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Summary

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References

The function of the LysR-type regulator LrhA of Escherichia coli was defined by comparing whole-genome mRNA profiles from wild-type E. coli and an isogenic lrhA mutant on a DNA microarray. In the lrhA mutant, a large number (48) of genes involved in flagellation, motility and chemotaxis showed relative mRNA abundances increased by factors between 3 and 80. When a representative set of five flagellar, motility and chemotaxis genes was tested in lacZ reporter gene fusions, similar factors for derepression were found in the lrhA mutant. In gel retardation experiments, the LrhA protein bound specifically to flhD and lrhA promoter DNA (apparent KD 20 nM), whereas the promoters of fliC, fliA and trg were not bound by LrhA. The expression of flhDC (encoding FlhD2C2) was derepressed by a factor of 3.5 in the lrhA mutant. FlhD2C2 is known as the master regulator for the expression of flagellar and chemotaxis genes. By DNase I footprinting, LrhA binding sites at the flhDC and lrhA promoters were identified. The lrhA gene was under positive autoregulation by LrhA as shown by gel retardation and lrhA expression studies. It is suggested that LrhA is a key regulator controlling the transcription of flagellar, motility and chemotaxis genes by regulating the synthesis and concentration of FlhD2C2.


Introduction

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References

The LysR-type regulator LrhA (LysRhomologue A) of Escherichia coli is encoded by lrhA, which is located upstream of the nuoA–N (NADH:quinone oxidoreductase) locus (Bongaerts et al., 1995). The LrhA protein has substantial homology to HexA from Erwinia carotovora (64% identity) and PecT from Erwinia chrysanthemi (61% identity). In these plant pathogenic bacteria, HexA and PecT function as repressors of motility and of virulence factors, such as exoenzymes required for lytic reactions (Surgey et al., 1996; Harris et al., 1998). Overexpression of the E. carotovora hexA gene also represses multiple virulence determinants in the opportunistic human pathogen Serratia (Harris et al., 1998). In hexA mutants of E. carotovora, expression of flagellar genes (fliA and fliC) is increased resulting in hypermotility (Harris et al., 1998). In the same organism, HexA seems to influence components of at least three global regulatory systems: (i) the expression of the regulatory RNA rsmB (a homologue of the E. coli csrB); (ii) the synthesis of the quorum-sensing pheromone OHL; and (iii) the stability of the stationary phase sigma factor RpoS (Mukherjee et al., 2000). A model suggests that HexA negatively regulates trans-cription of rsmB, a positive regulator of OHL production, which, in turn, is required for exoenzyme synthesis. Increased levels of RpoS protein in a hexA mutant appear to be the result of a different regulatory pathway, which might include SprE (Mukherjee et al., 2000). In E. coli, the stability or concentration of RpoS is controlled by LrhA via the SprE (or RssB)/ClpXP proteolytic pathway (Gibson and Silhavy, 1999).

LrhA also shows similarity (27% identity) to the transcriptional regulator CbbR of Ralstonia eutropha and Xanthobacter flavus. CbbR controls CO2 fixation in both bacteria and responds to phosphenolpyruvate in R. eutropha (Grzeszik et al., 2000) and to NADPH in X. flavus (van Keulen et al., 1998) as regulatory signals.

The lack of a clear phenotype for the lrhA mutation in E. coli (Bongaerts et al., 1995) and the similarity to the HexA and PecT proteins suggested that the major function of LrhA in E. coli has not been recognized so far. Therefore, in a genome-wide expression analysis (Wendisch et al., 2001), the target genes for regulation by LrhA were identified using gene expression profiling. The changes in relative mRNA abundances in E. coli were determined by pairwise comparison of the wild-type strain MG1655 and an isogenic lrhA mutant. These studies showed that one of the major functions of LrhA is transcriptional regulation of the flhDC genes and of the flhDC regulon. The flhDC regulon codes for a large number of structural, biosynthetic and regulatory genes of flagella, motility and chemotaxis. It consists of at least 13 operons that are regulated in a hierarchical mode. At the highest level is the flhDC operon (class I genes) encoding the (positive) master regulator FlhD2C2, a heterotetramer that controls the expression of the remaining genes of the regulon. The expression of the downstream genes depends either directly (class II genes) or indirectly via the FlhD2C2-regulated sigma factor FliA (class III genes) on the function of FlhD2C2.

Results

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References

Identification of target genes of LrhA by DNA microarray analysis

In order to identify the LrhA-regulated genes, the expression pattern of total mRNA from an lrhA mutant strain was compared with that of wild-type E. coli. The mRNA of wild-type MG1655 and the respective lrhA mutant was extracted after growth in mineral medium with glycerol. The mRNA from both strains was reverse transcribed and labelled with either green (Cy3) or red (Cy5) fluorescent-labelled nucleotides. The labelled cDNA probes were simultaneously hybridized to a DNA glass microarray containing 4108 genes corresponding to about 96% of the E. coli genome. After hybridization, fluorescence at 532 nm and 635 nm was determined quantitatively, and relative mRNA abundances were derived.

