. The active enzyme exists primarily as a
dimer, and the histidine-inhibited form is a hexamer. The structure
represents a new fold for a phosphoribosyltransferase, consisting of
three continuous domains. The inhibitor AMP binds in the active
site cavity formed between the two catalytic domains. A model for the mechanism of allosteric inhibition has been derived from conformational differences between the AMP:His-bound and apo structures.
The N-1-(5'-phosphoribosyl)-ATP transferase
(ATP-PRTase)1encoded by
the hisG locus catalyzes the condensation of ATP with PRPP, the first reaction in the histidine biosynthetic pathway. The reaction
is a Mg2+-dependent transfer of the
phosphoribosyl moiety from 5'-phosphoribosyl 1'-pyrophosphate (PRPP) to
the N1 nitrogen of adenosine ring of ATP yielding phosphoribosyl-ATP
and inorganic pyrophosphate (PPi) (Scheme 1) (1). The activity and the
expression of ATP-PRTase are regulated by feedback inhibition and by
repression of the his operon in response to host iron,
respectively (2, 3).
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS AND DISCUSSION
REFERENCES
View larger version (7K):
[in a new window]
Scheme 1.
Given the high energetic costs associated with the synthesis of a histidine molecule and the direct connections of the histidine pathway with purine, pyrimidine, and tryptophan biosynthesis, a multilevel regulatory control has been selectively retained in all bacteria studied to date. Whereas the transcriptional regulation based on nutrient conditions controls the steady-state level of enzyme over several bacterial generations, the feedback inhibition of ATP-PRTase serves as a fine-tuning control that provides rapid regulation of biosynthetic activity as a function of the available histidine.
The ATP-PRTase-catalyzed reaction has been studied for more than 4 decades and was originally believed to proceed via the formation of a 5'-phosphoribosyl enzyme covalent intermediate (4, 5). Detailed kinetic studies refuted the presence of such an intermediate (6). Steady-state studies of the enzymatic reaction in both directions were consistent with a sequential mechanism (7) where ATP binding precedes binding of PRPP (8). The ATP-PRTase reaction has also been shown to be completely reversible as addition of pyrophosphate to phosphoribosyl-ATP yields ATP and PRPP (9). The synergistic inhibition of the enzyme was demonstrated to occur allosterically by histidine and competitively by AMP, ADP, or guanosine tetraphosphate (10). AMP and ADP are both competitive inhibitors with respect to PRPP and ATP (1). Histidine inhibition was first thought to be "noncompetitive" with PRPP and ATP (1). A single histidine was later proposed to interact with more than one molecule of the enzyme, in a site shown to be allosteric in nature (11).
A clear understanding of the molecular basis of ATP-PRTase activity and
the mechanism of its regulation by histidine has been elusive due to
the lack of structural information. Although structures of several
PRTases are known (12), lack of sequence similarity precluded analyses
based on homology modeling. Based on their structural folds, the
PRTases have been subdivided into two groups (13, 14). The type I
PRTases have a central parallel five-stranded -sheet surrounded by
-helices. Type II PRTases, such as quinolinic acid PRTase, have a
modified
/
-barrel as the catalytic core. Association of alternate
structural motifs with PRTases has suggested a convergent evolution of
these enzymes.
In this study we report the structure of ATP-PRTase from
Mycobacterium tuberculosis (mtATP-PRTase) without bound
ligands (apo) and in a ternary complex with the inhibitors AMP and
histidine (AMP:His). These structures represent a new fold for a PRTase with a modular organization of the regulatory histidine binding domain
and catalytic PRTase domains. Comparison of the inhibitor-bound structure with the apo form reveals the structural basis of the allosteric regulation by histidine.
![]() |
EXPERIMENTAL PROCEDURES |
---|
![]() ![]() ![]() ![]() ![]() ![]() |
---|
Materials-- ATP, AMP, L-histidine, and 5,5'-dithiobis(2-nitrobenzoic acid) (DTNB) were purchased from Sigma. PRPP and lithium sulfate were purchased from Fluka. Standard proteins for calibrating gel filtration column were purchased from Amersham Biosciences.
Cloning, Expression, and Purification of mtATP PRTase--
The
hisG gene, Rv2121c from M. tuberculosis H37Rv
genome, was identified from the TubercuList sequence data base (15).
The hisG gene was amplified using M. tuberculosis
genomic DNA as a template. The PCR product was cloned into a pET28a
expression vector (Novagen) with N-terminal His tag and transformed
into Escherichia coli overexpression strain, BL21(DE3).
