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Applied and Environmental Microbiology, November 2008, p. 6703-6708, Vol. 74, No. 21
0099-2240/08/$08.00+0     doi:10.1128/AEM.00386-08
Copyright © 2008, American Society for Microbiology. All Rights Reserved.

Subfunctionality of Hydride Transferases of the Old Yellow Enzyme Family of Flavoproteins of Pseudomonas putida

Pieter van Dillewijn, Rolf-Michael Wittich, Antonio Caballero, and Juan-Luis Ramos^*

Departamento de Protección Ambiental, Estación del Zaidín, Consejo Superior de Investigaciones Científicas, Apdo. Correos 419, E-18008 Granada, Spain

Received 15 February 2008/ Accepted 3 September 2008

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To investigate potential complementary activities of multiple enzymes belonging to the same family within a single microorganism, we chose a set of Old Yellow Enzyme (OYE) homologs of Pseudomonas putida. The physiological function of these enzymes is not well established; however, an activity associated with OYE family members from different microorganisms is their ability to reduce nitroaromatic compounds. Using an in silico approach, we identified six OYE homologs in P. putida KT2440. Each gene was subcloned into an expression vector, and each corresponding gene product was purified to homogeneity prior to in vitro analysis for its catalytic activity against 2,4,6-trinitrotoluene (TNT). One of the enzymes, called XenD, lacked in vitro activity, whereas the other five enzymes demonstrated type I hydride transferase activity and reduced the nitro groups of TNT to hydroxylaminodinitrotoluene derivatives. XenB has the additional ability to reduce the aromatic ring of TNT to produce Meisenheimer complexes, defined as type II hydride transferase activity. The condensations of the primary products of type I and type II hydride transferases react with each other to yield diarylamines and nitrite; the latter can be further reduced to ammonium and serves as a nitrogen source for microorganisms in vivo.

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Gene duplication has long been recognized as a major source of new genes and functions (19). Until recently, it was assumed that gene duplication served as a way to evolve new functions (30). However, several recent case studies and comparisons of genome content have suggested that most new genes have no new functions (25). Instead, paralogous gene pairs are often subfunctionalized and may have overlapping activities. The duplication of genes encoding enzymes that participate in different cellular processes is of great interest for the evolution of a given microorganism because the two copies can evolve different functions that may be overlapping or not, allowing the complementation and maintenance of the original function.

We have centered our attention on the OYE family of proteins. This family of flavoproteins is based on the Old Yellow Enzyme (OYE), which was first isolated from brewers' bottom yeast (33). Members of this family of proteins have been found in other yeasts, bacteria, plants, and nematodes (35). Analyses of the three-dimensional structures of those OYE homologs, which have been crystallized, show that all consist of one or more monomers with a β/-barrel structure and a flavin mononucleotide (FMN) prosthetic group. In spite of detailed structural knowledge, the physiological role of most of these enzymes remains unknown (4, 35).

An activity often associated with OYE enzyme members is that they can reduce nitroaromatic compounds (10, 14, 36). These compounds are susceptible to the reduction of the nitro side groups to yield hydroxylamino groups. This type of activity is also called nitroreductase activity (6, 7, 8, 9, 20, 28, 34) and will be referred to here as type I hydride transferase activity. Other OYE members have been described to additionally catalyze the nucleophilic attack on the aromatic ring of 2,4,6-trinitrotoluene (TNT) by hydride ions to produce Meisenheimer monohydride and dihydride complexes (13, 23, 24, 29, 32, 37). These OYE member enzymes are classified as type II hydride transferases (31).

A search for OYE family enzymes in databases revealed that many bacteria harbor multiple OYE homologs, and to date, it appears that P. putida strains harbor the largest number of different OYE homologs (22). The reasons why P. putida KT2440 would harbor such a large number of these types of enzymes remains unknown but could be related to their physiological roles in the cell, as was previously proposed for the four different OYE homologs in Shewanella oneidensis (4).

A feature found in P. putida and also in other TNT-assimilating strains is the difficulty in obtaining mutants deficient in TNT utilization, which suggests that multiple proteins capable of releasing nitrogen from TNT are present in these microorganisms. Since OYE homologs are able to attack nitroaromatic compounds, we decided to analyze the OYE enzymes of P. putida KT2440 in detail to determine their potential contribution to the metabolism of TNT.

