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Infect Immun. Jul 2007; 75(7): 3305–3314.
Published online Apr 23, 2007. doi:  10.1128/IAI.00351-07
PMCID: PMC1932945

Identification of an Orphan Response Regulator Required for the Virulence of Francisella spp. and Transcription of Pathogenicity Island Genes[down-pointing small open triangle]

Abstract

Francisella tularensis is a category A agent of biowarfare/biodefense. Little is known about the regulation of virulence gene expression in Francisella spp. Comparatively few regulatory factors exist in Francisella, including those belonging to two-component systems (TCS). However, orphan members of typical TCS can be identified. To determine if orphan TCS members affect Francisella gene expression, a gene encoding a product with high similarity to the Salmonella PmrA response regulator (FTT1557c/FNU0663.2) was deleted in Francisella novicida (a model organism for F. tularensis). The F. novicida pmrA mutant was defective in survival/growth within human and murine macrophage cell lines and was 100% defective in virulence in mice at a dose of up to 108 CFU. In addition, the mutant strain demonstrated increased susceptibility to antimicrobial peptide killing, but no differences were observed between the lipid A of the mutant and the parental strain, as has been observed with pmrA mutants of other microbes. The F. novicida pmrA mutant was 100% protective as a single-dose vaccine when challenge was with 106 CFU of F. novicida but did not protect against type A Schu S4 wild-type challenge. DNA microarray analysis identified 65 genes regulated by PmrA. The majority of these genes were located in the region surrounding pmrA or within the Francisella pathogenicity island (FPI). These FPI genes are also regulated by MglA, but MglA does not regulate pmrA, nor does PmrA regulate MglA. Thus, the orphan response regulator PmrA is an important factor in controlling virulence in F. novicida, and a pmrA mutant strain is an effective vaccine against homologous challenge.

Francisella tularensis is a highly virulent gram-negative, facultative intracellular pathogen that causes tularemia in humans and other animals (17, 44). In nature, rabbits and rodents appear to be highly susceptible to F. tularensis infection and tularemia. It is assumed that vectors such as ticks and mosquitoes feed on the infected animals, sustaining the life cycle of F. tularensis and spreading the disease to humans (35). There are two clinically important subspecies: F. tularensis subsp. tularensis (type A strains) and F. tularensis subsp. holarctica (type B strains). Type A strains are found predominantly in North America, are highly infectious, and cause a life-threatening disease in humans, especially when inhaled (13, 14). Type B strains are found primarily in North America and Eurasia and are considered to be less virulent for mammals than type A strains (37). An attenuated live vaccine strain was derived from a type B strain (48), and it elicits a protective response in humans, monkeys, guinea pigs, and mice against systemic challenge with virulent type A F. tularensis (14, 16, 40, 41). Regardless of the port of entry, the bacterium eventually spreads to organs such as the liver and spleen and results in a fatal outcome. Francisella novicida, a close relative of type A F. tularensis, does not cause significant infection in humans (11, 15, 26) but causes a lethal infection of mice and is a good model for human tularemia.

F. tularensis survives and grows inside many types of cells, including macrophages (3, 18). The virulence mechanisms of this bacterium have not been clearly established, although the virulence regulator MglA and its Francisella pathogenicity island (FPI) targets, including iglC, iglD, and pdpA to -D, help Francisella to survive inside macrophages and cause disease (6, 20, 31, 36, 39). Electron microscopy studies have indicated that F. tularensis resides in a vacuolar compartment in phagocytic cells after entry, but within 6 h postinfection it is found primarily in the cytosol (3, 12). The mechanism of phagosomal escape is unclear, but MglA, IglC, and AcpA have each been shown to play a role in this process (6, 31, 32, 34, 39).

A large number of bacterial responses involve the function of two-component signal transduction systems (TCS). Currently more than 4,000 TCS have been identified in 145 sequenced bacterial genomes (49); thus, these TCS are ubiquitous in bacteria. Usually these regulatory systems consist of a membrane-bound sensor that monitors an environmental parameter and a cytoplasmic response regulator that mediates an adaptive response that is typically a change in gene expression. These systems nearly always function via a phosphorylation cascade, in which the sensor becomes autophosphorylated at a conserved His residue and subsequently transfers this phosphate to a conserved Asp in the response regulator. TCS regulate diverse responses, including nutrient acquisition, energy metabolism, virulence, adaptation to physical or chemical aspects of the environment, and complex developmental pathways (7). While most gram-negative bacteria possess numerous TCS, Francisella spp. have no complete paired TCS encoded in the published genome sequence (29). There are only 16 genes that are likely to be involved in environmental signal transduction, and of these, only 4 appear to be similar to genes for known TCS histidine kinases or response regulators. We hypothesize that, in order to coordinate efficient environmental responses, these orphan TCS members function in gene regulation.

