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Journal of Bacteriology logoLink to Journal of Bacteriology
. 2008 Jan 25;190(7):2496–2504. doi: 10.1128/JB.01670-07

Regulation of Swarming Motility and flhDCSm Expression by RssAB Signaling in Serratia marcescens▿

Po-Chi Soo 4,†, Yu-Tze Horng 3,†, Jun-Rong Wei 3, Jwu-Ching Shu 1, Chia-Chen Lu 2, Hsin-Chih Lai 1,*
PMCID: PMC2293207  PMID: 18223092

Abstract

Serratia marcescens cells swarm at 30°C but not at 37°C, and the underlying mechanism is not characterized. Our previous studies had shown that a temperature upshift from 30 to 37°C reduced the expression levels of flhDCSm and hagSm in S. marcescens CH-1. Mutation in rssA or rssB, cognate genes that comprise a two-component system, also resulted in precocious swarming phenotypes at 37°C. To further characterize the underlying mechanism, in the present study, we report that expression of flhDCSm and synthesis of flagella are significantly increased in the rssA mutant strain at 37°C. Primer extension analysis for determination of the transcriptional start site(s) of flhDCSm revealed two transcriptional start sites, P1 and P2, in S. marcescens CH-1. Characterization of the phosphorylated RssB (RssB∼P) binding site by an electrophoretic mobility shift assay showed direct interaction of RssB∼P, but not unphosphorylated RssB [RssB(D51E)], with the P2 promoter region. A DNase I footprinting assay using a capillary electrophoresis approach further determined that the RssB∼P binding site is located between base pair positions −341 and −364 from the translation start codon ATG in the flhDCSm promoter region. The binding site overlaps with the P2 “−35” promoter region. A modified chromatin immunoprecipitation assay was subsequently performed to confirm that RssB∼P binds to the flhDCSm promoter region in vivo. In conclusion, our results indicated that activated RssA-RssB signaling directly inhibits flhDCSm promoter activity at 37°C. This inhibitory effect was comparatively alleviated at 30°C. This finding might explain, at least in part, the phenomenon of inhibition of S. marcescens swarming at 37°C.

Swarming is a bacterial population surface translocation behavior demonstrated in a wide range of diverse bacterial genera and species (2, 12, 14). In Serratia spp., swarming requires close interactions between the environment and the bacterial cells, as well as among the cells, in order to develop a high degree of complex cell coordination within the swarming colony (2, 9, 13, 26, 33, 35). Previous studies on the regulation of swarming showed that bacterial flagellar, quorum-sensing, and two-component systems are important for swarming (3, 7, 33). Among these, flagellar motility, which is one of the essential factors for bacterial swarming, is controlled by the flagellar system, comprising large and complex regulons (4, 9). Studies with the flagellar systems of Escherichia coli and Salmonella enterica serovar Typhimurium have identified around 50 genes organized into three hierarchical transcriptional classes. At the top of the hierarchical cascade is the class I master operon flhDC (4). The FlhD2C2 complex is a transcriptional activator of σ70-dependent transcription from class II promoters (4). Thus, activation of the whole set of flagellar motility genes depends mainly on the expression of flhDC.

Serratia marcescens cells swarm at 30°C but not at 37°C (15). In a previous study utilizing a mini-Tn5 mutagenesis approach, we had discovered a group of S. marcescens mutant strains that demonstrated precocious swarming behavior not only at 30°C but also at 37°C (15). A pair of bacterial two-component signal transduction proteins, RssA and RssB, were subsequently identified as negative regulators for S. marcescens swarming at 37°C (15, 36). Although RssA-RssB His-Asp phosphorelay and signaling had already been proven in vitro (36), the underlying mechanism of the RssA-RssB signaling effect on the inhibition of swarming at 37°C remained undetermined. Previously we had also shown that expression of flhDCSm and hagSm (the flagellin structural gene in S. marcescens) was reduced when the incubation temperature was increased from 30 to 37°C in S. marcescens (18). In this study, we further report that RssA-RssB negatively regulates flhDCSm expression and flagellum production through its signaling status. This regulation is achieved through direct binding of RssB∼P with the flhDCSm P2 promoter region, leading to a reduction in the level of flhDCSm mRNA transcription. The RssA-RssB inhibitory effect is much more significant at 37°C than at 30°C, which might at least in part explain the underlying mechanism of inhibition of S. marcescens swarming at 37°C.

MATERIALS AND METHODS

Bacterial strains, mutants, and culture conditions.

S. marcescens CH-1 and the rssA mutant strain S. marcescens CH-1ΔA, in which rssA is interrupted by a HindIII-digested Smr gene cassette, were from a previous study (36). Escherichia coli DH5α (Invitrogen) was used as a host strain for the maintenance of recombinant DNA plasmids. E. coli BL21(DE3)pLysS (Novagen, Germany) was used for oversynthesis of recombinant proteins. All bacteria used in this study were grown in Luria-Bertani (LB) medium at 37°C (36) supplemented with adequate antibiotics when necessary, unless other conditions are specifically mentioned in the text.

Enzymes, chemicals, and primers.

DNA restriction and modification enzymes were purchased from Roche (Germany). Pfu polymerase and PCR-related products were from Stratagene and Perkin-Elmer. Other laboratory-grade chemicals were purchased from Sigma and Merck (Germany). The primers used in this study and summarized in Table 1 were purchased from MD Bio (Taiwan).

TABLE 1.

Bacterial strains, plasmids, and primers used in this study

Strain, plasmid, or primer Relevant characteristic(s) or sequence (5′→3′) Source or reference
Serratia marcescens strains
    CH-1 Clinical isolate 15
    CH-1ΔA rssA knockout mutant 15
Plasmids
    pGEX Expression vector, GST tag Pharmacia Biotech
    pGST-B pGEX::rssBSm; Ampr Smr This study
    pGST-B(D51E) pGEX::rssBSm(D51E); Ampr Smr This study
    pBG300 pACYC184 (PflhDCSm::luxCDABE); Cmr 26
    pBG301 pACYC184 (P1flhDCSm::luxCDABE); Smr Cmr This study
    pBG302 pACYC184 (P2flhDCSm::luxCDABE); Smr Cmr This study
    pBG pACYC184 (promoterless luxCDABE); Smr Cmr This study
Primers
    FlhD-PF CAGCCTCAGGCGGAGGG This study
    FlhD-PR ATTCCCCATCCCGACAGAGCTA This study
    F0-F TGCTAATGGTTCAGGGG This study
    F0-R DIG-ATTCCCCATCCCGACAGACTA This study
    Fa-F CAGCCTCAGGCGGAGGG This study
    Fa-R DIG-CTGAACCATTAGCACACA This study
    Fb-F CAGCCTCAGGCGGAGGG This study
    Fb-R TTGCTACGAGCGTAAACCAA This study
    FlhD-PE FAM-CGTACCCATATTCCCCATCCC This study
    flhD-SF TGTCGGGATGGGGAATATGG This study
    flhD-SR CGATAGCTCTTGCAGTAAATGG This study
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Swarming motility assay.

