JB Try JVI Online
Home Help [Feedback] [For Subscribers] [Archive] [Search] [Contents]
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Other Versions of this Article:
JB.00712-06v1
189/1/131    most recent
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Services
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrow Copyright Information
Right arrow Books from ASM Press
Right arrow MicrobeWorld
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Léonard, S.
Right arrow Articles by Letesson, J.-J.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Léonard, S.
Right arrow Articles by Letesson, J.-J.
Journal of Bacteriology, January 2007, p. 131-141, Vol. 189, No. 1
0021-9193/07/$08.00+0     doi:10.1128/JB.00712-06
Copyright © 2007, American Society for Microbiology. All Rights Reserved.

FtcR Is a New Master Regulator of the Flagellar System of Brucella melitensis 16M with Homologs in Rhizobiaceae{triangledown}

S. Léonard,1 J. Ferooz,1 V. Haine,1 I. Danese,2 D. Fretin,2 A. Tibor,1 S. de Walque,3 X. De Bolle,1 and J.-J. Letesson1*

Unité de Recherche en Biologie Moléculaire, Laboratoire d'Immunologie-Microbiologie, Facultés Universitaires Notre-Dame de la Paix, Namur, Belgium,1 Centre d'Etude et de Recherches Vétérinaires et Agronomiques, Brussels, Belgium,2 Service de Chimie Biologique, Institut de Biologie et de Médecine Moléculaires, Université Libre de Bruxelles, Gosselies, Belgium3

Received 18 May 2006/ Accepted 5 October 2006


   ABSTRACT
Top
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 ADDENDUM IN PROOF
 REFERENCES
 
The flagellar regulon of Brucella melitensis 16M contains 31 genes clustered in three loci on the small chromosome. These genes encode a polar sheathed flagellum that is transiently expressed during vegetative growth and required for persistent infection in a mouse model. By following the expression of three flagellar genes (fliF, flgE, and fliC, corresponding to the MS ring, hook, and filament monomer, respectively), we identified a new regulator gene, ftcR (flagellar two-component regulator). Inactivation of ftcR led to a decrease in flagellar gene expression and to impaired Brucella virulence. FtcR has a two-component response regulator domain as well a DNA binding domain and is encoded in the first flagellar locus of B. melitensis. Both the ftcR sequence and its genomic context are conserved in other related {alpha}-proteobacteria. During vegetative growth in rich medium, ftcR expression showed a peak during the early exponential phase that paralleled fliF gene expression. VjbR, a quorum-sensing regulator of the LuxR family, was previously found to control fliF and flgE gene expression. Here, we provide some new elements suggesting that the effect of VjbR on these flagellar genes is mediated by FtcR. We found that ftcR expression is partially under the control of VjbR and that the expression in trans of ftcR in a vjbR mutant restored the production of the hook protein (FlgE). Finally, FtcR binds directly to the upstream region of the fliF gene. Therefore, our data support the role of FtcR as a flagellar master regulator in B. melitensis and perhaps in other related {alpha}-proteobacteria.


   INTRODUCTION
Top
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 ADDENDUM IN PROOF
 REFERENCES
 
Flagella are highly complex bacterial organelles that are usually well conserved among diverse bacterial species (17, 39, 46). In addition to motility, the bacterial flagellum is involved in a variety of interactions between the bacterium and its environment (e.g., adhesion, biofilm formation, secretion, and modulation of the host reponse) (7, 16, 20, 22, 30, 36, 42, 50, 53, 57). Flagellar expression involves about 50 flagellar genes, distributed into three or four classes as observed in Escherichia coli (1) and Salmonella enterica serovar Typhimurium (31) or in Caulobacter crescentus (38), Pseudomonas aeruginosa (10), and Vibrio cholerae (41). Complex regulation processes control this hierarchical system, allowing for the expression of the genes of one downstream class if the genes from the upstream class have been expressed.

At the top of the flagellar regulatory cascade, one or more distinct master genes encode the transcriptional regulators responsible for turning on and off flagellar synthesis, in response to environmental factors and cell cycle-related signals. In {gamma}-proteobacteria, this so-called class I is represented by a heterotetrameric complex, FlhD2C2 (lateral flagellar systems), or a sigma 54-associated transcriptional activator of the NtrC family (polar flagellar systems) (4, 52). The situation is somewhat different in {alpha}-proteobacteria. In C. crescentus, a two-component response regulator called CtrA controls a part of the asymmetric cell cycle as well as the initiation of the flagellar cascade (49), whereas in Sinorhizobium meliloti, two LuxR type regulators, VisR and VisN, are at the top of the flagellar hierarchy (51). Direct binding of VisR or VisN to a promoter of a class II gene (e.g., fliF) has never been demonstrated, and so, the involvement of other unidentified effectors acting in concert with VisN/VisR at the top of the hierarchy cannot be excluded (51).

Recent studies have shown that Brucella melitensis possesses 31 flagellar genes and builds, under strictly defined conditions, a polar sheathed flagellum required for virulence in a mouse infection model (14). The flagellar hierarchy has not been characterized in Brucella. A comparison of the three Brucella flagellar clusters with the unique cluster of structural flagellar genes from S. meliloti (15) revealed extensive gene synteny, namely, a high conservation in the order of genes. Like the visNR operon of S. meliloti, two regulator genes are located downstream and in the reverse orientation from a set of flagellar genes in Brucella. Nevertheless, only one of these two, called VjbR (11), belongs to the LuxR family. The second regulator is of the TetR family, and its function is presently unknown (11). VjbR was recently described as a quorum-sensing regulator that activates the expression of the virB secretion system and the flagellar apparatus of B. melitensis, both required for virulence in BALB/c mice. A {Delta}vjbR strain does not express fliF (coding for the MS ring monomer) and does not produce the FlgE protein (hook monomer). Moreover, an excess of the quorum-sensing pheromone N-dodecanoyl homoserine lactone represses FlgE production, probably by inhibiting VjbR activity (11). VjbR could constitute a second example of the LuxR-type master flagellar regulator as in S. meliloti, but its effect is known for only two flagellar components (FliF and FlgE). The direct or indirect nature of this control has not yet been determined (11).

Besides the TetR-type and the VjbR regulators, there is a third gene coding for a predicted transcriptional regulator within the Brucella flagellar loci, with synteny in S. meliloti. This gene was called ftcR for "flagellar two-component regulator" (14). The FtcR regulator is in fact predicted to belong to the two-component response regulator family. Because the function of Brucella melitensis FtcR and its homologs in other {alpha}-proteobacteria is unknown, we set out to elucidate its role in flagellar regulation. In this paper, we demonstrate that FtcR is required in B. melitensis 16M for the transcription of the fliF gene during vegetative and intracellular growth and for the production of the two structural flagellar components FlgE and FliC during vegetative growth. We also show that an ftcR mutant has the same virulence phenotype as previously found with structural flagellar mutants. Furthermore, we show that FtcR directly and specifically binds to the fliF upstream region. Finally, we establish that the expression of ftcR in trans on a plasmid can complement a {Delta}vjbR strain for FlgE production and that ftcR is partially activated by VjbR. These data suggest that FtcR could mediate the action of VjbR on flagellar gene expression.

Taken together, these data indicate that FtcR could constitute a master regulator of the Brucella flagellar regulon and raises the possibility that FtcR homologs in other {alpha}-proteobacteria may play the same role.


