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. Author manuscript; available in PMC: 2008 Aug 13.
Published in final edited form as: Mol Microbiol. 2002 Jul;45(2):471–483. doi: 10.1046/j.1365-2958.2002.03023.x

Vibrio cholerae CytR is a repressor of biofilm development

Adam J Haugo 1, Paula I Watnick 1,2,*
PMCID: PMC2515492  NIHMSID: NIHMS56628  PMID: 12123457

Summary

Vibrio cholerae is both a human pathogen and a natural inhabitant of aquatic environments. In the aquatic environment, microorganisms are found attached to surfaces in structures known as biofilms. We have identified a transcriptional repressor in V. cholerae that inhibits exopolysaccharide synthesis and biofilm development. Our studies show that this repressor is the V. cholerae homologue of Escherichia coli CytR, a protein that represses nucleoside uptake and catabolism when nucleosides are scarce. We propose that the role of CytR in V. cholerae biofilm development is to co-ordinate bacterial biofilm accumulation with the presence of nucleosides. Thus, nucleosides may be a signal to planktonic cells to join the biofilm.

Introduction

Although surface adhesion or biofilm development is critical for the persistence and proliferation of all bacterial species in their natural habitats, the environmental signals that promote the surface-adherent mode of life are poorly understood. Vibrio cholerae, a native inhabitant of diverse aquatic environments and the human intestinal pathogen that causes the severe diarrhoeal disease cholera, is an ideal model for the study of biofilm development. Surface adhesion or biofilm development by V. cholerae is instrumental in both pathogenesis and colonization of aquatic environments. In the environment, V. cholerae has been identified on the surfaces of zooplankton, phytoplankton, crustaceans, insects and plants (Huq et al., 1983; 1986; Tamplin et al., 1990; Shukla et al., 1995). Because zoo-plankton and phytoplankton blooms precede cholera outbreaks, the association of V. cholerae with these organisms is hypothesized to play a role in the epidemiology of disease (Colwell, 1996; Lobitz et al., 2000). Thus, an understanding of the regulation of V. cholerae surface adhesion will not only increase our understanding of bacterial adaptation to aquatic habitats but may also suggest methods of improving our predictive models for cholera epidemics.

Vibrio cholerae biofilm development on abiotic surfaces has been described previously (Watnick and Kolter, 1999; Watnick et al., 2001). In batch biofilm experiments, two stages in V. cholerae biofilm development are observed during the first 24 h of growth (Watnick et al., 2001). In the initial period of 4–5 h, only transient association with the surface is observed. This association requires a functional polar flagellum. In the second stage of biofilm development, transient association with the surface continues, but now permanent immobilization and microcolony formation are observed. Permanent immobilization is dependent on the vps genes, which encode enzymes required for the synthesis of VPS, an exopolysaccharide produced by V. cholerae (Watnick and Kolter, 1999; Yildiz and Schoolnik, 1999).

Studies of both V. cholerae and Pseudomonas aeruginosa suggest that increased exopolysaccharide synthesis alters both the course of biofilm development and the three-dimensional architecture of the mature biofilm (Wai et al., 1998; Mizunoe et al., 1999; Yildiz and Schoolnik, 1999; Hentzer et al., 2001). In V. cholerae, spontaneously occurring ‘rugose’ variants have been isolated (White, 1938; Crutchley, 1968; Morris et al., 1996; Wai et al., 1998; Mizunoe et al., 1999; Yildiz and Schoolnik, 1999). These variants display a rough colony morphology that is correlated with increased transcription of the vps genes and increased VPS synthesis (Wai et al., 1998; Mizunoe et al., 1999; Yildiz and Schoolnik, 1999; Yildiz et al., 2001). Increased VPS synthesis by rugose variants also results in a higher profile biofilm than that formed by the parent V. cholerae strain (Yildiz and Schoolnik, 1999). The genetic basis of increased vps transcription in spontaneous rugose variants has not yet been ascertained, and one possibility is that these rugose variants represent a genotypically heterogeneous population.

The transcription profiles of biofilm-associated cells are quite different from their planktonic counterparts. Evidence from P. aeruginosa and Escherichia coli suggests that increased transcription of genes required for exopolysaccharide synthesis is a characteristic of cells that have been incorporated into the biofilm (Davies et al., 1993; Prigent-Combaret et al., 1999). More comprehensive genomics- and proteomics-based profiling of biofilm-associated cells is under way or completed in many bacterial backgrounds, and these studies are likely to yield a deeper understanding of the physiological state of the biofilm-associated cell (Sauer and Camper, 2001; Schoolnik et al., 2001; Whiteley et al., 2001; Sauer et al., 2002; Zhu et al., 2002). Complementary studies of the environmental signals and transcription factors that are operative in biofilm development will allow us to link signal transduction cascades with changes in gene transcription and protein expression within the biofilm.

