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J Bacteriol. Dec 2006; 188(23): 8109–8117.
Published online Sep 29, 2006. doi:  10.1128/JB.01262-06
PMCID: PMC1698186

The ColRS Two-Component System Regulates Membrane Functions and Protects Pseudomonas putida against Phenol[down-pointing small open triangle]

Abstract

As reported, the two-component system ColRS is involved in two completely different processes. It facilitates the root colonization ability of Pseudomonas fluorescens and is necessary for the Tn4652 transposition-dependent accumulation of phenol-utilizing mutants in Pseudomonas putida. To determine the role of the ColRS system in P. putida, we searched for target genes of response regulator ColR by use of a promoter library. Promoter screening was performed on phenol plates to mimic the conditions under which the effect of ColR on transposition was detected. The library screen revealed the porin-encoding gene oprQ and the alginate biosynthesis gene algD occurring under negative control of ColR. Binding of ColR to the promoter regions of oprQ and algD in vitro confirmed its direct involvement in regulation of these genes. Additionally, the porin-encoding gene ompAPP0773 and the type I pilus gene csuB were also identified in the promoter screen. However, it turned out that ompAPP0773 and csuB were actually affected by phenol and that the influence of ColR on these promoters was indirect. Namely, our results show that ColR is involved in phenol tolerance of P. putida. Phenol MIC measurement demonstrated that a colR mutant strain did not tolerate elevated phenol concentrations. Our data suggest that increased phenol susceptibility is also the reason for inhibition of transposition of Tn4652 in phenol-starving colR mutant bacteria. Thus, the current study revealed the role of the ColRS two-component system in regulation of membrane functionality, particularly in phenol tolerance of P. putida.

The bacterial cell membrane forms a permeability barrier between the cell and its external environment and should fulfill two antagonistic functions. First, it must protect the bacterium against external toxic compounds, and low membrane permeability can evidently contribute to this function. Second, as the membrane is responsible for steady uptake of nutrients, high permeability is useful for that purpose. Thus, membrane permeability has to be flexibly regulated to achieve an optimum balance between these two functions (10, 31).

The cell envelope of gram-negative bacteria possesses two membranes, and its permeability is largely regulated by modulation of the outer membrane. The lipopolysaccharide-containing outer membrane is essential for barrier function, as it restricts movement of both hydrophilic and lipophilic compounds. Therefore, gram-negative bacteria are more resistant to lipophilic dyes, detergents, and most lipophilic antibiotics than gram-positive bacteria (30). In addition, gram-negative bacteria, particularly members of the genus Pseudomonas, are relatively tolerant to several aromatic compounds (reviewed in references 20 and 35). Cyclic hydrocarbons are deleterious to microbes, as they easily dissolve in the bacterial cell membrane, inhibiting its function (45). The most obvious consequence of accumulation of an organic solvent in the cell envelope is permeabilization of the membrane, leading to leakage of cellular metabolites and ions (14, 20). Therefore, when bacteria encounter a hazardous solvent, adaptive changes should occur in the structure of the cell membrane. For instance, alterations in membrane phospholipids (15, 33, 36), lipopolysaccharides (2), and membrane-embedded efflux pumps (23, 27, 40) have previously been reported to participate in increased solvent tolerance of bacteria.

Many bacterial adaptive responses to environmental changes are controlled by two-component signal transduction systems (46). In a typical two-component system, an environmental stimulus is first detected by a transmembrane histidine sensor kinase, which is capable of autophosphorylation. The phosphoryl group is then transferred from a histidine to an aspartic acid residue in the second component of the pathway, a response regulator. Usually, the phosphorylated response regulator binds to the DNA, resulting in either activation or repression of target genes (see, e.g., reference 39). Two-component systems respond to a variety of environmental signals and regulate numerous functions, including cell division, sporulation, motility, metabolism, communication, virulence, stress adaptation, etc. There is also evidence of participation of two-component systems in regulation of membrane permeability of bacteria (12, 29, 34, 48).