Cells were incubated at 37 °C until the optical density reached 0.6 and induced with 1 mM
isopropyl-1-thio--D-galactopyranoside, and incubation was continued for an additional 4 h. The bacterial cells were harvested by centrifugation and resuspended in 20 mM sodium
phosphate (pH 7.5) containing 0.5 M NaCl and 0.1 M imidazole. The cells were lysed using a French press. The
cell extract was applied onto a 5-ml nickel-nitrilotriacetic acid
column (Amersham Biosciences), and the target protein was eluted using
an imidazole gradient. The eluate was concentrated by Centriprep
(Amicon) to 20 mg/ml and applied onto a Sephadex 200 gel-filtration
column (Amersham Biosciences) equilibrated with 20 mM HEPES
(1 mM EDTA and dithiothreitol (pH 7.5)) as a final step.
The protein was more than 95% pure as observed on an SDS-PAGE gel.
Selenomethionylated protein was prepared according to published methods
(16). The pET28a-hisG plasmid was transformed into E. coli B834(DE3) (Novagen) Met auxotroph strain. Cells were grown in
LB medium until an optical density of 0.6 was obtained. Cells were
pelleted by centrifugation, washed with LB medium, and resuspended in
M9 minimal medium lacking L-Met. SeMet was then added to a final concentration of 0.05 µg/ml along with 35 µg/ml kanamycin. Cultures were then induced with 1 mM
isopropyl-1-thio--D-galactopyranoside followed by
incubation for 4 h at 37 °C. The protein was purified using the
same methods as for the apoprotein.
Crystallization-- Initial crystallization conditions were obtained using Crystal Screen 2 from Hampton Research. Crystals were grown using the hanging drop vapor diffusion method at 16 °C. The apocrystals were obtained by mixing equal volumes (2-3 µl) of 20 mg/ml protein with a buffer containing 0.1 M MES (pH 6.5) and 1.8 M magnesium sulfate as a precipitant. AMP:His-crystals were obtained in condition number 15 of Crystal Screen 2 from Hampton Research (0.1 M sodium citrate (pH 5.6), 0.5 M ammonium sulfate, and 1.0 M lithium sulfate) in the presence of 5 mM AMP and 100 µM histidine.
Data Collection and Processing--
A complete and redundant
high resolution data set was collected at BioCARS beamline 14BMC at the
Advanced Photon Source, Argonne National Laboratory. Multiple anomalous
dispersion (MAD) data sets were collected for both the apocrystal
(MAD1) and AMP:His crystal (MAD2) (Table I). All data sets were indexed
and scaled using MOSFLM and SCALA of the CCP4 program suite (17). Unit cell dimensions for apocrystal were a = b = 132.5 Å, c = 110.5 Å, =
= 90, and
= 120. Space group was R32. The inhibitor complex crystallized
also in the space group R32, but the cell dimension changed by about 14 and 11% in a, b (113.8 Å), and c (124.3 Å),
respectively. Calculation of solvent content (18) indicated that for
both crystals the asymmetric unit contained one protomer of ATP-PRTase
and 58 (apo) or 48% (AMP:His) solvent.
Structure Determination of ATP-PRTase-- Selenium sites were located using SOLVE (19) with three different wavelength MAD data. The sites were refined using MLPHARE (20), and protein phases were calculated with SHARP (21) (30-3.0 Å) and improved by density modification using CNS (crystallography and NMR system) (22). A polyalanine backbone model was built into the electron density using O (23). Based on marker amino acids such as SeMet, Arg, and aromatic residues, polyalanines were converted to the original sequence. Initial refinement was performed by rigid body refinement, simulated annealing and individual B factor refinement. Initial Rfactor and Rfree were 35 and 42%, respectively. After an intensive series of manual rebuilding and refinement, the Rfactor and Rfree dropped down to 28 and 33%, respectively. Solvent molecules were picked using Xfit (24) and refined. As a final refinement step, the Restrained TLS refinement with Refmac5 (25) was used, and the R factors were 19.2 and 26.1% (Table I, bottom).
Structure Determination of AMP:His Form-- Molecular replacement of the AMP:His-bound data was attempted with the apo structure as a search model. However, any reasonable solution was not obtained from the whole molecule or separate domains. Therefore, another MAD experiment was performed. Four Se sites were determined by SOLVE, and phases were calculated with SHARP up to 2.6 Å. The MAD map was made after solvent flattening with DM (density modification) (26) of the CCP4 program suite. The apo structure was manually fitted into the electron density to make an initial model for the inhibitor-bound structure. Positional refinement and molecular dynamics were performed, and the Rfree was 30%. Shake & Warp (27) was used to remove phase bias from the model. Solvent molecules were picked and the restrained TLS refinement with Refmac5 was performed. The refinement statistics are shown in Table I, bottom.