Several environmental signals that influence bacterial biofilm development have been identified. These include quorum sensing, surface sensing and the nutritional content of the aquatic environment. In P. aeruginosa, autoinducer synthesis mutants form a flat, detergent-sensitive biofilm, suggesting that quorum sensing directs normal biofilm development (Davies et al., 1998; Parsek and Greenberg, 1999). The quorum-sensing systems of Burkholderia cepacia and V. cholerae have also been implicated in biofilm development (Huber et al., 2001; Zhu et al., 2002). Another signal that activates biofilm development is contact with the substratum. In both V. cholerae and P. aeruginosa, there is evidence that exopolysaccharide synthesis is activated by mutation of the flagellar structural genes but not the flagellar motor (Garrett et al., 1999; Watnick et al., 2001). One possibility is that, as is true for the closely related organisms Vibrio parahaemolyticus and Vibrio alginolyticus, the flagellar motor of V. cholerae is able to sense and respond to changes in torque on the polar flagellum that occur when a surface is encountered (Belas et al., 1986; McCarter et al., 1988; Kawagishi et al., 1996). For diverse bacteria, the nutritional composition of the aquatic environment is a key determinant of surface attachment (Bowden and Li, 1997; O’Toole and Kolter, 1998; Pratt and Kolter, 1998; Watnick et al., 1999; Danese et al., 2001). In particular, for many organisms, glucose and related carbohydrates greatly enhance exopolysaccharide production and, thus, biofilm development (Bryan et al., 1986; Bonet et al., 1993; Abbad Andaloussi et al., 1995; Kimmel et al., 1998; Degeest and De Vuyst, 1999; Looijesteijn et al., 1999; Petryet al., 2000; Degeest et al., 2001; Mozzi et al., 2001). Although transcription factors that regulate exopolysaccharide synthesis have been identified in various organisms, the environmental signals that these regulators sense is not known (Reeve et al., 1997; Chapman and Kao, 1998; Yildiz et al., 2001). This is the case for VpsR, the only known activator of V. cholerae biofilm development and VPS synthesis (Yildiz et al., 2001). By sequence homology, VpsR is the response element in a two-component system. Although activation of vps gene transcription by VpsR has been demonstrated, the environmental signals that govern activation by VpsR have not yet been delineated.

In E. coli, CytR has been shown to repress nucleoside uptake and catabolism in nucleoside-poor environments. In this paper, we provide evidence that V. cholerae CytR plays a role not only in nucleoside catabolism but also in the control of biofilm development and, specifically, in the synthesis of the V. cholerae exopolysaccharide, VPS. Our studies link nucleoside concentrations to transcriptional regulation of exopolysaccharide synthesis and biofilm development by V. cholerae. Thus, we suggest that nucleosides may be a signal to planktonic cells to join the nascent biofilm.

Results

Identification of a ‘super-biofilm’ mutant

A library of mini-Tn10 transposon-insertion mutants was constructed in the V. cholerae O139 strain MO10. This library was screened for mutants that formed increased or ‘super’ biofilms as described previously (Watnick and Kolter, 1999; Watnick et al., 2001). The transposon-insertion junctions of identified mutants were amplified by arbitrary polymerase chain reaction (PCR) and sequenced (O’Toole et al., 1999). Of the 6000 transposon-insertion mutants evaluated, three independent clones were found to be in the gene given the TIGR designation VC2677 (Heidelberg et al., 2000). Further evaluation demonstrated that all these mutants displayed a rugose colony morphology and formed a biofilm containing approximately twice as many cells as a wild-type V. cholerae biofilm, as determined by crystal violet staining and quantification. These mutants also displayed normal motility on swarm agar (data not shown). A transposon-insertion mutant in the same gene of V. cholerae O1 El Tor demonstrated a similar increase in biofilm development (data not shown), suggesting that this type of regulation is not strain specific.

The ‘super biofilm’ mutant is an E. coli CytR homologue

Sequence analysis demonstrated that the amino acid sequence of the gene containing the transposon insertion is 65% identical and 81% similar to the CytR protein of E. coli. No other protein in the V. cholerae genome showed this extent of similarity to E. coli CytR. As shown in Fig. 1A, this homology extends throughout the protein. Furthermore, the genomic environments of the E. coli CytR gene and the V. cholerae biofilm repressor are strikingly similar as shown in Fig. 1B. Thus, we term our biofilm repressor V. cholerae CytR and present additional evidence below that this protein is, in fact, V. cholerae CytR.

Figure 1.

Figure 1

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A comparison of the V. cholerae O1 El Tor CytR amino acid sequence and genomic environment with that of E. coli K-12 CytR.

A. Alignment of the amino acid sequences of V. cholerae and E. coli CytR. CLUSTALW was used to produce the sequence alignment (Thompson et al., 1994).