A swarming assay was performed on LB medium solidified with 0.8% Eiken agar (Eiken, Japan) by inoculating 3-μl portions of an overnight LB broth culture onto the centers of agar plate surfaces and incubating at 37°C.

Detection of luxCDABE reporter luciferase activity.

The Autolumat LB 953 luminometer (EG & G, Germany) with the program “replicates” was used for bioluminescence measurement. All procedures followed the protocols supplied by the manufacturer.

Gel mobility shift assay.

Promoter DNA fragments for gel mobility shift assays were amplified by PCR using primers that were 5′ end labeled with digoxigenin (DIG) (MWG Biotech, Germany). Reaction mixtures for the binding assay comprised 2 μM RssB protein with acetylphosphate treatment and 0.5 ng DIG-labeled promoter DNA fragments. For the serial dilution experiments, phosphorylated or unphosphorylated RssB proteins were serially diluted in binding reaction buffer (14 mM Tris-HCl [pH 7.4], 6.9 mM MgCl2, 69 mM KCl, and 10 mM EDTA). The binding reaction was performed in binding reaction buffer supplemented with 30 μg/ml poly(dI-dC) and 1 μg/μl bovine serum albumin. The reaction mixtures were incubated for 20 min at room temperature before being loaded onto 7% nondenaturing polyacrylamide gels containing 0.5× Tris-borate-EDTA buffer. Electrophoresis was performed at 100 V for 1 h. The DNA-protein complexes were then electroblotted onto a positively charged Hybond-N nylon membrane (Amersham, United Kingdom) and detected by alkaline phosphatase-conjugated anti-DIG antibodies (Roche, Germany). CSPD was added as the substrate as described by the manufacturer (Roche, Germany). Membranes were exposed to X-ray film at room temperature for 2 to 30 min.

RT-PCR assay.

Total bacterial RNA was extracted using a Trizol kit (Invitrogen). The relative amounts of transcripts from the flhDCSm gene were evaluated by the reverse transcription-PCR (RT-PCR) assay. RNA was isolated from strains CH-1 and CH-1ΔA grown aerobically on 0.8% LB agar plates for 2 or 3 h (early-logarithmic phase) and then reverse transcribed into cDNA with a SuperScript III first-strand synthesis system kit (Invitrogen). Equal amounts of total RNA (5 μg) were used to generate cDNA with random hexamer primers according to the manufacturer's protocol. The products were amplified by PCR with the primer pair flhD-SF-flhD-SR (Table 1). The cycle conditions for the PCR were as follows: 1 min at 96°C, 1 min at 60°C, and 50 s at 72°C for 15 or 25 cycles. The number of cycles was decided according to the comparison performed in the linear range of amplification. The RT-PCR products were analyzed by electrophoresis on 2% agarose gels, and then the amount of transcript was quantified by densitometry using the Scion Imager. 16S rRNA was used as the internal control to confirm that equal amounts of total RNA were used in each reaction (11).

Identification of the transcriptional start site(s) in flhDCSm and quantification of promoter activity.

The transcriptional start sites were identified by a primer extension assay using a 6-carboxyfluorescein (FAM)-labeled primer and the GeneScan analysis system (Applied Biosystems) described by Lloyd et al. (19) with some modifications. Briefly, purified RNA was reverse transcribed into cDNA with ImProm-II reverse transcriptase (Promega) by use of primer FlhD-PE to form FAM-labeled cDNA (Table 1). Samples containing 1 μg of RNA and 1 μM primers were heated at 70°C for 5 min before being placed on ice for 1 min. After addition of 0.5 mM deoxynucleoside triphosphates, 25 mM MgCl2, 1 μl reverse transcriptase, and 1× RT buffer (supplied by the manufacturer) in a 20-μl final volume adjusted with diethylpyrocarbonate water, samples were incubated at 55°C for another 60 min. The reaction was stopped by addition of 100 μl of H2O and 100 μl of phenol-chloroform-isoamyl alcohol (25:24:1). After centrifugation at 12,000 × g for 4 min, the supernatant containing cDNA was precipitated with 0.1 volume of solution III (60% sodium acetate, 11.5% glacial acetic acid [pH 5.2]) and 2.5 volumes of absolute ethanol. After incubation at −70°C for more than 30 min, cDNA was centrifuged at 12,000 × g for 15 min. The pellet was washed with 70% ethanol and dissolved in 4 μl of deionized H2O, followed by addition of 5 μl of deionized formamide containing 0.5 μl of the ROX-500 molecular size standard (Applied Biosystems). Samples were denatured at 95°C for 5 min and then chilled quickly on ice. Electrophoresis was performed on 6% polyacrylamide-8 M urea gels by using the ABI Prism 3100 capillary DNA genetic analyzer equipped with Avant Genetic Analyzer Data Collection, version 2.0 (both from Applied Biosystems). The data were analyzed using GeneMapper software, version 3.5 (Applied Biosystems). The cDNAs of the flhDCSm transcripts were reverse transcribed with a FAM-labeled primer. The length of the FAM-labeled cDNA primer extension product was then analyzed by using an ABI 3100 automated sequencer and GeneScan software (both from Applied Biosystems).

DNase I footprinting.