The ColRS two-component signal transduction pathway was first characterized in Pseudomonas fluorescens as a system involved in the ability of bacteria to colonize plant roots (7). A publication by Ramos-González et al. (37) showed that it can also be true for P. putida, as the promoter of colRS was activated in the maize rhizosphere. In a recent paper, we demonstrated that the P. putida two-component system ColRS is necessary for the accumulation of phenol-utilizing (Phe+) mutants, which arise due to transposition of Tn4652 (18). However, the checkpoint of response regulator ColR in both plant root colonization and regulation of Tn4652 transposition is yet unclear. Therefore, to determine ColR-controlled cell functions in P. putida we performed a search for ColR-regulated genes by using a promoter library from total chromosomal DNA of P. putida. The screen identified several target genes of ColR and revealed the role of ColR in regulation of membrane functionality, particularly its involvement in phenol tolerance of P. putida.

MATERIALS AND METHODS

Bacterial strains, plasmids, and media.

All P. putida strains used in this study originated from PaW85, which is isogenic to fully sequenced KT2440 (www.tigr.org). Bacteria were grown in Luria-Bertani (LB) medium (28) or in M9 minimal medium (1) containing either 10 mM glucose or 10 mM citrate. The phenol concentrations in minimal medium are specified in the text, as they varied between the experiments. When necessary, the growth medium was supplemented with ampicillin (100 μg ml−1), chloramphenicol (20 μg ml−1), kanamycin (50 μg ml−1), or streptomycin (20 μg ml−1) for Escherichia coli and with carbenicillin (1,500 μg ml−1), chloramphenicol (300 μg ml−1), kanamycin (50 μg ml−1), or streptomycin (500 μg ml−1) for P. putida. P. putida was incubated at 30°C and E. coli at 37°C. E. coli was transformed with plasmid DNA as described by Hanahan (13). P. putida was electrotransformed according to the protocol of Sharma and Schimke (44).

Construction of plasmids and strains.

Plasmids and strains are listed in Table Table11 and oligonucleotides used in PCR amplifications in Table Table2.2. The plasmid for the promoter library was designed on the basis of the low-copy-number plasmid pPR9TT. First, the lacZ gene, excised from pPR9TT as a HindIII-NotI fragment, was replaced by gusA, which was obtained from pKS/gusA by use of the same restriction enzymes. The resulting promoter probe plasmid, p9TTgusA, possesses HindIII, SalI, XhoI, KpnI, and BglII as unique restriction sites for promoter cloning. In order to add the second reporter gene, pheA, into p9TTgusA, it was first excised from pEST1332 as a PstI-NheI fragment and subcloned into PstI-SmaI-opened pUC18Not. The pheA-containing NdeI-KpnI fragment from pUCNot/pheA was inserted into HindIII-KpnI-cleaved p9TTgusA, resulting in the plasmid p9TTpheAgusA (Fig. (Fig.11).

FIG. 1.
Map of p9TTpheAgusA and schematic representation of a strategy used to select ColR-regulated promoters. The plasmid p9TTpheAgusA, employed for the construction of a promoter library of P. putida PaW85 genomic DNA, contains promoterless phenol monooxygenase ...
TABLE 1.
Bacterial strains and plasmids
TABLE 2.
Oligonucleotides

In order to construct a low-copy-number lacZ-based promoter probe plasmid, we employed plasmid pPR9TT. However, as pPR9TT contained lacZ without the ATG codon (42), we deleted it by using BamHI. The resulting plasmid, p9TTB, opened with Ecl136II-XbaI, was used for cloning of full-length lacZ obtained from pKRZ-1 as a PstI-XbaI fragment. The promoter probe plasmid p9TTBlacZ was used to clone the promoters of oprQ, algD, ompAPP0773, and csuB. The 613-bp-long oprQ promoter region was amplified from the chromosomal DNA of P. putida PaW85 by use of oligonucleotides 267Sma and oprE3Kpn. The PCR-amplified DNA fragment was digested with SmaI and KpnI and ligated into BglI-KpnI-cleaved p9TTBlacZ, yielding p9TToprQ. The algD, ompAPP0773, and csuB promoters were cut from the respective library plasmids and subcloned in front of lacZ in p9TTBlacZ as follows: the 695-bp-long Acc65I-SmaI-generated algD promoter-containing fragment was inserted into Acc65I-BglII-cleaved p9TTBlacZ, resulting in plasmid p9TTalgD; the 668-bp-long Acc65I-generated ompAPP0773 promoter fragment was inserted into Acc65I-opened p9TTBlacZ, resulting in plasmid p9TTompA; and the 443-bp-long Ecl136II-Eco47III-generated csuB promoter fragment was inserted into SmaI-opened p9TTBlacZ, resulting in plasmid p9TTcsuB.