B. Genomic environment of V. cholerae O1 El Tor cytR and E. coli K-12 cytR.

Construction of a V. cholerae cytR deletion mutant, cloning of V. cholerae cytR and complementation of the mutant phenotype

To confirm that the phenotype demonstrated by the transposon-insertion mutant was indeed the result of a mutation in cytR, a strain carrying a complete deletion of the cytR gene was constructed (strain PW324 denoted ΔcytR). The planktonic growth curves of wild-type V. cholerae and the ΔcytR mutant in LB broth were measured and found to be essentially superimposable before the onset of biofilm development (data not shown). As shown in Fig. 2, the deletion mutant demonstrated a rugose colony morphology and ‘super-biofilm’ phenotype that was similar to that of the transposon-insertion mutant. Measurements of total protein in wild-type V. cholerae and ΔcytR mutant biofilms indicated an increase in cell mass within the ΔcytR mutant biofilm that was consistent with crystal violet quantification (data not shown). Furthermore, we constructed a ΔcytR mutant harbouring a deletion in a large vps operon containing six open reading frames (ORFs) including vpsL (Yildiz et al., 2001; strain PW329 denoted ΔcytR Δvps). The ΔcytR Δvps mutant formed a smooth colony and did not accumulate on surfaces. This result suggests that the phenotype of the ΔcytR mutant results from derepression of exopolysaccharide synthesis rather than activation of a new mechanism of surface adhesion.

Figure 2.

Figure 2

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Comparison of wild-type V. cholerae and ΔcytR mutant colony morphology and biofilm development.

A. Colony morphology of wild-type V. cholerae (MO10), a ΔcytR mutant (PW324) and a ΔcytRΔvps double mutant (PW329).

B. Biofilm accumulation by wild-type V. cholerae (MO10), a Δvps mutant (PW328), a ΔcytR mutant (PW324) and a ΔcytRΔvps double mutant (pw329). Biofilms stained with crystal violet are shown above, and quantification of crystal violet staining is shown below.

We also confirmed the correlation of the ΔcytR mutation with the ‘super-biofilm’ phenotype by complementation with the V. cholerae cytR gene in trans. We amplified the cytR gene with its native promoter by PCR, sequenced the product fully and cloned the resulting PCR product into the moderate-copy-number plasmid pBR322. The cloned cytR promoter and coding sequence from the V. cholerae O139 strain, MO10, had a nucleotide sequence identical to that of the V. cholerae O1 El Tor strain N16961. As shown in Fig. 3, when the cytR gene was provided in trans, the ΔcytR mutant displayed a smooth colony morphology and biofilm accumulation similar to that of wild-type V. cholerae.

Figure 3.

Figure 3

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Complementation of the rugose colony morphology and increased biofilm accumulation phenotype of a V. cholerae ΔcytR mutant by the V. cholerae cytR gene provided in trans.

A. Colony morphology of the V. cholerae ΔcytR mutant (PW324) transformed with the cloning vector pBR322, wild-type V. cholerae transformed with the cloning vector pBR322 and a V. cholerae ΔcytR mutant (PW324) transformed with the pBR322-based plasmid pAJH3, denoted pcytR, carrying the promoter and coding sequence of V. cholerae cytR.

B. Quantification of biofilm accumulation by wild-type V. cholerae (MO10), a V. cholerae ΔcytR mutant (PW324), a V. cholerae ΔcytR mutant transformed with the plasmid pAJH3, denoted pcytR, and a V. cholerae ΔcytR mutant transformed with the cloning vector pBR322.

Vibrio cholerae CytR is able to regulate transcription of the E. coli udp gene, a member of the E. coli CytR regulon

When cytidine is scarce in the environment, the E. coli CytR protein represses the transcription of genes encoding proteins involved in nucleoside uptake and catabolism. One of these genes, udp, encodes uridine dephosphorylase, an enzyme involved in the conversion of uridine to ribose-1-phosphate and uracil (Barbier and Short, 1992; Brikun et al., 1996; Neidhardt, 1996; Gavigan et al., 1999). Uridine dephosphorylase is operative in the catabolism of both uridine and cytidine. To confirm that the gene we identified as V. cholerae cytR had a function similar to that of the E. coli cytR gene, we tested whether V. cholerae cytR, provided in trans, could repress transcription of the udp gene of E. coli. We compared transcription of a udp–lacZ fusion in a parent E. coli strain containing a wild-type copy of cytR as well as in a derivative cytR mutant transformed with either a control plasmid or a plasmid carrying the wild-type V. cholerae cytR gene (Barbier and Short, 1992). As shown in Fig. 4A, udp–lacZ transcription was derepressed in the E. coli cytR mutant. When the V. cholerae cytR gene was supplied in trans to an E. coli cytR mutant, udp–lacZ transcription was repressed.

Figure 4.

Figure 4

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Functional similarity of E. coli CytR and V. cholerae CytR.

A. Normalized β-galactosidase activity of an E. coli strain containing a chromosomal fusion of lacZ to the E. coli udp promoter (SS6005) transformed with pBR322, an E. coli ΔcytR mutant (SS6018) transformed with pBR322 and the same E. coli ΔcytR mutant transformed with the plasmid pAJH3, denoted pVcytR, carrying the promoter and coding sequence of V. cholerae cytR.