The DNase I footprinting protocol was modified from that described by Yindeeyoungyeon and Schell (39). Briefly, the 500-bp flhDCSm promoter region was PCR amplified from genomic DNA. The primer pairs used for PCR were Fb-F and Fb-R. Depending on which strand was analyzed, one primer was labeled with FAM (MD Bio, Taiwan) at the 5′ end and the other was not. The FΑΜ-labeled DNA fragment was incubated with 28 μM phosphorylated RssB in a 50-μl solution containing 11 mM Tris-HCl (pH 7.4), 0.1 mM EDTA, 5.5 mM MgCl2, 20 μg/ml of poly(dI-dC) (Pharmacia, Sweden), and 0.2 mg/ml of bovine serum albumin. After 20 min of incubation at room temperature, 50 μl of DNase I (1 × 10−3 U/μl, freshly prepared by diluting the stock [Promega] in D buffer, consisting of 10 mM Tris-HCl [pH 7.5], 10 mM MgCl2, and 5 mM CaCl2) was added, and the mixture was further incubated at 26°C for 3 or 5 min. The digestion was stopped by adding 100 μl of stop solution containing 0.2 M NaCl, 40 mM EDTA, 1% sodium dodecyl sulfate, and 125 μg/ml of tRNA. After incubation at 37°C for 20 or 30 min, samples were extracted with a phenol-chloroform-isoamyl alcohol solution (25:24:1), precipitated with absolute ethanol, washed with 70% ethanol, and dissolved in 10 μl deionized formamide. After addition of 0.5 μl of the ROX-500 molecular size standard (Applied Biosystems), samples were denatured at 95°C for 5 min and then quickly chilled on ice. Electrophoresis using the ABI Prism 3100 capillary DNA genetic analyzer and data analysis with GeneMapper software, version 3.5 (both from Applied Biosystems), were performed according to the instructions supplied by the manufacturer.

Modified chromatin immunoprecipitation (ChIP) assay.

The immunoprecipitation assay described by Shin and Groisman (23) was used with modifications. Briefly, cultures of S. marcescens were treated with 1 M sodium phosphate (final concentration, 10 mM) and 37% formaldehyde (final concentration, 1%). After 15 min, cross-linking was quenched by the addition of glycine (final concentration, 125 mM). Cultures of 10 ml were collected by centrifugation and washed twice with 10 ml of phosphate-buffered saline. Cells were lysed in 0.6 ml of lysis solution (20 mM Tris [pH 8.0], 150 mM NaCl, 1 mM EDTA [pH 8.0], 1 mM phenylmethylsulfonyl fluoride, 1% Triton X-100, 10 mg/ml of lysozyme), and 0.6 ml of 2× IP solution (100 mM Tris [pH 8.0], 300 mM NaCl, 2% Triton X-100) was then added. Cell extracts were then sonicated to produce DNA fragments with an average size of 500 to 1,000 bp. The extract (200 μl) was removed for total-DNA preparation. For pull-down of glutathione S-transferase (GST)-RssB-cross-linked DNA, a portion of the extracts (600 μl) was incubated with 100 μl glutathione-Sepharose 4B beads (Amersham, United Kingdom) at 4°C for 1 h. GST-RssB-cross-linked DNA was then pulled down with the glutathione-Sepharose beads by centrifugation. The beads were washed twice with 1× IP solution and then twice with a LiCl-detergent solution (10 mM Tris [pH 8.0], 250 mM LiCl, 1 mM EDTA, 0.5% Nonidet P-40, and 0.5% sodium deoxycholate). The beads were then resuspended in a 200-μl solution of 50 mM Tris-HCl (pH 8.0), 1 mM EDTA, and 0.67% sodium dodecyl sulfate. This was followed by incubation at 65°C for 16 h to reverse the cross-links. DNA samples were purified using phenol extraction, precipitated with absolute ethanol, and resuspended in TE buffer (10 mM Tris, 1 mM EDTA [pH 8.0]).

RESULTS

RssA negatively regulates the flagellar system on LB swarming plates.

Previously we had shown that inactivation in S. marcescens CH-1 of rssA or rssB, a sensor kinase gene and a cognate response regulator gene, respectively, making up a bacterial two-component system, resulted in precocious swarming behavior on a 0.8% LB swarming agar plate at 37°C (Fig. 1A) (15). To further characterize the underlying mechanism of this precocious swarming behavior, the amounts of flagellin synthesized by S. marcescens strain CH-1 and the rssA mutant strain S. marcescens CH-1ΔA seeded on LB swarming plates at 37°C were first quantified by Western blot analysis after 3 h of bacterial growth (early-logarithmic phase). The amounts of flagellin synthesized by CH-1 and CH-1ΔA differed significantly (Fig. 1A); relative intensities were determined to be 20 ± 5 and 78 ± 10 arbitrary units, respectively (a 3.9-fold increase for CH-1ΔA) (Fig. 1A). These results suggest that CH-1ΔA produces many more flagella than CH-1 at 37°C, indicating derepression of the flagellar synthesis system as a potential underlying mechanism for the precocious swarming phenotype observed in S. marcescens CH-1ΔA.

FIG. 1.

FIG. 1.

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Swarming behavior, flagellum production, and transcriptional activity of flhDCSm in S. marcescens CH-1 and CH-1ΔA on LB swarming plates at 37°C. (A) (i) Swarming assay and determination of the amount of flagellin 3 h after inoculation onto a swarming plate by Western blotting of whole-cell lysates with an antiflagellin monoclonal antibody. (ii) Relative intensities of flagellin synthesized were measured using Scion Imaging software. (B) Transcriptional activities of flhDCSm (pBG300) in CH-1ΔA (▪) and CH-1 (▴) measured by a luxCDABE luciferase reporter activity assay after seeding onto a swarming plate. (C) Determination by RT-PCR of flhDCSm mRNA and 16S rRNA levels in S. marcescens cells incubated for 2 h on a seeding plate. Hatched bar, CH-1; open bar, CH-1ΔA; *, P < 0.01. Results are means ± standard deviations from three independent experiments.

To compare the transcriptional levels of flhDCSm in CH-1 and CH-1ΔA at 37°C, the recombinant plasmid pBG300 (PflhDCSm::luxCDABE) containing luxCDABE reporter genes transcriptionally fused with the potential flhDCSm promoter region [a 655-bp DNA region upstream of the A(+1)TG translational start site (see Fig. 5A)] was constructed for real-time monitoring of flhDCSm promoter activity. The flhDCSm promoter activity in CH-1ΔA was ca. 3.6-fold elevated over that in CH-1 on LB plates seeded and incubated for 2 h at 37°C (for bacterial growth to the early-logarithmic phase) (Fig. 1B). For further confirmation of the difference in flhDCSm expression levels, RT-PCR using total cellular RNA as the template was performed for quantification. As bacterial cells were grown to the early-logarithmic phase (2 h) on LB plates at 37°C, CH-1ΔA cells showed a 53% increase in the flhDCSm expression level over that of CH-1 cells (72 ± 18 versus 47 ± 16 relative intensity units) (Fig. 1C). These findings suggested that the inhibition of swarming by RssA at 37°C in S. marcescens CH-1 was correlated with the repression of flhDCSm expression and flagellum production by RssA.