To disrupt the oprB1 gene in P. putida PaW85 and PaWcolR, the coding region of oprB1 was PCR amplified from chromosomal DNA of PaW85 with oligonucleotides oprB1ylem and oprB1all. The oprB1-containing PCR product was cloned into HincII-opened pBluescript KS, resulting in pKS/oprB1. The central 590-bp region of oprB1 in pKS/oprB1 was excised with HincII and EheI and replaced with the Smr gene cut from pUTmini-Tn5Sm/Sp with VspI. The obtained oprB1::Sm sequence was liberated from pKS/oprB1::Sm with EcoRI and inserted into EcoRI-opened pGP704L. The interrupted oprB1 gene was inserted into the chromosome of P. putida PaW85 and PaWcolR by homologous recombination. Plasmid pGP704L/oprB1::Sm was conjugatively transferred from E. coli CC118 λpir (17) into P. putida PaW85 and PaWcolR by use of a helper plasmid, pRK2013 (11). The oprB1 knockout strains were verified by PCR analysis.

Construction of promoter library and selection of colR-regulated promoters.

Chromosomal DNA of P. putida PaW85 was partially digested with Sau3AI and separated by agarose gel electrophoresis. Fragments of ~1 kb were purified and cloned into BglII-opened and shrimp alkaline phosphatase-treated p9TTpheAgusA in front of the promoterless pheA and gusA genes (Fig. (Fig.1).1). The pool of fusions was electrotransformed into E. coli DH5α, and after 1 h of growth without the antibiotic the bacteria were inoculated into 2 liters of LB supplied with chloramphenicol for the amplification of library plasmids. Bacteria were grown to an optical density at 580 nm of 0.6, and total plasmid DNA was isolated. Analysis of the bacterial culture before the amplification step revealed that about 130,000 clones were obtained. PCR analysis of a set of these clones, done using oligonucleotides pheA and T1T2 (Table (Table2),2), demonstrated that over 80% of library clones had an insert in the plasmid. Half of the inserts were about 1 kb in size, and the rest contained inserts of 2 to 3 kb.

For the selection of ColR-regulated promoters, the library DNA was electroporated into P. putida PaWRtaccolRD51A and bacteria were plated onto minimal media containing phenol (2.5 mM) and X-Gluc (5-bromo-4-chloro-3-indolyl-β-d-glucuronic acid) with and without 0.5 mM IPTG (isopropyl-β-d-thiogalactopyranoside). Only cells that carry plasmids with a promoter in their chromosomal insert can grow on phenol and yield blue colonies. In strain PaWRtaccolRD51A, expression of phosphorylation-deficient ColR can be induced with IPTG (18). Therefore, we replica plated the obtained colonies (altogether about 10,000) onto phenol plates with and without IPTG to reveal ColR-regulated promoters (Fig. (Fig.1).1). Colonies which were apparently affected by ColR were picked up, and their plasmid DNA was isolated. In order to prove that the library plasmid indeed contained a ColR-regulated promoter, wild-type and colR knockout P. putida strains were transformed with the plasmid of interest and recombinant colonies were tested on phenol plates containing X-Gluc.

Enzyme assay.

All enzyme activities presented in this paper were measured from solid-medium-grown bacteria. Bacteria grown on either glucose or citrate containing M9 minimal medium were scraped off from the plates using toothpicks and suspended in M9 solution. β-Galactosidase activity was assayed according to a previously described protocol (47).

MIC assay.