A large number of genes showed increased relative mRNA abundances in the mutant, and genes or gene clusters with ratios ≥ 2 and a signal amplitude of at least threefold over background are shown in Table 1. Most of the genes are located in three clusters on the E. coli chromosome and are related to either flagellation or chemotaxis. The flagella-specific genes include structural genes (flg and fli genes) and genes required for their tran-scriptional and functional control. The latter include the flagellum-specific σ factor FliA, an anti-σ factor, motor switch components and proteins for the assembly and export of the flagellar proteins. The chemotaxis genes encode typical chemotaxis proteins (che and mot genes), regulators of flagellar operons (flhDC and others) and the chemoreceptors for dipeptides, asparate, galactose and aerotaxis (tap, tar, trg and air genes). In addition to the genes shown in Table 1, the mRNA levels of the cadA gene (2.5-fold) encoding the pH regulator CadA and genes related to fimbriae synthesis (fimE gene and the fimAICDFG operon) were increased. Most of the LrhA-regulated genes are part of larger operons. The genes within the operons all showed increased expression in the mutant, although the factors for the individual genes could vary.

Table 1. Genes with increased mRNA abundances in the lrhA mutant.Thumbnail image of

LysR-type regulators often control the expression of adjacent genes (Schell, 1993). The nuoA–N, ackA and pta genes, however, which are adjacent to lrhA on the chromosome, showed no significantly changed mRNA levels. Similarly, the concentration of rpoS mRNA, which had previously been related to LrhA (Gibson and Silhavy, 1999), was not significantly increased.

Expression of LrhA-regulated genes in translational lacZ fusions

To study the role of LrhA on the expression of flagellar and chemotactic genes, translational lacZ fusions were constructed with genes from the three classes of LrhA-regulated motility genes, flhDC (class I), fliA and flgB (class II), fliC and trg (class III). The flhDC and fliA genes code for the transcriptional regulators FlhD2C2 and FliA. FlhD2C2 is the master regulator of flagellar operons, whereas FliA is a σ factor for class III flagellar gene expression. The flgB, fliC and trg genes code for a subunit of the flagellar basal body, flagellin, and the galactose chemoreceptor respectively.

The fusions were tested for β-galactosidase activity after aerobic growth (Table 2). The expression of the class I gene flhDC was increased by a factor of 3.5 in the mutant, compared with an increase in the flhD mRNA by a factor of 3.3 in the DNA microarray experiment (Table 1). In the class II genes fliA and flgB, the expression was raised by factors of 13 and 8, respectively, in the lrhA mutant, with corresponding increases in the mRNA levels. In the class III genes fliC and trg, the derepression was even higher with regard to expression and mRNA levels. The expression studies with the translational lacZ fusions therefore confirm the observed changes in the mRNA levels and suggest that the regulation occurs mainly at the transcriptional level.

Table 2. Expression of translational fusions of flagellar or chemotaxis genes in wild-type (lrhA+) and lrhA backgrounds.
  β-Galactosidase activity (Miller units) 
FusionStrains Wild type/lrhAWtlrhARepression by LrhA (-fold)
  • The strains were grown under aerobic conditions in supplemented M9 medium with glycerol. The β-galactosidase activities were obtained from three or more independent growth experiments.

  • a. 

    Wild type with respect to lrhA.

  • b. 

    LrhA-overproducing strain.

flhDClacZCP992/IMW355 88 ± 11 303 ± 9 3.5
fliAlacZLJ10a/IMW331a258 ± 63483 ± 14213
flgBlacZIMW353/IMW354 38 ± 3 311 ± 20 8
trglacZIMW334/IMW335 16 ± 1 270 ± 1217
fliClacZIMW356/IMW336108 ± 255968 ± 25255
lrhAlacZIMW326aa/IMW327 16 ± 5  9 ± 3 0.56
lrhAlacZIMW326bb/IMW327 67 ± 10  9 ± 3 0.13

The expression of the fliA, trg and fliC genes in response to lrhA inactivation was also analysed in a flhD background (strain MC4100) that is deficient in the regulator FlhD2C2 (Table 3). In the flhD mutant, expression of fliAlacZ and trglacZ was at background levels (Table 3) and negligible compared with the flhD+ strain (Table 2). Inactivation of lrhA caused no significant derepression. Expression of the class III gene fliClacZ from the low-copy plasmid in the flhD negative background (Table 3) was comparable with the expression of (monocopy) fliClacZ in the flhD+ strain (Table 2). Inactivation of lrhA in the flhD+ background caused a strong derepression of fliC (61-fold), whereas in the flhD background, only a weak derepression was found (factor 1.6). This suggests that the derepression by lrhA inactivation only works in a flhD+ background, and that LrhA affects motility mainly via FlhD2C2. For practical reasons, therefore, LrhA function cannot be studied in the flhD strain MC4100, which is frequently used for expression studies in E. coli.

