Gene Cluster of Arthrobacter ilicis Rü61a Involved in the Degradation of Quinaldine to Anthranilate

CHARACTERIZATION AND FUNCTIONAL EXPRESSION OF THE QUINALDINE 4-OXIDASE qoxLMS GENES*

Katja Parschat {ddagger} §, Bernhard Hauer ¶, Reinhard Kappl ||, Roswitha Kraft ||, Jürgen Hüttermann || and Susanne Fetzner {ddagger} § **

From the {ddagger}AG Mikrobiologie, Institut für Chemie und Biologie des Meeres, Carl von Ossietzky Universität Oldenburg, D-26111 Oldenburg, the §Institut für Molekulare Mikrobiologie und Biotechnologie, Westfälische Wilhelms-Universität Münster, D-48149 Münster, BASF AG, ZHFD-B009, D-67056 Ludwigshafen, and the ||Fachrichtung Biophysik und Physikalische Grundlagen der Medizin, Universität des Saarlandes, D-66421 HomburgSaar, Germany

Received for publication, February 6, 2003 , and in revised form, April 30, 2003.


    ABSTRACT
 TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 CONCLUSIONS
 REFERENCES
 
A genetic analysis of the anthranilate pathway of quinaldine degradation was performed. A 23-kb region of DNA from Arthrobacter ilicis Rü61a was cloned into the cosmid pVK100. Although Escherichia coli clones containing the recombinant cosmid did not transform quinaldine, cosmids harboring the 23-kb region, or a 10.8-kb stretch of this region, conferred to Pseudomonas putida KT2440 the ability to cometabolically convert quinaldine to anthranilate. The 10.8-kb fragment thus contains the genes coding for quinaldine 4-oxidase (Qox), 1H-4-oxoquinaldine 3-monooxygenase, 1H-3-hydroxy-4-oxoquinaldine 2,4-dioxygenase, and N-acetylanthranilate amidase. The qoxLMS genes coding for the molybdopterin cytosine dinucleotide-(MCD-), FeSI-, FeSII-, and FAD-containing Qox were inserted into the expression vector pJB653, generating pKP1. Qox is the first MCD-containing enzyme to be synthesized in a catalytically fully competent form by a heterologous host, P. putida KT2440 pKP1; the catalytic properties and the UV-visible and EPR spectra of Qox purified from P. putida KT2440 pKP1 were essentially like those of wild-type Qox. This provides a starting point for the construction of protein variants of Qox by site-directed mutagenesis. Downstream of the qoxLMS genes, a putative gene whose deduced amino acid sequence showed 37% similarity to the cofactor-inserting chaperone XdhC was located. Additional open reading frames identified on the 23-kb segment may encode further enzymes (a glutamyl tRNA synthetase, an esterase, two short-chain dehydrogenases/reductases, an ATPase belonging to the AAA family, a 2-hydroxyhepta-2,4-diene-1,7-dioate isomerase/5-oxopent-3-ene-1,2,5-tricarboxylate decarboxylase-like protein, and an enzyme of the mandelate racemase group) and hypothetical proteins involved in transcriptional regulation, and metabolite transport.


    INTRODUCTION
 TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 CONCLUSIONS
 REFERENCES
 
The genetic diversity and flexibility of prokaryotes has led to the evolution of an impressive variety of metabolic pathways to transform or degrade natural as well as numerous xenobiotic compounds. The genes coding for enzymes involved in degradative pathways are often organized as operons and supraoperonic clusters comprising "pathway modules" (14).

N-Heteroaromatic compounds are metabolized and even mineralized by various bacteria (for a review, see Ref. 5 and references therein). Quinaldine (2-methylquinoline) is utilized by Arthrobacter ilicis Rü61a as a source of carbon, nitrogen, and energy; its degradation proceeds via the "anthranilate pathway" (5, 6). The initial step, the hydroxylation of quinaldine in para position to the N-heteroatom, is catalyzed by the inducible enzyme quinaldine 4-oxidase (Qox).1 Qox is a molybdo-iron/sulfur-flavoprotein with an (LMS)2 subunit structure and has been classified to belong to the xanthine oxidase family of molybdenum enzymes (710; for reviews on molybdenum enzymes, see Refs. 1113). Like many other bacterial molybdenum hydroxylases, e.g. quinoline 2-oxidoreductase (Qor) from Pseudomonas putida 86 (14, 15), isoquinoline 1-oxidoreductase (Ior) from Brevundimonas diminuta 7 (16), CO dehydrogenase from Oligotropha carboxidovorans (17), and aldehyde dehydrogenases from Desulfovibrio gigas and Desulfovibrio desulfuricans (1821), Qox contains the molybdopterin cytosine dinucleotide form (MCD) of the molybdenum pyranopterin cofactor (7).

X-ray crystal structures of molybdenum hydroxylases have enabled the identification of amino acid residues that might possibly be involved in substrate positioning and/or catalytic turnover (1723). The catalytic relevance of these residues can be assessed by constructing protein variants carrying amino acid replacements and their biochemical, spectroscopic, and structural characterization. Replacement of a distinct amino acid residue in a protein can be performed by site-directed mutagenesis. However, a prerequisite for such a mutagenesis approach is the availability of a suitable system for the genetic manipulation and for the regulated, functional expression of the genes coding for the enzyme to be investigated. Whereas genes coding for molybdenum hydroxylases containing molybdopterin or the molybdopterin guanine dinucleotide form of the cofactor have been successfully expressed in Escherichia coli (2426), attempts to produce MCD-containing enzymes in E. coli clones failed (27, 28).2 We have recently been working at the construction of expression clones for the synthesis of Qor and Ior. Synthesis of catalytically fully competent Qor was only achieved when using a {Delta}qorMSL mutant of the donor strain P. putida 86 as recipient for the expression plasmid (29). Ior protein showing minor activity was synthesized from the respective expression plasmid when using the Qor-producing strain P. putida 86 as a host, suggesting that accessory gene(s) encoding product(s) essential for the synthesis or assembly of the enzyme is(are) part of the quinoline regulon in P. putida 86 (30).

Here we report on a gene cluster from A. ilicis Rü61a that comprises several genes coding for enzymes of the anthranilate pathway of quinaldine degradation. The amino acid sequences deduced from the qox genes are compared with those of other molybdenum hydroxylases. Due to the broad substrate specificity of Qox, which in addition to different N-heteroaromatic compounds oxidizes aromatic aldehydes (8), sequence comparisons to the crystallized aldehyde oxidoreductases are of special interest. Moreover, we present the functional heterologous expression of the qoxLMS genes in P. putida KT2440 pKP1.


    MATERIALS AND METHODS
 TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 CONCLUSIONS
 REFERENCES
 
Bacterial Strains, Plasmids, and Growth Conditions—The bacterial strains and plasmids used are listed in Table I. A. ilicis Rü61a was grown in mineral salts medium (8) at 30 °C with 0.5 ml/liter quinaldine. E. coli HB101 (31), which served as host strain for recombinant cosmids, and both E. coli DH5{alpha} (32) and E. coli XL1-Blue MRF' (Stratagene), used for cloning procedures with pUC18, were grown in Luria-Bertani (LB) broth (32) at 37 °C. If appropriate, E. coli and P. putida KT2440 cultures contained the following antibiotics: ampicillin (60 and 500 µg/ml for E. coli DH5{alpha} and P. putida KT2440 pKP1, respectively), tetracycline (15 µg/ml), and kanamycin (50 µg/ml). To investigate functional expression of the qox genes in the cosmid clone E. coli HB101 pVK55B/5, it was grown at 37 °C on mineral salts medium containing (per liter) 0.2 g of MgSO4 x 7H2O, 4 g of (NH4)2SO4, 5.25 g of K2HPO4, 2.25 g of KH2PO4, 2.7 mg of FeCl3 x 6H2O, and 8 g of glucose as carbon source. The medium was supplemented with 15 µg/ml of each proline and leucine, and 2 ml/liter vitamin solution (33); 0.1 ml/liter quinaldine was added when the culture reached an optical density (600 nm) of about 0.8. P. putida KT2440 pKP1 was grown in the mineral salt medium described by Tshisuaka et al. (15) with 8 mM benzoate as carbon and energy source and as XylS effector, and with 1 g/liter (NH4)2SO4. As an additional XylS effector, 2 mM 2-methylbenzoate was added at an optical density (600 nm) of 0.8–1.2. To generate biomass for protein purification, cells were grown in two glass fermenters (4-liters each) to which benzoate was added repeatedly. At an optical density (600 nm) of about 3, cells were harvested by centrifugation at 14,000 x g for 15 min at 4 °C. For the preparation of electrocompetent cells, E. coli DH5{alpha}, E. coli XL1-Blue MRF', and P. putida KT2440 were cultured in TB medium (32).


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TABLE I
Bacterial strains and plasmids

 

For transfection of E. coli HB101, 20 ml of LB broth with 0.2 ml of 1 M MgSO4 and 0.2 ml 20% (w/v) maltose were inoculated with a single bacterial colony and grown to an optical density (600 nm) of 0.8. After harvesting the cells by centrifugation at 2000 x g and 4 °C for 10 min, cells were diluted to an optical density (600 nm) of 1 with ice-cold, sterile 10 mM MgSO4.

Analysis of the Degradative Potential of P. putida KT2440 pVK55B/5 and P. putida KT2440 pVK55/11—To determine Qox activity and to identify metabolites of quinaldine catabolism, recombinant cosmids were transferred to P. putida KT2440 (34) by electroporation. The clones P. putida KT2440 pVK55B/5 and P. putida KT2440 pVK55/11 were grown in mineral salt medium (8) with 30 mM succinate, 1 g/liter (NH4)2SO4, and 1 ml/liter vitamin solution (33) at 30 °C. At an optical density (600 nm) of about 0.8, 0.1 ml/liter quinaldine was added. Quinaldine conversion was monitored spectrophotometrically in the culture supernatant. Spectra were compared with those of authentic references diluted in the same medium. Qox activity was measured in the cell free extracts, obtained after cell disruption by sonification, and centrifugation at 48,000 x g for 40 min at 4 °C.

DNA Techniques—Genomic DNA of A. ilicis Rü61a was isolated according to Hopwood et al. (35). Plasmid and cosmid DNA was isolated with the Qiagen Plasmid Mini- and Midi kits, respectively (Qiagen, Hilden, Germany). Gel extraction of DNA fragments for cloning was done with the Nucleo Spin® extraction kit of Macherey-Nagel (Düren, Germany); however, DNA fragments larger than 10 kb were size-fractionated in 0.5% low melting agarose gels and extracted by agarase treatment. DNA restriction, dephosphorylation, and ligation and agarose gel electrophoresis were carried out using standard procedures (32). Electrocompetent cells were generated according to Dower et al. (36) and Iwasaki et al. (37).

