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Applied and Environmental Microbiology, April 2003, p. 2298-2305, Vol. 69, No. 4
0099-2240/03/$08.00+0     DOI: 10.1128/AEM.69.4.2298-2305.2003
Copyright © 2003, American Society for Microbiology. All Rights Reserved.

Characterization of TsaR, an Oxygen-Sensitive LysR-Type Regulator for the Degradation of p-Toluenesulfonate in Comamonas testosteroni T-2

Tewes Tralau,{dagger} Jörg Mampel,{ddagger} Alasdair M. Cook, and Jürgen Ruff*

Department of Biology, The University of Konstanz, D-78457 Konstanz, Germany

Received 3 October 2002/ Accepted 23 January 2003


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ABSTRACT
 
TsaR is the putative LysR-type regulator of the tsa operon (tsaMBCD) which encodes the first steps in the degradation of p-toluenesulfonate (TSA) in Comamonas testosteroni T-2. Transposon mutagenesis was used to knock out tsaR. The resulting mutant lacked the ability to grow with TSA and p-toluenecarboxylate (TCA). Reintroduction of tsaR in trans on an expression vector reconstituted growth with TSA and TCA. The tsaR gene was cloned into Escherichia coli with a C-terminal His tag and overexpressed as TsaRHis. TsaRHis was subject to reversible inactivation by oxygen, which markedly influenced the experimental approaches used. Gel filtration showed TsaRHis to be a monomer in solution. Overexpressed TsaRHis bound specifically to three regions within the promoter between the divergently transcribed tsaR and tsaMBCD. The dissociation constant (KD) for the whole promoter region was about 0.9 µM, and the interaction was a function of the concentration of the ligand TSA. A regulatory model for this LysR-type regulator is proposed on the basis of these data.


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INTRODUCTION
 
p-Toluenesulfonate (TSA) is widely used in the metal and chemical industries and in laundry detergents in the home (2). TSA is degraded via p-sulfobenzoate (PSB) and protocatechuate (PCA) by the bacterium Comamonas testosteroni T-2 (Fig. 1) (9); this pathway is encoded in part on two plasmids, pTSA and pT2L, and in part on the chromosome (PCA). pTSA (12) is widespread (42).



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  		FIG. 1. Degradation of TSA and TCA to amphibolic intermediates by C. testosteroni T-2 and the four regulons involved (R1 to R4) (9, 29, 36). Reactions of gene products that are encoded by chromosomal genes are in insets. Regulatory units R1 (pTSA) and R3 (pT2L) are plasmid encoded (42; Ruff, unpublished). The tsa regulon is sketched schematically, with the directions of transcription indicated by shaded triangles. TsaMB, p-toluenesulfonate methylmonooxygenase (oxygenase M, reductase B); TsaC, p-sulfobenzylalcohol dehydrogenase; TsaD, p-sulfobenzaldehyde dehydrogenase; PszA(C), p-sulfobenzoate-3,4-dioxygenase.

The first three metabolic enzymes in the degradation of TSA by strain T-2 are encoded on pTSA in the operon tsaMBCD (Fig. 1); tsaR is located upstream of tsaMBCD in the reverse orientation (Fig. 1) (13). TsaR, a hypothetical protein of 298 amino acids (32.7 kDa) (13), was proposed to be a LysR-type regulator (35) of the tsa operon with amino acid sequence identities of 27.8% to Nac (nitrogen assimilation control regulator; 37) and 27.2% to ClcR (chlorocatechol operon regulator; 7). Most of the LysR-type regulatory proteins are known to bind specifically to their target DNA (for examples, see references 5, 8, and 35). The putative regulatory region between tsaR and tsaMBCD has all the elements characteristic of regulatory regions, including bacterial {sigma}70 promoters, putative ribosomal binding sites, and transcriptional start, Pribnow box, and -35 regions (13).

We now confirm that TsaR, examined as the His-tagged TsaRHis, is the major regulator of expression of tsaMBCD in C. testosteroni T-2, and we describe some of its properties and interactions with the promoter region.


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MATERIALS AND METHODS
 
Bacteria, mutants, and growth conditions.
C. testosteroni T-2 (DSM 6577) was grown in minimal medium as described previously (36, 40). Mutants 4A10 and TT6 (Table 1) were grown with minimal medium supplemented with 50 to 70 µg of tetracycline/ml. The compounds used as growth substrates were pure (>99%). Growth was estimated as optical density at 580 nm and quantified according to levels of Lowry-type protein (15). Substrate utilization was determined after separation on reversed-phase high-performance liquid chromatography columns (17), and sulfate was quantified as a suspension of barium sulfate (39).


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  		TABLE 1. C. testosteroni T-2 and some mutants and the substrates they utilize

Escherichia coli S17-I{lambda}pir[pUT-Tc] was kindly provided by M. Kertesz and grown as described previously (10). E. coli DH5{alpha}[pJB866], as the donor of the broad-host-range vector pJB866 (GenBank accession no. U8200), was kindly provided by J. M. Blatny and grown as described previously (3). The E. coli host strain M15[pREP4] (Qiagen, Hilden, Germany), which contains an isopropyl-ß-D-thiogalactopyranoside (IPTG)-inducible expression system, and the derived clone M10 (see below) were grown with Luria-Bertani (LB) medium (33) with 25 µg of kanamycin/ml and 100 µg of ampicillin/ml, respectively, according to the manufacturer's recommendations.

