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Applied and Environmental Microbiology, January 2008, p. 336-341, Vol. 74, No. 1
0099-2240/08/$08.00+0 doi:10.1128/AEM.01522-07
Copyright © 2008, American Society for Microbiology. All Rights Reserved.
Genomic Markers for Differentiation of Francisella tularensis subsp. tularensis A.I and A.II Strains
,
Claudia R. Molins-Schneekloth,1,2
John T. Belisle,2 and
Jeannine M. Petersen1*
Division of Vector-Borne Infectious Diseases, Centers for Disease Control and Prevention, Fort Collins, Colorado,1
Department of Microbiology, Immunology, and Pathology, Colorado State University, Fort Collins, Colorado2
Received 28 June 2007/
Accepted 30 October 2007
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ABSTRACT
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Tularemia is caused by two subspecies of Francisella tularensis, F. tularensis subsp. tularensis (type A) and F. tularensis subsp. holarctica (type B). F. tularensis subsp. tularensis is further subdivided into two genetically distinct populations (A.I and A.II) that differ with respect to geographical location, anatomical source of recovered isolates, and disease outcome. Using two human clinical isolates, suppression subtractive hybridization was performed to identify 13 genomic regions of difference between A.I and A.II strains. Two PCR assays, one to identify A.I and A.II as well as to discriminate between F. tularensis subsp. holarctica and F. novicida and another specific for A.I, were developed. This is the first report to identify and characterize conserved genomic differences between A.I and A.II.
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INTRODUCTION
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Francisella tularensis is a gram-negative bacterial pathogen that causes the zoonotic disease tularemia (5). Humans acquire tularemia through arthropod bites (ticks and deerflies), handling of infected animal tissues, ingestion of contaminated water or food, and inhalation of infective aerosols. The clinical presentation of tularemia varies depending upon the route of infection. Ulceroglandular tularemia is the most common form of disease, while pneumonic tularemia is the most severe (5). F. tularensis is classified as a category A agent due to its past development as a biological weapon and also its extreme infectivity (4).
Two subspecies of F. tularensis, F. tularensis subsp. tularensis (type A) and F. tularensis subsp. holarctica (type B), are of clinical relevance. These two subspecies differ with respect to geographic location, virulence, and biochemical properties (13). Tularemia infections caused by F. tularensis subsp. tularensis are documented only in North America, whereas F. tularensis subsp. holarctica causes infections in North America, Europe, and Asia.
Molecular subtyping has further divided F. tularensis subsp. tularensis into two genetically distinct populations (6, 14). Pulsed-field gel electrophoresis (PFGE) defined these two clusters as type A-east and type A-west, while multiple-locus variable-number tandem repeat analysis (MLVA) identified these populations as A.I and A.II. Comparative data suggest that type A-east is equivalent to A.I and type A-west is equivalent to A.II; thus, the A.I and A.II terminology is used in this report. A molecular and epidemiologic analysis of A.I and A.II isolates from human cases of tularemia in the United States showed a geographical separation, with A.I isolates found primarily in the central and eastern portions of the United States and A.II isolates localized to the western part of the country (14). Significant differences in disease outcome between infections caused by A.I and A.II were also noted, with case fatality rates of 14% and 0%, respectively. A.I isolates were associated more frequently with blood and lung infections than were A.II isolates, while A.II isolates were primarily isolated from lymph nodes. Thus, infections caused by A.I and A.II were found to differ with respect to geographical location, anatomical source of recovered isolates, and disease outcome (6, 14).
The goal of this study was to identify and characterize genomic regions of difference (RDs) between a panel of A.I and A.II strains for the development of a molecular diagnostic. Suppression subtractive hybridization (SSH) was used in this study to identify RDs between the genomes of A.I and A.II (1, 20). The A.I strain used in this study, MA00-2987, is a clinical isolate obtained from the blood of a human case of pneumonic tularemia in Martha's Vineyard in 2000 (7). WY96-3418 was picked as the A.II strain and is a clinical isolate recovered from a finger wound of a human ulceroglandular case of tularemia in Wyoming in 1996. Thirteen conserved regions of difference were identified and used for the development of PCR assays to identify A.I and A.II. The conserved RDs also identify genes that may be linked to biological differences between A.I and A.II.
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Thirteen conserved regions of difference between A.I and A.II strains.