Construction of Genomic Libraries—To generate an enriched gene library for A. ilicis Rü61a, genomic DNA, restricted with SmaI, was separated in agarose gels and vacuum-blotted to nylon membranes (parablot NY plus from Macherey-Nagel, Düren, Germany). Fragments in the size of 4–5 kb showing positive hybridization signals with the probe "b-DIG" (see below) were extracted from an agarose gel and ligated to the SmaI-digested, dephosphorylated vector pUC18 (38). E. coli XL1-Blue MRF' transformants were screened by colony blotting and identified by Southern hybridization of SmaI-restricted plasmids, using the probe b-DIG.

For construction of a cosmid library, genomic DNA of A. ilicis Rü61a was partially restricted with HindIII. DNA fragments ranging in size from 15 to 25 kb were extracted from a 0.5% low melting agarose gel and ligated to the HindIII-digested, dephosphorylated cosmid vector pVK100 (39). The cosmids were packaged in vitro into lambda phage particles using a commercial extract (DNA Packaging kit from Roche Applied Science, Mannheim, Germany). The preparation was used to infect E. coli HB101, which was selected for tetracycline resistance (Tcr) and kanamycin sensitivity (Kms). The clone library was screened by colony blotting and hybridization with a probe described below as "1.1 DpnI."

DNA Probes and Hybridization—The oligonucleotide probe b-DIG, which was 5'-end-labeled with a digoxigenin derivative, was a degenerated 29-mer: 5'-TTY ATG CAY CCN TTY CAR TTY ATH ACN CC-3' (following the IUPAC ambiguity code), deduced from the N-terminal amino acid sequence of the medium-sized subunit of Qox (FMH-PFQFITP) (7). Prehybridization for 2 h and hybridization overnight with b-DIG was carried out at 54.5 °C. The membranes were stringently washed for 2 x 15 min in 2x SSC, 0.1% SDS at room temperature, 2 x 15 min in 0.5x SSC, 0.1% SDS at 54.5 °C, and 2 x 15 min in 0.2x SSC, 0.1% SDS at 54.5 °C. Screening an enriched gene library of A. ilicis Rü61a established in the vector pUC18 with b-DIG revealed a clone containing an insert of 4580 bp (pUC55/4.5). Isolation of a 1052-bp DpnI fragment from this insert led to the specific probe 1.1 DpnI, which was used to screen the cosmid clone library. 1.1 DpnI hybridizes with the 5'-terminal half of qoxL (Fig. 1). After prehybridization for 2 h and hybridization overnight with the probe 1.1 DpnI at 68 °C, the membranes were washed twice for 15 min in 2x SSC, 0.1% SDS at room temperature and twice for 15 min in 0.5x SSC, 0.1% SDS at 68 °C. Random primed labeling of the probe 1.1 DpnI using the DIG-High Prime DNA labeling kit (Roche Applied Science, Mannheim, Germany), Southern and colony blotting, hybridization, and colorimetric detection with nitroblue tetrazolium salt and 5-bromo-4-chloro-3-indolyl phosphate were performed following the DIG System User's Guide for Filter Hybridization (40).



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FIG. 1.
23-kb DNA fragment of A. ilicis Rü61a cloned into the cosmid pVK100, yielding pVK55B/5. Genes coding for the subunits of Qox and for Hod are named accordingly. Genes coding for proteins proposed to be involved in quinaldine degradation are in light gray. For a detailed description of the ORFs, see text. The arrows indicate the DNA fragments subcloned for sequencing. Striped boxes give the positions of the probes 1.1 DpnI and b-DIG.

 

Subcloning Procedures—Restriction of the recombinant cosmid pVK55B/5 with HindIII produced two fragments of 12,203 and 10,812 bp, respectively, beside the 23-kb vector. These two fragments were separately inserted into the HindIII cleavage site of pUC18, generating pUC55/12 and pUC55/11. The 10.8-kb fragment was also inserted into the cosmid vector pVK100 (forming pVK55/11) to propagate it in P. putida KT2440. When pUC55/11 was restricted with SmaI, three fragments were generated, the internal 4.58-kb fragment showing the positive signal with probe b-DIG, and two fragments of 2,978 and 3,254 bp, respectively (Fig. 1). The 4.58-kb fragment was cloned into the SmaI site of pUC18 (forming pUC55/4.5). The 2.97-kb fragment and the 3.25-kb fragment were removed from pUC55/11 by SmaI-HindIII and SmaI digestion, respectively. Both fragments were separately cloned into the multiple cloning site of pUC18, yielding pUC55/3 and pUC55/3.2. All pUC derivatives were transferred to E. coli DH5{alpha} by electroporation.

Expression Cloning of qoxLMS Genes—Using the cosmid pVK55B/5 as template, the qoxLMS genes were amplified with Pfu polymerase. The forward primer was chosen to contain the assumed Shine-Dalgarno sequence (in italics) preceding the qox genes, and an EcoRI recognition site (underlined): 5'-ACGCGAATTCGTGACGAAGTTAAGGAGACC-3'; the nucleotides set in boldface are complementary to nucleotides 17,492–17,511 of EMBL accession number AJ537472 [GenBank] . The reverse primer was completely complementary to nucleotides 21,401–21,375 of EMBL accession number AJ537472 [GenBank] : 5'-TTTGGAATGCGCAGTGAGGAGATTTGC-3'. After EcoRI restriction the PCR product was ligated into the EcoRI- and SmaI-restricted plasmid pJB653 (41), generating pKP1. The recipient P. putida KT2440 was transformed by electroporation (36).

DNA Sequencing and Comparative Sequence Analysis—The genes and open reading frames shown in Fig. 1 were deduced from computer-assisted analysis of sequences obtained from single strand sequencing of the inserts of pUC55/12, pUC55/3, pUC55/4.5, and pUC55/3.2; sequences of the qox genes were verified by sequencing both strands. Sequencing was carried out according to the method of Sanger et al. (42) on a sequencer 377 from Applied Biosystems.

Analysis of the DNA sequences were performed with the HUSAR 4.0 program package (EMBL, GENIUSnet, DKFZ Heidelberg, Germany) using the BLAST family of programs (43) for data base searches, GAP for calculating similarities and identities, and ClustalW (44) for calculating multiple alignments. Conserved protein domain sequences and fingerprint motifs were found at Pfam (Sanger Institute, Hinxton, Cambridge, UK) and PRINTS (45). Gene-coding sequences were identified by the program FRAMES. Sequence data on the P. putida KT2440 genome were obtained from The Institute for Genomic Research through the website at www.tigr.org.

Assay for Qox Activity and PAGE—The activity of Qox was assayed spectrophotometrically by measuring the quinaldine-dependent reduction of the artificial electron acceptor iodonitrotetrazolium chloride (INT) as described by de Beyer and Lingens (7). Protein concentrations were estimated by the method of Bradford as modified by Zor and Selinger (46) using bovine serum albumin as standard protein. The Qox preparation used for the determination of Km app and kcat app for quinaldine and for INT showed a specific activity of 5 units/mg.

Non-denaturing PAGE was performed using the high pH discontinuous system according to Hames (47), preparing resolving gels containing a final acrylamide concentration of 7.5% (w/v). SDS-PAGE (48) was used to check the homogeneity of the Qox preparations; resolving gels contained 12.5% (w/v) acrylamide, and 20 mM 1,4-dithio-DL-threitol was added to the samples as reducing agent. Proteins were stained in Coomassie Blue G-250 (0.1% (w/v) in 50% (w/v) trichloroacetic acid). For activity staining of Qox in non-denaturing PA gels, gels were immersed in the same buffer as used for the spectrophotometric assay, containing the substrate quinaldine and INT as electron acceptor.

Purification of Qox from P. putida KT2440 pKP1—22 g of cells (wet weight), suspended in 33 ml of 100 mM Tris-HCl buffer, pH 7, containing 10 µM phenylmethylsulfonyl fluoride, and 1.25 units/ml benzon nuclease, were immersed in an ice bath and disrupted by sonification. Crude extract, obtained by centrifugation at 48,000 x g for 40 min at 4 °C, was loaded onto a 20-ml DEAE-Sepharose CL-6B column (Amersham Biosciences, Freiburg, Germany) that had been equilibrated in 50 mM Tris-HCl buffer, pH 7. Proteins were eluted with a linear gradient from 0 to 1 M NaCl in the equilibration buffer. Fractions showing Qox activity were pooled, and ammonium sulfate was added to a final concentration of 0.75 M. After centrifugation for 30 min at 20,000 x g and 4 °C, the supernatant was loaded onto a 5-ml column containing Phenyl-Sepharose CL-4B (Amersham Biosciences), which had been equilibrated in 100 mM Tris-HCl buffer, pH 7, containing 0.75 M (NH4)2SO4. After washing the column with 85 mM Tris-HCl buffer, pH 7, containing 0.55 M (NH4)2SO4, and elution with a linear gradient from the washing buffer to 50 mM Tris-HCl, pH 7, the active fractions were pooled and applied to an anion exchange column UNOTM-Q1 (Bio-Rad, München, Germany) that had been equilibrated in 50 mM Tris-HCl buffer, pH 7. The proteins were eluted with a linear gradient from 0.15 to 1 M NaCl in the equilibration buffer after washing the column with 0.15 M NaCl in the same buffer. For gel filtration, the enriched Qox was loaded onto the HiLoad 26/60 Superdex 200 prep grade column (Amersham Biosciences). The column was equilibrated and run at 1 ml/min with 50 mM Tris-HCl, pH 7, containing 0.25 M NaCl. Fractions showing Qox activity were pooled, concentrated by ultrafiltration (membrane cut-off 10 kDa), and stored at –80 °C.

Determination of the Nucleotide Moiety of the Molybdenum Cofactor—For this identification, the enzyme was incubated at 95 °C for 10 min in the presence of sulfuric acid (3%, by volume); hydrolysis leads to the release of nucleotides from MCD and FAD. After centrifugation for 10 min at 20,000 x g, the supernatant was analyzed by isocratic HPLC on a Lichrosorb C-RP-18 column (5-µm particle size, 4 x 250 mm) at a flow rate of 1 ml/min with 0.2% acetic acid, 0.5% methanol (by volume) in water as eluent. The compounds were identified by their retention times, as well as their UV spectra (obtained with a photodiode array detector, Waters 996), and by cochromatography with authentic reference compounds (CMP, cytidine, AMP, and GMP).

