Identification of Differentially Regulated Francisella tularensis Genes by Use of a Newly Developed Tn5-Based Transposon Delivery System

Blake W Buchan; Molly K McLendon; Bradley D Jones

doi:10.1128/AEM.02882-07

. 2008 Mar 14;74(9):2637–2645. doi: 10.1128/AEM.02882-07

Identification of Differentially Regulated Francisella tularensis Genes by Use of a Newly Developed Tn5-Based Transposon Delivery System^▿

Blake W Buchan ¹, Molly K McLendon ^1,2, Bradley D Jones ^1,^*

PMCID: PMC2394869 PMID: 18344342

Abstract

Francisella tularensis is the etiologic agent of an intracellular systemic infection of the lymphatic system in humans called tularemia. The organism has become the subject of considerable research interest due to its classification as a category A select agent by the CDC. To aid genetic analysis of this pathogen, we have constructed a temperature-sensitive Tn5-based transposon delivery system that is capable of generating chromosomal reporter fusions with lacZ or luxCDABE, enabling us to monitor gene expression. Transposition is catalyzed by the hyperactive Tn5 transposase, whose expression is driven by the Francisella groES promoter. When high-temperature selection (42°C) is applied to a bacterial culture carrying the transposon delivery plasmid, ∼0.1% of the population is recovered with Tn5 insertions in the chromosome. Nucleotide sequence analysis of a sample of mutants revealed that the insertions occur randomly throughout the chromosome. The kanamycin-selectable marker of the transposon is also flanked by FLP recombination target sequences that allow deletion of the antibiotic resistance gene when desired. This system has been used to generate transposon mutant libraries for the F. tularensis live vaccine strain as well as two different virulent F. tularensis strains. Chromosomal reporters delivered with the transposon were used to identify genes upregulated by growth in Chamberlain's defined medium. Genes in the fsl operon, reported to be involved in iron acquisition, as well as genes in the igl gene cluster were among those identified by the screen. Further experiments implicate the ferric uptake regulator (Fur) protein in the negative regulation of fsl but not igl reporters, which occurs in an iron-dependent manner. Our results indicate that we have created a valuable new transposon that can be used to identify and characterize virulence genes in F. tularensis strains.

Francisella tularensis is the etiologic agent of the human infectious disease tularemia. This bacterial pathogen has been categorized as a category A select agent due to its high virulence at a very low infectious dose (53, 55). A key pathogenic feature of this organism is its ability to grow within host cells, including macrophages, hepatocytes, and epithelial cell lines (1, 2, 11, 20, 23, 31, 50). Research efforts have demonstrated that F. tularensis strains grow in phagocytic cells by escaping from the phagosomal compartment into the host cell cytosol (10, 23, 32, 38), which allows the organisms to circumvent normal immune responses. While significant advances have been made in understanding aspects of Francisella infection, the extreme pathogenicity of this organism hints that many virulence factors remain unidentified and uncharacterized.

The development of genetic techniques and tools for studying Francisella pathogenesis has driven much of the progress that is being made in understanding the molecular pathogenesis of this organism. Some early efforts to identify virulence factors of Francisella relied upon nonspecific mutagenesis techniques (7, 39, 49) or unstable transposon insertions (Tn10 or Tn1721) (3-6, 12, 25, 29, 40). Additionally, Escherichia coli-Francisella shuttle vectors constructed for site-directed mutagenesis experiments (27, 35, 36, 43, 48) or for screening a promoter fusion library with a chloramphenicol acetyltransferase reporter gene (27) have been constructed. Other groups have used EZ::TN (Epicentre) transposon-transposase complexes to obtain stable Tn5 insertions in the chromosome of F. tularensis LVS (26, 51), F. tularensis subsp. novicida (21, 54), or F. tularensis Schu S4 (47). Recently, Maier et al. reported the development of a Himar1-based transposon system for creating mutants of F. tularensis (37).

The development of microarray and proteomic technologies provides alternative approaches relative to traditional transposon promoter fusion constructions for the study of gene and protein expression in bacteria. An advantage of a microarray or proteomic approach is that the expression of every bacterial gene or gene product can be compared under two or more conditions. A comprehensive view of the bacterial genome or proteome is difficult to achieve with transposon promoter fusions since one must simultaneously examine thousands of individual mutant strains. However, a transposon-based reporter library has unique and important advantages over either of the more global approaches. For instance, insertion of a transposon reporter into a given gene creates a mutant strain, in addition to creating a promoter reporter strain. With the mutant isolate in hand, work to characterize the mutated gene can be initiated very quickly. Another significant advantage is the ability to remutagenize a reporter strain to identify regulatory elements that govern expression of the gene. This allows regulatory pathways to be uncovered and characterized, which significantly increases our understanding of bacterial gene expression and signal transduction.

In this study, we report the construction of a highly efficient Francisella tularensis mutagenesis system that employs the hyperactive Tn5 transposase. Expression of the transposase has been placed under the dual control of the Francisella groES promoter and the lac operator and LacI repressor. The transposase, which resides outside the insertion sequences (mosaic ends), catalyzes insertion of the transposable element that carries a kanamycin resistance gene, flanked by FLP recombination target (FRT) sequences, and the pir-dependent R6K origin of replication into the Francisella chromosome. The transposon is delivered from a conditionally replicating (temperature-sensitive) F. tularensis plasmid (36) that, at high temperatures, allows the selection of insertions into the Francisella chromosome due to the loss of replication of the temperature-sensitive plasmid. Our results indicate that this transposon mutagenesis system produces single, random, stable insertions into the chromosomes of F. tularensis strains and is capable of creating a saturating Tn5 insertion library in a single experiment. In addition, derivatives of this system have been engineered to allow the creation of chromosomal luxCDABE or lacZ as transcriptional reporters.

We have utilized this system to create and screen a transposon library of F. tularensis LVS for differential gene expression when grown on modified Mueller-Hinton (MMH) or Chamberlain's defined medium (CDM). Reports in the literature indicate that growth of F. tularensis on CDM results in increased capsule production as well as increased type IV pilus expression (9, 22). In addition, it has been reported that growth on CDM causes a general increase in virulence of Francisella in a mouse model (9). Based upon the idea that CDM may upregulate virulence gene expression, we screened F. tularensis LVS transcriptional reporter libraries on MMH and CDM growth media. These screens have successfully identified established virulence genes as well as new genes that may play a role in the pathogenic lifestyle of F. tularensis. Some of the genes identified are involved in iron acquisition, suggesting that low iron availability is one of the signals sensed by Francisella on CDM agar that lead to upregulation of gene expression. Other groups have also reported iron availability as a signal resulting in differential expression of genes in F. tularensis (17, 30, 52). Classically, the ferric uptake regulator (Fur) functions as an iron-dependent transcriptional repressor by binding to DNA in the presence of ferrous iron (19). Sequences resembling Fur binding sites (Fur boxes) have been identified upstream of iron-regulated genes in F. tularensis (17, 52), although data linking Fur to regulation of these genes have not been presented. Based upon our observations and data published by others, we examined the role of Fur in the transcriptional regulation of two Francisella gene clusters that respond to iron concentration. We present evidence that suggests that Fur may regulate these two gene clusters by different mechanisms.

In summary, in this work we describe the construction and use of a Tn5 transposon delivery system that is capable of creating random, stable insertions in F. tularensis spp. We also present data demonstrating that chromosomal lacZ transcriptional reporters can be used to identify differentially regulated genes and quantify gene expression in F. tularensis. Finally, we have preliminary data that Fur is a negative regulator of transcription of the fslABCD operon but not the iglABCD operon.

MATERIALS AND METHODS

Bacterial strains, plasmids, growth conditions, and antibiotics.

F. tularensis LVS (ATCC 29684) and F. tularensis subsp. tularensis Schu S4 (Table 1) were grown in Mueller-Hinton broth (Becton Dickinson, Sparks, MD) or on Mueller-Hinton agar (Acumedia, Lansing, MI) supplemented with 1% glucose (wt/vol), 0.025% ferric pyrophosphate, and 2% IsoVitaleX. Spectinomycin and kanamycin were added to the growth media to give a final concentration of 25 μg/ml, when appropriate. CDM was prepared as described previously (8) or with 28 μM or 350 nM FeSO₄, as dictated by experiment. Plasmid pMKM219 is a derivative of the temperature-sensitive plasmid pFNLTP9 (36), in which the kanamycin resistance gene has been replaced with a spectinomycin resistance gene. The F. tularensis Fur-expressing plasmid was constructed by amplifying the fur gene from F. tularensis LVS, using oligonucleotide primers tailed such that the full-length fur gene could be cloned downstream of the groES promoter sequence in pBB103, a derivative of pKK202, to create pBB110. All cultures were grown at 30°C, 37°C, or 41°C as dictated by experiment. F. tularensis strain Schu S4 was handled within a BSL3 laboratory in accordance with all CDC/NIH regulatory and safety guidelines.

TABLE 1.