In this study, we have demonstrated that the F. novicida pmrA gene, encoding an orphan response regulator, affects the expression of numerous Francisella genes, including those within the pathogenicity island. In addition, we have demonstrated the efficacy of the F. novicida ΔpmrA mutant as an attenuated vaccine candidate against intranasal wild-type F. novicida challenge.

MATERIALS AND METHODS

Bacterial strain, plasmid construction, and molecular biology techniques.

F. novicida U112 was routinely grown at 37°C in cysteine heart agar (Hi-Media Laboratories, Mumbai, India) and modified tryptic soy broth (TSB) (Difco Laboratories, Detroit, MI) containing 135 μg/ml ferric pyrophosphate and 0.1% cysteine hydrochloride. Bacteria were also frequently plated onto chocolate II agar (BBL, Becton Dickinson and Co., Franklin Lakes, NJ). When required, the growth medium was supplemented with kanamycin (Kan) (25 μg/ml) or tetracycline (15 μg/ml). All manipulations with F. novicida were performed in a class II biological safety laboratory. Escherichia coli DH5α was grown at 37°C aerobically in Luria-Bertani (LB) medium (Difco Laboratories, Detroit, MI) supplemented with Kan (15 μg/ml), tetracycline (12.5 μg/ml), or ampicillin (100 μg/ml) as required.

Standard DNA isolations (plasmid and chromosomal) were performed with QIAGEN kits according to the manufacturer's instructions. E. coli electroporations and standard genetic manipulations (e.g., ligations) were performed according to methods described by Sambrook et al. (38). To construct the pPmrA-Kan plasmid, a fragment of 3,262 bp that included pmrA plus approximately 1 kb upstream and downstream was amplified with primers JG1167 (5′cggaattcCCAGTCGGTGCAAAGACAGGAAAACTT3′) and JG1168 (5′acatgcatgcACCGCCCTGTTCACTAGCACC3′) from F. novicida genomic DNA (uppercase letters indicate sequences in the F. novicida genome). The amplified fragment was digested with EcoRI and HindIII, blunt ended with Klenow fragment, and cloned into blunt-ended and dephosphorylated EcoRI- and HindIII-digested pUC18 vector. The resulting plasmid was renamed pPmrAUpDn. An F. tularensis outer membrane protein (YP_169847) promoter was incorporated into primer JG868, and this primer was used with JG1048 to amplify a Kan cassette from plasmid pDSK519 (JG868, 5′cggaattcggatccctgcagatcgattgttgtttcaagttttgataatgattaaaaataataggagttaaaaATGAGCCATATTCAACGGGAAACG3′; JG1048, 5′aactgcagTTAGAAAAACTCATCGAGCATCAAATGAAACTGC3′). The amplified fragment containing the Kan cassette driven by the YP_169847 promoter was digested with KpnI and PstI, blunt ended with Klenow fragment, and ligated into HindIII- and NheI-digested and blunt-ended pPmrApUpDn. This replaces the pmrA gene with the Kan cassette. HindIII and NheI fortuitously digest at the ends of the prmA gene such that no other gene is affected by the deletion of this fragment. The resulting plasmid was renamed pPmrA-Kan. Plasmid pPmrA-Kan was transformed into F. novicida by cryotransformation as previously described (34). Kan-resistant and ampicillin-sensitive colonies were identified and confirmed by PCR to have the Kan cassette in place of the pmrA open reading frame. This resulted in the creation of a F. novicida ΔpmrA::Kan strain (JSG2845). For complementation in trans, plasmid pKK214 containing the groEL promoter of the F. tularensis live vaccine strain was used (1). The pmrA gene was amplified by PCR using primers JG1184 (5′cgggatccATGAGAATATTGTTGGCTGAAGATGATCTT3′) and JG1185 (5′aactgcagTTACTTAATTACTTTATCCTTTTGTACAAAGTAACCAAC3′) and cloned into pKK214groEL such that pmrA was expressed from the groEL promoter. The resulting plasmid, pPmrA, was introduced into JSG2845 by cryotransformation, creating strain JSG2847. Details of the strains, plasmids, and primers are shown in Table Table11.

TABLE 1.
Strains, plasmids, and primers

Antimicrobial peptide (AMP) assay.

