Figures
Abstract
Vibrio cholerae biofilms contain exopolysaccharide and three matrix proteins RbmA, RbmC and Bap1. While much is known about exopolysaccharide regulation, little is known about the mechanisms by which the matrix protein components of biofilms are regulated. VrrA is a conserved, 140-nt sRNA of V. cholerae, whose expression is controlled by sigma factor σE. In this study, we demonstrate that VrrA negatively regulates rbmC translation by pairing to the 5′ untranslated region of the rbmC transcript and that this regulation is not stringently dependent on the RNA chaperone protein Hfq. These results point to VrrA as a molecular link between the σE-regulon and biofilm formation in V. cholerae. In addition, VrrA represents the first example of direct regulation of sRNA on biofilm matrix component, by-passing global master regulators.
Citation: Song T, Sabharwal D, Gurung JM, Cheng AT, Sjöström AE, Yildiz FH, et al. (2014) Vibrio cholerae Utilizes Direct sRNA Regulation in Expression of a Biofilm Matrix Protein. PLoS ONE 9(7): e101280. https://doi.org/10.1371/journal.pone.0101280
Editor: Thomas Rudel, University of Würzburg, Germany
Received: January 8, 2014; Accepted: June 5, 2014; Published: July 23, 2014
Copyright: © 2014 Song et al. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Funding: SNW was supported by the Swedish Research Council project grants 2006-4702 (VR-NT) and 2013-2392 (VR-MH). BEU was supported by the Swedish Research Council project grants 2010-3031 (VR-MH) and 2012-4638 (VR-NT). The Laboratory for Molecular Infection Medicine Sweden (MIMS) is supported by Umeå University and the Swedish Research Council (353-2010-7074). This work was performed as part of the Umeå Centre for Microbial Research (UCMR) Linnaeus Program supported by Umeå University and the Swedish Research Council (349-2007-8673). Funding website: URL: http://www.vr.se. FHY was supported by the National Institutes of Health (NIH) RO1-AI055987. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.
Competing interests: The authors have declared that no competing interests exist.
Introduction
Vibrio cholerae inhabits aquatic environments and when it enters the human intestine, e. g., through ingestion of contaminated food or water, it causes the severe diarrheal disease, cholera. Vibrios are shown to form biofilms on zooplankton, insects and intestines [1]–[5]. Compared to planktonic cells, bacteria within biofilms are more resistant to stress conditions, e. g., osmotic and oxidative stress, acidity, antibiotics exposure and immune clearance [6]–[12]. Biofilm structures are constructed of and maintained by biofilm matrix components [13]. In V. cholerae, formation of biofilm requires production of exopolysaccharide (VPS) and the biofilm matrix proteins RbmA, RbmC and Bap1 [14]–[18]. These matrix proteins appear to be involved at particular steps during the biofilm formation process. RbmA is involved in the initial cell-cell adhesion step and serves as a tether, forming flexible linkages between cells and the extracellular matrix [18], [19]; Bap1 facilitates adherence of the developing biofilm to surfaces; and the heterogeneous mixtures of VPS, RbmC and Bap1 appear to form envelopes to encase the cell clusters [18]. Without RbmC, incorporation of VPS through the biofilms is significantly reduced, suggesting an essential role for RbmC in maintaining the mature biofilm structure [18].
To date, studies on the regulation of biofilm formation have been mainly focused on VPS synthesis. A complex regulatory network controls transcription of the vps gene in response to multiple environmental signals, such as signals from quorum-sensing bacterial autoinducers [20], polyamines [21], [22], nucleosides [23], [24], indole [25] and nutrient scarcity [26]. Recently, glucose-specific enzyme IIA has also been shown to regulate biofilm formation through binding to a carbon storage regulator homolog MshH, demonstrating a link between the phosphoenolpyruvate phosphotransferase system and biofilm formation [27], [28]. In contrast to the vast body of knowledge about VPS regulation, very little is known about regulation of the matrix proteins (RbmA, RbmC and Bap1). Fong et al [29] has demonstrated the involvement of two factors: the cyclic AMP (cAMP)-cAMP receptor protein (CRP) complex and a transcriptional regulator VpsR. While VpsR positively regulates transcription of the rbm genes, cAMP-CRP appears to negatively regulate rbm expression, both mediated by and independently of VpsR [29].
In the past decade, an increasing body of evidence has highlighted the important and complex roles of small regulatory RNAs (sRNAs) in bacterial physiology and pathogenesis [30], [31]. Many sRNAs are produced in response to specific environmental signals/stresses. They act by base-pairing with target sequences, resulting in up- or down-regulating gene expression through modulating the translation or the turnover of target mRNAs (see review [32]). This mechanism of regulation often requires the RNA chaperone protein Hfq that facilitates base pairing between sRNAs and their target mRNAs [33], [34]. In Vibrio, a σE-dependent sRNA, VrrA, has been shown to be induced by envelope stress and to repress the outer membrane proteins OmpA and OmpT through base pairing to the 5′ untranslated regions (UTR) of the corresponding mRNAs. When the OmpA level decreases, envelope stress is reduced by releasing outer membrane vesicles (OMVs) [35], [36]. These OMVs further protect bacteria against environmental hazards such as UV damage [37]. Using the infant mouse model, VrrA was demonstrated to attenuate V. cholerae virulence [35], which could be partially explained by the VrrA-mediated down-regulation of TcpA, a major V. cholerae virulence factor essential for host colonization. In this study, we provide evidence that VrrA down-regulates the biofilm matrix protein RbmC by base-pairing with the 5′-UTR of rbmC mRNA. Because RbmC is essential for maintaining the mature structure of biofilms, this VrrA-mediated suppression of RbmC might be an additional mechanism of biofilm regulation in V. cholerae.