PCR, quantification of DNA, cycle sequencing, and Southern hybridization.
PCR was done in a final volume of 20 to 50 µl. All reaction mixtures contained 10% (vol/vol) dimethyl sulfoxide. Genomic DNA (180 to 200 ng), prepared by cetyltrimethylammonium bromide precipitation (1), or a bacterial culture was used as the template. Templates (<=5 kb) were amplified with Taq polymerase (MBI Fermentas) in buffer system 2 of the Expand Long PCR system (Roche) or (>5 kb) with the Expand Long PCR System (Roche). The latter has a proofreading activity and was also used for the generation of PCR products designed for cloning. Reactions were done according to the manufacturers' recommendations. DNA was quantified fluorimetrically (DyNA Quant 200; Hoefer) according to the manufacturer's instructions. Primers used for PCR are listed in Table 2. The Tn5 insertions were localized in strain 4A10 with primer pair TsaReg1-TsaReg2. The template for sequencing the Tn5 insertion in strain 4A10 was amplified with primer pair TsaRu1-AntiRu1; the sequencing primers were TsaIG1, TsaReg4, and Tn55-Out.


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  		TABLE 2. PCR primers used in this study

Primers 866-MB1 and 866-21 or tsaRHisHindIII were used to detect by PCR those clones carrying pJB866 with the His-tagged tsaR gene; primer pair pQE70-1 and pQE70-2 was used for E. coli transformants carrying the His-tagged tsaR gene on pQE-70. Templates for sequencing the inserts of pJB866 and pQE70 were generated with primer pair 866-MB2 and 866-22 and primer pair pQE70-1 and pQE70-2, respectively.

The amplified template for cycle sequencing was purified with the Qiaquick PCR purification kit (Qiagen), and the fragment (85 ng of DNA/kb) was added to the sequencing reaction mixture (ABI Prism Big Dye terminator kit; Applied Biosystems) and subjected to the following PCR program: 70 s at 95°C and then 26 cycles of 20 s at 95°C, 30 s at 50°C, and 4 min at 60°C. The sequence was determined by capillary electrophoreses (GATC, Konstanz, Germany). Using standard software (Edit View [Perkin Elmer], a GCG program package, and DNA-STAR [Lasergene]) and the neural network analysis for prokaryotic promoters software of M. G. Reese (http://searchlauncher.bcm.tmc.edu/seq-search/gene-search.html) (11; M. G. Reese and F. H. Eeckman, abstract from the Proceedings of the Seventh International Genome Sequencing and Analysis Conference, vol. 1, no. 1, p. 45, 1995, and M. G. Reese, N. L. Harris, and F. H. Eeckman, poster from the Proceedings of the 1996 Pacific Symposium, Biocomputing, 1996), we analyzed the sequence data on the basis of characterized E. coli promoters.

Southern blot hybridization was done as described elsewhere (24). Gene probes were generated according to the manufacturer's instructions by digestion with EcoRI (New England BioLabs) of pUT-Tc carrying Tn5. The 2.1-kb fragment was labeled with a DIG system (Roche) and used as a gene probe to confirm the presence of the Tn5 insertion on pTSA.

The DNA markers used were the 0.1- and 1-kb ladders (MBI Fermentas), with ranges of 0.1 to 3 kb and 0.25 to 10 kb, respectively.

Pulsed-field gel electrophoresis.
DNA was prepared from whole cells embedded in low-melting-point agarose as described elsewhere (38). Electrophoresis was done as described previously (42).

Transposon mutagenesis, competent cells, and cloning.
The mini-Tn5 transposon mutagenesis system (10) was used to knock out tsaR in C. testosteroni T-2. E. coli S17-I{lambda}pir[pUT-Tc] (6 x 109 cells; donor) was mated with C. testosteroni T-2 (2 x 109 cells; recipient) on LB agar at 30°C for 48 h. Prior to mating, cells were centrifuged separately, washed twice, resuspended in 0.9% (wt/vol) sterile NaCl solution, and mixed before a 50-µl volume was applied to the LB agar plate. Cells were then scraped off the agar and washed once with 0.9% (wt/vol) NaCl. Dilution series were plated on minimal agar with PCA or succinate as the carbon and energy source (antidonor selection) and 15 µg of tetracycline/ml (selection for the Tn5 marker). A total of 7,500 mutants were characterized for growth with TSA, TCA, PSB, or terephthalate (TER) in microtiter plates. Mutants unable to grow on TSA or TCA were further analyzed, because this phenotype is expected from a knockout of tsaR (Fig. 1) (or of tsaMBCD). A total of 5{per thousand} of the mutants corresponded to this phenotype, but only one tsaR-negative mutant was obtained.

Vectors with inserts were introduced into target cells either by electroporation or by chemical transformation. Cells of C. testosteroni were prepared for electroporation as described previously (23, 33, 43) and electroporated with 0.2 to 1 µg of DNA at 15 kV/cm in a Gene Pulser II (Bio-Rad) with 125 µF of capacity, 200 {Omega} of resistance, and a 4.8 x 10-3-s pulse. Chemical transformation was done with E. coli M15[pREP4] according to the supplier's instructions (QIAexpress Kit Type ATG; Qiagen).