B. Normalized β-galactosidase activity of wild-type V. cholerae (PW387) and V. cholerae ΔcytR mutant (PW386) cells containing a chromosomal fusion of lacZ to the V. cholerae udp promoter grown in LB broth, LB broth supplemented with 0.3% cytidine, LB broth supplemented with 0.3% glucose and LB broth supplemented with 0.3% glycerol.

Vibrio cholerae CytR regulates the V. cholerae udp gene in response to cytidine

We questioned whether CytR regulates nucleoside catabolism in V. cholerae as it does in E. coli. We identified an ORF in the V. cholerae genome (TIGR locus VC1034) whose putative protein product is 75% identical and 89% similar to the Udp protein of E. coli K-12. We constructed a lacZ fusion to the V. cholerae udp promoter and inserted this fusion into the lacZ locus of both wild-type V. cholerae and a ΔcytR mutant. We then assayed β-galactosidase activity in extracts of planktonic cells grown in LB broth alone as well as in LB broth with added cytidine, glucose and glycerol. In all the media used, the growth curves of wild-type V. cholerae and the ΔcytR mutant were similar (data not shown). As shown in Fig. 4B, when these strains were grown in LB broth, udp–lacZ transcription was greater in the V. cholerae ΔcytR mutant than in wild-type V. cholerae. As predicted, the addition of cytidine to LB broth increased udp–lacZ transcription in wild-type cells almost to the level of the V. cholerae ΔcytR mutant. The addition of glucose to the growth medium decreased the difference in udp–lacZ transcription between wild-type V. cholerae and a ΔcytR mutant slightly, whereas the addition of glycerol to the growth medium had no significant effect on udp–lacZ transcription in wild-type V. cholerae and ΔcytR mutant cells.

Phase-contrast microscopy of wild-type V. cholerae and ΔcytR mutant biofilms

Once the biofilm repressor was identified as the V. cholerae CytR protein, the mechanism of biofilm repression and its role in biofilm development was explored further. It has been shown previously that biofilm development by V. cholerae O139 consists of an initial phase of several hours when only transient surface association is seen. After this initial phase, immobilization and accumulation of the cells in a biofilm is observed. This step requires exopolysaccharide synthesis (Watnick et al., 2001).

The colony morphology and biofilm phenotype of the V. cholerae ΔcytR mutant suggests that exopolysaccharide synthesis is derepressed in this mutant. We were interested in determining how the course of biofilm development was altered by this derepression. We hypothesized that increased exopolysaccharide synthesis might (i) shorten or eliminate the initial phase of transient attachment; (ii) increase deposition of bacteria on the surface during the biofilm accumulation phase; and/or (iii) alter the mature biofilm architecture. Video and time-lapse microscopy of the early stages of biofilm development demonstrated that the initial phase of transient attachment was indistinguishable from that of wild-type V. cholerae both in duration and in the types of interactions observed between the bacterium and the surface. In Fig. 5A, we show representative micrographs of early surface immobilization by wild-type V. cholerae and a ΔcytR mutant. These micrographs represent simultaneous experiments in which wells were incubated with similar numbers of either wild-type V. cholerae or ΔcytR mutant cells for 5 h at 27°C. These micrographs demonstrate that the early stages of biofilm development by wild-type V. cholerae and a ΔcytR mutant were indistinguishable. However, as shown in Fig. 5B, after 18 h of incubation with a surface, the biofilm formed by the V. cholerae ΔcytR mutant was confluent and dense, whereas the wild-type V. cholerae biofilm had not yet reached confluence. Thus, biofilm development by the ΔcytR mutant deviates from that by wild-type V. cholerae in the surface accumulation phase but not in the initial phase of transient association.

Figure 5.

Figure 5

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Phase-contrast micrographs comparing batch biofilm development in LB broth by wild-type V. cholerae (MO10) and a V. cholerae ΔcytR mutant (PW324) after 5 h and 18 h of exposure to a polystyrene surface. Bar = 10 µm.

Three-dimensional architecture of V. cholerae ΔcytR mutant biofilms

Previous studies have suggested that bacterial variants or mutants that exhibit increased exopolysaccharide production may form three-dimensional biofilms with an altered architecture (Hentzer et al., 2001). To determine whether the architecture of a V. cholerae ΔcytR mutant biofilm is grossly different from that of wild-type V. cholerae, we used confocal microscopy to visualize the three-dimensional structures of wild-type V. cholerae and ΔcytR mutant biofilms after 2 days of growth. As shown in Fig. 6, horizontal and vertical sections through both wild-type V. cholerae and ΔcytR mutant biofilms demonstrate pillars of bacteria with water channels between. In contrast to the wild-type V. cholerae biofilm, however, the water channels observed in the ΔcytR mutant biofilm are much narrower, and the bacterial pillars are almost confluent. Thus, the density of cells in the ΔcytR mutant biofilm is much greater than that in wild-type biofilms.

Figure 6.

Figure 6

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Confocal micrographs of 2-day-old wild-type V. cholerae (MO10) and V. cholerae ΔcytR mutant (PW324) biofilms. XY-sections through the biofilms are shown on the left (bar = 25 µm), and vertical sections through the biofilms are shown on the right (bar = 10 µm).