The MIC of phenol was measured on agar plates containing 10 mM glucose or citrate as the carbon source. Bacterial culture grown overnight in LB medium was serially diluted into M9 medium, and 5 μl of diluted cultures was spotted onto agar plates supplemented with 4, 6, or 8 mM phenol and incubated at 30°C.

Gel mobility shift assay.

For the gel mobility shift assay, ColR and N-terminally truncated ColS were purified as His-tagged proteins by a previously described protocol (18). The DNA probes were generated by PCR as follows: the 453-bp oprQ promoter region was amplified from plasmid p9TToprQ by use of oligonucleotides 267start and 268gatc, the 461-bp algD promoter region was amplified from p9TTalgD by use of oligonucleotides algDprom and PP1289stop, the 373-bp ompA promoter region was amplified from p9TTompA by use of oligonucleotides ompAATG and ompASma, and the 443-bp csuB promoter region was amplified from p9TTcsuB by use of oligonucleotides Fw and T1T2 (Table (Table2).2). Before PCR amplification, one oligonucleotide was end labeled by phosphorylation with [γ-32P]ATP. The labeled DNA fragments were purified by native 5% polyacrylamide gel electrophoresis, eluted, and resuspended in water. For the binding reaction, different amounts of purified His-tagged ColR protein were incubated (for about 20 min) with 1,000 cpm of labeled DNA fragment in 20 μl of buffer A (25 mM Tris-HCl [pH 7.5], 10 mM MgCl2, 1 mM CaCl2, 0.05 mM EDTA, 50 mM KCl, 50 μg ml−1 of bovine serum albumin, 100 μg ml−1 of salmon sperm DNA, and 5% glycerol). To test binding of phosphorylated ColR with the DNA probe, ColS was first autophosphorylated by incubation in buffer A containing 0.1 mM ATP for 15 min. After addition of ColR to the phosphorylated ColS and further incubation for 15 min, labeled DNA was added to the reaction mixture. After 20 min of further incubation, the reaction mixtures were applied to a 5% nondenaturing polyacrylamide gel buffered with 0.5× Tris-borate-EDTA (pH 7.5). Electrophoresis was carried out at 4°C at 10 V/cm for 3 h. The gels were dried under vacuum and exposed to a PhosphorImager screen (Molecular Dynamics).

Transposition assay.

Transposition of native Tn4652 was examined in an assay that involves starvation of bacteria on phenol minimal plates (19). The transposition assay was carried out using P. putida PaW85 wild-type and colR mutant strains carrying the target plasmid pEST1332. Bacteria were grown overnight in LB medium at 30°C and washed in M9 solution. Approximately 1 × 108 cells from at least four independent overnight cultures of both P. putida strains were plated onto phenol minimal plates, and accumulation of mutant Phe+ colonies was monitored upon incubation of plates at 30°C for 7 days. Generally, 2.5 mM phenol was used for selection. However, to test the phenol sensitivity of P. putida PaW85 and PaWcolR, 0.25, 0.5, 1, 2.5, and 5 mM phenol concentrations were used.

RESULTS

Screening of P. putida promoter library to detect ColR-regulated genes.

