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Infect Immun. Aug 2005; 73(8): 4982–4992.
PMCID: PMC1201253

Characterization of the cciIR Quorum-Sensing System in Burkholderia cenocepacia


Several transmissible Burkholderia cenocepacia strains that infect multiple cystic fibrosis patients contain a genomic island designated as the cenocepacia island (cci). The cci contains a predicted N-acylhomoserine lactone (AHL) synthase gene, cciI, and a predicted response regulator gene, cciR. AHL production profiles indicated that CciI catalyzes the synthesis of N-hexanoyl-l-homoserine lactone and minor amounts of N-octanoyl-l-homoserine lactone. The cciI and cciR genes were found to be cotranscribed by reverse transcription-PCR analysis, and the expression of a cciIR::luxCDABE fusion in a cciR mutant suggested that the cciIR system negatively regulates its own expression. B. cenocepacia strains also have a cepIR quorum-sensing system. Expression of cepI::luxCDABE or cepR::luxCDABE fusions in a cciR mutant showed that CciR negatively regulates cepI but does not regulate cepR. Expression of the cciIR::luxCDABE fusion in a cepR mutant indicated that functional CepR is required for cciIR expression. Phylogenetic analysis suggested that the cciIR system was acquired by horizontal gene transfer from a distantly related organism and subsequently incorporated into the ancestral cepIR regulatory network. Mutations in cciI, cciR, cepI cciI, and cepR cciR were constructed in B. cenocepacia K56-2. The cciI mutant had greater protease activity and less swarming motility than the parent strain. The cciR mutant had less protease activity than the parent strain. The phenotypes of the cepI cciI and cepR cciR mutants were similar to cepI or cepR mutants, with less protease activity and swarming motility than the parent strain.

Over the past 20 years a group of closely related Burkholderia species termed the B. cepacia complex (Bcc) have emerged as opportunistic pathogens of particular importance in people with cystic fibrosis (CF) and chronic granulomatous disease (31, 33). Bcc infections are a major concern in CF because some of the Bcc strains have high patient-to-patient transmissibility, multidrug resistance, and the potential to cause inflammation and fatal invasive disease (47). Bcc bacteremia has been documented in CF and non-CF patients (59). The Bcc is currently comprised of nine species, each of which has been isolated from CF patients; however, they vary in their frequency of colonization, transmissibility, and geographic distribution. B. cenocepacia is the most commonly reported Bcc species isolated from patients with CF (26, 48). The majority of transmissible or epidemic Bcc strains are B. cenocepacia (26, 31, 48, 59), and epidemiological studies have shown this species is highly virulent, causing significant mortality among CF patients (28, 30).

N-Acylhomoserine lactone (AHL)-mediated quorum-sensing systems are comprised of a luxI homologue that encodes an AHL synthase that catalyzes the synthesis of AHL signal molecule(s) and a luxR homologue that encodes a sensor/response regulator, which binds its cognate AHL and alters gene expression at the level of transcription. Some organisms have multiple AHL quorum-sensing systems which may be integrated with each other as well as with other bacterial control circuitry (13). For example, Pseudomonas aeruginosa possesses two quorum-sensing systems, lasIR and rhlIR. These two systems are responsible for the regulation of a large number of genes, including many that play a role in virulence (39, 56). LasR and RhlR regulate some genes in a hierarchical manner (34).

The cepIR AHL-mediated quorum-sensing system is widely distributed among Bcc strains (15, 27). CepI directs the synthesis of N-octanoyl-l-homoserine lactone (OHL) and of N-hexanoyl-l-homoserine lactone (HHL) (23, 24). Recent proteomic and transcriptional analysis studies have indicated that the cepIR system is involved in the regulation of numerous target genes (2, 36). In B. cenocepacia CepR positively regulates cepI and negatively controls its own expression (24). The cepIR quorum-sensing system in B. cenocepacia is involved in the regulation of swarming motility, mature biofilm development, chitinase production, extracellular protease production, and the biosynthesis of the siderophore ornibactin (18, 23-25). Transcriptional analysis was used to demonstrate that a zinc metalloprotease gene, zmpA, is positively regulated by cepIR (46) and that the cepIR system is involved in the negative regulation of the ornibactin biosynthesis gene, pvdA (24). Ornibactin is a hydroxamate siderophore which has been shown to play a role in virulence in both chronic and acute models of respiratory infection (44). The cepIR quorum-sensing system positively regulates swarming motility in B. cenocepacia, possibly by controlling biosurfactant production (18, 25).

Animal, nematode, and plant infection models have shown that the cepIR quorum-sensing system contributes to virulence in B. cenocepacia and B. cepacia (1, 20, 46). Using a rat chronic respiratory infection model, Sokol et al. (46) demonstrated that infections with cepIR mutants caused significantly less lung histopathological changes despite similar lung bacterial counts compared to the wild type. A cepI mutant was less virulent in both wild-type and Cftr/ mice. The cepI mutant was recovered from the lungs in significantly lower numbers than the parent strain and was not invasive in the Cftr/ mice (46).

The cepIR system has also been shown to be required for efficient killing of Caenorhabditis elegans by B. cenocepacia H111 (20). A B. cepacia cepI mutant was attenuated in its ability to macerate onion tissue (1). Thus, a role for the cepIR system in the regulation of Bcc virulence traits in a wide range of infection models has been established.

A novel set of luxIR homologues was identified during characterization of the genomic region surrounding the Burkholderia cepacia epidemic strain marker (BCESM) (4). The BCESM was originally identified as a 1.4-kb DNA fragment amplified from transmissible Bcc strains during random amplified polymorphic DNA typing (29), and the marker has been widely applied in infection control as an indicator of clinical risk (28, 30, 48). Baldwin et al. (4) recently demonstrated that the BCESM was a part of a 31.7-kb low-GC content genomic island termed the cenocepacia island (cci), which contained 37 open reading frames involved in both virulence and metabolism. The luxIR homologues identified within the island and, therefore, associated with epidemic strains of B. cenocepacia were designated cciIR. A cciI mutant was shown to have attenuated virulence in a rat chronic respiratory infection model (4). The objectives of the current study were to determine if cciIR is a functional quorum-sensing system, if it is involved in the regulation of virulence-associated phenotypes, and if a regulatory hierarchy exists between the cepIR and cciIR quorum-sensing systems in epidemic strains of B. cenocepacia.


Bacterial strains and growth conditions.

The bacterial strains and plasmids used in this study are listed in Table Table1.1. For genetic manipulations, including RNA isolation, cultures were routinely grown at 37°C in Luria broth (Invitrogen, Burlington, Ontario, Canada) with shaking (200 rpm) or on 1.5% LB agar plates. When appropriate, the following concentrations of antibiotics were used: 100 μg/ml of trimethoprim (Tp) and 200 μg/ml of tetracycline (Tc) for B. cenocepacia and 1.5 mg/ml Tp, 15 μg/ml of Tc, and 50 μg/ml of kanamycin (Km) for Escherichia coli. Antibiotics were purchased from Sigma-Aldrich Canada Ltd. (Oakville, Ontario, Canada). For protease, β-galactosidase, and luminescence assays, cultures were grown in 0.25% Trypticase soy broth (TSB; Difco, Franklin Lakes, NJ) with 5% Bacto-Peptone (Difco) (PTSB) at 37°C. For chrome azurol S (CAS) assays, cultures were grown at 32°C in succinate medium supplemented with ornithine (10 mM) (32). For examination of swarming motility, cultures were grown in nutrient broth (Difco) supplemented with 0.5% glucose. For AHL extractions, cultures were grown in 100% TSB at 37°C.

