FREE
This Article
FREE
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Supplemental material
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Services
Right arrow Related articles in JB
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 arrowReprints and Permissions
Right arrow Copyright Information
Right arrow Books from ASM Press
Right arrow MicrobeWorld
Citing Articles
Right arrow Citing Articles via HighWire
Google Scholar
Right arrow Articles by Shikuma, N. J.
Right arrow Articles by Yildiz, F. H.
PubMed
Right arrow PubMed Citation
Right arrow Articles by Shikuma, N. J.
Right arrow Articles by Yildiz, F. H.

 Previous Article  |  Next Article 

Journal of Bacteriology, July 2009, p. 4082-4096, Vol. 191, No. 13
0021-9193/09/$08.00+0     doi:10.1128/JB.01540-08
Copyright © 2009, American Society for Microbiology. All Rights Reserved.

Identification and Characterization of OscR, a Transcriptional Regulator Involved in Osmolarity Adaptation in Vibrio cholerae{triangledown} ,{dagger}

Nicholas J. Shikuma and Fitnat H. Yildiz*

Department of Microbiology and Environmental Toxicology, University of California, Santa Cruz, Santa Cruz, California 95064

Received 30 October 2008/ Accepted 21 March 2009


arrow
ABSTRACT
 
Vibrio cholerae is a facultative human pathogen. In its aquatic habitat and as it passes through the digestive tract, V. cholerae must cope with fluctuations in salinity. We analyzed the genome-wide transcriptional profile of V. cholerae grown at different NaCl concentrations and determined that the expression of compatible solute biosynthesis and transporter genes, virulence genes, and genes involved in adhesion and biofilm formation is differentially regulated. We determined that salinity modulates biofilm formation, and this response was mediated through the transcriptional regulators VpsR and VpsT. Additionally, a transcriptional regulator controlling an osmolarity adaptation response was identified. This regulator, OscR (osmolarity controlled regulator), was found to modulate the transcription of genes involved in biofilm matrix production and motility in a salinity-dependent manner. oscR mutants were less motile and exhibited enhanced biofilm formation only under low-salt conditions.


arrow
INTRODUCTION
 
The halotolerant bacterium Vibrio cholerae causes the disease cholera. V. cholerae inhabits aquatic environments, and its occurrence strongly correlates with salinity (15, 24, 33, 37). Interannual climate variation is thought to have a significant impact on the environmental conditions governing V. cholerae occurrence and the incidence of cholera outbreaks (29, 47, 52). Studies have shown that cholera outbreaks occur seasonally and correlate with the monsoon in areas where cholera is endemic (11, 20, 46). Furthermore, seasonal fluctuations in rainfall as well as changes in sea surface height brought by tides and storm surges can cause wide variations in salinity concentrations, which affect the growth and distribution of the pathogen (15, 20, 57). Salinity is therefore a significant driving force in the occurrence of V. cholerae and cholera outbreaks.

Microcosm experiments show that the growth and survival of V. cholerae are dependent on salinity, and the optimal salt concentration for growth is similar to salinities of brackish water and estuarine environments (38, 56, 57). Additional factors synergize with salinity to affect the growth and survival of V. cholerae. Increased concentrations of organic nutrients can compensate for a growth defect caused by high or low salinity concentrations (25, 38, 57). The organic carbon level was shown to be a key parameter in the growth of V. cholerae in natural freshwater (61) and seawater (43) microcosms. Additionally, lower temperatures (≤15 to 20°C) were shown to slow or stop the growth of V. cholerae if salinity concentrations were above or below an optimal level (56). Adaptation to high-salinity conditions in many microorganisms involves the synthesis and transport of compatible solutes, which act to stabilize intracellular levels of water and turgor pressure without disturbing cellular function (6). V. cholerae can synthesize the compatible solute ectoine by using a four-gene operon which includes ectABC and a putative aspartokinase gene (49). V. cholerae can also import the compatible solutes proline and glycine betaine by using two transporters, OpuD and PutP (27). Multiple environmental and cellular factors can therefore influence the growth and persistence of V. cholerae in aquatic environments.

In addition to salinity changes in the aquatic environment, V. cholerae must cope with fluctuations in salinity and osmolarity as it is ingested, adheres to the epithelial lining of the intestinal tract, produces cholera toxin, inducing the production of large volumes of watery diarrhea, and is flushed into the environment. It was hypothesized that osmolarity might be one signal triggering the expression of virulence factors when V. cholerae is within the human gut (40). Indeed, the intestinal lumen is thought to have an osmolarity equivalent to 0.3 M NaCl or higher (19), while rice water stool has a sodium concentration of approximately 0.14 M and an osmolarity of approximately 0.30 osM (35). As expected, virulence factors in V. cholerae are regulated at multiple levels in response to salinity/osmolarity (36). Furthermore, genes induced in the latter stages of infection are known to help V. cholerae to survive in stool and pond water, indicating that the bacterium might be programmed to endure environments of low osmotic pressures after intestinal passage (54). V. cholerae is therefore subject to a range of salinities throughout its infectious cycle.

In this study, we analyzed the genome-wide transcriptional profile of V. cholerae grown at different NaCl concentrations and determined that the expression of compatible solute biosynthesis and transporter genes, virulence genes, and genes involved in adhesion and biofilm formation is differentially regulated. We also determined that the expression of a set of regulatory genes is controlled in a salinity/osmolarity-dependent manner. Mutational analysis and phenotypic characterization of one such regulator, OscR, revealed that OscR regulates the expression of genes involved in biofilm formation and motility in an osmolarity-dependent manner. Whole-genome expression profiling of V. cholerae grown in different salinities therefore revealed one mechanism by which this halotolerant bacterium regulates salinity/osmolarity adaptation.


arrow
MATERIALS AND METHODS
 
Bacterial strains, plasmids, and culture conditions. The bacterial strains and plasmids used in this study are listed in Table 1. All V. cholerae and Escherichia coli strains were grown aerobically, at 30°C and 37°C, respectively. Growth medium consisted of Luria-Bertani (1% tryptone, 0.5% yeast extract) medium (LB), pH 7.5, supplemented with 0, 0.1, 0.2, 0.3, 0.4, or 0.5 M NaCl (75, 257, 449, 625, or 1,024 mosmol/kg, respectively), 0.2 or 0.5 M KCl (447 or 1,007 mosmol/kg, respectively), or 0.1, 0.2, 0.3, 0.4 or 0.5 M D-lactose (184, 292, 406, 526, or 698 mosmol/kg, respectively). D-Lactose did not promote the growth of our prototype V. cholerae strain (FY_Vc_1) when provided at 0.5% in the defined artificial seawater (ASW) medium described below and was therefore used as a nonmetabolizable osmolyte. The growth rate and yield of V. cholerae were similar in LB medium of different salinities. LB-agar and LB-soft agar plates contained 1.5% (wt/vol) and 0.3% (wt/vol) granulated agar (Difco), respectively. Concentrations of antibiotics used were as follows, except that medium used for β-galactosidase assays was supplemented with 50 µg/ml ampicillin: ampicillin, 100 µg/ml; rifampin (rifampicin), 100 µg/ml; and gentamicin, 30 µg/ml. ASW used in this study (adapted from references 34 and 51) contained the following components, regardless of the salt base concentration: 50 mM Tris (pH 7.5), 18.5 mM NH4Cl, 10 mM CaCl2, 0.333 mM K2HPO4, 0.5% DL-lactate, 1x MEM vitamin solution (Mediatech), and trace metals (1 ml/liter of 5% MgSO4, 0.5% MnCl2·4H2O, 0.5% FeCl3, 0.4% trinitriloacetic acid) (7). The salt base contained 300 mM NaCl, 50 mM MgSO4, and 10 mM KCl, and these components were used at 5%, 10%, 50%, and 75% concentrations (283, 313, 573, and 715 mosmol/kg, respectively). The growth rate and yield of V. cholerae were similar in different ASW-based media. The low-osmolarity minimal medium (LO-MM) used in this study contained 50 mM MgSO4, 10 mM CaCl2, 0.333 mM K2HPO4, trace metals (defined above), MEM vitamin solution (Mediatech), 0.2% DL-lactate, 0.1% L-asparagine, 0.1% L-serine, 0.1% L-arginine, and 0.1% L-glutamic acid, and the pH was adjusted to 7.5 with NaOH. Minimal medium was used without added NaCl or with 0.2 M NaCl (96 or 444 mosmol/kg, respectively). The growth rate and yield of V. cholerae were similar in LO-MM with and without NaCl. Osmolalities of LB, ASW, and LO-MM used in this study were measured using a vapor pressure osmometer (model 5100B; Wescor).


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

  		TABLE 1. Bacterial strains and plasmids used in this study

Recombinant DNA techniques. DNA manipulations were carried out by standard molecular techniques (53). Restriction and DNA modification enzymes were obtained from New England Biolabs. PCRs were carried out using primers purchased from Operon Technologies and the Expand high-fidelity PCR system (Roche). Primers used in this study are listed in Table 2. DNA sequencing was carried out by the UC Berkeley DNA Sequencing Facility.


