RETRACTED: Interference of flagellar rotation up-regulates the expression of small RNA contributing to Bordetella pertussis infection

Abstract

Bacterial small RNAs (sRNAs) posttranscriptionally regulate gene expressions involved in various biological processes, including pathogenicity. Our previous study identified sRNAs, the expression of which was up-regulated in Bordetella pertussis, the causative agent of whooping cough, upon tracheal colonization of the bacteria; however, their roles in bacterial infection remain unknown. Here, we found that one sRNA, Bpr4, contributes to B. pertussis infection by posttranscriptionally up-regulating filamentous hemagglutinin (FHA), a major adhesin of the bacteria. Bpr4 bound to the 5′ untranslated region of fhaB mRNA encoding FHA and inhibited its degradation mediated by RNaseE. Our results demonstrated that Bpr4 up-regulation was triggered by the interference of flagellar rotation, which caused the disengagement of MotA, a flagellar stator. Subsequently, MotA activated a diguanylate cyclase to generate cyclic di-GMP, which plays a role in Bpr4 up-regulation through the RisK/RisA two-component system. Our findings indicate that a flagellum-triggered sensory system contributes to B. pertussis infection.

Bordetella pertussis senses the host cells via flagella and upregulates small RNA contributing to the bacterial infection.

INTRODUCTION

Pathogenic bacteria sense environmental cues in the host and express virulence factors in a regulated manner to establish infection. These responses are initiated by bacterial sensing for iron, bacterial cell density, carbon dioxide, and host molecules, such as hormones and serum components, and result in alterations of gene expressions that are regulated by phosphorelay two-component systems, σ factors, small RNAs (sRNAs), and other factors (1–4). Because appropriate gene expression is imperative for bacterial colonization/infection in the host, understanding bacterial sensing mechanisms teach the development of effective protective measures.

Bordetella pertussis, which causes the highly contagious respiratory disease pertussis (whooping cough), produces multiple virulence factors, including adhesins, such as filamentous hemagglutinin (FHA) and fimbriae, and toxins, such as pertussis toxin (PTx) and adenylate cyclase toxin, at appropriate periods in the course of infection (1). The expression of these virulence factors was previously considered to be largely regulated by the BvgAS two-component system (1), which is activated when the bacteria invade the host; however, recent studies indicated that the profiles of BvgAS-regulated genes differ substantially between in vitro and in vivo conditions (5, 6), suggesting a more complex network between BvgAS and other systems including the PlrSR two-component system and BspR/BtrA, an anti-σ factor (7, 8). In addition, accumulating studies have implied that sRNAs are likely involved in gene regulation in B. pertussis as well as other pathogenic bacteria during the course of infection (9–13). sRNAs, functional noncoding RNAs, are known to form base pairs with one or more target mRNAs to affect the latter’s stability and translational efficiency and ultimately fine-tune the expression of the target proteins (4). Studies using in silico and/or transcriptome analyses and Northern blotting revealed sRNA candidates that may participate in the gene regulation of B. pertussis (9–11). It was also reported that Hfq, a chaperone protein that stabilizes the interaction between sRNA and mRNA, influences B. pertussis virulence, indicating the involvement of sRNAs in bacterial pathogenicity (14).

We previously identified nine distinct sRNAs of B. pertussis that were highly transcribed upon tracheal colonization in mice, and the expression of eight of them was independent of the BvgAS system (15), implying that these sRNAs are up-regulated in response to environmental cues within the host in a BvgAS-independent manner and participate in downstream genes that support bacterial infection. Here, we explored the functions of sRNAs in B. pertussis infection and found the contribution of one, Bpr4, which is up-regulated up to 120-fold upon tracheal colonization (15), to bacterial infection by posttranscriptionally up-regulating FHA, a major adhesin of the bacteria (16). Bpr4 was up-regulated upon the interference of flagellar rotation after the interaction of flagellin and gangliosides on the host cells. We also demonstrate the mechanism by which the flagellum-mediated mechanosensing leads to sRNA up-regulation.

RESULTS

Bpr4 up-regulates FHA expression

Among the nine sRNAs identified in our previous study (15), four B. pertussis sRNAs (Bpr4, Bpr5, Bpr8, and Bpr9) were more highly expressed in bacteria colonizing mouse tracheas than in the bacteria grown in Stainer-Scholte (SS) broth (17). In the present study, we investigated the contributions of sRNAs to B. pertussis infection by examining Bpr4-, Bpr8-, and Bpr9-deficient mutants (the construction of Bpr5-deficient mutants was unsuccessful). The deletion of Bpr8 and Bpr9 did not influence the colonization ability of the bacteria. In contrast, the Bpr4-deficient mutant (Δbpr4) was found to less efficiently colonize mouse tracheas than the wild-type (WT) strain, while Δbpr4 complemented with a bpr4-carrying plasmid, pbpr4, showed equivalent tracheal colonization compared to WT (Fig. 1A). Thus, we further characterized Bpr4. SDS–polyacrylamide gel electrophoresis (SDS-PAGE) of bacterial culture supernatants revealed that the intensity of a protein band corresponding to approximately 200 kDa was reduced in Δbpr4 compared to WT (Fig. 1B). This reduction was reversed in Δbpr4 harboring pbpr4. The WT strain harboring a Ptac-bpr4 plasmid to overexpress Bpr4 produced a greater amount of the corresponding protein than WT (Fig. 1B and fig. S1A). Subsequent liquid chromatography–electrospray ionization–tandem mass spectrometry (LC-ESI-MS/MS) analysis identified the protein of this band as FHA. Immunoblot analyses confirmed that the expression levels of FHA were accordingly changed in the bacterial lysates and culture supernatants of the tested strains (Fig. 1C); however, the transcript levels of fhaB were largely unaffected by the presence of Bpr4 (fig. S1B). In addition, Δbpr4 ectopically expressing FHA by the plasmid Ptac-fhaB colonized mouse tracheas to a similar extent as WT harboring the empty vector (Fig. 1D and fig. S1C). We also confirmed that an FHA-deficient mutant (ΔfhaB) colonized mouse tracheas markedly less (0.003-fold) than WT (fig. S1D). These results indicate that Bpr4 contributes to bacterial colonization by posttranscriptionally up-regulating the expression of FHA.

Fig. 1. Involvement of Bpr4 in FHA expression of B. pertussis for colonization of mouse tracheas.

(A, D, and H) Colonization of B. pertussis 18323 WT and mutant strains in mouse tracheas. B. pertussis strains were intranasally inoculated into mice, and the bacteria recovered from mouse tracheas were counted on day 4. Each horizontal bar represents the geometric mean and geometric SD (n = 8 biological replicates). Data were obtained from two independent experiments and statistically analyzed by the Kruskal-Wallis test with Dunn’s multiple-comparison test. *P < 0.05, **P < 0.01, and ***P < 0.001. (B) SDS-PAGE analysis of culture supernatants of B. pertussis 18323 WT and mutant strains grown in SS broth for 24 hours. Arrow indicates the band that was identified as FHA by LC-ESI-MS/MS analysis. The image is a representative of two independent experiments. (C, G, and I) Immunoblotting for FHA, FtsZ, and PTx in bacterial lysates (C, G, and I) and culture supernatants (Sup.) (C) of B. pertussis 18323 WT and mutant strains. FtsZ and PTx were used as internal controls for each sample of the bacterial lysates and culture supernatants, respectively. The blot images are representatives of two independent experiments. The band intensity of FHA was measured using Fiji software (60) and is presented as a ratio relative to those of FtsZ and PTx, respectively. (E) Binding site between fhaB-5′-UTR and Bpr4 predicted by IntaRNA. Bpr4_m contains guanine-to-cytosine substitutions at nucleotide positions 5 and 6 in Bpr4. (F) Binding of Bpr4 to fhaB-5′-UTR. Bpr4 and Bpr4_m were incubated with fhaB-5′-UTR in the presence of rHfq or BSA (negative control), and RNA-RNA binding was determined by the electrophoretic mobility shift assay. The images are representatives of two independent experiments.

Since sRNAs are known to form base pairs with target mRNAs and regulate their translation, we examined the interaction of Bpr4 with fhaB mRNA. An RNA-RNA interaction prediction tool, IntaRNA (18), predicted that Bpr4 at positions 1 to 19 nucleotides (nt) forms base pairs with the 5′ untranslated region (5′UTR) of fhaB mRNA at positions 32 to 48 nt downstream of the transcription start site (Fig. 1E) (19). The direct binding of Bpr4 to fhaB-5′-UTR (positions 1 to 79 nt downstream from the transcription start site) was detected in the presence of recombinant Hfq (rHfq), while the binding was abolished in a mutated Bpr4 (Bpr4_m) (Fig. 1F), which was designed to break the base pairs with fhaB-5′-UTR (Fig. 1E). Consistently, FHA expression levels were not altered by the overexpression of Bpr4_m driven by Ptac-bpr4_m (Fig. 1G and fig. S1E), indicating that Bpr4 up-regulates FHA expression by binding to fhaB-5′-UTR. The Δbpr4 complemented with a bpr4_m-carrying plasmid, pbpr4_m, was recovered less from mouse tracheas than WT (Fig. 1H).

