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Journal of Bacteriology, January 2004, p. 98-103, Vol. 186, No. 1
0021-9193/04/$08.00+0     DOI: 10.1128/JB.186.1.98-103.2004
Copyright © 2004, American Society for Microbiology. All Rights Reserved.

NpdR, a Repressor Involved in 2,4,6-Trinitrophenol Degradation in Rhodococcus opacus HL PM-1

Dang P. Nga,1 Josef Altenbuchner,2 and Gesche S. Heiss1*

Institute of Microbiology,1 Institute of Industrial Genetics, University of Stuttgart, 70550 Stuttgart, Germany2

Received 12 June 2003/ Accepted 2 October 2003


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ABSTRACT
 
Rhodococcus opacus HL PM-1 utilizes 2,4,6-trinitrophenol (picric acid) as a sole nitrogen source. The initial attack on picric acid occurs through two hydrogenation reactions. Hydride transferase II (encoded by npdI) and hydride transferase I (encoded by npdC) are responsible for the hydride transfers. Database searches with the npd genes have indicated the presence of a putative transcriptional regulator, npdR. Here, the npdR gene was expressed in Escherichia coli, and the protein was purified and shown to form a complex with intergenic regions between open reading frames A and B and between npdH and npdI within the npd gene cluster. A change in DNA-NpdR complex formation occurred in the presence of 2,4-dinitrophenol, picric acid, 2-chloro-4,6-dinitrophenol, and 2-methyl-4,6-dinitrophenol. By constructing a promoter-probe vector, we demonstrated that both intergenic regions caused the expression of reporter gene xylE. Hence, both of these regions contain promoters. A deletion mutant of R. opacus HL PM-1 was constructed in which part of npdR was deleted. The expression of npdI and npdC was induced by 2,4-dinitrophenol in the wild-type strain, while in the mutant these genes were constitutively expressed. Hence, NpdR is a repressor involved in picric acid degradation.


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INTRODUCTION
 
The xenobiotic 2,4,6-trinitrophenol (TNP) contaminates industrial effluents, water systems, and soil. Its industrial and commercial versatility (as components of pesticides or azo dyes or as an explosive) has led to accumulation in the environment. Since the compound is toxic, there is interest in removing it from contaminated sites. Several bacteria have the capacity to use TNP as a nitrogen or carbon source under aerobic conditions (3, 17, 19, 23).

Rhodococcus opacus (originally misclassified as Rhodococcus erythropolis) HL PM-1 was originally isolated on the basis of its ability to grow on 2,4-dinitrophenol (DNP) as a sole nitrogen source; it also utilizes TNP as a sole nitrogen source (19). The capacity to degrade TNP or DNP is unusual in that initial conversion takes place through hydrogenation. The enzymes involved in these hydrogenation steps are hydride transferases (hydride transferase I [HTI], encoded by npdC, and hydride transferase II [HTII], encoded by npdI), which transfer hydride ions to the aromatic ring of TNP and subsequently to the hydride Meisenheimer complex of TNP (H--TNP) in two consecutive steps, forming the dihydride Meisenheimer complex of TNP (2H--TNP). Both hydrogenation steps are dependent on an NADPH-dependent F420 reductase (NDFR; encoded by npdG) to supply the hydride ions in the form of F420H2 (7, 8, 16).

Generally, the regulation of nitroaromatic degradation at the molecular level is not well understood. DNP has been shown to be an inducer of TNP degradation in R. opacus HL PM-1 (35). Not much more is known about how this unusual pathway is regulated. Understanding the regulation of TNP degradation is an important step toward increasing the efficiency of TNP degradation and may assist in the development of biological systems for efficient treatment of contaminated sites and effluents.

The TNP degradation gene cluster in R. opacus HL PM-1 was shown to contain an apparent transcriptional regulator, npdR (24). Here, we showed that NpdR is a repressor, negatively regulating the expression of the npd genes. We showed further that NpdR binds to two intergenic regions (IGRs) in the npd gene cluster and that these regions contain promoters.