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

  		TABLE 2. Primers used in this study

Generation of in-frame deletion and insertion mutants. Deletion and insertion mutants were generated in the V. cholerae wild-type strain (FY_Vc_1) according to previously published protocols (14, 16, 40). Sequences of the constructed plasmids were verified by DNA sequencing. Deletion and insertion mutants in V. cholerae were verified by PCR.

Creation of single-copy chromosomal promoter-lacZ fusions. We created three promoter-lacZ fusions that were inserted into the lacZ locus (VC2338). To this end, regions flanking the lacZ gene were amplified. The 5' region was amplified using primers knockin_a and knockin_b, and the 3' region was amplified using primers knockin_c and knockin_d. The two products were joined by splicing by overlap extension PCR (30) and inserted into the pGP704-sacB28 vector, generating pknockin. Promoter regions were amplified by PCR, ligated to the lacZ gene derived from pBAD102/D/lacZ (Invitrogen), and cloned into pknockin. The promoter-lacZ fusions were inserted in the reverse orientation into the endogenous lacZ locus to ensure that no expression was derived from the native promoter. Insertions were then carried out as described above.

Generation of gfp-tagged strains. V. cholerae deletion strains were tagged with the green fluorescent protein gene (gfp) according to a previously described procedure (12). Briefly, triparental matings were carried out with donor E. coli S17-1 ({lambda} pir) carrying pMCM11, helper E. coli S17-1 ({lambda} pir) harboring pUX-BF13, and V. cholerae deletion strains. Transconjugants were selected on thiosulfate-citrate-bile salts-sucrose (Difco) agar medium containing gentamicin at 30°C. gfp-tagged V. cholerae strains were verified by PCR and used in flow cell experiments.

β-Galactosidase assays. For V. cholerae cultures grown in LB medium, strains were grown overnight aerobically in LB containing 0.2 M NaCl, diluted 1:200 in LB containing the indicated NaCl, KCl, or lactose concentration to an optical density at 600 nm (OD600) of 0.3 to 0.4, and diluted again 1:200 in LB containing the corresponding addition. The assay was then performed once cultures reached an OD600 of 0.3 to 0.4. Strains that required antibiotics for plasmid maintenance were grown overnight in medium supplemented with 100 µg/ml ampicillin and grown to mid-exponential phase in medium containing 50 µg/ml ampicillin. For cultures grown in ASW medium, V. cholerae strains were grown overnight aerobically in 50% ASW medium and then diluted 1:200 in ASW containing the indicated ASW salt concentration. The assay was then performed when the culture reached an OD600 of 0.1 to 0.2. β-Galactosidase assays were carried out in MultiScreen 96-well microtiter plates fitted onto a MultiScreen filtration system (Millipore) or in 2- to 4-ml aliquots, using previously published procedures (12, 13) similar to that described by Miller (39). The assays were conducted with at least four technical replicates per strain and repeated with at least three biological replicates. Results for one representative biological replicate are shown.

Initial colonization on glass surfaces. The initial colonization of V. cholerae on glass surfaces was analyzed in cultures grown in LB supplemented with 0, 0.1, 0.2, or 0.5 M NaCl. Overnight cultures grown in LB medium supplemented with various NaCl concentrations were diluted to an OD600 of 0.10 in fresh LB supplemented with the corresponding NaCl concentration. Two-milliliter aliquots of the diluted culture were then inoculated into a Lab-Tek II chambered cover glass system (Nunc). Phase-contrast images of cells on the glass surface were captured using a Zeiss Axiovert 200 inverted microscope fitted with a x63 objective and with Optovar magnification of x1.6, using AxioCam MRm and AxioVision software, version 4.6.3.0 (Zeiss). Images of cells on the glass surface were captured 3, 30, and 60 min after initial inoculation. The number of adhered cells per field of view was determined at each time interval, with at least three biological replicates for each treatment.

RNA isolation. For cultures grown in LB medium, V. cholerae cells were grown aerobically overnight in LB supplemented with 0.2 M NaCl. Cultures were diluted 1:200 in fresh LB medium containing the indicated concentrations of NaCl, KCl, or lactose and grown aerobically at 30°C with shaking at 200 rpm to an OD600 of 0.3 to 0.4. To obtain a homogenous population of exponential-phase cells, mid-exponential-phase cultures were diluted 1:200 once more in fresh medium containing the corresponding NaCl concentration and harvested at an OD600 of 0.3 to 0.4. For cultures grown in ASW medium, V. cholerae cells were grown aerobically overnight in 50% ASW. Cultures were diluted 1:200 in fresh medium containing the indicated ASW concentration and grown aerobically at 30°C with shaking at 200 rpm to an OD600 of 0.1 to 0.2. For cultures grown in LO-MM, V. cholerae cells were grown aerobically overnight in LO-MM with 0.2 M NaCl. Cultures were diluted 1:200 in fresh medium containing the indicated NaCl concentration and grown aerobically at 30°C with shaking at 200 rpm to an OD600 of 0.1 to 0.2. Two-milliliter aliquots of cultures in LB, ASW, or LO-MM were collected by centrifugation for 2 min at room temperature. Cell pellets were immediately resuspended in 1 ml Trizol reagent (Invitrogen) and stored at –80°C. Total RNA was isolated from the cell pellets according to the manufacturer's instructions (Invitrogen). Contaminating DNA was removed by incubating RNA with RNase-free DNase I (Ambion), and an RNeasy Mini kit (Qiagen) was used to clean up RNA after DNase digestion.

Gene expression profiling. Microarrays used in this study were composed of spotted 70-mer oligonucleotides representing the open reading frames present in the V. cholerae strain N16961 genome and were printed at the University of California, Santa Cruz (3). Whole-genome expression analyses were performed using a common reference RNA. Reference RNA was obtained by growing V. cholerae overnight in LB supplemented with 0.2 M NaCl, diluting the culture 1:200 in fresh medium, growing it to mid-exponential phase at an OD600 of 0.3 to 0.4, diluting it 1:200 again, and harvesting samples at mid-exponential phase at an OD600 of 0.3 to 0.4. RNAs from the test and reference samples were subjected to cDNA synthesis, microarray hybridization, and scanning as described previously (3). Normalized signal ratios were obtained with LOWESS print-tip normalization, using the Bioconductor packages (18) in the R environment. Differentially regulated genes were determined using three biological replicates and two technical replicates for each treatment (six data points for each spot), using the Significance Analysis of Microarrays (SAM) program (59), with a 1.5- or 2-fold difference in gene expression and a 3% false discovery rate (FDR) as cutoff values.

qPCR. Quantitative PCR (qPCR) was performed by first synthesizing cDNA from 0.25 to 1 µg of each RNA sample, using an iScript cDNA synthesis kit (Bio-Rad). The product was then diluted 1:4 with water, and 4 µl was used as a template with 12 pmol of each primer in a subsequent PCR using the Expand high-fidelity PCR system (Roche). PCR conditions were as follows: 94°C for 2 min; 25 cycles of 94°C for 30 s, 55°C for 30 s, and 72°C for 30 s; and a final step at 72°C for 2 min. The amplified products were analyzed in a 1.5% agarose gel and quantified using ImageQuant software (Molecular Dynamics). The intensity of each DNA band was normalized to that of the corresponding gyrA band. At least three biological replicates were conducted for each treatment tested, and reaction mixtures lacking reverse transcriptase were used as negative controls.

Flow cell experiments and CLSM. Flow cell experiments were carried out according to a previously described procedure (22). Briefly, chambers were sterilized with 0.5% (vol/vol) hypochlorite overnight, followed by sterile MilliQ water and full-strength (10 g/liter tryptone, 5 g/liter yeast extract) or 2% (0.2 g/liter tryptone, 0.1 g/liter yeast extract) LB supplemented with 0, 0.1, 0.2, 0.3, or 0.5 M NaCl at a flow rate of 4.5 ml/h. Overnight cultures of gfp-tagged V. cholerae strains were diluted to an OD600 of 0.1, and 350-µl aliquots of the diluted cultures were inoculated by injection into flow cell chambers. After inoculation, the chambers were allowed to stand inverted, with no flow, for 1 h. Flow was resumed at a rate of 4.5 ml/h with the chambers standing upright. Flow cell experiments were carried out at room temperature. Confocal laser scanning microscopy (CLSM) images of the biofilms were captured with an LSM 5 Pascal system (Zeiss), using 488-nm excitation and 543-nm emission wavelengths. Three-dimensional images of the biofilms were reconstructed using Imaris software (Bitplane) and quantified using COMSTAT (23). Flow cell experiments were carried out with at least two biological replicates.

Motility assays. LB-soft agar (0.3% agar), with and without 0.2 M NaCl (supplemented with 100 µg/ml ampicillin to maintain plasmids when appropriate), was inoculated from a colony grown overnight on LB agar at 30°C. After incubation for 18 to 20 h at 25°C, the migration zone diameter was measured and compared between strains.


arrow
RESULTS AND DISCUSSION
 
Overview of the transcriptional profile of V. cholerae grown in different salinities. To determine the effects of different salinities on the V. cholerae transcriptional profile, cells were grown to mid-exponential phase in LB medium containing 0, 0.1, 0.2, 0.3, or 0.5 M NaCl. Transcriptional profiling was conducted on these cells, and data were analyzed using SAM software, with a ≥2-fold change in gene expression and an FDR of ≤3% as criteria (59). A total of 333 genes were significantly differentially regulated in response to NaCl when pairwise comparisons were conducted between expression profiles of V. cholerae cultures grown at each salinity concentration (Fig. 1; see Tables S1 and S2 in the supplemental material). Of these genes, 12.3% were annotated as metabolism-related genes, 23.1% were involved in cellular processes, and 42.9% were hypothetical (see Table S1 in the supplemental material).


