The SsrA/B TCRS is activated by environmental stimuli such as pH and osmolality, stimulating the expression of virulence genes essential for intra-cellular growth and survival of Salmonella (Feng et al., 2003; 2004). Thus, it was surprising that SsrA/B and SPI-2 were implicated in the multi-cellular lifestyle of Salmonella (Hamilton et al., 2009). Furthermore, the SsrA kinase was required, but SPI-2 genes were down-regulated. At SPI-2, SsrB~P de-represses H-NS and also activates transcription (Walthers et al., 2007; 2011). We therefore wanted to explore this seeming paradox during biofilm regulation.

In order to quantify the defect in biofilm-forming ability resulting from the absence of ssrB, the wild type strain 14028s, an ssrA null strain and an ssrB null strain were grown in 96-well polystyrene microtiter plates for 2 days. The surface-attached cells in 14028s, ssrA and ssrB strains were stained by crystal-violet and quantified. As shown in the figure, biofilm levels in the ssrB mutant strain decreased by around 60% compared to the wild type or an ssrA null strain (Figure 1A), and it could be complemented by over-expressing SsrBc in trans. Thus, we establish a new role for SsrB, but not the SsrA kinase in positively regulating biofilm formation. The ssrB strain was not compromised for planktonic growth, as measured by total viable counts of the wild type, ssrA and ssrB cultures (Figure 1—figure supplement 1 and Supplementary methods).

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The ability to form biofilms was analysed for three strains, wild type 14028s, an ssrA null strain and an ssrB null strain, by observing appearance of the rough dry and red (rdar) morphotype on Congo Red plates. After seven days, each of the 14028s and ssrA mutant plates showed the appearance of rough, dry and red macrocolonies. However, the ssrB null strain exhibited a smooth, wet and pale brown morphotype which showed a poor ability to exhibit a multicellular phenotype (Figure 1B). The absence of Congo Red staining in the ssrB strain also indicated that there were greatly reduced levels of extracellular curli fibers in the macrocolony. These fibers form one of the key components of the extra-cellular matrix of a Salmonella biofilm. To examine the levels of cellulose, the other main component of the extra-cellular matrix, the three strains were grown on Calcofluor plates. The macrocolonies formed by 14028s and ssrA appeared white under UV, as cellulose binds the Calcofluor fluorescent dye (Figure 1C). The dull and non-white macrocolony produced by the ssrB null strain was starkly different, indicating again a poor extracellular matrix.

We visualized the nature of biofilms formed by the wild type and ssrB strains by Confocal Fluorescence Imaging of solid-surface biofilms formed under a continuous flow system in a micro-fluidic chamber. Wild type and the ssrB null strain were grown in minimal M9 medium and inoculated in their respective flow cells. After allowing the cells to attach for 1h, flow commenced and the growing biofilms were monitored daily for six days. Each day, the flow cell was removed, and the glass-attached cells were stained with SYTO-9 green, a fluorescent dye that stains nucleic acids. The cells were then imaged using fluorescence microscopy (Figure 1D). The figure clearly shows that for the wild type strain, the attached biomass not only increased progressively in size, but also in its complexity of design. This correlated with the appearance of typical mushroom-like constituent microcolonies by around 2 days. Such type of mature biofilms were never observed in the ssrB null strain and even after 6 days it failed to form biofilms, as evident by fewer and thinly attached clusters of SYTO-9 green-labeled cells.

To compare the differences in the structural properties of biofilms formed by the ssrB strain with the wild type and the ssrA mutant, cells were grown on APTES-coated cover slips in 16-well polystyrene microtiter plates for two days. The surface-attached communities were fixed, dehydrated and dried for Scanning Electron Microscopy (SEM). As expected, the wild type and ssrA mutant showed the presence of tightly knit communities of cells containing an extensive network of curli fibres surrounding large groups of cells (Figure 1E). In contrast, the ssrB null strain formed sparsely spaced groups of few cells with a large reduction in the surrounding matrix and poor network properties.

It was possible that the ssrB strain was just slower in biofilm formation and needed more time to exhibit its biofilm capability or that it caused a growth defect. We ruled out this possibility by measuring growth curves (Figure 1—figure supplement 1) and by monitoring the wild type and ssrB strains by crystal-violet staining every 12 hr for around 4 days. This determined that the ssrB mutant was not deficient for growth and remained incapable of forming biofilms up to 84 h, while the wild type formed large biofilms by 2 days (Figure 1F).