Bacterial strains and plasmids used in this study

DNA manipulations.

DNA manipulations were performed using standard techniques as described by Sambrook et al. (38). Genomic DNA was isolated as described by Ausubel et al. (3). Restriction endonucleases and T4 DNA polymerase were purchased from Invitrogen. T4 DNA ligase was purchased from New England Biolabs (Mississauga, Ontario, Canada). Oligonucleotide primers were purchased from Invitrogen or from University of Calgary Core DNA and Protein Services (Calgary, Alberta, Canada). Plasmids were introduced into B. cenocepacia by electroporation using a Gene Pulser (Bio-Rad, Richmond, CA) as previously described (8). Nucleotide sequencing was performed by Macrogen Inc. (Seoul, Korea) or by University of Calgary Core DNA and Protein Services. The nucleotide sequence for the cciIR locus was obtained from the B. cenocepacia sequencing project (http://www.sanger.ac.uk/Projects/B_cenocepacia/).

Cloning of cciI.

The cciI gene was PCR amplified from K56-2 using the oligonucleotide primers CCIF (5′-GCCTCATTGTGCACTCGTG-3′) and CCIR (5′-GGTGGCACTGACATCGAAAG-3′), and the 850-bp PCR fragment was cloned into pCR2.1Topo (Invitrogen). The nucleotide sequence was determined to confirm the absence of PCR errors. The resulting cciI clone was designated pRM1T4.

Construction of cciR, cepI cciI, and cepR cciR mutants by allelic exchange.

The 2.0-kb fragment containing the cciR gene and flanking region was amplified from K56-2 using the oligonucleotide primers CCRBam (5′-CATCGCGGATCCCGCCAT-3′) with a BamHI site (underlined) and CCRHin (5′-ATGTGCTAAGCTTGATCGACC-3′) with a HindIII site and cloned into the BamHI/HindIII sites of pEX18Tc (17). The cciR gene was disrupted by incorporation of the Tp cassette from p34E-Tp (10) into the MluI site of cciR, resulting in pRM186Tp. The vector was transferred into K56-2 using pRK2013 as the mobilizing plasmid (12). Transconjugants were plated onto Pseudomonas isolation agar (Difco) containing Tp to select for single-crossover events. The cciR::Tp mutant (K56-2cciR) was identified by screening for Tc sensitivity.

To construct an unmarked ΔcciI ΔcciR mutant, a 4.3-kb DNA fragment containing cciIR was shotgun cloned from XhoI fragments separated by sucrose gradient fractionation. The fraction containing the cciIR fragment was identified by Southern hybridization with a 449-bp cciR probe and a 450-bp cciI probe and cloned into the XhoI site of pCR2.1Topo (Invitrogen). The cciI and cciR probes were PCR amplified using the oligonucleotide primers INcciIF (5′-CATCTTCGCTGGCAGTTTCG-3′) with INcciIR (5′-AACGCTGGTAAAGCCGTGC-3′) and INcciRF (5′-TTACGCGCAACGAGACTACG-3′) with INcciRR (5′-ATCTTCATCGCTTCGGCG-3′), respectively. Positive clones were identified using colony hybridization assays (60). The 4.3-kb cciIR fragment was cloned into pEX18Tc, and cciR was disrupted by deleting a 637-bp NcoI-to-MluI fragment, resulting in pRM187-KO. This deletion removed the 5′ end of cciR and the cciIR promoter region. Allelic exchange was conducted as described above, resulting in a ΔcciI ΔcciR double mutant (K56-2cciIR). A cepR::Tp ΔcciI ΔcciR mutant (K56-2cepR cciIR) was constructed in K56-2cciIR with pEXCEPR (24). A ΔcepI cciI::Tp mutant (K56-2cepI cciI) was constructed in K56-dI2 by using pCciI-Tp (4). K56-dI2 is a cepI allelic exchange mutant with a 290-bp NruI fragment deletion in cepI (C. E. Chambers, P. Law, M. B.Visser, E. I. Lutter, and P. A. Sokol, Abstr. 2nd ASM Conf. Cell-Cell Commun. Bacteria, abstr. 119B, 2004). Two independent ΔcepI cciI::Tp mutants, designated K56-2cepI cciIa and K56-2cepI cciIb, were isolated. K56-2cepI cciIa appeared to have a spontaneous mutation that affected ornibactin biosynthesis. Allelic exchange was confirmed in all mutants by PCR and Southern hybridization.

AHL extraction and TLC-AHL bioassays.

AHLs were extracted from culture supernatants twice with equal volumes of acidified ethyl acetate as described elsewhere (24). Thin-layer chromatogrpahy (TLC)-AHL bioassays were performed as described previously using Agrobacterium tumefaciens A136 (pCF218) (pCF372) as a reporter strain (24). This reporter strain is able to identify AHLs with 3-oxo-, 3-hydroxy-, and 3-unsubsituted side chains ranging from 6 to 16 carbons in length (42). Synthetic HHL and OHL (Sigma-Aldrich) were used as reference standards.

Construction of luxCDABE transcriptional fusions.

A cepR::luxCDABE transcriptional fusion, pRM432, was constructed by amplifying the cepR promoter region as described by Lewenza et al. (24) by PCR with the oligonucleotide primers CPRxho (5′-GTTCCGGCTCGAGCGGCG-3′) and CPRbam (5′-CATGAAGCGGATCCTCAGCG-3′). These primers were designed with incorporated XhoI and BamHI sites, respectively. The 1.8-kb cepR promoter fragment was subsequently cloned into the XhoI/BamHI cloning site of pMS402 (11). The cepI::luxCDABE transcriptional fusion, pCP300, contains a 300-bp fragment containing the cepI promoter region (Chambers et al., Abstr. 2nd ASM Conf. Cell-Cell Commun. Bacteria, 2004). The promoter regions for cciI and cciR were predicted in silico using SoftBerry BPROM (http://www.softberry.com). For construction of the cciI and cciR promoter fusions, a 391-bp SalI/SacII fragment and an 813-bp ClaI fragment from pRM4.3 containing the respective predicted promoter regions were subcloned into the EcoRV site of pCR2.1Topo (Invitrogen). A XhoI/BamHI fragment containing the cciI or cciR promoter region was subsequently cloned into pMS402 (11) and designated pRM446 or pRM445, respectively.

Luminescence assays.

Overnight cultures were subcultured to an initial absorbance at 600 nm of 0.02 in 20 ml medium. At selected times, 200-μl aliquots were removed and the luminescence in counts per second (cps) and turbidity at an absorbance of 600 nm were measured using a Wallac Victor2 Multi-label counter (Perkin-Elmer Life Sciences, Woodbridge, Ontario, Canada). The samples were read in black, clear-bottom 96-well microtiter plates (Corning Inc., Corning, NY). The level of promoter expression is reported as the ratio of luminescence to turbidity or relative luminescence.


