vivek sharma‡¶ and james c. sacchettini‡¶‡‡ · 02/01/2003 · yoonsang cho‡§, vivek...

Crystal structure of ATP Phosphoribosyltransferase from Mycobacterium tuberculosis

Yoonsang Cho‡§, Vivek Sharma‡¶ and James C. Sacchettini‡¶‡‡

From the ‡Department of Biochemistry & Biophysics, Texas A&M University, College

Station, TX 77843-2128, USA

§ Graduate School of Biomedical Sciences, Texas A&M University System Health Science

Center, College Station, TX 77843-1114, USA

¶ Center for Structural Biology, Institute of Biosciences and Technology, Houston, TX

77030-3303, USA

‡‡Corresponding author: Phone:(979) 862-7636, Fax:(979) 862-7638

E-mail: [email protected]

Data deposition:

The coordinates of apo (PDB ID:1NH7) and AMP:His bound form (1NH8) have been

deposited in the Protein Data Bank.

Running title: Crystal Structure of ATP Phosphoribosyltransferase

Copyright 2003 by The American Society for Biochemistry and Molecular Biology, Inc.

JBC Papers in Press. Published on January 2, 2003 as Manuscript M212124200 by guest on O

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SUMMARY

The N-1-(5'-phosphoribosyl)-ATP transferase (ATP-PRTase) 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 ATP-PRTase 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 while the

histidine-inhibited form is a hexamer. The structure represents a new fold for a

phosphoribosyltransferase, consisting of 3 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. by guest on October 11, 2020

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The N-1-(5'-phosphoribosyl)-ATP transferase (ATP-PRTase) encoded 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 (PR-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).

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. While the transcriptional regulation based on nutrient conditions control 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 four 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 PR-ATP yields ATP and PRPP (9). The synergistic inhibition of the

enzyme was demonstrated to occur allosterically by histidine and competitively by AMP,

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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. While 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�� /�-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.


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EXPERIMENTAL PROCEDURES

Materials-ATP, AMP, L-histidine, 5,5'-Dithio-bis(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 database (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 E. coli over-expression strain, BL21(DE3). Cells were incubated at 37oC

until the O.D. reached 0.6 and induced with 1 mM IPTG, and incubation continued for

another 4 hours. The bacterial cells were harvested by centrifugation and resuspended in 20

mM Sodium Phosphate (pH7.5) 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 Ni-NTA 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 DTT, pH7.5) as a final step. The protein was more than 95% pure as observed on

a 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


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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 �g/ml along

with 35 �g/ml of kanamycin. Cultures were then induced with 1 mM IPTG followed by

incubation for 4 hours at 37°C. The protein was purified using the same methods as for the

apo protein.

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 apo-crystals were obtained in by mixing equal volumes (2-�� 20 mg/ml of

protein with a buffer containing 0.1 M MES (pH6.5) and 1.8 M Magnesium Sulfate as a

precipitant. AMP:His-crystals were obtained in condition #15 of Crystal Scrren 2 from

Hampton Research (0.1 M Sodium Citrate (pH5.6), 0.5 M Ammonium Sulfate and 1.0 M

Lithium Sulfate) in the presence of 5 mM AMP and 100 ��.

Data collection and processing-A complete and redundant high-resolution data set was

collected at BioCARS beam line 14BMC at the Advanced Photon Source, Argonne National

Laboratory. MAD (Multiple Anomalous Dispersion) data sets were collected for both the

apo crystal (MAD1) and AMP:His-crystal (MAD2) (Table 1). All datasets were indexed and

scaled using MOSFLM and SCALA of the CCP4 program suite (17). Unit cell dimensions

for apo crystal 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


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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-Se 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 (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, poly-alanines

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 1b).

Structure determination of AMP:His form-Molecular replacement was tried to AMP:His

bound data 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


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DM (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 Rfee 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 at Table 1b.

Cysteine modification experiments -We followed a previously described (28) experimental

procedure 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 5,5'-

Dithiobis(2-nitrobenzoic acid) (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 AU for the apo and AMP:His form

respectively) and molar absorption coefficient of TNB (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 of TNB).