Next, we explored the mechanism by which Bpr4 up-regulates FHA expression. In general, sRNAs that bind to the 5′UTRs of the target mRNAs up-regulate their translation in two distinct manners (20): (i) sRNAs prevent the formation of a translation-inhibitory hairpin in the 5′UTR of target mRNAs and allow ribosomes to access the Shine-Dalgarno sequence, and (ii) the base pairs of sRNAs with 5′UTRs prevent ribonuclease E (RNaseE)–mediated degradation of the target mRNAs. Since secondary structure prediction by IPknot (http://rtips.dna.bio.keio.ac.jp/ipknot/) did not detect the translation-inhibitory hairpin in the fhaB-5′-UTR, we examined the latter possibility by using a B. pertussis mutant (ΔPrne) that barely expresses RNaseE because of a defect in the promoter region of the gene (fig. S1F). ΔPrne produced a larger amount of FHA than WT. This increase in FHA production was not reduced by the deletion of Bpr4 (Fig. 1I), indicating that RNaseE degrades fhaB mRNA in B. pertussis. In addition, Bpr4 overexpression by Ptac-bpr4 did not augment the FHA expression of ΔPrne (Fig. 1I and fig. S1G). These data indicate that Bpr4 prevents the RNaseE-mediated degradation of fhaB by binding to the 5′UTR of fhaB to ultimately up-regulate FHA expression.

Bpr4 is up-regulated upon cell adherence of B. pertussis

We previously reported that Bpr4 was highly expressed in B. pertussis colonizing mouse tracheas 4 days after infection (15). In the present study, we further found that the up-regulation of Bpr4 occurred as early as 1 hour after infection and persisted at similar levels at least for 10 days (Fig. 2A), implying that B. pertussis responds to certain conditions in the host environment and up-regulates Bpr4. In addition, the bacteria adhering to Calu-3 or THP-1 cells expressed Bpr4 more strongly than bacteria grown in SS broth (Fig. 2, B and C). Similar results were obtained for different bacterial strains and Calu-3 cells fixed with paraformaldehyde (PFA) or methanol (Fig. 2, D and E). In contrast, bacteria adhering to collagen-, gelatin-, or poly-l-lysine–coated culture plates did not up-regulate Bpr4 expression (Fig. 2F). These results suggest that B. pertussis recognizes cell surface factor(s) regardless of cell viability and up-regulates Bpr4.

Fig. 2. Bpr4 up-regulation upon cell adherence of B. pertussis.

(A) Bpr4 up-regulation in B. pertussis colonizing mouse tracheas. B. pertussis 18323 was intranasally inoculated into mice, and total RNA was extracted from mouse tracheas at the indicated times. Data represent fold changes in Bpr4 expression (mean + SEM) compared to the bacteria grown in SS broth (n = 8 biological replicates). (B) Schematic representation of free and cell-adhering bacteria. (C to F) Bpr4 up-regulation in B. pertussis adhering to cells. Calu-3 (C and D) and differentiated THP-1 cells (C) were incubated with B. pertussis strains 18323 (C and D), Tohama (a laboratory strain), BP139, BP140, BP141, and BP144 (clinical strains) (D) for 1 hour. B. pertussis 18323 was incubated with live and PFA- or methanol-fixed Calu-3 cells (E) or incubated on the wells of tissue culture–treated (normal), collagen type I–coated, gelatin-coated, and poly-l-lysine–coated plates (F) for 1 hour. Total RNA was extracted from free and cell- and plate-adhering bacteria that were separated as described in Materials and Methods. Data represent fold changes in Bpr4 expression (mean + SEM) compared to each strain grown in SS broth (n = 6 biological replicates). The number of bacteria that adhered to the cells and plates is represented as the percentage of CFU of the input bacteria (E and F) (bottom). Each bar represents the mean + SEM (n = 6 biological replicates). Data were obtained from two independent experiments and statistically analyzed by unpaired t test. *P < 0.05, **P < 0.01, ***P < 0.001, and ****P < 0.0001.

The RisK/RisA two-component system regulates Bpr4 expression

Because Bpr4 expression is independent of the BvgAS two-component system (15), the master regulator for genes contributing to B. pertussis infection (1), we examined the involvement of other two-component systems in Bpr4 expression. Among the 13 mutants deficient in sensor kinases of the two-component system that we tested, one RisK-deficient mutant (ΔrisK) did not increase Bpr4 expression upon bacterial adherence to Calu-3 cells (Fig. 3A and fig. S2A). The basal levels of Bpr4 expression were similar in WT and ΔrisK grown in SS broth (fig. S2B). A mutant deficient in RisA (ΔrisA), the partner response regulator for RisK (21), showed no increase in Bpr4 expression upon cell adherence and tracheal colonization (Fig. 3A), indicating that B. pertussis up-regulates Bpr4 expression through the RisK/RisA two-component system upon tracheal colonization. A previous report speculated that RisA regulates downstream genes in the presence of cyclic di-guanosine monophosphate (c-di-GMP) synthesized by diguanylate cyclase (Dgc) (21, 22). Therefore, we next examined the involvement of Dgc in Bpr4 expression. In the B. pertussis 18323 genome sequence [National Center for Biotechnology Information (NCBI) accession no. NC_018518.1], two distinct Dgcs are annotated; we designated them DgcA (locus tag: BN118_RS06950) and DgcB (locus tag: BN118_RS14135). The ectopic expression of DgcB, but not of DgcA, increased the levels of c-di-GMP and Bpr4 expression in bacteria grown in SS broth (Fig. 3B and fig. S2C). Furthermore, the deletion of dgcB, but not of dgcA, abolished the up-regulation of Bpr4 in response to bacterial adherence to Calu-3 cells and the colonization of mouse tracheas (Fig. 3C), whereas WT, ΔrisA, ΔdgcA, and ΔdgcB grown in SS broth equally expressed Bpr4 (fig. S2D). ΔrisA and ΔdgcB, but not ΔdgcA, exhibited less colonization than WT, which was restored by Bpr4 overexpression (Fig. 3, D and E). These results indicate that Bpr4 functions downstream of RisA and DgcB and exacerbates bacterial infection.

Fig. 3. c-di-GMP–mediated up-regulation of Bpr4.

(A and C) Bpr4 up-regulation in B. pertussis adhering to cells or colonizing mouse tracheas. Calu-3 cells were incubated with B. pertussis 18323 WT and mutant strains for 1 hour (left). Mice were intranasally inoculated with B. pertussis 18323 WT and mutant strains for 4 days (right). Total RNA was extracted from free and cell-adhering bacteria and bacteria recovered from mouse tracheas. Data represent fold changes in Bpr4 expression (mean + SEM) compared to each strain grown in SS broth (left, n = 6; right, n = 8 biological replicates). (B) Involvement of c-di-GMP in Bpr4 up-regulation. B. pertussis WT and mutant strains were grown in SS broth for 12 to 14 hours. c-di-GMP concentrations in the bacterial lysates were measured by ELISA (n = 6 biological replicates). Data are normalized to protein concentrations. Dashed line indicates the lower limit of detection (0.28 pmol/mg of protein) (top). Total RNA was extracted from bacteria, and the relative amount of Bpr4 was measured. Data represent fold changes in Bpr4 expression (mean + SEM) compared to WT (bottom) (n = 6 biological replicates). (D and E) Colonization of B. pertussis 18323 WT and mutant strains in mouse tracheas. The number of bacteria recovered from the tracheas of mice 4 days after intranasal inoculation was counted. Each horizontal bar represents the geometric mean and geometric SD (n = 8 biological replicates). Data were obtained from two independent experiments and statistically analyzed by unpaired t test (A and C) (left), one-way analysis of variance (ANOVA) with Tukey’s multiple-comparison test (B and C) (right), or Kruskal-Wallis test with Dunn’s multiple-comparison test (D and E). *P < 0.05, **P < 0.01, ***P < 0.001, and ****P < 0.0001.