We next investigated the ability of Salmonella to form biofilms, on cholesterol-coated eppendorf tubes. The cholesterol-attached biofilms were estimated by crystal violet staining for the strains 14028s, ssrA null and ssrB null. There was a drastic reduction in the amount of biofilms formed by the ssrB strain on cholesterol-coated eppendorf tubes (Figure 1G). If cholesterol biofilms are indeed an indicator of an ability to form biofilms on gallstones (Crawford et al., 2008), then SsrB may be crucial for establishment of the carrier state in Salmonella.

Taken together, our results modify the initial hypothesis that SsrA was involved in the formation of biofilms. SsrB, but not its cognate kinase SsrA, is involved in the switch from planktonic growth to a multi-cellular lifestyle, in addition to its role in regulating genes required for pathogenesis.

Phospho-relay between the sensor kinase, SsrA, and the response regulator, SsrB, was crucial for activation of SPI-2 virulence genes (Feng et al., 2003). SsrA is a tripartite sensor kinase that is presumably autophosphorylated by ATP at His405, followed by intra-molecular phosphorylation reactions at Asp739 and His867, based on homology to other tripartite kinases. His867 would then participate in the transfer reaction of the phosphoryl group to Asp56 in the N-terminus of SsrB. To confirm that SsrA-dependent phosphorylation was not required for SsrB-mediated regulation of biofilms (Figure 1A–G), we examined biofilm formation of Salmonella strains possessing H405Q, D739A and H867Q mutations in ssrA (Walthers and Kenney unpublished) and compared them to the wild type, ssrA and ssrB null mutants. Figure 3A indicates that the extent of biofilms formed by the three kinase point mutants was comparable to that of the wild type and the ssrA null mutant. A complete loss of ssrB was the only genetic change which adversely affected biofilm formation.

In the absence of SsrA kinase activity, small inorganic phosphate donors such as acetyl phosphate can phosphorylate SsrB, albeit with a lower efficiency (Feng et al., 2004). We therefore constructed strains that were deficient in both SsrA kinase activity as well as in the production of acetyl phosphate and assayed their biofilm capabilities. Strains H405Q ack pta, D739A ack pta and H867Q ack pta, were all capable of biofilm formation (Figure 2A), demonstrating that none of the known phosphate donors for SsrB were required for regulating biofilms. In contrast, SPI-2 activation required ssrA, as measurement of β-galactosidase activity of a sifA-lacZ transcriptional fusion confirmed that activation of sifA required SsrA/B phosphorylation (Figure 2C).

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SsrB is in the NarL subfamily of response regulators. NarL requires phosphorylation to relieve inhibition of C-terminal DNA binding by the N-terminus (Baikalov et al., 1996). Therefore, we substituted the conserved Asp56 residue with Ala (D56A), and determined its ability to form biofilms (Figure 2B). The D56A strain was as competent as the wild type, ssrA null and ssrA kinase mutants in biofilm formation. The ability of unphosphorylated SsrB to regulate biofilm formation was unique, as it was unable to activate sifA-lacZ levels. The beta-galactosidase activity was significantly lower in the ssrB null and D56A strains (8.7% and 14.9%, respectively) compared to the wild type (Figure 2C). The D56A mutant was also able to form rdar macrocolonies on Congo Red plates. In addition, SEM imaging of D56A biofilms revealed that they maintained their intricate architecture and wild type structure (data not shown).

When cultures of the wild type, ssrA, ssrB and D56A strains were separated as a non-adherent (or free-swimming) sub-population and an adherent sub-population (or multicellular aggregates) (MacKenzie et al., 2015), we observed that the ssrB strain had a greater number of cells in the non-adherent fraction and fewer in the adherent fraction as compared to the other three strains. This confirms that the absence of SsrB, and not SsrB~P, led to a defect in the ability of Salmonella to form surface-attached communities (Figure 1—figure supplement 2 and 3).

Taken together, these results indicate that SsrB can activate biofilm formation in the absence of any phosphorylation signals. This is the first evidence for a role of unphosphorylated SsrB in gene regulation. These results also strongly advocate for a larger contribution of SsrB in Salmonella pathogenesis owing to its dual regulation of intracellular (i.e., the SCV) as well as mutli-cellular lifestyles (the carrier state).