Cysteine Modification Experiments--
We followed an
experimental procedure described previously (28) for characterizing the
number of free cysteines per molecule of protein. Briefly, 0.1 ml of a
protein solution was added to 3.1 ml of reaction buffer containing 0.3 mM DTNB to achieve a final concentration of 0.3 mg/ml (9.4 µM) of freshly prepared reaction mixture. The absorbance
of 2-nitro-5-thiobenzoate anion (TNB2) was measured at
412 nm until it reached a plateau. The numbers of free cysteines were
calculated from the absorbance (0.18 and 0.35 absorbance units for the
apo and AMP:His form, respectively) and molar absorption coefficient of
TNB2
(14,150 M
1
cm
1) covalently linked to free cysteines. The numbers of
the free cysteines corresponding to the obtained absorbance were 2 and 1 (equivalent to 9.3 and 21.3 µM TNB2
).
Gel Filtration Experiments-- A Superdex 200 gel filtration column (24-ml bed volume, Amersham Biosciences) was used to estimate the molecular weight of ATP-PRTase and to observe the effect of different ligands on oligomerization. The column was calibrated using low and high molecular standard proteins (from Amersham Biosciences) in 20 mM HEPES (pH 7.5), 0.1 M NaCl, 1 mM EDTA, and dithiothreitol. 100 µM histidine and 1 mM AMP were added in the same buffer to observe the change of oligomeric status in the presence of the inhibitors.
In the absence of histidine at 4 °C, more than 99% of the apoenzyme
eluted as a dimer at a low protein concentration (less than 50 µg/ml). We were not able to detect the dimer when the protein was
preincubated at 10 µM histidine; only hexamers and higher
oligomers were detected.
![]() |
RESULTS AND DISCUSSION |
---|
![]() ![]() ![]() ![]() ![]() ![]() |
---|
Overall Structure of mtATP-PRTase-- The x-ray structure of the recombinant mtATP-PRTase was solved from electron density maps calculated by MAD methods using crystals of selenomethionylated protein formed in the space group R32. Crystals were produced in the absence of any ligands or after incubation of protein with two inhibitors, adenosine monophosphate and histidine (AMP:His). The structures have been refined to Rfactors of 19.2 (apo) and 19.8% (AMP:His) at resolutions of 2.7 and 1.8 Å, respectively (Table I). In both cases, the refined structure contains 276 of the 284 residues present in mtATP-PRTase. The residues 186-193 were disordered and omitted from the final model.
|
mtATP-PRTase is an elongated molecule consisting of 10 -helices and 15
-strands (Fig.
1a) composed in 3 domains.
Domain I (residues 1-90, 175-184, and 194-211) contains a central
-sheet consisting of four parallel
-strands (
1,
3,
4,
and
5) and two anti-parallel strands (
2 and
11). The
-sheet
is surrounded by 3
-helices,
1 on one side and
2 and
3 on
the other side. Domain II (residues 91-174) is also an
/
-structure composed of four (
7-10) parallel
-strands and
one (
6) anti-parallel
-strand with two
-helices on each side
(
4 and
5 on one side and
6 and
7 on the other side). Domain
III (residues 212-284) has one
-sheet consisting of four
anti-parallel
-strands (
12-15) with two
-helices (
9 and
10) on one side of the
-sheet.
|
Domains I and II form the catalytic core of ATP-PRTase. The competitive
inhibitor AMP binds in a cleft located between the two domains (Fig.
1b) and makes the most of its bonding interactions with
residues from domain II. The feedback inhibitor histidine was located
far from the active site in domain III (Fig. 1). The electron density
of both inhibitors is shown in Fig. 1, c and d.
The catalytic core of ATP-PRTase (domains I and II) is similar to the
E. coli glutamine-binding protein (Protein
Data Bank code 1WDN; r.m.s.d. 3.4 for 172 C atoms) (29)
(Fig. 2a), an E. coli histidine-binding protein (Protein Data Bank code
1HSL; r.m.s.d. 3.2 for 164 C
atoms) (30) as well as the ligand
binding core of a glutamate receptor from
Synechocystis sp. (Protein Data Bank code
1IIW) (31) and that of rat (Protein Data Bank code 1LB8)
(32).
|
A VAST2 structural similarity
search using domain III found that the domain shares a high degree of
similarity with the E. coli signal transducing protein PII
(Protein Data Bank code 2PII; r.m.s.d. 1.4 for 63 C atoms)
and the guanine nucleotide exchange factor domain from human elongation
factor-1
(Protein Data Bank code 1B64; r.m.s.d. 2.0 for 57 C
atoms). Whereas all the four
-strands and two
-helices are
conserved between structures of PII and domain III of the ATP-PRTase,
the two differ in the length of their connecting loops (i.e.