EPR Spectroscopy and Sample Preparation—The samples were filled into EPR quartz tubes (Wilmad) and immediately frozen in liquid nitrogen within 1 min. Each sample of Qox protein from P. putida KT2440 pKP1 was first reacted with a 5- to 10-fold excess of substrate (quinaldine) and measured at 77 K in a nitrogen finger Dewar flask or between 10 and 65 K in a continuous helium flow ESR900 cryostat (Oxford). In a second step a 60-fold excess of substrate was added to the sample in the quartz tube and measured again. Finally, the sample was exposed to an excess of dithionite (20-fold) for complete reduction. The samples were handled under anaerobic conditions. EPR spectra at X-band frequencies (9.5 GHz) were recorded on a Bruker ESP300 spectrometer. The magnetic field and the microwave frequency were determined with a NMR gaussmeter and a microwave counter, respectively. The modulation amplitude for spectra recording generally was 0.5 millitesla (mT). For measurements at 65 and 77 K, several spectra at different microwave powers (0.2–10 milliwatts) were recorded to avoid saturation broadening. The spectra below 25 K were recorded at about 10-milliwatt microwave power. To improve the signal/noise ratio X-band spectra were accumulated up to 50 times. All spectra were baseline-corrected.

Nucleotide Sequence Accession Number—The nucleotide sequence of the 23,015-bp insert of pVK55B/5, which includes the genes coding for QoxL, QoxM, and QoxS, is deposited in the EMBL Nucleotide Sequence Data base under the accession number AJ537472 [GenBank] .


    RESULTS
 TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 CONCLUSIONS
 REFERENCES
 
Degradative Capacities of P. putida KT2440 Cosmid Clones and Synthesis of Active Quinaldine 4-Oxidase by P. putida KT2440 Cosmid Clones and by P. putida KT2440 pKP1— P. putida KT2440 containing the cosmid pVK100 is able to grow on catechol, but it does not utilize quinaldine or the subsequent intermediates of the anthranilate pathway, namely, 1H-4-oxoquinaldine, 1H-3-hydroxy-4-oxoquinaldine, N-acetylanthranilate, and anthranilate. After transformation of P. putida KT2440 with the recombinant cosmids that hybridized with 1.1 DpnI, five of the resulting P. putida KT2440 pVK100 (Tcr and Kms) clones were able to convert quinaldine to anthranilate cometabolically, suggesting that the genes coding for the enzymes catalyzing the first four steps of the anthranilate pathway are located on the inserts of the cosmids. One of these clones, designated P. putida KT2440 pVK55B/5, was chosen for further investigation. Its cosmid pVK55B/5 harbors the 23-kb insert shown in Fig. 1. P. putida KT2440 pVK55/11, which contains the 10.8-kb region depicted in the lower part of Fig. 1, also shows cometabolic conversion of quinaldine to anthranilate. The specific activity of Qox in cell-free extracts of both P. putida KT2440 pVK55B/5 and P. putida KT2440 pVK55/11 was 0.04 units/mg. In contrast, an E. coli HB101 pVK55B/5 clone did not transform quinaldine, and crude extracts did not show Qox activity. This observation is in accordance with other reports on futile attempts to express genes coding for MCD-containing hydroxylases in E. coli (27, 28).

The expression plasmid pKP1 has been constructed from the broad host range expression vector pJB653 (41) and a fragment comprising the qoxLMS genes preceded by their putative Shine-Dalgarno sequence. Whereas the Pseudomonas cosmid clones mediate expression of the qox genes and further catabolic genes from their own promoters, expression of qoxLMS on pKP1 is regulated by the plasmid-encoded XylS protein that activates the operator region of the Pm promoter of the plasmid. Quinaldine 4-oxidase indeed was produced by P. putida KT2440 pKP1 when growing the strain in the presence of XylS effectors. The specific activity of Qox in crude extracts was 0.06 units/mg of protein. With respect to its electrophoretic mobility in non-denaturing PAGE, the Qox protein produced by the expression clone did not differ from Qox produced by the wild-type strain A. ilicis Rü61a (Fig. 2A).



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FIG. 2.
Polyacrylamide gels of Qox proteins. A, non-denaturing polyacrylamide gel of crude extracts stained for Qox activity. Lane 1, P. putida KT2440 pKP1; lane 2, A. ilicis Rü61a; lane 3, P. putida KT2440 pJB653. B, SDS-PAGE of purified quinaldine 4-oxidase (lane 1) from P. putida KT2440 pKP1 dissociated into its three subunits (82, 35, and 22 kDa), indicating electrophoretic homogeneity of the preparation. Lane 2, standard proteins: {beta}-galactosidase (116 kDa), bovine serum albumin (66.2 kDa), ovalbumin (45 kDa), lactate dehydrogenase (35 kDa), restriction endonuclease Bsp98I (25 kDa), and {beta}-lactoglobulin (18.4 kDa).

 

Purification of Qox from P. putida KT2440 pKP1—Qox was purified in a four-step procedure to near electrophoretic homogeneity (Fig. 2B). The results of the enzyme purification are listed in Table II. The enzyme was purified 85-fold from the crude extract with a yield of 7%. Qox from P. putida KT2440 pKP1 consists of three subunits with molecular masses of about 20, 35, and 80 kDa, as observed for the wild-type enzyme (8).


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TABLE II
Purification of quinaldine 4-oxidase from P. putida KT2440 pKP1 Starting material was 22 g of wet biomass.

 

Properties of the Qox Protein from P. putida KT2440 pKP1— The UV-visible spectrum of Qox purified from P. putida KT2440 pKP1 (Fig. 3) is characteristic for molybdo-iron/sulfur flavoproteins. The ratios E280 nm/E450 nm and E450 nm/E550 nm were 5.6 and 3.3, respectively, indicating stoichiometric amounts of FAD:Fe:S of 1:4:4 in the nearly pure enzyme (49).



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FIG. 3.
UV-visible spectrum of Qox purified from P. putida KT2440 pKP1 (1.2 mg/ml in 50 mM Tris-HCl, pH 7, containing 0.25 M NaCl).

 

HPLC analysis of the non-protein part of the enzyme after acidic hydrolysis indicated the presence of CMP and AMP with retention times of 4.6 and 7.9 min, respectively. GMP or free cytidine were not detected in the sample. The CMP is released from the molybdopterin cytosine dinucleotide cofactor, while AMP derives from the FAD cofactor. The apparent Km values of Qox purified from the expression clone were very similar to those of the wild-type Qox; its kcat values were even higher (Table III).


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TABLE III
Kinetic parameters of the Qox proteins

 

Analysis of Redox-active Centers in Qox from P. putida KT2440 pKP1 by EPR Spectroscopy—EPR spectroscopy is capable to selectively monitor the paramagnetic states of the various redox centers present in the Qox enzymes. In this way typical fingerprint spectra of redox centers (e.g. the rapid or very rapid species of the Mo(V)-cofactor) can be compared between the enzyme preparations to yield information on the presence and integrity of these centers. Here the comparison focuses on Qox of the wild-type specimen A. ilicis Rü61a and the enzyme produced by P. putida KT2440 pKP1. The spectra obtained at 65 K after addition of substrate are shown in Fig. 4. For Qox isolated from P. putida KT2440 pKP1 (Fig. 4, B and C), addition of substrate in small excess leads mainly to the formation of the FAD signal and small traces of the very rapid (vr) signal (Fig. 4B). At higher substrate concentration, very rapid signals comparable to those of wild-type Qox are observed (Fig. 4C). The EPR parameters (g1 = 2.024, g2 = 1.945, and g3 = 1.935) are identical to those of Qox from A. ilicis Rü61a (Fig. 4A) indicating that very similar Mo(V) conformations are prevalent in both enzymes. Only minor traces of a rapid (r) species are detectable. In comparison, the g2 and g3 components of the very rapid signals found in Qor from P. putida 86 and Ior from B. diminuta are clearly different (9). The FAD signals in all Qox samples display a line width of 1.6 mT typical for the anionic (red) FAD radical. The neutral (blue) FAD radical species found in Qor or in xanthine oxidase shows a larger line width of about 1.9 mT. The FAD signal intensity was shown to depend critically on the oxygen status of the Qox preparation (9).



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FIG. 4.
EPR spectra of Qox from A. ilicis Rü61a (A) and P. putida KT2440 pKP1 (B and C) measured at 65 K. The sample in B was exposed to a ~10-fold excess of substrate; in A and C to a 60-fold excess (vr = very rapid species; r = rapid species).

 

When the Qox samples reacted with substrate were measured at lower temperatures (25 K) the broad features of FeS clusters appeared. Interestingly, mainly the signals of FeSII characterized by a larger g anisotropy with respect to FeSI signals could be observed as shown in Fig. 5A for the enzyme from P. putida KT2440 pKP1. Complete reduction with dithionite generally led to a loss of FAD radical signals as well as of Mo(V) species, but the FeS centers were fully converted to their paramagnetic reduced states. The spectrum of fully reduced Qox from P. putida KT2440 pKP1 obtained at 25 K was compared with that of Qox from A. ilicis Rü61a in Fig. 5 (B and C, respectively). It is evident that the axial spectrum of the FeSI center typical for Qox from A. ilicis Rü61a was also present in the enzyme produced by P. putida KT2440 pKP1. The overall spectral pattern was very similar for the enzymes. There were, however, some subtle differences visible. The FeSI1 component was identical, whereas the axial signal FeSIax of Qox from P. putida KT2440 pKP1 displayed a slightly altered line shape caused by a small but unresolved increase in rhombicity of g2 and g3. The most prominent difference concerns the g3 component of FeSII, which was shifted to higher g factors for Qox from P. putida KT2440 pKP1 (Fig. 5B), indicated by the arrow. In addition, its line shape and that of the g1 component were clearly asymmetric (marked by asterisks) as compared with the corresponding features of Qox from A. ilicis Rü61a (Fig. 5, trace C). It is noted that for the partially reduced enzyme (Fig. 5A) the related lines appear more symmetric with the g3 component also slightly shifted to higher g factors. For other related enzymes (Qor and Ior), small shifts of the g factors of FeS clusters have been reported depending on the mode of reduction (9). An indication of a magnetic interaction between the FeS centers as found for Ior was not observed.



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FIG. 5.
EPR spectra of Qox from P. putida KT2440 pKP1 reduced with a 10-fold excess of substrate (A) and fully reduced by subsequent addition of dithionite (B) in comparison to the spectrum of Qox from A. ilicis Rü61a (C). Spectrum A was recorded at 15 K, B and C were recorded at 25 K. The assignment of EPR lines to the two 2Fe-2S centers I and II is indicated.