Table 3. Expression of translational fusions of flagellar or chemotaxis genes in flhD (MC4100) and flhD lrhA (IMW41) negative backgrounds.
  β-Galactosidase activity (MU)
FusionsPlasmidsMC4100 (flhD)IMW41(flhDlrhA)
  1. The strains were grown under aerobic conditions in mineral medium with glycerol. The β-galactosidase activities were obtained from two or more independent growth experiments.

fliAlacZpMW198 1 ± 1 4 ± 1
trglacZpMW196 3 ± 2 3 ± 2
fliClacZpMW197112 ± 29182 ± 43

Effect of the lrhA mutation on swimming and chemotaxis

The mRNA expression profile of the lrhA mutant and the expression studies suggested an effect of LrhA on swimming and chemotactic behaviour. Soft tryptone swarm agar plates were used to evaluate motility or swimming of the strains. Aerotaxis was tested in soft minimal swarm agar (MSA) plates complemented with succinate. In the tryptone swarm agar and the aerotaxis plates (Fig. 1), the lrhA mutant (IMW325) was hypermotile and formed swarm rings with significantly increased diameter compared with the wild type (MG1655). Similarly, the lrhA mutant showed swarm rings with increased diameter on aspartate chemotaxis plates (not shown). When the lrhA mutant was complemented with a cloned lrhA gene on a low-copy plasmid (pMW213), the motility was decreased in the swarm, chemotaxis and aerotaxis tests to levels below those of the wild type (Fig. 1), presumably because of the increased lrhA gene dose. The empty vector (pME6010) had no influence on the motility of the lrhA mutant (not shown). Thus, the lrhA mutant showed improved chemotactic and aerotactic abilities, and the effect was reversed in the presence of the lrhA expression plasmid.

image

Figure 1. Motility and aerotaxis in E. coli lrhA+ and lrhA strains. A sample of 15 μl of suspended E. coli wild-type MG1655, the lrhA mutant IMW325 and the complemented lrhA mutant IMW325(pMW213) were spotted onto tryptone swarm agar (A) or aerotaxis MSA agar plates with succinate (B) and incubated for 4– 6 h at 37°C (A), or overnight at 30°C (B). The wild-type MG1655 and the lrhA mutant IMW325 contained the control plasmid pME6010.

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Binding of isolated LrhA to target promoters: gel retardation experiments

Gel retardation experiments were performed with promoter fragments of the class I gene flhD, the class II genes fliA and flgB and the class III genes fliC and trg that had also been studied by the lacZ reporter gene fusions. When labelled DNA fragments with the promoter region of flhDC (Fig. 2) were incubated with LrhA protein and subjected to native gel electrophoresis, the band of free promoter DNA disappeared, and a retarded DNA band with decreased mobility turned up, which presumably represents the LrhA–DNA complex. The decrease in free DNA started in the presence of 5 nM LrhA and, at 40–50 nM LrhA, only the retarded band was detectable. An apparent KD of about 20 nM for the binding of LrhA can be estimated. The retardation was not inhibited by a large excess of unspecific DNA, suggesting a specific binding and direct transcriptional regulation of the flhDC genes by LrhA.

image

Figure 2. Gel retardation assay for binding of LrhA to flhDC (A) and trg (B) promoter DNA. The radioactively labelled promoter DNA (5 nM in each lane) was incubated in the presence of a 625-fold excess of competitor DNA with increasing concentrations of isolated His6–LrhA protein and subjected to non-denaturing DNA PAGE (5% acrylamide). The positions of free promoter DNA (766 bp and 478 bp fragment of flhD and trg respectively) and of the retarded DNA–LrhA complexes are indicated.

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The DNA fragments containing the complete promoter regions of class II and III genes [trg (Fig. 2) and fliC, flgB, fliA (not shown)] were tested in the same way as for gel retardation by LrhA. None of the class II and III promoter fragments was retarded, even at high concentrations of LrhA. Thus, the LrhA-responsive promoters can be assigned to two groups: one that binds LrhA (flhDC genes), and a second group (fliA, flgB, fliC, trg genes and presumably other class II and class III genes) that does not bind LrhA.

Footprint analysis of the LrhA binding site at the flhD promoter

An flhD promoter DNA fragment was analysed for LrhA binding by DNase I footprinting. The promoter DNA was incubated with increasing concentrations of isolated (His6)-LrhA and, after partial digestion with DNase I, the resulting fragments were analysed by denaturing gel electrophoresis (Fig. 3). In the presence of LrhA, the region from –89 to –129 bp upstream of the transcriptional start site (Soutourina et al., 1999) was protected from DNase I and produced no or decreased amounts of fragments compared with incubation without LrhA. The protection of the site started at concentrations of 100 nM LrhA. In addition, at positions +28 and +46 of the flhD promoter, single nucleotides were protected from DNase I. At positions +52, on the other hand, single additional nucleotides became (hyper)sensitive towards DNase I in the presence of LrhA. Therefore, the flhDC promoter contains sites that become protected in the presence of LrhA, whereas sites downstream of the promoter become hypersensitive (Fig. 4).

image

Figure 3. DNase I footprint of LrhA at the flhD promoter (A) and in the 5′ end of the transcribed region of flhD (B). pMW212 containing the complete flhD promoter region was digested with EcoRI and KpnI, labelled at the 5′ end with 33P, incubated with increasing amounts of His6–LrhA and DNase I and subjected to denaturing electrophoresis.