According to the sequence analysis, ColR is a potential transcription factor (7, 18). However, no information on target genes of ColR is available yet. In order to find promoters regulated by ColR, we constructed a plasmid library of transcriptional fusions between P. putida PaW85 chromosomal DNA and promoterless reporter genes. Based on our previous results showing that deficiency of ColR affects transpositional activity of Tn4652 in P. putida starving on phenol (18), we used the promoterless phenol monooxygenase (pheA) gene as a primary reporter in the library vector. pheA enables promoter selection on phenol minimal plates, i.e., conditions under which we had previously detected the effect of ColR on transposition of Tn4652. The second reporter, gusA, encoding β-glucuronidase, locates immediately downstream of pheA in the plasmid p9TTpheAgusA, enabling evaluation of the promoter strength (Fig. (Fig.1).1). The promoter library constructed in p9TTpheAgusA was introduced into a P. putida colR knockout strain harboring an extra copy of a mutated colR gene, colR(D51A), in the chromosome under the control of the Ptac promoter and the lacIq repressor (strain PaWRtaccolRD51A). colR(D51A) encodes a phosphorylation-deficient ColR with alanine at position 51 instead of aspartate. The choice of reporter strain was based on our previous observation that the induction of phosphorylation-deficient ColR(D51A) caused a stronger effect on Tn4652 transposition than did overexpression of wild-type ColR (18). Using the selection strategy described in Materials and Methods and illustrated in Fig. Fig.1,1, we obtained over 100 clones potentially containing a ColR-regulated promoter. Plasmid DNA from these clones was isolated and reintroduced into wild-type and colR mutant strains, and four library plasmids with a ColR-regulated promoter were verified. Partial sequencing of chromosomal inserts of the plasmids revealed upstream regions of P. putida genes PP0268, PP0773, PP1288, and PP2357 just in front of the reporter pheA. PP0268 and PP0773 are annotated as porin-encoding oprQ and ompA family genes according to P. putida KT2440 genome data in TIGR (www.tigr.org). PP1288 (algD), encoding a GDP-mannose 6-dehydrogenase, is the first gene of the algD-8-44-KEGXLIJFA operon which is responsible for alginate synthesis. PP2357 (csuB) is the first gene of the operon putatively encoding for type I pilus proteins. Thus, all four genes detected in our screen are responsible for cell surface and membrane functions.

Transcriptional analysis of oprQ, ompAPP0773, algD, and csuB promoters.

To test whether oprQ, ompAPP0773, algD, and csuB are indeed controlled by ColR, the upstream regions of these genes were cloned into a low-copy-number promoter probe plasmid, p9TTBlacZ. The β-galactosidase activity assay revealed moderate repression of the transcription from oprQ and algD promoters by ColR (Fig. 2A and B), while no ColR dependency was observed in the case of the ompAPP0773 and csuB promoters (Fig. 2C and D). These results contradicted the initial data of the library screen on phenol plates, which showed remarkable ColR dependency of these promoters, in particular, those of algD and ompAPP0773 (data not shown). As the library selection was performed on phenol plates, we presumed that phenol may affect the promoters identified as ColR regulated. To test the hypothesis, β-galactosidase activity was measured in bacteria grown on glucose supplemented with 4 mM phenol. The assay revealed that phenol indeed influenced all of these promoters (Fig. (Fig.2).2). However, the effect of phenol on promoters depended significantly on genetic background of the strains. In wild-type bacteria, no effect or only a slight effect of phenol was observed, while in a colR mutant strain, the promoters strongly responded to phenol. For example, in a P. putida colR mutant strain, the algD promoter was 60-fold induced in the presence of phenol, while in the wild type no induction by phenol was detected (Fig. (Fig.2B).2B). In contrast to promoters of oprQ, ompAPP0773, and algD, which were activated by phenol in a colR mutant strain, the csuB promoter was downregulated (Fig. (Fig.22).

FIG. 2.
β-Galactosidase (β-Gal) activities measured in wild-type (wt) and colR knockout (colR) P. putida strains carrying the reporter plasmids with promoters of oprQ, algD, ompAPP0773, or csuB (p9TToprQ, p9TTalgD, p9TTompA, or p9TTcsuB, respectively). ...

Thus, transcriptional analysis of oprQ, ompAPP0773, algD, and csuB promoters revealed that ColR can affect these promoters in two different ways. First, ColR appears to be a negative transcriptional regulator of the oprQ and algD genes. Additionally, all four promoters are influenced by phenol, the effect being especially prominent in the colR knockout background.

Purified ColR binds to the promoters of oprQ and algD in vitro.

The effect of ColR on the promoters of oprQ and algD was quite weak in glucose-grown bacteria (Fig. 2A and B). To test whether ColR can directly interact with these promoters, we performed a gel mobility shift assay using the purified ColR protein and the DNA fragments with promoter regions of oprQ or algD. The results (Fig. (Fig.3)3) demonstrate binding of ColR to both promoter DNAs. We also tested the possibility that the phosphorylation signal from the sensor kinase ColS may influence the binding affinity of ColR. Data in Fig. Fig.33 show that lower amounts of ColR were needed to shift the DNA in the presence of ColS and ATP, indicating that phosphorylation of ColR increases its affinity to oprQ and algD promoters. Thus, the in vitro binding assay suggests that ColR is directly involved in regulation of oprQ and algD and that the phosphorylation signal from ColS can influence the extent of this regulation.