Gel filtration experiments-Superdex200 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 pH7.5,


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0.1 M NaCl, 1 mM EDTA and DTT. 100 µM of histidine and 1 mM of 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 25 °C, more than 99% of the apo-enzyme eluted as a dimer at

low protein concentration (less than 50ug/ml). We were not able to detect the dimer when

the protein was preincubated at 10uM histidine; only hexamers and higher oligomers were

detected.


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RESULTS AND DISCUSSION

Overall structure of mt ATP-PRTase-The X-ray structure of the recombinant mtATP-

PRTase was solved from electron density maps calculated by MAD (Multiple Anomalous

Dispersion) 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 2.7Å and 1.8 Å, respectively (Tables 1 & 2). In both cases, the refined structure contains

276 out of the 284 residues present in mtATP-PRTase. The residues 186 to 193 were

disordered and omitted from the final model.

mt ATP-PRTase is an elongated molecule consisting of ��-�� -strands

(Fig. 1a) composed in 3 domains. Domain I (residues1-90, 175-184, and 194-211) contains a

central β sheet consisting of four parallel � strands (�1, �3,��4 and �5) and two anti-parallel

strands (�2 and �11). The ��sheet is surrounded by �� 1 on one side and �2 and

�3 on the other side. Domain II (residues 91-174) �� four

(�7-10) parallel ��strands and one (�6) anti-parallel ��strand with two�� helices on each side

(�4 and �5 on one side, �6 and �7 on the other side). Domain III (residues 212-284) has one

� sheet consisting of four anti-parallel � strands ��-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 most of its


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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. 1c&d. The catalytic core of ATP-PRTase (domains I and II) is similar to the

E. coli glutamine binding protein (1WDN; RMSD 3.4 for 17��(29) (Fig. 2a), an E.

coli� �� !"#� $�!%� �&�� '(� �� (30) as well as the

ligand binding core of a glutamate receptor from Synechocystis sp (1IIW) (31) and that of

rat (1LB8) (32).

A VAST (33) structural similarity search using domain III found that the domain

shares a high degree of similarity with the E. coli signal transducing protein PII (2PII;

$�!%� �&(� �� '�� he guanine nucleotide exchange factor domain from

human elongation factor-�� )'(#� $�!%� �&�� *� �� &� +� �� -

�� ,�� -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 PROCEDURE). In general, ATPase hexamers are more abundant at

concentrations of enzyme higher than 1 mg/ml or in the presence of stoichiometric AMP,


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PR-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.

While alternate roles and mechanisms for regulation of HisZ may not be ruled out,


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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 owing to crystallographic 3-fold and 2-fold symmetry axes in

the R32 spacegroup 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 apo-protein (Fig.

3d&e). In the case of the AMP:His hexamer, the subunit accessible surface area buried is

3078 Å2, it is only 2417 Å2 in the apo-form. The dimer interface buries 1203 Å2 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),

while those involved at the hexamer interface are mainly from domain III. The most

prominent structural feature of the AMP:His hexamer is the extended β-sheet for domain III

formed by the C-terminal β strand (residues 280-284) with the penultimate β strand (β15,

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. 1a&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


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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. 4). 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

towards 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


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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 apo- and that of the AMP:His-enzyme were superimposed, the

RMSD of the domain I was only 1.46 Å and that of domain II was 2.19 Å. The RMSD of

the domain III, however, was 12.89 Å, due to a solid body movement of the β 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 III’s suggesting that direct interactions


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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 as well as 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


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(Fig. 5��&�-��,��,��.��/��,��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). 5,5'-Dithio-bis(2-nitrobenzoic acid) (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 412nm suggested that the disulfide was

present in only two of the four cysteines. However, when the enzyme was incubated with

100 �M of AMP or histidine, or 2 mM AMP with 100 �M of histidine, the molar ratio of

cysteines modified per protomer was reduced to about 0.5 (see EXPERIMENTAL

PROCEDURE). 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 apo- 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


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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.