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Genomic SSH was performed using the PCR-Select bacterial genome subtraction kit (BD Biosciences Clontech, Palo Alto, CA). F. tularensis strains were grown from frozen stocks on cysteine heart agar with 9% chocolatized sheep blood at 35°C, and genomic DNA was isolated using the Puregene DNA isolation kit (Gentra Systems, Minneapolis, MN). To generate DNA fragment sizes of 
1 kb, the restriction enzyme AluI, a four-base, blunt-end restriction enzyme, was used. The subtraction of WY96-3418 (A.II) DNA from MA00-2987 (A.I) DNA worked efficiently at an annealing temperature of 58°C, whereas the reverse subtraction (MA00-2987 from WY96-3418) was most effective when 60°C was the annealing temperature (data not shown). Subtraction efficiency was verified by PCR amplification of three control genes (23S, rpoH, and dnaK) present in both the tester and driver genomic DNA (see Table S1 in the supplemental material). A marked reduction of PCR product for two of the three control genes was considered efficient subtraction.
To identify tester-specific fragments, DNA was amplified by both primary and nested PCR. Nested PCR products were cloned into pGEM-T Easy vector (Promega, Madison, WI) and transformed into competent DH5
cells (Invitrogen, Carlsbad, CA). Of the resulting β-galactosidase– Ampr Escherichia coli clones, 100 clones were randomly selected from both subtractions (A.I-A.II and A.II-A.I) and plasmid DNA recovered using the Qiagen mini prep kit (Qiagen, Inc., Valencia, CA). To verify the presence of tester-specific DNA inserts, plasmids were digested with BstZI; those with an insert (137 clones) were sequenced, and internal primers were designed (see Table S1 in the supplemental material).
PCR amplification with the internal primers, MA00-2987 DNA, and WY963418 DNA identified 24 RDs. In this study, an RD is defined as a genomic region present in one population and absent from the other population. Of these 24 RDs, 12 (RD-2, RD-3, RD-7, RD-8, RD-11, RD-14, RD-16, RD-17, RDR-14, TN-1, TN-2, and TN-4) were present in MA00-2987 (A.I) and another 12 (RD-1, RD-4, RD-5, RD-6, RD-9, RD-10, RD-12, RD-13, RD-15, RDR-5, TN-3, and TN-5) were present in WY96-3418 (A.II). The primers used to define RD-1, RD-2, RD-7, RD-8, RD-9, RD-11, RD-12, RD-14, RD-16, RD-17, RDR-14, TN-2, and TN-4 yielded PCR products from both A.I and A.II genomic DNA, but with different amplicon sizes. Thus, in some cases, one or both of the internal primers fell within the region of genomic DNA absent from one population, and in other cases, the internal primers flanked the region of DNA absent from one population. In cases where there was a difference in product size, both PCR products were cloned and sequenced.
To develop molecular diagnostics for A.I and A.II as well as identify genes that may be linked to biological differences, it was critical for us to focus analyses on RDs present among diverse strains because both PFGE and MLVA have shown diversity among A.I and A.II strains (6, 14). Therefore, to verify that the 24 RDs represented conserved differences between A.I and A.II strains, nine A.I and nine A.II clinical strains from diverse geographical locations (Table 1) were PCR amplified using the internal primer sets (see Table S1 in the supplemental material). All strains on the panel were identified as either A.I or A.II by PFGE molecular subtyping. Of the 24 RDs, 19 (RD-1 to RD-13, RDR-5, and TN-1 to TN-5) were present within all A.I or A.II strains tested. The five RDs (RD-14, RD-15, RD-16, RD-17, and RDR-14) not present within all A.I or A.II strains were dropped from further analyses.
To map the verified RDs in the F. tularensis genome, BLAST analyses (blastn) (www.ncbi.nlm.nih.gov/), along with sequence alignments using LALIGN (www.ch.embnet.org/software/LALIGN_form.html) and ClustalW (www.ebi.ac.uk/clustalw/#), were performed. Due to differences in annotation, RDs were mapped within multiple whole-genome sequences of F. tularensis subspecies, including F. tularensis subsp. tularensis SCHU S4 (A.I) (AJ749949) and WY96-3418 (A.II) (NC_009257) and F. tularensis subsp. holarctica LVS (AM233362) and OSU18 (NC_008369) (type B), and F. novicida U112 (NC_008601) (Table 2) (11, 12). At the time this study was initiated, no whole-genome sequence was available for an A.II strain; however, the genome sequence for WY96-3418 became available later during the course of this study and was used to map the RDs.