   MATERIALS AND METHODS
Top
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 ADDENDUM IN PROOF
 REFERENCES
 
Bacterial strains and culture conditions. All strains and plasmids used in this study are listed in Table 1 and oligonucleotides in Table 2. B. melitensis 16M strains were grown with shaking at 37°C in 2YT medium (1% yeast extract, 1.6% peptone, 0.5% NaCl) from an initial optical density at 600 nm (OD600) of 0.05. C. crescentus strains were grown at 30°C in PYE medium (40). E. coli strains were grown at 37°C in Luria-Bertani (LB) broth (44). Antibiotics were used at the following final concentrations: ampicillin, 100 µg/ml; chloramphenicol, 20 µg/ml; kanamycin, 30 µg/ml; nalidixic acid, 25 µg/ml; and nourseothricin, 50 µg/ml. DNA was transferred into B. melitensis or into C. crescentus using the E. coli S17-1 strain harboring the mobilization plasmid RP4. The E. coli strain DH10B was used for the propagation of the plasmids used in this study. BL21(DE3)pLysS was the host strain for the T7 expression system.


View this table:
[in this window]
[in a new window]

  		TABLE 1. Strains and plasmids used in this study

 

View this table:
[in this window]
[in a new window]

  		TABLE 2. Oligonucleotides used in this study

 
Construction of the ftcR mutant. The ftcR mutant was constructed by integrative disruption. An internal 0.315-kb fragment from nucleotide 16 to nucleotide 340 of the ftcR predicted coding sequence (pCDS) was amplified by PCR from genomic DNA of the B. melitensis 16M Nalr strain with the F2C2 and R2C2 primers and cloned into the EcoRV site of pSKKan (14) to generate pSKKanftcR. The pSKKanftcR plasmid was inserted into the ftcR pCDS of the B. melitensis 16M Nalr strain by homologous recombination. Transconjugants were selected by selecting for kanamycin resistance. Southern blot analysis was carried out on genomic DNA by using a probe that hybridized to the kanamycin resistance gene to confirm that one plasmid copy had stably integrated into the target gene, as described by Haine et al. (19).

Plasmid construction. Recombinant DNA techniques were carried out using standard protocols (44).

For construction of the reporter pftcR-lacZ fusion pBBCmpftcR-lacZ, a 0.437-kb fragment containing the ftcR promoter (pftcR) (including 24 bp of the 3' end of the upstream hypothetical pCDS, 327 bp of the noncoding region, and 68 bp of the predicted 5' end of ftcR) was PCR amplified from genomic DNA of the B. melitensis 16M Nalr strain with primers p2c2Xamont and p2c2Baval containing XbaI and BamHI sites, respectively. The PCR product was first subcloned into pGEM-T Easy vector (Promega) to generate pGEMTpftcR and then inserted into pBBCm-lacZ (14) in frame upstream of the promoterless lacZ reporter gene, generating pBBCmpftcR-lacZ.

The nourseothricin-resistant reporter fusion pBBNrspfliF-lacZ was obtained by cloning a 1.028-kb region containing the nourseothricin acetyltransferase gene (recovered from the pFA6a-natMX6 vector [Werner BioAgents] by FspI digestion) into the DraI site of the cat gene of pBBCmpfliF-lacZ (14).

The FtcR expression vectors pET15bftcR, pMRftcR, and pMRpftcRftcR were constructed using Gateway Technology (Invitrogen). Briefly, ftcR pCDS was amplified by PCR from B. melitensis 16M genomic DNA and tailed with attB1 and attB2 sequences using the 1038-attB1 and 1038-attB2 primers. The product was cloned by BP reaction (recombination reaction using BP clonase enzyme mix) into the pDONR201 vector (Invitrogen), generating the pDONRftcR entry vector. In the same way, a 1.132-kb fragment containing the ftcR pCDS and its upstream pftcR promoter region (pftcR-ftcR) was amplified by PCR from B. melitensis 16M genomic DNA and tailed with attB1 and attB2 sequences using the attB2-pbifR and attB1-bifR primers and then cloned by BP reaction into the pDONR201 plasmid, generating pDONRpftcR-ftcR. All pDONR constructs were verified by sequencing the insert. The ftcR pCDS was transferred by LR reaction (recombination reaction using LR clonase enzyme mix) from pDONRftcR into pMR10-GW under the control of the plac promoter, generating the pMRftcR plasmid, and into pET15b-GW under the control of the T7 promoter and in frame with an N-terminal histidine tag (His6) coding sequence, generating the pET15bftcR plasmid. The pftcR-ftcR insert was transferred from pDONRpftcR-ftcR into pMR10-GW in an orientation opposite of that for the plac promoter, generating pMRpftcR-ftcR.

ß-Galactosidase assays. ß-Galactosidase assays were performed in line with the protocol of Miller (34) as described by Fretin et al. (14).

SDS-PAGE and immunoblotting. Sodium dodecyl sulfate (SDS)-polyacrylamide gel electrophoresis (PAGE) and immunoblotting were performed as previously described (11, 14). Briefly, B. melitensis cultures were grown for various times until they reached the appropriate phase of growth (an OD600 of 0.1 for the end of latent phase and an OD600 of 0.25 for the beginning of exponential phase). Total bacterial cell samples were pelleted and resuspended in SDS sample buffer. The proteins were resolved on a 12% polyacrylamide gel and transferred to Hybond ECL nitrocellulose membranes (Amersham). Immunodetection of proteins was performed using anti-FliC or anti-FlgE polyclonal antibody (14) and an anti-Omp1 monoclonal antibody, A5310B2 (5), as a loading control. The detection of antibodies was performed using donkey anti-rabbit (Amersham) and goat anti-mouse (Dako) horseradish peroxidase-conjugated secondary antibodies as appropriate, and proteins were visualized using the ECL system (Amersham).

Virulence assays. Evaluation of the intracellular survival of the B. melitensis 16M wild-type strain and the ftcR mutant in cellular models (HeLa cells and bovine macrophages) was performed as previously described by Delrue et al. (12). Briefly, the strains were grown at 37°C in 2YT medium and then inoculated at a multiplicity of infection of 300 to subconfluent monolayers of HeLa cells or bovine macrophages in 96-well tissue culture plates. After a 10-min centrifugation at 1,000 rpm at room temperature in a Jouan centrifuge, the preparations were placed in a 5% CO2 atmosphere at 37°C for 1 h. The wells were then washed, and 50 µg/ml gentamicin was added to the cells until the end of the infection time (48 h). Cell monolayers were washed and lysed with 0.1% Triton X-100 in phosphate-buffered saline (PBS) for 20 min, and the lysates were serially diluted and plated onto 2YT plates to count viable bacteria.

Virulence assays with the BALB/c mouse model were performed as described by Fretin et al. (14). Briefly, 8-week-old female mice were inoculated intraperitoneally with a PBS suspension containing approximately 104 CFU of the appropriate bacterial strain. At different intervals after inoculation (1, 4, 8, and 12 weeks), four mice from each treatment group were sacrificed for spleen collection. Each spleen was homogenized in 2 ml of distilled water, and serial dilutions of it were plated on 2YT agar to determine bacterial survival. Data were expressed as the logarithm (log) of the number of CFU per spleen.