Bpr4 is up-regulated through a flagellum-mediated pathway

We then explored the mechanism underlying Bpr4 up-regulation in B. pertussis colonizing mouse tracheas by using reporter strains carrying a bpr4 promoter-driven gfp gene (Pbpr4-gfp). The up-regulation of the reporter gfp upon tracheal colonization was observed in WT, but not in ΔrisA (Fig. 4A), indicating that Bpr4 is up-regulated by the promoter activity level. We next generated transposon-based random mutants from B. pertussis 18323 harboring Pbpr4-gfp and screened them for genes that are involved in the activation of Pbpr4 in bacteria adhering to Calu-3 cells. We confirmed that the Pbpr4 activity of ΔrisA in the cell-adhering state was markedly lower than that of WT, but the activity was comparable between WT and ΔrisA grown in SS broth (Fig. 4, B and C). Screening of the 800 transposon-integrated mutants revealed 2 distinct mutants, designated 2-19 and 5-33, which scarcely expressed green fluorescent protein (GFP) (0.09- and 0.32-fold) upon cell adherence, similar to ΔrisA (Fig. 4B and fig. S2E). DNA sequencing indicated that the mutants 2-19 and 5-33 carry the transposon in the bvgS and flaA genes encoding BvgS and FliC, a flagellin, respectively (Fig. 4D). The low activity of Pbpr4 in mutant 2-19 was likely due to a loss of adhesion to Calu-3 cells, which results from the defect in bvgS, the sensor kinase of the BvgAS system (fig. S2F). Since mutant 5-33 and a FliC-deficient mutant (ΔflaA) retained the ability to adhere to Calu-3 cells, we focused on FliC as a bacterial factor required for Bpr4 up-regulation. No increase in Bpr4 expression was observed in ΔflaA adhering to cells or colonizing tracheas (Fig. 4E). FliC composes the filament of the flagellar complex, which includes stators, such as MotAB, MotCD, MotPS, and PomAB, embedded in the inner membrane (23, 24). These stator molecules, which play a critical role in flagellar rotation, have also been reported to stimulate c-di-GMP production by interacting with Dgc, a membrane protein (24–27). Thus, we investigated the involvement of MotAB in Bpr4 up-regulation, because the genome of B. pertussis 18323 harbors genes encoding MotAB but not other flagellar stators. The deletion of motA, but not of motB, abolished the up-regulation of Bpr4 in B. pertussis adhering to Calu-3 cells and colonizing mouse tracheas (Fig. 5A), but Bpr4 was equally expressed in WT, ΔflaA, ΔmotA, and ΔmotB grown in SS broth (fig. S2D). ΔflaA and ΔmotA, but not ΔmotB, exhibited less colonization than WT, which was restored by Bpr4 overexpression (Fig. 5, B and C). Because DgcB was involved in Bpr4 up-regulation (Fig. 3, B and C), we examined the interaction of MotA with DgcB by using a bacterial two-hybrid system, in which interactions between different proteins respectively fused with the T25 and T18 subunits of adenylate cyclase toxin are detected by an increase in cyclic adenosine monophosphate (cAMP) levels in Escherichia coli (24, 28). The coexpression of T25-fused MotA and T18-fused DgcB in E. coli increased intrabacterial cAMP and c-di-GMP (Fig. 5D), indicating that DgcB interacted with and was activated by MotA. Bpr4 was likely up-regulated through a pathway from FliC (flagellin) and MotA to DgcB-generating c-di-GMP that takes part in the signaling cascade of the RisK/RisA system.

Fig. 4. Flagellin-mediated up-regulation of Bpr4.

(A) Increase in Pbpr4 activity of B. pertussis colonizing mouse tracheas. B. pertussis 18323 WT and ΔrisA strains harboring Pbpr4-gfp were intranasally inoculated into mice, and total RNA was extracted from mouse tracheas on day 4. Data represent fold changes in gfp mRNA expression (mean + SEM) compared to each strain grown in SS broth (n = 8 biological replicates). (B) GFP reporter assay for Pbpr4 activity. Calu-3 cells were incubated with B. pertussis 18323 WT, ΔrisA, and the transposon-integrated mutants 2-19 and 5-33 (5 × 10⁶ CFU/5 × 10⁴ cells per well), which harbor Pbpr4-gfp, and GFP expression levels were measured after a 1-hour incubation. Data represent device-specific fluorescence units (mean + SEM) (n = 6 biological replicates). (C) GFP expression of Pbpr4-gfp–harboring B. pertussis 18323 WT and ΔrisA strains grown in SS broth (1 × 10⁸ CFU per well). Each bar represents the mean + SEM (n = 6 biological replicates). (D) Transposon insertion sites in mutants 2-19 and 5-33. The transposons in 2-19 and 5-33 were located at nucleotide positions 1079 of bvgS and 480 of flaA, respectively. (E) Bpr4 up-regulation of B. pertussis adhering to Calu-3 cells and colonizing mouse tracheas. B. pertussis 18323 WT and ΔflaA strains were incubated with Calu-3 cells for 1 hour (left) or intranasally inoculated into mice for 4 days (right). Total RNA was extracted from free and cell-adhering bacteria and from bacteria recovered from mouse tracheas. Data represent fold changes in Bpr4 expression (mean + SEM) compared to each strain grown in SS broth (left, n = 6; right, n = 8 biological replicates). Data were obtained from two independent experiments and statistically analyzed by unpaired t test (A, C, and E) or one-way ANOVA with Tukey’s multiple-comparison test (B). ****P < 0.0001.

Fig. 5. Involvement of MotA in flagellum-mediated Bpr4 up-regulation.

(A) Bpr4 up-regulation of B. pertussis adhering to Calu-3 cells and colonizing mouse tracheas. B. pertussis 18323 WT and mutant strains were incubated with Calu-3 cells for 1 hour (left) or intranasally inoculated into mice for 4 days (right). Total RNA was extracted from free and cell-adhering bacteria and from bacteria recovered from mouse tracheas. Data represent fold changes in Bpr4 expression (mean + SEM) compared to each strain grown in SS broth (left, n = 6; right, n = 8 biological replicates). (B and C) Colonization of B. pertussis 18323 WT and mutant strains in mouse tracheas. The number of bacteria recovered from the tracheas of mice intranasally inoculated with B. pertussis strains was counted on day 4. Each horizontal bar represents the geometric mean and geometric SD (n = 8 biological replicates). (D) Interaction between MotA and DgcB analyzed by the bacterial two-hybrid system. cAMP and c-di-GMP concentrations in the lysates of E. coli strains expressing T25 and T18 fusion proteins were measured by ELISA and normalized to the protein concentrations of each sample. Dashed line indicates the lower limit of detection (0.28 pmol/mg of protein) (bottom). Each bar represents the mean + SEM (n = 6 biological replicates). Data were obtained from two independent experiments and statistically analyzed by unpaired t test (A, left), one-way ANOVA with Tukey’s multiple-comparison test (A, right, and D), or Kruskal-Wallis test with Dunn’s multiple-comparison test (B and C). *P < 0.05, **P < 0.01, ***P < 0.001, and ****P < 0.0001.

A recent study reported that B. pertussis, which was once considered nonflagellated and nonmotile, produces flagella when grown in the presence of 40 mM MgSO₄ or 10% fetal bovine serum (FBS) (29). We confirmed that B. pertussis 18323 produces FliC and exhibits motility in the presence of bovine serum albumin (BSA) (Fig. 6, A and B). The expression of FliC was independent of MotA. The increased expression of flaA and motA mRNA was also observed in B. pertussis incubated in the presence of BSA and colonizing mouse tracheas (Fig. 6, C and D). In addition, Bpr4 was not up-regulated in B. pertussis adhering to Calu-3 cells in the absence of BSA, and the ability of B. pertussis to adhere to the cells increased in the presence of BSA (Fig. 6, E and F), indicating that flagella are required for Bpr4 up-regulation.

Fig. 6. Expression of flagella in B. pertussis.

(A) Immunoblotting for FliC (flagellin) in B. pertussis. B. pertussis 18323 WT and mutant strains were incubated in SS broth and Hanks’-Hepes buffer containing BSA or FBS at 37°C with (left) or without (all) 5% CO₂ for 1 hour. FliC and FtsZ (internal control) in the bacterial lysates were detected by immunoblotting. The blot images are representatives of two independent experiments. (B) Motility of B. pertussis 18323. A representative picture (left) and diameters of the motility zone (right) are shown. Each bar represents the mean + SEM (n = 6 biological replicates). (C and D) flaA and motA mRNA expression of B. pertussis. Total RNA was extracted from B. pertussis 18323 incubated in Hanks’-Hepes buffer containing BSA for 1 hour (C) and the bacteria recovered from mouse tracheas 4 days after infection (D). Data represent fold changes in mRNA expression (mean + SEM) compared to bacteria incubated in SS broth (n = 6 biological replicates). (E and F) Bpr4 up-regulation of B. pertussis cultivated in the presence of BSA. B. pertussis 18323 was suspended in Hanks’-Hepes buffer with or without BSA (1 mg/ml) and incubated with Calu-3 cells for 1 hour. Total RNA was extracted from free and cell-adhering bacteria (E). Data represent fold changes in Bpr4 expression (mean + SEM) compared to bacteria grown in SS broth (n = 6 biological replicates). The number of bacteria that adhered to the cells is represented as the percentage of CFU of the input bacteria (F). Each bar represents the mean + SEM (n = 6 biological replicates). Data were obtained from two independent experiments and statistically analyzed by one-way ANOVA with Tukey’s multiple-comparison test (B and C) or unpaired t test (E and F). *P < 0.05 and ****P < 0.0001.