7 residues longer in the case of PII) (Fig. 2a).
Interestingly, proteins of PII family (34) and GlnK (35) form a trimer
similar to that observed for domain III of ATP-PRTase (35, 36).
Quaternary Structure-- Gel filtration, sedimentation velocity ultracentrifugation, and light scattering experiments on the E. coli enzyme have demonstrated that the ATP-PRTase exists in equilibrium between its active dimeric form (Fig. 2c) and inactive higher oligomeric forms (37-39). Gel filtration experiments showed similar behavior for the mt ATP-PRTase (see "Experimental Procedures"). In general, ATPase hexamers are more abundant at concentrations of enzyme higher than 1 mg/ml or in the presence of stoichiometric AMP, phosphoribosyl-ATP, and histidine and particularly in the combination of one of the nucleotides and histidine (37). On the other hand, low enzyme concentrations (50 µg/ml) or the presence of the substrate PRPP seems to dissociate the hexamers, or higher oligomers, into active dimers (38). Thus regulation of the oligomeric state of ATP-PRTase appears to be an efficient way of controlling the enzyme activity by sensing the intracellular concentrations of both enzyme and histidine. At low in vivo intracellular histidine levels and enzyme concentrations, ATP-PRTase most likely exists as active dimers and constitutively replenishes the histidine pool. Under conditions of high histidine demand, such as active assimilation of nitrogen, transcriptional derepression of the hisG gene perhaps allows even higher intracellular concentration of ATP-PRTase that may be hexameric. However, once the histidine level exceeds the demand, the expression of hisG gene is reduced, and the existing ATP-PRTase is inhibited by histidine.
Some proteobacteria have a shorter version of the ATP-PRTase, missing about 100 residues from the C terminus (domain III). In these bacteria, HisG can associate with another protein HisZ, a parahomolog of aminoacyl-tRNA synthetase that is functionally unknown (40). Recent equilibrium sedimentation studies on HisG and HisZ from Lactococcus lactis show that they individually form stable homodimers. However, together the two proteins form an octameric structure that can be destabilized by allosteric regulators AMP and histidine (41). No homolog of HisZ is found in M. tuberculosis genome. However, given that HisZ is required for the activity and regulation of the truncated HisG, it is tempting to speculate that it may be compensating for some of the functions of the missing domain III. Whereas alternate roles and mechanisms for regulation of HisZ may not be ruled out, quaternary associations of HisG, both homo- or heteromeric, seem to have direct influence on the function and regulation of these enzymes.
In both the apo and AMP:His structures of ATP-PRTase the packing in the
crystal is consistent with a hexamer because of crystallographic 3- and
2-fold symmetry axes in the R32 space group that generates a "trimer
of dimers" (Fig. 3a).
However, comparison of the intersubunit interactions in the two
structures showed that the hexamers are different with the
histidine-containing complex being much more compact than the
apoprotein (Fig. 3, d and e). In the case of the
AMP:His hexamer, the subunit-accessible surface area buried is 3078 Å2, and it is only 2417 Å2 in the apo form.
The dimer interface buries 1203 and 965 Å2 of accessible
surface of each subunit in apo or AMP:His forms, respectively. The
interactions at the dimer interface are primarily from the catalytic
core (domains I and II), whereas those involved at the hexamer
interface are mainly from domain III. The most prominent structural
feature of the AMP:His hexamer is the extended -sheet for domain III
formed by the C-terminal
-strand (residues 280-284) with the
penultimate
-strand (
15, residues 273-276) of the adjacent
subunit (Fig. 3b).
|
Catalytic Site of ATP-PRTase--
The catalytic site of ATP-PRTase
is formed by a cleft located between domains I and II (Fig. 1,
a and b). The substrate-binding sites could be
identified from highly negative electrostatic potential of the protein,
presumably involved in binding to the Mg2+ ions required
for catalysis and by the presence of sulfate ions from the
crystallization buffer, marking the probable binding sites of phosphate
groups of the substrates. The inhibitor AMP (competitive with respect
to ATP) was located in clear electron density from omit maps calculated
from diffraction data collected from crystals of HisG that were
incubated with AMP and His prior to crystallization. AMP bound to the
expected ATP-phosphoribosyltransferase signature sequence region
(Glu141-Leu162), which was identified from
PROSITE (42) (documentation number PDOC01020). Residues from both
domains I and II contributed to the binding of AMP (Fig.
