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Osmoregulation  in Lactococcus lactis:  BusR,  a transcriptional repressor  of the glycine betaine uptake system  BusA

SUMMARY

The busA (opuA) locus of Lactococcus lactis encodes a glycine betaine uptake system. Transcription of busA is osmotically inducible and its induction after an osmotic stress is reduced in the presence of glycine betaine. Using a genetic screen in CLG802, an Escherichia coli strain carrying a lacZ transcriptional fusion expressed under the control of the busA promoter, we isolated a genomic fragment from the L. lactis subsp. cremoris strain MG1363, which represses transcription from busA _p . The cloned locus responsible for this repression was identified as a gene present upstream from the busA operon, encoding a putative DNA binding protein. This gene was named busR. Electrophoretic mobility shift and footprinting experiments showed that BusR is able to bind a site that overlaps the busA promoter. Overexpression of busR in L. lactis reduced expression of busA. Its disruption led to increased and essentially constitutive transcription of busA at low osmolarity. Therefore, BusR is a major actor of the osmotic regulation of busA in L. lactis.

INTRODUCTION

In natural environments, bacteria frequently face variations in the osmolarity of the surrounding medium. At elevated osmolarity, bacterial cells restore turgor, thought to be the motor of their elongation (Bremer and Krämer, 2000), by accumulating osmolytes in the cytoplasm (Csonka and Hanson, 1991; Kempf and Bremer, 1998; Bremer and Krämer, 2000). These include ions, such as K+, and a number of osmoprotectant organic compounds, the so-called compatible solutes, which can accumulate to very high levels in the cytoplasm (Galinski and Trüper, 1994). The specific induction of a number of genes by elevated osmolarity is a key step in adaptation to hyperosmotic conditions (Csonka and Hanson, 1991; Lucht and Bremer, 1994; Burg et al., 1996). The mechanisms of osmotic induction of transcription have been extensively studied in the Gram-negative bacteria Escherichia coli and Salmonella typhimurium. The systems that are best understood at present are: kdp, a high affinity K⁺ uptake system (Laimins et al., 1981); ompC/ompF, encoding major outer membrane porins (Hall and Silhavy, 1981); and proP and proU, encoding uptake systems for the compatible solute glycine betaine (Cairney et al., 1985; Dunlap and Csonka, 1985; Gowrishankar, 1986; May et al., 1989). Regulation of kdp involves a two-component phosphotransfer system composed of KdpD, a membrane-bound histidine kinase receptor, and the response regulator KdpE (Nakashima et al., 1992; Walderhaug et al., 1992). It has been proposed that KdpD senses turgor (Malli and Epstein, 1998). However, alternative mechanisms have also been proposed and this issue remains a matter of debate (Asha and Gowrishankar, 1993; Sugiura et al., 1994; Frymier et al., 1997; Jung et al., 2000; 2001). Transcription of ompC is induced in media of elevated osmolarity, whereas ompF is induced in low-osmolarity media (Hall and Silhavy, 1981). This osmotic regulation system also depends on a two-component regulatory system, involving the transmembrane sensor EnvZ and the transcriptional regulator OmpR (Forst and Roberts, 1994). EnvZ senses the osmotic signal, but how this is achieved is still unclear. OmpR and EnvZ control the osmotic regulation of the plasmid-encoded vir genes in Shigella flexneri (Bernardini et al., 1990) but they do not seem to affect the expression of other osmotically stimulated systems in E. coli K12 (Cairney et al., 1985; May et al., 1986; Gutierrez et al., 1987). Transcription of proU increases proportionally to the osmolarity, even in conditions in which turgor is thought to be maintained. Osmotic induction of proU is dependent on accumulation of K+ in the cytoplasm (Sutherland et al., 1986), but independent of the accumulation of its major physiological counter-ion glutamate (Csonka et al., 1994). Despite of extensive searches, no trans-acting mutation able to abolish proU osmoregulation has been found. Therefore, it has been proposed that proU regulation is not achieved by a specific regulatory protein, but by a more global phenomenon. In an in vitro reconstituted system, K⁺ glutamate was identified as a signal that stimulates proU expression (Jovanovich et al., 1989; Ramirez et al., 1989), and results obtained with purified components suggested direct action of K⁺ glutamate on the transcription complex (Prince and Villarejo, 1990). Although this is an attractive hypothesis, as accumulation of K⁺ glutamate in the cytoplasm is the primary response of enterobacteriae subjected to high osmolarity (Dinnbier et al., 1988), the magnitude of the K⁺ glutamate effect is not sufficient to explain the almost 100-fold osmotic induction of proU. Other studies indicated that osmolarity-dependent variations in DNA supercoiling are involved in proU osmotic regulation (Higgins et al., 1988). A number of studies established that a major element of proU osmotic regulation is a so-called 'transcriptional silencer' localized within proV, the first gene of the operon, which is able to repress transcription at low osmolarity (Fletcher and Csonka, 1995). Binding of the nucleoid-associated protein H-NS to this region exerts an inhibitory effect on proU transcription and modulation of this repression by DNA supercoiling or K+ glutamate probably contributes to the osmoregulation of proU transcription (Ueguchi and Mizuno, 1993; Lucht and Bremer, 1994). The proP gene is transcribed under the control of two promoters, P1 and P2 (Mellies et al., 1995). Both promoters can be induced by osmotic shock, but through different mechanisms. Transcription from P2 is dependent on the ^s sigma factor and activated by the Fis protein (Mellies et al., 1995; Xu and Johnson, 1995; 1997a). Its osmotic induction is probably mediated by the accumulation of ^s that follows an increase in osmolarity (Lange and Hengge-Aronis, 1994; Xu and Johnson, 1995). The proP P1 promoter is independent of ^s and repressed by binding of the CRP-cAMP complex to a site overlapping the 35 region of the promoter (Xu and Johnson, 1997b). proP P1 exhibits a strong induction after an osmotic shock that occurs through an original mechanism, the release of the repression exerted by CRP-cAMP (Landis et al., 1999). However, there is a major difference between the osmotic induction of proP P1 and that of proU, because the induction of proP P1 is only transient after an osmotic shock and stops when cells have adapted to the new osmotic conditions (Landis et al., 1999). The bet system that allows E. coli to synthesize glycine betaine from choline (Landfald and Strom, 1986; Lamark et al., 1991) is also osmotically inducible. The bet promoters are subject to a complex regulation, involving response to oxygen, through the ArcA regulator, induction by choline, through the action of the specific repressor BetI and osmotic induction through an ArcA and BetI-independent non-characterized mechanism (Lamark et al., 1996; Rokenes et al., 1996).

