https://orcid.org/0000-0002-8404-5702
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Figures
Abstract
The bactericidal/permeability-increasing protein (BPI)-inducible protein A (BipA) is a highly conserved protein in Gram-negative bacteria that is structurally similar to translational GTPases such as IF2, EF-Tu, and EF-G. Our previous research showed that deleting bipA in Escherichia coli at 20 °C leads to a defect in 50S ribosomal assembly and impaired lipopolysaccharide (LPS) synthesis. This LPS defect activates the Regulator of Capsule Synthesis (Rcs) pathway, resulting in an overproduction of capsular polysaccharides, a reduction in biofilm formation, and decreased flagella-mediated motility. In this study, we aimed to elucidate the role of BipA in the pathogenicity of Salmonella enterica serovar Typhimurium. We constructed bipA deletion mutants in two pathogenic S. Typhimurium strains, SL1344 and 14028, as well as in the attenuated strain LT2. Our ribosome profiling experiments using the mutant S. Typhimurium strains revealed a defect in ribosome assembly at 20 °C, with the accumulation of abnormal 50S ribosomal subunits. We further demonstrated that the absence of BipA in S. Typhimurium impaired LPS biosynthesis at 20 °C, compromising membrane integrity and presumably activating the Rcs pathway. This activation altered virulence factors, including reduced biofilm formation, particularly in the 14028ΔbipA strain. Furthermore, the SL1344ΔbipA and 14028ΔbipA strains exhibited significantly decreased swimming motility at 20 °C compared to 37 °C, confirmed by microscopic observation showing fewer flagella at 20 °C. Subsequently, both strains exhibited a significant reduction in invasion capability and cytotoxicity toward human intestinal epithelial cells (HCT116). This functional attenuation was corroborated by the decrease in virulence observed in the 14028ΔbipA strain in a mouse model. Our findings suggest that, in S. Typhimurium, BipA functions as a bacterial fitness factor, contributing to ribosome assembly, LPS synthesis, and virulence-related processes, particularly under stress conditions relevant to host environments.
Author summary
Pathogenic bacteria, such as Salmonella enterica serovar Typhimurium, utilize specialized proteins and mechanisms to facilitate infection and evade the host’s immune response. A critical aspect of these mechanisms involves modifying the cell envelope structures to protect the bacteria from host defenses. Understanding these processes is essential for developing new strategies to combat bacterial infections. Our study examines the role of the BipA protein in S. Typhimurium, which is crucial for ribosome assembly and the formation of surface structures, such as lipopolysaccharide, biofilm, and flagella. These structures are essential for bacterial motility, survival under stresses, and pathogenicity. We found that deleting bipA disrupted ribosome assembly at low temperatures. BipA is pivotal for lipopolysaccharide and flagella production under stressed conditions and is necessary for bacterial movement and host cell invasion. These findings highlight the significance of BipA in bacterial adaptation to stress and its regulatory role in key virulence determinant. Our study offers new insights into bacterial survival strategies and identifies potential targets for antimicrobial treatment development.
Citation: Choi E, Ryu E, Kim D, Byun J-W, Kim K, Lee M, et al. (2025) The dual functions of the GTPase BipA in ribosome assembly and surface structure biogenesis in Salmonella enterica serovar Typhimurium. PLoS Pathog 21(4): e1013047. https://doi.org/10.1371/journal.ppat.1013047
Editor: Samuel Wagner, University of Tübingen, GERMANY
Received: October 21, 2024; Accepted: March 16, 2025; Published: April 9, 2025
Copyright: © 2025 Choi et al. This is an open access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Data Availability: All relevant data are within the manuscript and its Supporting Information files.
Funding: This work was supported by a Two-Year Research Grant of Pusan National University (to JH). The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.
Competing interests: The authors have declared that no competing interests exist.
Introduction
Bacterial GTPases play crucial regulatory roles in various cellular processes, such as protein synthesis and adaptation to environmental stress [1,2]. Among these, bactericidal/permeability-increasing protein (BPI)-inducible protein A (BipA), also known as Tyrosine phosphorylation protein A (TypA), is a member of the translational GTPase family [3], and its structure closely resembles those of EF-G and LepA within this group [4]. BipA is upregulated in Salmonella enterica serovar Typhimurium when exposed to the BPI antimicrobial protein [5], which is released by innate immune phagocytic cells, particularly neutrophils [6–8].
BPI is a ~55 kDa single-chain protein, first identified in the primary granules of human neutrophils, known for its antimicrobial efficacy and high selectivity toward Gram-negative bacteria [9]. BPI consists of a highly cationic N-terminal domain that shares a 45% sequence identity with lipopolysaccharide (LPS)-binding protein and an acidic C-terminal domain [10–12]. The cationic N-terminal domain of BPI has a strong affinity for the lipid A-inner core region of LPS [13]. LPS, a key component of the outer membrane of Gram-negative bacteria, is stabilized by divalent ions, such as calcium and magnesium [14,15]. When BPI binds to lipid A, it displaces these ions [16], leading to outer membrane destabilization, increased permeability [17], and disruption of cell division [18]. While the N-terminal domain exhibits antibiotic activity, the C-terminal domain promotes opsonophagocytosis by directing BPI-coated bacteria to neutrophils [19].
In response to BPI cytotoxicity, bacteria trigger the expression of various genes [20]. A well-known strategy for counteracting BPI involves activating genes responsible for modifying or repairing the LPS structure [21]. Research has shown that increasing the length of the polysaccharide chain in LPS enhances resistance to BPI in E. coli, S. Typhimurium, and Proteus mirabilias [22–24]. This elongation may hinder the access of BPI to the lipid A-inner core region. Consequently, the expression of bipA upon BPI exposure may serve as a mechanism for Salmonella to respond to BPI by orchestrating structural changes in LPS. This phenomenon has been observed in bipA-deleted Yersinia pestis and enteropathogenic E. coli, both of which exhibit increased sensitivity to BPI [25,26].
Furthermore, our previous study revealed that E. coli BipA is promptly induced at low temperatures in a cAMP-CRP-dependent manner [27]. Deleting bipA in E. coli K-12 led to a growth defect at low temperatures due to the loss of BipA binding to the 50S ribosomal subunit, leading to a defect in ribosome assembly. Additionally, we observed defects in LPS biosynthesis, which activated the Rcs signaling pathway at low temperatures. This activation enhanced capsule production, inhibited biofilm formation, and decreased flagella-mediated motility [28]. Similarly, bipA deletion in Pseudomonas aeruginosa resulted in reduced biofilm formation and increased antibiotic susceptibility [29]. In Y. pestis, BipA enhances virulence and resistance to neutrophil attacks [26]. Similarly, in P. aeruginosa, bipA deletion weakens biofilm formation and downregulates the type III secretion system (T3SS), reducing bacterial uptake by human macrophages [29]. These findings establish BipA as a crucial regulator of temperature-dependent colony morphology, motility, and pathogenicity associated with surface structures.
In this study, we comprehensively investigated the diverse roles of BipA in ribosome assembly, stress adaptation, virulence-associated functions, and pathogenesis in two pathogenic S. Typhimurium strains, SL1344 and 14028, as well as in the attenuated pathogenic strain LT2, by deleting the bipA gene in each strain. Our data indicate that the role of Salmonella enterica BipA in 50S ribosome biogenesis is temperature-dependent, with bipA mutant strains showing defects in surface structures, such as LPS, biofilm, flagella, and T3SS at low temperatures. Conversely, at 37 °C, reduced ATP levels and increased sensitivity to reactive oxygen species (ROS) likely diminish the pathogenicity of the SL1344 and 14028 strains. Our findings provide insights into BPI resistance through BipA-mediated changes in LPS and highlight BipA as a key fitness factor at low temperatures, facilitating adaptation to stress conditions, such as cold, ROS, and exposure to antimicrobial effectors.
Materials and methods
Ethics statement
All animal study procedures were approved by the Institutional Animal Care and Use Committee of Hallym University (Hallym 2024-2).
Bacterial strains and growth conditions
All bacterial strains used in this study are listed in Table 1. The E. coli and S. Typhimurium strains were cultured at either 37 °C or 20 °C in Luria–Bertani (LB) medium supplemented with ampicillin (Amp, 100 μg/mL), chloramphenicol (Cm, 50 μg/mL), or kanamycin (Kan, 50 μg/mL) as required. To delete the bipA gene in the Salmonella strains, the SebipA::kan cassette was amplified through PCR using pKD13 as a template. The primers are listed in S1 Table. This cassette was introduced into the SL1344, 14028, and LT2 S. Typhimurium strains—which contained the pKD46 plasmid—through electroporation, following the methodology described by Datsenko and Wanner [30]. After transformation, kanamycin-resistant cells were confirmed through PCR to verify gene deletion. To delete rcsA and lon, MG1655 and ESC26 (ΔbipA) cells were subjected to P1 transduction using lysates from JW1935 (rcsA::kan) or JW0429 (lon::kan) cells, resulting in the MG1655ΔrcsA, MG1655ΔbipAΔrcsA, and MG1655Δlon strains. For growth monitoring, the optical density at 600 nm (OD600) was measured using a BioTek Epoch2 microplate spectrophotometer (Agilent Technologies).
