This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Services
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrowReprints and Permissions
Right arrow Copyright Information
Right arrow Books from ASM Press
Right arrow MicrobeWorld
Citing Articles
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Pope, S. D.
Right arrow Articles by Stewart, V.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Pope, S. D.
Right arrow Articles by Stewart, V.

 Previous Article  |  Next Article 

Journal of Bacteriology, February 2009, p. 1006-1017, Vol. 191, No. 3
0021-9193/09/$08.00+0     doi:10.1128/JB.01281-08
Copyright © 2009, American Society for Microbiology. All Rights Reserved.

Purine Utilization by Klebsiella oxytoca M5al: Genes for Ring-Oxidizing and -Opening Enzymes{triangledown}

Scott D. Pope,{dagger} Li-Ling Chen, and Valley Stewart*

Department of Microbiology, University of California, Davis, California 95616-8665

Received 11 September 2008/ Accepted 25 November 2008


arrow
ABSTRACT
 
The enterobacterium Klebsiella oxytoca uses a variety of inorganic and organic nitrogen sources, including purines, nitrogen-rich compounds that are widespread in the biosphere. We have identified a 23-gene cluster that encodes the enzymes for utilizing purines as the sole nitrogen source. Growth and complementation tests with insertion mutants, combined with sequence comparisons, reveal functions for the products of these genes. Here, we report our characterization of 12 genes, one encoding guanine deaminase and the others encoding enzymes for converting (hypo)xanthine to allantoate. Conventionally, xanthine dehydrogenase, a broadly distributed molybdoflavoenzyme, catalyzes sequential hydroxylation reactions to convert hypoxanthine via xanthine to urate. Our results show that these reactions in K. oxytoca are catalyzed by a two-component oxygenase (HpxE-HpxD enzyme) homologous to Rieske nonheme iron aromatic-ring-hydroxylating systems, such as phthalate dioxygenase. Our results also reveal previously undescribed enzymes involved in urate oxidation to allantoin, catalyzed by a flavoprotein monooxygenase (HpxO enzyme), and in allantoin conversion to allantoate, which involves allantoin racemase (HpxA enzyme). The pathway also includes the recently described PuuE allantoinase (HpxB enzyme). The HpxE-HpxD and HpxO enzymes were discovered independently by de la Riva et al. (L. de la Riva, J. Badia, J. Aguilar, R. A. Bender, and L. Baldoma, J. Bacteriol. 190:7892-7903, 2008). Thus, several enzymes in this K. oxytoca purine utilization pathway differ from those in other microorganisms. Isofunctional homologs of these enzymes apparently are encoded by other species, including Acinetobacter, Burkholderia, Pseudomonas, Saccharomyces, and Xanthomonas.


arrow
INTRODUCTION
 
Purines and purine derivatives comprise a large portion of biomass and are involved in almost every step of life. Not only a major constituent of nucleic acids, they also are central to energy transfer and storage (ATP) as well as protein synthesis and signaling (GTP). Plants, animals, and many microorganisms use purines and purine derivatives to store and translocate nitrogen for assimilation or excretion (96).

Salvage pathways operate to recycle purines, including hypoxanthine and xanthine, back into nucleoside pools (107). Additionally, some organisms can utilize purines as the sole source of nitrogen and carbon. Adenine and guanine are deaminated to form hypoxanthine and xanthine, respectively, which then are oxidized to form uric acid (urate at physiological pH) (Fig. 1). These oxidation steps are catalyzed by xanthine dehydrogenase, a well-studied molybdoflavoenzyme that is conserved from bacteria to humans (51). Two sequential ring-opening steps convert urate via allantoin to allantoate (Fig. 1). Subsequent steps, which comprise different pathways in different microorganisms (96), convert allantoate to ammonium, which is assimilated.


Figure 1
View larger version (28K):
[in this window]
[in a new window]

  		FIG. 1. Purine ring oxidation and opening steps. The enzyme proposed to catalyze each step is shown. The K. oxytoca gene for adenine deaminase was not identified in this study. Dashed lines show reactions that can occur spontaneously.

Some organisms express only the latter portion of the purine utilization pathway and cannot use purines or urate as sole sources of nitrogen. For example, Escherichia coli K-12 can use allantoin and its catabolites as the sole nitrogen source, albeit only under anaerobic conditions (21). Saccharomyces cerevisiae uses allantoin as a nitrogen storage compound (17). However, the complete pathway is present in other bacterial and fungal species, including Bacillus subtilis (84) and Aspergillus nidulans (83).

Molybdoenzymes (excepting dinitrogenase) contain the molybdenum cofactor Mo-molybdopterin (42). Thus, mutations in genes for molybdenum cofactor biosynthetic enzymes (mol genes in bacteria and cnx in A. nidulans) confer pleiotropic phenotypes: these mutants can utilize neither nitrate nor purines, due to lack of the molybdoenzymes nitrate reductase and xanthine dehydrogenase (74). We previously reported that Klebsiella oxytoca mol mutants cannot assimilate nitrate but can utilize xanthine as the sole nitrogen source (32). This suggested, as one possibility, that K. oxytoca uses a molybdenum-independent enzyme in place of conventional xanthine dehydrogenase. Results reported here demonstrate that this is correct, as insertion mutants blocked specifically in xanthine and hypoxanthine utilization define the structural genes for an apparent two-component Reiske nonheme iron oxygenase.

Here, we report analysis of 12 genes whose products catalyze conversion of purines to allantoate. Our investigation of the remaining genes, whose products catalyze allantoate utilization, is ongoing. Results show that several steps in the overall pathway are catalyzed by previously undescribed enzymes.

While this paper was in review, the paper by de la Riva et al. (24), describing the hpxDE, hpxR, hpxO, and hpxPQT genes from Klebsiella pneumoniae W70, was posted in the "JB Accepts" section of the Journal of Bacteriology online edition. Results and conclusions concerning these seven genes are congruent between the two studies.

(Some of the work presented here was submitted by Danielle Carl in 1994 as part of an undergraduate thesis to the Cornell University Division of Biological Sciences Honors Program.)


arrow
MATERIALS AND METHODS
 
Media. Defined, complex, and indicator media for routine genetic manipulations were used as described previously (57). Nitrogen-free medium contained 0.2% (wt/vol) glucose, 1% (wt/vol) sodium citrate, 0.74% (wt/vol) sodium phosphate (pH 8), and 1 mM MgSO4 (53). This medium was supplemented with additional nitrogen sources (10 mM NaNO3 or NH4Cl and 2.5 mM hypoxanthine, xanthine, urate, allantoin, or potassium allantoate) as indicated. Urate solutions were prepared as described previously (79). 5-Bromo-4-chloro-3-indolyl-β-D-galactopyranoside (X-Gal; 40 µg/ml) was included at 40 µg ml–1 to score the Lac phenotypes of MudJ insertion mutants. All nitrogen sources were from Sigma-Aldrich (St. Louis, MO).

Selection for K. oxytoca transformants carrying bla-containing plasmids was accomplished with a combination of carbenicillin and ampicillin (Ap) at 800 and 60 µg ml–1, respectively (54). Streptomycin (Sm) was used at 500 µg ml–1 for selecting K. oxytoca Smr segregants (102). Other antibiotics used were chloramphenicol (Cm) at 50 µg ml–1, kanamycin (Km) at 100 µg ml–1, spectinomycin (Sp) at 50 µg ml–1, and tetracycline (Tc) at 20 µg ml–1.

Defined medium used to grow cultures for β-galactosidase assays and for growth yield tests was buffered with 3-[N-morpholino]propanesulfonic acid (MOPS) as previously described (69). Glucose (40 mM) was used as the sole carbon source, and nitrogen sources were added as indicated.

Culture conditions. Liquid cultures and plates were incubated at 30°C (41). Cultures for β-galactosidase assays were aerated at 240 rpm in 10 ml of medium in 125-ml sidearm flasks. Culture densities were monitored with a Klett-Summerson photoelectric colorimeter (Klett Manufacturing Co., New York, NY) equipped with a no. 66 (red) filter. Cultures in the mid-exponential phase (about 40 Klett units) were harvested, chilled on ice, and washed with saline. Cell pellets were stored overnight at –20°C prior to the assay.

Cultures for nitrogen assimilation tests were grown in 5 ml of MOPS medium in 16- by 175-mm culture tubes, aerated on a roller drum. Nitrogen sources were added over a range of growth-limiting concentrations: for nitrate, 0.4 to 1.8 mM; for histidine, 0.2 to 0.9 mM; for xanthine, 0.1 to 0.45 mM; and for guanine, 0.08 to 0.36 mM. Culture densities (optical densities at 420 nm) were measured after 24 and 48 h.

Phenotypes were determined by growth on plates made with nitrogen-free medium and microbiological agar (Marine BioProducts, Delta, British Columbia) supplemented with the test nitrogen source at a final concentration of 10 mM N atoms. Strains were first streaked for single colonies on plates with added nitrate, which then were replica printed to plates with test compounds (49). Plates for anaerobic nitrogen assimilation tests were incubated in Brewer-Allgeier jars (11).

β-Galactosidase assays. Activity in CHCl3-sodium dodecyl sulfate-permeabilized cells was measured at room temperature, approximately 21°C, by monitoring the hydrolysis of o-nitrophenyl-β-D-galactopyranoside. Specific activities are expressed in arbitrary (Miller) units (63). Cultures were assayed in duplicate, and reported values are averaged from two independent experiments.

Strains. Table 1 lists the strains used in this study. Previously, strain M5al was classified under Aerobacter aerogenes and subsequently under K. pneumoniae. However, phenotypic properties (such as positive reaction in the indole test) place this strain in the species K. oxytoca (see reference 102). Genetic crosses were performed by P1 kc-mediated generalized transduction (90).


View this table:
[in this window]
[in a new window]

  		TABLE 1. Strains and plasmids

MudJ, originally termed Mu dI 1734, is a transposase-null bacteriophage transposon that encodes resistance to Km and is used for isolating lacZ operon fusions (14). Mu cts hP1, a P1 host range variant of bacteriophage Mu (19), was used as a helper phage for K. oxytoca as described previously (13). Tn10d(Tc) and Tn5d(Sp) are transposase-null transposons encoding resistance to Tc and Sp, respectively (25, 47).

