Phosphoribosyl Diphosphate (PRPP):Biosynthesis, Enzymology, Utilization,and Metabolic Significance

Bjarne Hove-Jensen,a,b Kasper R. Andersen,b Mogens Kilstrup,a

Jan Martinussen,a Robert L. Switzer,c Martin Willemoësd

DTU Systems Biology, Technical University of Denmark, Kongens Lyngby, Denmarka; Department of MolecularBiology and Genetics, Aarhus University, Aarhus, Denmarkb; Department of Biochemistry, University of Illinoisat Urbana-Champaign, Illinois, USAc; Department of Biology, University of Copenhagen, Copenhagen,Denmarkd

SUMMARY . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2INTRODUCTION . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3CHEMISTRY OF PRPP . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3BIOSYNTHESIS OF PRPP. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4

PRPP Synthase and Phosphoribosyl Bisphosphate Phosphokinase . . . . . . . . . . . . . . . . . . . . . . . . . 4CLASSIFICATION AND PROPERTIES OF PRPP SYNTHASES . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5

Class I PRPP Synthases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5Eubacterial PRPP synthases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5

(i) Three-dimensional structure of B. subtilis PRPP synthase . . . . . . . . . . . . . . . . . . . . . . . . . . . 6(ii) M. tuberculosis PRPP synthase. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 13(iii) Other bacterial PRPP synthases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 13

Yeast PRPP synthases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 14(i) S. cerevisiae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 14(ii) S. pombe . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 18(iii) PRPP synthases of other fungi . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 18

Mammalian PRPP synthases and PRPP synthase-associated proteins . . . . . . . . . . . . . . . . . . 19(i) PRPP synthase. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 20(ii) PAP . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 22

Other class I PRPP synthases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 23Class II PRPP Synthases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 23Archaeal PRPP Synthases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 25

M. jannaschii . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 25T. volcanium. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 26S. solfataricus. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 27Other archaeal PRPP synthases. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 27

MECHANISM OF CATALYSIS. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 28REGULATION OF PRPP SYNTHASE ACTIVITY . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 30PHOSPHORIBOSYL BISPHOSPHATE PHOSPHOKINASE . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 33DIPHOSPHORYL, NUCLEOTIDYL, AND PHOSPHORYL TRANSFER . . . . . . . . . . . . . . . . . . . . . . . . 34

Substitution Reactions at �-, �-, or �-Phosphates . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 34Diphosphoryl transfer . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 34

(i) Hydroxymethyldihydropterin diphosphokinase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 35(ii) Thiamine diphosphokinase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 36(iii) GTP/GDP 3=-diphosphokinase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 36(iv) MutT . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 38

Nucleotidyl transfer . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 38Phosphoryl transfer. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 39Catalytic strategies . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 40

UTILIZATION OF PRPP . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 42Reactions at the Anomeric Carbon of PRPP . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 42

Comparison of phosphoribosyltransferase and PRPP synthase structures . . . . . . . . . . . . . 42N-Glycosidic bond formation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 44

(i) Purine nucleotide biosynthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 45(ii) Pyrimidine nucleotide biosynthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 46(iii) NAD biosynthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 48(iv) Histidine biosynthesis.. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 50

(continued)

Published 28 December 2016

Citation Hove-Jensen B, Andersen KR, KilstrupM, Martinussen J, Switzer RL, Willemoës M.2017. Phosphoribosyl diphosphate (PRPP):biosynthesis, enzymology, utilization, andmetabolic significance. Microbiol Mol Biol Rev81:e00040-16. https://doi.org/10.1128/MMBR.00040-16.

Copyright © 2016 American Society forMicrobiology. All Rights Reserved.

Address correspondence to Bjarne Hove-Jensen,hove@mbg.au.dk.

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(v) Tryptophan biosynthesis. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 50(vi) Phosphoribosyl transfer without participation of PRPP . . . . . . . . . . . . . . . . . . . . . . . . . . 52

C-Glycosidic bond formation: tetrahydromethanopterin biosynthesis . . . . . . . . . . . . . . . . . 52O-Glycosidic bond formation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 54

(i) Tetrahydromethanopterin biosynthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 54(ii) Arabinosyl monophosphodecaprenol biosynthesis. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 55(iii) Aminoglycoside antibiotic biosynthesis. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 55

Reactions at the Diphosphoryl Moiety of PRPP. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 56Cofactor Biosynthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 56

Thiamine diphosphate . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 56Flavins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 57Pterins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 57

Carbon-Phosphorus Lyase Pathway. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 57PRPP AS MEDIATOR OF METABOLIC REGULATION . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 57

The Enigma of PRPP as a Regulator of Metabolic Activity. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 57Regulation of Pyrimidine Metabolism by the RNA-Binding PyrR Protein . . . . . . . . . . . . . . . . . 58

Three-dimensional structure of PyrR . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 58Mechanism of PRPP-mediated regulation by PyrR. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 58

Regulation of Purine Metabolism by PRPP-Responsive, DNA-Binding PurR Proteins . . . . 60Three-dimensional structure of B. subtilis PurR . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 60Mechanism of PRPP-mediated regulation of purine biosynthesis in B. subtilis . . . . . . . . 61Mechanism of PRPP-mediated regulation of purine biosynthesis in L. lactis . . . . . . . . . . 61Evolution of PurR . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 62Search for a common mechanism for PRPP modulation of PurR conformation . . . . . . 63

Allosteric Activation of Carbamoylphosphate Synthetase Activity by PRPP . . . . . . . . . . . . . . 64prs GENES, MUTANTS, AND GENE REGULATION . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 65

E. coli and S. enterica . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 65E. coli prs-1 and prs-2. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 65In vitro-generated knockout prs alleles . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 65dnaR . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 66ssrA . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 66S. enterica prs-100 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 67Thiamine biosynthesis. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 67

prs Gene Organization and Regulation of prs Gene Expression . . . . . . . . . . . . . . . . . . . . . . . . . . . 67E. coli and S. enterica . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 67Gram-positive organisms. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 68

FUTURE PERSPECTIVES . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 69SUPPLEMENTAL MATERIAL. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 70ACKNOWLEDGMENTS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 70REFERENCES . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 70AUTHOR BIOS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 82

SUMMARY Phosphoribosyl diphosphate (PRPP) is an important intermediate incellular metabolism. PRPP is synthesized by PRPP synthase, as follows: ribose5-phosphate � ATP ¡ PRPP � AMP. PRPP is ubiquitously found in living organ-isms and is used in substitution reactions with the formation of glycosidic bonds.PRPP is utilized in the biosynthesis of purine and pyrimidine nucleotides, the aminoacids histidine and tryptophan, the cofactors NAD and tetrahydromethanopterin,arabinosyl monophosphodecaprenol, and certain aminoglycoside antibiotics. Theparticipation of PRPP in each of these metabolic pathways is reviewed. Central tothe metabolism of PRPP is PRPP synthase, which has been studied from all king-doms of life by classical mechanistic procedures. The results of these analyses areunified with recent progress in molecular enzymology and the elucidation of thethree-dimensional structures of PRPP synthases from eubacteria, archaea, and hu-mans. The structures and mechanisms of catalysis of the five diphosphoryltrans-ferases are compared, as are those of selected enzymes of diphosphoryl transfer,phosphoryl transfer, and nucleotidyl transfer reactions. PRPP is used as a substrateby a large number phosphoribosyltransferases. The protein structures and reac-tion mechanisms of these phosphoribosyltransferases vary and demonstrate the ver-satility of PRPP as an intermediate in cellular physiology. PRPP synthases appear tohave originated from a phosphoribosyltransferase during evolution, as demonstratedby phylogenetic analysis. PRPP, furthermore, is an effector molecule of purine andpyrimidine nucleotide biosynthesis, either by binding to PurR or PyrR regulatory pro-teins or as an allosteric activator of carbamoylphosphate synthetase. Genetic analy-

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ses have disclosed a number of mutants altered in the PRPP synthase-specifyinggenes in humans as well as bacterial species.

KEYWORDS amino acid metabolism, diphosphoryl transfer, nucleotide metabolism,phosphoribosyl pyrophosphate, protein structure-function

INTRODUCTION

The compound 5-phospho-D-ribosyl-�-1-diphosphate (PRPP) is an important metab-olite required in the biosynthesis of purine and pyrimidine nucleotides, the amino

acids histidine and tryptophan, and the cofactors NAD and NADP (1–3). Furthermore,PRPP is utilized in the biosynthesis of methanopterin in certain archaeal species (4) andin the biosynthesis of polyprenylphosphate pentoses in Mycobacterium tuberculosis (5).By far the most abundant class of reactions using PRPP as a substrate result in theformation of N-glycosidic bonds, whereas a few reactions result in the formation of O-or C-glycosidic bonds. Kornberg and coworkers discovered PRPP in the mid-1950s whilesearching for a reaction that converted orotate to uridylate as well as the enzymescatalyzing these reactions (6, 7). The enzyme catalyzing the synthesis of PRPP, PRPPsynthase (ATP:D-ribose 5-phosphate diphosphotransferase; EC 2.7.6.1) is ubiquitousamong free-living organisms. Thus, only certain obligate intracellular parasites, such assome Chlamydia and Rickettsia species, lack a gene encoding a PRPP synthase. There-fore, in general, an organism contains at least one gene specifying PRPP synthase. A fewbacterial species contain more than one prs gene. In contrast, many eukaryotic organ-isms contain more than one PRPP synthase-specifying gene. The yeasts Saccharomycescerevisiae and Schizosaccharomyces pombe appear to contain five and three PRPPsynthase orthologs, respectively (8–10), mammals (humans and rats) contain threePRPP synthase-encoding genes (11–13), and the plants Spinacea oleracea and Arabi-dopsis thaliana contain four and five PRPP synthase genes, respectively (14–16).

Although prs is believed to be an essential gene (17), a number of prs mutants havebeen isolated, primarily in Escherichia coli and Salmonella enterica serovar Typhimurium,that are conditional or specify mutant variants of PRPP synthase with altered enzymaticproperties. Additionally, a few knockout mutant (�prs) strains have been constructed invitro. Insertion of these �prs mutant alleles into strains with a specific genetic back-ground revealed that an organism can be viable in the absence of PRPP synthase activity.Also, a number of naturally occurring variant PRPP synthases have been identified amongpatients with gout or uric acid overproduction (18).

The present review deals with all aspects of PRPP metabolism with an emphasis onbiochemical, genetic, and physiological aspects of PRPP synthase and the utilization ofPRPP in biosynthesis, primarily in microorganisms. The elucidation of high-resolutionstructures of PRPP synthase from a number of organisms makes this review timely.Previously, a few reviews have been published on PRPP synthases of microbial (1, 19),mammalian (18, 20), and plant (416) origins. These reviews are complemented bythe present review.

Sequence analysis was performed with BLAST (21), the program packages of theIntegrated Microbial Genomes website (http://img.jgi.doe.gov/) (22), and the NationalCenter for Biotechnology Information data (http://www.ncbi.nlm.nih.gov/) (23), whereasamino acid sequence analysis and alignments were performed with Multalin (http://multalin.toulouse.inra.fr/multalin/) and DNA Strider (24, 25). Figures showing the three-dimensional structures were prepared with the Pymol molecular viewer (http://www.pymol.org) (26).

CHEMISTRY OF PRPP

PRPP is synthesized by transfer, in a single step, of the �,�-diphosphoryl group ofATP to the C-1 hydroxyl of �-D-ribose 5-phosphate, with the simultaneous formation ofAMP by the following deceptively simple reaction: ribose 5-phosphate � ATP ¡ PRPP �

AMP (Fig. 1). The reaction is catalyzed by PRPP synthase (6, 7), which is encoded by aprs, prsA, or PRPS gene (13, 27, 28). PRPP may be regarded as an activated form of ribose5-phosphate. Essentially all of the reactions utilizing PRPP as a substrate are substitu-

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tions, where usually nitrogen-containing aromatic bases replace the diphosphorylgroup with simultaneous inversion of configuration at C-1 of the ribosyl moiety (Fig. 1).In the majority of these reactions, a ribonucleoside 5=-monophosphate is formed. Thus,“all” that happens in reactions such as those shown in Fig. 1 is an attachment of thenitrogenous compound to the ribosyl moiety and the formation of diphosphate (PPi).PPi, i.e., phosphoric acid anhydride, with a high negative free energy of hydrolysis, inturn is hydrolyzed to phosphate (Pi), which makes the phosphoribosyl transfer reactionthermodynamically irreversible.

Immediately after its discovery, it was noted that PRPP was a stable compound aslong as it was stored in the cold at neutral pH, whereas acidic or basic conditions andheating readily decomposed the compound. Divalent ions such as Mg2�, Mn2�, andBa2� encouraged decomposition. The degradation of PRPP, presumably by hydrolysis,appears to follow one of three pathways: (i) PRPP � H2O ¡ ribose 5-phosphate � PPi,(ii) PRPP � H2O ¡ 5-phosphoribosyl 1,2-cyclic phosphate � Pi (6, 7, 29), or (iii) PRPP �

H2O ¡ ribosyl 1,5-cyclic phosphate � PPi, with pathways i and ii representing the majorpathways (30, 31). Degradation through pathway ii may furthermore continue accord-ing to the following scheme: 5-phosphoribosyl 1,2-cyclic phosphate ¡ ribosyl1-phosphate ¡ ribose (32). The binding of divalent ions to PRPP is convenientlymeasured by 31P nuclear magnetic resonance, and this revealed that Mg2� binds toboth the 5-phosphate and the 1-diphosphate moieties of PRPP (33).

The standard free energy (ΔG°=) for the hydrolysis of PRPP (PRPP � H2O ¡ ribose5-phosphate � PPi) has been calculated to be approximately �35 kJ mol�1. This valueshould be compared to �47 kJ mol�1 for the hydrolysis of the �,�-phosphoanhydridebond of ATP (34). Consistently, the equilibrium constant for the diphosphoryl transferreaction of PRPP synthase has been determined to be 29 (35).

Despite the lability of PRPP, procedures have been developed for the quantitativeisolation of the compound. These include extraction of PRPP from cells with cold formicacid (36–38), cold perchloric acid (39), or boiling followed by immediate cooling (40),followed by chromatography or phosphoribosyltransferase-catalyzed conversion ofPRPP to ribonucleoside 5=-monophosphate.

A chemically stable analog of PRPP has been synthesized for use as a ligand forvarious PRPP-binding proteins: 1-�-diphosphoryl-2,3-�-dihydroxy-4-�-cyclopentane-methanol 5-phosphate (cPRPP) (41, 42). Although this analog is an inhibitor of PRPPsynthase activity, the compound has proven valuable as a PRPP analog in X-ray analysesof PRPP synthase.

BIOSYNTHESIS OF PRPPPRPP Synthase and Phosphoribosyl Bisphosphate Phosphokinase

The synthesis of PRPP is catalyzed by two enzymes, PRPP synthase and phosphori-bosyl bisphosphate phosphokinase. On a quantitative basis, PRPP synthase is by farthe most important. Phosphoribosyl bisphosphate phosphokinase is a componentof the phosphonate catabolic pathway and is present only in cells that thrive onphosphonate as a Pi source. Here, we shall deal first with PRPP synthase and then laterwith phosphoribosyl bisphosphate phosphokinase.

FIG 1 Biochemical function of PRPP. The �,�-diphosphoryl group of ATP is transferred to ribose 5-phosphate with the generation of PRPPin a reaction catalyzed by PRPP synthase, which is encoded by a prs gene. PRPP is a substrate in a number of substitution reactions, mostof which involve a nitrogen-containing compound. The reaction occurs at C-1 of the ribosyl moiety and proceeds with inversion of theconfiguration of this carbon and with PPi as a leaving group.

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CLASSIFICATION AND PROPERTIES OF PRPP SYNTHASES

Alignment of the amino acid sequences from a variety of organisms revealed a highsimilarity among PRPP synthases. For example, the overall identity of PRPP synthaseamino acid sequences of E. coli and humans is 47% (20), and that of PRPP synthases ofE. coli and the Gram-positive organism Bacillus subtilis is 51% (43). In addition, a15-amino-acid sequence is highly conserved among PRPP synthase and adenine phos-phoribosyltransferase (44, 45). The latter enzyme catalyzes the following reaction:adenine � PRPP ¡ AMP � PPi. This 15-amino-acid sequence of adenine phosphori-bosyltransferase was designated the PRPP-binding site and later the ribose5-phosphate-binding loop in PRPP synthase. Subsequently, studies of cDNA libraries ofthe plants S. oleracea and A. thaliana revealed the presence of four genes that specifya PRPP synthase. Two of these enzymes (isozymes 1 and 2) had primary structuressimilar to those of E. coli and humans, whereas two (isozymes 3 and 4) had primarystructures quite different from those of E. coli and humans. The amino acid sequenceidentity of S. oleracea isozymes 1 and 2 is 88%, and that of isozymes 3 and 4 is 75%. Incontrast, the amino acid sequence identity of isozymes 1 or 2 to isozymes 3 or 4 ismodest, 22 to 25%. These deviations of amino acid sequence identities of S. oleraceaPRPP synthase isozymes 3 and 4 from isozymes 1 and 2 and other “classical” PRPPsynthases are also reflected in their physico-chemical properties (see below). It wastherefore suggested that two different classes of PRPP synthases exist, classes I and II(46, 47). Additionally, PRPP synthases of archaeal origin show low amino acid sequenceidentity with class I or class II PRPP synthases. Thus, the amino acid sequence identitiesof B. subtilis and the thermophilic, methanogenic archaeon Methanocaldococcus jann-aschii, the thermoacidophilic archaeon Sulfolobus solfataricus, or the thermoacidophilic,facultatively anaerobic, organotrophic archeaon Thermoplasma volcanium PRPP syn-thases are 26 to 28%. Analysis of the PRPP synthase of M. jannaschii revealed biochem-ical properties that appeared to be a mixture of the properties of class I and II PRPPsynthases, and it was suggested to belong to a novel class of PRPP synthases, class III(48). In the present review, we instead designate these class III enzymes “archaeal PRPPsynthases.” An alignment of representatives of PRPP synthases of class I (B. subtilis),class II (S. oleracea isozyme 4), and archaea (M. jannaschii) is shown in Fig. 2. Overall, 36amino acid residues are identical among the three sequences, i.e., the amino acidsequence identity is 13% or less. The inclusion of an additional 26 conserved amino acidresidues results in a similarity of approximately 21%. As indicated below, the conser-vation of amino acid residues identified as responsible for catalysis is very high, whereasamino acid residues involved in allosteric regulation of activity of B. subtilis PRPPsynthase are nonconserved among the three classes, consistent with the fact that onlyclass I PRPP synthases are allosterically regulated.

Class I PRPP Synthases

The classical PRPP synthases, i.e., class I, are by far the most widely phylogeneticallydistributed PRPP synthases. These enzymes contain approximately 315 amino acidresidues. PRPP synthase of the bacterial species S. enterica, E. coli, and B. subtilis, as wellas those of humans and rats, are the best-studied PRPP synthases. Among the class Ienzymes, B. subtilis and E. coli PRPP synthases and human PRPP synthase isozyme 1have been crystallized, and high-resolution structures have been determined (49–52).A three-dimensional structure has been determined also for the PRPP synthase fromthe Gram-negative bacterium Burkholderia (Pseudomonas) pseudomallei strain 1710b(PDB code 3dah) (53). However, there are no biochemical data available for the latterenzyme. The crystal forms of PRPP synthase from the various organisms are summa-rized in Table 1. Some kinetic properties of PRPP synthases of various organisms arelisted in Table 2.

Eubacterial PRPP synthases. In this section, we focus on the properties of PRPPsynthase of the bacterial species B. subtilis, S. enterica, E. coli, and M. tuberculosis. Amongthese, a three-dimensional structure has been determined for B. subtilis and E. coli PRPPsynthases. The biochemical properties of B. subtilis PRPP synthase are much less studied

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than are those of S. enterica and E. coli PRPP synthases. The latter two enzymes areidentical except for two conservative amino acid substitutions (E. coli PRPP synthaseserines 278 and 283 are threonine and alanine, respectively, in the S. enterica enzyme).Furthermore, the S. enterica and E. coli PRPP synthase amino acid sequences are 51%identical to that of the B. subtilis enzyme. It is therefore very likely that these threeenzymes share identical three-dimensional structures, biochemical properties, andcatalytic and regulatory properties. Below, we describe the three-dimensional structureof B. subtilis PRPP synthase and, when applicable, the properties of the enterobacterialPRPP synthases with reference to the structure of the B. subtilis enzyme.

(i) Three-dimensional structure of B. subtilis PRPP synthase. A number of crystalforms, summarized in Table 1, are utilized for the description of the structure andcatalysis of B. subtilis PRPP synthase: (a) SO4

2� PRPP synthase with one SO42� bound

in the active site at the position corresponding to the phosphate moiety of ribose5-phosphate and a second SO4

2� bound at the position corresponding to the �-phosphateof ADP at the regulatory site. No divalent metal ion was present, and the structure wasdetermined to 2.3 Å resolution (PDB code 1dkr). (b) mADP PRPP synthase with meth-ylene ADP bound at the active and regulatory sites. As before, no divalent metal ionwas present, and the structure was determined to 2.2 Å resolution (PDB code 1dku)

FIG 2 Alignment of amino acid sequences of PRPP synthases from B. subtilis (B.s.), S. oleracea isozyme 4 (S.o.), and M. jannaschii (M.j.). TheB. subtilis PRPP synthase amino acid sequence is numbered from the N-terminal serine, as the original N-terminal methionine is removedin the mature protein. Functional elements of the B. subtilis PRPP synthase are indicated by bars above the B. subtilis amino acid sequence.Amino acid residues identical in all three sequences are indicated by asterisks below the M. jannaschii PRPP synthase amino acid sequence,whereas conserved amino acid residues are indicated by dots. Conserved amino acids are valine, leucine, and isoleucine; phenylalanineand tyrosine; aspartate and glutamate; arginine and lysine; serine and threonine; and glycine and alanine. The division of the N- andC-terminal domains (150-Leu-Met-151) of B. subtilis PRPP synthase is indicated by a vertical line. B. subtilis PRPP synthase amino acidresidues, which are located at the active site, are shown in red and those which are located at the allosteric, regulatory site are shownin blue, whereas those involved in subunit-subunit interactions are shown in green. Amino acid residues involved in formation of the bentdimer are shown in bold, whereas those involved in formation of the parallel dimer are shown in italics. Whenever amino acid residuesof the PRPP synthase of S. oleracea or M. jannaschii are identical to those of the B. subtilis enzyme, the color code of the latter enzymeis applied to the S. oleracea and M. jannaschii residues as well. The underlined amino acid residues Val178-Asp196, Arg198, andAsn209-Val211 are involved in the formation of a tightly packed interface necessary for allosteric inhibition. Amino acid residues wereselected on the basis of the three-dimensional structures previously published (49, 50, 54). Vertical arrowheads point to B. subtilis PRPPsynthase amino acids, which are homologous to amino acids altered in the human PRPP synthase isozyme 1 due to point mutations inthe PRSP1 gene. Red arrowheads point to amino acid alterations resulting in increased PRPP synthase activity, whereas blue arrowheadspoint to amino acid alterations resulting in decreased PRPP synthase activity. The amino acid alterations and properties of the human PRPPsynthase variants are described further in the text and are summarized in Table 4.

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(49); (c) Cd2� PRPP synthase with two Cd2� bound in each monomer, presumably atthe two Mg2� sites, AMP bound at the active site, and SO4

2� bound in place of thephosphate moiety at the ribose 5-phosphate site. The structure was resolved to 2.8 Åresolution (PDB code 1ibs) (50). (d) AlF3 PRPP synthase; the crystal form contained in theactive site an analog of ATP pieced together by three molecules: (i) AMP representingthe AMP moiety of ATP, (ii) AlF3 with Al3� representing the �-phosphorus, and thethree F� representing the three oxygens, and (iii) a second AMP molecule whose

TABLE 1 Crystal forms of PRPP synthases and PRPP synthase-associated proteins

Organism Designation Resolution (Å) Ligands PDB code Reference(s)

B. subtilis SO42� 2.3 Two SO4

2� (phosphate of ribose 5-phosphate,�-phosphate of ADP)

1dkr 49

mADP 2.2 AMP (active site), methylene ADP (allosteric site) 1dku 49Cd2� 2.8 2 Cd2� (Mg2� sites), AMP (active site), SO4

2�

(phosphate of ribose 5-phosphate)1ibs 50

AlF3 2.0 2 AMP, AlF3, 2 Mg2�, SO42� (�-phosphate of ADP in

allosteric site); ATP analog at active site is piecedtogether by 3 molecules: AMP (�-phosphate)-AlF3

(�-phosphate)-AMP (�-phosphate), with adenosylmoiety protruding into space between subunits

54

mADP-R5P 2.1 Methylene ADP and ribose 5-phosphate (active site),2 Mg2�, SO4

2� (�-phosphate of ADP in allosteric site)54

GDP 1.8 4 Mg2�, Ca2�, GDP, GTP, 2 �,�-methylene ATP, 2 ribose5-phosphate, glycerol

54

mGDP 1.9 5 Mg2�, methylene GDP, 3 �,�-methylene ATP, 2 ribose5-phosphate, 2 glycerol

54

B. pseudomallei 2.3 None 3dah 53E. coli PRPPS 2.7 Mg2� 4s2u 52, 407

M. jannaschii Apo 2.7 None 1u9y 48Ternary 2.9 AMP, ribose 5-phosphate 1u9z 48

S. solfataricus 2.8 AMP 4twb 139

T. volcanium ADP-SO42� 1.5 ADP, SO4

2� 3lrt 138ADP-SO4

2� 1.8 ADP, SO42� 3nag 138

mATP-SO42� 1.9 mATP, SO4

2� 3lpn 138ADP-Mg2�-R5P 1.9 ADP, Mg2�, Pi, ribose 5-phosphate 3mbi 138

Human hPRS1 wild typea 2.6 SO42� 2h06 51

hPRS1-ATP-SO42�-Cd2�a 2.2 AMP, SO4

2�, Cd2� 2hcr 51hPRS1 S132Ab 2.2 SO4

2� 2h07 51hPRS1 Y146Mc 2.5 SO4

2� 2h08 51—a 2.6 SO4

2� 3efh—a 2.0 SO4

2� 3s5j—d 2.3 Mg2�, SO4

2� 4f8e 124PRS1a 2.0 SO4

2� 3s5j 118E43Te 3.0 SO4

2� 4lyg 118D65Nf 2.1 SO4

2� 4lzn 118A87Tg 3.3 SO4

2� 4LZO 118M115Th 2.1 SO4

2� 4M0P 118Q133Pi 2.7 SO4

2� 4M0U 118PAP39j 2.7 SO4

2� 2c4kPAP41k 2.6 2ji4

aPRPP synthase isozyme 1.bSer132Ala mutant variant of PRPP synthase isozyme 1.cTyr146Met mutant variant of PRPP synthase isozyme 1.dAsp52His mutant variant of PRPP synthase isozyme 1.eGlu43Thr mutant variant of PRPP synthase isozyme 1.fAsp65Asn mutant variant of PRPP synthase isozyme 1.gAla87Thr mutant variant of PRPP synthase isozyme 1.hMet115Thr mutant variant of PRPP synthase isozyme 1.iGln133Pro mutant variant of PRPP synthase isozyme 1.jPRPP synthase-associated protein 39.kPRPP synthase-associated protein 41.

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phosphate represents the �-phosphate of ATP. The remaining adenosyl moiety of thelatter AMP molecule protrudes into an “empty space” between two subunits. Further-more, the crystal form contained two Mg2� (one ligated to the “triphosphate chain”),one ribose 5-phosphate at the active site, and one SO4

2� located at the position of the�-phosphate of ADP in the allosteric site. This structure was resolved to 2.8 Å resolution.Altogether, this complex is thought to resemble the transition state of PRPP synthase.(e) mADP-R5P PRPP synthase with one methylene ADP representing the ADP moiety ofATP as well one ribose 5-phosphate in the active site, and two Mg2� and one SO4

2�

representing the �-phosphate of ADP at the allosteric site in each monomer. Thisstructure was resolved to 2.1 Å (54). The structure of the active site is described in moredetail, including stereoscopic views, in the section “Mechanism of Catalysis,” below.Two additional crystal forms (GDP PRPP synthase and mGDP PRPP synthase) have beendetermined and have been particularly useful in elucidating the mechanism of alloste-ric regulation; these are also further described below.

(a) Tertiary structure. The PRPP synthase monomer is composed of two domains in ahead-to-tail arrangement. The tertiary structures of the two domains are remarkablysimilar, although the amino acid sequence identity of the two domains (amino acidresidues 1 to 150 and 152 to 292) is only 11%, with an additional 10% similar amino acidresidues. Thus, the similarities in tertiary structure of the two domains are not at allpredictable from their primary structures. Each domain possesses an �/� structure witha five-stranded parallel �-sheet at the center surrounded by four �-helices as well asone 310-helix in the N-terminal domain (Fig. 3A). Additionally, short antiparallel�-sheets, designated flag regions, flank both domains. This structure resembles that oftype I phosphoribosyltransferases. It is likely, therefore, that type I phosphoribosyltrans-ferases and PRPP synthase originated from the same ancestral gene and that “modern”PRPP synthases may have evolved from duplication of that ancestral gene that wouldhave been half the size of the contemporary prs gene (54).

A number of regions along the amino acid sequence have been highlighted on thebasis of their functions. These are called the regulatory flexible loop (Tyr97 to Thr113),the diphosphate (PP) loop (Asp174 to Gly177), the catalytic flexible loop (Lys197 toMet208), and the ribose 5-phosphate-binding loop (previously called the PRPP-bindingsite) (Gly216 to Thr231) (49, 50, 54) (Fig. 2).

There are two types of subunit interactions in PRPP synthase. First, interactions ofthe �3N and �4N helices of the N-terminal domain result in the formation of a benthead-to-head arrangement of two subunits, referred to as the bent dimer (Fig. 3B).

TABLE 2 Kinetic parameters of PRPP synthases of various organisms

Organism (andenzyme)

PRPPsynthase class

Km (�M)Vmax or sp act(�mol/min/mg of protein) Reference(s)R5P ATP dATP GTP CTP UTP

B. subtilis I 480 660 190 70B. caldolyticus I 530 310 400 55S. enterica I 160 50 130 56E. coli I 203 113 181 57M. tuberculosis I 8.2–71 1–25 1.4–530 77–79B. amyloliquefaciens I 105 50 37 149Human, isozyme 1 I 52 21 25 73Human, isozyme 2 I 83 70a 36 73Rat, isozyme 1 I 40 44 39 71Rat, isozyme 2 I 73 60 35 71Rat, liver enzyme I 64 49 16 143S. oleracea, isozyme 3 II 110 170 233 650 116 137 13.1b 46S. oleracea, isozyme 4 II 48 7 84 490 680 500 16.2c 47M jannaschii Archaea 2,800 2,600 2,200 48T. kodakarensis Archaea 182 140P. calidifontis Archaea 60 80 480 422aS0.5 value.bValue with ATP as diphosphoryl donor. Values varied between 1.2 and 6.9 with GTP, CTP, or UTP as the diphosphoryl donor.cValue with ATP as diphosphoryl donor. Values varied between 4.2 and 6.6 with GTP, CTP, or UTP as the diphosphoryl donor.

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The relevant amino acid residues involved in this interaction are Asn69, Glu70, Ile72, Met73, Leu76, Ile77 (�3N), and Leu116 and Leu120 (�4N), many of which arehighly conserved (Table 3, left column). Second, a subunit is aligned in a parallelmanner with a neighboring subunit in an arrangement involving residues from boththe N- and the C-terminal domains (Fig. 3B). These interactions include hydrophobicinteractions and salt bridges of the �1C helix and the flag region of the C-terminaldomain as well as interactions of the 310 helices and salt bridges of the N-terminaldomain. Important residues in the formation of this type of dimer, referred to here asthe parallel dimer, are Lys115, Gln138, Ile139, Phe142, Asp144, Val178, Asp186, Ile192,Ala193, Ile194, Arg198, and Val211 (Table 3, left column) (49). The two types of subunitinteractions are shown in Fig. 3B, and it is easy to see how this “trimer” may be formallyassembled into the hexameric, propeller-like structure shown in Fig. 3C. The enzymethus consists of a trimer of dimers with all of the N-terminal domains forming an innercircle and the C-terminal domains forming the propeller blades at the outside, resultingin a 3-fold symmetry axis with perpendicular 2-fold axes (49).

(b) The active site. The active site must accommodate the substrates ribose5-phosphate and MgATP. In addition, an overwhelming volume of research data hasshown that an additional so-called free Mg2� is required for activity, because maximalactivity is only obtained when Mg2� is added to the reaction mixture in excess of theMgATP concentration for both bacterial (35, 45, 55–58) and mammalian (59, 60) PRPPsynthases. Furthermore, most PRPP synthases may accept other divalent metal ions inplace of Mg2�, and thus Mg2� may be regarded as a pseudosubstrate (45, 56, 58, 61).In contrast, Ca2� is an inhibitor of PRPP synthase activity, even in the presence of Mg2�

(45). In the crystal structure one Mg2� (the MG1 site) coordinates to Asp174, a highlyconserved residue of the C-terminal domain, to the oxygens of the hydroxyls of C-1,C-2, and C-3 of ribose 5-phosphate, to an oxygen of the �-phosphate of ATP, and to awater molecule, which forms hydrogen bonds to Asp174 and Asp223. Thus Mg2�

ligates both of the substrates. A second Mg2� (MG2) coordinates to His135 of theN-terminal domain, to oxygens of the �-, �-, and �-phosphates of ATP, and to two watermolecules, which form hydrogen bonds to the side chains of Asp103 and Arg104 andthe carbonyl oxygen of Arg101. The effect of this intricate binding is a perfectalignment of ribose 5-phosphate and ATP for an in-line attack of the hydroxyl of C-1 of

FIG 3 Three-dimensional structure of B. subtilis PRPP synthase. (A) Monomer drawn on the basis of the SO42� PRPP synthase structure (PDB code 1dkr) (49).

