Deletion of flgD Enhances the Expression of Virulence Genes Through hapR.

To investigate how flagellar synthesis affects the expression of virulence genes, we constructed strains with an in-frame deletion of flgD, which encodes one of the flagellar rod proteins. As expected, flgD mutants are nonmotile ((Fig. S4A). Although TcpA and CT production was abolished in the luxO mutant, wild-type levels of TcpA and CT were detected in both flgD and flgD luxO mutants (Fig. S2A). To determine whether the deletion of flgD alters aphA expression in the luxO mutant, we measured the corresponding activity levels of an aphA-lacZ transcriptional reporter. As expected, deletion of luxO results in repression of aphA (Fig. S2B). However, mutation of flgD not only results in wild-type levels of aphA transcription but also restores aphA expression in the luxO mutant. This explains why the inhibition of the virulence regulon in the luxO mutant was restored in the flagellar mutants. To further characterize the role of flgD in quorum sensing, we transformed wild-type and flgD strains with pBB1, which contains the luxCDABE operon from V. harveyi that can be regulated directly by HapR (6). Although lux expression depends on cell density in both cases, overall expression is lower in the flgD strain than in the wild-type strain (Fig. S2C). We attribute this reduction in lux expression to lower levels of HapR because the expression of a hapR-lacZ transcriptional reporter was also inhibited in the flgD mutant (Fig. S2D). Taken together, these results indicate that flagellar synthesis modulates the quorum sensing output, including the expression of virulence factors, by regulating hapR transcription.

The Alternative σ-Factor FliA (σ²⁸) Represses hapR Expression.

The regulatory hierarchy that determines the order for expressing flagellar genes in V. cholerae ends with the σ²⁸-dependent activation of particular flagellar genes (e.g., flaBCD) (16). Because constitutive expression of fliA, which encodes σ²⁸, inhibits regulation by quorum sensing (Table S1), we hypothesized that the repression of hapR observed in flgB, flgC, and flgD mutants is due to increased activity of FliA (Fig. 1A). The activity of FliA increases upon secretion of the anti-σ²⁸ factor FlgM through the flagellar export apparatus (15, 16). To determine whether this process plays a role in the altered hapR regulation of the flgD mutant, we compared the level of FlgM in supernatants of wild-type and flgD cultures. The level of FlgM present in the supernatant of the flgD mutant is higher than that of wild-type V. cholerae (Fig. 1B), suggesting that disrupting flagellar rod assembly leads to higher FliA activity by increasing FlgM secretion. Consistent with this hypothesis, the expression of the FliA-activated gene flaD was enhanced in a flgD mutant (Fig. S3). To further investigate the relationship between FliA–FlgM interaction and hapR expression, we measured hapR transcription and HapR production in various mutants grown to midlog phase. Whereas hapR expression is inhibited in the flgM and flgD mutants, hapR is overexpressed in the fliA mutant (Fig. 1C). More importantly, deletion of fliA in the flgD mutant restores HapR production, indicating that the effect of hapR repression in flgD mutants is mediated through FliA. Furthermore, a fliA mutant complemented by a plasmid that constitutively expresses fliA restores the repression of hapR. Mutations in flaD and motY, which are known to be regulated by FliA (16), did not affect FlgM secretion or hapR expression (data not shown). Taken together, these results suggest that the high activity of FliA in either flgM or flgD mutants inhibits hapR expression. At this time, we do not know whether FliA regulates hapR directly or through another regulatory mechanism. The alternative σ-factor σ²⁸ could activate the expression of unknown repressor to repress the hapR expression or activate small RNAs to regulate hapR mRNA. In fact, hapR has been shown to be regulated by RNA-binding protein Hfq and a set of sRNAs activated by LuxO and σ⁵⁴ (5). However, if sRNAs are involved in σ²⁸-mediated hapR repression, they are different from those sRNAs induced by LuxO because σ²⁸ represses hapR in a luxO mutant (Fig. S2). Furthermore, because σ²⁸ also represses hapR in an hfq mutant, this regulation is independent of Hfq (data not shown).

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Fig. 1.

FliA represses quorum sensing in flgD mutants. (A) Proposed model for repression of the hapR promoter by secretion of FlgM. (B) FlgM secretion is increased in the flgD mutant. Strains containing a plasmid expressing a functional flgM-his₆ were grown to midlog. Culture pellets (P) and TCA-precipitated supernatants (S) were isolated and subjected to Western blot analysis using anti-His-6 antiserum. All samples were normalized to contain 10⁹ bacterial cells. (C) Reduced production of HapR is due to hapR repression by FliA. Strains harboring hapR-lacZ transcriptional fusions were grown to midlog in LB and harvested to measure β-galactosidase activity (Upper). Results, reported in Miller units, are means of three experiments ± standard deviations. Whole-cell extracts were subjected to Western blot analysis using anti-HapR antiserum (Lower). All samples were normalized to contain 10⁹ bacterial cells.

Repression of Quorum Sensing by FliA Is Important for Proper Intestinal Colonization.

Flagella are thought to help V. cholerae swim through mucosal layers to colonize the intestinal surface (13). Consistent with this hypothesis, flgD, fliA, and flgM mutants, which have reduced motility (15) (Fig. S4A), do not colonize the intestines of infant mice as well as motile wild-type cells (Fig. 2A). The observation that a fliA mutant displays a severe colonization defect was intriguing because FliA represses hapR expression (see Fig. 1). Colonization efficiency was significantly increased in a fliA hapR mutant (Fig. 2A) as compared with the fliA single mutant (P value <0.01). Consistent with previous reports (7), the hapR mutant colonized mice as well as wild-type. In addition, an in vivo colonization competition assay with a fliA single mutant and a fliA hapR double mutant demonstrated that the double-mutant colonizes ≈6-fold better than the fliA mutant (Fig. S4B). These results suggest that the colonization defect of the fliA mutant is partially due to insufficient repression of hapR.