Figure 1
View larger version (36K):
[in this window]
[in a new window]

  		FIG. 1. Expression profiles of V. cholerae cells grown in 0, 0.1, 0.2, 0.3, and 0.5 M NaCl. Clustered expression profiles of 333 differentially expressed genes are given on a log2-based color scale. Differentially expressed genes were identified using SAM software by making pairwise comparisons of cells grown at each salinity (with criteria of a ≥2-fold change in gene expression and an FDR of ≤3%). Profiles are ratios of gene expression in cells grown in LB with 0.1 M, 0.2 M, 0.3 M, and 0.5 M NaCl relative to that in cells grown with 0 M NaCl. Yellow segments, induced expression; blue segments, repressed expression. Expression profiles are shown separately for genes encoding Na+/H+ antiporters (A), compatible solute biosynthesis/transport proteins (B), toxin-coregulated pilus proteins (C), cholera toxin (D), outer membrane proteins OmpU and OmpT (E), VPS and biofilm matrix proteins (F), membrane-derived oligosaccharides (G), polyamine biosynthesis proteins (H), and the OscR regulator (I).

The expression profiles of significantly differentially expressed genes could be grouped into three classes according to their responses to salinity (Fig. 1). Class I genes were upregulated at high salinities, class II genes exhibited their highest expression at middle salinities (0.1 M to 0.2 M NaCl), and class III genes were induced at low salinities. The relationship between salinity and these genes and their associated cellular processes are discussed below.

Na+/H+ antiporter and compatible solute biosynthesis/transport class I genes are regulated by osmolarity. The V. cholerae genome encodes multiple characterized and putative Na+/H+ antiporters. In Escherichia coli, Na+/H+ antiporters establish an electrochemical Na+ gradient across the membrane, protect against Na+ toxicity, and regulate intracellular pH (44, 45, 50). In our studies, three antiporter genes exhibited greater expression in cells grown in media with higher NaCl concentrations and were grouped as class I genes (Fig. 1A). These included nhaA (VC1627), VCA0193, and VC2037. Expression of nhaA was previously shown to be Na+, Li+, and K+ concentration dependent (60), and a similar trend was observed in the present study, where expression increased between 0 M NaCl and 0.5 M NaCl (1.7-fold). Expression of VCA0193 also increased as salinity increased. VC2037 is homologous to yqkI of Bacillus subtilis (63) and was annotated as yqkI by The Institute of Genomic Research (TIGR). Expression of VC2037 was increased at 0.2 M NaCl (1.9-fold) and 0.5 M NaCl (2.8-fold) relative to that at 0 M NaCl.

Exposure to high salinity increased the transcription of genes required for compatible solute transport and production. V. cholerae can import compatible solutes by using two transporters, OpuD and PutP (27). The expression of putP (VCA1071) was not modulated in response to salinity, but the expression of opuD (VC1279) increased with increasing concentrations of NaCl (Fig. 1B). The compatible solute biosynthesis genes for ectoine, ectABC (VCA0825 to VCA0823), and a putative aspartokinase gene (VCA0822) increased in expression as the NaCl concentration increased (Fig. 1B). The deletion of ectA was previously shown to negatively impact V. cholerae growth in high-osmolarity media (49), and our results are congruent with the notion that V. cholerae utilizes compatible solutes under high-salinity conditions.

To confirm these transcriptome results, we constructed strains containing a chromosomal ectA or VC2037 promoter-lacZ transcriptional fusion (ectAp-lacZ or VC2037p-lacZ) and grew these cells at various concentrations of osmolytes. Expression of ectAp-lacZ increased as NaCl, KCl, or lactose was added to LB medium (Fig. 2A), indicating that ectA expression is dependent on osmolarity and not on NaCl or ionic strength specifically. Since V. cholerae is a natural inhabitant of aquatic environments, we wondered whether the expression of ectA would be similar in a defined medium containing different concentrations of seawater salts (ASW). As expected, the expression of ectA positively correlated with increased salinity in ASW (Fig. 2B). Similarly, VC2037 expression increased in response to added NaCl, KCl, or lactose and in response to increased ASW salinity (Fig. 2C and D). These results support the transcriptome data and give us confidence that our genome-wide analysis identified genes important for salinity/osmolarity adaptation in V. cholerae.


Figure 2
View larger version (27K):
[in this window]
[in a new window]

  		FIG. 2. Osmolarity regulates ectA and VC2037 expression. A comparison of ectA and VC2037 expression is shown, using β-galactosidase assays with chromosomal ectAp-lacZ and VC2037p-lacZ reporter fusion strains grown to mid-exponential phase in various osmolytes. ectAp-lacZ (A) and VC2037p-lacZ (C) reporter strains were grown in NaCl, KCl, or lactose at the indicated concentrations. ectAp-lacZ (B)- and VC2037p-lacZ (D)-containing strains were also grown in ASW at the indicated concentrations. The results of one representative experiment with three biological replicates are shown. Error bars indicate standard deviations for eight technical replicates.

Class II genes. (i) Pathogenesis-related genes are modulated by NaCl concentration. Multiple genes associated with pathogenesis were found to be expressed differentially when V. cholerae was grown in medium supplemented with different NaCl concentrations, and these fell into the class II gene category, where genes were upregulated under mid-salinity conditions. Expression of two main virulence factors of V. cholerae, cholera toxin and toxin-coregulated pilus (TCP), is well known to be controlled osmotically in the classical biotype (17, 40, 41). In the V. cholerae O1 El Tor strain used here, genes necessary for TCP biogenesis (tcp), cholera toxin subunit A and B production (ctxA and ctxB), and associated regulatory proteins (tcpP, tcpH, and toxT) showed maximal expression in 0.2 M NaCl (Fig. 1C and D). In a complex regulatory network, multiple proteins regulate the expression of ctx and tcp genes (reviewed in reference 36). Differences in osmolarity are thought to be sensed by the periplasmic domain of the transmembrane transcriptional regulator ToxR, which binds to the promoter of ctxAB (48) and regulates expression in an osmolarity-dependent manner (41). The ToxR-regulated ompU (VC0633) and ompT (VC1854) genes were also differentially regulated in our transcriptional profiling experiments (Fig. 1E), and these genes were shown previously to be regulated in response to salinity (40).

(ii) vps gene expression is modulated by salinity and osmolarity. The production of extracellular matrix components is essential for mature biofilm formation in V. cholerae, and these components act to bind cells to each other and to the surface. One component of the extracellular matrix is Vibrio polysaccharide (VPS), which is required for mature biofilm formation in V. cholerae (62, 66). The vps genes are organized into vps I (vpsU, vpsA to vpsK, VC0916, and VC0917 to VC0927) and vps II (vpsL to vpsQ and VC0934 to VC0939) coding regions, separated by an intergenic region containing six additional open reading frames (rbmABCDEF; VC0928 to VC0933) (12, 14, 66). We determined that the expression of vps and rbm genes was differentially regulated by salinity (Fig. 1F). vps and rbm gene expression was highest at median salinity, and the genes were therefore grouped into the class II gene category.

To verify these microarray results, a strain containing a vpsL (the first gene in the vps II cluster) promoter-lacZ fusion inserted into the lacZ locus (vpsLp-lacZ) was created. When this strain was grown to mid-exponential phase in LB medium supplemented with 0, 0.1, 0.2, 0.3, or 0.5 M NaCl, β-galactosidase activity was greatest at 0.1 M NaCl and decreased at the other NaCl concentrations tested (Fig. 3A). To determine whether the differential expression of vpsL was in response to NaCl specifically or in response to medium osmolarity, the vpsLp-lacZ reporter strain was grown to mid-exponential phase in LB supplemented with 0.1, 0.2, 0.3, 0.4, or 0.5 M lactose, without NaCl. β-Galactosidase activity was greatest at 0.1 M lactose (Fig. 3B). These results indicate that medium osmolarity, not NaCl specifically, causes the modulation in vpsL expression. We also analyzed the expression of vps genes in a defined ASW medium which mimics the natural aquatic environment of V. cholerae. Similar to growth in LB medium, expression of vpsL was greatest at a median 10% ASW concentration (Fig. 3C).


Figure 3
View larger version (116K):
[in this window]
[in a new window]

  		FIG. 3. Osmolarity affects vps expression and biofilm architecture. A comparison of vpsL expression was performed using β-galactosidase assays of a strain harboring a chromosomal vpsLp-lacZ reporter grown to mid-exponential growth phase in NaCl (A), lactose (B), or ASW (C) at the indicated concentrations. Results for one representative experiment with three biological replicates are shown. Error bars indicate standard deviations for eight technical replicates. (D) CLSM images of horizontal (xy) and vertical (xz) projections of biofilm structures formed by wild-type V. cholerae in a flow cell system with 2% LB supplemented with 0.1, 0.2, 0.3 and 0.5 M NaCl at 24, 48, and 72 h postinoculation. Bars, 30 µm.