Osmotically inducible systems in Gram-positive bacteria have also been described. In Bacillus subtilis, uptake systems for the osmoprotectant compounds glycine-betaine, ectoin or proline exhibit increased transcription at high osmolarity (Kempf and Bremer, 1995; Spiegelhalter and Bremer, 1998; Kappes et al., 1999). However, much less is known about the regulatory mechanisms than in Enterobacteriae. Recently, an uptake system for glycine-betaine of Lactococcus lactis was described and named BusA (Obis et al., 1999) or OpuA (Bouvier et al., 2000). This system, which appeared as the unique glycine betaine uptake system in L. lactis, is organized as an operon encoding an ABC transporter. The BusA betaine uptake activity is triggered when cells face an osmotic imbalance (Obis et al., 1999). The modifications of the ionic strength on the cytoplasmic side of the membrane and in the physical properties of the membrane have been shown to control the transport activity of BusA (van der Heide and Poolman, 2000; van der Heide et al., 2001).

In addition, the expression of busAis osmotically inducible in L. lactis (Obis et al., 1999; Bouvier et al., 2000). However, the osmotic regulation is lost upon transfer of the promoter region to the heterologous host E. coli, suggesting the existence of a specific machinery able to sense and transduce the osmotic signal in L. lactis (Bouvier et al., 2000). Our goal in the present work was to investigate the mechanisms of osmotic induction of the busA (opuA) operon of L. lactis. We report the identification of busR, a regulatory gene encoding a DNA-binding protein involved in this process.

RESULTS

Osmodependent transcription of busA and repression upon betaine accumulation

It has been previously shown that the betaine transport capacity of L. lactis is regulated at the transcriptional level (Obis et al., 1999; Bouvier et al., 2000). The osmodependent promoter of busA presents an optimal 17 bp spacing between two highly conserved 10 and 35 boxes (Bouvier et al., 2000) (shown in Fig. 2B). To study the expression of the busA promoter, we constructed a transcriptional busAA-luxAB fusion at the busA locus on the L. lactis chromosome. The resulting strain, TIL452 (Table 1), harbours a single copy of the transcriptional fusion and a functional copy of the busA locus. The growth properties and betaine transport activity of strain TIL452 were similar to that of the wild-type strain (data not shown).

We measured the luciferase activity in TIL452 cells challenged by the addition of 0.5 M NaCl in the presence or absence of betaine (Fig. 1A). At high osmolarity and in absence of betaine, the luciferase activity increased linearly during the time of the experiment. In the presence of betaine, the level of activity paralleled the levels observed in the absence of the osmoprotectant during the first 60 min but then it ceased abruptly to increase and reached a plateau. This effect was likely due to the cytoplasmic betaine accumulation and the restoration of osmotic balance. At low osmolarity (0.3 mosM), at which no betaine was accumulated, the presence of osmoprotectant in the medium did not act on the low, basal activity of luciferase. The same experiment was performed with the strain TIL451 in which the busAA gene was disrupted by the luxAB genes insertion leading to a fourfold reduction of the betaine transport activity (data not shown). In TIL451, the downregulation effect observed in the presence of betaine was suppressed (Fig. 1B). This indicated that the betaine accumulation and/or turgor restoration was responsible for the arrest of busA induction.

To verify that the transcriptional effects described above were reflected at the protein level, we performed immunoblot experiments to detect the membrane component of the BusA transport system. As shown in Fig. 1A, the amount of BusAB increased after an osmotic upshift. The protein remained at a high concentration in the membrane fraction of cells cultivated in absence of betaine while it was progressively diluted when the osmoprotectant was available. Therefore, the amount of transporter synthesized was in good agreement with the transcriptional data.