Overnight K56-2 cultures were subcultured into 20 ml medium at an initial optical density at 600 nm (OD600) of 0.02 and grown for 10 h. Total RNA was isolated from approximately 1 × 109 cells with a QIAGEN RNeasy mini kit (QIAGEN, Mississauga, Ontario, Canada). RNA was treated with amplification-grade DNase I (Invitrogen) before use. Reverse transcription-PCR (RT-PCR) was performed using a Titan One-tube RT-PCR kit (Roche, Mannheim, Germany) according to the manufacturer's instructions. For each reaction mixture, 75 ng of RNA was used. cDNA was synthesized by reverse transcription at 50°C for 40 min. Denaturation was performed for 2 min at 96°C followed by 35 cycles of PCR as suggested by the manufacturer. A final elongation step at 68°C for 7 min was conducted. The primers and annealing temperatures used were as follows: cciI (64°C), cciIRTF (5′-TCGCGGTCGTACGATTCAC-3′) and cciIRTR (5′-TTGCACCGATCAGGTAGGC-3′); cciR (58°C), cciRRTF (5′-ACGCGAGGCACTCTTGTTG-3′) and cciRRTR (5′-GCCGACATCAGAGGCTTGAA-3′); cciIR intergenic region (58°C), irBRRTF (5′-ATTGATCCCATTCGGTCAGG-3′) and irBRRTR (5′-CGCCTCCATTGTTGGCATA-3′). To ensure that there was no DNA contamination in the RNA samples, Platinum Taq polymerase (Invitrogen) was used instead of the reverse transcriptase enzyme mixture under the same reaction conditions.

Phylogenetic analysis of quorum-sensing systems.

The AHL synthase and response regulator protein sequences from B. cenocepacia strain K56-2 (CepI, CciI, CepR, and CciR) were used to search the protein databases with the PHI-BLAST search tool at NCBI (http://www.ncbi.nlm.nih.gov/BLAST/). The 100 matches with greatest identity were collected for each synthase and response regulator protein. These were combined with the recently described quorum-sensing genes in B. pseudomallei (51) (obtained from the Sanger Institute [http://www.sanger.ac.uk/Projects/B_pseudomallei/]) and subjected to phylogenetic analysis as follows. Multiple amino acid alignments were constructed using CLUSTAL W (50), and phylogenetic trees were drawn using genetic distance-based neighbor-joining algorithms within Treecon for Windows version 1.3 (55). The following parameters were used for each tree: sequence input order was randomized, the Poisson correction algorithm was applied, 1,000 data sets were examined by bootstrapping resampling statistics, and each tree was rooted with the appropriate synthase or response regulator sequence originally described in Vibrio fischeri. Sequences representative of each major phylogenetic cluster were then selected for final analysis. Corresponding synthase and response regulator protein sequences for each individual quorum-sensing system were concatenated, and their combined sequences were used to construct a final phylogenetic tree as described above.

β-Galactosidase assays.

β-Galactosidase activity was measured as previously described (35). Overnight cultures were subcultured to an initial absorbance at 600 nm of 0.02 in 20 ml of PTSB. Throughout the time course, 1-ml aliquots were removed and assayed for β-galactosidase activity, and this activity was expressed as Miller units (35).

Siderophore, protease, and swarming motility assays.

Siderophore activities present in the culture supernatant fluid were measured by CAS assays (23). Overnight cultures were subcultured (1/100) into fresh medium. The cultures were grown for 40 h at 32°C, and CAS assays were performed on 100 μl of supernatant fluid. The absorbance at 630 nm was measured and divided by the culture turbidity (absorbance at 600 nm) to normalize for cell density, and this ratio was reported as CAS activity (41).

Protease activity was determined using skim milk as a substrate. Overnight cultures were grown and subcultured (1/100) into fresh medium. At the mid-log phase of growth, the cultures were normalized to an optical density of 0.3 at 600 nm and spot inoculated (3 μl) in triplicate onto dialyzed 1.5% brain heart infusion agar (Difco) containing 1.5% skim milk (45). The plates were incubated at 37°C for 24 h, and the zone of clearing around the growth was measured. For AHL add-back assays, 2.5 nM and 10.0 nM concentrations of synthetic HHL and 2.5 nM of synthetic OHL dissolved in 20% acetonitrile were spread on the agar prior to spot inoculation.

Swarming motility was evaluated using semisolid agar (0.5%) motility assays as previously described (25). Briefly, overnight cultures were normalized, and a 1-μl was spot inoculated in the center of the nutrient broth (Difco), 0.5% glucose, and 0.5% agar surface. The plates were incubated at 37°C for 24 or 48 h, and the diameter of the swarming zone was measured.


Characterization of AHL production by CciI.

K56-2 has previously been reported to produce OHL and HHL. K56-I2 (cepI) produces small amounts of HHL but no detectable OHL (24). To determine if cciI encodes an AHL synthase that is responsible for the production of HHL in K56-I2 (cepI), the cciI gene was cloned and expressed in E. coli DH5α. The AHLs present in the culture supernatant of E. coli DH5α(pRM1T4), which contains cciI, were analyzed using the A. tumefaciens TLC-AHL bioassay. E. coli DH5α containing cciI produced AHLs that comigrated with synthetic HHL (Fig. (Fig.1A).1A). An AHL that comigrated with synthetic OHL was weakly detectable only after the plate was allowed to develop for more than 72 h. E. coli DH5α(pSLS225), which contains cepI, produced AHLs that comigrated on the TLC plate with both HHL and OHL (Fig. (Fig.1A1A).

FIG. 1.
TLC-AHL bioassay. Ethyl acetate extracts were chromatographed on C18 reverse-phase TLC plates developed with methanol-water (70:30, vol/vol). The spots were visualized using the A. tumefaciens reporter strain. (A) Heterologous expression of cciI in E. ...

To confirm that cciI directs the synthesis of HHL and minor amounts of OHL, extracts of K56-2cciI and K56-2cepI cciIa were analyzed using the A. tumefaciens TLC-AHL bioassay. K56-2cciI produced slightly less HHL than the parent strain. HHL production was restored to parental levels when cciI was present in trans (Fig. (Fig.1B).1B). There were no detectable AHLs in extracts from K56-2cepI cciIa, indicating that cepI and cciI are collectively responsible for AHL synthesis in K56-2 (Fig. (Fig.1C).1C). HHL or OHL production was restored in K56-2cepI cciIa by complementation with cepI or cciI, confirming the results obtained by heterologous expression in E. coli (Fig. (Fig.1A).1A). K56-R2 (cepR) has previously been shown to produce small amounts of HHL (24). The K56-2cciR and the K56-2cciIR AHL production profiles were similar to K56-2 (Fig. (Fig.1D),1D), indicating that cciR is not required for AHL production. K56-2cepR cciIR produced very small amounts of HHL and OHL that were weakly detectable only when the plate was overdeveloped (Fig. (Fig.2D),2D), confirming that cepR is required for optimum production of both HHL and OHL.

FIG. 2.
Physical and genetic map of the XhoI cciIR locus of epidemic strains of B. cenocepacia. (A) Map of the 4.3-kb insert of pRM4.3 containing cciIR. The arrows indicate the location and orientation of the genes. The Tp cassette was incorporated into the MluI ...

Transcriptional analysis of cciIR.