Of the 19 verified RDs, one was identified more than once (RD-5/RDR-5), 5 were associated with insertion sequence (IS) elements (TN-1 through TN-5), 12 were gene/pseudogene associated (RD-1, RD-2, RD-3, RD-4, RD-5, RD-6, RD-8, RD-9, RD-10, RD-11, RD-12, and RD-13), and 1 mapped to an intergenic region (RD-7) (Table 3). IS-associated RDs were not pursued in this study, since these elements are more likely to be associated with additional polymorphisms (8). Further characterization of RDs between A.I and A.II strains focused only on the 13 that were gene/pseudogene associated or intergenic (RD-1 through RD-13). Five such RDs (RD-2, RD-3, RD-7, RD-8, and RD-11) were present in A.I strains and eight (RD-1, RD-4, RD-5, RD-6, RD-9, RD-10, RD-12, and RD-13) were present in A.II strains. The sizes of these 13 RDs ranged from 15 bp to 1,181 bp (Table 2).
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TABLE 3. Predicted genes within RDs in F. tularensis subsp. tularensis strains SCHU S4 (A.I) and WY96-3418 (A.II)
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Differences in hypothetical proteins accounted for much of the genomic variability associated with the defined RDs (Table 3). Multiple hypothetical proteins are absent from or have limited homology in either A.I (FTW_0888 and FTW_0889) or A.II (FTT1733) strains or are predicted to be pseudogenes in one population relative to the other population (FTT1195c, FTT1429c, FTW_1356, FTW_1384, and FTW_2084). For hypothetical proteins encoded within the RDs, bioinformatic analyses were performed to predict protein localization (PSORT), signal sequences (SigP), transmembrane helix domains (TMHMM), and protein families (Pfam). RDs within intergenic regions were analyzed for putative promoter sequences (BPROM). See Table S2 in the supplemental material for a summary of this analysis.
Of the five RDs present in A.I strains and absent from A.II strains, four (RD-2, RD-3, RD-8, and RD-11) were within genes encoding hypothetical proteins (Tables 2 and 3). RD-2 (148 bp) is localized within the sequence encoding a hypothetical protein (FTT1733) predicted to be functional in A.I with no homologue in A.II. RD-3 is an 1,181-bp region that lies within a gene encoding a conserved hypothetical lipoprotein (FTT1194c), the adjacent intergenic region, and a downstream pseudogene encoding a conserved hypothetical protein (FTT1195c). RD-8 (264 bp) falls within a region encoding a hypothetical protein (FTT0096) predicted to localize to the cytoplasm and to contain an ATP_GRASP 3 domain (ATP-binding domain). RD-11 is 15 bp and its absence in A.II results in an in-frame deletion of five amino acids within a hypothetical protein (FTT1078c) predicted to be functional in A.I and a pseudogene in A.II (FTW_2084). By BLAST analyses, FTT1078c showed similarity to several regions (E values ranging from 4e-36 to 7e-33) of the FrpA/C-related proteins of Neisseria meningitidis (16, 17). The significance of this protein in A.I is unclear. Frp-like proteins are linked to virulence in a broad range of pathogenic gram-negative bacteria, and the predicted protein in A.I may be of interest for further study (21). The remaining RD (RD-7) (45 bp) mapped to an intergenic region.
Of the eight RDs present in A.II strains and absent in A.I strains, three (RD-1, RD-4, and RD-5) are associated with hypothetical proteins (Tables 2 and 3). Of interest, RD-1 falls within a gene for a hypothetical protein that is predicted to contain an N-terminal signal sequence and one transmembrane helix domain and is also predicted to localize to the outer membrane. The absence of RD-1 in A.I causes an in-frame deletion of 380 bp, deleting the N-terminal signal sequence and shifting the start codon for this hypothetical protein in A.I. The resulting prediction is that this protein is secreted in A.II (FTW_1826), but not in A.I (FTT0267). RD-4 and RD-5 lie within regions spanning both gene-carrying and intergenic regions. RD-4 is 674 bp and encompasses an intergenic region and two adjacent downstream regions encoding hypothetical proteins (FTW_0888 and FTW_0889). RD-5 spans a region of 310 bp that encompasses the coding region of a hypothetical protein (FTW_0430), an intergenic region, and a downstream pseudogene for a methyltransferase.