Isolation of a gene encoding a repressor of the busA promoter

To look for regulators of busA transcription, we designed a genetic screen in the heterologous host E. coli. CLG802 is an E. coli strain that carries a busA-lacZ transcriptional fusion expressed under the control of a 170 bp DNA fragment from L. lactis carrying the busA promoter. This transcriptional fusion is present as a single copy at the malA locus on the chromosome (Bouvier et al., 2000). Expression of this fusion confers to CLG802 a Lac⁺ phenotype (white colonies) on lactose tetrazolium agar. We used this property to isolate a locus of L. lactis encoding a repressor of the busA promoter. Chromosomal DNA of L. lactis subsp. cremoris strain MG1363 (Gasson, 1983) was subjected to partial digestion with Sau3AI and ligated with BamHI-digested vector pJPB209 (Pichoff et al., 1995). CLG802 was transformed with this genomic library and plated on lactose tetrazolium agar supplemented with spectinomycin (100 µg ml ¹). Among approximately 20 000 independent transformants, three exhibited a Lac phenotype (red colonies). The recombinant plasmids purified from these clones were shown by restriction mapping to carry different overlapping parts of a unique region of the L. lactis chromosome. The plasmid carrying the smallest insert (1.9 kb) was named pBUS6 and further characterized.

The repressor is encoded by busR, a gene located upstream of busA

Sequencing of the chromosomal insert of pBUS6 revealed a putative open reading frame (ORF) overlapping a unique XmnI site. Insertion of a kanamycin-resistance cassette at this XmnI site on pBUS6 abolished repression of busA _p in CLG802, indicating that the corresponding gene encodes a repressor of busA _p . We named this gene busR. The main features of its sequence are shown in Fig. 2. The position of the 5'-end of the busR transcript was identified by primer elongation experiments in wild-type and busR genetic backgrounds (data not shown). These experiments identified a single 5'-end, preceded by a putative promoter (Fig. 2A). In addition, these experiments demonstrated that the amount of busR transcript was essentially independent of the osmolarity of the growth medium and of the BusR protein (data not shown). busR is located immediately upstream of busAA, the first gene of the busA operon, and oriented in the same direction (Fig. 2B). However, the busR ORF is followed by two inverted repeats (underlined with arrows in Fig. 2B) that can form a stem-loop structure (free energy of 6 kcal at 25°C) and precede a run of thymine residues. This sequence harbours all the distinctive features of Rho-independent transcription terminators (Brendel et al., 1986) and transcription of busA is probably not coupled to that of busR.

The structural gene starts with a GTG codon preceded by a potential Shine-Dalgarno sequence (AGGAG) and encodes a protein of 206 amino acids with a calculated molecular mass of 23133 Da. A search for similarities with the BusR protein was performed using the blast network service ( http://www.ncbi.nlm.nih.gov/blast/ ), the PFAM collection of multiple alignments ( http://www.sanger.ac.uk/Software/Pfam/ ) and the Prodom protein domains library ( http://protein.toulouse.inra.fr/prodom.html ). One protein is highly similar to BusR through its entire length and is most probably the BusR homologue of Streptococcus pyogenes. In addition, two domains of the protein exhibited similarity to other proteins (Fig. 2C). The amino-terminal third of BusR is similar (E = 1e ⁵) to the DNA-binding domain of transcriptional regulators ofthe GntR family (Haydon and Guest, 1991; Reizer et al., 1991) (PFAM accession no. PF00392), the closest homologue being the fatty-acyl responsive repressor FarR of E. coli (Quail et al., 1994). The carboxy-terminal third of BusR shares significant similarity (E = 2e ⁰⁸) with the TrkA-C domain of the cytoplasmic subunit of Trk K⁺/H⁺ symporter of E. coli (PFAM accession no. PF02080).

Binding of BusR to the busA promoter region

Radioactively labelled DNA fragments encompassing various segments of the busA promoter region extending from 16 nucleotides upstream of the 35 motif to the TTG translation start codon were amplified by polymerase chain reaction (PCR) (Fig. 3A). These DNA probes were incubated with crude extracts prepared from E. coli strains carrying either pBUS6 or the vector pJPB209, and run on polyacrylamide gels. The most abundant retarded band observed in these band-shift experiments was obtained only with extracts containing BusR (Fig. 3B). This retarded band was observed with DNA fragments containing the entire busA promoter (probes 1 and 4), or lacking the 35 hexamer and its upstream region (probe 2). It was absent with probe 3, which lacked in addition most of the spacer of busA _p . These data indicated that BusR is able to bind the busA promoter region, at a site overlapping the spacer sequence of the promoter. In similar band-shift experiments, we added glycine-betaine (up to 600 mM) or proline (up to 120 mM) in the binding buffer: these conditions were not able to relieve BusR binding (not shown).