Gangliosides are host factors required for Bpr4 up-regulation

Our results demonstrate that bacterial adherence to host cells triggers Bpr4 up-regulation, which is mediated through flagellin, MotA, DgcB, and RisA. Among these bacterial factors, only flagellin is outside the outer membrane, implying its role in recognizing bacterial adherence and initiating signal transduction to up-regulate Bpr4. We therefore explored host factors that mediate Bpr4 up-regulation through flagellin. Toll-like receptor 5 (TLR5), the receptor for flagellin (30), was excluded from candidates for the host factor, because bacterial adherence to TLR5-knockout Calu-3 cells induced Bpr4 up-regulation similar to WT cells (fig. S3, A and B). We then examined gangliosides, which reportedly bind to bacterial flagellin (31, 32). The treatment of Calu-3 cells with dl-threo-1-phenyl-2-palmitoylamino-3-morpholino-1-propanol (PPMP), an inhibitor of ganglioside synthesis, markedly reduced Bpr4 up-regulation in cell-adhering B. pertussis (Fig. 7A). Similar results were obtained in cells treated with neuraminidase (NA), which removes sialic acids from gangliosides (Fig. 7B) (33). Furthermore, the level of Bpr4 up-regulation was reduced in bacteria adhering to cells, of which the ganglioside-rich lipid raft on the cell membrane was disrupted by methyl-β-cyclodextrin (MβCD) (Fig. 7C) (33). The binding of B. pertussis WT, but not ΔflaA, and recombinant FliC (rFliC) to gangliosides was confirmed by enzyme-linked immunosorbent assay (ELISA; Fig. 7, D and E). These results indicate that sialylated gangliosides on the lipid raft are recognized by flagellin to trigger Bpr4 up-regulation.

Fig. 7. Involvement of gangliosides in Bpr4 up-regulation upon adherence of B. pertussis to host cells.

(A to C) Bpr4 up-regulation of B. pertussis adhering to host cells. Calu-3 cells were pretreated with PPMP (A), NA (B), or MβCD (C) and then incubated with B. pertussis 18323 for 1 hour. Total RNA was extracted from the free and cell-adhering bacteria. Data represent fold changes in Bpr4 expression (mean + SEM) compared to the bacteria grown in SS broth (n = 6 biological replicates). (D and E) Binding of B. pertussis 18323 WT and ΔflaA strains (D) and rFliC (E) to gangliosides analyzed by ELISA. Heat treatment of rFliC was carried out at 98°C for 5 min. Each bar represents the mean + SEM (n = 6 biological replicates). Data were obtained from two independent experiments and statistically analyzed by one-way ANOVA with Tukey’s multiple-comparison test. *P < 0.05, ***P < 0.001, and ****P < 0.0001.

Extracellularly added gangliosides also stimulated Bpr4 up-regulation only when the bacteria were immobilized on the culture plate (Fig. 8A). Neither lewis X and heparan sulfate, which reportedly bind to flagellin (34, 35), nor other lipids composing the cell membrane stimulated Bpr4 up-regulation (fig. S3C). These results indicate that both bacterial immobilization and stimulation with gangliosides are required for Bpr4 up-regulation.

Fig. 8. Bpr4 up-regulation by interference of flagellar rotation in B. pertussis adhering to abiotic surfaces.

(A to C) Bpr4 up-regulation in immobilized B. pertussis. B. pertussis 18323 was adhered to wells of culture plates by centrifugation, which were then treated with gangliosides for 1 hour (A). The bacteria were also added to wells of plastic plates coated with gangliosides, anti-FHA antiserum, and/or anti-flagellin antiserum (B and C). Total RNA was extracted from the free (A) and plate-adhering bacteria (A to C). Data represent fold changes in Bpr4 expression (mean + SEM) compared to the bacteria grown in SS broth (n = 6 biological replicates). Data were obtained from two independent experiments and statistically analyzed by two-way ANOVA with Sidak multiple-comparison test (A) or one-way ANOVA with Tukey’s multiple-comparison test (B and C). ***P < 0.001 and ****P < 0.0001. (D) Subcellular localization of MotA in B. pertussis. B. pertussis 18323-dsRed-motA/Ptac-gfp was washed and resuspended in Hanks’-Hepes BSA. The bacterial suspensions were seeded on a glass-based dish and subjected to confocal microscopy after incubation at 37°C for 1 hour. Fluorescent images of the bacteria were captured at 2-min intervals after the addition of gangliosides (1 μg/ml) or ethanol (1 μl/ml; vehicle). Magenta and cyan indicate DsRed-MotA and GFP, respectively. Note that DsRed-MotA was observed as fluorescent puncta, because multiple MotAB stators consisting of the MotA pentamer and the MotB dimer are assembled into the flagellar motor in functional flagella (36). The fluorescent images are representatives of two independent experiments. Scale bars, 2 μm.

Interference of flagellar rotation triggers Bpr4 up-regulation

To explore bacterial immobilization as an essential requirement for Bpr4 up-regulation, we examined bacteria immobilized by different agents. To avoid nonspecific bacterial attachment, plastic plates were first coated with gangliosides, anti-FHA antiserum, and/or anti-flagellin antiserum, and then the bacteria were applied to the wells of the plate. Bpr4 up-regulation in B. pertussis WT was observed only when the plate was coated with both gangliosides and anti-FHA antiserum (Fig. 8B). Similar results were obtained in bacteria immobilized to plates coated with both anti-FHA and anti-flagellin (Fig. 8C), indicating that anti-flagellin could substitute for gangliosides. Bacteria on plates coated with one of gangliosides, anti-FHA, or anti-flagellin did not exhibit Bpr4 up-regulation. In these experiments, the bacterial cell bodies and flagella were likely immobilized on the plastic plate by anti-FHA, and gangliosides or anti-flagellin, thus interfering with the flagellar rotation. Therefore, we concluded that interference of the flagellar rotation triggers Bpr4 up-regulation.

Flagellar stators, including MotAB, are assembled into the flagellar motor in functional flagella (23, 36). Previous reports demonstrated that the stators of dysfunctional flagella are disengaged from the flagellar complex, and the cytoplasmic region of the disengaged flagellar stator interacts with Dgc on the inner membrane to stimulate Dgc’s enzymatic activity (24, 37, 38). Therefore, we hypothesized that the immobilization of both bacterial cell bodies and flagella leads MotA to dissociate from the flagellar complex and activate DgcB. We examined this point by confocal microscopy. DsRed-MotA was observed as fluorescent puncta in bacteria adhering to the abiotic surface of the culture plates, indicating the incorporation of MotA into the basal flagellar complex of B. pertussis, which reportedly forms a single flagellum (29). After the addition of gangliosides, the fluorescent signals disappeared within 8 min (Fig. 8D), implying MotA diffusion. We therefore concluded that gangliosides, which are adhesive to the abiotic plastic surface and bind to flagellin (Fig. 8A, illustration), disturb flagellar rotation by cross-linking flagellin with the plastic surface to trigger the dissociation of MotA from the basal body. We further examined the role of MotA in Bpr4 up-regulation using B. pertussis ΔmotB, MotA of which is no longer assembled into the flagellar motor, because MotB binds to peptidoglycan and serves to anchor the MotAB stator around the flagellar complex (23, 36). Bpr4 up-regulation was observed in ΔmotB, but not ΔmotA, adhering to plastic plates coated with anti-FHA without gangliosides and anti-flagellin (fig. S3, D and E), indicating that the diffusion of MotA due to the defect in MotB initiates signal transduction to up-regulate Bpr4. These results also imply that the bacteria need to be immobilized to up-regulate Bpr4 even when their flagella are dysfunctional, because ΔmotB did not show Bpr4 up-regulation without anti-FHA.

DISCUSSION

Here, we demonstrate that Bpr4, an sRNA, is up-regulated through flagellum-mediated sensing and contributes to B. pertussis infection by up-regulating the expression of FHA (Fig. 9). The interaction of flagellin with gangliosides on cells to which the bacteria adhere likely interferes with flagellar rotation and provides a cue to up-regulate Bpr4. The interaction between flagella and gangliosides is specific, as other carbohydrates and lipids composing the cell membrane did not induce Bpr4 up-regulation. Gangliosides are known to serve as cell surface receptors for bacterial toxins to be internalized and for pathogenic bacteria to adhere to or invade host cells (32, 39–42). In addition, the present study indicates that gangliosides serve as a host environmental cue leading bacteria to enhance colonization. B. pertussis was recently found to produce flagella and become motile under certain conditions (29); however, it remained unclear whether flagella play any role in bacterial infection. Our results revealed that flagella are produced by the bacteria, function as a mechanosensor for the host environment, and contribute to the bacterial infection. B. pertussis was reported to exhibit motility when incubated in the presence of BSA, implying that the bacteria may produce functional flagella in response to albumin during the course of infection; however, the mechanism through which albumin stimulates flagellar biosynthesis in the bacteria remains to be elucidated.