The N-terminal domain is at the top. Shown are the five-stranded parallel �-sheets (red), helices (blue), flag region (green), regulatory flexible (RF) loop, theribose 5-phosphate (R5P) loop, and the PP loop (yellow). The unresolved catalytic flexible (CF) loop is shown as a dotted line. (B) Bent and parallel dimers drawnon the basis of the Cd2� PRPP synthase structure (PDB code 1ibs) (50). Subunit A is colored similar to the monomer in panel A. Shown are the Cd2� (black),AMP of the active site (red), and sulfate bound at the position of the phosphate moiety of ribose 5-phosphate and at the position of the �-phosphate of ADPof the allosteric site (red). (C) Hexameric propeller structure drawn on the basis of the mADP PRPP synthase structure (PDB code 1dku) (49). Subunit A (as wellas subunits C and E) are colored as described for the monomer in panel A. Shown are the positions of the methylene ADP moieties (red) and methylene ADPmolecules (green), both modeled to only AMP, of the ATP binding sites and the allosteric sites, respectively.

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ribose 5-phosphate at the �-phosphorus of ATP (54), as described in detail below in“Mechanism of Catalysis.”

As described above, the two active site Mg2� ligate to Asp174 of the C-terminaldomain and His135 of the N-terminal domain. Similarly, the ATP-binding site is locatedat the interface of the N- and C-terminal domains of each subunit, but with contribu-tions of amino acid residues of the N-terminal domain of a neighbor subunit of theparallel dimer, for example, subunit A and subunit D (Fig. 3B). The specific aminoacid residues that are important in the binding of ATP in B. subtilis PRPP synthaseare listed in Table S1, left column, in the supplemental material. In contrast, theribose 5-phosphate-binding site is located within the C-terminal domain and isformed exclusively by amino acid residues of this domain. A subset of amino acidresidues of the ribose 5-phosphate-binding site, Gly216 to Thr231, has been shownto directly interact with hydroxyls 2 and 3 (Asp223 and Asp224) or the phosphatemoiety (Asp227, Thr228, Ala229, Thr231, and Ile232) (50, 54) (see Table S1, leftcolumn).

Chemical modification studies have confirmed the importance of some of the aminoacid residues mentioned above. Thus, chemical modification of S. enterica PRPP syn-thase with 5=-(4-fluorosulfonylbenzoyl)adenosine completely inactivated the enzyme ina 1:1 molar ratio. ATP protected the enzyme against inactivation, and the site ofmodification was His130, which corresponds to His135 in B. subtilis PRPP synthase (62),providing evidence for the importance of this residue. Similarly, affinity labeling of E.coli PRPP synthase has been performed with the ATP analog 2=,3=-dialdehyde ATP.Three lysine residues were labeled, Lys181, Lys193, and Lys230. Only Lys193 is con-served, and it corresponds to Lys197 of B. subtilis PRPP synthase (63). As we discussbelow, this Lys197 residue plays a very important role in the catalysis of PRPP synthase.

TABLE 3 Comparison of amino acid residues involved in dimer association of B. subtilisPRPP synthase with amino acids of PRPP synthases of other organisms

Dimer type

Residue involved in indicated dimer association for speciesa

B. subtilis E. coli M. tuberculosisSpinachisozyme 4 M. jannaschii T. volcanium S. solfataricus

Bent dimerb N69 N64 N72 I73 N60 E61 D61E70 D65 R73 F74 D61 V62 K62I72 L67 L75 Q76 I63 E64 L64M73 M68 M76 L77 V64 M65 I65L76 L70 L79 I80 I67 T68 F68I77 V72 I80 Y81 L68 L69 L69L116 V112 L119 L122 A107 I106 T108L120 F116 L123 V127 I111 I110 I112

Parallel dimerc K115 K111 R118 R120 R106 Q105 K107Q138 Q134 Q141 Q144 H128 T128 E130I139 I135 I142 E145 I129 L129 E131F142 F138 F145 F139 F132 S132 Y134D144 D140 D147 S150 T134 V144 K136V178 V174 V182 G185 V167 L165 L169D186 K182 D190 Q183 K175 A173 E177I192 M189 L197 M199 Y181 H179 Y183A193 A190 A198 V200 D182 F180 S184I194 I192 F199 V201 Y183 F181 Y185R198 R195 R201 V205 T187 K185 E189V211 I208 V218 E207 T200 N198 A202

aThe annotated amino acid residues involved in the formation of the two types of dimer (bent and parallel)(see text for details) of B. subtilis PRPP synthase are listed in the left column according to the previouslypublished three-dimensional structure (49). The B. subtilis PRPP synthase amino acid sequence was thenaligned pairwise with the amino acid sequences of PRPP synthases of E. coli (accession no. U00096) (313);M. tuberculosis (accession no. AL123456) (408); S. oleracea isozyme 4 (15); M. jannaschii (accession no.L77117) (409); T. volcanium GSS1 (accession no. BA000011) (410), and S. solfataricus P2 (accession no.AE006641) (411). Amino acid residues at similar positions of PRPP synthases of the latter six organisms arelisted in the other columns. Amino acid residues that are identical or conserved relative to the B. subtilisenzyme are shown in bold, whereas nonconserved residues are shown in lightface.

bInteractions of the B. subtilis PRPP synthase N-terminal domain.cInteractions of the B. subtilis PRPP synthase N- and C-terminal domains.

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Chemical modification with sulfhydryl reagents has been reported to cause inactivationof PRPP synthase, which is protected by the presence of ATP and Pi (64, 65), butcysteine residues have not been identified from the structural studies to be importantin catalysis, which suggests that these treatments result in nonspecific inactivation ofthe enzyme.

The geometry of the two Mg2�-binding sites confirms a wealth of information onthe binding properties of divalent cations to PRPP synthases of, in particular, S. entericaand E. coli. First, kinetic analysis of the S. enterica enzyme revealed the binding of bothMgATP and free, i.e., enzyme-bound, Mg2� (56). It was proposed that the enzyme-bound Mg2� ligates to the �-phosphate of ATP, which is provided to the enzyme as�,�-MgATP (58, 66). However, according to the crystal structure of B. subtilis PRPPsynthase, the true substrate of the enzyme is the �,�,�-tridentate complex of MgATP,which is consistent with the MG2 site described above. Additionally, analysis of E. coliPRPP synthase with an altered ribose 5-phosphate-binding site (Asp220Glu, Asp220Phe,and Asp221Ala) revealed that the effects on the values for the apparent maximalvelocity (Vapp) and Km for ribose 5-phosphate were dependent on the divalent cationpresent, suggesting that the binding of ribose 5-phosphate also occurs via interactionwith Mg2� (61), which is consistent with the MG1 site described above. The crystalstructure, furthermore, completes nuclear magnetic resonance analyses that attemptedto elucidate the conformation of ATP at the active site of S. enterica PRPP synthase.Thus, paramagnetic line broadening of the C-1 proton or of 31P of ribose 5-phosphateby Cr(III) bound to ATP as the exchange-stable �,�,�-tridentate complex was used toestimate the distances from the Cr atom (and presumably Mg2�) to the two atoms, at6.7 to 8.0 Å. This is consistent with proximity of the ribose 5-phosphate C-1 hydroxyland the �-phosphorus of ATP to the enzyme-bound divalent cation (67).

Steady-state kinetic analysis of the inhibition of S. enterica or E. coli PRPP synthaseby substrate analogs revealed an ordered Bi-Bi mechanism with binding of Mg2� first,followed by MgATP and then ribose 5-phosphate (41, 56, 57). The ordered kineticmechanism was further confirmed by equilibrium dialysis. Thus, radioactive ATP andthe inactive analog �,�-methylene ATP bound well to the free enzyme with dissociationconstants in the micromolar range, whereas ribose 5-phosphate binding could not bedetected unless �,�-methylene ATP was also included (68). Kinetic parameters of PRPPsynthases from various bacilli and enteric organisms are listed in Table 2.

(c) The allosteric site. In addition to the crystal forms SO42� PRPP synthase and mADP-

PRPP synthase described above, two additional crystal forms were found useful inelucidating the structure of the allosteric site. (f) GDP PRPP synthase (with four Mg2�,one Ca2�, one GDP, one GTP, two �,�-methylene ATP, and two ribose 5-phosphatemolecules bound per asymmetric unit) resolved to 1.8 Å resolution; (g) mGDP PRPPsynthase (with five Mg2�, one methylene GDP, three �,�-methylene ATP, and tworibose 5-phosphate molecules bound per asymmetric unit) resolved to 1.9 Å resolution(54). The allosteric site of B. subtilis PRPP synthase is composed of amino acid residuescontributed by three subunits, such as A, B, and D (Fig. 3B) (49). The amino acid residuesinvolved in the binding of the �-phosphate and the adenyl moiety of ADP are providedby subunit D (Ser52, Arg54, Ala85, and Ser86) and subunit B (Ser310, Val311, Ser312,and Phe315). Subunit B also provides amino acids for the so-called hydrophobic pocket(Leu134, Ile139, Gln140, Asp148, and His149). Finally, subunit A contains the regulatoryflexible loop (Lys105, Ala106, Arg107, Ser108, and Arg109) (49, 54). These amino acidresidues are listed in Table S2 in the supplemental material. A stereoscopic view ofthe allosteric site is shown in Fig. 4. We shall return to the mechanism of allostericregulation below in the section “Regulation of PRPP Synthase Activity.”

In general, the activity of class I PRPP synthases is regulated by the presence ofribonucleoside diphosphates, primarily ADP, which inhibits the enzyme in competitionwith ATP as well as by binding to a second, allosteric site. In some cases, GDP is also aninhibitor, although GDP only binds to the allosteric site. Thus, a dual mechanism,involving competitive and allosteric inhibition, appears to control the activity of class IPRPP synthases. The complex inhibition pattern by ADP of the enzyme from S. enterica

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has been studied in particular detail. Kinetic analysis revealed that ADP competes withATP at subsaturating ribose 5-phosphate concentrations and that ADP causes substrateinhibition by ribose 5-phosphate due to the binding at a site different from the activesite (69). Direct binding was demonstrated by equilibrium dialysis. Thus, in the absenceof ribose 5-phosphate, ADP binds to the same site as ATP, whereas in the presence ofribose 5-phosphate ADP binds to two sites per monomer (68). Presumably, these arethe active and allosteric sites. Similarly, PRPP synthases from E. coli (45), B. subtilis, andmammalian sources have been shown to be subject to allosteric regulation by ADP, andthe enzymes from B. subtilis, Bacillus caldolyticus, and mammals are also inhibited byGDP (55, 70 –72).

As mentioned above, the activity of class I PRPP synthases requires the presence ofPi. In some cases, sulfate may replace Pi, although a 10-fold-higher concentration ofsulfate is required (49, 71, 73). Removal of Pi from E. coli and S. enterica PRPP synthasecauses irreversible inactivation or aggregation (35, 45, 74), although the presence ofMgATP or MgPRPP stabilizes the enzyme from E. coli or S. enterica, respectively (35, 75).B. subtilis and mammalian PRPP synthases are stable but inactive upon Pi removal (49,71, 73). Indeed, Pi and ADP have been shown to compete for the same binding site inB. subtilis and human PRPP synthases, with Pi acting as an allosteric activator and ADPas an allosteric inhibitor. Thus, certain mutant variants of PRPP synthase desensitized ininhibition by ribonucleoside diphosphates are reciprocally activated at lower concen-trations of Pi (72), and the antitumor agent aminopyrimidopyrimidine ribonucleoside5=-phosphate binds to the allosteric site of human PRPP synthase isozymes 1 and 2.The concentration needed for half-maximal binding increases with increasing Pi

concentration (76). Finally, a sulfate ion was found in the B. subtilis SO42� PRPP

synthase structure. Superimposition of the SO42� PRPP synthase and mADP PRPP

structures revealed that the sulfate ion was located similarly to the �-phosphate ofthe methylene ADP molecule of the regulatory or allosteric ADP-binding site (49).Altogether, the data demonstrate that Pi and ADP compete for binding to the samesite. Steady-state kinetic analysis revealed that Pi binds randomly to E. coli PRPPsynthase, i.e., either before or after the ordered binding of Mg2�, MgATP, and ribose

FIG 4 Allosteric site of B. subtilis PRPP synthase. Stereo view based on the mADP PRPP synthase structure (PDB code 1dku) (49). Thesite is occupied by methylene ADP. Amino acid residues contributing to methylene ADP binding are provide by three subunits, labeledA, B, and D, as in Fig. 3. Amino acid residues of subunit A are shown in blue, amino acid residues of subunit B are in light gray, andamino acid residues of subunit D are shown in dark gray.

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5-phosphate. Similarly, ADP may bind to PRPP synthase in a random fashion, butwith significantly different rate constants constituting the equilibria, so that thepathway where Mg2�, MgATP, and ribose 5-phosphate bind in the absence of Pi ismuch slower than when Pi is bound first. This phenomenon provokes what istermed kinetic cooperativity and thus the sigmoid saturation curves frequentlyobtained for Pi activation with class I PRPP synthases (75).

As expected from the 51% identity of the amino acid sequences of B. subtilis and E.coli PRPP synthases, the amino acid residues involved in the formation of dimers as wellas the active site and the allosteric site (Table 3; see also Tables S1 and S2 in thesupplemental material) are highly conserved. The identity of this subset of amino acidsis 61%. These data were furthermore confirmed by analysis of the three-dimensionalstructure of E. coli PRPP synthase (52).

(ii) M. tuberculosis PRPP synthase. Some attention has been devoted to studies ofPRPP synthase of M. tuberculosis in searching for possible targets for drug treatment oftuberculosis. Three research groups have characterized this enzyme (77–79). All threegroups agree that the quaternary structure of the active enzyme is a hexamer and thatallosteric inhibition is exhibited by the ribonucleoside diphosphates ADP and GDP,whereas there is some disagreement about the kinetic properties of the enzyme (Table2). In addition, one research group established GTP, UTP, and CTP as diphosphoryldonors, with kcat/Km ratios of 3.8, 3.0, and 2.5 M�1 s�1, respectively, compared to 26M�1 s�1 for ATP (78). Interestingly, analytical ultracentrifugation revealed that theapo-enzyme exists as a trimer as well as a hexamer. The presence of ADP greatly shiftedthe oligomeric state toward the hexamer, whereas the presence of ATP had no effecton the oligomerization state (79). A comparison of the amino acid residues involved indimer formation, bent as well as parallel dimers, of M. tuberculosis and B. subtilis PRPPsynthases is shown in Table 3. It is evident that essentially all of the amino acidsinvolved in dimer formation of B. subtilis PRPP synthase are retained in the M. tuber-culosis enzyme. Additionally, a comparison of the amino acid residues involved in thebinding of the substrates MgATP, ribose 5-phosphate, and Mg2� for the two enzymesreveals a high degree of conservation, as does a comparison of the amino acid residuesinvolved in the formation of the allosteric site between the two enzymes (see Tables S1and S2 in the supplemental material). All of these data are consistent with theidentification of the M. tuberculosis enzyme as a class I PRPP synthase. The amino acidsinvolved in the binding of GTP and the pyrimidine ribonucleoside 5=-triphosphates CTPand UTP as substrates for PRPP synthase have not been mapped, so the propensity ofM. tuberculosis PRPP synthase to utilize these latter ribonucleoside 5=-triphosphatesremains unexplained. Other class I PRPP synthases do not utilize GTP, CTP, or UTP.

(iii) Other bacterial PRPP synthases. Although most prokaryotic species contain asingle gene encoding PRPP synthase, the genomes of almost 100 species have beenannotated as containing two PRPP synthase orthologs. Biochemical data for PRPP synthaseare available for only a few of these organisms. One of these is Lactococcus lactis, whichcontains two PRPP synthase-encoding orthologs, prsA and prsB, and for which somephysiologic data exist. The two genes are separated by approximately 300 kbp. Thesequence identities of the prsA- and prsB-specified amino acid sequences with that of B.subtilis PRPP synthase are 65% (L. lactis PrsA), and 52% (L. lactis PrsB), whereas the aminoacid sequence identity between the two L. lactis Prs sequences is 49%. Comparison ofamino acid sequence of L. lactis prsA- and prsB-specified gene products and the establishedactive site amino acid residues of B. subtilis PRPP synthase reveals that PrsA and B. subtilisPRPP synthase are nearly identical, whereas in PrsB the amino acid residues of the catalyticflexible loop vary considerably from those of B. subtilis PRPP synthase. Specifically, thecatalytically important Lys197 and Arg199 residues of B. subtilis PRPP synthase are replacedby Tyr198 and Asp200 in L. lactis PrsB (see Table S3 in the supplemental material).Complementation analysis revealed that L. lactis prsA complements an E. coli prs allele,whereas prsB does not, which demonstrated that prsA encodes a functional PRPP synthase.The function of the prsB gene product remains to be established (P. Bennedsen and J.Martinussen, The Technical University of Denmark, unpublished results).

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The Gram-negative bacterium Thermus thermophilus also contains two PRPPsynthase-specifying orthologs. However, both of these genes specify active PRPPsynthases (417).

A few bacterial species have three annotated PRPP synthase orthologs. In Roseiflexuscastenholzii (DSM 13941), one amino acid sequence has a histidine residue correspond-ing to B. subtilis Arg199 as well as an additional six amino acid residues in the catalyticflexible loop, and there is 43% amino acid sequence identity with B. subtilis PRPPsynthase. The other two amino acid sequences have 42% and 51% amino acid se-quence identity with B. subtilis PRPP synthase. Similarly, in Sphingobium japonicumstrain UT26S, one sequence has an aspartate at the position of the B. subtilis Lys197 andthere is a 9-amino-acid insertion in the catalytic flexible loop. In Pseudomonas stutzeristrain A 1501, the amino acid sequence identity with B. subtilis PRPP synthase variesbetween 23% and 49% and in Clostridium beijerinckii strain NCIMB 8052 these values are27 and 62%, whereas in Rhodoferax ferrireducens strain T118 these values are 24 and54%. No biochemical studies of any of these putative PRPP synthases are currentlyavailable.

Yeast PRPP synthases. Among yeast species, PRPP synthases have been analyzedonly in S. cerevisiae and S. pombe, but presumably many of the physiologic propertiesof PRPP metabolism for these two species are similar to those of other yeast species.Although yeast PRPP synthases share typical class I properties, they also contain propertiesdifferent from the PRPP synthases described so far.

(i) S. cerevisiae. S. cerevisiae contains five PRPP synthase-specifying orthologs, PRS1to PRS5 (8, 9). Inspection of the amino acid sequences reveals that PRS2, PRS3, and PRS4appear to encode typical class I enzymes. Also, PRS1 and PRS5 may encode class Ienzymes, but PRS1- and PRS5-specified amino acid sequences contain so-called non-homologous regions (NHRs), which are insertions relative to the sequences specified byPRS2, PRS3, or PRS4 (Fig. 5). Prs1 contains a single NHR (NHR1) of 105 amino acids inlength, whereas Prs5 contains two NHRs of 110 amino acids (NHR5-1) and 63 amino

FIG 5 Positions of nonhomologous regions. Each line of bars represents polypeptides of a PRPP synthase,with the amino terminus at the left end. S.c.1, S. cerevisiae Prs1; S.c.3, S. cerevisiae Prs3; S.c.5, S. cerevisiaePrs5; S.p.1, S. pombe Prs1; S.p.2, S. pombe Prs2; S.p.3, S. pombe Prs3; PAP39, human PRPP synthase-associated protein 39; PAP41, human PRPP synthase-associated protein 41. S. cerevisiae and S. pombePrs3 are shown at the top as a bar consisting of three segments (I, II, and III) represented in differentshades of blue. This bar could represent most class I PRPP synthases. The left and right ends of segmentII are located within the regulatory and catalytic flexible loops, respectively. An NHR is shown in red. Thenumber of amino acid residues of each NHR is shown below the bars. The three NHRs of S. cerevisiae Prs1and -5 are designated NHR1, NHR5-1, and NHR5-2. According to this nomenclature, S. cerevisiae Prs1 hasthe structure segment 1–segment 2–NHR1–segment 3; S. cerevisiae Prs5 has the structure segment1–NHR5-1–segment 2–NHR1–segment 3; S. pombe PRPP synthase 1 has the structure segment I–segmentII–NHR–segment III; S. pombe PRPP synthase 2 has the structure segment I–NHR–segment II–segment III;human PAP39 and -41 have the structure segment I–segment II–NHR–segment III.

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acids (NHR5-2). For Prs1, immunochemical analysis confirmed that the NHR1 is actuallypresent in the polypeptide found in yeast cells (8, 80, 81). The Prs1 NHR1 is insertedafter Ser199, whereas the Prs5 NHR5-2 is inserted after Ser306. These two positions,Ser199 and Ser306, correspond to Arg202 of the B. subtilis PRPP synthase, and thus theNHRs are inserted in the catalytic flexible loop. In contrast, the Prs5 NHR5-1 sequenceis inserted after Pro100, which corresponds to Asp103 of the B. subtilis PRPP synthase,and thus the NHR5-2 is inserted in the regulatory flexible loop. The identities of theamino acid sequences specified by PRS2, PRS3, and PRS4 with that of B. subtilis PRPPsynthase range between 42 and 46%, whereas for the PRS1- and PRS5-specifiedsequences the identities are 29 and 24%, respectively. After removal of the NHR aminoacid sequences in silico, the latter values increase to 41% (Prs1 ΔNHR1) and 39% (Prs5ΔNHR5-1 ΔNHR5-2).

(a) Interaction of PRS gene products. Unlike most other cloned prs genes, the geneproducts of each of the five S. cerevisiae PRS genes have no PRPP synthase activity invivo when their genes are harbored and expressed individually in an E. coli host strain(82). In contrast, the formation of active PRPP synthase requires the expression of twoor more PRS genes, as established by deletion analysis of PRS genes in S. cerevisiae orby expression of PRS genes in E. coli. Single-knockout mutations in PRS1, PRS2, PRS3,PRS4, or PRS5 result in viable phenotypes (80, 83). Additionally, all 10 combinations oftwo PRS gene knockouts result in viable phenotypes except the Δprs1 Δprs5 and Δprs3Δprs5 double knockout mutants, both of which are lethal (83). The latter doublemutant, Δprs3 Δprs5 strain, is lethal because the Prs1 polypeptide is unstable in a Δprs3genetic background. Thus, strains with the genotype Δprs3 Δprs5 are phenotypicallyprs1 prs3 prs5 (84). Among the five possible combinations of deletion of three PRSgenes, only Δprs1 Δprs2 Δprs3, Δprs1 Δprs3 Δprs4, and Δprs2 Δprs4 Δprs5 deletants areviable, whereas Δprs1 Δprs2 Δprs4 and Δprs2 Δprs3 Δprs4 mutants are lethal; thus, S.cerevisiae cannot exist with only a single PRS gene. These results were confirmed byanalysis of complementation by relevant combinations of PRS genes of an E. coli Δprsallele. This analysis furthermore showed that the simultaneous expression of PRS1 andPRS2 or the simultaneous expression of PRS1 and PRS4 resulted in PRPP synthase withvery low activity in vivo, presumably too low to promote growth of S. cerevisiae Δprs3Δprs4 Δprs5 or Δprs2 Δprs3 Δprs5 deletants. Although complementation of the E. coliΔprs allele by combinations of two PRS genes revealed the production of active PRPPsynthase in vivo, in vitro PRPP synthase activity could be detected only in cell extract ofthe strain that expressed both PRS1 and PRS3 (82). The conclusions from this analysisare that active PRPP synthase requires either PRS1 or PRS5 in addition to at least onemore PRS gene product, but not just any PRS gene product. Thus, an active enzymeconsists of the PRS1 and PRS3 gene products, or an active enzyme consists of the PRS5PRS2, the PRS5 PRS4, or the PRS5 PRS2 PRS4 gene products (83).

Physical interactions of Prs polypeptides was furthermore demonstrated by yeasttwo-hybrid analysis. The results of this analysis showed interactions between Prs1 andPrs3, between Prs1 and Prs2, and between Prs1 and Prs4. These interactions weredemonstrated in a reciprocal manner, i.e., with each PRS gene fused to the bindingdomain as well as to the activating domain. Additionally, some nonreciprocal interac-tions were also observed between Prs5 and Prs2 and between Prs5 and Prs4 (83).Studies of synthetic lethality of combinations of prs deletions also demonstratedinteractions between Prs1 and Prs3 and between Prs1 and Prs5 (84, 85). These Prspolypeptide interactions, demonstrated by genetic and physical analyses after manu-ally manipulating the relevant DNA molecules, have been by and large confirmed byhigh-throughput analyses with two-hybrid analysis (86–88), protein fragment comple-mentation (89, 90), or affinity capture mass spectrometry (91, 92).

The protein-protein interaction analyses revealed an extensive interaction betweenthe various PRS gene products. Evidently, all of the Prs polypeptides interact with oneanother. Presumably, each Prs polypeptide may be able to form a hexameric structurelike that of B. subtilis PRPP synthase, as well as mixed hetero-hexamers and eventually

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larger molecular structures similar to those described for mammalian PRPP synthases(see below).

(b) Enzyme structure. S. cerevisiae PRPP synthase appears to be a large enzyme. PRPPsynthase activity of wild-type S. cerevisiae strain YN94-2 eluted as a single symmetricalpeak corresponding to a molecular mass of at least 900 kDa based on exclusionchromatography (see Fig. S1A in the supplemental material). Also, the enzyme elutedas a single peak during ion exchange chromatography (Fig. S1B). These results mayindicate the presence of a single enzyme form containing all five PRS gene products,although the existence of multiple forms with nearly identical size and charge prop-erties cannot be excluded. Additionally, PRPP synthase activities of extracts of theΔprs2, Δprs3, or Δprs4 strains eluted with molecular masses identical to that of the wildtype, whereas no PRPP synthase activity could be determined in the exclusion chro-matography eluates of either a Δprs1 or a Δprs5 strain, even though activity could beeasily determined in the cell extracts before loading them to the column (Fig. S1A).Thus, PRPP synthase lacking either of the Prs1 and Prs5 polypeptides may be unstableunder the conditions of exclusion chromatography, and the presence of an additionalPRS gene product(s) apparently stabilizes the enzyme complex (93).

Amino acid residues important in active site formation, i.e., residues involved in thebinding of the adenyl moiety and the triphosphate chain of ATP as well as binding ofribose 5-phosphate, and the catalytic flexible loop of B. subtilis PRPP synthase werecompared to those of each of the five S. cerevisiae PRS-specified amino acid sequences(Table S3). The similarity of these selected B. subtilis PRPP synthase amino acid residueswith each of those of S. cerevisiae was 71% (PRS1-specified amino acid residues), 88%(PRS2-, PRS3-, and PRS4-specified amino acid residues), or 50% (PRS5-specified aminoacid residues). Little is known of the enzymatic properties of S. cerevisiae PRPP synthaseactivity. Some activity remains following dialysis of an S. cerevisiae crude extract againstPi-free buffer and in assays under Pi-free conditions, and only 18% of the activityremained when the enzyme was assayed in the presence of 2 mM ADP, relative to100% in the absence of ADP (82). These properties differ from those of bacterialPRPP synthases.

(c) Physiological function. The complexity of S. cerevisiae PRPP synthase polypeptidesmay reflect their physiological function. Evidence for involvement of PRS-specifiedpolypeptides in processes other than catalysis of PRPP synthesis originates fromcharacterization of (i) prs mutants and (ii) mutants defective in other processes andfrom analyses of (iii) protein-protein interactions with PRS-specified polypeptides. First,Δprs3 mutant strains are highly pleiotropic and it was concluded that PRS3 is requiredfor cell wall integrity, cell cycle arrest following deprivation of nutrients, proper ionhomeostasis, and proper actin cytoskeleton organization and cell size homeostasis(94–96). The cell wall integrity signaling pathway includes the protein Slt2 and thetranscription factor Rlm1 of the protein kinase C (Pkc1) mitogen-activated proteinkinase cascade (97). Strains with prs lesions are sensitive to the purine analog 1,3,7-trimethylxanthine (caffeine), which interferes with Pkc1 activated cell wall integritysignaling and the mitogen-activated protein kinase cascade (98), providing evidence forPrs polypeptides being involved in cell wall integrity signaling (99). Second, the pps1-1lesion of S. cerevisiae, isolated after screening for an elongated cell morphology, is anallele of PRS1 (100). Third, evidence of Prs polypeptides being involved in the stressand cell wall integrity signaling pathway comes from yeast two-hybrid analysis ofPrs polypeptides with Slt2 (85, 101, 102) and by coimmunoprecipitation analysis(418).

A number of other protein-protein interactions with Prs polypeptides as partnershave been identified. Rim11, a homolog of mammalian glycogen synthase kinase-3�,interacts with Prs2, Prs3, and Prs5, as shown by affinity-capture mass spectrometry (103)and confirmed by manual yeast two-hybrid analysis with the RIM11 gene and each ofthe PRS genes (104). High throughput analyses revealed a number of protein-proteininteractions of Prs polypeptides with other polypeptides (86, 88), although theirphysiological functions have not been established (102). Interestingly, the interaction of

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the protein kinase Rim11 with Prs5 may be responsible for the phosphorylation of thelatter protein (104). Phosphoproteome analysis by mass spectrometry revealed threephosphorylation sites in Prs5, 361-KTTpSTSpSTpSS-370 (where p indicates phosphory-lation of the following serine residues) (105). This sequence is located within NHR5-2.

From the data provided above, it appears very likely that yeast PRS-specifiedpolypeptides share functions in addition to the catalysis of PRPP formation. Furtherinvestigations are necessary to establish the detailed mechanisms of this behavior.

(d) Function of NHRs. The function of the NHRs has been studied to some extent.Deletion of NHR1 prevented the interaction with Slt2 (84). It is therefore possible thatyeast PRPP synthase may be a bifunctional enzyme involved in nucleotide and amino acidbiosynthesis as well as in maintaining cell wall integrity, the latter function involving NHR1(84, 85, 101, 102).

Finally, the deletion of NHR5-1 resulted in a polypeptide with altered interactionwith other Prs polypeptides. The simultaneous expression in an E. coli Δprs host strainof PRS3 and the prs5 ΔNHR5-1 mutant or of PRS3 and the prs5 ΔNHR5-1 ΔNHR5-2 mutantresulted in complementation of the E. coli Δprs allele, and thus production of activePRPP synthase, whereas no complementation was observed with the simultaneousexpression of the wild-type alleles PRS3 and PRS5. This result may indicate that theNHR5-1 prevents the formation of an active enzyme rather than preventing interactionof Prs3 and Prs5, as interaction of these polypeptides has been previously established(89, 106) (B. Hove-Jensen, unpublished result).

(e) Ribonucleotide pool sizes and PRPP synthase activity. The apparent interaction of Prspolypeptides with proteins of other cellular processes suggests an involvement of Prspolypeptides in linking metabolism to various stress situations. It remains to beestablished, however, if other physiological phenomena, such as perturbation of thepool sizes of ribonucleotides or PRPP, are also involved in the transmission of thesemetabolic effects. Single PRS knockouts are either unaffected in growth rate, asobserved for the mutant strains harboring �prs2, �prs4, and �prs5 or reduced ingrowth rate, as observed for mutant strains with �prs1 and �prs3 (8, 81, 106). Alongwith reduced growth rates, the pool sizes of adenylate nucleotides (AMP, ADP, and ATP)were reduced to approximately 50% of those of wild-type strains in �prs1 and �prs3strains, whereas less-reduced adenylate nucleotide pool sizes were observed in �prs2,�prs4, and �prs5 strains (106). Correspondingly, PRPP synthase activity was lower in�prs1 or �prs3 strains than in �prs2 or �prs4 strains (81). A �prs1 �prs3 strain (i.e.,synthesizing Prs2, Prs4, and Prs5) contained approximately 40% of the adenylatenucleotides and 2% to 3% of the PRPP synthase activity found with the wild-typestrain. A �prs2 �prs4 �prs5 strain (i.e., synthesizing Prs1 and Prs3) had an almost-normal growth rate, 90% adenylate nucleotide pools, and 16% PRPP synthaseactivity compared to the values for the wild-type strain (83). The relatively low PRPPsynthase activities determined in the various mutant strains may indicate that nonop-timal assay conditions were employed or that PRPP synthase may be unstable in vitrowhen one or more PRS gene products are absent, and thus these results may notaccurately reflect PRPP synthase activity in vivo. Altogether, a hierarchy exists amongprs lesions. Thus, �prs3 or �prs1 deletants appear to be more severely affected than�prs3, �prs4, or �prs5 deletants.

In summary, much focus has been devoted to the analysis of Prs polypeptideinteractions with one another as well as with other polypeptides. However, the enzy-mology of yeast PRPP synthase is far from solved. The characterization in the future ofa few mutants suggested from studies of the well-characterized bacterial PRPP syn-thases might contribute significantly to our understanding of the oligomerization ofthis enzyme. The B. subtilis PRPP synthase amino acid residues His135, Asp174, andLys197 are all very important for catalysis. These amino acid residues are identical in thefive yeast Prs polypeptides except for Prs1, which has Asn171 and Arg196 rather thanaspartate and lysine residues, respectively, that are found in B. subtilis PRPP synthase(Table S3). Thus, a replacement of the Lys197 analogs with alanine followed bycomplementation and enzyme activity analysis might prove valuable in establishing

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which Prs polypeptide, i.e., PRPP synthase subunit, is important for enzyme activity. Itis generally believed that an active yeast PRPP synthase is composed of Prs1 and Prs3or of Prs2, Prs4, and Prs5 (83). It is furthermore very likely that these polypeptides forma propeller-like hexameric structure similar to that of B. subtilis PRPP synthase. It wouldtherefore be of interest to analyze variants, such as Prs2Lys197Ala Prs4 Prs5, Prs2Prs4Lys197Ala Prs5, Prs2 Prs4 Prs5Lys308Ala, Prs1Arg196Ala Prs3, or Prs1 Prs3Lys196Ala, as wellas some double variants for complementation and enzymatic properties. Indeed, themultiple, heterologous subunits composition of yeast PRPP synthase provide a uniquesystem to study alterations in a single subunit of the hexameric enzyme, which is notpossible with other PRPP synthases, which are also hexamers but are composed of asingle type of subunit.