Results
VrrA down-regulates RbmC independently of Hfq
In our previous studies, VrrA was shown to down-regulate bacterial structural proteins such as OmpA, TcpA and OmpT [35], [36]. When we analyzed the profile of secreted proteins by SDS-PAGE and Coomassie-brilliant-blue staining, we noticed that a protein band at ≈100 kDa was more abundant in the Δhfq background than in the wild-type background (Fig. 1, compare lanes 5–8 to lanes 1–4). Further, this protein appeared to be more abundant in the ΔhfqΔvrrA strain than in the Δhfq strain, and the lower level was restored in the vrrA complemented strain (Fig. 1A, lanes 7 and 8). The protein band, marked with asterisk in Fig. 1A lane 6, was excised from the gel, subjected to mass spectrometry analysis, and identified as the biofilm matrix protein RbmC (VC0930).
(A) Coomassie-brilliant-blue-stained SDS-PAGE gel. (B) Western blot detecting RbmC (upper panel); proteins on the Western blot membrane were stained with Coomassie-brilliant-blue and shown as loading control (lower panel). Supernatant samples were prepared from bacteria cultivated in LB medium at 30°C. Lane 1, A1552 (wild type); lane 2, DNY7 (ΔvrrA); lane 3, DNY11 (ΔvrrA+pvrrA); lane 4, DNY12 (ΔvrrA+pMMB66HE); lane 5, DNY8 (Δhfq); lane 6, DNY9 (ΔhfqΔvrrA); lane 7, DNY16 (ΔhfqΔvrrA+pvrrA); lane 8, DNY17 (ΔhfqΔvrrA+pMMB66HE); lane 9, DHS196 (ΔrbmC). LaProtein marker sizes (lane M) are given to the left in kDa. The asterisk indicates the protein band that was excised from the gel and subjected to mass spectrometry analysis.
In order to detect the low levels of RbmC in the wild-type background, we performed Western blot analysis using anti-RbmC polyclonal antiserum [38]. As expected, the antiserum could detect RbmC in the wild-type strain (Fig. 1B, upper panel, lanes 1–4) while no band was detected in a ΔrbmC mutant (Fig. 1B, upper panel, lane 9), confirming antibody specificity. Similar to what was earlier noticed in the Δhfq background strains, the RbmC level was elevated in the absence of VrrA in the wild-type background strains and this elevated level was also reduced when the ΔvrrA strain was complemented with VrrA expressed from a plasmid (Fig. 1B, upper panel, lanes 1–4). A SDS-PAGE Coomassie blue staining gel was shown (Fig. 1B, lower panel) as a sample loading control. These data indicated that the VrrA-mediated regulation of RbmC expression did occur in the absence of Hfq. This suggests that Hfq is not essential for RbmC repression by VrrA although it is also feasible that Hfq can enhance the repression. We also observed that in the hfq mutant the basal RbmC protein level was higher (compare lane 1 with lane 5 in Fig. 1B, upper panel). The apparent repression by Hfq was presumably not strictly dependent on VrrA and could possibly also be mediated by some other sRNA. The higher basal level of the RbmC protein in the hfq mutant could also be an indirect effect through transcriptional control by a transcriptional regulator that is affected by Hfq.
The 5′ region of rbmC mRNA is responsive to VrrA regulation
In order to further study the interaction between VrrA and the rbmC mRNA, we first determined the transcriptional start site of rbmC by 5′ RACE analysis. After sequencing analysis as described in Material and Methods, the rbmC transcriptional start site was determined to be 125 nt upstream from the AUG start codon.
Our earlier studies on the interaction between VrrA and its targets demonstrated that VrrA represses translation initiation by base-pairing with the 5′-UTR of target mRNAs (ompA, tcpA and ompT). We hypothesized that VrrA would interact similarly with the rbmC mRNA. To test this hypothesis, we used a publically available prediction program, the RNAhybrid algorithm [39], to predict possible RNA duplexes formed by VrrA and the 5′ region of the rbmC mRNA. The query sequence used for rbmC mRNA included the region from the transcriptional start site to 30 nt into the rbmC coding region. As shown in Fig. 2A, RNAhybrid algorithm predicted duplex formation between the residues 91–106 of VrrA and the −8 to −25 region of rbmC mRNA (numbering of rbmC is relative to the AUG start codon). This 13-bp duplex is interrupted by a bulge dividing the stretch into a 7-bp and a 6-bp duplex, with the latter masking the Shine-Dalgarno (SD) region required for translation initiation.