Fig. 9. The mechanism of sequential up-regulation of Bpr4 and FHA upon bacterial adherence to the host cell.

(A) MotAB is incorporated into the flagellar basal body in motile B. pertussis. (B) The bacterial cell body and flagellin are anchored on the cell via FHA and gangliosides on the cell surface, respectively, resulting in the interference of flagellar rotation, which causes disengagement of MotAB in the inner membrane. MotA interacts with and activates DgcB to generate c-di-GMP. The accumulated c-di-GMP induces Bpr4 up-regulation under the control of the RisK/RisA two-component system. It remains unknown whether RisA directly up-regulates Bpr4 expression. Bpr4 binds to the 5′UTR of fhaB with the aid of Hfq and protects it from RNaseE-mediated degradation, resulting in the posttranscriptional up-regulation of FHA, which facilitates bacterial colonization.

We demonstrated that interference of the flagellar rotation caused MotA to dissociate from the flagellar basal body. The free MotA then interacted with and stimulated DgcB to generate c-di-GMP. Previous studies also indicated that Dgc activation and c-di-GMP generation are controlled by a component of the flagellar stator in Caulobacter crescentus, Pseudomonas aeruginosa, and Vibrio cholerae (25–27, 43). These signal pathways commonly regulate the motile-to-sessile transition of the bacteria. Similarly, B. pertussis up-regulated FHA expression through this pathway and made itself more adhesive to the host cell, resulting in enhanced colonization. In addition, c-di-GMP has also been reported to repress bacterial motility by dissociation of flagellar stators, including MotAB, proposing a positive feedback loop, in which the disengagement of MotA from the flagellar motor stimulates the production of c-di-GMP, which further causes MotAB to disengage (24, 44). The present study also showed that the flagellum- and c-di-GMP–mediated pathway regulates the expression of a virulence factor through the up-regulation of sRNA. The up-regulation of the sRNA Bpr4 was dependent on the RisK/RisA two-component system. Previous studies suggested that the intrabacterial concentration of c-di-GMP influences RisA activity to regulate downstream genes (21, 22); however, the Dgc involved in this system had not been identified. The present study identified DgcB as the enzyme providing c-di-GMP to activate RisA. A BLAST search (https://blast.ncbi.nlm.nih.gov/Blast.cgi) revealed that DgcB is shared by Bordetella species, including B. bronchiseptica and B. parapertussis (99% identical at the amino acid level), suggesting similar signaling cascades conserved in the bacteria.

The flagellum-to-RisA pathway that we found likely regulates the expression of not only Bpr4 but also many other genes, including virulence-repressed genes (vrgs), which are known to reside downstream of RisA (45, 46). Previous studies revealed some vrgs that are up-regulated during B. pertussis infection (5, 6). In addition, activated RisA is considered to down-regulate flagellar biosynthesis (22). These observations together with ours suggest that the RisK/RisA system plays a crucial role in the motile-to-sessile transition: Bacteria in the motile stage recognize host cells via the interference of flagellar rotation, then they firmly colonize via the up-regulation of FHA, and finally they turn into the sessile stage by shutting down the flagellar biosynthesis. Moreover, we showed that Bpr4 was up-regulated immediately after bacterial inoculation into mice and persisted at higher levels for at least 10 days after inoculation, leading to the up-regulation of FHA. FHA, which is commonly produced by closely related “classical Bordetella,” B. pertussis, B. parapertussis, and B. bronchiseptica, plays a role in bacterial adhesion to airway epithelial cells, bacterial shedding and transmission to new hosts, and evasion from immune-mediated clearance (47, 48). Therefore, Bpr4–up-regulated FHA likely contributes to various stages throughout the infection.

Previous studies revealed that hundreds of sRNA candidates are involved in gene expressions in B. pertussis (9–11). Of these, only one sRNA, RgtA, has been functionally analyzed and found to down-regulate the expression of a protein involved in glutamate transport by forming base pairs with the 5′UTR of its mRNA (11). Meanwhile, we identified nine sRNAs, including Bpr4, which are highly transcribed in B. pertussis upon the colonization of mouse tracheas (15), and Bpr4 as the first sRNA contributing to the pathogenicity of B. pertussis. Given the role of Bpr4 in bacterial colonization, the remaining eight sRNAs may also contribute to the establishment of B. pertussis infection. Notably, the construction of Bpr5-deficient mutants was unsuccessful in this study, implying that this sRNA is indispensable for the viability of the bacteria. Further work should consider the involvement of these sRNAs in B. pertussis pathogenicity, metabolism, and environmental responses.

Pathogenic bacteria sense the host cells in various ways and induce transcriptional changes to establish infection. The present study proposes an additional system involving flagellin, MotA, DgcB, and c-di-GMP, which are widely conserved in flagellated bacteria, to recognize the host cells and contribute to bacterial infection. Since c-di-GMP regulates genes involved in bacterial pathogenicity and/or adaptation to host environments (49), the flagellum-triggered sensory system may function as a gene regulatory system for a wide variety of pathogenic bacteria and serve as a potential target for control of bacterial infections.

MATERIALS AND METHODS

Bacterial strains and culture conditions

B. pertussis strains 18323 and Tohama were maintained in our laboratory, and clinical strains BP139, BP140, BP141, and BP144 were provided by K. Kamachi (National Institute of Infectious Diseases). B. pertussis was grown on Bordet-Gengou (BG) agar (Becton Dickinson, 248200) containing 1% HIPOLYPEPTON (Nihon Pharmaceutical, 396-02118), 1% glycerol, 15% defibrinated horse blood, and ceftibuten (10 μg/ml) (BG plate). The bacteria recovered from colonies on BG plates were suspended in SS broth [sodium hydrogen l(+)-glutamate monohydrate (10.7 g/liter), l(−)-proline (0.24 g/liter), NaCl (2.5 g/liter), KCl (0.2 g/liter), KH₂PO₄ (0.5 g/liter), MgCl₂·6H₂O (0.1 g/liter), CaCl₂·2H₂O (20 mg/liter), tris (hydroxymethyl) aminomethane (11.1 g/liter), casamino acids (10 g/liter), 2,6-di-O-methyl-β-cyclodextrin (1 g/liter), l-cysteine hydrochloride monohydrate (40 mg/liter), iron(II) sulfate heptahydrate (10 mg/liter), l(+)-ascorbic acid (0.4 g/liter), nicotinic acid (4 mg/liter), and glutathione (0.15 g/liter; reduced form) (pH 7.4)] to make an optical density at 650 nm (OD₆₅₀) of 0.2 and incubated at 37°C with shaking for 12 to 24 hours. The number of colony-forming units (CFU) was estimated from OD₆₅₀ values of fresh cultures according to the following equation: 1 OD₆₅₀ = 3.3 × 10⁹ CFU/ml. The bacterial lysates were prepared as described previously (15). The culture supernatants of B. pertussis were harvested by centrifugation at 8000g for 10 min and filtered through 0.2-μm pore membranes (Thermo Fisher Scientific, 720-1320). Proteins in the culture supernatants were precipitated with 10% trichloroacetic acid (TCA). E. coli used in this study was grown on Luria-Bertani agar or broth. The growth media were supplemented with antibiotics when necessary at the following concentrations: ampicillin (50 μg/ml), gentamicin (10 μg/ml), kanamycin (25 μg/ml), and tetracycline (20 μg/ml).

Cell cultures

Calu-3 lung adenocarcinoma cells that had been obtained from the American Type Culture Collection were maintained in Minimum Essential Medium Eagle (Sigma-Aldrich, M4655) supplemented with 10% FBS (HyClone; Cytiva, SH30910.03) at 37°C under 5% CO₂ and seeded at 1 × 10⁶ and 5 × 10⁴ cells per well into each well of 6- and 96-well plates (IWAKI, 3810-006 and 3860-096). Unless otherwise specified, the tissue culture–treated plates, which are generally used for adhesive cell culture, were used in this study. In some experiments, the cells were fixed with 4% PFA in Dulbecco’s modified phosphate-buffered saline (PBS) or methanol for 10 min or treated with 10 μM PPMP (Sigma-Aldrich, P4194) for 3 days, NA (1 U/ml) from Clostridium perfringens (Sigma-Aldrich, N2876) for 3 hours, and 10 mM MβCD (Sigma-Aldrich, C4555) for 2 hours. THP-1 cells were maintained in RPMI 1640 medium (Thermo Fisher Scientific, 11875093) supplemented with 10% FBS at 37°C under 5% CO₂ (50), seeded at 1 × 10⁶ cells per well into each well of a six-well plate, and allowed to differentiate into macrophage-like cells by treatment with phorbol 12-myristate 13-acetate (100 ng/ml; Sigma-Aldrich, P1585) at 37°C under 5% CO₂ for 48 hours.