4a). The phosphate of the AMP
is coordinated by the P-loop motif (residues from Asp154 to
Thr161) found in the domain II. One of the phosphate
oxygens of the AMP, O1P, forms hydrogen bonds with backbone
amides of Gly159 and Gly157. O2P makes a
hydrogen bond with OG1 of Thr161, and O3P hydrogen bonds
with backbone amides of Thr161 and Arg160 as
well as with OG of Ser158. O5 of the AMP interacts with the
backbone nitrogen of Ser158. N1 of the adenosine base forms
hydrogen bonds with three ordered water molecules. One of them is a
water-mediated interaction between the N1 of AMP and OD2 of
Asp70. N1 also hydrogen bonds via an ordered water
molecule, with OD1 of Asp70 and OG of Ser90. N6
of the adenosine ring forms a hydrogen bond with OH of
Tyr116. Of these, only the interactions of residues 70 and
90 are from domain I and the others are from domain II. The 2-OH and
3-OH of the AMP are close to domain I of the neighboring subunit and interact with the side chain carboxyls of Asp30' and
Asp33' of that subunit. As these interactions would
contribute to the compactness of the hexamer, they may be responsible
for the synergistic behavior of AMP toward inhibition with
histidine.
|
Analysis of the structure suggests that ATP would bind in a manner very similar to AMP binding. The presence of a tightly associated sulfate ion close to the AMP-binding site along with several basic residues (Arg49, Lys9, Lys32, and Arg160) indicate that PRPP may bind in a region adjacent to the AMP-binding site (Fig. 4b). Moreover, consideration of the catalytic reaction would require that PRPP be oriented such that the C1 carbon of the ribosyl group of PRPP is in close proximity to N1 of the adenine ring of the ATP. The 5'-phosphate of PRPP is more likely to occupy the location occupied by the sulfate ion bound to residues Lys9 and Arg49. In this model, the leaving pyrophosphate group would interact with residue Lys32. The location of probable PRPP-binding site at the dimer interface suggests that the PRPP bound to one subunit of the dimer would be located close to the ATP bound to the other subunit of the dimer. In this model PRPP would bury the bound ATP that is consistent with the sequential mechanism observed in other PRTases where binding of base precedes binding of PRPP.
Allosteric Inhibition by L-Histidine--
The major
conformational change observed in the histidine-bound form is a large
twist of the domain III relative to the domain I and II (Fig.
2b). When domain I of the apoenzyme and that of the AMP:His
enzyme were superimposed, the r.m.s.d. of the domain I was only 1.46 Å and that of domain II was 2.19 Å. The r.m.s.d. of the domain III,
however, was 12.89 Å, due to a solid body movement of the -sheet of
the domain III induced by residues involved in binding to histidine
(Fig. 2b). Six histidines bind to the domain III clusters at
both ends of three dimers, stabilizing the hexamer (Fig.
3a). These histidines are completely embedded in the domain
III cluster (Fig. 3c). Molecular surface representations of
the hexamers show this conformational change from an "open" cluster
(Fig. 3d) to a "closed" cluster (Fig. 3e).
The residues involved in binding to each histidine are contributed by
the two adjacent domain IIIs suggesting that direct interactions with histidine (Fig. 5a) are
responsible for bringing the three dimers together to form the hexamer.
The interactions include a well ordered hydrogen bonding network with
residues Asp218 and Ala273 from one subunit,
residues Leu234, Gly235, Ser236,
Thr238, and Leu253 of the second subunit, and
an ordered water molecule.
|
Inhibition resulting from hexamer formation is somewhat reminiscent of the allosteric mechanism observed in ribonucleotide reductase (43). Feedback inhibitor-based oligomerization, resulting in either altered topology or reduced access to the active site, is emerging as a way of regulating enzymes. In the case of ATP-PRTase, the allosteric inhibition by histidine can be synergistically favored by the competitive inhibitor AMP, thus adding yet another dimension to the regulation of activity. Maximal inhibition is observed when both inhibitors AMP:His are bound (11). The structures suggest that the reason for the synergistic behavior is that binding of histidine reorients some key active site residues (Tyr116, Arg135, Arg137, Asp154, and Arg160) in the active site, and in return binding of AMP establishes additional inter-subunit interactions that stabilize the histidine-bound hexamer. These interactions are only possible with the global conformational change triggered by histidine.
A Disulfide Bond and Its Potential Role in Regulation--
The
presence of disulfide bonds in prokaryote intracellular enzymes has not
been well documented, although crystallographic studies have shown the
existence of disulfide bonds in a handful of prokaryotic enzymes (44,
45). In the ATP-PRTase structure we not only observe a disulfide bond
between Cys73 and Cys175 but also found that it
was not present in the PRTase with AMP and histidine (Fig.