(A) Graphical presentation of the proposed interaction of VrrA sRNA with the 5′-UTR of rbmC sequence, and of compensatory base-pair changes. Numbering for rbmC is relative to start codon AUG (A is +1), and that for VrrA is relative to the +1 transcription start site. The predicted SD sequence of rbmC (AGGGAGU) is underlined. Vertical arrows denote nucleotides introduced into rbmC and VrrA for compensatory base-pair change experiment. (B) Sequences of wild-type VrrA (pTS2) and its nucleotide substitution mutants. Nucleotides that were substituted are underlined.
In order to dissect interacting base pairs, we introduced point mutations into VrrA (Fig. 2B). Plasmid pTS2 is a ColE1-based plasmid expressing vrrA from its own promoter [35]. Substitution of A91C92U93C94C95U96 with U91G92A93G94G95A96, A91C92U93 with U91G92A93, C100U101U102G103U104G105U106 with G100A101A102C103A104C105A106, and C100U101U102 with G100A101A102 generated plasmids pTS2-M7, pTS2-M8, pTS2-M9 and pTS2-M10, respectively. Each plasmid was introduced by transformation into strain DNY7 (ΔvrrA) and sRNA expression from the resulting plasmids were confirmed by Northern blot analysis (Fig. 3A, upper panel). The 5S rRNA was probed as internal control (Fig. 3A, lower panel). Interestingly, the VrrA-M7 level appeared higher than other VrrA variants. To compare the potential structures of these VrrA variants, RNA folding and pattern examination were performed using the Mfold web server [40]. The predicted structure of VrrA-M7 was found to be somewhat different from the predicted structures of the other variants (Fig. 3B). A feasible explanation would be that the VrrA-M7 might be more stable than wild-type VrrA, VrrA-M8, VrrA-M9, and VrrA-M10 due to a structural alteration. Another possible explanation for the higher levels of the VrrA-M7 mutant might be that this mutation could disrupt binding and co-degradation of the sRNA with another target.
(A) Wild-type VrrA is expressed from plasmid pTS2. Mutant variants VrrAM7 to VrrAM10 are expressed from corresponding basepair-substituted plasmids pTS2-M7 to pTS2-M10. All plasmids were transformed into V. cholerae strain DNY7 (ΔvrrA). pJV300 is used as plasmid control for pTS2. The 5S rRNA was probed as an internal control for Northern blot analysis. (B). Potential structures of VrrA variants. RNA folding was performed using the Mfold algorithm [40].
Supernatant proteins of the different sRNA-expressing strains were then analyzed to compare the production of RbmC. As shown in Fig. 4A (upper panel), compared to the wild-type VrrAWT (expressed from pTS2), VrrAM7 (expressed from pTS2-M7) partially lost its ability to repress RbmC production whereas VrrAM8 (expressed from pTS2-M8) could repress RbmC production to the same extent as VrrAWT. In contrast, VrrAM9 and VrrAM10 (expressed from pTS2-M9 and pTS2-M10, respectively) completely lost their ability to repress RbmC production. A SDS-PAGE Coomassie blue staining gel (Fig. 4A, lower panel) was included as the sample loading control. These results show that C100U101U102 in VrrA are important for regulating expression of RbmC.
Western blot analyses of RbmC levels in the culture supernatants of the wild type V. cholerae strain A1552 and vrrA mutant derivatives: (A) Detection of RbmC (upper panel) in the supernatants from DNY7 (ΔvrrA) carrying different plasmids expressing either the wild-type VrrA (from plasmid pTS2) or mutant variants VrrAM1 to VrrAM6 are expressed from plasmids pTS2-M1 to pTS2-M6. A SDS-PAGE Coomassie blue stained gel is shown as the sample loading control (lower panel). (B) Western blot analysis of the RbmC levels in supernatants isolated from DNY7 (ΔvrrA) and DNY189 (ΔvrrA rbmC*) carrying different plasmids (upper panel). A SDS-PAGE gel stained with Coomassie blue is shown as loading (lower panel). Protein marker sizes are given to the left in kDa.
We next introduced mutations in the rbmC 5′-UTR (A−21A−20G−19 to U−21U−20C−19, Fig. 2A), generating the compensatory rbmC* allele. This rbmC* allele was introduced into the chromosome of DNY7 (ΔvrrA) by site-directed mutagenesis. As shown in Fig. 4B, the VrrAM10 variant expressed from plasmid pTS2-M10 lost its ability to repress RbmC production (Fig. 4B, upper panel, lane 3). Likewise, rbmC* was resistant to regulation by the wild-type VrrA expressed from plasmid pTS2 (Fig. 4B, upper panel, lane 5). However, regulation of rbmC* was restored upon expression of the compensatory VrrAM10 allele (Fig. 4B, upper panel, lane 6). A SDS-PAGE Coomassie blue stained gel was used as a sample loading control (Fig. 4B, lower panel). These data suggest that VrrA acts directly as an antisense RNA to repress rbmC mRNA in vivo.
In our earlier study, VrrA mutant variants (VrrAM1 to VrrAM6) expressed from plasmids pTS2-M1 to pTS2-M6 (Fig. 2B) were constructed to study the interaction between VrrA and the ompT mRNA. We showed that VrrA mutant variants covering the VrrA region from residues 69–78 was responsible to base-pair with 5′ UTR of ompT mRNA [36]. In order to see whether these residues would be important for RbmC regulation as well since the residues 69–78 were closed to the interacting region, we monitored RbmC levels in the strains expressing VrrAM1 to VrrAM6 by Westen blot analysis. As shown in Fig. 5 (upper panel) none of these variants lost its ability to repress RbmC, suggesting that ompT- and rbmC-regulating regions in VrrA do not overlap.