B. Electrophoresis was performed for a shorter period of time compared with (A). Arrows indicate hypersensitive or changed bands.

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image

Figure 4. Promoter region of flhDC showing regulatory sites and the DNase I protected or hypersensitive sites in the presence of LrhA. The DNA fragment and the nucleotides ([DOWNWARDS ARROW]) that are protected (–) or become hypersensitive (+) in the presence of LrhA are indicated. The positions of the binding sites for the catabolite gene activator protein (CAP), the osmoregulator OmpR and the H-NS protein (black lines below sequence) are taken from Shin and Park (1995) and Soutourina et al. (1999). The putative LysR- type consensus is printed in bold.

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Autoregulation of lrhA expression

Many genes encoding LysR-type regulators are subject to autoregulation. The response of lrhA expression to the presence of LrhA was tested using a lrhAlacZ gene fusion in lrhA-positive and -negative backgrounds (Table 2). The expression of lrhAlacZ in the wild-type background (lrhA+) was low and decreased further by a factor of about 2 in the lrhA mutant. When a plasmid carrying the cloned lrhA gene was introduced into the wild type to increase the concentration of LrhA, the expression of lrhAlacZ increased and exceeded that of the lrhA mutant by a factor of 7.4, suggesting that lrhA expression is under positive autoregulation by LrhA.

Binding of LrhA to the promoter of lrhA was tested by gel retardation (Fig. 5). Inclusion of LrhA in the assay caused a strong retardation of the lrhA promoter DNA. The shifted band appeared already in the presence of 10 nM LrhA with an apparent KD for binding of 40 nM, suggesting high affinity of the protein for the promoter. Similar to the flhDC promoter, the shifted band was diffuse, which could result from the lack of an appropriate effector of LrhA. In a DNase I footprint experiment (Fig. 5B), a site of about 30 bp in the supposed promoter region of lrhA was protected by LrhA from DNase I digestion. Within and downstream of the protected site, additional sites became hypersensitive to DNase I in the presence of LrhA.

image

Figure 5. Gel retardation of lrhA promoter DNA by LrhA (A) and DNase I footprint of LrhA at the lrhA promoter (B). For gel retardation, the lrhA promoter fragment (676 bp, 5 nM in each lane) was incubated with increasing amounts of His6–LrhA protein; other conditions are described in the legend to Fig. 2. DNase I footprinting was performed as described in the legend to Fig. 3, but the 1018 bp lrhA promoter fragment was used. The black bar on the righthand side indicates the site protected by LrhA. The numbering refers to the translational start site of lrhA. Arrows indicate hypersensitive or changed bands.

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Discussion

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References

LrhA-regulated genes

Gene expression profiling of the E. coli genome shows that LrhA is a global transcriptional regulator of genes related to motility, chemotaxis and flagella synthesis. Among these, only the flhDC and lrhA promoters interact directly with LrhA, whereas the others are part of a hierarchical system under the control of the master regulator FlhD2C2. The FlhD2C2-regulated genes (Bartlett et al., 1988; Liu and Matsumura, 1994; Prüßet al., 2001) are essentially the same as the LrhA-regulated genes shown in Table 1. Both regulators control further genes that are not in common: FlhD2C2, the genes of anaerobic electron transport and sugar transport (Prüßet al., 2001); and LrhA, fimbrial genes. The same set of genes is also regulated by SdiA (Wei et al., 2001a), which is a transcriptional activator of cell cycle-related proteins (Wang et al., 1991).

Role of LrhA in the control of motility

FlhD2C2 is the master regulator of a three-step hierarchical control of flagella, motility and chemotaxis gene expression in E. coli (for reviews, see Macnab, 1996; Chilcott and Hughes, 2000; Kalir et al., 2001). The top level (class I) is taken by the flhDC genes encoding FlhD2C2. FlhD2C2 recognizes the second-level (or class II) flagellar promoters (Liu and Matsumura, 1994). The second-level genes encode proteins for the basal body and the hook of the flagellum, and for the sigma factor FliA, which is required for the transcription of class III genes. The class III genes are needed for assembly of the flagellar filament, motor activity and chemotaxis, and encode the anti-sigma factor FlgM. FlgM accumulates upon completion of the flagellum and inhibits FliA activity (Hughes et al., 1993). It is suggested that LrhA affects this hierarchy at the top level by controlling the expression of flhDC (Fig. 6). In accordance with the interaction of LrhA at the top level, the factors of regulation increase with increasing levels. The factors are low for class I genes (flhDC, factor about 3), intermediate for class II genes (e.g. flgB and fliA, factors 8 and 13) and highest for class III genes (e.g. fliC, factor 61) (Table 2).

image

Figure 6. LrhA and other regulators in the control of synthesis and concentration of the master regulator FlhD2C2 of flagella synthesis in E. coli. Regulators that affect flhDC expression by controlling flhDC transcription (LrhA, OmpR-P, H-NS, cAMP/CAP) or mRNA stability and translation (CsrA) positively (+) or negatively (–) are shown. In addition autoregulation of lrhA expression by LrhA is indicated. See text for more details.