Nomenclature. The genetic nomenclature extends that used by de la Riva et al. (24). The mnemonic hpx (hypoxanthine) is applied to all genes whose products are involved in utilization of hypoxanthine or its catabolites.

We use Hpx as a general phenotypic designation without reference to a particular step in the pathway. Specific phenotypes are denoted according to the blocked step (Fig. 1); for example, Hxn– mutants utilize neither hypoxanthine nor xanthine but grow normally with urate and downstream purine catabolites. Similarly, downstream blocks are denoted Urt– (urate), Aln– (allantoin), and Alt– (allantoate).

Transposon insertions. Table 2 lists insertion alleles used in this study. Insertion mutagenesis with transposons MudJ (13), Tn5d(Sp) (35), and Tn10d(Tc) (57) was performed as described previously. Insertions were selected on antibiotic plates, with nitrate as the sole nitrogen source, to preclude isolation of mutants with general defects in nitrogen assimilation. Colonies were then replica printed to antibiotic plates, with purine utilization intermediates (hypoxanthine, xanthine, urate, allantoin, or allantoate) as the sole nitrogen source.


View this table:
[in this window]
[in a new window]

  		TABLE 2. hpx insertions

Plasmids. Table 1 lists the plasmids used in this study. Standard methods (57) were used for restriction endonuclease digestion, ligation, transformation, and PCR amplification (80). Plasmid pVJS3982 was constructed by excising the Kmr determinant from plasmid pEG5005 with restriction endonuclease EcoRI and inserting the {Omega}-Sp integron (30) in its place.

Molecular cloning. In vivo cloning utilized the mini-Mu cloning vectors pEG5005 and pVJS3928 as described previously (53). Lysates prepared from the hpx+ donor strains VJSK038 and VJSK3047 were used to transduce hpx insertion mutants lysogenic for Mu cts hP1. Strain VJSK650 (hpxO; Tcr) was the recipient for cloning with vector pEG5005 (Kmr), and strain VJSK3046 (hpxE; Kmr) was the recipient for cloning with vector pVJS3982 (Spr). Antibiotic-resistant transformants were selected on complex medium, and colonies were replica printed to duplicate nitrogen-free plates, one containing nitrate and the other xanthine or hypoxanthine. Hpx+ colonies were isolated, and plasmid DNA was extracted and retransformed to confirm the phenotype.

This in vivo cloning method generates essentially random inserts whose boundaries are delimited by the ends of bacteriophage Mu DNA (38). Large inserts from the pEG5005-based clones were subcloned into the pHG165 or pSU18 medium-copy-number vector to generate the plasmids pVJS1435, pVJS1436, pVJS1454, and pVJS3941 (Table 1). Subcloning utilized HindIII or SalI restriction endonuclease sites near the Mu DNA ends. These sites are indicated in parentheses in Table 1. All other sites are native to the K. oxytoca hpx DNA region. Sites are denoted according to the nucleotide position within the sequenced region (see below).

DNA sequencing. Automated DNA sequencing reactions (82) were performed by the DNA Sequencing Facility, University of California—Davis. Double-stranded templates were sequenced on a model 373A stretch DNA sequencer by using dye terminator chemistry and AmpliTaq-FS DNA polymerase (Perkin Elmer/Applied Biosystems Division, Foster City, CA). Templates were prepared by using QIAprep spin plasmid kits (Qiagen, Inc., Chatsworth, CA).

Several subclones were generated by deleting or by recloning internal restriction fragments. Sequences were primed from flanking polylinker sequence in subclones or from oligonucleotide primers synthesized to match the ends of sequence reads (primer walking). The DNA sequence was determined from multiple reads on both strands. Sequence information was compiled and analyzed through programs from DNASTAR, Inc. (Madison, WI).

DNA sequence analysis. Gene identifications were based primarily on the results from BLAST searches (1) conducted through servers operated by the National Center for Biotechnology Information at the National Library of Medicine. Proteins with similar sequences are denoted by their locus tags or GenInfo Identifier (GI) accession numbers, as appropriate. Domains (58) are indicated by their database accession numbers: conserved domain (cd), clusters of orthologous groups (COG), or protein family (pfam).

Analysis was facilitated as unpublished whole-genome sequence data for K. pneumoniae strain MGH78578 were released by the Washington University Genome Center. These data were particularly useful for assigning likely promoters and sites for translation initiation and termination, which most likely are conserved between the two species. The K. pneumoniae MGH78578 chromosome sequence is available in GenBank under accession number NC_009648.

Allelic replacement. Two insertion alleles were constructed directly. The hpxR113::{Omega}-Cm allele was made by cloning a BamHI-released {Omega}-Cm interposon (30) into the unique BclI site within the hpxR gene. This insertion was cloned into the conditionally replicating vector pKAS46 and transplanted to the chromosome by integration-excision allelic exchange as described previously (102). The guaD114::pAH144 allele was made by cloning an internal fragment spanning guaD codons 125 to 334 into the conditionally replicating vector pAH144. The single-crossover integration was selected by resistance to Sp. The veracity of both replacements was confirmed by PCR analysis.

{Phi}(hpxD-lacZ) and {Phi}(hpxR-lacZ) operon fusions. The divergent hpxD-hpxR transcription control region was cloned as two approximately 320-nucleotide (nt) PCR-generated fragments with introduced XbaI and BamHI restriction endonuclease sites at either end. The endpoints lie within the codons encoding Ala-30 and Gln-37 of the hpxD and hpxR coding regions, respectively. Fragments were subcloned into BamHI- and XbaI-digested vector pVJS2354 to create {Phi}(hpxD-lacZ) and {Phi}(hpxR-lacZ) operon fusions. The resulting constructs were transplanted into the rha locus by integration-excision allelic exchange as described previously (103).

Nucleotide sequence accession number. The DNA sequence reported in this paper is available from the GenBank nucleotide sequence database under accession number EU884423. The sequence spans 23,740 nt, including 287 nt comprising the 3' portion of a gene homologous to the Serratia marcescens fbpC gene (2) (adjacent to the hpxE gene) and 581 nt comprising the 3' portion of a gene homologous to Erwinia carotovora gene ECA2135 (adjacent to the hpxK gene).


arrow
RESULTS AND DISCUSSION
 
Purine utilization. K. oxytoca M5al can use purines, pyrimidines, and most amino acids as sole sources of nitrogen (66). The purine rings contain four N atoms (Fig. 1), so we wished to determine if all four can be assimilated. We measured the growth yield of K. oxytoca cultured with limiting amounts of nitrogen, as described in Materials and Methods. Nitrate (one assimilable N atom) and histidine (two assimilable N atoms) (56) served as calibration controls. Results indicate that all four N atoms were assimilated from xanthine and that all five N atoms were assimilated from guanine (Table 3 and data not shown). We conclude that K. oxytoca quantitatively assimilates all N atoms from purines. An identical conclusion was drawn by de la Riva et al., who used adenine instead of guanine (24).


View this table:
[in this window]
[in a new window]

  		TABLE 3. Growth yields from nitrogen sources

Insertion mutants. We conducted several screens for transposon insertion mutants with specific blocks in purine utilization, as described in Materials and Methods. Mutants were tested for growth with hypoxanthine, xanthine, urate, allantoin, and allantoate (Fig. 1). Urea was not tested, since urease is encoded at a distinct, well-characterized locus (64). Four classes of mutants were identified. Mutants with the Hxn– phenotype utilized neither hypoxanthine nor xanthine but did utilize urate, allantoin, and allantoate. Thus, these mutants are specifically blocked in the conversion of hypoxanthine and xanthine to urate (Fig. 1). Similarly, mutants with the Urt–, Aln–, and Alt– phenotypes were specifically blocked in the utilization of urate, allantoin, and allantoate, respectively.

Our analysis of genes for utilizing allantoate (hpxFGHIJK), downstream purine catabolites (hpxWXYZ), and an associated regulator (hpxU) will be reported separately. Provisional functions for these genes are included in the deposited GenBank file.

We employed bacteriophage P1-mediated generalized transduction to backcross each insertion to the wild type in order to demonstrate linkage between the transposon and the Hpx– phenotype. Genetic crosses (105) with pairs of insertions encoding different antibiotic resistance phenotypes established the order hpxD-hpxO-hpxB-hpxF (Fig. 2A). Additional crosses not depicted in Fig. 2A yielded congruent results. Subsequent molecular genetic analysis confirmed this order (see below and Fig. 2B).


Figure 2
View larger version (17K):
[in this window]
[in a new window]

  		FIG. 2. Klebsiella genes for purine utilization. (A) Genetic map as determined by generalized transduction. Map distances are calculated as 100 minus percent cotransduction. The arrow points to the selected marker. (B) Physical map as determined by DNA sequence analysis. Genes are denoted by arrows according to their encoded products: gray, enzymes; open, transporters; filled, regulators. Filled circles show sites of transposon insertion, and filled squares show directed allelic replacements. Line arrows indicate predicted RpoN-dependent transcripts. The five genetic modules are denoted by the substrate upon which their products act. (C) Gene organization in K. pneumoniae MGH78578. See also Table 4. Locus tags (KPN_) delimiting the boundary of each unit are shown. Arrowheads indicate relative orientation in the genome.

As we completed the DNA sequence of the hpx locus (see below), we employed whole-colony PCR amplification and DNA sequencing to determine the exact insertion site for each of the transposons. The results revealed 11 distinct insertions (Table 2 and Fig. 2B). Thus, the insertion mutagenesis fell far short of saturating the hpx locus. We used allelic replacement methods to construct two additional null alleles as described below. Overall, analysis of mutant phenotypes in conjunction with predictions from DNA sequence analysis allows functional assignments for each of the genes as described below.

Gene product identification. We cloned the hpx+ genes by complementation as described in Materials and Methods. Further subcloning and complementation analyses localized specific hpx functions to defined regions of the restriction map as described below. We determined the DNA sequence for the 22,872-nt hpx region, and concomitant computer-assisted analyses identified 23 genes (Fig. 2B).