S. cerevisiae PRPP synthase is an intriguing enzyme, with its expression of more thanone ortholog necessary for PRPP synthase activity. Also, the presence of NHRs imposesinteresting questions about the structure and function of PRPP synthase. Do NHRs haveordered structures or are they intrinsically disordered regions? Could they confer a Hubprotein character to PRPP synthase, thus facilitating heterologous protein interactionssuch as those described above?

(ii) S. pombe. S. pombe contains three PRPP synthase orthologs, PRS1, PRS2, andPRS3, which presumably specify class I PRPP synthases, as judged from their deducedamino acid sequences. The three genes are unlinked with PRS1 located in chromosomeI, PRS2 in chromosome II, and PRS3 in chromosome III. As PRS-specified polypeptides ofS. cerevisiae, two of the PRS-specified polypeptides of S. pombe contain NHRs (Fig. 5).Similar to the NHRs of S. cerevisiae Prs1 and Prs5, the NHRs of S. pombe are locatedwithin the catalytic flexible loop (Prs1 NHR1) or within the regulatory flexible loop (Prs2NHR2). Amino acid sequence alignments with B. subtilis PRPP synthase and S. pombePRS-specified amino acid sequences revealed an identity for Prs1 of 29% (37% when theNHR1 is removed), for Prs2 of 39% (42% when the NHR2 is removed), and for Prs3, 45%.A comparison of amino acid residues presumed to be important for active site forma-tion and catalysis is included in Table S3. Specifically, these amino acid residues of Prs3are highly conserved, with 88% identity compared to 63% and 75% of those of Prs1 andPrs2, respectively. In addition, only Prs2 and Prs3 contain both of the important lysineand arginine residues of the catalytic flexible loop.

The three S. pombe PRS genes have been cloned in a plasmid vector and expressedin an E. coli Δprs strain for complementation analysis (Table S4). The results showedthat neither PRS1, PRS2, nor PRS3 alone complemented the prs null allele. In contrast,simultaneous expression of PRS3 and PRS1 or PRS2 resulted in complementation,whereas simultaneous expression of PRS1 with PRS2 did not result in complementation.Furthermore, very low PRPP synthase activity could be measured in extracts of cellsharboring PRS1 PRS3 or PRS2 PRS3. The activity, however, increased more than 10-foldin extracts of cells harboring PRS1 PRS3 in one plasmid and PRS2 in a second compatibleplasmid or, vice versa, PRS2 PRS3 in one plasmid and PRS1 in a second compatibleplasmid. These data suggest that the presence of all three PRS gene products may havea stabilizing effect on the enzyme activity (Hove-Jensen, unpublished data).

Heterozygous diploid deletion mutations have been constructed for essentially all ofthe genes of S. pombe. Strains containing deletion mutations in PRS1 (also designatedSPAC4A8.14) or PRS2 (also designated SPBC3D6.06c) are viable. In contrast, PRS3 (alsodesignated SPCC1620.06c) is essential (107). Additionally, two amino acids, Tyr168 andSer172, of the PRS3-specified amino acid sequence were shown to be phosphorylated(108). Ser172 corresponds to Ser172 of B. subtilis PRPP synthase, a fairly conservedamino acid residue. It remains to be established if phosphorylation affects the activityof PRPP synthase.

The combined conclusion of the complementation and deletion analyses with S.pombe PRS genes is that PRPP synthase activity requires the PRS3 gene product as wellas the gene product of either PRS1 or PRS2.

(iii) PRPP synthases of other fungi. The association of multiple PRPP synthasesubunits occurs in other species of fungi as well. The filamentous fungus Aspergillus

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nidulans contains a mycelium of multinucleate cells that are partitioned by septa(reviewed by Harris [109]). Briefly, the formation of septa is regulated by a proteinkinase cascade that involves a sepH-specified protein kinase, which is a positiveregulator of the septation initiation network. Thus, a phenotype of sepH strain muta-tions is a reduction or lack of septation. A number of suppressors of sepH mutation hasbeen isolated and characterized, among which are altered levels of PRPP synthaseactivity. Those authors offered a somewhat speculative model, in which competition ofPRPP synthase and SepH for ATP might be the trigger of septation (110).

A. nidulans contains three PRPP synthase orthologs: PRS1 (located on chromosomeI), PRS2 (on chromosome VI), and PRS3 (on chromosome VII). Prs1 (GenBank accessionno. CBF69369.1; 489 amino acid residues) resembles S. cerevisiae Prs1, as there is anaddition of 169 amino acid residues inserted within the catalytic flexible loop; Prs2 (435amino acid residues; GenBank no. CBF83270.1) has an insertion of 116 amino acidresidues at position 115, i.e., far from the catalytic flexible loop, and Prs2 may representa new variant of PRPP synthases; and Prs3 (320 amino acid residues; GenBank no.CBF85908.1) appears to be a “normal” PRPP synthase polypeptide. Each of the threeamino acid sequences reveals all of the characteristic features for class I PRPP synthases,i.e., amino acids involved in subunit-subunit interactions, substrate binding and catal-ysis, and allosteric regulation. Yeast two-hybrid analyses revealed interactions of Prs1with Prs2, Prs1 with Prs3, and Prs2 with Prs3. Thus, the enzyme may contain all threepolypeptides in an unknown stoichiometry (110).

The filamentous fungus Eremothecium (Ashbya) gossypii contains four PRPPsynthase-specifying orthologs (111, 112). Two of these genes, both located on chro-mosome VII, have been analyzed: Agl080c (accession no. NP_986586), a homolog of S.cerevisiae PRS2 or PRS4, and Agr371c (accession no. NP_987037), a homolog of S.cerevisiae PRS3. The two other orthologs are homologs of S. cerevisiae PRS1 (accessionno. NP_984943) and PRS5 (accession no. NP_984410). Both of these latter E. gossypiigene products have NHRs at positions similar to those of S. cerevisiae Prs1 and Prs5,although their lengths vary. All four E. gossypii genes may specify typical class I PRPPsynthases (112). Similar to the situation with S. cerevisiae, the knockout of the PRS3homolog had more dramatic effects on various growth parameters than that ofknockout of the PRS2/PRS4 homolog. The presence of only four PRS orthologs in E.gossypii, contrary to five in S. cerevisiae, may indicate some redundancy in the PRS2 andPRS4 genes of S. cerevisiae, as only one of these is present in E. gossypii. It wasfurthermore shown that introduction of the amino acid alteration Leu133Ile orLeu132Ile into the PRS2/PRS4 or the PRS3 homolog, respectively, resulted in reductionof feedback inhibition of PRPP synthase activity. Similar effects were observed withHis196Gln or His195Gln alterations of the PRS2/PRS4 or the PRS3 homolog, respectively.These amino acid alterations of PRPP synthase were introduced to desensitize thepathway for riboflavin biosynthesis for which purine nucleotides, and thus PRPP, areprecursors (112). We shall return to explain the effect of the amino acid alterations onPRPP synthase activity in the section “Regulation of PRPP Synthase Activity.”

It is obvious from the description above that the physiology of yeast PRPP synthasediffers from that of bacteria. Although also belonging to the class I PRPP synthases,major differences exist, such as the presence of the amino acid sequences of NHR,the requirement of heteromeric oligomerization, and the involvement of PRS-specifiedpolypeptides in cellular processes other than PRPP synthesis.

Mammalian PRPP synthases and PRPP synthase-associated proteins. As describedabove, yeasts contain PRPP synthase-specifying homologs with NHRs inserted at spe-cific positions as well as “ordinary” PRPP synthase-specifying homologs without theseinsertions. Similar proteins are found in mammalian organisms, where apparent PRPPsynthases have insertions similar to the yeast NHRs. In mammals, PRPP synthasepolypeptides with an insertion have been designated PRPP synthase-associated pro-teins, or PAP. The descriptions below for each type of PRPP synthase and PAP com-plement previously published reviews (18, 20).

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(i) PRPP synthase. Among PRPP synthases of mammalian origin, only those ofhumans and rats have been characterized. Both organisms contain three PRPP synthasegenes, PRPS1 to PRPS3, that encode PRPP synthase isozymes 1 to 3. PRPS1 and PRPS2are located on the X chromosome, whereas PRPS3 is located on chromosome 7. Thelatter gene is expressed only in testes, whereas both of the former genes are expressedin all the tissues examined (11).

(a) Enzyme characteristics. Human and rat PRPP synthases also belong to the class IPRPP synthases, and as such their properties resemble the microbial class I PRPPsynthases with respect to their restricted diphosphoryl donor capacity (ATP and dATP)and allosteric regulation by Pi and ADP (18, 20). The amino acid sequence identityamong the three human PRPP synthase isozymes is 91% to 95%, whereas the identitiesof B. subtilis PRPP synthase and the three human isozymes are 45% or 46%. Exclusionchromatography of PRPP synthase activity isolated from rat liver revealed a proteincomplex with a molecular mass of 1,000 kDa or more. The protein complex consistedof four different polypeptides, the two catalytic PRPP synthase isozymes 1 and 2 as wellas the two so-called PAP-39 and PAP-41 peptides, and was estimated to contain 20molecules of isozyme 1, five of isozyme 2, and eight and one of PAP-39 and PAP-41,respectively (113). Previous characterization of human PRPP synthase may have beenperformed on mixtures of isozyme 1 and 2, although erythrocytes, the source of theenzyme in those early studies, contain predominantly isozyme 1 (114), and similar tothe rat liver enzyme, human PRPP synthase was isolated as large oligomeric structures(115, 116). Human isozymes 1 and 2 differ in 15 amino acid residues, and human andrat isozyme 1 are identical, whereas human and rat isozyme 2 differ in only 3 amino acidresidues.

Although native PRPP synthase isolated from tissues may better represent thephysiological state of the enzyme, efforts have been spent also to characterize theindividual isozymes. Isozymes 1 and 2 of both humans and rats have been purified andcharacterized after expression of their recombinant genes in E. coli (71, 73). As expectedfrom the close identity of the four enzymes, they shared many similarities in physico-chemical properties. They all oligomerize to very large structures with molecular massesaround 1,000 kDa (human isozymes 1 and 2, rat isozyme 1) or approximately 550kDa (rat isozyme 2), as determined by size exclusion chromatography. Even thoughthey are quite similar in primary structure, there are also differences betweenhuman isozymes 1 and 2. Different purification procedures were employed for thetwo isozymes: isozyme 2 was much more labile to heat inactivation than isozyme 1,isozyme 1 was saturated by lower concentrations of ATP, ribose 5-phosphate, andMg2� than isozyme 2, and isozyme 1 was more prone to inhibition by ribonucle-oside diphosphates than isozyme 2. Comparison of human and rat enzymes re-vealed very few differences (71, 73). Some kinetic properties of PRPP synthases ofhumans and rats are listed in Table 2.

(b) Three-dimensional structure. The human PRPP synthase isozyme 1 three-dimensionalstructure has been determined from a number of crystal forms (Table 1). (i) hPRS1wild-type PRPP synthase has three SO4

2� ions bound, one at the phosphate position ofribose 5-phosphate, one at the position of the �-phosphate of ADP at the allosteric site,and one at a new, third position at the parallel dimer interface; its structure wasdetermined at 2.0 Å resolution (PDB code 2h06). (ii) ATP-SO4

2�-Cd2� PRPP synthase(also wild type), with three SO4

2� bound as in the hPRS1 wild-type PRPP synthasecrystal form, ATP (modeled to only AMP) at the active site, and one Cd2� per monomer;its structure was determined at 2.2 Å resolution (PDB code 2hcr) (51). (iii) PRS1(wild-type PRPP synthase) with one SO4

2� bound; its structure was determined at 2.0Å resolution (PDB code 3s5j) (117, 118). In addition, three-dimensional structures ofsome mutant variants have been determined as well and are described below. From the46% identity among B. subtilis and human PRPP synthase isozyme 1 amino acidsequences, it comes as no surprise that the three-dimensional structures of the twoenzymes are essentially identical. Superimposition of the hPRS1 wild-type PRPP syn-thase structure and various B. subtilis PRPP synthase structures revealed root mean

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square deviation values of 1.0 to 1.2 Å. As expected, also the human PRPP synthaseisozyme 1 has the hexameric propeller structure of B. subtilis PRPP synthase (Fig. 3C)(51). In addition, the structure of human PRPP synthase isozyme 1 has been visualizedby negative stain electron microscopy. The apo- and the ADP-bound forms have athree-leaf clover structure, into which the crystal structure of isozyme 1 can dock (118).

(c) Mutant PRPP synthases. A number of variants of human PRPP synthase isozyme 1have been discovered because of their severe clinical effects on the affected individuals.As reviewed previously (119), these pathological effects are caused either by increasedactivity of PRPP synthase isozyme 1, i.e., “superactivity” of the enzyme or overexpres-sion of the PRPS1 gene, which results in an array of physiological effects, includinghyperuricemia, gouty arthritis, or neurosensory defects, or by reduced activity of PRPPsynthase isozyme 1, resulting in neuropathy, deafness, or intellectual disabilities. His-torically, these altered PRPP synthases were described by the somewhat-imprecisedesignations “gain of function” or “loss of function.” Sixteen mutant variants affecting15 amino acid positions have been discovered and are summarized in Table 4. (Nomutant forms of PRPP synthase isozyme 2 or PAP have been described). The loss offunction of PRPS1 variants (i.e., reduced activity in Table 4) are the variants Asp64Asn,Ala86Thr, Met114Thr, Gln132Pro, Val141Leu, Leu151Pro, Ile289Thr, and Gly305Arg,which had 40 to 70% of the specific activity of normal PRPP synthase (120–122). In Fig.2, each of these amino acid positions is indicated by blue arrowheads, which point to

TABLE 4 Human PRPP synthase isozyme 1 variantsa

Effect and amino acidalteration

Homologous position inB. subtilis PRPP synthase Enzyme property Reference(s)

Increased activityb

Asp51His Asp57 Reduced inhibition by ADP, increased stimulation by Pi 72Asn113Ser Asn119 Reduced inhibition by ADP, increased stimulation by Pi 7Leu128Ile Leu134 Reduced inhibition by ADP, increased stimulation by Pi 72Asp182His Asp186 Reduced inhibition by ADP, increased stimulation by Pi 72Ala189Val Ala193 Reduced inhibition by ADP, increased stimulation by Pi 72His192Leu Asp196 Reduced inhibition by ADP, increased stimulation by Pi,

reduced stability72, 123

Reduced activityGlu42Asp Asn48 Not characterized 120Asp64Asn Glu70 Reduced sp act in crude erythrocyte extract 121Ala86Thr Ile92 Reduced sp act in crude erythrocyte extract 121Met114Thr L119 Reduced sp act in crude fibroblast extract 120Gln132Pro Gln138 Reduced sp act in crude erythrocyte and fibroblast extracts 122Val141Leu Ile147 Reduced activation by Pi 412Leu151Pro Leu156 Reduced sp act in crude erythrocyte and fibroblast extracts 122Ile289Thr Ser293 Reduced sp act in crude erythrocyte extract 121Gly305Arg Gln308 Reduced sp act in crude erythrocyte extract 121

Acute lymphoblastic leukemiarelapse specificc

Val52Ala Cys58 148Ile71Val Ile77 148Cys76Ser Leu82 148Ser102Asn Ser108 148Asn113Asp Asn119 148Asp138Gly Asp144 148Asn143Ser Asp148 148Gly173Glu Gly177 148Lys175Asn Thr179 148Asp182Glu Asp186 148Ala189Thr/Val Ala193 148Leu190Ile Ile194 148Thr302Ser Val305 148Tyr310Cys Tyr313 148

aAmino acid residues are numbered from the N-terminal proline residue, as the original N-terminal methionine is removed in the mature protein (72).bA comprehensive review of the various diseases caused by mutations in the human PRPP synthase isozyme 1-encoding gene was published previously (413).cRelapse-specific PRPP synthase variants have not been characterized enzymatically.

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the homologous position in B. subtilis PRPP synthase. It is evident that none of theseamino acid alterations are within positions that are directly involved in catalysis, whichpresumably would produce inactive enzyme forms. Asp64 and Met114, equivalent to B.subtilis PRPP synthase Glu70 and Leu120, respectively, are located at stretches of aminoacids involved in the formation of the bent dimer. Val141, equivalent to B. subtilis PRPPsynthase Ile147, is located close to sequences that are involved the formation of theparallel dimer, and Gln132, Ile289, and Gly305, equivalent to B. subtilis PRPP synthaseGln138, Ser293, and Gln308 are within or close to positions involved in allostericregulation of PRPP synthase activity. Finally, Ala86, equivalent to B. subtilis PRPPsynthase Ile92, is located close to the regulatory flexible loop. Altogether, the aminoacid positions causing reduced activity of PRPP synthase isozyme 1 are located atpositions secondary to catalysis, and their effect instead may be to alter the secondarystructure, resulting in poorer enzymatic function.

The gain-of-function variants (i.e., with increased activity in Table 4) are the variantsAsp51His, Asn113Ser, Leu128Ile, Asp182His, Ala189Val, and His192Leu. These variantforms of PRPP synthase have increased activity primarily because of their reciprocaleffects on allosteric regulation by ADP and Pi, namely, reduced inhibition by the formerand increased stimulation by the latter (72, 123). Studies of these PRPP synthase mutantvariants have proven valuable in the elucidation of the allosteric regulation of theenzyme, and we shall return to a further description of them in the section “Regulationof PRPP Synthase Activity.”

The structure of three mutant variants has been determined. One is the Asp51Hisvariant, which has SO4

2� bound at the position of the �-phosphate of ADP at theallosteric site as well as Mg2� (PDB code 4f8e) (124). Patients containing the Asp51Hisvariant suffer from hyperuricemia and severe gout. The other structures are those of theSer131Ala (PDB code 2h07) and Tyr145Met (PDB code 2h08) variants, each with threeSO4

2� bound as described for the hPRS1 wild-type PRPP synthase structure. Theauthors used the observation of these SO4

2� ions as indicative of a second allostericPi-binding site (51).

(d) PRPP synthase isozyme 2 and Myc-driven cancers. As mentioned above, no mutants ofhuman PRPP synthase isozyme 2, specified by the PRPS2 gene, have been described.Remarkably, however, the activity of isozyme 2 has been shown to be critical indeveloping and maintaining cancers caused by Myc transcription factor. The eukaryotictranslation initiation factor 4E is also involved in the development of this type of cancer.The PRPS2 gene, but not the PRPS1 gene, contains a pyrimidine-rich translationalelement in the 5=-untranslated region. Myc-driven hyperactivation results in an inter-action of the eukaryotic translation initiation factor 4E and possibly other factors withthe pyrimidine-rich translational element and results in an increase of translation ofPRPS2-specific mRNA, which results in increased PRPP synthase isozyme 2 synthesisand, thus, increased nucleotide production. The increase in nucleotide synthesis inhib-its PRPP synthase isozyme 1, whereas isozyme 2 is less sensitive to feedback inhibitionby ribonucleotides. Also, the lack of the pyrimidine-rich translational element within thePRPS1 mRNA prevents the stimulation of synthesis of isozyme 1. Altogether, compellingevidence shows that PRPP synthase isozyme 2 plays an important role in the metab-olism of cells with Myc-driven cancers (125).

In addition, PRPP synthase isozyme 2, but not isozyme 1, has been shown to beprone to arginylation (419). This process has severe effects impacting PRPP synthaseactivity and stability (420).

(ii) PAP. Mammalian PAP-39 and PAP-41 may be variants of the S. cerevisiae and S.pombe PRS1-specified gene products described previously, i.e., PRPP synthase-likeproteins containing an NHR. The amino acid sequence of human PRPP synthaseisozyme 1 is 41% identical to both the PAP-39 and PAP-41 sequences; the amino acidsequence of PAP-39 is 76% identical to that of PAP-41. The NHRs are both locatedwithin the catalytic flexible loop and are 29 and 30 amino acid residues in length,respectively. Thus, the additions are located at a position similar to NHR1 of S. cerevisiaePrs1, NHR5-2 of S. cerevisiae Prs5, and the NHR of S. pombe Prs1, but they differ in length

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(Fig. 5). The identity of B. subtilis PRPP synthase and PAP-39 and PAP-41 is 31% and 34%,respectively. After removal of the additions, these values increase to only 34% and 37%,respectively. Inspection of important amino acid residues revealed that the ribose5-phosphate-binding site is well-preserved in PAP-39 and PAP-41. In contrast, residuesinvolved in the binding of ATP and residues important for catalysis are poorly con-served. Specifically, the two important residues of the catalytic flexible loop (Lys197 andArg199 in B. subtilis) are absent in the two PAPs. Data for PAP-39 are included in TableS3 in the supplemental material. Originally, the PAPs were identified as subunits thatcopurified with rat PRPP synthase. Interaction of PRPP synthase and PAP has beendemonstrated by several methodologies, such as cross-linking and coimmunoprecipi-tation. The PAP could be removed from PRPP synthase isolated from rat liver byexclusion chromatography in the presence of 1 M magnesium chloride. Both of theselatter processes yielded PRPP synthase with increased activity, which suggested thatthe PAPs play a negative regulatory role on PRPP synthase activity (113). Furthermore,a protein devoid of PRPP synthase activity was obtained after expression of thePAP-39-specifying gene in E. coli. Partial reconstitution of rat PRPP synthase activitycould be achieved by coexpression of recombinant genes specifying PRPP synthaseisozymes 1 and 2 and PAP-39 in E. coli, which resulted in an active oligomeric enzymeeluting near the void volume, i.e., an enzyme consisting of a large number of subunits(126).

Mammalian and yeast PRPP synthase structures resemble one another in interestingways. Both mammalian and yeast PRPP synthases have constituent subunits of “ordi-nary” PRPP synthase polypeptides, i.e., isozymes 1 and 2 of humans or rats and Prs2,Prs3, and Prs5 of S. cerevisiae, as well as the NHR-containing polypeptides PAP-39 andPAP-41 of humans or rats and Prs1 and Prs5 of S. cerevisiae, as described above. Indeed,the presence of multiple, heterogenous subunits, including subunits with NHR se-quences, may be widespread among eukaryotic organisms. The studies with yeastsuggest that such heterologous structures may play roles in cellular regulation that arenot yet well defined.

The crystal structures of human PAP-39 and PAP-41 have been determined at 2.7and 2.6 Å resolution (PDB codes 2c4k and 2ji4, respectively). We found that superim-position of these two structures with that of B subtilis PRPP synthase (PDB code 1dku)revealed root mean square deviation values of 1.1 and 0.97 Å for PAP-39 and PAP-41,respectively, which indicates that the three-dimensional structures of the PAPs andhuman and B. subtilis PRPP synthases are essentially identical.

Other class I PRPP synthases. Some interest has been devoted to PRPP synthase ofthe mosquito Culex pipiens pallens. This mosquito is the major vector for variousroundworms causing lymphatic filariasis (elephantiasis) and for the Japanese enceph-alitis virus. An important insecticide used in controlling C. pipiens pallens and otherinsects as well is the pyrethroid ester deltamethrin. Many insect species developdeltamethrin resistance by knockout mutations, and increased expression of specificgenes may assist in providing resistance. Expression of the PRPS1 gene is one exampleof this phenomenon. The PRPS1-encoded enzyme is a typical class I PRPP synthase. Theexact mechanism of the contribution of overexpression of PRPS1 to deltamethrinresistance remains to be established (127).

Planarians, with Dugesia japonica as an example, are studied because of theirremarkable ability to regenerate from injuries such as lost body parts. A PRPP synthase-specifying ortholog (DjPRPS) of D. japonica has been cloned. The deduced amino acidsequence reveals a typical class I PRPP synthase, with 37% identity to the amino acidsequence of B. subtilis PRPP synthase (128).

Class II PRPP Synthases

S. oleracea isozymes 3 and 4 were proposed to constitute a second class of PRPPsynthases, class II, based on a comparison of the biochemical properties of the enzymes(14, 15). Both isozymes 3 and 4 are active in the absence of Pi, and both enzymes areinhibited by the ribonucleoside diphosphates ADP and GDP in a competitive manner,

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i.e., they compete with ATP for binding at the active site. Thus, no allosteric inhibitionby either of these ribonucleoside diphosphates was detected. Kinetic parameters aresummarized in Table 2. Both of the S. oleracea isozymes are able to use ATP, dATP, GTP,CTP, and UTP as diphosphoryl donors. The apparent Michaelis-Menten constants for theother diphosphoryl donors for isozyme 3 and isozyme 4 varied between 85 and 680�M. Exclusion chromatography of isozyme 4 revealed a molecular mass of 110 kDa.Identical results were obtained in Tris- or phosphate ion-based buffer and in thepresence or absence of MgATP. Recombinant S. oleracea isozyme 3 eluted identically toisozyme 4 in exclusion chromatography. With a calculated molecular mass of thesubunit as 35.4 kDa, this value of 110 kDa is consistent with a homotrimeric quaternarystructure (46, 47). Some controversy exists on this point, as characterization of sugar-cane class II PRPP synthase revealed a molecular mass of 214 kDa, i.e., a value consistentwith a hexameric quaternary structure. Like S. oleracea PRPP synthase isozymes 3 and4, the activity of sugarcane PRPP synthase is independent of Pi (129).

Amino acid sequence alignments of PRPP synthase isozyme 3 or 4 with the aminoacid sequence of B. subtilis PRPP synthase revealed a conservation of amino acids alongthe polypeptide. Thus, a small part of the regulatory flexible loop (Phe97 to Thr114 ofisozyme 4), the PRPP-binding site consisting of the PP loop (Asp183 to Ala186 ofisozyme 4) and the ribose 5-phosphate-binding loop (Gly222 to Thr237 of isozyme 4)are extensively conserved. The catalytic flexible loop (Lys204 to Leu215) is less con-served, but the apparently indispensable residues, Lys204 and Arg206, are retained. Theamino acid residues important for binding of the adenine moiety of ATP, Phe41, Asp43,and Gln47, and the catalytically important residue His141 are also conserved (Fig. 2 andTables S1 and S2). Altogether, it is therefore likely that the mechanism of catalysis ofclass II PRPP synthases is identical or similar to that of class I PRPP synthases.

The amino acid residues involved in the formation of the two types of subunit-subunit interactions identified in B. subtilis PRPP synthase, i.e., the bent dimer and theparallel dimer, are poorly conserved in class II PRPP synthases (Table 3). In class II PRPPsynthases represented by the S. oleracea PRPP synthase isozyme 4, only 3 of the 8residues involved in the formation of the B. subtilis PRPP synthase bent dimer areconserved, and only 4 of the 12 residues involved in the formation of the B. subtilis PRPPsynthase parallel dimer are conserved. It is likely, therefore, that the quaternarystructure of class II PRPP synthases differs from that of class I enzymes. Also, the aminoacid residues identified as involved in allosteric regulation of B. subtilis PRPP synthaseare poorly conserved in S. oleracea PRPP synthase isozyme 4; among the 20 residueslisted in Table S2, only 6 are conserved, which is consistent with the lack of allostericregulation of class II PRPP synthases.

The amino acid sequences of S. oleracea PRPP synthase isozyme 2 (class I) andisozyme 3 (class II) contain extensions of 76 and 87 amino acids, respectively, at the Nterminus compared to isozyme 4. For isozyme 2, this extension contains an amino acidsequence that indicates its localization in chloroplasts, whereas for isozyme 3 theextension contains an amino acid sequence that indicates its localization in mitochon-dria. Indeed, in vitro expression of PRS2 and synthesis of isozyme 2 in the presence ofchloroplasts showed that this isozyme was transported to chloroplasts (15). Unlike S.oleracea PRPP synthase isozymes 2 and 3, the amino acid length of isozyme 4 is similarto that of other PRPP synthases (318 amino acids compared to 316 of B. subtilis PRPPsynthase), and thus isozyme 4 is believed to be located in the cytoplasm (15).

Interestingly, previous analyses of plant PRPP synthases provide some additionalinformation on the class II enzymes. PRPP synthase from spinach leaves has beenpartially purified. The activity was independent of Pi. In fact, Pi inhibited the activitywith half-maximal inhibition at 28 �M. The apparent Km values for ATP, ribose5-phosphate, and Mg2� were 36 �M, 10 �M, and 1 mM, respectively. ATP was by far thebest diphosphoryl donor, but GTP, UTP, and CTP also were diphosphoryl donors. ThePRPP synthase activity was found predominantly in the cytosol (�95% of the totalactivity), with minor contributions by the nuclear, chloroplast, and mitochondrialfractions (1 to 2%) (130). These data have to be reevaluated in light of the present

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knowledge of multiple isozymes in spinach. Rubber tree (Hevea brasiliensis) latex PRPPsynthase has been partially purified and characterized. Exclusion chromatographyrevealed a molecular mass of 200 kDa. The authors estimated the subunit molecularmass as 56 kDa following gel electrophoresis under denaturing conditions. Actually, aband corresponding to a polypeptide of molecular mass 35 kDa was also visible in thephotograph of the gel. This size, 35 kDa, is the “standard” size of PRPP synthases.Activity was obtained in the absence of Pi. ATP functioned as a diphosphoryl donor,whereas neither GTP, CTP, nor UTP functioned as a diphosphoryl donor. Apparent Km

values for ATP and ribose 5-phosphate were determined to be 0.2 M and 40 �M,respectively (131). As before, the data should be reevaluated with the current knowl-edge of the multiple forms of PRPP synthases. It remains unknown if the latex fractioncontains a specific PRPP synthase. Finally, native PRPP synthase has been characterizedin partially purified or crude preparations from black gram (Vigna mungo) hypocytols(416, 421).

BLAST analysis with the amino acid sequences of S. oleracea or A. thaliana PRPPsynthase isozyme 3 or 4 as the queries revealed the presence of these enzymes also ingreen algae species, such as Chlamydomonas reinhardtii (accession no. XP_001694483[an isozyme 3-like sequence]) (132), Chlorella variabilis (XP_005848636 [isozyme 3-like])(133), Coccomyxa subellipsoidea (XP_005643144.1 [353-amino-acid sequence, isozyme4-like]) (134), Ostreococcus lucimarinus (XP_001421441.1 [305-amino-acid sequence,isozyme 4-like] and XP_001421590 [304-amino-acid sequence, isozyme 4-like]) (135),Ostreococcus tauri (XP_003083594.1 [396-amino-acid sequence, isozyme 3-like] andXP_003083246.1 [552-amino-acid sequence]) (136), and Volvox carteri (XP_002955183[398-amino-acid sequence, isozyme 3-like]) (137). BLAST analysis with S. oleracea PRPPsynthase isoenzyme 1 as the query revealed the presence of class I PRPP synthase alsoin O. lucimarinus, C. variabilis, C. subellipsoidea, and V. carteri. Thus, in general the classII PRPP synthases seem to be an addition to the classical, class I PRPP synthases. Itshould be added that the completion of the A. thaliana genome sequencing revealedthe presence of a gene encoding a class I PRPP synthase isozyme 5 (16).

Archaeal PRPP Synthases

The biochemical characterization of PRPP synthase of M. jannaschii persuaded theauthors to propose the existence of a third class of PRPP synthases. The activity of thisenzyme is activated by Pi and it has restricted diphosphoryl donor capacity (ATP anddATP only), which are properties of class I PRPP synthases. However, ADP binds to onlythe active site, and the enzyme appears to be unregulated by an allosteric mechanisminvolving purine ribonucleoside diphosphates, which is a property of class II PRPPsynthases. Additionally, the quaternary structure of M. jannaschii PRPP synthase istetrameric (48). In light of the three-dimensional structures of PRPP synthase of T.volcanium and S. solfataricus, which are dimers, it seems more reasonable to define thisclass of PRPP synthases as “archaeal PRPP synthases” rather than class III synthases.Amino acid sequence alignments of PRPP synthase of the three archaeal species, M.jannaschii, T. volcanium, and S. solfataricus, with the amino acid sequence of B. subtilisPRPP synthase revealed conservation of amino acids along the polypeptide. Theregulatory flexible loop, the PP loop and the ribose 5-phosphate-binding loop areextensively conserved; the catalytic flexible loop is less conserved, but the apparentindispensable lysine and arginine residues are retained. The amino acid residuesimportant for binding of the adenine moiety and the catalytically important histidineresidues are also conserved (Table S1). As with the class II PRPP synthases, it is likely thatthe archaeal PRPP synthases and class I PRPP synthases share identical catalyticmechanisms.

M. jannaschii. The amino acid sequences involved in various functions are con-served in M. jannaschii PRPP synthase, i.e., the regulatory flexible loop, Leu88 to Ser104,the PP loop, Asp163 to Ala166, the catalytic flexible loop, Lys186 to Ala197, and theribose 5-phosphate-binding loop, Asp205 to Thr220 (Table S1). Also, the two-domain

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structure of the M. jannaschii PRPP synthase subunit resembles that of the B. subtilisenzyme.

Biochemical characterization of PRPP synthase from M. jannaschii revealed proper-ties unlike those described above for class I and class II PRPP synthases (Table 2).Apparently, the poor affinity for both substrates may be compensated by a very largeVmax value. Additionally, the activity of M. jannaschii PRPP synthase is stimulated by Pi,diphosphoryl donors are ATP and dATP, ADP is a competitive inhibitor, and ADP bindsto a single site per subunit, hence, the active site. These properties partly point to aclass I enzyme, i.e., restricted diphosphoryl donor capacity and activation by Pi, andpartly to a class II enzyme, i.e., lack of allosteric regulation. The quaternary structure ofM. jannaschii PRPP synthase was furthermore found to be tetrameric (48).