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Motility and chemotaxis in E. coli are affected by numerous environmental conditions and global regulators. FlhD2C2 plays a central role as the site for the input of environmental signals. The factors required for full flhDC expression (and thus motility) include cyclic AMP and the cAMP receptor protein CAP (Kutsukake, 1997; Soutourina et al., 1999), the carbon storage regulator Csr (Wei et al., 2001b), the nucleoid-structuring protein H-NS (Bertin et al., 1994; Ko and Park, 2000), the heat shock response proteins DnaK, DnaJ and GrpE (Shi et al., 1992), the regulators Pss and Psd of phosphatidyl ethanolamine synthesis (Shi et al., 1993) and inorganic polyphosphate (Rashid et al., 2000). In contrast, motility and flhDC expression are inhibited under conditions of high acetyl phosphate levels and high osmolarity by the phosphorylated form of OmpR (Prüß and Wolfe, 1994; Shin and Park, 1995). As shown here, LrhA is a further important regulator of this network. Control of motility is an important function of LrhA in addition to the supposed role in the control of RpoS stability (Gibson and Silhavy, 1999). In this way, LrhA has a function similar to homologues HexA and PecT in Erwinia (Surgey et al., 1996; Harris et al., 1998).

The LrhA binding site

LysR-type regulators commonly bind at consensus sites (CTGA-N7-TCAG, most conserved residues underlined; Schell, 1993). The LrhA binding sites at the flhDC and lrhA promoters include a sequence with low similarity (AT-N9-AT) (Fig. 4), which could serve this function. Sites with low similarity to the general consensus are also known for OxyR (regulator of oxidative stress), CysB (regulation of cysteine biosynthesis) and NahR (naphthalene biosynthesis) (Schell, 1993). The flhDC promoter contains multiple sites for the binding of regulatory proteins (Fig. 4). These include a CAP (‘catabolite gene activator protein’) binding site centred around –71.5 (class I site), two OmpR (osmoregulation) binding sites at positions –100 to –163 and +1 to +66, respectively, and multiple binding sites for the H-NS protein from position –178 up to the coding region (Shin and Park, 1995; Soutourina et al., 1999). The LrhA binding site partially overlaps with the upstream OmpR site, whereas the hypersensitive sites are located within the downstream OmpR site.

Control of LrhA function and autoregulation

The function of most LysR-type regulators is controlled by effectors (Schell, 1993); however, the effector of LrhA and of the close homologues HexA and PecT from Erwinia is not known (Surgey et al., 1996; Harris et al., 1998). DNA binding by the homologous CbbR proteins is affected by the presence of phosphoenolpyruvate or NADPH (Van Keulen et al., 1998; Grzeszik et al., 2000). However, LrhA does not bind these or related compounds, nor does presence of such compounds affect DNA binding of LrhA (D. Lehnen and G. Unden, unpublished results). Expression of lrhA is subject to autoregulation, which could be important for the control of LrhA function. Whereas most LysR-type regulators are under negative autoregulation, lrhA is positively autoregulated similar to hexA and pecT (Van Keulen et al., 1998; Grzeszik et al., 2000). The positive autoregulation contrasts with the function of LrhA as a repressor of the other target genes (Fig. 6). However, the autoregulation of LrhA does not respond significantly to the growth phase or the C source (D. Lehnen and G. Unden, unpublished). Therefore, autoregulation and its role in the control of LrhA function is not clear so far.

Experimental procedures

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References

Bacteria and media

The bacteria, phage and plasmids used are shown in Table 4. Plasmids were maintained in E. coli JM105 (Yanisch-Perron et al., 1985). Luria–Bertani broth (LB) and LB agar were used for bacterial growth for strain and plasmid construction. For expression studies, M9 medium (Miller, 1992) supplemented with acid-hydrolysed casamino acids (0.1%) and tryptophan (0.005%) with 20 mM glycerol was used. Cultures were incubated aerobically at 37°C in flasks with shaking at 180 r.p.m. Antibiotics were added at 100 μg ml−1 ampicillin, 50 μg ml−1 kanamycin, 50 μg ml−1 spectinomycin and 15 μg ml−1 tetracycline. Cell densities were measured as the OD at 578 nm.