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MATERIALS AND METHODS
 
Bacterial strains, plasmids, and culture conditions. The bacterial strains and plasmids used are shown in Table 1. Recombinant Escherichia coli strains were grown at 37°C in Luria-Bertani (LB) medium supplemented with ampicillin (100 µg ml-1) or kanamycin (80 µg ml-1). For the expression of npdR, isopropyl-ß-D-thiogalactopyranoside (IPTG) (1 mM) was added to cultures of E. coli BL21(DE3)/pNGA1 or E. coli BL21(DE3)/pNGA5 at an optical density at 600 nm of 0.4 to 0.5, and culturing was continued for a further 4 h at 30°C.


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  		TABLE 1. Bacterial strains and plasmids used in this study

Rhodococcus strains were cultured at 30°C in LB medium or in minimal medium (50 mM KH2PO4-Na2HPO4 buffer (pH 7.4) containing 0.45 mM CaCl2, 40 mM acetate, and mineral salts solution [6]). For induction, 0.5 mM DNP was added at an optical density at 600 nm of 0.4 to 0.5, and culturing was continued for 3 h. Plasmids in Rhodococcus were selected for on kanamycin (80 µg ml-1). Sensitivity to sucrose was tested on LB agar supplemented with 10% (wt/vol) sucrose.

PCR and DNA manipulations. PCR was performed with a T Gradient Thermocycler 96 (Biometra GmbH, Göttingen, Germany). Primers were purchased from Eurogentec. Reaction mixtures (total volume, 25 µl) contained 25 pmol of each primer, 1.5 mM MgCl2, 4% dimethyl sulfoxide (DMSO), 0.2 mM each deoxynucleoside triphosphate, 0.5 to 1 U of Taq polymerase (Eppendorf-Netheler-Hinz GmbH, Hamburg, Germany), PCR buffer, and 10 ng of genomic DNA. Reactions performed with Pwo DNA polymerase (Peqlab Biotechnologie GmbH, Erlangen, Germany) were carried out as described above, except that 1 U of enzyme and 2.0 mM Mg2SO4 were used.

For heterologous expression of npdR, the gene was amplified to incorporate NdeI and BamHI sites at the 5' and 3' ends, respectively. The primers used were as follows: 5'-CGACATATGCCCGCCATCTCGCGC-3' and 5'-CGCGGATCCTCAGCCGCGCCCGGCGCCGAG-3' (NdeI and BamHI sites are underlined). The PCR fragment was purified, restricted with BamHI and NdeI, and ligated with pET11a or pAC28 (Table 1) which had been cut with the same restriction enzymes. E. coli DH5{alpha} was transformed with the resulting plasmid, pNGA1 or pNGA5, respectively. The plasmids were isolated from the strains and used to transform E. coli BL21(DE3).

For measurement of IGRI'-driven expression (IGRI' is part of IGRI [see Fig. 1]), pNGA6 was constructed. IGRI' was amplified by PCR to introduce EcoRI and BamHI sites. The primers used were 5'-CGGAATTCCTTTCGTTTCGCGTTGCTGC-3' and 5'-CGGGATCCCATCACAAGCTCCGTTCAC-3' (EcoRI and BamHI sites are underlined). The appropriately cut PCR product was inserted upstream of the promoterless xylE gene in pJOE814.2 (which had been cut with EcoRI and BamHI) to produce pNGA6. IGRI' plus xylE was subsequently cut out of pNGA6 as a single HindIII fragment and ligated with shuttle vector pK4 (which had been cut with XbaI) to produce pNGA7. Rhodococcus rhodochrous ATCC 12674 was transformed with pNGA7, creating R. rhodochrous ATCC 12674/pNGA7.



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  		FIG. 1. Diagram of npd gene cluster (GenBank accession number AF323606) showing IGRs. IGRI and IGRIV are enlarged. Inverted repeats and imperfect direct repeats are indicated by arrows; -35 and -10 hexamers are indicated by bold type and underlining; putative Shine-Dalgarno sequences are indicated by bold type. Brackets demarcate IGRI' and IGRIV', which were used for the experiments.

Reporter shuttle vector pNGA8 was constructed from pK4 as follows. xylE was cut out of pJOE814.2 as an XbaI fragment and ligated with pK4 to produce pNGA8. R. rhodochrous ATCC 12674 was transformed with pNGA8, forming R. rhodochrous ATCC 12674/pNGA8. IGRIV' (IGRIV plus the 3' end of npdH) was then introduced into pNGA8 to produce pNGA9 as follows. IGRIV' was amplified by PCR with primers 5'-CCGGAATTCCTGATCACCCCGTCATACGC-3' and 5'-GGGGTACCGAACTTCTCTTCATGATGTTGAAC-3' (EcoRI and KpnI sites are underlined). The PCR fragment and pNGA8 were each restricted with the enzymes and ligated to create pNGA9. R. rhodochrous ATCC 12674 was transformed with pNGA9, forming R. rhodochrous ATCC 12674/pNGA9.