Plasmid construction
To construct pACYC177BipA, pACYC177SeBipA, and pACYC184PagP, DNA fragments containing the open reading frame and its flanking regions were obtained through PCR using the E. coli and S. Typhimurium genomes, with the primers listed in S1 Table. The PCR-generated DNA fragments were ligated into the pACYC177 vector at the PsiⅠ site or into the pACYC184 vector at the EcoRV site. Using these clones and mutant primers, site-directed mutagenesis was conducted to generate pACYC177BipAN128D and pACYC177SeBipAN128D. To construct the overexpression vectors pET28SeBipA and pET28SeBipAN128D, the DNA fragments amplified from pACYC177SeBipA and pACYC177SeBipAN128D were digested with the restriction enzymes NdeⅠ and HindⅢ and then inserted into the corresponding sites of the expression vector pET28a. Similarly, the fliCs and fljB DNA fragments were amplified using E. coli or S. Typhimurium genomes as templates, with the primer sets listed in S1 Table. These fragments were subsequently digested with NdeⅠ and HindⅢ and ligated into the corresponding sites of the pIN-A vector, resulting in the construction of pINFliC, pINSeFliC, and pINSeFljB. All plasmids utilized in this study are listed in Table 1.
Quantitative real-time PCR
Total RNA was extracted from 6 × 108 cells using the hot phenol method, as previously described [36]. Fifty micrograms of purified RNAs were incubated with RNase-free DNase I (TaKaRa Bio) at 37 °C for 30 min. After DNase I removal through phenol extraction and ethanol precipitation, RNA quality was assessed using agarose gel electrophoresis, and concentration was measured with the NanoDrop One Spectrophotometer (Thermo Fisher Scientific). For cDNA synthesis, 1 μg of total RNA was mixed with 20 pmol of gene-specific primer in a total volume of 14 μL and heated to 65 °C for 5 min. After cooling at 4 °C for 2 min, a reaction buffer, dithiothreitol, and Moloney murine leukemia virus RNase H–RTase (BioFact RT-Kit) were added. The reverse transcription reaction was conducted at 50 °C for 30 min and then inactivated at 95 °C for 5 min. A quantitative real-time polymerase chain reaction (qRT-PCR) was performed using a QuantStudio 3 Real-Time PCR Instrument (Applied Biosystems), as previously described [27]. The 16S rRNA gene served as an endogenous control to normalize the expression levels of bipA, cspA, gmd, fimD, flagella-related genes, and Salmonella pathogenicity island 1 (SPI-1) genes associated with the T3SS. Relative quantification values were calculated using the comparative CT method [37].
Western blot
After SDS-PAGE, the gel was immersed in transfer buffer (20% methanol in 1X SDS running buffer) and agitated for 20 min. The proteins were then transferred to a PVDF membrane (GE Healthcare) at 10 V for 30 min using a semidry transfer system. The membrane was then immersed in 5% skim milk in Tween-Tris-buffered saline (TTBS) and blocked for 3 h at room temperature. Primary antibodies, including anti-BipA, anti-CspA, anti-OmpA, and anti-FliC, were diluted to ratios ranging from 1:5,000 to 1:20,000 in 3% skim milk in TTBS and applied to the membrane. After overnight incubation with the antibodies at 4 °C, the membrane was washed with TTBS and incubated with alkaline phosphatase–conjugated antirabbit antibody (1:10,000) in 3% skim milk in TTBS at 4 °C for 1 h. Protein detection was conducted using 5-bromo-4-chloro-3-indolyl-phosphate and nitro blue tetrazolium.
Protein overexpression and purification
E. coli BL21(DE3) cells were transformed with a pET28a-derived vector and cultured at 37 °C until the OD600 reached 0.5–0.6. To induce protein expression, isopropyl β-D-thiogalactopyranoside was added to a final concentration of 0.5 mM, and the cultures were incubated at 15 °C overnight. The induced cells were harvested by centrifugation at 4 °C, and the cell pellets were washed with 10 mM Tris-HCl (pH 6.8), followed by another round of centrifugation. For the purification of wild-type and mutant His6-tagged BipA proteins, the cell pellets were resuspended in 20 mL of Buffer A (20 mM Tris-HCl [pH 7.5], 300 mM NaCl, and 2 mM β-mercaptoethanol [BME]), lysed by sonication, and centrifuged at 10,000 × g at 4 °C for 25 min to remove the insoluble fraction. The supernatant was then ultracentrifugated at 70,000 × g at 4 °C for 1 h using a Beckman 70Ti rotor to remove the membrane fraction. The soluble fractions were applied to a Ni-NTA agarose resin (Qiagen) column pre-equilibrated with Buffer A. The column was washed with 40 column volumes of Buffer A, and the proteins were eluted using Buffer A containing 250 mM of imidazole. The eluted proteins were dialyzed overnight twice, each against 2 L of Buffer A.
GTPase assay.
The release of free inorganic phosphate was quantified using a Malachite Green phosphate assay kit (BioAssay Systems). The enzymatic reaction was conducted in a 100 μL mixture containing 20 mM Tris-HCl (pH 7.5), 400 mM KCl, 5 mM MgCl2, 5 mM BME, and 1 mM GTP, along with 1 μM protein, and incubated at 37 °C for 2 h. The reaction was halted by adding 20 μL of Working Reagent to 80 μL of the sample solution, followed by a 30-min incubation at room temperature for color development. Optical absorbance was then measured at 620 nm using a Multiskan GO microplate spectrophotometer (Thermo Fischer Scientific).
Sucrose density gradient sedimentation
Wild-type and bipA-deleted strains were inoculated in LB or LB broth supplemented with Kan. The cultures were grown at 37 °C or 20 °C until the OD600 reached 0.5. Subsequently, the cultures were treated with Cm and further incubated at 37 °C or 20 °C for 3 min. After the Cm treatment and incubation, 50 mL of the cells were harvested at 3,000 rpm for 10 min and resuspended in 1 mL of Buffer BP (20 mM Tris-HCl [pH 7.5], 10 mM MgCl2, 100 mM NH4Cl, and 5 mM BME) for polysome analysis or Buffer BS (20 mM Tris-HCl [pH 7.5], 1 mM MgCl2, 100 mM NH4Cl, and 5 mM BME) for subunit profiling. The cells were treated with 5 μL of lysozyme and subjected to three freeze-thaw cycles for lysis. The cleared cell lysates were obtained after centrifugation in a Beckman ultracentrifuge tube. For polysome analysis, 320 μg of RNA was loaded onto a 10 mL 5%–40% sucrose gradient in Buffer BP, while for subunit profiling, 160 μg of RNA was loaded onto a 10 mL 5%–25% sucrose gradient in Buffer BS. The cell lysates were fractionated by ultracentrifugation at 4 °C for 2.5 h for polysome profiling or 3.5 h for subunit profiling at 37,000 rpm [27].
Bile salt sensitivity test
Overnight cultures incubated at 37 °C were diluted 1:100 in fresh medium and grown at 37 °C until they reached an OD600 of 0.5. The culture was then diluted to an OD600 of 0.2 and subjected to serial dilutions down to 10−2. Subsequently, 2.5 μL of the diluted samples were streaked onto bile salt gradient agar plates with concentrations ranging from 20 to 27 g/L.
Lipopolysaccharide extraction
Colonies grown on LB agar plates incubated at 37 °C for 24 h or at 20 °C for 2 weeks were scraped into 10 mL of phosphate-buffered saline (PBS, pH 7.2). Based on the OD600 reading of the 10 mL suspension, a cell suspension was prepared and harvested at an OD600 of 0.5. LPS extraction was performed following the method described by Kenyon et al. [38]. The cells were resuspended in 200 μL of TAE buffer (40 mM Tris, 20 mM acetic acid, 1 mM EDTA [pH 8.0]) and 400 μL of LPS lysis buffer (50 mM Tris, 100 mM SDS, 0.128 mM NaCl). Subsequently, 600 μL of phenol solution was added to each sample, and the samples were vortexed for 10 s. The mixtures were incubated at 65 °C for 15 min. After cooling to room temperature, the samples were centrifuged at 13,000 rpm for 10 min. LPS was extracted from the aqueous phase and concentrated through ethanol precipitation. The resulting residue was then dissolved in 20 mM Tris-HCl (pH 6.8) and sequentially treated with DNase I, RNase A, and Proteinase K. LPS was re-extracted from the aqueous phase through ethanol precipitation and resuspended in 40 μL of distilled water. LPS samples were analyzed using 15% SDS-PAGE or 16% tricine-SDS-PAGE and visualized with silver staining, as previously described [39].
Crystal violet staining of the biofilm
The wild-type and bipA-deleted strains were grown overnight in LB or LB broth containing Kan at 37 °C. The overnight cultures were inoculated 1:100 into a fresh medium in either glass or polypropylene tubes and incubated at 37 °C for 24 h. The cultures in glass tubes were incubated at 20 °C for 48 h, while those in polypropylene tubes were incubated for 96 h. After incubation, the glass or polypropylene tubes were rinsed with a saline solution and stained with 0.1% crystal violet. The tubes were washed three times with distilled water, and the crystal violet was then dissolved in 95% ethanol. The absorbance was measured at 540 nm.
Swimming motility assays
The cultures with an OD600 of 0.5, incubated at 37 °C, were concentrated 10-fold by centrifugation, and 5 μL of the suspension was injected onto an LB plate containing 0.3% Bacto agar for the swimming assay. The incubation times for the wild-type and bipA-deleted strains were as follows: E. coli for 18 h, SL1344 and 14028 for 4 h, and LT2 for 8 h at 37 °C; and E. coli for 144 h, SL1344 and 14028 for 22 h, and LT2 for 72 h at 20 °C. The migration speed was assessed by measuring the distance traveled from the injection site. The plates were photographed using the Azure C200 Gel Imaging System (Azure Biosystems).