Reporter measurements of vps gene transcription

To investigate further the observation that a cytR mutation affects the surface accumulation phase of biofilm development, we measured transcription of one of the identified vps operons in the planktonic and biofilm-associated phases. We inserted a lacZ fusion to the vpsL promoter into the chromosomal lacZ gene of wild-type V. cholerae and a ΔcytR mutant. vpsL is the first gene in one of two large putative operons involved in VPS synthesis (Yildiz et al., 2001). vpsL–lacZ transcription was measured in planktonic and biofilm-associated wild-type V. cholerae and ΔcytR mutant cells. As shown in Fig. 7, transcription of vpsL–lacZ in both wild-type V. cholerae and ΔcytR mutant planktonic cells was low. However, vpsL–lacZ transcription in biofilm-associated cells was seven and 14 times, respectively, that in planktonic cells with the same genetic background. Furthermore, vpsL–lacZ transcription in the V. cholerae ΔcytR biofilm was three times that of vpsL–lacZ transcription in the wild-type V. cholerae biofilm.

Figure 7.

Figure 7

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Normalized β-galactosidase activity of biofilm-associated and planktonic wild-type V. cholerae (PW357) and V. cholerae ΔcytR mutant (PW358) cells carrying a chromosomal fusion of the vpsL promoter to lacZ. Black bars represent β-galactosidase measurements of wild-type V. cholerae, and grey bars represent β-galactosidase measurements of V. cholerae ΔcytR mutants.

Discussion

We have identified a repressor of exopolysaccharide synthesis and biofilm development in V. cholerae and have shown that this repressor is the V. cholerae CytR protein. The E. coli CytR protein is a member of the LacI family of repressors. These repressors respond to increased concentrations of effector molecules by lifting repression of the catabolic pathways relevant to these effector molecules. The previously defined function of CytR in E. coli and other Gram-negative bacteria was repression of the transcription of genes encoding proteins involved in nucleoside uptake and catabolism in response to low nucleoside concentrations (Neidhardt, 1996; Thomsen et al., 1999). Our experiments demonstrate that V. cholerae CytR functions in a manner similar to that of E. coli CytR. When provided in trans, V. cholerae CytR is able to complement an E. coli cytR mutant. The udp gene product, encoding uridine dephosphorylase, is involved in cytidine and uridine catabolism and is regulated by CytR in E. coli. We have shown that V. cholerae CytR represses transcription of the V. cholerae udp gene in response to low environmental cytidine concentrations. Thus, we provide strong evidence that the mechanism of regulation and binding site specificity of V. cholerae CytR is similar to that of E. coli CytR and that V. cholerae CytR is also involved in the repression of nucleoside catabolism in response to low nucleoside concentrations.

We draw two conclusions from our studies of vpsL transcription in wild-type V. cholerae and ΔcytR mutant planktonic and biofilm-associated cells. The first is that, in both planktonic and biofilm-associated cells, V. cholerae CytR regulates VPS synthesis at the level of vps gene transcription. Furthermore, for wild-type V. cholerae, vpsL transcription in the biofilm is greater than vpsL transcription in the planktonic phase. We hypothesize that, as has been observed for other Gram-negative bacteria, surface contact activates transcription of exopolysaccharide synthesis genes in wild-type V. cholerae (Davies et al., 1993; Prigent-Combaret et al., 1999; Sauer and Camper, 2001; Whiteley et al., 2001; Sauer et al., 2002). Because surface contact activation of vpsL is also observed in ΔcytR mutant cells, we conclude that CytR is not a component of the surface-sensing signal transduction cascade. Instead, an independent regulatory circuit must be operative in surface sensing.

Previous studies suggest that wild-type V. cholerae biofilm development includes a period of transient surface association followed by surface immobilization and accumulation (Watnick and Kolter, 1999; Watnick et al., 2001). The initial period of transient surface association by the V. cholerae ΔcytR mutant is indistinguishable from that of wild-type V. cholerae in both its character and duration. Transient surface association has previously been postulated to be important for surface sensing (Watnick et al., 2001). The observation that transient surface association is unaltered in biofilm development by the V. cholerae ΔcytR mutant is consistent with the conclusion stated above that surface contact activation, which occurs during the period of transient surface activation, is functional in the V. cholerae ΔcytR mutant. The period of biofilm accumulation is altered, however, in a V. cholerae ΔcytR mutant. Once biofilm accumulation by the V. cholerae ΔcytR mutant begins, it proceeds rapidly and produces a structure in which cells are much more densely packed together than they are in the wild-type V. cholerae biofilm. We hypothesize therefore that signals that maintain water channels between the pillars of the biofilm, such as those involved in quorum sensing, are over-ridden in the ΔcytR mutant.