Bacterial strains and plasmids used in this study

Strain or plasmid	Description	Source or reference
F. tularensis strains
LVS	Live vaccine strain	K. L. Ekins
Schu S4	F. tularensis subsp. tularensis Schu S4 “type A” strain	BEI Resources
Plasmids
pRL27	Suicide plasmid carrying hyperactive transposase and Tn5 transposable element	28
pBDJ303	pRL27 modified for use in F. tularensis subsp.	This study
pMKM219	E. coli-F. tularensis shuttle vector with temp-sensitive F. tularensis origin of replication	This study; 36
pBB107	Fusion of pBDJ303 and pMKM219, final Tn5 delivery plasmid for mutagenesis of F. tularensis subsp.	This study
pBB108	E. coli-F. tularensis shuttle vector containing cloned FLP gene downstream of F. tularensis groES promoter	This study
pBB110	E. coli-F. tularensis shuttle vector containing cloned LVS fur gene downstream of F. tularensis groES promoter	This study

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Construction of Tn5 delivery vector.

The DNA fragment carrying the Francisella groES promoter, the lac operator, and lacI^q was first constructed in pCR2.1 by cloning PCR fragments that were amplified with primers (sequences will be supplied upon request) that introduced the desired restriction sites at the end of lacI^q and the groES-lac operator fragment. This DNA fragment was removed from pCR2.1 by digestion with BmgBI and NdeI sites and ligated into the BmgBI and NdeI sites of pRL27 (28). These genetic elements were oriented in the pRL27 vector such that the lac operator sequence was positioned immediately downstream of groESp in the same orientation as the hyperactive transposase, while lacI^q was upstream of groESp on the complementary strand (Fig. 1). This plasmid intermediate was next modified by PCR amplifying and cloning the Francisella omp26 promoter sequence upstream of the aphA3 gene within the transposable element. This modification ensures expression of kanamycin resistance independent of the position of the chromosomal insertion. Finally, the aphA3 gene was amplified from pRL27 by using primers that included the FRT recognition sequence for FLP recombinase, and the DNA fragment was used to replace the existing aphA3 gene. These combined modifications resulted in the creation of pBDJ303, a stable, kanamycin-resistant transposon delivery vector suitable for use in F. tularensis. One feature in pBDJ303 worth mentioning is the presence of unique EcoRI and KpnI sites immediately inside the right side mosaic end of the transposable element. These sites allow the directional cloning of reporter genes that can be used to create transcriptional promoter fusions. For conditional (temperature-sensitive) maintenance in F. tularensis, pBDJ303 was joined to a pFNLTP9-derived E. coli-Francisella shuttle vector (pMKM219) at unique SpeI sites present in each vector to create the final transposon delivery vector, pBB107.

FIG. 1. — Construction of plasmids carrying a modified mini-Tn5 transposon. The plasmid pBDJ303 was derived from pRL27. Plasmid pRL27 carries a hyperactive Tn5 transposase outside the mosaic ends, which define the transposed element. Within the mosaic ends are the R6K plasmid origin and the kanamycin resistance gene *aphA3*. This plasmid was modified by cloning a DNA fragment carrying *lacI*^q, the *Francisella groES* promoter, and the *lac* operator upstream of the hyperactive transposase gene *tnp*. In addition, the *Francisella omp26* promoter was cloned upstream of the *aphA3*, gene which was modified by flanking with FRT sequences. The original features of plasmid pRL27 are shown in black, and the modifications are gray. Plasmid pMKM219 (features are shown in white) was digested with SpeI and ligated to SpeI-cut pBDJ303 to form the *E. coli*-*Francisella* temperature-sensitive Tn5 delivery plasmid pBB107. Plasmid pBB107 confers kanamycin and spectinomycin resistance and is 12.4 kb in size.

Cryotransformation.

Plasmid DNA was introduced into F. tularensis strains by a cryotransformation protocol (42). Briefly, 500 ng of DNA was added to ∼10⁸ CFU F. tularensis LVS that were suspended in Francisella transformation buffer (0.2 M MgSO₄, 0.1 M Tris-acetate, pH 7.5), frozen in liquid nitrogen, and then thawed. The transformed bacteria were grown in either MMH broth or on MMH agar without selection at 30°C for 7 h. Dilutions of the transformed bacteria were plated on MMH agar with 25 μg/ml spectinomycin at 30°C to select for F. tularensis containing the transposon delivery plasmid pBB107.

Transposon selection protocol.

Colonies obtained after ∼3 days of growth at 30°C on MMH agar containing 25 μg/ml spectinomycin were inoculated into 5 ml MMH broth with 25 μg/ml spectinomycin and were grown at 30°C with agitation to an optical density at 600 nm (OD₆₀₀) of ∼0.1. Cultures of LVS were serially diluted and plated on MMH agar with no selection to quantitate the viable cells or on MMH agar with 25 μg/ml kanamycin at 41°C to select for Tn5 insertions into the F. tularensis chromosome with concomitant loss of the transposon delivery plasmid. Selection of F. tularensis Schu S4 transposition events was performed at 40°C, because the strain grew poorly at 41°C; the frequency of transposition was similar to those obtained with LVS at 41°C. F. tularensis Tn5 mutants were arrayed to 96-well cell culture plates in 100 μl MMH broth and were incubated at 37°C until turbid. Freezer stocks were made by adding 100 μl of 2× freezing medium (1.0 M sucrose, 20% glycerol).

Identification of transposon insertion sites.

To identify the sites of Tn5 insertions, genomic DNA was isolated from individual colonies and digested with EcoRI (no reporter), PciI (lux reporter), or NdeI (lacZ reporter) to create a DNA fragment containing the oriR6K origin, the aphA3 gene, and flanking chromosomal sequence. The digested DNA was ligated, transformed into a pir⁺ E. coli strain, and plated onto agar plates with kanamycin to select for transformants that carried the plasmid of interest. Plasmid DNA was isolated and sequenced using a primer with the sequence 5′CATGCAAGCTTCAGGGTTGAG 3′ that anneals to the 3′ end of the aphA3 gene and produces sequence of the flanking chromosomal DNA. Sequence data were used to search the sequenced bacterial chromosomal database, using NCBI BLAST to identify Tn5 insertion sites within the F. tularensis chromosome.

Screening lacZ and luxCDABE mutants for reporter activity.

Tn5 mutants were recovered from freezer stocks and plated on MMH agar at 37°C using a 96-prong replicator (Boekel, Feasterville, PA). After ∼24 h, reporter enzyme activity was detected using a 60-min exposure time with a Fujifilm LAS-1000 luminescence imager (lux reporters) or was visualized by overlaying Whatman no. 1 filter paper presoaked with 20 mg/ml X-Gal (5-bromo-4-chloro-3-indolyl-β-d-galactopyranoside) in dimethylformamide diluted 1:4 in water (lacZ reporters). Quantitation of lacZ activity was done according to the method of Miller (41). Duplicate cultures of tested strains were grown to mid log phase (OD₆₀₀, 0.5 to 0.6) or late log phase (OD₆₀₀ ≈ 0.9 to 1.1), and β-galactosidase assays were performed on triplicate samples of each culture.

Cloning and expression of FLP recombinase in F. tularensis.

The gene encoding FLP recombinase was amplified from pFT-K plasmid template (45) DNA by using upstream (5′AGCAGCGGTACCCAAGGGGTGTTATGCCACAATTTGATATATTATGTAAACACC 3′) and downstream (5′ATCGATCGGTCGACTTATATGCGTCTATTTATGTAGGATG 3′) oligonucleotide primers. At the 5′ end of the upstream primer, a Shine-Dalgarno sequence from aphA3 was included to ensure effective translation of the FLP gene in F. tularensis, and a KpnI site was included to facilitate cloning. At the 5′ end of the downstream primer, a SalI site was included in the sequence of the primer. This PCR-amplified fragment was subcloned into pCR2.1 before being moved to an E. coli-Francisella shuttle vector containing the temperature-sensitive origin of replication. Expression of FLP is driven by the Francisella groES promoter. The shuttle vector containing FLP recombinase, pBB111, was introduced into Tn5 insertion mutants of Francisella strains by cryotransformation, and transformants were selected on MMH agar containing 25 μg/ml spectinomycin. Spectinomycin-resistant colonies were passaged once on MMH agar containing spectinomycin, and isolated colonies were streaked to MMH agar with or without 25 μg/ml kanamycin to screen for FLP-mediated deletion of the aphA3 gene. Southern blot confirmation of the loss of the aphA3 gene was conducted using a digoxigenin-labeled DNA probe generated using the oriR6K region of the transposon as template DNA.

RESULTS

Construction of a Tn5 transposon delivery system in Francisella.

Our initial efforts to create Tn5 kanamycin-resistant transposon mutants of Francisella strains by electroporation or conjugation of the R6K pir-dependent plasmid pRL27 (28) into F. tularensis were unsuccessful. Factors that may have contributed to our inability to initially obtain mutants include poor transformation or conjugation efficiency, low expression of the transposase, and/or low expression of the kanamycin resistance marker. In response to our failed experiments, we created a new Francisella Tn5 transposon delivery system that overcomes these experimental concerns. First, we created plasmid pBDJ303 by optimizing several features of pRL27 for use in Francisella (Fig. 1). The Francisella groES promoter was placed upstream of the hyperactive Tn5 transposase gene to drive expression of the gene in Francisella strains. In the process of making this modification, it became apparent that the plasmid was quite unstable, probably due to the high activity level of the transposase. To alleviate plasmid instability, we placed the expression of the Tn5 transposase under the control of the lac operator and cloned the lacI^q gene onto the plasmid so that, in E. coli, transposase expression is repressed by LacI^q binding to the lac operator sequence between the groES promoter and the transcriptional start site of the transposase. These modifications resolved the plasmid stability issues and also significantly reduced the likelihood of transposition occurring in E. coli during cloning. In addition, two other modifications were made to pBDJ303 before it was fused to a conditionally replicating E. coli-Francisella shuttle plasmid. We placed expression of the kanamycin resistance gene in the transposon under the control of the Francisella omp26 promoter to ensure high-level expression of kanamycin resistance from chromosomal insertions. Also, the kanamycin resistance gene was amplified by PCR with oligonucleotide primers that contain FRT sequences and recloned into the vector. This modification provides the option of deleting the kanamycin resistance gene from a Francisella Tn5 mutant when desired. Finally, we fused pBDJ303 to pMKM219 (a derivative of pFNLTP9) (36), an E. coli-Francisella shuttle vector containing a Francisella conditional origin of replication, to produce the Tn5 transposon delivery vector pBB107 (Fig. 1).