Immunofluorescence assays. Bovine macrophages infected with the B. melitensis 16M wild-type strain and the ftcR mutant harboring the pBBpfliF-gfp plasmid (14) were fixed at various times after infection. Bacteria were detected using primary anti-Brucella lipopolysaccharide O-chain monoclonal antibody 12G12 (6) and secondary Alexa 568-conjugated anti-mouse immunoglobulin G antibodies (Molecular Probes) as previously described (14).

Expression and purification of recombinant FtcR. FtcR was overproduced under 1 mM isopropyl-ß-D-thiogalactopyranoside induction in E. coli BL21(DE3)plysS (Novagen) from the pET15bftcR plasmid as a fusion protein containing an N-terminal six-histidine tag. Protein purification was performed on a 2.5-ml nickel chelation resin column (His-Bind; Novagen) with elution by 6 M urea as recommended by the manufacturer, followed by renaturation of the protein by dialysis using 1x PBS and 0.03 M Tris, pH 7.9. The protein was concentrated to 0.3 mg/ml using a 15-ml Amicon Ultra centrifugal filter unit (Millipore) and finally stored at –20°C in 50% glycerol.

Electrophoretic mobility shift assay (EMSA). Four 40-bp DNA fragments (synthesized and PAGE purified by Sigma-Genosys) corresponding to the upstream region of fliF pCDS (probes 1, 2, 3, and 4) (see Fig. 4) were 5' end radiolabeled using T4 polynucleotide kinase (Biolabs) and [{gamma}-32P]ATP (Amersham). The promoter probes were purified with Sephadex G50 micro columns (ProbeQuant G-50 micro columns; Amersham) and then by QIAEX II gel extraction (QIAGEN) after separation on a 12% polyacrylamide gel. The protein-DNA binding reactions were carried out as follows. Various amounts of recombinant FtcR (from 0 to 200 ng) were first incubated for 10 min at room temperature in the absence of a probe in a 17-µl reaction mixture containing 1 µl dithiothreitol (20 mM), 1 µl bovine serum albumin (10 mg/ml), and 4 µl 5x binding buffer (100 mM Tris-HCl, pH 7.5, 5 mM MgCl2, 0.5 mM EDTA, 50% glycerol). A total of {approx}30,000 cpm of probe was then added to the mixture to a final volume of 20 µl with or without a molar excess of an unlabeled competitor probe, and the mixture was incubated for 20 min at room temperature. Samples were then separated on a 6% nondenaturing polyacrylamide gel in 1x TGE buffer (25 mM Tris-acetate, pH 8.3, 190 mM glycine, 1 mM EDTA). To determine the specificity of the binding of recombinant FtcR to the pfliF probes, unlabeled probe 1 (data not shown) or 2 or heterologous DNA containing a mammalian Oct binding site (CV1048, 5'-TGTCGAATGCAAATCACTAGAA-3') was included in the competition assay.


Figure 4
View larger version (48K):
[in this window]
[in a new window]

  		FIG. 4. Analysis of FtcR binding on the fliF promoter. (A) Representation of the ~100-nucleotide genomic region containing the fliF promoter. The ATG of the fliF pCDS is in bold. The four labeled pfliF fragments used for the EMSA experiments are indicated (probes 1, 2, 3, and 4). //, predicted transcriptional terminator for the preceding fliC gene. (B) Binding of FtcR to pfliF. Four different end-labeled probes corresponding to fragments of the pfliF genomic region (see panel A) were incubated with increasing quantities of recombinant FtcR for 20 min at room temperature and then subjected to electrophoresis in a nondenaturating polyacrylamide gel. The FtcR quantities used were 0 ng (lanes 1, 5, and 9), 50 ng (lanes 2, 6, and 10), 100 ng (lanes 3, 7, and 11), and 200 ng (lanes 4, 8, and 12) (data for probe 3 not shown). (C) Competitive binding of FtcR to probe 2. End-labeled probe 2 was mixed with unlabeled probe (competitive cold probe) 1 or 2, or a nonspecific (NS) probe and incubated with 200 ng of recombinant FtcR. Samples were then subjected to an electrophoretic mobility shift assay. A competitive cold probe was used in excess of 2.5 ng (lanes 3, 7, and 11), 5 ng (lanes 4, 8, and 12), 15 ng (lanes 5, 9, and 13), and 45 ng (lanes 6, 10, and 14). As controls, labeled probe 2 was tested alone with FtcR (lane 2) or without FtcR (lane 1) and labeled aspecific probe alone was tested with FtcR (lane 16) or without FtcR (lane 15).

 
Statistical analysis. Data were analyzed by one-way analysis of variance after testing of the homogeneity of variance. Post hoc comparisons were performed by a pair-wise Scheffé test.


   RESULTS
Top
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 ADDENDUM IN PROOF
 REFERENCES
 
The ftcR gene encodes a predicted two-component response regulator conserved in Rhizobiaceae. BMEII0158 on the B. melitensis 16M small chromosome corresponds to a pCDS of unknown function. This sequence is located downstream from a predicted operon encoding the MS ring monomer (FliF), three motor proteins (MotB, MotC, and MotD), and two hypothetical proteins and upstream of the flgE pCDS encoding the hook monomer (Fig. 1A) (14). The deduced 227-residue polypeptide sequence (25.7 kDa) of FtcR is well conserved in other members of the {alpha}-subgroup of proteobacteria (Fig. 1B). In Sinorhizobium meliloti, Agrobacterium tumefaciens, and Mesorhizobium loti, the corresponding pCDSs exhibit 64%, 62%, and 58% identity with FtcR, respectively, at the amino acid level. The ftcR genomic context is also conserved in several {alpha}-proteobacteria (Fig. 1A). The deduced sequence contains two Pfam domains (PF00486, Trans_reg C; and PF00072, Response reg) which are characteristic of regulators belonging to two-component response regulators (55). This prompted us to rename BMEII0158 "ftcR" for "flagellar two-component regulator." However, FtcR is atypical because it lacks the aspartate residue usually conserved as a phosphorylation site in the receiver domain (Fig. 1B). Actually, in Brucella spp. and in FtcR homologs in S. meliloti and A. tumefaciens, the aspartate residue is replaced at position 50 by a glutamate residue, while in the FtcR homolog of M. loti, it is replaced by an arginine residue. In addition, an inventory of all genes predicted to encode histidine kinases and response regulators in the B. melitensis genome did not reveal the presence of a cognate sensor partner (putative histidine protein kinase) for FtcR (29).


Figure 1
View larger version (80K):
[in this window]
[in a new window]

  		FIG. 1. Analysis of the FtcR sequence. (A) Synteny between B. melitensis 16M, A. tumefaciens, M. loti, and S. meliloti 1021 in the genomic region containing the ftcR CDS. The ftcR CDS is indicated by a white arrow, as is the homologous CDS encoding a predicted response regulator from a two-component system (ftcR like). The names of the CDSs are written in the arrows. The length in amino acid residues of each CDS is indicated under the arrows. The CDS regions conserved without predicted function (hypothetical conserved) are labeled with a question mark. This genomic region is conserved in Brucella suis and Brucella abortus. fliK*, fliK was previously called motD. (B) CLUSTALW multiple alignment of the deduced amino acid sequences of the pCDSs located upstream of the flgE gene in B. melitensis (Bme; AAL53399.1), S. meliloti (Sme; AAG48153.1), A. tumefaciens (Atu; NP_353599.1), and M. loti (Mlo; NP_104158.1) (GenBank accession numbers are in parentheses). Residues marked with an asterisk and shaded black correspond to identical residues. Residues marked with points belong to a conserved group of amino acids defined as weak (one point) or strong (two points) by CLUSTALW 1.83. The frames indicate the response regulator domain (continuous line) and the transcriptional regulatory domain (discontinuous line) as recognized in Pfam. The white inverted triangle indicates the classical position of the conserved aspartate in the phosphorylation site replaced by another residue in B. melitensis, S. meliloti, A. tumefaciens (E), and M. loti (R).

