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. Author manuscript; available in PMC: 2014 May 1.
Published in final edited form as: Environ Microbiol. 2012 Jun 12;15(5):1387–1399. doi: 10.1111/j.1462-2920.2012.02805.x

The Transcriptional Regulator, CosR, Controls Compatible Solute Biosynthesis and Transport, Motility and Biofilm Formation in Vibrio cholerae

Nicholas J Shikuma 1, Kimberly R Davis 1, Jiunn N C Fong 1, Fitnat H Yildiz 1,*
PMCID: PMC3597767  NIHMSID: NIHMS387729  PMID: 22690884

SUMMARY

Vibrio cholerae inhabits aquatic environments and colonizes the human digestive tract to cause the disease cholera. In these environments, V. cholerae copes with fluctuations in salinity and osmolarity by producing and transporting small, organic, highly soluble molecules called compatible solutes, which counteract extracellular osmotic pressure. Currently, it is unclear how V. cholerae regulates the expression of genes important for the biosynthesis or transport of compatible solutes in response to changing salinity or osmolarity conditions. Through a genome-wide transcriptional analysis of the salinity response of V. cholerae, we identified a transcriptional regulator we name CosR for compatible solute regulator. The expression of cosR is regulated by ionic strength and not osmolarity. A transcriptome analysis of a ΔcosR mutant revealed that CosR represses genes involved in ectoine biosynthesis and compatible solute transport in a salinity-dependent manner. When grown in salinities similar to estuarine environments, CosR activates biofilm formation and represses motility independently of its function as an ectoine regulator. This is the first study to characterize a compatible solute regulator in V. cholerae and couples the regulation of osmotic tolerance with biofilm formation and motility.

INTRODUCTION

All microorganisms must cope with variable environments and one important condition that microbial cells must adapt to is osmolarity. Many bacteria adapt to increased osmolarity by importing or producing compatible solutes to counteract osmotic pressure (Wood, 1999). Compatible solutes are small, highly soluble organic molecules that act to stabilize intracellular levels of water and turgor pressure without disturbing cellular function (Brown, 1976). Compatible solutes can be divided into various groups including carbohydrates, polyols, heterosides, amino acids and amino acid derivatives (da Costa et al., 1998; Sleator and Hill, 2002). Common amino acids used as compatible solutes are proline and glutamate. The amino acid derivatives glycine betaine and ectoine are also widely used by bacteria to counteract high osmolarity (Lucht and Bremer, 1994; Pflughoeft et al., 2003; Kapfhammer et al., 2005; Wood, 2007).

The halo-tolerant bacterium V. cholerae causes the disease cholera and thrives in aquatic environments. Salinity is a significant factor affecting the growth and distribution of V. cholerae in the environment (Singleton et al., 1982a; Miller et al., 1984; Louis et al., 2003; Huq et al., 2005), and cholera epidemics correlate with increased salinity in riverine and estuarine habitats (Faruque et al., 1998; Lobitz et al., 2000; Pascual et al., 2002). V. cholerae growth and survival is optimal in the range of salinities present in estuarine environments (Singleton et al., 1982a; Louis et al., 2003), although V. cholerae is also capable of growth in both fresh and seawater salinities (Singleton et al., 1982b; Alsina and Blanch, 1994; Vital et al., 2007). During infection, V. cholerae must adapt to a range of salinities or osmolarities as it passes through the human digestive tract and is released back into the environment (Barua and Burrows, 1974; Gupta and Chowdhury, 1997). Indeed, osmolarity is one factor hypothesized to induce virulence factor production during V. cholerae infection. Additionally, virulence factor gene expression and production is modulated by various salinity concentrations (Tamplin and Colwell, 1986; Miller and Mekalanos, 1988; Shikuma and Yildiz, 2009). Thus, salinity is a significant factor V. cholerae must cope with as a facultative human pathogen.

To survive estuarine conditions and passage through the human digestive tract where salinity concentrations often vary, V. cholerae synthesizes and imports compatible solutes. V. cholerae possesses ectoine biosynthesis genes including ectA, ectB, ectC and a putative aspartokinase (VCA0822) that are similar to genes in the ectoine biosynthesis pathway determined for several moderate halophiles (Peters et al., 1990; Louis and Galinski, 1997; Canovas et al., 1998; Ono et al., 1999; Kuhlmann and Bremer, 2002; Pflughoeft et al., 2003). In V. cholerae, ΔectA mutants exhibit a growth defect in defined media lacking compensatory compatible solutes glutamate, proline, and aspartate (Pflughoeft et al., 2003). V. cholerae also possesses homologs of opuD from Bacillus species and putP from Vibrio vulnificus, both of which were shown to transport proline and glycine betaine and facilitate growth (Kapfhammer et al., 2005). Compatible solutes are therefore important for V. cholerae to grow and persist in environments of variable osmolarities.

Another factor that facilitates V. cholerae survival and transmission is biofilm formation. Through a genome-wide transcriptome analysis, we showed previously that salinity affects the expression of biofilm genes in V. cholerae (Shikuma and Yildiz, 2009). Furthermore, salinity modulates V. cholerae surface attachment and biofilm architecture (Shikuma and Yildiz, 2009). In microcosm experiments, the attachment to zooplankton enhances V. cholerae survival (Huq et al., 1983) and biofilm growth improves resistance to protozoan grazing (Matz et al., 2005). Furthermore, mutants defective in biofilm formation are attenuated in the colonization of an infant mouse colonization model (Fong et al., 2010) and V. cholerae biofilm aggregates are more infectious than their free-living counterparts (Colwell et al., 2003; Faruque et al., 2006).

Although compatible solute transport and biosynthesis are important for V. cholerae survival as a facultative human pathogen, it remains unclear how V. cholerae regulates compatible solute genes in response to salinity or osmolarity. In this work, we identify and characterize a previously undescribed MarR-type transcriptional regulator, which we call CosR for Compatible Solute Regulator. cosR expression is controlled specifically by ionic strength and not osmolarity. Whole genome transcriptome profiles revealed that CosR represses the expression of the compatible solute transporter, opuD and the ectoine biosynthesis genes. Furthermore, CosR represses motility and activates biofilm formation independently of its regulatory role in ectoine biosynthesis. V. cholerae therefore utilizes CosR to adapt to environments of varying salinity and for activating biofilm growth in the marine environment.

RESULTS AND DISCUSSION

Identification of CosR

Salinity is an important factor in V. cholerae environmental occurrence and incidence of infection, and we previously described the V. cholerae transcriptional response to salinity (Shikuma and Yildiz, 2009). To expand our insight into the mechanisms by which V. cholerae responds to salinity, we searched for transcriptional regulators whose expression correlated with salinity concentration. Expression of one such regulator, encoded by VC1278, increased in a salinity-dependent manner (Fig. 1A) (Shikuma and Yildiz, 2009). We named this gene cosR (compatible solute regulator) because our results indicate that CosR regulates genes involved in compatible solute production and transport. cosR expression was lowest in cells grown in 0 M NaCl, and increased in 0.2 M NaCl (2.4-fold) and 0.5 M NaCl (6.2-fold).

Fig 1. cosR expression is regulated by salinity.

Fig 1

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(A) Relative cosR expression in wild type V. cholerae grown at the indicated NaCl concentration relative to cells grown at 0 M NaCl in LB media identified through transcriptome analysis. Expression was not significantly different between 0 M and 0.1 M NaCl treatments (Shikuma and Yildiz, 2009). (B) Quantitative PCR of cosR expression in total RNA isolated from cells grown in LB supplemented with NaCl, KCl or lactose at the indicated concentrations. Reactions without reverse transcriptase (RT) were included as a negative control. Graph depicts an average of three biological replicates of cosR/gyrA message levels and error bars indicate standard deviations, p value determined by student’s t test (*p < 0.05). (C) Arrangement of genes encoding the transcriptional regulator, CosR, the compatible solute transporter OpuD and proteins required for ectoine biosynthesis, EctA, EctB, EctC and aspartokinase (VCA0822), in V. cholerae.