Transformation/transposition and rescue of Tn5 insertions.

F. tularensis LVS was transformed with the E. coli-Francisella Tn5 transposon shuttle plasmid pBB107 by using a cryotransformation protocol (42-44). In preliminary experiments, no kanamycin-resistant colonies were obtained by directly plating transformed F. tularensis LVS and selecting at 41°C, indicating that the combined transformation and transposition frequencies were below a detectable threshold. Accordingly, the creation of transposon mutants was performed in two steps. First, transformants were selected at 30°C on plates containing spectinomycin which yielded ∼100 spectinomycin-resistant transformants (frequency of 1 transformant per 10⁶ recipient bacteria). The efficiency of transformation was not significantly altered by broth or plate outgrowth (data not shown). A single spectinomycin-resistant colony was inoculated into MMH broth containing 25 μg/ml spectinomycin and grown at 30°C to an OD₆₀₀ of ∼0.1. To determine the frequency of transposition from the pBB107 plasmid, cultures were serially diluted and plated on MMH agar with or without 25 μg/ml kanamycin at 41°C (40°C for Schu S4) to simultaneously cure the delivery plasmid and select for Tn5 insertion mutants. Our results indicated that ∼1 in 1,000 organisms containing pBB107 gave rise to a kanamycin-resistant Tn5 mutant (i.e., transposition frequency of 10⁻³) (Table 2). Fifteen kanamycin-resistant LVS mutants were randomly selected for identification of Tn5 insertion sites. Of the 15 Tn5 insertions that were recovered, each mapped to a unique location on the F. tularensis chromosome (data not shown), which is consistent with the findings of others (26, 47). This same frequency of transposition (1 × 10⁻³ to 4 × 10⁻³) has also been observed repeatedly in F. tularensis Schu S4, as part of the process of constructing various F. tularensis Schu S4 transposon libraries. Sequencing the transposon insertion site of individual mutants from these libraries revealed that the insertions are random. The Schu S4 libraries are the focus of work beyond the scope of the experiments reported in this paper. The frequency of transposition in Francisella is virtually identical to the transposition frequency observed in other bacterial species by using the Tn5 hypertransposase (24, 28). A clear advantage of making Tn5 mutants with this method over the EZ::TN transposome system is that the number of mutants that can be created in a single selection is virtually unlimited.

TABLE 2.

Transposition frequency of Tn5 in F. tularensis

Trial	No. of CFU plated^a	No. of Tn5 insertions^b	Frequency of transposition
1	8.45 × 10⁷	1.94 × 10⁵	2.29 × 10⁻³
2	3.51 × 10⁸	4.55 × 10⁵	1.29 × 10⁻³
3	2.85 × 10⁸	2.11 × 10⁵	7.40 × 10⁻⁴

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Total number of CFU determined by plating on MMH agar with no added antibiotics at 41°C.

Total number of Tn5 insertions determined by plating on MMH agar with 25 μg/ml kanamycin at 41°C.

Creation of unmarked Tn5 mutations in F. tularensis by use of FLP recombinase.

To improve the utility of the Tn5 transposon delivery system, the 18-base-pair FRT sequence was added to each end of the aphA3 gene present within the Tn5 transposable element by amplification of the aphA3 gene with FRT-tailed oligonucleotide primers. Following mutagenesis with the transposon containing the aphA3 gene flanked by FRT sites, we sought to remove the antibiotic marker by expressing FLP recombinase. A conditionally replicating (temperature-sensitive) E. coli-F. tularensis shuttle plasmid (pBB111) that expresses FLP recombinase from the F. tularensis groES promoter was cryotransformed into an F. tularensis LVS Tn5 mutant strain, and transformants were selected on solid agar with spectinomycin. Single transformants were purified, and isolated colonies were patched to MMH agar with and without kanamycin. Twenty of twenty F. tularensis colony restreaks grew on plates with no antibiotics but failed to grow on plates with kanamycin, indicating that FLP recombinase was extremely efficient at deleting the kanamycin resistance gene present in the Tn5 mutant. Additionally, Southern blotting was performed on a selected mutant and an ∼1-kb deletion was detected, compared to the parent strain, which corresponds to the loss of the aphA3 gene (Fig. 2).

FIG. 2. — FLP recombinase, expressed from a temperature-sensitive shuttle vector, deletes the kanamycin resistance gene flanked by FRT sequences. (A) Southern blot analysis of EcoRI-digested chromosomal DNA from a Tn5 insertion mutant before (lane 1) and after (lane 2) FLP-mediated deletion of the kanamycin resistance gene. The ∼1.0-kb loss of size in the hybridizing band is the expected deletion size. The probe used for this experiment hybridizes to the oriR6K DNA, contained in the 0.8-kb region of the chromosome as depicted in the figure. (B) Depiction of the Tn5 transposon inserted into the *F. tularensis* LVS chromosome. Features in gray represent the *F. tularensis* chromosome, while features in white represent Tn5 elements. The two black boxes represent the FRT recognition sites for FLP recombinase.

Screening for regulated promoter activity by using luxCDABE and lacZ reporters.

Separate libraries have been constructed with derivatives of pBB107 that create luxCDABE promoter fusions or lacZ promoter fusions with genes on the F. tularensis chromosome. Following selection of Tn5lux or Tn5lacZ mutants, individual mutant isolates were arrayed into the wells of a 96-well master plate and then replica plated onto large MMH or CDM agar plates. The objective of these screens was to identify genes that were differentially regulated by growth on the different media.

The activity of lux reporters in randomly generated F. tularensis strains was analyzed using a Fujifilm LAS-1000 luminescence imager. Our experimental results revealed that detection of lux activity had several technical concerns. First, the activity of bacterial luciferase, encoded by the Vibrio harveyi luxCDABE operon, was extremely low at 37°C compared with the activity of the luciferase complex at the optimal temperature of 25°C (18). This concern was magnified by the relatively low sensitivity of the photoimager, which was unable to detect the luminescence of strains grown at 37°C. These detection issues could be partially overcome by first incubating the F. tularensis lux strains at 25°C for 4 h, followed by a relatively long exposure time (1 h) in the photoimager. However, even with this relatively elaborate detection method, fewer than 1% of F. tularensis strains carrying the Tn5lux transposon insertion produced detectable luciferase activity. Despite these difficulties, we identified three strains carrying luciferase reporters that were upregulated when grown on CDM agar (Fig. 3). Sequence analysis of the transposon insertion sites in these strains revealed that the Tn5 insertions were in genes carrying a 16S rRNA (FTL_R0003), fslD (FTL_1835), and iglC (FTL_0113/1159). Quantitation of the luciferase reporter activity levels in these strains, using a luminometer, revealed significant upregulation of each of these genes when grown in CDM, although the highest level of luciferase activity was near the lower limit of detection of the luminometer.

FIG. 3. — *F. tularensis* LVS strains containing the *luxCDABE* reporter created by Tn5 mutagenesis. Strains containing transcriptional *lux* fusions in the ISFtu1 gene (control) (1), *fslD* (2), FTL_R0003 (3), or *iglC* (4) were streaked to either MMH or CDM agar and were photographed in a light field (top panels) or by use of a photoimager (bottom panels).

F. tularensis mutants, generated using the lacZ reporter, did not grow when plated directly onto differential media containing X-Gal; however, the lacZ expression of individual isolates could be detected by first growing the strains in the absence of X-Gal and then exposing the colony to a filter soaked with X-Gal. Following a short incubation period (15 to 30 min) at 37°C, lacZ⁺ strains were readily detectable by the characteristic blue precipitate observed when X-Gal is cleaved. When β-galactosidase filter assays were performed with randomly selected colonies, ∼30% of the strains expressed β-galactosidase to various degrees. Of ∼1,500 individual mutants screened in this manner, 24 were identified as carrying lacZ reporters in genes that were differentially expressed on the two media (Fig. 4). When quantitative β-galactosidase assays were performed after growth in MMH or CDM broth to quantify gene expression, results for several strains did not match the plate-grown lacZ expression phenotypes. However, when gene expression was compared after growth in MMH versus CDM with only 350 nM FeSO₄ (instead of 7 μM FeSO4), the expression profiles of broth-grown strains were found to mirror those of their plate-grown counterparts (Table 3). This finding led us to conclude that iron starvation was responsible for the observed increase in expression of at least some of the reporter strains, which was detectable on plates due to local depletion of iron around the colonies.

FIG. 4. — *F. tularensis* LVS strains containing the *lacZ* reporter created by Tn5 mutagenesis. Random mutants containing the *lacZ* reporter were arrayed to 96-well plates and were replica plated to MMH (A) or CDM (B) agar. Following ∼24 h of growth, plates were overlaid with filter paper presoaked in X-Gal substrate. After ∼20 min, a characteristic blue precipitate was observed in strains expressing *lacZ*. Several strains contained insertions in genes apparently causing auxotrophy for growth on CDM. Mutants that demonstrated increases in *lacZ* activity when grown on CDM agar were identified (box).