To locate more precisely the BusR binding site, we performed footprinting experiments. Band-shift polyacrylamide gels were treated with 1,10-phenantroline-copper, and the DNAs extracted from the gel were analysed on sequencing gels (Fig. 4A and B). Comparison of the cleavage pattern of free and retarded DNA fragments demonstrated that the presence of BusR in the crude extract resulted in protection against cleavage extending from the 35 region to 10 bp after the transcription start (Fig. 4C).

Inhibition of BusA synthesis by BusR overexpression

Genetic data obtained in E. coli suggested that BusR acts as a repressor of busA transcription. To confirm these observations in L. lactis, the busR coding sequence was cloned under the control of its own promoter in pJIM2279, a L. lactis high-copy number plasmid (Table 1) and introduced into the L. lactis wild-type strain NCDO763. The growth rate under osmotic constraint and the betaine transport activity were measured in the resulting strain, TIL455. At low osmolarity the growth of TIL455 (µ = 0.54 h ¹) was indistinguishable from that of TIL456, the wild-type strain transformed with pJIM2279 (µ = 0.52 h ¹) (Fig. 5). In the presence of 0.5 M NaCl, the growth was strongly inhibited for both strains. When betaine was added to the high-osmolality medium, the growth of the wild-type strain was partially restored (µ = 0.21 h ¹). In contrast, TIL455 barely took an advantage of the presence of the osmoprotectant (µ = 0.08 h ¹). This phenotype was directly linked to the betaine transport activity. A 1 h osmotic up-shift stimulated the betaine transport activity sixfold in wild-type cells (5.7-30.4 nmol of betaine min ¹ mg of protein ¹). In TIL455, the basal level of betaine transport was much lower (1.2 nmol of betaine min ¹ mg of protein ¹) and weakly increased upon the osmotic up-shift (2.1 nmol of betaine min ¹ mg of protein ¹), suggesting that, even when overexpressed, BusR is still sensitive to elevated osmolarity. An immunodetection experiment of BusAB protein in membrane fraction of wild-type and BusR overexpressing strains was performed before and after a 1 h osmotic up-shock. The results demonstrated that the defect in betaine accumulation capacity was due to a lower amount of BusAB protein in TIL455 cells (Fig. 5B). Altogether, these data confirmed that the busR gene product was acting as a negative regulator on the expression of the busA operon.

Inactivation of busR causes an increase in basal busA expression

To characterize further the mechanism of osmodependent activation of busA _p , we constructed strain TIL470, in which the busR gene was inactivated (see Experimental procedures). The growth of TIL470 followed under various situations (CDM, CDM-0.3 M NaCl and CDM-0.3 M NaCl-betaine) was identical to that measured for the parental strain (data not shown). The betaine transport activity was measured in strains TIL470 and NCDO763. After growth in CDM or CDM + 0.2 M NaCl, strain NCDO763 (busR⁺) exhibited betaine transport activities of 10.3 and 17.6 nmol of betaine min ¹ mg of protein ¹ respectively. Strain TIL470 (busR ) exhibited values of 27.6 and 16.3 nmol of betaine min ¹ mg of protein ¹ in the same conditions. Therefore, the absence of BusR resulted in a 2.7-fold increased uptake at low osmolarity. In contrast with the wild-type situation, TIL470 did not exhibit an osmotic induction of the BusA transport activity, rather, we measured a lower activity in the presence of NaCl. The reason of this behaviour is not explained at present.

We then analysed the busA mRNA produced in TIL470 or NCDO763. Primer extension experiments demonstrated that inactivation of busR resulted in an increased amount of busA mRNA in rich medium of low osmolarity (M17), and again, within the limit of this semiquantitative experiment, an abolition of the osmotic induction (Fig. 6). In addition, this experiment showed that the busA mRNA was starting from the same busA _p promoter in both busR ⁺ and busR backgrounds.

DISCUSSION

We have identified a gene, busR, which encodes a regulator of the expression of the osmotically inducible glycine-betaine uptake system BusA of L. lactis. Expression of cloned busR results in repression of busA transcription in the heterologous host E. coli. Placing busR on a multicopy plasmid in L. lactis strongly inhibited the synthesis of the betaine transporter and increased the osmosensitivity of the cells in the presence of betaine (Fig. 5). Furthermore, deletion of busR in L. lactis results in an increased expression of busA at low osmolarity, and abolition of the osmotic inducibility during equilibrated growth, as shown by betaine uptake measurements and primer extension analysis of the busA mRNA in busR ⁺ and busR strains (Fig. 6). Altogether, these data show that BusR is a major actor of busA osmotic regulation through its repressor role on the busA promoter. Transcription of the proU operon in E. coli is also controlled by a repressor, the nucleoid-associated protein H-NS (discussed in Lucht and Bremer, 1994). However, H-NS is not the sole element of the osmotic responsiveness of proU _p but rather appears as one component of a complex mechanism involving additional elements such as the level of DNA supercoiling, and most probably an intrinsic sensitivity of the RNA polymerase-promoter interaction to the cytoplasmic concentration of K⁺ glutamate. Although the osmotic control of busA expression seems lost in the absence of BusR during equilibrated growth at different osmolarities, it will be interesting to know whether DNA supercoiling or cytoplasmic ionic strength are able to affect the transcription from busA _p .