Polymyxin B (8,050 units/mg; Invitrogen, Carlsbad, CA) and protegrin 1 (IntraBiotics Pharmaceuticals, Lafayette, CA) were dissolved in double-distilled deionized sterile water to a final concentration of 1 mg/ml and stored at −20°C. Overnight cultures grown in TSB with 0.1% cystine hydrochloride were diluted to 1 × 105 CFU/ml in TSB. Diluted cultures (50 μl) were placed in 96-well microtiter dishes and mixed with various amounts of peptide (50 μl, 2× concentration) diluted in phosphate-buffered saline (PBS). Cells and peptides were incubated at 37°C overnight in a humid environment. Wells were evaluated for growth visually, and a portion of each sample was diluted in PBS and plated on chocolate II agar plates to assess bacterial viability. Controls contained PBS in place of peptide. Data are presented as percent survival relative to the original inoculum and are from triplicate determinations.

Intramacrophage survival assay.

F. novicida or F. novicida ΔpmrA was used to infect J774.1 murine macrophages and phorbol myristate acetate (PMA) (10 ng/ml)-induced THP-1 human macrophages at a multiplicity of infection of ~50:1. Wells were seeded with ~2 × 105 macrophages, and ~1.0 × 107 to 1.1 × 107 bacteria were added to each well. After 2 h of incubation at 37°C and 5% CO2, gentamicin (50 μg/ml) was added to the medium to eliminate extracellular organisms. After 30 min, wells were washed twice with PBS and incubated with their respective media supplemented with 10 μg/ml gentamicin. The macrophages were lysed with 0.1% sodium dodecyl sulfate (SDS) at 2 h, 12 h, and 24 h postinfection, and the lysates were serially diluted in PBS and plated on chocolate II agar plates for determination of viable counts.

Mouse survival studies.

Francisella strains were given intranasally to groups of five anesthetized female 4- to 6 week-old BALB/c mice (Harlan Sprague, Indianapolis, IN) at a dose of ~100 to ~1 × 108 CFU/20 μl PBS. Actual bacterial counts delivered were determined by plate counts from the inoculum. Mice were monitored for 5 weeks postinfection. The F. novicida ΔpmrA-infected mice were challenged intranasally with F. novicida at 5 weeks postvaccination and monitored for 3 weeks postchallenge. As a control, five unvaccinated mice were infected with equal doses of F. novicida. To determine organ burdens, the livers and spleens were harvested from three mice that were sacrificed at specific time points. Organs were macerated, diluted, and plated onto chocolate II plates to enumerate the bacteria. Type A Schu S4 challenge experiments were performed in CDC-approved BSL3/aBSL3 suites at The Ohio State University.

Microarray.

Bacterial RNA (10 μg) was prepared according to protocols supplied in the Affymetrix GeneChip Expression Analysis Prokaryotic manual (Santa Clara, CA). Briefly, TSB-cysteine broth was inoculated with a colony from a 48-h chocolate II agar plate and grown to an optical density at 600 nm of 0.5. The cells were harvested, and RNA was isolated with the RNeasy kit (QIAGEN, Valencia, CA). The assay utilizes reverse transcriptase and random hexamer primers to generate DNA complementary to the RNA. The cDNA is then fragmented by DNase I and labeled with terminal transferase and biotinylated GeneChip DNA labeling reagent at the 3′ termini. The labeled samples were hybridized to a custom GeneChip containing probes to the 1,804 open reading frames of the F. tularensis Schu S4 genome (supplied by Battelle). Washing and staining of the chips with streptavidin-phycoerythrin were performed using the Affymetrix Fluidics Station 450. Scanning of the chips was done using the Affymetrix Genechip Scanner 3000.

Microarray data analysis.

The GeneSifter microarray data analysis system was used to identify changes in gene expression. The CHP files (Affymetrix MAS 5 normalized) for all samples were loaded into GeneSifter. Using a filtering criterion of a 1.5-fold or greater change in expression and a P value of <0.05 from analysis of variance, a list of differentially expressed genes was generated. The data were corrected using the method of Hochberg and Benjamini (25a) to derive a false discovery rate estimate from the raw P values, and a false discovery rate of 5% was used as a cutoff.

Quantitative real-time PCR (qRT-PCR).

RNA was extracted from mid-log-phase (optical density at 600 nm, 0.4 to 0.5) F. novicida and F. novicida ΔpmrA bacteria by use of the RNeasy Kit (QIAGEN, Valencia, CA). The RNA quality and quantity were determined with the Experion automated electrophoresis system (Bio-Rad, Hercules, CA). One microgram of total RNA was reverse transcribed to cDNA with Superscript II RNase H reverse transcriptase (Invitrogen, Carlsbad, CA) and normalized according to the concentration. Two nanograms of the converted cDNA was used for quantitative PCR with the SYBR green PCR master mixture in the Bio-Rad iCycler apparatus (Bio-Rad, Hercules, CA). Relative quantification was used to evaluate the expression of chosen genes. All primers were designed to give 200- to 220-nucleotide amplicons, have a G+C range of 30 of 50%, and a melting temperature of 58 to 60°C. Relative copy numbers and expression ratios of selected genes were normalized to the expression of two housekeeping genes (the 16S rRNA gene and dnaK) and calculated as described by Gavrilin et al. (19).