DNA manipulations were carried out as described by Sambrook et al. (26) and Ausubel et al. (2). Plasmids were isolated by using a Micro Plasmid Prep kit (Amersham Biosciences).

npdR gene deletion. To delete npdR from Rhodococcus, the 3' end of open reading frame (ORF) E, ORF F, and the 5' end of npdR were amplified. Further, the 3' end of npdR, npdG, and the 5' end of npdH were amplified. The primers used were 5'-GAGAATTCGGCGGAACTCCGTGAACTCG-3' and 5'-TGAGGTACCCGTCCGGCATCGGCTGG-3' (EcoRI and KpnI sites are underlined) and 5'-ATAGGTACCGGAACTCAACGTCGTGG-3' and 5'-GGGGATCCTGCGGTGCAGGTCCTCG-3' (KpnI and BamHI sites are underlined). The PCR fragments were ligated, creating a 2.2-kb fragment with a deletion in npdR (nucleotides 317 to 526). The ligated fragment was restricted with EcoRI and BamHI and ligated into the EcoRI and BamHI sites of pK18mobsacB to produce pNGA20. E. coli S17.1 was transformed with pNGA20 and then conjugated into R. opacus HL PM-1. Southern analysis of kanamycin-resistant colonies demonstrated that both expected types of single crossover events had taken place. One clone of each type was replica plated on LB medium plus sucrose. PCR and Southern hybridization of sucrose-resistant colonies showed the second homologous recombination event. The gene deletion mutant was named R. opacus ND1.

Transformation and conjugation. E. coli was transformed as described by Sambrook et al. (26) and Ausubel et al. (2). Rhodococcus spp. were transformed by electroporation as described by Hashimoto et al. (15). For conjugal transfer from E. coli S17.1 to R. opacus HL PM-1, cells were harvested in the exponential growth phase, resuspended in 400 µl of LB medium, and mixed at a ratio of 2:1, 1:1, or 1:2. From the mixture, 200 µl was spotted onto a filter (25-mm diameter; Sartorius) on an LB agar plate and incubated for 2 days (30°C). The membrane was washed with salts solution (0.9% [wt/vol] NaCl, 0.01% [wt/vol] Triton X-100), and the suspension was spread on LB medium containing kanamycin at 80 µg ml-1 and nalidixic acid at 20 µg ml-1.

Gel retardation assays. DNA binding reactions were performed with 10-µl mixtures consisting of gel mobility assay buffer (10 mM Tris-HCl [pH 7.5], 50 mM KCl, 4.35% glycerol, 5 µg of fish sperm DNA ml-1, 50 µg of bovine serum albumin ml-1), 0.001 to 10 mM DNP, 3 to 4 ng of labeled DNA fragment, competitor DNA (160 ng), and cell extract (100 to 300 ng of protein) containing the His tag fusion to NpdR (NpdR-His) or 100 to 200 ng of purified NpdR-His. The DNA fragments were end labeled with Cy5 by PCR with primers 5'-Cy5-CACAAGCTCCGTTCACTA-3' and 5'-Cy5-TTGCTGCGCGCCCGCCATTTCC-3' for IGRI', 5'-Cy5-TGACAGCATTCGCACGAC-3' and 5'-Cy5-CAGCTGCTCGCTGGATTG-3' for IGRII, 5'-Cy5-CCGAGCCCCCGATTTCA-3' and 5'-Cy5-GTCTGTCTCCTACACATTG-3' for IGRIII, and 5'-Cy5-GCACCGAGAGCGACGGGCCGC-3' and 5'-Cy5-CGAACTTCTCTTCATGATGTTGAAC-3' for IGRIV'. The reaction mixtures were incubated for 1 h at 4°C and then subjected to electrophoresis (1 to 2 h, 130 V, 4°C) in 8% polyacrylamide gels with native Tris-acetate-EDTA buffer. The gels were analyzed with a PhosphorImager (Storm 860; Amersham Biosciences) and ImageQuant 5.2 software.