Western blot analysis of supernatants from DNY7 (ΔvrrA) carrying the indicated plasmids (upper panel). A SDS-PAGE Coomassie blue staind gel was shown as a sample loading control (lower panel).
VrrA modulates biofilm formation
The findings about the ability of VrrA to down-regulate RbmC levels prompted us to analyze the impact of VrrA on biofilm formation by V. cholerae. We compared the biofilm forming ability using a once-through flow cell system and analysis by confocal laser scanning microscopy (CLSM). The over-expression of VrrA from a plasmid clone in the wild-type strain markedly decreased biofilm formation at 48 h when compared to that of the same strain containing the plasmid vector (Fig. 6A, c and f). Although initial stages of biofilm formation at 2 h and 24 h were not markedly altered by vrrA gene overexpression (Fig. 6A, a and d; b and e), COMSTAT analysis of biofilms developed 48 h post inoculation revealed that total biomass, average and maximum thicknesses of the wild-type strain overexpressing vrrA were markedly decreased after 48 h compared to those of the wild-type strain harboring only the plasmid vector after 48 h although the growth rate and yield were similar between control and over-expression strains. These results show that over-expressing VrrA impairs the ability of V. cholerae to form biofilms.
Cells were grown for 2(a and d), 24 h (b and e); 48 h (c and f) in 2% LB medium in the presence of ampicillin and 0.05% arabinose at room temperature. (B) COMSTAT analysis of biofilms formed by wild-type strains harboring a plasmid vector or vrrA over-expression plasmid.
Discussion
V. cholerae transits between fundamentally different habitats the aquatic environment and the human digestive tract. Such transitions require rapid acquisition and integration of environmental cues in order to coordinate adequate genetic programs and adapt to the new niche. One such adaptation program involves the switch between a planktonic, motile lifestyle and a biofilm-based sessile lifestyle. To date, numerous regulator proteins have been found to affect biofilm formation in V. cholerae, such as those described in the Introduction. Results from this study add a new class of regulators, sRNAs, as a direct regulator of a biofilm matrix component. Through down-regulation of RbmC, VrrA weakens the stability of the mature biofilm structure and might therefore facilitate dispersal of bacteria from a sessile to a planktonic life style. In addition, because expression of VrrA is controlled by sigma factor σE, VrrA serves as a molecular link between the σE-regulon and biofilm formation in V. cholerae.
Several sRNAs have been shown to be involved in biofilm formation in E. coli and Salmonella, e. g. OmrA/B [41], McaS [42], [43], RprA [44] and GcvB [42]. In contrast to VrrA, these sRNAs do not target biofilm matrix components directly, instead they target biofilm master regulators such as CsgD, which in turn regulates biofilm components. This generates a hierarchical regulatory network and enables csgD mRNA to serve as a hub for complex signal integration via multiple sRNAs [45], [46]. Similarly in Vibrio, sRNAs Qrr1-4 and CsrB/C/D regulate the biofilm master regulator HapR or the regulatory molecule cyclic di-GMP (through diguanylate cyclase) [47], [48], and thus are indirectly involved in biofilm formation.
VrrA belongs to a growing family of sRNAs that regulate multiple targets [48], [49]. VrrA uses unique pairing regions to differentially regulate different mRNA targets. Compensatory base pair change experiments revealed that residues C100U101U102 (numbers relative to the +1 transcriptional start site) in VrrA are essential for base-pairing with rbmC mRNA, while those required for the regulation of ompT mRNA are G73C74U75 in VrrA [36].
In addition to the target-specific regulating regions in VrrA, dependency on the chaperon protein Hfq differs among mRNA targets as well. Although deletion of hfq abolishes the interaction between VrrA and ompT mRNA, Hfq is not absolutely required for the regulation on ompA [35] or rbmC mRNAs (this study). The observation that OmpA and RbmC levels were elevated in the Δhfq strain and that VrrA could only partially repress this elevated expression suggests that additional sRNAs are involved in the regulation. The combination of target-specific regions in VrrA and differentiated requirement of Hfq allows VrrA to modulate multiple targets differentially.
According to the RNAhybrid prediction, as shown in Fig. 2, A91C92U93C94C95U96 in VrrA base pairs to the potential SD sequence (AGGGAGU) of rbmC. We therefore expected to see the most drastic change in RbmC level in strains expressing VrrAM7 (substitution of A91C92U93C94C95U96 with U91G92A93G94G95A96) and VrrAM8 (substitution of A91C92U93 with U91G92A93). However, our results showed that VrrAM9 and VrrAM10, which base pairs to the region upstream of the SD sequence, had more impact on the regulation of RbmC. This unexpected result might be due to the fact that the SD sequence was predicted based on the consensus sequence and therefore might not be the exact SD site. Future studies using e. g. toeprint analyses will hopefully identify the actual interaction site(s) between VrrA and rbmC mRNA. Nevertheless, the present results from the compensatory base pair substitution experiment demonstrate that there is a direct interaction between VrrA and rbmC at the region upstream of the putative SD sequence (Fig. 4B).