TABLE 3.

F. tularensis LVS genes upregulated by growth on CDM

Gene^a	No. of Miller units on indicated agar^b		Fold increase	Annotation^c
Gene^a	MMH	CDM	Fold increase	Annotation^c
FTL_1834	24	237	9.9	fslC
FTL_1835	13	101	7.8	fslD
FTL_0122	21	99	4.7	Hypothetical in FPI
FTL_1832	51	215	4.2	fslA
FTL_0112	211	665	3.1	iglB
FTL_1576	7	21	3.0	DNA mismatch repair
FTL_1576	15	38	2.6	DNA mismatch repair
FTL_1863	9	20	2.2	Glutamate decarboxylase
FTL_1352	7	14	2.0	tatD
FTL_1492	22	38	1.7	Fructokinase
FTL_0515	38	57	1.5	ABC transporter

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Gene interrupted by the Tn5 transposon.

Numbers of Miller units are averages for at least two experiments.

Annotation according to gene homology with F. tularensis strain Schu S4. FPI, Francisella pathogenicity island.

Our differential growth condition screen identified three of the four genes in the fsl operon to be among the most highly upregulated when grown on CDM agar or in CDM broth. It is likely that strains carrying reporters in these genes were identified on CDM plates, because local iron levels in the iron-limiting CDM agar were depleted, resulting in induction of these iron-regulated genes. This mechanism would be consistent with low reporter activity in liquid media with the same iron concentration because effective iron concentrations would remain higher in liquid, due to mixing. When CDM broth was used with 350 nM iron, we saw induction of genes in the fsl operon as well as other genes identified by the plate screen. Two other strains identified by the screen to be highly regulated were those containing lacZ fusions in iglB (FTL_0112/1158) as well as another hypothetical protein (FTL_0122/1168) located on the Francisella pathogenicity island. Because of these results, we believe that low iron concentration is likely one factor that contributes to the increased pathogenicity reported for Francisella grown on CDM agar.

Expression of Fur in fsl and igl reporter strains grown in low- and high-iron media.

Since low-iron growth conditions resulted in the induction of genes in the fsl operon as well as both iglB and iglC, we examined the role of Fur in the regulation of these genes. The fur gene was PCR amplified from the F. tularensis chromosome, cloned into the E. coli-Francisella shuttle plasmid pBB110, and introduced into the fslC-lacZ and iglB-lacZ reporter strains. Since we wanted to use the iglB-lacZ reporter as an indicator of gene expression of the entire igl gene cluster, we first demonstrated that the four genes are likely transcribed as a single mRNA by using reverse transcription-PCR (RT-PCR) (Fig. 5). F. tularensis strains with lacZ reporters in fslC or iglB carrying pBB110 for expression of Fur, or without the expression vector as a control, were grown in CDM containing 28 μM (high-iron condition) or 350 nM (restricted-iron condition) FeSO₄. Miller assays were conducted on the strains after growth to mid-log and late log growth phases.

FIG. 5. — Genes in the *iglABCD* cluster are operonic. (A) DNA or RNA was isolated from wild-type *F. tularensis* LVS, and RT-PCR was conducted using primer sets that amplified DNA spanning intragenic regions between *iglA-iglB* (300 bp; arrow A), *iglB-iglC* (350 bp; arrow B), or *iglC-iglD* (400 bp; arrow C). For lanes 1, 4, and 7, DNA was used as a template; for lanes 2, 5, and 8, RNA was used as a template without addition of reverse transcriptase; for lanes 3, 6, and 9, RNA was used as a template with the addition of reverse transcriptase. (B) Schematic drawing of the *iglABCD* gene cluster and location of primers used to amplify intragenic regions of DNA.

When the fslB-lacZ reporter strain was grown in CDM containing a high iron concentration, the reporter produced ∼15 Miller units of activity regardless of growth phase. In contrast, the same strain showed a ∼5-fold increase in expression in mid-log phase and a ∼10-fold increase during late log phase when grown under iron-limiting conditions (Fig. 6). This result is consistent with an increase in gene expression as a result of iron depletion in the growth medium. When the fslB-lacZ reporter strain harboring the F. tularensis Fur expression plasmid was examined, it was found to exhibit little lacZ expression (<5 Miller units) when grown in high-iron media and a ∼10-fold reduction in activity compared to that of the parent strain when grown in iron-restricted media (Fig. 6). The observed repression of fslB by overexpression of Fur was not surprising, given the strong consensus Fur box that overlaps the predicted fsl promoter region. Similar experiments with Vibrio vulnificus demonstrated that overexpression of Fur in bacteria grown in iron-replete or -depleted media can have a repressing effect on Fur-regulated genes (33).

FIG. 6. — Overexpression of *F. tularensis* LVS *fur* in chromosomal *lacZ* reporter strains. (A) *F. tularensis* LVS carrying a *lacZ* reporter in *fslC* alone (parent strain; gray bars) or harboring the *fur* expression plasmid pBB110 (white bars) was grown to mid-log phase in CDM broth with either 28 μM (high) or 350 nM (low) FeSO₄. (B) *F. tularensis* LVS carrying a *lacZ* reporter in *iglB* alone (parent strain; gray bars) or harboring the *fur* expression plasmid pBB110 (white bars) was grown to mid-log phase in CDM broth with either 28 μM (high) or 350 nM (low) FeSO₄. Numbers of Miller units are averages for six independent samples.

When similar experiments were conducted using the iglB reporter strain, we also observed iron-dependent induction of the gene. Regardless of growth phase, the parent strain produced ∼200 Miller units of activity when grown in CDM broth containing a high iron concentration. When the strain was grown in iron-limiting media, we observed an ∼1.5-fold induction in mid log phase and a ∼3-fold induction in late log phase. These data are similar to those obtained from the fslC reporter, in that as iron is depleted from the growth medium, induction of iglB is increased. Unexpectedly, when we expressed the F. tularensis Fur protein in this strain we observed a modest increase in activity from the reporter in both iron-rich and iron-limiting media (Fig. 6). These results provide surprising preliminary evidence that Fur acts as a repressor for the fsl operon but not the igl operon. These data allow the possibility that Fur regulates the igl operon by a mechanism different from that of fslABCD or not at all. In fact, our findings suggest that an additional factor or factors may be responsible for repression of the igl operon genes in high iron concentrations. We intend to explore this potentially interesting regulatory mechanism in more detail.

DISCUSSION

Our effort to create an efficient Tn5 transposon delivery system has relied upon the observation and work of other research groups. Tn10 and Tn1721 have both been found to produce unstable insertions when used for mutagenesis (24), and the commercially available Tn5-based in vitro system used to create mutant libraries in different Francisella strains (21, 26, 47, 51, 54) has a reported efficiency of transposition of ∼1 mutant per 10⁸ CFU (26). Our own efforts to use the in vitro transposition system yielded small numbers of mutants per reaction that quickly consumed resources and made it difficult to obtain enough mutants for a saturating library. Combined, this information led us to create a more efficient system for creating transposon mutants of Francisella strains.

Here, we have described the construction of a Tn5 mutagenesis system that has been optimized for use with Francisella tularensis. This approach takes advantage of the hyperactive Tn5 transposase, which increases transposition ∼1,000-fold compared to that obtained with wild-type Tn5 (28). Transcription of the transposase gene and the kanamycin resistance gene has been placed under the control of Francisella promoters to achieve sufficient expression in Francisella strains for activity and detection. In addition, expression of the transposase gene has been placed under the control of the lac operator and LacI repressor to stabilize the transposon delivery plasmid. We have also increased the utility of the system by flanking the kanamycin resistance gene with FRT sequences to allow the creation of unmarked mutations. A key aspect of our system is the use of a temperature-sensitive F. tularensis plasmid origin of replication that was described by Maier et al. (36) as the delivery platform for the Tn5 transposon. The use of this plasmid overcomes the problem of low frequencies of plasmid transformation into F. tularensis strains because a single transformant, recovered at 30°C, can be grown to provide sufficient numbers of bacteria to obtain virtually limitless numbers of transposon mutants. Furthermore, the temperature-sensitive replicon provides a strong selection against maintenance of the plasmid, allowing mutants to be recovered with ease at 41°C. Our experimental results have validated the usefulness of this approach.

In addition to creating the Francisella Tn5 transposon mutagenesis system, we have also made two derivatives of Tn5 that create promoter fusions with luxCDABE or lacZ when inserted into the F. tularensis chromosome. The experimental data indicate that both reporters can be used to detect promoter activity in F. tularensis, although lacZ cleavage of the X-Gal substrate is much more sensitive and able to produce more consistent results than light production from the luxCDABE gene fusions. We have used strains carrying randomly inserted Tn5lacZ reporters to identify genes differentially regulated when F. tularensis is grown on MMH and CDM agar. Results from the qualitative plate screen were corroborated by Miller assays performed on broth-grown bacteria, and it was determined that iron depletion was responsible for upregulation of several genes. Among the genes found to be most highly regulated were genes in the fsl and igl operons.