Three recombinant organisms were generated. First, C. testosteroni 4A10 (Table 1) was generated from C. testosteroni T-2 by Tn5 transposon mutagenesis; the insertion was in tsaR. Second, C. testosteroni TT6 was generated from mutant 4A10 (Table 2); it contained wild-type tsaR in trans. The tsaR gene from strain T-2 was amplified with primers CT1 and CT2 (Table 2) and modified by nested PCR to encode a C-terminal, sixfold His tag (tsaRhis). For that purpose, the reverse primer, tsaRHisHindIII, encoded a C-terminal His tag upstream of a stop codon and a restriction site for HindIII. A restriction site for AflIII was generated in the forward primer, tsaRAflIII, by replacing the original start codon, GTG, with ATG plus a subsequent TTG (leucine) codon. The construct generated by PCR was cloned into the broad-host-range vector pJB866 under the control of a promoter inducible by 2 mM m-toluate. This plasmid was electroporated into mutant 4A10 to yield mutant TT6 (Table 1). Third, E. coli M10 was constructed from E. coli M15[pREP4] to express TsaRHis. The tsaR gene from strain T-2 was ligated into pQE-70, which encodes a His tag. The insert was generated from the PCR-amplified template (primers CT1 and CT2; see above) and modified by nested PCR. The primer pair was tsaRSphI, in which the original start codon was replaced with ATG and a subsequent CTC (leucine) codon to generate a restriction site for SphI, and tsaRBglII, in which the original stop codon was replaced with TCT and AGA (serine and arginine) to fuse TsaR to the C-terminal His tag of pQE-70 and create a BglII restriction site. This plasmid was introduced into E. coli M15 by chemical transformation.

Preparation of TsaRHis, Western blotting, and determination of molecular mass under native and denaturing conditions.
Suspensions of chilled E. coli M10 were disrupted by sonication (3 bursts for 1 s at 70 W, on ice), and TsaR was purified under native conditions as described elsewhere (QIAexpress kit type ATG; Qiagen) and stored in 20% glycerol at -20°C. Immunological detection of purified TsaRHis after sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) was done by Western blotting following standard protocols (32), which were modified as necessary according to the antibody's manufacturer. Monoclonal antibodies (QIAexpress detection kit and Tetra-His antibody) against a fourfold His epitope were provided by Qiagen; anti-mouse immunoglobulin coupled to alkaline phosphatase (Roche) was used as secondary antibody, and detection was done with a disodium 3-(4-methoxyspiro (1,2-dioxetane-3,2'-(5'-chloro) tricyclo [3.3.1.13,7] decan)-4-yl)phenyl phosphate system (CSPD; Roche). The molecular mass of TsaRHis was estimated by gel filtration chromatography on a Superose 12 column calibrated with chymotrypsin, ovalbumin, bovine serum albumin, and aldolase (molecular mass values of 25, 43, 67, and 158 kDa, respectively). The mobile phase was 10 mM Tris, pH 7.0, containing 150 mM NaCl; the loading buffer was the elution buffer of the QIAexpress kit.

Denatured proteins were separated on SDS-PAGE (34) and stained with either standard (33) or colloidal (27) Coomassie brilliant blue. The protein markers used were a low-range marker (6.5, 14.4, 21.5, 31, 45, 66.2, and 97.4 kDa; Bio-Rad) and a 10-kDa protein ladder (10, 20, 30, 40, 50, 60, 70, 80, 90, 100, 110, 120, and 200 kDa; Gibco). Soluble protein was quantified according to Bradford (4).

DNA-binding assays (20-µl volumes).
Binding of TsaRHis to DNA was assayed by incubation of freshly prepared TsaRHis with 100 ng of PCR-amplified target DNA in binding buffer (5 mM Tris/HCl, 20 mM NaCl, 8 mM MgCl2, 10% glycerol, 1.5 M fresh 2-mercaptoethanol, 30 µg of freshly added bovine serum albumin/ml, pH 8.0) for 40 min on ice. Binding assays of TsaRHis were done with 2 µg of protein, and assays to determine KD, the dissociation constant, were done with 0.3 to 1.5 µg of protein. KD was defined as the protein concentration at which 50% of the target DNA molecules were occupied, as assayed by band shift. The influence of TSA (0.01 to 6 mM) on the binding of TsaRHis to DNA was measured at constant protein concentration (30.6 µg/ml). Samples for analysis were supplemented (10% [vol/vol]) with a solution of 15% Ficoll containing 0.25% bromophenol blue and subjected to standard agarose gel electrophoresis (33) in 1.5% agarose at 14.7 V/cm. Staining with ethidium bromide and destaining (33) were done after electrophoresis to exclude intercalation into the DNA during separation. Stained gels were documented (Gel Doc 1000; Bio-Rad) and subject to densitometric analysis (Multi-Analyst software for Macintosh, version 1.02; Bio-Rad Laboratories, Hercules, Calif.).

Nucleotide sequence accession number.
The putative promoter region sequence and partial gene sequence of pszA are available in the National Center for Biotechnology Information GenBank library under accession number AY044257.


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RESULTS
 
Transposon mutagenesis and complementation confirm TsaR as a regulator.
Junker et al. (13) suggested that TsaR is a LysR-type regulator of the tsa operon in C. testosteroni T-2 (Fig. 1). Transposon mutagenesis of tsaR (see below) yielded mutant 4A10 (Table 1), which was phenotypically tsa and tca negative but was genotypically tsa positive and tetracycline resistant (the resistance marker on the Tn5 minitransposon). The transposon was located by PCR and cycle sequencing. It was inserted between two cytosines at positions 563 and 564 on the tsa locus (U32622) in reverse transcriptional orientation to the tsaR gene, thus truncating 33 C-terminal amino acids of TsaR. pTSA was separated by pulsed-field gel electrophoresis, and Southern hybridization against the Tn5 transposon confirmed the prediction that the latter was inserted in the plasmid (data not shown).