Transcriptional analysis of cciIR was performed to determine if these genes regulate their own transcription. The promoter regions of cciI and cciR were predicted in silico, and cciI::luxCDABE and cciR::luxCDABE transcriptional fusions were constructed based on these predictions (Fig. (Fig.2B).2B). The expression of cciI::luxCDABE (pRM446) and cciR::luxCDABE (pRM445) was compared in K56-2 (data not shown), and expression was only observed for the cciR::luxCDABE transcriptional fusion, pRM445. Additional cciI transcriptional fusion constructs were made by cloning various lengths of the region upstream of cciI in front of the promoterless luxCDABE operon. None of these constructs resulted in the expression of the luxCDABE operon (data not shown).

RT-PCR was used to determine if cciI and cciR are cotranscribed. Three primer sets were designed to internally amplify cciI, cciR, and the intergenic region between cciR and cciI (Fig. (Fig.2C).2C). A product was amplified for each gene as well as the intergenic region, indicating that cciI and cciR are on the same transcript (Fig. (Fig.2D).2D). Therefore, cciI is transcribed from a promoter located upstream of cciR.

The expression of cciIR::luxCDABE was compared in K56-2 and K56-2cciIR over a 24-h time course. The expression of cciIR::luxCDABE was significantly greater in K56-2cciIR than in K56-2 from 12 to 16 h (P < 0.05, analysis of variance [ANOVA]). The twofold increase in expression indicates that the cciIR system negatively regulates its own expression (Fig. (Fig.3A3A).

FIG. 3.
Transcriptional analysis of cciIR cepR and cepI expression. Gene expression was monitored throughout growth in PTSB plus 100 μg/ml of trimethoprim. All values are the means ± SD of triplicate cultures. [filled square], K56-2; [filled triangle], K56-2 ...

Regulatory relationship between cciIR and cepIR quorum-sensing systems.

To determine if there is a regulatory relationship between the cepIR and the cciIR genes, expression levels of the cepI::luxCDABE(pCP300) and cepR::luxCDABE(pRM432) transcriptional fusions were compared in K56-2 and K56-2cciIR. There was a significant increase in cepI::luxCDABE expression in K56-2cciIR compared to K56-2 (P < 0.02; t test), indicating that cciIR is involved in the negative regulation of cepI (Fig. (Fig.3B).3B). The expression of cepR::luxCDABE in K56-2cciIR was the same as K56-2, demonstrating that cciIR does not regulate cepR expression (Fig. (Fig.3C).3C). Expression of cciIR::luxCDABE (pRM445) was compared in K56-2 and K56-R2 (cepR) (Fig. (Fig.3A).3A). The expression of cciIR::luxCDABE was reduced to almost background levels in K56-R2 (cepR), indicating that cepR is required for cciIR expression.

Evolutionary analysis of B. cepacia complex quorum-sensing systems.

Phylogenetic analysis of the CepIR and CciIR systems of B. cenocepacia was carried out to determine if the evolutionary origins of each system could be traced. Construction of separate trees for synthase or response regulator proteins demonstrated that there was significant congruence in the phylogeny of proteins belonging to a single quorum-sensing system (data not shown). The overlapping tree topology of AHL synthase and response regulator proteins showed that coevolution occurs for each given system and that phylogenetic relationships between strains could be inferred from the analysis. Therefore, corresponding synthase and response regulator protein sequences were concatenated and analyzed to produce a tree consisting of 28 systems that was representative of the phylogenetic diversity observed during analysis of the 100 most closely related sequences (Fig. (Fig.4).4). The B. cenocepacia CepIR proteins clustered closely with other B. cepacia complex and B. pseudomallei group quorum-sensing systems, indicating that this type of quorum-sensing system may be ancestral to the Burkholderia genus. From an evolutionary perspective, the pathogenicity island system CciIR was highly distinct from CepIR, and phylogenetic analysis did not specifically locate its potential origin to bacterial sequences currently available in the genetic databases (Fig. (Fig.4).4). CciIR was most closely related to one of two novel systems present in the draft genome of B. xenovorans strain LB400; neither of the B. xenovorans quorum-sensing systems was associated with a genomic island (data not shown). One of the multiple systems in B. pseudomallei, BpmIR3 (51) also clustered on the same evolutionary arm as CciIR. The B. vietnamiensis BviIR quorum-sensing system (27) was distantly related to CepIR, CciIR, and all other Burkholderia genus quorum-sensing systems. This phenomenon of two evolutionary distinct quorum-sensing systems being present in a single strain was also observed for other genera, such as P. aeruginosa (RhlIR and LasIR) (Fig. (Fig.44).

FIG. 4.
Phylogenetic analysis of the B. cenocepacia cepIR and cciIR quorum-sensing systems. A phylogenetic tree of concatenated AHL synthase and receptor protein sequences related to the B. cenocepacia quorum-sensing systems is shown. The tree was rooted with ...

Characterization of cciI and cciR mutants.

Previously, cepI and cepR mutants were shown to be protease deficient (23). To evaluate the effect of mutations in the cciI and cciR quorum-sensing genes on the protease activity of K56-2, skim milk assays were performed (Table (Table2).2). K56-2cciI produced significantly greater protease activity than the parent strain (P < 0.01). This phenotype is opposite to that observed for K56-I2 (cepI) and K56-2cepI cciIa, which do not produce detectable protease activity. K56-2cciR produces less protease activity than the wild type but more protease activity than K56-R2 (cepR) or K56-2cepR cciIR.

Extracellular protease activity and swarming motility of B. cenocepacia quorum-sensing mutants

The cepIR system was previously shown to positively regulate the expression of the zinc metalloprotease gene zmpA (46). To determine if zmpA is also regulated by the cciIR system, the expression of a zmpA::lacZ transcriptional fusion was examined in K56-2, K56-I2 (cepI), K56-2cciI, and K56-2cciR (Fig. (Fig.5).5). The expression of zmpA::lacZ in K56-2cciR was significantly less than in the parent strain in stationary phase at 24 and 32 h, and the expression of zmpA::lacZ in K56-2cciI was significantly less than in the parent strain during the entire growth curve (P < 0.01; ANOVA). The respective twofold and fivefold differences in zmpA expression indicate that the cciIR system is involved in the positive regulation of zmpA.

FIG. 5.
Effect of cepI cciI and cciR on zmpA expression in K56-2. β-Galactosidase activity was monitored throughout growth in PTSB and is reported in Miller units. [filled square], K56-2(pSG208); [filled triangle], K56-2cciR(pSG206); •, K56-2cciI(pSG206); ×, ...

The abilities of the cciI gene to complement a mutation in cepI and of the cepI gene to complement a mutation in cciI were investigated with respect to protease activity. Plasmids containing cciI or cepI were introduced into the heterologous AHL synthase mutant, and the protease activity of each strain was determined on skim milk agar (Table (Table3).3). Introduction of cciI into K56-I2 (cepI) or cepI into K56-2cciI did not restore protease activity to parental levels. Plasmids containing either cciI or cepI were introduced into K56-2cepI cciIa. The protease activity of the mutant was restored when a plasmid containing cepI was introduced, but not upon the introduction of a plasmid containing cciI. These experiments indicate that cepI is unable to compensate for the mutation in cciI, and cciI is unable to compensate for the mutation in cepI. Complementation of K56-I2 (cepI) or K56-2cciI with cepI or cciI in trans, respectively, restored protease activity to the parental phenotype.