The Fur protein is associated with regulation of genes that respond to iron concentration in the growth medium. Ferrous iron binds to Fur as a corepressor, causing an allosteric change in the protein that results in Fur binding to conserved nucleotide sequences which often overlap the promoter regions of Fur-regulated genes (13, 16). When iron becomes limiting, Fur adopts a non-DNA binding conformation and repression is relieved at these promoters. Fur-regulated genes are often involved in iron acquisition, but specific virulence factors in several pathogens have also been shown to be directly or indirectly regulated by Fur (34, 46). Given the strong consensus Fur box upstream of the fsl operon, it was not surprising that overexpression of F. tularensis-Fur resulted in superrepression of transcription of the fsl operon, under iron-replete growth. Interestingly, overexpression of Fur during iron-restricted growth also resulted in significant repression of the fslB-lacZ reporter. These data strongly implicate the F. tularensis Fur protein as a repressor of the fsl operon in the presence of iron.

We and others have also found that genes in the igl operon, which are essential for intracellular survival and fundamental to the virulence of this pathogen, are upregulated when F. tularensis is grown in iron-restricted medium (17, 30, 52). Deng et al. (17) proposed that a Fur box that shares 11 of 19 nucleotides with the consensus FUR box resides upstream of iglC. However, it is difficult to reconcile how a functional FUR box upstream of iglC could control iron regulation of other genes in the igl operon. To determine if Fur from F. tularensis plays a role in the regulation of iglABCD similar to that which we have observed for fslABCD, we expressed the Fur protein in an F. tularensis iglB-lacZ reporter strain. Surprisingly, overexpression of Fur in this strain resulted in a slight increase in reporter activity. While it is unclear from these experiments how Fur expression induces an increase in iglB transcription, we believe that these data clearly indicate that Fur is not acting as an iron-dependent repressor of the igl operon.

Two models are proposed to explain these data. First, F. tularensis Fur could positively regulate the expression of iglB in the absence of iron either directly, through productive contacts with RNA polymerase or by bending the DNA to favor transcription, or indirectly, through repression of a transcriptional activator elsewhere on the chromosome. A mechanism of direct activation by Fur would involve the binding of Fur upstream of the iglA promoter in the absence of iron. This model fits our data and would explain the lack of an obvious Fur box upstream of iglA. Since iron binding to Fur causes an allosteric change, the DNA binding site for Fur not bound by iron would be expected to be different from the canonical Fur-Fe consensus binding site (15). The second model holds that overexpression of Fur simply allows Fur to act as an intracellular chelator of iron, which would trigger the activation of a second iron-sensitive system that then regulates the expression of the igl operon. This model also fits our data, although the annotated LVS genome lacks an obvious alternative iron regulator.

Genetic approaches have been, and continue to be, invaluable in identifying and characterizing a wide range of bacterial characteristics, including mechanisms of pathogenesis. In particular, transposon mutagenesis and the creation of chromosomal reporters of transcriptional activity are valuable techniques for identifying bacterial virulence genes and studying their regulation. Tn5-based transposons are well characterized and widely used because of their high frequency of transposition, functionality in many gram-negative bacterial species, low sequence specificity for insertion, and stability when inserted into the host genome (14). The Tn5 transposon delivery system described in this report supplies an additional tool for work aimed at identifying and characterizing virulence factors, and their regulation, in F. tularensis strains. Our data indicate that this transposon mutagenesis system produces virtually limitless numbers of single, random, stable insertions in the chromosomes of F. tularensis strains. Modifications to the transposon provide additional features that are actively being used by our research group to explore and characterize F. tularensis pathogenesis. In addition, we have been able to demonstrate that reporters delivered by this transposon can be used to identify virulence genes (e.g., igl genes) and to study the regulation of various Francisella genes (iron regulation). Future work in our laboratory will focus on utilizing these new genetic tools to identify virulence genes and regulatory pathways that have been, until recently, inaccessible for characterization.

Acknowledgments

We thank Steve Lindemann for helpful insights and careful review of the manuscript. We thank Thomas Zahrt for providing the temperature-sensitive Francisella ori plasmid, contained within pFNLTP9, which was used for the derivative Francisella shuttle plasmids described in this report. We acknowledge the expertise and assistance of the Carver College of Medicine BSL3 Laboratory Core Facility.

This work was supported by U.S. Public Health Service grant PO1 (AI044642).

Footnotes

^▿

Published ahead of print on 14 March 2008.