BusR is a member of the GntR family of bacterial transcriptional regulators (Haydon and Guest, 1991; Reizer et al., 1991) that control the expression of genes involved in various metabolic pathways: gluconate metabolism and transport in E. coli and B. subtilis (Tong et al., 1996), malonate transport and its conversion to AcylCoA in Rhizobium sp. (Lee et al., 2000), glycolate oxidation (Pellicer et al., 1996), trehalose and arabinose metabolism in B. subtilis (Schock and Dahl, 1996; Sa-Nogueira and Mota, 1997), fatty acids synthesis (DiRusso et al., 1993). These repressors bind specific sites on DNA through a helix-turn-helix motif located in the amino-terminal part of the protein. Our band shift and footprinting experiments indicate that BusR is able to bind to a site overlapping the spacer of the busA promoter (Figs 3 and 4). The closest homologue of BusR is the fatty acyl responsive regulator FarR of E. coli. The two proteins exhibit 50% identity in the region of the helix-turn-helix motif of FarR (Haydon and Guest, 1991) (see Fig. 7A), suggesting that they may bind their DNA targets in a similar fashion. As shown in Fig. 7B, alignment of the sequences of busA _p of L. lactis and farR _p of E. coli reveals similarities within the region where FarR is known to bind to its own promoter (Quail et al., 1994). A 5 bp direct repeat (TATTT) is present within the region of farR _p protected by FarR. A very closely related direct repeat (TATTG) is present at the same location at the centre of the region protected by BusR on busA _p . This sequence is likely to contribute to BusR binding site. BusR bound at this site, overlapping the 10 region of the promoter, can then repress transcription by preventing either the binding of the RNA polymerase or the occurrence of a subsequent step in open complex formation.

Under osmotic constraint and in absence of betaine, the cells were found to accumulate the transporter (Fig. 1), whereas the presence and the cytoplasmic accumulation of betaine lead to a repression of busA transcription. At the saturating concentration of betaine used in this experiment, the initial burst of busA transcription allows the cells to synthesize a level of transporter sufficient to keep the cytoplasmic betaine at a concentration compatible with growth. This experiment shows that it is not the external osmolarity per se which is sensed by the busA promoter, but rather the osmotic imbalance.

An interesting question raised by the discovery of BusR is that of how repression is modulated by osmolarity. Elevated osmolarity could displace BusR from its site on busA _p , in a manner analogous to the displacement of CRP-cAMP from the proP P1 promoter (Landis et al., 1999). However, the osmotic induction of busA _p is not transient and lasts much longer than that of proP P1 after an osmotic shock (compare data in Fig. 1 with those in Landis et al., 1999), suggesting that release of BusR binding should be permanent at elevated osmolarity. It must be kept in mind that displacement from the binding site is not a necessary consequence of inducer interaction with regulator proteins. For instance, although choline is an inducer of the bet operon, Rokenes et al. (1996) have shown by band-shift experiments that not only does choline not displace the repressor BetI from its target, but that it actually enhances binding. We note that the direct repeat that is likely to constitute the BusR binding site core is largely on the side of the DNA opposite to that of RNA polymerase binding. Therefore, it is conceivable that BusR could remain on the DNA even in induction conditions.

The similarity observed between the carboxy-terminal region of BusR and a component of a K⁺/H⁺ symporter suggests the very attractive hypothesis that BusR could be a sensor of intracellular ionic strength. An increase in cytoplasmic ionic concentration at high osmolarity could then be the inducing signal of the regulatory network. It is well established in enterobacteria that potassium influx is a primary and transitory response to osmotic upshock that triggers the induction of osmodependent operon (Bremer and Krämer, 2000). A similar mechanism is not established in Gram-positive bacteria, which maintain a high turgor, and the studies available in B. subtilis (Whatmore and Reed, 1990) or Lactobacillus plantarum (Glaasker et al., 1996) lead to opposite conclusions.

It should however, be stressed that the homology found in the C-terminal domain of BusR is with a cytoplasmic resident protein (TrkA) that interacts with the transmembrane K⁺/H⁺ symporter TrkH subunit (Durell et al., 1999). Another mode of regulation of BusR activity could be through sequestration at the membrane level, which is the seat of important changes during osmotic stress. Biochemical experiments currently underway in our laboratory will test these possibilities.

EXPERIMENTAL PROCEDURES

Bacterial strains and cultures

Lactococcus lactis subsp. cremoris NCDO763 and its plasmid-free derivative MG1363 (Gasson, 1983) were used in this work. The L. lactis were grown in M17 broth (Difco; Terzaghi and Sandine, 1975) or in chemically defined medium (CDM; Molenaar et al., 1993) modified as described (Obis et al., 1999). The media were supplemented with 0.5% glucose.

The Escherichia coli strains used are described in Table 1. EC101 (Kan^r) is a derivative of strain TG1 containing a chromosomal copy of pWV01 repA gene (Law et al., 1995). Growth of E. coli was performed at 37°C in Luria-Bertani (LB) medium (Difco) or on lactose tetrazolium agar plates (Miller, 1992).