The structure of M. jannaschii PRPP synthase, as deduced from X-ray crystallographicanalysis, is considerably more compact than that of B. subtilis PRPP synthase, primarilybecause of truncations at the N- and C-terminal ends as well as one shorter loop in theinterior of the sequence. As with B. subtilis PRPP synthase, the structure of M. jannaschiiPRPP synthase revealed two different types of subunit-subunit interactions; these wererevealed by the structure of the apo-PRPP synthase structure (PDB code 1u9y) and thestructure of the AMP-R5P PRPP synthase complex (with AMP and ribose 5-phosphatebound; PDB code 1u9z). The tetrameric M. jannaschii PRPP synthase could be super-imposed easily with two of the subunits of B. subtilis PRPP synthase to form almostidentical bent dimers. Consistent with the bent dimer organization of M. jannaschiiPRPP synthase, 6 of the 8 amino acid residues selected (Table 3) were identical orconserved among B. subtilis and M. jannaschii PRPP synthases. Interestingly, the inter-face of the bent dimer contains residues involved in the binding of the adenine moietyof ATP, namely, Phe32B (Phe40D of B. subtilis PRPP synthase), Asp34B (Asp42D), Glu36B(Glu44D), and Arg92A (Arg101A). The rest of the active site, i.e., the binding of theremaining part of ATP and ribose 5-phosphate, is provided entirely by one subunit ofa bent dimer, indicating that substrate recognition and catalytic machinery of B. subtilisand M. jannaschii PRPP synthases are identical. The conservation of amino acid residuesinvolved in substrate binding and catalysis is apparent from the amino acid comparisonsummarized in Table S1. In contrast, the other subunit-subunit interaction of M.jannaschii did not superimpose with the linear dimers of the B. subtilis enzyme. Only 4of 12 amino acid residues involved in the parallel dimer formation of B. subtilis PRPPsynthase were identical or conserved (Table 3). The allosteric effectors of B. subtilis PRPPsynthase, ADP and Pi, bind to residues at the interface of the parallel dimer, which is notconserved among the two enzymes, and very few if any amino acid residues involvedin allosteric regulation of B. subtilis PRPP synthase are conserved in M. jannaschii PRPPsynthase (Table S2). This is consistent with the lack of allosteric regulation in vitro of M.jannaschii PRPP synthase, which binds only a single molecule of ADP per subunit (48).

T. volcanium. As described above for M. jannaschii PRPP synthase, amino acidsinvolved in the formation of the bent dimer are relatively well conserved relative to thehomologous amino acid residues of B. subtilis PRPP synthase, whereas the amino acidresidues involved in the formation of the parallel dimer are much less conserved. Aminoacid residues involved in substrate binding and catalysis are well conserved (Table S1),whereas only a few amino acid residues involved in allosteric regulation of B. subtilisPRPP synthase are conserved in T. volcanium PRPP synthase (Table S2).

Structures of several crystal forms of T. volcanium PRPP synthase have been de-scribed. (i) ADP-SO4

2� PRPP synthase, with ADP and SO42� present at the active site

representing ATP and the phosphoryl moiety of ribose 5-phosphate, determined at 1.5Å resolution (PDB code 3lrt). The active site is located in the cleft between the twodomains as in the B. subtilis enzyme. The binding of ADP is well defined; the adeninemoiety is stacked between Arg91 and Phe32 and N6 is bound to Asp34 and Glu36, andthe ribose hydroxyls are bound to Glu92, His93, and Tyr96. In addition, ADP interactswith the side chains of amino acid residues Asp34, Glu36, Arg91, and Ser214 throughwater molecules. The crystal form has an open conformation, i.e., the catalytic flexibleloop is open, resulting in full accessibility of the active site for substrates. The side chain

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nitrogens of the catalytic flexible loop Lys184 are located 14 to 16 Å away from the�-phosphate oxygen of ADP (138). (ii) mATP-SO4

2� PRPP synthase, determined at 1.8Å resolution (PDB code 3lpn). In this crystal form, the ATP analog �,�-methylene ATPreplaces ADP of the ADP-SO4

2� PRPP synthase described above. The binding of�,�-methylene ATP, including the triphosphate chain, is well defined and shows His93interacting with the oxygens of the �- and �-phosphates and Tyr96, His124, and Asp125interacting with the �-phosphate. The catalytic flexible loop remained open (138). (iii)ADP-Mg2�-R5P PRPP synthase, determined at 1.8 Å resolution (PDB code 3mbi). Here,MgADP and ribose 5-phosphate are located at the active site. Mg2� coordinates thepolypeptide through His124, the �- and �-phosphates, as well as three water mole-cules. Thus, the binding of this Mg2� resembles that of the MG2 site of B. subtilis PRPPsynthase, except for the coordination of Mg2� to the �-phosphate rather than water inthe B. subtilis enzyme. Furthermore, the catalytic flexible loop is closed, i.e., it is bentapproximately 45° toward the bound ligands, MgADP and ribose 5-phosphate, com-pared to the open form; this results in the placement of Lys184 and Arg186 close to the�-phosphate of ADP (and presumably of ATP), and in shielding of the active site and thetransition state from the solvent (138). The importance of this closure of the catalyticflexible loop for catalysis is discussed below in the section “Mechanism of Catalysis.”

The crystal structure as well as molecular sieving revealed that T. volcanium PRPPsynthase is a dimer, which does not form higher-symmetry oligomers (hexamers ortetramers) like those described for the class I PRPP synthases (138). Also, the crystalstructure with �,�-methylene ATP/ADP and ribose 5-phosphate present revealed adimeric quaternary structure. However, molecular sieving in the presence of ATP (or�,�-methylene ATP) and ribose 5-phosphate has not been performed, so formation ofa tetramer resembling that of M. jannaschii PRPP synthase under some conditionscannot be ruled out.

S. solfataricus. S. solfataricus PRPP synthase resembles PRPP synthases from M.jannaschii and T. volcanium with respect to conservation or lack of conservation ofamino acid residues involved in dimer formation (Table 3), substrate binding andcatalysis (Table S1), and allosteric regulation (Table S2). Structural information for S.solfataricus PRPP synthase is available from analysis of a crystal form with AMP andsulfate representing the phosphoryl group of ribose 5-phosphate at 2.8 Å resolution(PDB code 4twb). The enzyme appears to be active as a dimer like that described for T.volcanium PRPP synthase. Amino acid sequence identity of S. solfataricus and T.volcanium PRPP synthases is 29%. The fold of the subunit resembles that of other PRPPsynthases, and the binding of the nucleotide substrate ATP (modeled to only the AMPportion) is highly conserved in comparison to the PRPP synthases of T. volcanium, M.jannaschii, and B. subtilis (139).

Other archaeal PRPP synthases. The gene encoding PRPP synthase of the hyper-thermophilic archaeon Thermococcus kodakarensis has been expressed in E. coli andPRPP synthase purified to near homogeneity. Recombinant T. kodakarensis PRPP syn-thase accepts ATP, CTP, and GTP as diphosphoryl donors, with ATP as the preferreddiphosphoryl donor. The enzyme eluted as a monomer in exclusion chromatography(140). Chromatography was performed without the presence of substrates. It is there-fore possible that the active enzyme consists of more than one subunit. Indeed,comparison of amino acid residues involved in the formation of the bent dimer of M.jannaschii with those of T. kodakarensis revealed an almost exact match, suggestingthat active T. kodakarensis PRPP synthase may constitute a tetramer similar to that ofM. jannaschii PRPP synthase. PRPP synthase from the hyperthermophilic archaeonPyrobaculum calidifontis has been characterized and shown to resemble that of T.kodakarensis (422). The similarities of the amino acid sequence of T. kodakarensis PRPPsynthase with those of M. jannaschii, S. sulfolobus, and T. volcanium are 52 to 58%(identity, 32 to 40%), whereas with B. subtilis PRPP synthase the values are 47%(similarity) and 28% (identity).

The genomes of the following archaeal species appear to contain two prs genes:Archaeoglobus fulgidus, Archaeoglobus veneficus, Ferroglobus placidus, Halorhabdus uta-

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hensis, Methanocella conradii, Thermofilum pendens, and Thermogladius cellulolyticus. Nobiochemical information is available on the properties of the corresponding proteins.Amino acid sequence comparisons of two putative A. fulgidus PRPP synthases and B.subtilis PRPP synthase revealed a high degree of conservation of amino acid residuesinvolved in substrate binding and catalysis. Specifically, the two important lysine andarginine residues are present in both sequences: Lys172 and Arg175 in sequence 1,Lys186 and Arg 188 in sequence 2 (Table S3). Indeed, amino acid sequence compari-sons revealed that A. veneficus, F. placidus, M. conradii, T. pendens, and T. cellulolyticusmay encode two bona fide PRPP synthases, as evaluated by the presence of conservedor identical amino acids at positions homologous to B. subtilis PRPP synthase residuesPhe40, His135, and Asp174 of the ribose 5-phosphate-binding loop and Lys197 andArg199 of the catalytic flexible loop. In contrast, one of the putative PRPP synthases ofH. utahensis has aspartate and threonine rather than lysine and arginine in the catalyticflexible loop, and thus it very likely does not represent an active PRPP synthase.

In spite of large variations in protein structure among class I, class II, and archaealPRPP synthases as a result of the different oligomerization properties described above,it is obvious that the catalytic mechanism essentially is the same among the three PRPPsynthase classes, as evaluated by the conservation of the catalytically important aminoacid residues in the three classes.

MECHANISM OF CATALYSIS

The three-dimensional structure of the important catalytic flexible loop, Lys197 toMet208 of B. subtilis PRPP synthase, Lys186 to Ala197 of the M. jannaschii enzyme,Lys188 to Ile199 of the S. solfataricus enzyme, and Lys194 to Met205 of human PRPPsynthase isozyme 1, remained unresolved in studies of the structures of these enzymes(48–51, 139). However, the solution of the T. volcanium PRPP synthase structurerevealed that the binding of the substrate ribose 5-phosphate and the substrate analogMgADP resulted in a closed conformation of the catalytic flexible loop, in which strands�10 and �11 move 17 Å toward the substrates and bring the important catalyticresidues, Lys184 and Arg186, into proximity with ribose 5-phosphate and the triphos-phate chain of the nucleotide (Fig. 6A). According to this structure, Lys184 and Arg186bind to oxygen(s) of the �-phosphate of ATP. Additionally, Mg2� coordinates to anitrogen of His124 as well as to oxygens of the �- and �-phosphates of ADP and,presumably, of ATP. In the open conformation, the C-5=, the �-phosphorus, and the�-phosphorus of ADP or ATP are essentially linearly arranged, whereas in the closedconformation the �-phosphate is bent approximately 90° relative to the C-5= and the�-phosphorus and thus locates the �-phosphorus in an ideal position for nucleophilicattack by the O-1 of ribose 5-phosphate (Fig. 6B) (138). Only a single Mg2� wasidentified at the active site of T. volcanium PRPP synthase, and biochemical informationas to the number of Mg2� necessary for catalysis has not been published. This isimportant because dual roles for Mg2 have been identified in the reaction mechanismof other PRPP synthases.

Additional information was obtained from studies of two structures of activated B.subtilis PRPP synthase. In the first of these structures, a transition-state analog ofphosphoryl transfer reactions, AlF3, was employed in the presence of the product AMPand the substrate ribose 5-phosphate, which generated a transition-state bound com-plex (AlF3 PRPP synthase structure) (Fig. 6C). Here, Al3� represents the �-phosphorus,whereas the three F� ions represent the three oxygens of the �-phosphate, and the �-and �-phosphates were provided by two AMP molecules as described before. In thesecond of these structures, the substrate analog �,�-methylene ATP and the substrateribose 5-phosphate were employed to generate an active-state quaternary complex(mATP-R5P PRPP synthase structure). In the former structure containing AlF3, thecatalytic flexible loop was closed and revealed contacts between Lys197 and Arg199with the �-phosphoryl group of ATP, whereas in the second, the mATP-R5P structure,the catalytic flexible loop was open. Thus, the contacts between Lys197 and Arg199observed in the AlF3 structure appear to be lost in the active-state quaternary complex

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FIG 6 Catalytic mechanism of PRPP synthase. (A) Closure of the catalytic flexible loop of T. volcaniumPRPP synthase by superimposition of the open and closed structures (PDB codes 3lrt and 3mbi,respectively). Structural elements are colored as described for Fig. 3A. A 17-Å movement of the catalyticflexible loop, consisting of the �10 and �11 strands, results in the closed conformation necessary forcatalysis. (B) Close-up view of the binding of substrates at the active site of T. volcanium PRPP synthase,with open and closed catalytic flexible loops. In the open conformation, the triphosphate chain of ATP,modeled here to only ADP, forms a more or less linear arrangement. In the closed conformation, thetriphosphate chain, again modeled to only ADP, bends with the �-phosphate, resulting in a position idealfor attack of O-1 of ribose 5-phosphate on the �-phosphorus. An Mg2� of the closed conformation isshown as a black sphere (138). (C) Stereo view of the binding of ribose 5-phosphate, Mg2�, and thetransition state analog AlF3 to the active site of B. subtilis PRPP synthase, AlF3 PRPP synthase. (Reproducedfrom reference 54 with permission.) � indicates the �-phosphate of ATP provided by an AMP molecule;� indicates the �-phosphate of ATP provided by Al3� (bound to three F� ions); � indicates the�-phosphate of ATP provided by the phosphate of a second AMP molecule. The two Mg2� are indicatedby MG1 and MG2. Relevant amino acid residues His135, Asp174, Lys197, and Arg199 are included as well.(D) Stereo view of the binding of ribose 5-phosphate, �,�-methylene ATP, Mg2�, and Ca2� to the activesite of B. subtilis PRPP synthase in the GDP PRPP synthase complex. (Reproduced from reference 54 withpermission.) Ca2� (designated CA1) coordinates to the hydroxyls at C-1, C-2, and C-3 of ribose5-phosphate, oxygen of the �- and �-phosphates of �,�-methylene ATP, Asp174, as well as a watermolecule. The Mg2� (designated MG2) coordinates to the oxygen of C-2= of the ribosyl moiety as wellas oxygen of the �- and �-phosphates of �,�-methylene ATP, as well as to three water molecules. Thus,there is no coordination to oxygen of the �-phosphate of �,�-methylene ATP.

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(mATP-R5P), where the flexible catalytic loop remains unresolved. Thus, loop closureappears to be transient and specific for only the transition state. Al3� of the AlF3

structure has an octahedral coordination with the C-1 oxygen of ribose 5-phoshate, thethree F� ions, and also one oxygen from each of the �- and �-phosphoryl groups. Thedistances from the Al3�, which corresponds to the �-phosphorus of ATP, to the C-1oxygen of ribose 5-phosphate and the oxygen of the �-phosphoryl group are 2.0 and1.9 Å, respectively, and thus these coordinations have partial bond character. In the AlF3

structure, Mg2� is located on both sides of the triphosphate chain. As observed with T.volcanium PRPP synthase, the binding of Mg2� at the MG2 site of B. subtilis PRPPsynthase results in bending of the triphosphate chain of ATP. This bent conformationis caused by the tridentate conformation, �,�,�-MgATP. Furthermore, the Mg2� of theMG1 site coordinates to the �-phosphate (as well as to ribose 5-phosphate andAsp174), and thus the two Mg2� are responsible for a tight conformation of thetriphosphate chain of ATP, and a perfect inline geometry of the C-1 oxygen of ribose5-phosphate, the �-phosphate, and the �-phosphate is created. Furthermore, the bentconformation of the triphosphate chain results in electrostatic repulsion among thephosphoryl groups, whereas the coordination of Mg2� to a nitrogen of His135 maywithdraw charge from the phosphoryl groups coordinated to the Mg2� at the MG2 site(54).

Although the Al3�-generated transition state adopts a hexacoordinated octahedralbipyrimidal structure rather than the pentacoordinated trigonal-bipyrimidal transitionstate expected for a phosphotransfer mechanism, the data are in agreement with anassociative nucleophilic substitution, SN2, mechanism. This mechanism is supported bythe perfect inline orientation of the nucleophile (the C-1 oxygen of ribose 5-phosphate),the partial bond character between the attacking nucleophile and the transferredgroup (represented by Al3�), and a transient charge of �2 (provided by the two “free”�-phosphoryl oxygens and represented by fluorides). An effect of the two basic aminoacid residues of the catalytic flexible loop, Lys197 and Arg199, may be stabilization ofthe transient negative charge. The SN2 mechanism is supported by previous studies ofS. enterica PRPP synthase, which showed that the product of the reaction of ATP with[1-18O]ribose 5-phosphate was [18O]PRPP rather than [18O]AMP (66, 141).

REGULATION OF PRPP SYNTHASE ACTIVITY

All PRPP synthases analyzed, class I from microbes (S. enterica [69], B. subtilis [70], E.coli [45, 75], humans [72, 142], and rats [143, 144]), class II (spinach isozymes 3 and 4 [46,47]), and also the archaeon M. jannaschii (48), are inhibited in a competitive manner byADP, which binds in competition with ATP at the active site. Superimposed on thecompetition of ADP and ATP at the active site is allosteric inhibition by ADP. Thisallosteric inhibition is well documented. However, let us first mention some results ofa steady-state kinetic analysis of the binding of allosteric effectors to E. coli PRPPsynthase, which showed that Pi and ADP compete for binding to the same site (75).Additionally, Pi is required for the activity of the class I PRPP synthases of S. enterica (35,56), E. coli (45), B. subtilis (70), B. caldolyticus (55), humans (73), and rats (71), as well asfor the stability in the absence of MgATP of S. enterica and E. coli PRPP synthases (35,45). Finally, Pi can be replaced as an activator by high concentrations of SO4

2� (49, 71,73). Altogether, it is therefore more correct to say that that PRPP synthase is alloster-ically regulated with Pi as an allosteric activator and ADP as an allosteric inhibitor (75).

Allosteric inhibition of PRPP synthase by ADP was demonstrated by detailed steady-state kinetic analysis. A distinctive feature of allosteric binding by ADP is that it requiresthe active site to be fully occupied by both ATP or ADP and ribose 5-phosphate. Thiswas concluded from the observation that ADP inhibition becomes stronger as theenzyme approaches saturation with ribose 5-phosphate; this is explained by theordered binding of substrates in which ribose 5-phosphate binds last, i.e., ADP bindingat the active (ATP) site induces apparent substrate inhibition by ribose 5-phosphate(69). The presence of a separate ADP-binding site was demonstrated directly byequilibrium binding dialysis. Thus, ATP (actually the unreactive analog �,�-methylene

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ATP) binds to a single site per molecule, the active site, in the presence of ribose5-phosphate. In the absence of ribose 5-phosphate, ADP binds to the same site as ATP,whereas in the presence of ribose 5-phosphate ADP binds to the additional allostericsite to yield two bound ADPs per monomer (68).

The synthesis of PRPP was further studied in vivo. Purine starvation of S. entericaresulted in accumulation of PRPP, whereas purine nucleotides were depleted, suggest-ing that these conditions caused reduction of an inhibitor of PRPP synthesis. Thisinhibitor could very well be ADP (145). Also, the addition of adenine to cultures of S.enterica caused a dramatic reduction in the PRPP pool, which only slowly returned tothe previous level. Adenine and PRPP react in vivo to form AMP followed by theformation ADP, which causes inhibition of PRPP synthase. Therefore, the PRPP pool onlyslowly returns to the level before adenine addition (146).

PRPP synthase from some organisms such as B. subtilis, B. caldolyticus, and mammalsare also allosterically inhibited by GDP (55, 70, 71, 73).

Characterization of several mutants of human PRPP synthase isozyme 1 demon-strated the existence and physiological importance of the allosteric site in human PRPPsynthase. The mutations were identified in the human PRPS1 gene in individuals withinherited defects in purine metabolism (72). The positions of the amino acid replace-ments identified, Asp51His, Asn113Ser, Leu128Ile, Asp182His, Ala189Val, andHis192Gln, are indicated by arrows above the deduced B. subtilis PRPP synthase aminoacid sequence in Fig. 2. The affected individuals exhibit hyperuricemia, gouty arthritis,and in some cases neurosensory defects, with an X-linked pattern of inheritance. Sixdifferent mutations in human PRPP synthase isozyme 1 have been characterized at themolecular level. The kinetics of inhibition of the purified recombinant mutant enzymesby ADP and GDP confirmed that they were much less sensitive to inhibition than thewild-type isozyme and that the pattern of inhibition by ADP with respect to ATP wasaltered from a complex pattern of inhibition of the wild-type enzyme to simplecompetition. Thus, the existence of a distinct allosteric mode of inhibition by ADP andGDP was demonstrated. The fact that desensitization to allosteric inhibition results inmetabolic defects that are readily explained by overproduction of PRPP in human cellsis strong evidence that this inhibition is critical for normal regulation of PRPP synthesis.The mutations that destroy allosteric inhibition in human PRPP synthase are scatteredthroughout the primary sequence but are localized in the three-dimensional structurealmost exclusively in the interface between dimers in the hexamer.

In molecular terms, the mechanism of allosteric inhibition may be described bycomparing the three-dimensional structures of various PRPP synthase complexes. Thus,the B. subtilis SO4

2�, ADP, GDP, and mGDP PRPP synthase complexes are characterizedby tight packing of both the N- and C-terminal domains and represent an inactive form(the T-state) of the enzyme, whereas the B. subtilis AlF3 and mATP-R5P PRPP synthasecomplexes are less tightly packed and represent an active form (the R-state) of theenzyme. Of particular importance is a comparison of the crystal structures of thetransition-state complex, AlF3 PRPP synthase, and the inhibited-state quaternary com-plex, mGDP PRPP synthase. The latter complex contained Mg2�, �,�-methylene ATP,and ribose 5-phosphate in the active site as well as methylene GDP in the allosteric site.The binding of ribose 5-phosphate, the adenine base of ATP, the �-phosphate, and alsoMg2� of the MG1 site is virtually identical in the two complexes. In contrast, Mg2� atthe MG2 site was displaced, and the torsional angle of the glycosidic bond of ATPdiffered by a 55° rotation among the two structures, 72° (in mGDP PRPP synthase)versus 127° (in AlF3 PRPP synthase). This rotation displaces the �- and �-phosphates ofthe nucleotide bound to the active site and appears to prevent the inline arrangementof the C-1 oxygen of ribose 5-phosphate and the �- and �-phosphates of ATP necessaryfor nucleophilic attack. The important amino acid residues involved in these alterationsare located within the regulatory flexible loop (Tyr97 to Thr113) (Fig. 2). Thus, for thebinding of the ribose moiety of ATP, a shift in hydrogen bonding of the C-2= oxygenand Gln102 to hydrogen bonding of the C-3= oxygen and Glu110 occurred. Addition-ally, the MG2-coordinated water molecules changed from coordinating with Arg101,

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Asp103, and Arg104 to coordinate only with Asp103, whereas Arg104 changed to formhydrogen bonds to oxygens of the �-phosphoryl. Arg101 no longer formed hydrogenbonds in the T-state (54).

As expected from the analysis of the mutant variants of human PRPP synthaseisozyme 1 described above, these altered substrate-binding capabilities are accompa-nied by transitions in the hexameric structure. Reciprocal interactions between twoC-terminal domain amino acid residues, Val178 to Asp195, Arg198, and Asn209 toVal211 result in a tightly packed interface. This stretch of amino acid residues straddlesthe catalytic flexible loop (Lys197 to Met208). Particularly important is a hydrophobicpocket consisting of Ile192, Ile194, and Val211. Altogether, the interactions preventloop closure. The binding of substrates at the active site as well as SO4

2� (and hencepresumably Pi) at the allosteric site decouples these C-terminal amino acid interactionsand permits loop closure. On the other hand, when ADP or GDP is bound at theallosteric site, interactions of the C-2= hydroxyl of the bound ADP or GDP with Gln140,which interacts with the imidazole moiety of His149 through a water molecule andhence interacts with Leu134, results in maintaining the hydrophobic pocket and thetight C-terminal domain interaction. All these interactions are lost in the transition-state-like complex, AlF3 (54).

Site-directed mutagenesis of the B. subtilis prs gene was performed to generatemutant variants with amino acid alterations similar to the naturally occurring mutationsin human PRPP synthase isozyme 1 describe above (54). As observed for the humancounterparts, the kinetic properties of Pi activation among the mutant variants weregenerally shifted toward an increased affinity for Pi. The mutant B. subtilis enzymesdisplayed an approximate 10-fold decrease or more with the half-saturation concen-trations for Pi compared to the wild-type form, except for the Asp57His variant(homologous to the human Asp51His variant), whose Pi dependence on activity wassimilar to that of the wild type, and Asp196Gln (homologous to human His192Gln),whose activity appeared independent of Pi within the range of Pi concentrationsapplied.

The B. subtilis PRPP synthase variants described above were also analyzed withrespect to inhibition by the ribonucleoside diphosphate GDP, which is less complexthan ADP inhibition, as GDP only binds to the regulatory site and not the active site.When inhibition by GDP of each of the mutant PRPP synthases was analyzed at a Pi

concentration corresponding to the individual half-saturation concentration value forPi, similar Ki

GDP values were obtained for the wild type and the following variants:Asn119Ser (homologous to human Asn113Ser), Leu134Ile (homologous to humanLeu128Ile), Asp186His (homologous to human Asp182His), and Ala193Val (homologousto human Ala189Val). This observation indicates that PRPP synthase may represent theV-system of the Monod, Wyman and Changeux model for allosteric systems (147),where the activator (Pi) and the inhibitor (GDP) compete for binding to the same siteto alter the equilibrium between the R and T conformation of the enzyme. The mutantenzymes described above that have an increased affinity for Pi all have changedequilibria between conformers that further the R-conformer compared to the wild-typeenzyme.

Mapping of each of the amino acid substitutions in the structure (Fig. 2) thatinfluence this equilibrium between the active and inactive conformer shows that threeamino acid residues, Asp186, Ala193, and Asp186 are located within the region of theC-terminal domain interaction, which is important for transition of the allosteric signal,i.e., the binding of ribonucleoside diphosphate at the allosteric site. Additionally,Asn199 is located at the N-terminal interface of the bent dimer, and likewise mayperturb the binding of ribonucleoside diphosphate at the allosteric signal, as describedabove. Finally, Leu134 is involved in the transmission of the allosteric signal. Almost allof the amino acid substitutions are located on the interface within a dimer or betweendimers in the hexamer. These positions are in accordance with the existence of anequilibrium between the active and the inactive form of class I PRPP synthase that can

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be altered by side chain substitutions that stabilize any of the two conformers, theactive R or the inactive T conformer.

A large number of mutations have furthermore been identified in the genespecifying human PRPP synthase isozyme 1 of patients suffering from relapse ofacute lymphoblastic leukemia. Negative feedback-defective PRPP synthase resultedfrom mutations that arose after treatment with 6-mercaptopurine and 6-thioguanine aschemotherapeutics. Twenty-nine lesions affecting 14 amino acid positions have beenidentified. Some of these positions were already known from the previous analysis ofPRPP synthase from patients with gout or uric acid overproduction, Asn113, Asp182,and Ala189. At the latter position, two variants were identified (Ala189Thr [10 patients]and Ala189Val [1 patient]). Mapping of the amino acid alterations on the three-dimensional structure of human PRPP synthase isozyme 1 revealed the location of thevariations at the allosteric site (Ser102, Asn143, Thr302) or at the dimer interface(Asn113, Gly173, Lys175, Asp182, Ala189, Leu190). The conclusion from this analysis isthat the cytotoxic effect of administering compounds such as 6-mercaptopurine and6-thioguanine may be counteracted by mutations that accelerate the production ofPRPP and hence accelerate the de novo synthesis of purine nucleotides, which com-petes with the cytotoxic compounds (148).

Fortuitously, the presence of Ca2� during crystallization of B. subtilis PRPP synthaseallowed important conclusions about the mechanism of inhibition by this ion (54). Ca2�

was identified as a hepta-coordinating metal ion at the MG1 site of the GDP PRPPsynthase complex, whereas Mg2� occupied the MG2 site (Fig. 6D). In contrast to thisMg2�/Ca2� arrangement, two Mg2� were found at the other active site of the asym-metric unit of the GDP PRPP synthase complex. The ligands coordinated to Ca2�

resembled those coordinated to Mg2�: Asp174, oxygens of C-1, C-2, and C-3 of ribose5-phosphate, oxygen of the �-phosphate, and a water molecule, with an additionalseventh coordination to oxygen of the �-phosphate of ATP. In particular, the lattercoordination of Ca2� causes perturbation of the inline arrangement of the C-1 oxygenof ribose 5-phosphate and the �- and �-phosphates of ATP described above (54).Previous kinetic analysis revealed that Ca2�, which does not support PRPP synthaseactivity alone, is an inhibitor of PRPP synthase activity even in the presence of a verylarge excess of Mg2� (56, 57).

The detailed molecular description of the regulation of PRPP formation has promptedthe application of this knowledge for the improvement of commercially importantproducts of biochemical pathways for which PRPP is an intermediate. The previouslydiscussed improvement of riboflavin synthesis by E. gossypii harboring feedback-resistant variants of PRPP synthase is one example (112). Another example is theproduction of purine nucleosides by Bacillus amyloliquefaciens. The following PRPPsynthase variants were constructed (with the equivalent variant of human PRPP syn-thase isozyme shown in parentheses): Asp58His (Asp51His), Asn119Ser (Asn113Ser),and Leu134Ile (Leu128Ile). As expected, the B. amyloliquefaciens PRPP synthase variantsAsn119Ser and Leu134Ile were activated at lower Pi concentrations and were relativelymore resistant to inhibition by ADP and GDP than the wild type, and indeed, both B.amyloliquefaciens and B. subtilis strains carrying the mutations specifying Asn119Ser orLeu134Ile had improved production of purine nucleosides, presumably due to in-creased purine de novo synthesis. Kinetic constants of B. amyloliquefaciens PRPP syn-thase are summarized in Table 2 (149).

PHOSPHORIBOSYL BISPHOSPHATE PHOSPHOKINASE

Mutants that are defective in the prs gene have been isolated in E. coli and S. entericaas described below. Strains with knockout mutations (Δprs) have a simultaneousrequirement for five metabolites: purine and pyrimidine nucleosides, histidine, trypto-phan, and NAD (2, 3). Mutants that suppress the requirement for NAD can be easilyisolated. These mutants retain the other four requirements; their genetic defect mapsto the pstSCAB-phoU operon (150, 151). Lesions in this operon result in constitutiveexpression of the Pho regulon operons, which are involved in the assimilation of Pi and

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the acquisition of Pi from other sources (152). Genetic analysis revealed that suppres-sion of the NAD requirement phenotype originated from increased expression of thephnN cistron of the tetrakaidecacistronic phnC-P operon (150). The gene products ofthis operon are involved in the catabolism of phosphonates (152). It was furthermoreshown that the phnN cistron specifies a phosphokinase with the ability to use ribosyl1,5-bisphosphate as a substrate: ribosyl 1,5-bisphosphate � ATP ¡ PRPP � ADP, i.e., aribosyl bisphosphate phosphokinase (EC 2.7.4.23) (Fig. 7A). It was shown that theenzyme was able to convert ribosyl 1,5-bisphosphate to a product that functioned asa substrate for purified xanthine phosphoribosyltransferase and that the product wasXMP according to the following chemical equation: xanthine � PRPP ¡ XMP � PPi

(151). Subsequently, it was shown that ribosyl 1,5-bisphosphate is an intermediate inphosphonate catabolism (153). However, ribosyl 1,5-bisphosphate is also a metabolitein cells that are not thriving on phosphonate, although the origin and metabolic roleof the compound under these conditions remain unclear. Even though the reactionproduct of ribosyl bisphosphate phosphokinase is PRPP, the amount synthesized isinsufficient to replace the activity of PRPP synthase, even in cells growing withphosphonate as their Pi source (B. Hove-Jensen, unpublished result) or in cells thatexpress phnN in a multicopy plasmid (151). The amino acid sequences of ribosylbisphosphate phosphokinase and PRPP synthase show only low similarity, approxi-mately 25%. Among the amino acid residues of B. subtilis PRPP synthase with annotatedfunctions from the crystal structure, very few, if any at all, can be recognized in theamino acid sequence of ribosyl bisphosphate phosphokinase. Likewise, the similarity ofribosyl bisphosphate phosphokinase to type I phosphoribosyltransferase and type IIphosphoribosyltransferase is very low, 16% with adenine phosphoribosyltransferase(type I) and with quinolinate phosphoribosyltransferase (type II). Thus, the evolutionaryorigin and mechanism of phosphoribosyl bisphosphate phosphokinase remain enig-matic; a deeper understanding awaits the determination of the three-dimensionalstructure of the enzyme. The physiologic importance of phosphoribosyl bisphosphatephosphokinase in the context of phosphonate catabolism has been discussed andpreviously reviewed (154, 155).

DIPHOSPHORYL, NUCLEOTIDYL, AND PHOSPHORYL TRANSFER

In spite of a large number of di- or triphosphate-containing compounds, very fewenzymes catalyze the transfer of an intact diphosphoryl group, i.e., attack the�-phosphorus of ribonucleoside triphosphates, in contrast to enzymes that catalyze thetransfer of a phosphoryl group, i.e., attack the �-phosphorus, and enzymes that catalyzethe transfer of a nucleotidyl group, i.e., attack the �-phosphorus. These latter twogroups of enzymes are much more abundant than the diphosphoryltransferases. Below,we give examples of the mechanism of catalysis of each of these three enzyme classesas a means of comparing catalytic strategies for nucleophilic substitutions on the �-, �-,and �-phosphorus atoms of nucleoside triphosphate substrates.

Substitution Reactions at �-, �-, or �-PhosphatesDiphosphoryl transfer. The diphosphoryltransferases other than PRPP synthase

are 2-amino-4-hydroxy-6-hydroxymethyldihydropterin diphosphokinase (ATP:2-amino-4-hydroxy-6-hydroxymethyl-7,8-dihydropterin 6=-diphosphotransferase; EC 2.7.6.3)encoded by folK in E. coli (156), the stringent factor or GTP/GDP 3=-diphosphokinase(ATP:nucleoside-5=-phosphate pyrophosphotransferase; EC 2.7.6.5) encoded byrelA in E. coli (157, 158), and in some organisms thiamine diphosphokinase (ATP:thiamine diphosphotransferase; EC 2.7.6.2) (159) and nucleotide diphosphokinase(ATP:nucleoside-5=-phosphate diphosphotransferase; EC 2.7.6.4) (160) (Fig. 7B to F).In addition, a number of other enzymes, primarily diphosphohydrolases, catalyzereactions with nucleophilic attacks on the �-phosphorus of (deoxy)ribonucleosidetriphosphates with the release of PPi and the corresponding (deoxy)ribonucleoside 5=-monophosphate, and these are members of the so-called Nudix enzyme superfamily, asreviewed previously (161–163).