Mutant 4A10 grew with neither TSA nor TCA, and growth with PSB was poor (Table 1). In contrast, construct TT6 (Table 1) showed utilization of TSA, TCA, or PSB in the presence of 2 mM m-toluate to induce TsaRHis; substrate utilization was determined by high-performance liquid chromatography. Inducible expression of TsaRHis in strain TT6 was confirmed by SDS-PAGE. Using antibodies against the His tag, separated proteins from crude extracts of cells grown in the presence or absence of m-toluate were subjected to Western blotting; only induced cells showed cross-reactivity, with a single band at about 33 kDa (Fig. 2B and data not shown). This proved that TsaR is a gene product which is essential for expression of tsaMBCD in C. testosteroni T-2. It also showed that the function of TsaR is not significantly affected by the C-terminal His tag.



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  		FIG. 2. Expression and purification of TsaRHis from E. coli strain M10 monitored by SDS-PAGE (A) and Western blot analysis (B). E. coli M10 was grown in the absence and presence of IPTG, the inducer of expression of tsaRhis. Soluble proteins from disrupted cells were compared; TsaRHis from induced cells was purified, and the His tag was identified immunologically. Lanes 1 and 8, calibration proteins with the molecular mass values given; lane 2, extract of noninduced cells; lane 3, extract of induced cells; lane 4, supernatant fluid after addition of nickel-agarose; lane 5, supernatant fluid from wash 1 of the purification protocol; lane 6, supernatant fluid from wash 2 of the purification protocol; lane 7, purified TsaRHis.

Overexpression in and purification of TsaRHis from E. coli M10 and specific binding of TsaRHis to the tsa promoter region.
No His-tagged protein was detected in noninduced cells of E. coli M10 (Fig. 2B, lane 2). Cells grown in the presence of 1 mM IPTG expressed a 33-kDa polypeptide (Fig. 2A, lane 3), which represented some 6% of soluble protein and carried a His tag (Fig. 2B, lane 3). This molecular mass value corresponded to that of the wild-type TsaR, 32.7 kDa (13), with a His tag of 0.7 kDa, as predicted by the DNA sequence encoding the recombinant protein. This protein was purified (Fig. 2, lane 7). The apparent molecular mass was 30 kDa (gel filtration), which implied that TsaRHis is a monomer in solution.

Preliminary binding studies showed that TsaRHis rapidly lost its ability to bind DNA within 5 h of cell disruption, unless it was stored under an atmosphere of nitrogen. Under the latter conditions, binding capacity was retained for at least 1 month. Samples with low levels of binding capacity were able to be restored to full capacity by incubation under nitrogen. Reproducible binding of TsaRHis to its target DNA in vitro required the presence of high concentrations of 2-mercaptoethanol (1.5 M instead of the millimolar concentrations which are often used; for an example, see reference 25). Binding assays had to be done quickly, because even under optimal conditions, all activity was lost after 20 min. We presume that TsaR is sensitive to oxygen.

This rapid loss of binding capacity, and the requirement for 1.5 M mercaptoethanol, ruled out the use of standard footprint assays. We chose to avoid radiochemical techniques, so standard band shift assays of DNA binding (1) were not applicable and other approaches were developed.

DNA-binding assays with TsaRHis and with specific fragments of DNA from promoter and coding regions were done in parallel, which compensated for the addition of competitor DNA normally used in radioactive DNA-binding assays to ensure sequence-specific binding of the protein. TsaRHis did not bind significantly to a coding region of the tsa operon (tsaMcod) (Table 3 and Fig. 3A); the mobility of the DNA fragments was unaltered by the presence of TsaRHis. The protein did cause a band shift when the putative promoter region was examined (tsaprom) (Table 3 and Fig. 3), and the DNA fragment was shifted completely by addition of 1.5 µM TsaRHis (tsaprom; Fig. 3B). We were concerned that TsaRHis might interact with a putative promoter region(s), so we explored the interaction with the appropriate region of the locus encoding PSB dioxygenase (Fig. 1) (pszAprom; Table 3). TsaRHis had no interaction with that fragment of DNA (pszAprom; Fig. 3B). TsaR is thus a DNA-binding protein specific for the tsa promoter region. The dissociation constant (KD [Materials and Methods]) of TsaRHis for the tsa promoter region (tsaprom; Table 3) was found to be 0.9 µM.


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  		TABLE 3. PCR products used for DNA-binding assays



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  		FIG. 3. Band shift assays with TsaRHis and specific fragments of DNA from C. testosteroni T-2. (A) tsaMcod, coding region of tsaMB (Table 3) used as a negative control for nonspecific DNA binding; (B) tsaprom, promoter region between tsaR and tsaM (Table 3); pszAprom, promoter of pszA (Table 3) used as a control for promoter specificity of DNA binding of TsaRHis.

Binding of TsaR to tsaprom depends on the ligand (TSA) concentration.
Changes in the affinity of binding (Fig. 4) of TsaRHis to tsa promoter DNA (tsaprom; Table 3) as a function of the substrate (TSA) concentration were observed and quantified as the amount of DNA subject to a band shift at various concentrations of TSA, the putative inducer of the tsa operon (13, 20). At low concentrations of TSA (0.01 mM), about 80% of the promoter DNA was shifted. When TSA concentrations were intermediate (0.1 to 1 mM), the amount of DNA shifted was lower (about 50%), which indicates a lower affinity of TsaR to its DNA-binding site(s). At higher concentrations of TSA (above 3 mM), more promoter DNA (about 80%) was shifted. Binding of TsaR to the tsa promoter region thus seems to depend on the substrate concentration, which suggests the presence of a regulatory system sensitive to levels of the growth substrate.