Protein purification and Western blotting.

A pmrA His tag fusion was constructed in the pQE30Xa vector (QIAGEN, Valencia, CA). This fusion was generated by PCR amplification of pmrA using primers JG1184 (5′cgggatccATGAGAATATTGTTGGCTGAAGATGATCTT3′) and JG1185 (5′aactgcagTTACTTAATTACTTTATCCTTTTGTACAAAGTAACCAAC3′). This fragment was then cloned between the BamHI and PstI sites to obtain plasmid pQE30Xa-PmrA. The His-tagged PmrA protein was purified using a Ni-nitrilotriacetic acid resin affinity chromatography column according to the QIAexpressionist purification protocol (QIAGEN, Valencia, CA). The eluted protein was exhaustively dialyzed against 20 mM Tris-HCl (pH 7.9)-50 mM KCl. The protein concentration of purified PmrA was determined with the bicinchoninic acid assay kit (Pierce Biotechnology Inc., Rockford, IL), and the purified PmrA-H6 protein was analyzed by SDS-polyacrylamide gel electrophoresis (SDS-PAGE). Antiserum against PmrA was raised in rabbits at Alpha-Diagnostics (San Antonio, TX). The titer of the sera was determined by enzyme-linked immunosorbent assay and Western blotting.

Whole-cell lysate preparation and 2D gel electrophoresis.

Whole-cell lysates were prepared essentially as described by Hubalek et al. (27). Briefly, the whole-cell lysate was prepared from 25-ml overnight cultures of F. novicida and F. novicida ΔpmrA grown in TSB-0.1% cysteine hydrochloride medium. The pellet was collected at 16,000 × g for 15 min at 4°C and suspended in 1 ml of cold PBS. Bacteria were then centrifuged, and the pellets were resuspended in 1 ml of lysis buffer composed of 137 mM NaCl, 10% glycerol, 1% p-octyl-β-d-glucopyranoside, 50 mM NaF, 1 mM Na3VO4, and protease inhibitor cocktail (Sigma-Aldrich, St. Louis, MO). The bacterial suspension was then subjected to 10 cycles of freeze-thawing in liquid nitrogen, and undisrupted microbes were removed by centrifugation at 16,000 × g for 15 min at 4°C. Bacterial proteins were precipitated by using the ReadyPrep two-dimensional (2D) cleanup kit (Bio-Rad, Hercules, CA) and then solubilized in rehydration buffer (2 M thiourea, 6 M urea, 4% (wt/vol) 3-[(3-cholamidopropyl)-dimethylammonio]-1-propanesulfonate (CHAPS), 40 mM Tris base, 2 mM tributylphosphine, 0.003% (wt/vol) bromophenol blue, and 1% (vol/vol) Biolyte pH 3-10). The protein concentration was determined with the reducing agent-compatible-detergent-compatible protein assay kit II (Bio-Rad, Hercules, CA).

2D gel electrophoresis was performed basically as previously described (25). The isoelectric-focusing 11-cm, pH 5 to 8 gradient Readystrip immobilized pH gradient (IPG) strips were rehydrated overnight and equilibrated for 20 min with buffer containing 6 M urea, 2 M thiourea, 2% SDS, 0.375 M Tris-HCl (pH 8.8), 20% glycerol, and 2% dithiothreitol, followed by 20 min of equilibration in the same buffer plus 4% iodoacetamide. The proteins extracted from whole-cell lysates (300 μg) were separated by use of rehydrated IPG strips run at 200 V for 30 min, 500 V for 2 h, and 8,000 V for 24 h (Bio-Rad, Hercules, CA). Second-dimension gel electrophoresis was carried out in an 8 to 16% gradient SDS-PAGE ready gel (Bio-Rad, Hercules, CA) at 30 mA. Gels were fixed for 1 h in 10% methanol and 7% acetic acid, followed by overnight staining with Sypro Ruby (Bio-Rad, Hercules, CA) or Coomassie blue. Background staining was removed by 10% methanol and 7% acetic acid washes, and a gel picture was captured in the Bio-Rad gel Doc system. At least five replicates of the gel were run for each sample and analyzed with PD-Quest software (Bio-Rad, Hercules, CA). Gel spots were matched by this software as well as manually. Twelve spots of interest were excised from a gel, washed, rehydrated, and dried before trypsin digestion. The in-gel digests of the protein spots were analyzed by liquid chromatography-tandem mass spectrometry.

