Crystal Structure of ATP Phosphoribosyltransferase from Mycobacterium tuberculosis*

Yoonsang ChoDagger §, Vivek SharmaDagger , and James C. SacchettiniDagger ||

From the Dagger  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
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ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS AND DISCUSSION
REFERENCES

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.

    INTRODUCTION
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ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS AND DISCUSSION
REFERENCES

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).


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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 beta -sheet surrounded by alpha -helices. Type II PRTases, such as quinolinic acid PRTase, have a modified alpha /beta -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
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ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS AND DISCUSSION
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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-beta -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-beta -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, alpha  = beta  = 90, and gamma  = 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
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS AND DISCUSSION
REFERENCES

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.

                              
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Table I
Data collection and refinement statistics

mtATP-PRTase is an elongated molecule consisting of 10 alpha -helices and 15 beta -strands (Fig. 1a) composed in 3 domains. Domain I (residues 1-90, 175-184, and 194-211) contains a central beta -sheet consisting of four parallel beta -strands (beta 1, beta 3, beta 4, and beta 5) and two anti-parallel strands (beta 2 and beta 11). The beta -sheet is surrounded by 3 alpha -helices, alpha 1 on one side and alpha 2 and alpha 3 on the other side. Domain II (residues 91-174) is also an alpha /beta -structure composed of four (beta 7-10) parallel beta -strands and one (beta 6) anti-parallel beta -strand with two alpha -helices on each side (alpha 4 and alpha 5 on one side and alpha 6 and alpha 7 on the other side). Domain III (residues 212-284) has one beta -sheet consisting of four anti-parallel beta -strands (beta 12-15) with two alpha -helices (alpha 9 and alpha 10) on one side of the beta -sheet.


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Fig. 1.   The overall fold of the mtATP-PRTase. a, stereo view of the ribbon representation of the mtATP-PRTase protomer. Ribbon is colored by secondary structure with yellow for helices, cyan for sheets, and gray for coils. The ligand AMP and His are shown in ball-and-stick representation colored by type of atom. The ribbon diagram was prepared by MOLSCRIPT (47) and Raster3d (48). b, molecular surface of an ATP-PRTase protomer colored by electrostatic potential. AMP was located in the cleft between domains I and II and the histidine on the allosteric regulatory domain. c and d, electron density of bound AMP (c) and histidine (d). Shake&Warp (26) electron density map was averaged from six independent refinements of a composite model. This and all the remaining figures were prepared by SPOCK (quorum.tamu.edu/spock/).

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 Calpha atoms) (29) (Fig. 2a), an E. coli histidine-binding protein (Protein Data Bank code 1HSL; r.m.s.d. 3.2 for 164 Calpha 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).


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Fig. 2.   Structural comparison of mtATP-PRTase. a, superimposition of AMP:His-mtATP-PRTase (yellow) with GlnBP (blue) and a PII homolog, GlnK (red) using the local-global-alignment program (A. Zemla, PredictionCenter.I1nI.gov/local/lga). b, superimposition of apo mtATP-PRTase (green) and AMP:His-form (yellow). Domain I of both apo and AMP:His form of the enzyme structures were superimposed by InsightII (www.accelrys.com). c, molecular surface of the apo dimer by electric potential. The location of the ATP-binding site in the dimer is based on the superimposed competitive inhibitor, AMP (stick model), from the AMP:His structure.

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 Calpha atoms) and the guanine nucleotide exchange factor domain from human elongation factor-1beta (Protein Data Bank code 1B64; r.m.s.d. 2.0 for 57 Calpha atoms). Whereas all the four beta -strands and two alpha -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 beta -sheet for domain III formed by the C-terminal beta -strand (residues 280-284) with the penultimate beta -strand (beta 15, residues 273-276) of the adjacent subunit (Fig. 3b).


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Fig. 3.   Mechanism of allosteric inhibition in mtATP-PRTase hexamer. a, ribbon representation of the hexamer form observed in the crystal, showing the 2- and 3-fold symmetry axes relating the protomers in three dimers (colored red, blue, and yellow). The cross line and the arrow indicate clipping plane and viewing direction for the clipped view. b and c, hexamer interface with three embedded histidines in holoenzyme. The hexamerization interfaces of domain III are marked by arrows (b). d and e, molecular surface of the two ATP-PRTase structures viewed along the 3-fold axis and colored according to electrostatic potential. The domain III cluster is more opened in apo structure (d) and closed in the AMP:His-structure (e). Domain I and II undergo conformational shift (marked by arrows) upon binding of inhibitors that causes steric hindrance in the active site.

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.


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Fig. 4.   AMP in the active site and possible PRPP binding model. a, AMP bound to the ATP binding P-loop in a highly negatively charged cavity. The regions interacting with AMP are represented by yellow or red ribbons. b, proposed binding of ATP and PRPP were modeled based on the location of AMP, sulfates, and several positively charged residues in the active site.

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 beta -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.


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Fig. 5.   Binding of histidine in the allosteric site and its affect on lysis of disulfide. a, ribbon representation of the histidine-binding region. Residues involved in binding histidine in domain III from two adjacent subunits are shown in blue and red ribbons. The residues involved in direct interactions with histidine are shown in a stick representation. b, superimposition of the apo form (cyan ribbon) and AMP:His form (red ribbon) showing the conformational differences observed near the disulfide bond. The bound AMP in the active site is shown as a ball-and-stick representation.

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 Calpha s 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
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS AND DISCUSSION
REFERENCES

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