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  		FIG. 4. The effect of TSA concentration on the band shift of tsaprom caused by TsaRHis.

Location of several binding regions for TsaRHis within the tsa promoter region.
The putative tsa promoter region (positions 1363 to 1476 in U32622) is presumed to regulate expression of the divergently oriented tsaR and tsaMBCD genes (13). Newer software (30) allowed us to suggest transcriptional start sites for tsaR and tsaMBCD at positions 1389 and 1453, respectively (Fig. 5). We have now been able to explore the interaction of TsaRHis with five defined, overlapping DNA subfragments in this region (tsaprom) (Fig. 5 and Tables 3 and 4). In the absence of TSA, TsaRHis bound strongly to only one fragment (subfragment 4; Table 4). In the presence of 6 mM TSA, about a quarter of subfragment 1 was shifted (Table 4), whereas two shifts were visible with subfragment 2. Subfragments 3 and 5 gave no significant shift, whereas subfragment 4 gave two strong shifts. We conclude from Table 4 that there are two experimentally confirmed, TSA-dependent binding sites for TsaRHis (BS I on subfragment 1 and BS II on subfragment 2) (Fig. 5), whereas a third binding site (BS III, at the 5' end of tsaM), which was derived from the consensus sequence (35) and may contribute to the double shift of subfragment 4, has negligible direct experimental support (subfragment 5; Table 4). In the absence of TSA, in contrast, there is only one binding site for TsaRHis (BS IV; Fig. 5).



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  		FIG. 5. Sequence and structure near the tsa promoter region. The nucleotide positions are those of U32622 (13). Data from genes tsaR (both strands) and tsaM are in uppercase characters, with the start codons in bold characters; amino acids are in uppercase italic characters. The putative ribosomal binding site (*1), transcriptional start (*2), Pribnow box (*3), and -35 (*4) regions are in bold characters and are underlined for tsaR and shaded grey for tsaM. Subfragments 1 to 5 (Table 3) are indicated with pairs of arrows. The dashed line indicates putative DNA-binding sites for TsaRHis in the presence of TSA (I and III), and the dotted line indicates the putative binding site for TsaRHis in absence of TSA (IV); the binding region II is also shown with a dotted line with terminal arrows. An overview depicting subfragments 1 to 5 with the binding sites and the requirement of TSA is sketched below the sequence.


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  		TABLE 4. Band shifts with TsaRHis and subfragments of the tsaR/tsaMBCD regulona


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DISCUSSION
 
The hypothesis that TsaR regulates expression of the tsa operon (13) (Fig. 1) has been confirmed. Mutant 4A10 (Table 1) was generated by insertion of the Tn5 minitransposon in the tsaR gene, and the effect of this specific knockout (Table 1) was complemented by providing an inducible copy of tsaR in trans (mutant TT6; Table 1). Further, TsaRHis interacts specifically with the divergent tsa promoter region (Fig. 3) and its affinity is modulated by the putative inducer (Fig. 4) as required of a regulatory protein.

Knockout mutant 4A10 failed to grow with both TSA and TCA (Fig. 1), and utilization of both substrates was restored in mutant TT6 (Table 1). We thus confirmed the prediction that TsaR regulates the degradation of both compounds (13, 36); the gene products from tsaMBCD are used in the initial attacks on each substrate (Fig. 1) (21). Tralau et al. (unpublished data) showed that mutants in TSA transport prevent neither growth with TCA, which has a different transport system (22), nor normal growth with PSB, for which TsaB is required to complement the missing PszC (9, 14; J. Ruff, unpublished data) (Fig. 1). We thus conclude that tsaMBCD is directly affected by the knocking out of tsaR and that failure to grow is not a result of, e.g., positional effects.

Mutant 4A10 grew poorly with PSB (Table 1), and the deleterious effect of the mutation was corrected on complementation of TsaR (mutant TT6; Table 1). The surprise is that mutant 4A10 grew at all with PSB, given that TsaB is required to complement the missing PszC. We presume that an unidentified reductase is able to complement PszC partially.

Gel filtration chromatography indicated that active TsaR is a monomer in solution, though le Maire et al. (18) warn of errors sometimes encountered with this method. TsaR as the active monomer differs from several LysR-type regulators. One example of a monomeric regulator exists, AmpR (31), whereas ClcR is a dimer (6) and CysB and OxyR are tetramers (16, 44).

TsaR is sensitive to oxygen. Thus, gel filtration chromatography could not be done with TsaRHis bound to DNA. So it was impossible to determine whether the dissolved monomeric TsaRHis binds to its target sequence in a multimeric form, as LysR-type regulators usually do. We examined band shifts in agarose gels instead of polyacrylamide gels for the same reason, because separation is faster, thus allowing us to assay the interaction with DNA before TsaR lost its binding capacity.

It is unclear what interaction causes this oxygen sensitivity. Redox sensitivity has been described previously for the DNA-binding regulator FurS (ferric iron uptake) in Streptomyces reticuli, for which the effect is attributed to the redox state of cysteine residues (28). There was no sequence similarity between TsaR and FurS, so we presume that a different interaction occurs between oxygen and TsaR. Furthermore, reactivation of TsaR under nitrogen occurs in the absence of a reducing agent; the biochemical mechanism of this oxygen sensitivity is unknown. Given that TsaR regulates an operon encoding an oxygenase (TsaMB; Fig. 1) (13, 19) and that it seems to be a negative autoregulator (Tralau et al., unpublished), this sensitivity to oxygen may be useful to the cell. TsaR is very stable in the cell (Tralau et al., unpublished). One possible explanation is that TsaR in its active conformation is stable under conditions in which oxygen levels were low. This would reduce the turnover of active TsaR, and the transcription of the tsa operon would be repressed efficiently, because all binding sites would be occupied. Alternatively, the stability of TsaR would decrease in the presence of oxygen. This would increase the turnover of TsaR and allow transcription of the tsa operon to start more easily. The mechanism might thus help to compensate for the relatively high KD value (0.9 µM). This value is of the same magnitude as that for CatR, the regulator for catechol degradation at its internal binding site in catB in Pseudomonas putida (5), whereas the DNA-binding regulator ModE (26) shows higher affinity for its target promoter (24 nM).