Transmission electron microscopy
Overnight cultures were inoculated 1:100 into fresh LB medium and cultured statically at 37 °C for 1 d or at 20 °C for 3 d. The cells were collected by centrifugation at 3,000 rpm and resuspended in PBS. A 10-μL sample was placed on a 200-mesh formvar/carbon-coated copper grid for 10 min and then blotted with filter paper. The grid containing the sample was washed with PBS and stained with 2% phosphotungstic acid for 2 s. The cells were then examined using transmission electron microscopy (TEM, H-7600, HITACHI), and images were captured at 12,000× magnification.
Human cell lines and culture conditions
Human colorectal intestinal epithelial carcinoma cells (HCT116) were cultured in McCoy’s 5A medium (ATCC), supplemented with 10% (v/v) heat-inactivated fetal bovine serum (Avantor) and 1% (v/v) penicillin-streptomycin (Gibco). The cells were cultured at 37 °C in a humidified atmosphere with 5% CO2.
Cell invasion assay
Wild-type strains were transformed using pACYC177, while the bipA-deleted strains were transformed with either pACYC177 or pACYC177(Se)BipA. The transformants were cultured at 37 °C or 20 °C until they reached an OD600 of 0.6 and then harvested through centrifugation. The pellets were washed, resuspended in PBS, and mixed with HCT116 cells at a multiplicity of infection (MOI) of 100. After incubation under 5% CO2 at 37 °C for 1 h, the extracellular bacteria were eliminated by treating the cells with gentamicin (50 μg/mL) for 2 h under the same conditions. The cells were rinsed with PBS, and intracellular bacteria were released using 0.25% sodium deoxycholate. The number of intracellular bacteria was determined by measuring viable counts on LB agar plates.
In vitro cell viability test
The bacterial cells in the cell invasion assay were cultured overnight at 37 °C. HCT116 cells were initially plated at a density of 1 × 104 cells per well and incubated for 24 h. Before the experiment, the growth medium (DMEM) supplemented with antibiotics was replaced with antibiotic-free DMEM. The HCT116 cells were then infected with bacterial cells at a MOI of 50 and incubated for 1, 2, 3, and 4 h. After infection, the HCT116 cells were washed with antibiotic-free DMEM containing 50 μg/mL gentamicin and PBS. The cells were then detached using trypsin-EDTA, and the viable and dead cell counts were determined using trypan blue staining.
Animal studies
Mouse feeding and experimental procedures were conducted as previously described [40]. Specific pathogen-free 6-week-old female BALB/c mice (n = 70 for the mouse survival assay, n = 30 for the organ invasion assay). The mice were intraperitoneally infected with 104 colony-forming units (CFUs) of S. Typhimurium strains in 100 μL of PBS and euthanized after 3 d. The spleen, liver, and mesenteric lymph nodes were aseptically removed, and viable intracellular bacterial cells were quantified as previously described [41]. The log-rank (Mantel-Cox) test was used to analyze the difference in survival rates [42].
Statistical analysis
Individual data points are represented by white circles, with error bars indicating the standard deviation (SD) from three independent repetitions, otherwise specified. Data were analyzed using an unpaired two-tailed t-test. Significance levels were defined as follows: NS (not significant), *p < 0.05, **p < 0.01, ***p < 0.001.
Results
The bipA deletion in S. Typhimurium leads to defective growth at low temperatures
Although the bipA gene is not essential for optimal growth conditions, previous studies have indicated its critical role in the cold-shock response, enabling cells to adapt to low temperatures in E. coli [27]. Thus, the bipA gene in the three S. Typhimurium strains—SL1344, 14028, and LT2—were deleted to determine whether BipA was required for the normal growth of Salmonella strains at low temperatures. As shown in Fig 1A, the bipA-deleted E. coli (MG1655ΔbipA) and S. Typhimurium strains exhibited growth defects at low temperatures compared to their wild-type counterparts. Notably, the bipA-deleted S. Typhimurium LT2 (LT2ΔbipA) showed the most significant defects among the S. Typhimurium strains at low temperatures (Fig 1A). To quantitatively compare the growth between the strains, growth was monitored by measuring the optical density, consistently showing defective growth of the bipA-deleted cells (Fig 1B and 1C). However, when grown at 37 °C, both wild-type and bipA-deleted strains exhibited similar growth in liquid and solid media, without noticeable differences (S1A-S1C Fig).
The growth of wild-type and bipA-deleted strains of E. coli (MG1655) and S. Typhimurium (SL1344, 14028, and LT2) was evaluated at 20 °C. (A) Overnight cultures were diluted to an OD600 of 0.2 in the same medium and further diluted from 10−1 to 10−5-fold dilutions. Three microliters of the diluted samples were spotted on LB agar plates and incubated at 20 °C. (B, C) Cells grown at 37 °C to an OD600 of 0.5 were inoculated into fresh LB medium and incubated at 20 °C for 24 h. Error bars represent the SD. (D) Upregulation of bipA transcription after cold shock. The transcript levels of bipA and cspA were determined using qRT-PCR for the wild-type E. coli and S. Typhimurium strains at 0, 0.5, 1, and 2 h after cold shock. The relative quantities of the transcripts were calculated against the mean of the reference gene (rrsA) and normalized by those obtained for the cells at 0 h. (E) Western blot analyses and (F) densitometry-based quantification of the BipA, CspA, and OmpA protein levels after cold shock were conducted. Samples obtained at the indicated time points were subjected to western blot analysis, and representative images from triplicate experiments are shown. Band intensities were measured using ImageJ software. M, PageRuler-prestained protein ladder (Thermo Fisher Scientific).
To further investigate BipA-dependent growth at low temperatures, we conducted qRT-PCR and western blot analyses to assess the transcriptional and translational changes of BipA following cold shock. As shown in Fig 1D, exposure to cold shock triggered a 4.57- to 8.67-fold increase in bipA mRNA levels within 0.5 h in both the E. coli and S. Typhimurium strains. Additionally, we assessed cspA mRNA levels as a positive control, revealing significant increases of 24.02-fold in E. coli and 65.30- to 99.85-fold in S. Typhimurium strains after cold shock. These increases subsequently returned to the baseline due to auto-repression, consistent with previous findings [43]. The increased expression level of bipA transcripts was further confirmed by western blot analysis, which revealed increased BipA expression at the protein level (Fig 1E). Densitometric analysis showed a 2.00- to 2.95-fold increase in BipA protein levels across all strains (Fig 1F). As previously demonstrated [27], the cold-inducibility of bipA expression likely involves cAMP-CRP regulation, given the presence of a consensus CRP-binding site upstream of the SebipA promoter (S1D Fig). These findings indicate that while BipA expression is moderately induced at low temperatures compared to CspA, its expression is crucial for the growth of both E. coli and S. Typhimurium under low-temperature conditions and may be regulated by a conserved mechanism in both strains.
The GTPase activity of BipA is essential for normal growth at low temperatures
Previous studies on E. coli BipA revealed that GTPase activity is essential for normal growth and ribosome association at low temperatures [27]. To investigate the GTPase-dependent function of S. Typhimurium BipA (SeBipA), we introduced an inhibitory mutation (Asn128 to Asp) in SeBipA, which disrupts hydrogen bond formation with the 2-amino group of the guanine ring [1]. The bipA gene sequences were identical across the three S. Typhimurium strains—SL1344, 14028, and LT2. Moreover, primary sequence alignment showed that E. coli and S. Typhimurium BipA proteins share 96% identity and 99% similarity, with conservation of the Asn128 residue in both proteins (S2A Fig). We constructed the mutant clones pET28SeBipAN128D and pACYC177SeBipAN128D using pET28SeBipA and pACYC177SeBipA as templates. To compare the GTPase activity of the wild-type and mutant BipA, His6-tagged BipA proteins from E. coli and S. Typhimurium were overexpressed and purified, as described in the Materials and Methods section. In vitro GTPase assays using these purified proteins revealed that the mutant BipA exhibited approximately 5-fold lower GTPase activities than the wild-type BipA (Fig 2A).
(A) GTPase assays were conducted using the wild-type and mutant BipA proteins of E. coli and S. Typhimurium. The GTP hydrolysis reaction was conducted as described in the Materials and Methods section. (B) Cross-complementation of the bipA-deleted strains restored the growth defect. The bipA-deleted E. coli and S. Typhimurium strains were transformed with the plasmid as shown. The transformants were spotted and grown at 20 °C, as described in Fig 1. (C–F) The growth of transformants at 20 °C was assessed as outlined in Fig 1.