LPS and lipid A isolation from bacteria and MALDI mass spectrometry.

Lipopolysaccharide (LPS) of F. novicida and F. novicida ΔpmrA grown in TSB-0.1% cysteine hydrochloride medium was purified from overnight-grown stationary-phase cells by using the hot phenol-water method as previously described (4). Various concentrations of LPS were loaded on 18% deoxycholine (DOC)-polyacrylamide gels and electrophoresed at a 30-mA constant current. Completed gels were treated with alcain blue and silver stained. Lipid A was isolated after hydrolysis in 1% SDS at pH 4.5 as described previously (10). Briefly, 500 μl of 1% SDS in 10 mM Na-acetate (pH 4.5) was added to a lyophilized sample. Samples were incubated at 100°C for 1 h, frozen, and lyophilized. The dried pellets were resuspended in 100 μl of water and 1 ml of acidified ethanol (100 μl 4 N HCl in 20 ml 95% ethanol). Samples were centrifuged at 5,000 rpm for 5 min. The lipid A pellet was further washed (three times) in 1 ml of 95% ethanol. The entire series of washes was repeated twice. Samples were resuspended in 500 μl of water, frozen on dry ice, and lyophilized. Lipid A was used for matrix-assisted laser desorption ionization (MALDI) mass spectrometry analysis as described by Gunn et al. (22).

Microarray accession number.

The microarray data associated with this paper can be found at ArrayExpress under accession number A-AFFY-83.

RESULTS

Francisella TCS members.

To determine the role of TCS in Francisella gene regulation, we examined the genome sequence of F. tularensis Schu S4 (type A) for complete or partial members. As suggested previously (29), Francisella spp. have no intact TCS. However, single components identified were histidine kinases FTT0094c and FTT1736c and response regulators FTT1543 and FTT1557c. One of the orphan response regulators that was identified possessed 44% identity over its entire length to the PmrA protein of Salmonella. Thus, we have named the gene (FTT1557c/FNU0663.2) encoding this protein pmrA. In Salmonella, this TCS is involved in LPS modification, survival in high-iron and low-magnesium environments, AMP resistance, and mouse virulence (21-23, 33, 45, 52).

To determine the function of pmrA in F. novicida, the open reading frame was replaced by a Kan gene cassette. The deletion and Kan gene insertion were confirmed by PCR and DNA sequencing (data not shown), as well as by Western blot analysis (Fig. (Fig.1).1). The resulting pmrA::Kan strain (JSG2845) had no obvious morphological differences from the parental strain and growth characteristics identical to those of the parental strain (data not shown).

FIG. 1.
Western blot image of the PmrA protein of F. novicida. Ten micrograms of total protein was loaded in each well and separated by 12% SDS-PAGE. Electrophoresis was performed at constant voltage (80 V). Proteins were transferred to nitrocellulose ...

The pmrA mutant has increased susceptibility to AMP.

Antimicrobial peptides (AMPs) are ubiquitous in nature and represent a host defense strategy in mammals, amphibians, and insects (2). They are produced at mucosal sites (including those of the airway and lung) and within phagocytes, which represent anatomic locations/cells likely encountered during Francisella infection of a host. In Salmonella spp., resistance to the AMP polymyxin B is regulated by PmrAB (21). To determine if the Francisella pmrA mutant was altered in AMP resistance, MIC and time-kill assays were performed with polymyxin B and protegrin 1. Overall, F. novicida was highly resistant to AMP killing and was much more so than gram-negative enteric bacteria such as E. coli, which have MICs of less than 10 μg/ml. The MIC assay revealed that the F. novicida ΔpmrA mutant was 32-fold more susceptible than its F. novicida parental strain against polymyxin B and protegrin 1 (MICs of 800 μg/ml versus 25 μg/ml to polymyxin B and of 50 μg/ml versus 1.56 μg/ml to protegrin 1). Therefore, the loss of pmrA dramatically affects resistance to AMP killing.

DOC-PAGE and MALDI mass spectrometry analysis of LPS and lipid A.

Because PmrA affects AMP resistance in various gram-negative bacteria by LPS modification, LPS was isolated from bacteria grown in various media and at various temperatures. DOC-PAGE analysis of the LPSs from F. novicida and F. novicida ΔpmrA showed that they had identical banding patterns, indicating no gross changes in size or mobility (data not shown). The results of the MALDI-time-of-flight mass spectrometry analyses of the lipid A isolated by very mild acid hydrolysis of the purified LPS preparations showed no significant difference (data not shown).

F novicida ΔpmrA is defective in intramacrophage survival.