The modulation by TSA of the interaction of TsaRHis with the whole intergenic tsaprom fragment (Fig. 4) involves regulatory sites for the expression of both tsaR and tsaMBCD, and we observed single shifts with this fragment only (Fig. 3). When we used shorter fragments of DNA, we obtained evidence for four binding regions and some double shifts, whose nature we do not understand (Table 4). Presumably the resolution of the agarose gels allowed double shifts to be detected with short fragments only. We realize that the multiple binding regions prevent a quantitative evaluation of the data presented in Fig. 4, but they imply that the system can react in a finely tuned manner. We also do not know how the ligands TSA and oxygen interact during the binding of TsaR to its target sites. However, regulatory complexity is already apparent at the physiological level, e.g., in defined mixed cultures utilizing multiple substrates (41).

Up to four binding sites for TsaR can be deduced in the sequence of tsaprom (Table 3 and Fig. 5), three of which may be derived from the consensus sequences (T-N11-A) presented by Schell for LysR-type regulators (35). BS I at positions 1376 to 1389 was proposed previously (13) and now has experimental support (subfragment I; Table 4). The location of BS II was not defined experimentally (Table 4 and Fig. 5), but of two possible binding motifs (35) at positions 1397 to 1409 and positions 1434 to 1446, the latter could bind TsaRHis without blocking the putative ribosomal binding site, the transcriptional start, the Pribnow box, or the -35 region for gene tsaR. We tentatively located BS III at positions 1562 to 1574 near the 5' end of tsaM; this site, derived from the consensus sequence, may contribute to the double shift with subfragment 2 (Table 4), but this could not be confirmed experimentally (subfragment 5; Table 4). BS IV, at positions 1384 to 1396, has clear experimental support in the absence of TSA (subfragment 4; Table 4).

We propose that in the absence of TSA, TsaR interacts at BS IV, which blocks the transcription of tsaR. In the presence of TSA, the regulator would shift to BS II (presumably position 1434 to 1446), thereby releasing the repression of transcription for tsaR and activating transcription of tsaMBCD. The existence of two additional binding sites is uncommon for LysR-type regulatory systems (35). The marginal activity of BS III resembles a situation seen with the CatR system in the catBCA operon of P. putida, which possibly represents a mechanism to save energy by reducing enzyme synthesis (5); induced TsaM represents 5% of soluble protein (19). The role of BS I may be involved with synthesis of TsaR, which reaches micromolar concentrations during growth (Tralau et al., unpublished).


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ACKNOWLEDGMENTS
 
We are grateful to M. Kertesz and J. M. Blatny, who kindly made bacteria available to us.

T.T. and J.M. were funded by the Deutsche Forschungsgemeinschaft (through A.M.C. and J.R.), the EU project SUITE (to A.M.C.), the University of Konstanz, and the Fonds der Chemischen Industrie.


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FOOTNOTES
 
* Corresponding author. Mailing address: Department of Biology, The University of Konstanz, Universitätsstr. 10, D-78457 Konstanz, Germany. Phone: 49 7531 88 2100. Fax: 49 7531 88 2966. E-mail: juergen.ruff{at}uni-konstanz.de. Back

{dagger} Present address: School of Biological Sciences, University of Manchester, Manchester, United Kingdom. Back

{ddagger} Present address: Institute of Biotechnology, Swiss Federal Institute of Technology, Zürich, Switzerland. Back