Some functional assignments are based on homology to known proteins. In other instances, however, the proteins had not previously been characterized. In these cases, we initially attempted to identify orthologs present in genomes of other species (48). However, orthologous proteins do not necessarily perform identical functions (75); furthermore, it can be difficult to ascertain orthology (48). Therefore, we have identified what we presume to be "isofunctional homologs" in other species, based not only on overall sequence similarity and conservation of critical residues but also on genetic clustering of structural genes with those of other enzymes in the same or related pathways (i.e., synteny) (43). These assignments are described in detail below and are summarized in Table 4.


View this table:
[in this window]
[in a new window]

  		TABLE 4. hpx genes, proposed functions, and isofunctional homologs

Transcriptional organization. Transcription of most K. oxytoca genes encoding nitrogen assimilation functions is regulated by general nitrogen control (Ntr) acting through the NtrC or Nac activator proteins (7, 55, 62). The NtrC protein activates promoters dependent upon the RpoN ({sigma}54) form of RNA polymerase, which recognizes promoters whose consensus sequence is TGGYRYRRNNNYYGCW (where R represents purine, Y represents pyrimidine, and W represents A or T) (55). We identified six potential promoters by visual inspection, each of which is conserved in the K. pneumoniae MGH78578 sequence. These promoters are indicated in Fig. 2B as directing synthesis of the following hypothetical transcripts, shown by arrows: hpxO, hpxPQT, guaD, hpxC, hpxWXYZ, and hpxFGHIJK. Nucleotide sequence coordinates for these potential promoters are included in the deposited GenBank file. Thus, transcription of most hpx genes is likely activated directly by phospho-NtrC protein. We emphasize that we have not examined operon structure through direct experiments.

Modularity of hpx gene organization. Genes apparently involved in each step of the overall pathway are clustered together, so the overall hpx locus appears to comprise a supraoperonic cluster (100) containing five distinct modules of hpx genes plus the guaD and hpxU genes (Fig. 2B). The gene content and order within modules is essentially identical in the K. pneumoniae MGH78578 genome (Table 4). However, in K. pneumoniae, the five modules are separated into three distinct groups: hpxEDR-hpxOPQT- ~100 kb - hpxKJIHGF- hpxU-hpxWXYZ - ~16 kb - hpxBASC-guaD (Fig. 2C). Note that the relative orientations between modules also differ in the two species.

For K. oxytoca M5al, both the genetic and the physical maps indicate that these modules are immediately adjacent. For example, the cotransduction frequency between the hpxF111::Tn5d(Sp) and hpxD103::MudJ alleles corresponds to a physical distance of roughly 30 kb, as estimated from the formula of Sanderson and Roth (81), approximately twice the actual distance of about 16 kb. Similarly, the calculated distance between the hpxO112::Tn10d(Tc) and hpxB105::MudJ alleles, roughly 15 kb, is approximately twice the actual distance of about 8 kb. Thus, at least for the region encompassing the hpx genes, the genomes of these two Klebsiella species appear to be organized differently.

Guanine to xanthine: GuaD guanine deaminase. Guanine is deaminated to form xanthine (Fig. 1). The guaD gene product is most similar to E. coli K-12 guanine deaminase (60% sequence identity over 97% of its length), the product of the guaD (ygfP; cd01303) gene (61). The GuaD sequence includes a conserved nine-residue motif (P-G-X-V/I-D-X-H-T/V/I-H) also shared by members of the cyclic amidohydrolase family. This motif has been implicated in Mn2+ or Zn2+ binding in members of the family (46, 101). E. coli guanine deaminase contains Zn2+ (61).

We constructed a guaD disruption strain to determine the role of this gene in purine utilization. The resulting mutant failed to grow on plates with guanine as the sole nitrogen source but exhibited wild-type growth with adenine, hypoxanthine, xanthine, and urate. The guaD coding sequence is preceded by an apparent RpoN-dependent promoter and thus likely constitutes a single-gene operon (Fig. 2B).

K. oxytoca M5al also uses adenine as the sole nitrogen source (66). However, the hpx locus does not include the gene for adenine deaminase.

(Hypo)Xanthine to urate: the hpxR-hpxDE module. Hypoxanthine is hydroxylated to form xanthine, which also is hydroxylated to form urate (Fig. 1). Xanthine dehydrogenase (purine hydroxylase), a well-studied molybdoflavoenzyme, is the previously described enzyme catalyzing the sequential oxidation of hypoxanthine and xanthine to urate (51). Structural genes for this enzyme have been characterized for Rhodobacter capsulatus (52) and Bacillus subtilis (84). However, no such genes are present in the K. oxytoca hpx locus, and none were identified in the K. pneumoniae MGH78578 genome sequence.

Three MudJ insertion mutants exhibited the Hxn– phenotype, failing to grow with hypoxanthine or xanthine as the sole nitrogen source but displaying wild-type growth with urate and downstream purine catabolites. Two of these insertions disrupt the hpxE gene, whereas the third lies 20 nt upstream of the hpxD initiation codon (Table 2) and therefore blocks hpxDE operon transcription. We also constructed an hpxR insertion to determine the role of this gene in regulation (see below). This hpxR::{Omega}-Cm mutant also exhibited the Hxn– phenotype. All four insertion mutants were complemented to the Hxn+ phenotype by plasmid pVJS3958, a subclone that contains only the hpxRAB region (Table 1). Therefore, the hpxD, hpxE, and hpxR genes encode functions required for conversion of (hypo)xanthine to urate, a conclusion reached independently by de la Riva et al. (24). The hpxD and hpxE genes are separated by only 13 nt and thus likely comprise an operon.

(i) HpxD and HpxE: (hypo)xanthine hydroxylase. Sequence comparisons indicate that the hpxE and hpxD gene products comprise the reductase and oxygenase, respectively, of a two-component Rieske nonheme iron aromatic-ring-hydroxylating oxygenase system. The HpxD sequence is that of a Rieske nonheme iron oxygenase, categorized in the phthalate family of Gibson and Parales (34), group I of Nam et al. (68), and class IA of Batie et al. (5). These oxygenases are homomultimeric aromatic-ring-activating mono- and dioxygenases with broad substrate specificity. They share relatively low sequence identity and range in length from 329 to 446 aminoacyl residues (the HpxD enzyme is 345 residues). The HpxD sequence includes a Rieske-type iron-sulfur cluster (consensus sequence, C-X-H-X15-26-C-X2-H; pfam00355; COG2146) near the amino terminus and a likely nonheme Fe(II)-binding 2-His-1-carboxylate facial triad (consensus sequence, H-X4-5-H-Xn-D) near the center (reviewed in references 31 and 73). The HpxD sequence shares 29% identity over a 160-residue span with that of the well-studied phthalate dioxygenase (OphA2 enzyme; GI 4128221; 443 residues) from Burkholderia cepacia DBO1 (6, 16).

The HpxE sequence (322 residues) is that of a corresponding reductase, categorized in class IA of Batie et al. (5), and includes the three defining cofactor domains: an amino-terminal FMN binding domain (consensus sequences, R-X-Y-S-L and G/S-R-G-G-S), a central NAD binding domain (consensus sequence, G-G-I-G-I-T-P; pfam00175) (in HpxE and KshB [95], the first Gly residue is replaced with Ala), and a carboxyl-terminal plant-type iron-sulfur cluster (consensus sequence, C-X2-G-X-CG-X-C-X6-G-X3-HRD-X2-L-X5-A-X7-C-X-S; pfam00111; COG0633) (in HpxE, L-X5 is Q-X6) (16, 18, 67, 95; reviewed in reference 60). The HpxE sequence shares 34% identity with the sequence of the phthalate dioxygenase reductase (OphA1 enzyme; GI 13432205; 322 residues) from B. cepacia DBO1 (6, 16) and about 40% identity with those of the vanillate O-demethylase reductases from a variety of species (e.g., the VanB enzyme from Acinetobacter sp. strain ADP1; GI 2271499; 318 residues) (86).

Purines, which resemble two fused aromatic rings, exhibit some aromatic properties (87). We suggest that the HpxE and HpxD enzymes comprise (hypo)xanthine hydroxylase reductase and bifunctional (hypo)xanthine hydroxylase, respectively. This enzyme complex likely functions by the same mechanisms as those described for homologous aromatic-ring-hydroxylating oxygenases, hydroxylating first the C-2 position of hypoxanthine to yield the enol form of xanthine and second the C-8 position to yield the enol form of urate (Fig. 1). Purines undergo tautomeric interconversions to shift between the relatively unstable enol form and the more stable keto form (8, 23, 87).

(ii) Transcriptional regulation in the hpxR-hpxDE module. The two hpxE::MudJ insertion mutants expressed LacZ activity, indicating that they form {Phi}(hpxE-lacZ) operon fusions. We cultured these strains in defined medium, with glutamine as a neutral nitrogen source, and measured LacZ-specific activity. Both strains expressed approximately 4,000 Miller units after growth with added xanthine but less than 50 units after growth with added urate, allantoin, or allantoate and about 10 units after growth with added xanthine plus ammonium. This indicates that hpxDE operon transcription is subject to dual control by global nitrogen regulation (Ntr) and by pathway-specific induction by (hypo)xanthine (see also below) but not downstream purine catabolites.

The hpxDE operon shares a 125-nt intergenic region with the divergently transcribed hpxR gene (Fig. 2B). The HpxR sequence (312 residues) is similar to those of LysR family transcriptional regulators, many of which regulate degradation pathways for aromatic compounds (93). The HpxR sequence includes an amino-terminal helix-turn-helix motif (pfam00126) and a carboxyl-terminal LysR-type substrate-binding domain (pfam03466).

We constructed an hpxR::{Omega}-Cm insertion strain to determine the role of this gene in purine utilization. We also constructed monocopy {Phi}(hpxD-lacZ) and {Phi}(hpxR-lacZ) operon fusions at a separate locus in order to evaluate expression in hpx+ strains. We cultured both hpxR+ and hpxR::{Omega}-Cm derivatives of these strains in defined medium, with nitrate as a neutral nitrogen source, and measured LacZ-specific activity. The results are shown in Table 5. Expression from the {Phi}(hpxD-lacZ) hpxR+ strain was subject to both Ntr and pathway-specific control, consistent with the results from the hpxE::MudJ strains summarized above. In contrast, expression from the {Phi}(hpxD-lacZ) hpxR::{Omega}-Cm strain was undetectable, demonstrating that the HpxR protein is essential for activating hpxDE operon transcription. This is consistent with the Hxn– phenotype conferred by the hpxR::{Omega}-Cm insertion noted above. Expression from the {Phi}(hpxR-lacZ) strain was essentially indifferent to added ammonium or hypoxanthine but was elevated about threefold in the hpxR::{Omega}-Cm insertion strain relative to the level for the hpxR+ strain. This suggests that hpxR transcription is subject to weak negative autoregulation.