It is noteworthy that there are only a few functional homologs to VrrA in other Gram-negative bacteria. One such example is the MicA sRNA in Salmonella and E. coli [50], [51]. Both MicA and VrrA are σE-dependent and are capable of down-regulating multiple outer membrane proteins by base-pairing mechanisms [35], [52]. Interestingly, Kint et al [45] observed that MicA in Salmonella was involved in biofilm formation, although the molecular mechanism remains unknown. Systematic searches for MicA targets using bioinformatics prediction tools have not identified yet any biofilm-related genes. Future work will be needed to examine possible interactions between MicA and Salmonella biofilm components such as curli and fimbriae.
In summary, VrrA is the first example of an sRNA molecule that directly targets expression of a biofilm matrix component. Given the similarities between VrrA and its homologs in other Gram-negative bacteria, it is plausible that similar direct regulation exists in other bacteria as well. Because VrrA weakens the stability of the mature biofilm structure, strategies directed towards mechanisms or levels of sRNAs to disturb bacterial biofilm formation may potentially be used to combat biofilm-related infections. Furthermore, in our earlier studies, we showed that the TcpA, one of the colonization factors of V. cholerae, was down-regulated by VrrA (Song et al. 2008). In this study, we demonstrated that the expression of one of the extracellular matrix proteins, RbmC that is important for the biofilm formation by V. cholerae was modulated by VrrA. We hypothesize that at the later stage of V. cholerae infection in the host, bacteria can move away from the epithelial surface and into the fluid-filled lumen of the intestine. During this time, the bacteria may undergo a switch from attachment to the epithelial surface to detachment. This process may be associated with up-regulation of VrrA. We suggest that this transition prepares the bacteria to leave the intestine, for survival in the environment, and for eventual transmission to a new host. This process might be orchestrated by VrrA that can modulate expression of both a colonization factor (Tcp) and attachment factor (RbmC).
Materials and Methods
Oligonucleotides
The complete list of DNA oligonucleotides used for cloning and generating probes in hybridization is provided in Table 1.
Bacterial strains and growth conditions
Strains used in this study are listed in Table 2. V. cholerae El Tor Inaba strain A1552 is referred to as the wild-type throughout this study. V. cholerae strains were grown in LB at 37°C or 30°C, as indicated. Carbenicillin was supplemented at 100 µg ml−1 when appropriate.
DNA manipulations
An in-frame deletion of rbmC in A1552 resulting in strain DHS196 was performed using the method described by Skorupski and Taylor [53]. Primer sequences are summarized in Table 1. The rbmC* allele was introduced into the chromosome of DNY7 (ΔvrrA) by site-directed mutagenesis, resulting in strain DNY189. The site-directed mutagenesis experiment was performed as previously described [36], with the addition of an intermediate step using strain DNY188. Primers TIS-96 and TIS-97 were used to introduce a nucleotide change (from −21AAGGT to −21AAGCT) into DNY7, resulting in strain DNY188; primers TIS98 and TIS-99 were used to introduce nucleotide changes (from −21AAGCT to −21TTCGT) into DNY188, resulting in strain DNY189. The intermediate strain DNY188 contains an AluI restriction site (AGCT), which allows for mutant screening. Generation of GFP-tagged V. cholerae wild-type strain A1552 was performed as described in the earlier studies [16]. A DNA fragment (304 bp) containing the vrrA gene including its putative promoter region was amplified from the A1552 genome and cloned into pBAD18 vector [54]at the EcoR1/Xba1 sites. The resulting plasmid pBAD/vrrA and its vector control (pBAD18) were introduced by transformation into the wild type V. cholerae strain A1552-gfp, resulting WT-gfp/pBAD and WT-gfp/pBAD-vrrA respectively.
Plasmid pTS2 is a ColE1-based plasmid expressing wild-type VrrA from its own promoter [35]. This plasmid served as template for the construction of plasmids pTS2-M7, pTS2-M8, pTS2-M9 and pTS2-M10 that carry the nucleotide changes shown in Fig. 2B. Procedures were performed as described earlier [55], and primers used to introduce nucleotide change are summarized in Table 2.
SDS-PAGE and Western blot analysis
Protein samples were prepared from equal amounts of bacteria cells after overnight growth at 30°C. Bacteria were harvested by centrifugation at 10,000×g for 10 min at 4°C. The culture supernatant fluid was precipitated with 10% trichloroacetic acid (TCA). Briefly, 1 volume (250 µl) of 50% TCA stock was added to 4 volumes (1 ml) of protein sample. The protein-TCA mixture was kept on ice for 15 min, and subsequently the tube was centrifuged at 15,000×g for 5 min. The supernatant was removed and the protein pellet was washed with 200 µl of cold acetone. Finally, the tube was centrifuged at 15,000×g for 5 min and the resulting pellet was dissolved in sample buffer containing 10% glycerol, 0.05% bromophenol blue, 2% SDS, 5% 2-mercaptoethanol, and 10 mM Tris-HCl, pH 6.8. Proteins with known molecular masses (Fermentas) were used as molecular mass markers. SDS-PAGE and Western blotting were carried out according to the methods of Laemmli [56] and Towbin et al. [57]. HRP-conjugated donkey anti-rabbit IgG (Promega, USA) was used as secondary antibody. Detection was performed using ECL Prime Western Blotting Detection Reagent (Amersham or GE Life Sciences, USA). Pre-stained Protein Ladder (SM0679, Fermentas) was used as size standards. Gels were stained with Coomassie brilliant blue.