Complementation of the extracellular protease activity and swarming motility of the AHL synthase mutants with cepI or cciI in trans

The effects of exogenous HHL and OHL concentrations on the protease activities of K56-I2 (cepI), K56-2cciI, and K56-2cepI cciIa were examined by adding 2.5 to 10.0 nM synthetic HHL or 2.5 nM synthetic OHL to the skim milk agar prior to inoculation. The protease zones of K56-I2 (cepI) and K56-2cepI cciIa were restored to wild-type size when 2.5 nM OHL was added to the agar. Restoration of the protease zones in K56-I2 (cepI) was also observed when 2.5 nM HHL was added to the agar; however, K56-2cepI cciIa required more HHL to restore the protease zones to wild-type levels (Table (Table4).4). The protease zone size of K56-2cciI was reduced to parental levels when 2.5 nM HHL was added to the agar but was not affected by either 10 nM HHL or 2.5 nM OHL (Table (Table44).

Effect of various concentrations of exogenous AHL on extracellular protease activity

Previously, cepI and cepR mutants were shown to have severely impaired swarming motility. K56-2cciI, K56-I2 (cepI), and K56-2cepI cciIa had significantly less swarming motility than the parent strain (Table (Table2)2) (P < 0.01, ANOVA), indicating that AHL production is essential for maximal swarming motility in K56-2. The zones of swarming motility for K56-I2 (cepI) and K56-2cciI were increased to parental levels when their respective mutations were complemented with the genes in trans. When the cciI gene was introduced into K56-I2 (cepI) the swarming motility phenotype was not restored; however, when cepI was introduced into K56-2cciI the zone of swarming increased to parental levels (Table (Table3).3). The zone of swarming motility for K56-2cepI cciIa was only fully complemented when cepI was present in trans (Table (Table33).

The cepR mutant, K56-R2, and K56-2cepR cciIR also exhibited significantly less swarming motility than the parent strain (P < 0.01; ANOVA) (Table (Table2);2); however, the abilities of K56-2cciR (Table (Table2)2) and K56-2cciI cciR (data not shown) to swarm were not altered. These data confirm that AHL production is essential for maximal swarming motility, as K56-R2 (cepR) and K56-2cepR cciIR do not produce significant amounts of AHLs, whereas K56-2cciR produces parental AHL levels (Fig. (Fig.1D1D).

The role of cciIR in the regulation of ornibactin production was monitored using a quantitative CAS assay (Fig. (Fig.6).6). There was little difference in the amount of CAS activity present in culture supernatants with the exception of K56-I2 (cepI), which has previously been shown to produce more ornibactin (23), and K56-2cepI cciIa, which had no CAS activity. Attempts to complement CAS activity in K56-2cepI cciIa with either cciI or cepI in trans were unsuccessful (data not shown). Therefore a second cepI cciI mutant was constructed (designated K56-2cepI cciIb). This mutant had parental levels of CAS activity (Fig. (Fig.6),6), suggesting that K56-2cepI cciIa has a secondary mutation that affects ornibactin biosynthesis. K56-2cepI cciIb and cepI cciIa have the same protease and swarming phenotypes (data not shown). These data, together with the complementation data in Table Table3,3, confirm that both cepI and cciI influence protease production and swarming motility, but only cepI influences ornibactin biosynthesis.

FIG. 6.
Effect of cepIR and cciIR mutations on ornibactin biosynthesis in B. cenocepacia K56-2. Cultures were grown in succinate medium supplemented with 10 mM ornithine for 40 h. Spent supernatants were added to the CAS dye complex, and the activity was evaluated ...


The cciIR quorum-sensing system associated with epidemic strains of B. cenocepacia is involved in the regulation of known virulence factors and is a part of a regulatory network with the cepIR quorum-sensing system, since CepR is required for cciIR expression. Phylogenetic analysis demonstrated that the cepIR and cciIR systems are highly distinct. The cepIR system that is widely distributed among the Bcc was predicted to be the ancestral system, whereas the cciIR system was likely acquired by horizontal gene transfer when cci was incorporated into the B. cenocepacia genome. The presence of an ancestral and a more-recently acquired AHL-dependent quorum-sensing system has been observed in other species. The P. aeruginosa lasIR system is considered to be the ancestral system, and the rhlIR system is considered to be more recently acquired (22). The rhlIR system has been incorporated into the regulatory pathway of the lasIR system and is dependent on the lasIR system for expression (21, 34). Both of these pathways have been shown to be involved in the virulence of the organism in a variety of infection models (43), and it is believed that the acquisition of the rhlIR system has aided P. aeruginosa in honing regulation of virulence traits during evolution (22).

In the current study, transcriptional analysis of cciIR in the presence and absence of functional CepR and phenotypic characterization of double AHL synthase and response regulator mutants demonstrated that the B. cenocepacia cciIR system has been incorporated into the cepIR regulatory network. The predicted cciIR promoter region does not contain a cepR binding box similar to the lux-type consensus sequence identified upstream of cepI by Lewenza et al. (23). The delayed initiation of cciIR expression compared to the expression observed for cepI and cepR in the wild type (Fig. (Fig.4)4) is consistent with the observation that active CepR is required for expression of cciIR. The lack of AHL production by K56-R2 (cepR) (Fig. (Fig.1D)1D) also confirms that CepR is required for the expression of cciIR. Since both the cepIR and cciIR systems have been shown to be involved in the virulence of B. cenocepacia (4, 46) and B. cenocepacia strains that possess the cci are often associated with transmissibility and increased severity of clinical outcome (4), the evolution of the cepIR and cciIR quorum-sensing regulatory network may contribute to the invasiveness and transmissibility observed with B. cenocepacia.

Pathogenicity islands tend to contain genes encoding regulatory elements (16). These regulatory elements can be specific for genes encoded on the pathogenicity island and may also regulate genes in the rest of the genome. As well, regulatory systems encoded on pathogenicity islands may be regulated by systems encoded outside of the genomic region. Each of these regulatory arrangements is observed with the cciIR quorum-sensing system. The cciIR system was shown to be involved in negative self-regulation as well as positive regulation of the zmpA gene, which is not located within the cci. The cepIR quorum-sensing system is involved in the positive regulation of the cciIR quorum-sensing system. There are most likely additional regulatory genes involved in the cepIR hierarchy in epidemic strains of B. cenocepacia. Three regulators of the H111 cepIR quorum-sensing system (yciR, suhB, and yciL) have been identified and are postulated to influence cepIR by posttranscriptional control of cepR expression or by affecting the activity status of the response regulator (19). It is yet to be determined if these regulatory elements are also involved in the quorum-sensing cascade of K56-2.

The B. cenocepacia genome sequence (http://www.sanger.ac.uk/Projects/B_cenocepacia/) contains one other open reading frame with homology to luxR response regulators that may also be involved in the quorum-sensing cascade of K56-2. This luxR homologue is without a proximal luxI homologue. The LuxR homologue contains both a predicted AHL binding domain and a DNA binding motif typical of LuxR response regulators (data not shown) and is 38% identical to SolR of Ralstonia solanacearum (accession no. NP-521406) (37), 37% identical to BviR of B. vietnamiensis (accession no. AAK35156) (27), and 36% identical to B. cenocepacia CepR (accession no. AF019654.1) (23). This LuxR homologue could be involved in gene regulation employing the AHLs produced by CepI and CciI. B. pseudomallei possesses at least three sets of luxIR homologues, pmlIR, bpmIR2, and bpmIR3, and two additional luxR homologues, bpmR4 and bpmR5 (51, 54). Ulrich et al. (51) demonstrated that each gene of the B. pseudomallei quorum-sensing network is involved in animal pathogenicity. Although the AHL specificity of each of the LuxR homologues has yet to be determined, it is conceivable that BpmR4 and BpmR5 employ the AHLs produced by the three AHL synthases in order to regulate virulence gene expression.