Table 4. Strains of E. coli K-12, phages and plasmids.
 Genotype and characteristicsReference
Strains  
 MC4100FaraD139 Δ(argF-lac)U169 rpsL150 relA1 flhD530 deoC1 ptsF25 rbsR Silhavy et al. (1984)
 BL21(DE3) E. coli B, F′hsdS gal1 DE3 Studier and Moffat (1986)
 JM105 thi strA endA sbcB15 hsdR4 (lac-proAB) F′(traD36 proAB+lacIqλαχZΔM15) Yanisch-Perron et al. (1985)
 CP992ΔlacλSS10(flhDClacZ) ampR Shin and Park (1995)
 MG1655Wild-type E. coli K-12 Jensen (1993)
 LJ10MG1655, but Δ(lacA–lacZ) 515(::cat) lacIp-4000(lacIQ)K. Jahreis (Osnabrück)
 IMW334LJ10, but λ(Φ(trglacZ)hyb)This work
 IMW353LJ10, but λ(Φ(flgBlacZ)hyb)This work
 IMW356LJ10, but λ(Φ(fliClacZ)hyb)This work
 IMW331LJ 10, but lrhA::spcRP1(IMW41) × lJ10
 IMW335LJ 10, but lrhA::spcRλ(Φ(trglacZ)hyb)This work
 IMW336LJ 10, but lrhA::spcRλ(Φ(fliClacZ)hyb)This work
 IMW354LJ 10, but lrhA::spcRλ(Φ(flgBlacZ)hyb)This work
 LJ10aLJ10 with pMW198This work
 IMW331aLJ 10, but lrhA::spcR with pMW198This work
 IMW355CP992, but lrhA::spcRP1(IMW41) × CP992
 IMW325MG1655, but lrhA::spcRP1(IMW41) × MG1655
 IMW41MC4100, but lrhA::spcR Bongaerts et al. (1995)
 IMW326MC4100, but λ(Φ(lrhAlacZ)hyb)This work
 IMW326aMC4100, but λ(Φ(lrhAlacZ)hyb) with pME6010This work
 IMW326bMC4100, but λ(Φ(lrhAlacZ)hyb) with pMW213 (lrhA+)This work
 IMW327MC4100, but lrhA::spcRλ(Φ(lrhAlacZ)hyb)This work
Plasmids  
 pMW196pJL28 derivative carrying the trg promoter (nt –458 to +20)This work
 pMW197pJL30 derivative carrying the fliC promoter (nt –656 to +101)This work
 pMW198pJL28 derivative carrying the fliA promoter (nt –501 to +15)This work
 pMW211pJL29 derivative carrying the flgB promoter (nt –223 to +249)This work
 pME6010Cloning vector, pVS1 shuttle vector, tetR Heeb et al. (2000)
 pMW213pME6010, but lrhA and promoter (–359 to end of lrhA gene)This work
 pJL28p(‘lacZ, bla+), protein fusion vectorE. Bremer (Marburg)
 pJL29p(‘lacZ, bla+), protein fusion vectorE. Bremer (Marburg)
 pJL30p(‘lacZ, bla+), protein fusion vectorE. Bremer (Marburg)
 pMW212pKS derivative carrying the flhD promoter (nt –733 to +33)This work
 pMW132pET28a, but with complete lrhA gene for LrhA expressionThis work
 pMW179pJL29 derivative carrying the lrhA promoter (nt –964 to +30)This work
 pMW226pKS derivative carrying the lrhA promoter (nt –964 to +30)This work
Phage  
 P1kc  Miller (1992)
 λRZ5λ‘bla, ‘lacZ, lacY+ Ostrow et al. (1986)

RNA preparation

Total RNA was extracted from bacteria grown aerobically in 35 ml of M9 mineral medium supplemented with small amounts of amino acids (Miller, 1992; Tran et al., 1997) and 20 mM glycerol to an OD578 of 0.3–0.45. Cells were harvested and disrupted mechanically (Wendisch et al., 2001), and total RNA was isolated using Qiagen RNeasy mini columns. RNA was treated with 30 U of RNase-free DNase I (Roche) and extracted with phenol–chloroform–isoamyl alcohol (25:24:1, v/v/v) and chloroform–isoamyl alcohol (24:1, v/v). RNA was precipitated with 0.1 volume of 3 M sodium acetate, pH 5.2, and 3 volumes of ethanol. The RNA concentration and purity were determined by A260 and A280 measurement (A260/A280 > 1.8). RNA was reverse transcribed and fluorescently labelled (Wendisch et al., 2001). Briefly, 30 μg of total RNA was reverse transcribed with 500 ng of random hexanucleotide primers (Amersham Pharmacia), 0.1 mM Cy3 or Cy5-dUTP, 50 mM Tris-HCl, pH 8.3, 75 mM KCl, 3 mM MgCl2, 10 mM dithiothreitol (DTT), 0.5 mM dATP, 0.5 mM dGTP, 0.5 mM dCTP, 0.2 mM dTTP and 400 U of Superscript II reverse transcriptase (Life Technologies). After 2 h at 42°C, reactions were stopped with 0.05 M NaOH, incubated for 10 min at 65°C and neutralized with 0.05 M HCl. Using Microcon-30 microconcentrators (Amicon), the labelled cDNA was concentrated and deoxynucleotides removed.