FIG. 3.
Specific binding of purified ColR to oprQ and algD promoter DNA. Shown is a gel mobility shift analysis of binding of purified ColR to oprQ, algD, and ompA promoter DNA. Approximately 0.5 ng (1,000 cpm) of the DNA probe was incubated with different amounts ...

The promoters of ompAPP0773 and csuB did not bind ColR when assayed under the same conditions (Fig. (Fig.33 [only results with ompAPP0773 DNA are presented]). Therefore, we concluded that ColR affects the transcription from ompAPP0773 and csuB promoters indirectly.

Glucose transport through OprB1 porin enhances ColR-dependent phenol sensitivity of P. putida.

During the analysis of oprQ, ompAPP0773, algD, and csuB promoter fusions, we observed that the colR background-dependent phenol effect on promoters depended on a growth substrate. The effect of phenol was clearly evident for glucose-grown colR mutant bacteria but was undetectable in the case of the cells grown on other carbon sources (citrate, succinate, fructose, Casamino Acids, or benzoate). For example, if a colR mutant strain grows on citrate plus phenol medium, no induction of the ompAPP0773 promoter occurs (Fig. (Fig.4A).4A). Similar results were also obtained with oprQ, algD, and csuB promoter fusions (data not shown). Thus, glucose-growing colR mutant bacteria responded to phenol in a more pronounced way than respective bacteria grown on other substrates. From these results, we hypothesized that glucose transport into colR mutant bacteria may also facilitate the entry of phenol. The hypothesis is supported by literature data by Wierckx and coauthors (50) indicating that glucose transport may contribute to phenol entrance in P. putida. These authors used a solvent-tolerant P. putida S12 strain for bioproduction of phenol from glucose. As disruption of the gene for glucose transport porin OprB1 (PP1019) caused enhanced phenol production, they suggested that phenol may reenter the cell via that porin. Therefore, we asked whether the glucose-inducible porin B1 (43) may be implicated in the phenol response of promoters in a glucose-growing P. putida colR mutant strain. To answer the question, the promoters were analyzed in the colR oprB1 double mutant background. A β-galactosidase activity assay demonstrated that in a colR oprB1 double mutant strain, phenol did not influence transcription from the ompAPP0773 promoter (Fig. (Fig.4B).4B). Analysis of oprQ, algD, and csuB promoter fusions gave similar results (data not shown). These data indicate that glucose porin B1 may be involved in increased phenol entry into P. putida.

FIG. 4.
β-Galactosidase (β-Gal) activities measured in wild-type (wt) and colR, oprB1, and colR oprB1 knockout P. putida strains carrying ompAPP0773 promoter-containing plasmid p9TTompA. Bacteria were grown either on citrate (citr) and citrate ...

P. putida colR mutant strain has enhanced susceptibility to phenol.

The results presented above indicate that phenol may enter into colR mutant bacteria more easily than into the wild type. To test the assumption, phenol tolerance of P. putida wild-type and colR mutant strains was assayed on glucose and citrate plates supplemented with different concentrations of phenol. Results presented in Fig. Fig.55 demonstrate that the colR mutant bacteria are more sensitive to high phenol concentrations than the wild type on both carbon sources. However, the MIC of phenol for the colR mutant strain was dependent on the growth substrate, being 8 mM on glucose medium and 6 mM on citrate medium (Fig. (Fig.5).5). At the same time, the colR knockout strain demonstrated retarded growth on glucose medium already in the presence of 4 mM phenol, while bacteria growing on citrate were not affected by that low phenol concentration (Fig. (Fig.5).5). Notably, the sensitivity of colR mutant bacteria to phenol on glucose medium was partially alleviated by the oprB1 knockout (Fig. (Fig.55).