The cciIR system regulates protease production and swarming motility. The hyperproduction of protease by K56-2cciI was surprising, since cciI positively influences zmpA expression. Corbett et al. (6) showed that a zmpA mutant of K56-2 produce very little protease activity in the skim milk agar assay. The decrease in protease production observed for K56-2cciR is most likely due to the decrease in zmpA expression in this mutant. The predicted result for the absence of functional CciI would also be a decrease in protease production. The most plausible explanation for the observed increase in protease production is that cciI is involved in the negative regulation of an additional, uncharacterized protease, or that cciI is involved in the regulation of a regulator of a protease gene and the disruption of cciI affected downstream events in a regulatory cascade resulting in the unexpected phenotype.

Our data demonstrate that cciIR is involved in the positive regulation of swarming motility through the production of AHLs by CciI. The regulation of swarming motility by quorum-sensing systems is believed to allow optimal dissemination of the bacteria when the population is saturating a particular niche and may allow bacteria to progress from local infection sites to other organs (49, 58).

The phenotypes of the cciI and cciR mutants differed with respect to AHL production, protease activity, and swarming motility, although transcriptional analysis of zmpA suggests that the cciIR system operates as a unit to positively regulate the expression of zmpA (Fig. (Fig.5).5). Presumably, both CepR and CciR respond to HHL and OHL, as their cognate AHL synthases produce these signaling molecules. The difference in phenotypes of K56-2cciR and K56-2cciI could be due to the parental levels of AHLs produced by K56-2cciR (Fig. (Fig.1D)1D) compared to K56-2cciI, which produces less HHL (Fig. (Fig.1B).1B). The expected decrease in HHL production in K56-2cciIR is likely compensated by the absence of negative regulation of cepI by CciR.

Both CciI and CepI are responsible for production of the same AHLs; however, the ratios of HHL to OHL production by the two synthases are different. The quantity of each AHL present appears to be critical, as K56-2cepI cciIa required more HHL to restore protease production to parental levels than K56-I2 (cepI), which produces minor amounts of HHL (Table (Table4).4). Protease activity in the K56-2cciI mutant was only reduced by the addition of HHL to the medium and not OHL. The AHL synthases cepI and cciI are not redundant, since cciI did not restore protease activity or the swarming phenotype in the cepI mutant (Table (Table3),3), and both cepI and cciI mutants are reduced in virulence (4, 46).

B. thailendensis, B. mallei, B. pseudomallei, and B. vietnamiensis possess multiple luxIR quorum-sensing systems (27, 51-53). The regulatory relationships, if any, between these systems have yet to be described. There is a diversity of signals produced by Burkholderia AHL synthases. B. pseudomallei PmlI, BpmI2, and BpmI3 have been shown to direct the synthesis of OHL, N-decanoyl-l-homoserine lactone (DHL), N-(3-hydroxyoctanoyl)-l-homoserine lactone (3-hydroxy-OHL), and N-(3-hydroxydecanoyl)-l-homoserine lactone (3-hydroxy-DHL)(51, 54). B. pseudomallei BpmI2 and BpmI3 direct the synthesis of N-(3-oxotetradecanoyl)-l-homoserine lactone (51). B. thailandensis BtaI1, BtaI2, and BtaI3 are responsible for the production of OHL, DHL, and HHL, respectively (53). B. mallei BmaI1 and BmaI3 produce OHL, DHL, and 3-hydroxy-OHL and OHL, DHL, and 3-hydroxy-DHL, respectively (52). B. vietnamiensis possesses the cepIR and bviIR systems (9). BviI synthesizes HHL, OHL, DHL, and N-dodecanoyl-homoserine lactone (5, 27). The phenotypes regulated by the AHL-mediated quorum-sensing systems vary among different Burkholderia species, but in the pathogenic species, B. cenocepacia, B. mallei, and B. pseudomallei, these systems have been shown to contribute to virulence (46, 51, 52, 54). Further studies are needed to determine the specific role of these quorum-sensing systems in the regulation of virulence factors.


We thank M. B. Visser for her excellent technical assistance.

This study was supported by a grant from the Canadian Cystic Fibrosis Foundation to P.A.S. R.J.M. is the recipient of an Alberta Heritage Foundation for Medical Research studentship. E.M. acknowledges the support of the UK Cystic Fibrosis Trust (grant PJ501) and Wellcome Trust (grant 075586).