SsrB is encoded on SPI-2 and regulates SPI-2 genes involved in assembly and function of the type three secretion system encoded by SPI-2, as well as effectors that are encoded on SPI-2 and outside of SPI-2 (Feng et al., 2004; Walthers et al., 2007). To determine whether the SsrB target of biofilm regulation was dependent on the SPI-2 injectisome or any of its secreted SPI-2 effectors, we examined ssaC and ssaJ null strains for their ability to form biofilms (Figure 3A). SsaC is an outer ring component of the injectisome and SsaJ forms the inner ring of the SPI-2 needles. Both of these strains formed biofilms to an extent similar to the wild type and hence, we ruled out any involvement of the SPI-2 secretory apparatus or its related secreted proteins in the SsrB-dependent regulation of Salmonella biofilms.

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Biofilm formation in Escherichia coli and Salmonella Typhimurium is governed by the master regulator, CsgD, that acts as a transcriptional activator of genes involved in curli biogenesis and cellulose synthesis. Some of the environmental conditions that favor the formation of biofilms in Salmonella such as low salt and acidic pH also up-regulate levels of SsrB in the cell (Feng et al., 2003). An obvious null hypothesis was therefore to test whether SsrB (in its unphosphorylated state), activated the expression of csgD. We first over-expressed csgD from a plasmid in the ssrB null mutant and determined whether it could rescue the defect in biofilm formation. The presence of csgD in trans restored the biofilm capability to wild type levels as measured by crystal-violet staining of 2 day old biofilms (Figure 3B).

We next examined the levels of csgD transcripts in 2 day macrocolonies formed by the wild type, ssrA null, ssrB null and D56A strains by quantitative real time RT-PCR (qRT-PCR). As shown in Figure 3C, there was around a 60-fold decrease in csgD transcripts when ssrB was deleted (Figure 3C). We also corroborated our previous findings (Figure 1A and 2B), as csgD transcripts in the ssrA null and D56A strains were maintained at levels similar to the wild type. Furthermore, we probed the whole-cell lysates obtained from such macrocolonies with a monoclonal anti-CsgD antibody in order to measure the CsgD protein levels by western blotting. CsgD levels were undetected in the ssrB null compared to the wild type, ssrA null or D56A strains (Figure 3D). Hence, unphoshorylated SsrB was able to up-regulate the expression of csgD and positively influence the shift to a sessile lifestyle.

Both in E. coli and S. Typhimurium, the expression of csgD is sensitive to various environmental stimuli such as starvation, temperature, pH and osmolality due to the action of upstream global regulators at the intergenic region of the csgDEFG and csgBAC operons. We noted that the SsrA/B system also responded to similar environmental conditions such as acidic pH and low osmolality. Furthermore, H-NS, a known repressor of SPI-2 genes (Walthers et al., 2007), was also known to regulate the expression of csgD in E. coli and S. Typhimurium (Ogasawara et al., 2010; Gerstel et al., 2003). In regulating SPI-2 genes, SsrBc and H-NS work in an opposing fashion (Walthers et al., 2011). SsrBc antagonises the repressive action of H-NS at regulatory regions upstream of the promoters, while also acting as a direct transcriptional activator. We therefore examined whether or not this paradigm for SsrB-mediated transcriptional activation was applicable to csgD regulation.

In order to test this, we completely deleted hns from the ssrB null strain and determined if its deletion rescued biofilm formation. Crystal-violet staining of static biofilms formed by the wild type, ssrB and ssrB hns strains was performed; the amount of biofilms formed by ssrB hns was the same as the wild type. In contrast, the ssrB null strain remained at around 35% of these levels (Figure 4A). Macrocolonies formed by the ssrB hns strain also displayed a rugose morphology, reminiscent of the wild type strain after 2 days (Figure 4B). In addition, SEM images of 2 day-old biofilms formed by the ssrB hns strain showed a typical ‘biofilm’ architecture, i.e., densely packed communites of cells surrounded by biofilm matrix (Figure 4C). This result indicated that H-NS was functioning to repress csgD (see Discussion). Thus, we strengthened our prediction of a role for SsrB in the activation of csgD expression by acting as an anti-H-NS molecule.

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We next examined the levels of csgD transcripts in mature biofilms formed by the wild type, ssrB null, hns null and ssrB hns null strains by qRT-PCR (Figure 4D). Normalized levels of csgD transcripts were highly up-regulated in the ssrB hns double mutant compared to the ssrB null strain, indicating that the loss of H-NS repression rescued the defect in csgD expression in the absence of SsrB. Expression of csgD was slightly higher than the wild type levels in the hns single mutant and the ssrB hns double mutant, which also correlated with their biofilm capabilities (Figure 4A). Since the levels of csgD transcripts were equal in the hns and ssrB hns null strains, it seemed likely that SsrB influenced csgD expression by silencing H-NS-mediated repression and not by direct activation of transcription, which requires phosphorylation for a productive interaction with RNA polymerase (Walthers et al., 2011).