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(i) Hydroxymethyldihydropterin diphosphokinase. In addition to PRPP synthase,the crystal structure of a number of enzymes catalyzing diphosphoryl transfer reactionshas been determined. These allow the comparison of the active sites and catalyticmechanism of these enzymes. Hydroxymethyldihydropterin diphosphokinase of E. coli

FIG 7 Reactions catalyzed by diphosphoryltransferases and alternative biosynthesis of PRPP. Insome cases, ATP may be replaced by dATP. The diphosphoryl and phosphoryl moieties of the productsare shown in red and blue, respectively. (A) Reaction catalyzed by phosphoribosyl bisphosphatephosphokinase. The substrate is ribosyl 1,5-bisphosphate, the product is PRPP. (B) Reaction catalyzed byPRPP synthase. The substrate is ribose 5-phosphate, the product is PRPP. (C) Reaction catalyzed by2-amino-4-hydroxy-6-hydroxymethyldihydropterin diphosphokinase. The substrate is 2-amino-4-hydroxy-6-hydroxymethyldihydropterin, and the product is 2-amino-4-hydroxy-6-hydroxymethyldihydropterin diphos-phate. (D) Reaction catalyzed by GTP/GDP 3=-diphosphokinase (stringent factor). R may be a hydrogenor a phosphoryl group, i.e., the substrate is GDP or GTP, respectively, and the product is guanosine3=-diphosphate 5=-diphosphate (ppGpp) or guanosine 3=-diphosphate 5=-triphosphate (pppGpp), respec-tively. (E) Reaction catalyzed by nucleotide diphosphokinase. R1 may be an adenyl, a guanyl, or ahypoxanthyl univalent radical, whereas R2 may be a hydrogen, a phosphoryl, or a diphosphoryl moiety.(F) Reaction catalyzed by thiamine diphosphokinase. The substrate is thiamine, and the product isthiamine diphosphate.

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contains 159 amino acid residues, is monomeric with a molecular mass of 18 kDa, andcatalyzes the diphosphorylation of 6-hydroxymethyl 7,8-dihydropterin, the committedstep in folate biosynthesis (Fig. 7C). The structure of E. coli hydroxymethyldihydropterindiphosphokinase with a variety of ligands has been determined: these include theternary complex with a substrate analog and a substrate, enzyme-Mg2�-�,�-methyleneATP-hydroxydihydropterin complex (PDB code 1q0n) (164). Additional reports on crystalor solution structures of hydroxymethyldihydropterin diphosphokinase of E. coli andHaemophilus influenzae have been published (165–167). Structural and mechanistic prop-erties of this enzyme have been previously reviewed (168). Only 23 amino acid residues areidentical between E. coli hydroxymethyldihydropterin diphosphokinase and PRPP synthase,and the folds of the two polypeptides are very different. The former enzyme adopts a foldcomprised of four �-helices and six �-strands, �1�1�2�3�2�4�5�6�3�4. Substrate recogni-tion is mediated by three loop regions located between �1 and �1 (loop L1), between �2

and �3 (loop L2), and between �2 and �4 (loop L3). ATP binds first to hydroxymethyldihy-dropterin diphosphokinase, which results in large structural changes in the three loops, inparticular L2 and L3, and allows the binding of the second substrate, hydroxymethyldihy-dropterin (164, 169, 170). Furthermore, two Mg2� are bound within the active site (Fig. 8A).One Mg2�, MgA, coordinates to oxygens of the �- and �-phosphates of ATP, to Asp95 andAsp97 as well as to two water molecules. A second Mg2�, MgB, coordinates to oxygens ofthe �- and �-phosphates of ATP, to the hydroxyl of the substrate hydroxy-methyldihydropterin, to Asp95 and Asp97, as does the first Mg2�, and finally to a watermolecule. MgB thus coordinates both substrates. This arrangement of Mg2�, substrate, andamino acid residues places the hydroxymethyldihydropterin hydroxyl at a distance of 3.2 Åfrom the �-phosphorus of ATP (164). The triphosphate chain of ATP and the hydroxyl ofhydroxymethyldihydropterin are organized in a head-to-head arrangement, much like thatof ribose 5-phosphate and ATP of PRPP synthase, with the �-phosphate “bent” away ratherthan in a linear arrangement, which allows attack of the hydroxymethyldihydropterinhydroxyl on the �-phosphate of ATP (Fig. 8A).

(ii) Thiamine diphosphokinase. Thiamine diphosphate is the active cofactor in anumber of COC bond formation and scission processes (171). Some organisms syn-thesize thiamine diphosphate by diphosphoryl transfer of the �,�-diphosphoryl groupof ATP to thiamine (Fig. 7F), whereas others phosphorylate thiamine phosphate in asingle phosphoryl transfer reaction. The crystal structure of mouse thiamine diphos-phokinase, which utilizes the former mechanism, has been solved; the enzyme structurecontains AMP and pyrithiamine diphosphate, i.e., the products of the reaction of ATPwith pyrithiamine, a thiamine analog which contains a substituted pyrimidine moietyrather than a substituted thiazole moiety. The structure also contains one Mg2�, whichcoordinated with oxygens of the AMP-phosphate to the �-phosphate of pyrithiaminediphosphate and to the carboxyls of Asp46, Asp71, Asp73, and Asp100 (PDB code 2f17).The coordination of Mg2� to the �-phosphate of pyrithiamine diphosphate is consis-tent with (but does not demonstrate) a bent structure of the triphosphate chain of thesubstrate ATP, as was observed with PRPP synthase and hydroxymethyldihydropterindiphosphokinase (172). However, those authors also suggested that the data permitteda linear arrangement of the triphosphate chain and a more or less perpendicularorientation of the pyrithiamine, with the hydroxyl group oriented for nucleophilicattack on a �-phosphate oxygen (172). The structures of thiamine diphosphokinasesfrom a few other organisms have been solved. These include the structure of theStreptococcus mutans enzyme with thiamine diphosphate and two Mg2� present butwithout an adenylyl moiety present (PDB code 3ihk), Candida albicans with thiaminylimidodiphosphate present (173), and S. cerevisiae with thiamine present (PDB code1ig0) (174). The exact architecture of the active site with substrates and Mg2� cannotpresently be clearly established.

(iii) GTP/GDP 3=-diphosphokinase. The structure of the third diphosphoryltrans-ferase, GTP/GDP 3=-diphosphokinase (Fig. 7D), is poorly characterized, presumablybecause the C terminus associates with the ribosome. This association may hampercrystallization because that region may be disordered when not bound. However,

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important conclusions have been obtained from the studies of the bifunctional RelA/SpoT homolog (RelSeq) of Streptococcus dysgalactiae subsp. equisimilis, in which thediphosphoryltransferase (RelA) and diphosphohydrolytic (SpoT) activities are located inthe same polypeptide but in different domains (175). The diphosphoryltransferasedomain of RelSeq bears some structural resemblance to the so-called palm domain ofhuman DNA polymerase �, in which two Mg2� intricately position the substrates,primer, and deoxyribonucleoside triphosphate through coordination to the 3=-hydroxyl

FIG 8 Diphosphoryl, nucleotidyl, and phosphoryl transfer reactions. Wat, water molecules. (A) The activesite of the ternary complex of E. coli hydroxymethyldihydropterin diphosphokinase. AMPCPP, �,�-methylene ATP; HP, hydroxydihydropterin (PDB code 1q0n) (164). (B) The active site of DNA polymerase� (PDB code 1bpy) (176). (C) The active site of cAMP protein kinase A. AMPPNP, �,�-imido ATP (PDB code4hpu) (201).

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of the nucleophilic primer and to oxygens of each of the �-, �-, and �-phosphates ofthe incipient deoxyribonucleoside triphosphate, and by means of coordination to threeinvariably conserved aspartate residues (Asp190, Asp192, and Asp256) (176). UnlikeDNA polymerase � and PRPP synthase, RelSeq appears to carry out catalysis by theassistance of only a single Mg2�. Thus, the dependence of the diphosphoryl transferreaction on the Mg2� concentration parallels the ribonucleoside triphosphate concen-tration; that is, there is no stimulation of diphosphoryl transfer activity by excess Mg2�

(177). RelSeq contains only two of the three acidic residues: Asp264, which is homolo-gous to Asp190 of DNA polymerase, and Glu323, which is homologous to Asp256. Thelack of a homolog of the third aspartate residue may imply an inability to bind a secondMg2� (175). However, this view that GTP/GDP 3=-diphosphokinase requires only asingle Mg2� has been challenged. The activity of monofunctional E. coli GTP/GDP3=-diphosphokinase is stimulated by Mg2� in excess of the ribonucleoside triphos-phate (ATP and GTP) concentration; this is unlike the activity of the RelA/SpoTbifunctional M. tuberculosis GTP/GDP 3=-diphosphokinase, which was inhibited atincreased Mg2� concentrations. Analysis of mutant variants of these two GTP/GDP3=-diphosphokinases revealed the importance of the amino acid sequence 306-Glu-Any-Asp-Asp-309 in E. coli GTP/GDP 3=-diphosphokinase (homologous to 348-Arg-Any-Lys-Asp-351 in M. tuberculosis GTP/GDP 3=-diphosphokinase). After swapping thesemotifs, the activity of E. coli GTP/GDP 3=-diphosphokinase is inhibited by excess Mg2�,whereas the activity of M. tuberculosis GTP/GDP 3=-diphosphokinase chimera is stimu-lated by Mg2� (178). It is possible that the mechanism of diphosphoryl transfer ofbifunctional GTP/GDP 3=-diphosphokinases, such as those of S. dysgalactiae or M.tuberculosis, may deviate from that of PRPP synthase and hydroxymethyldihydropterindiphosphokinase by the apparent requirement of only a single Mg2�, whereas themechanism of diphosphoryl transfer of monofunctional GTP/GDP 3=-diphosphokinases,such as that of E. coli, may resemble other diphosphoryltransferases in requiring twoMg2� for activity.

(iv) MutT. For the sake of completeness, we include a brief description of onemember of the Nudix hydrolases, MutT, encoded by the mutT gene in E. coli. Like thediphosphoryltransferases, MutT catalyzes a nucleophilic substitution reaction on the�-phosphorus but uses water as the nucleophile. MutT contains 129 amino acidresidues, has a molecular mass of 15 kDa, and is active as a monomer. The solutionstructure has been determined (179). The enzyme is also designated 8-oxo-dGTPdiphosphohydrolase, because 8-oxo-dGTP appears to be the best substrate in vitro andis probably also the most important substrate in vivo. Other (deoxy)ribonucleosidetriphosphates are also hydrolyzed, albeit at much higher Km values (180). 8-oxo-dGTPresults from oxidation of dGTP and may be misincorporated into DNA opposite ofdeoxyadenylate residues, which would result in AT ¡ GC transversions (181). Thefrequency of such transversions increased by 100- to 1,000-fold in a mutT strain (182).Like PRPP synthase and other diphosphoryltransferases, MutT requires two divalentmetal ions (Mg2� or Mn2�, with a preference for Mg2�) for activity. One ion is enzymebound, and the other is bound to the (deoxy)ribonucleoside triphosphate. The enzyme-bound Mg2�/Mn2� coordinates to the nucleophilic water, to Gly38, Glu56, Glu57, andGlu98 and to a second water molecule, whereas the nucleotide-bound Mg2�/Mn2�

coordinates to oxygens of the �- and �-phosphates (162, 183). Catalysis involves theabstraction of a proton of the nucleophilic water by Glu53, followed by attack on the�-phosphorus and displacement of 8-oxo-dGMP and MgPPi (162). The biochemistry,physiology, and genetics of MutT as well as other members of the Nudix enzyme familyhave been previously reviewed (161, 162).

Nucleotidyl transfer. Nucleotidyltransferases catalyze reactions of the followingtype: acceptor � (deoxy)ribonucleoside-P-P-P ¡ acceptor-P-(deoxy)ribonucleoside �

PPi. Examples include reactions catalyzed by nucleic acid polymerases (DNAn �

dNTP ¡ DNAn�1 � PPi, or RNAn � NTP ¡ RNAn�1 � PPi,) or glucose 1-phosphateuridylyltransferase (glucose 1-phosphate � UTP ¡ UDP-glucose � PPi). Indeed, nucleicacid polymerases and sugar phosphate nucleotidyltransferases (sometimes called

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sugar nucleoside diphosphate diphosphorylases) are the most prominent members ofthis class of enzymes. These reactions have substitutions with nucleophilic attack on the�-phosphorus of ribonucleoside triphosphate or deoxyribonucleoside triphosphate incommon. The structure of the active sites of RNA and DNA polymerases are quitesimilar. Here, we return to the human DNA polymerase �, briefly described above, tofurther describe the active site and the mechanism of catalysis, which involves theintricate positioning of two Mg2� by means of coordination to three conservedaspartate residues, as originally suggested by Steitz and Steitz for DNA polymeraseI and other nucleic acid polymerases (184). Figure 8B shows the binding of the twoMg2� to the active site of DNA polymerase �. MgA coordinates to oxygens of the�-, �-, and �-phosphates to Asp190 and Asp192 and to a water molecule, whereasMgB coordinates to an oxygen of the �-phosphate to the two carboxylate oxygensof Asp190, and to Asp192 and Asp256. The sixth coordination of MgB was notdetected. Presumably, this coordination normally involves the 3=-hydroxyl of thedeoxyribosyl that receives the nucleotidyl moiety. This 3=-hydroxyl is missing in2=,3=-dideoxyribonucleotides and, possibly, the crystal packaging does not allowspace for a water molecule (176). Similar geometric arrangements are found in otherDNA polymerases, such as DNA polymerase II (185). Extensive reviews of the structureand function of DNA polymerase � and other DNA polymerases have been previouslypublished (186, 187). The binding of two Mg2� by a triad of aspartate residues is foundalso in RNA polymerases. Reviews of the importance of Mg2� in DNA and RNA synthesishave been previously published (188, 189).

An example of a sugar phosphate nucleotidyltransferases is the glmU gene product, abifunctional enzyme that catalyzes the acetylation of 2-amino-glucose 1-phosphate and theuridylylation of N-acetyl-2-aminoglucose 1-phosphate to yield UDP-N-acetylglucosamine.The two activities are located in different domains, with the uridylyltransferase allo-cated to the N-terminal domain (190). UDP-N-acetylglucosamine is a precursor of cellwall synthesis in many bacterial and fungal species as well as a substrate in glycosy-lation reactions in eukaryotic species (191). The crystal structure of GlmU of M. tuber-culosis contains two Mg2�. One (MgA

2�) coordinates to Asn239 and Asp114, to anoxygen of the phosphate of N-acetylglucosamine 1-phosphate, and to an oxygen of the�-phosphate of UTP, and thus MgA

2� bridges the two substrates. The other (MgB2�)

coordinates to oxygens of the �-, �-, and �-phosphates of UTP and to three watermolecules. Thus, MgB

2� apparently does not coordinate to any amino acid residue(and, formally, there is no MgB

2�-binding site). Rather, the uridylylation domain ofGlmU appears to harbor a single enzyme-bound Mg2� (MgA

2�), whereas the secondMg2� enters as MgUTP during the catalytic cycle. This two-Mg2�-mediated mechanismis believed to be general in sugar nucleotide synthesis (192), as similar active sitearrangements are observed for 2-keto-3-deoxymanno-octulonate cytidylyltransferaseof E. coli (PDB code 3k8d) (193), glucose 1-phosphate uridylyltransferase of Corynebac-terium glutamicum (PDB code 2pa4) (194), 2-C-methylerythritol 4-phosphate cytidylyl-transferase of Mycobacterium smegmatis (PDB code 2xwl), and M. tuberculosis (PDB code2xwn) (195).

An exception to this two-Mg2�-mediated mechanism is N-acetylglucosamine uridy-lyltransferase of C. albicans, which contains only a single Mg2�, presumably coordi-nated with oxygens of the �-, �-, and �-phosphates of UTP, i.e., MgB

2� in analogywith M. tuberculosis GlmU. Furthermore, the function of MgA

2� may be performed byLys421, because the �-N of this residue coordinates oxygens of the �- and�-phosphates of UDP-N-acetylglucosamine after product formation and, thus, it alsopresumably coordinates to oxygens of the phosphate of N-acetylglucosamine and ofthe �-phosphate of UTP during catalysis. Arg116 may also participate in the functionperformed by MgA

2� with other sugar phosphate nucleotidyltransferases (196).Phosphoryl transfer. In general, enzymes catalyzing phosphoryl transfer reactions

also require two Mg2�, as demonstrated from studies of Pyrococcus furiosus UMP kinase(PDB code 2bmu) (197), Staphylococcus aureus tagatose 6-phosphate kinase (PDB code2jg1) (198), Bifidobacterium longum N-acetylhexosamine kinase (PDB code 4wh3) (199),

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mammalian cyclin-dependent kinase 2 (PDB code 3qhw) (200), or cyclic AMP (cAMP)-dependent protein kinase A (PDB codes 1l3r and 4hpu) (201, 202). We shall use proteinkinase A for a more detailed description. Crystals of the enzyme were obtained as acomplex with reaction-inert �,�-imido ATP, Mg2�, and a serine-containing peptidesubstrate. Of the two Mg2�, MgA

2� coordinates to oxygens of the �- and�-phosphates, to the bridging N of the �- and �-phosphates of �,�-imido ATP, toAsn171 and Asp184, as well as to a water molecule, whereas the other, MgB

2�,coordinates to oxygens of the �- and �-phosphates, to both of the oxygens ofAsp184, as well as to two water molecules (Fig. 8C). Unlike the examples ofdiphosphoryl and nucleotidyl transfer reactions described above, there is apparently nocoordination between Mg2� and the nucleophilic hydroxyl of the serine residue of thepeptide substrate. However, this oxygen appears to be held firmly in place by hydrogenbonding to Lys168 and Asp166 (202).

The number of Mg2� involved in kinase-catalyzed reactions may vary from enzymeto enzyme. Typically, nucleoside diphosphokinases contain only a single Mg2�. Ade-nylate kinase catalyzes the reversible reactions AMP � ATP ¡ 2 ADP or dAMP � ATP¡ dADP � ADP (203). The enzyme contains a loop, which closes upon substratebinding and thus prevents nucleophilic attack by water on the ATP triphosphate chain.The structure of the E. coli enzyme has been probed with �,�-imido ATP (PDB code1ank). The phosphoryl group of AMP and the phosphate chain of �,�-imido ATP faceone another in an almost colinear arrangement. One oxygen of the phosphoryl groupof AMP is in close proximity to the �-phosphorus of �,�-imido ATP, and thus it is ideallyplaced for an inline attack with the subsequent formation of ADP (204). This colinearstructure of the phosphoryl groups resembles the structure of adenylate kinase ligatedto the inhibitor di(adenosine-5=)pentaphosphate, except for the presence of an addi-tional phosphoryl group (205). Mg2�, which is necessary for adenylate kinase activity,does not coordinate to side chains of the enzyme but rather coordinates to twooxygens of di(adenosine-5=)pentaphosphate as well as to four water molecules (206).This binding of Mg2� to di(adenosine-5=)pentaphosphate is consistent with �,�-MgATPas the substrate for adenylate kinase (207) and Mg2� coordination to oxygens of bothof the �-phosphates of two ADP molecules of M. tuberculosis adenylate kinase, i.e., thereaction in the direction of ATP synthesis (PDB code 2cdn) (208). Furthermore, incomparison with diphosphoryl and nucleotidyl transfer reactions described above,adenylate kinase may represent an alternative means of performing the function playedby the second Mg2� in other phosphotransferases. The stabilizing and neutralizingeffect of a second Mg2� may be replaced by one or more arginine and/or lysineresidues, such as those found near the triphosphate chain and AMP in adenylate kinase(PDB codes 1ank and 2cdb) (204, 208).

Catalytic strategies. The triphosphate chain of (deoxy)ribonucleoside triphosphatesis quite electron rich and, hence, negatively charged, so that chemical reactionsinvolving this chain require special conditions. In addition, an activated, i.e., deproto-nated, nucleophile may contribute additional negative charge to the transition state. Asdescribed above, these difficulties are circumvented at least in part by the presence ofMg2� in the active site. Figure 9 shows a schematic representation of part of the activesites of enzymes or groups of enzymes that catalyze substitution reactions at the �-, �-,or �-phosphates of (deoxy)ribonucleoside triphosphates and that contain two Mg2�.The triphosphate chain, the coordination of each Mg2�, and the nucleophile are shownfor each enzyme in Fig. 9. Additional amino acid residues may be involved in thebinding of the substrates but are not shown in Fig. 9. The two Mg2� may have multiplefunctions that include exact positioning of the substrates for transition-state formation,electrostatic stabilization of the nucleophile and the transition state, and electrostaticstabilization of the product(s) or leaving group(s). As described before for PRPPsynthase, one Mg2� (MgB [Fig. 9]) presumably is involved in the stabilization of thenucleophile, i.e., the acceptor of the diphosphoryl group, the C-1 hydroxyl of ribose5-phosphate. Similarly, MgB of nucleotidyltransferases may participate in stabilization ofthe nucleophiles, i.e., hexose phosphate (Fig. 9C) and deprotonated DNA primer

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FIG 9 Catalytic strategies. The coordination of Mg2� (shown as blue spheres) of the active site to thesubstrates, to amino acid residues, and to water molecules (shown as red spheres) is schematicallyillustrated by blue lines. (A) cAMP-dependent protein kinase A (PDB code 4hpu) (201). (B) PRPP synthase.R5P, ribose 5-phosphate. MgA was designated MG2, and MgB was designated MG1 before (54). (C) Hexose1-phosphate nucleotidyltransfrease, based on the structure of N-acetylglucosamine 1-phosphate uridy-lyltransferase (PDB code 4g87). MgB coordinates to aspartate and asparagine residues (192). (D) DNApolymerase based on the structure of DNA polymerase � (PDB code 1bpy) (176). The three acidic residuesare usually aspartates, but occasionally one aspartate is replaced by a glutamate (185, 186). The structureis valid also for RNA polymerases (187).

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(Fig. 9D) (189). In contrast, no coordination is observed between MgB and the�-phosphate-attacking nucleophilic serine residue in protein kinase A (Fig. 9A), andnucleophile positioning involves amino acid residues. As in the case of MgB andPRPP synthase, MgB of nucleotidyltransferases binds the nucleophile and maystabilize the deprotonated nucleophile (Fig. 9C and D). Interestingly, MgB of PRPPsynthase and the nucleotidyltransferases coordinate both of the substrates, i.e.,ribose 5-phosphate and ATP in PRPP synthase (Fig. 9B), N-acetylglucosamine1-phosphate and UTP in N-acetylglucosamine 1-phosphate uridylyltransferase (Fig. 9C),and the DNA template and deoxyribonucleoside triphosphate in DNA polymerases (Fig.9D). The �-phosphate of nucleoside triphosphates carries two negative charges atphysiologic pH, whereas the �- and �-phosphates carry one negative charge each.These charges are neutralized by Mg2�. The second Mg2�, MgA, in the enzymes underdiscussion typically binds the substrate (deoxy)ribonucleoside triphosphate. Thus, theform bound to the enzyme in all the cases shown appears to be the tridentate�,�,�-MgATP. The MgA therefore not only partly neutralizes the triphosphate chain butalso provides the proper conformation of the substrate to the enzyme. The generalbelief is that the functioning of MgA includes a stabilization of the transition state. It isnoteworthy that the two MgA and MgB are localized on both sides of the triphosphatechain, except in the nucleic acid polymerases, where the two Mg2� are located on thesame side of the triphosphate chain.

UTILIZATION OF PRPP

The concentration of PRPP is approximately 0.5 mM in cells of S. enterica growingexponentially in minimal salts medium with glucose as carbon source. This value can becompared to the concentrations of ATP, GTP, UTP and CTP, which are 3, 0.9, 0.9, and 0.5mM, respectively (19). PRPP is utilized in nucleotide, amino acid, and cofactor biosyn-thetic pathways. The relative consumption of PRPP for each of these processes in E. colihas been estimated. Thus, purine and pyrimidine nucleotide biosyntheses each con-sume 30 to 40% of the PRPP synthesized, whereas histidine and tryptophan biosyn-theses each consume 10 to 15% of the PRPP synthesized, and NAD and NADPbiosyntheses consume approximately 1% of the PRPP synthesized (1).

Reactions at the Anomeric Carbon of PRPP

The reactions occurring at the anomeric carbon of PRPP greatly outnumber thereactions occurring at the diphosphoryl chain. The reactions at the anomeric carbon arecatalyzed by phosphoribosyltransferases, whereas hydrolases catalyze the reactions atthe diphosphoryl chain. Phosphoribosyltransferases catalyze reactions in which N-, C-,or O-glycosidic bonds are formed with the concomitant formation of PPi (Fig. 1). Weshall review first here the reactions occurring at the anomeric carbon and then thereactions occurring at the diphosphate chain.

Comparison of phosphoribosyltransferase and PRPP synthase structures. Thestructure of phosphoribosyltransferases vary to some extent. The most widespreadstructure, present in the so-called type I phosphoribosyltransferases, consists of a coreof a five-stranded �-sheet surrounded by a number of helices and additionally, a loopstructure, a hood, which participates in the binding of the nucleobase substrate, and a(catalytic) flexible loop, which remains unresolved until the binding of PRPP, when it closesthe active site for catalysis (209, 210). The type I phosphoribosyltransferases utilize purineor pyrimidine bases or glutamine-derived ammonia as the phosphoribosyl acceptor.Remarkably, in spite of only approximately 20% amino acid sequence identity amongtype I phosphoribosyltransferases and PRPP synthase, the three-dimensional structures oftype I phosphoribosyltransferases strongly resemble those of each domain of PRPPsynthase, as shown in Fig. 10A for T. volcanium PRPP synthase and Toxoplasma gondiihypoxanthine-guanine phosphoribosyltransferase. We shall use the T. gondiihypoxanthine-guanine phosphoribosyltransferase as a structural “prototype” of type Iphosphoribosyltransferases, as many of the structural features of this enzyme apply tomost other type I phosphoribosyltransferases as well. The homology of the central

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five-stranded �-sheet and the surrounding helices is evident in both structures, as isalso the homologous position of the catalytic flexible loop of PRPP synthase and theflexible loop of the phosphoribosyltransferase.

A comparison of amino acid residues of PRPP synthase and various type I phospho-ribosyltransferases reveals only a single conserved region, a series of hydrophobicresidues, usually three, followed by one or two aspartate residues, two or threehydrophobic residues, and a four-residue sequence characteristic of sharp turns in thepolypeptide chain, such as Thr-Gly-Gly-Thr (44, 45, 211). This region in B. subtilis PRPPsynthase (216-Gly-Lys-Thr-Ala-Ile-Leu-Ile-Asp-Asp-Ile-Ile-Asp-Thr-Ala-Gly-Thr-232) is theribose 5-phosphate-binding loop described above (45), whereas the correspondingregion in T. gondii hypoxanthine-guanine phosphoribosyltransferase (139-Asp-Lys-His-Val-Leu-Ile-Val-Glu-Asp-Ile-Val-Asp-Thr-Gly-Phe-Thr-154) is the PRPP-binding loop (212).Additionally, the PP loop 79-Lys-Glu-80 of T. gondii hypoxanthine-guanine phosphori-bosyltransferase and amino acid residues of the flexible catalytic loop (residues 114 to134 in T. gondii hypoxanthine-guanine phosphoribosyltransferase) are involved in thebinding of PRPP or MgPRPP in other phosphoribosyltransferases; collectively, thesesequences constitute the PRPP-binding site (209, 210). The three-dimensional struc-tures of the ribose 5-phosphate-binding loop of PRPP synthase and the PRPP-bindingloop of phosphoribosyltransferases are very much alike, as shown in Fig. 10B, which

FIG 10 Similarity of the folds of the PRPP synthase domain and type I phosphoribosyltransferases. (A)Superimposition of the C-terminal domain (amino acids 151 to 286) of T. volcanium PRPP synthase (blue;PDB code 3mbi) (138) and T. gondii hypoxanthine-guanine phosphoribosyltransferase (green; PDB code1fsg) (212). The PP, ribose 5-phosphate (R5P), and catalytic flexible (CF) loops are indicated. (B) Substratebinding at the active sites of B. subtilis PRPP synthase and T. gondii hypoxanthine-guanine phosphori-bosyltransferase. (Reproduced from reference 54 with permission.) The B. subtilis PRPP synthasetransition-state analog (Fig. 6C) consisting of the phosphoryl moiety of AMP (�), AlF3 (�), the phosphorylmoiety of a second AMP molecule (�), and ribose 5-phosphate, as well as Asp223 and Asp224 are shownas thick lines, and the two Mg2� are shown as green spheres (54). Residues from the structure of T. gondiihypoxanthine-guanine phosphoribosyltransferase in complex with 9-deazaguanine (not shown), PRPP,and Mg2�, as well as Glu146 and Asp147 (PDB code 1fsg) (212), are shown as thin yellow lines or spheressuperimposed on the AlF3 PRPP synthase structure.

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shows a superimposition of parts of the active site of B. subtilis PRPP synthase andT. gondii hypoxanthine-guanine phosphoribosyltransferase: the binding of ribose5-phosphate, the �- and �-phosphates of ATP, and two Mg2� at the active site of B.subtilis PRPP synthase and the binding of PRPP and Mg2� at the active site of T. gondiihypoxanthine-guanine phosphoribosyltransferase (54, 212).

N-Glycosidic bond formation. By far the most abundant reactions involving PRPPare formation of N-glycosidic bonds. The prototype of reactions involving PRPP isshown in Fig. 1, and examples of the products containing carbons of PRPP are shownin Fig. 11. A nitrogen-containing compound displaces the diphosphoryl group ofPRPP in a nucleophilic reaction with inversion of the anomeric carbon (for example,adenine � PRPP ¡ AMP � PPi). In general, the nitrogen-containing compounds arearomatic bases. In purine and pyrimidine nucleotide biosynthesis, the ribosyl group ofPRPP remains the ribosyl or deoxyribosyl group of the nucleotides. Similarly, in cofactor(NAD and NADP) biosynthesis, the ribosyl group of PRPP remains the ribosyl group ofthe cofactors. In contrast, in tryptophan and histidine biosynthesis, only some of theribosyl carbons are built into the products. Thus, in tryptophan biosynthesis, C-2 andC-3 of the indolyl moiety originate from C-1 and C-2, respectively, of the ribosyl moiety

FIG 11 N-Glycosidic bond formation with PRPP. PRPP and atoms of the products derived from PRPP are shown in red. Eachreaction produces an N-glycosidic 5=-phosphoribosyl compound and PPi. The reaction with quinolinate also producescarbon dioxide. The compounds a to e are products of de novo reactions: a, 5-phosphoribosyl 1-amine; b, orotidine5=-monophosphate; c, 5=-phosphoribosylnicotinate; d, 5=-phosphoribosyl-ATP; e, 5=-phosphoribosylanthranilate. The com-pounds f to k are products of salvage reactions: f, AMP; g, IMP; h, XMP; i, GMP; j, UMP; k, 5=-phosphoribosylnicotinate.

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of PRPP, whereas five of the six carbons of histidine originate from the ribosyl moietyof PRPP with the carboxyl group originating from C-5 of PRPP. The nitrogen-containingcompounds utilized by phosphoribosyltransferases may be the purine bases adenine,hypoxanthine, xanthine, or guanine, the pyrimidine bases may be orotate or uracil, thepyridine bases may be quinolinate or nicotinate, or the bases may be anthranilate orATP. A variant among phosphoribosyltransferases, amidophosphoribosyltransferase,the first enzyme of de novo purine biosynthesis, utilizes glutamine-derived ammonia.Altogether, 10 enzymes utilize PRPP for N-glycosidic bond formation, with 11 productsin organisms, including E. coli and B. subtilis (Fig. 11).

As we shall see below, PRPP is a very versatile compound for the delivery ofphosphoribosyl moieties. A large number of phosphoribosyltransferases have beendescribed. Although they all use PRPP as phosphoribosyl donor and, presumably, theyfollow similar chemical mechanisms, they differ widely in their three-dimensionalstructures. The enzymes range from the type I phosphoribosyltransferases, which areprimarily involved in purine and pyrimidine nucleotide biosynthesis, type II phospho-ribosyltransferases, which are involved in pyridine nucleotide biosynthesis, to type IIIand type IV phosphoribosyltransferases, which are involved in histidine and tryptophanbiosynthesis, respectively. The three-dimensional structures of a large number ofphosphoribosyltransferases have been determined. Detailed reviews on the biochem-istry of phosphoribosyltransferases have been published (209, 210, 213).

(i) Purine nucleotide biosynthesis. Nucleotide biosynthesis occurs via two sets ofreactions: de novo reactions, in which nucleotides are generated by the addition ofatoms one by one to the starting material, and the so-called salvage or auxiliaryreactions, by which preformed nucleobases are utilized for nucleotide biosynthesis.

(a) Purine de novo synthesis. De novo purine nucleotide biosynthesis commences withPRPP. The purine ring is built on the phosphoribosyl moiety, atom by atom, beginningwith the addition to C-1 of the ribosyl moiety of the nitrogen of glutamine-derived NH3,which subsequently becomes N-9 of the complete purine moiety. This reaction iscatalyzed by amidophosphoribosyltransferase (5-phospho-�-D-ribosylamine:diphos-phate phospho-�-D-ribosyltransferase [glutamate-amidating]; EC 2.4.2.14), which is en-coded by purF in E. coli: glutamine � PRPP � H2O ¡ glutamate � 5-phosphoribosyl1-amine � PPi (Fig. 11, compound a) (214). Amidophosphoribosyltransferase containsa domain that catalyzes the hydrolysis of glutamine to form glutamate and ammonia.Subsequently, the ammonia is transferred through a channel 20 Å long to the phos-phoribosyltransferase domain active site, where PRPP is already positioned for furtherreaction (215). The process of ammonia channeling in amidophosphoribosyltrans-ferases has been previously reviewed (216). Although it is unusual for a phosphoribo-syltransferase to have an additional enzyme activity, such as glutaminase, amidophos-phoribosyltransferase has a typical type I phosphoribosyltransferase fold with thefive-stranded �-sheets surrounded by two helices on either side, a hood, a catalyticflexible loop, as well as an MgPRPP-binding site consisting of the PRPP-binding loopand a PP loop. The enzyme binds PRPP in a manner similar to that described above forT. gondii hypoxanthine-guanine phosphoribosyltransferase (Fig. 10B).