To determine whether the expression of the mutant BipA could restore the cold sensitivity of MG1655ΔbipA and S. Typhimurium cells with bipA deletions were transformed with pACYC177, pACYC177(Se)BipA, or pACYC177(Se)BipAN128D. The transformants were cultured in solid or liquid LB media containing ampicillin at 37 °C or 20 °C. At 20 °C, E. coli and S. Typhimurium cells harboring the mutant plasmids pACYC177BipAN128D or pACYC177SeBipAN128D failed to restore the cold sensitivity, while the cells complemented with the wild-type did (Fig 2B–2F). Western blot analysis showed no significant difference in the expression levels of the wild-type and mutant BipA proteins across the four mutant strains (S2B Fig). This indicates that the inhibitory effect of SeBipAN128D in 14028ΔbipA and its promotive effect in LT2ΔbipA strains were not attributable to differences in expression levels. At 37 °C, all transformants exhibited normal growth without defects (S2C Fig). These findings indicate that the GTPase activity of BipA is essential for its function and that E. coli BipA can replace SeBipA, and vice versa. Furthermore, the growth-defective phenotype resulting from bipA deletion was not due to a polar effect.
The critical role of BipA in the ribosome assembly of S. Typhimurium
In E. coli, bipA-deleted cells accumulate abnormal 50S ribosomal subunits at low temperatures. Wild-type BipA associates with both 70S ribosomes and 50S ribosomal subunits in a GTP-dependent manner, whereas BipAN128D does not exhibit this association [27]. To investigate the interaction between SeBipA and the 50S ribosome and its implications, we conducted polysome and subunit profiling analyses using sucrose gradient sedimentation. Cleared cell lysates were prepared and subjected to ultracentrifugation, as described in the Materials and Methods section. Polysome analyses of cells grown at 37 °C revealed that all strains had normal polysomes and ribosomal subunits, with or without BipA. However, in MG1655ΔbipA cells cultured at 20 °C, the peak of the 50S subunit collapsed, while the peak of the 30S increased (Fig 3A). Consistent with our previous report, abnormal particles (~44S, indicated by the red arrowhead in Fig 3A) accumulated between the 50S and 30S peaks [27]. Among S. Typhimurium strains SL1344, 14028, and LT2, S. Typhimurium 14028ΔbipA exhibited a pattern most similar to MG1655ΔbipA (indicated by the blue arrowhead in Fig 3A). Conversely, the SL1344ΔbipA and LT2ΔbipA strains did not show an abnormal 50S ribosomal peak; instead, there was a significant increase in the 30S peak compared to the wild-type. Ribosomal subunits analyzed under lower sucrose gradient concentrations revealed highly superimposable subunit profiles between the wild-type and mutant strains at 37 °C (Fig 3B). However, similar to the polysome profiling results, a consistent abnormality was observed in the 50S subunits, either through the accumulation of ~44S particles or a diminished quantity. Notably, bipA-deleted S. Typhimurium strains exhibited a reduced abundance of intact 50S subunits at 20 °C compared to the MG1655ΔbipA strain. Unlike E. coli MG1655, these S. Typhimurium strains possess rRNA genes that contain one or two intervening sequences in the 23S rRNA portion (S3 Fig). These intervening sequences are excised by RNase III during maturation, resulting in ribosomes with two or three cleaved segments of 23S rRNA [44]. While cleavage within the 23S rRNA does not affect ribosomal function, it accelerates ribosomal degradation by creating more targets for ribonucleases [45]. Consequently, in the absence of BipA, nascent 50S ribosomes likely undergo structural instability, making them more susceptible to degradation. This ultimately results in significantly low levels of intact 50S ribosomal subunits in bipA-deleted S. Typhimurium strains at low temperatures (Fig 3B).
(A) Polysome profiles and (B) subunit profiles of wild-type and bipA-deleted strains of E. coli and S. Typhimurium. Wild-type and mutant strains were inoculated into LB or LB broth containing Kan (50 μg/mL). Cells were grown at 37 °C or 20 °C until the OD600 reached 0.5 and then harvested by centrifugation at 3,000 rpm for 10 min. Cleared cell lysates were prepared, and sucrose gradient sedimentation was performed as described in the Materials and Methods section. The red and blue arrowheads indicate abnormal 50S ribosome particles in the E. coli and S. Typhimurium strains.
Previous studies have shown that His6-tagged SeBipA binds to the 70S ribosome in the presence of GTP. Furthermore, under stringent response conditions or the presence of ppGpp, His6SeBipA exhibited an increased affinity for the 30S ribosomal subunit [46]. Therefore, we further investigated how SeBipA modulates its ribosome binding mode, depending on the nucleotides present. To precisely determine the ribosome binding mode of SeBipA, we mixed wild-type cells cultured at 37 °C or 20 °C with ppGpp (a stringent response alarmone), GDP, and GMPPNP (a nonhydrolyzable analog of GTP) at a final concentration of 100 μM. Subsequently, we performed sucrose gradient sedimentation and western blotting to locate the endogenously expressed SeBipA proteins in the polysome profiling fractions. Both BipA and SeBipA bound to 70S ribosomes and 50S ribosomal subunits at both temperatures in the presence of GMPPNP, indicating a temperature-independent association (Fig 4A and 4B). For the 30S ribosome association, both BipA and SeBipA were detected in the trail fraction near the soluble portion, indicating that this association may not be GMPPNP-specific and requires further investigation. Notably, adding GDP or ppGpp did not enhance the interaction between E. coli BipA and ribosomal fractions at either temperature. Conversely, SeBipA from pathogenic S. Typhimurium SL1344 and 14028 appeared to cosediment with polysomes and 70S ribosomes in the presence of ppGpp and GDP at low temperatures. This suggests that SeBipA may have a different binding mechanism than E. coli BipA. Nevertheless, our findings indicate that the specific interaction of BipA proteins with 70S and 50S ribosomes is GMPPNP-dependent. Although BipA binds to 50S ribosomal subunits at 37 °C, ribosome biogenesis occurs without defects, indicating that BipA is dispensable.
Wild-type E. coli (MG1655) and S. Typhimurium (SL1344, 14028, and LT2) strains were cultured at (A) 37 °C or (B) 20 °C, as outlined in Fig 3. Cell pellets were resuspended in buffer BP containing ppGpp, GDP, or GMPPNP, each at a final concentration of 100 μM. Sucrose gradient sedimentation was conducted as described in the Materials and Methods section. Fractions from polysome to free RNAs in polysome profiling were subjected to western blot analysis using anti-BipA antibodies. The control “C” indicates wild-type MG1655 cells harboring pACYC177(Se)BipA. M, molecular marker.
Increased bile salt sensitivity and LPS synthesis defects in bipA-deleted cells at low temperatures
Exposure to BPI with LPS decomposition activity results in an approximately 7-fold upregulation of bipA expression in S. Typhimurium [5], and its deletion makes bacterial cells more sensitive to BPI [26]. This suggests that BipA may influence the LPS structure or composition in response to BPI. Our previous findings further support this, showing that MG1655ΔbipA cells accumulate an abnormal lipid A-core oligosaccharide (lipooligosaccharides, LOS) at low temperatures [28]. This LPS perturbation, whether due to bipA deletion or other outer membrane damage, is known to signal through the Rcs pathway [47]. The outer membrane protein RcsF initiates this process by detecting stress signals, which are then transmitted through the inner membrane proteins RcsC-RcsD to the response regulator RcsB through a phosphorelay system [48]. This signaling cascade ultimately regulates the expression of various genes. The phosphorylated RcsB can form a heterodimer with RcsA or a homodimer with itself. The former activates the transcription of the cps gene cluster for colanic acid synthesis and represses the expression of flhDC, which encodes the primary regulator of flagella biosynthesis genes [49,50]. Conversely, the latter dimer stimulates the expression of rprA, which encodes a negative regulatory sRNA of curli and cellulose production [51]. Therefore, the Rcs two-component regulatory system is a key signaling pathway for protecting cells from environmental challenges and intrinsic sources of envelope stress.
Therefore, we examined how bipA deletion in each strain affects cell surface structures, such as LPS, capsule, biofilm, and flagella, at both temperatures. To determine whether BipA of S. Typhimurium strains is involved in LPS biosynthesis, we streaked wild-type MG1655 and the three S. Typhimurium strains (SL1344, 14028, and LT2), along with their bipA-deleted strains on solid media containing bile salt at concentrations ranging from 20 to 27 g/L. The plates were incubated at either 37 °C or 20 °C. Bacterial strains with LPS defects are known to be more susceptible to bile salts [52,53]; thus, we indirectly assessed the LPS status by examining bile salt sensitivity. As shown in Fig 5A, while BipA is dispensable for ribosome synthesis at 37 °C, the colony formation of MG1655ΔbipA cells was inhibited at a bile salt concentration of 25 g/L at 37 °C. Conversely, wild-type cells failed to grow at a bile salt concentration of 27 g/L. Note that the concentration of bile salts in human bile varies depending on the source and can reach to 3.5% in the duodenum and ≤0.005% in the gastrointestinal tract [54]. Among the S. Typhimurium strains, LT2 was more sensitive to bile salts than the other two strains, and the bipA-deleted cells were more sensitive than the wild-type cells. At 20 °C, the four bipA-deleted strains lost nearly all growth ability, failing to form colonies at a bile salt concentration of 22 g/L, while the wild-type cells continued to grow at bile salt concentrations ranging from 24 to 25 g/L. These results indicate that the deletion of bipA in both E. coli and S. Typhimurium significantly influences LPS biosynthesis at 20 °C, and, to a minimal extent, at 37 °C.