F. tularensis and F. novicida can survive and replicate intracellularly in human and mouse macrophages (3, 5, 8, 18, 46, 47), and this characteristic is key to dissemination and virulence. Here we examined the intracellular growth kinetics of F. novicida strains in J774.1 murine macrophages and PMA-induced THP-1 cells. After 24 h of growth in PMA-induced THP-1 cells and J774.1 cells, the mutant showed 123- and 73-fold reductions, respectively, in survival compared to F. novicida (Fig. 2A, and B). While the mutant cell numbers were reduced over time in J774.1 cells, there was little or no net growth or killing over time in THP-1 cells. Complementation experiments resulted in intramacrophage survival of the mutant in both macrophage cell lines that was similar to that of F. novicida (Fig. 2A and B). We also examined survival in HeLa epithelial cells. While the bacteria were able to enter these nonphagocytic cells in equal numbers (mutant versus wild type), entry was much less efficient than in macrophages. The data show that in HeLa cells, the mutant strain showed a 310-fold reduction in survival at 24 h postinfection (Fig. (Fig.2C).2C). These results demonstrate that pmrA plays a major role in intracellular survival of F. novicida.

FIG. 2.
Intracellular survival assays performed with the THP-1 (A), J774.1 (B), and HeLa (C) cell lines. Standard gentamicin assays were conducted with a multiplicity of infection of ~50:1 (bacteria to eukaryotic cells). After cell lysis, bacteria were ...

The F. novicida ΔpmrA has a virulence defect in the mouse model.

To examine virulence in the mouse model, BALB/c mice were infected intranasally with various doses of either F. novicida or F. novicida ΔpmrA. F. novicida-infected mice (n = 30) were sick between days 2 and 3 postinfection and died before 9 days (lowest dose, 10 CFU); however, all of the F. novicida ΔpmrA-infected mice (n = 80 total mice over multiple experiments) survived more than 9 weeks postinfection at all doses up to the highest tested, 1 × 108 CFU (Fig. (Fig.3).3). Complementation of the ΔpmrA strain restored virulence (Fig. (Fig.3).3). The bacterial burdens in the livers and spleens of F. novicida-infected mice were examined at various time points postinfection, and bacterial numbers were shown to increase by 105 to 107 CFU over the initial inoculum of 10 or 1,000 CFU (Fig. (Fig.4A).4A). However, very few bacteria (100 to 1,000 CFU) were recovered from the livers and spleens of F. novicida ΔpmrA-infected mice, and no further replication was observed after 48 h postinfection (Fig. (Fig.4B4B).

FIG. 3.
Mouse virulence assays. BALB/c mice (n = 5 in each group) were anesthetized and infected with various doses of F. novicida and F. novicida ΔpmrA by the intranasal route: F. novicida, 10 CFU (diamonds) and 103 CFU (squares); F. novicida ...
FIG. 4.
Bacterial numbers in the livers and spleens of BALB/c mice (n = 3 at each time point) infected with 10 or 1,000 CFU of F. novicida (A) or F. novicida ΔpmrA (B). Organs were removed at various times postinfection and the CFU in these organs ...

To determine the protective capacity of this attenuated strain, mice vaccinated with various doses of the F. novicida ΔpmrA mutant were challenged with F. novicida (106 CFU/mouse). All vaccinated mice survived the F. novicida challenge, including those vaccinated with only 10 CFU. At days 40, 45, and 50 postchallenge, mice from each group were sacrificed and their organ burdens were determined. Few bacteria could be recovered in the liver and spleen after 50 days postchallenge, with the numbers steadily decreasing over days 40 to 50 (Fig. 5A and B). A vaccine dose of 106 CFU was not protective against an extremely high challenge dose of 1011 CFU, and though this was not unexpected, F. novicida ΔpmrA-vaccinated mice (106 CFU) were not protected against wild-type type A (Schu S4) challenge (challenge dose of 100 CFU) (data not shown). Thus, these results demonstrate that the F. novicida ΔpmrA mutant is highly attenuated in the mouse model and is 100% protective versus homologous bacterial challenge at nearly all vaccine-challenge doses.

FIG. 5.
Bacterial loads in livers (A) and spleens (B) of BALB/c mice (n = 5) challenged intranasally with F. novicida 5 weeks postvaccination with F. novicida ΔpmrA. Mice were sacrificed at various time points after F. novicida infection to determine ...

DNA microarray identification of PmrA-regulated genes.