To the well-characterized roles of the CytR protein in repression of nucleoside uptake and catabolism, we have now added a novel role for V. cholerae CytR in the repression of an anabolic process, namely synthesis of the V. cholerae exopolysaccharide VPS. The co-regulation of a catabolic process and an anabolic process is particularly thought provoking. One possibility is that derepression of exopolysaccharide synthesis gene transcription in a V. cholerae ΔcytR mutant is simply the indirect result of increased uptake and catabolism of nucleosides, leading to increased carbon and energy stores within the cell. This seems unlikely given the growth medium used in our experiments. The primary carbon and energy sources in LB broth are amino acids, and nucleosides represent ≈ = 0.1% of the dry weight of amino acids in LB. Thus, the carbon and energy stores of a bacterium growing in LB broth would not be significantly altered by derepression of nucleoside uptake and catabolism. Rather, we propose that elevated intracellular cytidine levels may be a signal for surface immobilization and that this signal is transduced by the V. cholerae CytR protein.

Our studies build a bridge between increased levels of cytidine and synthesis of an extracellular polysaccharide matrix leading to surface attachment. These findings are particularly intriguing in view of a recent report that DNA is an important component of the extracellular matrix of P. aeruginosa biofilms and may actually be required for initial biofilm development (Whitchurch et al., 2002). Although it seems improbable that nucleosides would be present in abundance in the aqueous phase of an aquatic environment, we hypothesize that DNA and its catabolites may accumulate within the extracellular matrix of the biofilm. Thus, nucleoside concentrations in and around biofilms may be increased, providing an environmental signal that recruits new bacteria into the biofilm.

Experimental procedures

Bacterial strains and media

The bacterial strains and plasmids used in this study are listed in Table 1. All experiments used Luria–Bertani (LB) broth. Cytidine, glucose and glycerol were used at concentrations of 3 g l−1 where noted. The wild-type V. cholerae strain, MO10, is resistant to streptomycin, as are all mutants derived from this strain. Thus, most biofilm experiments were performed in LB broth supplemented with 100 µg ml−1 streptomycin. Furthermore, biofilm experiments involving strains harbouring a plasmid encoding ampicillin resistance were performed in LB broth supplemented with 150 µg ml−1 ampicillin. Procedures used to construct strains and plasmids for this study are detailed below.

Table 1.

Bacterial strains and plasmids.

Strains or plasmids Genotype Reference/source
E. coli
  β2155 ThrB1004 pro thi strA hsdS lacZΔM15 (F' lacZΔM15 lacIq trajD36 proA+ proB+) ΔdapA::erm pir.:RP4, Ermr, Kmr Dehio and Meyer (1997)
  SM10λpir Thi thr leu tonA lacY supE recA::RP4-2-Tc::Mu λpirR6K; Kmr Miller and Mekalanos (1988)
  SS6005 F− thi leu rpsL Δ(argF-lac)U169 Φ(udp-lac)8 (λRS45) Barbier and Short (1992)
  SS6018 SS6005, cytR::Tn10dTet Barbier and Short (1992)
V. cholerae
  MO10 1992 clinical isolate of V. cholerae O139 from India, Smr Waldor and Mekalanos (1994)
  PW324 MO10ΔcytR, Smr This study
  PW328 MO10Δvps, Smr This study
  PW329 MO10ΔcytRΔvps, Smr This study
  PW357 MO10lacZ::vpsLp→lacZ, Smr This study
  PW358 PW324lacZ::vpsLp→lacZ, Smr This study
  PW386 PW324lacZ::udp1p→lacZ, Smr This study
  PW387 MO10lacZ::udp1p→lacZ, Smr This study
Plasmids
  pBSL180 OriR6K mobRP4 lacI pTac tnp mini-Tn10Km; Kmr, Apr Alexeyev and Shokolenko (1995)
  pWM91 OriR6K OriT sacB lacZα, Apr Metcalf et al. (1996)
  pAJH1 pWM91 carrying a fragment of cytR harbouring an internal, unmarked deletion This study
  pAJH5 pWM91 carrying a fragment of a vps operon harbouring an internal, unmarked deletion This study
  pAJH3 A pBR322 derivative containing the promoter and coding sequence of cytR; Tcr Apr This study
  p6891MCS pBR327 with 8 kbp Sau3A fragment of V. cholerae lacZ gene interrupted by a multiple cloning site Butterton et al. (1995)
  PAJH8 p6891MCS derivative with udp1p→lacZ inserted in the multiple cloning site This study
  pAJH2 p6891MCS derivative with vpsLp→lacZ inserted in the multiple cloning site This study
  pSMC2 pKEN carrying a bright mutant of gfp and 1.8 kb stabilizing fragment from pUC181.8; Apr Bloemberg et al. (1997)
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Construction of Δvps and ΔcytR mutants

A VPS locus containing two large operons required for VPS synthesis has been identified. (Yildiz et al., 2001). One of these, containing vpsL as well as five additional putative ORFs, was selected for deletion. A 400 bp fragment located just upstream of the start codon of vpsL and a 346 bp fragment located 184 bp downstream from the start codon of the sixth ORF were amplified by PCR using the primer pairs P62, P63 and P64, P65 respectively (see Table 2). Primers P63 and P64 included a complementary 15 bp sequence at their 3′ and 5′ ends respectively. These two fragments were joined using the splicing by overlap extension (SOE) technique, resulting in the construction of a fragment with a 6 kb deletion in the vpsL operon (Horton et al., 1990; Lefebvre et al., 1995). The fragment containing the deletion was purified and ligated into pCR2.1TOPO (Invitrogen). This fragment was then removed from pCR2.1TOPO by digestion with SpeI and XhoI and ligated into pWM91 to create the suicide plasmid pAJH5. This plasmid was used to create vpsL operon deletions in the relevant strains by double homologous recombination and sucrose selection as described previously (Donnenberg and Kaper, 1991).