Preparation of cell extracts and purification and analysis of proteins. Cell extracts were prepared by using a French press as previously described (16). Sodium dodecyl sulfate (SDS)-polyacrylamide gel electrophoresis (PAGE) and determination of protein concentrations were performed as described previously (16). NpdR was purified as a His tag fusion protein from induced cultures of E. coli BL21(DE3)/pNGA5 by Ni-nitrilotriacetic acid metal affinity chromatography by following the instructions of the supplier (Qiagen GmbH). The purity of the protein was estimated by SDS-PAGE. Gel retardation assays were used to test whether the protein was active or not. NDFR was purified as described before (16).

Enzyme assays. Enzyme assays were performed with a Varian Cary 50 Bio spectrophotometer controlled by Cary WinUV Biopackage software. The reactions (total volume, 1 ml) were carried out with 100 mM KH2PO4-K2HPO4 buffer (pH 7.5) for XylE assays and 50 mM KH2PO4-K2HPO4 buffer (pH 7.5) for NpdI and NpdC assays.

The enzyme activity of catechol-2,3-dioxygenase (C23DO; XylE) was measured by the addition of 0.2 mM catechol (Ferak, Berlin, Germany) plus 8 to 15 µg of protein. The increase in the absorbance was monitored at 375 nm for 2 min. Reaction rates were calculated by using an extinction coefficient of 36,000 M-1 cm-1 (25). For screening of clones expressing xylE, plates were sprayed with a 20 mM solution of catechol.

The conversion of TNP to H--TNP (detection of HTII) or H--TNP to 2H--TNP (detection of HTI) was monitored at 485 nm for 30 s. The following were added to the enzyme assays: 0.1 mM TNP or H--TNP, 125 µM NADPH, 13 µM F420, and 5 µg of purified NDFR. Reaction rates were calculated by using an extinction coefficient of 8,535 M-1 cm-1. H--TNP was synthesized and the compounds were identified as described before (16).

Reactions were initiated by the addition of cell extracts. One unit of enzyme activity was taken as the amount of enzyme which converted 1 µmol of substrate per min.

Sequence analyses. Database searches were performed with Blastn, Blastp, and Blastx. Pairwise and multiple alignments were carried out with Blast2 (http://www.ncbi.nlm.nih.gov/gorf/bl2.html) and ClustalW (1, 29, 30; http://www.ebi.ac.uk/clustalw/). Translations were achieved by using Translation Machine (http://www2.ebi.ac.uk/; EMBL Outstation European Bioinformatics Institute). Motif searches were performed with the tools CD-search (http://www.ncbi.nlm.nih.gov), Panal (http://mgd.ahc.umn.edu/panal/), Motif (http://motif.genome.ad.jp/), and Network Protein Sequence Analysis (http://npsa-pbil.ibcp.fr/cgi-bin/npsa_automat.pl?page=/NPSA/npsa_hth.html) (5). Promoter regions were identified by using the program Promoter Finder (http://wwwmgs.bionet.nsc.ru/mgs/programs/bdna/tata_bdna.html).

Chemicals. TNP, 2-chloro-4,6-dinitrophenol, 2-methyl-4,6-dinitrophenol, 4-nitrophenol, 4-nitrobenzoate, 2,6-dinitrophenol, trinitrotoluene (TNT), and DNP were purchased from Bayer and Fluka.


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RESULTS AND DISCUSSION
 
Expression of npdR. SDS-PAGE analysis of cell extracts from E. coli BL21(DE3)/pNGA1 expressing NpdR showed the presence of an IPTG-induced predominant polypeptide band at about 26.9 kDa that was absent from cell extracts from E. coli BL21(DE3)/pET11a not expressing NpdR and that coincided with the size of the translated DNA sequence of NpdR (26,867.82 Da). SDS-PAGE analysis of cell extracts from E. coli BL21(DE3)/pNGA5 expressing NpdR-His showed the presence of a predominant protein band that was absent from crude extracts derived from E. coli BL21(DE3)/pAC28 not expressing NpdR. More than half of the protein resided in the cells as inclusion bodies. NpdR-His, with the His tag at the N-terminal end, was purified from cell lysates. SDS-PAGE analysis of the purified fraction showed a thick, single polypeptide band. Hence, the protein was >95% pure. The size of the purified fusion protein was about 28 kDa.