Erythromycin (Em) and Kanamycin (Kan) were used at 1-5 µg ml ¹ and at 50-100 µg ml ¹ for L. lactis and E. coli respectively.

To measure the growth rates of the L. lactis strains, a Bioscreen Microbiology Reader (Labsystems) was used (Fig. 5). Cultures were performed in microtitre plates with 300 µl of M17 or CDM containing 0.5% (v/v) glucose inoculated with 5 µl of exponentially growing cultures (OD₆₀₀ = 0.5-1). The optical density was measured at 600 nm every 30 min. Cultures were performed in triplicates.

Construction of strains

To create chromosomal transcriptional fusions between busA promoter region and luxAB in L. lactis, the integrative vector pJIM2374 (Em^R) was used (Delorme et al., 1999). pJIM2374 lacks the repA gene required for its replication in L. lactis and carries the luxA and luxB genes of Vibrio harveyi. A 1.2 kb polymerase chain reaction (PCR) fragment was generated from the L. lactis NCDO763 chromosomal DNA with the bus44f-SacI (5'-CGGGCGAGCTCGCGGAAGTGGGCGAT GTGGATAGAT-3') and EPTr-KpnI (5'-GGGCGGTACCGT TCAATCAATCGATTAAGC-3') primers. The amplified region contains busR coding sequence, busA promoter region and 238 bp from the busAA coding sequence corresponding to the first 79 codons. This fragment was cloned in pGEM-T Easy vector (Promega), yielding pGEM-T452. The 1.2 kb SacI-SalI fragment containing the lactococcal sequence was subcloned in pJIM2374, using E. coli EC101 as a recipient and selecting for resistance to 100 µg ml ¹ of Em and Kan (Law et al., 1995), yielding pIL452. L. lactis NCDO763-competent cells containing the helper plasmid pVE6007 Cm^R plasmid (Maguin et al., 1996) were transformed by electroporation with pIL452. In this strain, the helper plasmid supplied in trans the RepA protein, allowing the replication of pIL452 at 28°C. Transformants were selected at 28°C on M17-glu containing 1 µg of Em per ml. A few colonies were chosen and cultivated at 28°C overnight in M17-Glu containing 5 µg Em per ml. The culture was then diluted 1000-fold, cultivated for 2 h at 30°C and plated at 37°C on preheated M17-Glu plates containing 1 µg of Em per ml. The replication of the helper plasmid is conditional and 37°C is the non-permissive temperature. This procedure allows selecting colonies in which the helper plasmid has been lost and pIL452 has been inserted into the chromosome. The Em^R, Cm^S phenotype of the clones was verified, and chromosomal integration of pIL452 was confirmed by PCR and Southern blot experiments.

In the resulting strain, TIL452, the lux genes have been introduced behind the duplicated busA promoter region. The strain still possesses a fully functional busA operon, and the betaine transport activity was found to be indistinguishable from that of the wild-type strain. L. lactis strain TIL354 possessing luxAB genes inserted at the non-osmodependent locus bcaT was used as a control (Yvon et al., 2000).

A similar strategy was used to construct strain TIL451 except that a internal busAA fragment generated by PCR with the two following primers: EPT-Kpn (5'-CCGGGTACCCT TAATCGATTGATTGAACC-3') and FLM-Kpn (5'-GGGCGG TACCCTATTTCTTCTACTTCATCAG-3') was used to insert the lux genes at the busA locus. In the resulting strain, the disruption of the busAA gene leads to a fourfold reduction in the betaine transport activity.

Deletion of busR in L. lactis strains NCDO763 was performed as follows. A 1.8 kb DNA fragment carrying the entire busR gene and a large part of the busA operon, was purified from plasmid pBUS4 cleaved with HindIII and cloned in the replication thermosensitive vector pG⁺host9 (Maguin et al., 1996), using E. coli EC101 strain as a recipient and selecting for resistance to 100 µg ml ¹ of Em, yielding pBUS98. A deletion of 556 bp internal to busR was generated on pBUS98 by cleavage at AhdI and NspV sites (shown in Fig. 2), resulting in pBUS99. Competent L. lactis cells were electrotransformed with 10 µg of pBUS99, and plated on M17 agar plates with 5 µg ml ¹ of Em at 30°C. Deletion of the chromosomal copy of busR was carried out by insertion-excision of the thermosensitive vector as described in Maguin et al. (1996). The resulting strain, derived from NCDO763, was named TIL470. The presence of the correct deletion was verified by PCR amplification using chromosomal DNA of control and recombinant strains and appropriate oligonucleotides derived from sequences of busR, busA or pG⁺host9.