TLR5-knockout cells

TLR5-knockout Calu-3 cells were generated by deleting the full length of TLR5 gene with the CRISPR-Cas9 system as described previously with slight modifications (51). The single-guide RNA target sequences for the 5′ and 3′ regions of TLR5 gene were 5′-GGCCGGGGGCCCTAAGTCGT-3′ and 5′-GCTGTTATTAGACTTTCCAC-3′, respectively. These sequences were inserted into the Bbs I site of pX330, which was provided by M. Ikawa (Osaka University), using T4 DNA ligase. The resultant plasmids were transfected with Calu-3 cells using Lipofectamine 3000 (Thermo Fisher Scientific, L3000001) according to the manufacturer’s instructions. The clones obtained by limiting dilution were genotyped by polymerase chain reaction (PCR) using the primers TLR5-check-S1, TLR5-check-S2, and TLR5-check-AS (table S1). The absence of TLR5 protein in the cell lysates from the clones was confirmed by immunoblotting for TLR5.

Construction of plasmids and mutant strains

Mutant strains derived from B. pertussis 18323 were constructed by double-crossover homologous recombination as described previously (15, 52). The primers used in this study are listed in table S1. For the generation of 18323-Δbpr4, Δbpr8, Δbpr9, ΔrisA, ΔdgcA, ΔdgcB, ΔflaA, ΔmotA, ΔmotB, and ΔPrne, ~1–kilo–base pair (kbp) DNA fragments of the up- and downstream regions of bpr4, bpr8, bpr9, risA, dgcA, dgcB, flaA, motA, and motB genes and Prne were amplified by PCR using the genomic DNA (gDNA) from B. pertussis 18323 as the template with the appropriate primers. The DNA fragments of the up- and downstream regions were ligated and inserted into the Sac I site of pABB-CRS2-Gm (53), which was provided by A. Abe (Kitasato University), using the In-Fusion HD Cloning Kit (TaKaRa Bio, 639649). The resultant plasmids, designated Δbpr4-, Δbpr8-, Δbpr9-, ΔrisA-, ΔdgcA-, ΔdgcB-, ΔflaA-, ΔmotA-, ΔmotB-, and ΔPrne-pABB-CRS2-Gm, were introduced into E. coli DH5α λpir and transconjugated into B. pertussis 18323 by triparental conjugation with a helper E. coli HB101 harboring pRK2013, which was provided by K. Minamisawa (Tohoku University).

The two-component sensor kinase–deficient mutants and ΔfhaB were generated as follows. DNA fragments (~1.2 kbp) of the up- and downstream regions of bvgS, kdpD, phoR, plrS, risK, bp0157, bp0992, bp1092, bp2206, bp2548, bp3137, bp353, bp3866, and fhaB genes were amplified by PCR using gDNA from B. pertussis 18323 as the template with the appropriate primers. A DNA fragment of the chloramphenicol-resistant (Cm^r) gene was also amplified by PCR using pKD3 (54) as the template with primers CmR-S and CmR-AS. The PCR products of the up- and downstream regions of each gene were ligated to the 5′ and 3′ ends of the amplified Cm^r, respectively, and the resultant fragments containing Cm^r were ligated to an inverse PCR product, which was generated by PCR using pABB-CRS2-Gm as the template with primers pABB-inverse-S and pABB-inverse-AS, using the In-Fusion HD Cloning Kit. The resultant plasmids, designated ΔbvgS, ΔkdpD, ΔphoR, ΔplrS, ΔrisK, Δbp0157, Δbp0992, Δbp1092, Δbp2206, Δbp2548, Δbp3137, Δbp3535, Δbp3866, and ΔfhaB-pABB-CRS2-Gm, were introduced into E. coli DH5α λpir and transconjugated into B. pertussis 18323 by triparental conjugation.

For the complementation experiments, the bpr4 gene and its up- and downstream regions were amplified by PCR using the gDNA from B. pertussis 18323 as the template with primers Pbpr4-BamHI-S and Tbpr4-EcoRI-AS. The PCR product was inserted into the Bam HI–Eco RI sites of pBBR1MCS5 (55) using the In-Fusion HD Cloning Kit. The resultant plasmid was designated pbpr4. The Bpr4_m expression vector, designated pbpr4_m, was constructed by site-directed mutagenesis of the bpr4 gene. Two distinct DNA fragments of ~0.3 and ~0.2 kbp composing bpr4 with the mutation were amplified by PCR using pbpr4 as the template with primers Pbpr4-BamHI-S and bpr4m-AS, and bpr4m-S and Tbpr4-EcoRI-AS, respectively. The DNA fragments were ligated and inserted into the Bam HI–Eco RI sites of pBBR1MCS5 using the In-Fusion HD Cloning Kit. Plasmids for the ectopic expression of Bpr4, Bpr4_m, FHA, DgcA, and DgcB were constructed on the basis of a Ptac-driven GFP expression plasmid, Ptac-gfp, which was pBBR1MCS5 carrying Ptac, gfp, and trpA terminator (52). DNA fragments of bpr4, bpr4_m, fhaB, dgcA, and dgcB genes were amplified by PCR using pbpr4, pbpr4_m, and the gDNA from B. pertussis 18323 as the templates with primers bpr4-SmaI-S and bpr4-EcoRI-AS, fhaB-SmaI-S and fhaB-EcoRI-AS, dgcA-SmaI-S and dgcA-EcoRI-AS, and dgcB-SmaI-S and dgcB-EcoRI-AS. The DNA fragments were inserted into the Sma I–Eco RI sites of Ptac-gfp using the In-Fusion HD Cloning Kit, and the obtained plasmids were designated Ptac-bpr4, -bpr4_m, -fhaB, -dgcA, and -dgcB. Last, these resultant plasmids and the empty vector (pBBR1MCS5) were introduced into E. coli DH5α and transconjugated into B. pertussis 18323 WT, Δbpr4, ΔPrne, ΔrisA, ΔdgcB, ΔflaA, and ΔmotA strains by triparental conjugation.

For the GFP reporter assay, the reporter plasmid harboring the gfp gene downstream of Pbpr4 was generated as follows. A DNA fragment containing the 0.3-kbp upstream region of bpr4 gene was amplified by PCR using the gDNA from B. pertussis 18323 as the template with primers Pbpr4-BamHI-S and Pbpr4-AS. A DNA fragment of the gfp gene was also amplified by PCR using Ptac-gfp as the template with primers gfp-S and gfp-AS. The PCR products were ligated and inserted into the Bam HI–Eco RI sites of Ptac-gfp using the In-Fusion HD Cloning Kit. The resultant plasmid, designated Pbpr4-gfp, was introduced into E. coli DH5α and transconjugated into B. pertussis 18323 WT and ΔrisA strains by triparental conjugation.

The plasmid pMariT was generated for the delivery of the mariner transposon containing the tetracycline resistance gene (Tc^r) in transposon mutagenesis. A DNA fragment carrying Tc^r was amplified by PCR using pBBR1MCS3 (55) as the template with primers pBBR1-TcR-S and pBBR1-TcR-AS. Inverse PCR was also performed using pMariK (56) as the template with primers pMariK-S and pMariK-AS. These PCR products were then ligated using the In-Fusion HD Cloning Kit. The resultant plasmid, designated pMariT and carrying Tc^r in place of the kanamycin resistance gene in the pMariK backbone, was introduced into E. coli S17-1 λpir and transconjugated into B. pertussis 18323 WT strain harboring Pbpr4-gfp by biparental conjugation.

A B. pertussis mutant strain producing DsRed fused to the N terminus of MotA was generated as follows. Approximately 1-kbp DNA fragments covering the upstream and 5′ regions of motA gene were amplified by PCR using the gDNA from B. pertussis 18323 as the template with primers motA-U-S and moA-U-AS2, and motA-5′-S and motA-5′-AS, respectively. A DNA fragment of the dsRed gene was also amplified by PCR using pDsRed2-Nuc (TaKaRa Bio, 632408) as the template with primers dsRed-S and dsRed-AS. The PCR products of the upstream and 5′ regions of motA gene were ligated to the 5′ and 3′ ends of the amplified dsRed, and the resultant fragment was inserted into the Sac I site of pABB-CRS2-Gm using the In-Fusion HD Cloning Kit. The resultant plasmid, designated dsRed-motA-pABB-CRS2-Gm, was introduced into E. coli DH5α λpir and transconjugated into B. pertussis 18323 by triparental conjugation. Ptac-gfp was then transconjugated into the B. pertussis 18323 mutant strain producing DsRed-MotA by triparental conjugation for visualization of the bacterial body. The resultant strain was designated 18323-dsRed-motA/Ptac-gfp and used for confocal microscopy.