FIG. 5.

FIG. 5.

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The flhDCSm promoter region and its interaction with RssB. (A) Diagram of each flhDCSm promoter DNA fragment. Nucleotides are numbered by labeling the translation initiation site of the flhDSm gene as +1. (B) Determination by EMSA of the interaction between RssB∼P and the Fa fragment, the rssB promoter region (positive control), the ygfFSm promoter region (negative control) (36), or the F0 fragment. (C) Competition assay. A 0.5-ng portion of labeled Fa, either without RssB∼P or with 4 μM RssB∼P, was mixed with unlabeled DNA. −, no unlabeled DNA; + Fa, 0.5 ng unlabeled Fa; ++ Fa, 50 ng unlabeled Fa; ++ ygfF, 50 ng unlabeled ygfFSm promoter fragment. (D) An aspartate residue (D51) is essential for interaction between RssB and Fa. EMSA were performed using (from left to right) a labeled Fa fragment either alone, together with RssB∼P, or together with RssB(D51E), which had lost the ability to be phosphorylated.

Characterization of P1 and P2 promoters in flhDCSm.

The complexity of the regulation of flhDC expression has been well studied in many bacterial species, including E. coli and Salmonella serovar Typhimurium (4, 28). Compared to the characterized flhDC upstream promoter region in Salmonella serovar Typhimurium (28, 38), potential multiple transcriptional start sites were identified in S. marcescens CH-1 flhDCSm. To identify the transcriptional start site(s) of flhDCSm in S. marcescens at 37°C, primer extension assays were performed. Total RNA was extracted from S. marcescens seeded onto LB swarming plates and incubated for 2 h after inoculation at 37°C. The flhDCSm transcripts were first reverse transcribed into cDNA by primer extension using a FAM-labeled DNA primer designed from 9 bp downstream of the ATG translational initiation site. This was followed by length analysis of synthesized single-stranded cDNAs by using the ABI 3100 automated DNA sequencer. Two major flhDCSm transcripts were identified in S. marcescens CH-1 and S. marcescens CH-1ΔA (Fig. 2). The start sites of these two transcripts were mapped 68 bp and 306 bp upstream of the flhDCSm ATG initiation codon, and the transcripts were designated P1 and P2, respectively (Fig. 2 and 3).

FIG. 2.

FIG. 2.

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Characterization of transcriptional initiation sites of flhDCSm in S. marcescens CH-1 (A) and CH-1ΔA (B). Primer extension analysis of in vivo transcripts was performed with primer FlhD-PE. RNA was isolated from S. marcescens CH-1 and the rssA mutant strain (CH-1ΔA) grown on swarming plates at 37°C for 2 h. The fluorescence intensities of the DNA fragments (ordinate) were plotted against the sequence lengths of the fragments (abscissa). P1 and P2 indicate the potential transcription starts that map 68 and 306 bases upstream of the translational initiation codon of the flhD gene, respectively.

FIG. 3.

FIG. 3.

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Transcriptional start sites and RssB∼P binding site in the upstream promoter region of the flhDCSm operon in S. marcescens CH-1. Open arrowheads and boldfaced letters indicate the transcriptional start sites P1 and P2. The RssB∼P binding region (see Fig. 6) is boxed. The potential Shine-Dalgarno sequence is underlined; the translational start site of flhDSm is double underlined. Wavy underlining indicates the promoter consensus sequences. Nucleotides are numbered by labeling the translation initiation site of the flhDSm gene as +1. The flhDCSm promoter sequence was deposited in GenBank under accession number AF077334 (18).

To further confirm the potential P1 and P2 flhDCSm promoter activities, the P1 (bp −60 to −210) and P2 (bp −299 to −423) promoter regions were fused with the luxCDABE genes in the pACYC184 vector to form recombinant plasmids pBG301 and pBG302, respectively (Fig. 4A). These two plasmids were then transformed into S. marcescens CH-1 and CH-1ΔA for measurement of P1 and P2 promoter activities. Both P1 and P2 displayed significant promoter activities in S. marcescens CH-1 and CH-1ΔA compared with the pBG vector control (promoterless luxCDABE) (data not shown). However, the activities of the two promoters were lower in S. marcescens CH-1 than in CH-1ΔA, although a more significant reduction was observed for P2 than for P1 (Fig. 4B and C). To sum up, two flhDCSm promoters were identified in S. marcescens CH-1, and the transcriptional level of the P2 promoter was more significantly inhibited by RssA.

FIG. 4.

FIG. 4.

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Effects of RssA on promoter activities of PflhDCSm in S. marcescens CH-1. (A) flhDCSm promoter region. P1 and P2 indicate the two transcriptional start sites, and each potential promoter was fused with the luxCDABE reporter genes in the pACYC184 vector to form pBG301 and pBG302, respectively. (B and C) P1 and P2 promoter activities, respectively, at 37°C. Triangles and squares, growth curves in CH-1 and CH-1ΔA, respectively; solid and open bars, transcription activities of strains CH-1 and CH-1ΔA, respectively. Results are means ± standard deviations from three independent experiments.

Phosphorylated RssB interacts with the flhDCSm upstream promoter region.