Identification of consensus sequences in the ORF A-ORF B and npdH-npdI IGRs. To identify possible regulatory regions, the npd gene cluster (Fig. 1) was examined for IGRs. A 275-bp IGR between ORF A and ORF B (IGRI) was found to contain a putative promoter region with a score of 0.92 (Fig. 1). Two further IGRs, IGRII, between ORF F and npdR (98 bp), and IGRIII, between npdR and npdG (131 bp), exist. No promoter regions were identified in these IGRs. Analysis of a fourth IGR (IGRIV), between npdH and npdI (102 bp), revealed a putative promoter region with a score of 0.62 (Fig. 1).

Part of IGRI (referred to here as IGRI') and IGRIV plus the 3' end of npdH (referred to here as IGRIV') shared a sequence identity of 57%. In both the putative -35 and the putative -10 hexamers, five out of the six nucleotides were identical. Interestingly, IGRI' contained nucleotides identical to the 3' end of npdH. Perfect and imperfect inverted repeats and imperfect direct repeats were detected (Fig. 1).

NpdR binds to IGRI' and IGRIV'. Gel retardation assays were used to assess the binding of NpdR to IGRs. For IGRI', a complex, C1, appeared with cell extracts containing NpdR-His from E. coli BL21(DE3)/pNGA5 (Fig. 2A, lane 3) or NpdR from E. coli BL21(DE3)/pNGA1 (data not shown) or purified NpdR-His. These findings demonstrated NpdR-His-IGRI' complex formation. No DNA-protein complex formation was observed with cell extracts lacking NpdR [from E. coli BL21(DE3)/pAC28] (Fig. 2A, lane 2).



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  		FIG. 2. Gel retardation assays showing DNA-NpdR-His complex formation with IGRI' (A) and IGRIV' (B). Assays were performed as described in Materials and Methods. The amount of DNA was 3 ng. (A) Lanes: 1, IGRI' DNA only; 2, IGRI' plus 300 ng of cell extract from E. coli BL21(DE3)/pAC28 not expressing NpdR-His; 3, IGRI' plus 300 ng of NpdR-His; 4, IGRI' plus 300 ng of NpdR-His and 120 ng of unlabeled IGRI'; 5, IGRI' plus 300 ng of NpdR-His and 120 ng of unlabeled npdH. Cell extracts from E. coli BL21(DE3)/pNGA5 expressing NpdR-His were used. (B) Lanes: 1, IGRIV' DNA only; 2, IGRIV' plus 200 ng of NpdR-His; 3, IGRIV' plus 200 ng of NpdR-His and 10 mM DNP; 4, IGRIV' plus 200 ng of NpdR-His and 0.001 mM DNP. Purified NpdR-His was used.

Complex formation was not prevented by the presence of an arbitrary, unlabeled sequence (npdH) 40-fold in excess of labeled IGRI' (Fig. 2A, lane 5). Binding was prevented by the addition of a 40-fold excess of unlabeled IGRI' (Fig. 2A, lane 4). These findings indicated that NpdR-His bound specifically to IGRI'. The same results were obtained with IGRIV' (Fig. 2B, lane 2). NpdR-His did not form a complex with either IGRII or IGRIII (data not shown). This finding coincided with the inability to detect putative promoter regions in these DNA segments.

A second complex, C2, formed with both IGRI' and IGRIV' in the presence of DNP. The results are shown for IGRIV' only (Fig. 2B, lanes 3 and 4). To show that C2 was specifically caused by DNP, gel retardation assays were performed with various concentrations of DNP (0.001 to 10 mM) for IGRI' and for IGRIV'. C2 formation was reduced with decreasing DNP concentrations. At 10 mM DNP, C2 was the predominant complex, and free DNA was visible (Fig. 2B, lane 3). At 0.001 mM DNP, C1 predominated (Fig. 2B, lane 4). These results indicated that C2 was a true complex formed in the presence of DNP.

As the protein concentration in the binding assay mixture was reduced, C2 was observed even in the absence of DNP. Figure 3 shows the results for IGRI' only. We hypothesize that NpdR binds to two sites in the IGRs, forming C1. The presence of C2 indicates that one of the sites is occupied due to a decrease in the affinity or concentration of NpdR. Adding DNP to the reaction mixture causes a decrease in the affinity of NpdR, forming C2. At lower NpdR concentrations, the two sites would not be expected to be saturated, also forming C2. The formation of C2 in the presence of a decreasing protein concentration suggests a lack of cooperative binding (i.e., NpdR molecules bind to the sites independently).