The expression of cosR correlated with NaCl concentration; however, cosR expression could be dependent on NaCl concentration, ionic concentration or medium osmolarity. To determine whether cosR expression would change in response to the concentration of other solutes in the medium, we cultured wild-type V. cholerae cells in NaCl, KCl, or the non-metabolizable osmolyte, lactose, and compared the levels of cosR mRNA using quantitative PCR (qPCR). Lactose was used previously to increase the osmolarity of the medium without altering ionic strength to observe the differential production of virulence factors (TcpA and cholera toxin) in V. cholerae (Miller and Mekalanos, 1988). In agreement with microarray data, cosR levels increased in cells grown in 0.2 M NaCl (3.0-fold) and 0.5 M NaCl (4.6-fold) when compared to 0 M NaCl (Fig. 1B). Similarly, levels of cosR increased as KCl concentrations increased from 0 M KCl to 0.2 M KCl (2.7-fold) and 0.5 M KCl (4.1-fold). In contrast, cells grown in media containing 0.4 M lactose exhibited similar levels of cosR expression when compared to 0 M NaCl. The osmolarity of 0.2 M NaCl or KCl in solution is equivalent to 0.4 M lactose. Expression of cosR is therefore dependent on the ionic strength of the medium, and appears not to be regulated by medium osmolarity.

CosR is predicted to be a MarR-type (multiple antibiotic resistance) transcriptional regulator and homologs exist in other related Vibrio species (Fig S1). Moreover, the genomes of other Vibrios also encode similar osmoadaptation genes (Naughton et al., 2009), leaving open the possibility that this regulator might perform similar functions in related bacteria. A recent study identified another MarR-type regulator, EctR1, from the halotolerant obligate methanotroph, Methylomicrobium alcaliphilum, which was also shown to regulate ectoine biosynthesis genes (Mustakhimov et al., 2010). EctR1 exhibits a 51% sequence identity to CosR (Fig S1) and both EctR1 and CosR repress ectoine biosynthesis genes in response to NaCl concentration (see below). While ectR1 is divergently transcribed from the ectABC and aspartokinase genes of M. alcaliphilum, cosR is divergently transcribed from opuD encoding a compatible solute transporter in V. cholerae (Fig 1C). Furthermore, the ectABC and aspartokinase operon is located on chromosome-II of V. cholerae, while cosR resides on chromosome-I.

CosR represses compatible solute transport and biosynthesis genes

To determine the function of CosR, we generated a strain with an in-frame deletion of cosR. Growth of wild type and ΔcosR strains were similar when grown in LB with 0.2 M NaCl or 0.5 M NaCl (Fig S2). To identify the CosR regulon, we compared transcriptional profiles of the ΔcosR strain to that of the wild-type strain when grown in LB media supplemented with 0.2 M NaCl or 0.5 M NaCl. Thirty eight genes were differentially regulated ≥2-fold between ΔcosR and wild type (Table 1). One of these genes was previously identified to encode a compatible solute transporter, OpuD (Kapfhammer et al., 2005). opuD expression in the mutant was 6.2- and 1.9-fold upregulated in 0.2 M NaCl and 0.5 M NaCl, respectively, when compared to the wild type (Table 1). In the sequenced genome of V. cholerae strain N16961 (Heidelberg et al., 2000), cosR and opuD are divergently transcribed and have a 357 bp intergenic region between them. Genes in the ectoine biosynthesis operon, consisting of ectA, ectB, ectC and VCA0822 (Pflughoeft et al., 2003), were also upregulated in the cosR mutant in both 0.2 M and 0.5 M NaCl, indicating that CosR is a repressor of genes involved in production/transport of compatible solutes (Table 1). All genes in the ectoine biosynthesis operon were upregulated in the mutant compared to the wild type in both 0.2 M and 0.5 M NaCl treatments. However, we did observe variation in the magnitude of expression between genes predicted to be the same operon, which we suspect could be due to inherent variability in microarray data collection (Beckman et al., 2004; Morey et al., 2006). Altogether, these results suggest that CosR is a repressor of compatible solute transport and biosynthesis genes.

Table 1.

Genes differentially expressed in the cosR mutant versus wild type grown to exponential phase in 0.2 M NaCl and 0.5 M NaCl-containing LB.

ORF Gene Function ΔcosR
0.2M /
Wt
0.2M		 ΔcosR
0.5M /
Wt
0.5M
Amino Acid Biosynthesis
VC0162 ilvC Ketol-acid reductoisomerase 2.02
VC2746 glnA Glutamate-ammonia ligase 2.11
Biosynthesis of Cofactors, Prosthetic Groups and Carriers
VCA0712 pncA Pyrazinamidase/nicotinamidase 2.27
Cell Envelope
VC2633 Fimbrial assembly protein PilN, putative 3.43
Ectoine Biosynthesis
VCA0822 ask Aspartokinase putative 73.61 34.67
VCA0823 ectC Ectoine synthase 85.37 33.86
VCA0824 ectB Diaminobutyrate pyruvate aminotransferase 6.48 44.71
VCA0825 ectA L-2, 4-diaminobutyric acid acetyltransferase 42.69 95.24
Energy Metabolism
VC1690 Alpha-1, 6-galactosidase putative 0.37
VC1827 manA-2 Mannose-6-phosphate isomerase 0.47
VCA0828 phhA Phenylalanine-4-hydroxylase 4.82
Hypothetical Proteins
VC0316 Hypothetical protein 2.25
VC1020 Hypothetical protein 2.43
VC1389 Hypothetical protein 2.20
VC1586 Hypothetical protein 0.41
VC1657 Hypothetical protein 2.18
VC1696 Hypothetical protein 0.40
VC1764 Hypothetical protein 0.48
VC1787 Hypothetical protein 0.21
VC1808 Hypothetical protein 0.45
VC2151 Hypothetical protein 2.05
VCA0284 Hypothetical protein 0.45
VCA0326 Hypothetical protein 0.38
VCA0467 Hypothetical protein 2.17
VCA0839 Hypothetical protein 0.30
VCA0849 Hypothetical protein 2.90
VCA0973 Hypothetical protein 0.36
VC1645 Conserved hypothetical protein 0.47
VC2473 Conserved hypothetical protein 0.50
Regulatory Functions
VC2692 cpxR Transcriptional regulator CpxR 2.01
Transport and Binding Proteins
VC0173 peptide ABC transporter permease protein 3.45
VC1092 oppB oligopeptide ABC transporter permease protein 2.07
VC1279 opuD transporter BCCT family 6.17
VC1854 ompT ompT protein 0.45
VC1929 dctP-2 C4-dicarboxylate-binding periplasmic protein 3.08
VCA0516 fruA-2 PTS system fructose-specific IIBC component 0.40
VCA0745 glpF Glycerol uptake facilitator protein 0.48
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Differentially expressed genes were determined using SAM software, with criteria of a ≥2-fold change in gene expression and an FDR of ≤3%.