REFERENCES

1.Abd, H., T. Johansson, I. Golovliov, G. Sandstrom, and M. Forsman. 2003. Survival and growth of Francisella tularensis in Acanthamoeba castellanii. Appl. Environ. Microbiol. 69:600-606. [DOI] [PMC free article] [PubMed] [Google Scholar]
2.Anthony, L. D., R. D. Burke, and F. E. Nano. 1991. Growth of Francisella spp. in rodent macrophages. Infect. Immun. 59:3291-3296. [DOI] [PMC free article] [PubMed] [Google Scholar]
3.Anthony, L. S., M. Z. Gu, S. C. Cowley, W. W. Leung, and F. E. Nano. 1991. Transformation and allelic replacement in Francisella spp. J. Gen. Microbiol. 137:2697-2703. [DOI] [PubMed] [Google Scholar]
4.Baron, G. S., and F. E. Nano. 1998. MglA and MglB are required for the intramacrophage growth of Francisella novicida. Mol. Microbiol. 29:247-259. [DOI] [PubMed] [Google Scholar]
5.Baron, G. S., T. J. Reilly, and F. E. Nano. 1999. The respiratory burst-inhibiting acid phosphatase AcpA is not essential for the intramacrophage growth or virulence of Francisella novicida. FEMS Microbiol. Lett. 176:85-90. [DOI] [PubMed] [Google Scholar]
6.Berg, J. M., K. E. Mdluli, and F. E. Nano. 1992. Molecular cloning of the recA gene and construction of a recA strain of Francisella novicida. Infect. Immun. 60:690-693. [DOI] [PMC free article] [PubMed] [Google Scholar]
7.Bhatnagar, N., E. Getachew, S. Straley, J. Williams, M. Meltzer, and A. Fortier. 1994. Reduced virulence of rifampicin-resistant mutants of Francisella tularensis. J. Infect. Dis. 170:841-847. [DOI] [PubMed] [Google Scholar]
8.Chamberlain, R. E. 1965. Evaluation of live tularemia vaccine prepared in a chemically defined medium. Appl. Microbiol. 13:232-235. [DOI] [PMC free article] [PubMed] [Google Scholar]
9.Cherwonogrodzky, J. W., M. H. Knodel, and M. R. Spence. 1994. Increased encapsulation and virulence of Francisella tularensis live vaccine strain (LVS) by subculturing on synthetic medium. Vaccine 12:773-775. [DOI] [PubMed] [Google Scholar]
10.Clemens, D. L., B. Y. Lee, and M. A. Horwitz. 2004. Virulent and avirulent strains of Francisella tularensis prevent acidification and maturation of their phagosomes and escape into the cytoplasm in human macrophages. Infect. Immun. 72:3204-3217. [DOI] [PMC free article] [PubMed] [Google Scholar]
11.Conlan, J. W., and R. J. North. 1992. Early pathogenesis of infection in the liver with the facultative intracellular bacteria Listeria monocytogenes, Francisella tularensis, and Salmonella typhimurium involves lysis of infected hepatocytes by leukocytes. Infect. Immun. 60:5164-5171. [DOI] [PMC free article] [PubMed] [Google Scholar]
12.Cowley, S. C., C. J. Gray, and F. E. Nano. 2000. Isolation and characterization of Francisella novicida mutants defective in lipopolysaccharide biosynthesis. FEMS Microbiol. Lett. 182:63-67. [DOI] [PubMed] [Google Scholar]
13.Coy, M., and J. B. Neilands. 1991. Structural dynamics and functional domains of the fur protein. Biochemistry 30:8201-8210. [DOI] [PubMed] [Google Scholar]
14.de Bruijn, F. J., and J. R. Lupski. 1984. The use of transposon Tn5 mutagenesis in the rapid generation of correlated physical and genetic maps of DNA segments cloned into multicopy plasmids—a review. Gene 27:131-149. [DOI] [PubMed] [Google Scholar]
15.Delany, I., G. Spohn, R. Rappuoli, and V. Scarlato. 2001. The Fur repressor controls transcription of iron-activated and -repressed genes in Helicobacter pylori. Mol. Microbiol. 42:1297-1309. [DOI] [PubMed] [Google Scholar]
16.de Lorenzo, V., S. Wee, M. Herrero, and J. B. Neilands. 1987. Operator sequences of the aerobactin operon of plasmid ColV-K30 binding the ferric uptake regulation (fur) repressor. J. Bacteriol. 169:2624-2630. [DOI] [PMC free article] [PubMed] [Google Scholar]
17.Deng, K., R. J. Blick, W. Liu, and E. J. Hansen. 2006. Identification of Francisella tularensis genes affected by iron limitation. Infect. Immun. 74:4224-4236. [DOI] [PMC free article] [PubMed] [Google Scholar]
18.Dorn, J. G., R. J. Frye, and R. M. Maier. 2003. Effect of temperature, pH, and initial cell number on luxCDABE and nah gene expression during naphthalene and salicylate catabolism in the bioreporter organism Pseudomonas putida RB1353. Appl. Environ. Microbiol. 69:2209-2216. [DOI] [PMC free article] [PubMed] [Google Scholar]
19.Escolar, L., J. Perez-Martin, and V. de Lorenzo. 1999. Opening the iron box: transcriptional metalloregulation by the Fur protein. J. Bacteriol. 181:6223-6229. [DOI] [PMC free article] [PubMed] [Google Scholar]
20.Fortier, A. H., S. J. Green, T. Polsinelli, T. R. Jones, R. M. Crawford, D. A. Leiby, K. L. Elkins, M. S. Meltzer, and C. A. Nacy. 1994. Life and death of an intracellular pathogen: Francisella tularensis and the macrophage. Immunol. Ser. 60:349-361. [PubMed] [Google Scholar]
21.Gallagher, L. A., E. Ramage, M. A. Jacobs, R. Kaul, M. Brittnacher, and C. Manoil. 2007. A comprehensive transposon mutant library of Francisella novicida, a bioweapon surrogate. Proc. Natl. Acad. Sci. USA 104:1009-1014. [DOI] [PMC free article] [PubMed] [Google Scholar]
22.Gil, H., J. L. Benach, and D. G. Thanassi. 2004. Presence of pili on the surface of Francisella tularensis. Infect. Immun. 72:3042-3047. [DOI] [PMC free article] [PubMed] [Google Scholar]
23.Golovliov, I., V. Baranov, Z. Krocova, H. Kovarova, and A. Sjostedt. 2003. An attenuated strain of the facultative intracellular bacterium Francisella tularensis can escape the phagosome of monocytic cells. Infect. Immun. 71:5940-5950. [DOI] [PMC free article] [PubMed] [Google Scholar]
24.Goryshin, I. Y., J. Jendrisak, L. M. Hoffman, R. Meis, and W. S. Reznikoff. 2000. Insertional transposon mutagenesis by electroporation of released Tn5 transposition complexes. Nat. Biotechnol. 18:97-100. [DOI] [PubMed] [Google Scholar]
25.Gray, C. G., S. C. Cowley, K. K. Cheung, and F. E. Nano. 2002. The identification of five genetic loci of Francisella novicida associated with intracellular growth. FEMS Microbiol. Lett. 215:53-56. [DOI] [PubMed] [Google Scholar]
26.Kawula, T. H., J. D. Hall, J. R. Fuller, and R. R. Craven. 2004. Use of transposon-transposase complexes to create stable insertion mutant strains of Francisella tularensis LVS. Appl. Environ. Microbiol. 70:6901-6904. [DOI] [PMC free article] [PubMed] [Google Scholar]
27.Kuoppa, K., A. Forsberg, and A. Norqvist. 2001. Construction of a reporter plasmid for screening in vivo promoter activity in Francisella tularensis. FEMS Microbiol. Lett. 205:77-81. [DOI] [PubMed] [Google Scholar]
28.Larsen, R. A., M. M. Wilson, A. M. Guss, and W. W. Metcalf. 2002. Genetic analysis of pigment biosynthesis in Xanthobacter autotrophicus Py2 using a new, highly efficient transposon mutagenesis system that is functional in a wide variety of bacteria. Arch. Microbiol. 178:193-201. [DOI] [PubMed] [Google Scholar]
29.Lauriano, C. M., J. R. Barker, F. E. Nano, B. P. Arulanandam, and K. E. Klose. 2003. Allelic exchange in Francisella tularensis using PCR products. FEMS Microbiol. Lett. 229:195-202. [DOI] [PubMed] [Google Scholar]
30.Lenco, J., M. Hubalek, P. Larsson, A. Fucikova, M. Brychta, A. Macela, and J. Stulik. 2007. Proteomics analysis of the Francisella tularensis LVS response to iron restriction: induction of the F. tularensis pathogenicity island proteins IglABC. FEMS Microbiol. Lett. 269:11-21. [DOI] [PubMed] [Google Scholar]
31.Lindemann, S. R., M. K. McLendon, M. A. Apicella, and B. D. Jones. 2007. An in vitro model system used to study adherence and invasion of Francisella tularensis live vaccine strain in nonphagocytic cells. Infect. Immun. 75:3178-3182. [DOI] [PMC free article] [PubMed] [Google Scholar]
32.Lindgren, H., I. Golovliov, V. Baranov, R. K. Ernst, M. Telepnev, and A. Sjostedt. 2004. Factors affecting the escape of Francisella tularensis from the phagolysosome. J. Med. Microbiol. 53:953-958. [DOI] [PubMed] [Google Scholar]
33.Litwin, C. M., and B. L. Byrne. 1998. Cloning and characterization of an outer membrane protein of Vibrio vulnificus required for heme utilization: regulation of expression and determination of the gene sequence. Infect. Immun. 66:3134-3141. [DOI] [PMC free article] [PubMed] [Google Scholar]
34.Litwin, C. M., and S. B. Calderwood. 1993. Role of iron in regulation of virulence genes. Clin. Microbiol. Rev. 6:137-149. [DOI] [PMC free article] [PubMed] [Google Scholar]
35.LoVullo, E. D., L. A. Sherrill, L. L. Perez, and M. S. Pavelka, Jr. 2006. Genetic tools for highly pathogenic Francisella tularensis subsp. tularensis. Microbiology 152:3425-3435. [DOI] [PubMed] [Google Scholar]
36.Maier, T. M., A. Havig, M. Casey, F. E. Nano, D. W. Frank, and T. C. Zahrt. 2004. Construction and characterization of a highly efficient Francisella shuttle plasmid. Appl. Environ Microbiol. 70:7511-7519. [DOI] [PMC free article] [PubMed] [Google Scholar]
37.Maier, T. M., R. Pechous, M. Casey, T. C. Zahrt, and D. W. Frank. 2006. In vivo Himar1-based transposon mutagenesis of Francisella tularensis. Appl. Environ. Microbiol. 72:1878-1885. [DOI] [PMC free article] [PubMed] [Google Scholar]
38.McCaffrey, R. L., and L. A. Allen. 2006. Francisella tularensis LVS evades killing by human neutrophils via inhibition of the respiratory burst and phagosome escape. J. Leukoc. Biol. 80:1224-1230. [DOI] [PMC free article] [PubMed] [Google Scholar]
39.McDonald, M. K., S. C. Cowley, and F. E. Nano. 1997. Temperature-sensitive lesions in the Francisella novicida valA gene cloned into an Escherichia coli msbA lpxK mutant affecting deoxycholate resistance and lipopolysaccharide assembly at the restrictive temperature. J. Bacteriol. 179:7638-7643. [DOI] [PMC free article] [PubMed] [Google Scholar]
40.Mdluli, K. E., L. S. Anthony, G. S. Baron, M. K. McDonald, S. V. Myltseva, and F. E. Nano. 1994. Serum-sensitive mutation of Francisella novicida: association with an ABC transporter gene. Microbiology 140:3309-3318. [DOI] [PubMed] [Google Scholar]
41.Miller, J. H. 1972. Experiments in molecular genetics. Cold Spring Harbor Laboratory, Cold Spring Harbor, NY.
42.Mokrievich, A. N., V. V. Fursov, and V. M. Pavlov. 1994. Plasmid cryotransformation. Problemy Osobo Opasnyh Infektsy Saratov 4:186-190. (In Russian.) [Google Scholar]
43.Norqvist, A., K. Kuoppa, and G. Sandstrom. 1996. Construction of a shuttle vector for use in Francisella tularensis. FEMS Immunol. Med. Microbiol. 13:257-260. [DOI] [PubMed] [Google Scholar]
44.Pavlov, V. M., A. N. Mokrievich, and K. Volkovoy. 1996. Cryptic plasmid pFNL10 from Francisella novicida-like F6168: the base of plasmid vectors for Francisella tularensis. FEMS Immunol. Med. Microbiol. 13:253-256. [DOI] [PubMed] [Google Scholar]
45.Pósfai, G., M. D. Koob, H. A. Kirkpatrick, and F. R. Blattner. 1997. Versatile insertion plasmids for targeted genome manipulations in bacteria: isolation, deletion, and rescue of the pathogenicity island LEE of the Escherichia coli O157:H7 genome. J. Bacteriol. 179:4426-4428. [DOI] [PMC free article] [PubMed] [Google Scholar]
46.Prince, R. W., D. G. Storey, A. I. Vasil, and M. L. Vasil. 1991. Regulation of toxA and regA by the Escherichia coli fur gene and identification of a Fur homologue in Pseudomonas aeruginosa PA103 and PA01. Mol. Microbiol. 5:2823-2831. [DOI] [PubMed] [Google Scholar]
47.Qin, A., and B. J. Mann. 2006. Identification of transposon insertion mutants of Francisella tularensis tularensis strain Schu S4 deficient in intracellular replication in the hepatic cell line HepG2. BMC Microbiol. 6:69-80. [DOI] [PMC free article] [PubMed] [Google Scholar]
48.Rasko, D. A., C. D. Esteban, and V. Sperandio. 2007. Development of novel plasmid vectors and a promoter trap system in Francisella tularensis compatible with the pFLN10 based plasmids. Plasmid 58:159-166. [DOI] [PMC free article] [PubMed] [Google Scholar]
49.Sandström, G., S. Löfgren, and A. Tärnvik. 1988. A capsule-deficient mutant of Francisella tularensis LVS exhibits enhanced sensitivity to killing by serum but diminished sensitivity to killing by polymorphonuclear leukocytes. Infect. Immun. 56:1194-1202. [DOI] [PMC free article] [PubMed] [Google Scholar]
50.Schulert, G. S., and L. A. Allen. 2006. Differential infection of mononuclear phagocytes by Francisella tularensis: role of the macrophage mannose receptor. J. Leukoc. Biol. 80:563-571. [DOI] [PMC free article] [PubMed] [Google Scholar]
51.Su, J., J. Yang, D. Zhao, T. H. Kawula, J. A. Banas, and J. R. Zhang. 2007. Genome-wide identification of Francisella tularensis virulence determinants. Infect. Immun. 75:3089-3101. [DOI] [PMC free article] [PubMed] [Google Scholar]
52.Sullivan, J. T., E. F. Jeffery, J. D. Shannon, and G. Ramakrishnan. 2006. Characterization of the siderophore of Francisella tularensis and role of fslA in siderophore production. J. Bacteriol. 188:3785-3795. [DOI] [PMC free article] [PubMed] [Google Scholar]
53.Tarnvik, A., and L. Berglund. 2003. Tularaemia. Eur. Respir. J. 21:361-373. [DOI] [PubMed] [Google Scholar]
54.Weiss, D. S., A. Brotcke, T. Henry, J. J. Margolis, K. Chan, and D. M. Monack. 2007. In vivo negative selection screen identifies genes required for Francisella virulence. Proc. Natl. Acad. Sci. USA 104:6037-6442. [DOI] [PMC free article] [PubMed] [Google Scholar]
55.Wu, T. H., J. A. Hutt, K. A. Garrison, L. S. Berliba, Y. Zhou, and C. R. Lyons. 2005. Intranasal vaccination induces protective immunity against intranasal infection with virulent Francisella tularensis biovar A. Infect. Immun. 73:2644-2654. [DOI] [PMC free article] [PubMed] [Google Scholar]