For the generation of expression vectors for poly-histidine affinity tag (HAT)–tagged recombinant proteins, DNA fragments of hfq and flaA genes encoding Hfq and FliC proteins were amplified by PCR using gDNA from B. pertussis 18323 as the template with primers hfq-XhoI-S and hfq-EcoRI-AS, and flaA-XhoI-S and flaA-EcoRI-AS, respectively. Each DNA fragment was inserted into the Xho I–Eco RI sites of pColdII-HAT (57) using an In-Fusion HD Cloning Kit. The resultant plasmids, designated pColdII-HAT-hfq and -flaA, were transformed into E. coli DH5α.

Template plasmids for the in vitro transcription of Bpr4, Bpr4_m, and fhaB-5′-UTR were constructed to examine the binding of the RNAs. DNA fragments of bpr4, bpr4_m, and fhaB-5′-UTR were amplified by PCR using pbpr4, pbpr4_m, and gDNA from B. pertussis 18323 as the templates with primers bpr4-EcoRI-S and bpr4-BamHI-AS, bpr4m-EcoRI-S and bpr4-BamHI-AS, and fhaB-5′-UTR-EcoRI-S and fhaB-5′-UTR-BamHI-AS, respectively. Each DNA fragment was inserted downstream of the T7 promoter on the Eco RI–Bam HI sites of pSPT18 (Sigma-Aldrich) using the In-Fusion HD Cloning Kit, and the plasmids designated pSPT18-bpr4, -bpr4_m, and -fhaB-5′-UTR were obtained.

For the bacterial two-hybrid assay, DNA fragments encoding the T18 and T25 regions of adenylate cyclase toxin were amplified by PCR using gDNA from B. pertussis 18323 as the template with primers T18-S and T18(dgcA)-AS, T18-S and T18(dgcB)-AS, T25-S and T25(motA)-AS, and T25-S and T25(motB)-AS, respectively. DNA fragments of dgcA, dgcB, motA, and motB genes were also amplified by PCR using Ptac-dgcA and -dgcB, and gDNA from B. pertussis 18323 as the templates with primers dgcA-S and dgcA-HindIII-AS, dgcB-S and dgcB-HindIII-AS, motA-S and motA-HindIII-AS, and motB-S and motB-HindIII-AS, respectively. The T18-coding gene was ligated to the 5′ ends of the amplified dgcA and dgcB genes, and the resultant fragments were inserted into the Nco I–Hind III sites of pET-21d(+) (Merck Millipore, 69743) using the In-Fusion HD Cloning Kit. The T25-coding gene was ligated to the 5′ ends of the amplified motA and motB genes, and the resultant fragments were inserted into the Nco I–Hind III sites of pET-28b(+) (Merck Millipore, 69865) using the In-Fusion HD Cloning Kit. These obtained plasmids were designated pET21d-T18-dgcA and -dgcB and pET28b-T25-motA and -motB and transformed into E. coli BL21(DE3).

Purification of recombinant proteins

The expressions of rHfq and rFliC from E. coli DH5α harboring pColdII-HAT-hfq and -flaA were induced by incubating with 1 mM isopropyl-β-d-thiogalactopyranoside (IPTG) at 15°C for 24 hours. The bacteria were disrupted by sonication in 50 mM sodium phosphate buffer and 300 mM NaCl (pH 8.0) (buffer A) containing 7.5 mM imidazole. The sonicated suspensions were centrifuged at 12,000g for 5 min, and the supernatants were independently applied to a column of HIS-Select Nickel Affinity Gel (Sigma-Aldrich, P6611) equilibrated with buffer A containing 7.5 mM imidazole. After nonabsorbed substances had been washed out of the column with buffer A containing 7.5 mM imidazole, the rHfq and rFliC proteins were eluted with buffer A containing 300 mM imidazole. Imidazole in the rHfq and rFliC fractions was removed by dialysis against PBS.

Protein identification by LC-ESI-MS/MS

TCA-precipitated culture supernatants of B. pertussis were separated by SDS-PAGE, and the gels were stained with Coomassie Brilliant Blue R-250. A protein band of interest was excised, digested by trypsin, and then subjected to LC-ESI-MS/MS analysis using an L-column octa decyl silyl (0.1 × 150 mm; Chemical Evaluation and Research Institute) on a nanoflow system (Advance UHPLC; Bruker Daltonics) coupled with a linear trap quadrupole Orbital Velos plus electron transfer dissociation mass spectrometer (Thermo Fisher Scientific). Tandem mass spectra were acquired automatically and searched against the B. pertussis 18323 database from NCBI using the Mascot server (Matrix Science).

RNA-RNA binding assay

RNA-RNA binding assays were performed as described previously with slight modifications (58). Bpr4, Bpr4_m, and fhaB-5′-UTR were prepared by in vitro transcription using pSPT18-bpr4, -bpr4_m, and -fhaB-5′-UTR as the templates with T7 RNA polymerase (Roche, 10881767001) according to the manufacturer’s instructions. Ten micromolars Bpr4 or Bpr4_m was incubated with 10 μM fhaB-5′-UTR in the presence of 100 μM rHfq or BSA (FUJIFILM Wako Chemicals, 015-27053) in a total of 10 μl of 20 mM tris-acetate buffer (pH 7.6), containing 100 mM sodium acetate and 5 mM magnesium acetate, at 37°C for 15 min. After the addition of RNA-loading buffer (FUJIFILM Wako Chemicals, 182-02571), 10 μl of each solution was subjected to 3% nondenaturing agarose gel electrophoresis with ethidium bromide staining.

Bacterial infection in mice

Seven-week-old male C57BL/6J mice (CLEA Japan and Japan SLC) were anesthetized with a mixture of medetomidine (Kyoritsu Seiyaku), midazolam (Teva Takeda Pharma), and butorphanol (Meiji Seika Pharma) at final doses of 0.3, 2, and 5 mg/kg of body weight, respectively. B. pertussis was intranasally inoculated at 1 × 10⁷ CFU/25 μl (SS broth) into anesthetized mice using a micropipette with a needle-like tip. The amounts of bacteria were confirmed by counting colonies after cultivation of the inocula on BG plates. On the indicated times and days after inoculation, the mice were euthanized with pentobarbital, and the tracheas were collected. For quantitative reverse transcription PCR (qRT-PCR), total RNA was extracted and purified from the tracheas using RNase-free deoxyribonuclease I (DNase I; TaKaRa Bio, 2270) and the PureLink RNA Mini Kit (Thermo Fisher Scientific, 12183020) as described previously (15). In independent experiments, the tracheas were aseptically excised, minced, and homogenized in PBS with BioMasher I (Nippi, 49118-52). The resultant tissue extracts were serially diluted with PBS and spread onto BG plates. The bacteria on the plates were incubated at 37°C for 3 to 4 days, and the number of CFU was determined. All animal experiments were approved by the Animal Care and Use Committee of the Research Institute for Microbial Diseases, Osaka University, and carried out according to the Regulations on Animal Experiments at Osaka University.

Bacterial adhesion to cells

B. pertussis was washed and resuspended in Hanks’ balanced salt solution (Sigma-Aldrich, H6136) containing 20 mM Hepes-NaOH (pH 7.4) and BSA (1 mg/ml; Hanks’-Hepes BSA) at a concentration of 5 × 10⁷ CFU/ml. The media of Calu-3 and differentiated THP-1 cells that had been seeded in each well of a six-well plate were replaced with the bacterial suspensions to make a multiplicity of infection (MOI) of 100. The bacteria were allowed to attach to the cells by centrifugation at 500g for 5 min and then incubated at 37°C under 5% CO₂. After 1 hour, supernatants were collected, and the cells were washed three times with Hanks’-Hepes BSA. For qRT-PCR, total RNA was then extracted and purified from the bacteria in the supernatant (free bacteria) and those adhering to the cells (cell-adhering bacteria) using the PureLink RNA Mini Kit and RNase-free DNase according to the manufacturer’s instructions. In independent experiments, the cells were treated with 0.1% saponin in Hanks’-Hepes BSA to detach the bacteria. The resultant suspensions were serially diluted with Hanks’-Hepes BSA, and the CFU was enumerated as described above.