The remaining five RDs present in A.II strains and absent from A.I strains fall within genes with putative function, or pseudogenes (RD-6, RD-9, RD-10, RD-12, and RD-13) (Tables 2 and 3). RD-6 (138 bp) maps to the C4-dicarboxylate anaerobic carrier pseudogene (FTW_1709). RD-10 (33 bp) falls within a region encoding a DNA processing protein, DprA. RD-13 (246 bp) is within a putative drug transporter pseudogene (FTW_0298). RD-9 (16 bp) lies within a magnesium chelatase pseudogene (FTW_0121). Of interest, RD-12 (179 bp) falls within a chitinase family 18 protein pseudogene (homologue of chiA) in A.II. The absence of RD-12 in A.I results in an in-frame deletion of 44 amino acids, with the shortened product (760 amino acids) annotated as functional. A.II contains a premature stop codon (198 amino acid protein) not present in A.I and is annotated as a pseudogene. In a previous study, Twine et al. demonstrated an increase in chitinase (ChiA homologue) expression of more than 20-fold when grown in vivo compared to the in vitro level for type A (18). Additional studies demonstrate a role for chitinases in parasite transmission (during in vivo infections by Legionella pneumophila) and in environmental survival (3, 9). A comparison of chitinase-associated proteins of F. tularensis subsp. tularensis (A.I and A.II), F. tularensis subsp. holarctica, and F. novicida revealed differences in the ChiA and ChiB homologues and a predicted chitinase-binding protein. Thus, these proteins may have some role in host vectors, environmental survival, or human infections. While further studies are required to determine the true biological relevance of any of the gene products encoded by the RDs, the genomic differences between A.I and A.II identified in this study provide a basis for the development of hypotheses that can be experimentally tested.
None of the 13 conserved RDs or predicted genes associated with the RDs identified were restricted to either the A.I or the A.II populations; they were also found in the genomes of either F. tularensis subsp. holarctica or F. novicida (Tables 2, 3, and 4; see Table S2 in the supplemental material). This result is consistent with the study by Svensson et al. that compared genomic differences between F. tularensis subsp. tularensis, F. tularensis subsp. holarctica, F. tularensis subsp. mediasiatica, and F. novicida (15). The differences identified in this previous study were considerably larger (300 bp to 
11,500 bp) than those identified here, and none of the RDs described here were previously identified. RDs could result from genetic deletions or insertions. Direct repeat-mediated homologous recombination was likely the mechanism leading to the RDs identified in this study, since direct repeats such as those described for other deletions within the F. tularensis genomes (2, 15) were identified flanking the RDs (data not shown).
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PCR assays for identification of A.I and A.II.
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Based on the verified non-IS-associated RDs, diagnostic PCR assays were developed. Since all identified RDs were present within F. tularensis subsp. holarctica and/or F. novicida, two regions of difference (RD-3 and RD-6) were targeted for a multiplex diagnostic PCR that could identify A.I and A.II as well as discriminate them from F. tularensis subsp. holarctica and F. novicida. The RD-3 primer set amplifies a product in A.I (570 bp) and not A.II, whereas the RD-6 primer set results in an amplicon for A.II (396 bp) and not for A.I. The multiplex assay consisted of PuReTaq Ready-To-Go PCR beads, 2.5 mM MgCl2, and 1 µM each of primers RD-3F (5'-ATATATGGTGATGCTAA AGAC-3'), RD-3R (5'-TCTATTGAGATCTAAATCTGC-3'), RD-6F (5'-TGACCTGAGCCAGAGGTTAC-3'), and RD-6R (5'-ACTTGCCAGCCTAATAATTAC). PCR conditions were 95°C for 5 min, followed by 30 cycles of 95°C for 45 s, 53°C for 45 s, and 72°C for 45 s.
The multiplex PCR assay was used to amplify DNA from a panel of 24 strains representing F. tularensis subsp. tularensis (A.I and A.II), F. tularensis subsp. holarctica, and F. novicida (Table 1). Although both primer sets amplify the corresponding regions in F. tularensis subsp. holarctica and F. novicida, when used in a multiplex PCR, three different banding patterns are observed: one pattern for A.I strains, one pattern for A.II strains, and one pattern for F. tularensis subsp. holarctica and F. novicida strains (Fig. 1A). Thus, the multiplex assay can distinguish the A.I and A.II populations of F. tularensis subsp. tularensis and further differentiate these two populations from F. tularensis subsp. holarctica and F. novicida.