RssA and RssB form a cognate two-component sensing system in S. marcescens CH-1. Once activated, phosphorelay signaling from RssA is transferred to RssB (15, 36). The possibility that RssB in its phosphorylated form (RssB∼P) binds directly to the flhDCSm promoter region was evaluated. Electrophoretic mobility shift assays (EMSA) were performed by using purified RssB∼P (36) mixed with either the F0 or the Fa flhDCSm promoter fragment. The 291-bp F0 DNA fragment, spanning the translational start site, ATG, and its upstream 291-bp region, contained the P1 promoter (Fig. 3 and 5A) (GenBank accession number AF077334) (18). For comparison, the promoter regions of ygfFSm (370-bp promoter DNA fragment) and rssB (104-bp promoter DNA fragment) were used as negative and positive controls, respectively (36). No sign of a DNA shift was observed (Fig. 5B), indicating no RssB∼P binding onto the F0 flhDCSm promoter fragment. Subsequently, the 378-bp (bp −278 to −655 from ATG) Fa DNA fragment, upstream of F0 and containing the P2 promoter (Fig. 5A), was PCR amplified for EMSA. A clear DNA shift was observed when RssB∼P (2 μM) and Fa (0.5 ng) were used (Fig. 5B). Furthermore, a competition assay through addition of unlabeled Fa DNA fragments (0.5 ng to 50 ng) inhibited the DNA shift and verified the specific binding of RssB∼P to Fa (Fig. 5C). These results suggested specific interaction between RssB∼P and the Fa flhDCSm promoter region. To confirm that phosphorylation of RssB was necessary for binding to the Fa region, RssB(D51E), where RssB cannot be phosphorylated (36), was used in the EMSA. No DNA shift was observed (Fig. 5D), indicating that phosphorylation was essential for the binding of RssB to the Fa fragment. Thus, RssB∼P specifically binds to the upstream promoter region of flhDCSm, and the potential binding site is located between bp −278 and −655. These results suggested that the binding site of phosphorylated RssB is located in the Fa fragment, which contains P2, but not in F0, containing the P1 region.

Determination of the specific RssB∼P binding site within the Fa region.

To determine the specific binding site(s) of RssB∼P in the Fa region, DNase I footprinting experiments using an automated capillary DNA sequencer (ABI Prism 3100) were performed. The Fb DNA fragment, which spanned bp −156 to −655 and was 122 bp longer than the Fa fragment (Fig. 5A), was PCR amplified for DNase I footprinting assays. Either the “template” or the “nontemplate” strand of Fb was modified by the fluorescent dye FAM before the assay. The results, as shown in Fig. 6, revealed that for the “template” and “nontemplate” strands, the DNA region spanning bp −364 to −341 in the flhDCSm promoter region was protected by RssB∼P. In contrast, no region within the Fb fragment was protected by RssB(D51E) in the DNase I footprinting assay (data not shown). These results confirmed that phosphorylated, and not unphosphorylated, RssB binds the region of the flhDCSm promoter from bp −364 to −341. Detailed analysis of the flhDCSm P2 promoter sequence revealed that the RssB binding site partially overlaps with the −35 region of P2 (Fig. 3).

FIG. 6.

FIG. 6.

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Identification of the RssB∼P binding site in the flhDCSm promoter region by a DNase I footprinting assay. (A and B) FAM was used to label the template (A) or nontemplate (B) strand of the PCR-amplified Fb fragment (see Fig. 5). (i and ii) Each strand was incubated in the absence or presence of RssB∼P before treatment with DNase I. (iii and iv) Expanded views of the RssB∼P binding region, selected from panels i and ii. The fluorescence intensity of the FAM-labeled DNA fragment (ordinate) was plotted against the sequence length of the fragment. (C) Partial sequence of the flhDCSm promoter region. Nucleotide positions are numbered by labeling the translation initiation site of the flhDCSm gene as +1. Nucleotides protected by RssB∼P are shadowed.

In vivo RssAB signaling and binding of RssB∼P with the flhDCSm promoter.

To determine whether RssAB signaling was important for the interaction of RssB∼P with the flhDCSm promoter in vivo, a modified ChIP assay (23) was used to evaluate the binding of RssB∼P to the flhDCSm promoter region in vivo at 37°C and 30°C. DNA binding proteins in actively growing S. marcescens CH-1 cells (early-logarithmic phase) were cross-linked onto DNAs by formaldehyde, followed by extraction and shearing of the genomic DNA into fragments of 500 to 1,000 bp. Glutathione-Sepharose was then used to pull down the complexes comprising recombinant GST fusion proteins [GST, GST-RssB, and GST-RssB(D51E)] together with the cross-linked DNA fragments. After separation of cross-linked protein-DNA complexes by heat, the presence of DNA in the pulled-down complexes was analyzed by PCR using primers designed to amplify the 378-bp Fa fragment. The results in Fig. 7A show that while no amplified Fa fragment was detected in either the GST-only or the GST-RssB(D51E) group, it was clearly detected in the GST-RssB group. These results indicated that RssB phosphorylation and the aspartate residue in RssB are essential for in vivo Fa binding. Further comparison of the amount of the Fa fragment captured by RssB in S. marcescens CH-1 showed that less Fa fragment was bound by RssB at 30°C than at 37°C (Fig. 7A).

FIG. 7.

FIG. 7.

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In vivo identification of RssB∼P binding to the flhDCSm promoter region. (A) Oversynthesis of the GST-tagged fusion proteins in S. marcescens CH-1 grown at 37°C or 30°C was used to capture the flhDCSm promoter DNA fragment, followed by a modified ChIP assay. The precipitated DNA and total (input) DNA were subjected to PCR using primers FlhD-PF and FlhD-PR, specific to the flhDCSm promoter region. (B) The modified ChIP assay was performed against the pull-down extracts from S. marcescens CH-1(RssB) (i.e., CH-1 cells containing pGEX::rssBSm), CH-1ΔA(RssB), and CH-1ΔA[RssB(D51E)] [i.e., CH-1ΔA containing pGEX::rssBSm(D51E)] cells at 37°C or 30°C. S. marcescens CH-1 is the parent strain, and S. marcescens CH-1ΔA is the rssA deletion strain.

To determine whether intact RssA-RssB signaling is essential for the binding of RssB∼P to the Fa region, a modified ChIP assay was performed using the pull-down extracts from S. marcescens CH-1(RssB) (i.e., CH-1 containing pGST-RssB), S. marcescens CH-1ΔA(RssB), and S. marcescens CH-1ΔA[RssB(D51E)] [i.e., CH-1ΔA containing pGST-RssB(D51E)] cells. Only S. marcescens CH-1(RssB) showed a positive binding result (Fig. 7B). Thus, RssA is essential for RssB∼P binding with the Fa fragment, indicating that under in vivo conditions, phosphorylated RssB binds directly to the flhDCSm promoter region only when its cognate sensor, RssA, is present. Thus, a complete RssA-RssB signaling pathway is important for RssB∼P binding and inhibition of flhDCSm promoter activity on LB swarming plates at 37°C.

Role of RssA signaling in flhDCSm promoter activity at 30 and 37°C.