RNA isolation and Northern blot analysis
RNA samples were prepared as previously described [36] from bacterial cultures grown overnight (14 hr) at 37°C. The RNA was treated with DNase I and quantified on a NanoDrop ND-1000 Spectrophotometer (NanoDrop Technologies, USA). For Northern blot analysis, 10 µg RNA sample was resolved in a polyacrylamide gel and transferred to a Hybond-XL membrane (GE Healthcare, USA) by electro-blotting (1 h, 50 V, 4°C) in a tank blotter. Radiolabeled probes were used to visualize the required mRNA or sRNA. Northern blots were exposed to a phosphorimager screen and scanned on a Storm™ phosphorimager (Molecular Dynamics, USA). Quantification was performed using Quantity One software (Roche, USA). For VrrA and 5S rRNA detection, radio labeled (γ-P32-ATP) oligo probe JVO-8109 and JVO-8106 was used respectively.
5′ RACE analysis
5′ RACE was performed as previously described [55] to determine the transcription start site of rbmC. Total RNA isolated from the wild-type V. cholerae strain A1552 was used to generate cDNA. Oligo TIS-79 (Table 1) was used as rbmC-specific primer in PCR. PCR products were separated on a 2% agarose gel, gel-eluted and used as template for sequencing.
Mass spectrometry peptide sequencing
Proteins of interest were excised from the Coomassie-stained SDS-PAGE gel and analyzed by Alphalyse (Denmark) for mass spectrometry.
Biofilm analysis
Flow cell experiments were carried out according to the procedure previously described [58]. Briefly, overnight-grown cultures of gfp-tagged V. cholerae strains were diluted to an optical density at 600 nm (OD600) of 0.02 in 2% LB (0.02% tryptone, 0.01% yeast extract, 1% NaCl; pH 7.5) containing 100 µg/ml of ampicillin and used to inoculated flow chambers. Flow cell experiments were carried out at room temperature with 2% LB containing ampicillin (100 µg/ml) and arabinose (0.2%, wt/vol). CLSM images of the biofilms were captured with a LSM 5 PASCAL system (Zeiss) at 488 nm excitation and 543 nm emission wavelengths. Three dimensional images of the biofilms were reconstructed using Imaris software (Bitplane) and quantified using COMSTAT (Heydorn and Molin, 2000). Flow cell experiments were carried out with at least two biological replicates.
Acknowledgments
This work was performed within the Umeå Centre for Microbial Research (UCMR) Linnaeus Program and was supported by grants from the Swedish Research Council, the Faculty of Medicine at Umeå University and the National Institutes of Health (NIH).
Author Contributions
Conceived and designed the experiments: TS DS JMG AES ATC FHY BEU SNW. Performed the experiments: TS DS JMG AES ATC. Analyzed the data: TS DS JMG AES ATC FHY BEU SNW. Contributed reagents/materials/analysis tools: FHY BEU SNW. Wrote the paper: TS DS ATC FHY BEU SNW.
References
- 1. Huq A, Small EB, West PA, Huq MI, Rahman R, et al. (1983) Ecological relationships between Vibrio cholerae and planktonic crustacean copepods. Appl Environ Microbiol 45: 275–283.
- 2. Huq A, Xu B, Chowdhury MA, Islam MS, Montilla R, et al. (1996) A simple filtration method to remove plankton-associated Vibrio cholerae in raw water supplies in developing countries. Appl Environ Microbiol 62: 2508–2512.
- 3. Yildiz FH, Visick KL (2009) Vibrio biofilms: so much the same yet so different. Trends Microbiol 17: 109–118.
- 4. Purdy AE, Watnick PI (2011) Spatially selective colonization of the arthropod intestine through activation of Vibrio cholerae biofilm formation. Proc Natl Acad Sci U S A 108: 19737–19742.
- 5. Broza M, Halpern M (2001) Pathogen reservoirs. Chironomid egg masses and Vibrio cholerae. Nature 412: 40.
- 6. Parsek MR, Singh PK (2003) Bacterial biofilms: an emerging link to disease pathogenesis. Annu Rev Microbiol 57: 677–701.
- 7. Islam MS, Jahid MI, Rahman MM, Rahman MZ, Islam MS, et al. (2007) Biofilm acts as a microenvironment for plankton-associated Vibrio cholerae in the aquatic environment of Bangladesh. Microbiol Immunol 51: 369–379.
- 8. Faruque SM, Biswas K, Udden SM, Ahmad QS, Sack DA, et al. (2006) Transmissibility of cholera: in vivo-formed biofilms and their relationship to infectivity and persistence in the environment. Proc Natl Acad Sci U S A 103: 6350–6355.
- 9. Costerton JW, Stewart PS, Greenberg EP (1999) Bacterial biofilms: a common cause of persistent infections. Science 284: 1318–1322.