(b) Purine salvage synthesis. The purine salvage or auxiliary reactions consist of severalenzymes that phosphoribosylate adenine, or the 6-oxopurines hypoxanthine, xanthine,and guanine. The products of these phosphoribosylation reactions are AMP, IMP, XMP,and GMP, respectively (Fig. 11, compounds f to i). In general, in most organisms thereis a single phosphoribosyltransferase that is specific for adenine and at least oneadditional phosphoribosyltransferase with specificity toward 6-oxopurine. E. coliand S. enterica both contain an apt-specified adenine phosphoribosyltransferase,hpt-specified hypoxanthine phosphoribosyltransferase, and gpt-specified xanthine-guanine phosphoribosyltransferase (19), whereas S. solfataricus contains a pgT-1-specified adenine phosphoribosyltransferase and a pgT-2-specified 6-oxopurinephosphoribosyltransferase with affinity for hypoxanthine, xanthine, and guanine(217). The purine phosphoribosyltransferase-catalyzed reactions are responsible for

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the utilization of the purine compounds adenine, hypoxanthine, xanthine, andguanine as purine sources for purine auxotrophic strains.

The specificities of purine phosphoribosyltransferases are not readily deduced fromtheir amino acid sequences. The PRPP-binding loop of adenine phosphoribosyltrans-ferase (EC 2.4.2.7) (adenine � PRPP ¡ AMP � PPi) of E. coli is 126-Asp-Asp-Leu-Leu-Ala-Thr-131. The sequence Asp-Asp-hyd-hyd-Ala-Thr (hyd indicates hydrophobic resi-dues) is found in a large majority of adenine phosphoribosyltransferases. An exceptionis S. solfataricus adenine phosphoribosyltransferase, whose sequence is 95-Asp-Asp-Ile-Thr-Asp-Thr-100. On the other hand, hypoxanthine and hypoxanthine-guanine phos-phoribosyltransferases contain the homologous sequence Glu-Asp-Ile-Ile-Asp-Thr/Ser.That is, a glutamate residue replaces the first acidic residue of the Asp-Asp dipeptideand the underlined Asp replaces an alanine residue of adenine phosphoribosyltrans-ferase, exemplified by Ala130 of the E.coli adenine phosphoribosyltransferase. E. coligpt-specified xanthine-guanine phosphoribosyltransferase instead has the sequence88-Asp-Asp-Leu-Val-Asp-Thr-93, i.e., an Asp-Asp dipeptide as in adenine phosphoribo-syltransferase and Asp92 as in oxopurine phosphoribosyltransferase (217).

The lengths of the adenine phosphoribosyltransferases show some heterogeneity.The enzyme has a basal size of approximately 180 amino acid residues, as seen in theenzymes from E. coli, the purine de novo synthesis-lacking protozoan parasite Giardialamblia, S. cerevisiae, and humans, whereas the enzymes from the protozoan parasiteLeishmania donovani and the archeon S. solfataricus have 237 and 210 amino acidresidues, respectively. Nevertheless, all of them share the common core structure oftype I phosphoribosyltransferases (218–220). The additional amino acid residues of S.solfataricus and L. donovani adenine phosphoribosyltransferases protrude from boththe N and C termini relative to the 180-amino-acid adenine phosphoribosyltransferasesand may contribute to structural variations of the hood (221, 222). The structures havebeen determined with the apo-forms or with adenine or AMP bound in the active site.Particularly interesting is the structure of the G. lamblia enzyme with the purine analog9-deazaadenine and MgPRPP bound at the active site. In this structure, the hydroxylsof the ribose moiety coordinate to Mg2� and the 124-Glu-Asp-125 acidic dipeptide,whereas the 5-phosphate is bound to the 128-Ala-Thr-Gly-Gly-Thr-132 subset of thePRPP-binding loop, and the diphosphate chain ligates to Mg2� and to Arg63, which ispart of the PP loop. Superimposition of the 9-deazaadenine and MgPRPP-containingstructure and an AMP-containing structure revealed a 2.1 Å movement of the anomericcarbon of PRPP during catalysis, whereas the 5- or 5=-phosphate and adenine moietiesessentially remained fixed (218).

As already mentioned, the specificity of phosphoribosyltransferases with affinity for6-oxopurine varies depending on the organism. Any given 6-oxopurine phosphoribo-syltransferase has some affinity for all three 6-oxopurines (hypoxanthine, xanthine, andguanine), but in general has a preference for one or two of the compounds. Thediscrimination between 6-amino and 6-oxo purines is based on hydrogen bonding tothe nitrogen versus oxygen of the purine ring (209, 213). Thus, the allocation of aphosphoribosyltransferase to the EC 2.4.2.8 (hypoxanthine phosphoribosyltransferase)class as opposed to the EC 2.4.2.22 (xanthine phosphoribosyltransferase) class maybe a somewhat superficial and misleading description. The binding of PRPP to the6-oxypurine phosphoribosyltransferases is similar to that described above for adeninephosphoribosyltransferase; that is, it involves a PRPP-binding site consisting of aPRPP-binding loop with a carboxylate dipeptide, Asp-Asp or Glu-Asp, a PP loop, and aThr/Ser-Gly-Glu-Thr consensus sequence involved in the binding of the 5-phosphatemoiety.

(ii) Pyrimidine nucleotide biosynthesis. Unlike de novo purine nucleotide biosyn-thesis, with de novo pyrimidine nucleotide biosynthesis the pyrimidine moiety is firstsynthesized and the product, orotate, and PRPP are assembled to form orotidine5=-monophosphate (Fig. 11, compound b) in a reaction catalyzed by orotate phospho-ribosyltransferase (orotidine-5=-phosphate:diphosphate phospho-�-D-ribosyl-transferase,EC 2.4.2.10; orotate � PRPP ¡ orotidine 5=-monophosphate � PPi). Orotate phospho-

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ribosyltransferase, a typical type I phosphoribosyltransferase, contains the “usual”structural elements, a five-stranded �-sheet surrounded by a number of helices, usuallyfour, a hood structure involved in the binding of the substrate orotate, and a flexiblecatalytic loop, which closes the active site on catalysis. The structures of orotatephosphoribosyltransferase from a variety of sources, including S. enterica (223), E. coli(224), S. mutans (225), Plasmodium falciparum (226), and L. donovani (227) have beenpublished.

The three-dimensional structure of S. enterica orotate phosphoribosyltransferase incomplex with orotate and MgPRPP revealed an asymmetric structure of the dimer,because both substrates bind to the active site of one subunit (B), whereas only orotatewas bound to the A subunit. The flexible catalytic loop of subunit A closes the activesite of subunit B, whose flexible catalytic loop remains unstructured and presumablyopen. Orotate is bound by amino acids of the hood, and PRPP is bound in thePRPP-binding loop, which includes the 124-Asp-Asp-125 dipeptide (coordinating tohydroxyls of ribose C-2 and C-3), the sequence 128-Thr-Ala-Gly-Thr-131 (binding the5-phosphate), and Tyr72 and Lys73 (the PP loop). A single Mg2� coordinates theoxygens of ribosyl C-1, C-2, and C-3, an oxygen of the �-phosphate as well as two watermolecules. On catalytic flexible loop closure, lysine, arginine, and histidine residues ofthis loop bind to the diphosphate chain, and, together with the Mg2� they participatein neutralizing the diphosphate chain during catalysis (228). The structure reveals thepresence of two shared active sites at the interface of the dimer, which has been alsodemonstrated by formation of a heterodimer composed of one subunit defective inPRPP binding and one subunit defective in the catalytic flexible loop (229).

The structure of S. cerevisiae orotate phosphoribosyltransferase resembles that of S.enterica (230). As mentioned, the dimeric structure is asymmetric. This asymmetry,together with the movement of the catalytic flexible loops during (i.e., before and after)catalysis, as well as kinetic analysis results prompted the authors to suggest analternating active site catalytic mechanism, i.e., only one active site is functional at anygiven time (230, 231).

Unlike the purine salvage reactions, pyrimidine salvage involves the activity of onlya single phosphoribosyltransferase: uracil phosphoribosyltransferase (UMP:diphosphatephospho-�-D-ribosyltransferase; EC 2.4.2.9; uracil � PRPP ¡ UMP � PPi) (Fig. 11,compound j). Uracil phosphoribosyltransferase allows organisms to utilize uracil formedendogenously by nucleic acid catabolism or supplied by the environment. Uracilphosphoribosyltransferase requires Mg2� for activity. In general, the activity of theenzyme is allosterically activated by GTP. GTP causes association of subunits to tetram-ers, as observed for uracil phosphoribosyltransferase from E. coli (232) and T. gondii(233). Alternatively, GTP functions as an allosteric activator of the oligomeric uracilphosphoribosyltransferase of Sulfolobus species (234–237). The enzyme from M. tuber-culosis differs from the aforementioned in being unaffected by GTP (238, 239), and theenzyme from Giardia intestinalis dimerizes in the presence of GTP (240).

The PRPP-binding motif of uracil phosphoribosyltransferase differs markedly fromthat of other phosphoribosyltransferases or PRPP synthase in that the typical acidicdipeptide (Asp-Asp or Glu-Asp) is replaced by Asp-Pro (highlighted in bold in thesequence below), as illustrated here with the E. coli uracil phosphoribosyltransferaseamino acid sequence: 123-Glu-Arg-Met-Ala-Leu-Ile-Val-Asp-Pro-Met-Leu-Ala-Thr-Gly-Gly-Ser-138 (241). The Asp-Pro dipeptide appears to be universal among uracil phos-phoribosyltransferases, as it is present in all the enzymes from the organisms men-tioned above as well as yeasts (242), bacilli (243, 244), and lactococci (245). Analysis ofa mutant variant that had this particular proline residue replaced by an aspartaterevealed a 100-fold increase in the Km value for uracil and a 50-fold reduction in the kcat

value. The Km value for PRPP was reduced in the mutant, whereas the kcat/KmPRPP value

was almost unchanged. It was therefore concluded by the authors that the prolineresidue of the PRPP-binding loop of uracil phosphoribosyltransferase “is of little im-portance for binding of PRPP to the free enzyme, but is critical for binding of uracil tothe enzyme-PRPP complex and for the catalytic rate” (246).

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Two variant forms of uracil phosphoribosyltransferase are found in Gram-positiveorganisms, such as B. caldolyticus and B. subtilis. Both genes were shown to comple-ment E. coli upp lesions and therefore encode active enzymes. One of these (upp)specifies uracil phosphoribosyltransferase, whereas the other (pyrR) specifies a regula-tory protein involved in regulation of pyrimidine gene expression, PyrR (247, 248). Weshall return to PyrR below in the section “PRPP as Mediator of Metabolic Regulation.”

Finally, a phosphoribosyltransferase activity for dioxotetrahydropyrimidine (dioxo-tetrahydropyrimidine phosphoribosyltransferase; EC 2.4.2.20) has been characterized(249). Whether it has a separate activity or is a variant of other pyrimidine phosphori-bosyltransferases remains unresolved.

(iii) NAD biosynthesis. The phosphoribosyltransferases of NAD biosynthesis aremembers of the so-called type II phosphoribosyltransferases. De novo pyridine nucle-otide biosynthesis resembles de novo pyrimidine nucleotide biosynthesis in that thearomatic base is synthesized prior to phosphoribosylation (250). The reaction with PRPPand quinolinate is catalyzed by quinolinate phosphoribosyltransferase (�-nicotinate-D-ribonucleotide:diphosphate phospho-�-D-ribosyltransferase [carboxylating]; EC 2.4.2.19)encoded by nadC in E. coli: quinolinate � PRPP ¡ 5=-phospho �-D-ribosylnicotinate �

PPi � CO2. 5=-Phosphoribosylnicotinate is also designated nicotinate mononucleotide(Fig. 11, compund c). The reaction is unusual in producing carbon dioxide (251, 252). Inpyridine cofactor salvage synthesis, nicotinate phosphoribosyltransferase (5-phospho-�-D-ribose 1-diphosphate:nicotinate ligase [ADP, diphosphate-forming]; EC 6.3.4.21),encoded by pncB in E. coli, catalyzes the reaction nicotinate � PRPP (� ATP) ¡5=-phospho-�-D-ribosyl-nicotinate � PPi (� ADP � Pi). This reaction is unusual in thatthe enzyme is stimulated by ATP in a stoichiometric manner. That is, the enzyme isphosphorylated by ATP at His219, which results in the formation of a high-affinity formwith Km values for nicotinate and PRPP 200-fold lower than those of the unphosphor-ylated enzyme. Also, the coupling of ATP hydrolysis to the phosphoribosyltransferasereaction causes a change in the equilibrium value, from 0.67 to 1,100 (253–255).Mutational replacement of His219 completely abrogates phosphorylation and stimu-lation by ATP (256). Mammalian organisms such as mice and humans contain yet adifferent pyridine phosphoribosyltransferase, nicotinamide phosphoribosyltransferase(nicotinamide-D-ribonucleotide:diphosphate phosphor-�-D-ribosyltransferase; EC 2.4.2.12):nicotinamide � PRPP ¡ 5=-phospho-�-D-ribosyl-nicotinamide (nicotinamide mononu-cleotide, NMN) � PPi (257, 258). As with purine and pyrimidine nucleotide biosynthesis,PRPP is used in both the de novo reactions of NAD synthesis (quinolinate phosphori-bosyltransferase) and in NAD salvage reactions (nicotinate and nicotinamide phospho-ribosyltransferases).

(a) Quinolinate phosphoribosyltransferase. The structures of quinolinate phosphoribo-syltransferase from a number of sources have been determined. These sources includeM. tuberculosis (259), S. enterica (211), Helicobacter pylori (260), Thermotoga maritima(261), and S. cerevisiae (262). The structures of these enzymes resemble one another.The structure of the ternary complex of M. tuberculosis quinolinate phosphoribosyl-transferase with a quinolinate analog (phthalate) and a PRPP analog (5-phosphoribosyl-1-[�-methylene]diphosphate, i.e., the two phosphorus atoms are connected by amethylene moiety) reveals the binding of quinolinate and PRPP together with twodivalent metal ions (in this case, Mn2�). In the ternary complex, the ribosyl moiety ofPRPP is hydrogen bonded to Glu201 and to Asp222 through the C-2 and C-3 hydroxyls.The two divalent metal ions coordinate to the same two hydroxyls, to the diphosphatechain or to water molecules; the ions do not coordinate to enzyme amino acid residues.The diphosphate chain forms hydrogen bonds to arginine, lysine, and aspartate resi-dues. The 5-phosphate is hydrogen bonded to, among other residues, Gly270 andGly249. Thus, although the fold of quinolinate phosphoribosyltransferase (a type IIphosphoribosyltransferase) is very different from the fold of type I phosphoribo-syltransferases, some similarities are evident. PRPP binding involves two acidicresidues (Glu201 and Asp222), which is reminiscent of the Asp-Asp dipeptide of type Iphosphoribosyltransferases and of PRPP synthase; the 5-phosphate binding involves

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glycine residues, and diphosphate binding involves several basic residues. Althoughdifferent in structure, similar arrangements have evolved for the binding of the com-mon substrate PRPP, which suggests a common theme in the design of active sites.

(b) Nicotinate phosphoribosyltransferase. The structure of nicotinate phosphoribosyl-transferase of the archaeon Thermoplasma acidophilum revealed a three-domain ar-rangement, where the structures of the N-terminal and central domains resemble thetwo domains of quinolinate phosphoribosyltransferases and a unique C-terminal do-main is added. Only a few differences were observed in the number of �-strandsbetween nicotinate and quinolinate phosphoribosyltransferases. The phosphate groupof PRPP binds to T. acidophilum nicotinate phosphoribosyltransferase in a mode similarto the binding of the PRPP analog to M. tuberculosis quinolinate phosphoribosyltrans-ferase, and the structures of the active sites of the two enzymes are very similar in spiteof little sequence conservation between them (263).

The C-terminal domain, approximately 100 amino acid residues in length, is com-posed of seven �-strands and one helix. Enzymological data for T. acidophilum nicoti-nate phosphoribosyltransferase are not available, so it remains unknown if the activityof this enzyme is stimulated by ATP. It is possible that the C-terminal domain partici-pates in phosphorylation and ATP hydrolysis and, by analogy, the phosphorylation sitemay be present in the central domain, although a His219 homolog is not found in T.acidophilum nicotinate phosphoribosyltransferase. The C-terminal domain of S. entericanicotinate phosphoribosyltransferase is exposed to the solvent; trypsin treatmentremoved 24 C-terminal amino acid residues and inactivated the ATPase activity. Thisinactivation was prevented by ATP binding, which provides evidence for the involve-ment of the C-terminal domain in energy coupling (264).

(c) Nicotinamide phosphoribosyltransferase. Vertebrates lack nicotinamide deamidaseand, thus, do not produce nicotinate from nicotinamide. Rather, they salvage nicotin-amide by means of nicotinamide phosphoribosyltransferase.

The sandwich and �/�-barrel structures described above for M. tuberculosis quino-linate phosphoribosyltransferase are repeated in human nicotinamide phosphoribosyl-transferase. The position of 5=- or 5-phosphate maps identically in structures with5=-phospho �-D-ribosylnicotinamide or with PRPP bound, whereas the ribose moietiesare displaced relative to one another. The 5-phosphate of PRPP is hydrogen bound toGly383 and Gly384 (both of the A subunit), which are part of the 381-Gly-Ser-Gly-Gly-Gly-385 loop that is similar to the corresponding loop of T. acidophilum nicotinatephosphoribosyltransferase (263). The diphosphate of PRPP is hydrogen bonded toArg39 (B subunit), Arg40 (B subunit), Arg196 (A subunit), and Lys400 (B subunit), andthe C-2 and C-3 hydroxyls are hydrogen bonded to Arg311 and Asp313 (both of the Asubunit). Thus, the binding of PRPP to human nicotinamide phosphoribosyltransferaseis homologous to the binding of PRPP to the other type II phosphoribosyltransferases(265).

As expected, nicotinamide phosphoribosyltransferases from mouse and rat havestructures similar to that of the human enzyme (266, 267). There is extensive structuralresemblance between the type II phosphoribosyltransferases, namely, quinolinate,nicotinate, and nicotinamide phosphoribosyltransferases. Chappie and coworkers haveoffered a theory for the evolution of the type II phosphoribosyltransferases, in whichnicotinamide phosphoribosyltransferase evolved from nicotinate phosphoribosyltrans-ferase, which in turn evolved from quinolinate phosphoribosyltransferase (268).

Studies of human nicotinamide phosphoribosyltransferase have attracted interest,because the enzyme has been identified as a growth factor for early-stage B cells(hence it is called “pre-B cell colony-enhancing factor”) and has been associated withbinding to and activation of the insulin receptor and has been also called “visfatin.” Fora review of the many apparent functions of human nicotinamide phosphoribosyltrans-ferase, consult the article by Garten and colleagues (269). It is unclear if the manyfunctions may be ascribed simply to variations in the NAD pool. However, because ofthe increased synthesis of nicotinamide phosphoribosyltransferase in tumor tissues,studies have been directed toward the isolation of inhibitors of the enzyme. Several of

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these inhibitors could be converted enzymatically to phosphoribosylated derivatives.Analysis of the structure of a ribosylated inhibitor bound to nicotinamide phosphori-bosyltransferase confirmed the binding of its 5-phosphoribosyl moiety (270).

(iv) Histidine biosynthesis. In histidine biosynthesis, all of the five carbons of PRPPare retained. The carboxylate, the �-C and the �-C of the amino propanoic side chainof histidine are generated from C-5, C-4, and C-3, respectively, of PRPP, whereas C-4 andC-5 of the imidazole moiety are derived from C-2 and C-1 of PRPP. In the first step ofhistidine biosynthesis, PRPP and ATP react with the formation of PPi and N-(5=-phospho-�-D-ribosyl)-ATP, which has an N-glycosidic bond of C-1 of the ribosyl moietyto N-1 of ATP (Fig. 11, compound d) (271). The reaction is catalyzed by ATP phospho-ribosyltransferase (ATP phosphoribosyltransferase; EC 2.4.2.17), which is encoded byhisG in E. coli (272, 273).

There are two subfamilies of ATP phosphoribosyltransferases. They have differentquaternary structures (hexa- or octameric) (274–276) but similar molecular architecturesof their catalytic domains. The E. coli ATP phosphoribosyltransferase polypeptidecontains three domains. Domain I consists of four parallel and two antiparallel �-strandsflanked by two �-helices on each side; domain II is constructed of four parallel and oneantiparallel �-strand flanked by one helix on one side and two helices on the other side.Domain III is a histidine-binding domain (277). Domains I and II constitute the catalyticfold of ATP phosphoribosyltransferase. Thus, although this fold appears to be reminis-cent of the type I phosphoribosyltransferase and PRPP synthase domain fold, it repre-sents a unique fold. Furthermore, ATP phosphoribosyltransferases lack the hood andthe catalytic flexible loop characteristic of type I phosphoribosyltransferases. Addition-ally, the active form of both types of enzymes are dimers or higher-order oligomers.

The active form of E. coli ATP phosphoribosyltransferase is a dimer of the three-domain polypeptide. Activity is inhibited by AMP, which prevents the binding of ATPand PRPP. The binding of the inhibitors histidine and eventually also guanosine3=-diphosphate 5=-diphosphate result in the formation of a hexameric, inactive quater-nary structure (277–279). There is only 27% sequence identity between E. coli guanine-xanthine phosphoribosyltransferase and ATP phosphoribosyltransferase. A PRPP-binding loop located within domain II, however, is easily identified in the ATPphosphoribosyltransferase sequence 162-Gly-Leu-Ala-Asp-Ala-Ile-Cys-Asp-Leu-Val-Ser-Thr-Gly-Ala-Thr-176. The structure of ATP phosphoribosyltransferase from M. tubercu-losis is similar to that of E. coli; in this case, the PRPP -binding loop was identified as147-Gly-Val-Ala-Asp-Ala-Ile-Ala-Asp-Val-Val-Gly-Arg-Thr-Leu-Ser-162 (280). ATP phos-phoribosyltransferases, such as those of E. coli and M. tuberculosis, are sometimesdesignated HisGL, with L designating “long” form. This is in contrast to HisGS, with Sdesignating “short” form in, for example, L. lactis. HisGS ATP phosphoribosyltransferaseslack domain III and instead associate with a distinct polypeptide, HisZ, to form activeoctameric oligomers, whereas the isolated HisG and HisZ polypeptides associate intoinactive dimers. The catalytic fold of HisGS ATP phosphoribosyltransferases is almostidentical to that of HisGL ATP phosphoribosyltransferases, and the active site is located,once again, in a cleft between domain I and II. PRPP-binding loops have been identifiedin ATP phosphoribosyltransferases of L. lactis (148-Gly-Leu-Ala-Asp-Ala-Ile-Val-Asp-Ile-Val-Glu-Thr-Gly-Asn-Thr-162) (281) and of the hyperthermophilic bacterium T. maritima(140-Gly-Leu-Ser-Asp-Leu-Ile-Val-Asp-Ile-Thr-Glu-Thr-Gly-Arg-Thr-155) (282). Mutationsthat replaced Thr159 or Thr162 of L. lactis ATP phosphoribosyltransferase confirmed theimportance of this sequence in PRPP binding (283).

(v) Tryptophan biosynthesis. Contrary to the situation in histidine biosynthesis, onlytwo carbons of PRPP are retained in tryptophan. C-2 and C-3 of the indole moiety oftryptophan originate from C-1 and C-2, respectively, of PRPP. The tryptophan biosyn-thetic intermediate anthranilate reacts with PRPP in a reaction catalyzed by anthranilatephosphoribosyltransferase (EC 2.4.2.18), encoded by trpD, which results in the forma-tion of N-(5-phospho-�-D-ribosyl)-anthranilate (Fig. 11, compound e) and PPi. Theanthranilate phosphoribosyltransferase isolated from E. coli is bifunctional: it alsocontains glutamine chorismate amidotransferase activity. Consequently, the E. coli trpD

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gene product is considerably larger, a total of 531 amino acid residues, than mono-functional anthranilate phosphoribosyltransferases, such as those from S. solfataricus orM. tuberculosis, which have 345 or 370 amino acid residues, respectively (284, 285).Other tryptophan biosynthetic enzymes from various species have been shown toharbor two activities.

The overall three-dimensional structure of anthranilate phosphoribosyltransferase issimilar among all of the species so far analyzed, i.e., the enterobacterium Pectobacte-rium carotovorum (286), S. solfataricus (284), M. tuberculosis (285), Xanthomonas camp-estris (PDB code 4hkm), T. thermophilus (PDB code 1v8g), and a cyanobacterial species(Nostoc sp.) (PDB code 1vqu). The two-domain structure consists of a smaller N-terminaldomain of six helices (the �-domain) and a larger C-terminal domain of six parallel�-sheets and one antiparallel �-sheet, as well as eight or more helices (the �/�-domain). Although the five-stranded parallel �-sheet characteristic of type I phospho-ribosyltransferases is recognizable, anthranilate phosphoribosyltransferase contains ad-ditional structural features and thus constitutes a separate fold sometimes designatedtype IV phosphoribosyltransferase (287). The enzyme is active as a homodimer; the�-domain is responsible for the dimerization. Binding of PRPP and anthranilate ismediated in a cleft of the �/�-domain. The binding of PRPP is promoted by two Mg2�

or Mn2�, which accounts for a requirement of Mg2� for activity (288). The binding ofPRPP has been well mapped in the structures of S. solfataricus and M. tuberculosisanthranilate phosphoribosyltransferase. The amino acid residues involved are thesequence 79-Gly-Thr-Gly-Gly-Asp-83, Lys106 and 223-Asp-Glu-224 of the S. solfataricusenzyme (107-Gly-Thr-Gly-Gly-Asp-111, Lys136 and 251-Asp-Glu-252, respectively, of theM. tuberculosis enzyme). Remarkably, in spite of this sequence conservation, the bind-ing of PRPP in the two enzymes is very different. In common, one divalent cation (MG1)coordinates to the �- and �-phosphates of PRPP. Another divalent cation (MG2)coordinates to the two acidic residues (Asp-Glu). The Gly-Thr-Gly-Gly-Asp seqence“wraps around” PRPP with the amino-terminal glycine binding to the diphosphate chainand the carboxy-terminal aspartate or glycine binding to the 5-phosphate. Specifically,Gly79 forms hydrogen bond to the �-phosphate in the S. solfataricus enzyme. Lys106of the latter enzyme binds to the �-phosphate, whereas Lys135 of the M. tuberculosisenzyme binds to the �-phosphate. Thus, this sequence may be compared to the PPloop of type I phosphoribosyltransferases and of PRPP synthases. Among the differ-ences in PRPP binding between the two enzymes are the binding properties of Asp-Glu,which bind to MG2 and to the oxygen of C-5 (Asp residue only) in S. solfataricusanthranilate phosphoribosyltransferase. MG2 furthermore coordinates to the5-phosphate and to the oxygens of C-4 and C-5 of ribose 5-phosphate (289). Thecorresponding aspartate and gluatmate residues of the M. tuberculosis enzyme arecoordinated to MG1 (Glu252) or to MG2 (Asp251). The PRPP divalent cation coordina-tion activities of the latter enzyme are directed exclusively toward the diphosphatechain (285). The conserved glycine-rich sequence 79-Gly-Thr-Gly-Gly-Asp-83 resemblesthe subsequence Asp-Asp-Xxx-Xxx-Thr-Gly-Gly-Thr of several phosphoribosyltrans-ferases and the glycine-containing subsequence of the PRPP-binding loop of PRPPsynthase (228-Thr-Ala-Gly-Thr-232, 234-Ser-Gly-Gly-Thr-237 or 217-Thr-Gly-Gly-Thr-220of B. subtilis, S. solfataricus, or M. jannaschii PRPP synthase, respectively). These se-quences of phosphoribosyltransferases and PRPP synthases are involved in the bindingof the 5-phosphate moiety of PRPP or ribose 5-phosphate.

Each subunit of anthranilate phosphoribosyltransferase binds two anthranilate mol-ecules in addition to one PRPP molecule and two Mg2�. Three sites for anthranilate areresponsible for tunnel movement of anthranilate, with a concomitant 180° flipping ofthe compound toward MgPRPP, which at this point is presumably already present.However, anthranilate may bind to the outer end of the tunnel before MgPRPP isbound, and binding of the latter compound may participate in tunnel formation (290).Interestingly, in S. solfataricus anthranilate phosphoribosyltransferase Gly79 also formsa hydrogen bond to the anthranilate nitrogen. Thus, this residue may be particularly

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important in positioning anthranilate and PRPP for inline attack of anthranilate on C-1of PRPP (289).

Tryptophan biosynthesis in M. tuberculosis has attracted much attention, as trpmutant strains of this organism are avirulent (291). Since the organism’s host, humans,are unable to synthesize tryptophan, a search for inhibitors of tryptophan biosyntheticenzymes, including for anthranilate phosphoribosyltransferase, has been conducted inhopes of identifying an antimicrobial agent (287, 290, 292).

(vi) Phosphoribosyl transfer without participation of PRPP. For the sake of com-pleteness, the cobT-encoded nicotinate nucleotide 5,6-dimethylbenzimidazole phos-phoribosyltransferase (EC 2.4.2.21) of S. enterica and E. coli should be mentioned at thispoint. Although this enzyme is a phosphoribosyltransferase, it does not utilize PRPP asa substrate. Rather, the 5-phosphoribosyl moiety is detached from the nicotinamidemononucleotide and is added to the 5,6-dimethylbenzimidazole, and thus oneN-glycoside bond is replaced by a second N-glycoside bond (293, 294). A variant of thisenzyme (EC 2.4.2.55) has been characterized from the bacterium Sporomusa ovata. Incontrast to nicotinate nucleotide:5,6-dimethylbenzimidazole phosphoribosyltrans-ferase, the S. ovata enzyme (EC 2.4.2.55) also utilizes phenol derivatives as acceptors inthe transfer of the 5-phosphoribosyl moiety (295).

C-Glycosidic bond formation: tetrahydromethanopterin biosynthesis. Tetrahydro-methanopterin is one of a number of cofactors involved in methanogenesis and sulfatereduction in a variety of archaeal species (296, 297) and methylotrophic bacteria (298,299). The pathway of tetrahydromethanopterin biosynthesis has been deduced fromthe structures of a series of metabolites meticulously solved by White and colleagues(4, 300 –303). The function of methanopterin is analogous to that of dihydrofolate, i.e.,it is a carrier of one-carbon molecules, including formyl, methylene, and methyl groups.Both methanopterin and folic acid contain a substituted pteridyl moiety as well as anaryl component apparently derived from 4-aminobenzoate. However, methanopterincontains both deoxyribulosyl and phosphoribosyl moieties that are not present infolate. Both of these pentose moieties are derived from PRPP. The two PRPP-utilizingbiochemical reactions of methanopterin biosynthesis are shown in Fig. 12A. Initially, and inwhat is regarded as the first committed step in tetrahydromethanopterin biosynthesis,4-hydroxybenzoate condenses with PRPP with the formation of 5=-phospho-�-D-ribosyl4-hydroxybenzene as well as PPi and carbon dioxide (Fig. 12A, reaction 1). The product5=-phospho-�-D-ribosyl 4-hydroxybenzene contains a carbon-carbon bond betweenthe ribosyl and aryl moieties (Fig. 12A, compound m). The enzyme catalyzing this reactionhas affinity for both 4-hydroxy- and 4-aminobenzoate (304, 305). In vitro and in vivoanalyses used 4-aminobenzoate to elucidate the metabolic pathway. The enzyme has beendesignated 4-(�-D-ribofuranosyl)aminobenzene-5=-phosphate synthase, and in M. jann-aschii it is encoded by the MJ1427 gene. Although 4-hydroxybenzoate rather than4-aminobenzoate appears to be the origin of the aryl moiety of methanopterin, at least inM. jannaschii (304), we shall use the designation aminobenzoate phosphoribosyltransferasefor this enzyme, because most studies have used 4-aminobenzoate as the substrate.Indeed, the product of the aminobenzoate phosphoribosyltransferase-catalyzed reactionwith 4-hydroxybenzoate as the substrate, 5=-phospho-�-D-ribosyl 4-hydroxybenzene, istransformed in a subsequent enzymatic step to the amino derivative 5=-phospho-�-D-ribosyl 4-aminobenzene (Fig. 12A, compound n).

The aminobenzoate phosphoribosyltransferases of the methanogenic archaeaMethanosarcina thermophila, M. jannaschii, and Methanothermobacter thermautotrophi-cus as well as the hyperthermophilic, nonmethanogenic (sulfate-reducing) archaeon A.fulgidus have been purified and studied. Edman degradation of aminobenzoate phos-phoribosyltransferase isolated from cell extracts of M. thermophila strain TM-1 was usedto establish the sequence of the 20 N-terminal amino acid residues, which were thenused to identify aminobenzoate phosphoribosyltransferase-encoding genes in a varietyof organisms (299). The molecular mass of the native enzyme is 63.5 kDa, and that ofthe subunit is 32.6 kDa, i.e., the enzyme is a homodimer. PPi and 5=-phospho-�-D-ribosyl4-aminobenzene have been demonstrated to be products of the reaction. The reaction

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FIG 12 C- and O-glycosidic bond formation with PRPP. The phosphoribosyl donor PRPP is shown in red, and atoms derived from PRPP inthe intermediates and products are also shown in red. (A) Biosynthesis of tetrahydromethanopterin. Compound labels: l, 4-hydroxybenzoate;m, 5=-phospho-�-D-ribosyl 4-hydroxybenzene; n, 5=-phospho-�-D-ribosyl 4-aminobenzene; o, N-[(7,8-dihydropterin-6-yl)methyl]-4-(1-deoxy-D-ribulosyl)aminobenzene; p, 1-(4-{N-[(7,8-dihydropterin-6-yl)methyl]amino}phenyl)-5-(5-phospho-�-D-ribulosyl)-1-deoxyribitol. Enzyme1, 4-aminobenzoate phosphoribosyltransferase (5-phospho-�-D-ribose 1-diphosphate:4-aminobenzoate 5-phospho-�-D-ribofuranosyltransferase [decarboxylating], EC 2.4.2.54); enzyme 2, 1-(4-{N-[(7,8-dihydropterin-6-yl)methyl]amino}phenyl)-5-(5-phospho-�-D-ribulosyl)-1-deoxyribitol synthase. This enzyme activity has not been identified. The four reactions leading fromcompound m to compound n in effect convert a hydroxy group to an amino group and involve the formation of phosphate ester andthe addition of an aspartyl residue and the removal of Pi and fumarate (304). The three reactions leading from compound n tocompound o involve the attachment of a pterin derivative (R � N-[7,8-dihydropterin-6-yl]methyl) to the nitrogen of compound nfollowed by opening of the ribosyl moiety and isomerization to a ribulose derivative and dephosphorylation to form compound o. The

(Continued on next page)

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requires Mg2� or Mn2�, the latter ion being the most efficient. The optimal pH is 4.8,and the optimal temperature is 50°C. Some kinetic constants were determined; theapparent Km

4-aminobenzoate was 58 �M and the apparent KmPRPP was 3.6 mM. Pyridoxal

phosphate is suggested to be involved in the reaction, but data are inconclusive in thisrespect, and it has been suggested that pyridoxal phosphate may be involved in thereaction in a way that does not require the carbonyl of pyridoxal phosphate (301, 305).