 

Sequence Analysis of the qox Genes—The genes and potential open reading frames identified on the 23-kb insert of pVK55/5 are presented in Fig. 1 and Table IV. Genes coding for the three subunits of Qox were identified by comparing their N-terminal amino acid sequences deduced from the nucleotide sequence with those determined by Edman degradation (7). The calculated molecular weights for the three peptides were 84,115 for QoxL, 30,608 for QoxM, and 18,539 for QoxS. The calculated mol % G+C contents of qoxL, qoxM, and qoxS were 60.1, 62.2, and 58.8, respectively. For the whole 23-kb fragment, a G+C content of 61.8 mol % was calculated. This value is in good agreement with the mol % G+C content of 61.5 reported for A. ilicis (50). Strain P. putida KT2440, used as a host for expression cloning of the qox genes, shows a similar G+C content of 61.6 mol % (51).


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TABLE IV
Survey of the ORFs identified on a 23-kb fragment of the genome of A. ilicis Rü61a

 

The transcriptional order of the genes coding for the subunits of Qox is 5'-qoxL-M-S-3'. Among the heterotrimeric molybdoenzymes, no conservation in gene arrangement was obvious. Although in many cases the genes coding for the three subunits of these enzymes are transcribed in the order 5'-medium-small-large-3' (28, 5254), other enzymes are known whose genes are arranged in the order 5'-large-small-medium-3' (55) or even with a gap between the gene for the large subunit and the genes for the medium and the small subunit (54). These divergences may lead to the assumption that there is no ancestral common transcriptional unit for these enzymes.

The qoxL gene is 2388 bp in length, coding for a protein of 795 amino acids (aa). A potential ribosome-binding site (AAGGAGA) is located 14 nucleotides upstream of the start codon ATG. 146 nucleotides upstream of the qoxL start codon a putative –35 region was detected (TTGACG), which, however, is not followed by a recognizable –10 region in the usual distance of 16–19 nucleotides (56). QoxL exhibits the well conserved motifs assumed to be involved in binding the pyranopterin cofactor (MoCoI-MoCoV) (19, 57) (Fig. 6A). The glutamate residue Glu736 of QoxL corresponds to Glu869 and Glu869 of the D. gigas and D. desulfuricans aldehyde oxidoreductases (MOP and MOD) (19, 21), respectively, to Glu730 of the B-subunit of xanthine dehydrogenase from Rhodobacter capsulatus (XDHBRc) (23) and to Glu1261 of bovine xanthine dehydrogenase/oxidase (XOb) (22). This glutamate residue is conserved in all enzymes of the xanthine oxidase family; it is assumed to activate the water ligand by proton abstraction and to form a transient bond to the metal during catalysis (19, 21). Residues forming hydrogen bonds to the pyranopterin (MOP: Arg533 and Gln807; MOD: Arg535 and Gln807; XDHBRc: Arg342 and Gln663) and to the water ligand of the molybdenum (MOP: Gly697; MOD: Gly699; XOb: Ala1079; XDHBRc: Ala529; QorL: Ala546) are also conserved in QoxL (Arg362, Gln671, and Gly526) (Fig. 6A).



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FIG. 6.
Comparison of sequence stretches of different molybdenum hydroxylases. A, conserved motifs presumed to be involved in binding the pyranopterin cofactor or in positioning of the substrate. Sequence stretches of QoxL are compared with corresponding sequences of the large subunit of quinoline 2-oxidoreductase from P. putida 86 (QorL) (accession number CAA66828 [GenBank] ), the large subunit of CO dehydrogenase from O. carboxidovorans (CoxL) (accession number P19919 [GenBank] ), aldehyde oxidoreductases from D. gigas (MOP) (accession number A57429 [GenBank] ), and D. desulfuricans (MOD) (accession number CAB64929 [GenBank] ), xanthine oxidase from cow's milk (XOb) (accession number P80457 [GenBank] ), and xanthine dehydrogenase from R. capsulatus (XDHBRc) (accession number CAA04470 [GenBank] ). Residues that are completely conserved are in shaded boxes. Residues supposed to be involved in coordination of the molybdenum pyranopterin cofactor and in catalysis, which are discussed in the text, are marked with an asterisk below the sequence. Based on our sequencing results, residues 465 and 466 in QorL were corrected (EV, instead of DC reported previously (28)/accession number CAA66828 [GenBank] ). B, sequence stretches of the medium sized subunit QoxM comprising the loop-forming FAD-binding sites as described for the vanillyl-alcohol oxidases, compared with the corresponding segments of QorM, CoxM, XDHARc, and XOb (for an explanation of the abbreviations, see A); NdhM: medium sized subunit of the nicotine dehydrogenase from A. nicotinovorans (accession number CAD47954 [GenBank] ).

 

QoxM starts at position 19,901 (EMBL accession number AJ537472 [GenBank] ); its start codon ATG overlaps with the TGA stop codon of qoxL. QoxM ends after 873 nucleotides with the stop codon TAG, coding for a protein of 290 aa. Thirteen nucleotides upstream of the start codon a putative ribosome-binding site was identified (AAGGAGA).

The motifs 30AGGQT34 and 109TIGG112, which correspond to the typical loop-forming FAD-binding sites of the vanillyl-alcohol oxidase family (58), were identified in QoxM, indicating that it harbors the FAD cofactor (Fig. 6B). The motif TIGG is described to create a pocket for the adenosine and to contact the pyrophosphate moiety of the FAD molecule in XDH from Rhodobacter capsulatus (23). In XDHARc, Thr206 of the N-terminal motif (first motif in Fig. 6B) as well as the double glycine of this motif also interact with the pyrophosphate (23). In QoxM, Gln33 corresponds to this residue; the medium-sized subunits of Qor from P. putida 86 (28) and of nicotine dehydrogenase from Arthrobacter nicotinovorans (59) likewise contain a glutamine residue in this position. Tyrosine 193 of the medium-sized subunit of the CO dehydrogenase from O. carboxidovorans (CoxM), shielding the central part of the FAD isoalloxazine ring from the solvent (17), is not conserved in QoxM (Pro192 by sequence comparison), but in some other molybdenum hydroxylases.

QoxS starts two nucleotides downstream of the stop codon of qoxM with the start codon ATG at position 20,776 and ends with the stop codon TGA at position 21,300. Fourteen nucleotides upstream a putative ribosome-binding site was detected (AAGGGAG). QoxS, which consists of 174 aa, shows the two cysteine rich motifs strictly conserved in the prokaryotic molybdenum hydroxylases, which probably bind the two different [2Fe2S] clusters (60). The N-terminal motif following the sequence 40CX4CGXCX11C60 is homologous to the plant-type ferredoxin signature pattern; the second motif 117CGXCX31CXC154 has a binding motif that is typical for molybdenum hydroxylases. Whereas the latter motif is presumed to bind the FeSI center, which is "proximal" to the molybdenum cofactor, the first motif may bind the "distal" FeSII center (57).

Sequence analysis of the qoxLMS Flanking Regions—Within the 23-kb region, fifteen putative ORFs were identified upstream of the qoxLMS gene cluster, and one ORF downstream of qoxS (Fig. 1 and Table IV). The hypothetical protein encoded by the nearly complete ORF 1, which starts 574 nucleotides downstream of qoxS but lacks some C-terminal residues, is related to chaperone-like proteins presumed to be involved in recruitment of the molybdenum cofactor. The deduced protein shows 37% similarity to XdhC from R. capsulatus, which has been proven to be required for insertion of MPT into xanthine dehydrogenase (61). 41% similarity was found to PucA, which is part of the purine catabolic gene cluster of Bacillus subtilis; PucA and XdhC show 22% aa identity in their C-terminal half (62).

1958 nucleotides upstream of the qoxL start codon the gene coding for 1H-3-hydroxy-4-oxoquinaldine-2,4-dioxygenase (Hod) was identified, which is transcribed in the opposite direction to the qox gene cluster. Hod catalyzes the third step in the anthranilate pathway of quinaldine degradation, namely the cleavage of 1H-3-hydroxy-4-oxoquinaldine to N-acetylanthranilate and carbon monoxide; it belongs to a unique group of oxygenases without requirement for cofactors or metal ions (6366).

ORF 2, which starts 1257 nucleotides upstream of the hod start codon, is identical to ORF 491 reported by Betz et al. (67). The 42,216-Da hypothetical protein (388 aa) deduced from this ORF shows high similarities to monooxygenases belonging to the single component flavoproteins, e.g. the 2-methyl-3-hydroxypyridine-5-carboxylic acid oxygenase from Pseudomonas sp. MA1 (68) and a salicylate hydroxylase of Sphingomonas sp. (accession number BAA19150 [GenBank] ). Its N-terminal region includes a typical ADP-binding site that may bind the ADP portion of an FAD cofactor (69, 70). Upstream of the ORF 2 start codon a putative –35 region (TTGACG), identical to that found in front of qoxL, was identified, which is followed by a putative –10 region (TATATAA) in a distance of 16 bp. No possible –35/–10 promoter regions upstream of the hod gene were obvious, so we may speculate that ORF 2 and the hod gene form an operon.

Seven nucleotides downstream of the hod stop codon starts ORF 4, which codes for a hypothetical protein of Mr 32,002. The 293-aa protein exhibits 49% similarity to a putative protein from Pseudomons sp. CA10 (accession number BAB32459 [GenBank] .1), which is assumed to belong to a family of esterases/lipases/thioesterases. The ORF 4 amino acid segment spanning positions 59–158 is related to a domain of type-B carboxylesterases (Pfam signature PF00135), although the described consensus pattern for this family (Prosite accession number PDOC00112) is not completely conserved. On the 10.8-kb fragment of pVK55/11, which confers to P. putida KT2440 the ability to convert quinaldine to anthranilate, ORF 4 is the sole hypothetical gene supposed to code for a hydrolase, therefore, it apparently acts as an N-acetylanthranilate amide hydrolase in quinaldine degradation. No potential –35/–10 promoter region was identified upstream of ORF 4, so the putative operon comprising ORF 2 and hod might also involve ORF 4.