- 10. Stewart PS, Costerton JW (2001) Antibiotic resistance of bacteria in biofilms. Lancet 358: 135–138.
- 11. Anderson GG, O'Toole GA (2008) Innate and induced resistance mechanisms of bacterial biofilms. Curr Top Microbiol Immunol 322: 85–105.
- 12. Tamayo R, Patimalla B, Camilli A (2010) Growth in a biofilm induces a hyperinfectious phenotype in Vibrio cholerae. Infect Immun 78: 3560–3569.
- 13. Flemming HC, Wingender J (2010) The biofilm matrix. Nat Rev Microbiol 8: 623–633.
- 14. Yildiz FH, Schoolnik GK (1999) Vibrio cholerae O1 El Tor: identification of a gene cluster required for the rugose colony type, exopolysaccharide production, chlorine resistance, and biofilm formation. Proc Natl Acad Sci U S A 96: 4028–4033.
- 15. Fong JC, Yildiz FH (2007) The rbmBCDEF gene cluster modulates development of rugose colony morphology and biofilm formation in Vibrio cholerae. J Bacteriol 189: 2319–2330.
- 16. Fong JC, Karplus K, Schoolnik GK, Yildiz FH (2006) Identification and characterization of RbmA, a novel protein required for the development of rugose colony morphology and biofilm structure in Vibrio cholerae. J Bacteriol 188: 1049–1059.
- 17. Fong JC, Syed KA, Klose KE, Yildiz FH (2010) Role of Vibrio polysaccharide (vps) genes in VPS production, biofilm formation and Vibrio cholerae pathogenesis. Microbiology 156: 2757–2769.
- 18. Berk V, Fong JC, Dempsey GT, Develioglu ON, Zhuang X, et al. (2012) Molecular architecture and assembly principles of Vibrio cholerae biofilms. Science 337: 236–239.
- 19. Giglio KM, Fong JC, Yildiz FH, Sondermann H (2013) Structural Basis for Biofilm Formation via the Vibrio cholerae Matrix Protein RbmA. J Bacteriol 195: 3277–3286.
- 20. Hammer BK, Bassler BL (2003) Quorum sensing controls biofilm formation in Vibrio cholerae. Mol Microbiol 50: 101–104.
- 21. Karatan E, Duncan TR, Watnick PI (2005) NspS, a predicted polyamine sensor, mediates activation of Vibrio cholerae biofilm formation by norspermidine. J Bacteriol 187: 7434–7443.
- 22. McGinnis MW, Parker ZM, Walter NE, Rutkovsky AC, Cartaya-Marin C, et al. (2009) Spermidine regulates Vibrio cholerae biofilm formation via transport and signaling pathways. FEMS Microbiol Lett 299: 166–174.
- 23. Haugo AJ, Watnick PI (2002) Vibrio cholerae CytR is a repressor of biofilm development. Mol Microbiol 45: 471–483.
- 24. Lim B, Beyhan S, Yildiz FH (2007) Regulation of Vibrio polysaccharide synthesis and virulence factor production by CdgC, a GGDEF-EAL domain protein, in Vibrio cholerae. J Bacteriol 189: 717–729.
- 25. Mueller RS, Beyhan S, Saini SG, Yildiz FH, Bartlett DH (2009) Indole acts as an extracellular cue regulating gene expression in Vibrio cholerae. J Bacteriol 191: 3504–3516.
- 26. Liang W, Silva AJ, Benitez JA (2007) The cyclic AMP receptor protein modulates colonial morphology in Vibrio cholerae. Appl Environ Microbiol 73: 7482–7487.
- 27. Pickering BS, Smith DR, Watnick PI (2012) Glucose-specific enzyme IIA has unique binding partners in the Vibrio cholerae biofilm. MBio 3: e00228-00212.
- 28. Houot L, Watnick PI (2008) A novel role for enzyme I of the Vibrio cholerae phosphoenolpyruvate phosphotransferase system in regulation of growth in a biofilm. J Bacteriol 190: 311–320.
- 29. Fong JC, Yildiz FH (2008) Interplay between cyclic AMP-cyclic AMP receptor protein and cyclic di-GMP signaling in Vibrio cholerae biofilm formation. J Bacteriol 190: 6646–6659.
- 30. Bartel DP (2009) MicroRNAs: target recognition and regulatory functions. Cell 136: 215–233.
- 31. Battesti A, Majdalani N, Gottesman S (2011) The RpoS-mediated general stress response in Escherichia coli. Annu Rev Microbiol 65: 189–213.
- 32. Hoe CH, Raabe CA, Rozhdestvensky TS, Tang TH (2013) Bacterial sRNAs: Regulation in stress. Int J Med Microbiol 303: 217–229.
- 33. Panja S, Schu DJ, Woodson SA (2013) Conserved arginines on the rim of Hfq catalyze base pair formation and exchange. Nucleic Acids Res
- 34. De Lay N, Schu DJ, Gottesman S (2013) Bacterial small RNA-based negative regulation: Hfq and its accomplices. J Biol Chem 288: 7996–8003.
- 35. Song T, Mika F, Lindmark B, Liu Z, Schild S, et al. (2008) A new Vibrio cholerae sRNA modulates colonization and affects release of outer membrane vesicles. Mol Microbiol 70: 100–111.