Taken together, these results identified a regulatory role for unphosphorylated SsrB by orchestrating anti-silencing at the H-NS-repressed csgD locus. This work clearly shows that, as observed in E. coli, H-NS represses expression of csgD in Salmonella (see Discussion).

SsrB is a NarL family member and X-ray crystallography suggested that phosphorylation was required for DNA binding (Baikalov et al., 1996). Thus, it was of interest to determine how SsrB relieved H-NS silencing at csgD. We used atomic force microscopy (AFM) to visualize the csgDEFG-csgBAC intergenic region and examined the effect of addition of full-length SsrB, D56A SsrB and SsrBc (DNA binding domain alone). Surprisingly, we observed binding of SsrB, D56A SsrB and SsrBc to distinct regions of the csgD regulatory region (Figure 5B and Figure 5—figure supplement 1A and B). Binding occured at low protein concentrations, suggesting high affinity interactions and showed that unphosphorylated SsrB was capable of binding csgD. An SsrB mutant, K179A, that was incapable of binding DNA (Carroll et al., 2009) did not bind csgD even at 300 nM, indicting that binding was specific (Figure 5E and F). Closer examination revealed a sharp curvature at the regions where SsrB was bound to csgD (Figure 5B, arrows). This result indicates that, like its NarL homologue (Maris et al., 2002), SsrB bends DNA upon binding, as predicted (Carroll et al., 2009). We estimate that on average, binding of SsrB to the csgDEFG-csgBAC intergenic sequence occurred with a bending angle of around 82° (Figure 5D and Figure 5—figure supplement 2 for naked DNA). The observation that SsrB bending was more severe than NarL (82° compared to the 42° of NarLc at its cognate palindrome) (Maris et al., 2002), likely contributes to its ability to promote anti-silencing.

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When the SsrB concentration was increased to 300 nM, we detected large-scale condensation of the DNA-protein complexes by AFM (Figure 5C). Condensation at the csgD regulatory region was similar when incubated with 300 nM D56A SsrB or SsrBc, indicating that protein binding led to DNA structural changes, irrespective of whether SsrB was phosphorylated (Figure 5—figure supplement 1C and D). Biochemical analysis of the protein-DNA complexes by an electrophoretic mobility shift assay using a shorter fagment of the csgD regulatory region (Figure 6—figure supplement 4), also indicated the presence of an SsrB-DNA complex. This complex was dissociated when an unlablelled csgD fragment was added as a competitor. In contrast, K179A SsrB failed to form a complex with DNA (Figure 5—figure supplement 3). The DNA bending ability of SsrB is important for SsrB relief of H-NS-mediated transcriptional silencing at csgD (see below and [Winardhi et al., 2015]). The DNA binding behavior of SsrB was recapitulated in solution AFM imaging experiments, indicating that binding and condensation were not an artifact of drying the samples (Figure 6—figure supplement 1C).

Our previous work established that H-NS silenced genes by forming a rigid filament on DNA (Liu et al., 2010; Walthers et al., 2011; Lim et al., 2012). We purified H-NS, incubated it with the csgD regulatory region, and immobilised it on a glass coverslip. Subsequent AFM imaging indicated the presence of a straight and rigid nucleoprotein filament (Figure 6A(i)), which was distinct from the random conformation adopted by naked DNA (Figure 5A). Thus, we reaffirmed our earlier observations that H-NS repressed expression of csgD by filament formation, leading to transcriptional silencing (Figure 4D).

When we pre-formed stiffened filaments by addition of H-NS and then added SsrB, evidence of SsrB condensation was immediately apparent (Figure 6A(ii)). For example, areas of SsrB binding to DNA led to the formation of condensed nucleoprotein complexes and abolished the prior structural rigidity introduced by H-NS binding (see arrows, Figure 6A(ii)). Interestingly, Figure 6A(ii) also indicates that H-NS continued to form straight nucleoprotein complexes in regions devoid of SsrB (yellow line), i.e. H-NS was still bound to some regions of DNA when SsrB was also bound. Similar binding behavior with D56A SsrB and SsrBc at the H-NS-bound csgD regulatory region was also observed (Figure 6—figure supplement 2A and B), and similar binding patterns were observed in solution AFM (Figure 6—figure supplement 1B–D). This combined nucleoprotein complex was also detected as a supershift by electrophoretic mobility shift assay using a shorter fragment of the csgD regulatory region and specific concenterations of H-NS, SsrB and anti-SsrBc serum (Figure 6—figure supplement 3). This csgD regulatory element harbored the H-NS binding region (Gerstel et al., 2003) as well as an SsrB binding motif (Feng et al., 2004; refer Figure 6—figure supplement 4). Thus, unphosphorylated SsrB can bind to the csgD regulatory region when it has been coated with the repressor H-NS. Anti-silencing results from SsrB-induced local topological changes in the DNA, in part as a result of its bending ability (Figure 6B). This likely provides enough free DNA to enable access to the promoter for RNA polymerase to activate transcription.