[r1] 1.Abd, H., T. Johansson, I. Golovliov, G. Sandstrom, and M. Forsman. 2003. Survival and growth of Francisella tularensis in Acanthamoeba castellanii. Appl. Environ. Microbiol. 69:600-606. [DOI] [PMC free article] [PubMed] [Google Scholar]

[r2] 2.Anthony, L. D., R. D. Burke, and F. E. Nano. 1991. Growth of Francisella spp. in rodent macrophages. Infect. Immun. 59:3291-3296. [DOI] [PMC free article] [PubMed] [Google Scholar]

[r3] 3.Anthony, L. S., M. Z. Gu, S. C. Cowley, W. W. Leung, and F. E. Nano. 1991. Transformation and allelic replacement in Francisella spp. J. Gen. Microbiol. 137:2697-2703. [DOI] [PubMed] [Google Scholar]

[r4] 4.Baron, G. S., and F. E. Nano. 1998. MglA and MglB are required for the intramacrophage growth of Francisella novicida. Mol. Microbiol. 29:247-259. [DOI] [PubMed] [Google Scholar]

[r5] 5.Baron, G. S., T. J. Reilly, and F. E. Nano. 1999. The respiratory burst-inhibiting acid phosphatase AcpA is not essential for the intramacrophage growth or virulence of Francisella novicida. FEMS Microbiol. Lett. 176:85-90. [DOI] [PubMed] [Google Scholar]

[r6] 6.Berg, J. M., K. E. Mdluli, and F. E. Nano. 1992. Molecular cloning of the recA gene and construction of a recA strain of Francisella novicida. Infect. Immun. 60:690-693. [DOI] [PMC free article] [PubMed] [Google Scholar]

[r7] 7.Bhatnagar, N., E. Getachew, S. Straley, J. Williams, M. Meltzer, and A. Fortier. 1994. Reduced virulence of rifampicin-resistant mutants of Francisella tularensis. J. Infect. Dis. 170:841-847. [DOI] [PubMed] [Google Scholar]

[r8] 8.Chamberlain, R. E. 1965. Evaluation of live tularemia vaccine prepared in a chemically defined medium. Appl. Microbiol. 13:232-235. [DOI] [PMC free article] [PubMed] [Google Scholar]

[r9] 9.Cherwonogrodzky, J. W., M. H. Knodel, and M. R. Spence. 1994. Increased encapsulation and virulence of Francisella tularensis live vaccine strain (LVS) by subculturing on synthetic medium. Vaccine 12:773-775. [DOI] [PubMed] [Google Scholar]

[r10] 10.Clemens, D. L., B. Y. Lee, and M. A. Horwitz. 2004. Virulent and avirulent strains of Francisella tularensis prevent acidification and maturation of their phagosomes and escape into the cytoplasm in human macrophages. Infect. Immun. 72:3204-3217. [DOI] [PMC free article] [PubMed] [Google Scholar]

[r11] 11.Conlan, J. W., and R. J. North. 1992. Early pathogenesis of infection in the liver with the facultative intracellular bacteria Listeria monocytogenes, Francisella tularensis, and Salmonella typhimurium involves lysis of infected hepatocytes by leukocytes. Infect. Immun. 60:5164-5171. [DOI] [PMC free article] [PubMed] [Google Scholar]

[r12] 12.Cowley, S. C., C. J. Gray, and F. E. Nano. 2000. Isolation and characterization of Francisella novicida mutants defective in lipopolysaccharide biosynthesis. FEMS Microbiol. Lett. 182:63-67. [DOI] [PubMed] [Google Scholar]

[r13] 13.Coy, M., and J. B. Neilands. 1991. Structural dynamics and functional domains of the fur protein. Biochemistry 30:8201-8210. [DOI] [PubMed] [Google Scholar]

[r14] 14.de Bruijn, F. J., and J. R. Lupski. 1984. The use of transposon Tn5 mutagenesis in the rapid generation of correlated physical and genetic maps of DNA segments cloned into multicopy plasmids—a review. Gene 27:131-149. [DOI] [PubMed] [Google Scholar]

[r15] 15.Delany, I., G. Spohn, R. Rappuoli, and V. Scarlato. 2001. The Fur repressor controls transcription of iron-activated and -repressed genes in Helicobacter pylori. Mol. Microbiol. 42:1297-1309. [DOI] [PubMed] [Google Scholar]

[r16] 16.de Lorenzo, V., S. Wee, M. Herrero, and J. B. Neilands. 1987. Operator sequences of the aerobactin operon of plasmid ColV-K30 binding the ferric uptake regulation (fur) repressor. J. Bacteriol. 169:2624-2630. [DOI] [PMC free article] [PubMed] [Google Scholar]

[r17] 17.Deng, K., R. J. Blick, W. Liu, and E. J. Hansen. 2006. Identification of Francisella tularensis genes affected by iron limitation. Infect. Immun. 74:4224-4236. [DOI] [PMC free article] [PubMed] [Google Scholar]

[r18] 18.Dorn, J. G., R. J. Frye, and R. M. Maier. 2003. Effect of temperature, pH, and initial cell number on luxCDABE and nah gene expression during naphthalene and salicylate catabolism in the bioreporter organism Pseudomonas putida RB1353. Appl. Environ. Microbiol. 69:2209-2216. [DOI] [PMC free article] [PubMed] [Google Scholar]

[r19] 19.Escolar, L., J. Perez-Martin, and V. de Lorenzo. 1999. Opening the iron box: transcriptional metalloregulation by the Fur protein. J. Bacteriol. 181:6223-6229. [DOI] [PMC free article] [PubMed] [Google Scholar]

[r20] 20.Fortier, A. H., S. J. Green, T. Polsinelli, T. R. Jones, R. M. Crawford, D. A. Leiby, K. L. Elkins, M. S. Meltzer, and C. A. Nacy. 1994. Life and death of an intracellular pathogen: Francisella tularensis and the macrophage. Immunol. Ser. 60:349-361. [PubMed] [Google Scholar]

[r21] 21.Gallagher, L. A., E. Ramage, M. A. Jacobs, R. Kaul, M. Brittnacher, and C. Manoil. 2007. A comprehensive transposon mutant library of Francisella novicida, a bioweapon surrogate. Proc. Natl. Acad. Sci. USA 104:1009-1014. [DOI] [PMC free article] [PubMed] [Google Scholar]

[r22] 22.Gil, H., J. L. Benach, and D. G. Thanassi. 2004. Presence of pili on the surface of Francisella tularensis. Infect. Immun. 72:3042-3047. [DOI] [PMC free article] [PubMed] [Google Scholar]

[r23] 23.Golovliov, I., V. Baranov, Z. Krocova, H. Kovarova, and A. Sjostedt. 2003. An attenuated strain of the facultative intracellular bacterium Francisella tularensis can escape the phagosome of monocytic cells. Infect. Immun. 71:5940-5950. [DOI] [PMC free article] [PubMed] [Google Scholar]

[r24] 24.Goryshin, I. Y., J. Jendrisak, L. M. Hoffman, R. Meis, and W. S. Reznikoff. 2000. Insertional transposon mutagenesis by electroporation of released Tn5 transposition complexes. Nat. Biotechnol. 18:97-100. [DOI] [PubMed] [Google Scholar]

[r25] 25.Gray, C. G., S. C. Cowley, K. K. Cheung, and F. E. Nano. 2002. The identification of five genetic loci of Francisella novicida associated with intracellular growth. FEMS Microbiol. Lett. 215:53-56. [DOI] [PubMed] [Google Scholar]

[r26] 26.Kawula, T. H., J. D. Hall, J. R. Fuller, and R. R. Craven. 2004. Use of transposon-transposase complexes to create stable insertion mutant strains of Francisella tularensis LVS. Appl. Environ. Microbiol. 70:6901-6904. [DOI] [PMC free article] [PubMed] [Google Scholar]