(A) Bile salt-sensitive phenotypes of (a) MG1655, (b) MG1655ΔbipA, (c) SL1344, (d) SL1344ΔbipA, (e) 14028, (f) 14028ΔbipA, (g) LT2, and (h) LT2ΔbipA at 37 °C or 20 °C. Overnight cultures were diluted to an OD600 of 0.2 in the same medium and further diluted 10−2-fold. Two and a half microliters of the diluted samples were spread onto LB agar plates containing a gradient concentration of bile salt and incubated overnight at 37 °C or 20 °C for 1 week. (B) Schematic LPS structure of E. coli K-12. The action sites of WaaF, WaaG, WaaQ, and PagP were indicated. Kdo, 3-deoxy-d-manno-oct-2-ulosonic acid. (C) LPS profile analysis of the E. coli and S. Typhimurium strains. LPS was extracted and analyzed using SDS-PAGE followed by silver staining, as described in the Materials and Methods section. M, molecular marker. (D) LPS profile analysis of colanic acid-overproducing strains. LPS samples were extracted from cells grown on LB agar plates as in (C). (E) Lipid A-core oligosaccharide profile analysis. Twenty-five-fold diluted LPS samples were separated by tricine-SDS-PAGE followed by silver staining.
To further investigate LPS abnormalities in bipA-deleted cells, we extracted LPS from the wild-type and bipA-deleted strains grown on LB agar plates at 37 °C or 20 °C. The LT2ΔbipA cells did not adequately grow at 20 °C, resulting in insufficient biomass for LPS extraction. As controls, we extracted LPS from three E. coli mutants with deletions in the waaF, waaG, or waaQ genes, which are responsible for adding the second heptose, the first glucose, and the side-branch third heptose, respectively (Fig 5B) [55–57]. The LPS samples were then analyzed using SDS-PAGE or tricine-SDS-PAGE followed by silver staining. In the waaF- and waaG-deleted strains, which exhibit severe defects in the LPS core oligosaccharide, mature LPS was not observed, whereas the waaQ mutant showed a normal LPS ladder (Fig 5C). As shown in S4A Fig, no discernible differences were observed between the wild-type and bipA-deleted strains at 37 °C. However, at 20 °C, a significant reduction in LPS levels was observed in the bipA-deleted E. coli and S. Typhimurium strains. Notably, additional LPS ladder bands were observed between the repeated O-antigen units in the MG1655ΔbipA strain (indicated by the red arrowheads in Fig 5C). These bands were absent in the MG1655 strain, suggesting potential alterations in the LOS moiety or variations in the size of the O-antigen unit. It has been reported that when colanic acid, also referred to as the M-antigen (mucoid antigen), is overproduced, the M-antigen can replace the O-antigen on the LPS, resulting in the formation of M-LPS [58,59]. To investigate this possibility, LPS was extracted from the MG1655ΔrcsA, MG1655ΔbipAΔrcsA, and MG1655Δlon strains and compared with that from MG1655ΔbipA. Lon is a protease that degrades RcsA, the activator of the cps cluster, and in its absence, colanic acid was overproduced at 20 °C (S4B Fig) [49,60]. As shown in Fig 5D, the aberrant LPS ladder was observed in both MG1655ΔbipA and MG1655Δlon strains, while no such ladder was observed in the rcsA-deleted cells, suggesting the formation of M-LPS. In the case of S. Typhimurium strains, the LPS ladders were more closely spaced due to the smaller size of the O-antigen units compared to those of E. coli [61].
To further investigate potential perturbations in the LOS moiety, we analyzed the LPS extracts using tricine-SDS-PAGE. An examination of the LOS from the three control mutants showed that the absence of a branched heptose in the inner core did not result in a substantial size difference compared to the LOS from the wild-type cells (band “f” in Fig 5E). However, the absence of five sugars due to the waaG deletion significantly reduced LOS size (band “e” in Fig 5E), and the LOS in the waaF mutant was barely detectable (Fig 5E).
At 37 °C, both MG1655 and MG1655ΔbipA cells exhibited two major LOS forms (bands “a” and “b” in Fig 5E), and they are likely to be a hexa-acylated form based on the position of the hepta-acylated form by PagP (band “g”) [62]. Likewise, no significant LOS alterations were observed between the wild-type and mutant S. Typhimurium strains. Notably, LOS + 1 O-antigen unit was predominant in S. Typhimurium compared to E. coli (band “c” in Fig 5E). However, at 20 °C, LOS forms without O-antigen unit became prevailing, and their overall amount was diminished in the mutant S. Typhimurium.
More interestingly, the larger LOS (band “a”) disappeared in MG1655ΔbipA cells grown at 20 °C, along with a concomitant reduction in the smaller LOS (band “b”). This overall decrease in LOS levels likely explains the observed reduction in total LPS. Unexpectedly, a band slightly larger than the LOS + 1 O-antigen unit of S. Typhimurium appeared (band “d”), which was also observed in MG1655Δlon cells. Considering that the molecular weight of M-antigen (1,106.0 Da) is larger than that of O-antigen in S. Typhimurium (672.6 Da), the band “d” likely represents LOS + 1 M-antigen unit [63,64]. Furthermore, it is also possible that the missing band “a” in MG1655ΔbipA cells at 20 °C was used to synthesize the M-LPS, consistent with the result in Fig 5D.
Taken together, at 20 °C, the bipA-deleted S. Typhimurium produced less LOS, whereas bipA deletion in E. coli resulted in significant changes in LOS and LPS structures, consequently impairing outer membrane integrity.
The effect of bipA deletion on the capsule and biofilm
To assess changes in capsule synthesis, diluted cell cultures were either spread on LB agar plates or spotted onto MacConkey agar plates. Following incubation at 37 °C or 20 °C, we examined the morphologies of the colonies and macrocolonies. On LB agar plates, MG1655ΔbipA colonies merged with neighboring colonies, resulting in a mucoid appearance at 20 °C—a phenomenon not observed in any other strains at either temperature (S5A Fig). Similarly, only MG1655ΔbipA cells exhibited a transparent halo around the macrocolony on MacConkey agar plates at 20 °C (S5B Fig). Although the regulatory mechanism of the cps cluster through the Rcs pathway is conserved in S. Typhimurium [65], the bipA-deleted S. Typhimurium strains, unlike MG1655ΔbipA cells, did not exhibit capsule overproduction at either temperature under the specified growth conditions.
The Rcs signaling pathway enhances the production of the small regulatory RNA RprA [66]. RprA represses the translation of csgD, which encodes a positive regulator of curli synthesis and cellulose production, resulting in decreased biofilm formation [67,68]. Additionally, it has been reported that the overproduction of colanic acid inhibits biofilm formation by reducing surface adhesion [69]. Therefore, we compared the biofilm formation abilities of the wild-type and bipA-deleted strains. Biofilm formation on a glass tube, representing a hydrophilic surface, was assessed using the crystal violet staining method as described in the Materials and Methods section. As shown in Fig 6A, none of the wild-type or mutant strains formed substantial biofilm at 37 °C, whereas biofilm formation in the wild-type E. coli strain increased approximately 5.31-fold at 20 °C compared to 37 °C. Notably, MG1655ΔbipA cells showed a significant reduction in biofilm formation (0.09-fold) (Fig 6B). The SL1344ΔbipA and 14028ΔbipA strains exhibited a 0.44-fold and 0.46-fold decrease in biofilm-forming ability, respectively, while no significant difference was observed in the LT2 strain (Fig 6B).
Biofilm formation of wild-type E. coli (MG1655) and S. Typhimurium (SL1344, 14028, and LT2), along with their bipA-deleted strains at 37 °C or 20 °C. Cell culturing and crystal violet staining were performed as described in the Materials and Methods section. The cultures in glass tubes were incubated at (A) 37 °C or (B) 20 °C. The cultures in Eppendorf tubes were incubated at (C) 37 °C or (D) 20 °C.
Given that S. Typhimurium biofilms preferentially attach to hydrophobic surfaces [70], we conducted a parallel experiment using polystyrene tubes with hydrophobic surfaces. Consistent with the results observed with glass tubes, minimal biofilm formation was observed at 37 °C in both the wild-type and bipA-deleted strains, with no significant difference in the biofilm-forming ability between them (Fig 6C). Note that conventional laboratory conditions for Salmonella biofilm formation involve low osmolarity (low- salt LB) and a temperature of 28 °C, which differ from our experimental conditions. These conditions promote RpoS-dependent activation of biofilm-related genes such as csgD, adrA, and agfA [71,72]. However, at 20 °C, the wild-type 14028 cells formed a substantial amount of biofilm, which significantly reduced in the absence of BipA (0.03-fold) (Fig 6D).
These findings indicate that the distinct cell surface properties of E. coli and S. Typhimurium result in opposite patterns of bacterial attachment depending on whether the surface is hydrophobic or hydrophilic. Nevertheless, bipA deletion resulted in reduced biofilm formation in both E. coli and S. Typhimurium 14028.
Reduced motility of bipA-deleted strains with fewer flagella
Given that the Rcs pathway inhibits the expression of flhDC, the master regulator of flagella biosynthesis, we investigated how bipA deletion affects motility and flagella synthesis in S. Typhimurium strains.
To assess swimming motility, cells in the early exponential phase were inoculated onto 0.3% agar plates and incubated at 37 °C or 20 °C. As shown in Fig 7A and 7B, the wild-type pathogenic S. The Typhimurium SL1344 and 14028 strains exhibited significantly faster motility at 37 °C than the E. coli MG1655 and S. Typhimurium LT2 strains. In all four strains, bipA-deleted strains resulted in a moderate reduction in motility (0.72–0.76-fold) (Fig 7B). While the overall swimming motility decreased at 20 °C compared to 37 °C, the two pathogenic strains maintained higher migration speeds than their nonpathogenic counterparts (Fig 7A and 7C). Notably, the reduction in the migration speed of the bipA-deleted strains, particularly, SL1344ΔbipA and 14028ΔbipA, became more pronounced at 20 °C (0.16–0.46 fold) compared to 37 °C (Fig 7C).