5b). In this structure the distance between the two Cs of
the cysteines was 8.6 Å, too far for disulfide bond formation. Two
possible scenarios can explain this observation. First, the lack of a
disulfide bond could be from the strain imposed by the conformational
changes observed in the AMP:His structure possibly due to a closure
between domains I and II (see Fig. 5b). It could also be due
to exposure of crystals to synchrotron radiation. Structurally and
functionally significant disulfide bonds have been shown as broken in
crystals exposed to synchrotron radiation (46). DTNB was used to
determine the presence or absence of the disulfide bond between
Cys73 and Cys175. In the absence of any
ligands, the absorbance of TNB2
at 412 nm suggested that
the disulfide was present in only two of the four cysteines. However,
when the enzyme was incubated with 100 µM AMP or
histidine, or 2 mM AMP with 100 µM of
histidine, the molar ratio of cysteines modified per protomer was
reduced to about 0.5 (see "Experimental Procedures"). We believe
the reduction was due to the formation of hexamer that reduces exposure
of all cysteines. When the same experiments were performed in the
presence of 6 M guanidinium hydrochloride, all protein
forms again showed only two free cysteines. These results suggest that
the disulfide is present in both the apoprotein and AMP:His protein and
that the observed broken disulfide was the result of the radiation, although we cannot rule out the possibility that in the inhibitor-bound protein the disulfide rapidly reforms upon denaturation.
The structure described here provides an explanation of the molecular
basis of feedback inhibition of the histidine biosynthetic pathway by
allosteric regulation of ATP-PRTase by histidine. The binding of
histidine seems to influence activity both by stabilizing the inactive
hexameric form and by sterically hindering the binding of substrates to
the catalytic site. Although the presence of an allosteric domain that
binds the end product of the pathway has been observed in several
enzymes, to our knowledge this represents the first example for the
PRTases. ATP-PRTase also appears to be another example of the
convergent evolution of the PRTases.
![]() |
ACKNOWLEDGEMENTS |
---|
We thank the scientists of BioCARS beamlines at Advanced Photon Source, Argonne National Laboratory, for help in data collection. The use of the Advanced Photon Source was supported by the United States Department of Energy, Basic Energy Sciences, Office of Science, under Contract W-31-109-Eng-38. The use of the BioCARS Sector 14 was supported by the National Institutes of Health, National Center for Research Resources, Grant RR07707. We thank Dr. Bernhard Rupp (Lawrence Livermore National Laboratory) for help in performing the Shake&Warp program and for comments on the manuscript.
![]() |
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.
The atomic coordinates and structure factors (codes 1NH7 and 1NH8 ) have been deposited in the Protein Data Bank, Research Collaboratory for Structural Bioinformatics, Rutgers University, New Brunswick, NJ (http://www.rcsb.org/).
To whom correspondence should be addressed. Tel.:
979-862-7636; Fax: 979-862-7638; E-mail: sacchett@tamu.edu.
Published, JBC Papers in Press, January 2, 2003, DOI 10.1074/jbc.M212124200
2 On-line address, www.ncbi.nlm.nih.gov/structure/VAST/vastsearch.html.
![]() |
ABBREVIATIONS |
---|
The abbreviations used are:
ATP-PRTase, N-1-(5'-phosphoribosyl)-ATP transferase;
DTNB, 5,5'-dithiobis(2-nitrobenzoic acid);
PRPP, 5'-phosphoribosyl
1'-pyrophosphate;
MES, 4-morpholineethanesulfonic acid;
MAD, multiple
anomalous dispersion;
TNB2, thiobenzoate anion;
r.m.s.d., root mean square deviation;
mtATP-PRTase, ATP-PRTase from
M. tuberculosis.
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() |
---|
1. |
Martin, R. G.