The M. jannaschii aminobenzoate phosphoribosyltransferase, encoded by the geneMJ1427, resembles that of M. thermophila (306). Product inhibition studies establishedthe reaction as an ordered Bi-Ter mechanism with binding of PRPP followed by4-aminobenzoate, and release of CO2, then 4-(�-D-ribofuranosyl)aminobenzene-5=-phosphate, and finally PPi. As with the enzyme from M. thermophila, the affinity for PRPPis low (Km

PRPP, 1.5 mM), whereas that for 4-aminobenzoate is lower (Km4-aminobenzoate, 0.15

mM). A kcat value was determined to be 0.23 s�1. Phosphoribosyltransferase-catalyzedreactions are thought to proceed with the intermediate formation of a ribooxocarbeniumion as well as a negatively charged nucleophile. Aminobenzoate phosphoribosyltransferaseposes a special problem for this generalization. How does the enzyme stabilize the negativecharge at C-1 of the nucleophile when the substrate already carries a negatively chargedcarboxyl group (4-aminobenzoate or 4-hydrozybenzoate)? No prosthetic group, neitherpyridoxal 5-phosphate nor a pyruvoyl moiety, appear to be present, indicating that enzymeside chains are only involved in the catalytic process (306).

In A. fulgidus, the gene AF2089 encodes aminobenzoate phosphoribosyltransferase.Following cloning and expression of the gene in E. coli, the enzyme was partly purified.It has a temperature optimum at 70°C and maximal activity at pH 5.3. Gel filtrationrevealed a molecular mass of 57 kDa, corresponding to a homodimer, because themolecular mass of the subunit was established as 35.5 kDa (307). Finally, a procedurefor the purification of aminobenzoate phosphoribosyltransferase from M. thermau-totrophicus has been published (307).

Some archaeal species appear to contain two aminobenzoate phosphoribosyl-transferase-encoding genes. These are Pyrococcus abyssi, where the aminobenzoatephosphoribosyltransferases (sequence designations PAB0141 and PAB1694) show 30%amino acid sequence identity, Pyrococcus horikoshii (sequence designations PH0227and PH1228), whose enzymes show 29% amino sequence identity, and Aeropyrumpernix (sequence designations APE2425 and APE1512), whose enzymes show 28%amino acid sequence identity (299).

None of the aminobenzoate phosphoribosyltransferases contains a classical PRPP-binding site, as is the case for type II phosphoribosyltransferases. As mentioned, thebinding of PRPP to type II phosphoribosyltransferases occurs by interaction of a fewamino acid residues to the phosphate moieties of PRPP. Amino acid sequence com-parison of aminobenzoate phosphoribosyltransferases with M. tuberculosis or T. acido-philum nicotinate and quinolinate phosphoribosyltransferases did not reveal conservedamino acid residues between these phosphoribosyltransferases.

O-Glycosidic bond formation. (i) Tetrahydromethanopterin biosynthesis. Anotherreaction of archaeal methanopterin biosynthesis utilizes PRPP as a substrate; interest-

FIG 12 Legend (Continued)boxed compound is the product of the pathway for 5,6,7,8-tetrahydromethanopterin (with a complete structure of the pteridylmoiety), the active cofactor in transformation of carbon dioxide to methane in methanogenic Archaea (301), and is formed fromcompound p by attachment of a glutamyl moiety to the phosphate group followed by dehydrogenation of the pteridyl moiety. (B)Biosynthesis of arabinosyl monophosphodecaprenol. Compound labels: q, decaprenyl phosphate; r, 5-phospho-�-D-ribosyl 1-O-monophosphodecaprenol (decaprenylphospho-�-D-ribosyl 5-phosphate). Enzyme 3, decaprenyl phosphate phosphoribosyltransferase(5-phospho-�-D-ribosyl 1-diphosphate:decaprenyl-phosphate 5-phosphoribosyltransferase; EC 2.4.2.45). The boxed compound is thearabinosyl donor arabinosyl monophosphodecaprenol, which is formed from compound r by dephosphorylation of the ester at C-5of the ribosyl moiety followed by epimerization. The latter two reactions occur outside the cell (5). (C) Biosynthesis of butirosin.Compound labels: s, neamine; t, 5�-phosphoribostamycin. Enzyme 4, neamine phosphoribosyltransferase. The boxed compound isbutirosin. Two isomers are synthesized; one contains a ribosyl moiety (shown) and a second contains an arabinosyl moiety (notshown). Other aminoglycoside antibiotics derived from neamine, i.e., neomycin B, paromomycin, and lividomycin B, lack the4-amino-2-hydroxybutyryl side chain but contain an N-acetylaminoglucosyl moiety attached to C-3� of the ribosyl moiety, and thepseudodisaccharides of the three compounds are differently decorated with hydroxyl and amino groups (318).

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ingly, this reaction results in the formation of an O-glycosidic bond (Fig. 12A, reaction2). The phosphoribosyl acceptor substrate of this reaction is N-[(7,8-dihydropterin-6-yl)methyl]-4-(1-deoxy-D-ribulosyl)aminobenzene (Fig. 12A, compound o), and the prod-uct is 1-(4-{N-[(7,8-dihydropterin-6-yl)methyl]amino}phenyl)-5-(5-phospho-�-D-ribulosyl)-1-deoxyribitol (Fig. 12A, compound p), i.e., C-1 of the phosphoribosyl moiety is attachedto a hydroxyl group of the ribulosyl moiety of compound o, with inversion of theanomeric carbon of the phosphoribosyl moiety, as usual (301). The enzyme catalyzingthis reaction has not been identified. One additional reaction is necessary to convertcompound p to the final cofactor (Fig. 12A) and it is described in the figure legend.

(ii) Arabinosyl monophosphodecaprenol biosynthesis. Arabinogalactan is an im-portant constituent of the cell wall of mycobacteria, such as M. tuberculosis and M.smegmatis. Arabinogalactan contains D-arabinosyl and D-galactosyl moieties. Polymer-ization of the arabinosyl moieties is catalyzed by a number of arabinosyltransferases,for which the donor is a lipid-linked arabinosyl donor, �-D-arabinosyl monophos-phodecaprenol (reviewed previously in references 5 and 308). Tracer methodologyapplied to membrane preparation of M. smegmatis to study the origin of thearabinosyl derivatives revealed that ribulose 5-phosphate was a precursor, but thatribulose 5-phosphate was not converted directly to arabinose 5-phosphate (309).Rather, the pentosyl phosphate donor was shown to be PRPP. Thus, followingincubation of M. smegmatis membrane preparations with [14C]PRPP, the productionof [14C]ribosyl phosphopolyprenol, [14C]arabinosyl phosphopolyprenol, 5-phospho-[14C]ribosyl phosphopolyprenol, and 5-phospo-[14C]arabinosyl phosphopolyprenolwas demonstrated, which showed the involvement of PRPP in the synthesis of arabi-nose derivatives for cell wall formation in this organism Fig. 12B (310). Subsequently, anM. tuberculosis gene (Rv3806c) encoding an enzyme that phosphoribosylates deca-prenyl phosphate was identified, cloned, and expressed in E. coli. The enzyme is anintegral membrane protein; membrane fractions of an E. coli strain hosting Rv3806cwere able to convert PRPP and decaprenyl phosphate into 5-phospho-�-D-ribosyl1-phosphoryldecaprenol (311). The enzyme is designated 5-phospho-�-D-ribosyl-1-diphosphate:decaprenyl-phosphate 5-phosphoribosyltransferase. In analogy withother phosphoribosyltransferases, we shall use the name decaprenyl phosphate phos-phoribosyltransferase.

Inspection of the amino acid sequence of decaprenyl phosphate phosphoribosyl-transferase does not reveal the classical PRPP-binding site that is characteristic of PRPPsynthases and type I phosphoribosyltransferases. As mentioned, decaprenyl phosphatephosphoribosyltransferase is a membrane-bound enzyme, which is consistent withboth a membrane-soluble substrate (decaprenyl phosphate) and a soluble substrate(PRPP). A topology with nine transmembrane domains was suggested for the structureof M. tuberculosis decaprenyl phosphate phosphoribosyltransferase by molecular mod-eling studies. According to this model, the active site may involve the cytoplasmic loopII. From a search of amino acid sequence alignments with other phosphoribosyltrans-ferases, a sequence, 73-Asn-Asp-x-x-Asp-77 (where x indicates any amino acid), wassuggested as important for PRPP binding and enzymatic function (312). Indeed, the M.tuberculosis decaprenyl phosphate phosphoribosyltransferase amino acid sequence69-Val-Tyr-Leu-Val-Asn-Asp-Val-Arg-Asp-77 resembles a large part of the PRPP bindingloop of E. coli hypoxanthine-guanine phosphoribosyltransferase, 95-Val-Leu-Ile-Val-Glu-Asp-Ile-Ile-Asp-103, as well as the corresponding amino acid sequence of other phos-phoribosyltransferases (213, 313). These amino acids are located within the cytoplasmicloop II and the junction of this loop and the transmembrane 2-domain. Analysis of thedecaprenyl phosphate phosphoribosyltransferase replacement mutants Asn73Gln andAsp77Glu revealed significantly increased Km values for PRPP, providing evidence thatthis sequence is involved in PRPP binding (312).

(iii) Aminoglycoside antibiotic biosynthesis. A third reaction that forms anO-glycosidic bond with PRPP as the phosphoribosyl donor occurs in the biosynthesis of theaminoglycoside antibiotic butirosin. The 26-ORF btr gene cluster specifies the biosyntheticpathway for buritosin in Bacillus circulans (314, 315). The btrL cistron encodes neamine

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phosphoribosyltransferase (neamine:5-phospho-�-D-ribose 1-diphosphate phosphoribo-syltransferase; EC 2.4.2.49), which phosphoribosylates neamine with the formation of5�-phosphoribostamycin (Fig. 12C) (316). The B. circulans neamine phosphoribosyltrans-ferase contains a well-conserved PRPP-binding site, 212-Asp-Ile-Val-Ile-Leu-Glu-Asp-Gln-Pro-His-Thr-Gly-Gly-Thr-225, which may be compared to the E. coli hypoxanthinephosphoribosyltransferase sequence, 94-Asp-Val-Leu-Ile-Val-Glu-Asp-Ile-Ile-Asp-Ser-Gly-Asn-Thr-107. Neamine phosphoribosyltransferase contains 604 amino acid residuesand is much larger than type I phosphoribosyltransferases (316), such as E. colihypoxanthine and guanine-xanthine phosphoribosyltransferases, which contain 178and 152 amino acids, respectively. It is also larger than type II phosphoribosyltrans-ferases, such as E. coli quinolinate and nicotinate phosphoribosyltransferases, whichcontain 297 and 400 amino acids, respectively. It is possible that the neamine phos-phoribosyltransferase is a bifunctional enzyme, although a second activity has not beendiscovered. This putative second activity is not the phosphatase that hydrolyzes thephosphate ester of 5�-phosphoribostamycin to ribostamycin, as this activity is encodedby the btrP cistron (316).

In addition to butirosin, a number of other aminoglycoside antibiotics contain apseudodisaccharide resembling that of butirosin as well as a ribosyl moiety. Examplesof these antibiotics are neomycin B, paromomycin, and lividomycin B. The neomycin Bbiosynthesis-specifying gene cluster of Streptomyces fradiae contains the neoL readingframe (accession number CAF33322), whose deduced amino acid sequence shows only22% amino acid sequence identity with that of neamine phosphoribosyltransferase.However, certain stretches of amino acids along the sequences appear highly con-served, among these is the PRPP-binding site (317). Similarly, a putative neaminephosphoribosyltransferase may be present in Streptomyces lividus, because an ORF(accession number CAG38706) of the lividomycin B biosynthetic cluster of S. lividus is29% identical to the B. circulans neamine phosphoribosyltransferase amino acid se-quence. Details on the genetics of aminoglycoside antibiotic biosynthesis have beenpreviously reviewed (318).

Reactions at the Diphosphoryl Moiety of PRPP

A few enzymes have been shown to hydrolyze the phosphoric anhydride bond ofthe diphosphoryl moiety of PRPP: PRPP � H2O ¡ ribosyl 1,5-bisphosphate � Pi. Anumber of archaeal species, such as M. jannaschii, Methanosarcina acetivorans, A.fulgidus, and T. kodakarensis, contain a form III ribulose 1,5-bisphosphate carboxylase/oxygenase, which catalyzes the carboxylation of ribulose 1,5-bisphosphate. Usuallyribulose 1,5-bisphosphate is formed by ATP-dependent phosphorylation of ribulose5-phosphate. However, in M. jannaschii and M. acetivorans, ribulose 1,5-bisphosphateoriginates from PRPP, and it has been suggested that PRPP is hydrolyzed to ribose5-phosphate followed by NAD�-dependent oxidation to ribulose 1,5-bisphosphate.Alternatively, PRPP may be transformed to ribosyl 1,2-cyclic phosphate and then toribulose 1,5-bisphosphate. The physiologic function of this pathway remains to beestablished (319). In addition, certain diphosphoryl (Nudix) hydrolases are able tohydrolyze PRPP with the formation of ribosyl 1,5-bisphosphate and Pi (320). Finally, aPRPP pyrophosphatase activity has been identified in macrophages exposed to hypoxia(321).

Cofactor BiosynthesisThiamine diphosphate. Thiamine diphosphate synthesis utilizes 5=-phosphoribosyl

5-aminoimidazole as a precursor. This compound is also a precursor of purine nucle-otide biosynthesis. 5=-Phosphoribosyl 5-aminoimidazole is converted to a pyrimidinemoiety by thiC-encoded phosphomethylpyrimidine synthase. As such, some of thecarbons of PRPP end up in the pyrimidine moiety of thiamine diphosphate. Thus, C-2of PRPP becomes the methyl group of the pyrimidine moiety, C-4 of PRPP becomes C-5of the pyrimidine moiety, and C-5 of PRPP becomes the methylene group that connects

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the pyrimidine and thiazole moieties of thiamine diphosphate. C-1 and C-3 of PRPP arelost as formate and carbon dioxide (322).

Flavins. The ribitol moiety of the flavin-containing cofactors FAD and FMN arederived from the ribosyl moiety of GTP, and thus originally from PRPP (323, 324).

Pterins. The pterin moiety of folate derivatives are synthesized from GTP. In tetra-hydrofolate, the C-7, C-6, and the methylene attached to C-6 of the pterin moiety arederived from C-1, C-2, and C-3 of PRPP. C-4 and C-5 of PRPP are lost as glyoxylate. Thesynthesis of pterin and methanopterin follow identical pathways at this stage (325,326). Also, the molybdopterin is synthesized from GTP with similar utilization of carbonsof PRPP (327).

Carbon-Phosphorus Lyase Pathway

As mentioned previously, PRPP can be synthesized in a reaction catalyzed byphosphoribosyl bisphosphate phosphokinase. This reaction is part of the carbon-phosphorus lyase pathway, by which phosphonates are catabolized by means of aradical-mediated process. This catabolic pathway is specified by the 14-cistron operonphnCDEFGHIJKLMNOP found in numerous microorganisms, including E. coli (154). Phos-phoribosyl bisphosphate phosphokinase is specified by the phnN cistron and catalyzesthe following reaction: ribosyl 1,5-bisphospate � ATP ¡ PRPP � AMP, which is the finalchemical reaction specified by the phnC-D operon (151, 153). Thus, catabolism ofphosphonates by the carbon-phosphorus lyase pathway produces PRPP, in which thephosphorus of phosphonate is represented by the �-phosphate at the C-1 position. Theutilization of phosphonate-derived phosphorus requires one or more of the above-described phosphoribosyltransferases, which produces PPi, which can by hydrolyzed toPi by diphosphatase (154, 328).

PRPP AS MEDIATOR OF METABOLIC REGULATION

Until now this review has presented enzymes that produce PRPP and enzymes thatconsume PRPP with the formation of N-, O-, or C-glycosidic bonds. The followingsection is dedicated to the regulatory functions played by PRPP. These functionsinclude binding to the PyrR regulator, which is involved in translational control ofpyrimidine biosynthesis gene expression, binding to the PRPP responsive PurR regu-lator of Gram-positive organisms with low GC content, and allosteric activation by PRPPof carbamoylphosphate synthetase.

The Enigma of PRPP as a Regulator of Metabolic Activity

Bacterial enzymes that utilize PRPP generally have Km values at least 1 order ofmagnitude lower than the normal intracellular PRPP concentration. Thus, the PRPPconcentration is unlikely to be a rate-determining factor for phosphoribosyltrans-ferases. No regulatory function, such as allosteric inhibition, of phosphoribosyltrans-ferase activity by PRPP at a metabolic level under normal growth conditions hasbeen identified experimentally. Traditionally, PRPP pool sizes are expressed asmillimoles per gram (dry weight); this unit is dependent on the cell compositionand is not directly translatable into the intracellular molar concentration for com-parisons between different species. However, it appears safe to state that the PRPPconcentration in bacterial cells is in the millimolar range (329, 330). Bacteriaregulate their PRPP pools in response to exogenous purine sources. Thus, the PRPPpool of B. subtilis, L. lactis, E. coli, or S. enterica is reduced 2- to 4-fold by the additionof adenine or hypoxanthine (Table 5). This reduction in PRPP pool size may becaused by allosteric and isosteric regulation of PRPP synthase by purine nucleo-tides, i.e., ADP and GDP, as described above.

The situation is different in mammals, where the PRPP concentration is rate limitingfor purine biosynthesis de novo and for salvage reactions (331), which suggests that therates of PRPP-utilizing enzymes may be determined by substrate saturation kinetics.The PRPP concentration in mammalian cells is much lower than that of bacteria, 5 to30 �M, although there are variations throughout the cell cycle (331, 332).

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We are therefore faced with the enigma of explaining how the high intracellularPRPP levels of microbial organisms are sensed by regulator proteins, which presumablyshould be saturated with PRPP under most physiological conditions. The enigma hasbeen only partly resolved, and further research will be needed to fill in the missinginformation. Details of the mechanism of PRPP-mediated regulation of pyrimidine andpurine nucleotide synthesis by the PyrR and PurR regulatory proteins are described inthe next section.

Regulation of Pyrimidine Metabolism by the RNA-Binding PyrR Protein

Pyrimidine biosynthetic genes are present in most organisms, and their expressionis subject to control by different mechanisms. Regulation of bacterial pyrimidinemetabolism has been extensively reviewed previously (333–335). Here we shall con-centrate on a description of the structure of PyrR and the involvement of PRPP inPyrR-mediated regulation of pyrimidine biosynthesis in B. subtilis, B. caldolyticus, and L.lactis.

Three-dimensional structure of PyrR. The three-dimensional structure of PyrR fromthree microbial sources has been published: B. subtilis (336), B. caldolyticus (337), and M.tuberculosis (338). The three-dimensional structure of T. thermophilus PyrR has also beensolved (PDB code 1ufr). The overall structures of the four PyrR proteins are identical andcontains the typical type I phosphoribosyltransferase fold, a five-stranded �-sheetsurrounded by three helices, and the typical sequence elements are easily identified: aPRPP-binding site, i.e., a PRPP-binding loop (100-Val-Ile-Leu-Val-Asp-Asp-Val-Leu-Phe-Thr-Gly-Arg-Thr-112 of B. caldolyticus PyrR), a PP loop (37-Gly-Ile-Lys-Thr-Arg-41 of B.caldolyticus PyrR), a flexible loop (Thr71 to Asn83 of B. caldolyticus PyrR), and a hoodstructural element. The binding of UMP and GMP to B. caldolyticus PyrR involvesidentical amino acid residues, Asp104, Asp105, Thr109, and Arg137. The 104-Asp-Asp-105 dipeptide of the PRPP-binding loop binds to the nucleotide via Mg2�. Altogether,the binding of PRPP and nucleotides to PyrR is similar to that of type I phosphoribo-syltransferases. PyrR from B. caldolyticus has been proposed to exist as a dimer ortetramer (337, 339, 340), although B. subtilis PyrR has been shown to exist in ahexameric state (341). The dimerization pattern of PyrR is different from that of type Iphosphoribosyltransferases, and the RNA-binding activity of PyrR has been proposed tobe due to the exposure of a region of basic amino acid residues in the dimer. Effectorsof the quaternary structure are PRPP, uridine ribonucleotides, GTP, and pyr leadermRNA. A close resemblance of the crystal structure of GMP-bound B. caldolyticus PyrRto an evolutionarily distant hypoxanthine-guanine phosphoribosyltransferase suggeststhat the two proteins share ancestry (337). However, PyrR from both B. subtilis and B.caldolyticus have a low uracil phosphoribosyltransferase activity that can support slowgrowth of a pyrimidine requiring strain on uracil (248, 342).

Mechanism of PRPP-mediated regulation by PyrR. The B. subtilis pyr gene clusterencodes all of the enzymes required for the biosynthesis of UMP, a uracil transporter,

TABLE 5 PRPP pool sizes in various bacterial species

Bacterial species Purine added PRPP pool sizea Reference

B. subtilis None 1.1 � 0.1 356Adenine 0.6b 356

L. lactis None 2.6 � 0.6 414Hypoxanthine 0.7 � 0.2 414

E. coli None 4.4 415Hypoxanthine 1.2 415

S. enterica None 1.2 36Hypoxanthine 0.3 36

aThe pool size units (means � standard errors) are millimoles per gram dry weight.bNo repression was observed with hypoxanthine for this species.

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and the regulatory protein PyrR (342). Transcription is controlled by pyrimidine avail-ability through an attenuation mechanism. There are three terminator-antiterminatorstructures in pyr mRNA. PyrR binds to a conserved region in the pyr operon leaderresulting in disruption of an antiterminator that is the dominant RNA structure in theabsence of PyrR binding. The negative effect of PyrR is observed only when pyrimidineprecursors are available in the growth medium (341–343).

The active RNA binding form of PyrR appears to be a dimer (Fig. 13A) which iscapable of recognizing and binding a specific RNA structure present in the leadermRNA transcripts of PyrR-regulated genes (340). Dimer formation and RNA bindingoccurs when UMP, UTP, or PRPP is bound to PyrR, resulting in transcriptional termina-tion. GTP counteracts the binding of UTP (and possibly UMP), resulting in multimerformation and presumably transcriptional readthrough because the RNA-binding sitesare sequestered in the tetramer. UMP, UTP, and GTP bind to PyrR at physiologicalconcentrations, while PRPP binding has an apparent dissociation constant at subphysi-ological concentrations, as discussed above (340).

The physiological relevance of PRPP binding to L. lactis PyrR was demonstrated bytranscriptomic analysis of nucleotide metabolism. L. lactis pyr gene expression isregulated by PyrR-dependent attenuation with a mechanism similar to that of B. subtilis(344–346). The mRNA levels under PyrR attenuation control showed an inverse corre-lation with the intracellular PRPP concentrations (347), in accordance with an increased

FIG 13 Mechanism of PRPP-mediated regulation of transcription by PyrR and PurR. PyrR and PurR are shown as orange spheres. (A) Model of PyrRregulation based on the combined data from B. subtilis and B. caldolyticus. RNA (green hairpin) binds to dimeric PyrR, which is stabilized by UMP,UTP, or PRPP. The tetrameric conformation is stabilized by GMP and does not bind RNA. See the text for details. (B) Model of PurRBs repression.DNA (blue line) containing one strong (solid green line) and one weak (green punctuated line) PurBox binds to two PurRBs dimers (forming a weaktetramer) in the absence of PRPP. In the presence of PRPP, the DNA binding is prevented and the tetramerization is lost. (C) Model of PurRLl

activation. PurRLl binds to PurBox sequences, irrespective of PRPP binding, presumably as a dimer. Binding of PRPP is hypothesized to expose abinding site for RNA polymerase. RNA polymerase is positioned correctly relative to the �10 region of the promoter.

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RNA binding and attenuation upon PRPP binding to PyrR. It is reasonable to suspectthat PyrR from L. lactis also binds RNA as a dimer and that dimers are stabilized inrelation to tetramers by binding of PRPP. An increase in PRPP concentration is a conse-quence of purine shortage and hence a diminished demand for pyrimidine synthesis; thus,the inverse correlation of PRPP and pyr gene expression serves a valuable physiologicalfunction.

Proteins with homology to PyrR are found among many bacterial species. With fewexceptions, PyrR is found among Firmicutes, Cyanobacteria, and Actinobacteria. Appar-ently, PyrR is absent from the alphaproteobacteria, but it is present in some orders ofthe other classes of proteobacteria. PyrR is absent from Enterobacteriales but present inthe Pseudomonadales order. B. subtilis has two different enzymatic activities thatcatalyze the formation of UMP from uracil and PRPP, PyrR and uracil phosphoribosyl-transferase, encoded by upp (248). PyrR and uracil phosphoribosyltransferase share 23%identity, which suggests that PyrR, aside from its close ancestry to the hypoxanthine-guanine phosphoribosyltransferases, also shares ancestry with uracil phosphoribosyl-transferases.

Regulation of Purine Metabolism by PRPP-Responsive, DNA-Binding PurRProteins

PRPP-responsive PurR transcriptional regulators are found only in low-GC Gram-positive bacteria, but the members of this small protein family differ widely in theirmechanism of regulation. The B. subtilis PurR, here designated PurRBs, is a negativeregulator, whereas the L. lactis PurR, here designated PurRLl, is an activator of purinemetabolism gene expression. In contrast to the PRPP-responsive PurR regulators fromGram-positive bacteria, a LacI-type PurR repressor protein is present in E. coli and avariety of other bacterial species. The LacI-type PurR proteins do not respond to PRPP,but they repress the transcription of purine genes when bound to the corepressorshypoxanthine or guanine (348, 349). In the following sections, we shall describe in moredetail the structure and function of B. subtilis and L. lactis PurR.

Three-dimensional structure of B. subtilis PurR. B. subtilis PurR contains 285 aminoacid residues and has a homodimeric quaternary structure (350). A C-terminal domainof the subunit (residues 77 to 285) has the characteristic fold of type I phosphoribo-syltransferases and PRPP synthases described previously, a central parallel � sheetflanked by �-helices. Hood and flexible loop structural elements are also present. TheC-terminal domain of PurRBs also contains a PRPP-binding site, i.e., a PRPP-binding loop(196-Gly-Ser-Asn-Val-Leu-Ile-Ile-Asp-Asp-Phe-Met-Lys-Ala-Gly-Gly-Thr-211) and a PPloop (139-Thr-Lys-Gly-Ile-142) (351).

Gel mobility shift analysis showed that binding of PRPP to PurRBs changes its affinitytoward its DNA-binding sites (350, 352). X-ray structural analysis of PurRBs crystalscontaining the PRPP analog cPRPP revealed that the compound binds to the PRPP andPP loops analogous to PRPP binding in type 1 phosphoribosyltransferases. Contactingamino acid residues were Lys140 and Asp203/Asp204 of the PP- and PRPP-bindingloop, respectively, as well as residues in the flexible loop (353). Curiously, the bindingof cPRPP occurred without the participation of Mg2�, which may suggest relativelyweak binding of cPRPP. DNA binding of two mutant variants of PurRBs, Asp203Ala andAsp204Ala, could not be inhibited by PRPP in vitro, and the alterations caused asuperrepression phenotype of pur gene expression consistent with poor binding ofPRPP to the altered PurRBs proteins (352). Altogether, binding of PRPP to PurRBs stronglyresembles the binding of PRPP to type I phosphoribosyltransferases.

In spite of the high sequence similarity to type I phosphoribosyltransferases, PurRBs

is inactive as a phosphoribosyltransferase and does not even bind purine bases. Thisdeficiency is explained by the interactions of the large side chains of well-conservedTyr102 and Phe205, which prevent the binding of a nucleophile in proper positionrelative to C-1 of PRPP. The presence of these bulky amino acid side chains also explainsthe inability of compounds containing nucleobases to inhibit PurRBs-PurBox interac-tions (351). Interestingly, xanthine phosphoribosyltransferases contain phenylalanine in

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the analogous position to Phe205, which indicates that the Phe205-Tyr102 interactionis crucial to blocking access of purines to the active site of PurR.

The N-terminal domain (residues 1 to 74) of PurRBs forms a winged-helix domain, asubdivision of the DNA binding helix-turn-helix domains (351), and is attached to theC-terminal domain by a three-residue linker. This domain is responsible for the bindingof PurRBs to so-called PurBox sites located upstream of the PurR-regulated operons(353). Functionally, PurRBs may be regarded as a crippled phosphoribosyltransferasewhich has lost the catalytic activity but retained the PRPP-binding capability and hasgained an N-terminal DNA-binding domain.

Mechanism of PRPP-mediated regulation of purine biosynthesis in B. subtilis. Theaddition of adenine to cultures of B. subtilis causes repression of transcription of a largenumber of operons, including genes for purine nucleotide de novo biosynthetic en-zymes, genes specifying purine transport proteins, genes for enzymes in purine nucle-otide interconversion, and certain cofactor biosynthetic genes (350, 354, 355). Similar tothe situation in enteric organisms (146), the addition of adenine to cultures of B. subtiliscauses depletion of the PRPP pool (356). Induction of expression of PurR-regulatedgenes by PRPP binding has been evaluated in PurRBs-DNA-binding assays in vitro (350).PurRBs binds to a region 20 to 150 bp upstream of the B. subtilis pur operon (357). ThisDNA region contains the operator region �81AAACACGAACATTA (PurBox1)-16 nucleotides-TATCGTTCGATAAT�38 (PurBox2), numbered relative to the transcription start site. Pur-Box1 and PurBox2 have similarity to the consensus PurBox (AWWWCCGAACWWT),although PurBox2 is inverted (354, 358). Functional dissection has defined a minimalcontrol region and revealed the importance of the two PurBoxes in the overallregulatory process. Based on the affinity of PurRBs for the PurBoxes, PurBox1 wasdesignated “strong” and PurBox2 was designated “weak.” High-affinity binding of PurRto the control region was shown to require at least one strong PurBox, whereas theinduction of PurRBs repression by PRPP requires at least one weak PurBox. The bindingof PurRBs to the operator is cooperative with a stoichiometry of two PurRBs dimersbinding to the operator, which suggests that one PurRBs dimer binds to PurBox1 andanother PurRBs dimer to PurBox2, with wrapping of the DNA around the two PurRBs

dimers (Fig. 13B) (353).In B. subtilis a guanine-responsive attenuation mechanism is superimposed on the

repression of transcription initiation by the PurR regulatory system, which results in acomplex regulatory pattern. Thus, the addition of guanine to cultures of B. subtiliscauses premature transcription termination in the mRNA leader region of purE (theupstream cistron of the major 12-cistron pur operon) (359). This premature transcriptiontermination is caused by the purine riboswitch mechanism, which regulates the pur,xpt, pbuG, and yxjA operons (360). This riboswitch mechanism involves the binding ofguanine or hypoxanthine to the untranslated leader sequences but does not involvePRPP. Details of the mechanisms of riboswitches have been previously reviewed (361,362).

Mechanism of PRPP-mediated regulation of purine biosynthesis in L. lactis.Overall, the effect of purine addition has the same effect on L. lactis as that on B. subtilis:the PRPP level drops and the expression levels of genes of the PurR regulon are lowered(363). However, a different mechanism of action became apparent when the L. lactispurR gene was identified among purine auxotrophic strains obtained after transposonmutagenesis. The purR insertion mutation caused reduced transcription of purD, whichwas complemented by purR�. These and other data showed that L. lactis purR encodesan activator of purine gene expression. PurRLl contains 271 amino acid residues, closeto the length of PurRBs (285 amino acids). Remarkably, PurRLl and PurRBs are 48%identical (69% similar), and yet their mechanisms are very different. Like PurRBs, PurRLl

contains a helix-turn-helix domain at the N-terminal end. It also contains the typicaltype I phosphoribosyltransferase structural elements, a PRPP-binding site, i.e., a PRPP-binding loop (194-Gly-Gln-Asn-Val-Leu-Ile-Val-Asp-Asp-Phe-Met-Lys-Gly-Ala-Gly-Thr-209), a PP loop (137-Thr-Lys-Gly-Ile-140), and a flexible loop (358).

PurRLl recognizes PurBox sequences similar to the consensus PurBox sequence,

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AWWWCCGAACWWT (364). PurRLl binds to PurBoxes under both activating (high PRPP)and nonactivating conditions. Activation of transcription requires that the PurBoxsequence is located at an exact position relative to the �10 promoter region, presum-ably in order to bind and position RNA polymerase correctly for open complexformation (358) (Fig. 13C). The standard �35 promoter elements of constitutive pro-moters are usually absent from PurR-activated promoters. In L. lactis, different topolo-gies of PurBoxes have been identified in which the binding sites are located in tandemor head to head, which suggests that PurR may form different multimeric conforma-tions (363).

In vivo kinetics of the PRPP-modulated activation of transcription by PurRLl wasstudied by a combination of temporal mRNA and intracellular PRPP quantificationsduring a metabolic downshift (347). Saturation curves for the coupled process of PRPPbinding to PurRLl and subsequent activation of mRNA synthesis were obtained for allgenes belonging to the PurR regulon in L. lactis. Interestingly, most saturation curvesappeared sigmoid, suggesting mechanisms employing cooperativity, but more classicalMichaelis-Menten curve forms were apparent for some PurR-activated genes involvedin purine salvage and folate metabolism (347). To our knowledge, this work representsthe first example of a kinetic analysis of the action of a regulatory protein performedin a living organism.