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Chemicals, microorganisms, and culture conditions.
TNT was obtained from the Unión Española de Explosivos (Madrid, Spain). 2-Hydroxylamino-4,6-dinitrotoluene (2HADNT), 4-hydroxylamino-2,6-dinitrotoluene (4HADNT), 2-amino-4,6-dinitrotoluene, 4-amino-2,6-dinitrotoluene, and glycerol trinitrate were obtained from AccuStandard (New Haven, CT). N-Ethylmaleimide and 2-cyclohexen-1-one were obtained from Fluka (Buchs, Switzerland). Standards of diarylamines and diarylhydroxylamines were prepared as described previously (37). Pseudomonas putida KT2440 was grown routinely at 30°C in M9 minimal medium supplemented with 0.5% (wt/vol) glucose or 10 mM sodium benzoate as a carbon source (1).

Analytical methods.
TNT, its transformation products, and nitrite were analyzed by high-performance liquid chromatography (HPLC) using a Waters-Alliance chromatograph equipped with a photodiode array detector (model 2996) and a 5-µm C₈ reversed-phase column (Novapak, 150 by 3.9 mm; Waters S.A., Barcelona, Spain) and temperature settings at 25°C. To detect TNT, hydroxylamine, and amino derivatives of TNT, 25-µl samples were run for 20 min in a 35% (vol/vol) methanol-water solution at an 0.85-ml/min flow rate with the detector set at 230 nm. To separate nitrite and the isoforms of TNT hydride adducts and the diaryl adducts from each other, samples (25 µl) were run with 5 mM tetrabutylammonium phosphate (Fluka) at pH 7.0 and acetonitrile as the respective mobile phases at a flow rate of 0.85 ml/min, and the detector was set at 210, 230, and 450 nm. The method consisted of a linear gradient from 0% to 25% (vol/vol) acetonitrile during 20 min and from 25% to 62% acetonitrile for another 23 min. The column was then reequilibrated with 100% 5 mM tetrabutylammonium solution for another 9 min.

Purification of OYE family members.
The xenA to xenF genes of P. putida were amplified using appropriate primers (sequences will be provided upon request) with BamHI and HindIII sites and P. putida KT2440 chromosomal DNA as a template. After digestion with these restriction enzymes, the PCR products were ligated into the pET28b(+) vector (Novagen) previously digested with BamHI-HindIII. The resulting plasmids contained each coding sequence in frame with a DNA sequence encoding a His₆ tag at its 3' end, which resulted in a hexahistidine tail at the C termini of the proteins, which was expected to have no effect on activity based on the three-dimensional structures of homologous proteins. Cloned genes were sequenced to confirm that no point mutations were introduced during the amplification step. For protein-His₆ purification, each respective pET28b(+) derivative plasmid was transformed into Escherichia coli BL21(pLysS) cells (Novagen). The cells were grown in several 1-liter batches at 30°C in 2x YT culture medium with 50 µg/ml kanamycin to an A₆₆₀ of between 0.5 and 0.7 and then induced with 1 mM IPTG (isopropyl-β-D-thiogalactopyranoside). Cells were harvested after overnight induction at 20°C; resuspended in a solution containing 25 mM sodium phosphate buffer (pH 7.0), 0.5 M NaCl, 5% (vol/vol) glycerol, and protease inhibitor cocktail (Complete; Roche); and disrupted by treatment with 20 µg/ml of lysozyme and French press (1000 lb/in²). Following centrifugation at 20,000 x g for 30 min, the protein was found predominantly (more than 80%) in the soluble fraction. Each His₆-tagged protein was purified by nickel affinity chromatography and eluted with a continuous imidazole gradient. Peak fractions were pooled and dialyzed against the gel filtration buffer (25 mM sodium phosphate buffer [pH 7.5], 0.1 mM EDTA, 500 mM NaCl, 10% [vol/vol] glycerol, and 1 µM FMN). Protein concentrations were determined using the Bio-Rad protein assay kit, and molecular size was determined by sodium dodecyl sulfate-polyacrylamide gel electrophoresis as well as gel filtration after calibration using a 12- to 200-kDa molecular marker kit (Sigma).