View this table:
[in this window]
[in a new window]

  		TABLE 5. Effects of hypoxanthine and ammonium on expression from {Phi}(hpxD-lacZ) and {Phi}(hpxR-lacZ) constructs

These results indicate that the HpxR protein is activated by binding (hypo)xanthine to stimulate hpxDE operon transcription initiation. The source of Ntr regulation is less clear. No RpoN-dependent promoter is evident in the hpxD-hpxR intergenic region in either K. oxytoca M5al or K. pneumoniae MGH78578. Expression of the {Phi}(hpxR-lacZ) fusion was not regulated by Ntr, indicating that Ntr regulation does not act by controlling HpxR protein synthesis.

Very similar results were reported by de la Riva et al. (24), who further demonstrated specific binding of HpxR protein to the intergenic regulatory region. They additionally concluded that Ntr control is mediated by neither the NtrC nor the Nac proteins (7).

(iii) HpxD, HpxE, and HpxR isofunctional homologs in other species. The HpxD and HpxE sequences are most closely related to sequences from Xanthomonas spp. (63% and 47% identity, respectively), currently annotated as vanillate O-demethylase. Likewise, the HpxR sequence is most closely related to sequences from Xanthomonas spp. (46% identity). The genes encoding these proteins are adjacent to several other genes involved in purine catabolism (Table 4). Therefore, these genes comprise the hpxR-hpxDE module in Xanthomonas spp.

Urate to S-(+)-allantoin: the hpxO-hpxPQT module. Urate is hydroxylated to form 5-hydroxyisourate (HIU), which is hydrolyzed to form 2-oxo-4-hydroxy-4-carboxy-5-ureidoimidazoline (OHCU), which is decarboxylated to form S-(+)-allantoin (92) (Fig. 1). HIU also undergoes spontaneous conversion to racemic allantoin (45). These reactions comprise the first ring-opening step in purine utilization.

Uricase (urate oxidase) is the previously described enzyme catalyzing the initial oxidation of urate to HIU. Structural genes for this enzyme have been characterized for Bacillus subtilis (84) and for a range of eukaryotic species (71). However, no such genes are present in the K. oxytoca hpx locus, and none were identified in the K. pneumoniae MGH78578 genome sequence.

Two insertion mutants exhibited the Urt– phenotype, failing to grow with hypoxanthine, xanthine, or urate as the sole nitrogen source but displaying wild-type growth with allantoin and allantoate. Both insertions disrupt the hpxO gene (Table 2). Therefore, the hpxO gene encodes a function required for utilization of urate, a conclusion reached independently by de la Riva et al. (24).

The hpxP and hpxQ genes overlap (23 nt), as do the hpxQ and hpxT genes (4 nt), and thus likely comprise the hpxPQT operon (Fig. 2B). Apparent RpoN-dependent promoters for both the hpxO gene and the divergently transcribed hpxPQT operon are present in the 346-nt intergenic region.

(i) HpxO: urate hydroxylase. Sequence comparisons classify the hpxO gene product (42 kDa) in the {approx}45-kDa aromatic-ring flavoprotein monooxygenase family (pfam01360), members of which share relatively low sequence similarity (40). For example, the HpxO sequence shares 28% sequence identity over 84% of its length with salicylate hydroxylase (NahG enzyme; GI 4104764; 437 residues) from Pseudomonas stutzeri AN10 (9).

Enzymes in this family cleave dioxygen without the use of a metal ion and incorporate one oxygen atom as a hydroxyl group ortho or para to an existing hydroxyl group while reducing the other oxygen atom to water (33; reviewed in reference 40). The HpxO sequence (384 residues) comprises three defining cofactor domains: an amino-terminal domain that binds the ADP moiety of flavin adenine dinucleotide (FAD) (consensus sequence, G-X-G-X2-G-X3-A/G-X6-G; pfam01494), a central domain that binds the pyrophosphate moiety of NAD(P)H and interacts indirectly with the pyrophosphate moiety of FAD (consensus sequence, D-X5-DG-X5-R; pfam01360), and a carboxyl-terminal domain that binds the ribose moiety of FAD. Sequences in the last domain vary considerably, save for a conserved G-D sequence (10, 28, 29, 59; reviewed in reference 40).

p-Hydroxybenzoate hydroxylase (PobA enzyme) is the archetype for the {approx}45-kDa aromatic-ring flavoprotein monooxygenase family (3). The reaction mechanism initiates with binding of the substrate (p-hydroxybenzoate). This leads to the reduction of the flavin isoalloxazine ring by NADPH, forming reduced FAD. Dioxygen then binds to the complex and is reduced by two single-electron transfers. The resulting C4a-flavin (hydro)peroxide hydroxylates the substrate (33; reviewed in reference 40). The HpxO enzyme likely uses a similar mechanism to hydroxylate urate to form HIU (Fig. 1).

(ii) HpxT and HpxQ. Sequence comparisons indicate that the HpxT and HpxQ proteins are HIU hydrolase and OHCU decarboxylase, respectively (15, 44, 77). The HpxT sequence (108 residues; COG2351) is 32% identical to that of HIU hydrolase (PucM protein; 121 residues) from Bacillus subtilis 168 (44, 84). The HpxQ sequence (166 residues; COG3195) is 27% identical to that of the amino-terminal 165 residue OHCU decarboxylase domains of urate oxidase (uricase) from Bacillus sp. strain TB-90 (GI 21431959) (104) and B. subtilis 168 (PucL protein) (84). The Bacillus uricase is synthesized as a precursor of about 500 residues, from which the amino-terminal OHCU decarboxylase is proteolytically cleaved (70).

(iii) HpxP purine permease. The HpxP sequence (460 residues) shares 32% identity over 95% of its length with the high-affinity, high-capacity urate-xanthine permease UapA (GI 6136091; 615 residues) (36) and 30% identity over 90% of its length with the general purine permease UapC (GI 1351342; 580 residues), both from A. nidulans (26). Additionally, the HpxP sequence is 34% identical to those of the urate permeases PucJ (449 residues) and PucK (430 residues) from B. subtilis 168. The HpxP sequence meets the criteria for inclusion in the nucleobase ascorbate transporter family (COG2233) (37), including the nucleobase ascorbate transporter signature motif (Q/E/P-N-X-G-X4-T-R/K/G), the QH motif in the middle of transmembrane segment 1, and a conserved alanyl residue important for function but not specificity (Ala-404 in UapA and Ala-272 in HpxP).

The region spanning residues 378 to 446 in UapA and 336 to 404 in UapC (analogous to the region spanning residues 246 to 314 in HpxP) determines uptake specificity, with UapA residues Glu-412 and Arg-414 and UapC residues Gln-370 and Glu-372 possibly forming part of a purine binding site (27). The corresponding residues in the HpxP sequence are more similar to those of UapC than to those of UapA: Ser-280HpxP and Gln-370UapC are both uncharged, with polar side chains, whereas Glu-412UapA is acidic, and Asp-282HpxP and Glu-372UapC are both acidic, whereas Arg-414UapA is basic. This suggests that HpxP may be a general purine permease, an appealing hypothesis since this is the only hpx cluster-encoded permease for transport of guanine, hypoxanthine, xanthine, and urate.

(iv) HpxO isofunctional homologs in other species. The HpxO sequence is most similar (53% identical) to a protein sequence annotated as "putative flavoprotein monooxygenase acting on aromatic compound" from Acinetobacter sp. strain ADP1 (locus tag ACIAD3540; GI 49532471; 385 residues). This gene is adjacent to those encoding HIU hydrolase and OHCU decarboxylase (Table 4), supporting its assignment as encoding the isofunctional homolog of HpxO protein. Intriguingly, Xanthomonas genomes contain a gene for a presumptive flavin-containing monooxygenase (e.g., XCC0279), not homologous to HpxO, adjacent to those encoding HIU hydrolase and OHCU decarboxylase (Table 4). Thus, it appears that at least two different monooxygenases have been recruited into the urate hydroxylase step of the purine utilization pathway.

R-(–)-allantoin to S-(+)-allantoin to allantoate: the hpxC-hpxSAB module. Conversion of allantoin to allantoate is the second of the two ring-opening steps in purine utilization (Fig. 1). Five insertion mutants exhibited the Aln– phenotype, failing to grow with allantoin or upstream purine catabolites as the sole nitrogen source but displaying wild-type growth with allantoate. The insertions disrupt the hpxA and hpxB genes (Table 2). Therefore, the hpxAB operon encodes functions involved in allantoin utilization. However, the hpxA::Tn10d(Tc) insertion was complemented by subclones carrying the hpxB but not the hpxA gene, indicating that the hpxA gene is not essential for growth on allantoin. Presumably, the hpxA::Tn10d(Tc) insertion is polar on hpxB gene expression.

Allantoinase is the previously described enzyme catalyzing the conversion of allantoin to allantoate (91). Structural genes for this enzyme have been characterized for Bacillus subtilis (84) and E. coli (21) and for a range of eukaryotic species (12). However, no such genes are present in the K. oxytoca hpx locus, and none were identified in the K. pneumoniae MGH78578 genome sequence.

Most characterized allantoinases are specific for the S isomer. However, the nonenzymatic conversion of HIU to allantoin creates a racemic mixture of R- and S-allantoin (45), and allantoin is present as the racemic mixture in decaying organic matter (94). Allantoin racemase has been characterized for bacteria (94) and for Candida utilis (72). To our knowledge, the structural gene for an allantoin racemease has not been identified.