 
Inactivation of the ftcR gene downregulates the activity of the flagellar fliF promoter region and the production of both the FlgE and FliC flagellar proteins during vegetative growth. In order to test whether FtcR plays a role in the regulation of flagellar gene expression, we generated an ftcR mutant by integrative disruption as described in Materials and Methods. We knew from a previous study that the noncoding region just upstream of fliF (pfliF), a gene encoding the flagellar MS ring monomer, is transiently active during vegetative growth of the wild-type (WT) B. melitensis 16M strain in 2YT medium. In fact, the pfliF activity is maximal at the very beginning of the exponential growth phase, and concomitantly, there is transient production of the FlgE and FliC proteins, which correspond to monomers of the flagellar hook and filament, respectively (14). To investigate the contribution of FtcR to flagellar expression during vegetative growth in rich medium, we used a pfliF-lacZ translational fusion (plasmid pBBCmpfliF-lacZ) to measure pfliF activity in the ftcR mutant compared to that in the WT strain. A 20- to 160-fold reduction of pfliF activity was observed, depending on the growth stage of pfliF in the mutant strain (Fig. 2A). The same experiment with a {Delta}vjbR mutant strain showed a 5- to 40-fold reduction of pfliF activity as previously described (Fig. 2A) (11). In the same way, FlgE and FliC protein levels were analyzed in WT and ftcR mutant strains by Western blot analysis. We used a {Delta}vjbR mutant strain harvested at the same time as a negative control of flagellar protein expression (11). The Omp1 protein detected by a monoclonal antibody was used as a loading control (Fig. 3).


Figure 2
View larger version (24K):
[in this window]
[in a new window]

  		FIG. 2. Activity of the fliF promoter region in different Brucella strains. (A) Activity of the fliF promoter region in the wild-type strain (black squares), the ftcR mutant strain (black circles), and the vjbR mutant strain (black triangles). A pfliF-lacZ fusion plasmid was transmitted by conjugal transfer into the three strains, each was cultivated in 2YT nutrient broth, and samples were recovered at intervals during bacterial growth to test the pfliF activity using ß-galactosidase assays. The ß-galactosidase activity is expressed as the mean ± standard deviation from three replicates. White squares, circles, and triangles correspond to the optical density at 600 nm for the wild-type strain, the ftcR mutant, and the vjbR mutant, respectively. (B) Same results as those in panel A shown with a larger scale for the abscissa axis to facilitate the distinction between results for the ftcR and vjbR mutants. Only the activity of the fliF promoter in the ftcR mutant strain and the vjbR mutant strain are visible.

 

Figure 3
View larger version (25K):
[in this window]
[in a new window]

  		FIG. 3. Western blot of FlgE and FliC expression in the ftcR mutant strain, the vjbR mutant strain, and the wild-type strain. The strains were cultivated in 2YT nutrient broth, and extracts were prepared from samples harvested at the end of the latent phase of growth (OD600 of 0.1) (lanes 1) and at the beginning of the exponential phase of growth (OD600 of 0.25) (lanes 2). The extracts were separated by electrophoresis, transferred to nylon membranes, and probed with FlgE-specific (A) or FliC-specific (B) antiserum. A monoclonal anti-Omp1 antibody was used as a loading control. CftcR, mutant ftcR containing a complementation pMRpftcR-ftcR plasmid.

 
A clear increase in the production of the hook and the flagellin proteins can be seen in the WT extracts between the end of the latent phase of growth and the beginning of the exponential phase. However, almost no FlgE or FliC production was detected in either the ftcR mutant or the {Delta}vjbR mutant strain (Fig. 3). The expression in trans of the ftcR pCDS (using the plasmid pMRpftcR-ftcR) in the ftcR mutant restores the production of FlgE, indicating that the mutation has no polar effect on downstream genes (Fig. 3A). Taken together, these data show that FtcR is required for the production of at least three flagellar components that are found in distinct locations in the flagellar structure (base, hook, and filament). These results strongly point to FtcR as an important regulatory protein for flagellar gene expression, at least during vegetative growth.

FtcR also regulates the fliF promoter region intracellularly. Using a pfliF-gfp fusion, it was recently demonstrated that pfliF is active during macrophage infection (14). We transferred this pfliF-gfp reporter system (pBBpfliF-gfp) by conjugation to the ftcR mutant and used this strain to infect a monolayer of bovine macrophages. Postinfection observations at different times contrasted with those for the parent strain, with the ftcR mutant failing to exhibit pfliF activity (data not shown). This result confirms that FtcR is also a transcriptional activator of pfliF ex vivo.

The ftcR mutant has the same phenotype as structural flagellar mutants in both cellular and murine models of infection. The flagellum of B. melitensis is involved in persistence in a murine model of infection, whereas there was no demonstrable need of this appendage during the course of cellular infection (14). Because FtcR positively regulates the production of at least three flagellar components related to distinct locations at the flagellar structural level (see above), it is probable that the absence of functional FtcR leads to the absence of a flagellar structure. Accordingly, an ftcR mutant should be attenuated as much as the strains mutated in genes encoding flagellar structural components. To test this hypothesis, cellular and murine infections were carried out with the ftcR mutant (see Materials and Methods). In HeLa cells and bovine macrophages, we observed no attenuation of the ftcR mutant compared to that in the WT parental strain (data not shown). In BALB/c mice, the ftcR mutant was not attenuated after 1 week of infection but was attenuated after 4 weeks of infection (Table 3). These results are in agreement with data obtained with Brucella strains mutated in genes encoding various structural flagellar components (14).


View this table:
[in this window]
[in a new window]

  		TABLE 3. Virulence of Brucella melitensis wild-type and flagellar mutant strains in BALB/c mice

 
All of the above data strengthen the evidence that FtcR plays a major role in flagellar gene regulation.

FtcR is a direct activator of the fliF gene. Like Brucella, Caulobacter crescentus belongs to the {alpha}-subgroup of proteobacteria, but no predicted ortholog of the ftcR gene is found in its genome. This fact allowed us to exploit the natural cellular context of C. crescentus to study the potential direct transcriptional effect of FtcR on pfliF, with both partners being imported from Brucella.

The strain NA1000 of C. crescentus (Table 1) was first conjugated with a plasmid bearing a pfliF-lacZ fusion (pBBNrspfliF-lacZ) and then with a second plasmid which either had no insert as a negative control (pMR10CmKm) or harbored ftcR under the control of a constitutive promoter (pMRplac-ftcR). ß-Galactosidase assays were performed, and the averages from three independent experiments ± standard errors were calculated to determine pfliF activity in the two strains. In the absence of ftcR in trans, 67.2 ± 3.0 Miller units was obtained. In contrast, 620.9 ± 12.8 Miller units was calculated in the presence of ftcR. These data correspond to a ninefold induction of pfliF, in comparison to what was seen with the negative control, which suggests that FtcR might interact directly with the fliF promoter.