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FIG. 1. PCR assays for identification of F. tularensis subsp. tularensis (A.I and A.II). The nine A.I (lanes 2 to 10), nine A.II (Lanes 11 to 19), three F. tularensis subsp. holarctica (type B) (lanes 20 to 22), and three F. novicida (lanes 23 to 25) strains are indicated. Lane 1, 1-kb DNA ladder; lane 2, NE03-1457; lane 3, MA00-2987; lane 4, SCHU S4; lane 5, OK01-2528; lane 6, SD00-3147; lane 7, NY04-2526; lane 8, VA00-1000; lane 9, MO02-1911; lane 10, AR99-3448; lane 11, WY01-3847; lane 12, WY96-3418; lane 13, GA02-5453; lane 14, ID04-2687; lane 15, WY03-1228; lane 16, CA02-0099; lane 17, AZ01-4999; lane 18, UT02-1927; lane 19, WY01-3911; lane 20, KY99-3387; lane 21, LVS; lane 22, OR96-0246; lane 23, GA99-3448; lane 24, GA99-3449; lane 25, GA99-3550; lane 26, negative control; lane 27, 1-kb DNA ladder. (A) PCR assay differentiating A.I from A.II using two RD primer sets: RD-3 and RD-6. (B) A.I-specific PCR assay using primer set RD-10J-F and RD-10J-R.
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For an A.I-specific PCR, a junction region (representing the deleted RD in A.I) was targeted. For several A.I junction regions (RD-4, RD-9, and RD-10), nucleotide differences occurred and degenerate primers specific for A.I could be designed. Thus, an A.I-specific PCR was developed by targeting the junction region resulting from the deletion of RD-10 in A.I. The forward primer RD-10J-F was designed with 3-bp changes in order to ensure specificity for A.I. The A.I-specific PCR assay consisted of PuReTaq Ready-To-Go PCR beads and 1 µM each of primers RD-10J-F (5'-ATCATTGATTGGAAATCAAAAATTG-3') and RD-10J-R (5'-ATCCTGATATTCCTCCTAACACAA-3'). PCR conditions were 95°C for 5 min, followed by 30 cycles of 95°C for 45 s, 55°C for 45 s, and 72°C for 45 s.
When DNA from the panel of 24 strains (Table 1) was tested with the A.I PCR, all A.I strains were amplified and no cross-reactivity was observed with A.II, F. tularensis subsp. holarctica, or F. novicida strains (Fig. 1B), thus demonstrating the use of this primer set as a diagnostic specific for A.I strains. An A.II-specific PCR can also be developed using the same principle to target A.II junction regions where RDs have been deleted.
These two PCR assays provide new diagnostic tools for the identification of A.I and A.II. Currently, the only methods available for distinguishing these populations are PFGE and MLVA, both of which are time consuming and labor intensive. The conserved RDs identified here are also amenable for future development of a real-time PCR assay. Such an assay could be used in combination with existing F. tularensis real-time PCR assays (10, 19) so that in a single run, identification can be provided at the subspecies level (F. tularensis subsp. tularensis or F. tularensis subsp. holarctica), with further delineation of F. tularensis subsp. tularensis into A.I or A.II.
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ACKNOWLEDGMENTS
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We thank Aimee Janusz for her assistance with DNA isolation, Kiersten Kugeler for contributing DNA, and Darragh Heaslip, Torsten Eckstein, and Herbert Schweizer for their helpful suggestions.
The work was additionally supported by National Institutes of Health Cooperative Agreement U54 AI-065357.
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FOOTNOTES
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* Corresponding author. Mailing address: Centers for Disease Control and Prevention, Division of Vector-Borne Infectious Diseases, Bacterial Diseases Branch, 3150 Rampart Road, Fort Collins, CO 80521. Phone: (970) 266-3524. Fax: (970) 494-6631. E-mail: JPetersen{at}cdc.gov 
Published ahead of print on 16 November 2007.
Supplemental material for this article may be found at http://aem.asm.org/.
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Applied and Environmental Microbiology, January 2008, p. 336-341, Vol. 74, No. 1
0099-2240/08/$08.00+0 doi:10.1128/AEM.01522-07
Copyright © 2008, American Society for Microbiology. All Rights Reserved.
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ABSTRACT
INTRODUCTION