An apparent RpoN-dependent promoter for the hpxC gene (but not for the hpxSAB operon) is present in the 52-nt hpxC and hpxSAB intergenic region. The hpxS and hpxA genes overlap (1 nt; TAATG), and the hpxA and hpxB genes are separated by only 16 nt, so these likely comprise the hpxSAB operon (Fig. 2B).

(i) HpxB: allantoinase. Sequence comparisons suggested that the HpxB enzyme catalyzes conversion of allantoin to allantoate. The HpxB sequence (310 residues; COG0726) shares 26% identity over 74% of its length with an imidase (291 residues; GI 14040045) from Ralstonia eutropha (97). The imidase catalyzes the hydrolysis of hydantoin and related compounds. Since allantoin is a 5-substituted hydantoin (ureidohydantoin), this suggests that the HpxB enzyme acts similarly to the imidase by hydrolyzing a cyclic C-N bond in the hydantoin ring to form allantoate (Fig. 1).

This conjecture was confirmed by Ramazzina et al., who recently identified the HpxB homolog from Pseudomonas fluorescens DSM 50090 as an (S)-allantoin-specific allantoinase (76). They denote this enzyme PuuE.

(ii) HpxA: allantoin racemase. Sequence comparisons indicate that the HpxA protein is allantoin racemase. The HpxA sequence (247 residues; COG 4126; pfam01177) shares 45% identity over 96% of its length with 5-substituted hydantoin racemase (HyuE enzyme; 229 residues; GI 266318) from Pseudomonas sp. strain NS671 (98). The HyuE enzyme catalyzes racemization of a variety of 5-substituted hydantoins (99). Since HpxB allantoinase is specific for the S enantiomer (76), allantoin racemase could be important for efficient allantoin utilization in natural environments. However, allantoin racemase likely is not required for colony formation on plates with abundant (racemic) allantoin, explaining why the hpxB+ gene was able to complement the polar hpxA insertion (see above).

(iii) HpxC: allantoin permease. The HpxC sequence (495 residues; COG1953) shares 22% identity over 85% of its length with allantoin permease from Saccharomyces cerevisiae (Dal4p protein; 635 residues) and 33% identity over 92% of its length with allantoin permease from Bacillus subtilis W168 (PucI protein; 490 residues).

(iv) HpxS. Sequence comparisons indicate that the HpxS protein is a member of the GntR superfamily of transcriptional regulators (COG1802). For example, the HpxS sequence (238 residues) shares 28% identity over 58% of its length with the MatR repressor (222 residues; GI 6840861), which controls expression of the matABC operon responsible for the uptake and conversion of malonate to acetyl-coenzyme A in Rhizobium leguminosarum (50).

The GntR family is made up of bacterial transcriptional regulators with similar amino-terminal helix-turn-helix DNA-binding domains and divergent carboxyl-terminal effector-binding and oligomerization domains (93). The GntR superfamily has been subdivided into four subfamilies on the basis of secondary structure in the C-terminal domain (78). On the basis of these considerations, the HpxS protein can be classified into the VanR subgroup of the FadR subfamily (COG2186). Many proteins in the FadR subfamily regulate operons involved in amino acid metabolism. Allantoin resembles a dipeptide, and the bond cleaved to yield allantoate resembles a peptide bond (Fig. 1).

(v) Transcriptional regulation in the hpxC-hpxSAB module. The hpxB105::MudJ mutant expressed LacZ activity, indicating that this insertion forms a {Phi}(hpxB-lacZ) operon fusion. We cultured this strain in defined medium, with glutamine as a neutral nitrogen source. LacZ-specific activity was approximately 1,600 Miller units after growth with added xanthine and about 10 units after growth with added xanthine plus ammonium. This indicates that hpxSAB operon transcription is subject to global nitrogen regulation (Ntr). LacZ-specific activity was about 1,050 units after growth with added allantoin, about 775 units with urate, and about 400 to 500 units with allantoate or with no addition, suggesting possible pathway-specific induction by allantoin or an upstream intermediate. Presumably, the HpxS protein controls pathway-specific transcriptional regulation of the hpxSAB operon.

(vi) HpxA and HpxB isofunctional homologs in other species. The HpxB allantoinase sequence is most similar (about 70% identity) to protein sequences typically annotated as "polysaccharide deacetylase" (76), and the HpxA sequence is most similar (up to 62% identity) to protein sequences typically annotated as "hydantoin racemase." Representative examples are shown in Table 4. In the case of Burkholderia spp. and related species, the hpxA and hpxB homologs are present as an hpxSCAB module within a larger cluster of purine utilization genes, including those for HIU hydrolase and OHCU decarboxylase. In contrast, the Acinetobacter sp. hpxB homolog is clustered with the hpxO, hpxT, and hpxQ homologs, encoding enzymes of urate catabolism, whereas the hpxA and hpxC homologs are adjacent to each other at a separate location. This indicates that the HpxA allantoin racemase may function physiologically to scavenge exogenous, racemic allantoin. Consistent with this notion, the Xanthomonas spp. have hpxB homologs but none for hpxA or hpxC.

Van der Drift et al. identified allantoin racemase in P. fluorescens and Pseudomonas putida (among others) but not in Pseudomonas aeruginosa or P. stutzeri (94). Database searches reveal that hpxA homologs are present in genomes of P. fluorescens, P. putida, and Pseudomonas syringae but not in P. aeruginosa or P. stutzeri (locus tag Pput_1557 in P. putida F1). This reinforces the conclusion that hpxA encodes allantoin racemase. In these genomes, the hpxA and hpxC genes are adjacent to each other but not to other purine utilization genes, as described above for the Acinetobacter sp. (The P. syringae clusters include an hpxS homolog as well.) In contrast, the hpxB gene, encoding allantoinase, is present in the genomes of all five Pseudomonas species examined (locus tag Pput_1582 in P. putida F1), clustered with several other genes involved in purine utilization.

The HpxA sequence shares 25% identity with the Dcg1p proteins of S. cerevisiae (244 residues; locus tag YIR030C) and Candida albicans (232 residues; locus tag CaO19.244). The S. cerevisiae DCG1 gene was identified as a nitrogen-regulated gene within the DAL cluster, encoding allantoinase, allantoin permease, and allantoicase (106), but the function of the Dcg1p protein has not been determined. Allantoin racemase activity has been characterized for Candida utilis (72). The analysis presented here indicates that Dcg1p may function as allantoin racemase.

Conclusions. Microbial genome sequencing projects reveal large numbers of genes with unknown functions. Here, Klebsiella purine utilization (Fig. 1) provides numerous examples of unpredicted enzymes catalyzing an established pathway (96).

Several enzymes were evidently recruited from pathways for catabolism of aromatic compounds. Aromatic compounds and the pathways for their catabolism are diverse and widespread, and so, in retrospect, it is unsurprising that such enzymes are involved in oxidizing purines (Fig. 1).

Oxidation of (hypo)xanthine to form urate is catalyzed by a two-component Rieske nonheme iron aromatic-ring-hydroxylating oxygenase system (HpxE-HpxD enzyme), and oxidation of urate to form 5-hydroxyisourate is catalyzed by an {approx}45-kDa aromatic-ring flavoprotein monooxygenase (HpxO enzyme). These enzymes were identified independently by de la Riva et al. (24). Genome sequence comparisons suggest that a distinct flavin-containing monooxygenase catalyzes the latter step in Xanthomonas spp. (Table 4).

Yet another enzyme is involved in xanthine utilization by A. nidulans, for which purine utilization has long been studied (83). Early work established that mutants lacking xanthine dehydrogenase (hxA) or molybdenum cofactor (cnx) exhibit leaky growth with xanthine (but not hypoxanthine) as the sole nitrogen source (22). This indicated the presence of a molybdenum cofactor-independent xanthine alternative pathway, dependent on the xanA gene (85). Recently, it was established that XanA protein, a member of the taurine dioxygenase group of enzymes active on aromatic compounds, is an {alpha}-ketoglutarate-dependent dioxygenase that hydroxylates xanthine to form urate (20, 65).

Finally, our analysis reveals that Klebsiella spp. employ HpxA allantoin racemase, related to hydantoin racemases (72, 94), as well as the HpxB (PuuE) allantoinase, related to polysaccharide deacetylases and described recently by Ramazzina and coworkers (76). Our work in progress indicates further that other previously undescribed enzymes are involved in allantoate catabolism (Fig. 2).


arrow
ACKNOWLEDGMENTS
 
We thank Ryan Feathers for his substantial contributions to determining the hpx DNA sequence, Becky Parales and Sydney Kustu for helpful discussion and critique of the manuscript, and Juan Aguilar for providing details on the hpx genetic nomenclature. Cornell University undergraduate students Jeff Hanna, Ann-Marie Roy, and Wendy Pyo participated in the early stages of this work. V.S. is grateful to Robert Rabson, Gregory Dilworth, James Tavares, and Sharlene Weatherwax (Energy Biosciences Program; Chemical Sciences, Geosciences and Biosciences Division) for funding basic research in Klebsiella nitrogen assimilation.