(iii) Biofilms are modulated by salinity. The differential expression of vps genes led us to hypothesize that biofilm formation would be different between cells grown at different NaCl concentrations. We therefore grew V. cholerae biofilms in a once-through flow cell system in 0.1, 0.2, 0.3, and 0.5 M NaCl. Flow cell systems are normally conducted with 2% LB to facilitate the observation of biofilm development. Cells grown in 2% LB without added NaCl in a flow cell system were not viable due to low nutrient concentrations of the medium, and this treatment was excluded from this experiment. As expected, biofilm architecture was markedly different when biofilms were grown using media at different salinities. Differences in biofilm structure were slightly apparent at 24 h postinoculation, where biofilms appeared more heterogeneous in lower-salinity media (Fig. 3D). At 48 and 72 h postinoculation, more dramatic changes in biofilm architecture took place. COMSTAT analysis revealed that the roughness coefficient, an indicator of how much the biofilm thickness varies, was greater for 0.2 M NaCl than for 0.5 M NaCl (1.7-fold) after 72 h (Table 3). Biomass and average thickness were 1.7- and 1.5-fold greater, respectively, in 0.5 M NaCl than in 0.2 M NaCl at 72 h. These results indicate that V. cholerae biofilm formation is modulated by salinity.


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

  		TABLE 3. COMSTAT analysis of biofilm structures formed by wild-type V. cholerae in 0.2 M and 0.5 M NaCl-containing 2% LBa

The two-component signal transduction systems CpxA-CpxR and EnvZ-OmpR are known osmolarity-responsive signal transduction systems, and both systems regulate biofilm formation in E. coli (26). We therefore hypothesized that the CpxA-CpxR and EnvZ-OmpR systems might affect biofilm formation of V. cholerae in an osmolarity-dependent manner. Biofilm formation in once-through flow cell systems of cpxA (VC2693) and envZ (VC2713) in-frame deletion mutants showed only minor differences compared to the wild type grown in LB medium supplemented with 0.2 M NaCl and 0.5 M NaCl (see Fig. S1 in the supplemental material). These results indicate that CpxA and EnvZ play a minimal role in biofilm formation in V. cholerae under the conditions examined.

Genes encoding proteins involved in modulating cell surface properties of V. cholerae are important for adhesion to and biofilm formation on biotic and abiotic surfaces. Because many genes that are important for cell surface properties were differentially expressed in response to salinity, we hypothesized that initial attachment properties of V. cholerae cells would also differ as the medium salinity was varied. To test this hypothesis, we analyzed surface attachment to a glass substratum over 60 min. As the NaCl concentration of the medium increased, attachment rates of cells increased (see Fig. S2 in the supplemental material). Sixty minutes after exposure to the surface, the number of attached cells at 0 M NaCl was increased at 0.1 M (7.3-fold), 0.2 M (10.1-fold), and 0.5 M (11.2-fold) NaCl. These results indicate that attachment rates are greater for cells grown at higher NaCl concentrations, and cell surface properties might be important for these effects.

(iv) Salinity modulates vps expression through VpsR and VpsT. Since vps gene expression is regulated by salinity, we then questioned whether previously identified regulators of vps genes were responsible for modulating expression in response to extracellular salinity/osmolarity. Two positive regulators of vps expression are VpsR and VpsT (9, 64). While VpsR is essential for the expression of vps genes and the formation of a typical three-dimensional biofilm structure (64), VpsT plays an accessory role and modulates vps gene expression and biofilm formation (2, 9).

Microarray analysis revealed that only vpsT, not vpsR, was significantly differentially expressed in response to salinity (Fig. 1F). Expression of vpsT was increased at 0.2 M (2.6-fold), 0.3 M (2.4-fold), and 0.5 M NaCl (3.4-fold) relative to that at 0 M NaCl. To confirm the array results, the transcriptional modulation of vpsR and vpsT was analyzed using qPCR. Message levels of vpsR were similar across all NaCl treatments, paralleling the microarray results (Fig. 4A). vpsT levels were lowest in 0 M NaCl and were upregulated in 0.1 M, 0.2 M, 0.3 M, and 0.5 M NaCl. These results indicate that salinity does not modulate the level of vpsR but does modulate the level of vpsT.


Figure 4
View larger version (124K):
[in this window]
[in a new window]

  		FIG. 4. Salinity modulates vpsL expression and biofilm formation through VpsT and VpsR. (A) qPCR analysis of gyrA, vpsR, and vpsT message levels in wild-type cells grown to mid-exponential phase in LB supplemented with NaCl at the indicated concentrations. Gel panels are representative images of qPCRs. cDNA synthesis reactions without reverse transcriptase (RT) were included as controls. The graph shows averages for three biological replicates of vpsR/gyrA and vpsT/gyrA message levels, and error bars indicate standard deviations. (B) vpsL expression of wild-type, {Delta}vpsR, and {Delta}vpsT strains harboring a chromosomal vpsLp-lacZ fusion. β-Galactosidase activity was measured in cells grown to mid-exponential phase at the indicated salinities. Results of one representative experiment with three biological replicates are shown. Error bars indicate standard deviations for eight technical replicates. (C) CLSM images of horizontal (xy) and vertical (xz) projections of biofilm structures formed by wild-type, {Delta}vpsR, and {Delta}vpsT strains in a flow cell system with 2% LB supplemented with 0.2 and 0.5 M NaCl at 24, 48, and 72 h postinoculation. Bars, 30 µm.

To further characterize how these regulators might influence vps expression at different salinities, we deleted the vpsR or vpsT gene in the strain carrying the chromosomal vpsLp-lacZ fusion. The strains were grown in 0 M to 0.5 M NaCl, and β-galactosidase activity was measured when cells reached mid-exponential phase. While the {Delta}vpsR mutant showed little to no expression at all salinities examined, the {Delta}vpsT mutant showed decreased vpsL expression compared to the wild type at all salinities tested (Fig. 4B). vpsR is the most downstream regulator of vps gene expression identified to date (2), and vpsR is also essential for vps expression at different salinities. Although not essential, vpsT also positively regulates vps expression. The {Delta}vpsT mutant exhibited less vps expression, but in this mutant vps expression was still modulated by salinity.

The quorum sensing master regulator, HapR (65, 67), the alternative sigma factor RpoS (65), and the cyclic AMP-CRP complex (32) were previously shown to negatively regulate vps expression. To determine whether these regulators were important for vps expression at different salinities, we analyzed vpsL expression, using a plasmid-borne vpsLp-lacZ transcriptional fusion, in hapR, rpoS, and crp mutants in both 0.1 M and 0.5 M NaCl. Similar to the wild-type case, vpsL expression was higher in cells grown in 0.1 M NaCl than in cells grown in 0.5 M NaCl (data not shown), indicating that these regulators do not regulate vps expression in a salinity/osmolarity-dependent manner under the conditions tested.

(v) Salinity modulates biofilm formation through VpsR and VpsT. Transcription of vps genes is positively regulated by VpsR and VpsT (2, 9, 64). We therefore hypothesized that vpsR or vpsT mutations would modulate the effects of salinity on the development of a mature biofilm structure. To test this hypothesis, biofilms of {Delta}vpsR and {Delta}vpsT mutants were grown in 2% LB supplemented with 0.2 M or 0.5 M NaCl in a flow cell system, and biofilm images were captured at 24, 48, and 72 h (Fig. 4C). COMSTAT analyses revealed that total biomass, average thickness, and maximum thickness were markedly lower for biofilms of both the {Delta}vpsT and {Delta}vpsR mutants than for wild-type biofilms in both 0.2 M and 0.5 M NaCl (Table 4). It should be noted that the transcription of vps genes, and hence VPS production, was impaired in the vpsR mutant. Thus, for the vpsR mutant, changes in biofilm structure due to VPS production in response to salinity could not be evaluated. For the vpsR mutant, surface coverage was decreased and the roughness coefficient was increased for biofilms formed in 0.5 M NaCl compared to those formed in 0.2 M NaCl. For the vpsT mutant, total biomass was increased and the roughness coefficient was decreased for biofilms formed in 0.5 M NaCl compared to those formed in 0.2 M NaCl. Taken together, these results suggest that VpsR and VpsT are involved in the modulation of biofilm formation in response to salinity.


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

  		TABLE 4. COMSTAT analysis of biofilm structures formed by V. cholerae wild-type, {Delta}vpsR, and {Delta}vpsT strains in 0.2 M and 0.5 M NaCl-containing 2% LBa

Class III genes. (i) MDO and polyamine production genes are regulated by salinity. One set of genes whose expression was increased at low salinity were the mdoG and mdoH genes (VC1288 and VC1287). Both mdoG and mdoH exhibited maximal expression in cells grown in 0 M NaCl and sequentially decreased as salinity increased (Fig. 1G). MdoG and MdoH mediate the biosynthesis of membrane-derived oligosaccharides (MDOs), and their activity in other bacteria is known to be regulated osmotically at the transcriptional and/or posttranscriptional level (5). MDOs may play a structural role in the cell envelope and were shown previously to affect biofilm formation in an osmolarity-dependent manner in Pseudomonas aeruginosa (31).