(A) Swimming motility assays of wild-type E. coli (MG1655) and S. Typhimurium (SL1344, 14028, and LT2), along with their bipA-deleted strains. Overnight cultures were diluted 10−2-fold in fresh medium and incubated until the OD600 reached 0.5. The cultures were then concentrated 10-fold and injected onto LB agar plates, followed by incubation at 37 °C or 20 °C. (B and C) The migration speeds of the wild-type and bipA-deleted strains were measured at (B) 37 °C or (C) 20 °C by recording the distance traveled per incubation time. Error bars represent the SD from four independent replicates and each data point is indicated by a white circle. (D) Fewer flagella were observed in bipA-deleted pathogenic S. Typhimurium strains at low temperatures. Overnight cultures were diluted 10−2-fold in fresh medium and incubated at 37 °C for 1 d or 20 °C for 3 d. The cells were harvested at 3,000 rpm to minimize flagella loss, deposited onto carbon-copper grids, and negatively stained. TEM was used to visualize the flagella of the wild-type and bipA-deletion strains. Flagella count was quantified from 10 individual cells grown at (E) 37 °C or (F) 20 °C (F), represented in the bar graphs.
This flagella-mediated motility was further confirmed through a swarming assay. As shown in S6A Fig, neither the wild-type nor the mutant E. coli strains exhibited motility at either temperature. Consistent with the swimming motility results, the two pathogenic S. Typhimurium strains exhibited higher motility than the attenuated LT2 strain. However, the mutant S. Typhimurium strains completely lost motility at 20 °C, highlighting the role of BipA in flagella function.
To correlate the reduced mobility with the diminished expression of flagella, the flagellated states of the respective strains were examined using TEM. Cells cultured in the stationary phase at 37 °C or 20 °C were stained with phosphotungstic acid on a carbon-copper grid and analyzed using TEM, as described in the Materials and Methods section. As shown in Fig 7D and 7E, the SL1344 and 14028 strains exhibited approximately 6–8 flagella on their surfaces at 37 °C, while LT2 had fewer (approximately 4), and the MG1655 had the least (only 1 flagellum). At 20 °C, the flagella counts for the four strains followed the same pattern; however, the mutant strains produced half the number of flagella compared to the wild-type strains (Fig 7F). At 37 °C, there was no significant difference in flagella count between the wild-type and mutant strains (Fig 7D and 7E). This finding was further confirmed by western blot analysis targeting the deflagellated FliC proteins, as outlined in S1 Supporting Information. In the MG1655 and LT2 strains, the FliC protein was undetectable in the deflagellated soluble fraction at both temperatures, likely due to the lower flagella count. Conversely, a similar amount of deflagellated FliC protein was detected in both wild-type and mutant SL1344 and 14028 strains at 37 °C. However, at 20 °C, these FliC proteins diminished in the SL1344ΔbipA and 14028ΔbipA strains (S6B Fig).
These phenotypes were further validated by analyzing the expression levels of several key flagella-related genes (flgM, fliE, fliK, fliM, fliS, and fliZ) in all four strains at both temperatures using qRT-PCR. The deletion of bipA significantly reduced gene expression in all strains, particularly at 20 °C (S7 Fig). Given the phenotypic effects of bipA deletion in S. Typhimurium, it is likely that the outer membrane damage from reduced LPS in bipA-deleted strains results in reduced flagella production, thereby diminishing motility.
Effect of bipA deletion on the pathogenicity of S. Typhimurium SL1344 and 14028 strains
In many pathogens, flagella-related phenotypes are closely linked to infection processes, such as host cell invasion and virulence in mouse models [40,73–75]. Our findings revealed that bipA deletion affected LPS, biofilm, and flagella formation (Figs 5, 6, and 7). To determine whether bipA deletion affects the pathogenicity of S. Typhimurium, we examined its influence on host cell invasion. HCT116 cells were infected with wild-type (WT; + pACYC177), bipA-deleted (ΔbipA; + pACYC177), and bipA-complemented [Complemented; + pACYC177(Se)BipA] strains cultured at 37 °C or 20 °C. After infection, gentamicin was used to eliminate bacterial cells that did not infect the host cells. Bacterial invasion was then assessed by measuring the CFUs. As anticipated, neither the E. coli nor S. Typhimurium LT2 strains showed any invasion of HCT116 cells at either temperature. Conversely, the bipA-deleted pathogenic S. Typhimurium SL1344 and 14028 strains produced approximately 45% fewer CFUs in HCT116 cells than the wild-type and complemented strains at both temperatures (Fig 8A and 8B). The invasion assay indicated reduced pathogenicity, a finding further supported by an in vitro cytotoxicity assay, in which dead and viable cells were differentiated using trypan blue staining. Consistent with the invasion assay results, E. coli and S. Typhimurium LT2 strains did not significantly affect host cell viability (Fig 8C and 8F). However, the number of viable HCT116 cells increased when infected with the S. Typhimurium SL1344ΔbipA and 14028ΔbipA strains compared to those infected with wild-type or complemented cells (Fig 8D and 8E).
Effect of bipA expression on epithelial cell invasion by E. coli (MG1655) and S. Typhimurium strains (SL1344, 14028, and LT2). Epithelial cells (HCT116) were infected with the wild-type and bipA-deleted strains of E. coli and S. Typhimurium at an MOI of 100, and the number of intracellular bacteria was counted as CFUs at (A) 37 °C or (B) 20 °C. Error bars represent the SD from nine independent replicates and each data point is indicated by a white circle. (C–F) Effect of bipA expression on E. coli (MG1655) and S. Typhimurium strains (SL1344, 14028, and LT2) epithelial cell viability. HCT116 cells were infected with E. coli and S. Typhimurium cultures from (B) at an MOI of 100. Cell viability was determined by counting viable cells using the trypan blue staining assay at the indicated time points. (G) and (H) Survival rates and durations of BALB/c mice infected with S. Typhimurium (14028 and LT2), bipA-deleted strains, and PBS as a control (uninfected) were monitored for 10 d (n = 10 per group). (I) Colonization assay in BALB/c mice (n = 5 per group) infected with S. Typhimurium 14028 and bipA-deleted strains, with bacterial CFUs counted in the liver, mesenteric lymph nodes, and spleen. Data were expressed as the mean ± standard error of the mean from at five independent experiments, with individual data points indicated by white circles.
Next, we investigated the role of BipA-related S. Typhimurium pathogenicity using a mouse model infected with S. Typhimurium 14028 and LT2 strains, which exhibited distinct differences in host cell invasion. Mice infected with either the wild-type or complemented S. Typhimurium 14028 strains succumbed to infection within 5 and 8 d, respectively. Conversely, 40% of the mice infected with the 14028ΔbipA strain survived 8 d postinfection (Fig 8G). Although the wild-type LT2 strain exhibited a slight mortality rate, bipA deletion did not influence the pathogenicity of the S. Typhimurium LT2 strain (Fig 8H). We also assessed the bacterial load in the liver, mesenteric lymph nodes, and spleen of the infected mice. The bacterial load in the 14028ΔbipA strain was significantly lower than that of the wild-type S. Typhimurium 14028 strain across all three organs. Notably, this colonization defect was restored to the wild-type levels upon complementation with bipA (Complemented; Fig 8I). These findings clearly demonstrate the role of BipA in regulating the pathogenicity of the Salmonella Typhimurium SL1344 and 14028 strains.
Discussion
In this study, we comprehensively assessed the roles of BipA in ribosome assembly and virulence factors across two different bacterial species, E. coli and S. Typhimurium, evaluating its impact on pathogenicity of the strain S. Typhimurium 14028 strain. In our previous study, we demonstrated that BipA exhibits protein-chaperone activity by refolding denatured proteins. We also reported that the incorporation of the ribosomal protein L6 into the 50S ribosomal subunit in E. coli was hindered at low temperatures in the absence of BipA, indicating the role of BipA as a 50S ribosome assembly factor [27]. Our results revealed that the GTP-dependent association of BipA with the 50S ribosome is crucial for ribosome assembly in S. Typhimurium at low temperatures. Deletion of bipA reduced the levels of the 50S ribosomal subunit and resulted in the accumulation of abnormal 50S ribosomal particles at 20 °C (Fig 3). This finding indicates that the role of BipA in ribosome assembly is conserved across various bacterial species.
Notably, while BipA is not essential for ribosome assembly at 37 °C, bipA-deleted E. coli and S. Typhimurium cells exhibited increased sensitivity to higher concentrations of bile salts at 37 °C. This sensitivity was also evident at lower concentrations of bile salts at low temperatures (Fig 5). Additionally, the bipA mutants were more susceptible to the cationic antimicrobial peptide polymyxin B, which targets the outer membrane, at low temperatures compared to wild-type cells (S8 Fig).