We had previously shown that the promoter activities of PflhDCSm and PhagSm are reduced when the incubation temperature is changed from 30 to 37°C (18). The amount of flagellin synthesized in S. marcescens CH-1 on LB seeding plates was measured at both 30 and 37°C for comparison. A significant, 2.61-fold increase in the amount of flagellin at 30°C over that at 37°C was observed in CH-1 (Table 2). Further quantification by primer extension of flhDCSm promoter activity in S. marcescens CH-1 grown to early-log phase showed that the P1 and P2 promoter activities at 30°C were 1.96-fold and 2.31-fold higher than those at 37°C, respectively (Table 2). In contrast, the amount of flagellin synthesized and the level of flhDCSm expression in S. marcescens CH-1ΔA showed no statistically significant difference when the incubation temperature was changed from 30 to 37°C.

TABLE 2.

Quantification of flhDCSm promoter activity and flagellin synthesis in S. marcescens CH-1 and CH-1ΔA at 30 and 37°Ca

Strain and temp P1
P2
Flagellin synthesis
Activity Ratio Activity Ratio Amt Ratio
CH-1
    30°C 1,351 ± 51 1.96 2,539 ± 105 2.31 60 ± 2 2.61
    37°C 688 ± 31 1,099 ± 43 23 ± 4
CH-1ΔA
    30°C 1,444 ± 30 1.14 3,271 ± 173 1.15 87 ± 4 1.12
    37°C 1,268 ± 19 2,847 ± 185 78 ± 9
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a

RNA was isolated from S. marcescens CH-1 and the rssA mutant strain (CH-1ΔA) grown on swarming plates for 2 h. flhDCSm P1 and P2 promoter activities were evaluated by primer extension using a FAM-labeled primer followed by electrophoresis on an ABI Prism 3100 capillary DNA genetic analyzer. The promoter activity was scored for relative fluorescence intensity by using the peak area with GeneMapper software, version 3.5. The amount of flagellin was evaluated by calculation of the relative intensities of Western blotting results using Scion Imaging software after incubation for 3 h on seeding plates. The promoter activities and amounts of flagellin are presented as arbitrary units. Each result is the mean ± standard error from three independent experiments, with triplicate samples in each assay. Ratios were calculated as the promoter activity or flagellin amount of a strain incubated at 30°C divided by that at 37°C.

DISCUSSION

In this communication, we characterized the underlying mechanism of the RssA-RssB effect on the regulation (inhibition) of swarming in S. marcescens CH-1 at 37°C. When rssA was mutated, significant increases in flagellum production and flhDCSm promoter activity, and in accompanying precocious swarming behavior, were observed. Further studies showed that a complete RssA-RssB signaling system is required for inhibition of S. marcescens CH-1 swarming. The underlying molecular mechanism of the RssA-RssB effect lies in direct interaction of RssB∼P with the flhDCSm promoter region located between bp −341 and −364 from the translational initiation codon ATG (Fig. 6C). Such interaction reduced the levels of transcription and expression of flhDCSm. P1 promoter activity was also affected by RssAB signaling, albeit to a lesser extent (Fig. 4B). To date, however, no evidence has been obtained showing that RssB binds to the flhDCSm P1 promoter region. Thus, the possibility exists that P1 promoter activity is indirectly affected by RssAB signaling through a steric hindrance effect. Although the growth phase-dependent phosphorylation (and thus activation) of RssA-RssB signaling in vivo remains to be characterized, our experimental results indicated that when S. marcescens is grown to the early-logarithmic phase on LB swarming plates at 37°C, RssB∼P shows evidence of binding to the promoter region of flhDCSm, subsequently inhibiting its level of transcription (Table 2; Fig. 7). These results suggested that RssA-RssB signaling modulates flhDCSm expression and that when this signaling is activated, flhDCSm expression is inhibited.

Synthesis of FlhD and FlhC from flhDC is complicated, and multiple levels of intracellular regulation exist, including transcriptional and posttranscriptional control in E. coli and even posttranslational control in Proteus mirabilis (5, 28) and Salmonella serovar Typhimurium (32). The expression of flhDC is both positively and negatively regulated by the histone-like nucleoid-structuring H-NS protein, and H-NS-binding sites have been identified upstream and downstream of the promoter region in E. coli (27, 29). flhDC expression is also positively regulated by the cAMP-CAP (catabolite gene activator protein) complex, and the CAP-binding site is found upstream of the flhDC promoter in E. coli and Salmonella serovar Typhimurium (1, 25, 27, 40). A LysR-type regulator, LrhA, also negatively regulates flhDC expression in E. coli (16). In P. mirabilis, the flhDC operon is up-regulated by four unlinked genes, umoA, umoB, umoC, and umoD, encoding putative membrane or periplasmic proteins (8). Moreover, besides transcriptional activity, the stability of flhDC mRNA is controlled by the RNA binding regulator, CsrA, in E. coli (21, 34). These results indicate that the regulation of flhDC expression is under complicated and stringent controls.

The regulation of flhDC expression is complicated by influences from numerous environmental signals, such as temperature, osmolarity, and pH. The two-component signal transduction systems (17, 22, 29) play important roles in mediating these regulatory processes (31, 37). High osmolarity inhibits flhDC expression through the response regulator OmpR, and two OmpR-binding sites have been found in the flhDC promoter region in E. coli (24). Expression of flhDC in Xenorhabdus nematophila is also repressed by EnvZ-OmpR in response to high environmental osmolarity (20). A report by Sperandio et al. showed that flhDC expression was positively regulated by quorum sensing through QseBC in E. coli (30). QseB binds directly to the flhDC promoter at two sites, the high- and low-affinity binding sites, respectively (6). In E. coli, the RcsCDB His-Asp phosphorelay system is a negative regulator of the flhDC operon. The site of binding of RcsAB to the flhDC promoter was mapped as the RcsAB box, located downstream of the promoter in E. coli (10, 20). These results indicated that two-component systems either positively or negatively regulate the expression of flhDC, depending on the environmental signals and binding sites of the response regulators in the promoter regions.