Table 2.

Primers used for PCR.

Primer Location in chromosome I Sequence
Construction of V. cholerae vps deletion
  P62 998815–998834 GGCTGGTCTATGTGG CTTGT
  P63 999215–999195 TTACGAGCGGCCGCATCCCGAAGAATAATTGGAATG
  P64 1005215–1005235 TGCGGCCGCTCGTAATGGTTGGTGATTGAACATCTG
  P65 1005561–1005542 CAACGTCAGCAACATGGCTA
Construction of V. cholerae cytR deletion
  P38 2843448–2843465 CGGATATGGCTGCTGATG
  P39 2844012–2843994 TTACGAGCGGCCGCACTTGAAAGGTTTGAACTCG
  P40 2845064–2845081 TGCGGCCGCTCGTAACTCTATCGAGTGATGTTG
  P41 2845571–2845554 GCTGCAATCCTTGGTTTG
Cloning of V. cholerae cytR
  P55 2845374–2845354 ATTGATCCGCAAGATTTGAGC
  P56 2843961–2843978 TCTCAGCGCTGCGACAGG
Construction of V. cholerae vpsLp→lacZ
  P75 998382–998403 TAGTTCAATCGCGGCTTGTACCACA
  P76 999402–999382 TAGACCCCTAGCAAGGCAACCGAA
Construction of V. cholerae udp1p→lacZ
  P95 1099912–1099931 TAGCGTCTGCTTAAA TTCCGCTC
  P96 1100416–1100435 TAGCACCAAGGTGGA ATACGGTT
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ΔcytR mutants were constructed in a manner similar to that described for the construction of Δvps mutants. Briefly, the primer pairs P38, P39 and P40, P41 were used to amplify a 564 bp fragment immediately upstream of the cytR gene and a 507 bp fragment beginning 23 bp downstream from the putative cytR stop codon. These fragments were ligated using the SOE technique and used to create a deletion in the V. cholerae cytR gene, resulting in strain PW324.

Construction of pAJH3 for complementation studies

In order to determine whether we could complement the phenotype of our ΔcytR mutant with the cytR gene provided in trans, we cloned cytR into the cloning vector pBR322. The cytR gene is 309 bp downstream of the priA gene. Thus, it is most probably transcribed by a promoter positioned directly upstream of its own coding sequence. The primers P55 and P56, which are positioned 20 bp upstream of the stop codon of priA and 80 bp downstream of the stop codon of cytR, respectively, were used to amplify the cytR gene promoter and coding sequence. The amplification product was ligated into pCR2.1-TOPO (Invitrogen) to yield pCR2.1-TOPO:: cytR. This plasmid was digested with EcoRI, and the fragment containing the cytR gene was purified and ligated into pBR322.

Construction of chromosomally based operon fusions of the vpsL promoter (vpsLp)and the udp1 promoter (udp1p) to lacZ

The intergenic region between the divergently transcribed genes, VC0933 and VC0934, was amplified by PCR using the primer pair P75 and P76. This produced a DNA fragment (vpsLp) including 31 bp of the coding region of VC0933 and 140 bp of the coding region of VC0934. The primers were designed with stop codons at either end of the fragment to avoid the generation of chimeric proteins. vpsLp was recovered by cloning into pCR2.1-TOPO (Invitrogen) to create PCR2.-TOPO::vpsLp. PCR2.-TOPO::vpsLp and pUJ10 were digested with XhoI and BamHI. The resulting vpsLp promoter fragment and the pUJ10 fragment harbouring a promoterless lacZ gene were purified and ligated to yield a plasmid-based fusion of vpsLp to the lacZ gene. The resulting plasmid and p6891 multiple cloning site (MCS) were digested with NotI. The liberated vpsLp–lacZ fusion was purified and ligated with p6891MCS to yield the vpsL–lacZ fusion inserted between two homologous fragments of the V. cholerae lacZ gene and oriented opposite to the direction of transcription of the wild-type V. cholerae lacZ gene. The promoter region of this construct was fully sequenced to confirm the absence of mutations introduced by PCR or cloning. This fusion was crossed into the chromosomal locus of the V. cholerae lacZ gene of relevant strains as described previously (Watnick et al., 2001).

The chromosomal udp1p–lacZ fusion was constructed as described for the vpsLp–lacZ fusion except that the region between the putative ORFs with the TIGR designation VC1033 and VC1034 was amplified by PCR using the primer pair P95 and P96.