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When the SsrA kinase is present and activated by acid stress, SsrB is phosphorylated and SsrB~P de-represses H-NS and activates transcription at SPI-2 and SPI-2 co-regulated genes, including: sifA (Walthers et al., 2011), ssaB, ssaM, sseA and ssaG (Walthers et al., 2007). In the absence of the SsrA kinase, SsrB is not phosphorylated, but it can counter H-NS silencing at csgD (Figure 4A–D and Figure 6A). SsrB binding and bending at the csgD promoter causes a sufficient change in the DNA secondary structure (Figure 5B,C) that likely enables access for RNA polymerase, stimulating csgD transcription. It is interesting to note that SsrB is located on the SPI-2 pathogenicity island, and thus was acquired as Salmonella enterica diverged from Salmonella bongori. However, the capability to form biofilms is an ancestral trait, as phylogeny studies have shown that most of the natural or clinical isolates of Salmonella belonging to all the five sub-groups form rdar colonies (White and Surette, 2006). The SsrB response regulator can control two distinct lifestyle choices: the ability to assemble a type three secretory system and survive in the macrophage vacuole or the ability to form biofilms on gallstones in the gall bladder to establish the carrier state.

What then controls the presence or activation of the kinase SsrA? Our early experiments indicated that SsrA and SsrB were uncoupled from one another (i.e., SsrB was present in the absence of SsrA) and ssrA transcription was completely dependent on OmpR (Feng et al., 2004). The EnvZ/OmpR system is stimulated by a decrease in cytoplasmic pH when Salmonella enters the macrophage vacuole (Chakraborty et al., 2015). This may also be the stimulus for activating SsrA, since the Salmonella cytoplasm acidifies to pH 5.6 during infection and the cytoplasmic domain of EnvZ (EnvZc) was sufficient for signal transduction (Wang et al., 2012; Chakraborty et al., 2015). Previous reports also identified a role for PhoP in ssrA translation (Bijlsma and Groisman, 2005), which would further add to fluctuating SsrA levels. The present work describes a novel role for the unphosphorylated response regulator SsrB in de-repressing H-NS (Figure 6B). We show that under biofilm-inducing conditions, unphosphorylated SsrB is sufficient to activate the expression of csgD. There are only a few such examples of unphosphorylated response regulators playing a role in transcription such as DegU (Dahl et al., 1992) in Bacillus subtilis and RcsB (Latasa et al., 2012) in S. Typhimurium.

In recent years, it has become apparent that H-NS silences pathogenicity island genes in Salmonella (Lucchini et al., 2006; Navarre et al., 2006; Walthers et al., 2007; 2011). Understanding how H-NS silences genes and how this silencing is relieved is an active area of research (Will et al., 2015; Winardhi et al., 2015). Because the anti-silencing style of gene regulation is indirect and does not rely on specific DNA interactions, searching for SsrB binding sites has not been informative in uncovering this type of regulation (Tomljenovic-Berube et al., 2010; Worley et al., 2000; Shea et al., 1996). Even a recent report in which the proteomes of wild type, hilA null (a transcriptional regulator of SPI-1 genes) and ssrB null were analyzed by SILAC and compared with an existing CHIP dataset failed to identify csgD as an SsrB-regulated locus (Brown et al., 2014), as sequence gazing alone does not help in identifying mechanisms of transcriptional regulation.

SsrB is well suited to this style of regulation, because it does not recognize a well-defined binding site (Feng et al., 2004; Walthers et al., 2007; Tomljenovic-Berube et al., 2010), it has a high non-specific binding component (Carroll et al., 2009) and it bends DNA upon binding (Carroll et al., 2009; Figure 6B, this work). Furthermore, previous microarray studies disrupted both ssrA and ssrB, which would not uncover a distinct role for SsrB in gene regulation under non SPI-2-inducing conditions in the absence of the SsrA kinase. It is worth mentioning here that in our AFM images, it was apparent that H-NS was still bound to some regions of the csgD promoter when SsrB condensed the DNA (Figure 6A(ii)). Thus, H-NS does not have to be completely stripped off the DNA for de-repression to occur, a finding that was also evident in our previous studies (Liu et al., 2010) and others (Will et al., 2014).