This hypothetical interaction was tested using the EMSA technique. In the first instance, increasing amounts of FtcR recombinant protein were mixed with four labeled pfliF fragments that were 40 bp in length (designated probes 1, 2, 3, and 4). The probes overlap each other by half of their length and altogether cover a genomic region of 100 nucleotides localized between a predicted transcriptional Rho-independent terminator for the preceding fliC gene and the beginning of the fliF pCDS (Fig. 4A). Following native gel electrophoresis, a shifted DNA band of increasing intensity appeared when probe 1 or 2 was incubated with increasing amounts of the FtcR protein (Fig. 4B). No clear binding was observed using probe 3 (data not shown) or 4 (Fig. 4B). The specificity of the binding assay was evaluated in a second experiment. A nonspecific DNA probe caused no mobility shift and did not, when unlabeled, compete with labeled probe 1 or 2 for FtcR binding. On the contrary, both cold probes 1 and 2 were able to compete with labeled probe 2 (Fig. 4C) and probe 1 (data not shown) for FtcR binding. This indicates that FtcR binds directly and specifically to the pfliF promoter region.

FtcR mediates the activator effect of VjbR on flagellar genes. VjbR was demonstrated to be the Brucella functional counterpart of the S. meliloti flagellar master regulator VisR (51), and it is also encoded at the same genetic locus. Notably, the expression levels of both fliF and flgE were strongly reduced in a vjbR mutant (11). A similar result for FliC production was also observed in this study (Fig. 3B). Because this phenotype was globally shared with the ftcR mutant described here, we wanted to estimate the relative contribution of FtcR and VjbR with respect to flagellar expression.

As demonstrated, in this study pfliF activity was almost abolished in both the vjbR and the ftcR mutants compared to that in the WT strain but was two- to fourfold higher in the vjbR mutant than in the ftcR mutant (Fig. 2B). Thus, the absence of FtcR seems to be more deleterious than the absence of VjbR regarding the activation of pfliF.

To gain further insight into the respective roles of these two regulators in flagellar expression, we tested the ability of VjbR or FtcR to cross-complement an ftcR or a vjbR mutant, respectively. To achieve this, an expression plasmid of FtcR (pMRpftcR-ftcR) or VjbR (pBBR-vjbR) (11) or plasmids without inserts as negative controls (pMR10CmKm and pBBR1MCS-4) were conjugated in the vjbR mutant and in the ftcR mutant. FlgE protein levels were analyzed in the resulting strains by Western blot analysis. Both the ftcR mutant and the vjbR mutant produced little or no detectable FlgE with the plasmids used as negative controls, but both were complemented by the expression in trans of the corresponding WT gene (positive control). Importantly, while the expression of vjbR in trans failed to restore FlgE production in the ftcR mutant, the expression of ftcR in trans was able to complement the vjbR mutant (Fig. 5). This strongly indicates that the absence of flagellar gene expression in the vjbR mutant is actually caused by a reduction in the amount of active FtcR.


Figure 5
View larger version (15K):
[in this window]
[in a new window]

  		FIG. 5. Expression of FlgE in the vjbR and ftcR mutants containing expression plasmid vjbR pBBR-vjbR (vjbR), expression plasmid FtcR pMRpftcR-ftcR (ftcR), or the corresponding empty plasmids pBBR1MCS-4 and pMR10CmKm(–). All strains were cultivated in 2YT broth, and whole-cell extracts were recovered for each strain at the end of the latent phase of growth (OD600 of 0.1) (lanes 1) and at the beginning of the exponential phase of growth (OD600 of 0.25) (lanes 2). Recombinant FlgE was used as a positive control (lane 3). The extracts were separated by electrophoresis, transferred to nylon membranes, and probed with FlgE-specific antiserum. A monoclonal anti-Omp1 antibody was used as a loading control.

 
In this context, VjbR could be an activator of ftcR gene expression. To test this hypothesis, we used a reporter plasmid (pBBCmpftcR-lacZ) to compare pftcR activity between the vjbR mutant and the WT strain of B. melitensis 16M along a growth curve. In the WT strain, pftcR activity reached a maximum value at the beginning of the exponential phase and then decreased slowly. The same profile was observed in the vjbR mutant, but pftcR activity was reduced by about half compared to that in the WT strain (Fig. 6). This observation confirms that the expression of ftcR is affected (twofold) by the loss of VjbR.


Figure 6
View larger version (15K):
[in this window]
[in a new window]

  		FIG. 6. Activity of the ftcR promoter in the vjbR mutant (white) and in wild-type B. melitensis 16M (black). A pftcR-lacZ fusion plasmid was conjugated into the vjbR mutant and into the wild-type strain. Both strains were cultivated in 2YT nutrient broth, and samples were harvested along the growth curve to test pftcR activity by ß-galactosidase assays. Circles, optical density at 600 nm; squares, ß-galactosidase activity expressed as the mean ± standard deviation from three replicates.

 

   DISCUSSION
Top
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
ADDENDUM IN PROOF
 REFERENCES
 
The present work focused on B. melitensis, describing for the first time a transcriptional regulator called FtcR that directly activates flagellar expression. FtcR is conserved in several symbiotic and pathogenic {alpha}-proteobacterial Rhizobiales. A B. melitensis ftcR mutant is unable to produce the FlgE and FliC flagellar proteins and has almost no pfliF activity during vegetative growth or during macrophage infection compared to the parental WT strain. Moreover, observations of this mutant by electron microscopy did not reveal any flagellar structures under growth conditions in which the WT strain was flagellated (14; data not shown). In addition, the ftcR mutant is not attenuated in cellular infection models but exhibited a marked virulence defect after 4 weeks or more of infection in BALB/c mice. These virulence phenotypes were reminiscent of the phenotypes observed for strains mutated in genes encoding various flagellar components (14). Taken together, these results suggest the total absence of flagella in the ftcR mutant and point to FtcR as a key regulator in switching on the expression of flagella in Brucella.

This study also shows that FtcR binds directly to the region just upstream of the fliF pCDS. This region is localized between the transcriptional Rho-independent terminator for the preceding fliC pCDS and the beginning of the fliF pCDS and so is predicted as a noncoding region. In addition, cloning this sequence upstream of a reporter gene such as gfp or lacZ provides an expression profile which is modified in response to ftcR or vjbR mutation in the same way as other flagellar components. All of this evidence suggests that the fliF promoter (pfliF) is located in this genomic region between the fliC and the fliF pCDS. In the EMSA, a clear positive and specific signal was observed for probes 1 and 2 (Fig. 4). No signal was observed with probes 3 and 4, suggesting that the FtcR binding site is located in the 20-nucleotide-sized region common to probes 1 and 2. Additional experiments should be carried out to be able to identify and characterize the FtcR binding sequence relative to the transcriptional start of the fliF gene.

It has been noted that in all known flagellar hierarchies, the fliF gene is considered to be a class II gene, encoding the monomer for the MS ring formation, which is one of the first steps of a classical flagellar assembly. A hierarchical expression of flagellar genes has not yet been demonstrated in Brucella, although this is the most probable situation as demonstrated for all other bacterial flagellar systems (except rare spirochetes [3]) investigated to date. The binding of FtcR to pfliF, along with several lines of evidence for the control of flagellar gene expression by this regulator, suggests that FtcR could be considered a flagellar class I master regulator (32, 52).