[r27] 27.Kuoppa, K., A. Forsberg, and A. Norqvist. 2001. Construction of a reporter plasmid for screening in vivo promoter activity in Francisella tularensis. FEMS Microbiol. Lett. 205:77-81. [DOI] [PubMed] [Google Scholar]

[r28] 28.Larsen, R. A., M. M. Wilson, A. M. Guss, and W. W. Metcalf. 2002. Genetic analysis of pigment biosynthesis in Xanthobacter autotrophicus Py2 using a new, highly efficient transposon mutagenesis system that is functional in a wide variety of bacteria. Arch. Microbiol. 178:193-201. [DOI] [PubMed] [Google Scholar]

[r29] 29.Lauriano, C. M., J. R. Barker, F. E. Nano, B. P. Arulanandam, and K. E. Klose. 2003. Allelic exchange in Francisella tularensis using PCR products. FEMS Microbiol. Lett. 229:195-202. [DOI] [PubMed] [Google Scholar]

[r30] 30.Lenco, J., M. Hubalek, P. Larsson, A. Fucikova, M. Brychta, A. Macela, and J. Stulik. 2007. Proteomics analysis of the Francisella tularensis LVS response to iron restriction: induction of the F. tularensis pathogenicity island proteins IglABC. FEMS Microbiol. Lett. 269:11-21. [DOI] [PubMed] [Google Scholar]

[r31] 31.Lindemann, S. R., M. K. McLendon, M. A. Apicella, and B. D. Jones. 2007. An in vitro model system used to study adherence and invasion of Francisella tularensis live vaccine strain in nonphagocytic cells. Infect. Immun. 75:3178-3182. [DOI] [PMC free article] [PubMed] [Google Scholar]

[r32] 32.Lindgren, H., I. Golovliov, V. Baranov, R. K. Ernst, M. Telepnev, and A. Sjostedt. 2004. Factors affecting the escape of Francisella tularensis from the phagolysosome. J. Med. Microbiol. 53:953-958. [DOI] [PubMed] [Google Scholar]

[r33] 33.Litwin, C. M., and B. L. Byrne. 1998. Cloning and characterization of an outer membrane protein of Vibrio vulnificus required for heme utilization: regulation of expression and determination of the gene sequence. Infect. Immun. 66:3134-3141. [DOI] [PMC free article] [PubMed] [Google Scholar]

[r34] 34.Litwin, C. M., and S. B. Calderwood. 1993. Role of iron in regulation of virulence genes. Clin. Microbiol. Rev. 6:137-149. [DOI] [PMC free article] [PubMed] [Google Scholar]

[r35] 35.LoVullo, E. D., L. A. Sherrill, L. L. Perez, and M. S. Pavelka, Jr. 2006. Genetic tools for highly pathogenic Francisella tularensis subsp. tularensis. Microbiology 152:3425-3435. [DOI] [PubMed] [Google Scholar]

[r36] 36.Maier, T. M., A. Havig, M. Casey, F. E. Nano, D. W. Frank, and T. C. Zahrt. 2004. Construction and characterization of a highly efficient Francisella shuttle plasmid. Appl. Environ Microbiol. 70:7511-7519. [DOI] [PMC free article] [PubMed] [Google Scholar]

[r37] 37.Maier, T. M., R. Pechous, M. Casey, T. C. Zahrt, and D. W. Frank. 2006. In vivo Himar1-based transposon mutagenesis of Francisella tularensis. Appl. Environ. Microbiol. 72:1878-1885. [DOI] [PMC free article] [PubMed] [Google Scholar]

[r38] 38.McCaffrey, R. L., and L. A. Allen. 2006. Francisella tularensis LVS evades killing by human neutrophils via inhibition of the respiratory burst and phagosome escape. J. Leukoc. Biol. 80:1224-1230. [DOI] [PMC free article] [PubMed] [Google Scholar]

[r39] 39.McDonald, M. K., S. C. Cowley, and F. E. Nano. 1997. Temperature-sensitive lesions in the Francisella novicida valA gene cloned into an Escherichia coli msbA lpxK mutant affecting deoxycholate resistance and lipopolysaccharide assembly at the restrictive temperature. J. Bacteriol. 179:7638-7643. [DOI] [PMC free article] [PubMed] [Google Scholar]

[r40] 40.Mdluli, K. E., L. S. Anthony, G. S. Baron, M. K. McDonald, S. V. Myltseva, and F. E. Nano. 1994. Serum-sensitive mutation of Francisella novicida: association with an ABC transporter gene. Microbiology 140:3309-3318. [DOI] [PubMed] [Google Scholar]

[r41] 41.Miller, J. H. 1972. Experiments in molecular genetics. Cold Spring Harbor Laboratory, Cold Spring Harbor, NY.

[r42] 42.Mokrievich, A. N., V. V. Fursov, and V. M. Pavlov. 1994. Plasmid cryotransformation. Problemy Osobo Opasnyh Infektsy Saratov 4:186-190. (In Russian.) [Google Scholar]

[r43] 43.Norqvist, A., K. Kuoppa, and G. Sandstrom. 1996. Construction of a shuttle vector for use in Francisella tularensis. FEMS Immunol. Med. Microbiol. 13:257-260. [DOI] [PubMed] [Google Scholar]

[r44] 44.Pavlov, V. M., A. N. Mokrievich, and K. Volkovoy. 1996. Cryptic plasmid pFNL10 from Francisella novicida-like F6168: the base of plasmid vectors for Francisella tularensis. FEMS Immunol. Med. Microbiol. 13:253-256. [DOI] [PubMed] [Google Scholar]

[r45] 45.Pósfai, G., M. D. Koob, H. A. Kirkpatrick, and F. R. Blattner. 1997. Versatile insertion plasmids for targeted genome manipulations in bacteria: isolation, deletion, and rescue of the pathogenicity island LEE of the Escherichia coli O157:H7 genome. J. Bacteriol. 179:4426-4428. [DOI] [PMC free article] [PubMed] [Google Scholar]

[r46] 46.Prince, R. W., D. G. Storey, A. I. Vasil, and M. L. Vasil. 1991. Regulation of toxA and regA by the Escherichia coli fur gene and identification of a Fur homologue in Pseudomonas aeruginosa PA103 and PA01. Mol. Microbiol. 5:2823-2831. [DOI] [PubMed] [Google Scholar]

[r47] 47.Qin, A., and B. J. Mann. 2006. Identification of transposon insertion mutants of Francisella tularensis tularensis strain Schu S4 deficient in intracellular replication in the hepatic cell line HepG2. BMC Microbiol. 6:69-80. [DOI] [PMC free article] [PubMed] [Google Scholar]

[r48] 48.Rasko, D. A., C. D. Esteban, and V. Sperandio. 2007. Development of novel plasmid vectors and a promoter trap system in Francisella tularensis compatible with the pFLN10 based plasmids. Plasmid 58:159-166. [DOI] [PMC free article] [PubMed] [Google Scholar]

[r49] 49.Sandström, G., S. Löfgren, and A. Tärnvik. 1988. A capsule-deficient mutant of Francisella tularensis LVS exhibits enhanced sensitivity to killing by serum but diminished sensitivity to killing by polymorphonuclear leukocytes. Infect. Immun. 56:1194-1202. [DOI] [PMC free article] [PubMed] [Google Scholar]

[r50] 50.Schulert, G. S., and L. A. Allen. 2006. Differential infection of mononuclear phagocytes by Francisella tularensis: role of the macrophage mannose receptor. J. Leukoc. Biol. 80:563-571. [DOI] [PMC free article] [PubMed] [Google Scholar]

[r51] 51.Su, J., J. Yang, D. Zhao, T. H. Kawula, J. A. Banas, and J. R. Zhang. 2007. Genome-wide identification of Francisella tularensis virulence determinants. Infect. Immun. 75:3089-3101. [DOI] [PMC free article] [PubMed] [Google Scholar]

[r52] 52.Sullivan, J. T., E. F. Jeffery, J. D. Shannon, and G. Ramakrishnan. 2006. Characterization of the siderophore of Francisella tularensis and role of fslA in siderophore production. J. Bacteriol. 188:3785-3795. [DOI] [PMC free article] [PubMed] [Google Scholar]

[r53] 53.Tarnvik, A., and L. Berglund. 2003. Tularaemia. Eur. Respir. J. 21:361-373. [DOI] [PubMed] [Google Scholar]

[r54] 54.Weiss, D. S., A. Brotcke, T. Henry, J. J. Margolis, K. Chan, and D. M. Monack. 2007. In vivo negative selection screen identifies genes required for Francisella virulence. Proc. Natl. Acad. Sci. USA 104:6037-6442. [DOI] [PMC free article] [PubMed] [Google Scholar]

[r55] 55.Wu, T. H., J. A. Hutt, K. A. Garrison, L. S. Berliba, Y. Zhou, and C. R. Lyons. 2005. Intranasal vaccination induces protective immunity against intranasal infection with virulent Francisella tularensis biovar A. Infect. Immun. 73:2644-2654. [DOI] [PMC free article] [PubMed] [Google Scholar]

PERMALINK

Identification of Differentially Regulated Francisella tularensis Genes by Use of a Newly Developed Tn5-Based Transposon Delivery System^▿

Blake W Buchan

Molly K McLendon

Bradley D Jones

Abstract

MATERIALS AND METHODS