67 nucleotides downstream of ORF 4 starts ORF 5 coding for a putative protein of 423 aa with Mr 43,695. For the ORF 5 gene product, a transmembrane protein belonging to the family of general substrate transporters is predicted (InterPro entry IPR005828). These proteins share a common structural feature of 12 transmembrane {alpha}-helices with a cytoplasmatic loop after the sixth transmembrane {alpha}-helix. Prediction of transmembrane regions of the ORF 5 protein by the program SOSUI (71) indeed revealed twelve {alpha}-helices corresponding to those of the transport proteins; the conserved feature RXGR(R/K), which is proposed to form a {beta}-bend that links two {alpha}-helices (72), was found at positions 80–84 (RWGLK). This bend is located between {alpha}-helix two and {alpha}-helix three as suggested by the secondary structure prediction (using SOSUI (71)). However, contrary to representative proteins of this family, the RXGR(R/K) motif is not duplicated in the deduced ORF 5 protein. A hydropathy plot (73) of the ORF 5 protein also suggested that it is a transmembrane protein. One subfamily of the general substrate transporter family comprises the benzoate transporters, including, e.g. the 4-hydroxybenzoate transporters PcaK from P. putida PRS2000 and Acinetobacter calcoaceticus ADP1 (74, 75), the cis,cis-muconate transporter MucK from A. calcoaceticus ADP1 (76), and the benzoate transporter BenK from the same strain (77, 78). All these transporters are located in or close to operons coding for the enzymes and proteins of the respective aromatic degradation pathway. The proximity of ORF 5 to other genes involved in quinaldine degradation may suggest that this putative transmembrane protein could be involved in transport of quinaldine or of one of the metabolites across the cell membrane.

ORF 6 encodes a protein of Mr 32,349 composed of 295 aa, which shows the 5-element fingerprint for glutamyl-tRNA synthetases (PRINTS accession number PR00987). The segments 16HVGN19 and 233RLAKR237 of the deduced protein correspond to the conserved motifs HIGH and KMSKS (or KLSKR), respectively, that in the class I aminoacyl-tRNA synthetases are involved in ATP binding (79, 80).

Twenty-five nucleotides downstream of the ORF 6 stop codon starts ORF 7 coding for a hypothetical protein of 372 aa, which has 87% similarity and 82% identity to the ethyl chrysanthemate-hydrolyzing esterase from Arthrobacter globiformis SC-6–98-28 (81). Like this esterase the putative ORF 7 protein shows significant similarities to {beta}-lactamases, other esterases, and the DD-carboxypeptidase from Streptomyces R61 (82). The serine and the lysine residues in the N-terminal motif SXXK, suggested to be involved in substrate binding and proton transfer during catalysis in {beta}-lactamases and {beta}-lactam-sensitive enzymes (82, 83), are conserved in the ORF 7 protein in positions 59 and 62, respectively.

When comparing the putative protein encoded by ORF 8 of 203 aa (deduced Mr of 21,472) to proteins in databases, a high degree of similarity to transcriptional regulators of the TetR family was detected. Analysis of the ORF 8 protein predicted a helix-turn-helix (HTH) motif at the N-terminal part comprising residues 32–53. Moreover, the signature pattern (Prosite accession number PS01081) described to surround this motif in proteins of the TetR family is nearly completely conserved.

The hypothetical proteins encoded by ORF 9 and ORF 14 with a length of 279 and 253 aa, respectively, are assumed to be members of the large and diverse superfamily of short chain dehydrogenases/reductases (SDR), which is defined by a common folding pattern rather than by function (84). They can be assigned to the "classical family" of SDRs according to the classification of Kallberg et al. (84). The members of this family catalyze NAD(P)(H)-dependent oxidation/reduction reactions on a wide spectrum of substrates, e.g. alcohols, steroids, or aromatic compounds. The binding sites for the cosubstrates NAD(H) or NADP(H) in the N-terminal part of the SDR proteins are represented by the conserved pattern GXXXGXG; this motif is indeed found in the sequences deduced from ORF 9 and ORF 14. The motif YXXXK, comprising the conserved tyrosine and lysine residues involved in catalysis, is also present in both amino acid sequences. The conserved residues Arg39 in the ORF 9 protein and Asp42 in the ORF 14 protein suggest that the cosubstrate of the former protein is NADP(H), whereas the latter may utilize NAD(H) (84).

ORF 10, which is located downstream of ORF 9, probably codes for a protein of 178 aa with Mr 20,331 and shows up to 40% similarity to the N-terminal part of (p)ppGpp 3'-pyrophosphohydrolases belonging to the RelA/SpoT family. This N-terminal part of the RelA/SpoT family enzymes includes a so-called HD domain (Pfam accession number PF01966) (85) that contains highly conserved histidine and aspartate residues; these are presumed to coordinate divalent cations. It has been proposed that all the enzymes harboring an HD domain catalyze divalent-cation-dependent phosphohydrolase reactions (85). The HD superfamily comprises not only multidomain enzymes like the RelA/SpoT proteins, but also "stand-alone" proteins that essentially consist of a single HD domain (85). Because about half of the ORF 10 protein (residues 30–120 out of 178 aa) resemble the HD domain, it may belong to the latter group.

ORFs 11 and 12 are transcribed in the reverse direction to ORF 10 and code for putative proteins of Mr 43,306 and 37,043, respectively. The N-terminal part (aa 18–306) of the ORF 11 protein (414 aa) shares conserved sequence stretches with putative membrane proteins of unknown function grouped by sequence homology in an orthologous group (COG4292; available at www.ncbi.nlm.nih.gov/COG/new/release/cow.cgi?cog=COG4292). Prediction of putative transmembrane regions using the program SOSUI (70) indicated 10 membrane-spanning helices for the ORF 11 protein. BLAST searches for the ORF 12 protein showed high similarities to ATPases associated with diverse cellular activities (AAA ATPases) from a variety of organisms. Members of the AAA ATPases family usually share a ring-shaped oligomeric structure and represent a type of molecular chaperone. They are involved either in protein folding and assembly, in protein transport, or in disassembly or degradation of proteins (86, 87). The hypothetical ORF 12 protein consists of 336 aa and harbors one AAA domain where the two strictly conserved Walker motifs A and B are located. Walker motif A or "P-loop" is found in position 105–113 and Walker motif B is in position 160–167 of the ORF 12 protein. These two sequences are responsible for ATP binding and hydrolysis (86, 88). Additionally, the consensus pattern for AAA ATPases (Prosite accession number PS00674) was detected in the central part of the AAA domain in position 204–224.

The 376-aa hypothetical protein encoded by ORF 13 with Mr 40,153 is located upstream of ORF 12 in the opposite orientation. Beside the 43% similarity to the C-terminal part of an ATP/GTP-binding protein (ORF 666) of A. nicotinovorans (accession number CAD47885 [GenBank] ), the amino acid sequence shows slight similarities to different putative proteins from Streptomyces coelicolor A3 (2, 89), but no hypothetical function can be deduced for these proteins.

ORF 15 codes for a hypothetical protein with Mr 31,820. The 302-aa protein may be identified as a member of the family of fumaracetoacetate hydrolases; this family also includes the bifunctional enzyme 2-hydroxyhepta-2,4-diene-1,7-dioate (HHDD) isomerase/5-oxopent-3-ene-1,2,5-tricarboxylate (OPET) decarboxylase, which catalyzes two steps in the catabolic pathway of homoprotocatechuate (90). The ORF 15 protein indeed exhibits a high degree of similarity to known HHDD isomerases/OPET decarboxylases (see Table IV).

ORF 16 codes for a putative protein composed of 434 aa with Mr 47,172 that shows significant similarity to members of the enolase superfamily. These enzymes catalyze diverse overall reactions, which, however, are initiated by a common step, i.e. abstraction of the {alpha}-proton of a carboxylic acid to form an enolic intermediate (91, 92). The members of the enolase superfamily were assigned to three subgroups by Babbitt et al. (91): (i) mandelate racemases (MR); (ii) muconate lactonizing enzymes (MLE), and (iii) enolases. All members of the superfamily show a two-domain structure, one common N-terminal domain and one catalytic "TIM barrel" domain that contains residues acting as general acid/base catalysts and residues that coordinate one (MR, MLE) or two (enolase) metal ion(s). Both domains were predicted for the ORF 16 protein; residues 4–137 and 174–424 resemble the common N-terminal domain (Pfam accession number PF02746) and the TIM barrel domain (Pfam accession number PF01188), respectively. Residues that in the catalytic domain of MR and MLE from P. putida coordinate a metal ion are also conserved in the ORF 16 protein (MR, accession number P11444 [GenBank] : Asp195, Glu221, and Glu247; MLE, accession number AAA66202 [GenBank] : Asp198, Glu224, and Asp249; ORF 16 protein: Asp248, Glu274, and Glu301) (91, 92). Moreover, amino acid residues described to be involved in catalysis of MR and MLE were also identified in the ORF 16 protein. In the MR reaction, Lys166 acts as the (S)-specific general base that abstracts the {alpha}-proton from (S)-mandelate. One carboxylate oxygen of the resulting enolic intermediate is thought to be stabilized by functioning as a ligand to the Mg2+ and by binding to Lys164, whereas the second carboxylate oxygen is stabilized through a strong hydrogen bond to Glu317. In MLE, these functions probably are taken over by the residues Lys169, Lys167, and Glu327 (91). Sequence alignment with the ORF 16 protein revealed Lys220, Lys218, and Glu382 as homologous residues. Because enolases are devoid of the KXK motif of MR and MLR (91), it is unlikely that the ORF16 protein belongs to the enolase subgroup. In the reverse MR reaction, His297 is the (R)-specific catalytic base that abstracts the {alpha}-proton from (R)-mandelate, assisted by Asp270 (91, 93). These two residues are not conserved in MLE, but they actually were found in the ORF 16 protein in positions His351 and Asp324. Based on the pattern of conserved residues, we suggest that the gene product of ORF 16 belongs to the mandelate racemase subgroup of enzymes. This subgroup apart from MR includes D-galactonate dehydratase, D-glucarate dehydratase, L-rhamnoate dehydratase, and some reading frames with unassigned functions (91, 93).