Two genes involved in the biosynthesis of polyamines, speA and speB (VCA0815 and VCA0814), were also expressed at high levels in cells grown at 0 M NaCl, and their expression decreased as salinity increased (Fig. 1H). SpeA is an arginine decarboxylase responsible for the conversion of arginine to agmatine, while SpeB is an agmatinase, catalyzing the conversion of agmatine to putrescine. In E. coli, polyamines are required for normal cell growth, and their levels were shown to change in response to osmolarity (58). The role of MDOs and polyamines in response to salinity/osmolarity in V. cholerae has yet to be examined.

(ii) A class III transcriptional regulator is regulated by salinity and osmolarity. To further characterize the transcriptional response of V. cholerae to salinity, we focused on a putative gene encoding a transcriptional regulator (annotated by TIGR) with mRNA transcript levels that decreased as salinity increased and which was grouped into the class III category. This gene was VCA0029, and its expression was greatest in cells grown in 0 M NaCl and decreased in 0.2 M NaCl (5.5-fold) and 0.5 M NaCl (7.0-fold) (Fig. 1I). We named VCA0029 oscR (osmolarity-controlled regulator) because our results suggest that the expression of this gene is modulated by medium osmolarity (see below). OscR is annotated as an IclR-type (isocitrate lyase) regulator, and it contains a helix-turn-helix DNA binding domain. IclR-type regulators are widely distributed in a variety of bacterial species and can act as repressors, activators, or both (42). Interestingly, IclR-type regulators have domains that exhibit structural similarities to GAF domains, which are known to bind a variety of small molecules, including cyclic nucleotides and sodium ions (8). OscR showed homology to proteins from a range of bacterial species, including other Vibrio species, such as Vibrio vulnificus, as well as other bacterial pathogens, such as P. aeruginosa strain PA14 and E. coli (see Fig. S3 in the supplemental material).

Transcriptional profiling results showed that oscR expression was upregulated in 0 M NaCl and decreased as the NaCl concentration increased. We then asked whether this effect was dependent on the NaCl concentration specifically or was a response to medium osmolarity. To this end, we cultured the wild-type strain in LB medium supplemented with NaCl, KCl, or lactose and quantified oscR mRNA levels by qPCR. oscR expression levels paralleled the microarray results for increasing NaCl concentrations, where expression was highest in 0 M NaCl and lower in 0.2 M (2.5-fold) and 0.5 M NaCl (2.0-fold) (Fig. 5A). A similar downregulation trend was observed as 0.2 M KCl (3.8-fold), 0.5 M KCl (3.6-fold), or 0.4 M lactose (4.4-fold) was included in the medium.


Figure 5
View larger version (26K):
[in this window]
[in a new window]

  		FIG. 5. oscR expression is modulated by osmolarity. qPCR analysis was performed to determine oscR and gyrA message levels in cells grown in LB supplemented with NaCl, KCl, or lactose at the indicated concentrations (A) or in LO-MM without NaCl or with 0.2 M NaCl (B). Gel panels are representative images of qPCR results. Reactions without reverse transcriptase (RT) were included as negative controls. Graphs show average oscR/gyrA message levels for three biological replicates, and error bars indicate standard deviations.

To better determine the conditions under which oscR was expressed, we grew V. cholerae cells in defined ASW medium of different salinities or in M9 minimal medium supplemented with different NaCl concentrations. These media did not induce the expression of oscR (data not shown). The osmolalities of 5% ASW and M9 media without NaCl were 283 and 309 mosmol/kg, respectively. We reasoned that the osmolarities of these media might be too high to induce oscR expression, and therefore we formulated LO-MM to analyze oscR expression. Indeed, the oscR level, quantified by qPCR, was induced in LO-MM without added NaCl (96 mosmol/kg) compared to that in LO-MM supplemented with 0.2 M NaCl (444 mosmol/kg) (Fig. 5B). Our results show that oscR expression is regulated by osmolarity.

(iii) OscR regulates gene expression in a salinity-dependent manner. To determine whether oscR regulates gene expression in a salinity-dependent manner, a strain with an in-frame deletion of oscR was created. The oscR mutant showed similar growth to that of the wild type at 0 M NaCl and 0.2 M NaCl (data not shown). The transcriptional profile of the {Delta}oscR mutant was compared to that of the wild type grown to mid-exponential phase in medium supplemented with 0 M or 0.2 M NaCl, using the previously indicated criteria. Seventy-three genes were differentially regulated ≥2-fold between the {Delta}oscR mutant and the wild type in 0 M NaCl, while only two genes were differentially regulated ≥2-fold in 0.2 M NaCl (Table 5). A select set of these genes are discussed below.


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

  		TABLE 5. Genes differentially expressed in the oscR mutant versus the wild type grown to exponential phase in 0 M and 0.2 M NaCl-containing LBa

(iv) OscR negatively regulates vps expression under low-salinity conditions only. Multiple genes important for biofilm matrix production, including the vps, rbm, and bap-1 genes, were upregulated in the {Delta}oscR mutant compared to the wild type, but only in 0 M NaCl (Table 5). We further confirmed this finding by assessing the expression of a vpsLp-lacZ chromosomal fusion in the wild type and the {Delta}oscR mutant (Fig. 6A).


Figure 6
View larger version (104K):
[in this window]
[in a new window]

  		FIG. 6. OscR regulates biofilm matrix genes and biofilm formation under low-salinity conditions. (A) Comparison of vpsL expression in β-galactosidase assays of strains harboring a single-copy chromosomal vpsLp-lacZ fusion in wild-type or {Delta}oscR mutant cells. (B) Comparison of vpsL expression in β-galactosidase assays of strains harboring a single-copy chromosomal vpsLp-lacZ fusion in the wild-type strain or the {Delta}oscR, VCA0030::pGP704, or {Delta}oscR VCA0030::pGP704 mutant. β-Galactosidase activity was measured in cells containing a chromosomal vpsLp-lacZ fusion grown to mid-exponential phase in 0 M or 0.2 M NaCl. Results are shown for one representative experiment with three biological replicates. Error bars indicate standard deviations for at least six technical replicates. (C) CLSM images of horizontal (xy) and vertical (xz) projections of biofilm structures formed by wild-type and {Delta}oscR V. cholerae cells in a flow cell system with 0 M and 0.2 M NaCl in full-strength LB at 8 and 24 h postinoculation. Bars, 30 µm.

oscR precedes VCA0030, encoding a hypothetical protein, in a predicted two-gene operon (TIGR). Transcriptional profiling of the {Delta}oscR mutant revealed that VCA0030 was downregulated 7.5-fold in 0 M NaCl compared to the wild-type level but was not significantly different in 0.2 M NaCl. We therefore hypothesized that the effects of OscR at low salinities could be modulated in part by VCA0030. To test this hypothesis, we inactivated VCA0030 by insertional mutagenesis (VCA0030::pGP704) and created a {Delta}oscR VCA0030::pGP704 construct in a strain carrying the chromosomal vpsLp-lacZ fusion. vpsL expression in the VCA0030::pGP704 strain was similar to that in the wild type in both 0 M and 0.2 M NaCl (Fig. 6B). Furthermore, the β-galactosidase activity of the {Delta}oscR VCA0030::pGP704 mutant was similar to that of the {Delta}oscR mutant in both 0 M NaCl and 0.2 M NaCl. These results indicate that OscR regulates vps expression independently of VCA0030.

The vps genes are known to be regulated by the second messenger 3',5'-cyclic diguanylic acid (c-di-GMP). We therefore hypothesized that OscR regulates the expression of proteins that control intracellular c-di-GMP levels, which consequently modulates vps gene expression. A search for GGDEF, EAL, or HD-GYP domain-containing proteins that might be linked to the {Delta}oscR phenotype yielded VC1353 (encoding a GGDEF protein), which was upregulated 1.8-fold in the {Delta}oscR mutant compared to the wild type in 0 M NaCl and was not significantly different in 0.2 M NaCl. {Delta}VC1353 and {Delta}VC1353 {Delta}oscR mutants were therefore constructed in the vpsLp-lacZ reporter strain, and β-galactosidase activity was measured in cells grown in 0 M and 0.2 M NaCl. The β-galactosidase activity of the {Delta}VC1353 strain was similar to the wild-type level in both 0 M and 0.2 M NaCl, while the activity of the {Delta}oscR {Delta}VC1353 mutant was similar to that of the {Delta}oscR strain in both 0 M NaCl and 0.2 M NaCl (see Fig. S4 in the supplemental material). These results indicate that VC1353 is not required for OscR to exert its effect on vps expression in a salinity-dependent manner.