Bacterial adhesion to abiotic surfaces

The bacterial suspensions of B. pertussis (5 × 10⁷ CFU/ml in Hanks’-Hepes BSA) were added at 1 × 10⁸ and 1.5 × 10⁷ CFU per well into each well of collagen type I–coated (IWAKI, 4810-010), gelatin-coated (IWAKI, 4810-020), or poly-l-lysine–coated (IWAKI, 4810-040) 6-well plates and a 96-well microplate (ELISA Plate H; Sumitomo Bakelite, MS-8596F), which was coated with gangliosides (5 μg/ml) (ganglioside mixture, ammonium salt, and bovine brain; Merck Millipore, 345717) and/or rabbit anti-flagellin (FliC) (Abcam, ab93713) and anti-FHA antisera (59) diluted 50-fold with PBS at 4°C overnight. After centrifugation of the plates at 500g for 5 min, the bacteria in the wells were incubated at 37°C for 1 hour and washed three times with Hanks’-Hepes BSA. The plate-adhering bacteria were then treated with gangliosides (1 μg/ml), lewis X (FUJIFILM Wako Chemicals, L395000), heparan sulfate (Sigma-Aldrich, H4777), phosphatidic acid (PA; Sigma-Aldrich, P9511), phosphatidylcholine (PC; Sigma-Aldrich, P5763), phosphatidylethanolamine (PE; Sigma-Aldrich, P7943), phosphatidylinositol (PI; Sigma-Aldrich, P2517), phosphatidylserine (PS; Sigma-Aldrich, P6641), and cholesterol (Sigma-Aldrich, C8867) for 1 hour. In an independent experiment, B. pertussis in the wells was treated with gangliosides (1 μg/ml) without washing, and free and plate-adhering bacteria were collected after a 1-hour incubation. Total RNA was then extracted and purified from the free and plate-adhering bacteria using the PureLink RNA Mini Kit and RNase-free DNase according to the manufacturer’s instructions. The CFU of bacteria adhering to abiotic surfaces was calculated similarly to that for bacterial adhesion to cells.

Quantitative reverse transcription polymerase chain reaction

Total RNA samples of B. pertussis extracted and purified from mouse tracheas, free and cell- and plate-adhering bacteria, and the bacteria grown in SS broth were reverse-transcribed into complementary DNA (cDNA) using the PrimeScript RT Reagent Kit (TaKaRa Bio, RR037) with random hexamers. The relative expression levels of Bpr4 and fhaB, rne, dgcA, dgcB, gfp, flaA, motA, and recA transcripts were determined using Fast SYBR Green Master Mix (Thermo Fisher Scientific, 4385612) and the primers listed in table S1 with the StepOnePlus Real-Time PCR System (Applied Biosystems) under the following conditions: initial denaturation for 1 min at 95°C and 40 cycles at 95°C for 15 s and 60°C for 1 min. The relative amount of each transcript was calculated using the ΔΔCt method normalized relative to that of recA mRNA as an internal control for each sample.

GFP reporter assay for Pbpr4 activity

B. pertussis strains and transposon-integrated mutants harboring Pbpr4-gfp were diluted with Hanks’-Hepes BSA at a concentration of 1.67 × 10⁷ CFU/ml, and the media of Calu-3 cells that had been seeded in each well of a 96-well plate were replaced with 0.3 ml of the bacterial suspensions (MOI of 100). After centrifugation of the plates at 500g for 5 min, the cells were incubated with the bacteria for 1 hour. In an independent experiment, B. pertussis strains harboring Pbpr4-gfp were diluted with SS broth at 1 × 10⁸ CFU/300 μl and added to each well of an empty 96-well plate. The GFP fluorescence intensity of each well was measured using the Glomax-Multi Detection System (Promega).

Determination of transposon insertion sites

The transposon insertion sites were determined as described with slight modification (50, 56). Briefly, gDNA was extracted from the transposon-integrated mutants using the DNeasy Blood and Tissue Kit (Qiagen, 69504) and digested with Sau 3AI. The digested products were circulated with T4 DNA ligase (Promega, M1801) and used as a template for PCR with primers TcR-S and Mari5. The PCR products were subjected to direct sequencing using the same primers. The obtained sequences were aligned to the B. pertussis 18323 genome sequence, and the transposon insertion sites were identified.

Confocal microscopy

The bacterial suspensions of B. pertussis 18323-dsRed-motA/Ptac-gfp mutant (5 × 10⁷ CFU/ml in Hanks’-Hepes BSA) were seeded on a 35-mm glass-based dish (IWAKI, 3910-035), incubated at 37°C for 1 hour, and then washed three times with Hanks’-Hepes BSA to remove nonadhered bacteria. Fluorescent images of the bacteria, which were treated with gangliosides (1 μg/ml) or ethanol (1 μl/ml; vehicle), were captured at 2-min intervals using an LSM880 confocal laser scanning microscope with ZEN2.3 software (ZEISS).

Motility assay

B. pertussis was washed and resuspended in Hanks’-Hepes BSA at a concentration of 2 × 10⁹ CFU/ml and incubated at 37°C for 1 hour. One microliter of bacterial suspensions was then stabbed into motility agar plates [0.45% potato infusion powder (Sigma-Aldrich, 52424), 0.55% NaCl, 1% HIPOLYPEPTON, 1% glycerol, ceftibuten (10 μg/ml), BSA (10 mg/ml), and 0.4% agar]. After incubation at 37°C for 3 days, diameters of the motility zone were measured.

Bacterial two-hybrid system

Interactions between MotA and DgcB derived from B. pertussis were analyzed by the bacterial two-hybrid system as described previously with modifications (24, 28). T18 and T25 fusion proteins were coexpressed in E. coli BL21(DE3) harboring pET21d-T18-dgcA and -dgcB, and/or pET28b-T25-motA and -motB by incubating with 1 mM IPTG at 37°C for 3 hours. The bacteria were collected by centrifugation at 8000g for 10 min, resuspended in PBS, and disrupted by five rounds of 2-min sonication with a Bioruptor (Cosmo Bio). The sonicated suspensions were centrifuged at 12,000g for 5 min, and the bacterial lysates were obtained after filtration of the supernatants using 0.2-μm pore membranes. The concentrations of cAMP and c-di-GMP in the bacterial lysates were measured by ELISA.

Ganglioside ELISA

Wells of a 96-well ELISA Plate H were coated with 0.5 μg per well of gangliosides at 4°C overnight and then blocked with PBS containing 10% skim milk at 37°C for 1 hour. B. pertussis and rFliC were serially diluted in SS broth and PBS, respectively; added to the wells of the plates; and incubated at 37°C for 2 hours. The bacteria and rFliC bound to gangliosides were probed with rabbit anti-FHA antiserum and anti–HAT-tag antibody (GenScript, A00636), respectively, followed by goat anti-rabbit immunoglobulin G (IgG)–horseradish peroxidase (HRP) (Jackson ImmunoResearch Laboratories, 111-035-003) at 37°C for 2 hours. After each step, the wells were washed five times with PBS containing 0.05% Tween 20. One hundred microliters of 0.1% TMBZ (3,3′,5,5′-tetramethylbenzidine; Dojindo Laboratories, T022) in 0.1 M citrate-acetate buffer (pH 6.0) containing 0.01% H₂O₂ was added to each well and allowed to react at room temperature for 30 min. The reactions were stopped by the addition of 100 μl per well of 1 M H₂SO₄. The OD₄₅₀ value of each well was measured using the Glomax-Multi Detection System.

Other methods

The concentrations of total proteins, c-di-GMP, and cAMP in the tested materials used in this study were measured using the Micro BCA Protein Assay Kit (Thermo Fisher Scientific, 23235), the c-di-GMP ELISA Kit (MyBioSource, MBS288159), and the cAMP EIA System (Cytiva, RPN2251), respectively, according to the respective manufacturer’s instructions. For immunoblotting, the target proteins were visualized by enhanced chemiluminescence using Immobilon Western (Merck Millipore, WBKLS0500) and LAS-4000mini Luminescent Image Analyzer (GE Healthcare) or Amersham Imager 600 UV (GE Healthcare). Rabbit anti-FtsZ and anti-PTx antisera were prepared as reported previously (52, 59). The following antibodies were purchased from the indicated vendors: rabbit anti-TLR5 (Thermo Fisher Scientific, PA1-41139), mouse anti-GAPDH (glyceraldehyde-3-phosphate dehydrogenase) (FUJIFILM Wako Chemicals, 014-25524), and goat anti-mouse IgG-HRP (Jackson Immune Research Laboratories, 115-035-003).

Statistical analysis

Statistical analyses were performed using Prism 9 (GraphPad Software). Significance is expressed as follows: *P < 0.05, **P < 0.01, ***P < 0.001, and ****P < 0.0001. In all analyses, P < 0.05 was taken to indicate statistical significance.

Supplementary Materials

This PDF file includes:

Figs. S1 to S3

Table S1

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