Analysis of the FtcR-predicted amino acid sequence revealed two domains in the protein: a response regulator receiver domain at the N terminus and a C-terminal transcriptional regulatory domain that classifies FtcR as a response regulator belonging to the two-component system (55). The closest homologs of FtcR (more than 30% identity) found in the NCBI nonredundant database are exclusively from Rhizobiaceae and are most often encoded by genes located in flagellar loci as in Brucella. A unique feature of the FtcR regulator is the absence of a classical phosphorylation site in the response regulator domain. Interestingly, upon multiple alignment (CLUSTALW) of FtcR and its closest homologs with the well-known response regulators CheY and NtrC, the aspartate (D57 in CheY and D54 in NtrC), which has been shown to be the phosphorylated residue of the active site, aligned with the glutamate E50 residue of FtcR (45, 56). The replacement of the classical aspartate residue by a glutamate residue is also observed in A. tumefaciens and S. meliloti. It has to be noted that the molecular structure of glutamate is very similar to that of aspartate, and like aspartate, glutamate is negatively charged. The E50 glutamate residue in FtcR and its homologs could thus mimic constitutive phosphorylation. Indeed, it has been shown for other transcriptional regulators, such as NtrC (25, 37), CtrA (47), OmpR (28), and RcsB (18), that the experimental replacement of aspartate by a glutamate residue in the phosphorylation active site allows these regulators to bypass, at least partially, the requirement for phosphorylation.

Apart from FtcR and its closest homologs, some other rare bacterial regulators have been discovered which lack the conserved aspartate residue in the phosphorylation site. Examples include FleQ of Pseudomonas aeruginosa (9) and Legionella pneumophila (21), AdnA of Pseudomonas fluorescens (2), FlrA of Vibrio cholerae and Vibrio fischeri (24, 35), and FlaK and LafK of Vibrio parahaemolyticus (23, 54). All of these belong to the {sigma}54-dependent response regulator family, and all are at the top of a flagellar hierarchy without known sensor histidine kinase partners. The absence of a sensor histidine kinase encoded in Brucella flagellar loci argues in favor of the hypothesis that FtcR does not need phosphorylation to be active. In fact, genes of two-component systems are often coupled in genomes, as demonstrated in a study by Koretke et al. (26). The hypothesis that FtcR is nonphosphorylated and constitutively active raises the question of how the activity of FtcR is modulated. In P. aeruginosa, nonphosphorylated FleQ is regulated at the transcriptional level by the Vfr repressor and its activity is repressed posttranslationally upon binding with the FleN antiactivator (8, 9). The first regulator of FtcR activity in Brucella, VjbR, was identified in the present study.

The LuxR-type transcriptional regulator VjbR, which also activates the type IV secretion system, caught our attention principally because it was shown to positively control the activity of the pfliF promoter (11), FlgE production (11), and FliC production (this study) in B. melitensis. This phenotype, also observed for the ftcR mutant, suggests the involvement of FtcR and VjbR in the same flagellar regulatory pathway, making it probable that one of them modulates the expression or the activity of the other to control flagellar expression. In this study, we observed a more attenuated pfliF activity in the ftcR mutant than in the vjbR mutant. In addition, FlgE production was shown to be restored in the vjbR mutant when ftcR was expressed in trans under the control of its own promoter, while the expression in trans of vjbR did not restore FlgE production in the ftcR mutant. These results suggest a more direct regulation of flagellar genes by FtcR. This led us to investigate the influence of VjbR on pftcR activity in Brucella. While the expression of flagellar genes is almost extinguished in a vjbR mutant, the expression of ftcR is only twofold reduced. This mild activation of pftcR by VjbR could be direct or indirect. In addition, it should also be noted that our data do not exclude a mild activating effect from VjbR directly on pfliF, leaving the possibility of a three-gene feed-forward loop motif, including FtcR. This motif has frequently been observed in regulation networks and was defined by Mangan and Alon as a three-gene pattern composed of two input transcription factors, one of which regulates the other and both of which jointly regulate a target gene (33). That way, a regulation model in which vjbR activates ftcR, which in turn activates fliF and other flagellar genes, could be proposed, without excluding the possibility of a more direct VjbR effect on flagellar genes. More experiments will have to be carried to clarify the situation.

Finally, a comparison of pfliF and pftcR activities, both under the influence of VjbR, reveals a more abrupt decrease of pfliF activity along the growth curve (compare Fig. 2A and 6). This observation, in addition to the partial maintenance of pftcR activity in a vjbR mutant, suggests the existence of yet-to-be-identified regulators acting beside VjbR. Extra efforts will have to be made to gain further insight into the regulation of the flagellar expression in Brucella. Flagellar expression is a complex process that must be precisely tied to the intricacies of the bacterial life cycle. The discovery of the key flagellar regulator, FtcR, is a first step in leading to a better understanding of Brucella and, by extension, other {alpha}-proteobacteria.


   ADDENDUM IN PROOF
Top
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 ADDENDUM IN PROOF
REFERENCES
 
Rotter et al. recently published the description of the ftcR homologue in Sinorhizobium meliloti (C. Rotter, S. Mühlbacher, D. Salamon, R. Schmitt, and B. Scharf, J. Bacteriol. 188:6932-6942, 2006).


   ACKNOWLEDGMENTS
 
We thank C. Didembourg for technical assistance. We are grateful to the Brucella team of the URBM for helpful discussions.

This work was supported by the Commission of the European Communities, contract no. QLK2-CT-1999-00014. V. Haine, S. Léonard, and J. Ferooz hold a specialization grant from the Fonds pour la Formation à la Recherche dans l'Industrie et dans l'Agriculture (FRIA).


   FOOTNOTES
 
* Corresponding author. Mailing address: Laboratoire d'Immunologie-Microbiologie, Unité de Recherche en Biologie Moléculaire (URBM), Facultés Universitaires Notre-Dame de la Paix, rue de Bruxelles 61, 5000 Namur, Belgium. Phone: (32) 81 72 44 02. Fax: (32) 81 72 44 20. E-mail: jean-jacques.letesson{at}fundp.ac.be. Back

{triangledown} Published ahead of print on 20 October 2006. Back


   REFERENCES
Top
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 ADDENDUM IN PROOF
 REFERENCES
 