S.D.P. was the recipient of a University of California President's Undergraduate Fellowship. This study was supported by U.S. Department of Energy grant DE-FG03-99ER20326 from the Office of Basic Energy Sciences.


arrow
FOOTNOTES
 
* Corresponding author. Mailing address: Department of Microbiology, University of California, One Shields Ave., Davis, CA 95616-8665. Phone: (530) 754-7994. Fax: (530) 752-9014. E-mail: vjstewart{at}ucdavis.edu Back

{triangledown} Published ahead of print on 5 December 2008. Back

{dagger} Present address: Molecular Biology Interdepartmental Ph.D. Program and Department of Microbiology, Immunology & Molecular Genetics, University of California, Los Angeles, CA 90095-1489. Back


arrow
REFERENCES
 
    1
  1. Altschul, S. F., T. L. Madden, A. A. Schaffer, J. Zhang, Z. Zhang, W. Miller, and D. J. Lipman. 1997. Gapped BLAST and PSI-BLAST: a new generation of protein database search programs. Nucleic Acids Res. 25:3389-3402.[Abstract/Free Full Text]
    2. 2
  2. Angerer, A., S. Gaisser, and V. Braun. 1990. Nucleotide sequences of the sfuA, sfuB, and sfuC genes of Serratia marcescens suggest a periplasmic-binding-protein-dependent iron transport mechanism. J. Bacteriol. 172:572-578.[Abstract/Free Full Text]
    3. 3
  3. Ballou, D. P., B. Entsch, and L. J. Cole. 2005. Dynamics involved in catalysis by single-component and two-component flavin-dependent aromatic hydroxylases. Biochem. Biophys. Res. Commun. 338:590-598.[CrossRef][Medline]
    4. 4
  4. Bartolome, B., Y. Jubete, E. Martinez, and F. de la Cruz. 1991. Construction and properties of a family of pACYC184-derived cloning vectors compatible with pBR322 and its derivatives. Gene 102:75-78.[CrossRef][Medline]
    5. 5
  5. Batie, C. J., D. P. Ballou, and C. C. Correll. 1991. Phthalate dioxygenase reductase and related flavin-iron-sulfur containing electron transferases, p. 543-556. In F. Müller (ed.), Chemistry and biochemistry of flavoenzymes,vol. 3. CRC Press, Boca Raton, FL.
    6. 6
  6. Batie, C. J., E. LaHaie, and D. P. Ballou. 1987. Purification and characterization of phthalate oxygenase and phthalate oxygenase reductase from Pseudomonas cepacia. J. Biol. Chem. 262:1510-1518.[Abstract/Free Full Text]
    7. 7
  7. Bender, R. A. 1991. The role of the NAC protein in the nitrogen regulation of Klebsiella aerogenes. Mol. Microbiol. 5:2575-2580.[CrossRef][Medline]
    8. 8
  8. Bendich, A. 1955. Chemistry of purines and pyrimidines, p. 81-136. In E. Chargaff and J. N. Davidson (ed.), The Nucleic Acids,vol. 1 . Academic Press, Inc., New York, NY.
    9. 9
  9. Bosch, R., E. Garcia-Valdes, and E. R. Moore. 2000. Complete nucleotide sequence and evolutionary significance of a chromosomally encoded naphthalene-degradation lower pathway from Pseudomonas stutzeri AN10. Gene 245:65-74.[CrossRef][Medline]
    10. 10
  10. Bosch, R., E. R. Moore, E. Garcia-Valdes, and D. H. Pieper. 1999. NahW, a novel, inducible salicylate hydroxylase involved in mineralization of naphthalene by Pseudomonas stutzeri AN10. J. Bacteriol. 181:2315-2322.[Abstract/Free Full Text]
    11. 11
  11. Brewer, J. H., and D. L. Allgeier. 1966. Safe self-contained carbon dioxide-hydrogen anaerobic system. Appl. Microbiol. 14:985-988.[Medline]
    12. 12
  12. Buckholz, R. G., and T. G. Cooper. 1991. The allantoinase (DAL1) gene of Saccharomyces cerevisiae. Yeast 7:913-923.[CrossRef][Medline]
    13. 13
  13. Cali, B. M., J. L. Micca, and V. Stewart. 1989. Genetic regulation of nitrate assimilation in Klebsiella pneumoniae M5al. J. Bacteriol. 171:2666-2672.[Abstract/Free Full Text]
    14. 14
  14. Castilho, B. A., P. Olfson, and M. J. Casadaban. 1984. Plasmid insertion mutagenesis and lac gene fusion with mini-Mu bacteriophage transposons. J. Bacteriol. 158:488-495.[Abstract/Free Full Text]
    15. 15
  15. Cendron, L., R. Berni, C. Folli, I. Ramazzina, R. Percudani, and G. Zanotti. 2007. The structure of 2-oxo-4-hydroxy-4-carboxy-5-ureidoimidazoline decarboxylase provides insights into the mechanism of uric acid degradation. J. Biol. Chem. 282:18182-18189.[Abstract/Free Full Text]
    16. 16
  16. Chang, H. K., and G. J. Zylstra. 1998. Novel organization of the genes for phthalate degradation from Burkholderia cepacia DBO1. J. Bacteriol. 180:6529-6537.[Abstract/Free Full Text]
    17. 17
  17. Cooper, T. G. 1996. Regulation of allantoin catabolism in Saccharomyces cerevisiae, p. 139-170. In R. Brambl and G. A. Marzluf (ed.), The mycota. A comprehensive treatise on fungi as experimental systems for basic and applied research,vol. III . Biochemistry and molecular biology. Springer-Verlag, Berlin, Germany.
    18. 18
  18. Correll, C. C., C. J. Batie, D. P. Ballou, and M. L. Ludwig. 1992. Phthalate dioxygenase reductase: a modular structure for electron transfer from pyridine nucleotides to [2Fe-2S]. Science 258:1604-1610.[Abstract/Free Full Text]
    19. 19
  19. Csonka, L. N., M. M. Howe, J. L. Ingraham, L. S. Pierson III, and C. L. Turnbough, Jr. 1981. Infection of Salmonella typhimurium with coliphage Mu d1 (Apr lac): construction of pyr::lac gene fusions. J. Bacteriol. 145:299-305.[Abstract/Free Full Text]
    20. 20
  20. Cultrone, A., C. Scazzocchio, M. Rochet, G. Montero-Moran, C. Drevet, and R. Fernandez-Martin. 2005. Convergent evolution of hydroxylation mechanisms in the fungal kingdom: molybdenum cofactor-independent hydroxylation of xanthine via alpha-ketoglutarate-dependent dioxygenases. Mol. Microbiol. 57:276-290.[CrossRef][Medline]
    21. 21
  21. Cusa, E., N. Obradors, L. Baldoma, J. Badia, and J. Aguilar. 1999. Genetic analysis of a chromosomal region containing genes required for assimilation of allantoin nitrogen and linked glyoxylate metabolism in Escherichia coli. J. Bacteriol. 181:7479-7484.[Abstract/Free Full Text]
    22. 22
  22. Darlington, A. J., and C. Scazzocchio. 1968. Evidence for an alternative pathway of xanthine oxidation in Aspergillus nidulans. Biochim. Biophys. Acta 166:569-571.[Medline]
    23. 23
  23. Davidson, J. N. 1957. The biochemistry of the nucleic acids, 3d ed. Methuen, London, United Kingdom.
    24. 24
  24. de la Riva, L., J. Badia, J. Aguilar, R. A. Bender, and L. Baldoma. 2008. The hpx genetic system for hypoxanthine assimilation as a nitrogen source in Klebsiella pneumoniae: gene organization and transcriptional regulation. J. Bacteriol. 190:7892-7903.[Abstract/Free Full Text]
    25. 25
  25. de Lorenzo, V., M. Herrero, U. Jakubzik, and K. N. Timmis. 1990. Mini-Tn5 transposon derivatives for insertion mutagenesis, promoter probing, and chromosomal insertion of cloned DNA in gram-negative eubacteria. J. Bacteriol. 172:6568-6572.[Abstract/Free Full Text]
    26. 26
  26. Diallinas, G., L. Gorfinkiel, H. N. Arst, Jr., G. Cecchetto, and C. Scazzocchio. 1995. Genetic and molecular characterization of a gene encoding a wide specificity purine permease of Aspergillus nidulans reveals a novel family of transporters conserved in prokaryotes and eukaryotes. J. Biol. Chem. 270:8610-8622.[Abstract/Free Full Text]
    27. 27
  27. Diallinas, G., J. Valdez, V. Sophianopoulou, A. Rosa, and C. Scazzocchio. 1998. Chimeric purine transporters of Aspergillus nidulans define a domain critical for function and specificity conserved in bacterial, plant and metazoan homologues. EMBO J. 17:3827-3837.[CrossRef][Medline]
    28. 28
  28. Eppink, M. H., H. A. Schreuder, and W. J. Van Berkel. 1997. Identification of a novel conserved sequence motif in flavoprotein hydroxylases with a putative dual function in FAD/NAD(P)H binding. Protein Sci. 6:2454-2458.[Medline]
    29. 29
  29. Eppink, M. H., H. A. Schreuder, and W. J. van Berkel. 1998. Interdomain binding of NADPH in p-hydroxybenzoate hydroxylase as suggested by kinetic, crystallographic and modeling studies of histidine 162 and arginine 269 variants. J. Biol. Chem. 273:21031-21039.[Abstract/Free Full Text]
    30. 30
  30. Fellay, R., J. Frey, and H. Krisch. 1987. Interposon mutagenesis of soil and water bacteria: a family of DNA fragments designed for in vitro insertional mutagenesis of Gram-negative bacteria. Gene 52:147-154.[CrossRef][Medline]
    31. 31
  31. Ferraro, D. J., L. Gakhar, and S. Ramaswamy. 2005. Rieske business: structure-function of Rieske non-heme oxygenases. Biochem. Biophys. Res. Commun. 338:175-190.[CrossRef][Medline]
    32. 32
  32. Garzón, A., J. Li, A. Flores, J. Casadesus, and V. Stewart. 1992. Molybdenum cofactor (chlorate-resistant) mutants of Klebsiella pneumoniae M5al can use hypoxanthine as the sole nitrogen source. J. Bacteriol. 174:6298-6302.[Abstract/Free Full Text]
    33. 33
  33. Gatti, D. L., B. A. Palfey, M. S. Lah, B. Entsch, V. Massey, D. P. Ballou, and M. L. Ludwig. 1994. The mobile flavin of 4-OH benzoate hydroxylase. Science 266:110-114.[Abstract/Free Full Text]