Transposon mutagenesis and screen for biofilm-altered mutants

Transposon mutagenesis and the screen for biofilm-altered mutants were performed as described previously (Watnick and Kolter, 1999). Briefly, wild-type V. cholerae and the E. coli strain β2155(pBSL180) were crossed on an LB agar plate supplemented with 0.3 mM diaminopimelic acid for 2 h at 37°C. Transposon-insertion mutants were selected on LB agar plates containing kanamycin. The resulting V. cholerae transposon-insertion mutants were replica plated onto LB agar plates and then transferred to LB-filled polyvinylchloride microtitre dish wells using a multipronged device. The bacteria were incubated in these microtitre dish wells at 27°C for 18 h. Wells were then rinsed and stained with crystal violet. Wells that demonstrated an enhanced crystal violet ring were stored at −80°C in LB broth supplemented with 15% glycerol.

Biofilm assays

For observation and quantification of biofilms by crystal violet staining, biofilms were formed either in wells of sterile polystyrene microtitre dishes or in borosilicate glass tubes. Very little difference in bacterial surface accumulation was observed between these two surfaces. Again, biofilms were incubated at 27°C for 18 h. Quantification of surface-adherent cells was measured by crystal violet staining as described previously (Watnick et al., 2001). Briefly, biofilms were stained by incubation with a 1 mg ml−1 aqueous solution of crystal violet for 6 min. Dimethyl sulphoxide (DMSO) was then added immediately to disrupt the biofilm, and an OD595 of the resulting solution was measured.

Phase-contrast and confocal microscopy

Biofilms intended for observation by time-lapse phase-contrast microscopy were formed in 24-well polystyrene microtitre dishes. Wells were filled with 300 µl of LB broth, which allowed adequate aeration of the base of the well and, thus, the formation of a robust biofilm. Biofilm development on the bottoms of wells was recorded either at discrete times, by video microscopy, or at intervals by time-lapse microscopy using an Eclipse TE-200 inverted phase-contrast microscope (Nikon) equipped with an Orca digital CCD camera (Hamamatsu) and VVM-D1 shutter drivers (Uniblitz). A computer equipped with METAMORPH imaging software (Universal Imaging) was used for image acquisition and processing.

For confocal microscopy, bacteria harbouring the plasmid pSMC2 were incubated with coverslips placed vertically in Falcon tubes containing LB broth supplemented with ampicillin as described previously (Watnick et al., 2001). For these experiments, the medium was changed daily. Confocal micrographs were obtained using an Odyssey confocal microscope (Noran) equipped with a graphics work station.

β-Galactosidase measurements

For measurements of gene transcription by the vpsLp promoter in both planktonic and biofilm-associated cells, strains of interest were incubated in 96-well plates at 27°C for 18 h. The planktonic cells from four wells were combined in a separate tube, wells were rinsed with LB broth, and biofilms from these four wells were removed by mechanical disruption and pooled. All experiments were done in triplicate. Planktonic and biofilm-associated cells were gently pelleted and then resuspended in 1.5 ml of Z-buffer (Miller, 1992). Borosilicate beads (1 mm; BioSpec) were added to the cell suspension, and this suspension was agitated using a mini-Beadbeater (BioSpec) for 10 s at 2500 r.p.m. to separate cell aggregates. Although this agitation step was specifically included to disperse biofilm-associated cells, planktonic and biofilm-associated cells were treated similarly. Cells were then lysed by three freeze–thaw cycles at −80°C. A sample of this lysate (200 µl) was set aside for subsequent protein determination using the Coomassie Plus protein assay (Pierce), and ONPG (Sigma), a colorimetric substrate for β-galactosidase, was added to 1 ml of the remaining mixture. All samples were incubated for 18.5 h at 37°C to allow a yellow colour to develop, samples were centrifuged to remove cell debris, and the OD405 of each sample was measured (model 550 microplate reader; Bio-Rad). To obtain relative β-galactosidase activity measurements for each sample, the OD405 was multiplied by 1000 and divided by the calculated protein concentration.

For all measurements of udp–lacZ transcription in E. coli and V. cholerae strains, relevant cells were grown overnight in LB broth at 37°C. Cells were lysed by three freeze–thaw cycles at −80°C. Subsequent measurements of β-galactosidase activity followed the protocol above.

Acknowledgements

We would like to thank Drs Matthew Waldor and Carol Kumamoto for many stimulating discussions and for careful reading of this manuscript. We would also like to acknowledge Dr Anne Kane of the GRASP Center and her staff for expert advice and the preparation of many reagents, which greatly accelerated the course of these experiments. We would like to thank Dr Stephen Short for the generous gift of several E. coli strains and plasmids. Confocal micrographs were obtained at the Tufts Confocal Microscope Facility with the assistance of Dr Robert Willson. This work was supported by NIH K08 AI01588 and an award from the Ellison Medical Foundation to P.I.W., and by the New England Medical Center GRASP Center NIH/NIDDK, P30 DK34928.

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