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  		FIG. 3. Gel retardation assay showing DNA-protein complex formation with IGRI' and NpdR-His at different protein concentrations. Lanes: 1, IGRI' DNA only; 2, IGRI' plus 200 ng of NpdR-His; 3, IGRI' plus 100 ng of NpdR-His; 4, IGRI' plus 50 ng of NpdR-His.

Retardation assays were performed with other aromatic compounds besides DNP. In the presence of TNP (Fig. 4, lane 4), 2-chloro-4,6-dinitrophenol (lane 8), and 2-methyl-4,6-dinitrophenol (lane 10), C2 formed with IGRI' or IGRIV' (data are shown for IGRI' only). Strain HL PM-1 has the ability to use TNP and 2-chloro-4,6-dinitrophenol as sole carbon and nitrogen sources (20). Hence, it was not unexpected that NpdR responded to these analogous compounds. Strain HL PM-1 converts 2-methyl-4,6-dinitrophenol to dead-end products (20). It is possible that NpdR binds to the compound, leading to the induction of enzymes, although they may not convert it as a true substrate. 4-Nitrophenol (Fig. 4, lane 6), 4-nitrobenzoate (lane 7), 2,6-dinitrophenol (lane 5), and TNT (lane 9) did not affect the binding of NpdR to either IGR (data are shown for IGRI' only). Strain HL PM-1 does not grow on any of these substrates (21), a fact which may explain why no complex formation occurred. TNT can be converted to the hydride and the dihydride Meisenheimer complex of TNT, but no further conversion takes place (33, 34).



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  		FIG. 4. Gel retardation assay showing DNA-protein complex formation with IGRI'and NpdR-His in the presence of various nitrogenous compounds. Lanes: 1,IGRI'DNA only; 2,IGRI'DNA with 200ng of NpdR-His; 3 to 10,IGRI'DNA with 200ng of NpdR-His and a mM concentration of the indicated aromatic compound.

Reporter gene xylE is expressed when under the control of IGRI' and IGRIV'. To determine that the promoters identified within IGRI' and IGRIV' are functional, the respective regions were cloned upstream of the promoterless reporter gene xylE in pK4, creating pNGA7 and pNGA9, respectively (Table 1). Plasmid pNGA8 (containing the promoterless xylE gene in pK4 only) was used for comparing expression levels. R. rhodochrous ATCC 12674 was used as a host for the plasmids. This strain was used instead of R. opacus HL PM-1 because it does not grow on TNP or DNP as a sole nitrogen source. Hence, interference with endogenous IGRI', IGRIV', and NpdR would be less likely to occur with this strain than with wild-type strain HL PM-1. Cell extracts were prepared from mid-log-phase cultures of R. rhodochrous ATCC 12674 containing pNGA7 (xylE under the control of IGRI'), pNGA8 (containing promoterless xylE), or pNGA9 (xylE under the control of IGRIV'). The activities of C23DO in cell extracts of R. rhodochrous ATCC 12674/pNGA7 (1,030 mU/mg) or R. rhodochrous ATCC 12674/pNGA9 (1,513 mU/mg) were about 50- or 75-fold higher than those in cell extracts of R. rhodochrous ATCC 12674/pNGA8 (20 mU/mg), respectively. These results showed that IGRI' and IGRIV' both contain a promoter.

NpdR is a repressor. To assess the implications from the in vitro binding studies, namely, that NpdR is a repressor, enzyme assays were performed with wild-type R. opacus HL PM-1 or an npdR deletion derivative thereof, R. opacus ND1. The strains were cultured in minimal medium and induced with DNP. Cell extracts were prepared, and enzyme assays were performed to detect HTII (conversion of TNP to H--TNP) and HTI (conversion of H--TNP to 2H--TNP) (Table 2). Cell extracts from cultures of R. opacus HL PM-1 induced with DNP exhibited approximately 50- to 60-fold greater activity for HTII or for HTI than did those from noninduced cultures. These findings coincide with earlier observations that npdC, npdG, and npdI are induced by DNP (35). Cell extracts from R. opacus ND1 showed even higher and very similar enzyme activities for HTII and HTI, irrespective of whether they had been induced. These results indicate that the expression of npdI or npdC was constitutive in the deletion mutant. Further, they demonstrate that NpdR represses the expression of npdI or npdC in R. opacus HL PM-1.