Plasmid pIL455 was constructed by inserting a 840 bp PCR fragment containing the busR gene in the high-copy-number plasmid pJIM2279, a derivative of enterococcal pAM1 (Renault et al., 1996). pJIM2279 was digested with EcoRV and SacI. The busR PCR fragment was generated by Pfu polymerase using bus44f-SacI and P1rev (5'-TTCT CATTTAAAGTGACCAC-3') primers with L. lactis chromosomal DNA as a template. The purified PCR fragment was digested with SacI and cloned in linearized pJIM2279. Plasmid pIL455 was used to transform L. lactis wild-type strain NCDO763 and transformants were selected on M17 containing 5 µg ml ¹ of Er. The resulting strain was named TIL455.

Methods used with nucleic acids

Isolation of plasmid DNA, digestion with restriction enzymes, ligation with T4-DNA ligase and transformation of E. coli were carried out as described by Sambrook et al. (1989). Preparation of competent cells and electrotransformation of L. lactis were as described by Holo and Nes (1988). Extraction of chromosomal DNA from L. lactis cells was performed with the DNeasy Tissue Kit (Qiagen) according to the manufacturer's protocols for Gram-positive bacteria. Total RNAs were extracted from L. lactis by the hot-phenol method (Aiba et al., 1981). Determination of the 5'-end of the busR and busA mRNA were performed by primer extension experiments as described previously (Bouvier et al., 2000). The primers used in these experiments were bus7 (5'-CAGCTATTTGCAGGG-3') for busR mRNA and bus37 (5'-CTCTCCTTCATTCTAT TACTCATGAGC-3') for busA mRNA.

Preparation of E. coli crude extracts

Bacterial cells, grown in LB medium with spectinomycin (100 µg ml ¹), were harvested by centrifugation, washed in 'B' buffer (20 mM Hepes, pH 8.0; 1 mM EDTA; 7 mM -mercaptoethanol; 10% (v/v) glycerol) and lysed by sonication. Lysates were centrifuged for 1 h at 12 000 g at 4°C, and the supernatants were mixed with an equal volume of saturated ammonium sulphate and incubated for 30 min at 4°C. After centrifugation at 12 000 g at 4°C, pellets were resuspended in B buffer and adjusted to 1 µg µl ¹ of protein after assaying the total protein content by Bio-Rad protein assay kit (Bio-Rad).

Band-shift experiments

Plasmid pOPU2 (Bouvier et al., 2000) carries the entire busA operon cloned in the vector pJPB209 (Pichoff et al., 1995). DNA fragments carrying various parts of the busA promoter region (Fig. 3A) were synthesized by PCR amplification in the presence of 20 µCi of [-³²P]-dATP, using pOPU2 as a template and the following couples of oligonucleotides: probe 1, bus47 (5'-CAAAATAACTCTCCTTCATTC-3') and bus56 (5'-GTTCAAATTTAGTTAGTTGACATAG-3'); probe 2, bus47 and bus62 (5'-GTGACTACATATTGTTATTATTGAG-3'); probe 3, bus47 and bus57 (5'-GTTATTATTGAGTGGTCACTTTAAA TG-3'); and probe 4, bus56 and bus58 (5'-CATTTAAAGT GACCACTCAATAATAAC-3'). Then, 10 ng of the labelled DNA fragment was incubated for 15 min at room temperature with crude extract (1.5 µg of protein) in 20 µl of B buffer supplemented with 50 mM NaCl and 1 µg of poly-dI-dC)/poly-dI-dC) (Pharmacia) competitor DNA. The binding mix was loaded on 5% polyacrylamide gels under 6 V cm ¹ tension, and run under 12 V cm ¹ tension. The wet gel was autoradiographed at 70°C in the presence of intensifying screens.

In situ footprinting with 1,10-phenantroline-copper (OP-Cu)

In situ assays were performed as described by Sigman et al. (1991). DNA probes were synthesized by PCR amplification using plasmid pOPU2 as a template and the couple of oligonucleotides bus83 (5'-CGATGCTTTTTTTTAAGTTC-3') and bus84 (5'-GGTTACACAAGTGATTTTC-3'). Probes labelled on the upper or lower DNA strand were obtained with 5'-end-³²P-labelled bus83 and non-labelled bus84, or 5'-end-³²P-labelled bus84 and non-labelled bus83 respectively. Then, 10 ng of the labelled DNA fragments was incubated with crude extract (10 µg of protein), and DNA-protein complexes were separated from free DNA by electrophoresis as described above. The wet gel was immersed in 200 ml of 10 mM Tris-HCl pH 8, followed by the addition of 20 ml of solution A (0.45 mM CuSO₄, 2 mM 1,10-phenantroline). In situ digestion of the DNA-protein complexes was initiated by the addition of 20 ml of solution B (1:200 dilution of 3-mercaptopropionic acid in H₂O). The reactions were stopped after 6 min at room temperature by the addition of 20 ml of solution C (5.8 mg ml ¹ neocuproine in ethanol). Free DNA and DNA-protein complexes were localized by autoradiography of the wet gel and excised from the gel. The DNA was recovered by elution overnight at 37°C in 10 mM Tris-HCl pH 8, 1 mM EDTA, 300 mM NaCl, 0.2% SDS followed by filtration on acetate cellulose 0.2 µm microspin filter (PolyLabo), phenol-chloroform extraction and ethanol precipitation. The purified DNA samples were resuspended in 33% formamide, 0.025% xylene cyanol, heated at 95°C for 3 min and separated by electrophoresis in 6% denaturing polyacrylamide gel. Sequence standards for footprinting were prepared with the thermosequenase cycle sequencing kit (Amersham). The gel was dried on Whatman DE81 paper and exposed to a phosphor screen for quantification on a PhosphorImager. The cleavage pattern for the complex was scanned and compared with the cleavage pattern of free DNA fragments isolated from the same gel.