It is noteworthy that CosR did not solely regulate genes involved in compatible solute production or transport (Table 1). Other genes regulated by CosR included the porin gene, ompT (which was 2.2-fold downregulated in the mutant in 0.2 M NaCl) and the transcriptional regulator cpxR (which was 2.0-fold upregulated in the mutant in 0.5 M NaCl). The Cpx two-component system is well studied in E. coli and medium osmolarity induced by NaCl or sucrose was shown to stimulate this regulon (Prigent-Combaret et al., 2001). In contrast, the V. cholerae Cpx system is specifically activated by chloride ions and not osmolarity (Slamti and Waldor, 2009). We show that NaCl and KCl regulate cosR expression and CosR regulates cpxR (Table 1). We therefore speculate that the Cpx system and CosR might regulate each other’s expression.

To substantiate the transcriptional profiling results of the ΔcosR mutant, qPCR was conducted using RNA isolated from wild type and ΔcosR strains grown in 0.2 M NaCl and 0.5 M NaCl. In both salinities, the ectA and VCA0822 genes had greater message levels in the ΔcosR mutant when compared to wild type (Fig. 2A and 2B). De-repression of the ectA operon was most apparent in the cosR mutant grown in 0.2 M NaCl, where ectoine biosynthesis genes are normally repressed. The difference in ectA and VCA0822 expression between ΔcosR and wild type strains as quantified by qPCR were not as pronounced as those observed by microarray analyses. We suspect that differences in quantification and normalization methods factor into differences we observed in gene expression. Nonetheless, we do observe similar trends in gene expression with both methods. Altogether, these results corroborate microarray expression profile results of the ΔcosR mutant, where cosR represses the ectoine biosynthesis operon.

Fig 2. CosR represses compatible solute biosynthesis/transporter genes.

Fig 2

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Quantitative PCR analysis of (A) ectA/gyrA and (B) VCA0822/gyrA message levels from wild type and ΔcosR mutant cells grown in 0.2 M or 0.5 M NaCl in LB. Reactions without reverse transcriptase (RT) were included as a negative control. Bars represent the average of three biological replicates and error bars indicate standard deviations, p value determined by student’s t test (*p < 0.01). (C) Quantification of ectoine levels in wild type, ΔcosR and ΔectA strains grown in M9 minimal media supplemented with 0.2 M NaCl. Ectoine was quantified using high-performance liquid chromatography-tandem mass spectrometry (HPLC-MS/MS) as described in the materials and methods and normalized to total protein levels. Bars are an average of 3 biological replicates and error bars indicate standard deviations, p value determined by student’s t test (*p < 0.001). Not detected (ND).

To support the gene expression results, we quantified the cellular levels of ectoine in wild type, ΔcosR and ΔectA strains using high-performance liquid chromatographytandem mass spectrometry (HPLC-MS/MS). In cells grown in M9 minimal media supplemented with 0.2 M NaCl, ectoine levels were 3.5-fold greater in a ΔcosR mutant when compared to wild type cells, while ectoine levels were undetectable in a ΔectA strain (Fig 2C). These results indicate that CosR represses the biosynthesis of ectoine in V. cholerae.

CosR complements a ΔcosR mutation

To verify that a cosR deletion caused derepression of the ect operon, the ΔcosR mutant was complemented in trans with a plasmid containing the putative cosR promoter and cosR gene. Introduction of cosR to the ΔcosR mutant in trans decreased the expression of ectA and VCA0822 by 2.0- and 1.9-fold, respectively, compared to the ΔcosR strain with the vector alone (Fig 3). These results indicate that the addition of cosR in trans is able to partially complement the expression levels of the ectA operon in a ΔcosR mutant. The activity of some transcriptional regulators is known to be modulated by small molecule ligands. Since CosR regulates compatible solute production in V. cholerae, we hypothesized that CosR activity might be modulated by ectoine, proline or glycine betaine, however we observed no difference in expression of opuD upon addition or removal of these compatible solutes (Fig S3).

Fig 3. Complementation of ΔcosR.

Fig 3

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Quantitative PCR analysis of (A) ectA/gyrA and (B) VCA0822/gyrA message levels from wild type and ΔcosR mutant grown in 0.2 M NaCl. Strains contained the pACYC177 plasmid (vector) or pACYC177 harboring the putative cosR promoter and coding region (pcosR). Reactions without reverse transcriptase (RT) were included as a negative control. Graphs are an average of three biological replicates and error bars indicate standard deviations, p value determined by student’s t test (*p < 0.05).

One puzzling aspect of CosR regulation is that cosR and the compatible solute genes are upregulated in high salinity, yet CosR is a repressor of the compatible solute genes. We speculate that CosR is involved in a negative autoregulatory loop, which facilitates rapid response times and reduces variation in gene expression (Fig 6) (Yosef and Regev, 2011). This regulatory system makes sense in the context of the V. cholerae lifestyle, as rapid response times are likely important to counteract salinity or osmotic shifts in estuarine environments due to tide and storm surge, or during passage through the human digestive tract. The mechanism by which CosR controls compatible solute biosynthesis/transport genes appears to be complex and we are currently working to elucidate this regulatory pathway.

Fig 6. Model of the V. cholerae salinity/osmolarity response.

Fig 6

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The expression of cosR, encoding a MarR-type transcriptional regulator, is activated by ionic strength. CosR represses expression of the genes encoding the compatible solute transporter, OpuD, and the ectoine biosynthesis proteins. Salinity activates the same set of genes and opposes CosR regulation. CosR additionally activates biofilm formation and represses motility independently of its role as an ectoine regulator. The expression of oscR, encoding an IclR-type transcriptional regulator, is upregulated in low salinity/osmolarity conditions (Shikuma and Yildiz, 2009). OscR represses biofilm formation through the repression of the vps genes and activates motility.

CosR activates biofilm formation in artificial seawater

V. cholerae depends on its ability to form biofilms for its lifestyle as a facultative human pathogen and normal inhabitant of aquatic environments. We therefore asked whether CosR is important for V. cholerae to form biofilms. Wild type and ΔcosR strains were inoculated into once-through flow-cell chambers containing either LB supplemented with 0.2 M NaCl or 50% artificial seawater media (ASW), which mimics conditions found in estuarine environments. In ASW, wild-type V. cholerae formed robust biofilms after 72 h, while the ΔcosR mutant was severely defective in biofilm growth as observed by Confocal Laser Scanning Microscopy (CLSM) (Fig 4A). These results were confirmed using image analysis software to quantify biofilm properties (Table 2). The growth rate of wild type and ΔcosR was similar in 50% ASW liquid cultures (Fig S2), suggesting that the biofilm defect of the ΔcosR mutant was not due to differences in growth in this medium. In contrast to ASW-grown biofilms, wild type and ΔcosR strains did not exhibit differences in biofilm formation when grown with LB medium in flow-cell chambers (Fig 4B, Table 2). These results suggest that CosR activates biofilm formation when grown in ASW.

Fig 4. CosR activates biofilm formation in seawater microcosms.

Fig 4

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Representative CLSM micrographs showing horizontal (xy) and vertical (xz) projections of biofilm structures formed by wild type, ΔcosR, ΔectA and ΔcosRΔectA V. cholerae in 50% artificial seawater (A) or LB supplemented with 0.2 M NaCl (B). Strains were grown in a once-through flow cell system at room temperature and images were acquired at the indicated timepoints. Scale bar is 50 μm.

Table 2.

COMSTAT analysis of biofilm structures formed by wild-type, ΔcosR, ΔectA and ΔcosRΔectA strains in a once-through flow cell system in 50% artificial seawater (ASW) or LB media supplemented with 0.2 M NaCl (LB).