SsrB binds and bends DNA, resulting in highly curved DNA conformations. This DNA binding property of SsrB is distinct from H-NS, which forms rigid nucleoprotein filaments and thus straight DNA conformations (Figure 6A(i)). Bent DNA is therefore an energetically unfavorable substrate for H-NS binding, and a likely mechanism of SsrB-mediated anti-silencing of H-NS repressed genes. SsrB-dependent displacement of H-NS is more energetically favored to occur predominantly at the ends of H-NS-bound filaments, which requires disruption of fewer H-NS protein-protein interactions (Winardhi et al., 2015 and Figure 6B). In an equal mixture of H-NS and SsrB (Figure 6A(ii)), we do not see evidence of sharply bent filaments. This is expected because H-NS dissociation is likely restricted to the filament ends. Such events occur due to the cooperative nature of H-NS binding that results in a chain of linked H-NS proteins. Hence, H-NS displacement by SsrB likely occurs progressively from the filament end. This behavior has been observed in our single-molecule stretching experiments with H-NS filaments in the presence of SsrB. This ability of H-NS to re-orient on the DNA without being released would also promote its re-binding and silencing when SsrB or other anti-silencers are released (Figure 6B).

Response regulators are grouped into subfamilies on the basis of the structures of their DNA binding domains. SsrB is in the NarL/FixJ subfamily, which possess a helix-turn-helix (HTH) motif in the C-terminus (Baikalov et al., 1996). NarL was the first full-length structure of a response regulator and it showed that the N-terminal phosphorylation domain physically blocked the recognition helix in the HTH motif (Maris et al., 2002). Thus, phosphorylation is required to relieve the inhibition of the N-terminus. In the results presented herein, it is apparent that SsrB has adapted to relieving H-NS-silencing and that phosphorylation is not required for this behavior, nor is it required for DNA binding (Figure 5B).

In summary, we showed that the response regulator SsrB is required for biofilm formation because it can de-repress H-NS at the csgD promoter (Figure 6B). This leads to the production of CsgD, the master regulator of biofilms. It is noteworthy that a laterally acquired gene product, SsrB, has evolved the job of regulating the levels of csgD, a transcriptional regulator encoded by the core genome. For this activity, phosphorylation of SsrB was not required, which is rare amongst response regulators. Furthermore, we identify H-NS as a repressor of csgD in Salmonella, instead of an activator (Gerstel et al., 2003). This unifies the regulation of CsgD by H-NS in E. coli (Ogasawara et al., 2010) and Salmonella. This work places SsrB at a unique decision point in the choice between lifestyles by Salmonella and makes it crucial for the entire gamut of pathogenesis, i.e., biofilms and virulence.

The bacterial strains and plasmids used in this study are listed in Table 1. Salmonella enterica serovar Typhimurium strains were grown in LB medium with shaking at 37°C in the presence of 100 μg/ml ampicillin, 12.5 μg/ml Tetracycline (Tet) or 50 μg/ml Kanamycin (Km) when necessary. For observing the rdar morphotype, plates of LB medium (without salt) containing 1% Tryptone and 0.5% Yeast Extract supplemented with congo red (40 μg/ml) (Sigma-Aldrich, Singapore) were prepared and kept at 30°C after inoculation. To detect cellulose production, LB without salt medium plates were supplemented with Fluorescent Brightener 28 (200 μg/ml) (Sigma-Aldrich), stored under darkness and observed under UV light after incubating at 30°C. For the SsrBc complementation test using the plasmid pKF104, 0.2% arabinose was added to the medium.

All DNA manipulation procedures were carried out according to (Sambrook, 1989) using reagents procured from Qiagen, Invitrogen or Fermentas, Singapore. Transformation in S. Typhimurium strains was performed by standard electroporation protocols (Sambrook, 1989). Polymerase chain reaction (PCR) was carried out using oligonucleotides as listed in the Table 2 following standard protocols (Sambrook, 1989).

The S. Typhimurium strain harboring a D56A mutation in SsrB was generated by the homologous recombination technique as decribed in (Karlinsey, 2007). Plasmid pKF46 (Feng et al., 2004) was used to construct the necessary linear DNA fragment by a two-step overlapping PCR (Sambrook, 1989). Homologous recombination was also used to construct the hns and ssrA deletion mutants in the respective strain backgrounds. The sifA-lacZ marker was transduced into the lambda attP site of strains DW748 (ssrAH1), DW749 (ssrAD) and DW750 (ssrAH2) using standard P22 transduction protocols (Davis, 1980).