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  		TABLE 2. Specific activities of HTII and HTI in R. opacus HL PM-1 and R. opacus ND1a

To show that NpdR directly affected expression from IGRI' or IGRIV', pNGA7 (expressing xylE from IGRI') or pNGA9 (expressing xylE from IGRIV') was transformed into R. opacus HL PM-1 or R. opacus ND1. Cell extracts from DNP-induced cultures of recombinant strains of R. opacus HL PM-1 possessed C23DO activities of about 1,400 mU/mg of protein. Cell extracts from induced or noninduced cultures of recombinant R. opacus ND1 strains possessed comparable activities (data not shown). Cell extracts from noninduced cultures of recombinant R. opacus HL PM-1 strains possessed activities four- to fivefold lower (about 300 mU/mg of protein). The results show that xylE expression was induced by DNP when under the control of IGRI' or IGRIV' in the wild-type strain and that expression was constitutive in the npdR deletion mutant. A possible reason for the low level of induction is that not enough NpdR is expressed from strain HL PM-1 to saturate the binding sites in IGRI' or IGRIV' to completely repress xylE expression from pNGA7 or pNGA9.

NpdR is a helix-turn-helix (HTH) IclR-type regulator. Database comparisons with npdR or NpdR showed sequence similarities of 40 to 42% to transcriptional regulators of the IclR family of Bacillus halodurans, Salmonella enterica serovar Typhimurium, Yersinia pestis, and E. coli (expect values: 2e-11 to 1e-09). Sequence similarities of 40 to 42% to KdgR (LacI family) transcriptional regulators of S. enterica, Pectobacterium carotovorum, and E. coli also were identified (expect values: 9e-10 to 6e-10).

An IclR HTH motif was detected at the N-terminal end of NpdR as well as in PcaR and CatR from R. opacus (9), in putative regulators from Mycobacterium smegmatis, and in IclR-like regulators from Streptomyces spp. An HTH motif was identified at positions 21 to 42 with a 90% probability and a score of 4.28 at position 21 (5; Network Protein Sequence Analysis). Hence, NpdR is probably an IclR-type transcriptional regulator containing an HTH motif at the N-terminal end.

In this work, we showed that NpdR is a repressor of the IclR family of transcriptional regulators and is involved in TNP degradation. IclR-type regulators have been described mostly for E. coli and S. enterica serovar Typhimurium, in which the IclR regulator is a repressor of the glyoxylate bypass operon during growth on acetate or fatty acids (10, 11, 13, 14). Hence, it was not unexpected that NpdR was found to be a repressor. Members of the PobR subfamily of the IclR family of regulators (PcaR, CatR, PobR, and PcaU) are involved in the regulation of protocatechuate or catechol degradation in Pseudomonas, Acinetobacter, and Rhodococcus (9, 12, 31). Of these regulators, PcaU has been shown to be a transcriptional activator (12). IclR-type regulators involved in the regulation of nitroarene degradation have not yet been described. The putative regulators DntR (2,4-dinitrotoluene degradation) and NbzR (nitrobenzene degradation) are both LysR-type regulators (22). Hence, it seems likely that LysR-like proteins will appear more predominantly in future studies of the regulation of nitroaromatic catabolism.


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ACKNOWLEDGMENTS
 
We thank Robert van der Geize (University of Groningen, Haren, The Netherlands) for the gift of pK18mobsacB.

This work was supported by the German Research Foundation (DFG).


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FOOTNOTES
 
* Corresponding author. Mailing address: Institute of Microbiology, University of Stuttgart, Allmandring 31, 70550 Stuttgart, Germany. Phone: 49 711 685 5491. Fax: 49 711 685 5725. E-mail: gesche.heiss{at}po.uni-stuttgart.de. Back


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REFERENCES
 
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Journal of Bacteriology, January 2004, p. 98-103, Vol. 186, No. 1
0021-9193/04/$08.00+0     DOI: 10.1128/JB.186.1.98-103.2004
Copyright © 2004, American Society for Microbiology. All Rights Reserved.



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