Editor: J. T. Barbieri


1. Aguilar, C., I. Bertani, and V. Venturi. 2003. Quorum-sensing system and stationary-phase sigma factor (rpoS) of the onion pathogen Burkholderia cepacia genomovar I type strain, ATCC 25416. Appl. Environ. Microbiol. 69:1739-1747. [PMC free article] [PubMed]
2. Aguilar, C., A. Friscina, G. Devescovi, M. Kojic, and V. Venturi. 2003. Identification of quorum-sensing-regulated genes of Burkholderia cepacia. J. Bacteriol. 185:6456-6462. [PMC free article] [PubMed]
3. Ausubel, F., R. Brent, R. Kingston, D. Moore, J. Seidman, J. Smith, and K. Struhl. 1989. Current protocols in molecular biology, vol. 1. John Wiley & Sons, Inc., New York, N.Y.
4. Baldwin, A., P. A. Sokol, J. Parkhill, and E. Mahenthiralingam. 2004. The Burkholderia cepacia epidemic strain marker is part of a novel genomic island encoding both virulence and metabolism-associated genes in Burkholderia cenocepacia. Infect. Immun. 72:1537-1547. [PMC free article] [PubMed]
5. Conway, B. A., and E. P. Greenberg. 2002. Quorum-sensing signals and quorum-sensing genes in Burkholderia vietnamiensis. J. Bacteriol. 184:1187-1191. [PMC free article] [PubMed]
6. Corbett, C. R., M. N. Burtnick, C. Kooi, D. E. Woods, and P. A. Sokol. 2003. An extracellular zinc metalloprotease gene of Burkholderia cepacia. Microbiology 149:2263-2271. [PubMed]
7. Darling, P., M. Chan, A. D. Cox, and P. A. Sokol. 1998. Siderophore production by cystic fibrosis isolates of Burkholderia cepacia. Infect. Immun. 66:874-877. [PMC free article] [PubMed]
8. Dennis, J. J., and P. A. Sokol. 1995. Electrotransformation of Pseudomonas. Methods Mol. Biol. 47:125-133. [PubMed]
9. Dennis, J. J., and G. J. Zylstra. 1998. Plasposons: modular self-cloning minitransposon derivatives for rapid genetic analysis of gram-negative bacterial genomes. Appl. Environ. Microbiol. 64:2710-2715. [PMC free article] [PubMed]
10. DeShazer, D., and D. E. Woods. 1996. Broad-host-range cloning and cassette vectors based on the R388 trimethoprim resistance gene. BioTechniques 20:762-764. [PubMed]
11. Duan, K., C. Dammel, J. Stein, H. Rabin, and M. G. Surette. 2003. Modulation of Pseudomonas aeruginosa gene expression by host microflora through interspecies communication. Mol. Microbiol. 50:1477-1491. [PubMed]
12. Figurski, D. H., and D. R. Helinski. 1979. Replication of an origin-containing derivative of plasmid RK2 dependent on a plasmid function provided in trans. Proc. Natl. Acad. Sci. USA 76:1648-1652. [PMC free article] [PubMed]
13. Fuqua, C., and E. P. Greenberg. 2002. Listening in on bacteria: acyl-homoserine lactone signalling. Nat. Rev. Mol. Cell Biol. 3:685-695. [PubMed]
14. Fuqua, C., and S. C. Winans. 1996. Conserved cis-acting promoter elements are required for density-dependent transcription of Agrobacterium tumefaciens conjugal transfer genes. J. Bacteriol. 178:435-440. [PMC free article] [PubMed]
15. Gotschlich, A., B. Huber, O. Geisenberger, A. Togl, A. Steidle, K. Riedel, P. Hill, B. Tummler, P. Vandamme, B. Middleton, M. Camara, P. Williams, A. Hardman, and L. Eberl. 2001. Synthesis of multiple N-acylhomoserine lactones is wide-spread among the members of the Burkholderia cepacia complex. Syst. Appl. Microbiol. 24:1-14. [PubMed]
16. Hacker, J., and J. B. Kaper. 2000. Pathogenicity islands and the evolution of microbes. Annu. Rev. Microbiol. 54:641-679. [PubMed]
17. Hoang, T. T., R. R. Karkhoff-Schweizer, A. J. Kutchma, and H. P. Schweizer. 1998. A broad-host-range Flp-FRT recombination system for site-specific excision of chromosomally-located DNA sequences: application for isolation of unmarked Pseudomonas aeruginosa mutants. Gene 212:77-86. [PubMed]
18. Huber, B., K. Riedel, M. Hentzer, A. Heydorn, A. Gotschlich, M. Givskov, S. Molin, and L. Eberl. 2001. The cep quorum-sensing system of Burkholderia cepacia H111 controls biofilm formation and swarming motility. Microbiology 147:2517-2528. [PubMed]
19. Huber, B., K. Riedel, M. Kothe, M. Givskov, S. Molin, and L. Eberl. 2002. Genetic analysis of functions involved in the late stages of biofilm development in Burkholderia cepacia H111. Mol. Microbiol. 46:411-426. [PubMed]
20. Kothe, M., M. Antl, B. Huber, K. Stoecker, D. Ebrecht, I. Steinmetz, and L. Eberl. 2003. Killing of Caenorhabditis elegans by Burkholderia cepacia is controlled by the cep quorum-sensing system. Cell. Microbiol. 5:343-351. [PubMed]
21. Latifi, A., M. Foglino, K. Tanaka, P. Williams, and A. Lazdunski. 1996. A hierarchical quorum-sensing cascade in Pseudomonas aeruginosa links the transcriptional activators LasR and RhIR (VsmR) to expression of the stationary-phase sigma factor RpoS. Mol. Microbiol. 21:1137-1146. [PubMed]
22. Lerat, E., and N. A. Moran. 2004. The evolutionary history of quorum-sensing systems in bacteria. Mol. Biol. Evol. 21:903-913. [PubMed]
23. Lewenza, S., B. Conway, E. P. Greenberg, and P. A. Sokol. 1999. Quorum sensing in Burkholderia cepacia: identification of the LuxRI homologs CepRI. J. Bacteriol. 181:748-756. [PMC free article] [PubMed]
24. Lewenza, S., and P. A. Sokol. 2001. Regulation of ornibactin biosynthesis and N-acyl-l-homoserine lactone production by CepR in Burkholderia cepacia. J. Bacteriol. 183:2212-2218. [PMC free article] [PubMed]
25. Lewenza, S., M. B. Visser, and P. A. Sokol. 2002. Interspecies communication between Burkholderia cepacia and Pseudomonas aeruginosa. Can. J. Microbiol. 48:707-716. [PubMed]
26. LiPuma, J. J., T. Spilker, L. H. Gill, P. W. Campbell III, L. Liu, and E. Mahenthiralingam. 2001. Disproportionate distribution of Burkholderia cepacia complex species and transmissibility markers in cystic fibrosis. Am. J. Respir. Crit. Care Med. 164:92-96. [PubMed]
27. Lutter, E., S. Lewenza, J. J. Dennis, M. B. Visser, and P. A. Sokol. 2001. Distribution of quorum-sensing genes in the Burkholderia cepacia complex. Infect. Immun. 69:4661-4666. [PMC free article] [PubMed]
28. Mahenthiralingam, E., A. Baldwin, and P. Vandamme. 2002. Burkholderia cepacia complex infection in patients with cystic fibrosis. J. Med. Microbiol. 51:533-538. [PubMed]
29. Mahenthiralingam, E., D. A. Simpson, and D. P. Speert. 1997. Identification and characterization of a novel DNA marker associated with epidemic Burkholderia cepacia strains recovered from patients with cystic fibrosis. J. Clin. Microbiol. 35:808-816. [PMC free article] [PubMed]
30. Mahenthiralingam, E., P. Vandamme, M. E. Campbell, D. A. Henry, A. M. Gravelle, L. T. Wong, A. G. Davidson, P. G. Wilcox, B. Nakielna, and D. P. Speert. 2001. Infection with Burkholderia cepacia complex genomovars in patients with cystic fibrosis: virulent transmissible strains of genomovar III can replace Burkholderia multivorans. Clin. Infect. Dis. 33:1469-1475. [PubMed]
31. McDowell, A., E. Mahenthiralingam, K. E. Dunbar, J. E. Moore, M. Crowe, and J. S. Elborn. 2004. Epidemiology of Burkholderia cepacia complex species recovered from cystic fibrosis patients: issues related to patient segregation. J. Med. Microbiol. 53:663-668. [PubMed]