FIG. 5.
Plate assay for phenol tolerance of wild-type (wt) or oprB1, colR, and colR oprB1 knockout P. putida strains. The minimal media contain either glucose (glc) or citrate (citr) as the carbon source. Concentration of added phenol (phe) is indicated above ...

These results demonstrate that in general the P. putida colR mutant strain has reduced phenol tolerance but that the extent of the effect of phenol depends on the carbon source.

Transposition of Tn4652 in a colR mutant strain is affected by phenol concentration.

According to our recent data, the P. putida ColRS signal system is necessary for the emergence of Phe+ mutants arising due to transposition of chromosomal Tn4652 in front of the silent phenol monooxygenase pheA gene in the plasmid pEST1332 (18). As the present study suggests involvement of the ColRS system in phenol tolerance of P. putida, we asked whether the emergence of Phe+ mutants can be affected by phenol concentration in the medium. To test this, the wild-type and colR-defective strains harboring transposition target plasmid pEST1332 were starved on solid media containing different concentrations of phenol (0.25, 0.5, 1, 2.5, and 5 mM). Figure Figure6A6A shows that in the wild-type P. putida strain PaW85, phenol concentration had no effect on frequency of occurrence of Tn4652-linked Phe+ mutants. However, accumulation of Phe+ mutants in a colR knockout strain depended on phenol concentration: elevated concentrations clearly inhibited Tn4652 transposition (Fig. (Fig.6B).6B). At low (0.25 mM) phenol concentration, transposition was largely recovered, suggesting that a reduced tolerance of colR mutant bacteria to phenol may be the reason for inhibition of accumulation of Phe+ mutants.

FIG. 6.
Accumulation of Tn4652 transposition-dependent Phe+ mutants on plates with different concentrations of phenol in a P. putida wild-type (wt) strain (A) and in its colR knockout derivative (B). Each presented value represents the mean ± ...

DISCUSSION

Accumulation of organic solvents like alcohols or phenols in bacterial membranes increases membrane permeability, reduces energetic status of the cell, and affects function of many membrane-embedded proteins (20). Studies of bacterial solvent tolerance have revealed several mechanisms developed by bacteria to overcome solvent-caused toxic effects and to adapt to a solvent-containing environment (35). Here, we demonstrate that the ColRS two-component signal system regulates phenol tolerance of P. putida, most probably by adjusting permeability of the membrane to phenol.

The promoter library screen revealed the membrane as a target of the P. putida ColRS system. ColR directly regulated oprQ, encoding the outer membrane porin OprE3, and algD, the first gene of an operon for biosynthesis and degradation of surface polysaccharide alginate (www.tigr.org). In addition, ColR indirectly affected expression of two other membrane-related genes: ompAPP0773, encoding an outer membrane protein, and the type I pilus gene csuB. The finding that all ColR-regulated genes discovered by us contribute to membrane functions is in good accordance with the phenol-sensitive phenotype of the colR knockout P. putida strain. The MIC assay demonstrated that phenol tolerance of the colR knockout strain is generally reduced (Fig. (Fig.5).5). Transcriptional analysis of the promoters of phenol-responsive genes oprQ, algD, ompAPP0773, and csuB (Fig. (Fig.2)2) and assay of transpositional activity of Tn4652 (Fig. (Fig.6)6) also indicated that the P. putida colR mutant strain is more sensitive to phenol than the wild type. These data suggest that membrane permeability of colR knockout bacteria is altered, becoming particularly leaky to phenol. Lowered phenol tolerance is obviously also the reason for the enhanced response of oprQ, algD, ompAPP0773, and csuB promoters to phenol in colR knockout bacteria (Fig. (Fig.2).2). Among these promoters, the promoter of algD was particularly highly induced in the presence of phenol in the colR knockout strain. The role of alginate in P. putida has not been studied, but in P. aeruginosa it is implicated in biofilm formation and in resistance of bacteria to antibiotics and to the immune system of the host (16, 38). To draw parallels with the proposed protective function of alginate in P. aeruginosa (16, 25, 38), one may hypothesize that under phenol stress the increased synthesis of alginate may contribute to phenol tolerance of P. putida. In the current study, the csuB promoter was the only promoter downregulated by phenol in the colR mutant background. Most interestingly, the literature data on toluene stress in bacteria also show repression of the motility/chemotaxis and pilus/flagellum-related genes (9). Since motility is a highly energy-consuming function, downregulation of motility genes may serve as an adaptive response to save energy in coping with solvent-caused stress (9).