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REFERENCES
 
    1
  1. Ausubel, F. M., R. Brent, R. E. Kingston, D. D. Moore, J. G. Seidman, J. A. Smith, and K. Struhl. 1987. Current protocols in molecular biology. John Wiley & Sons, New York, N.Y.
    2. 2
  2. Behret, H., J. Ahlers, S. Ettel, E. Feicht, E. Futterer, I. Mangelsdorf, C. Pohlenz-Michel, H. Roß, H. Sterzl-Eckert, D. Vogel, L. Weis, and K. Widmann. 1991. p-Toluolsulfonsäure, Beratergremium für umweltrelevante Alstoffe (BUA)-Stoffberichte, vol. 63. Verlag Chemie, Weinheim, Germany.
    3. 3
  3. Blatny, J. M., T. Brautaset, H. C. Winther Larsen, P. Karunakaran, and S. Valla. 1997. Improved broad-host-range RK2 vectors useful for high and low regulated gene expression levels in gram-negative bacteria. Plasmid 38:35-51.[CrossRef][Medline]
    4. 4
  4. Bradford, M. 1976. A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Anal. Biochem. 72:248-254.[CrossRef][Medline]
    5. 5
  5. Chugani, S. A., M. R. Parsek, and A. M. Chakrabarty. 1998. Transcriptional repression mediated by LysR-type regulator CatR bound at multiple binding sites. J. Bacteriol. 180:2367-2372.[Abstract/Free Full Text]
    6. 6
  6. Coco, W. M., M. R. Parsek, and A. M. Chakrabarty. 1994. Purification of the LysR family regulator, ClcR, and its interaction with the Pseudomonas putida clcABD chlorocatechol operon promoter. J. Bacteriol. 176:5530-5533.[Abstract/Free Full Text]
    7. 7
  7. Coco, W. M., R. K. Rothmel, S. Henikoff, and A. M. Chakrabarty. 1993. Nucleotide sequence and initial functional characterization of the clcR gene encoding a LysR family activator of the clcABD chlorocatechol operon in Pseudomonas putida. J. Bacteriol. 175:417-427.[Abstract/Free Full Text]
    8. 8
  8. Collier, L. S., G. L. Gaines III, and E. L. Neidle. 1998. Regulation of benzoate degradation in Acinetobacter sp. strain ADP1 by BenM, a LysR-type transcriptional activator. J. Bacteriol. 180:2493-2501.[Abstract/Free Full Text]
    9. 9
  9. Cook, A. M., H. Laue, and F. Junker. 1999. Microbial desulfonation. FEMS Microbiol. Rev. 22:399-419.[CrossRef]
    10. 10
  10. de Lorenzo, V., and K. N. Timmis. 1994. Analysis and construction of stable phenotypes in gram-negative bacteria with Tn5- and Tn10-derived minitransposons. Methods Enzymol. 235:386-405.[Medline]
    11. 11
  11. Harley, C. B., and R. P. Reynolds. 1987. Analysis of E. coli promoter sequences. Nucleic Acids Res. 15:2343-2361.[Abstract/Free Full Text]
    12. 12
  12. Junker, F., and A. M. Cook. 1997. Conjugative plasmids and the degradation of arylsulfonates in Comamonas testosteroni. Appl. Environ. Microbiol. 63:2403-2410.[Abstract]
    13. 13
  13. Junker, F., R. Kiewitz, and A. M. Cook. 1997. Characterization of the p-toluenesulfonate operon tsaMBCD and tsaR in Comamonas testosteroni T-2. J. Bacteriol. 179:919-927.[Abstract/Free Full Text]
    14. 14
  14. Junker, F., E. Saller, H. R. Schläfli Oppenberg, P. M. H. Kroneck, T. Leisinger, and A. M. Cook. 1996. Degradative pathways for p-toluenecarboxylate and p-toluenesulfonate and their multicomponent oxygenases in Comamonas testosteroni strains PSB-4 and T-2. Microbiology (Reading) 142:2419-2427.[Abstract/Free Full Text]
    15. 15
  15. Kennedy, S. I. T., and C. A. Fewson. 1968. Enzymes of the mandelate pathway in bacterium N.C.I.B. 8250. Biochem. J. 107:497-506.[Medline]
    16. 16
  16. Kullik, I., J. Stevens, M. B. Toledano, and G. Storz. 1995. Mutational analysis of the redox-sensitive transcriptional regulator OxyR: regions important for DNA binding and multimerization. J. Bacteriol. 177:1285-1291.[Abstract/Free Full Text]
    17. 17
  17. Laue, H., J. A. Field, and A. M. Cook. 1996. Bacterial desulfonation of the ethanesulfonate metabolite of the chloroacetanilide herbicide metazachlor. Environ. Sci. Technol. 30:1129-1132.[CrossRef]
    18. 18
  18. le Maire, M., A. Ghasi, and J. V. Moller. 1996. Gel chromatography as an analytical tool for the characterisation of size and molecular mass of proteins. ACS Symp. Ser. 635.
    19. 19
  19. Locher, H. H., T. Leisinger, and A. M. Cook. 1991. 4-Toluene sulfonate methyl-monooxygenase from Comamonas testosteroni T-2: purification and some properties of the oxygenase component. J. Bacteriol. 173:3741-3748.[Abstract/Free Full Text]
    20. 20
  20. Locher, H. H., T. Leisinger, and A. M. Cook. 1989. Degradation of p-toluenesulphonic acid via sidechain oxidation, desulphonation and meta ring cleavage in Pseudomonas (Comamonas) testosteroni T-2. J. Gen. Microbiol. 135:1969-1978.[Abstract/Free Full Text]
    21. 21
  21. Locher, H. H., C. Malli, S. Hooper, T. Vorherr, T. Leisinger, and A. M. Cook. 1991. Degradation of p-toluic acid (p-toluenecarboxylic acid) and p-toluene sulphonic acid via oxygenation of the methyl sidechain is initiated by the same set of enzymes in Comamonas testosteroni T-2. J. Gen. Microbiol. 137:2201-2208.
    22. 22
  22. Locher, H. H., B. Poolman, A. M. Cook, and W. N. Konings. 1993. Uptake of 4-toluene sulfonate by Comamonas testosteroni T-2. J. Bacteriol. 175:1075-1080.[Abstract/Free Full Text]
    23. 23
  23. Mampel, J. 2000. Transport- und Regulationsphänomene beim Abbau von 4-Toluolsulfonat in Comamonas testosteroni. Ph.D. thesis. University of Konstanz, Konstanz, Germany.