DNA microarray analysis

Microarrays covering 4108 (about 96% of the genome) E. coli genes were prepared as described previously (Zimmer et al., 2000) with polymerase chain reaction (PCR) products generated using the Genosys ORFmer primer set (Sigma-Genosys) (Richmond et al., 1999). Fluorescently labelled cDNAs were mixed and hybridized to the DNA microarrays in 3× SSC, 25 mM HEPES, pH 7, 0.25% SDS after 2 min heat treatment at 100°C. After hybridization at 65°C for 5–16 h in a humid chamber, the DNA microarrays were washed in 1× SSC–0.03% SDS and in 0.05× SSC. Fluorescence was determined using an Axon Genepix 4000 laser scanner. Fluorescence intensities and background levels were calculated using GENEPIX software. Scanning and normalization were carried out as described previously (Khodursky et al., 2000).

Genetic procedures and DNA manipulation

The trglacZ fusion plasmid (pMW196) was constructed by PCR amplification of the promoter region of trg (–452 to +20) from genomic DNA of strain MG1655 with primers trgEcoRI (5′-CAGGAGTGTGAATTCCAGAATC) and trgHindIII (5′-GCAAAAAACCTAAGCTTTGTGAGG). The resulting 478 bp fragment comprising the complete intergenic region was cloned into the EcoRI and HindIII sites of the protein fusion vector pJL28. The fliClacZ (650 to +101) (plasmid pMW197), fliAlacZ (495 to +15) (plasmid pMW198) and flgBlacZ (223 to +249) (plasmid pMW211) fusions were constructed in the same way with primers fliCEcoRI (5′-CGC CTTGTTGAATTCTGAGTTTAC), fliCHindIII (5′-GTTAATACG CAAGCTTGAAGACAG), fliASalI (5′-GTTACCAACGTCGAC AACAC), fliAHindIII (5′-CATTAAAGCTTCAGCGGTAT AG), flgBEcoRI (5′-GTTGTGGAATTCCGATGTGAG) and flgBHindIII (5′-GGTATTGAAGCTTTGCGGTAG) respectively. Genetic constructions of lrhA derivatives were based on the lrhA sequence in Blattner et al. (1997) as the original lrhA sequence (Bongaerts et al., 1995) was altered by a cloning artifact (B. Wackwitz and G. Unden, unpublished) at the 3′ end of the gene. For constructing the lrhAlacZ fusion, the lrhA promoter region was amplified by PCR with primers lrhAHindIII (5′-CGAGGTCAAGCTTAATTATCGG-3′) and lrhABglII (5′-GGACCACAGATCTCATAACAGA-3′) from E. coli AN387 DNA. The resulting fragment was restricted with HindIII and BglII and cloned into the HindIII and BamHI sites of plasmid pJL29, resulting in plasmid pMW179. After verification by automated sequencing, the gene fusions were transferred into the genome of E. coli with phage λRZ5 or transferred by transformation (pMW179) into the receptor strain. For construction of strains, P1 transduction was used (Bongaerts et al., 1995).

Plasmid pMW213 containing the complete lrhA gene coding for (functional) LrhA (with an unintentional D150G exchange in the protein) and the promoter region up to position –357 (T [RIGHTWARDS ARROW] C exchange at position –230) was used for the lrhA complementation experiments. The plasmid was constructed with primers lrhA3 (5′-GTCTCAGGAATTCTCT ATCGTCCG) and lrhA6 (5′-GGACCACAGATCTCATAA CAG). The amplified fragment was cloned into pME6010 digested with EcoRI and BglII.

For overexpression and isolation of LrhA, a 957 bp fragment of chromosomal DNA (from E. coli AN387) containing the coding sequence of the lrhA gene was amplified using PCR with primers lrhA3 (5′-GTCTCAGGAATTCTCTA TCGTCCG) and lrhA4 (5′-GCCAGTAAGTGATAACATATGA TAAGTGC). After restriction, the fragment was cloned into the expression vector pET28a (Novagen), yielding pMW132 coding for LrhA with a His6 tag and a linker with a thrombin cleavage site close to the N-terminus of LrhA.

Gel retardation

For gel retardation assays, the DNA fragment (766 bp) containing the flhD promoter region was obtained by PCR with primers flhDHindIII and yecG from pMW212, digested with HindIII, purified and labelled with [α-33P]-dATP at both strands. The labelling mixture contained 0.1 pmol of DNA fragment, Klenow reaction buffer, 0.25 mM dNTP mix (without labelled nucleotide), 22 μCi of [α-33P]-dATP and 5 U of Klenow enzyme (exo fragment), and was incubated for 25 min at 30°C and for 10 min at 75°C (MBI Fermentas). The fragments for gel retardation of the fliC, fliA, trg and flgB promoter regions were prepared by PCR from a colony of MG1655 with primers fliCEcoRI, fliCHindIII, fliASalI, fliAHindIII, trgEcoRI, trgHindIII, flgBEcoRI and flgBHindIII, and are 757 bp, 516 bp, 478 bp and 472 bp in length respectively. The lrhA promoter fragment (676 bp) was amplified by PCR from pMW179 using primers lrhAHindIII and lrhABglII, and digested with HindIII and EcoRI. The gel retardation assays were performed essentially as described previously (Drapal and Sawers, 1995). The proteins were incubated with labelled DNA (5 nM) in binding buffer (10 mM Tris-HCl, pH 7.5, 10% glycerol, 2.5 mM EDTA, 50 mM KCl, 0.1 mM DTT, 4 mM spermidine, 12.5 μg of sonicated calf thymus DNA, 1 μg of BSA in a final volume of 20 μl) for 30 min at room temperature. After incubation, the reaction mix was applied to a non-denaturing polyacrylamide gel (5%) buffered with TBE (Sambrook et al., 1989).