Media Time
(h)		 Strain Total biomass
(μm3/μm2)		 Thickness (μm)
Average Maximum
ASW 24 Wild type 6.80 (1.51) 7.04 (1.70) 16.28 (7.32)
Δ cosR 3.24 (1.06) 3.52 (1.39) 8.30 (0.70)
Δ ectA 6.10 (0.83) 5.89 (0.73) 10.56 (4.35)
Δ cosR Δ ectA 1.87 (0.72) 1.87 (0.76) 5.28 (1.11)
ASW 48 Wild type 8.60 (2.45) 9.19 (2.81) 21.21 (2.47)
Δ cosR 4.65 (1.90) 4.78 (1.89) 16.72 (2.90)
Δ ectA 5.59 (2.24) 6.29 (2.96) 18.23 (3.51)
Δ cosR Δ ectA 0.93 (0.14) 0.98 (0.21) 8.95 (3.17)
ASW 72 Wild type 19.27 (3.33) 21.52 (4.61) 33.34 (4.89)
Δ cosR 8.47 (2.58) 8.71 (3.31) 20.11 (3.71)
Δ ectA 9.39 (2.95) 10.42 (3.95) 22.00 (6.05)
Δ cosR Δ ectA 4.19 (1.20) 4.75 (1.25) 14.52 (4.71)
LB 24 Wild type 10.6 (1.1) 9.7 (1.1) 16.4 (1.2)
Δ cosR 8.7 (1.3) 8.0 (1.1) 14.1 (1.9)
Δ ectA 10.0 (1.2) 9.2 (1.2) 15.5 (0.7)
Δ cosR Δ ectA 9.6 (0.6) 8.9 (0.6) 15.6 (1.7)
LB 48 Wild type 19.9 (2.3) 19.1 (2.4) 29.1 (3.4)
Δ cosR 15.8 (0.8) 15.0 (0.8) 26.1 (3.5)
Δ ectA 18.5 (1.8) 18.1 (1.9) 28.4 (3.7)
Δ cosR Δ ectA 17.0 (1.4) 16.4 (1.7) 26.7 (1.9)
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Values presented are averages of data from at least six z-series image stacks. The numbers in parentheses indicate standard deviations.

Since CosR is a repressor of ectoine biosynthesis and a ΔcosR mutant overproduces ectoine, we asked whether ΔectA or ΔcosRΔectA strains also exhibited biofilm formation phenotypes. When grown in ASW, the ΔectA strain was defective in biofilm formation when compared to wild type (Fig 4 and Table 2). Moreover, ΔcosRΔectA biofilms were more defective in biofilm formation when compared to wild type, ΔcosR or ΔectA strains (Fig 4 and Table 2), suggesting that mutations in ΔcosR and ΔectA additively affect biofilm formation in V. cholerae in ASW. Growth of ΔectA and ΔcosRΔectA were similar to wild type in ASW liquid cultures (Fig S2), suggesting that biofilm defects were not due to differential growth of the mutant strains. When grown in a flow cell in LB medium, ΔectA and ΔcosRΔectA strains did not show biofilm defects (Fig 4B, Table 2). These results suggest that the positive effect of CosR on biofilm formation in ASW is independent of ectoine production. Furthermore, the presence of ectoine in the cell, at normal wild type levels or elevated levels in a ΔcosR mutant, appears to be sufficient to activate biofilm formation. In contrast, above results suggest that the lack of ectoine in a ΔectA mutant affects biofilm formation. In agreement with our findings, Kapfhammer et al. (2005) showed previously that ectoine or another compatible solute, glycine betaine, could stimulate vps expression and biofilm formation in V. cholerae. Although no genes known to be directly involved in biofilm formation were differentially regulated between ΔcosR and wild-type grown in LB medium (Table 1), CosR might regulate genes important for biofilm formation in cells grown in ASW under flow cell conditions, or regulate biofilm formation through a post-transcriptional mechanism independently of its regulatory effect on ectoine production.

Biofilm formation is a key factor in both the environmental persistence and transmission of V. cholerae. We previously showed that salinity modulates biofilm architecture in V. cholerae (Shikuma and Yildiz, 2009), and we show here that CosR is important for V. cholerae to form biofilms in artificial seawater microcosms. CosR might therefore play a role in both the environmental survival and transmission throughout the lifecycle of V. cholerae.

CosR negatively regulates motility

Since biofilm production and motility are often inversely regulated and motility is important for environmental survival and infectivity of the human host (Richardson, 1991; Gardel and Mekalanos, 1996; Silva et al., 2006), we tested the motility phenotype of ΔcosR on LB soft agar motility plates containing a range of NaCl concentrations. Wild type showed the highest motility zone diameter at a median salinity of 0.2 M NaCl (Fig 5A). Furthermore, the motility zone of ΔcosR was greater than that of wild type in salinities between 0.1 and 0.3 M NaCl, while ΔcosR motility was similar to wild type in low (0 M) or high (0.5 M NaCl) salinities (Fig 5A). When grown at 0.2 M NaCl, the motility phenotype of ΔcosR could be complemented by cosR in trans (Fig 5B), confirming that cosR represses motility. Although we observed a decrease in motility diameter for both wild type and ΔcosR strains when ampicillin was included in the growth medium for plasmid maintenance, there was still a significant difference in motility between wild type and ΔcosR strains when each contained the empty vector. We then hypothesized that the motility phenotype of a ΔcosR mutant was due to the overproduction of ectoine, and therefore additionally tested the motility phenotype of ΔectA and ΔcosRΔectA strains in LB containing 0.2 M NaCl. The ΔectA motility zone diameter was no different than wild type, and ΔcosRΔectA motility zone diameter was no different than the ΔcosR mutant (Fig 5C). These results indicate that CosR is a salinity-dependent repressor of motility and this phenotype is not due to the overproduction of ectoine.

Fig 5. CosR represses motility in a salinity-dependent manner.

Fig 5

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(A) Graph of the motility zone diameter of wild type and ΔcosR V. cholerae strains inoculated in semisolid LB agar plates containing the indicated NaCl concentration after 16 h growth at 30°C. Bars indicate the average of 4 biological replicates and error bars indicate standard deviations, p value determined by student’s t test (*p < 0.0005). (B) Graph of the motility zone diameter of wild type or ΔcosR V. cholerae containing an empty pACYC177 plasmid (Vector) or pACYC177 containing the putative promoter and cosR coding region (pcosR). Strains were inoculated into semisolid LB agar containing 0.2 M NaCl and 100 μg/ml ampicillin and the motility diameter was measured after 16 h growth at 30°C. Bars indicate the average of 4 biological replicates and error bars indicate standard deviations, p value determined by student’s t test (*p < 0.0005). (C) Graph of the motility zone diameter of wild type, ΔcosR, ΔectA and ΔcosRΔectA V. cholerae strains inoculated in semisolid LB agar plates containing 0.2 M NaCl after 16 h growth at 30°C. Bars indicate the average of 4 biological replicates and error bars indicate standard deviations, p value determined by student’s t test (*p < 0.00005).

In some microbial pathogens, genes important for compatible solute transport are thought to facilitate infection by mediating stress encountered in the infection process (Sleator and Hill, 2002). In pathogenic Escherichia coli and Staphylococcus aureus, mutations in compatible solute transporters were shown to reduce colonization and infection in model virulence assays (Culham et al., 1998; Bayer et al., 1999). Deletions of multiple compatible solute transporters in Listeria monocytogenes attenuated virulence in a murine infection model (Wemekamp-Kamphuis et al., 2002). It is likely that V. cholerae also uses CosR for the regulation of compatible solute transport and biosynthesis in response to various osmotic stresses during its infectious lifecycle. Furthermore, this study implicates CosR as a link between compatible solute production/transport and the regulation of biofilm formation and motility.