    24. 24
  24. Mampel, J., J. Ruff, F. Junker, and A. M. Cook. 1999. The oxygenase component of the 2-aminobenzenesulfonate dioxygenase system from Alcaligenes sp. strain O-1. Microbiology (Reading) 145:3255-3264.[Abstract/Free Full Text]
    25. 25
  25. Martínez-Costa, O. H., A. J. Martín-Triana, E. Martínez, M. A. Fernández-Moreno, and F. Malpartida. 1999. An additional regulatory gene for actinorhodin production in Streptomyces lividans involves a LysR-type transcriptional regulator. J. Bacteriol. 181:4353-4364.[Abstract/Free Full Text]
    26. 26
  26. McNicholas, P. M., S. A. Rech, and R. P. Gunsalus. 1997. Characterization of the ModE DNA-binding sites in the control regions of modABCD and moaABCDE of Escherichia coli. Mol. Microbiol. 23:515-524.[CrossRef][Medline]
    27. 27
  27. Neuhoff, V., N. Arold, D. Taube, and W. Ehrhardt. 1988. Improved staining of proteins in polyacrylamide gels including isoelectric focusing gels with clear background at nanogram sensitivity using Coomassie brilliant blue G-250 and R-250. Electrophoresis 9:255-262.[CrossRef][Medline]
    28. 28
  28. Ortiz de Orué Lucana, D., and H. Schrempf. 2000. The DNA-binding characteristics of the Streptomyces reticuli regulator FurS depend on the redox state of its cysteine residues. Mol. Gen. Genet. 264:341-353.[CrossRef][Medline]
    29. 29
  29. Providenti, M. A., J. Mampel, S. MacSween, A. M. Cook, and C. R. Wyndham. 2001. Comamonas testosteroni BR6020 possesses a single genetic locus for extradiol cleavage of protocatechuate. Microbiology (Reading) 147:2157-2167.[Abstract/Free Full Text]
    30. 30
  30. Reese, M. G. 1998. NNPP-neural network promoter prediction. Baylor College of Medicine Human Genome Sequencing Center, Houston, Tex.
    31. 31
  31. Roh, I. K., I. J. Kim, J. H. Chung, and S. M. Byun. 1996. Affinity purification and binding characteristics of Citrobacter freundii AmpR, the transcriptional regulator of the ampC ß-lactamase gene. Biotechnol. Appl. Biochem. 23:149-154.
    32. 32
  32. Rosenberg, I. M. 1996. Protein analysis and purification. Birkhäuser, Boston, Mass.
    33. 33
  33. Sambrook, J., E. F. Fritsch, and T. Maniatis. 1989. Molecular cloning: a laboratory manual, 2nd ed. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y.
    34. 34
  34. Schägger, H., and G. von Jagow. 1987. Tricine-sodium dodecyl sulfate-polyacrylamide gel electrophoresis for the separation of proteins in the range from 1 to 100 kDa. Anal. Biochem. 166:368-379.[CrossRef][Medline]
    35. 35
  35. Schell, M. A. 1993. Molecular biology of the LysR family of transcriptional regulators. Annu. Rev. Microbiol. 47:597-626.[CrossRef][Medline]
    36. 36
  36. Schläfli Oppenberg, H. R., G. Chen, T. Leisinger, and A. M. Cook. 1995. Regulation of the degradative pathways from 4-toluenesulphonate and 4-toluenecarboxylate to protocatechuate in Comamonas testosteroni T-2. Microbiology (Reading) 141:1891-1899.
    37. 37
  37. Schwacha, A., and R. A. Bender. 1993. The product of the Klebsiella aerogenes nac (nitrogen assimilation control) gene is sufficient for activation of the hut operons and repression of the gdh operon. J. Bacteriol. 175:2116-2124.[Abstract/Free Full Text]
    38. 38
  38. Smith, C. L., S. R. Klco, and C. R. Cantor. 1988. Pulsed-field gel electrophoresis and the technology of large DNA molecules, p. 40-72. In K. E. Davies (ed.), Genome analysis: a practical approach. IRL Press, Oxford, United Kingdom.
    39. 39
  39. Tabatabai, M. A. 1974. Determination of sulfate in water sample. Sulphur Inst. J. 10:11-13.
    40. 40
  40. Thurnheer, T., T. Köhler, A. M. Cook, and T. Leisinger. 1986. Orthanilic acid and analogues as carbon sources for bacteria: growth physiology and enzymic desulphonation. J. Gen. Microbiol. 132:1215-1220.
    41. 41
  41. Tien, A. J. 1996. The physiology of a defined four-membered mixed bacterial culture during continuous cultivation with mixtures of three pollutants in synthetic sewage. Ph.D. thesis. Swiss Federal Institute of Technology, Zürich, Switzerland.
    42. 42
  42. Tralau, T., A. M. Cook, and J. Ruff. 2001. Map of the IncP1ß plasmid pTSA encoding the genes (tsa) for p-toluenesulfonate degradation in Comamonas testosteroni T-2 and conservation of these genes in different genetic backgrounds. Appl. Environ. Microbiol. 67:1508-1516.[Abstract/Free Full Text]
    43. 43
  43. Trevors, J. T., and M. E. Starodub. 1990. Electroporation of pKK1 silver-resistance plasmid from Pseudomonas stutzeri AG259 into Pseudomonas putida CYM318. Curr. Microbiol. 21:103-107.
    44. 44
  44. Tyrrell, R., K. H. G. Verschueren, E. J. Dodson, G. N. Murshudov, C. Addy, and A. J. Wilkinson. 1997. The structure of the cofactor-binding fragment of the LysR family member, CysB: a familiar fold with a surprising subunit arrangement. Structure 5:1017-1032.[Medline]

Applied and Environmental Microbiology, April 2003, p. 2298-2305, Vol. 69, No. 4
0099-2240/03/$08.00+0     DOI: 10.1128/AEM.69.4.2298-2305.2003
Copyright © 2003, American Society for Microbiology. All Rights Reserved.



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