  1. Aldridge, P., and K. T. Hughes. 2002. Regulation of flagellar assembly. Curr. Opin. Microbiol. 5:160-165.[CrossRef][Medline]
  2. Casaz, P., A. Happel, J. Keithan, D. L. Read, S. R. Strain, and S. B. Levy. 2001. The Pseudomonas fluorescens transcription activator AdnA is required for adhesion and motility. Microbiology 147:355-361.[Abstract/Free Full Text]
  3. Charon, N. W., and S. F. Goldstein. 2002. Genetics of motility and chemotaxis of a fascinating group of bacteria: the spirochetes. Annu. Rev. Genet. 36:47-73.[CrossRef][Medline]
  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. 64:694-708.[Abstract/Free Full Text]
  5. Cloeckaert, A., P. de Wergifosse, G. Dubray, and J. N. Limet. 1990. Identification of seven surface-exposed Brucella outer membrane proteins by use of monoclonal antibodies: immunogold labeling for electron microscopy and enzyme-linked immunosorbent assay. Infect. Immun. 58:3980-3987.[Abstract/Free Full Text]
  6. Cloeckaert, A., M. S. Zygmunt, G. Dubray, and J. N. Limet. 1993. Characterization of O-polysaccharide specific monoclonal antibodies derived from mice infected with the rough Brucella melitensis strain B115. J. Gen. Microbiol. 139:1551-1556.[Medline]
  7. Daniels, R., J. Vanderleyden, and J. Michiels. 2004. Quorum sensing and swarming migration in bacteria. FEMS Microbiol. Rev. 28:261-289.[CrossRef][Medline]
  8. Dasgupta, N., E. P. Ferrell, K. J. Kanack, S. E. H. West, and R. Ramphal. 2002. fleQ, the gene encoding the major flagellar regulator of Pseudomonas aeruginosa, is {sigma}70 dependent and is downregulated by Vfr, a homolog of Escherichia coli cyclic AMP receptor protein. J. Bacteriol. 184:5240-5250.[Abstract/Free Full Text]
  9. Dasgupta, N., and R. Ramphal. 2001. Interaction of the antiactivator FleN with the transcriptional activator FleQ regulates flagellar number in Pseudomonas aeruginosa. J. Bacteriol. 183:6636-6644.[Abstract/Free Full Text]
  10. Dasgupta, N., M. C. Wolfgang, A. L. Goodman, S. K. Arora, J. Jyot, S. Lory, and R. Ramphal. 2003. A four-tiered transcriptional regulatory circuit controls flagellar biogenesis in Pseudomonas aeruginosa. Mol. Microbiol. 50:809-824.[CrossRef][Medline]
  11. Delrue, R. M., C. Deschamps, S. Leonard, C. Nijskens, I. Danese, J. M. Schaus, S. Bonnot, J. Ferooz, A. Tibor, X. De Bolle, and J. J. Letesson. 2005. A quorum-sensing regulator controls expression of both the type IV secretion system and the flagellar apparatus of Brucella melitensis. Cell. Microbiol. 7:1151-1161.[CrossRef][Medline]
  12. Delrue, R. M., M. Martinez-Lorenzo, P. Lestrate, I. Danese, V. Bielarz, P. Mertens, X. De Bolle, A. Tibor, J. P. Gorvel, and J. J. Letesson. 2001. Identification of Brucella spp. genes involved in intracellular trafficking. Cell. Microbiol. 3:487-497.[CrossRef][Medline]
  13. Evinger, M., and N. Agabian. 1977. Envelope-associated nucleoid from Caulobacter crescentus stalked and swarmer cells. J. Bacteriol. 132:294-301.[Abstract/Free Full Text]
  14. Fretin, D., A. Fauconnier, S. Kohler, S. Halling, S. Leonard, C. Nijskens, J. Ferooz, P. Lestrate, R. M. Delrue, I. Danese, J. Vandenhaute, A. Tibor, X. DeBolle, and J. J. Letesson. 2005. The sheathed flagellum of Brucella melitensis is involved in persistence in a murine model of infection. Cell. Microbiol. 7:687-698.[CrossRef][Medline]
  15. Galibert, F., T. M. Finan, S. R. Long, A. Puhler, P. Abola, F. Ampe, F. Barloy-Hubler, M. J. Barnett, A. Becker, P. Boistard, G. Bothe, M. Boutry, L. Bowser, J. Buhrmester, E. Cadieu, D. Capela, P. Chain, A. Cowie, R. W. Davis, S. Dreano, N. A. Federspiel, R. F. Fisher, S. Gloux, T. Godrie, A. Goffeau, B. Golding, J. Gouzy, M. Gurjal, I. Hernandez-Lucas, A. Hong, L. Huizar, R. W. Hyman, T. Jones, D. Kahn, M. L. Kahn, S. Kalman, D. H. Keating, E. Kiss, C. Komp, V. Lelaure, D. Masuy, C. Palm, M. C. Peck, T. M. Pohl, D. Portetelle, B. Purnelle, U. Ramsperger, R. Surzycki, P. Thebault, M. Vandenbol, F. J. Vorholter, S. Weidner, D. H. Wells, K. Wong, K. C. Yeh, and J. Batut. 2001. The composite genome of the legume symbiont Sinorhizobium meliloti. Science 293:668-672.[Abstract/Free Full Text]
  16. Ghelardi, E., F. Celandroni, S. Salvetti, D. J. Beecher, M. Gominet, D. Lereclus, A. C. Wong, and S. Senesi. 2002. Requirement of flhA for swarming differentiation, flagellin export, and secretion of virulence-associated proteins in Bacillus thuringiensis. J. Bacteriol. 184:6424-6433.[Abstract/Free Full Text]
  17. Gophna, U., E. Z. Ron, and D. Graur. 2003. Bacterial type III secretion systems are ancient and evolved by multiple horizontal-transfer events. Gene 312:151-163.[CrossRef][Medline]
  18. Gupte, G., C. Woodward, and V. Stout. 1997. Isolation and characterization of rcsB mutations that affect colanic acid capsule synthesis in Escherichia coli K-12. J. Bacteriol. 179:4328-4335.[Abstract/Free Full Text]
  19. Haine, V., A. Sinon, F. Van Steen, S. Rousseau, M. Dozot, P. Lestrate, C. Lambert, J. J. Letesson, and X. De Bolle. 2005. Systematic targeted mutagenesis of Brucella melitensis 16M reveals a major role for GntR regulators in the control of virulence. Infect. Immun. 73:5578-5586.[Abstract/Free Full Text]
  20. Harshey, R. M. 2003. Bacterial motility on a surface: many ways to a common goal. Annu. Rev. Microbiol. 57:249-273.[CrossRef][Medline]
  21. Jacobi, S., R. Schade, and K. Heuner. 2004. Characterization of the alternative sigma factor {sigma}54 and the transcriptional regulator FleQ of Legionella pneumophila, which are both involved in the regulation cascade of flagellar gene expression. J. Bacteriol. 186:2540-2547.[Abstract/Free Full Text]
  22. Jeon, B., K. Itoh, N. Misawa, and S. Ryu. 2003. Effects of quorum sensing on flaA transcription and autoagglutination in Campylobacter jejuni. Microbiol. Immunol. 47:833-839.[Medline]
  23. Kim, Y. K., and L. L. McCarter. 2000. Analysis of the polar flagellar gene system of Vibrio parahaemolyticus. J. Bacteriol. 182:3693-3704.[Abstract/Free Full Text]
  24. Klose, K. E., and J. J. Mekalanos. 1998. Distinct roles of an alternative sigma factor during both free-swimming and colonizing phases of the Vibrio cholerae pathogenic cycle. Mol. Microbiol. 28:501-520.[CrossRef][Medline]
  25. Klose, K. E., D. S. Weiss, and S. Kustu. 1993. Glutamate at the site of phosphorylation of nitrogen-regulatory protein NTRC mimics aspartyl-phosphate and activates the protein. J. Mol. Biol. 232:67-78.[CrossRef][Medline]
  26. Koretke, K. K., A. N. Lupas, P. V. Warren, M. Rosenberg, and J. R. Brown. 2000. Evolution of two-component signal transduction. Mol. Biol. Evol. 17:1956-1970.[Abstract/Free Full Text]