While cosR expression is activated in response to ionic strength, the expression of oscR, encoding another regulator important for the salinity response of V. cholerae, is inversely correlated with medium osmolarity (Fig 6) (Shikuma and Yildiz, 2009). V. cholerae can therefore distinguish medium ionic strength from osmolarity to differentially activate these regulators. Furthermore, cosR and oscR are inversely expressed as salinity increases. Given the importance of salinity and osmolarity in the lifestyle of V. cholerae, it makes sense that this bacterium would have the means for differentiating between these two signals. CosR also activates biofilm formation and represses motility in salinities between 0.1 and 0.3 M NaCl. This regulation contrasts with OscR, which represses biofilm formation and activates motility under low osmolarity conditions (Shikuma and Yildiz, 2009). V. cholerae therefore modulates its behavior in different salinity/osmolarity conditions by differentially activating CosR and OscR.

Conclusion

V. cholerae depends on its ability to sense and respond to various salinities and osmolarities as an aquatic bacterium and human gastrointestinal pathogen. CosR represses compatible solute gene expression and production in a salinity dependent manner (Fig 6). This work is the first account of a key mechanism by which V. cholerae senses and responds to various salinities through the production of compatible solutes for environmental survival. Biofilm formation and motility are important for the environmental survival, transmission and infectivity of V. cholerae, and we also show that CosR is important for these phenotypes. CosR is therefore important for V. cholerae to cope with osmotic stress and is likely important for V. cholerae survival as a facultative human pathogen.

MATERIALS AND METHODS

Bacterial strains, plasmids, and culture conditions

The bacterial strains and plasmids used in this study are listed in Table S1. All V. cholerae and E. coli strains were grown aerobically, at 30°>C and 37°>C, respectively, unless otherwise noted. Luria Bertani (LB) growth medium (1% Tryptone, 0.5% Yeast Extract) pH 7.5 was supplemented with 0.2 or 0.5 M NaCl, 0.2 or 0.5 M KCl or 0.2 or 0.5 M D-lactose. D-lactose was used as a non-metabolizable osmolyte (Shikuma and Yildiz, 2009). LB-agar and soft agar plates contained 1.5% and 0.3% (wt/vol) granulated agar (Difco), respectively. Concentrations of antibiotics used were as follows: ampicillin (100 μg/ml), rifampicin (100 μg/ml), and gentamicin (30 μg/ml). M9 minimal medium was adapted from Gerhardt et al. (1994) and consisted of 42 mM Na2HPO4, 22 mM KH2PO4, 19 mM NH4Cl, 1 mM MgSO4, 0.1 mM CaCl2, 1X MEM Vitamin solution (Mediatech), 0.2% DL-lactate and 0.2 M NaCl. Artificial seawater was described previously (Shikuma and Yildiz, 2009), which was adapted from (Reichelt and Baumann, 1973; Lupp et al., 2002), and contained 300 mM NaCl, 50 mM MgSO4, 10 mM KCl, 10 mM CaCl2 and 0.333 mM K2HPO4 and these components were used at 50% concentrations to replicate estuarine conditions. Tryptone (100 mg/L) and yeast extract (50 mg/L) were used as carbon and nitrogen sources (Singleton et al., 1982a; Mourino-Perez et al., 2003; Shikuma and Yildiz, 2009).

DNA manipulations

Oligonucleotides used in the present study were purchased from Bioneer Corporation and are listed in Table S2. Restriction enzymes, DNA modification enzymes and Phusion High-Fidelity DNA polymerase were purchased from New England Biolabs. DNA sequencing was carried out by Sequetech Corporation.

Generation of in-frame deletion, chromosomal fusion and point mutation strains

In-frame deletion strains were generated according to previously published protocols (Miller and Mekalanos, 1988; Fullner and Mekalanos, 1999; Fong and Yildiz, 2007; Shikuma and Yildiz, 2009). All strains were verified by PCR. Plasmid sequences were verified by DNA sequencing by Sequetech Corporation.

β-galactosidase assays

β-galactosidase assays were performed and Miller units calculated as described previously (Shikuma et al., 2009) which is similar to the procedure described by Miller (Miller, 1972). The assays were repeated with three biological replicates and eight technical replicates.

RNA isolation

V. cholerae cells were grown aerobically overnight in LB supplemented with 0.2 M NaCl. Cultures were diluted 1:200 in fresh LB media containing concentrations of NaCl as indicated and grown aerobically at 30°C with shaking at 200 rpm to an OD600 of 0.3 to 0.4, diluted 1:200 once more into fresh media containing the same NaCl concentration and harvested at an OD600 of 0.3 to 0.4. Two milliliter aliquots of cultures 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 oligos representing the open reading frames present in the V. cholerae strain N16961 genome and were printed at UCSC (Beyhan et al., 2006b). Whole-genome expression analyses were performed as described previously (Beyhan et al., 2006a; Shikuma and Yildiz, 2009). Differentially regulated genes were determined using three biological replicates and two technical replicates for each treatment (6 data points for each spot) using a 2-fold difference in gene expression and 3% false discovery rate (FDR) as cut-off values.

Quantitative PCR (qPCR)

qPCR was performed as described previously (Shikuma and Yildiz, 2009). Briefly, cDNA was produced from 0.25 to 1 μg of each RNA sample using an iScript cDNA Synthesis Kit (Bio-Rad). The product was diluted with water (1:4), and 4 μl was used as a template with 12 pmol of each primer in a subsequent PCR reaction using Expand High Fidelity PCR System (Roche). PCR reaction conditions were as follows: 94°C for 2 min, then 25 cycles of 94°C for 30 sec, 55°C for 30 sec and 72°C for 30 sec, and a final 72°C for 2 min. The amplified products were analyzed on a 1.5% agarose gel and quantified using ImageQuant software (Molecular Dynamics). Intensities of each DNA band were normalized to the corresponding gyrA band. At least three biological replicates were conducted for each treatment tested and reactions lacking reverse transcriptase were used as negative controls.

Determination of cellular ectoine levels

Cell cultures were grown in 25 ml M9 minimal media supplemented with 0.2 M NaCl shaking at 200 rpm at 30°C for 20 h. Extraction of ectoine from cells was performed using a modified Bligh and Dyer technique (Bligh and Dyer, 1959; Kuhlmann and Bremer, 2002). Briefly, 1 ml of cell cultures was pelleted by centrifugation at 16,100 xg for 15 min at room temperature. Replicate samples were kept for protein quantification using a BCA Protein Assay Kit (Pierce). 400 μl of extraction mixture (methanol/chloroform/water 10:5:4, by vol) was added to samples and vigorously shaken for 45 min. 130 μl chloroform and 130 μl water were added followed by vigorous shaking for 30 min. Phase separation was performed by centrifugation at 16,100 xg for 30 min at room temperature. 300 μl of aqueous phase was recovered, lyophilized, and resuspended in 100 μl of HPLC grade water.

Samples were analyzed by high-performance liquid chromatography-tandem mass spectrometry (HPLC-MS/MS) on a Thermo-Electron Finnigan LTQ mass spectrometer (Thermo, San Jose, CA) coupled with a surveyor HPLC (Thermo, San Jose, CA). Twenty μl of each sample was injected to a Nucleodur NH2 column (4.6 mm × 150 mm, Macherey-Nagel). The gradient was started at 5% acetonitrile with 0.1 % formic acid and a linear gradient to 80% acetonitrile was achieved in 10 min at a flow rate of 400 μl/min at 25°C. Ectoine (143.2 m/z) was detected by selected ion-monitoring analyses (SRM) (m/z = 141.7 to 144.7) in positive mode and identified by the fragmentation pattern in the second MS (MS/MS). The electrospray voltage was set to 5.0 kV, and a data-dependent selection was monitored to select the most abundant three ions from SRM Scan for MS/MS analysis. The collision-induced dissociation (CID) at normalized collision energy of 35% was used for MS/MS. The retention time and standard curve were determined using commercially available ectoine (Sigma) with concentrations of 5, 10, 30, 70, 100, 300, 700, 1000 and 1200 μM. Ectoine levels were normalized to total protein per ml of culture. Quantification was performed with at least 3 biological replicates.