To determine the genes that were regulated by PmrA, we pursued a DNA microarray approach. This analysis identified 65 differentially expressed genes (F. novicida Δ pmrA versus F. novicida), with a 1.5-fold cutoff. Of these, 52 genes were PmrA activated and 13 genes were PmrA repressed. Thirteen of the total identified genes were pseudogenes, while 27 encoded hypothetical proteins (Table (Table2).2). Interestingly, 11 of the 52 PmrA-activated genes were located in the FPI (e.g., pdpD, iglC, and iglD). In addition, five genes in the local region around pmrA were shown to be regulated. Three genes were shown to be very highly PmrA repressed, and these are the F. novicida gene homologs to FTT1242, FTT0172, and FTT0272 (all encoding hypothetical membrane proteins).

TABLE 2.
Comparison of gene expression in F. novicida ΔpmrA and wild-type F. novicida, determined by microarray analysis and qRT-PCR

In addition to microarray analysis, we used 2D PAGE to compare the expression patterns of F. novicida and F. novicida ΔpmrA. The majority of the protein spots resolved in the pH range of 5 to 8. Computer analysis of the protein spot patterns and intensities showed 60 proteins regulated by PmrA, with 44 of these reduced by the loss of pmrA (putative PmrA-activated species) and 4 enhanced by the loss of PmrA (putative PmrA-repressed species). Twelve spots were analyzed by mass spectrometry, but only four of these spots resulted in a positive identification with a Francisella gene product. Two spots (spots 1 and 2 in Fig. Fig.6)6) contained the same peptide corresponding to the Schu S4 protein FTT1242, encoded by the above-mentioned most highly regulated gene identified by microarray analysis. However, while this is a single gene in type A strains, these two spots represent two near-identical genes in F. novicida (FNU466 and FNU467). The other identified spots correspond to FTT1377 and FTL0632 (F. tularensis subsp. holarctica), which are hypothetical proteins and were not among those genes identified as PmrA regulated by microarray analysis.

FIG. 6.
Representative Coomassie blue-stained 2D gels of the F. novicida and F. novicida ΔpmrA whole-cell protein fractions. The fractions were electrophoresed in IPG strips (pH 5 to 8), followed by SDS-PAGE (9 to 16% gradient). The numbered spots ...

To confirm the microarray results, we compared the expression of nine PmrA-activated genes and two PmrA-repressed genes (FTT1242/FNU466 and FNU467) by qRT-PCR. These data corroborated the microarray data but frequently showed greater levels of regulation compared to what was observed in the microarray analysis (Table (Table2).2). This is likely due to the necessary background subtraction in microarray analysis. Because MglA and PmrA both appeared to regulate identical genes within the pathogenicity island, we examined whether mglA could regulate pmrA and vice versa. These data demonstrate that under the growth conditions employed, which were the same as those for which bacteria were grown for RNA isolation for cDNA synthesis and microarray analysis, PmrA and MglA are not in a hierarchical pathway involving transcriptional regulation (the mglA fold change in F. novicida ΔpmrA versus F. novicida was 1.1; the pmrA fold change in F. novicida ΔmglA versus F. novicida was 1.1).

DISCUSSION

Francisella spp. have been shown to exist within amoebae in the environment and within macrophages in vertebrate hosts (1, 3, 18, 46, 47). Coping with this and other complex environments requires coordinated gene expression. Such responsiveness to the environment is often mediated by TCS; however, Francisella spp. do not express any tandemly arranged (sensor and response regulator genes), functional TCS. Orphan members of these systems do exist in the genome, and in this work, we determined that one of these orphan members functions in the regulation of F. novicida virulence.

The product of gene FTT1557c (F. tularensis Schu S4)/FNU0663.2 (F. novicida) is homologous to the S. enterica serovar Typhimurium PmrA response regulator. PmrA is part of the PmrA-PmrB TCS found in various gram-negative organisms, including Salmonella spp., E. coli, and Pseudomonas, and functions similarly in each organism to modify the LPS and affect resistance to AMP (28). F. novicida PmrA was shown to play a significant role in resistance to the AMP polymyxin B and protegrin 1. This result was perhaps a bit surprising, as PmrA-PmrB affects resistance to polymyxin B but not to protegrin 1 in various gram-negative organisms (J. S. Gunn, unpublished results). In addition, while PmrA-PmrB directs the modification of the LPS lipid A with phosphoethanolamine and 4-aminoarabinose groups, no differences were detected between the lipids A of F. novicida and F. novicida ΔpmrA. Perhaps the inherent high-level AMP resistance of Francisella spp. (even in the absence of pmrA) has eliminated the need for the complex regulated lipid alterations. Thus, while PmrA mediates enhanced resistance to AMPs of various structures, the mechanism for PmrA-mediated AMP resistance remains unknown.