(v) OscR modulates biofilm formation under low-salt conditions only. Since vps genes were differentially expressed in the {Delta}oscR strain in 0 M NaCl only, we hypothesized that oscR modulates biofilm formation in a NaCl concentration-dependent manner. Therefore, wild-type and {Delta}oscR strains were grown in once-through flow cell systems, and biofilm architecture was analyzed using CLSM and COMSTAT analysis. Since V. cholerae is unable to grow at 0 M NaCl without high organic nutrient concentrations, wild-type and {Delta}oscR strains were grown in 0 M and 0.2 M NaCl in full-strength LB. Eight hours after initial inoculation into both 0 M and 0.2 M treatments, biofilms of both the wild-type and {Delta}oscR strains appeared qualitatively similar by CLSM (Fig. 6C), and COMSTAT analysis showed no difference in the biofilm parameters examined (data not shown). At 24 h, wild-type and oscR mutant biofilms appeared similar in 0.2 M NaCl but were markedly different in 0 M NaCl. Total biomass, average thickness, and maximum thickness were greater for biofilms formed by the {Delta}oscR mutant than for those formed by the wild type at 0 M NaCl (Table 6). Additionally, the roughness coefficient was decreased in the {Delta}oscR strain at 0 M NaCl compared to that for the wild type. These findings are consistent with the observation that oscR mutants express higher vps levels at low NaCl concentrations and therefore have an enhanced capacity to form biofilms under these conditions.


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

  		TABLE 6. COMSTAT analysis of biofilm structures formed by V. cholerae wild-type and {Delta}oscR mutant strains in 0 M and 0.2 M NaCl-containing LBa

(vi) OscR requires VpsR and VpsT for vps expression but regulates vps in parallel with HapR. Transcriptional profiling showed that the expression of vpsT was 3.8-fold greater in the {Delta}oscR mutant than in the wild type when both strains were grown at 0 M NaCl, while vpsT expression was not significantly different at 0.2 M NaCl. We therefore asked whether OscR repression of vps expression was dependent on other known positive regulators, VpsR and VpsT, or on the negative regulator HapR. To this end, we constructed {Delta}vpsR {Delta}oscR, {Delta}vpsT {Delta}oscR, and {Delta}hapR {Delta}oscR mutants in a strain containing a chromosomal vpsLp-lacZ fusion. We then analyzed vpsL expression in wild-type and mutant strains grown in 0 M NaCl. In agreement with previous findings, {Delta}vpsR and {Delta}vpsT strains showed little to no vpsL expression, and similar expression levels were observed with double {Delta}vpsR {Delta}oscR and {Delta}vpsT {Delta}oscR strains (Fig. 7). These results indicate that vpsR and vpsT are epistatic to oscR. In contrast, OscR repression was independent of HapR, as the {Delta}hapR single-deletion strain exhibited increased expression and a {Delta}hapR {Delta}oscR double-deletion mutant exhibited a further increase in expression.


Figure 7
View larger version (30K):
[in this window]
[in a new window]

  		FIG. 7. vpsR and vpsT are epistatic to oscR. vpsL expression is shown for wild-type and {Delta}vpsR, {Delta}vpsT, {Delta}hapR, {Delta}oscR, {Delta}oscR {Delta}vpsR, {Delta}oscR {Delta}vpsT, and {Delta}oscR {Delta}hapR mutant cells containing a chromosomal vpsLp-lacZ fusion and grown to exponential phase in 0 M NaCl. Results of one representative experiment with three biological replicates are shown. Error bars indicate standard deviations for at least six technical replicates.

(vii) OscR regulates motility under low-salt conditions only. Motility genes were differentially expressed in the {Delta}oscR strain in 0 M NaCl only. These genes included the four alternate flagellin-encoding genes flaB (VC2142), flaC (VC2187; 1.9-fold repressed in the {Delta}oscR mutant), flaD (VC2143), and flaE (VC2144; 1.7-fold repressed in the {Delta}oscR mutant), as well as the putative flagellin chaperone-encoding gene fliS (VC2138; 1.6-fold repressed in the {Delta}oscR mutant) (Table 5). A previous study showed that multiple alternate flagellin mutations in a classical V. cholerae strain produced shortened flagella that sheared easily, although these mutants did not show an obvious motility defect (28).

We then questioned whether the differential regulation of motility genes in the {Delta}oscR mutant would correlate with motility rates on semisolid agar in a salinity-dependent manner. Therefore, wild-type and {Delta}oscR strains with either a control vector or a complementation plasmid harboring the wild-type copy of oscR were inoculated onto 0 M NaCl- or 0.2 M NaCl-containing LB-0.3% agar plates. The oscR mutant exhibited a decrease in motility compared to the wild type on 0 M NaCl motility plates (Fig. 8). Moreover, this difference in motility was abolished when the same strains were grown on 0.2 M NaCl motility plates. The {Delta}oscR motility defect could also be complemented by the oscR gene in trans, which included a 157-bp upstream region that contained the predicted promoter. These results indicate that an {Delta}oscR mutation decreases motility under low-salinity conditions only and that this phenotype can be complemented by oscR in trans.


Figure 8
View larger version (22K):
[in this window]
[in a new window]

  		FIG. 8. OscR induces motility in a salinity-dependent manner. Images show the motility phenotypes of strains harboring the pACYC177 plasmid (vector) or pACYC177 containing the oscR coding and promoter regions (poscR) on semisolid LB agar plates containing ampicillin (100 µg/ml), with or without supplemented 0.2 M NaCl. The graph shows the mean migration zone diameter of each strain. Means are averages for 12 replicates, and error bars indicate standard deviations.

Conclusions. V. cholerae experiences periodic changes in salinity in the natural environment, and these changes are linked with outbreaks of cholera. Therefore, understanding the effects of salinity on V. cholerae physiology is particularly important for understanding the environmental persistence and outbreak potential of a waterborne pathogen. In this study, we analyzed the transcriptional response of V. cholerae to salinity and determined that biofilm formation is regulated by salinity. To be advantageous to V. cholerae survival, some signals should induce biofilm formation and others should prevent it. Our findings suggest that it may not be beneficial for V. cholerae to form biofilms under low-salinity/low-osmolarity conditions, such as the conditions found in freshwater environments. In this study, we identified and characterized a transcriptional regulator (OscR) which inversely regulates motility and biofilm formation in an osmolarity-dependent manner (Fig. 9). We speculate that OscR or another regulator whose transcription is regulated by OscR might sense a low-osmolarity signal and prevent biofilm formation. In summary, our study shows that the transcription of genes involving multiple cellular processes is modulated by salinity/osmolarity in V. cholerae and that the transcriptional regulator OscR mediates one aspect of this response.


Figure 9
View larger version (25K):
[in this window]
[in a new window]

  		FIG. 9. Model of salinity/osmolarity response in V. cholerae. Under low-salinity conditions, oscR is upregulated, which in turn positively regulates motility and negatively regulates biofilm formation. At a median salinity, biofilm matrix gene expression is increased through the action of the transcriptional regulators VpsR and VpsT. At high salinity, ectoine biosynthesis genes are upregulated, leading to compatible solute production.


arrow
ACKNOWLEDGMENTS
 
This work was supported by a grant from the NIH (AI055987) to F.H.Y. and by grants and scholarships from the ARCS Foundation, the Coastal Environmental Quality Initiative (CEQI), the STEPS Institute at UC Santa Cruz, and the Friends of the Long Marine Lab to N.J.S.

We thank James Meir for constructing FY_Vc_616 and pFY-217, Bentley Lim for constructing pFY-393, Sinem Beyhan for constructing FY_Vc_1739, Lindsay Odell for constructing FY_Vc_2407, and Vanessa Soliven for constructing pFY-315 and pFY-591. We also thank Karen Ottemann and members of the Yildiz laboratory for their valuable comments on the manuscript.


arrow
FOOTNOTES
 
* Corresponding author. Mailing address: Department of Microbiology and Environmental Toxicology, University of California, Santa Cruz, Santa Cruz, CA 95064. Phone: (831) 459-1588. Fax: (831) 459-3524. E-mail: yildiz{at}metx.ucsc.edu Back

{triangledown} Published ahead of print on 27 March 2009. Back

{dagger} Supplemental material for this article may be found at http://jb.asm.org/. Back