    34. 34
  34. Gibson, D. T., and R. E. Parales. 2000. Aromatic hydrocarbon dioxygenases in environmental biotechnology. Curr. Opin. Biotechnol. 11:236-243.[CrossRef][Medline]
    35. 35
  35. Goldman, B. S., J. T. Lin, and V. Stewart. 1994. Identification and structure of the nasR gene encoding a nitrate- and nitrite-responsive positive regulator of nasFEDCBA (nitrate assimilation) operon expression in Klebsiella pneumoniae M5al. J. Bacteriol. 176:5077-5085.[Abstract/Free Full Text]
    36. 36
  36. Gorfinkiel, L., G. Diallinas, and C. Scazzocchio. 1993. Sequence and regulation of the uapA gene encoding a uric acid-xanthine permease in the fungus Aspergillus nidulans. J. Biol. Chem. 268:23376-23381.[Abstract/Free Full Text]
    37. 37
  37. Gournas, C., I. Papageorgiou, and G. Diallinas. 2008. The nucleobase-ascorbate transporter (NAT) family: genomics, evolution, structure-function relationships and physiological role. Mol. Biosyst. 4:404-416.[CrossRef][Medline]
    38. 38
  38. Groisman, E. A., and M. J. Casadaban. 1986. Mini-Mu bacteriophage with plasmid replicons for in vivo cloning and lac gene fusing. J. Bacteriol. 168:357-364.[Abstract/Free Full Text]
    39. 39
  39. Haldimann, A., and B. L. Wanner. 2001. Conditional-replication, integration, excision, and retrieval plasmid-host systems for gene structure-function studies of bacteria. J. Bacteriol. 183:6384-6393.[Abstract/Free Full Text]
    40. 40
  40. Harayama, S., M. Kok, and E. L. Neidle. 1992. Functional and evolutionary relationships among diverse oxygenases. Annu. Rev. Microbiol. 46:565-601.[CrossRef][Medline]
    41. 41
  41. Hennecke, H., and K. T. Shanmugam. 1979. Temperature control of nitrogen fixation in Klebsiella pneumoniae. Arch. Microbiol. 123:259-265.[CrossRef][Medline]
    42. 42
  42. Hille, R. 2002. Molybdenum and tungsten in biology. Trends Biochem. Sci. 27:360-367.[CrossRef][Medline]
    43. 43
  43. Huynen, M. A., and P. Bork. 1998. Measuring genome evolution. Proc. Natl. Acad. Sci. USA 95:5849-5856.[Abstract/Free Full Text]
    44. 44
  44. Jung, D. K., Y. Lee, S. G. Park, B. C. Park, G. H. Kim, and S. Rhee. 2006. Structural and functional analysis of PucM, a hydrolase in the ureide pathway and a member of the transthyretin-related protein family. Proc. Natl. Acad. Sci. USA 103:9790-9795.[Abstract/Free Full Text]
    45. 45
  45. Kahn, K., and P. A. Tipton. 2000. Kinetics and mechanism of allantoin racemization. Bioorg. Chem. 28:62-72.[CrossRef]
    46. 46
  46. Kim, G. J., and H. S. Kim. 1998. Identification of the structural similarity in the functionally related amidohydrolases acting on the cyclic amide ring. Biochem. J. 330:295-302.[Medline]
    47. 47
  47. Kleckner, N., J. Bender, and S. Gottesman. 1991. Uses of transposons with emphasis on Tn10. Methods Enzymol. 204:139-180.[Medline]
    48. 48
  48. Koonin, E. V. 2005. Orthologs, paralogs, and evolutionary genomics. Annu. Rev. Genet. 39:309-338.[CrossRef][Medline]
    49. 49
  49. Lederberg, J., and E. M. Lederberg. 1952. Replica plating and indirect selection of bacterial mutants. J. Bacteriol. 63:399-406.[Free Full Text]
    50. 50
  50. Lee, H. Y., J. H. An, and Y. S. Kim. 2000. Identification and characterization of a novel transcriptional regulator, MatR, for malonate metabolism in Rhizobium leguminosarum bv. trifolii. Eur. J. Biochem. 267:7224-7230.[Medline]
    51. 51
  51. Leimkuhler, S., R. Hodson, G. N. George, and K. V. Rajagopalan. 2003. Recombinant Rhodobacter capsulatus xanthine dehydrogenase, a useful model system for the characterization of protein variants leading to xanthinuria I in humans. J. Biol. Chem. 278:20802-20811.[Abstract/Free Full Text]
    52. 52
  52. Leimkuhler, S., M. Kern, P. S. Solomon, A. G. McEwan, G. Schwarz, R. R. Mendel, and W. Klipp. 1998. Xanthine dehydrogenase from the phototrophic purple bacterium Rhodobacter capsulatus is more similar to its eukaryotic counterparts than to prokaryotic molybdenum enzymes. Mol. Microbiol. 27:853-869.[CrossRef][Medline]
    53. 53
  53. Lin, J. T., B. S. Goldman, and V. Stewart. 1993. Structures of genes nasA and nasB, encoding assimilatory nitrate and nitrite reductases in Klebsiella pneumoniae M5al. J. Bacteriol. 175:2370-2378.[Abstract/Free Full Text]
    54. 54
  54. MacNeil, D., J. Zhu, and W. J. Brill. 1981. Regulation of nitrogen fixation in Klebsiella pneumoniae: isolation and characterization of strains with nif-lac fusions. J. Bacteriol. 145:348-357.[Abstract/Free Full Text]
    55. 55
  55. Magasanik, B. 1996. Regulation of nitrogen utilization, p. 1344-1356. In F. C. Neidhardt, R. Curtiss III, J. L. Ingraham, E. C. C. Lin, K. B. Low, B. Magasanik, W. S. Reznikoff, M. Riley, M. Schaechter, and H. E. Umbarger (ed.), Escherichia coli and Salmonella: cellular and molecular biology, 2nd ed. ASM Press, Washington, DC.
    56. 56
  56. Magasanik, B., and H. R. Bowser. 1955. The degradation of histidine by Aerobacter aerogenes. J. Biol. Chem. 213:571-580.[Free Full Text]
    57. 57
  57. Maloy, S., V. Stewart, and R. Taylor. 1996. Genetic analysis of pathogenic bacteria: a laboratory manual. Cold Spring Harbor Laboratory Press, Plainview, NY.
    58. 58
  58. Marchler-Bauer, A., J. B. Anderson, M. K. Derbyshire, C. DeWeese-Scott, N. R. Gonzales, M. Gwadz, L. Hao, S. He, D. I. Hurwitz, J. D. Jackson, Z. Ke, D. Krylov, C. J. Lanczycki, C. A. Liebert, C. Liu, F. Lu, S. Lu, G. H. Marchler, M. Mullokandov, J. S. Song, N. Thanki, R. A. Yamashita, J. J. Yin, D. Zhang, and S. H. Bryant. 2007. CDD: a conserved domain database for interactive domain family analysis. Nucleic Acids Res. 35:D237-D240.[Abstract/Free Full Text]
    59. 59
  59. Marin, E., L. Nussaume, A. Quesada, M. Gonneau, B. Sotta, P. Hugueney, A. Frey, and A. Marion-Poll. 1996. Molecular identification of zeaxanthin epoxidase of Nicotiana plumbaginifolia, a gene involved in abscisic acid biosynthesis and corresponding to the ABA locus of Arabidopsis thaliana. EMBO J. 15:2331-2342.[Medline]
    60. 60
  60. Mason, J. R., and R. Cammack. 1992. The electron-transport proteins of hydroxylating bacterial dioxygenases. Annu. Rev. Microbiol. 46:277-305.[CrossRef][Medline]
    61. 61
  61. Maynes, J. T., R. G. Yuan, and F. F. Snyder. 2000. Identification, expression, and characterization of Escherichia coli guanine deaminase. J. Bacteriol. 182:4658-4660.[Abstract/Free Full Text]
    62. 62
  62. Merrick, M. J., and R. A. Edwards. 1995. Nitrogen control in bacteria. Microbiol. Rev. 59:604-622.[Abstract/Free Full Text]
    63. 63
  63. Miller, J. H. 1972. Experiments in molecular genetics. Cold Spring Harbor Laboratory, Cold Spring Harbor, NY.
    64. 64
  64. Mobley, H. L., M. D. Island, and R. P. Hausinger. 1995. Molecular biology of microbial ureases. Microbiol. Rev. 59:451-480.[Abstract/Free Full Text]
    65. 65
  65. Montero-Moran, G. M., M. Li, E. Rendon-Huerta, F. Jourdan, D. J. Lowe, A. W. Stumpff-Kane, M. Feig, C. Scazzocchio, and R. P. Hausinger. 2007. Purification and characterization of the FeII- and alpha-ketoglutarate-dependent xanthine hydroxylase from Aspergillus nidulans. Biochemistry 46:5293-5304.[CrossRef][Medline]
    66. 66
  66. Nagatani, H., M. Shimizu, and R. C. Valentine. 1971. The mechanism of ammonia assimilation in nitrogen fixing bacteria. Arch. Mikrobiol. 79:164-175.[CrossRef][Medline]
    67. 67
  67. Nakatsu, C. H., N. A. Straus, and R. C. Wyndham. 1995. The nucleotide sequence of the Tn5271 3-chlorobenzoate 3,4-dioxygenase genes (cbaAB) unites the class IA oxygenases in a single lineage. Microbiology 141:485-495.[Abstract/Free Full Text]
    68. 68
  68. Nam, J. W., H. Nojiri, T. Yoshida, H. Habe, H. Yamane, and T. Omori. 2001. New classification system for oxygenase components involved in ring-hydroxylating oxygenations. Biosci. Biotechnol. Biochem. 65:254-263.[CrossRef][Medline]
    69. 69
  69. Neidhardt, F. C., P. L. Bloch, and D. F. Smith. 1974. Culture medium for enterobacteria. J. Bacteriol. 119:736-747.[Abstract/Free Full Text]
    70. 70
  70. Nishiya, Y., T. Hibi, and J. Oda. 2002. A purification method of the diagnostic enzyme Bacillus uricase using magnetic beads and non-specific protease. Protein Expr. Purif. 25:426-429.[CrossRef][Medline]
    71. 71
  71. Oestreicher, N., and C. Scazzocchio. 1993. Sequence, regulation, and mutational analysis of the gene encoding urate oxidase in Aspergillus nidulans. J. Biol. Chem. 268:23382-23389.[Abstract/Free Full Text]
    72. 72
  72. Okumura, I., and T. Yamamoto. 1978. Enzymic racemization of allantoin. J. Biochem. 84:891-895.[Abstract/Free Full Text]
    73. 73