(1963)
J. Biol. Chem.
238,
257-268 |
2. | Dall-Larsen, T. (1988) Int. J. Biochem. 20, 231-235[CrossRef][Medline] [Order article via Infotrieve] |
3. | Rodriguez, G. M., Gold, B., Gomez, M., Dussurget, O., and Smith, I. (1999) Tubercle Lung Dis. 79, 287-298[CrossRef][Medline] [Order article via Infotrieve] |
4. | Bell, R. M., and Koshland, D. E., Jr. (1970) Biochem. Biophys. Res. Commun. 38, 539-545[Medline] [Order article via Infotrieve] |
5. | Koshland, D. E., Jr. (1953) Biol. Rev. Camb. Philos. Soc. 28, 416-436 |
6. | Brashear, W. T., and Parsons, S. M. (1975) J. Biol. Chem. 250, 6885-6890[Abstract] |
7. | Cleland, W. W. (1970) in The Enzymes (Boyer, P. D., ed) , pp. 1-65, Academic Press, New York |
8. | Musick, W. D. (1981) CRC Crit. Rev. Biochem. 11, 1-34[Medline] [Order article via Infotrieve] |
9. | Ames, B. N., Martin, R. G., and Garry, B. J. (1961) J. Biol. Chem. 236, 2019-2026[Medline] [Order article via Infotrieve] |
10. | Morton, D. P., and Parsons, S. M. (1977) Arch. Biochem. Biophys. 181, 643-648[Medline] [Order article via Infotrieve] |
11. |
Bell, R. M.,
Parsons, S. M.,
Dubravac, S. A.,
Redfield, A. G.,
and Koshland, D. E., Jr.
(1974)
J. Biol. Chem.
249,
4110-4118 |
12. | Sinha, S. C., and Smith, J. L. (2001) Curr. Opin. Struct. Biol. 11, 733-739[CrossRef][Medline] [Order article via Infotrieve] |
13. | Scapin, G., Ozturk, D. H., Grubmeyer, C., and Sacchettini, J. C. (1995) Biochemistry 34, 10744-10754[Medline] [Order article via Infotrieve] |
14. | Eads, J. C., Ozturk, D., Wexler, T. B., Grubmeyer, C., and Sacchettini, J. C. (1997) Structure (Camb.) 5, 47-58 |
15. | Cole, S. T., Brosch, R., Parkhill, J., Garnier, T., Churcher, C., Harris, D., Gordon, S. V., Eiglmeier, K., Gas, S., Barry, C. E., III, Tekaia, F., Badcock, K., Basham, D., Brown, D., Chillingworth, T., Connor, R., Davies, R., Devlin, K., Feltwell, T., Gentles, S., Hamlin, N., Holroyd, S., Hornsby, T., Jagels, K., and Barrell, B. G. (1998) Nature 393, 537-544[CrossRef][Medline] [Order article via Infotrieve] |
16. | Davies, C., Heath, R. J., White, S. W., and Rock, C. O. (2000) Struct. Fold. Des. 8, 185-195[Medline] [Order article via Infotrieve] |
17. | Collaborative Computational Project 4. (1994) Acta Crystallogr. Sect. D Biol. Crystallogr. 50, 760-763[CrossRef][Medline] [Order article via Infotrieve] |
18. | Matthews, B. W. (1968) J. Mol. Biol. 33, 491-497[Medline] [Order article via Infotrieve] |
19. | Terwilliger, T. C., and Berendzen, J. (1999) Acta Crystallogr. Sect. D Biol. Crystallogr. 55, 849-861[CrossRef][Medline] [Order article via Infotrieve] |
20. | Otwinowski, Z. (1991) in Isomorphous Replacement and Anomalous Scattering: Daresbury Study Weekend Proceedings (Wolf, W. , Evans, P. R. , and Leslie, A. G. W., eds) , pp. 80-86, Daresbury Laboratory, Daresbury, UK |
21. | de La Fortelle, E., and Bricogne, G. (1997) Methods Enzymol. 276, 472-494 |
22. | Brünger, A. T., Adams, P. D., Clore, G. M., DeLano, W. L., Gros, P., Grosse-Kunstleve, R. W., Jiang, J.-S., Kuszewski, J., Nilges, M., Pannu, N. S., Read, R. J., Rice, L. M., Simonson, T., and Warren, G. L. (1998) Acta Crystallogr. Sect. D Biol. Crystallogr. 54, 905-921[CrossRef][Medline] [Order article via Infotrieve] |
23. | Jones, T. A., Zhow, J. Y., Cowan, S. W., and Kjeldgaard, M. (1991) Acta Crystallogr. Sect. A 47, 110-119[CrossRef][Medline] [Order article via Infotrieve] |
24. | McRee, D. E. (1999) J. Struct. Biol. 125, 156-165[CrossRef][Medline] [Order article via Infotrieve] |