    DISCUSSION
 TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 CONCLUSIONS
 REFERENCES
 
The qoxLMS genes from A. ilicis Rü61a were expressed in a Pseudomonas host, yielding catalytically competent Qox protein. Active quinaldine 4-oxidase was produced by cosmid clones containing the whole 23-kb insert, or a 10.8-kb fragment (Fig. 1), and by P. putida KT2440 pKP1, where expression of the qox genes is regulated by the Pm promoter, which in turn is induced by benzoate-activated XylS protein. The specific activity for Qox in the crude extract of P. putida KT2440 pKP1 was as high as that found in crude extracts of the wild-type strain A. ilicis Rü61a. The specific activity of Qox purified from P. putida KT2440 pKP1 even exceeded that determined by Stephan et al. (8) for preparations of wild-type Qox. The biochemical and spectroscopic properties of the Qox protein purified from P. putida KT2440 pKP1 were similar to those of the wild-type enzyme. EPR spectroscopy revealed identical spectral patterns of the FAD-radical signal and the catalytically competent very rapid species in both enzymes (Fig. 4), indicating that the environment of the centers and particularly the mode of substrate/product binding at the site of the Mo(V) cofactor are very similar. The sole appearance of the FeSII signals at lower temperatures in the enzyme of P. putida KT2440 pKP1 are in accordance with the findings of redox and rapid freeze experiments of Qox from A. ilicis Rü61a for which the redox potential of FeSII was 180 mV higher than that of FeSI. Consequently, the FeSII signal was observed first under single turnover conditions in kinetic EPR experiments (10). Hence, it is concluded that the difference of the redox potentials of both FeS centers in Qox from P. putida KT2440 pKP1 should be similar. The axial type FeSI signal is not found for other proteins of the xanthine oxidase family. Its presence in Qox from the expression clone points to a conserved structural environment of this cluster, which also has been shown to be proximal to the molybdenum cofactor (10, 57, 94). The spectral signature of the FeSII center of Qox of P. putida KT2440 pKP1 is similar to that of A. ilicis Rü61a but shows some slight differences in g factor and line shape. These changes seem to be related to the simultaneous presence of substrate and dithionite reduced clusters. Because this cluster presumably is located close to the surface of the domain, it is more susceptible to solvent effects and generally shows a relaxation behavior different from FeSI cluster (10, 11, 57, 95). On the whole, our results suggest that the Qox protein produced by the expression clone is identical to the wild-type enzyme, implying that the host strain P. putida KT2440 is able to provide all the accessory functions that are required for the assembly of this complex enzyme. Qox is in fact the first MCD-containing enzyme to be synthesized in a catalytically fully competent form by a heterologous host. There is no overexpression of the qox-LMS genes in P. putida KT2440 pKP1; however, overexpression was not our primary goal, because we intended to construct a system that allows the genetic manipulation of the qox genes by mutagenesis approaches and the production of protein variants of Qox.

Assembly of the Fe/S protein (QoxS) and the flavoprotein (QoxM) is thought to involve ubiquitously conserved pathways. However, biosynthesis of the MCD form of the molybdenum pyranopterin cofactor and its insertion into QoxL requires not only proteins involved in molybdenum uptake and MPT biosynthesis but also a tailoring enzyme forming MCD from MPT and maybe even a specific chaperone for MCD insertion (for reviews on molybdate uptake and biosynthesis of the molybdenum cofactor, see Refs. 96 and 97). Sequence comparisons of known genes of the moa, moe, and mod operons of E. coli with the P. putida KT2440 genome revealed corresponding sequences with significant similarities, suggesting that strain KT2440 is able to synthesize the MPT cofactor. The successful expression of fully active Qox and the release of CMP upon acidic hydrolysis of the enzyme indicated that P. putida KT2440 is also able to provide the MCD cofactor, and moreover, to insert it into the maturing Qox protein. It is remarkable to note that the gene product of ORF 1 (Fig. 1), which due to its similarity to the XdhC protein (61) is thought to be involved in cofactor insertion during Qox assembly in the wild-type strain, is not required for formation of functional Qox by the P. putida expression clone.

On the basis of x-ray crystal structure analyses of aldehyde oxidoreductases from D. gigas (MOP) and D. desulfuricans (MOD), CO dehydrogenase from O. carboxidovorans (Cox), xanthine oxidase/dehydrogenase from bovine milk (XOb) and xanthine dehydrogenase from R. capsulatus (XDHBRc), amino acid residues responsible for coordination of the molybdenum pyranopterin cofactor as well as residues probably involved in the catalytic mechanism were described (1723). Additionally, mutagenesis of xanthine dehydrogenase from Emericella (formerly Aspergillus) nidulans (HxA) defined residues contributing to substrate specificity and substrate positioning at the active site (98). Thus, comparison of Qox with these proteins may provide a first view of the molecular features of the active site of Qox. Non-polar residues described to give access to the active site in MOP and MOD (MOP: Phe425, Phe494, Leu497, and Leu626; MOD: Phe427, Phe496, Leu499, and Leu628) have counterparts in the QoxL sequence (Gly253, Phe323, Ile327, and Leu459). Although the preferred substrates of the aldehyde oxidases (i.e. aliphatic aldehydes) differ from those of Qox (N-heteroaromatic compounds and aromatic aldehydes) with regard to their molecular structure, a hydrophobic entrance to the active site appears to be necessary for both types of enzymes.

Glutamate 833 of HxA is conserved among xanthine dehydrogenases (XOb: Glu802; XDHBRc: Glu232) and is proposed to influence the substrate specificity of the enzymes. MOP and MOD have a phenylalanine in this position (Phe425 and Phe427, respectively). Alignments of MOP, MOD, QoxL, and QorL revealed Gly253 for QoxL and Ala259 for QorL as corresponding residues (Fig. 6A); thus small residues seem to be necessary at this position for the accommodation of the bicyclic azaarenes in the active site.

Of particular interest is a conserved arginine found in the vicinity of the MOP and MOD substrate binding sites (MOP: Arg501; MOD: Arg503); these residues correspond to Arg911 in the E. nidulans xanthine dehydrogenase HxA. This amino acid has been proposed to be involved in positioning the substrate relative to the molybdenum center, because mutations of Arg911 of HxA yielding Gly911 or Gln911 changed the hydroxylation position of 2-hydroxypurine from C-8 to C-6 (97). In QoxL and QorL this arginine is not conserved, it is exchanged to hydrophobic residues in both enzymes (QoxL: Ile330; QorL: Val339; Fig. 6A).

The regulation of catabolic operons coding for enzymes of aromatic degradation pathways has been the subject of intensive research (for a review, see Ref. 99). The regulatory proteins involved were found to belong to a variety of families, among them the family of tetracycline repressors (TetR), to which the protein encoded by ORF 8 seems to be related. Genes and operons regulated by TetR-like repressors encode proteins with very diverse functions (100103). An example is agmatine utilization by Pseudomonas aeruginosa, which is catalyzed by the products of aguA and aguB, transcribed in one operon and negatively regulated by AguR (104). The calculated Mr of the ORF 8 protein of 21,472 resembles the average Mr of 21,000 to 25,000 reported for the TetR proteins. In the case of the archetypal TetR repressor, its DNA-binding site is located between the tetR gene and the vicinal gene tetA that is transcribed in the opposite direction. The tetA gene, which codes for an energy-dependent tetracycline/Mg2+ antiporter, is the target of the TetR-mediated regulation (105). Such an arrangement has been found for other tetR-like genes and their targets. However, we have not been able to identify possible DNA-binding sites for the regulator between ORFs 8 and 9 or in front of qoxL or ORF 2. Nevertheless, involvement of the putative ORF 8 protein in regulation of transcription of these genes cannot be excluded, because the DNA sequences recognized by the HTH motifs appear to be very diverse among different TetR-like regulators.


    CONCLUSIONS
 TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 CONCLUSIONS
 REFERENCES
 
Identification, sequencing, cloning, and functional, heterologous expression of the qoxLMS gene cluster has been achieved in this work. The available expression system will allow the genetic manipulation of the qox genes by site-directed mutagenesis. Of special interest is the investigation of the functional role of the glutamate residue Glu736, which corresponds to the glutamate residues strictly conserved in enzymes of the xanthine dehydrogenase family and which has been predicted to be involved in the catalytic mechanism. Residues thought to be involved in substrate positioning are also important targets for mutagenesis studies; their alteration might give us an idea about the molecular basis of substrate specificity and regioselectivity of hydroxylation. The identification of a number of putative genes that might be functionally related to quinaldine oxidation may open up further investigations on the anthranilate pathway and its regulation.


    FOOTNOTES
 
The nucleotide sequence(s) reported in this paper has been submitted to the GenBankTM/EBI Data Bank with accession number(s) AJ537472 [GenBank] .

* This work was supported by the Deutsche Forschungsgemeinschaft (Grant FE 383/4-4) and the Fonds der Chemischen Industrie. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

** To whom correspondence should be addressed. Tel.: 49-251-833-9824; Fax: 49-251-833-8388; E-mail: fetzner{at}uni-muenster.de.

1 The abbreviations used are: Qox, quinaldine 4-oxidase; Qor, quinoline 2-oxidoreductase; Ior, isoquinoline 1-oxidoreductase; MCD, molybdopterin cytosine dinucleotide; Tc, tetracycline; Km, kanamycin; INT, iodonitrotetrazolium chloride; HPLC, high pressure liquid chromatography; aa, amino acid(s); MOP, aldehyde oxidoreductase from D. gigas; MOD, aldehyde oxidoreductase from D. desulfuricans; XDHRc, xanthine dehydrogenase from R. capsulatus; XOb, xanthine oxidase from cow's milk; ORF, open reading frame; Hod, 1H-3-hydroxy-4-oxoquinaldine 2,4-dioxygenase; TetR, tetracycline repressor; HTH, helix-turn-helix; SDR, short chain dehydrogenase/reductase; COG, cluster of orthologous groups; HHDD, 2-hydroxyhepta-2,4-diene-1,7-dioate; OPET, 5-oxopent-3-ene-1,2,5-tricarboxylate; MR, mandelate racemase; MLE, muconate lactonizing enzyme; MPT, molybdopterin cofactor; HxA, xanthine dehydrogenase from E. nidulans; Ap, ampicillin. Back

2 K. Parschat, U. Frerichs-Deeken, and S. Fetzner, unpublished results. Back


    ACKNOWLEDGMENTS
 
We thank Renate Gahl-Jan{beta}en for excellent technical assistance and Sonja Sielker for the kinetic data for Qox from A. ilicis Rü61a. Sequence data of the P. putida KT2440 genome were obtained from The Institute for Genomic Research through the website at www.tigr.org. We gratefully acknowledge the financial support of the Deutsche Forschungsgemeinschaft and the Fonds der Chemischen Industrie.