Compared with the findings for known regulatory systems for flhDC expression, the experimental results of this study showed a negative regulatory characteristic of RssAB in S. marcescens as the culture temperature is shifted from 30 to 37°C. When S. marcescens was cultured on LB swarming plates at 37°C, the transcriptional level of flhDCSm was reduced from that at 30°C. This might be due, at least in part, to interaction between RssB∼P and the upstream region of the flhDCSm P2 promoter (from bp −364 to −341) and subsequent inhibition of flhDCSm RNA transcription (Fig. 7; Table 2). Based on these results, it was reasoned that the extents of RssAB signaling and phosphorylation of RssB might be decreased when the culture temperature is shifted from 37 to 30°C. Indeed, ChIP results confirmed less Fa binding by RssB at 30°C than at 37°C in vivo (Fig. 7). Furthermore, such temperature-related inhibition of flhDCSm expression activities might be connected to S. marcescens pathogenesis in humans. Since increases in the expression of S. marcescens virulence factor genes, such as hemolysin, were observed for precocious-swarming mutants (15), and S. marcescens swarming is inhibited at the human body temperature, 37°C, it is reasoned that the expression of virulence factors in S. marcescens is reduced during human infection. In a rat model of acute pneumonia, S. marcescens strain CH-1ΔAB, in which rssA and rssB were deleted, showed a significantly more virulent phenotype and caused a much higher mortality rate than its parent strain, S. marcescens CH-1 (data not shown). In conclusion, swarming of S. marcescens CH-1 at 37°C is under strict regulation, and activation of the RssA-RssB signaling system and inhibition of the flagellar system and virulence factor expression contribute to such phenotype regulation. Whether similar regulatory effects occur at 30°C, leading to a density-dependent swarming phenomenon in S. marcescens, is currently under study.

Acknowledgments

This work was supported by grants from the National Science Council (NSC-95-2320-B-002-061-MY3 and NSC96-2320-B-155-001) and the Technology Development Program for Academia, Ministry of Economical Affairs (91-EC-17-A-10-S1-0013), which were greatly appreciated.

We are grateful to Yu-Huan Tsai for excellent technical assistance.

Footnotes

▿

Published ahead of print on 25 January 2008.