As BipA co-localizes with the 70S ribosome in the presence of GMPPNP, it may play a role in the temperature-dependent formation of specialized ribosomes that preferentially translate mRNAs involved in LPS synthesis or modification. Such specialized ribosomes can form through various mechanisms, including rRNA sequence variations, rRNA modification, r-protein modifications, r-protein paralogs, and ribosome-associated proteins [76]. The association of BipA with the ribosome may induce conformational changes in rRNAs and r-proteins, resulting in the formation of ribosomes with altered structural and functional properties. Alternatively, ribosome-associated BipA may directly recruit specific mRNAs to the ribosome, similar to the pro-apoptotic protein Reaper in Drosophila melanogaster [77] and the RACK1 protein in Saccharomyces cerevisiae [78–80], thereby promoting the specific translation of LPS-related mRNAs at low temperatures.
LPS, a glycolipid present on the cell surface of Gram-negative bacteria, consists of a conserved lipid A anchor, a core oligosaccharide, and an O-antigen. The synthesis of LPS begins at the cytoplasmic side of the inner membrane through the Raetz pathway, which involves a series of conserved enzymatic steps. This pathway initiates with UDP-N-acetylglucosamine, which is converted into Lipid IVA, then to Kdo2-lipid A, eventually forming LOS that includes core sugars. The LOS is flipped to the periplasmic side by the MsbA2 flippase. Subsequently, the O-antigen is added by the O-antigen ligase. Finally, the Lpt complex transports the complete LPS molecule to the outer leaflet of the outer membrane [81]. During or after the Raetz pathway, LPS can be further modified by various enzymes that typically alter the phosphate groups, the sugar backbone of lipid A, the core oligosaccharide, and the acyl chains.
In the MG1655ΔbipA strain at 20 °C, an M-LPS was observed, along with the absence of the larger LOS band (Fig 5C and 5E). Notably, in the waaG-deleted strain, only one LOS band was detected, suggesting that the missing band represents a LOS variant with a modified outer core. MG1655 is reported to predominantly contain glycoform I core oligosaccharide [82,83], which likely corresponds to the smaller LOS band (band “b” in Fig 5E). Given that hepta-acylated LOS is 256.42 Da larger than hexa-acylated LOS, and the missing band is positioned between these two forms, it is predicted that this missing LOS species is approximately 120 Da larger than glycoform I. Based on its size and modification position, it is likely glycoform II, which contains an additional N-acetylglucosamine attached to the fourth heptose, compared to glycoform I [84]. However, the enzyme responsible for this modification has not yet been definitively identified. Therefore, it is essential to explore the functional relationship between LOS-synthesizing enzymes and BipA at low temperatures.
The Rcs phosphorelay system is activated in response to cell envelope stresses, including defects in LPS, leading to increased transcription of the cps gene cluster and rprA and repression of the flhDC operon [48,50,66,85]. The phenotypic traits observed in the bipA-deleted E. coli—such as defects in LPS biosynthesis, capsule overproduction, impaired biofilm formation, and reduced flagella-mediated motility—indicate the activation of the Rcs pathway. Typically, perturbations in the cell envelope are detected by the outer membrane lipoprotein RcsF. The activated RcsF then interacts with the inner membrane protein IgaA, disrupting the inhibitory action of IgaA on RcsD. Subsequently, the phosphorelay occurs from RcsC to RcsD to RcsB [85]. This signal transmission can be described as an RcsF-dependent pathway. In S4B Fig, the MG1655ΔbipAΔrcsF appear to overproduce capsular polysaccharides, whereas the MG1655ΔbipAΔrcsA cells do not. This indicates that the activation of the Rcs pathway in the MG1655ΔbipA cells may be mediated in an RcsF-independent manner, while capsule overproduction relies on the RcsA. One potential trigger for activating the Rcs pathway in an RcsF-independent manner is the overexpression of djlA, which encodes the cochaperone DnaJ homolog. Overexpressed DjlA, along with DnaK, may alter the conformation of the inner membrane components involved in the Rcs pathway, potentially disrupting the inhibitory action of IgaA on RcsD [86]. As mentioned earlier, BipA exhibits chaperone-like activity and plays a role in the LPS core biosynthesis, which primarily occurs at the inner leaflet of the cytoplasmic membrane. This leads us to hypothesize that BipA may influence a specific conformation involving the LPS core-synthesizing enzymes and/or Rcs components. Further investigations are required to identify which RcsF-independent signals arise in the bipA-deleted strain and how they contribute to signaling activation.
In S. Typhimurium, overexpression of RcsA, a defective mutant IgaA, or constitutively activated RcsC leads to colanic acid overproduction, even at 37 °C, resulting in mucoid colony morphology and loss of motility [65,87,88]. Although the Rcs pathway is highly conserved between E. coli and S. Typhimurium, our finding that capsule synthesis was unaffected by the bipA deletion in S. Typhimurium strains is particularly intriguing. Notably, there are differences in the lipid A-core structure between the two species [56,89], and as shown in Fig 5A, wild-type E. coli and S. Typhimurium strains exhibited maximum resistance to bile salts at concentrations of 26 g/L and 25 g/L at 37 °C and 20 °C, respectively. Additionally, E. coli cells were more sensitive to bile salts than S. Typhimurium cells at low temperatures. This indicates that the defective LPS in the bipA-deleted strain may cause varying degrees of damage to the cell envelope. Furthermore, the formation of the RcsB-RcsA heterodimer, essential for activating the cps gene cluster, is regulated at multiple levels. In normally growing cells, RcsA is limiting for cps gene transcription because H-NS represses rcsA transcription [90]. This repression is alleviated by a small, 85-nucleotide RNA molecule, DsrA [91]. Additionally, the quantity of RcsA protein is tightly regulated by the Lon protease [60]. Consequently, the LPS defects in the bipA-deleted Salmonella strains may not produce sufficient RcsB-RcsA heterodimers formation to induce sufficient cps gene cluster expression (S5C Fig). However, we do not rule out the possibility of alternative regulatory mechanisms affecting capsular polysaccharide production in S. Typhimurium.
The deletion of bipA did not uniformly induce mucoid colony morphology in all four strains, leading to varied impacts on biofilm formation among the four strains. Biofilm formation disruption due to bipA deletion was observed only in the MG1655 and 14028 strains, while in the SL1344 and LT2 strains (Fig 6). S. Typhimurium primarily produces cellulose as the main exopolysaccharide compound for biofilm matrix formation [92]. RpoS positively regulates mlrA, whose gene product subsequently upregulates CsgD, a key player in curli synthesis. CsgD also stimulates the expression of AdrA, which regulates cellulose production. However, in the SL1344 and LT2 strains, the transcription of mlrA and the levels of RpoS were either defective or nearly undetectable, respectively [34,93–95]. These discrepancies in the expression of positive regulators resulted in the cellulose-deficient phenotype, explaining the lack of inherent biofilm formation in the SL1344 and LT2 strains under the tested conditions.
Unlike the cps gene, which regulates colanic acid production through the activation of the RcsB-RcsA heterodimer, this complex inhibits flagellum synthesis by repressing flhDC within the Rcs pathway. Notably, bipA-deleted Salmonella strains exhibited only a moderate motility defect despite having a similar number of flagella at 37 °C. Additionally, qRT-PCR and western blot analyses revealed no significant changes in the expression levels of flagellum structural genes and FliC proteins at 37 °C (S6 and S7 Figs). However, ΔbipA cells exhibited an increase in the expression of type 1 fimbriae at 37 °C (S9 Fig), whose overexpression partially reduced flagella-mediated motility [96]. Furthermore, this motility defect at 37 °C may be attributed to the reduced ATP levels observed in all bipA-deleted strains (S10 Fig). Apparently, a significant motility defect was observed in the bipA-deleted SL1344 and 14028 strains, with a clear reduction in flagella count at 20 °C (Fig 7). In the MG1655 and LT2 strains, although the transcriptional expression of flagellum-related genes significantly reduced in the corresponding mutant strains compared to the wild-type strain at 20 °C (S7 Fig), this did not result in a reduced flagella count, as these strains inherently have fewer flagella. Nevertheless, the mutant MG1655 and LT2 strains showed impaired motility at 20 °C.
Overall, our findings indicate that BipA is involved not only in flagellum biosynthesis but also in the quality and functionality of the flagellum. Notably, EptC, a phosphoethanolamine transferase, plays a dual role in modifying the flagellum rod protein FlgG, and the lipid A substructure of Campylobacter jejuni LPS with a phosphoethanolamine residue. The eptC-deficient stains exhibited decreased motility and flagella formation [97,98]. This suggests that the defects in motility and flagella count observed in the bipA mutant may result from the combined effects of the Rcs pathway and the absence of interaction of BipA with LPS-modifying or -synthesizing enzymes. Further studies are needed to elucidate the exact molecular mechanisms by which BipA influences these processes and to clarify its potential role in coordinating flagella formation and LPS biosynthesis.
LPS, capsules, biofilms, and flagella are critical virulence determinants, along with T3SSs and the bacterial energy state, both of which are crucial for virulence. The pronounced defects in motility observed in the pathogenic S. Typhimurium SL1344 and 14028 strains at 37 °C highlight the essential role of BipA in pathogenicity (Fig 8). Moreover, our data indicate that deletion of bipA leads to a significant reduction in the expression of T3SS genes only at lower temperatures (S11 Fig), suggesting that the observed virulence attenuation is not due to T3SS repression. Specifically, decreased motility and ATP levels in bipA-deleted strains (Figs 7 and S10) suggest that BipA contributes to energy homeostasis, which may, in turn, impact virulence-related functions in host infection models (Fig 8).