Evolution of PurR. PurR proteins from various organisms share extensive sequencesimilarity, but the differences in regulatory mechanisms between PurRBs and PurRLl

show that they have diverged considerably during evolution. Figure 14 shows thephylogenetic relationship among members of the PurR family. The branching patternof the PurR phylogeny resembles the branching pattern for the whole-genome bacte-rial phylogeny (365), indicating that the PurR protein was present during most of theevolution of the low-GC branch of Gram-positive bacteria. The Bacillus branch and theStreptococcus branch share an ancestor, which is not shared by any other species. It isapparent that the bacilli and streptococci use the B. subtilis type PurBox, while all theothers use the L. lactis type PurBox (Fig. 14A), except the clostridia, for which noinformation is available. According to the unrooted phylogenetic tree, it is likely thatthe activating PurRLl type evolved first and that the PurRBs type diverged early in theevolution of the low-GC Gram-positive bacteria. As a consequence, the L. lactis PurBoxmust simultaneously have evolved into the B. subtilis PurBox. When the sequences inthe logo plots for the two PurBox types are compared (Fig. 14B), it is easily recognizedthat the B. subtilis PurBox consists of two L. lactis type PurBox sequences located asinverted repeats with a spacing of 16 nucleotides, as previously described (354).

To further analyze the relationship of PurR with enzymes containing a type Iphosphoribosyltransferase fold, a phylogenetic analysis was performed for homologs ofB. subtilis class 1 phosphoribosyltransferases with all the major bacterial lineages,spanning an evolution of at least 3 billion years. Included in the analysis are also logoplots of conserved amino acids in the PP loop and the PRPP-binding loop (Fig. 15). Itis apparent that PurR and xanthine phosphoribosyltransferase are close relatives (366),that PyrR and hypoxanthine-guanine phosphoribosyltransferases are also closely re-lated (337), and that they share ancestry with uracil phosphoribosyltransferases (336).As noted in Fig. 15, PurR and xanthine phosphoribosyltransferase homologs are onlyfound in the low-GC Gram-positive bacterial lineage. PurR and adenine phosphoribo-syltransferase has previously been inferred to be structurally related, based upon thecommon structure of the flexible loop, which forms a two-stranded antiparallel�-ribbon structure (210). A similar structure is also present in the flexible loop of B.subtilis xanthine phosphoribosyltransferase (366). Thus, PurR, xanthine phosphoribosyl-transferase, and adenine phosphoribosyltransferase are all structurally related.

If PurR arose by evolution from a xanthine phosphoribosyltransferase, perhaps itcould have exploited some of the peculiarities of this enzyme. Of special interest is thefact that the active form of xanthine phosphoribosyltransferase is a dimer and thatbinding of PRPP is necessary for stabilization of the dimeric conformation (366). Mostother type I phosphoribosyltransferases have active tetrameric rather than dimeric

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structures. Dimer formation could be important for PRPP modulation of PurRLl as it isfor PurRBs, perhaps by exposing interaction sites needed for RNA polymerase capture.

Search for a common mechanism for PRPP modulation of PurR conformation.PurR is a family of proteins that share extensive similarity but which are divided into atleast two phylogenetic branches with different PurR functionalities, a B. subtilis-likebranch and an L. lactis-like branch. Members of both branches appear to have rigidDNA-binding domains with hydrophobic cores that are not readily modulated by PRPPbinding, and both appear to harbor extensive positively charged domains that maybind the DNA backbone and result in protection of large DNA stretches. Yet, regulatorsfrom one family represented by PurRLl appear to bind to single-PurBox motifs withunknown stoichiometry. Presumably, the binding of PRPP and closure of the flexibleloop expose a binding site that is used to recruit RNA polymerase. Regulators from the

FIG 14 Phylogenetic analysis of PurR from low-GC Gram-positive bacteria. (A) PurR sequences from representatives of the major bacterial lineages in the low-GCGram-positive bacteria were aligned using the Clustal Omega program, and a phylogenetic tree was constructed using the ClustalW2 phylogeny program. (B)The type of PurBox used by each species was identified at the RegPrecise website (http://regprecise.lbl.gov/RegPrecise/collection_tffam.jsp?tffamily_id�53),and the two types of logo plots were constructed from the PurBox sequences presented for Streptococcaceae and Bacillales, respectively, using the weblogoservice (http://weblogo.berkeley.edu/logo.cgi).

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second family, represented by PurRBs, recognize two PurBoxes in a stoichiometry of onePurR dimer per PurBox and in a conformation that involves formation of PurR tetramers(Fig. 13B and C). Occupancy of the PRPP-binding site reduces unspecific DNA bindingand prevents PurR repression of transcription initiation. It is likely that the closing of theflexible loop upon PRPP binding, which is also shared by all type I phosphoribosyl-transferases, is the mechanistic basis for both effects. There is currently no evidencethat all PurR regulators in one family share a mechanism, and it is not clear whetherPurRLl and PurRBs may substitute for one another. However, B. subtilis-like tandemPurBoxes are found only in the Bacillus and Staphylococcus branches and singlePurBoxes are found in the Lactococcus, Streptococcus, and Lactobacillus branches. Asearch for evolutionary “crossover” organisms with PurR proteins that possess bothrepression and activation abilities may be very useful in this respect.

Allosteric Activation of Carbamoylphosphate Synthetase Activity by PRPP

Carbamoyl phosphate is a precursor for the synthesis of pyrimidine nucleotides andthe amino acid arginine. While bacterial species like E. coli and S. enterica contain onlya single carbamoyl phosphate synthetase, other bacterial species, like B. subtilis, containspecific enzymes for each pathway, pyrimidine nucleotides and arginine. The activity ofpyrimidine nucleotide-specific enzyme was found to be stimulated by PRPP (367, 368).

In animals and fungi, two or three of the six enzymes of de novo pyrimidinenucleotide biosynthesis are grouped in di- or trifunctional enzymes. Animals containthe trifunctional fusion of carbamoylphosphate synthase, aspartate transcarbamoylase,and dihydroororase activities, called CAD. CAD thus catalyzes the ATP-dependentconversion of glutamine, bicarbonate, and aspartate to dihydroorotate. The function

FIG 15 Common phylogeny of PyrR, PurR, type 1 phosphoribosyltransferases, and class I PRPP synthasesin all major bacterial lineages. Homologous protein sequences were deduced and identified fromrepresentative genome sequences of all major bacterial lineages for the following proteins: adeninephosphoribosyltransferase (APRTase), hypoxanthine-guanine phosphoribosyltransferase (HPRTase), oro-tate phosphoribosyltransferase (OPRTase), uracil phosphoribosyltransferase (UPRTase), and xanthinephosphoribosyltransferase (XPRTase). The collected sequences were aligned using the Clustal Omegaprogram, and a phylogenetic tree was constructed using the ClustalW2 phylogeny program. (Left)Phylogenetic tree, with protein sequences belonging to same enzyme families grouped together astriangles, starting at the point of the first branching of the individual protein sequences and labeled withthe enzyme abbreviations shown above. The triangle marked OPRTase* is a group of orotate phospho-ribosyltransferase sequences that appear to form a separate branch with a distinct set of amino acidsequences in the PP- and PRPP-binding loops. Sequences with homology to the xanthine phosphori-bosyltransferase and PurR proteins were only found among the low-GC Gram-positive bacteria, asindicated. (Middle) Logo plot of the frequency of amino acids in the PP loop identified in the ClustalOmega alignment and constructed using the weblogo service (http://weblogo.berkeley.edu/logo.cgi).(Right) Logo plot of the frequency of amino acids in the PRPP-binding loop identified in the ClustalOmega alignment and constructed using the weblogo service.

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and structure of CAD have been previously reviewed (369, 370). Dihydroorotate in turn isoxidized to orotate, which is then phosphoribosylated to orotidine 5=-monophosphate, areaction catalyzed by orotate phosphoribosyltransferase. In the present context, theimportant activity of CAD is carbamoyl phosphate synthase. The activity of this enzymeis regulated by UTP and PRPP. UTP is a feedback inhibitor, whereas PRPP is an allostericactivator of the enzyme, as found for the B. subtilis pyrimidine-specific carbamoylphosphate synthetase (368, 371). CAD activity is modulated further by protein phos-phorylation mediated by the mitogen-activated protein kinase (372), cAMP-dependentprotein kinase (372), and mTORC1 (373). The expression of the cad genes is also subjectto regulation. Upon transition from G1 to S phase, the human cad mRNA level increasedapproximately 10-fold, which is mediated by the global regulator c-MYC (373, 374).

prs GENES, MUTANTS, AND GENE REGULATION

PRPP synthase is believed to be present in all free-living organisms. A number of prsmutants with alterations in the enzyme have been isolated and characterized. Inaddition, a few knockout mutant alleles have been constructed in vitro and, followingrecombination, gene conversions have yielded null alleles. Also, mutant variants ofPRPP synthase have been characterized in a wide variety of organisms, includinghumans. We review here the properties of these mutant variants of bacterial origin.

E. coli and S. entericaE. coli prs-1 and prs-2. A number of prs mutants have been isolated in E. coli. First,

the prs-1 allele appeared in a genetic selection designed to isolate mutants withimproved utilization of guanosine as a purine source (28). Briefly, five mutants werecharacterized; all of them had a lesion in the gsk gene as well as a lesion in a genespecifying proteins of an early part of the purine nucleotide biosynthesis de novopathway, i.e., mutants with the genotype purF gsk and purM gsk. The appearance of theprs-1 gsk mutant strain may be explained along this line by extending the purinenucleotide biosynthesis de novo pathway to include prs in addition to purF, purD, purL,etc. The prs-1 allele specifies Asp128Ala. This residue is only two residues away fromHis130, equivalent to His135 of B. subtilis PRPP synthase. As described before, His135 isimportant for the coordination of the MG2 site, i.e., maintaining the �,�,�-tridentatecomplex of MgATP. Consistent with this, PRPP synthase specified by prs-1 had a 27-foldincrease in the Km for ATP, whereas the Km for ribose 5-phosphate was essentiallyunaltered (375).

A temperature-labile PRPP synthase, specified by the prs-2 allele, was isolated bylocalized mutagenesis of the hemA region (2). The prs-2 allele contained two mutations,one responsible for a Gly8Ser alteration, the other a cytidylate-to-thymidylate transitionthat results in an alteration close to the Shine-Dalgarno sequence. The latter mutationresulted in increased synthesis of PRPP synthase and thus in part compensated for apoorly functioning enzyme. The Gly8Ser alteration was responsible for the temperature-labile PRPP synthase (376).

In vitro-generated knockout prs alleles. A prs knockout mutation would render thestrain prototrophic for purine, pyrimidine, and pyridine nucleotides as well as histidineand tryptophan. Any scheme to isolate such a strain must include the followingconsiderations. NAD, or its metabolite, nicotinamide mononucleotide, may be utilizedintact without catabolism to satisfy the pyridine nucleotide requirement (377). Therequirement of PRPP for tryptophan and histidine synthesis can be eliminated by theaddition of these two amino acids to the growth medium. In contrast, PRPP is normallyessential for both de novo and salvage purine and pyrimidine nucleotide reactions.PRPP consumption in de novo synthesis of purine and pyrimidine nucleotides ismediated by purF-specified amidophosphoribosyltransferase and pyrE-specified orotatephosphoribosyltransferase, whereas the salvage reactions of purine and pyrimidinenucleotides are mediated by adenine, hypoxanthine, xanthine, guanine, and uracilphosphoribosyltransferases. Some organisms, such as E. coli and S. enterica, also containnucleoside kinases, which phosphorylate nucleosides, such as inosine, guanosine, and

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uridine to IMP, GMP, and UMP, respectively. Uptake of these nucleosides via thesekinase reactions would eliminate the requirement for PRPP. However, nucleosides arepredominantly phosphorolyzed by nucleoside phosphorylases, such as purine nucleo-side phosphorylase I (deoD) with specificity for adenosine, inosine, and guanosine,purine nucleoside phosphorylase II (xapA) with specificity for inosine, xanthosine andguanosine, and uridine phosphorylase (udp) with specificity for uridine (19). Thus,knockout of deoD and udp is necessary to permit the salvage of nucleosides byphosphorylation of guanosine by guanosine kinase (gsk) and of uridine by uridinekinase (udk). With respect to guanosine utilization, yet another genetic lesion wasnecessary to permit isolation of a �prs strain. This lesion, for example gsk-3, improvedthe activity of guanosine kinase by rendering the enzyme insensitive to feedbackinhibition by GTP and presumably GDP (378; Hove-Jensen, unpublished data). Alto-gether, a background strain, a deoD udp gsk-3 mutant strain, was used for the successfulrecombination of the in vitro-generated, plasmid-borne �prs allele into the chromo-some of E. coli. (The growth medium included guanosine, uridine, histidine, tryptophan,and NAD.) Two alleles, �prs-3::Kanr and �prs-4::Kanr, were recombined into the chro-mosome (3). Also, the temperature sensitivity of the prs-2 allele could be rescued in thedeoD udp gsk-3 genetic background (2).

dnaR. PRPP synthase has been shown to be involved in DNA replication, but it isunclear whether the effect is direct or indirect. A genetic lesion of E. coli, dnaR130, wasshown to be an allele of the prs gene, here designated prs-130. A strain harboringprs-130 was temperature sensitive for initiation of DNA replication, whereas elongationwas unaffected. Only replication initiating at the chromosome-borne chromosomalorigin of replication, oriC, was affected by prs-130. Thus, replication of an oriC-harboringplasmid, stable DNA synthesis, i.e., replication independent of RNase HI (specified byrnhA), and replication of bacteriophage � were all unaffected by prs-130. The fact thatbacteriophage � DNA replication was unaffected by temperature and that identicalbacteriophage � burst sizes of prs-130 and prs� strains were observed prompted theresearcher to suggest that the temperature-sensitive DNA replication of the prs-130strain was not caused by lack of precursors for nucleic acids or proteins. Also, thedecline in PRPP synthase activity specified by prs-130 paralleled the decline in DNAreplication upon temperature shift. The prs-130 lesion is located within a 274-bpHindIII-AvaII DNA fragment of the prs coding sequence. The entire prs-130 allele hasbeen sequenced, but the sequence cannot be found in the available databases (379,380).

A number of mutant alleles of genes other than prs were able to suppress thetemperature sensitivity caused by the prs-130 allele. These extragenic suppressors ofprs-130 include dnaA (381), certain rpoB alleles specifying rifampin-resistant variants ofthe �-subunit of RNA polymerase (382), and rpe, specifying D-ribulose 5-phosphate3-epimerase. Suppression of prs-130 by mutations in the dnaA gene prompted thestudy author to suggest a direct physical interaction of DnaA and PRPP synthase,whereas the appearance of prs-130 suppressors in RNA polymerase suggested involve-ment of PRPP synthase in a transcriptional event in DNA replication at oriC. Remarkably,D-ribulose 5-phosphate 3-epimerase (specified by the rpe gene) also suppressed thetemperature-sensitive phenotype of the prs-130 strain (383). Ribulose phosphate epimerasecatalyzes the interconversion of ribulose 5-phosphate and xylulose 5-phosphate, and theenzyme is part of the nonoxidative pentose-phosphate pathway (384), which involvesribose 5-phosphate, the precursor of PRPP. This fact leads us to ask whether knockoutof rpe would result in an increase of ribose 5-phosphate isomerase activity? This activitycatalyzes the interconversion of ribulose 5-phosphate and ribose 5-phosphate, whichmight lead to improved PRPP synthesis.

Altogether, these various observations demonstrate an involvement of PRPP syn-thase in some aspects of DNA synthesis in E. coli, but the detailed biochemistry of theseeffects remain unclear.

ssrA. The ssrA gene specifies the SsrA RNA, which together with the SmpB proteinis involved in rescuing stalled ribosomes. SsrA RNA functions both as a tRNA and as an

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mRNA molecule; SsrA RNA function results in the synthesis of an Ssr polypeptide fusionthat is destined for degradation and the liberation of the stalled ribosome (385, 386).Two temperature-sensitive mutants were isolated in an �ssrA genetic background. Bothmutations mapped within the prs gene, one of which specified Cys215Tyr. Thetemperature-sensitive phenotype was suppressed by reinsertion of a wild-type ssrAgene (387). Similar mutants were isolated involving the thyA gene (388), and thus therestoration of the activity of SsrA is not restricted to misexpression of the prs gene (386).

However, using SmpB as bait in pulldown assays, SsrA RNA and a number ofpolypeptides were copurified. One of these polypeptides was PRPP synthase. Half-maximal binding of PRPP synthase to SsrA RNA was obtained at 1 to 2 �M PRPPsynthase, which was compared to half-maximal binding of ribosomal protein S1 to SsrARNA at 0.030 �M, which indicates relatively poor binding of PRPP synthase to SsrA RNA.The binding of PRPP synthase is therefore relatively unspecific, so that physiologicallysignificant binding of PRPP synthase to an SsrA RNA particle, if it occurs at all, mayrequire additional protein factors (389).

S. enterica prs-100. The prs-100 allele, previously designated prsB (390), was isolatedby combined screening of ethyl methanesulfonate-mutagenized cells for temperature-sensitive growth and lack of PRPP synthase activity (390). The prs-100 lesion causes anArg78Cys alteration and is located within a stretch of amino acid residues that areinvolved in the formation of the bent dimer (Table 3, amino acid residues Leu76 andIle77). This finding supported the conclusion of the authors, “that the mutation altersthe enzyme’s kinetic properties through substantial structural alterations rather than byspecific perturbation of substrate binding or catalysis” (27).

Thiamine biosynthesis. A genetic screen for mutants of S. enterica with improvedbiosynthesis of thiamine was conducted in a strain with purF and gnd lesions, i.e., astrain lacking amidophosphoribosyltransferase and glucose 6-phosphate dehydroge-nase activities. Normally, amidophosphoribosyltransferase (specified by purF) is essen-tial for thiamine diphosphate synthesis. Six thiamine-prototrophic mutants arose at alow frequency (approximately 4 10�8). All of the mutants had decreased activity ofPRPP synthase, which presumably resulted in an increase in the ribose 5-phosphatepool and thus stimulation of the amidophosphoribosyltransferase-independent, spon-taneous (i.e., nonenzymatic) amination of ribose 5-phosphate to 5-phosphoribosyl1-amine and subsequent thiamine diphosphate synthesis. One mutation mappedwithin the Shine-Dalgarno sequence, whereas the remaining five mapped within theprs coding region. The in vitro PRPP synthase activity of these latter mutants was 20 to60% of the wild-type activity. Interestingly, one PRPP synthase variant, Asp224Ala, i.e.,an alteration within the PRPP-binding loop, retained approximately 30% of the wild-type PRPP synthase activity. As pointed out by the study authors, the selectionprocedure yielded only mutants with residual activity of PRPP synthase. Because themutations were acquired by positive selection (growth in the absence of thiamine) andall of the mutations negatively affected the activity of PRPP synthase, this procedurecould be valuable in a screen for mutations that test the importance of various aminoacid residues in PRPP synthase activity (391).

prs Gene Organization and Regulation of prs Gene ExpressionE. coli and S. enterica. The prs gene is a member of the purine regulon, because

prs gene expression is under the control of the purR gene product (a LacI typeprotein). The PurBox sequence of the prs gene is identical and located at identicalpositions in E. coli and S. enterica: �56AAGAAAACGTTTTCGC�41 (with the firstcytidylate residue of the transcript numbered �1). Four nucleotides separate thePurBox and the �35 region. PurR caused 3-fold repression of prs gene expressionin vivo in experiments that measured expression of prs-lacZ operon fusions with orwithout the presence of purR. Furthermore, electrophoretic mobility shift analysis ofPurR binding to DNA containing the prs PurBox demonstrated that PurR bound tothe DNA. Finally, protection of the prs PurBox by PurR was demonstrated by DNaseI footprinting (392). The synthesis of PRPP synthase is derepressed approximately

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10-fold during pyrimidine starvation (393), and uridine derivatives specifically causerepression of PRPP synthase synthesis (394).

The genetic organization of the prs region in E. coli and S. enterica is identical. Theprs gene is located downstream of the lolB gene (encoding an outer membranelipoprotein required for localization of lipoproteins) (395) and ispE (encoding4-diphosphocytidyl 2-C-methylerythritol kinase) (396). The prs gene is transcribed fromtwo promoters. A larger transcript is initiated two genes upstream of prs and compriseslolB-ispE-prs. A smaller transcript initiates within the ispE coding sequence and containsthe last 152 nucleotides of the ispE coding sequence in both organisms. The transcrip-tion start site of the smaller transcript and a rho-independent termination site down-stream of the prs coding sequence have been mapped (45, 397). The smaller transcriptis 20-fold more abundant than the larger transcript. Pyrimidine-mediated prs generegulation occurs via increased amounts of the smaller transcript (397). The codingregion of the prs gene is preceded by a leader sequence 302 nucleotides long in E. coliand 417 nucleotides long in S. enterica (45, 397, 398). Comparison of the two leadersequences revealed 149 almost-identical nucleotides of the 5= end of the transcripts ofthe two organisms, followed by 115 nucleotides in the S. enterica prs leader that are notpresent in the E. coli prs leader and 152 nucleotides which are almost identical in theleaders of the two transcripts, i.e., an a-b-c arrangement in S. enterica and an a-carrangement in E. coli.

Altogether, the expression of the prs genes of E. coli and S. enterica is controlled byboth purine and pyrimidine compounds. Purine-mediated regulation occurs by meansof the PurR repressor binding to the PurBox located within the upstream ispE gene.Several lines of experimental results have provided evidence for the involvement of theprs leader in regulation of the synthesis of PRPP synthase. First, two prs-galK operonfusions were analyzed for production of galactokinase activity. The first gene fusioncontained the prs promoter as well as 24 of the 5= nucleotides of the prs transcript; thesecond contained the prs promoter, the entire prs leader, and 13 nucleotides of the prscoding sequence. Cells containing gene fusion one had 5 times as much galactokinaseactivity as cells containing gene fusion two. Thus, nucleotides of the prs leader containa structure(s) that reduces the production of galactokinase of the gene fusion, andpresumably of PRPP synthase under native conditions (399). In another study, a mutantvariant of RNA polymerase (rpoBC) of S. enterica that was selected as resistant topyrimidine fluoroanalogs was constitutive in expression of the pyrE gene specifyingorotate phosphoribosyltransferase. The activity of orotate phosphoribosyltransferasewas elevated 40-fold, and the activity of PRPP synthase was also elevated, but onlyapproximately 2-fold (400). The defective RNA polymerase was shown to increase thecoupling of transcription and translation at the pyrE attenuator (401). It is possible,therefore, that the increase in prs gene expression is also caused by increased couplingof transcription and translation within the prs leader. In other experiments, whole-transcriptome shotgun sequencing has revealed a short transcript originating from theE. coli prs promoter. Although the ends of this transcript have not been mapped, theanalysis provides additional evidence for the existence of a prematurely terminatedprs transcript (M. A. Sørensen, University of Copenhagen, personal communication).Indeed, a number of potential leaders may be formed from the prs leader, and quitea number of secondary structures, including hairpins and loops, are predicted fromthe nucleotide sequence. One possible secondary structure of the S. enterica prsleader is shown in Fig. 16. Unlike the pyrE attenuator, which is followed by eighturidylate residues and has been shown to be regulated by premature transcriptiontermination in the leader sequence (402), the putative E. coli prs leader stem-loopsare followed by none or a few uridylate residues. This may explain the difference inchanges in pyrimidine-mediated regulation of the two operons: 40-fold in pyrE geneexpression and 2.3-fold in prs gene expression.

Gram-positive organisms. Genetic mapping and nucleotide sequencing has shownthat the prs gene of B. subtilis is located between the gcaD (tms-26) and ctc genes (43,403). The former gene encodes the bifunctional enzyme �-D-glucosamine 1-phosphate

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acetyltransferase, N-acetylglucosamine 1-phosphate uridylyltransferase (404), whereasthe latter gene encodes a general stress protein similar to E. coli ribosomal protein L25(405). In vegetative cells of B. subtilis, gcaD, prs, and ctc are expressed as a tricistronicoperon from a promoter immediately upstream of gcaD (406). Similar arrangements ofgcaD-prs-cts are found in other Gram-positive organisms, Clostridiaceae, Corynebacteri-aceae, Listeriaceae, and Staphylococcaceae species. On the other hand, Streptococcaceaesuch as Streptococcus pyogenes, Enterococcus faecalis, and L. lactis have other moreheterogeneous genetic organization patterns for prs genes. Streptococcaceae generallycontain two prs genes, as described previously for L. lactis. In contrast to the situationin enterobacteria, the regulation of prs gene expression in Gram-positive organisms isessentially unknown.

FUTURE PERSPECTIVES

Although extensive efforts have been spent on elucidation of the various aspects ofPRPP metabolism, a number of important aspects remain to be disclosed. Perhaps most

FIG 16 Possible secondary structure of the S. enterica prs leader. The sequence ranges from nucleotide1, indicated by 5=, to nucleotide 417, which is immediately followed by the initiator codon-specifyingguanylate-uridylate-guanylate triplet and is indicated by 3= (45). The �G of the structure is �540 kJmol�1. The leader prs sequences of S. enterica and E. coli are identical except for the framed sequence,which is present only in the S. enterica prs leader. The nucleotides circled in red represent a possibleleader peptide, and the red arrows point to the nucleotides of a possible Shine-Dalgarno sequence.Possible leader peptide amino sequences are indicated for E. coli and S. enterica. Identical amino acidresidues of the two possible leader peptides are underlined.

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important is the function or functions of the NHR found in many eukaryotic PRPPsynthases and the functions of the PAPs in human and rat PRPP synthases. For example,purification of PRPP synthase from crude extracts of wild-type S. cerevisiae should bepossible and allow elucidation of the quaternary structure of the native enzyme orenzymes. Additionally, the fact that S. cerevisiae PRPP synthase is a multimeric enzymeconsisting of different subunits, such as Prs1 and Prs3 and possibly higher-orderstructures, makes possible studies of mutants with alterations of various amino acidresidues in one subunit while the other subunit(s) retains the wild-type phenotype.Candidates for such an analysis are the His135, Asp174, and Lys197 residues (aminoacid numbers refer to B. subtilis PRPP synthase positons). Similar experiments may bepossible with mammalian PRPP synthase(s). As mentioned previously, there is compel-ling evidence for the involvement of yeast PRPP synthase polypeptides in a variety ofcellular processes other than amino acid and nucleotide syntheses, and there is someevidence that NHRs play roles in this involvement. PAPs may also be involved in cellularprocesses other than nucleotide synthesis, and studies in mammalian cell lines mightreveal roles of the PAPs in other such physiological processes and cross-regulation.Studies of mouse cell lines (and live animals) with the PAPs knocked out, individuallyor together, could be of very great interest in this respect. As described above, a specificrole for human PRPP synthase isozyme 2 in cancer cells has been documented (125).Although PAPs were not implicated, they could very well participate in mediating theobserved effects.

Another aspect of interest is the three-dimensional structure of class II PRPPsynthases. The available data on their quaternary structure are conflicting. On onehand, circular dichroism and gel filtration analyses predict the structure of sugarcanePRPP synthase to be hexameric, similar to that of B. subtilis PRPP synthase (129),whereas recombinant spinach PRPP synthase isozymes 3 and 4 eluted as trimers in gelfiltration chromatography (47).

Furthermore, the three-dimensional structures of PRPP-utilizing enzymes other thanthose for purine, pyrimidine, pyridine, histidine, and tryptophan biosynthesis, i.e.,enzymes catalyzing the formation of O- and C-glycosidic bonds, are worth solving, asthey may contribute additional information regarding the versatility of PRPP in inter-mediary metabolism.

Regulation of PRPP synthase activity in vivo is presently poorly understood. In vitro,the activity is intricately regulated by the energy charge, with ATP being a substrate,AMP a product, and ADP a potent negative, allosteric, and isosteric (i.e., competitive atthe ATP site) regulator of PRPP synthase activity. Understanding how these propertiescontribute to physiological in vivo regulation requires further investigation.

SUPPLEMENTAL MATERIAL

Supplemental material for this article may be found at https://doi.org/10.1128/MMBR.00040-16.

TEXT S1, PDF file, 0.4 MB.

ACKNOWLEDGMENTSThe work of K.R.A. is funded by The Danish Diabetes Academy, which is supported

by the Novo Nordisk Foundation, a Sapere Aude grant from the Danish Council forIndependent Research, and a grant from the Lundbeck Foundation.

REFERENCES1. Jensen KF. 1983. Metabolism of 5-phosphoribosyl 1-pyrophosphate (PRPP)

in Escherichia coli and Salmonella typhimurium, p 1–25. In Munch-PetersenA (ed), Metabolism of nucleotides, nucleosides and nucleobases in micro-organisms. Academic Press, Inc., London, United Kingdom.

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Bjarne Hove-Jensen began graduate studies atthe Enzyme Division, Institute of BiologicalChemistry B, University of Copenhagen, in1976, where he worked on various aspects ofpurine nucleotide salvage metabolism in E. coli.He serendipitously discovered a mutant with analtered phosphoribosyl diphosphate synthase.After receiving his Ph.D. degree in Biochemistryin 1983 at the University of Copenhagen, hecontinued studies of nucleotide metabolism invarious organisms with an emphasis on thephysiological role of PRPP synthase. He became Associate Professor of Bio-chemistry in 1988 at the University of Copenhagen. Fortuitously, around2000 he discovered that PRPP is also an intermediate in the catabolic path-way by which phosphonic acid-phosphorus is assimilated by the carbonphosphorus-lyase pathway. Important parts of his research project were con-ducted on sabbaticals with Dr. Switzer at the University of Illinois at Urbana-Champaign, at Queen’s University, Kingston, Ontario, Canada, and at Aar-hus University. He is now Associate Professor Emeritus at Aarhus Universityand continues studies of the physiological role of PRPP.

Kasper R. Andersen received his Ph.D. degreein structural biology in 2009 at Aarhus Univer-sity. He worked with Nobel Laureate V. Ra-makrishnan and his laboratory at the MedicalResearch Council, Laboratory of Molecular Bi-ology, Cambridge, UK, in an effort to explorethe mechanism of mRNA cleavage by the bacte-rial toxin RelE. From the crystal structures ofRelE bound to the ribosomal A-site in both thepre- and postcleaved state, this group of re-searchers described how the RelE endonucleaseperforms its ribosome-dependent mRNA cleavage. In 2010, Dr. Andersenbegan postdoctoral studies at Massachusetts Institute of Technology, wherehe determined the structure of the largest nucleoporins, Nup188 andNup192, by using a combination of X-ray crystallography and electron mi-croscopy, and he revealed that the structures have distant similarity to nu-clear transport receptors. In 2014, he returned to Denmark as an AssistantProfessor at Aarhus University, where he studies kinase-mediated signaling.

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Mogens Kilstrup received his Master’s degreein Biochemistry in 1986 and his Ph.D. degree inmolecular biology in 1990, both at the Instituteof Biological Chemistry B, University of Copen-hagen. He worked on multivalent regulation ofpyrimidine gene expression in enteric bacteria.He discovered the guanine-responsive PurRregulator and characterized the first purR mu-tant of E. coli. After a short postdoctoral period,he was employed at the Technical University ofDenmark, where he studied mechanisms ofstress response in L. lactis. Dr. Kilstrup became Associate Professor andReader at DTU-Bioengineering, the Technical University of Denmark, in2000 and 2008, respectively. Dr. Kilstrup and Dr. Martinussen collaborateand have jointly discovered and characterized the first activating PRPP-responsive PurR regulator protein of L. lactis, and they have published thefirst in vivo kinetic analysis of allosteric interactions in bacteria by quantify-ing and correlating the intracellular concentrations of PRPP and all purmRNA levels during a purine downshift.

Jan Martinussen received his Ph.D. degreefrom the Department of Molecular Biology,University of Southern Denmark, in 1992. Hisresearch area was the control of gene expressionin E. coli by using the cAMP/CRP and CytR/DeoR regulons and the hok/sok antisense systemas model systems. From 1992 to 1997, he was apostdoctoral researcher at the Danish Centerfor Lactic Acid Bacteria, the Technical Univer-sity of Denmark, where he investigated pyrimi-dine nucleotide metabolism in L. lactis. He be-came Associate Professor at DTU-Bioengineering, Technical University ofDenmark, in 1997, and his work has ever since been focused on the geneticsand physiology of Gram-positive organisms, with an emphasis on nucleotidemetabolism in lactic acid bacteria.

Robert L. Switzer received his Ph.D. degree inBiochemistry from the University of California,Berkeley, in 1966 under the direction of H. A.Barker. He began his studies on the regulationand mechanistic enzymology of bacterial PRPPsynthase at the suggestion of Earl R. Stadtmanwhile a postdoctoral fellow at the National In-stitutes of Health from 1966 to 1968. After join-ing the Department of Biochemistry at the Uni-versity of Illinois at Urbana-Champaign in1968, he continued research on PRPP synthasefor more than 25 years, a period that included fruitful collaborations with Dr.Hove-Jensen and Dr. Michael Becker. He also led research on the regulationof numerous enzymes and genes of bacterial pyrimidine and purine biosyn-thesis. His group discovered the RNA-binding regulatory protein PyrR. Heremained on the University of Illinois Department of Biochemistry facultyuntil retirement as Professor Emeritus in 2002. He was a Guggenheim Fellowand is a Fellow of the American Academy of Microbiology.

Martin Willemoës received his Ph.D. degreefrom the University of Copenhagen in 1996while working with Dr. Hove-Jensen onstructure-function analysis of the PRPP-binding motif in PRPP synthase and the en-zyme kinetics of allosteric regulation of this en-zyme. As both Assistant and Associate Professorat the Centre for Crystallographic Studies, De-partment of Chemistry, University of Copenha-gen (1999 to 2003) and at the Section for Bio-molecular Sciences, Department of Biology(from 2005), he has worked with structure-function analyses of catalysis andregulation of enzymes of pyrimidine nucleotide metabolism. Over the last 7years, his research interests have gradually moved toward the area of proteinand enzyme design as a member of the Linderstrøm-Lang Centre for ProteinScience at the Department of Biology, University of Copenhagen.

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