Biofilms were grown in three-channel glass-bottomed flow cells (individual cell dimensions 1 × 4 × 40 mm³) with an individual-channel flow rate of 8 ml/h of 0.5X M9 glucose minimal medium. Each channel was inoculated with M9 minimal medium cultures adjusted to an OD₆₀₀ of 0.04. Flow was started after allowing initial attachment for 1 hr. After specific time points, each channel of the flow cell was stained with 200 μl SYTO-9 green (Invitrogen, Singapore) as per the manufacturer’s protocol. Subsequently, for each flow cell channel image acquisition was performed using a LSM 780 Carl Zeiss confocal fluorescence microscope at 480/500 nm (20X magnification). Five image stacks were acquired starting from the center of the channel to a distance of 5–10 mm from the inlet, approximately 5 mm apart. This experiment was repeated twice in duplicates for the wild type and ssrB mutant strains. All images were processed using the Image J software.

Static biofilms were grown by inoculating the bacterial strains on APTES-coated coverslips in 24-well Nunclon polystyrene plates in 500 μl LB without salt medium and kept shaking at 30°C, 100 rpm. After two days, the growth medium was removed; wells were washed with PBS and incubated with 200 μl of the fixative solution (4% para-formaldehyde+0.2% glutaraldehyde in filter-sterilized PBS) for 1 hr at room temperature. After washing twice with sterile water, 35%, 50%, 75%, 90%, 95% and 100% ethanol were added sequentially and kept for 10 min at room temperature. Dehydrated samples were stored in absolute ethanol and kept at 4°C until subjected to critical point drying. Images were obtained using a Carl Zeiss Merlin field emission scanning electron microscope by detecting secondary electrons (SE2) under low electron beam acceleration voltage (1 kV) and low probe current (76 pA).

To estimate the amount of biofilms typically a single colony was inoculated in LB broth medium and incubated at 37°C/250 rpm. 2 μl of this culture was added to 198 μl LB without salt medium in a 96-well polystyrene plate and kept at 30°C with gentle shaking of 100 rpm. After two days, the growth medium was removed and each well was washed twice with 200 μl of Phosphate-buffered Saline (PBS). The attached bacterial communities or biofilms in each well were then stained with 200 μl crystal-violet solution (0.1%) for a minimum of 1 min. This was followed by washing twice with PBS and addition of 200 μl absolute ethanol. Appropriate dilutions were measured for absorbance at 580 nm using a Tecan Infinite M200 plate reader. Each experiment was performed at least thrice in triplicates. For the time course experiment, the above procedure was performed twice at 12, 24, 36, 47, 60, 72 and 84 hr respectively in triplicates.

To study the capability of the Salmonella strains to form biofilms on cholesterol-attached surfaces, a tube biofilm assay was performed as described in (Crawford et al., 2008), but without the use of bile salts in the growth medium. LB without salt medium was used for growth at all steps. All experiments were performed at least thrice in triplicates for 7 days on a nutator shaker at room temperature. Biofilms formed at the end of seven days were estimated using the above described crystal violet staining protocol with appropriate controls.

Each of the bacterial strains were grown overnight in LB, washed twice and resuspended in 50 μl of PBS buffer. These were then transferred into 1 ml of fresh MgM medium, pH 5.7, as in (Feng et al., 2003) and kept shaking at 37°C for around 6–7 h, until growth reached OD₆₀₀ between 0.6–0.8. At this stage, 50 μl of the culture was removed in a 96-well microtiter plate and 145 μl of lysis buffer was added (0.01% SDS, 50 mM Beta Mercaptoethanol in Z-buffer) as performed previously (Feng et al., 2003). The β-galactosidase activity was represented in Miller Units and calculated as 1000 x [(OD₄₂₀-1.75 x OD₅₅₀)]/ t (min) x volume (ml) x OD₆₀₀). Measurements were made in a Tecan Infinite M200 plate reader and repeated thrice in triplicates.