DNase I footprinting

Plasmid pMW212 with the flhD promoter was obtained by PCR amplification of the promoter region from a colony of E. coli wild-type MG1655 with oligos flhDHindIII (5′-GATGT CATAAAGCTTTTTCAGCAA) and yecG (5′-GCGATAGATAC CGCTTTTGC). After restriction with HindIII, the 766 bp fragment containing the complete flhD–yecG intergenic region was cloned in the HindIII site of pBluescript KS. The flhD promoter fragment (818 bp) for the DNase I footprinting assays was prepared from plasmid pMW212 by restriction with EcoRI and KpnI. For the footprint at the lrhA promoter, the HindIII–ApaI lrhA promoter DNA (1018 bp) from plasmid pMW226 was used. The fragments were labelled on one strand with [α-33P]-dATP for 25 min at 30°C and for 10 min at 75°C with the mixture containing 2 pmol of DNA fragment, 1× Klenow reaction buffer (MBI Fermentas), 0.25 mM dNTP mix (without labelled nucleotide), 44 μCi of [α-33P]-dATP and 20 U of Klenow enzyme (exo fragment). DNase I footprinting was performed essentially as described previously (Drapal and Sawers, 1995). LrhA was incubated with labelled DNA (40 nM) for 30 min at room temperature in the footprinting buffer (10 mM Tris-HCl, pH 7.5, 10% glycerol, 2.5 mM EDTA, 50 mM KCl, 0.1 mM DTT, 4 mM spermidine, 20 μg of BSA in 50 μl). The footprinting reaction and electrophoresis were performed as described by Drapal and Sawers (1995). As size marker, plasmid DNA sequenced by the chain termination method (T7 sequencing kit, Pharmacia) was used.

Expression and purification of His6–LrhA

pMW132 was transformed into E. coli BL21(DE3) and cultured in LB with 50 μg ml−1 kanamycin at 37°C to an OD of 0.7. Then, 1 mM IPTG was added. After 2 h, the cells were harvested by centrifugation and suspended in buffer I [50 mM K-phosphate, pH 7.0, 20 mM imidazole, 300 mM KCl, 5% glycerol, 1 mM Pefabloc (AppliChem)]. After disruption in a French pressure cell and centrifugation at 11 000 r.p.m. for 30 min, the protein was purified by standard procedures with a Ni–NTA–agarose column (Qiagen). LrhA (>90% purity) was eluted in buffer I with 500 mM imidazole. All steps in the isolation procedure were performed at 4°C. Protein measurement and SDS gel electrophoresis were performed as described previously (Laemmli, 1970; Bradford, 1976).

Motility test of E. coli strains

Tryptone swarm plates (1% tryptone, 0.5% NaCl and 0.3% Difco Bacto agar) (Tisa and Adler, 1995) were spotted with 15 μl of cells toothpicked from colonies and resuspended in chemotaxis buffer (10 mM K+-phosphate at pH 7.0, 0.1 M K+ EDTA and 0.1 mM L-methionine) and incubated for 4 h at 37°C. For the chemotaxis assays, minimal swarm plates containing 10 mM K+-phosphate at pH 7.0, 1 mM (NH4)2SO4, 1 mM MgSO4, 0.3% Difco Bacto agar, 20 μg ml−1 each L-threonine, L-leucine, L-histidine, L-methionine and 1 μg ml−1 thiamine were incubated overnight at 30°C. For testing aspartate chemotaxis, the minimal swarm agar was supplemented with 100 μM aspartate, 0.5% NaCl and 1 mM glycerol (Weerasuriya et al., 1998). In the aerotaxis tests, the minimal swarm agar was supplemented with 30 mM succinate (Bibikov et al., 2000).

Assay of β-galactosidase

The strains were grown at 37°C in M9 medium supplemented with glycerol (20 mM). From exponentially growing bacteria (OD578 = 0.5–0.6), the specific activity was measured and calculated (Miller, 1992).

Acknowledgements

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References

This paper is dedicated to Professor H. Sahm (Jülich) on the occasion of his 60th birthday. The work was supported by grants from Deutsche Forschungsgemeinschaft and the Fonds der Chemischen Industrie to G.U., and from the Swiss National Foundation for Scientific Research (project 83EU-059835) to C.B. V.F.W. and T.P. wish to thank Hermann Sahm for constant support. We are grateful to K. Jahreis (Osnabrück), C. Park and E. Bremer (Marburg) for the supply of bacterial strains.

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  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
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