Motility assays

LB motility plates (0.3% agar) were used to determine the motility of bacterial strains (Wolfe and Berg, 1989). Single colonies were stabbed into LB motility plates containing the indicated NaCl concentrations and antibiotics, where appropriate, and incubated at 30°C for 16 h. The migration zone diameter was then measured for each strain analyzed. Quantification was performed with at least 4 biological replicates and repeated independently at least twice.

Flowcell biofilms

V. cholerae biofilms in flow chambers were produced as described previously (Heydorn et al., 2000a; Beyhan et al., 2007). Briefly, GFP-tagged V. cholerae strains were grown overnight in 25 ml of 50% artificial seawater (ASW) at 25°C or 0.2 M NaCl LB at 30°C and 200 rpm. Sterilized flow cell chambers were inoculated with 350 μl of cell suspension in 50% artificial seawater at an OD600 of 0.08 to 0.1 or 350 μl of cell suspension diluted with 0.2 M NaCl LB to an OD600 of 0.02. The chambers were then inverted and the cells were allowed to attach without flow for 1 h. Flow was then started at a rate of 4.5 ml/h. Flow cell experiments were conducted at room temperature and images were acquired at the timepoints indicated. Confocal images of biofilms were captured with a LSM 5 PASCAL laser scanning microscope (Zeiss) at 488 nm excitation and 543 nm emission wavelengths. Flow cell experiments were conducted with at least two biological replicates. Three dimensional images of biofilms were reconstructed using Imaris software (Bitplane) and quantified using COMSTAT (Heydorn et al., 2000b).

Supplementary Material

Supp Figure S1
Supp Figure S2
Supp Figure S3
Supp Table S1
Supp Table S2

ACKNOWLEDGEMENTS

This work was supported by a NIH grant AI055987 to FHY and 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 NJS. Ectoine quantification was performed at the UCSC Mass Spectrometry Facility, which is funded by NIH grant S10-RR20939 (MS equipment grant) and we thank Q. Zhang for help with the HPLC/MS-MS experiments and analysis. We also thank K. Ottemann, G. Hartzog, M. Parsek and members of the Yildiz group for helpful discussions and reading of this manuscript.