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ACKNOWLEDGEMENTS

We thank scientists of BioCARS beamlines at Advanced Photon Source, Argonne National

Laboratory for their help in data collection. Use of the Advanced Photon Source was

supported by the U.S. Department of Energy, Basic Energy Sciences, Office of Science,

under Contract No. W-31-109-Eng-38. Use of the BioCARS Sector 14 was supported by the

National Institutes of Health, National Center for Research Resources, under grant number

RR07707. We thank Dr. Bernhard Rupp (LLNL) for his help in performing Shake&Warp

program and for his comments on the manuscript.


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FIGURE LEGENDS

Scheme 1. Schematic representation of the reaction catalyzed by ATP-PRTase.

The figure was drawn using ISIS/Draw version 2.3 (MDL Information Systems, Inc.).

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 atom type.

The ribbon diagram was prepared by MOLSCRIPT (45) and Raster3d (46). b) Molecular

surface of an ATP-PRTase protomer colored by electrostatic potential. AMP was located in

the cleft between domain I and II, while the histidine on the allosteric regulatory domain. c)

& d) Electron density of bound AMP (c) and histidine (d). Shake&Warp (26) electron

density map, averaged from 6 independent refinements of a composite model. This and all

the remaining figures were prepared by SPOCK (49).

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 LGA (local-

global-alignment) program (50). (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


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apo dimer by electric potential. The location of the ATP binding site in the dimer based on

the superimposed competitive inhibitor, AMP (stick model) from the AMP:His structure.

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 holo enzyme. The

trimerization interfaces of domain III are marked by arrows (b). e and d) 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.

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.

Fig. 5. Binding of histidine in the allosteric site and its affect on lysis of disulfide.


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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 ribbon. 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-bound-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.


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TABLE 1. Data Collection and refinement statistics Data collectiona Wavelength (Å) Resolution (Å) % Complete Rsym

b MAD1 0.9642 (remote) 3.0 99.9 (99.8) 0.195 (0.569) 0.9795 (peak) 3.0 99.9 (99.8) 0.109 (0.306) 0.9798 (inflection) 3.0 93.7 (99.6) 0.171 (0.701) MAD2 0.9571 (remote) 2.6 99.1 (99.7) 0.088 (0.447) 0.9798 (peak) 2.6 99.1 (99.6) 0.081 (0.185) 0.9800 (inflection) 2.6 99.1 (99.7) 0.156 (0.207) Apo 1.0 2.7 99.1 (99.9) 0.067 (0.150) AMP:His 1.0 1.8 97.5 (99.5) 0.047 (0.134) Refinement statisticsb Apo AMP:His Resolution range(Å) 28-2.7 20-1.8 No. of reflections Working Set 9850 25850 Test set 547 1366 Number of atoms Protein 2073 2097 Solvent 199 251 Rcryst

c 19.2 19.8 Rfree

c 26.1 23.6 Average B (Å2) 18.3 17.9 r.m.s.d. from ideal Bond length 0.048 Å 0.014 Å Bond angle 3.6° 1.5°

a Values in parentheses are for the highest resolution shell: 3.17-3.0 Å for the MAD1 data sets and , 2.6-2.75 Å for the MAD2 data sets, 2.7-2.97 Å for the apo data sets and 1.8-1.93 Å for the AMP:His data sets. b Rsym = 100 × ΣhΣi |I(h)I - <I(h)>|/ΣhΣi <I(h)i>, where I is the observed intensity, and <I> is the average intensity of multiple observations of symmetry-related reflections. c R = Σ||Fo|- |Fc||/Σ|Fo|. Rcryst and Rfree were calculated using the working and the test reflection sets, respectively. 5% of the entire reflection was taken as test set.


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Yoonsang Cho, Vivek Sharma and James C. Sacchettinituberculosis

Crystal structure of ATP Phosphoribosyltransferase from Mycobacterium

published online January 2, 2003J. Biol. Chem.

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vivek sharma‡¶ and james c. sacchettini‡¶‡‡ · 02/01/2003 · yoonsang cho‡§, vivek...

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