arrow
REFERENCES
 
    1
  1. Bao, Y., D. P. Lies, H. Fu, and G. P. Roberts. 1991. An improved Tn7-based system for the single-copy insertion of cloned genes into chromosomes of gram-negative bacteria. Gene 109:167-168.[CrossRef][Medline]
    2. 2
  2. Beyhan, S., K. Bilecen, S. R. Salama, C. Casper-Lindley, and F. H. Yildiz. 2007. Regulation of rugosity and biofilm formation in Vibrio cholerae: comparison of VpsT and VpsR regulons and epistasis analysis of vpsT, vpsR, and hapR. J. Bacteriol. 189:388-402.[Abstract/Free Full Text]
    3. 3
  3. Beyhan, S., A. D. Tischler, A. Camilli, and F. H. Yildiz. 2006. Differences in gene expression between the classical and El Tor biotypes of Vibrio cholerae O1. Infect. Immun. 74:3633-3642.[Abstract/Free Full Text]
    4. 4
  4. Beyhan, S., A. D. Tischler, A. Camilli, and F. H. Yildiz. 2006. Transcriptome and phenotypic responses of Vibrio cholerae to increased cyclic di-GMP level. J. Bacteriol. 188:3600-3613.[Abstract/Free Full Text]
    5. 5
  5. Bohin, J. P. 2000. Osmoregulated periplasmic glucans in proteobacteria. FEMS Microbiol. Lett. 186:11-19.[Medline]
    6. 6
  6. Bremer, E., and R. Kramer. 2000. Coping with osmotic challenges: osmoregulation through accumulation and release of compatible solutes in bacteria, p. 79-97. In G. Storz and R. Hengge-Aronis (ed.), Bacterial stress responses. ASM Press, Washington, DC.
    7. 7
  7. Callahan, L. T., III, R. C. Ryder, and S. H. Richardson. 1971. Biochemistry of Vibrio cholerae virulence. II. Skin permeability factor-cholera enterotoxin production in a chemically defined medium. Infect. Immun. 4:611-618.[Abstract/Free Full Text]
    8. 8
  8. Cann, M. 2007. A subset of GAF domains are evolutionarily conserved sodium sensors. Mol. Microbiol. 64:461-472.[CrossRef][Medline]
    9. 9
  9. Casper-Lindley, C., and F. H. Yildiz. 2004. VpsT is a transcriptional regulator required for expression of vps biosynthesis genes and the development of rugose colonial morphology in Vibrio cholerae O1 El Tor. J. Bacteriol. 186:1574-1578.[Abstract/Free Full Text]
    10. 10
  10. de Lorenzo, V., and K. N. Timmis. 1994. Analysis and construction of stable phenotypes in gram-negative bacteria with Tn5- and Tn10-derived minitransposons. Methods Enzymol. 235:386-405.[Medline]
    11. 11
  11. Faruque, S. M., M. J. Albert, and J. J. Mekalanos. 1998. Epidemiology, genetics, and ecology of toxigenic Vibrio cholerae. Microbiol. Mol. Biol. Rev. 62:1301-1314.[Abstract/Free Full Text]
    12. 12
  12. Fong, J. C., K. Karplus, G. K. Schoolnik, and F. H. Yildiz. 2006. Identification and characterization of RbmA, a novel protein required for the development of rugose colony morphology and biofilm structure in Vibrio cholerae. J. Bacteriol. 188:1049-1059.[Abstract/Free Full Text]
    13. 13
  13. Fong, J. C., and F. H. Yildiz. 2008. Interplay between cyclic AMP-cyclic AMP receptor protein and cyclic di-GMP signaling in Vibrio cholerae biofilm formation. J. Bacteriol. 190:6646-6659.[Abstract/Free Full Text]
    14. 14
  14. Fong, J. C., and F. H. Yildiz. 2007. The rbmBCDEF gene cluster modulates development of rugose colony morphology and biofilm formation in Vibrio cholerae. J. Bacteriol. 189:2319-2330.[Abstract/Free Full Text]
    15. 15
  15. Fries, J., R. Noble, G. Kelly, and J. Hsieh. 2007. Storm impacts on potential pathogens in estuaries. Eos Trans. AGU 88:93.
    16. 16
  16. Fullner, K. J., and J. J. Mekalanos. 1999. Genetic characterization of a new type IV-A pilus gene cluster found in both classical and El Tor biotypes of Vibrio cholerae. Infect. Immun. 67:1393-1404.[Abstract/Free Full Text]
    17. 17
  17. Gardel, C. L., and J. J. Mekalanos. 1994. Regulation of cholera toxin by temperature, pH, and osmolarity. Methods Enzymol. 235:517-526.[Medline]
    18. 18
  18. Gentleman, R. C., V. J. Carey, D. M. Bates, B. Bolstad, M. Dettling, S. Dudoit, B. Ellis, L. Gautier, Y. Ge, J. Gentry, K. Hornik, T. Hothorn, W. Huber, S. Iacus, R. Irizarry, F. Leisch, C. Li, M. Maechler, A. J. Rossini, G. Sawitzki, C. Smith, G. Smyth, L. Tierney, J. Y. Yang, and J. Zhang. 2004. Bioconductor: open software development for computational biology and bioinformatics. Genome Biol. 5:R80.[CrossRef][Medline]
    19. 19
  19. Gupta, S., and R. Chowdhury. 1997. Bile affects production of virulence factors and motility of Vibrio cholerae. Infect. Immun. 65:1131-1134.[Abstract]
    20. 20
  20. Han, W., and P. Webster. 2002. Forcing mechanisms of sea level interannual variability in the Bay of Bengal. J. Phys. Oceanogr. 32:216-239.[CrossRef]
    21. 21
  21. Herrero, M., V. de Lorenzo, and K. N. Timmis. 1990. Transposon vectors containing non-antibiotic resistance selection markers for cloning and stable chromosomal insertion of foreign genes in gram-negative bacteria. J. Bacteriol. 172:6557-6567.[Abstract/Free Full Text]
    22. 22
  22. Heydorn, A., B. K. Ersboll, M. Hentzer, M. R. Parsek, M. Givskov, and S. Molin. 2000. Experimental reproducibility in flow-chamber biofilms. Microbiology 146:2409-2415.[Abstract/Free Full Text]
    23. 23
  23. Heydorn, A., A. T. Nielsen, M. Hentzer, C. Sternberg, M. Givskov, B. K. Ersboll, and S. Molin. 2000. Quantification of biofilm structures by the novel computer program COMSTAT. Microbiology 146:2395-2407.[Abstract/Free Full Text]
    24. 24
  24. Huq, A., R. B. Sack, A. Nizam, I. M. Longini, G. B. Nair, A. Ali, J. G. Morris, Jr., M. N. Khan, A. K. Siddique, M. Yunus, M. J. Albert, D. A. Sack, and R. R. Colwell. 2005. Critical factors influencing the occurrence of Vibrio cholerae in the environment of Bangladesh. Appl. Environ. Microbiol. 71:4645-4654.[Abstract/Free Full Text]
    25. 25
  25. Huq, A., P. A. West, E. B. Small, M. I. Huq, and R. R. Colwell. 1984. Influence of water temperature, salinity, and pH on survival and growth of toxigenic Vibrio cholerae serovar O1 associated with live copepods in laboratory microcosms. Appl. Environ. Microbiol. 48:420-424.[Abstract/Free Full Text]
    26. 26
  26. Jubelin, G., A. Vianney, C. Beloin, J. M. Ghigo, J. C. Lazzaroni, P. Lejeune, and C. Dorel. 2005. CpxR/OmpR interplay regulates curli gene expression in response to osmolarity in Escherichia coli. J. Bacteriol. 187:2038-2049.[Abstract/Free Full Text]
    27. 27
  27. Kapfhammer, D., E. Karatan, K. J. Pflughoeft, and P. I. Watnick. 2005. Role for glycine betaine transport in Vibrio cholerae osmoadaptation and biofilm formation within microbial communities. Appl. Environ. Microbiol. 71:3840-3847.[Abstract/Free Full Text]
    28. 28
  28. Klose, K. E., and J. J. Mekalanos. 1998. Differential regulation of multiple flagellins in Vibrio cholerae. J. Bacteriol. 180:303-316.[Abstract/Free Full Text]
    29. 29
  29. Koelle, K., X. Rodo, M. Pascual, M. Yunus, and G. Mostafa. 2005. Refractory periods and climate forcing in cholera dynamics. Nature 436:696-700.[CrossRef][Medline]
    30. 30
  30. Lefebvre, B., P. Formstecher, and P. Lefebvre. 1995. Improvement of the gene splicing overlap (SOE) method. BioTechniques 19:186-188.[Medline]
    31. 31
  31. Lequette, Y., E. Rollet, A. Delangle, E. P. Greenberg, and J. P. Bohin. 2007. Linear osmoregulated periplasmic glucans are encoded by the opgGH locus of Pseudomonas aeruginosa. Microbiology 153:3255-3263.[Abstract/Free Full Text]
    32. 32
  32. Liang, W., A. J. Silva, and J. A. Benitez. 2007. The cyclic AMP receptor protein modulates colonial morphology in Vibrio cholerae. Appl. Environ. Microbiol. 73:7482-7487.[Abstract/Free Full Text]
    33. 33
  33. Louis, V. R., E. Russek-Cohen, N. Choopun, I. N. Rivera, B. Gangle, S. C. Jiang, A. Rubin, J. A. Patz, A. Huq, and R. R. Colwell. 2003. Predictability of Vibrio cholerae in Chesapeake Bay. Appl. Environ. Microbiol. 69:2773-2785.[Abstract/Free Full Text]
    34. 34
  34. Lupp, C., R. E. Hancock, and E. G. Ruby. 2002. The Vibrio fischeri sapABCDF locus is required for normal growth, both in culture and in symbiosis. Arch. Microbiol. 179:57-65.[CrossRef][Medline]
    35. 35
  35. Mahalanabis, D., R. Watten, and C. Wallace. 1974. Clinical aspects and management of pediatric cholera, p. 221-233. In D. Barua and W. Burrows (ed.), Cholera. W. B. Saunders Company, Philadelphia, PA.
    36. 36
  36. Matson, J. S., J. H. Withey, and V. J. DiRita. 2007. Regulatory networks controlling Vibrio cholerae virulence gene expression. Infect. Immun. 75:5542-5549.[Free Full Text]
    37. 37
  37. Miller, C. J., B. S. Drasar, and R. G. Feachem. 1982. Cholera and estuarine salinity in Calcutta and London. Lancet i:1216-1218.
    38. 38
  38. Miller, C. J., B. S. Drasar, and R. G. Feachem. 1984. Response of toxigenic Vibrio cholerae O1 to physico-chemical stresses in aquatic environments. J. Hyg. (London) 93:475-495.[Medline]
    39. 39
  39. Miller, J. H. 1972. Assay of β-galactosidase, p. 352-355. In J. H. Miller (ed.), Experiments in molecular genetics. Cold Spring Harbor Laboratory, Cold Spring Harbor, NY.
    40. 40
  40. Miller, V. L., and J. J. Mekalanos. 1988. A novel suicide vector and its use in construction of insertion mutations: osmoregulation of outer membrane proteins and virulence determinants in Vibrio cholerae requires toxR. J. Bacteriol. 170:2575-2583.[Abstract/Free Full Text]
    41. 41
  41. Miller, V. L., R. K. Taylor, and J. J. Mekalanos. 1987. Cholera toxin transcriptional activator toxR is a transmembrane DNA binding protein. Cell 48:271-279.[CrossRef][Medline]
    42. 42
  42. Molina-Henares, A. J., T. Krell, M. Eugenia Guazzaroni, A. Segura, and J. L. Ramos. 2006. Members of the IclR family of bacterial transcriptional regulators function as activators and/or repressors. FEMS Microbiol. Rev. 30:157-186.[CrossRef][Medline]
    43. 43
  43. Mourino-Perez, R. R., A. Z. Worden, and F. Azam. 2003. Growth of Vibrio cholerae O1 in red tide waters off California. Appl. Environ. Microbiol. 69:6923-6931.[Abstract/Free Full Text]
    44. 44
  44. Padan, E., N. Maisler, D. Taglicht, R. Karpel, and S. Schuldiner. 1989. Deletion of ant in Escherichia coli reveals its function in adaptation to high salinity and an alternative Na+/H+ antiporter system(s). J. Biol. Chem. 264:20297-20302.[Abstract/Free Full Text]
    45. 45
  45. Padan, E., and S. Schuldiner. 1994. Molecular physiology of Na+/H+ antiporters, key transporters in circulation of Na+ and H+ in cells. Biochim. Biophys. Acta 1185:129-151.[Medline]
    46. 46
  46. Pascual, M., M. J. Bouma, and A. P. Dobson. 2002. Cholera and climate: revisiting the quantitative evidence. Microbes Infect. 4:237-245.[CrossRef][Medline]
    47. 47
  47. Pascual, M., X. Rodo, S. P. Ellner, R. Colwell, and M. J. Bouma. 2000. Cholera dynamics and El Nino-Southern Oscillation. Science 289:1766-1769.[Abstract/Free Full Text]
    48. 48
  48. Pfau, J. D., and R. K. Taylor. 1996. Genetic footprint on the ToxR-binding site in the promoter for cholera toxin. Mol. Microbiol. 20:213-222.[Medline]
    49. 49
  49. Pflughoeft, K. J., K. Kierek, and P. I. Watnick. 2003. Role of ectoine in Vibrio cholerae osmoadaptation. Appl. Environ. Microbiol. 69:5919-5927.[Abstract/Free Full Text]
    50. 50
  50. Pinner, E., E. Padan, and S. Schuldiner. 1994. Kinetic properties of NhaB, a Na+/H+ antiporter from Escherichia coli. J. Biol. Chem. 269:26274-26279.[Abstract/Free Full Text]
    51. 51
  51. Reichelt, J. L., and P. Baumann. 1973. Taxonomy of the marine, luminous bacteria. Arch. Microbiol. 94:283-330.
    52. 52
  52. Rodo, X., M. Pascual, G. Fuchs, and A. S. Faruque. 2002. ENSO and cholera: a nonstationary link related to climate change? Proc. Natl. Acad. Sci. USA 99:12901-12906.[Abstract/Free Full Text]
    53. 53
  53. Sambrook, J., E. F. Fritsch, and T. Maniatis. 1989. Molecular cloning: a laboratory manual, 2nd ed. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY.
    54. 54
  54. Schild, S., R. Tamayo, E. J. Nelson, F. Qadri, S. B. Calderwood, and A. Camilli. 2007. Genes induced late in infection increase fitness of Vibrio cholerae after release into the environment. Cell Host Microbe 2:264-277.[CrossRef][Medline]
    55. 55
  55. Simons, R. W., F. Houman, and N. Kleckner. 1987. Improved single and multicopy lac-based cloning vectors for protein and operon fusions. Gene 53:85-96.[CrossRef][Medline]
    56. 56
  56. Singleton, F. L., R. Attwell, S. Jangi, and R. R. Colwell. 1982. Effects of temperature and salinity on Vibrio cholerae growth. Appl. Environ. Microbiol. 44:1047-1058.[Abstract/Free Full Text]
    57. 57
  57. Singleton, F. L., R. W. Attwell, M. S. Jangi, and R. R. Colwell. 1982. Influence of salinity and organic nutrient concentration on survival and growth of Vibrio cholerae in aquatic microcosms. Appl. Environ. Microbiol. 43:1080-1085.[Abstract/Free Full Text]
    58. 58
  58. Tkachenko, A. G., O. Salakhetdinova, and M. R. Pshenichnov. 1997. Exchange of putrescine and potassium between cells and media as a factor in the adaptation of Escherichia coli to hyperosmotic shock. Mikrobiologiia 66:329-334.[Medline]
    59. 59
  59. Tusher, V. G., R. Tibshirani, and G. Chu. 2001. Significance analysis of microarrays applied to the ionizing radiation response. Proc. Natl. Acad. Sci. USA 98:5116-5121.[Abstract/Free Full Text]
    60. 60
  60. Vimont, S., and P. Berche. 2000. NhaA, an Na+/H+ antiporter involved in environmental survival of Vibrio cholerae. J. Bacteriol. 182:2937-2944.[Abstract/Free Full Text]
    61. 61
  61. Vital, M., H. P. Fuchslin, F. Hammes, and T. Egli. 2007. Growth of Vibrio cholerae O1 Ogawa El Tor in freshwater. Microbiology 153:1993-2001.[Abstract/Free Full Text]
    62. 62
  62. Watnick, P. I., and R. Kolter. 1999. Steps in the development of a Vibrio cholerae El Tor biofilm. Mol. Microbiol. 34:586-595.[CrossRef][Medline]
    63. 63
  63. Wei, Y., A. A. Guffanti, M. Ito, and T. A. Krulwich. 2000. Bacillus subtilis YqkI is a novel malic/Na+-lactate antiporter that enhances growth on malate at low protonmotive force. J. Biol. Chem. 275:30287-30292.[Abstract/Free Full Text]
    64. 64
  64. Yildiz, F. H., N. A. Dolganov, and G. K. Schoolnik. 2001. VpsR, a member of the response regulators of the two-component regulatory systems, is required for expression of vps biosynthesis genes and EPS(ETr)-associated phenotypes in Vibrio cholerae O1 El Tor. J. Bacteriol. 183:1716-1726.[Abstract/Free Full Text]
    65. 65
  65. Yildiz, F. H., X. S. Liu, A. Heydorn, and G. K. Schoolnik. 2004. Molecular analysis of rugosity in a Vibrio cholerae O1 El Tor phase variant. Mol. Microbiol. 53:497-515.[CrossRef][Medline]
    66. 66
  66. Yildiz, F. H., and G. K. Schoolnik. 1999. Vibrio cholerae O1 El Tor: identification of a gene cluster required for the rugose colony type, exopolysaccharide production, chlorine resistance, and biofilm formation. Proc. Natl. Acad. Sci. USA 96:4028-4033.[Abstract/Free Full Text]
    67. 67
  67. Zhu, J., and J. J. Mekalanos. 2003. Quorum sensing-dependent biofilms enhance colonization in Vibrio cholerae. Dev. Cell 5:647-656.[CrossRef][Medline]