REFERENCES

  1. Alsina M, Blanch AR. A set of keys for biochemical identification of environmental Vibrio species. J Appl Bacteriol. 1994;76:79–85. doi: 10.1111/j.1365-2672.1994.tb04419.x. [DOI] [PubMed] [Google Scholar]
  2. Barua D, Burrows W, editors. Cholera. W. B. Saunders Company; Philadelphia, London, Toronto: 1974. [Google Scholar]
  3. Bayer AS, Coulter SN, Stover CK, Schwan WR. Impact of the high-affinity proline permease gene (putP) on the virulence of Staphylococcus aureus in experimental endocarditis. Infect Immun. 1999;67:740–744. doi: 10.1128/iai.67.2.740-744.1999. [DOI] [PMC free article] [PubMed] [Google Scholar]
  4. Beckman KB, Lee KY, Golden T, Melov S. Gene expression profiling in mitochondrial disease: assessment of microarray accuracy by high-throughput Q-PCR. Mitochondrion. 2004;4:453–470. doi: 10.1016/j.mito.2004.07.029. [DOI] [PubMed] [Google Scholar]
  5. Beyhan S, Tischler AD, Camilli A, Yildiz FH. Transcriptome and phenotypic responses of Vibrio cholerae to increased cyclic di-GMP level. J Bacteriol. 2006a;188:3600–3613. doi: 10.1128/JB.188.10.3600-3613.2006. [DOI] [PMC free article] [PubMed] [Google Scholar]
  6. Beyhan S, Tischler AD, Camilli A, Yildiz FH. Differences in gene expression between the classical and El Tor biotypes of Vibrio cholerae O1. Infect Immun. 2006b;74:3633–3642. doi: 10.1128/IAI.01750-05. [DOI] [PMC free article] [PubMed] [Google Scholar]
  7. Beyhan S, Bilecen K, Salama SR, Casper-Lindley C, Yildiz FH. 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. 2007;189:388–402. doi: 10.1128/JB.00981-06. [DOI] [PMC free article] [PubMed] [Google Scholar]
  8. Bligh EG, Dyer WJ. A rapid method of total lipid extraction and purification. Can J Biochem Physiol. 1959;37:911–917. doi: 10.1139/o59-099. [DOI] [PubMed] [Google Scholar]
  9. Brown AD. Microbial water stress. Bacteriol Rev. 1976;40:803–846. doi: 10.1128/br.40.4.803-846.1976. [DOI] [PMC free article] [PubMed] [Google Scholar]
  10. Canovas D, Vargas C, Calderon MI, Ventosa A, Nieto JJ. Characterization of the genes for the biosynthesis of the compatible solute ectoine in the moderately halophilic bacterium Halomonas elongata DSM 3043. Syst Appl Microbiol. 1998;21:487–497. doi: 10.1016/S0723-2020(98)80060-X. [DOI] [PubMed] [Google Scholar]
  11. Colwell RR, Huq A, Islam MS, Aziz KMA, Yunus M, Khan NH, et al. Reduction of cholera in Bangladeshi villages by simple filtration. Proc Natl Acad Sci. 2003;100:1051–1055. doi: 10.1073/pnas.0237386100. [DOI] [PMC free article] [PubMed] [Google Scholar]
  12. Culham DE, Dalgado C, Gyles CL, Mamelak D, MacLellan S, Wood JM. Osmoregulatory transporter ProP influences colonization of the urinary tract by Escherichia coli. Microbiology. 1998;144:91–102. doi: 10.1099/00221287-144-1-91. [DOI] [PubMed] [Google Scholar]
  13. da Costa MS, Santos H, Galinski EA. An overview of the role and diversity of compatible solutes in Bacteria and Archaea. Adv Biochem Eng Biotechnol. 1998;61:117–153. doi: 10.1007/BFb0102291. [DOI] [PubMed] [Google Scholar]
  14. Faruque SM, Albert MJ, Mekalanos JJ. Epidemiology, genetics, and ecology of toxigenic Vibrio cholerae. Microbiol Mol Biol Rev. 1998;62:1301–1314. doi: 10.1128/mmbr.62.4.1301-1314.1998. [DOI] [PMC free article] [PubMed] [Google Scholar]
  15. Faruque SM, Biswas K, Udden SM, Ahmad QS, Sack DA, Nair GB, Mekalanos JJ. Transmissibility of cholera: in vivo-formed biofilms and their relationship to infectivity and persistence in the environment. Proc Natl Acad Sci U S A. 2006;103:6350–6355. doi: 10.1073/pnas.0601277103. [DOI] [PMC free article] [PubMed] [Google Scholar]
  16. Fong JC, Yildiz FH. The rbmBCDEF gene cluster modulates development of rugose colony morphology and biofilm formation in Vibrio cholerae. J Bacteriol. 2007;189:2319–2330. doi: 10.1128/JB.01569-06. [DOI] [PMC free article] [PubMed] [Google Scholar]
  17. Fong JC, Syed KA, Klose KE, Yildiz FH. Role of Vibrio polysaccharide (vps) genes in VPS production, biofilm formation and Vibrio cholerae pathogenesis. Microbiology. 2010;156:2757–2769. doi: 10.1099/mic.0.040196-0. [DOI] [PMC free article] [PubMed] [Google Scholar]
  18. Fullner KJ, Mekalanos JJ. Genetic characterization of a new type IV-A pilus gene cluster found in both classical and El Tor biotypes of Vibrio cholerae. Infect Immun. 1999;67:1393–1404. doi: 10.1128/iai.67.3.1393-1404.1999. [DOI] [PMC free article] [PubMed] [Google Scholar]
  19. Gardel CL, Mekalanos JJ. Alterations in Vibrio cholerae motility phenotypes correlate with changes in virulence factor expression. Infect Immun. 1996;64:2246–2255. doi: 10.1128/iai.64.6.2246-2255.1996. [DOI] [PMC free article] [PubMed] [Google Scholar]
  20. Gerhardt P, Murray RGE, Wood WA, Kreig NR. Methods for Genral and Molecular Bacteriology. American Society for Microbiology; Washington, DC: 1994. [Google Scholar]
  21. Gupta S, Chowdhury R. Bile affects production of virulence factors and motility of Vibrio cholerae. Infect Immun. 1997;65:1131–1134. doi: 10.1128/iai.65.3.1131-1134.1997. [DOI] [PMC free article] [PubMed] [Google Scholar]
  22. Heidelberg JF, Eisen JA, Nelson WC, Clayton RA, Gwinn ML, Dodson RJ, et al. DNA sequence of both chromosomes of the cholera pathogen Vibrio cholerae. Nature. 2000;406:477–483. doi: 10.1038/35020000. [DOI] [PMC free article] [PubMed] [Google Scholar]
  23. Heydorn A, Ersboll BK, Hentzer M, Parsek MR, Givskov M, Molin S. Experimental reproducibility in flow-chamber biofilms. Microbiology. 2000a;146(Pt 10):2409–2415. doi: 10.1099/00221287-146-10-2409. [DOI] [PubMed] [Google Scholar]
  24. Heydorn A, Nielsen AT, Hentzer M, Sternberg C, Givskov M, Ersboll BK, Molin S. Quantification of biofilm structures by the novel computer program COMSTAT. Microbiology. 2000b;146(Pt 10):2395–2407. doi: 10.1099/00221287-146-10-2395. [DOI] [PubMed] [Google Scholar]
  25. Huq A, Small EB, West PA, Huq MI, Rahman R, Colwell RR. Ecological relationships between Vibrio cholerae and planktonic crustacean copepods. Appl Environ Microbiol. 1983;45:275–283. doi: 10.1128/aem.45.1.275-283.1983. [DOI] [PMC free article] [PubMed] [Google Scholar]
  26. Huq A, Sack RB, Nizam A, Longini IM, Nair GB, Ali A, et al. Critical factors influencing the occurrence of Vibrio cholerae in the environment of Bangladesh. Appl Environ Microbiol. 2005;71:4645–4654. doi: 10.1128/AEM.71.8.4645-4654.2005. [DOI] [PMC free article] [PubMed] [Google Scholar]
  27. Kapfhammer D, Karatan E, Pflughoeft KJ, Watnick PI. Role for glycine betaine transport in Vibrio cholerae osmoadaptation and biofilm formation within microbial communities. Appl Environ Microbiol. 2005;71:3840–3847. doi: 10.1128/AEM.71.7.3840-3847.2005. [DOI] [PMC free article] [PubMed] [Google Scholar]
  28. Kuhlmann AU, Bremer E. Osmotically regulated synthesis of the compatible solute ectoine in Bacillus pasteurii and related Bacillus spp. Appl Environ Microbiol. 2002;68:772–783. doi: 10.1128/AEM.68.2.772-783.2002. [DOI] [PMC free article] [PubMed] [Google Scholar]
  29. Lobitz B, Beck L, Huq A, Wood B, Fuchs G, Faruque AS, Colwell R. Climate and infectious disease: use of remote sensing for detection of Vibrio cholerae by indirect measurement. Proc Natl Acad Sci U S A. 2000;97:1438–1443. doi: 10.1073/pnas.97.4.1438. [DOI] [PMC free article] [PubMed] [Google Scholar]
  30. Louis P, Galinski EA. Characterization of genes for the biosynthesis of the compatible solute ectoine from Marinococcus halophilus and osmoregulated expression in Escherichia coli. Microbiology. 1997;143(Pt 4):1141–1149. doi: 10.1099/00221287-143-4-1141. [DOI] [PubMed] [Google Scholar]
  31. Louis VR, Russek-Cohen E, Choopun N, Rivera IN, Gangle B, Jiang SC, et al. Predictability of Vibrio cholerae in Chesapeake Bay. Appl Environ Microbiol. 2003;69:2773–2785. doi: 10.1128/AEM.69.5.2773-2785.2003. [DOI] [PMC free article] [PubMed] [Google Scholar]