The loss of pmrA severely attenuated F. novicida. At the highest dose given, 108 CFU intranasally, all of the infected mice survived. Thus, this mutation attenuates Francisella more than other virulence gene mutations that have been reported, including mglA (50% lethal dose, 3.1 × 106) or iglC (50% lethal dose, 9.4 × 107) (31). The pmrA mutant was also defective for survival/growth in a variety of vertebrate macrophages. This appears to be a common trait of avirulence in Francisella, as all or nearly all avirulent mutants that have been identified also are defective for intramacrophage survival (43). Interestingly, the F. novicida ΔpmrA mutant was not eliminated quickly from the livers and spleens of infected mice, as it was maintained at ~1,000 CFU up to 7 days postinfection. Upon challenge of vaccinated animals with F. novicida, all of the mice survived (vaccine dose, 10 to 108 CFU; challenge dose 106 CFU). The vaccine dose of 106 CFU was not protective against an extremely high dose of F. novicida (1011 CFU), nor was it protective against Schu S4 challenge. This latter result was not surprising, though, since F. novicida does not protect against Schu S4 challenge (42; Gunn, unpublished results). Sterilizing immunity was not achieved within 50 days postchallenge, as ~100 CFU could be recovered at this time point from the spleens or livers of vaccinated mice (all were F. novicida and not the vaccine strain, which was eliminated from these organs by this time point). Thus, the F. novicida ΔpmrA mutant strain is a very effective live-attenuated vaccine against homologous challenge. Perhaps the reason why it worked so effectively as a vaccine was the fact that the bacterium was not eliminated quickly from the mouse, allowing sufficient time for an efficient protective immune response to develop.

We identified 65 genes that were regulated by PmrA in F. novicida (>1.5-fold). Of these, 80% were activated by PmrA. While over half of the activated genes encode hypothetical proteins of unknown function, a large proportion of activated loci are found within the FPI. This island contains a number of genes previously identified to play a role in intramacrophage survival and/or virulence (36). It is highly likely that the loss of expression of these genes resulted in the observed virulence attenuation and intramacrophage survival defects of the F. novicida ΔpmrA mutant. However, PmrA is not the only Francisella regulator required for FPI gene transcription. MglA, which is similar to the E. coli “stringent starvation” regulator SspA (50, 51), is also required. PmrA and MglA do not appear to be in a cascade, as they do not affect the transcription of one another. In addition, most of the recently identified MglA-regulated genes do not overlap with those regulated by PmrA (9). It remains possible, though, that they affect the abundance/activity of one another at a posttranscriptional level. In addition, a third Francisella regulator (called SspA) that shares considerable homology with MglA has been recently described. These two regulatory factors, like their counterpart in E. coli, have been described to physically interact with RNA polymerase as a heterodimer (24; J. C. Charity and S. L. Dove, presented at the 5th International Conference on Tularemia, Woods Hole, MA, 1 to 4 November 2006). Thus, we hypothesize that FPI gene transcription requires all three regulators such that PmrA binds to FPI promoter recognition sites and physically interacts with SspA and/or MglA bound to RNA polymerase to initiate transcription of genes within the pathogenicity island.

Of the genes determined to be PmrA repressed, FTT1242 was the strongest, and it was the most regulated gene identified in the microarray analysis. The product of this gene was also identified in the 2D gel mass spectrometry fingerprinting, not in one but in two spots with different isoelectric points (pI) but essentially the same molecular weight. The reason for this is that FTT1242 is a single gene in F. tularensis Schu S4 but is two linked and highly related genes (65% identity) in F. novicida (FNU466/FNU467). qRT-PCR analysis of these two genes shows only FNU466 to be repressed by PmrA. Given this result and the pI of FNU466, this protein is that in spot 1 in Fig. Fig.6.6. The proteins encoded by these genes are of unknown function, so their roles in the phenotypes elicited by PmrA are unclear.

The regulation of virulence gene expression in Francisella spp. is poorly understood, as no regulated promoters have been studied at the genetic level. The identification of PmrA as an orphan two-component response regulator is a first step in the study of other sensor kinases/response regulators of this bacterium and demonstrates that these rogue elements function in gene regulation. Ongoing work will determine the roles of other orphan regulatory and sensory factors as well as the role that phosphorylation cascades may play in the function of these regulators.

Acknowledgments

We thank the OSU Proteomics Core Lab in the Davis Heart and Lung Research Institute for assistance with 2D gels.

This work was supported by funding from The Region V “Great Lakes” Regional Center of Excellence in Biodefense and Emerging Infectious Diseases Consortium (NIH award 1-U54-AI-057153).

Notes

Editor: V. J. DiRita

Footnotes

[down-pointing small open triangle]Published ahead of print on 23 April 2007.

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