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Fig. 2.

The expression of hapR is repressed by FliA during the colonization of infant mice. (A) In vivo competition assays. Infant CD-1 mice were orally inoculated with ≈10⁶ wild-type and mutant bacteria. For each mutant, the competitive index is defined as the number of colony-forming units (CFU) for the mutant compared with the corresponding CFU number for a wild-type strain recovered from the intestines 18 h after inoculation. (B) Expression of hapR during colonization. Infant CD-1 mice were orally inoculated with ≈10⁶ bacteria that contain a hapR-K_m^r reporter. At each time point, mouse intestines were homogenized, treated with or without kanamycin (500 μg/ml) for 10 min, and plated on LB plates. The number of colonized bacteria per mouse (above each bar) was calculated based on the number of CFU recovered from samples not treated with kanamycin. Expression of hapR is defined as the percentage of K_m-resistant CFU of the total CFU. Results are means from experiments with three to five mice, and the bars represent the corresponding standard deviations.

To provide further in vivo evidence that hapR expression is higher in fliA mutants, we colonized infant mice using strains that contain a hapR-K_m^r transcriptional fusion, which confers kanamycin resistance to cells expressing hapR (24). At 4 h, hapR expression in wild-type cells was low, presumably because of the small number of cells that had colonized the small intestine (Fig. 2B). However, by 18 h, the number of cells in the small intestine had increased, and hapR expression was high. The hapR expression patterns of flgD, fliA, and flgM mutants in vivo were similar to those in vitro (compare Figs. 1C and ​and22B), with significantly higher hapR expression in fliA mutants early in colonization (Fig. 2B). Taken together, these results suggest that the repression of hapR by FliA is required for proper colonization of host intestines.

Transposon Screen for hapR Repression.

LZV8 (lacZ::tcpA-sh ble, ΔluxO) was mutagenized with the TnAraOut mariner transposon (19). The resulting libraries were inoculated into AKI medium containing 0.05% arabinose and incubated without shaking at 37°C for 4 h until the OD₆₀₀ was ≈0.1. Cultures were treated with zeocin (25 μg/ml) for 30 min at 37°C and plated on LB agar without zeocin. Arbitrary PCR (19) and DNA sequencing were performed to identify transposon insertion sites.

hapR Transcription Measurements.

The hapR-lacZ transcriptional reporter was integrated into the chromosomes of various flagellar mutants. β-Galactosidase assays were performed as described (7).

In the mouse colonization and mucin-penetration assays, hapR expression was measured by using strains harboring the hapR-K_m^r reporter, which is at the hapR locus and maintains an intact copy of hapR (24). Bacteria isolated from mouse intestines or mucin columns were added to fresh LB in the presence or absence of 500 μg/ml kanamycin and incubated for 10 min at 37°C. This treatment is sufficient to kill 100% of V. cholerae cells that do not carry any K_m^r gene or that contain the hapR-K_m^r construct and a constitutively active LuxO (40). After treatment, the samples were plated onto LB. Expression of hapR was defined as the number of kanamycin-resistant CFU normalized by the number of total CFU.

For the mucin-penetration assays, hapR transcription was measured by quantitative real-time PCR. Total RNA was isolated from column samples by using TRIzol reagent (Invitrogen) and cleaned with the RNeasy kit (Qiagen). RNA reverse transcription was performed by using the SuperScript II kit (Invitrogen) with 200 ng of RNA sample. Quantitative real-time PCR using primers specific for hapR was performed on an Opticon 2 system (MJ Research). The 16S ribosomal RNA was used for an internal control in all reactions.

Infant Mouse Colonization Assay.

The infant mouse colonization assay was performed as described (41). Briefly, 10⁶ V. cholerae cells were inoculated into 6-day-old CD-1 suckling mice. At different time points, intestines were homogenized, serially diluted, and plated onto LB agar. Bacterial numbers were determined from the number of CFU.

Mucin-Penetration Assay and Flagella Visualization.

Mucin columns were prepared by adding different concentrations of bovine submaxillary mucin (Sigma) to 1-ml syringes. Midlog bacterial cultures (100 μl)or 100 μl of cells premixed with 1% mucin were loaded on the top of the mucin columns and allowed to settle for 30 min at 37°C. Fractions (500 μl) were collected from the bottom of the mucin columns. Bacteria numbers were measured by serially diluting samples, plating onto LB agar, and counting CFU. A drop of the collected fractions was placed on slides to detect flagella by using the flagella stain kit (Remel, Lenexa) or by transmission electron microscopy (University of Pennsylvania Bioimaging Core).

FlgM Detection.

To detect FlgM protein, a plasmid containing the P_BAD-flgM-his6 was introduced into various strains of V. cholerae. All samples were normalized to contain 10⁹ (or 10⁸ in the case of samples treated with mucin) bacterial cells. Supernatants were concentrated by using TCA precipitation (10% trichloroacetic acid). Both extracellular and cytoplasmic proteins were separated by SDS/PAGE, transferred to a nitrocellulose membrane, and immunoblotted with affinity purified anti-6× His rabbit antiserum (Rockland).

This work was supported by National Institutes of Health (NIH) Grant R01AI072479 (to J.Z.), National Science Foundation Grant MCB0615957 (to M.G.), an American Heart Association predoctoral fellowship (to T.M.) and a NIH T32 Bacteriology training grant (to A.T.).