Enzyme assays.
All in vitro enzymatic assays were performed at 25°C using equivalent amounts of the corresponding enzyme (based on flavin concentrations), NAD(P)H, and substrates in a total volume of 1 ml in 50 mM phosphate buffer (pH 7.0). The reductase activity of each enzyme was determined by measuring the rate of NAD(P)H consumption in a spectrophotometer (UV-160A; Shimadzu) at 340 nm. Specific activity is given as µmol NAD(P)H consumed per minute and per mg of protein. The kinetic parameters of each enzyme were determined in assays with appropriate amounts of NADPH and TNT or other substrates. K_m and V_max values were determined from Lineweaver-Burk plots generated with Excel software (Microsoft).

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Identification of OYE family members in P. putida.
A search was performed within the genome of P. putida KT2440 (22) for sequences similar to those of OYE family members with hydride transferase activity using the yeast OYE enzyme as a query. This search revealed the presence of six open reading frames (Table 1), all of which could potentially code for an OYE family protein and corresponded to the amino acid sequences listed in GenBank for P. putida KT2440, under accession numbers AAN66878, AAN66545, AAN68098, AAN68101, AAN67100, and AAN68781. In the primary sequence of these proteins, we found conserved OYE active-site residues and a number of residues involved in interactions with the FMN cofactor (5, 12). The sequence under GenBank accession number AAN66878 showed 98% identity with xenobiotic reductase A of P. putida II-B (3) and 99.7% identity with XenA of P. putida 86 (15). The amino acid sequence under GenBank accession number AAN66545 has 88% identity with xenobiotic reductase B of Pseudomonas fluorescens I-C (3) (Table 1) and 100% identity with XenB purified from P. putida JLR11 (37). The high similarity of these two proteins to previously named xenobiotic reductases led us to maintain the same nomenclature for these proteins of P. putida KT2440. Furthermore, we kept this nomenclature for the remaining OYE homologs in P. putida KT2440, so they were called XenC, XenD, XenE, and XenF. In general, this set of enzymes exhibited relatively high identities among each other (Table 1). The genes encoding these xenobiotic reductases are dispersed throughout the genome of P. putida, with the exception of xenC and xenD, which are located within 3 kb of each other.

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TABLE 1. Percent identity at the amino acid level between members of the OYE family of flavoproteins identified in P. putida KT2440 and XenB of P. fluorescens, which shows type II hydride transferase activity with TNT

Purification and characterization of OYE-type proteins from P. putida KT2440.
All six genes were cloned into the pET28b(+) vector for the induced expression of terminal histidine-tagged products. The proteins were purified to near homogeneity by nickel affinity chromatography and gel filtration. The molecular masses of the six His₆-tagged proteins were in the range of 40.9 to 42.6 kDa, as determined by gel filtration. All of the enzymes were monomers in solution. Spectral analysis of each protein showed that all enzymes except XenD (which was colorless) shared an absorption peak at 470 nm, a characteristic of the FMN cofactor, which is responsible for the yellow color of each enzyme (data not shown).

In vitro substrate range of P. putida KT2440 OYE family enzymes.
In vitro reactions were performed to determine the specific activity of each of the xenobiotic reductases of P. putida KT2440 toward TNT as a model nitroaromatic substrate. All enzymes preferentially used NADPH as a cofactor except XenC, which showed a greater preference for NADH (data not shown). XenB showed the highest specific activity (1.88 U/mg protein). Much lower but measurable activity was found toward TNT with XenC, XenA, XenE, and XenF (activity was in the range of 0.1 to 0.2 U/mg protein). In the case of XenD, no activity could be measured with TNT or any other substrate irrespective of whether NADH or NADPH was used as a cofactor, nor could any activity be detected when 2.25 µM XenD had previously been incubated for 24 h with 20 µM FMN. If the association of a histidine tag was the cause for the impaired activity of XenD, as it was with YqjM of Bacillus subtilis (11), remains unknown. With the other five active enzymes, we determined the kinetic parameters K_m and V_max for NAD(P)H and TNT. K_m values were in the range of 18 to 77 µM for the pyridine nucleotide and in the range of 23 to 494 µM for the nitroaromatic compound (Table 2). With regard to K_m values, XenA showed the lowest affinity for TNT. On the other hand and in agreement with the specific activity determined, XenB exhibited the highest V_max values and the most favorable V_max/K_m relationship for TNT compared to those of the other active xenobiotic reductases of P. putida KT2440.