  27. Kovach, M. E., R. W. Phillips, P. H. Elzer, R. M. Roop II, and K. M. Peterson. 1994. pBBR1MCS: a broad-host-range cloning vector. BioTechniques 16:800-802.[Medline]
  28. Lan, C. Y., and M. M. Igo. 1998. Differential expression of the OmpF and OmpC porin proteins in Escherichia coli K-12 depends upon the level of active OmpR. J. Bacteriol. 180:171-174.[Abstract/Free Full Text]
  29. Letesson, J. J., and X. De Bolle. 2004. Brucella virulence: a matter of control, p. 117-158. In I. Lopez-Goni and I. Moriyon (ed.), Brucella: molecular and cellular biology. Horizon Bioscience, Norfolk, United Kingdom.
  30. Lupp, C., and E. G. Ruby. 2005. Vibrio fischeri uses two quorum-sensing systems for the regulation of early and late colonization factors. J. Bacteriol. 187:3620-3629.[Abstract/Free Full Text]
  31. Macnab, R. M. 1996. Flagella and motility, p. 123-145. In F. C. Neidhardt et al. (ed.), Escherichia coli and Salmonella: cellular and molecular biology, vol. 1. ASM Press, Washington, D.C.
  32. Macnab, R. M. 2003. How bacteria assemble flagella. Annu. Rev. Microbiol. 57:77-100.[Medline]
  33. Mangan, S., and U. Alon. 2003. Structure and function of the feed-forward loop network motif. Proc. Natl. Acad. Sci. USA 100:11980-11985.[Abstract/Free Full Text]
  34. Miller, J. H. 1972. Experiments in molecular genetics. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y.
  35. Millikan, D. S., and E. G. Ruby. 2003. FlrA, a {sigma}54-dependent transcriptional activator in Vibrio fischeri, is required for motility and symbiotic light-organ colonization. J. Bacteriol. 185:3547-3557.[Abstract/Free Full Text]
  36. Moens, S., and J. Vanderleyden. 1996. Functions of bacterial flagella. Crit. Rev. Microbiol. 22:67-100.[Medline]
  37. Moore, J. B., S. P. Shiau, and L. J. Reitzer. 1993. Alterations of highly conserved residues in the regulatory domain of nitrogen regulator I (NtrC) of Escherichia coli. J. Bacteriol. 175:2692-2701.[Abstract/Free Full Text]
  38. Muir, R. E., and J. W. Gober. 2004. Regulation of FlbD activity by flagellum assembly is accomplished through direct interaction with the trans-acting factor, FliX. Mol. Microbiol. 54:715-730.[CrossRef][Medline]
  39. Nguyen, L., I. T. Paulsen, J. Tchieu, C. J. Hueck, and M. H. Saier, Jr. 2000. Phylogenetic analyses of the constituents of type III protein secretion systems. J. Mol. Microbiol. Biotechnol. 2:125-144.[CrossRef][Medline]
  40. Poindexter, J. S. 1964. Biological properties and classification of the Caulobacter group. Bacteriol. Rev. 28:231-295.[Free Full Text]
  41. Prouty, M. G., N. E. Correa, and K. E. Klose. 2001. The novel sigma54- and sigma28-dependent flagellar gene transcription hierarchy of Vibrio cholerae. Mol. Microbiol. 39:1595-1609.[CrossRef][Medline]
  42. Ramos, H. C., M. Rumbo, and J. C. Sirard. 2004. Bacterial flagellins: mediators of pathogenicity and host immune responses in mucosa. Trends Microbiol. 12:509-517.[CrossRef][Medline]
  43. Robertson, G. T., M. E. Kovach, C. A. Allen, T. A. Ficht, and R. M. Roop II. 2000. The Brucella abortus Lon functions as a generalized stress response protease and is required for wild-type virulence in BALB/c mice. Mol. Microbiol. 35:577-588.[CrossRef][Medline]
  44. Sambrook, J., E. F. Fritsch, and T. Maniatis. 1989. Molecular cloning: a laboratory manual, 2nd ed. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y.
  45. Sanders, D. A., B. L. Gillece-Castro, A. M. Stock, A. L. Burlingame, and D. E. Koshland, Jr. 1989. Identification of the site of phosphorylation of the chemotaxis response regulator protein, CheY. J. Biol. Chem. 264:21770-21778.[Abstract/Free Full Text]
  46. Shapiro, L. 1995. The bacterial flagellum: from genetic network to complex architecture. Cell 80:525-527.[CrossRef][Medline]
  47. Siam, R., and G. T. Marczynski. 2003. Glutamate at the phosphorylation site of response regulator CtrA provides essential activities without increasing DNA binding. Nucleic Acids Res. 31:1775-1779.[Abstract/Free Full Text]
  48. Simon, R., U. Priefer, and A. Pühler. 1983. A broad host range mobilization system for in vivo genetic engineering: transposon mutagenesis in gram-negative bacteria. Bio/Technology 1:784-791.[CrossRef]
  49. Skerker, J. M., and M. T. Laub. 2004. Cell-cycle progression and the generation of asymmetry in Caulobacter crescentus. Nat. Rev. Microbiol. 2:325-337.[CrossRef][Medline]
  50. Song, Y. C., S. Jin, H. Louie, D. Ng, R. Lau, Y. Zhang, R. Weerasekera, S. Al Rashid, L. A. Ward, S. D. Der, and V. L. Chan. 2004. FlaC, a protein of Campylobacter jejuni TGH9011 (ATCC43431) secreted through the flagellar apparatus, binds epithelial cells and influences cell invasion. Mol. Microbiol. 53:541-553.[CrossRef][Medline]
  51. Sourjik, V., P. Muschler, B. Scharf, and R. Schmitt. 2000. VisN and VisR are global regulators of chemotaxis, flagellar, and motility genes in Sinorhizobium (Rhizobium) meliloti. J. Bacteriol. 182:782-788.[Abstract/Free Full Text]
  52. Soutourina, O. A., and P. N. Bertin. 2003. Regulation cascade of flagellar expression in Gram-negative bacteria. FEMS Microbiol. Rev. 27:505-523.[CrossRef][Medline]
  53. 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. 43:809-821.[CrossRef][Medline]
  54. Stewart, B. J., and L. L. McCarter. 2003. Lateral flagellar gene system of Vibrio parahaemolyticus. J. Bacteriol. 185:4508-4518.[Abstract/Free Full Text]
  55. Stock, A. M., V. L. Robinson, and P. N. Goudreau. 2000. Two-component signal transduction. Annu. Rev. Biochem. 69:183-215.[CrossRef][Medline]
  56. Weiss, V., and B. Magasanik. 1988. Phosphorylation of nitrogen regulator I (NRI) of Escherichia coli. Proc. Natl. Acad. Sci. USA 85:8919-8923.[Abstract/Free Full Text]
  57. Young, G. M., D. H. Schmiel, and V. L. Miller. 1999. A new pathway for the secretion of virulence factors by bacteria: the flagellar export apparatus functions as a protein-secretion system. Proc. Natl. Acad. Sci. USA 96:6456-6461.[Abstract/Free Full Text]

Journal of Bacteriology, January 2007, p. 131-141, Vol. 189, No. 1
0021-9193/07/$08.00+0     doi:10.1128/JB.00712-06
Copyright © 2007, American Society for Microbiology. All Rights Reserved.



This article has been cited by other articles:


This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Other Versions of this Article:
JB.00712-06v1
189/1/131    most recent
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Services
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrow Copyright Information
Right arrow Books from ASM Press
Right arrow MicrobeWorld
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Léonard, S.