    REFERENCES
 TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 CONCLUSIONS
 REFERENCES
 

  1. van der Meer, J. R., de Vos, W. M., Harayama, S., and Zehnder, A. J. (1992) Microbiol. Rev. 56, 677–694[Medline] [Order article via Infotrieve]
  2. Wyndham, R. C., Cashore, A. E., Nakatsu, C. H., and Peel, M. C. (1994) Biodegradation 5, 323–342[Medline] [Order article via Infotrieve]
  3. Romine, M. F., Stillwell, L. C., Wong, K. K., Thurston, S. J., Sisk, E. C., Sensen, C., Gaasterland, T., Fredrickson, J. K., and Saffer, J. D. (1999) J. Bacteriol. 181, 1585–1602[Abstract/Free Full Text]
  4. Parke, D., D'Argenio, D. A., and Ornston, L. N. (2000) J. Bacteriol. 182, 257–263[Free Full Text]
  5. Fetzner, S. (1998) Appl. Microbiol. Biotechnol. 49, 237–250[CrossRef]
  6. Hund, H.-K., de Beyer, A., and Lingens, F. (1990) Biol. Chem. Hoppe-Seyler 371, 1005–1008[Medline] [Order article via Infotrieve]
  7. de Beyer, A., and Lingens, F. (1993) Biol. Chem. Hoppe-Seyler 374, 101–110[Medline] [Order article via Infotrieve]
  8. Stephan, I., Tshisuaka, B., Fetzner, S., and Lingens, F. (1996) Eur. J. Biochem. 236, 155–162[Abstract]
  9. Canne, C., Stephan, I., Finsterbusch, J., Lingens, F., Kappl, R., Fetzner, S., and Hüttermann, J. (1997) Biochemistry 36, 9780–9790[CrossRef][Medline] [Order article via Infotrieve]
  10. Canne, C., Lowe, D. J., Fetzner, S., Adams, B., Smith, A. T., Kappl, R., Bray, R. C., and Hüttermann, J. (1999) Biochemistry 38, 14077–14087[Medline] [Order article via Infotrieve]
  11. Hille, R. (1996) Chem. Rev. 96, 2757–2816[CrossRef][Medline] [Order article via Infotrieve]
  12. Hille, R., Rátey, J., Bartlewski-Hof, U., Reichenbecher, W., and Schink, B. (1999) FEMS Microbiol. Rev. 22, 489–501[CrossRef]
  13. Hille, R. (2002) in Molybdenum and Tungsten: Their Roles in Biological Processes (Sigel, A., and Sigel, H., eds) Vol. 39, pp. 187–226, Marcel Dekker, New York
  14. Hettrich, D., Peschke, B., Tshisuaka, B., and Lingens, F. (1991) Biol. Chem. Hoppe-Seyler 372, 513–517[Medline] [Order article via Infotrieve]
  15. Tshisuaka, B., Kappl, R., Hüttermann, J., and Lingens, F. (1993) Biochemistry 32, 12928–12934[Medline] [Order article via Infotrieve]
  16. Lehmann, M., Tshisuaka, B., Fetzner, S., Röger, P., and Lingens, F. (1994) J. Biol. Chem. 269, 11254–11260[Abstract/Free Full Text]
  17. Dobbek, H., Gremer, L., Meyer, O., and Huber, R. (1999) Proc. Natl. Acad. Sci. U. S. A. 96, 8884–8889[Abstract/Free Full Text]
  18. Romão, M. J., Archer, M., Moura, I., Moura, J. J. G., LeGall, J., Engh, R., Schneider, M., Hof, P., and Huber, R. (1995) Science 270, 1170–1176[Abstract]
  19. Romão, M. J., and Huber, R. (1998) Structure and Bonding 90, 69–95
  20. Rebelo, J. M., Dias, J. M., Huber, R., Moura, J. J. G., and Romão, M. J. (2001) J. Biol. Inorg. Chem. 6, 791–800[CrossRef][Medline] [Order article via Infotrieve]
  21. Rebelo, J., Macieira, S., Dias, J. M., Huber, R., Ascenso, C. S., Rusnak, F., Moura, J. J. G., Moura, I., and Romão, M. J. (2000) J. Mol. Biol. 297, 135–146[CrossRef][Medline] [Order article via Infotrieve]
  22. Enroth, C., Eger, B. T., Okamoto, K., Nishino, T., Nishino, T., and Pai, E. F. (2000) Proc. Natl. Acad. Sci. U. S. A. 97, 10723–10728[Abstract/Free Full Text]
  23. Truglio, J. J., Theis, K., Leimkühler, S., Rappa, R., Rajagopalan, K. V., Kisker, C. (2002) Structure (Camb.) 10, 115–125[CrossRef][Medline] [Order article via Infotrieve]
  24. Pollock, V. V., and Barber, M. J. (1997) J. Biol. Chem. 272, 3355–3362[Abstract/Free Full Text]
  25. Garrett, R. M., and Rajagopalan, K. V. (1994) J. Biol. Chem. 269, 272–276[Abstract/Free Full Text]
  26. Temple, C. A., and Rajagopalan, K. V. (2000) J. Biol. Chem. 275, 40202–40210[Abstract/Free Full Text]
  27. Black, G. W., Lyons, C. M., Williams, E., Colby, J., Kehoe, M., and O'Reilly, C. (1990) FEMS Microbiol. Lett. 58, 249–254[Medline] [Order article via Infotrieve]
  28. Bläse, M., Bruntner, C., Tshisuaka, B., Fetzner, S., and Lingens, F. (1996) J. Biol. Chem. 271, 23068–23079[Abstract/Free Full Text]
  29. Frerichs-Deeken, U., Goldenstedt, B., Gahl-Jan{beta}en, R., Kappl, R., Hüttermann, J., and Fetzner, S. (2003) Eur. J. Biochem. 270, 1567–1577[Abstract/Free Full Text]
  30. Israel, I., Sohni, M., and Fetzner, S. (2002) FEMS Microbiol. Lett. 210, 123–127[Medline] [Order article via Infotrieve]
  31. Boyer, H. W., and Roulland-Dussoix, D. (1969) J. Mol. Biol. 41, 459–472[Medline] [Order article via Infotrieve]
  32. Sambrook, J., Fritsch, E. F., and Maniatis, T. (1989) Molecular Cloning: A Laboratory Manual, Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY
  33. Röger, P., Erben, A., and Lingens, F. (1990) Biol. Chem. Hoppe-Seyler. 371, 511–513[Medline] [Order article via Infotrieve]
  34. Bagdasarian, M., Lurz, R., Rückert, B., Franklin, F. C. H., Bagdasarian, M. M., Frey, J., and Timmis, K. N. (1981) Gene (Amst.) 16, 237–247[CrossRef][Medline] [Order article via Infotrieve]
  35. Hopwood, D. A., Bibb, M. J., Chater, K. F., Kieser, T., Bruton, C. J., Kieser, H. M., Lydiate, C. P., Ward, J. M., and Schrempf, H. (1985) Genetic Manipulation of Streptomyces, a Laboratory Manual, The John Innes Foundation, Norwich, UK
  36. Dower, W. J., Miller, J. F., and Ragsdale, C. W. (1988) Nucleic Acids Res. 16, 6127–6145[Abstract]
  37. Iwasaki K., Uchiyama, H., Yagi, O., Kurabayashi, T., Ishizuku, K., and Takamura, Y. (1994) Biosci. Biotechnol. Biochem. 58, 851–854[Medline] [Order article via Infotrieve]
  38. Vieira, J., and Messing, J. (1982) Gene (Amst.) 19, 259–268[CrossRef][Medline] [Order article via Infotrieve]
  39. Knauf, V. C., and Nester E. W. (1982) Plasmid 8, 45–54[Medline] [Order article via Infotrieve]
  40. Roche molecular biochemicals (1995) The DIG System User's Guide for Filter Hybridization (ISBN 3-88630-200-8) Boehringer Mannheim GmbH, Mannheim, Germany
  41. Blatny, J. M., Brautaset, T., Winther-Larsen, H. C., Haugan, K., and Valla, S. (1997) Appl. Environ. Microbiol. 63, 370–379[Abstract]
  42. Sanger, F., Nicklen, S., and Coulson, A. R. (1977) Proc. Natl. Acad. Sci. U. S. A. 74, 5463–5467[Abstract]
  43. Altschul, S. F., Madden, T. L., Schäffer, A. A., Zhang, J., Zhang, Z., Miller, W., Lipman, D. J. (1997) Nucleic Acids Res. 25, 3389–3402[Abstract/Free Full Text]
  44. Higgins, D. G., Thompson, J. D., and Gibson, T. J. (1996) Methods Enzymol. 266, 383–402[Medline] [Order article via Infotrieve]
  45. Attwood, T. K., Beck, M. E., Flower, D. R., Scordis, P., and Selley, J. N. (1998) Nucleic Acids Res. 26, 304–308[Abstract/Free Full Text]
  46. Zor, T., and Selinger, Z. (1996) Anal. Biochem. 236, 302–308[CrossRef][Medline] [Order article via Infotrieve]
  47. Hames, B. D. (1990) in Gel Electrophoresis of Proteins: a Practical Approach (Hames, B. D., and Rickwood, D., eds) 2nd Ed., pp. 1–147, IRL Press at Oxford University Press
  48. Laemmli, U. K. (1970) Nature 227, 680–685[Medline] [Order article via Infotrieve]
  49. Coughlan, M. P. (1980) in Molybdenum and Molybdenum-containing Enzymes (Coughlan, M. P., ed) pp. 119–185, Pergamon Press, Oxford
  50. Jones, D., and Collins, M. D. (1986) in Bergey's Manual of Systematic Bacteriology (Sneath, P. H. A., Mair, N. S., Sharp, M. E., and Holt, J. G., eds) Vol. 2, pp. 1261–1434, Williams & Wilkins, Baltimore
  51. Nelson, K. E., Weinel, C., Paulsen, I. T., Dodson, R. J., Hilbert, H., Martins dos Santos, V. A., Fouts, D. E., Gill, S. R., Pop, M., Holmes, M., Brinkac, L., Beanan, M., DeBoy, R. T., Daugherty, S., Kolonay, J., Madupu, R., Nelson, W., White, O., Peterson, J., Khouri, H., Hance, I., Chris Lee P., Holtzapple, E., Scanlan, D., Tran, K., Moazzez, A., Utterback, T., Rizzo, M., Lee, K., Kosack, D., Moestl, D., Wedler, H., Lauber, J., Stjepandic, D., Hoheisel, J., Straetz, M., Heim, S., Kiewitz, C., Eisen, J., Timmis, K. N., Dusterhoft, A., Tummler, B., and Fraser, C. M. (2002) Environ. Microbiol. 4, 799–808[CrossRef][Medline] [Order article via Infotrieve]