It is not clear how phenol crosses the cell membrane, and it is generally believed that phenol just dissolves in membranes. However, some literature data show that several aromatic compounds may enter the cell periplasm through porins in the outer membrane (21, 22, 26, 49). For example, transport of xylene across the outer membrane proceeds through the XylN porin, which is encoded as a part of the operon required for the catabolism of toluene and xylene (22). Intriguingly, our results indicate that phenol may enter the cells of P. putida through the glucose porin OprB1, as disruption of oprB1 significantly alleviated phenol sensitivity of colR mutant bacteria growing on glucose plus phenol (Fig. (Fig.5).5). Glucose-inducible OprB1 not only allows high-affinity glucose transport but also enables uptake of other sugars (43, 51). It has been hypothesized that the presence of a hydroxyl group may be sufficient to permit binding of a compound to OprB1 (51). Consistent with our results, a previous study of solvent-tolerant P. putida S12 also suggests entering of phenol into the cell via porin B1 (50). The presence of OprB1 affected both the MIC for phenol (Fig. (Fig.5)5) and phenol responsiveness of the promoters specifically in a colR mutant background (Fig. (Fig.4).4). Therefore, we suppose that membrane permeability of colR mutant bacteria to phenol is intrinsically increased and that additional phenol uptake through OprB1 porin may elevate the concentration of phenol in colR mutant bacteria over a threshold level, increasing physiological stress and enhancing the phenol response of the colR mutant strain.

Transposition of Tn4652, needed for generation of the Phe+ phenotype in phenol-starving P. putida, is severely inhibited in bacteria defective in the colRS two-component system (18). Moreover, ColR is also required for accumulation of other Phe+ phenotype-generating mutations in P. putida. Namely, the frequency of frameshift and base substitution mutations restoring the open reading frame of the pheA gene was seriously reduced in a colR mutant strain (data not shown). Thus, different mutational processes occurring under phenol selection seem to be repressed in P. putida colR mutant strains. The current study demonstrates that these phenotypes may be related to the reduced tolerance of colR knockout bacteria to phenol. However, though colR knockout bacteria demonstrated increased sensitivity to phenol, they did not die during starvation on phenol plates (18). Therefore, even if the concentration of phenol in colR mutant bacteria is higher than that in the wild type, it is not easy to understand the reason for repression of mutations. It is possible that phenol disturbs the DNA turnover in starving colR mutant bacteria. Indeed, studies of the stationary-phase (adaptive) mutation have revealed that a remarkable DNA turnover is needed for the emergence of mutations in starving populations of bacteria (6). As both transposition and point mutations require DNA synthesis, inhibition of DNA replication by phenol overdose should explain the suppression of mutational processes in starving bacteria. Growing evidence suggests that the bacterial membrane plays multiple crucial roles in regulation of chromosomal replication (5) and that destabilization of membrane integrity may impair DNA replication (3). Therefore, the DNA turnover in colR mutant bacteria may be disturbed due to phenol-caused damage of the membrane. Another conceivable reason for the reduction of DNA turnover in colR mutant bacteria may be the decrease of the energetic status of the cell, as accumulation of hydrophobic organics in the cell membrane causes leakage of protons and ATP (45). Hopefully, further experiments will lead us towards an understanding of the relationship between ColR-controlled membrane functions and regulation of mutational processes in P. putida.

Acknowledgments

We are grateful to Tiina Alamäe, Inga Sarand, and Niilo Kaldalu for critically reading the manuscript.

This work was supported by grant 5758 from the Estonian Science Foundation and grant HHMI 55000316 from the Howard Hughes Medical Institute International Research Scholars Program to M.K. and by grant 6025 from the Estonian Science Foundation to R.H.

Footnotes

[down-pointing small open triangle]Published ahead of print on 29 September 2006.

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