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TABLE 2. Kinetic parameters for NADPH and TNT for a set of OYE enzymes of P. putidaa

Other substrates such as N-ethylmaleimide, cyclohexenone, and glycerol trinitrate were also tested in vitro, and K_m and V_max values were determined (Table 3). V_max values revealed that the highest reaction rates were achieved with N-ethylmaleimide reductase as a substrate; in particular, with this substrate, XenC exhibited at least eightfold-higher rates than any other enzyme. To derive some other conclusions about substrate specificity, we took the derived V_max/K_m values into account. The data showed that all five enzymes were effective with N-ethylmaleimide, with the best ratio found for XenF. Cyclohexenone, a substrate often tested with OYE members, is a poor substrate for XenB and XenC but a good substrate for XenA and XenE (Table 3). Similarly, the explosive glycerol trinitrate is a better substrate for XenA and XenE than for the other xenobiotic reductases. XenC showed no activity with this substrate, which could be due to an inhibiting effect. Altogether, we observed that each enzyme has different activities toward different substrates, with some overlapping in substrate preferences between XenA and XenE. This supports a certain degree of subfunctionality within this group of enzymes for the bacteria. In this regard, it should also be noted that in response to stress mediated by toluene, the XenA cellular protein level increased about threefold, whereas the level of the other xenobiotic reductase enzymes did not change (27).

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TABLE 3. Kinetic parameters for xenobiotic reductases of P. putida with N-ethylmaleimide, 2-cyclohexen-1-one, and glycerol trinitrateb

Products formed from TNT reduction by xenobiotic reductases.
To determine the products formed from TNT by each enzyme, time course reactions were performed with 50 mM potassium phosphate buffer (pH 7) previously saturated with TNT containing 1 µM enzyme, 0.1 mM NAD(P)H, and the NAD(P)H recycling system (36). The transformation products formed by each xenobiotic reductase were analyzed by HPLC. The TNT derivatives that were successfully identified by HPLC (31) are shown in Fig. 1, in which we also show schematically the possible TNT biotransformation reactions that could lead to the formation of these products, whose chemical structures and masses were determined previously (37). Analysis of the products formed in vitro showed that the five functional P. putida OYE enzymes had type I hydride transferase activity since they could reduce the nitro side groups to form hydroxylamine derivatives of TNT. XenC and XenA showed a slight preference for the formation of 2HADNT over 4HADNT or vice versa, respectively. However, the other enzymes produced both hydroxylamine isomers at roughly equal amounts, which is in accordance with the findings described previously by Williams et al. (36) with OYE itself and morphinone reductase. However, contrary to the data reported previously by Williams et al. (36), the enzymes described here produced small but clearly detectable amounts of azoxynitrotoluenes (17) even when an NAD(P)H recycling system was put into place; moreover and in contrast with data described previously by Williams et al. (36), we were unable to detect the presence of 2,4-dihydroxylamino-6-nitrotoluene in any of the reactions.

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FIG. 1. Scheme showing pathways leading to the products detected in vitro in TNT biotransformations by the xenobiotic reductases of P. putida.

The XenB protein was also shown to have type II hydride transferase activity since it directly reduced the aromatic ring of TNT to yield Meisenheimer hydride complexes. The Meisenheimer monohydride complex produced by XenB was quickly reduced to various isoforms of the Meisenheimer dihydride complex (not shown). An interesting observation that we made is that with TNT, the combination of type I and type II hydride transferases in a single enzyme led to the release of significant amounts of nitrite (up to 300 µM), which resulted from the condensation of Meisenheimer dihydride complexes with hydroxylamine derivatives to produce nitrite and diarylamines (9.3% of TNT consumed). It was also found that P. putida KT2440 XenB produced larger amounts of N-(2-methyl-3,5-dinitrophenyl)-4-methyl-3,5-dinitroaniline (8.5% of TNT consumed) than of symmetric diarylamine (0.8%), which is in close agreement with the observation that initially more 2HADNT is produced than 4HADNT.