32. Meyer, J. M., and M. A. Abdallah. 1978. The fluorescent pigment of Pseudomonas fluorescens: biosynthesis, purification and physico-chemical properties. J. Gen. Microbiol. 107:319-328.
33. Mohr, C. D., M. Tomich, and C. A. Herfst. 2001. Cellular aspects of Burkholderia cepacia infection. Microbes Infect. 3:425-435. [PubMed]
34. Pesci, E. C., J. P. Pearson, P. C. Seed, and B. H. Iglewski. 1997. Regulation of las and rhl quorum sensing in Pseudomonas aeruginosa. J. Bacteriol. 179:3127-3132. [PMC free article] [PubMed]
35. Platt, T. B., B. Muller-Hill, and J. H. Miller. 1972. Analysis of the lac operon enzymes, p. 352-355. In J. H. Miller (ed.), Experiments in molecular genetics. Cold Spring Harbor Laboratory, Cold Spring Harbor, N.Y.
36. Riedel, K., C. Arevalo-Ferro, G. Reil, A. Gorg, F. Lottspeich, and L. Eberl. 2003. Analysis of the quorum-sensing regulon of the opportunistic pathogen Burkholderia cepacia H111 by proteomics. Electrophoresis 24:740-750. [PubMed]
37. Salanoubat, M., S. Genin, F. Artiguenave, J. Gouzy, S. Mangenot, M. Arlat, A. Billault, P. Brottier, J. C. Camus, L. Cattolico, M. Chandler, N. Choisne, C. Claudel-Renard, S. Cunnac, N. Demange, C. Gaspin, M. Lavie, A. Moisan, C. Robert, W. Saurin, T. Schiex, P. Siguier, P. Thebault, M. Whalen, P. Wincker, M. Levy, J. Weissenbach, and C. A. Boucher. 2002. Genome sequence of the plant pathogen Ralstonia solanacearum. Nature 415:497-502. [PubMed]
38. Sambrook, J., E. F. Fritsch, and T. Maniatis. 1989. Molecular cloning: a laboratory manual. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y.
39. Schuster, M., C. P. Lostroh, T. Ogi, and E. P. Greenberg. 2003. Identification, timing, and signal specificity of Pseudomonas aeruginosa quorum-controlled genes: a transcriptome analysis. J. Bacteriol. 185:2066-2079. [PMC free article] [PubMed]
40. Schweizer, H. P. 1992. Allelic exchange in Pseudomonas aeruginosa using novel ColE1-type vectors and a family of cassettes containing a portable oriT and the counter-selectable Bacillus subtilis sacB marker. Mol. Microbiol. 6:1195-1204. [PubMed]
41. Schwyn, B., and J. B. Neilands. 1987. Universal chemical assay for the detection and determination of siderophores. Anal. Biochem. 160:47-56. [PubMed]
42. Shaw, P. D., G. Ping, S. L. Daly, C. Cha, J. E. Cronan, Jr., K. L. Rinehart, and S. K. Farrand. 1997. Detecting and characterizing N-acyl-homoserine lactone signal molecules by thin-layer chromatography. Proc. Natl. Acad. Sci. USA 94:6036-6041. [PMC free article] [PubMed]
43. Smith, R. S., and B. H. Iglewski. 2003. P. aeruginosa quorum-sensing systems and virulence. Curr. Opin. Microbiol. 6:56-60. [PubMed]
44. Sokol, P. A., P. Darling, D. E. Woods, E. Mahenthiralingam, and C. Kooi. 1999. Role of ornibactin biosynthesis in the virulence of Burkholderia cepacia: characterization of pvdA, the gene encoding l-ornithine N5-oxygenase. Infect. Immun. 67:4443-4455. [PMC free article] [PubMed]
45. Sokol, P. A., D. E. Ohman, and B. H. Iglewski. 1979. A more sensitive plate assay for detection of protease production by Pseudomanas aeruginosa. J. Clin. Microbiol. 9:538-540. [PMC free article] [PubMed]
46. Sokol, P. A., U. Sajjan, M. B. Visser, S. Gingues, J. Forstner, and C. Kooi. 2003. The CepIR quorum-sensing system contributes to the virulence of Burkholderia cenocepacia respiratory infections. Microbiology 149:3649-3658. [PubMed]
47. Speert, D. P. 2002. Advances in Burkholderia cepacia complex. Paediatr. Respir. Rev. 3:230-235. [PubMed]
48. Speert, D. P., D. Henry, P. Vandamme, M. Corey, and E. Mahenthiralingam. 2002. Epidemiology of Burkholderia cepacia complex in patients with cystic fibrosis, Canada. Emerg. Infect. Dis. 8:181-187. [PMC free article] [PubMed]
49. Swift, S., J. A. Downie, N. A. Whitehead, A. M. Barnard, G. P. Salmond, and P. Williams. 2001. Quorum sensing as a population-density-dependent determinant of bacterial physiology. Adv. Microb. Physiol. 45:199-270. [PubMed]
50. Thompson, J. D., D. G. Higgins, and T. J. Gibson. 1994. CLUSTAL W: improving the sensitivity of progressive multiple sequence alignment through sequence weighting, position-specific gap penalties and weight matrix choice. Nucleic. Acids Res. 22:4673-4680. [PMC free article] [PubMed]
51. Ulrich, R. L., D. Deshazer, E. E. Brueggemann, H. B. Hines, P. C. Oyston, and J. A. Jeddeloh. 2004. Role of quorum sensing in the pathogenicity of Burkholderia pseudomallei. J. Med. Microbiol. 53:1053-1064. [PubMed]
52. Ulrich, R. L., D. Deshazer, H. B. Hines, and J. A. Jeddeloh. 2004. Quorum sensing: a transcriptional regulatory system involved in the pathogenicity of Burkholderia mallei. Infect. Immun. 72:6589-6596. [PMC free article] [PubMed]
53. Ulrich, R. L., H. B. Hines, N. Parthasarathy, and J. A. Jeddeloh. 2004. Mutational analysis and biochemical characterization of the Burkholderia thailandensis DW503 quorum-sensing network. J. Bacteriol. 186:4350-4360. [PMC free article] [PubMed]
54. Valade, E., F. M. Thibault, Y. P. Gauthier, M. Palencia, M. Y. Popoff, and D. R. Vidal. 2004. The PmlI-PmlR quorum-sensing system in Burkholderia pseudomallei plays a key role in virulence and modulates production of the MprA protease. J. Bacteriol. 186:2288-2294. [PMC free article] [PubMed]
55. Van de Peer, Y., and R. De Wachter. 1994. TREECON for Windows: a software package for the construction and drawing of evolutionary trees for the Microsoft Windows environment. Comput. Appl. Biosci. 10:569-570. [PubMed]
56. Wagner, V. E., D. Bushnell, L. Passador, A. I. Brooks, and B. H. Iglewski. 2003. Microarray analysis of Pseudomonas aeruginosa quorum-sensing regulons: effects of growth phase and environment. J. Bacteriol. 185:2080-2095. [PMC free article] [PubMed]
57. West, S. E., H. P. Schweizer, C. Dall, A. K. Sample, and L. J. Runyen-Janecky. 1994. Construction of improved Escherichia-Pseudomonas shuttle vectors derived from pUC18/19 and sequence of the region required for their replication in Pseudomonas aeruginosa. Gene 148:81-86. [PubMed]
58. Whitehead, N. A., A. M. Barnard, H. Slater, N. J. Simpson, and G. P. Salmond. 2001. Quorum-sensing in gram-negative bacteria. FEMS Microbiol. Rev. 25:365-404. [PubMed]
59. Woods, C. W., A. M. Bressler, J. J. LiPuma, B. D. Alexander, D. A. Clements, D. J. Weber, C. M. Moore, L. B. Reller, and K. S. Kaye. 2004. Virulence associated with outbreak-related strains of Burkholderia cepacia complex among a cohort of patients with bacteremia. Clin. Infect. Dis. 38:1243-1250. [PubMed]
60. Woods, D. 1984. Oligonucleotide screening of cDNA libraries. Focus 6:1-2.
61. Zhu, J., J. W. Beaber, M. I. More, C. Fuqua, A. Eberhard, and S. C. Winans. 1998. Analogs of the autoinducer 3-oxooctanoyl-homoserine lactone strongly inhibit activity of the TraR protein of Agrobacterium tumefaciens. J. Bacteriol. 180:5398-5405. [PMC free article] [PubMed]

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