Total RNA was isolated from two-day old macrocolonies using the Qiagen RNeasy kit (Qiagen, Singapore) as per the manufacturer’s protocol with some modifications. Briefly, the macrocolonies were selected and re-suspended in 1 ml Qiagen RNAprotect reagent followed by immediate total RNA isolation or storage at -80°C. Appropriate volume of lysis buffer, 15 mg/ml lysozyme in TE buffer, was used. After dissolving the RNA in 20 μl RNase-free water, samples were assessed by gel electrophoresis and quantified using the Nanodrop system (THERMO Scientific, Singapore). All RNA samples were treated with TURBO DNase as per the manufacturer’s protocol (Life Technologies, Singapore). PCR analysis using rrsA specific primers was performed to ensure the absence of genomic DNA in the RNA preparations.

Reverse transcription reaction was carried out using the iScript reverse transcription supermix for RT-qPCR (BIO-RAD, Singapore) according to the recommended protocol. This was followed by amplifying 5–10 ng cDNA by real-time qPCR using SsoFast EvaGreen Supermix (BIO-RAD) and csgD-specific primers. Similar reactions were set up using rrsA-specific primers for their use as normalization controls. All experiments were performed in triplicate with two independent RNA preparations. Relative transcript abundance was determined using the △C_T method using a reference gene as per the manufacturer’s guide.

Two-day old macrocolonies were resuspended in 400 μl Laemmli buffer (Sambrook, 1989) and total protein was separated by 10% SDS-PAGE. This was followed by electro-transfer to a PVDF membrane as described before (Feng et al., 2004). The membrane was incubated with anti-CsgD (1:10,000) or anti-GroEL (1:5000) antibodies in PBS buffer containing 0.05% Tween-20 and 3% BSA. Anti-rabbit secondary antibody (Santa Cruz Biotechnology Inc., Dallas, TX) was used for detection as described previously (Feng et al., 2004).

The E. coli BL21 (DE3) strain was used as a host for the overproduction of proteins His-SsrBc, His-SsrB, D56A His-SsrB, K179A His-SsrB and His-H-NS. The respective plasmids harboring the constructs have been listed in Supplementary Table 2. Detailed procedures for the purification of His-SsrBc, His-SsrB, D56A His-SsrB and K179A His-SsrB have been described before (Feng et al., 2004; Walthers et al., 2007; Carroll et al., 2009). The DW160 plasmid harboring His-tagged H-NS from Salmonella Typhimurium was used to overexpress and purify His-H-NS following the procedure described in (Walthers et al., 2011).

Glutaraldehyde-modified mica surface was prepared as decribed previously in (Liu et al., 2010) and references therein. A 755 bp sequence upstream to the +1 start site (Gerstel et al., 2003) of csgD was amplified and gel purified. This fragment also harbored a SsrB-specific site (Feng et al., 2004; Walthers et al., 2007), TTATAAT sequence (Figure 6—figure supplement 4). A typical 50 μl reaction contained 10 ng of this DNA (755 bp of the csgD regulatory region) mixed with an appropriate amount of SsrB, SsrBc or D56A SsrB and incubated for 15 min at room temperature. This mixture was then deposited on glutaraldehyde-modified mica for 15 min. Images were acquired on a Bruker Dimension FastScan AFM system using the tapping mode with a silicon nitride cantilever (FastScan C, Bruker). Raw AFM images were processed using Gwyddion software (http://gwyddion.net/). The bending angle was analysed using a home-written Matlab code as described in the Supplementary methods.

GraphPad Prism 6 version 6.00, GraphPad Software, La Jolla California, www.graphpad.com was used to make all the graphs and do statistical analysis by Student’s t-test wherever required.

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

We thank the following: Assoc Prof Scott Rice, SCELSE, Singapore, for his guidance with the flow cell experiments, Prof T Venky Venkatesan, NUSNNI, Singapore, for the use of SEM facilities, Dr. Aaron White, Vaccine and Infectious Disease Organization, University of Saskatchewan, Canada, for the monoclonal anti-CsgD antibody, Prof Iñigo Lasa, Universidad Pública de Navarra-CSIC-Gobierno de Navarra, Spain, for the csgD construct and Dr. Swaine Chen, GIS, Singapore, for the anti-GroEL antibody. We are grateful to the MBI Central facilities, especially Chen HongYing from the Protein Expression core and Zhang Bo, Cindy, from the Science Communications core for the artwork. We appreciate Prof Stephen Lory, Harvard Medical School, Boston, and the three anonymous reviewers for their critical comments. Supported by 5IOBX-000372 from the VA to LJK, Singapore Ministry of Education Academic Research Fund Tier 2 [MOE 2013-T2-1-154) to YJ and LJK and funding from the Research Centre of Excellence in Mechanobiology, NUS from the Ministry of Education.

© 2016, Desai et al.

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