In addition, BipA is important for bacterial resistance to host antimicrobial mechanisms. Given that Salmonella encounters ROS during infection, it must resist oxidative stress to survive. The bipA-deleted strains exhibited increased susceptibility to H2O2 at both 37 °C and 20 °C (S12 Fig), suggesting that BipA may contribute to bacterial defense mechanisms against oxidative damage in host cells.
Thus, our findings indicate that BipA functions as a key fitness factor, modulating surface-associated structures, including flagella, LPS, and T3SS, and plays a crucial role in maintaining bacterial energy balance and enhancing resistance to oxidative stress. Future research aimed at elucidating the mechanisms by which BipA regulates membrane integrity, responses to various stresses, and virulence will provide insights into bacterial adaptation and pathogenesis.
Supporting information
S1 Supporting Information. Supplementary Materials and Methods.
This section provides detailed descriptions of the experimental procedures, including cell morphology observation, the swarming motility assay, the deflagellation method, growth curve measurement, the Epsilometer test, and the ATP assay.
https://doi.org/10.1371/journal.ppat.1013047.s001
(DOCX)
S1 Fig. Normal growth of bipA-deleted E. coli and S. Typhimurium at 37 °C.
The growth of wild-type and bipA-deleted strains of E. coli and S. Typhimurium was assessed at 37 °C. (A) Overnight cultures were diluted and spotted, as described in Fig 1A, followed by incubation at 37 °C. (B and C) Cells grown at 37 °C to an OD600 of 0.5 were inoculated into fresh LB medium and further incubated at 37 °C for 12 h. Error bars indicate SD. (D) Conserved CRP-binding site upstream of the SebipA promoter. The upstream DNA sequences of E. coli and S. Typhimurium are aligned, with the consensus CRP-binding site highlighted in yellow, the −35 and −10 regions in blue, and the initiation codon GTG in green.
https://doi.org/10.1371/journal.ppat.1013047.s002
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S2 Fig. Sequence alignment of the BipA proteins.
(A) Sequence alignment of BipA from E. coli and S. Typhimurium was conducted using ClustalW and ESPript. The numbers indicate the corresponding residues, and the blue arrowhead indicates the Asn128 residue. (BipA, GenBank accession number: AAT48232; SeBipA, CAC14270). (B) Expression of wild-type and mutant BipA proteins in MG1655ΔbipA and mutant S. Typhimurium cells. The transformants shown in Fig 2B underwent Western blot analysis using an anti-BipA antibody. M: PageRuler Prestained Protein Ladder (Thermo Fisher Scientific). (C) Colony formation of cross-complemented bipA-deletion strains at 37 °C. The bipA-deleted E. coli and S. Typhimurium strains were transformed with the plasmid as shown. Transformants were spotted as described in Fig 2B, and the plates were incubated at 37 °C.
https://doi.org/10.1371/journal.ppat.1013047.s003
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S3 Fig. Multiple sequence alignment analysis of the intervening sequences (IVSs) in the 23S rRNA genes.
The nucleotide sequences of the 23S rRNA genes from E. coli MG1655 (A and B) and S. Typhimurium SL1344 (C and D), 14028 (E and F), and LT2 (G and H) were analyzed. The first IVSs in the helix 25 region (A, C, E, and G) and the second IVSs in the helix 45 region (B, D, F, and H) were aligned using Clustal Omega [99] and visualized using Jalview [100].
https://doi.org/10.1371/journal.ppat.1013047.s004
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S4 Fig. Analysis of LPS in E. coli and S. Typhimurium strains.
(A) LPS profile analysis of the E. coli and S. Typhimurium strains. LPS was extracted and analyzed, as described in Fig 5. (B) Colony morphology of bipA, rcsF, rcsA, and lon mutants. Overnight cultures were diluted to an OD600 of 0.04 in the LB medium and then further diluted 10−3-fold. One hundred microliters of the diluted culture were spread with glass beads on LB agar plates and incubated at 37 °C or 20 °C.
https://doi.org/10.1371/journal.ppat.1013047.s005
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S5 Fig. Effect of bipA deletion on capsule production.
(A) Colony morphology of wild-type and bipA-deleted strains of E. coli and S. Typhimurium. Overnight cultures were diluted and spread as in S4B Fig. (B) Macrocolony assay of the same strains at 37 °C for 18 h or 20 °C for 7 d. Overnight cultures were diluted to an OD600 of 0.02 in the same medium, and 3 μL of the diluted cultures were spotted on MacConkey agar plates. (C) Relative quantification of gmd mRNA levels in the cps cluster. Total RNA was extracted from cells grown to an OD600 of 0.5 and analyzed using qRT-PCR. The relative quantity was normalized to the rrsA gene and expressed relative to the transcript level of the wild-type strain grown at 37 °C.
https://doi.org/10.1371/journal.ppat.1013047.s006
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S6 Fig. Reduced swarming motility and flagellin production in bipA-deleted S. Typhimurium strains at low temperatures.
(A) Swarming motility assay of wild-type and bipA-deleted strains of E. coli and S. Typhimurium at 37 °C or 20 °C. The cells were grown and diluted, as described in Fig 7A. The cultures were concentrated 10-fold and spotted on an LB agar plate containing glucose. Plates were incubated at 37 °C for 14 h or 20 °C for 2 d. (B) Reduced flagellin production in the ΔbipA of S. Typhimurium at 20 °C. Overnight cultures were diluted 10−2-fold in fresh medium and incubated until the OD600 reached 0.5 at 37 °C or 20 °C. Deflagellation was performed as described in the Materials and Methods section. Western blot analysis of the wild-type and bipA-deleted strain samples was conducted using an anti-FliC antibody. To determine the locations of FliC and FljB, the lysates of MG1655 cells harboring pIN-A, pINFliC, pINSeFliC, or pINSeFljB were loaded onto SDS-PAGE.
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S7 Fig. Repression of flagella-related gene transcription in bipA-deleted strains at low temperatures.
Total RNA was extracted from the wild-type and bipA-deleted strains cultured in the early exponential phase at 37 °C or 20 °C and analyzed using qRT-PCR. The relative expression levels of flgM, fliE, fliK, fliM, fliS, and fliZ were normalized to the endogenous control gene rrsA.
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S8 Fig. Increased polymyxin B susceptibility in bipA-deleted strains at 20 °C.
(A) Growth curves of wild-type and bipA-deleted strains in the presence of polymyxin B. Overnight cultures were diluted 200-fold in fresh LB medium. A 180 μL aliquot of the diluted culture was added to each well of a 96-well plate, along with 20 μL of polymyxin B or distilled water as a control. Plates were incubated at 37 °C for 18 h or at 20 °C for 48 h with shaking. (B) ΔLag time between wild-type and bipA-deleted strains. Lag time was determined using Gen5 software (Agilent Technologies). ΔLag time was calculated as the lag time of the mutant strain minus the lag time of the wild-type strain. (C) Polymyxin B E-test for MIC determination. A polymyxin B MIC test strip was placed onto LB agar plates spread with diluted cultures of wild-type or bipA-deleted strains, followed by incubation at 37 °C or 20 °C.
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S9 Fig. Elevated expression of fimD in bipA-deleted strains.
Total RNA was extracted from wild-type and bipA-deleted strains in the early exponential phase at 37 °C or 20 °C and analyzed using qRT-PCR. The relative expression levels of the type 1 fimbriae gene fimD were normalized to the endogenous control gene rrsA.
https://doi.org/10.1371/journal.ppat.1013047.s010
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S10 Fig. ATP levels in wild-type and bipA-deleted strains at 37 °C and 20 °C.
Lysate samples were prepared from wild-type and bipA-deleted strains cultured in the early exponential phase at 37 °C or 20 °C, as described in S1 Supporting Information. A 50 μL sample was mixed with 50 μL of luciferase reagent, followed by luminescence measurement. ATP concentrations were calculated based on a standard curve ranging from 10-5-10-10 M.
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S11 Fig. Decreased expression levels of SPI-1 genes in bipA-deleted strain at 20 °C.
Total RNA was extracted from the wild-type and bipA-deleted cells incubated at 37 °C or 20 °C to the early exponential phase and analyzed using qRT-PCR. The relative expression levels of hilA, invA, prgH, and sipA were normalized to the endogenous control gene rrsA.
https://doi.org/10.1371/journal.ppat.1013047.s012
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S12 Fig. BipA is required for optimal growth in the presence of ROS.
(A) Growth curves of wild-type and bipA-deleted strains of E. coli and S. Typhimurium in the presence of H2O2. Growth was measured as described in S8A Fig. (B) Increased lag time in bipA-deleted strains in the presence of H2O2. ΔLag time was calculated as described in S8B Fig.
https://doi.org/10.1371/journal.ppat.1013047.s013
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S1 Table. Primers used in this study.
List of primers used for strains and plasmids construction, and qRT-PCR.
https://doi.org/10.1371/journal.ppat.1013047.s014
(DOCX)
S1 Data. Source data for graphs in this study.
This file contains the raw data used to generate all the graphs presented in the manuscript, as well as the data points extracted from images used in the analysis.
https://doi.org/10.1371/journal.ppat.1013047.s015
(XLSX)
Acknowledgments
We thank Dr. Ho Young Kang (Pusan National University) for generously providing the anti-FliC antibody.
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