  32. Lucht JM, Bremer E. Adaptation of Escherichia coli to high osmolarity environments: osmoregulation of the high-affinity glycine betaine transport system proU. FEMS Microbiol Rev. 1994;14:3–20. doi: 10.1111/j.1574-6976.1994.tb00067.x. [DOI] [PubMed] [Google Scholar]
  33. Lupp C, Hancock RE, Ruby EG. The Vibrio fischeri sapABCDF locus is required for normal growth, both in culture and in symbiosis. Arch Microbiol. 2002;179:57–65. doi: 10.1007/s00203-002-0502-7. [DOI] [PubMed] [Google Scholar]
  34. Matz C, McDougald D, Moreno AM, Yung PY, Yildiz FH, Kjelleberg S. Biofilm formation and phenotypic variation enhance predation-driven persistence of Vibrio cholerae. Proc Natl Acad Sci U S A. 2005;102:16819–16824. doi: 10.1073/pnas.0505350102. [DOI] [PMC free article] [PubMed] [Google Scholar]
  35. Miller CJ, Drasar BS, Feachem RG. Response of toxigenic Vibrio cholerae 01 to physico-chemical stresses in aquatic environments. J Hyg (Lond) 1984;93:475–495. doi: 10.1017/s0022172400065074. [DOI] [PMC free article] [PubMed] [Google Scholar]
  36. Miller JH. Assay of β-galactosidase. In: Miller JH, editor. Experiments in molecular genetics. Cold Spring Harbor Laboratory; Cold Spring Harbor, NY: 1972. pp. 352–355. [Google Scholar]
  37. Miller VL, Mekalanos JJ. 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. 1988;170:2575–2583. doi: 10.1128/jb.170.6.2575-2583.1988. [DOI] [PMC free article] [PubMed] [Google Scholar]
  38. Morey JS, Ryan JC, Van Dolah FM. Microarray validation: factors influencing correlation between oligonucleotide microarrays and real-time PCR. Biol Proced Online. 2006;8:175–193. doi: 10.1251/bpo126. [DOI] [PMC free article] [PubMed] [Google Scholar]
  39. Mourino-Perez RR, Worden AZ, Azam F. Growth of Vibrio cholerae O1 in red tide waters off California. Appl Environ Microbiol. 2003;69:6923–6931. doi: 10.1128/AEM.69.11.6923-6931.2003. [DOI] [PMC free article] [PubMed] [Google Scholar]
  40. Mustakhimov, Reshetnikov AS, Glukhov AS, Khmelenina VN, Kalyuzhnaya MG, Trotsenko YA. Identification and characterization of EctR1, a new transcriptional regulator of the ectoine biosynthesis genes in the halotolerant methanotroph Methylomicrobium alcaliphilum 20Z. J Bacteriol. 2010;192:410–417. doi: 10.1128/JB.00553-09. [DOI] [PMC free article] [PubMed] [Google Scholar]
  41. Naughton LM, Blumerman SL, Carlberg M, Boyd EF. Osmoadaptation among Vibrio species and unique genomic features and physiological responses of Vibrio parahaemolyticus. Appl Environ Microbiol. 2009;75:2802–2810. doi: 10.1128/AEM.01698-08. [DOI] [PMC free article] [PubMed] [Google Scholar]
  42. Ono H, Sawada K, Khunajakr N, Tao T, Yamamoto M, Hiramoto M, et al. Characterization of biosynthetic enzymes for ectoine as a compatible solute in a moderately halophilic eubacterium, Halomonas elongata. J Bacteriol. 1999;181:91–99. doi: 10.1128/jb.181.1.91-99.1999. [DOI] [PMC free article] [PubMed] [Google Scholar]
  43. Pascual M, Bouma MJ, Dobson AP. Cholera and climate: revisiting the quantitative evidence. Microbes Infect. 2002;4:237–245. doi: 10.1016/s1286-4579(01)01533-7. [DOI] [PubMed] [Google Scholar]
  44. Peters R, Galinski EA, Truper HG. The biosynthesis of ectoine. FEMS Microbiol Lett. 1990;71:157–162. [Google Scholar]
  45. Pflughoeft KJ, Kierek K, Watnick PI. Role of ectoine in Vibrio cholerae osmoadaptation. Appl Environ Microbiol. 2003;69:5919–5927. doi: 10.1128/AEM.69.10.5919-5927.2003. [DOI] [PMC free article] [PubMed] [Google Scholar]
  46. Prigent-Combaret C, Brombacher E, Vidal O, Ambert A, Lejeune P, Landini P, Dorel C. Complex regulatory network controls initial adhesion and biofilm formation in Escherichia coli via regulation of the csgD gene. J Bacteriol. 2001;183:7213–7223. doi: 10.1128/JB.183.24.7213-7223.2001. [DOI] [PMC free article] [PubMed] [Google Scholar]
  47. Reichelt JL, Baumann P. Taxonomy of the marine, luminous bacteria. Arch Microbiol. 1973;94:283–330. [Google Scholar]
  48. Richardson K. Roles of motility and flagellar structure in pathogenicity of Vibrio cholerae: analysis of motility mutants in three animal models. Infect Immun. 1991;59:2727–2736. doi: 10.1128/iai.59.8.2727-2736.1991. [DOI] [PMC free article] [PubMed] [Google Scholar]
  49. Shikuma NJ, Yildiz FH. Identification and characterization of OscR, a transcriptional regulator involved in osmolarity adaptation in Vibrio cholerae. J Bacteriol. 2009;191:4082–4096. doi: 10.1128/JB.01540-08. [DOI] [PMC free article] [PubMed] [Google Scholar]
  50. Shikuma NJ, Fong JC, Odell LS, Perchuk BS, Laub MT, Yildiz FH. Overexpression of VpsS, a hybrid sensor kinase, enhances biofilm formation in Vibrio cholerae. J Bacteriol. 2009;191:5147–5158. doi: 10.1128/JB.00401-09. [DOI] [PMC free article] [PubMed] [Google Scholar]
  51. Silva AJ, Leitch GJ, Camilli A, Benitez JA. Contribution of hemagglutinin/protease and motility to the pathogenesis of El Tor biotype cholera. Infect Immun. 2006;74:2072–2079. doi: 10.1128/IAI.74.4.2072-2079.2006. [DOI] [PMC free article] [PubMed] [Google Scholar]
  52. Singleton FL, Attwell RW, Jangi MS, Colwell RR. Influence of salinity and organic nutrient concentration on survival and growth of Vibrio cholerae in aquatic microcosms. Appl Environ Microbiol. 1982a;43:1080–1085. doi: 10.1128/aem.43.5.1080-1085.1982. [DOI] [PMC free article] [PubMed] [Google Scholar]
  53. Singleton FL, Attwell R, Jangi S, Colwell RR. Effects of temperature and salinity on Vibrio cholerae growth. Appl Environ Microbiol. 1982b;44:1047–1058. doi: 10.1128/aem.44.5.1047-1058.1982. [DOI] [PMC free article] [PubMed] [Google Scholar]
  54. Slamti L, Waldor MK. Genetic analysis of activation of the Vibrio cholerae Cpx pathway. J Bacteriol. 2009;191:5044–5056. doi: 10.1128/JB.00406-09. [DOI] [PMC free article] [PubMed] [Google Scholar]
  55. Sleator RD, Hill C. Bacterial osmoadaptation: the role of osmolytes in bacterial stress and virulence. FEMS Microbiol Rev. 2002;26:49–71. doi: 10.1111/j.1574-6976.2002.tb00598.x. [DOI] [PubMed] [Google Scholar]
  56. Tamplin ML, Colwell RR. Effects of microcosm salinity and organic substrate concentration on production of Vibrio cholerae enterotoxin. Appl Environ Microbiol. 1986;52:297–301. doi: 10.1128/aem.52.2.297-301.1986. [DOI] [PMC free article] [PubMed] [Google Scholar]
  57. Vital M, Fuchslin HP, Hammes F, Egli T. Growth of Vibrio cholerae O1 Ogawa Eltor in freshwater. Microbiology. 2007;153:1993–2001. doi: 10.1099/mic.0.2006/005173-0. [DOI] [PubMed] [Google Scholar]
  58. Wemekamp-Kamphuis HH, Wouters JA, Sleator RD, Gahan CG, Hill C, Abee T. Multiple deletions of the osmolyte transporters BetL, Gbu, and OpuC of Listeria monocytogenes affect virulence and growth at high osmolarity. Appl Environ Microbiol. 2002;68:4710–4716. doi: 10.1128/AEM.68.10.4710-4716.2002. [DOI] [PMC free article] [PubMed] [Google Scholar]
  59. Wolfe AJ, Berg HC. Migration of bacteria in semisolid agar. Proc Natl Acad Sci U S A. 1989;86:6973–6977. doi: 10.1073/pnas.86.18.6973. [DOI] [PMC free article] [PubMed] [Google Scholar]
  60. Wood JM. Osmosensing by bacteria: signals and membrane-based sensors. Microbiol Mol Biol Rev. 1999;63:230–262. doi: 10.1128/mmbr.63.1.230-262.1999. [DOI] [PMC free article] [PubMed] [Google Scholar]
  61. Wood JM. Bacterial osmosensing transporters. Methods Enzymol. 2007;428:77–107. doi: 10.1016/S0076-6879(07)28005-X. [DOI] [PubMed] [Google Scholar]
  62. Yosef N, Regev A. Impulse control: temporal dynamics in gene transcription. Cell. 2011;144:886–896. doi: 10.1016/j.cell.2011.02.015. [DOI] [PMC free article] [PubMed] [Google Scholar]

Associated Data

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Supplementary Materials

Supp Figure S1
NIHMS387729-supplement-Supp_Figure_S1.eps (7MB, eps)
Supp Figure S2
NIHMS387729-supplement-Supp_Figure_S2.eps (5MB, eps)
Supp Figure S3
NIHMS387729-supplement-Supp_Figure_S3.eps (4MB, eps)
Supp Table S1
NIHMS387729-supplement-Supp_Table_S1.doc (66.5KB, doc)
Supp Table S2
NIHMS387729-supplement-Supp_Table_S2.doc (34.5KB, doc)

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