REFERENCES

  • 1.Adler, J., and B. Templeton. 1967. The effect of environmental conditions on the motility of Escherichia coli. J. Gen. Microbiol. 46175-184. [DOI] [PubMed] [Google Scholar]
  • 2.Alberti, L., and R. M. Harshey. 1990. Differentiation of Serratia marcescens 274 into swimmer and swarmer cells. J. Bacteriol. 1724322-4328. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 3.Berg, H. C. 2005. Swarming motility: it better be wet. Curr. Biol. 15R599-R600. [DOI] [PubMed] [Google Scholar]
  • 4.Chilcott, G. S., and K. T. Hughes. 2000. Coupling of flagellar gene expression to flagellar assembly in Salmonella enterica serovar Typhimurium and Escherichia coli. Microbiol. Mol. Biol. Rev. 64694-708. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 5.Claret, L., and C. Hughes. 2000. Rapid turnover of FlhD and FlhC, the flagellar regulon transcriptional activator proteins, during Proteus swarming. J. Bacteriol. 182833-836. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 6.Clarke, M. B., and V. Sperandio. 2005. Transcriptional regulation of flhDC by QseBC and σ28 (FliA) in enterohaemorrhagic Escherichia coli. Mol. Microbiol. 571734-1749. [DOI] [PubMed] [Google Scholar]
  • 7.Daniels, R., J. Vanderleyden, and J. Michiels. 2004. Quorum sensing and swarming migration in bacteria. FEMS Microbiol. Rev. 28261-289. [DOI] [PubMed] [Google Scholar]
  • 8.Dufour, A., R. B. Furness, and C. Hughes. 1998. Novel genes that upregulate the Proteus mirabilis flhDC master operon controlling flagellar biogenesis and swarming. Mol. Microbiol. 29741-751. [DOI] [PubMed] [Google Scholar]
  • 9.Eberl, L., S. Molin, and M. Givskov. 1999. Surface motility of Serratia liquefaciens MG1. J. Bacteriol. 1811703-1712. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 10.Francez-Charlot, A., B. Laugel, G. A. Van, N. Dubarry, F. Wiorowski, M. P. Castanie-Cornet, C. Gutierrez, and K. Cam. 2003. RcsCDB His-Asp phosphorelay system negatively regulates the flhDC operon in Escherichia coli. Mol. Microbiol. 49823-832. [DOI] [PubMed] [Google Scholar]
  • 11.Greisen, K., M. Loeffelholz, A. Purohit, and D. Leong. 1994. PCR primers and probes for the 16S rRNA gene of most species of pathogenic bacteria, including bacteria found in cerebrospinal fluid. J. Clin. Microbiol. 32335-351. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 12.Harshey, R. M. 1994. Bees aren't the only ones: swarming in gram-negative bacteria. Mol. Microbiol. 13389-394. [DOI] [PubMed] [Google Scholar]
  • 13.Horng, Y. T., S. C. Deng, M. Daykin, P. C. Soo, J. R. Wei, K. T. Luh, S. W. Ho, S. Swift, H. C. Lai, and P. Williams. 2002. The LuxR family protein SpnR functions as a negative regulator of N-acylhomoserine lactone-dependent quorum sensing in Serratia marcescens. Mol. Microbiol. 451655-1671. [DOI] [PubMed] [Google Scholar]
  • 14.Kaiser, D. 2007. Bacterial swarming: a re-examination of cell-movement patterns. Curr. Biol. 17R561-R570. [DOI] [PubMed] [Google Scholar]
  • 15.Lai, H. C., P. C. Soo, J. R. Wei, W. C. Yi, S. J. Liaw, Y. T. Horng, S. M. Lin, S. W. Ho, S. Swift, and P. Williams. 2005. The RssAB two-component signal transduction system in Serratia marcescens regulates swarming motility and cell envelope architecture in response to exogenous saturated fatty acids. J. Bacteriol. 1873407-3414. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 16.Lehnen, D., C. Blumer, T. Polen, B. Wackwitz, V. F. Wendisch, and G. Unden. 2002. LrhA as a new transcriptional key regulator of flagella, motility and chemotaxis genes in Escherichia coli. Mol. Microbiol. 45521-532. [DOI] [PubMed] [Google Scholar]
  • 17.Li, C., C. J. Louise, W. Shi, and J. Adler. 1993. Adverse conditions which cause lack of flagella in Escherichia coli. J. Bacteriol. 1752229-2235. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 18.Liu, J. H., M. J. Lai, S. Ang, J. C. Shu, P. C. Soo, Y. T. Horng, W. C. Yi, H. C. Lai, K. T. Luh, S. W. Ho, and S. Swift. 2000. Role of flhDC in the expression of the nuclease gene nucA, cell division and flagellar synthesis in Serratia marcescens. J. Biomed. Sci. 7475-483. [DOI] [PubMed] [Google Scholar]
  • 19.Lloyd, A. L., B. J. Marshall, and B. J. Mee. 2005. Identifying cloned Helicobacter pylori promoters by primer extension using a FAM-labelled primer and GeneScan analysis. J. Microbiol. Methods 60291-298. [DOI] [PubMed] [Google Scholar]
  • 20.Park, D., and S. Forst. 2006. Co-regulation of motility, exoenzyme and antibiotic production by the EnvZ-OmpR-FlhDC-FliA pathway in Xenorhabdus nematophila. Mol. Microbiol. 611397-1412. [DOI] [PubMed] [Google Scholar]
  • 21.Romeo, T. 1998. Global regulation by the small RNA-binding protein CsrA and the non-coding RNA molecule CsrB. Mol. Microbiol. 291321-1330. [DOI] [PubMed] [Google Scholar]
  • 22.Shi, W., C. Li, C. J. Louise, and J. Adler. 1993. Mechanism of adverse conditions causing lack of flagella in Escherichia coli. J. Bacteriol. 1752236-2240. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 23.Shin, D., and E. A. Groisman. 2005. Signal-dependent binding of the response regulators PhoP and PmrA to their target promoters in vivo. J. Biol. Chem. 2804089-4094. [DOI] [PubMed] [Google Scholar]
  • 24.Shin, S., and C. Park. 1995. Modulation of flagellar expression in Escherichia coli by acetyl phosphate and the osmoregulator OmpR. J. Bacteriol. 1774696-4702. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 25.Silverman, M., and M. Simon. 1974. Characterization of Escherichia coli flagellar mutants that are insensitive to catabolite repression. J. Bacteriol. 1201196-1203. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 26.Soo, P. C., J. R. Wei, Y. T. Horng, S. C. Hsieh, S. W. Ho, and H. C. Lai. 2005. Characterization of the dapA-nlpB genetic locus involved in regulation of swarming motility, cell envelope architecture, hemolysin production, and cell attachment ability in Serratia marcescens. Infect. Immun. 736075-6084. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 27.Soutourina, O., A. Kolb, E. Krin, C. Laurent-Winter, S. Rimsky, A. Danchin, and P. Bertin. 1999. Multiple control of flagellum biosynthesis in Escherichia coli: role of H-NS protein and the cyclic AMP-catabolite activator protein complex in transcription of the flhDC master operon. J. Bacteriol. 1817500-7508. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 28.Soutourina, O. A., and P. N. Bertin. 2003. Regulation cascade of flagellar expression in Gram-negative bacteria. FEMS Microbiol. Rev. 27505-523. [DOI] [PubMed] [Google Scholar]
  • 29.Soutourina, O. A., E. Krin, C. Laurent-Winter, F. Hommais, A. Danchin, and P. N. Bertin. 2002. Regulation of bacterial motility in response to low pH in Escherichia coli: the role of H-NS protein. Microbiology 1481543-1551. [DOI] [PubMed] [Google Scholar]
  • 30.Sperandio, V., A. G. Torres, and J. B. Kaper. 2002. Quorum sensing Escherichia coli regulators B and C (QseBC): a novel two-component regulatory system involved in the regulation of flagella and motility by quorum sensing in E. coli. Mol. Microbiol. 43809-821. [DOI] [PubMed] [Google Scholar]
  • 31.Stock, A. M., V. L. Robinson, and P. N. Goudreau. 2000. Two-component signal transduction. Annu. Rev. Biochem. 69183-215. [DOI] [PubMed] [Google Scholar]
  • 32.Tomoyasu, T., T. Ohkishi, Y. Ukyo, A. Tokumitsu, A. Takaya, M. Suzuki, K. Sekiya, H. Matsui, K. Kutsukake, and T. Yamamoto. 2002. The ClpXP ATP-dependent protease regulates flagellum synthesis in Salmonella enterica serovar Typhimurium. J. Bacteriol. 184645-653. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 33.Van Houdt, R., M. Givskov, and C. W. Michiels. 2007. Quorum sensing in Serratia. FEMS Microbiol. Rev. 31407-424. [DOI] [PubMed] [Google Scholar]
  • 34.Wei, B. L., A. M. Brun-Zinkernagel, J. W. Simecka, B. M. Pruss, P. Babitzke, and T. Romeo. 2001. Positive regulation of motility and flhDC expression by the RNA-binding protein CsrA of Escherichia coli. Mol. Microbiol. 40245-256. [DOI] [PubMed] [Google Scholar]
  • 35.Wei, J. R., and H. C. Lai. 2006. N-Acylhomoserine lactone-dependent cell-to-cell communication and social behavior in the genus Serratia. Int. J. Med. Microbiol. 296117-124. [DOI] [PubMed] [Google Scholar]
  • 36.Wei, J. R., Y. H. Tsai, P. C. Soo, Y. T. Horng, S. C. Hsieh, S. W. Ho, and H. C. Lai. 2005. Biochemical characterization of RssA-RssB, a two-component signal transduction system regulating swarming behavior in Serratia marcescens. J. Bacteriol. 1875683-5690. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 37.West, A. H., and A. M. Stock. 2001. Histidine kinases and response regulator proteins in two-component signaling systems. Trends Biochem. Sci. 26369-376. [DOI] [PubMed] [Google Scholar]
  • 38.Yanagihara, S., S. Iyoda, K. Ohnishi, T. Iino, and K. Kutsukake. 1999. Structure and transcriptional control of the flagellar master operon of Salmonella typhimurium. Genes Genet. Syst. 74105-111. [DOI] [PubMed] [Google Scholar]
  • 39.Yindeeyoungyeon, W., and M. A. Schell. 2000. Footprinting with an automated capillary DNA sequencer. BioTechniques 291034-1041. [DOI] [PubMed] [Google Scholar]
  • 40.Yokota, T., and J. S. Gots. 1970. Requirement of adenosine 3′,5′-cyclic phosphate for flagella formation in Escherichia coli and Salmonella typhimurium. J. Bacteriol. 103513-516. [DOI] [PMC free article] [PubMed] [Google Scholar]

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