Journal of Bacteriology, July 2009, p. 4082-4096, Vol. 191, No. 13
0021-9193/09/$08.00+0     doi:10.1128/JB.01540-08
Copyright © 2009, American Society for Microbiology. All Rights Reserved.

Related articles in JB:

OscR, a New Osmolarity-Responsive Regulator in Vibrio cholerae
Paul V. Dunlap
JB 2009 191: 4053-4055. [Full Text]  



This article has been cited by other articles:

  • Shikuma, N. J., Fong, J. C. N., Odell, L. S., Perchuk, B. S., Laub, M. T., Yildiz, F. H. (2009). Overexpression of VpsS, a Hybrid Sensor Kinase, Enhances Biofilm Formation in Vibrio cholerae. J. Bacteriol. 191: 5147-5158 [Abstract] [Full Text]  
  • Dunlap, P. V. (2009). OscR, a New Osmolarity-Responsive Regulator in Vibrio cholerae. J. Bacteriol. 191: 4053-4055 [Full Text]  

This Article
FREE
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Supplemental material
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Services
Right arrow Related articles in JB
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 arrowReprints and Permissions
Right arrow Copyright Information
Right arrow Books from ASM Press
Right arrow MicrobeWorld
Citing Articles
Right arrow Citing Articles via HighWire
Google Scholar
Right arrow Articles by Shikuma, N. J.
Right arrow Articles by Yildiz, F. H.
PubMed
Right arrow PubMed Citation
Right arrow Articles by Shikuma, N. J.
Right arrow Articles by Yildiz, F. H.

Copyright © 2009 by the American Society for Microbiology.   For an alternate route to Journals.ASM.org, visit: http://intl-journals.asm.org | More Info»