Phylogenetic comparisons.
To gain insights on the basis of the above-described subfunctionality of the enzymatic activities associated with P. putida OYE enzymes, we created a phylogenetic tree based on CLUSTALW alignments (18) of the amino acid sequences of the six KT2440 enzymes and a number of type I and type II hydride transferases of the OYE family, including those for which the three-dimensional crystal structures have been determined (Fig. 2). The results showed that the xenobiotic reductases of P. putida KT2440 are spread over two clades with various subgroups (21). This suggests that the xenobiotic reductases present in P. putida KT2440 might have different origins, with perhaps the exception of XenD, which shares high identity with XenA and therefore could be the result of a relatively recent duplication event.

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FIG. 2. Phylogenetic tree constructed with amino acid sequences of some members of the OYE family. Neighbor-joining percentages are supported by 2,000 bootstrap runs. Underlined enzymes have been shown to have type I hydride transferase activity, and boxed enzymes have been shown to have type II hydride transferase activity. Asterisks indicate that the three-dimensional crystal structure is known.

XenA, XenD, and XenE fall into clade 2, and XenB, XenC, and XenF fall into clade 1, and this phylogenetic grouping correlates to a certain extent with the substrate preferences observed (Table 3). However, it did not facilitate the differentiation of type I and type II hydride transferases. We therefore took a close look at the structural elements and available mutants in the OYE family to attempt to find the residues responsible for the partitioning between nitroreduction (type I hydride transferase activity) and ring reduction (type II hydride transferase activity). However, the type II hydride transferases pentaerythritol tetranitrate (PETN) reductase of Enterobacter cloacae and N-ethylmaleimide reductase of Escherichia coli grouped more closely with the type I hydride transferase XenF than with xenobiotic reductase B of either P. putida or P. fluorescens. As a result, the phylogenetic grouping did not facilitate the differentiation of type I and type II hydride transferases. We therefore took a close look at the structural elements and available mutants in the OYE family to attempt to find residues responsible for the partitioning between nitroreduction (type I hydride transferase activity) and ring reduction (type II hydride transferase activity). In PETN reductase, the exchange of the active-site residue His184 with an asparagine residue causes this enzyme to lose its ability to convert TNT into the Meisenheimer monohydride complex (36). However, xenobiotic reductases B of both P. fluorescens and P. putida have an asparagine residue at this position but retain type II hydride transferase activity, which indicates that the simple replacement of histidine by asparagine cannot account for the loss of function and that the loss of type II hydride transferase activity in the PETN H184N mutant might be due to slight changes in the active-site environment rather than to the residue itself.

We cannot rule out that type I and type II hydride transferase activities may result from slightly different modes of interaction with TNT in the active site so that either only a lateral nitro group or the aromatic ring is reduced or both the lateral nitro group and the aromatic ring are reduced. Different modes of binding have been described for different chemicals in the recognition pocket of regulators that control the extrusion of xenobiotic compounds, such as QacR (26), TtgR (2), and TtgV (16). Although further analyses are needed to support this hypothesis, the subfunctionality of OYE family members in P. putida KT2440 allows this bacterium to confront a xenobiotic contaminant such as TNT by both redundancy (several type I hydride transferases) and divergent activity (type II hydride transferase).

 
This study was supported by a grant from the European Commission (MADOX QLRT-2001-00345), a grant from the Spanish Ministry of Science and Education (CSD2007-00005), and a grant from the Junta de Andalucia in Spain (Proyecto de Excelencia CIV344).

We thank José A. Paz and David Martín for technical assistance, Lourdes Sánchez of the Servicio de Instrumentación Científica of the Estación Experimental del Zaidín for help with HPLC analysis, and M. Mar Fandila and Carmen Lorente for secretarial assistance.

 
* Corresponding author. Mailing address: EEZ-CSIC, C/Profesor Albareda 1, 18008 Granada, Spain. Phone: 34 958 181608. Fax: 34 958 135740. E-mail: jlramos{at}eez.csic.es

Published ahead of print on 12 September 2008.

Present address: Göteborg University, Cmb. Microbiologi Box 462, 405 30 Göteborg, Sweden.

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Applied and Environmental Microbiology, November 2008, p. 6703-6708, Vol. 74, No. 21
0099-2240/08/$08.00+0     doi:10.1128/AEM.00386-08
Copyright © 2008, American Society for Microbiology. All Rights Reserved.

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