25. | Winn, M. D., Isupov, M. N., and Murshudov, G. N. (2000) Acta Crystallogr. Sect. D Biol. Crystallogr. 57, 122-133[CrossRef] |
26. | Cowtan, K. D. (1994) Joint CCP4 and ESF-EACBM Newsletter on Protein Crystallography 31, 34-38 |
27. | Kantardjieff, K. A., Höchtl, P., Segelke, B. W., Tao, F.-M., and Rupp, B. (2002) Acta Crystallogr. Sect. D Biol. Crystallogr. 58, 735-743[CrossRef][Medline] [Order article via Infotrieve] |
28. | Riddles, P. W., Blakeley, R. L., and Zerner, B. (1983) Methods Enzymol. 91, 49-60[Medline] [Order article via Infotrieve] |
29. | Sun, Y. J., Rose, J., Wang, B. C., and Hsiao, C. D. (1998) J. Mol. Biol. 278, 219-229[CrossRef][Medline] [Order article via Infotrieve] |
30. | Yao, N., Trakhanov, S., and Quiocho, F. A. (1994) Biochemistry 33, 4769-4779[Medline] [Order article via Infotrieve] |
31. | Mayer, M. L., Olson, R., and Gouaux, E. (2001) J. Mol. Biol. 311, 815-836[CrossRef][Medline] [Order article via Infotrieve] |
32. | Sun, Y., Olson, R., Horning, M., Armstrong, N., Mayer, M., and Gouaux, E. (2002) Nature 417, 245-253[CrossRef][Medline] [Order article via Infotrieve] |
33. | Deleted in proof |
34. | Cheah, E., Carr, P. D., Suffolk, P. M., Vasudevan, S. G., Dixon, N. E., and Ollis, D. L. (1994) Structure 2, 981-990[Medline] [Order article via Infotrieve] |
35. | Xu, Y., Cheah, E., Carr, P. D., van Heeswijk, W. C., Westerhoff, H. V., Vasudevan, S. G., and Ollis, D. L. (1998) J. Mol. Biol. 282, 149-165[CrossRef][Medline] [Order article via Infotrieve] |
36. |
van Heeswijk, W. C.,
Wen, D.,
Clancy, P.,
Jaggi, R.,
Ollis, D. L.,
Westerhoff, H. V.,
and Vasudevan, S. G.
(2000)
Proc. Natl. Acad. Sci. U. S. A.
97,
3942-3947 |
37. | Klungsoyr, L., and Kryvi, H. (1971) Biochim. Biophys. Acta 227, 327-336[Medline] [Order article via Infotrieve] |
38. | Tebar, A. R., Fernandez, V. M., Martin Del Rio, R., and Ballesteros, A. O. (1973) Experientia (Basel) 29, 1477-1479 |
39. | Lohkamp, B., Coggins, J. R., and Lapthorn, A. J. (2000) Acta Crystallogr. Sect. D Biol. Crystallogr. 56, 1488-1491[Medline] [Order article via Infotrieve] |
40. |
Sissler, M.,
Delorme, C.,
Bond, J.,
Ehrlich, S. D.,
Renault, P.,
and Francklyn, C.
(1999)
Proc. Natl. Acad. Sci. U. S. A.
96,
8985-8990 |
41. | Bovee, M. L., Champagne, K. S., Demeler, B., and Francklyn, C. S. (2002) Biochemistry 41, 11838-11846[CrossRef][Medline] [Order article via Infotrieve] |
42. | Appel, R. D., Bairoch, A., and Hochstrasser, D. F. (1994) Trends Biochem. Sci. 19, 258-260[CrossRef][Medline] [Order article via Infotrieve] |
43. | Kashlan, O. B., Scott, C. P., Lear, J. D., and Cooperman, B. S. (2002) Biochemistry 41, 462-474[CrossRef][Medline] [Order article via Infotrieve] |
44. |
Bourne, Y.,
Redford, S. M.,
Steinman, H. M.,
Lepock, J. R.,
Tainer, J. A.,
and Getzoff, E. D.
(1996)
Proc. Natl. Acad. Sci. U. S. A.
93,
12774-12779 |
45. | Wells, W. W., Yang, Y., Deits, T. L., and Gan, Z. R. (1993) Adv. Enzymol. Relat. Areas Mol. Biol. 66, 149-201[Medline] [Order article via Infotrieve] |
46. |
Weik, M.,
Ravelli, R. B.,
Kryger, G.,
McSweeney, S.,
Raves, M. L.,
Harel, M.,
Gros, P.,
Silman, I.,
Kroon, J.,
and Sussman, J. L.
(2000)
Proc. Natl. Acad. Sci. U. S. A.
97,
623-628 |
47. | Kraulis, P. J. (1991) J. Appl. Crystallogr. 24, 946-950[CrossRef] |
48. | Merrit, E., and Murphy, M. (1994) Acta Crystallogr. Sect. D Biol. Crystallogr. 50, 869-873[CrossRef][Medline] [Order article via Infotrieve] |