Measurement of luciferase activity

To measure luciferase activity, 1 ml of cell culture was withdrawn at the indicated times or at a given OD₆₀₀ and rapidly mixed with 5 µl non-yl aldehyde 95% (30 µM final concentration) (Sigma-Aldrich), the emission of light was immediately measured in a luminoskan II luminometer (Labsystems). In parallel, growth was monitored with a spectrophotometer at a wavelength of 600 nm and the emitted light value was standardized to the optical density (ULU/OD). The experiments were repeated at least twice.

Betaine transport activity

The betaine transport activity on energized cells resuspended in buffer was measured by the filter-binding assay as described (Obis et al., 1999). In brief, cells were grown in CDM at 30°C with or without 0.2 M NaCl. At OD₆₀₀ of 0.5, cells were harvested, washed and resuspended in reaction buffer containing 0.3 M NaCl at 30°C. The reaction was initiated by the addition of [¹⁴C]-betaine (20 µM final concentration, specific activity 0.25 mCi mmol ¹). The initial rate of [¹⁴C]-betaine uptake was calculated from kinetic experiments with a point every minute for 4 min, corresponding to the linear phase of transport.

Immunodetection of betaine transporter

Cells cultivated in CDM were harvested by centrifugation (5000 g, 5 min at 4°C). Cell pellets were resuspended in TE (final concentration, 10 mg of protein ml ¹) containing 20% sucrose (w/v) and 10 mg of lysozyme ml ¹ at a cell concentration of 5 OD ml ¹ (corresponding to a total protein concentration of 1 mg ml ¹). The cell suspension was incubated 30 min at 37°C and protoplasts were lysed by addition of SDS at 1% (w/v) final concentration. The sample was heated at 60°C for 5 min to facilitate the resuspension. Next, 50 units of benzonase (Merck) and MgCl₂ 2 mM were added and the incubation was continued for 30 min at 30°C. Then, 5 µg of proteins were loaded on SDS-PAGE performed in a Miniprotean II apparatus (Bio-Rad laboratories). The electrotransfer of proteins was performed in 25 mM Tris-HCl, 192 mM glycine, pH 8.3, methanol 20% (v/v), 0.02% SDS, at a constant current of 80 mA for 2 h at 4°C. The addition of SDS in the transfer buffer was found to be essential for an efficient transfer of membrane proteins on the nitrocellulose membrane (BA85, Schleicher and Schuell). Membrane was blocked for 1 h at room temperature in PBST (10 mM Na phosphate buffer, pH 7.4, 145 mM NaCl, 0.2% (v/v) Tween) containing 3% (w/v) low-fat milk (PLT). The membrane was then incubated for 1 h at room temperature with BusAB rabbit antiserum, diluted 1:1000 in PLT. The antiserum was obtained by immunization of rabbits with purified BusAB proteins (van der Heide and Poolman, 2000). Membranes were washed twice in PLT and incubated for 1 h at room temperature with horseradish peroxydase-labelled goat anti-rabbit antibodies (diluted 1:2500 in PLT). Membranes were washed twice in PBST and colorimetric detection was performed with Opti-4 CN detection kit following the instructions of the manufacturer (Bio-Rad).

Nucleotide sequence accession number

The nucleotide sequence of busR has been deposited in the GenBank nucleotide sequence database under accession no. AF393650.

FIGURES

Fig. 1. Osmotic induction of busA in Lactococcus lactis. Luciferase activity was assayed during growth...

Fig. 2. Partial sequence of the busR gene.

Fig. 3. Binding of BusR to the busA promoter region of L. lactis.

Fig. 4. Footprinting of the BusR binding site.

Fig. 5. Consequence of busR overexpression on osmoadaptability of L. lactis.

Fig. 6. Reverse transcriptase mapping of busA mRNA 5'-end.

Fig. 7. Comparison of BusR of L. lactis and FarR of E. coli.

Table 1 . Bacterial strains and plasmids.

ACKNOWLEDGEMENTS

Y.R. and D.O. were recipients of fellowships from the CNRS and the French Ministère de la Recherche respectively. We thank B. Poolman and T. van der Heide for the generous gift of purified BusAA and BusAB proteins, and M. Nardi, P. Renault, E. Guédon, P. Le Bourgeois and M. Mingot for the gift of plasmids, transfer of protocols and helpful discussions. Part of this work was supported by grants from the Région Midi-Pyrénées (no. 97002157) to C.G. and I.B., from the DGA (no. 0034058/DSP) to M-Y.M. and from the Institut Universitaire de France to C.G.

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