  73. Parales, R., and S. Resnick. 2006. Aromatic ring hydroxylating dioxygenases, p. 287-340. In J. L. Ramos and R. Levesque (ed.), Pseudomonas,vol. 4 . Springer, Berlin, Germany.
    74. 74
  74. Pateman, J. A., D. J. Cove, B. M. Rever, and D. B. Roberts. 1964. A common co-factor for nitrate reductase and xanthine dehydrogenase which also regulates the synthesis of nitrate reductase. Nature 201:58-60.[CrossRef][Medline]
    75. 75
  75. Price, M. N., P. S. Dehal, and A. P. Arkin. 2007. Orthologous transcription factors in bacteria have different functions and regulate different genes. PLoS Comput. Biol. 3:1739-1750.[Medline]
    76. 76
  76. Ramazzina, I., L. Cendron, C. Folli, R. Berni, D. Monteverdi, G. Zanotti, and R. Percudani. 2008. Logical identification of an allantoinase analog (puuE) recruited from polysaccharide deacetylases. J. Biol. Chem. 283:23295-23304.[Abstract/Free Full Text]
    77. 77
  77. Ramazzina, I., C. Folli, A. Secchi, R. Berni, and R. Percudani. 2006. Completing the uric acid degradation pathway through phylogenetic comparison of whole genomes. Nat. Chem. Biol. 2:144-148.[CrossRef][Medline]
    78. 78
  78. Rigali, S., A. Derouaux, F. Giannotta, and J. Dusart. 2002. Subdivision of the helix-turn-helix GntR family of bacterial regulators in the FadR, HutC, MocR, and YtrA subfamilies. J. Biol. Chem. 277:12507-12515.[Abstract/Free Full Text]
    79. 79
  79. Rouf, M. A., and R. F. Lomprey, Jr. 1968. Degradation of uric acid by certain aerobic bacteria. J. Bacteriol. 96:617-622.[Abstract/Free Full Text]
    80. 80
  80. Saiki, R. K., D. H. Gelfand, S. Stoffel, S. J. Scharf, R. Higuchi, G. T. Horn, K. B. Mullis, and H. A. Erlich. 1988. Primer-directed enzymatic amplification of DNA with a thermostable DNA polymerase. Science 239:487-491.[Abstract/Free Full Text]
    81. 81
  81. Sanderson, K. E., and J. R. Roth. 1988. Linkage map of Salmonella typhimurium, edition VII. Microbiol. Rev. 52:485-532.[Free Full Text]
    82. 82
  82. Sanger, F., S. Nicklen, and A. R. Coulson. 1977. DNA sequencing with chain-terminating inhibitors. Proc. Natl. Acad. Sci. USA 74:5463-5467.[Abstract/Free Full Text]
    83. 83
  83. Scazzocchio, C. 1994. The purine degradation pathway: genetics, biochemistry and regulation, p. 221-257. In S. D. Martinelli and J. R. Kinghorn (ed.), Aspergillus: 50 years on. Elsevier Science B. V., Amsterdam, The Netherlands.
    84. 84
  84. Schultz, A. C., P. Nygaard, and H. H. Saxild. 2001. Functional analysis of 14 genes that constitute the purine catabolic pathway in Bacillus subtilis and evidence for a novel regulon controlled by the PucR transcription activator. J. Bacteriol. 183:3293-3302.[Abstract/Free Full Text]
    85. 85
  85. Sealy-Lewis, H. M., C. Scazzocchio, and S. Lee. 1978. A mutation defective in the xanthine alternative pathway of Aspergillus nidulans: its use to investigate the specificity of uaY mediated induction. Mol. Gen. Genet. 164:303-308.[CrossRef][Medline]
    86. 86
  86. Segura, A., P. V. Bunz, D. A. D'Argenio, and L. N. Ornston. 1999. Genetic analysis of a chromosomal region containing vanA and vanB, genes required for conversion of either ferulate or vanillate to protocatechuate in Acinetobacter. J. Bacteriol. 181:3494-3504.[Abstract/Free Full Text]
    87. 87
  87. Shabarova, Z., and A. Bogdanov. 1994. Advanced organic chemistry of nucleic acids. VCH, Weinheim, Germany.
    88. 88
  88. Skorupski, K., and R. K. Taylor. 1996. Positive selection vectors for allelic exchange. Gene 169:47-52.[CrossRef][Medline]
    89. 89
  89. Stewart, G. S. A. B., S. Lubinsky-Mink, C. G. Jackson, A. Cassel, and J. Kuhn. 1986. pHG165: a pBR322 copy number derivative of pUC8 for cloning and expression. Plasmid 15:172-181.[CrossRef][Medline]
    90. 90
  90. Streicher, S., E. Gurney, and R. C. Valentine. 1971. Transduction of the nitrogen-fixation genes in Klebsiella pneumoniae. Proc. Natl. Acad. Sci. USA 68:1174-1177.[Abstract/Free Full Text]
    91. 91
  91. Syldatk, C., O. May, J. Altenbuchner, R. Mattes, and M. Siemann. 1999. Microbial hydantoinases—industrial enzymes from the origin of life? Appl. Microbiol. Biotechnol. 51:293-309.[CrossRef][Medline]
    92. 92
  92. Tipton, P. A. 2006. Urate to allantoin, specifically (S)-allantoin. Nat. Chem. Biol. 2:124-125.[CrossRef][Medline]
    93. 93
  93. Tropel, D., and J. R. van der Meer. 2004. Bacterial transcriptional regulators for degradation pathways of aromatic compounds. Microbiol. Mol. Biol. Rev. 68:474-500.[Abstract/Free Full Text]
    94. 94
  94. Van der Drift, L., G. D. Vogels, and C. Van der Drift. 1975. Allantoin racemase: a new enzyme from Pseudomonas species. Biochim. Biophys. Acta 391:240-248.[Medline]
    95. 95
  95. van der Geize, R., G. I. Hessels, R. van Gerwen, P. van der Meijden, and L. Dijkhuizen. 2002. Molecular and functional characterization of kshA and kshB, encoding two components of 3-ketosteroid 9{alpha}-hydroxylase, a class IA monooxygenase, in Rhodococcus erythropolis strain SQ1. Mol. Microbiol. 45:1007-1018.[CrossRef][Medline]
    96. 96
  96. Vogels, G. D., and C. Van der Drift. 1976. Degradation of purines and pyrimidines by microorganisms. Bacteriol. Rev. 40:403-468.[Free Full Text]
    97. 97
  97. Wang, Y., Y. Zhang, J. Ding, Y. Liu, J. Wang, and Z. Yu. 2002. Cloming, sequence analysis of imidase gene from Alcaligenes eatrophus and its expression in E. coli. Wei Sheng Wu Xue Bao 42:153-162. (In Mandarin.)[Medline]
    98. 98
  98. Watabe, K., T. Ishikawa, Y. Mukohara, and H. Nakamura. 1992. Identification and sequencing of a gene encoding a hydantoin racemase from the native plasmid of Pseudomonas sp. strain NS671. J. Bacteriol. 174:3461-3466.[Abstract/Free Full Text]
    99. 99
  99. Watabe, K., T. Ishikawa, Y. Mukohara, and H. Nakamura. 1992. Purification and characterization of the hydantoin racemase of Pseudomonas sp. strain NS671 expressed in Escherichia coli. J. Bacteriol. 174:7989-7995.[Abstract/Free Full Text]
    100. 100
  100. Wheelis, M. L., and R. Y. Stanier. 1970. The genetic control of dissimilatory pathways in Pseudomonas putida. Genetics 66:245-266.[Free Full Text]
    101. 101
  101. Williams, N. K., M. K. Manthey, T. W. Hambley, S. I. O'Donoghue, M. Keegan, B. E. Chapman, and R. I. Christopherson. 1995. Catalysis by hamster dihydroorotase: zinc binding, site-directed mutagenesis, and interaction with inhibitors. Biochemistry 34:11344-11352.[CrossRef][Medline]
    102. 102
  102. Wu, Q., and V. Stewart. 1998. NasFED proteins mediate assimilatory nitrate and nitrite transport in Klebsiella oxytoca (pneumoniae) M5al. J. Bacteriol. 180:1311-1322.[Abstract/Free Full Text]
    103. 103
  103. Wu, S. Q., W. Chai, J. T. Lin, and V. Stewart. 1999. General nitrogen regulation of nitrate assimilation regulatory gene nasR expression in Klebsiella oxytoca M5al. J. Bacteriol. 181:7274-7284.[Abstract/Free Full Text]
    104. 104
  104. Yamamoto, K., Y. Kojima, T. Kikuchi, T. Shigyo, K. Sugihara, M. Takashio, and S. Emi. 1996. Nucleotide sequence of the uricase gene from Bacillus sp. TB-90. J. Biochem. 119:80-84.[Abstract/Free Full Text]
    105. 105
  105. Yanofsky, C., and E. S. Lennox. 1959. Transduction and recombination study of linkage relationships among the genes controlling tryptophan synthesis in Escherichia coli. Virology 8:425-447.[CrossRef][Medline]
    106. 106
  106. Yoo, H. S., and T. G. Cooper. 1991. Sequences of two adjacent genes, one (DAL2) encoding allantoicase and another (DCG1) sensitive to nitrogen-catabolite repression in Saccharomyces cerevisiae. Gene 104:55-62.[CrossRef][Medline]
    107. 107
  107. Zalkin, H., and P. Nygaard. 1996. Biosynthesis of purine nucleotides, p. 561-579. In F. C. Neidhardt, R. Curtiss III, J. L. Ingraham, E. C. C. Lin, K. B. Low, B. Magasanik, W. S. Reznikoff, M. Riley, M. Schaechter, and H. E. Umbarger (ed.), Escherichia coli and Salmonella: cellular and molecular biology, 2nd ed. ASM Press, Washington, DC.

Journal of Bacteriology, February 2009, p. 1006-1017, Vol. 191, No. 3
0021-9193/09/$08.00+0     doi:10.1128/JB.01281-08
Copyright © 2009, American Society for Microbiology. All Rights Reserved.




This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Services
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrowReprints and Permissions
Right arrow Copyright Information
Right arrow Books from ASM Press
Right arrow MicrobeWorld
Citing Articles
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Pope, S. D.
Right arrow Articles by Stewart, V.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Pope, S. D.
Right arrow Articles by Stewart, V.

Copyright © 2009 by the American Society for Microbiology.   For an alternate route to Journals.ASM.org, visit: http://intl-journals.asm.org | More Info»