  52. Schübel, U., Kraut, M., Mörsdorf, G., and Meyer, O. (1995) J. Bacteriol. 177, 2197–2203[Abstract]
  53. Pearson, D. M., O'Reilly, C., Colby, J., and Black, G. W. (1994) Biochim. Biophys. Acta 1188, 432–438[Medline] [Order article via Infotrieve]
  54. Baitsch, D., Sandu, C., Brandsch, R., and Igloi, G. L. (2001) J. Bacteriol. 183, 5262–5267[Abstract/Free Full Text]
  55. Bilous, P. T., Cole, S. T., Anderson, W. F., and Weiner, J. H. (1988) Mol. Microbiol. 2, 785–795[Medline] [Order article via Infotrieve]
  56. Reznikoff, W. S., Siegele, D. A., Cowing, D. W., and Gross, C. A. (1985) Annu. Rev. Genet. 19, 355–387[CrossRef][Medline] [Order article via Infotrieve]
  57. Kappl, R., Hüttermann, J., and Fetzner, S. (2002) in Molybdenum and Tungsten: Their Roles in Biological Processes (Sigel, A., and Sigel, H., eds) Vol. 39, pp. 481–537, Marcel Dekker, New York
  58. Fraaije, M. W., Van Berkel, W. J. H., Benen, J. A. E., Visser, J., and Mattevi, A. (1998) Trends Biochem. Sci. 23, 206–207[CrossRef][Medline] [Order article via Infotrieve]
  59. Schenk, S., Hoelz, A., Krau{beta}, B., and Decker, K. (1998) J. Mol. Biol. 284, 1323–1339[CrossRef][Medline] [Order article via Infotrieve]
  60. Hughes, R. K., Doyle, W. A., Chovnick, A., Whittle, J. R., Burke, J. F., and Bray, R. C. (1992) Biochem. J. 285, 507–513[Medline] [Order article via Infotrieve]
  61. Leimkühler, S., and Klipp, W. (1999) J. Bacteriol. 181, 2745–2751[Abstract/Free Full Text]
  62. Schultz, A. C., Nygaard, P., and Saxild, H. H. (2001) J. Bacteriol. 183, 3293–3302[Abstract/Free Full Text]
  63. Fischer, F., Künne, S., and Fetzner, S. (1999) J. Bacteriol. 181, 5725–5733[Abstract/Free Full Text]
  64. Fischer, F., and Fetzner, S. (2000) FEMS Microbiol. Lett. 190, 21–27[CrossRef][Medline] [Order article via Infotrieve]
  65. Fetzner, S. (2000) Naturwissenschaften 87, 59–69[CrossRef][Medline] [Order article via Infotrieve]
  66. Fetzner, S. (2002) Appl. Microbiol. Biotechnol. 60, 243–257[CrossRef][Medline] [Order article via Infotrieve]
  67. Betz, A., Facey, S. J., Hauer, B., Tshisuaka, B., and Lingens, F. (2000) J. Basic Microbiol. 40, 7–23[Medline] [Order article via Infotrieve]
  68. Chaiyen, P., Ballou, D. P., and Massey, V. (1997) Proc. Natl. Acad. Sci. U. S. A. 94, 7233–7238[Abstract/Free Full Text]
  69. Wierenga, R. K., Terpstra, P., and Hol, W. G. J. (1986) J. Mol. Biol. 187, 101–107[Medline] [Order article via Infotrieve]
  70. Möller, W., and Amos, R. (1985) FEBS Lett. 186, 1–7[CrossRef][Medline] [Order article via Infotrieve]
  71. Hirokawa, T., Boon-Chieng, S., and Mitaku, S. (1998) Bioinformatics 14, 378–379[Abstract]
  72. Maiden, M. C. J., Davis, E. O., Baldwin, S. A., Moore, D. C. M., and Henderson, P. J. F. (1987) Nature 325, 641–643[CrossRef][Medline] [Order article via Infotrieve]
  73. Kyte, J., and Doolittle, R. F. (1982) J. Mol. Biol. 157, 105–132[Medline] [Order article via Infotrieve]
  74. Harwood, C. S., Nichols, N. N., Kim, M.-K., Ditty, J. L., and Parales, R. E. (1994) J. Bacteriol. 176, 6479–6488[Abstract]
  75. Kowalchuk, G. A., Hartnett, G. B., Benson, A., Houghton, J. E., Ngai, K.-L., and Ornston, L. N. (1994) Gene (Amst.) 146, 23–30[CrossRef][Medline] [Order article via Infotrieve]
  76. Williams, P. A., and Shaw, L. E. (1997) J. Bacteriol. 179, 5935–5942[Abstract]
  77. Collier, L. S., Nichols, N. N., and Neidle, E. L. (1997) J. Bacteriol. 179, 5943–5946[Abstract]
  78. Clark, T. J., Momany, C., and Neidle, E. L. (2002) Microbiology 148, 1213–1223[Abstract/Free Full Text]
  79. Perona, J. J., Rould, M. A., and Steitz, T. A. (1993) Biochemistry 32, 8758–8771[Medline] [Order article via Infotrieve]
  80. Freist, W., Gauss, D. H., Söll, D., and Lapointe, J. (1997) Biol. Chem. 378, 1313–1329[Medline] [Order article via Infotrieve]
  81. Nishizawa, M., Shimizu, M., Ohkawa, H., and Kanaoka, M. (1995) Appl. Environ. Microbiol. 61, 3208–3215[Abstract]
  82. Kelly, J. A., Knox, J. R., Moews, P. C., Hite, G. J., Bartolone, J. B., Zhao, H., Joris, B., Frère, J.-M., and Ghuysen, J.-M. (1985) J. Biol. Chem. 260, 6449–6458[Abstract/Free Full Text]
  83. Herzberg, O., and Moult, J. (1987) Science 236, 694–701[Medline] [Order article via Infotrieve]
  84. Kallberg, Y., Oppermann, U., Jörnvall, H., and Persson, B. (2002) Eur. J. Biochem. 269, 4409–4417[Abstract/Free Full Text]
  85. Aravind, L., and Koonin, E. V. (1998) Trends Biochem. Sci. 23, 469–472[CrossRef][Medline] [Order article via Infotrieve]
  86. Ogura, T., and Wilkinson, A. J. (2001) Genes to Cells 6, 575–597[Abstract/Free Full Text]
  87. Neuwald, A. F., Aravind, L., Spouge, J. L., and Koonin, E. V. (1999) Genome Res. 9, 27–43[Abstract/Free Full Text]
  88. Koonin, E. V. (1993) J. Mol. Biol. 229, 1165–1174[CrossRef][Medline] [Order article via Infotrieve]
  89. Bentley, S. D., Chater, K. F., Cerdeno-Tarraga, A. M., Challis, G. L., Thomson, N. R., James, K. D., Harris, D. E., Quail, M. A., Kieser, H., Harper, D., Bateman, A., Brown, S., Chandra, G., Chen, C. W., Collins, M., Cronin, A., Fraser, A., Goble, A., Hidalgo, J., Hornsby, T., Howarth, S., Huang, C. H., Kieser, T., Larke, L., Murphy, L., Oliver, K., O'Neil, S., Rabbinowitsch, E., Rajandream, M. A., Rutherford, K., Rutter, S., Seeger, K., Saunders, D., Sharp, S., Squares, R., Squares, S., Taylor, K., Warren, T., Wietzorrek, A., Woodward, J., Barrell, B. G., Parkhill, J., and Hopwood, D. A. (2002) Nature 417, 141–147[CrossRef][Medline] [Order article via Infotrieve]
  90. Roper, D. I., and Cooper, R. A. (1993) Eur. J. Biochem. 217, 575–580[Abstract]
  91. Babbitt, P. C., Hasson, M. S., Wedekind, J. E., Palmer, D. R., Barrett, W. C., Reed, G. H., Rayment, I., Ringe, D., Kenyon, G. L., and Gerlt, J. A. (1996) Biochemistry 35, 16489–16501[CrossRef][Medline] [Order article via Infotrieve]
  92. Hasson, M. S., Schlichting, I., Moulai, J., Taylor, K., Barrett, W., Kenyon, G. L., Babbitt, P. C., Gerlt, J. A., Petsko, G. A., and Ringe, D. (1998) Proc. Natl. Acad. Sci. U. S. A. 95, 10396–10401[Abstract/Free Full Text]
  93. Gerlt, J. A., and Babbitt, P. C. (2001) Annu. Rev. Biochem. 70, 209–246[CrossRef][Medline] [Order article via Infotrieve]
  94. Andrade, S. L., Brondino, C. D., Feio, M. J., Moura, I., and Moura, J. J. (2000) Eur. J. Biochem. 267, 2054–2061[Abstract/Free Full Text]
  95. Romão, M. J., Knäblein, J., Huber, R., and Moura, J. J. (1997) Prog. Biophys. Mol. Biol. 68, 121–144[CrossRef][Medline] [Order article via Infotrieve]
  96. Pau, R. N., and Lawson, D. M. (2002) in Molybdenum and Tungsten: Their Roles in Biological Processes (Sigel, A., and Sigel, H., eds) Vol. 39, pp. 31–74, Marcel Dekker, New York
  97. Mendel, R. R., and Schwarz, G. (2002) in Molybdenum and Tungsten: Their Roles in Biological Processes (Sigel, A., and Sigel, H., eds) Vol. 39, pp. 317–368, Marcel Dekker, New York
  98. Glatigny, A., Hof, P., Romão, M. J., Huber, R., and Scazzocchio, C. (1998) J. Mol. Biol. 278, 431–438[CrossRef][Medline] [Order article via Infotrieve]
  99. Díaz, E., and Prieto, M. A. (2000) Curr. Opin. Biotechnol. 11, 467–475[CrossRef][Medline] [Order article via Infotrieve]
  100. Lucas, C. E., Balthazar, J. T., Hagman, K. E., and Shafer, W. M. (1997) J. Bacteriol. 179, 4123–4128[Abstract]
  101. Schumacher, M. A., Miller, M. C., Grkovic, S., Brown, M. H., Skurray, R. A., and Brennan, R. G. (2002) EMBO J. 21, 1210–1218[Abstract/Free Full Text]
  102. Kojic, M., Aguilar, C., and Venturi, V. (2002) J. Bacteriol. 184, 2324–2330[Abstract/Free Full Text]
  103. Eaton, R. W. (1997) J. Bacteriol. 179, 3171–3180[Abstract]
  104. Nakada, Y., Jiang, Y., Nishijyo, T., Itoh, Y., and Lu, C.-D. (2001) J. Bacteriol. 183, 6517–6524[Abstract/Free Full Text]
  105. Kisker, C., Hinrichs, W., Tovar, K., Hillen, W., and Saenger, W. (1995) J. Mol. Biol. 247, 260–280[CrossRef][Medline] [Order article via Infotrieve]
  106. Dembek, G., Rommel, T., Lingens, F., and Höke, H. (1989) FEBS Lett. 246, 113–116