- 36. Song T, Sabharwal D, Wai SN (2010) VrrA mediates Hfq-dependent regulation of OmpT synthesis in Vibrio cholerae. J Mol Biol 400: 682–688.
- 37. Song T, Wai SN (2009) A novel sRNA that modulates virulence and environmental fitness of Vibrio cholerae. RNA Biol 6: 254–258.
- 38. Duperthuy M, Sjostrom AE, Sabharwal D, Damghani F, Uhlin BE, et al. (2013) Role of the Vibrio cholerae matrix protein Bap1 in cross-resistance to antimicrobial peptides. PLoS Pathog 9: e1003620.
- 39. Rehmsmeier M, Steffen P, Hochsmann M, Giegerich R (2004) Fast and effective prediction of microRNA/target duplexes. RNA 10: 1507–1517.
- 40. Zuker M (2003) Mfold web server for nucleic acid folding and hybridization prediction. Nucleic Acids Res 31: 3406–3415.
- 41. Holmqvist E, Reimegard J, Sterk M, Grantcharova N, Romling U, et al. (2010) Two antisense RNAs target the transcriptional regulator CsgD to inhibit curli synthesis. EMBO J 29: 1840–1850.
- 42. Jorgensen MG, Nielsen JS, Boysen A, Franch T, Moller-Jensen J, et al. (2012) Small regulatory RNAs control the multi-cellular adhesive lifestyle of Escherichia coli. Mol Microbiol 84: 36–50.
- 43. Thomason MK, Fontaine F, De Lay N, Storz G (2012) A small RNA that regulates motility and biofilm formation in response to changes in nutrient availability in Escherichia coli. Mol Microbiol 84: 17–35.
- 44. Mika F, Busse S, Possling A, Berkholz J, Tschowri N, et al. (2012) Targeting of csgD by the small regulatory RNA RprA links stationary phase, biofilm formation and cell envelope stress in Escherichia coli. Mol Microbiol 84: 51–65.
- 45. Kint G, De Coster D, Marchal K, Vanderleyden J, De Keersmaecker SC (2010) The small regulatory RNA molecule MicA is involved in Salmonella enterica serovar Typhimurium biofilm formation. BMC Microbiol 10: 276.
- 46. Mika F, Hengge R (2013) Small Regulatory RNAs in the Control of Motility and Biofilm Formation in E. coli and Salmonella. Int J Mol Sci 14: 4560–4579.
- 47. Lenz DH, Miller MB, Zhu J, Kulkarni RV, Bassler BL (2005) CsrA and three redundant small RNAs regulate quorum sensing in Vibrio cholerae. Mol Microbiol 58: 1186–1202.
- 48. Zhao X, Koestler BJ, Waters CM, Hammer BK (2013) Post-transcriptional activation of a diguanylate cyclase by quorum sensing small RNAs promotes biofilm formation in Vibrio cholerae. Mol Microbiol
- 49. Shao Y, Bassler BL (2012) Quorum-sensing non-coding small RNAs use unique pairing regions to differentially control mRNA targets. Mol Microbiol 83: 599–611.
- 50. Udekwu KI, Darfeuille F, Vogel J, Reimegard J, Holmqvist E, et al. (2005) Hfq-dependent regulation of OmpA synthesis is mediated by an antisense RNA. Genes Dev 19: 2355–2366.
- 51. Papenfort K, Pfeiffer V, Mika F, Lucchini S, Hinton JC, et al. (2006) SigmaE-dependent small RNAs of Salmonella respond to membrane stress by accelerating global omp mRNA decay. Mol Microbiol 62: 1674–1688.
- 52. Vogel J, Papenfort K (2006) Small non-coding RNAs and the bacterial outer membrane. Curr Opin Microbiol 9: 605–611.
- 53. Skorupski K, Taylor RK (1996) Positive selection vectors for allelic exchange. Gene 169: 47–52.
- 54. Guzman LM, Belin D, Carson MJ, Beckwith J (1995) Tight regulation, modulation, and high-level expression by vectors containing the arabinose PBAD promoter. J Bacteriol 177: 4121–4130.
- 55. Urban JH, Vogel J (2007) Translational control and target recognition by Escherichia coli small RNAs in vivo. Nucleic Acids Res 35: 1018–1037.
- 56. Laemmli UK (1970) Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227: 680–685.
- 57. Towbin H, Staehelin T, Gordon J (1979) Electrophoretic transfer of proteins from polyacrylamide gels to nitrocellulose sheets: procedure and some applications. Proc Natl Acad Sci U S A 76: 4350–4354.
- 58. Shikuma NJ, Fong JC, Odell LS, Perchuk BS, Laub MT, et al. (2009) Overexpression of VpsS, a hybrid sensor kinase, enhances biofilm formation in Vibrio cholerae. J Bacteriol 191: 5147–5158.
- 59. Yildiz FH, Schoolnik GK (1998) Role of rpoS in stress survival and virulence of Vibrio cholerae. J Bacteriol 180: 773–784.
- 60. Furste JP, Pansegrau W, Frank R, Blocker H, Scholz P, et al. (1986) Molecular cloning of the plasmid RP4 primase region in a multi-host-range tacP expression vector. Gene 48: 119–131.